Coupling of Mechanical and Electromagnetic Fields Stimulation for Bone Tissue Engineering

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Alyaa I. Aldebs Wright State University

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This Thesis is brought to you for free and open access by the Theses and Dissertations at CORE Scholar. It has been accepted for inclusion in Browse all Theses and Dissertations by an authorized administrator of CORE Scholar. For more information, please contact [email protected]. COUPLING OF MECHANICAL AND ELECTROMAGNETIC FIELDS STIMULATION FOR BONE TISSUE ENGINEERING

A thesis submitted in partial fulfillment of the

requirements for the degree of Master of Science

in Biomedical Engineering

ALYAA I. ALDEBS

B.Sc. Biomedical Engineering, University of Al-Nahrain, 2012

2018

Wright State University

WRIGHT STATE UNIVERSITY GRADUATE SCHOOL April 27, 2018 I HEREBY RECOMMEND THAT THE THESIS PREPARED UNDER MY SUPERVISION BY Alyaa I. Aldebs Entitled Coupling of Mechanical and Electromagnetic Fields Stimulation for Bone Tissue Engineering BE ACCEPTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF Master of Science in Biomedical Engineering.

______Jaime E. Ramirez-Vick, Ph.D. Thesis Director

______Jaime E. Ramirez-Vick, Ph.D. Department Chair

Committee on Final Examination

______Jaime E. Ramirez-Vick, Ph.D.

______Ulas Sunar, Ph.D.

______Nasim Nosoudi, Ph.D.

______Barry Milligan, Ph.D. Interim Dean of the Graduate School

ABSTRACT

ALDEBS, ALYAA I. M.S.B.M.E. Department of Biomedical, Industrial and Human Factors Engineering, Wright State University, 2018. Coupling of Mechanical and Electromagnetic Fields Stimulation for Bone Tissue Engineering.

Alternative bone regeneration strategies that do not rely on harvested tissue or exogenous

growth factors and cells are badly needed. However, creating living tissue constructs that

are structurally, functionally and mechanically comparable to the natural bone has been a

challenge so far. A major hurdle has been recreating the bone tissue microenvironment

using the appropriate combination of cells, scaffold and stimulation to direct

differentiation. This project presents a bone regeneration formulation that involves the use

of human adipose-derived mesenchymal stems cells (hASCs) and a 3D scaffold based on

a self-assembled peptide hydrogel doped with superparamagnetic nanoparticles (NPs).

Osteogenic differentiation of hASCs is achieved through the direct stimulation by

extremely-low frequency pulsed electromagnetic fields (pEMFs) and the indirect

mechanical stimulation, through NP vibration induced by the field. This 3D construct was

cultured for up to 21 days and its osteogenic capacity was assessed. Cellular morphology,

proliferation, viability, as well alkaline phosphatase activity, calcium deposition were

monitored during this time.

The results show that the pEMFs have no negative effect on cell viability and

induce early differentiation of hASCs to an osteoblastic phenotype when compared to a

iii

cell without biophysical stimulation. This effect results from the synergy between the pEMF and NP that acts as remote stimulation of the mechanotransduction pathways which activate biochemical signals between cells to go under differentiation or proliferation. The use of this approach offers a safe and effective treatment option for the treatment of non-union bone fractures. In addition, this formulation can be directly injected into the wound site, making it minimally invasive as well.

Table of Contents

ABSTRACT ...... iii

Table of Contents ...... v

List of Figures ...... vii

Acknowledgment ...... ix

CHAPTER I ...... 1

Introduction ...... 1

1.1 Project overview ...... 1 1.2 Objectives ...... 5 CHAPTER II ...... 7

Literature Review ...... 7

2.1 Bone structure ...... 7 2.2 Bone anatomy ...... 7 2.3 Bone composition ...... 11 2.3.1 Bone Extracellular Matrix ...... 11 2.3.2 bone cells ...... 14 2.3.3 growth factors ...... 17 2.4 Bone diseases ...... 19 2.5 Bone replacement and tissue engineering ...... 20 2.6 Cellular sources for bone tissue engineering...... 22 2.7 Three-dimensional scaffold for bone tissue engineering..……..………………25 2.7.1 Self-assembled peptide scaffold ...... 26 2.8 Biophysical stimulation ...... 30 2.8.1 Effect of mechanical stimulation ...... 30 2.8.2 Effect of electromagnetic fields ...... 34

2.9 Effect of magnetic scaffold under EMF stimulation for bone regeneration ...... 38 CHAPTER III ...... 46

Materials and Methods ...... 46

3.1 Cell culture ...... 46 3.2 Three-dimensional cells encapsulation and gel formation ...... 46 3.3 Cells viability assay ...... 47 3.4 Cells differentiation assay ...... 48 3.5 Measurement of mineralization ...... 49 3.6 Cells morphology ...... 49 3.7 Alkaline phosphatase staining ...... 50 3.8 Fourier transform infrared spectroscopy (FTIR) analysis ...... 51 3.9 Electromagnetic fields exposure system ...... 51 3.10 Statistical analysis ...... 53 CHAPTER IV...... 54

Results &Discussion ...... 54

4.1 Results & Discussion ...... 54 4.1.1 Cell viability ...... 54 4.1.2 Differentiation to osteoblasts ...... 59 4.1.3 Mineralization ...... 63 4.1.4 Cell morphology ...... 65 4.1.5 Alkaline phosphatase staining ...... 68 4.1.6 FTIR analysis ...... 70 4.2 Conclusion ...... 71 References ...... 74

Appendix I ...... 89

List of Figures

Figure Page

1. Three-dimensional structure of bone that shows the cortical and trabecular components…………………………………………………………………...... 10

2. Topographic relationships of different bone cells that initiate from different origins…………………………………………………………………….……...15

3. Amino acids sequence of self- assembled peptide (RADA-16) ………………...28

4. The mechanosensors that resulted from changes in ions channels, proteins, and cytoskeleton to activate the intercellular signals……………………………...... 32

5. Schematic illustration of using Helmholtz coil to generate pEMF……………...35

6. Electromagnetic exposure system: a) closed mu-metal box and b) Helmholtz coils with cells.…………………………………………………………………..……53

7. LDH assay. The proliferation of hASCs seeded within self-assembled peptide hydrogel with and without NPs under extremely low-frequency pEMF of 1 mT was quantified, after 7, 14, 21 d. *p < 0.05; **p < 0.001; ***p < 0.0001, indicate statistically significant differences between basal media group (control) and the other groups. The mean values are calculated from the average results of three samples, the results are represented as mean ± SD..…………………….……….56

8. Alkaline phosphatase activity. The differentiation of hASCs cells was assessed at 7, 14, 21 d. ALP values were normalized with the number of cells of each sample. *p < 0.05; **p < 0.001; ***p < 0.0001, indicate statistically significant differences between basal media group (control) and the other groups (one-way ANOVA followed by Dunnett test). The mean values are calculated from the average results of three samples, the results are represented as mean ± SD………………………………………………………………….………….…62

9. Mineralization assay. Calcium concentrations were quantified after 7,14, 21 days. Error bar represents the s.d. p < 0.05, indicate statistically significant differences between MSCs media gel+cells group (control) and the other groups (one-way ANOVA followed by Dunnett test). The mean values are calculated from the average results of three samples, the results are represented as mean ± SD…………………………………………………………………………..……65

vii

10. Phalloidin-labeled actin filaments stain (red) and DAPI stain (blue) for hASCs within hydrogel at 3 and 14 days. A) basal media, B) basal media + NPs, C) osteogenic media, D) osteogenic media + NPs, E) basal media + pEMF, F) basal media + NPs + pEMF, G) osteogenic media + pEMF, H) osteogenic media + NPs + pEMF. Scale bars at day 3 and 14 are 100 µm (10X) and 50 µm (20X), respectively. Biochemical stimulation – osteogenic induction media…………………………………………..……………………………...… 67

11. Alkaline phosphatase stain for hASCs cells seeded within hydrogel at day 14, scale bar = 200 µm (10X). A) basal media, B) basal media + NPs, C) osteogenic media, D) osteogenic media + NPs, E) basal media + pEMF, F) basal media + NPs + pEMF, G) osteogenic media + pEMF, H) osteogenic media + NPs + pEMF. Biochemical stimulation – osteogenic induction media . ……..………..69

12. FTIR analysis of self-assembled peptides with and without iron oxide nanoparticles……………………………………………………………………..71

viii

Acknowledgment

In the beginning, I would like to express my sincere thanks to my thesis advisor, Dr.

Ramirez-Vick for his support during my research time and for advising me to find the answer to all my concerns through his valuable comments. I would also like to thank Dr.

Nosoudi for helping and training me during the research. I have gratefully appreciated all the assistance she provided through giving me her time and her efforts. Also, I would like to thank my thesis committee member Dr. Sunar. Also, I would like to thank my colleague lab students.

In addition, I would like to thank my husband, Ali for making me follow my dream, for encouraging me, and for providing me the suitable atmosphere to focus on my study without him I could not be here. Also, I want to thank my mother and my father for their love and support.

CHAPTER I

Introduction

1.1 Project overview

Bone fractures represent a substantial incidence and cost burden among musculoskeletal injuries in the United States, with 15.3 million cases reported each year, of which 5-10% involve complications, such as delayed or non-unions (Nauth, et al., 2011).

Conventional surgical treatments for bone damaged that resulted from trauma, tumor, bone fracture and abnormalities when the defect is above the critical size of 1-mm by either using autografts, allografts or metallic or ceramic implants (Meng et al., 2010). Each of these bone grafts has some limitations, such as donor site morbidity, the risk of infections, pain, shortage in graft quantity and pathogen transmission (Brydone, et al., 2010; Meng, et al., 2010). Thus, recently, extensive efforts have been done to enhance bone tissue engineering to mimic the bone tissue microenvironment focusing on scaffolds as substitutes autologous or allogeneic bone grafts (Xu, et al., 2014)

Bone tissue engineering and regenerative medicine required three complementary and essential components to mimic the native tissue through culturing diverse cell types in three-dimensional (3D) microenvironments to generate a fully functional tissue. These components are osteoprogenitor cells that responsible for creating bone tissue, biomimetic scaffold to provide 3D structure that is osteoconductive, and stimulation which can be biochemical or biophysical in nature, to create an osteoinductive environment for

promoting cell differentiation (Horii, et al., 2007; Stevens, 2008; Neves, et al., 2017). To generate this microenvironment requires that the scaffold is manufactured in a way that mimics the nanostructure of the natural extracellular matrix (ECM; Li, et al., 2013). The

ECM is considered a vital component of any tissue, which works not only supporting cells but also by creating an appropriate microenvironment that influences cell-function (Horii, et al., 2007). Therefore, by using a biomimetic scaffold that has similar properties as natural tissue ECM seeded with suitable cell types under the presence of appropriate stimulation can guide tissue growth and regeneration.

However, creating living tissue constructs that are structurally, functionally and mechanically comparable to the natural bone has been a challenge (Polo-Corrales, et al.,

2014). For instance, recreating the bone tissue microenvironment using the appropriate combination of cells, scaffold and stimulation to direct differentiation (Polo-Corrales, et al., 2018). As an alternative, the use of extremely-low-frequency pulsed electromagnetic fields (pEMFs) as an adjuvant therapy for the treatment of bone disorders to reduce complications have been widely used in orthopedics since it was approved by the FDA almost four decades ago (Lohmann, et al., 2000). This allowed various clinical trials and production of commercial devices to promote bone fracture healing (Heckman, et al.,

1981). Since then, different effects of pEMF stimulation on in vitro differentiation and proliferation of osteogenic cell lines have been published in the literature (Daish, et al.,

2018). Researchers have indicated that the forced vibration of all the free ions on the surface of a cellular plasma membrane, changes in voltage, and conductivities are possible mechanisms in which EMF could regulate cell process (Panagopoulos, et al., 2002;

Markov, 2007; Ross, et al., 2015; Rubio Ayala, et al., 2018). Since then, man investigations

have focused on the use of this therapy to accelerate the cell proliferation and osteogenic differentiation of progenitor cells, such as mesenchymal stem cells (MSCs).

A strategy to improve fracture healing is to look more closely at the bone tissue microenvironment, in which mechanical stimulation is an important part and is essential for bone health and homeostasis (Voog and Jones, 2010). The importance of this type of stimulation is because it mediates an adaptive remodeling response in the bone at the cellular level, through a process known as Wolff’s law (Lanyon, 1974; Lanyon and

Baggott, 1976; Woo, et al., 1981). At the molecular level, the process by which cells transduce these force-induced signals into biochemical responses is known as mechanotransduction and it leads to variations in gene expression, cell function, morphology, and extracellular matrix (ECM) remodeling. Since the mechanisms involved during bone remodeling are the same as those found during fracture healing (McKibbin,

1978), it is thus reasonable to consider mechanical stimulation as a therapeutic strategy to induce healing when a bone fracture is present. A recent alternative that has shown great interest is the use of scaffolds that can provide mechanical stimulation through vibrations induced on superparamagnetic scaffolds by external EMF (Zeng, et al., 2012; Meng, et al.,

2013; Xu and Gu, 2014; Grant, et al., 2015). In addition, the presence of superparamagnetic nanoparticles (NPs) could also affect the mechanical properties of the scaffolds, by enhancing the compressive strength and elastic modulus. The improved mechanical properties in 3D scaffolds, particularly elastic modulus values, have been proven to promote MSC osteogenic differentiation (Jegal, et al. 2011). Early work on this type of scaffold was focused on polymer-based nanocomposites loaded with magnetic particles for drug delivery applications (Edelman, et al., 1984; Liu, et al., 2006; Zhao, et al., 2011).

Recently, the use of these superparamagnetic scaffolds and EMFs, to promote bone formation has been proposed (Kanczler, et al., 2010; Sapir-Lekhovitser, et al., 2016). The interesting aspect of this approach is that it couples two separate stimuli on the cells, mechanical, through the vibratory movement of the scaffold where the cells are attached to, and magnetic, where the forced-vibration of all the free ions on the surface of the plasma membrane, changes in voltage, and conductivities affecting cell function (Panagopoulos, et al., 2002; Deng, et al., 2007; Garner, et al., 2007).

In this research, we used a biomimetic scaffold made of the self-assembling peptide

RADA16, which consist of regular repeats of alternating ionic hydrophilic and hydrophobic amino acids and self-assemble to form stable β-sheet structures in water.

When exposed to physiological solutions they spontaneous assemble into a stable, macroscopic membranous matrix, composed of ordered filaments (~10 nm) forming pores

5–200 nm in size (Wang, et al., 2008; Zhang, et al., 1993; Zhang, et al., 1995). the self- assembled peptide is the spontaneous arrangements of amino acids that followed the state of thermodynamic equilibrium to get a well-defined structure with firm organizations through various noncovalent interactions to produce hierarchical structures. In spite the fact that these interactions are weak, but when they together it formed a firm and stable structure (Zhang, 2002). This type consists of an alternating amino acid (Arginine, Alanine, and Aspartic acid) that spontaneously assembling to produce microscopic and macroscopic matrices of interwoven nanofibers to form higher-order hydrogels scaffolds in the presence of monovalent cations (Zhang et al.,1995; Hauser et al., 2010; Zhao et al., 2006). Although these interactions are weak, together they form microscopic and macroscopic matrices of interwoven nanofibers to form higher-order hydrogels that are firm and stable (Zhang,

2002; Hauser et al., 2010; Zhao et al., 2006). Due to their nanometer scale, their network and biomechanical properties are comparable to the natural ECM, making them good candidates to generate biomimetic cell niches (Semino, 2008). In addition, their stiffness can be modulated with concentration. Studies have found that collagen I and RADA16 hydrogels have similar mechanical properties with mean G’ modulus values in the 10–1000

Pa range (Semino, 2008; Cunha, et al., 2011). The generation of superparamagnetic scaffolds involves either a dip-coating method of the scaffolds in aqueous ferrofluids comprising biocompatible and nontoxic superparamagnetic nanoparticles (NPs), allowing these to infiltrate to the scaffold, or using in situ method by mixing NPs during scaffold synthesis, reducing the number of processing steps and time. It is expected that the superparamagnetic NPs will chelate to the hydrogel matrix (Kantipuly, et al., 1990). In this manner, the resulting magnetic scaffolds are capable to take up seeded cells (Bañobre-

López, et al., 2011; Samba Sivudu, et al., 2009). Our focus on this study is on the use adipose-derived mesenchymal stem cells (hASCs) encapsulated with self-assembled peptides containing superparamagnetic iron oxide NPs and study their capacity to differentiate after being biochemically-stimulated with osteogenic induction media and/or extremely low-frequency pEMFs.

1.2 Objectives

The main goal of this research is to create biologically compatible superparamagnetic scaffolds suitable for hASC growth and differentiation by mimicking the bone microenvironment to provide a level of osteoconductive and osteoinductive properties equivalent native bone tissue. These goals will be achieved through:

• Synthesis of biomimetic hydrogels using based on self-assembled peptides

combined with superparamagnetic iron oxide nanoparticles to create a 3D porous

scaffold and evaluate its specific functional groups through FTIR analysis.

• Encapsulate the hASCs within the scaffold and evaluate their viability and

morphology within the scaffold through LDH analysis and F-actin staining.

• Evaluate the hASCs differentiation under pEMF stimulation in vitro by monitoring

the ALP activity and calcium deposition.

CHAPTER II

Literature Review

2.1 Bone structure

Bone is a vascularized, metabolically dynamic connective tissue that forms, along with cartilage, the skeletal system of the body. The main function of bone is to provide mechanical support and physiological protection. The mechanical support is necessary to provide protection to vital organs and offer a site for muscle attachment to control movement. While the physiological protection results from the providing of reservoirs for growth factors and minerals such as calcium, potassium, carbonate, magnesium, strontium, chloride and fluoride, phosphate to maintain blood hemostasis and a place for hematopoiesis to occur. Since bone is a heterogenic structure, these functions vary depending on its location. For example, the bone that is not exposed to loadings, such as in the skull and scapula have a dissimilar structure to long bones, which are exposed to applied external forces such as tension, compression, and bending (Rogel, et al., 2008).

2.2 Bone anatomy

The adult human skeleton, which consists of 213 bones, can be classified according to their location, shape, or size. In terms of shape, bones can be divided into flat bone

(i.e., skull, scapula, sternum, ribs, or mainly the facial skeleton bones) and tubular bone.

The latter can be further classified as long tubular bone (i.e., femur, tibia, humerus, clavicles, etc.), or short tubular bone as in the hand and feet (i.e., metacarpals, metatarsals, and phalanges) (Buck, et al., 2012). The anatomical structure of the tubular bone consists mainly of three different sections: diaphysis, metaphysis, and epiphysis. The diaphysis is located in the middle of the tubular bone and has the shape of the cylindrical hollow shaft, composed mainly of dense and hard cortical bone, which is filled by the hollow shaft, or medullary cavity, that contains bone marrow and fat. The metaphysis located between diaphysis and epiphysis, is cone-shaped and composed of a growth plate that calcifies with age. The epiphysis located at the end of the tubular bone, with rounded and wide sections, consists primarily a trabecular spongy meshwork filled with red marrow and covered from the exterior with a thin shell of the cortical bone (Clarke, et al., 2008; Buck, et al., 2012).

In contrast, flat bones have a varied structure consisting of either purely cortical bone or cortical bone with a thin central trabecular region.

At the macrostructure level, cortical and trabecular are the primary bone structures that formed from the same matrix composition but vary in their porosity, 3D structure, and their metabolic activities. The cortical (also known as compact bone) makes up 80% of human body skeletal system, with a porosity of 5-10%, that has a compressive strength with the ability to resist torsion and bending. Although the trabecular (also known as cancellous bone) makes up only 20% of the body, with a 50-90% porosity of interconnected pores filled with bone marrow, which gives the ability of deformation and force absorption.

Nevertheless, both bone structure shares the same metabolic activities since the trabecular

bone has a higher metabolic activity (Buck, et al., 2012). In contrast, the cortical bone is covered by two surfaces, the outer surface called periosteal, which plays a role in the appositional growth and fracture healing. promoting bone enlargement through the process of periosteal apposition. The inner surface, called endosteal surface, has a high remodeling activity, allowing the bone to be under resorption (Clarke, et al., 2008; Datta et al., 2008).

At the microstructure level, the main structure in cortical and trabecular bone consists of osteons (Figure 1). The osteons (also known as Haversian systems) in cortical bones are oriented longitudinally along long bones and consist of mineralized collagen fibers oriented in concentric layers. The osteons are approximately 200 µm in diameter and

10-20 µm in length, with concentric lamellae 3–7 µm in diameter. It is also composed of a central Haversian canal which is a hollow tube, 80 µm in diameter, located in the center of the lamellae, allowing blood vessels to pass through it to distribute the necessary nutrients to bone cells (Athanasiou, et al., 2000; Rho, et al., 1998; Sharir, et al., 2008). Mineralized collagen fibers that are in a disorganized pattern, known as woven bone, yield a mechanically weak structure. When the lamellae are oriented tangentially to the external surface of the bone without making osteons and along with woven bone, they form a larger plywood-type stacking of dense layer known as a lamellar bone (Rho, et al., 1998).

On the other hand, the trabecular osteons (also known as packets) have a semilunar shape formed from interconnected plates and rods with a thickness of 50-400 µm and with

300-1500 µm of space between them (Athanasiou, et al., 2000; Clarke, et al., 2008). The trabeculae distribution reflects the bone strength, shown as an alignment along lines of

stress. Consequently, trabecular bone is capable of enduring compressive stresses, an as a result, it shows a predominant presence in the vertebrae (Datta, et al., 2008).

Figure 1. Three-dimensional structure of bone that shows the cortical and trabecular components (Buck, et al., 2012).

2.3 Bone composition

2.3.1 Bone Extracellular Matrix

Bone is a compound that contains different materials that together constitute the bone matrix, which consists of about 90% of the total bone. The major components found in this 3D ECM are about 70% of inorganic components or minerals, 22% of organic components, mainly collagen, and about 8% lipids and water (Augat, et al., 2006; Buck, et al., 2012). The distribution of materials and their quality has a strong effect on bone strength; the composition of the mineral, which is mostly in the form of hydroxyapatite, gives the bone the ability to resist the compressive load and its quality affects bone stiffness. The collagen fibrils give the bone the ability to resist the tensile load and its quality affects bone toughness (Augat, et al., 2006; Athanasiou, et al., 2000). The mechanical properties of bone result from the interactions between these components, since the deposited mineral crystal and collagen are oriented in the longitudinal axis, giving bone its high stiffness and strength along its axis (Athanasiou, et al., 2000). The organic and inorganic components of the matrix interact together to produce a diverse set of mechanical and biological characteristics than those produce separately. Hence, the composition of the matrix plays a vital role in the calcified bone tissue whereas the elasticity is associated to the mineral phase of the bone and plasticity to the organic components of the matrix

(Landis,1995).

The organic component of mature bone that is considered its main building block.

It consists mainly of the fibrous protein type I collagen that represents 90% of ECM proteins, which is organized in a fibrillar structure and sometimes onto which, the mineral crystalizes through gaps between the collagen fibrils. The collagen fibril structure consists

of triple helical chains twisted around each other, consisting of two α1 polypeptide chains and one α2 chain. It structures composed of a sequence of three different amino acids glycine, proline and hydroxyproline that define the helix-forming repeated motif, which produces a rod shape molecule 1.4 nm wide and 300 nm long (Weiner, et al., 1998; Rogel, et al., 2008). During collagen synthesis, preprocollagen (i.e., a single chain of the polypeptide) becomes procollagen (i.e., a triple helix chain of the polypeptide) after hydroxylation and glycosylation, where hydrogen and disulfide bonds are formed. In addition, another bond is generated when tropocollagen (i.e., procollagen with a cut in the extra terminal amino acids) during which covalent bonds are formed between the terminal of the triple polypeptide helices (Last, et al., 1984; Zioupos, et al., 1999). The collagen fibril results from the self-assembling of tropocollagen by establishing a link to neighboring tropocollagen molecules through trivalent bonds known as hydroxypyridinium bonds (Zioupos, et al., 1999). The attachment between the NH2- terminus of one triple helical molecule and the COOH-terminus of the following molecules forms a gap or a hole. This arrangement forms an area with densely packed molecules and an area of less densely packed molecules in the 2D model arrangement, while the 3D arrangement shows transverse channels or grooves (Weiner, et al., 1998). The mineralized collagen fibrils diameter is approximately 80-100 nm, and the mineral crystals between these fibrils are around 50 nm by 25 nm and 2–3 nm in thickness (Sharir, et al., 2008). In contrast, noncollagenous proteins, which represent 10-15% of total bone protein, such as serum albumin and α2-HS-glycoprotein, have an impact on ECM mineralization through binding to the hydroxyapatite crystal through their acidic properties. These noncollagenous proteins can be divided into proteoglycans and glycosylated proteins, which affect the bone

mineral deposition and bone cell activity. Of these, osteonectin is considered the most widespread noncollagenous protein, representing around 2% of the entire protein in bone.

The main impact of this protein on the bone is through its effect on osteoblast proliferation and matrix mineralization (Clarke, et al., 2008).

The second major component of the bone matrix is bone mineral known as dahllite, which has a hexagonal crystallographic symmetry, although the microscopic results show it does not exhibit this type of symmetry instead it shows as a thin plate-shaped crystal

(Weiner et al., 1998). The main mineral component of bone that gives it its hardness resulted from the presence of mineralized calcium phosphate in form of hydroxyapatite,

[Ca₁₀(PO4)6(OH)2], with small amounts of carbonate, magnesium, and acid phosphate

(Datta, et al., 2008; Weiner, et al., 1998). The mineral crystal has a dimension of tens of nanometers in length and several nanometers in width, with an elongated morphology and a preferred crystallographic and morphological alignment with the main directions of stress

(Rogel, et al., 2008). The orientation of minerals has an influence on mechanical properties of bones, where the longitudinal alignment of hydroxyapatite crystals promotes transverse isotropy (Sasaki, et al.,1989). In addition, the presence of alkaline phosphatase and other non-collagenous proteins, such as osteocalcin, osteopontin, and bone sialoprotein help in

ECM maturation. Consequently, these calcium- and phosphate-binding proteins control the quantity and size of hydroxyapatite crystals, thus influencing the mineral deposition

(Clarke, et al., 2008).

The third main component of bone is water, which has a major influence on bone mechanical properties as it was found that the dried bone has different mechanical

properties than it is wet. The water exists mainly in the holes between the triple-helical polypeptide and within the fibril and between collagen fibers (Weiner, et al., 1998).

2.3.2 bone cells

Cells in bone tissue makeup 10% of its volume and are composed of different types that are at different stages of maturity (Figure 2). These cells migrate from different places, they can have a hematopoietic origin such as the osteoclast or be derived from local mesenchymal cells known as osteoprogenitor cells such as osteoblasts, lining cells, and osteocytes (Buck, et al., 2012).

Osteoblasts are mononuclear cells with diameter range between 15 to 30 µm that has a large spherical nucleus with high composition of rough endoplasmic reticulum and

Golgi apparatus, which promotes ECM synthesis, mainly collagen type I. The presence of actin, myosin, and other cytoskeletal proteins allows these cells to modify their shape and helps them during migration and binding to the ECM (Jayakumar, et al., 2010). Osteoblasts play a vital role in skeletal development by becoming highly specialized synthetic cells, that when mature, support and regulate hematopoiesis. Besides, they have the capacity to respond to many mechanical and systemic stimulations, which induce mineral deposition

(Taichman, 2005).

Osteoblasts cover the surfaces of bone and are closely aligned with each other.

Since they are very active, their shape is oval or polyhedral. In addition, during new matrix secretion, osteoblasts will consist large amounts of endoplasmic reticulum, Golgi apparatus

(for synthesizing proteins), and mitochondria. Thus, osteoblasts not only form and secrete the organic matrix of bone but also have a role in its mineralization through the control of

electrolyte fluxes between the extracellular fluid and the osseous fluid (Buckwalter, et al.,

1995). After becoming activated, these cells can either remain inactive osteoblasts, osteocytes, or return to osteoprogenitor cells (Buck, et al., 2012). Osteoblasts main protein secretion is collagen type I, which self-assembles into fibrils. Osteocytes are osteoblasts that have surrounded themselves with an organic matrix to live within vacuoles called lacunae. They develop cytoplasmic projections that move across bone to come in contact with adjacent osteocytes, allowing direct communication (Marks, et al., 1988).

Figure 2. Topographic relationships of different bone cells that initiate from different origins (Marks, et al., 1988).

Osteoclasts are the only bone cells responsible for the resorption of bone mineral and originate from CD34+ hematopoietic progenitor cells. They are located on the surface of bone close to the vascular channels and can be distinguished from the other bone cells

by its large and multinucleated cells shape (Taichman, 2005). Once activated, osteoclasts form specialized membrane structures, such as the sealing zone, where they attach to bone and the ruffled border, through which they release hydrogen ions to aid in the mineralized bone matrix dissolution. Cytoplasmic vacuoles, containing primary and secondary lysosomes can also be seen close to the ruffled border area (Marks, et al., 1988).

Osteocytes are the most abundant type of cells in bone tissue, making up to 95% of bone cells, being ten times more common than osteoblasts. In spite the fact that osteocytes are buried in the bone mineralized matrix, they can still communicate and connect with each other and with cells located on the bone surface using a network of cell processes that cross their path through canaliculi in the ECM (Franz‐Odendaal, et al.,2006). These osteocytes are in the interior of the bone tissue and reside within spaces known as lacunae.

In addition, these cells have the capability to make and resorb bone to change the volume of its lacunae (Marks et al., 1988).

Bone lining cells, also known as resting osteoblasts, cover the surface of the bone to provide a selective barrier between bone and other extracellular fluid compartments.

Besides, they represent one of the two states of terminal differentiation of osteoblasts.

These cells have flat elongated morphology separated from the marrow (Marks, et al.,

1988). These cells do not achieve any bone formation or resorption, and they act as gatekeepers, informing if the bone needs remodeling. They can receive and deliver signals to other bone cells, as well as to those in nearby soft tissue (Parfitt,1994). Since bone lining cells are inactive, they require fewer organelles and cytoplasm than osteoblasts. Their function still not clear, although some reports have shown they secrete enzymes to dissolve

the osteoid matrix on the bone surface, allowing osteoclasts to remove the bone in the presence of parathyroid hormone (Downey, et al., 2006).

2.3.3 Growth factors

Growth factors and cytokines provide biochemical stimulation to cells within bone tissue inducing bone growth and repair. They are soluble polypeptides that target specific cells, binding them through transmembrane receptors (Vo, et al., 2012). There are many growth factors with the capacity to affect bone cell function and provide the necessary induction for a bone formation, also known as osteoinduction. The way growth factors induce changes in cell function is through something called signal transduction and involves protein phosphorylation, ion fluxes, changes in metabolism, gene expression, and protein synthesis. They are different from the hormones in the way of delivery and response

(Lee, et al., 2011). Growths factors found in bone tissue, such as, bone morphogenic proteins (BMP), insulin-like growth factors (IGFs), fibroblast growth factor (FGF), transforming growth factor β (TGF-β), and platelet-derived growth factor (PDGF), affect both cartilage and bone cells (Wozney, et al., 1988).

One of these growth factors BMPs have over sixteen subtypes classified due to their structure similarity (Haarman, et al., 2005). These originate from the TGF-β superfamily and play an important role in bone growth during embryonic development and during adult bone healing. Therefore, the studied elucidates that BMP-2 through 7 and BMP-9 are the most widely studied osteogenic molecules, which can provide the required signal for the differentiation of MSCs cells into osteoblasts (Harman, et al., 2005). During bone healing,

BMPs molecules can stimulate progenitor cells in the bone-formation cascade to promote

healing of the damaged tissue (Calori, et al., 2009). BMPs are now available commercially in a form of recombinant human (rh)BMP-2 and rhBMP-7, which have been approved for restricted clinical use, such as, for non-union, bone defects, open tibial fractures, and spinal fusions (Nauth, et al., 2011).

It has been established that various members of the TGF-β superfamily are involved in many functions related to bone induction, such as embryonic growth, tissue morphogenesis, cell proliferation and differentiation (Vo, et al., 2012). Also, IGF-I and

IGF-II are known to be involved in maintaining osteoblast function and enhancing cell proliferation and differentiation through the stimulation of precursor cells and the upregulation type I collagen expression. Besides, studies detected that downregulation of

IGF-I expression leads to apoptosis of bone cells (Canalis, 2009)

Another growth factor is known to play a vital role in the proliferation of mesenchymal stem cells (MSCs) and preosteoblast differentiation to osteoblasts is PDGF

(Khojasteh, et al., 2013). This growth factor consists of a disulfide-bonded dimer of two homologous polypeptide chains, which acts as stimulating factor for the stimulation of angiogenesis, which is an essential for bone healing (Elangovan, et al., 2014).

The FGF is an extremely potent mitogen for mesodermal cells by initiating angiogenesis required for wound healing; hence, it is important in bone fracture healing by providing an early vascular response through the formation of new microvessels (Kigami, et al., 2014). FGF-1 (acidic FGF) and FGF-2 (basic FGF) helped to promote angiogenesis and osteoblast proliferation, enhancing fracture healing (Sela, et al., 2012).

2.4 Bone diseases

Bone tissue might get damaged either by disease, fail while bearing a mechanical load, or from hormonal deficiencies that make the bone tissue predisposed to damage.

Bone defects include fractures, osteoporosis, osteogenesis imperfecta, osteomalacia, cancer, and Paget's disease.

Fractures result from either trauma or excessive mechanical strain that leads to discontinuity of bone tissue (Velasco, et al., 2015). Osteoporosis is an age-related disorder of the bone tissue microarchitecture and results from a loss in bone density. There are multiple causes leading to osteoporosis, for instance, hormone deficiency, poor nutrition from calcium and vitamin D deficiency, sedentary physical activity, and several pharmacological agents (Downey, et al.,2006). Osteogenesis imperfecta is a genetic disease that makes the bone brittle and fragile, due to a collagen matrix disorder caused by a mutation in one of the two genes that encode the chains of collagen type 1 (i.e., COL1A1 and COL1A2) (Rauch, et al., 2004). Osteomalacia is disease resulting from nutritional deficiencies that allow the bone to lose mineral (Velasco, et al., 2015). Cancer of bone tissue causes a massive pain and bone destruction. Metastatic cancer causes various effects on bone, such as an increase in calcium ions in the blood, the release of inflammatory mediators, which leads to the disintegration of the cortical and trabecular bone, predisposing to bone fractures (O’Toole, et al., 2006). Paget’s disease is a common aging disease affecting bone by inducing a high rate of bone remodeling, which results in abnormal bone formation. The cause of this disease can be either genetic or environmental,

and the main sites that commonly affected are the pelvis, vertebrae, and the femur (Al

Nofal, et al., 2015).

2.5 Bone replacement and tissue engineering

Due to increasing the number of patients with bone disease in the United States, with osteoporosis affecting over 30% of the population have led to an annual cost burden of $200 B. Orthopedic surgeons estimate that 50% of women with age exceeding 65 years are at high risk to develop bone fractures (Luu, et al., 2009). The cases where the bone defects are larger than the critical size of 1 mm, in which they can heal without assistance, are of specific concern (Hollinger et al., 1990). The current treatment for such types of defects involves the use of bone grafts, mainly autografts, allografts, and synthetic grafts

(ceramics and metals). The use of these bone graft is estimated at around 600,000 procedures annually in the United States, and approximately 2.2 million worldwide (Panek, et al., 2015).

The autograft is a bone graft used to restore integrity through the transplantation of healthy bone tissue from the same patient into the defect site, which helps mitigate immune rejection issues. This type of graft is considered ideal since it incorporates both the osteogenic cells and the mineralized matrix from the individual. The most common place used to harvest the bone from is the iliac crest. However, autografts have some limitations which limit its use, for example, donor site morbidity, pain, bleeding, infection, the struggle in shaping the bone grafts to fill the defect, and it is not suitable for large bone defects since it needs a large bone mass (Brydone, et al., 2010; Rose, et al., 2002).

The allograft is a bone graft harvested from a donor usually a cadaver which is processed and preserved for its future use. Although the allograft has more flexibility to be shaped according to the defect site, it raises the potential problems, such as immune rejection and pathogen transmission (Salgado, et al., 2004; Rose, et al., 2002).

Nevertheless, some procedures sterilize the bone graft, which has a negative influence on the osteoinductive properties required for bone healing, leading to longer healing time than autografts (Parikh, 2002).

Another substitute used to treat large bone defects are synthetic grafts such as those made from metals and ceramics. Both grafts work to provide the necessary mechanical support, but they have limited healing performance. For instance, metal grafts have poor integration with the host tissue at the injury site, which can lead to infection. Ceramics have limited tolerance to torsion, bending, or shear stress due to its low tensile strength and are brittle (Salgado, et al., 2004).

Still, there still an unmet need of developing an ideal bone replacement, which will require further work in bone tissue engineering and regenerative medicine, to provide the ideal substitute for treating bone defects. Tissue engineering is a developing field focusing on repair, replace, or regenerate tissue by combing physics, chemistry, and biology to create effective materials (Griffin, et al., 2004). The objective of tissue engineering and regenerative medicine is to restore, maintain, enhance tissue function to make in vitro tissue to implanted later in vivo; this goal can be achieved by the presence of three main components, mainly, a biocompatible scaffold, a cell source, and growth/differentiation conducting cell culture conditions (Kannarkat, et al., 2010).

2.6 Cellular sources for bone tissue engineering

For optimal bone regeneration and tissue engineering applications, it is important to use a reliable cell source to ensure no immune rejection, with osteogenic potential, a strong proliferation rate, and that allows good integration with the host tissues while offering the ability for load-bearing and remodeling (Logeart-Avramoglou, et al., 2005).

The most common types of stem cells that used for bone tissue regeneration are MSCs, embryonic stem cells (ESTs), and induced pluripotent stem cells (iPSCs). The main sources of MSCs are bone marrow (BM-MSCs), and adipose tissue (ASCs), which are usually cultured within 3D scaffolds to generate a new bone tissue with osteoinductive cues

(Griffith, et al., 2004).

The most common type of cells used for bone regeneration is adult stem cells, such as MSCs, due to their capacity to self-renew and are readily available. Also, they can be broadly proliferated in vitro, besides under specific culture conditions, they can differentiate into various phenotypes, such as bone, cartilage, muscle, marrow stroma, tendon/ligament, fat, and different connective tissues (Campagnoli, et al., 2001; Caplan,

2005; Vo, et al., 2012). In addition, in vivo behavior makes them an excellent candidate for tissue engineering applications due to their regenerative response to injury or disease

(Caplan, 2005). Stem cells generate progenitor bone cells, which during development become pre-osteoblasts, and osteoblasts (Griffith, et al., 2004). Stem cells are undifferentiated cells which can become specialized cells and tissues in response to specific stimulation, which can be biochemical and biophysical in nature. However, the life cycle of stem cells from totipotent stem cells (morula stage), capable of differentiating into all

embryonic and extraembryonic tissues. Then, their potency reduces to pluripotent stem cells (blastocyst stage), creating all embryonic tissues and to multi- or unipotent adult stem cells, forming tissues within their specific germ layer (Panek, et al., 2015). MSCs are culture-adherent, multipotent progenitor cells, and can be isolated from numerous tissues, for example, bone marrow, adipose tissue, muscle, amniotic fluid, human placenta, periosteum, cord blood and even peripheral blood (Vo, et al., 2012).

Pluripotent human embryonic stem cells (hESCs) form during embryonic development, isolated from the inner cell mass of blastocysts, have the capacity of differentiating into any type of specialized cell found in the bone. Some studies demonstrate that hESC-derived mesenchymal progenitors could have the same cellular behavior as the adult BM-MSCs (Marolt, et al., 2012). Since they are pluripotent, they can differentiate into cell types from all three germ layers. However, hESC raised ethical concerns, which greatly limited their clinical use since it required sacrificing embryos

(Bitar, et al., 2014; Vo, et al., 2012). Some studies showed that using hESCs might lead to the formation of a teratoma and that it might also cause immune rejection (Liu, et al., 2014).

Induced pluripotent stem cells (iPSCs) are artificially made pluripotent by making an adjustment in the expression of specific genes. Some studies have shown that human iPSCs exhibit some featured like hESCs, for instance, their morphology, gene expression, surface antigens, and in vitro differentiation ability and pluripotency. Thus, making them a viable alternative to hESCs for tissue engineering, and thereby eliminating any restrictive issues or ethical concerns (Teng, et al., 2014). Nevertheless, it is still important to continue with further research to define what is the best starting somatic cell for iPSC generation for

human clinical research (Amini, et al., 2012). Until now, most of the bone tissue engineering methods developed are based on the use of easily-accessible BM-MSCs as a cellular source. These have shown the ability to differentiate into chondrocytes and osteoblasts in vitro (Colnot, 2011). BM-MSCs are currently considered the choice for bone tissue engineering and regeneration applications due to their high osteogenic ability

(Yousefi, et al., 2016). However, the BM-MSCs have been shown to decay with growing passages, requiring longer periods to proliferate, thus showing limited proliferative capacity (Bruder, et al., 1997).

The stem cells used in this research are human adipose tissue (hASCs) since these very accessible, requiring only local anesthesia with slight patient discomfort, through procedures like, which is less invasive than bone marrow aspiration. Also, it found that 1g of adipose tissue contains about 5×103 hASCs, which is 500-fold higher than in bone marrow (Yousefi, et al., 2016). In addition, different studies imply that hASCs can commit to osteogenesis quickly and with less biochemical stimulation (i.e., through exogenous cytokines), making them better candidates for bone tissue engineering (Levi, et al., 2011), with equivalent potential to BM-MSCs of differentiating into cells from the mesodermal layer (Mizuno, 2009). Moreover, increasing evidence suggests that hASCs and BM-MSCs are distinct cell populations that differ in their inherent properties (Macotela, et al., 2012;

Tchkonia, et al., 2005; 2006). This is evidence through their surface antigen expression profiles, with hASCs being STRO-1 negative, CD36 positive and CD106 negative in contrast to BM-MSCs. In general, different surface markers have been reported to define hASCs, such as CD44 (hyaluronate) and CD90, as well as integrin b1 (CD29), endoglin

(CD105), and integrin a4 (CD49) (Levi, et al., 2011). In addition, an in vitro study using

hASCs showed these could differentiate into osteoblasts as early as 3 days after using induction media (Wan, et al., 2006).

2.7 Three-dimensional Scaffold for bone tissue engineering

To mimic the native tissue structure, 3D scaffold designed which is a fundamental tool in tissue engineering able to mimic the bone tissue microenvironment by modifying its composition, structure, surface chemistry, and stiffness (Marolt, et al., 2012). This design must control cell response, such as adhesion, migration, and proliferation (Causa, et al., 2007). Therefore, 3D porous scaffolds must provide surface and void volume that allows proper cell attachment, migration, proliferation, and differentiation to generate the desired tissue (Griffith, et al., 2004). Thus, the scaffold should be designed following a specific criteria appropriate for bone tissue engineering, for instance, it should be biocompatible and not induce an immune response, be nontoxic, absorbable at a rate proportional to bone formation, be osteoconductive, capable of being sterilized, and easy to manufacture and handle (Logeart Avramoglou, et al., 2005; Gunatillake, et al., 2003).

Moreover, the scaffold should promote cellular attachment, provide a porous structure to allow cells to migrate, differentiate and secrete their ECM, allow cells to integrate with tissue, and be in contact with bioactive molecules (Caplan, 2005). However, in the past decades, many efforts have focused on the discovery of nobel scaffolds materials that can be better substitutes for autologous or allogeneic bone grafts in the bone regeneration (Xu, et al., 2014). These have been developed by using either synthetic, natural, or composite

materials from organic or inorganic sources. These include synthetics such as calcium phosphates, which are inorganic, while polymers like poly(phosphazenes) poly(tyrosine carbonates), poly(caprolactones), poly(propylene fumarates), and poly(hydroxy acids) are organic (Karp, et al., 2003). In spite the fact that using natural materials have many benefits, synthetics offer benefits many of the former cannot, such as not having the immunogenic problem of natural biomaterials, or have a much better control over material properties. However, a major deficiency of synthetics is due to their limited biological recognition capacity (Causa, et al., 2007). In search of a balance, in this study, we used the self-assembled peptides, which are synthetic but based on amino acids, which are natural biomaterials to provide a biomimetic 3D microenvironment for hASCs.

2.7.1 Self-assembled peptide scaffold

The main building block of self-assembly molecular system is chemical complementarity and structural compatibility connected through weak noncovalent bonds such as hydrogen bonds, hydrophobic bonds, van der Waals interactions, and ionic complementary bonds under thermodynamic equilibrium settings. These bonds gather the molecules into stable organized structures (Zhao, et al.,2006). Self-assembled peptides have diverse applications extending from models to evaluate protein folding and protein conformational diseases to their application in tissue engineering and regenerative medicine as 3D scaffolds that mimic the microenvironment of native tissues and drug delivery. Due to their capacity to generate a diversity of structures, properties, being easily synthesized at a reasonable price, allow these self-assembled peptides as candidates in numerous technological innovations (Zhao, et al.,2006). Also, due to the peptidic nature of

these provides a great opportunity for in vivo applications since the degradation products of this peptides have a lower immune response, when compared to other (Cormier, et al.,

2013).

In 1989, a study was done by Shuguang Zhang on yeast genetics and protein chemistry led to the discovery of a self-complementary peptide. He recognized a protein named zuotin with the 16-residue peptide sequence repetitive motif, n-

AEAEAKAKAEAEAKAK-c (EAK16-II), between a segment of interchanging alanine-X repeating 34 residues. Later, he performed different analysis to evaluate its structure and its biochemical properties to produce a class of simple β-sheet peptides (Hauser, et al.,

2010). In this context, different types of peptides have been developed from this class, such as RADA16-I, RAD16-II, EAK-I, and EAK16-II, which all self-assemble into stable β- sheet structures in aqueous solution to form a nanofiber that can be used as a 3D scaffold for tissue engineering (Hauser, et al., 2010).

Self-assembled peptides type I are those with alternating hydrophobic and hydrophilic sequences that connect through different noncovalent bonds, forming a β-sheet structure in aqueous solution that contains monovalent alkaline cations or in physiological media

(Zhang, et al., 1994). This behavior results from the hydrophobic side chains on one side of the charged amino acid and hydrophilic side chains on the other surface that enabled it to be assembled into specific structure as pegs and holes are known as ‘‘molecular Lego’’ according to its appearance at molecular level (Zhang, et al.,1993; Zhang, 2002). This alternating of amino acid residues results in the formation of β-sheets with a nanofiber with

a hydrophobic core and a hydrophilic surface resulting from the stacking of two β-sheets into a simple fibril unit (Cormier, et al., 2013). Also, some studies showed that combining different type of polymers into the alternating amphiphilic-peptide, reveal that these polymers can form β-sheet structures that can aggregate, depending upon pH, salt, and time

(Zhang, et al.,1993). In contrast, peptides that have the same amino acids composition but with a different sequence, they tend to form α-coils structure instead of stable β-sheets

(Zhang, et al.,1994). These stable self-assembled nanofibers are affected by the pH and ionic strength of the aqueous solution (Nagai, et al., 2006). Once the type I peptides are assembled, they remain stable under varying chemical and physical states. Therefore, they demonstrate resistance to degradation by several proteases, heat, and chemical denaturation agents (Zhang, et al.,1993; Zhang, et al.,1995).

One member of the type I family is RADA-16 (Figure 3), which is commercially available as PuraMatrixTM that has been used in this research (Zhao et al.,2006). This type of self-assembled peptide produces microscopic and macroscopic matrices of interwoven nanofibers that form higher-order hydrogels in the presence of monovalent cations (Zhang, et al.,1995). The resulting hydrogel is based on 10-20 nm diameter fibers, 5-200 nm pores and a 99% water content (peptide content 1-10 mg/ml) (Wang, et al., 2008; Zhang, 2002).

Figure 3. Amino acid sequence of self- assembled peptide (RADA-16) (Zhang, et

al., 1993).

The amino acid sequence of this type contains positively charged Arginine,

Alanine, and negatively charged Aspartic acid, forming the RADA16-I peptide (AcN-

RADARADARADARADA-CONH2). The RADA-16 peptide is ionic self-complementary due to the presence of an ionic pair between the positive amino acid residues (arginine) and the negative amino acid residues (aspartic acid). Consequently, this sequence of amino acids leads to the formation of two distinctive hydrophobic and hydrophilic sides, whereas the hydrophilic sides represent the outside of the peptide fiber that is in direct contact with water, and the other side form a double sheet inside the peptide fiber (Hauser, et al., 2010).

Due to the presence of both the polar and the non-polar surface, a stable β-sheet is formed, considered essential for peptide self-assembly and nanofiber formation (Wang, et al.,

2008).

In aqueous solution, the PuraMatrix forms hydrogen bond through its backbone; besides they contain two distinctive sides, one hydrophobic due to the overlapping of alanine, as with the spider silk or silk fibroin, with the other side of the backbone being hydrophilic due to arginine and aspartic acid (Yokoi, et al., 2005). Although the forces generated during sonication can disassemble the RADA-16 hydrogels, they will readily re- assemble the moment sonication ceases. Sonication acts to mechanically break the hydrogen, ionic, and hydrophobic bonds to produce peptide fragments. This result when the hydrophobic bonds that formed between alanine and water are disrupted mechanically, during which time their cohesive ends start to find each other by sliding diffusion in aqueous solution due to it being energetically unfavorable (Yokoi, et al., 2005). These 16-

residue peptides have a length between 2.5 and 5 nm, and after reassembling they form a longer nanofiber between a hundred nanometers to a few micrometers in length (Yokoi, et al., 2005). Self-assembled peptides are commercially synthesized either through a solid phase or solution peptide synthesis chemistry (Hauser, et al., 2010). Since this self- assembled peptide scaffold contain a large amount of water, whereas water molecules able to be arranged by surface tension to create clusters divided by nanofibers into compartments. The 3D nanofiber scaffold was able to create an environment that mimics the in vivo conditions when cells create molecular gradients within the scaffold (Hauser, et al., 2010).

2.8 Biophysical stimulation

2.8.1 Effect of mechanical stimulation

Bone has the capacity to sense and adapt to any skeletal loading making bone a dynamic tissue capable of modifying its mass, strength, and geometry to accommodate for any external mechanical stimulation. This stimulation can affect osteogenic cells by causing a local deformation of the ECM, generating a fluid flow that causes shear stresses, and the initiation of electric fields (Mauney, et al., 2004; Zimmerman, et al., 2000). In general, all eukaryotic cells are sensitive to mechanical and physical forces, such as gravity, tension, compression, and shear that lead to modulation of cell function. Hence, mechanical stimulation can initiate a biochemical signal that can be interpreted as a cellular reaction in a process called mechanotransduction. In this process, mechanical energy is transformed into electrical and/or biochemical signals (Burger, et al., 1999). Since bone is stiff, exposure to a physiological load results in a small 0.2% deformation, while in vitro a 1 to

3% deformation is required to initiate a cellular response (Burger, et al., 1999). This leaves shear stress due to hydrostatic pressure and fluid flow, as the main mechanical stimulating forces (Huang, et al., 2010). Mechanotransduction can be described as the conversion of mechanical forces to biochemical signals that alter cell function through four different phases, which are: mechanocoupling, biochemical coupling, transmission of a signal to the sensory cell, and the effector cell response (Huang, et al., 2010).

To understand the mechanical stimulation from a cellular perspective, the applied forces on bone cause bone cells to deform and expose them to shear stress from the interstitial fluid motion in the canalicular spaces. In addition, fluid flow causes streaming effects that generate electric potentials (Pavalko, et al., 2003). This theory, called mechanosomes, regards the multiprotein complexes that represent the focal and cell adhesion protein complexes, the cytoskeleton, the muscleoskeleton and adherents’ junction protein that connect the neighbor cells and how they respond to load-induced deformations.

This cause changes in ions channels which drive variations in protein conformation

(Figure 4). This leads to the release of protein complexes known as mechanosomes, able to transfer mechanical information into the nucleus, causing alterations in DNA geometry and mediating the formation or mobilization of signaling complexes as the mechanical load transforms into chemical energy (Pavalko, et al., 2003).

Figure 4) The mechanosensors that resulted from changes in ions channels,

proteins, and cytoskeleton to activate the intercellular signals (Rubin, et al., 2006)

Therefore, when macroscopic loads generated, osteocytes which, act as the main sensory cells in bone, senses the interstitial fluid flow generated within the lacuna and the canaliculi, sending signaling molecules to either osteoclasts for bone resorption or osteoblasts for bone formation (Pavalko, et al., 2003; Huang, et al., 2010). As fluid flows within the lacunar-canalicular porosity, load-induced mechanical strains on cells actin filament of the cytoskeleton which are 1-2 orders of magnitude larger than whole-tissue level strains, resulting in intracellular signaling (Han, et al., 2004). Hence, this hypothesis suggests that these strains resulting from canalicular fluid flow act as a local force, instead of loading-related strains. Thus, when force is applied on the bone, fluid would squeeze out of the unmineralized matrix adjacent to cell bodies into the Haversian or Volkmann channels generating fluid shear stress on osteocytes membrane (Burger, et al., 1999).

Moreover, canaliculi fluid flow detected on osteocyte surface can generate shear stresses of 0.8–3 Pa (Burger, et al., 1999). As osteocytes become mechanically stimulated, the start paracrine signaling to stimulate osteoblasts into the bone formation, including increased release of nitric oxide (NO) and prostaglandin PGE2 and PGI2, and IGFs (Orr, et al., 2006;

Burger, et al., 1999). The study the mechanotransduction on bone cells have recognized numerous candidates involved in the mechanosensing process. Mainly, mechanically gated ion channels, integrins and focal adhesions kinase, G proteins, and the interaction between the cytoskeleton and certain phospholipase C isoforms. Current researchers have shown that focal adhesion kinase plays a vital role in mechanically induced bone formation in vivo

(Morgan, et al., 2008).

Since osteoblasts act as an effector cell under proper physical stimulation, these can initiate osteogenesis. Mechanical stimulators such as stress, strain, and hydrostatic pressure have been proven to induce bone regeneration and fracture healing (Xu, et al., 2014). In vitro studies have shown that cyclic pressure improves osteoblast functions related to new bone development, using a custom-made system that provides cyclically oscillating pressure with specific amplitude and frequency. These studies demonstrate the upregulation of osteogenic biomarkers type-I collagen, osteocalcin, and TGFβ1 (Nagatomi, et al., 2003). Another study using combinations of the two key mechanical stimuli that affect mesenchymal tissue differentiation (i.e., shear strain and fluid flow), showed bone- healing through histological analyses (Morgan, et al., 2008). An in vivo study of using 1.2

Pa of fluid flow shear stress for mechanical stimulation revealed an upregulation of COX-

2 and c-Fos expression, which are important to maintain the osteoblast phenotype (Pavalko, et al., 1998).

2.8.2 Effect of electromagnetic fields

Electromagnetic fields play a vital role in tissues development through a cascade of processes for tissue regeneration that include cell interactions, ECM synthesis, cell migrations, differentiation, and proliferation. In 1974, electromagnetic fields were introduced as time-varying fields used for therapeutic purposes, specifically to treat nonunion bone fractures and congenital pseudarthrosis (Bassett, et al.,1987). After many studies analyzing the influence of magnetic fields on cells, FDA approved to use pulsed electromagnetic fields (pEMFs) in 1979 as an active and harmless mechanism for healing nonunion, congenital pseudarthrosis, and failed fusions (Carpenter, et al., 1994). The idea behind using EMF to induce osteogenesis come from the natural endogenous streaming potentials in bone during deformation. Clinical trials using this treatment were carried through direct conduction by using electrodes; then, they used a wire coil to the general magnetic field at fracture site through forcing electric currents on it. Later, time-varying magnetic field was used to generate the required electric field in bone via Faraday coupling

(Pilla, 2002; Funk, et al., 2006). Those devices used pEMFs with extremely low frequency ranging from 1 to 100 Hz, inducing fields on the microvolt/centimeter level at the fracture area (Funk, et al., 2006). The reason to use the time-varying magnetic fields is to induce an electrical field like the one that it generated in dynamically deformed bone tissues with a similar signal shape and amplitude. The generated fields are used to trigger specific cellular responses, by relying on the nature of cellular targets, its tissue environment, and its function (Carpenter, et al., 1994). Recently, pEMFs have been used as a therapeutic

procedure to treat pain, inflammation, and dysfunctions related to rheumatoid arthritis and osteoarthritis (Ganesan, et al., 2009).

FDA-approved noninvasive magnetic fields (pEMFs) used for bone healing and remodeling, are generated using coils in a Helmholtz arrangement (Figure 5), to initiate pulses repeated signal with extremely low frequency (15 Hz) and making millivolt per centimeter (mV/cm) electric fields at the treatment area (Pilla, 2002). Magnetic fields with a frequency range between 0 Hz up to several hundred GHz are considered as nonionizing radiation, and researchers have found that this nonionizing electromagnetic energy can generate different biological effects through interaction mechanisms that do not contain any macroscopic heating, when the field is applied to tissue samples, it found that the global temperature change is generally less than 0.001°C (Walleczek, 1991). The magnetic fields with frequency band between 3 Hz-3 kHz are considered as extremely low EMFs

(Ganesan, et al., 2009).

Figure 5. Schematic illustration of using Helmholtz coil to generate PEMF (Rauh, et al.,

2011).

In biological tissue, the effect of EMFs can generate coherently oscillating forces on charged molecules within the tissue, in a phase with the polarized field and on parallel planes between each other (Panagopoulos, et al., 2015). Even though the oscillation of the molecules is at high velocity which results from the thermal motion, it has no biological effect, the fact that these are coherently polarized allows them to initiate changes at the cellular level (Panagopoulos, et al., 2015).

To understand the effect of EMFs on tissue engineering and regenerative medicine, we need to understand the how of electric and magnetic fields interact with the native tissue. During early embryo development, cells move towards forming an organ, through migration driven by voltage gradients from by the distribution of charged ions, such as Na+,

Cl-, K+. The currents generated by passive sodium uptake from the environment leads to an internally positive transepithelial potential difference with an endogenous static electric field is in the order of 1–5 V/cm. This field is also generated in wounds resulting from disruption of this transepithelial potential in the epithelial layer.

In bone tissue, the therapeutic use of electric fields is derived from the observation that when bones are placed under mechanical load (stress) the deformation (strain) generates an electrical potential. This voltage gradient generates between liquid and solid from endogenous streaming potentials produced due to fluid motion (Kovacic, et al., 2010;

Funk, et al., 2006). Thus, an EMF is generated due to these processes, which could induce an electric current in bone tissue by Faraday coupling (Funk et al., 2006). It has been shown in various studies that there is very close correlation connected between EMF and mechanical vibration due to the piezoelectric properties of bone (Funk, et al.,2006).

At the molecular level, weak EMFs have more influence on cell biology than strong ones, and its bioeffect is seen in signal-transduction cascades, such as the Ca2+ transport system, gene expression, cell growth, and apoptosis (Kovacic, et al., 2010). In any typical tissue, there is intercellular space composed of small narrow fluid channels of 150 Å between cells that provide conduits for the cell to cell communication. Those channels have a distinct low impedance as compared to the cell membrane, making them a preferred path for environmental EMF-induced currents, since it can transmit around 90% of intrinsic current, leading ionic species to the membrane surface. These channels sense any weak electrochemical oscillations in the pericellular fluid through the charged tips of glycoproteins that detect any chemical and electrical signals in the surrounding fluid

(Kovacic, et al., 2010).

There are different theories describing the effect of weak EMFs on cells membrane and on ion channels. One of these theories is the forced vibration ion theory which states that there are various kinds of ions surrounding both sides of cell membranes, such as, K+,

Na+, Ca+2, Cl-, etc., which take part in the cell’s signal transduction, and in establishing the transmembrane electric potential. A flux of ions occurs due to the ion concentration or electrical potential gradient, which induce their movement through mechanically gated ion channels. Therefore, when a pulsed low external electric or magnetic field is applied, it will generate an oscillating force on the cell's membrane, hence on the free ions on both sides of the membrane and on the ion channel proteins. These vibratory forces will give the oscillating ions a false signal for gating channels that led to disorder the electrochemical balance of the cell's membrane and consequently the entire cell function. The ions will be

moved with a homogeneous motion where all exhibit the same value and phase. In addition, it has been suggested that the low-frequency magnetic fields are more bioactive than those with a higher frequency because there is an inverse relationship between the amplitude of the forced-vibration and the frequency of the field (Panagopoulos, et al. 2000;

Panagopoulos, et al., 2002).

2.9 Effect of magnetic scaffold under EMF stimulation for bone regeneration

Biophysical forces, mainly mechanical loading and electromagnetic signals are essential regulators of bone formation. Tagging superparamagnetic NPs to mechanosensitive cell membrane receptors in osteoprogenitor cells, allows the possibility of mechanically activating these cells with an external magnetic field enhancing their osteogenic potential (Kanczler, et al.,2010). Providing the correct stimulatory conditions that leads to a tissue promoting microenvironment in vitro and in vivo is considered a crucial goal for regenerative medicine (Sapir-Lekhovitser, et al., 2016). Over the last decades, superparamagnetic NPs (mainly iron oxide-based) have been widely used in many biomedical applications (Liu, et al., 2009). When the alternating magnetic fields are combined with superparamagnetic NPs embedded within 3D scaffold structures, it is possible to induce mechanical forces on the scaffold to which cells are attached, thus inducing mechanical forces on the cells themselves (Sapir-Lakhovister, et al., 2016). The design of novel superparamagnetic scaffolds for bone tissue engineering have generated much interest in recent years. An approach involves dip-coating of the scaffolds in aqueous ferrofluids comprising biocompatible and nontoxic superparamagnetic NPs, allowing them to infiltrate to the pores of the scaffold. In this manner, the resulting superparamagnetic

scaffolds are capable to up taking cells and growth factors (Bañobre-López, et al., 2011).

Also, an in situ method can be used to generate a superparamagnetic nanocomposite scaffold by mixing the superparamagnetic NPs during scaffold synthesis, reducing the number of processing steps and time (Sivudu, et al., 2009). Most of the superparamagnetic

NPs used are made from iron oxides, such as magnetite (Fe3O4) and maghemite (γFe2O3)

(Dobson, 2008). In bone regeneration, superparamagnetic NPs are widely used for in vitro as well as in vivo applications, with sizes in the 1–100 nm range. A special characteristic of interest is their superparamagnetic nature, in which they are only magnetized under the effect of the external magnetic field, allowing their control noninvasively (Kannarkat, et al., 2010). Superparamagnetic NP parameters such as size and shape have an impact on their properties, for instance, their coercivity and magnetization values and the ability to change the magnetic behavior from the ferromagnetic regime to the superparamagnetic regime (Jun, et al., 2008). Therefore, by applying a magnetic gradient field to a superparamagnetic scaffold, causes NP displacement within the scaffold where cells are bound, inducing compression and tensile forces on the cell membrane, leading to cytoskeleton deformation and cell dragging. Membrane receptors such as integrins act to transmit these forces applied on the cytoskeleton through activation of intracellular signaling pathways to regulate osteocyte and osteoblast function (Russo, et al., 2016).

These superparamagnetic NPs act as magneto-mechanical stimulators of cell arrays at the cellular level, impacting them through cell proliferation, differentiation, and migration

(Lima, et al., 2015). Therefore, superparamagnetic NPs act as a remote stress inducer without the need for invasive bioreactor system to mimic the in vivo environment (Dobson et al., 2006). Furthermore, researchers also utilize superparamagnetic NPs generating

magnetic force, using magnetic fields, for inducing drag and rotation. The magnetic drag technique require NPs coated with a cell-specific ligand that attaches to surface receptors, such as integrins, to allow vertical displacement when applying magnetic fields. These forces cause deformation of cell cytoskeleton that activate different mechanosensitive ion channels. The benefit of this technique is that it produces a localized force on the specific cell receptor. The other approach is the twisting or rotation method, which works by magnetizing the NPs in one direction for short pulse then applying a second weaker pulse perpendicular to the magnetic fields, making the NPs rotate. The advantage of this approach is that it produced a localized mechanical stress instead of a deformation of the entire cell membrane (Hughes, et al., 2005). Mechanical stress can also result from generating a dragging force by magnetic NPs with specific strength and frequency on cell cytoskeleton, resulting in increased phosphorylation of tyrosine kinases, including the

MAP1 kinase activity (Schmidt, et al.,1998).

Overall, the rate of bone cell growth, proliferation, and differentiation have been improved by incorporating superparamagnetic NPs into the scaffold in the presence of external magnetic fields (Kannarkat, et al., 2010). In recent years, nanoscale metals in the shape of metal oxides in polymer-based nanomaterials have been studied for their enhanced antimicrobial characteristics in numerous fields (Dhivya, et al., 2015). The research by

Bock and co-workers developed a composite scaffold based on collagen and hydroxyapatite (HA), the latter was included because of its high osteoinductive properties, lack of antigenicity or cytotoxicity, and low degradation rate. Collagen was added for its osteoinductive properties and biocompatibility. Superparamagnetic NPs were added to the scaffold using the dip-coating method (Bock, et al., 2010). In another study, Kannarkat and

his team developed a magnetic scaffold structure that mimics the natural bone tissue and

ECM using polycaprolactone-based scaffolds fabricated by electrospinning. They added

1–100 nm superparamagnetic NPs (Fe3O4) with the polymer precursor prior to electrospinning. These NPs along with an external magnetic field allow the induction of low-level mechanical stress within the scaffold, producing shear stresses at the cellular level. The mechanical stresses on the cells increase the expression levels of multiple genes, the production of the second messenger nitric oxide and cyclic adenosine monophosphate and increase in the activity of various proteins. The external magnetic field used was 1–6

Gauss at 15 Hz, applied for two hours daily on MC3T3-E1 mouse preosteoblasts. Their results showed that the attachment of the cells was similar for all scaffolds with and without

NPs. It was also observed that the cells formed clusters after nine days in culture as signs of proliferation, but it was slower than the control. They also noticed the cells migrate into the scaffold due to the high porosity, after cells adhered to the scaffold they exhibited an elongated shape as a sign of differentiation to osteoblasts (Kannarkat, et al., 2010).

Russo and co-workers used a collagen/HA composite scaffold with superparamagnetic NPs (mean diameter ~200 nm) for bone regeneration in an in vivo bone defect model using a rabbit femoral condyle under the effect of static magnetic fields. In their experiments, they considered four groups, in which the permanent magnet without superparamagnetic scaffolds was used as controls, while the other two groups fabricated scaffolds by two methods. The first method synthesized a scaffold made of type I collagen fibrils-HA mixed with uncoated ferromagnetic iron (II, III) oxide NPs (<50 nm), yielding freeze-dried cylindrical porous scaffolds. The second method allowed the infiltration of the

NPs into the collagen/HA hybrid porous scaffolds by capillary action. Their results showed

that after 12 weeks the bone regeneration noticed with both of magnetic scaffold and the permanent magnet was significantly higher than with a non-magnetic scaffold and that the scaffold made using the first method was more osteoinductive than the first Also, the nanoindentation results showed that the mechanical properties of newly-formed bone within the first scaffold were closer to that of native tissue and more mature bone trabeculae as compared to the other groups. In addition, they noticed a clear reorganization of the scaffold architecture under the static magnetic field in vivo, where the magnetized collagen fibers aligned in a similar way of the field lines generated by the permanent magnet (Russo et al., 2016).

In another study, Gloria, et al., (2013) developed a magnetic scaffold for bone tissue engineering, based on PCL, to which iron-HA NPs were added during polymerization. The aim was to induce mechanical stimulation on the seeded hMSCs using alternate external magnetic fields (27 mT, 260 kHz) to control cell function. The magnetic scaffold used three different polymer-to-particle weight ratios, i.e., 90/10, 80/20 and 70/30 w/w. Results indicate that the higher the number of NPs embedded in the scaffold the more hydrophilic and nanostructured would the surface be. They also observed in all groups a magnetically induced thermal response as a function of NPs during characterization. This is an expected result due to the high-frequency field used, which can be explained by the energy released through the Neel relaxation process. Osteogenic differentiation of the hMSCs cells in vitro by ALP activity showed significant differentiation after a week that the magnetic scaffolds supporting the osteogenic differentiation. It is important to mention that these magnetic scaffolds were only exposed to magnetic fields during characterization and not during cell growth, (Gloria et al., 2013). Also, Panseri and co-workers developed a biomimetic

magnetic scaffold, exposed to a rare earth permanent magnet (1.2 T) to assess in vivo bone regeneration using a rabbit model. Defects 2 mm in size were drilled on the lateral condyle of the distal femoral epiphysis and filled with one of two different scaffolds, based on collagen-HA and superparamagnetic iron oxide NPs. Their results showed that the static magnetic fields have no undesirable impact on tissue formation and these magnetic scaffolds seemed to have well integrated with an adjacent cancellous bone with no necrosis or inflammatory response to corrosion products and iron toxicity. After 4 weeks, they noticed inside scaffolds thin bone trabeculae while in the periphery of the scaffolds were more mature trabecular tissues. Furthermore, the histological analysis showed a newly formed woven bone through the scaffolds structure with the presence of normal osteocyte lacunae developed inside and around magnetic scaffolds, and typical mineralization gradient initiated from outside to the inside of scaffold. These results confirmed that the magnetic scaffold along with static magnetic fields has an influence on bone tissue remodeling and tissue regeneration (Panseri, et al., 2013).

In another study Panseri, et al. (2012) analyzed the effects of the adding different amounts of magnetic NPs to HA scaffolds (HA/NP 100/0, 95/5, 90/10 and 50/50 wt.%) in bone tissue regeneration, in an in vitro model using human osteoblast-like cells (Saos-2) and an in vivo rabbit model under the influence of applying a static magnetic field of 320 mT. Their results showed a significant increase in cell proliferation on the 90/10 scaffold.

ALP activity was measured for all groups, showing no significant increase between the groups, with no influence by the presence of the magnetic field. On the other hand, the in vivo experiments showed similar histocompatibility. Also, macroscopic evaluation displayed no infection or tissue necrosis in none of the scaffolds. Bone tissue was

observable around and inside the scaffold in both groups and some pores were full of new bone, verifying a high level of histocompatibility with the magnetic scaffold comparable to the control group (Panseri, et al., 2012).

Another in vivo study using superparamagnetic scaffolds combined with the external static magnetic field using a rabbit model. The scaffolds consisted of a composite of γ-Fe2O3 and HA NPs in poly(lactide acid) prepared by electrospinning. The magnetic scaffolds were implanted in New Zealand white rabbits, and to provide the magnetic field they fixed permanent magnets to the rabbit cages of opposite sides. Their results showed that at day 10 there were host-derived cells composed mainly of macrophages and fibroblasts that had migrated to the defect site. Over time, there was degradation of the implanted scaffold and showed the presence of osteoblast cells and ECM around the scaffold at day 20 as an indication of a new bone tissue formation. This behavior increased with time showing an increment in the new bone tissue as the scaffolds degraded. Also, the superparamagnetic scaffolds exposed to magnetic fields displayed significantly more collagen than those without magnetic stimulation, which indicates bone growth increased with magnetic field stimulation. In addition, the new bone tissue became connected and homogeneous with an organized morphology similar to the original tissue and with faster degradation rate, while the scaffolds group without external magnetic field exhibited non- homogeneous bone tissue with slower scaffolds degradation. Thus, the nanofibrous magnetic scaffolds with the static magnetic field stimulation were able to mimic the original ECM microenvironment in the defect, promoting osteogenic cell attachment and growth (Meng, et al., 2013).

Zeng, et al., (2012) studied the effect on adhesion, proliferation, and differentiation of osteoblastic cell lines by superparamagnetic NP content in HA scaffolds after being stimulated by magnetic fields (1 mT = 10 Oe in Air, 50 Hz). Rat osteoblasts (ROS 17/2.8) and mice preosteoblasts (MC3T3-E1) were chosen and grown in HA scaffolds with a 70-

80% porosity. with micro- and macro pores with interconnectivity comparable to native spongy bone tissue, and with different concentrations (0.2 to 2.0 wt.%) of 8 nm superparamagnetic NPs introduced by dip-coating. Their results showed that the ALP activity of both cell lines grown in superparamagnetic scaffolds was significantly higher under magnetic field stimulation than in the scaffolds without NPs. Also, they revealed that with the increase of NP content within scaffold affects cell adhesion, proliferation, and differentiation, giving the superparamagnetic scaffold the ability to attain intrinsic magnetic therapy and gain some synergistic effect to enhance the cell response when exposed to magnetic fields, allowing them to be used as a bone graft substitutes (Zeng, et al., 2012).

CHAPTER III

Materials and Methods

3.1 Cell culture

Human adipose-derived mesenchymal stem cells (hASCs) purchased from Lonza

(Walkersville, MD) were cultured under standard culture conditions in a sterile, humidified incubator at 37°C, and 5% CO2/95% air. Cells were cultured in T75 flasks at a density of

5.0×105 cells/flask, using mesenchymal stem cell growth proprietary kit purchased from

ScienCell (Carlsbad, CA) that consisted of 500 ml of basal medium, 5 ml of MSC osteogenic differentiation supplement and 5 ml of a penicillin/streptomycin solution. The appropriate amount of growth medium was added to each flask (0.2-0.4 ml/cm2) and was replaced every three days. The cells used in all experiments were from the fourth passage.

3.2 Three-dimensional cells encapsulation and gel formation

To prepare the three-dimensional (3D) cell culture, the self-assembled peptides

PuraMatrixTM composed of standard amino acids (1% w/v) and 99% water were purchased from Corning (New York, NY). The encapsulation of cells within the hydrogel scaffold was prepared following the manufacturer’s instructions. Briefly, PuraMatrixTM stock solution (1% w/v) was sonicated for 30 min in the ultrasonic bath to remove air bubbles

and reduce its viscosity. Then, the required amount of the PuraMatrixTM stock was aliquoted and mixed with 20% sterile sucrose at a 1:1 ratio to yield a 0.5% w/v concentration (half of the 20% sucrose was mixed with the gel and the other half with suspended cells). At this point, 10 nm carboxy-functionalized superparamagnetic iron oxide nanoparticles (Sigma-Aldrich, St. Louis, MO) were added at a concentration of 0.5% of the total volume of the mixed solution. Subsequently, the cell suspension was prepared by trypsinizing the flasks with a 0.25% trypsin/EDTA solution purchased from ScienCell

(Carlsbad, CA). Cells (1.5×105 cells/well) were resuspended in sterile 20% sucrose. The cells/sucrose mixture was mixed equally with hydrogel mixture, to be transferred quickly to the center of 24-well plates with total volume of 150 µL/well and 750 µL media was added to form the gel in each well. Since the PuraMatrixTM pH is 2-2.5, the media was changed twice to equilibrate to the physiological pH of 3D cell culture.

3.3 Cells viability assay

Lactate dehydrogenase (LDH) assay (CytoTox 96R Assay kit) purchased from Promega

(Madison, WI) was used to evaluate the cytotoxicity and proliferation of cells cultured within the hydrogel. LDH is a stable cytosolic enzyme that can be measured from the cells lysate, which reacts with a tetrazolium salt (i.e., iodonitrotetrazolium violet) to form a deep red formazan dye. The number of cells was assessed at specific intervals (7, 14, and 21 d) according to the manufacturer's protocol. Briefly, to prepare the lysis samples, the cells- gel constructs were treated with a collagenase to digest the collagen secreted from cells.

The samples were incubated in 50 µL of collagenase for 30 minutes after washing with

PBS. Later, each sample was mixed with 500 µL of lysis buffer (1% Triton X-100), then sonicated for 1 hour in a sonicator. The samples were then centrifuged at high speed. Next,

50 µL of the supernatant and 50 µL of CytoTox 96 Reagent were added to each well of a

96-well plate and covered with foil for 30 min at room temperature. Then 50 µL of the stop solution was added to each well. Absorbance at 492nm was recorded and using the calibration curve, cell number was quantified.

3.4 Cells differentiation assay

The alkaline phosphatase (ALP) assay was used to detect osteogenic differentiation of hASCs. ALP is early osteogenic marker expressed on the cell surface. The SensoLyte® pNPP Alkaline Phosphatase Assay Kit (AnaSpec, Fremont, CA) was used by measuring the absorbance at 405 nm. Briefly, after removing the growth medium from each sample and washing with PBS, 50 µL of collagenase was added and incubated for 30 minutes.

Then, 500 µL lysis buffer (1% Triton X-100) was added to each well. The cell lysate was ready after sonication for 1 hour and centrifuging at 10,000 rpm for 5 min. Then 50 µL of lysate cells were added to 96-well plate with a flat bottom, and 50 µL of pNPP substrate solution was added to detect the ALP. Upon dephosphorylation of pNPP, the lysate samples turn yellow. After incubation for 30 minutes at 37ºC, 50 µL of stop solution was added to stop the reaction. With the absorbance reading from the microplate reader, the ALP secretion was quantified using the calibration curve. Then, the data were normalized with

the total number of cells per well, expressing the ALP concentration in ng/cell. This assay was performed at 7, 14, 21 d of incubation in triplicate.

3.5 Measurement of mineralization

Mineralization on the constructs was quantified using with Inductively Coupling Plasma-

OE Spectroscopy (ICP-OES, Varian) detecting calcium at 7, 14, and 21 d of incubation

(n=3). The samples were decalcified in 35% HCl (trace metal™ grade, Fisher Chemical), followed by boiling for 5 h at 90ºC. Then, the samples were collected in liquid form, and calcium was measured with ICP-OES through emissions at 396 nm. From the emission intensity, the calcium concentration is obtained in mg/L (ppm) using a calibration curve.

3.6 Cells morphology

To investigate cell morphology in response to variations in the extracellular environment, cytoskeletal actin microfilaments (F-actin) were stained using fluorescent phalloidin conjugates (F-actin visualization Biochem kit). Also, 4',6-diamidino-2-phenylindole, dihydrochloride (DAPI) was used to stain the cell’s nucleus This experiment was performed at 3, 14 d and the results were visualized using by confocal microscopy

(Olympus FV1000) with an excitation/emission filter for F-actin (535/585 nm), and DAPI

(358/461 nm). The staining was done according to manufacturer's instructions. Briefly, after removing media from each sample and washing them with washing buffer, 200µL of

the fixative solution was added for 10 min. Then, the samples were washed twice with washing buffer to remove the fixative materials, 200 µl of permeabilization buffer was added to each fixative sample. After 5 min, each sample was washed twice with washing buffer. A 0.165 μM F-stain phalloidin and a 300 nM DAPI solution were prepared, then the stained sample was covered with foil to keep it dark for 30 min at room temperature. Then, each sample was washed three times to stop the reaction and covered with mounting medium and coverslip.

3.7 Alkaline phosphatase staining

For qualitative investigation of cells differentiation, Stemgent Alkaline Phosphatase (ALP)

Staining Kit II (Cambridge, MA) has been used to detect ALP activity within the samples at day 14. Briefly, after removing media, samples were washed with 0.05% concentration of PBS containing Tween-20 as a permeabilizing agent. 0.5ml of fixative solution have been added for 5 to 10 min. Then, the samples were washed twice, and 0.6 ml of ALP staining was added that composed of equal ratio 1:1:1 of AP substrate solution (a mixture of 0.2 ml of solution A and 0.2 ml of solution and 0.2 ml of solution C) for 15 minutes at room temperature. For stopping the reaction, the samples washed twice with PBS. Later, the samples were covered with mounting medium and coverslip to prevent drying. By using, the ALP expression was detected as a red or purple stain.

3.8 Fourier transform infrared spectroscopy (FTIR) analysis

The self-assembled peptides (PuraMatrix) with and without iron oxides nanoparticles scaffolds have been characterized by using FTIR device. Since FTIR spectrum provides information about specific functional groups presented in each of hydrogel type through measuring the transmittance frequency at which specific atoms will be vibrating. The characteristic of testing hydrogel will show the transmittance and wavelength that range between 400 to 4000 cm– 1.

3.9 Electromagnetic fields exposure system

The equipment that used to generate extremely low-frequency pEMFs consisted of a function generator, oscilloscope, Helmholtz coils, and µ-metal box. The function generator

(Agilent) generated the burst signal with specific frequency, amplitude, and shape. The test signal was equivalent to FDA approved signal for bone fracture healing, which consists of

15 Hz pulse burst of 20 pulses with magnetic field increased from 0 to 1 mT in 5 ms and then decreased to zero in 61 ms [Bassett, et al., 1982; Daish, et al., 2018, Polo-Corrales, et al., 2018]. An oscilloscope was used to display the generated signals in terms of voltage as a function of time. The EMF was generated using Helmholtz coils (3B Scientific®

Physics), which consist of a pair of copper coils that operate on alternating fields. When an alternating current ran through the Helmholtz coils, uniform electromagnetic fields are generated in space over a considerable volume. The samples were placed in the center of

the coils. The main feature of each coil is composed of 124 turns, outer coil diameter is

311 mm, inner coil diameter is 287 mm, mean coil radius is 150 mm, and coil resistance is

1.2 ohm. Thus, from coil featured and the current intensity, the magnitude of magnetic flux density (B) can be measured according to the following formula (Gupta, et al., 1991):

3 5 2 푛 퐵 = ( ) 휇 퐼 4 0 푅

where n is the number of turns in each coil, R is mean coil radius and 휇0 is permeability of free space (4π × 10−7 Tꞏm/A). which leads to B = 7.433×10‾⁴ I in T.

The Helmholtz coils were placed in the incubator inside a μ–metal enclosure, which shields against the earth static fields and low magnetic fields from the equipment around it. This assures that the only EMFs exposure comes from the Helmholtz coils. This μ–metal is composed mainly of nickel, iron, and some copper or chromium that gives a low reluctance path for the magnetic flux.

The experimental groups were divided into two, pEMF stimulated and non-stimulated.

Each experimental group consisted of four different formulations, all based on hASC- seeded scaffolds, with/without osteogenic induction media, and with/without NPs. The stimulated group was exposed to pEMFs for 8 h every day. The non-stimulated group was placed in the pEMF system for 8 h per day but it was kept off. This accounted for any differences in cell culture conditions between the pEMF system, and the cell culture incubator, where all cell cultures were kept the remaining time.

a b

Figure 6. Electromagnetic exposure system: a) closed mu-metal box and b) Helmholtz coils with cells.

3.10 Statistical analysis

All numerical data were evaluated statistically according to the ANOVA test followed

Dunnett test to identify the significant differences. Values of p < 0.05 were accepted to be used as a significant level of difference between the mean of the experimental and corresponding control groups.

CHAPTER IV

Results &Discussion

4.1 Results & Discussion

4.1.1 Viability and Proliferation

Cell proliferation was quantified using an LDH cytotoxicity assay using passage 4 hASCs for four different formulations that were either pEMF stimulated or non-stimulated.

The results were assessed after 7, 14, and 21 days of culture for all formulations. The results showed (Figure7) that cells start to grow rapidly for all groups. However, there was no statistically significant difference in cellular viability between groups with and without pEMF stimulation.

At day 7, within the non-stimulated (i.e., without pEMF or osteogenic induction media) group there was a significant increase (p<0.001) in the number of viable cells detected within the superparamagnetic (NP-doped) scaffold over those using the hydrogel alone. This revealed that the carboxy-functionalized iron oxide NPs not only do they have any cytotoxic effect on the cells but in addition promote their proliferation. It is known that carboxy functionalities increases hydrophilicity and significantly increases the proliferation and osteogenic differentiation of MSCs (Phillips, et al., 2010). It has also been shown that by using electrospun paramagnetic scaffolds based on γ-Fe2O3/nano- hydroxyapatite/poly (lactic acid), the proliferation of preosteoblasts was enhanced

the group of superparamagnetic scaffolds under pEMF stimulation (without osteogenic induction media) showed a similar proliferation rate to the non-stimulated group, which were significantly different (p<0.001) to the control (lacking NPs). The biochemical stimulation using osteogenic induction media alone was enough to significantly (p<0.001) enhance cell proliferation when compared to the non-stimulated control. Although this significant increase in the rate of proliferation was also seen on both pEMF stimulated and non-stimulated superparamagnetic scaffold groups, there was no significant difference between them, but only with respect to the control (p<0.0001).

At day 14, all the groups continue to grow, under basal media, with similar proliferation rates, but with no significant differences between pEMF stimulated and non- stimulated groups regardless of the presence of NP-doped superparamagnetic scaffolds, when compared to the control. When cells were stimulated with osteogenic induction media there were no significant differences between pEMF stimulated and non-stimulated groups when grown in the hydrogel alone, but significantly different with the control

(p<0.05). On the other hand, there was a significant difference (p<0.001) when these were grown in superparamagnetic scaffolds, with enhanced the cellular proliferation with pEMF stimulation. At day 21, we observe the same trend as for day 14. In addition, the group of stimulated with osteogenic induction media has a significant difference (p<0.05) as compared to control.

Figure 7. LDH assay. The proliferation of hASCs seeded within self-assembled peptide hydrogel with and without NPs under extremely low-frequency pEMF of 1 mT was quantified, after 7, 14, 21 d. *p < 0.05; **p < 0.001; ***p < 0.0001, indicate statistically significant differences between basal media group (control) and the other groups. The mean values are calculated from the average results of three samples, the results are represented as mean ± SD.

In this study, the results from the proliferation assay showed no cytotoxicity from the hydrogel with or without the presence of NPs towards hASCs, showing an appreciable proliferation rate. The assessment of biochemical, mechanical, and electromagnetic stimulation on hASC proliferation shows that the simultaneous presence of osteogenic induction media and superparamagnetic NPs has the strongest effect on cell proliferation, without any significant impact by pEMFs. This lack of effect due to pEMF stimulation on proliferation was also seen in the non-stimulated (i.e., biochemical and mechanical)

controls. The presence of either osteogenic induction media or superparamagnetic NPs had an equivalent positive effect on hASC proliferation. Although this positive effect was significantly less than the combined presence of osteogenic induction media and NPs. Also, during the time and after 21 days of culture the cells have the same trend of proliferation as the cells grown after 14 days. This behavior is due to cells that differentiated and stopped proliferating, whereas cells exhibit cell-cycle arrest following differentiation by the activation and deactivation of a collection of cyclin-dependent kinases, which regulate specific steps in the cell cycle (Myster, et al., 2000).

The objective of adding superparamagnetic NPs to the hydrogel was so that the matrix would be induced to mechanically vibrate with the application of an alternating magnetic field (Golovin, et al., 2017). The results show that although the presence of superparamagnetic NPs promotes hASC proliferation, the mechanical vibration induced by the pEMFs is either not required or that the force generated is too weak to elicit a response from the cells. It must thus be an intrinsic property of the NPs, which stimulates cell proliferation and that further boosts this effect in the presence of osteogenic induction media. This is consistent with published results showing the osteoinductive effects of superparamagnetic NPs without the application of external magnetic fields (Wu, et al.,

2010; Wei, et al., 2011; Yun, et al., 2015). Although many of the studies reporting this effect do not provide possible mechanisms to explain this phenomenon, some have explored different hypotheses (Castro, et al., 2017; Zhu, et al., 2017). One of these hypotheses demonstrates that the influence of the superparamagnetic NPs within the scaffold at the nanoscale level act as a single magnetic domain, initiating micromotions.

This movement is translated by the cells as a trigger for the mechanotransduction pathway by affecting the ions channels (Gil, et al., 2014). Other studies demonstrate the effect of superparamagnetic NPs on cell proliferation by significantly decreasing the intracellular

H₂O₂ and peroxidase-like activity. They also found that NPs had an impact on cells cycle through lysosomal metabolism of iron oxide particles that led to iron depletion, which might lead to cell cycle block at G1/S, and it affects the expression of regulating molecules vital for the cell cycle process and apoptosis (Huang, et al., 2009). Furthermore, a very thorough and interesting study proposed that the composition of the protein corona that forms on the superparamagnetic NPs could provide the necessary stimulation to promote the levels of proliferation measured. The study characterized the protein composition of the corona after being exposed to fetal bovine serum alone or in the presence of the proteins secreted by pre-osteoblasts (Zhu, et al., 2017). Their findings showed the presence of proteins related to calcium ions, G-protein coupled receptors, and MAPK/ERK cascades as compared with scaffolds not containing NPs. All these could be related to helping in the induction of cell proliferation and could thus explain the increase in hASC proliferation in the presence of superparamagnetic scaffolds without pEMF stimulation. This hypothesis supports the proliferation results (Figure 7) since the groups that were cultured in the presence of osteogenic media and superparamagnetic scaffolds showed a high proliferative rate due to the formation of protein corona composed by the components of the osteogenic media allowing these to have a higher stability and longer half-life.

It has been shown that the force necessary to stimulate of a single ion channel is of the order of 1-2 pN (Yoshimura, et al., 2010). Other studies have demonstrated that a mechanical force less than 0.2 pN is required to activate the TREK-1 channel and of 2 pN

to break the bond between fibronectin and the cytoskeleton (Sapir-Lekhovitser, et al.,

2016). However, our calculations showed that the magnetic force on our systems was about

0.148 pN per cell and its influence was noticed for a group of magnetic scaffolds under pEMF during the first week of culture to have a significant degree (p<0.001) as compared to the control. Nevertheless, after 14 and 21 d the magnetic force was not strong enough to activate the mechanotransduction process due to the larger number of cells as shown by the group of magnetic scaffolds under pEMF that did not exhibit significant changes as compared to the control. [APPENDIX]

4.1.2 Differentiation to osteoblasts

Osteoblast differentiation from hASCs was detected measuring the activity of the

ALP biomarker after 7, 14, 21 d of culture (Wang, et al., 2007). As before, the two experimental groups were tested, pEMF stimulated and non-stimulated, which were subdivided into four different formulations, all based on hASC-seeded scaffolds, with/without osteogenic induction media, and with/without NPs.

ALP activity at day 7 showed signs of early differentiation of hASCs cells into osteoblasts (Figure 8). However, the cells cultured within the group of superparamagnetic scaffolds under pEMF stimulation (without osteogenic induction media) showed a statistically similar low level of ALP activity as the non-stimulated group and the controls

(i.e., with and without pEMF stimulation and both lacking NPs). However, when the hASCs in hydrogels (no NPs) were simultaneously stimulated with osteogenic induction media and pEMF, demonstrated signs of an early differentiation to osteoblasts with a

significant increase in ALP activity (p<0.05) when compared to the control. In contrast, a lack of pEMF stimulation showed low levels of ALP activity with no significant change when compared to the control. Under these conditions, the presence of superparamagnetic scaffolds shows high levels of ALP activity with significant difference (p<0.001) without pEMF stimulation and even higher with pEMF stimulation significantly higher than the control (p<0.0001).

Although after 14 d of culture, the levels of ALP activity had significantly increased in all groups, those using superparamagnetic scaffolds (without osteogenic induction media) with and without pEMF stimulation showed no significant differences when compared to the control. In the case of the group under pEMF stimulation (without NPs or osteogenic induction media) had a slight increase of ALP activity, it was not significant when compared to the control.

However, when cells were stimulated with osteogenic induction media (not pEMF) there was a significant increase in differentiation as measured by ALP activity (p<0.001), with the addition of pEMF stimulation showing a significant increase. As with the results at day 7, the presence of superparamagnetic scaffolds led to elevated levels of ALP activity with significant difference (p<0.001) without pEMF stimulation and even significantly higher with pEMF stimulation (p<0.0001).

At day 21, we still observe an increase in the ALP activity with time, and we noticed that the groups with osteogenic induction stimulation regardless of the presence of pEMF stimulation were significantly different from the control (p<0.0001). Similarly, the presence of superparamagnetic scaffolds, regardless of the presence of pEMF stimulation, were significantly different from the control (p<0.0001). Moreover, ALP activity has

significantly increased (p<0.05) for the first time for the superparamagnetic scaffolds group without osteogenic induction media stimulation, but with pEMF stimulation, while the other groups remain unchanged.

The results from ALP assay showed that the hASCs start to differentiate into the osteoblast phenotype since day 7 when grown in the superparamagnetic scaffold and cultured with osteogenic induction media. Also, early differentiation at day 7 was noticed in the group cultured within osteogenic media, with and without pEMF stimulation. With the ALP activity being higher with the latter. These effects were not seen in the presence of osteogenic induction media alone, but only after 14 d of stimulation. At this same time, it can also be seen that hASCs biochemically stimulated with induction media continue to increase their level of differentiation, with a non-significant increase due to the presence of pEMF stimulation and a non-significant difference due to the presence of NPs. After 2 weeks a trend starts to emerge which involves a significant level of osteogenic differentiation by hASCs stimulated with osteogenic induction media, with a non- significant increase due to pEMF stimulation and no effect due to the superparamagnetic scaffolds. We also noticed that ALP activity, an early osteogenic marker, did not show a significant difference between groups with/without magnetic fields after 21 days of culture.

This could be since ALP is not a good late osteogenic biomarker. Perhaps the use of a late osteogenic biomarker such as osteocalcin is a better choice (Granéli, et al., 2014), to show significant differences between groups.

If the proposed NP protein corona mechanism (Zhu, et al., 2017) is correct, then the significant difference in ALP activity see after 7 days in the presence of superparamagnetic scaffolds is due to osteogenic proteins adsorbed onto the NPs and

inducing hASC differentiation. During this initial stage, there is a sustained presence of osteogenic proteins which allows an early commitment of hASC to the osteogenic lineage.

After lineage commitment, there is less need for osteogenic proteins to continue the differentiation process (Ferroni, et al., 2018). So, the presence of the superparamagnetic scaffolds helps improve the levels of hASC osteogenic differentiation by allowing an early commitment of these cells to the osteogenic lineage.

Figure 8. Alkaline phosphatase activity. The differentiation of hASCs cells was assessed at 7, 14, 21 d. ALP values were normalized with the number of cells of each sample. *p <

0.05; **p < 0.001; ***p < 0.0001, indicate statistically significant differences between basal media group (control) and the other groups (one-way ANOVA followed by Dunnett test). The mean values are calculated from the average results of three samples, the results are represented as mean ± SD.

4.1.3 Mineralization

While ALP is relatively an early differentiation marker that increases during the proliferation and matrix synthesis stage (Harris, 1990), the matrix calcium deposition defines the terminal stage in osteoblast maturation (Cormier, 1995). Therefore, the series of osteogenic differentiation assays, involving ALP activity and staining, and the calcium mineralization, clearly demonstrate the significant role of the 3D superparamagnetic scaffolds played in accelerating osteoblastogenesis of hASCs. The calcium depositions were quantified for all the groups over three weeks (at day 7, 14, and 21) as shown in

Figure 9.

At day 7, results revealed that four groups did not exhibit any early calcium depositions, these included the hASC-seeded scaffolds, with pEMF stimulation, with osteogenic induction media stimulation, and NP-containing scaffolds without any stimulation. However, the group with NP-containing (superparamagnetic) scaffolds were stimulated with induction media showed some early calcium deposition. Also, although the remaining three groups, which were all pEMF-stimulated, showed some early calcium deposition, it was not significantly different from the superparamagnetic scaffold only stimulated with osteogenic induction media. These groups were superparamagnetic scaffolds with and without induction media and hydrogel (no NPs) with induction media, all three being pEMF-stimulated.

After 7 d, non-significant traces of calcium in the extracellular matrix is observed.

This is true for groups with pEMF stimulation or osteogenic media and no pEMF. After 14

d, even the hASC-seeded control (without stimulation) start to show traces of calcium in the extracellular matrix. Meanwhile, the groups of superparamagnetic scaffolds stimulated with induction media and with and without pEMF stimulation, showed higher levels of calcium deposition, but not significantly higher than the control. After 21 d, there was a significant increase (p<0.05) in calcium deposition, in the two groups with superparamagnetic scaffolds being pEMF-stimulated, with and without osteogenic induction media stimulation. The remaining groups deposited equivalent amounts of calcium to the control.

Groups with a low initial (day 7 and 14) ALP activity did not form lots of mineralized matrix later in day 21, especially the control group. However, mineralization measurements support the ALP activity results, whereas the samples that were cultured in superparamagnetic scaffolds and osteogenic induction media showed an early calcium deposition after 7 d. In fact, this effect can be present during the first 2 weeks. These are the groups with early ALP increase.

In addition, there is a clear and significant increase in mineralization after 21 d due to pEMF stimulation. As the results show, the hASCs have already differentiated and do not need further stimulation from the induction media or NPs and are depositing mineral stimulated solely by pEMFs. A group with osteogenic media and nanoparticles had high

ALP from the very initial culture (day 7) and showed non-significant mineralization on day

7 compared to the control. We believe that the cells in this group are differentiated very early and secreting calcium earlier than groups with pEMF stimulation, so the ECM calcium is comparable to the group with pEMF stimulation on day 21.

P<0.05 1.6E-07 MSC media (gel+cells)

) 1.4E-07 Msc media (gel Np+ cells)

1.2E-07 osteo media (gel+cells)

0.0000001 osteo media (gel Np+cells)

8E-08 MSC media (gel+cells)+MF 6E-08 Msc media (gel Np+ cells)+MF

4E-08 Calcium concentration mg/L (ppm mg/L concentration Calcium osteo media (gel+cells)+MF 2E-08 osteo media (gel

Normalized Normalized 0 Np+cells)+MF 7d 14d 21d -2E-08 Time (day)

Figure 9. Mineralization assay. Calcium concentrations were quantified after 7,14, 21 days. Error bar represents the SD. p < 0.05, indicate statistically significant differences between MSCs media gel+cells group (control) and the other groups (one-way ANOVA followed by Dunnett test). The mean values are calculated from the average results of three samples, the results are represented as mean ± SD.

4.1.4 Cell morphology

A qualitative test used to test cell morphology is through of actin and DAPI staining, as shown in Figure 10, at day 3 and 14. At day 3, after culturing the cells within the hydrogel, it got a spherical shape at an early stage especially when they were cultured with mesenchymal media either with and without exposing to magnetic fields (Figure 10;

Day 3; A, B, E, F). Also, at the same time point when we used an induction media, the cells have a spindle and elongated shape morphology, and this was noticed with or without applying magnetic fields which is the sign of early osteoblastic differentiation (Figure 10;

Day 3; C, D, G, H). Others have shown the same where the elongated cells were the sign of differentiation or specialization into osteoblasts after following adherence to the scaffold

(Kannarkat et al., 2010). At Day 14, the images showed that the cells were entirely embedded in the nanofiber hydrogel by taking the shape of the fibers (Figure 10; Day 14).

Figure 10. Phalloidin-labeled actin filaments stain (red) and DAPI stain (blue) for hASCs within hydrogel at 3 and 14 days. A) basal media, B) basal media + NPs, C) osteogenic media, D) osteogenic media + NPs, E) basal media + pEMF, F) basal media + NPs + pEMF,

G) osteogenic media + pEMF, H) osteogenic media + NPs + pEMF. Scale bars at day 3 and 14 are 100 µm (10X) and 50 µm (20X), respectively. Biochemical stimulation – osteogenic induction media.

The results on hASC morphology showed that the cells intimately interacted with the ECM since the scaffold is mainly composed of water, allowing the cells to freely

migrate and interact. Also, the presence of NPs or EMFs did not affect cell morphology.

During the early days of seeding hASCs adopted a spherical shape, but later they started to elongate and got spindle morphology. Fan, et al., mention that the hASCs need to adhere to a surface to allow them to stretch and to get its original shape to be able to contact with other cells, if not they will become apoptotic (Fan and Wang, 2017).

4.1.5 Alkaline phosphatase staining

To do a qualitative evaluation of hASCs osteoblast differentiation, ALP staining was used at day 14 for all eight groups as shown in Figure 11. The results demonstrate that at day 14, hASCs had been differentiated into osteoblasts. Whereas the groups that were cultured with and without NPs, and without pEMF or induction media stimulation showed only marginal ALP staining (Figure 11, A and B), when these were pEMF-stimulated, they showed high levels of ALP staining (Figure 11, E and F). In addition, the results showed a higher level of ALP staining when stimulated with osteogenic induction media, with or without pEMF stimulation (Figure 11, C, D, G, and H).

Figure 11. Alkaline phosphatase stain for hASCs cells seeded within hydrogel at day 14, scale bar = 200 µm (10X). A) basal media, B) basal media + NPs, C) osteogenic media, D) osteogenic media + NPs, E) basal media + pEMF, F) basal media + NPs + pEMF, G) osteogenic media + pEMF, H) osteogenic media + NPs + pEMF. Biochemical stimulation

– osteogenic induction media.

As expected, the ALP staining result was consistent with the quantitative ALP activity results. At day 14, biochemically-stimulated cells contained high levels of alkaline phosphatase, with a non-significant increase due to the presence of pEMF stimulation and a non-significant difference due to the presence of NPs.

4.1.6 FTIR analysis

FTIR analysis of the self-assembled peptides scaffolds with or without iron oxide nanoparticles showed that the presence of nanoparticles did not cause much difference in the spectrum from the scaffold without nanoparticles. As shown in Figure 12, the hydrogel with nanoparticles has shown three major peaks, located at (3625-3100), 2100, 1625 cm–1.

The spectrum showed that there is a broad –OH stretching band between 3625 and 3100 cm–1, which is mainly resulted from water crystal and characteristic of the O-H stretch band of the hydroxyl group. A medium peak is shown on 2100 cm–1 that represent the C≡C bond.

Also, the spectrum displays at 1625 cm–1 peak which is the characteristic of amide I bond that link the amino acids of the hydrogel which resulted from stretching vibrations of the

C=O bond of the amide, which represented the existence of the β-sheet structures of the hydrogel. Also, FTIR spectrum for hydrogel alone showed almost the same spectrum with a slight shift. However, there is a strong stretching band of –OH at 3400 cm–1, and two medium peaks 2400 and 2100 cm–1 to represent the C≡C bond. Also, another peak in 1620 to represent the amid I of stretching vibrations of the C=O bond.

Figure 12. FTIR analysis of self-assembled peptides with and without iron oxide nanoparticles.

4.2 Conclusion

In this study, we developed a superparamagnetic scaffold based on a biomimetic hydrogel for the 3D culture of hASCs to study their response to biochemical, electromagnetic and mechanical stimulation. This was done by evaluating proliferation, osteoblastic differentiation, ECM mineralization, and morphology under a combination of these forms of stimulation. The 3D superparamagnetic scaffold was based on peptides that self-assembled to form nanofibers and an ECM-type structure that allows nutrients and oxygen to be effectively transported to the seeded cells in a manner to the natural condition

(Zhang, 2004). The specific peptide used was 16-mer that consisted of four repeats of the

RADA amino acid sequence. Hydrogels made from this peptide have demonstrated the

capacity to enhance the proliferation and differentiation of primary osteoblasts in vitro

(Bokhari, et al., 2005) and in vivo (Misawa, et al., 2006). The choice of hASC over bone marrow-derived MSCs is based on their capacity to proliferate faster and retain and enhanced longer an enhanced capacity for differentiation over their bone marrow counterparts (Burrow, et al., 2017)

Our results show no negative or cytotoxic effects on hASCs, due to the presence of pEMFs or superparamagnetic NPs. Also, the result showed that there was an early differentiation by incorporating osteogenic media either with the presence of superparamagnetic NPs or pEMFs. The principal results of the present study revealed several novel findings regarding the events involved in the induction of the osteogenic differentiation of hASCs by pEMFs and mechanical stimulation. For instance, that adding superparamagnetic NPs to the hydrogel induced a significant increase in proliferation, but not due to mechanical stimulation due to NP vibrations induced by the pEMFs. We propose that this effect is due to the proteins adsorbed onto the NPs, which help induce cell proliferation. Although a positive effect due to mechanical stimulation was expected, perhaps the use of extremely low-frequency field did actuate a level of vibration in the superparamagnetic NPs to mechanically stimulate the hASCs (Golovin, et al., 2017). In addition, it was seen that after two weeks there was a significant level of osteogenic differentiation by hASCs stimulated with osteogenic induction media, with a non- significant increase due to pEMF stimulation and no effect due to the superparamagnetic scaffolds. Perhaps, prior to that due to a sustained presence of osteogenic proteins on the corona of the NPs, there was an early commitment of hASC to the osteogenic lineage.

Committed cells then became significantly more responsive to pEMF stimulation not only

promoting osteogenic differentiation, as evidenced by ALP activity and staining, the extent of ECM mineralization (Ferroni, et al., 2018).

References

Amini, A., Laurencin, C., & Nukavarapu, S. (2012). Bone tissue engineering: Recent advances and challenges. Critical Reviews in Biomedical Engineering, 40(5), 363- 408.

Al Nofal, A. A., Altayar, O., BenKhadra, K., Agha, O. Q., Asi, N., Nabhan, M., Prokop, L. J., Tebben, P., & Murad, M. H. (2015). Bone turnover markers in Paget’s disease of the bone: a systematic review and meta-analysis. Osteoporosis International, 26(7), 1875-1891.

Athanasiou, K., Zhu, C., Lanctot, D., Agrawal, C., & Wang, X. (2000). Fundamentals of biomechanics in tissue engineering of bone. Tissue Engineering, 6(4), 361-81.

Augat, P., & Schorlemmer, S. (2006). The role of cortical bone and its microstructure in bone strength. Age and Ageing, 35(2), Ii27-Ii31.

Bañobre-López, M., Piñeiro-Redondo, De Santis, Gloria, Ambrosio, Tampieri, A., Dediu, V., & Rivas, J. (2011). Poly(caprolactone) based magnetic scaffolds for bone tissue engineering. Journal of Applied Physics, 109(7), 07B313.

Bassett, C. (1987). Low energy pulsing electromagnetic fields modify biomedical processes. BioEssays, 6(1), 36-42.

Bitar, K., & Zakhem, E. (2014). Design strategies of biodegradable scaffolds for tissue regeneration. Biomedical Engineering and Computational Biology, 6, 13-20.

Bock, N., Riminucci, A., Dionigi, C., Russo, A., Tampieri, A., Landi, E., Goranov, V. A., Marcacci, M., & Dediu, V. (2010). A novel route in bone tissue engineering: Magnetic biomimetic scaffolds. Acta Biomaterialia, 6(3), 786-796.

Bokhari, M. A., Akay, G., Zhang, S., & Birch, M. A. (2005). The enhancement of osteoblast growth and differentiation in vitro on a peptide hydrogel—polyHIPE polymer hybrid material. Biomaterials, 26(25), 5198-5208.

Bruder, S. P., Jaiswal, N., & Haynesworth, S. E. (1997). Growth kinetics, self‐renewal, and the osteogenic potential of purified human mesenchymal stem cells during

extensive subcultivation and following cryopreservation. Journal of Cellular Biochemistry, 64(2), 278-294.

Brydone, A. S., Meek, D., & Maclaine, S. (2010). Bone grafting, orthopedic biomaterials, and the clinical need for bone engineering. Proceedings of the Institution of Mechanical Engineers, Part H: Journal of Engineering in Medicine, 224(12), 1329-1343.

Buck, D., & Dumanian, G. (2012). Bone biology and physiology: Part I. The fundamentals. Plastic and Reconstructive Surgery, 129(6), 1314-1320.

Buckwalter, J. A., Glimcher, M. J., Cooper, R. R., & Recker, R. (1995). Bone Biology. The Journal of Bone & Joint Surgery, 77(8), 1276-1289.

Burger, E. H., & Klein-Nulend, J. (1999). Mechanotransduction in bone—role of the lacuno-canalicular network. FASEB Journal, 13, S101–S112.

Burrow, K. L., Hoyland, J. A., & Richardson, S. M. (2017). Human adipose-derived stem cells exhibit enhanced proliferative capacity and retain multipotency longer than donor-matched bone marrow mesenchymal stem cells during expansion in vitro. Stem cells international, 2017, 2541275.

Calori, G.M., Donati, D., Di Bella, C., & Tagliabue, L. (2009). Bone morphogenetic proteins and tissue engineering: Future directions. Injury, 40, S67-S76.

Campagnoli, C., Roberts, I. A., Kumar, S., Bennett, P. R., Bellantuono, I., & Fisk, N. M. (2001). Identification of mesenchymal stem/progenitor cells in human first- trimester fetal blood, liver, and bone marrow. Blood, 98(8), 2396-2402.

Canalis, E. (2009). Growth factor control of bone mass. Journal of Cellular Biochemistry, 108(4), 769-777.

Caplan, M., Schwartzfarb, E., Zhang, S., Kamm, R., & Lauffenburger, D. (2002). Effects of systematic variation of amino acid sequence on the mechanical properties of a self-assembling, oligopeptide biomaterial. Journal of Biomaterials Science, Polymer Edition, 13(3), 225-236.

Caplan, A. I. (2005). Review: mesenchymal stem cells: cell-based reconstructive therapy in orthopedics. Tissue Engineering, 11 (7-8): 1198-1211.

Carpenter, D. O., & Aĭrapet︠ia︡ n, S. N. (Eds.). (1994). Biological effects of electric and magnetic fields: sources and mechanisms (Vol. 2). Academic Press.

Castro, P. S., Bertotti, M., Naves, A. F., Catalani, L. H., Cornejo, D. R., Bloisi, G. D., & Petri, D. F. (2017). Hybrid magnetic scaffolds: The role of scaffolds charge on the

cell proliferation and Ca2+ ions permeation. Colloids and Surfaces B: Biointerfaces, 156, 388-396.

Causa, F., Netti, P. A., & Ambrosio, L. (2007). A multi-functional scaffold for tissue regeneration: the need to engineer a tissue analogue. Biomaterials, 28(34), 5093- 5099.

Clarke, B. (2008). Normal bone anatomy and physiology. Clinical Journal of the American Society of Nephrology, 3(Suppl 3), S131-S139.

Colnot, C. (2011). Cell sources for bone tissue engineering: insights from basic science. Tissue Engineering Part B: Reviews, 17(6), 449-457.

Cormier, C. (1995). Markers of bone metabolism. Current Opinion in Rheumatology, 7, 243–248

Cormier, A., Pang, X., Zimmerman, M., Zhou, H., & Paravastu, A. (2013). Molecular structure of RADA16-I designer self-assembling peptide nanofibers. ACS Nano, 7(9), 7562-7572.

Cunha, C., Panseri, S., Villa, O., Silva, D., & Gelain, F. (2011). 3D culture of adult mouse neural stem cells within functionalized self-assembling peptide scaffolds. International Journal of Nanomedicine, 6, 943-955.

Daish, C., Blanchard, R., Fox, K., Pivonka, P., & Pirogova E. (2018). The Application of Pulsed Electromagnetic Fields (PEMFs) for Bone Fracture Repair: Past and Perspective Findings. Annals of Biomedical Engineering, 46, 525–542.

Datta, H., Ng, W., Walker, J., Tuck, S., & Varanasi, S. (2008). The cell biology of bone metabolism. Journal of Clinical Pathology, 61(5), 577-587.

Deng, X. L., Lau, C. P., Lai, K., Cheung, K. F., Lau, G. K., & Li, G. R. (2007). Cell cycle-dependent expression of potassium channels and cell proliferation in rat mesenchymal stem cells from bone marrow. Cell Proliferation, 40, 656–670.

Dhivya, S., Ajita, J., & Selvamurugan, N. (2015). Metallic Nanomaterials for Bone Tissue Engineering. Journal of Biomedical Nanotechnology, 11(10), 1675-700.

Dobson, J., Cartmell, S., Keramane, A., & Haj, A. (2006). Principles and design of a novel magnetic force mechanical conditioning bioreactor for tissue engineering, stem cell conditioning, and dynamic in vitro screening. NanoBioscience, IEEE Transactions on, 5(3), 173-177.

Downey, P. A., & Siegel, M. I. (2006). Bone biology and the clinical implications for osteoporosis. Physical Therapy, 86(1), 77-91

Edelman, E. R., Brown, L., Kost, J., Taylor, J., Langer, R. (1984). Modulated release from polymeric drug delivery systems using oscillating magnetic fields: in vitro and in vivo characteristics. Transaction of the American Society of Artificial and Internal Organs, 30, 445-449.

Elangovan, S., D'Mello, S. R., Hong, L., Ross, R. D., Allamargot, C., Dawson, D. V., Stanford, C. M., Johnson, G. K., Sumner, D. R., & Salem, A. K. (2014). The enhancement of bone regeneration by gene activated matrix encoding for platelet derived growth factor. Biomaterials, 35(2), 737-747.

Fan, C., & Wang, D. A. (2017). Macroporous Hydrogel scaffolds for three-dimensional cell culture and tissue engineering. Tissue Engineering Part B: Reviews, 23(5), 451-461.

Ferroni, L., Gardin, C., Dolkart, O., Salai, M., Barak, S., Piattelli, A., Amir-Barak, H., & Zavan, B. (2018). Pulsed electromagnetic fields increase osteogenetic commitment of MSCs via the mTOR pathway in TNF-α mediated inflammatory conditions: an in-vitro study. Scientific reports, 8(1), 5108.

Franz‐Odendaal, T., Hall, B., & Witten, P. (2006). Buried alive: How osteoblasts become osteocytes. Developmental Dynamics, 235(1), 176-190.

Funk, R., & Monsees, T. (2006). Effects of electromagnetic fields on cells: physiological and therapeutical approaches and molecular mechanisms of interaction. Cells Tissues Organs, 182(2), 59-78

Ganesan, K., Gengadharan, A. C., Balachandran, C., Manohar, B. M., & Puvanakrishnan, R. (2009). Low frequency pulsed electromagnetic field—a viable alternative therapy for arthritis. Indian Journal of Experimental Biology, 47(12), 939-948.

Garner, A. L, Chen, G., Chen, N., Sridhara, V., Kolb, J. F., Swanson, R. J., Beebe, S. J., Joshi, R. P., & Schoenbach, K. H. (2007). Ultrashort electric pulse induced changes in cellular dielectric properties. Biochemical et Biophysical Research Communications, 362, 139–144.

Gil, S., & Mano, J. F. (2014). Magnetic composite biomaterials for tissue engineering. Biomaterials Science, 2(6), 812-818.

Gloria, A., Russo, T., D'Amora, U., Zeppetelli, S., D'Alessandro, T., Sandri, M., Bañobre-López, M., Piñeiro-Redondo, Y., Uhlarz, M., Tampieri, A., Rivas, J., Herrmannsdörfer, T., Dediu, V. A., Ambrosio, L., & De Santis, R. (2013). Magnetic poly(ε-caprolactone)/iron-doped hydroxyapatite nanocomposite substrates for advanced bone tissue engineering. Journal of the Royal Society, Interface, 10(80), 20120833.

Golovin, Y. I., Gribanovsky, S. L., Golovin, D. Y., Zhigachev, A. O., Klyachko, N. L., Majouga, A. G., Master, А. M., Sokolsky, M., & Kabanov, A. V. (2017). The dynamics of magnetic nanoparticles exposed to non-heating alternating magnetic field in biochemical applications: theoretical study. Journal of Nanoparticle Research, 19:59.

Grant, D. N., Cozad, M. J., Grant, D. A., White, R. A., & Grant, S. A. (2015) In vitro electromagnetic stimulation to enhance cell proliferation in extracellular matrix constructs with and without metallic nanoparticles. Journal of Biomedical Materials Research Part B: Applied Biomaterials, 103(8), 1532-1540.

Granéli, C., Thorfve, A., Ruetschi, U., Brisby, H., Thomsen, P., Lindahl, A., & Karlsson, C. (2014). Novel markers of osteogenic and adipogenic differentiation of human bone marrow stromal cells identified using a quantitative proteomics approach. Stem Cell Research, 12(1), 153-165.

Muschler, G. F., Nakamoto, C., & Griffith, L. G. (2004). Engineering principles of clinical cell-based tissue engineering. Journal of Bone and Joint Surgery, 86(7), 1541-1558.

Grüttner, C., Müller, K., Teller, J., Westphal, F., Foreman, A., & Ivkov, R. (2007). Synthesis and antibody conjugation of magnetic nanoparticles with improved specific power absorption rates for alternating magnetic field cancer therapy. Journal of Magnetism and Magnetic Materials, 311(1), 181-186.

Gunatillake, P. A., & Adhikari, R. (2003). Biodegradable synthetic polymers for tissue engineering. European Cells & Materials, 5(1), 1-16.

Termaat, M. F., Den Boer, F. C., Bakker, F. C., Patka, P., & Haarman, H. J. (2005). Bone morphogenetic proteins. development and clinical efficacy in the treatment of fractures and bone defects. Journal of Bone and Joint Surgery, 87(6), 1367-1378.

Han, Y., Cowin, S. C., Schaffler, M. B., & Weinbaum, S. (2004b). Mechanotransduction and strain amplification in osteocyte cell processes. Proceedings of the National Acaddemy of Sciences USA, 101, 16689–16694.

Harris, H. (1990). The human alkaline phosphatases: what we know and what we don't know. Clinica chimica acta, 186(2), 133-150.

Hauser, C., & Zhang, S. (2010). Designer self-assembling peptide nanofiber biological materials. Chemical Society Reviews, 39(8), 2780-2790.

Heckman, J. D., Ingram, A. J., Loyd, R. D., Luck, J. V., & Jr, Mayer, P. W. (1981). Nonunion treatment with pulsed electromagnetic fields. Clinical Orthopaedics and Related Research, 161, 58-66.

Henstock, J., Rotherham, M., Rashidi, H., Shakesheff, K., & El Haj, A. (2014). Remotely activated mechanotransduction via magnetic nanoparticles promotes mineralization synergistically with bone morphogenetic protein 2: applications for injectable cell therapy. Stem Cells Translational Medicine, 3(11), 1363-1374.

Hollinger, J. O., & Kleinschmidt, J. C. (1990). The critical size defect as an experimental model to test bone repair materials. The Journal of craniofacial surgery, 1(1), 60- 68. Horii, A., Wang, X., Gelain, F., & Zhang, S. (2007). Biological designer self-assembling peptide nanofiber scaffolds significantly enhance osteoblast proliferation, differentiation and 3-D migration. PloS one, 2(2), e190.

Huang, C., & Ogawa, R. (2010). Mechanotransduction in bone repair and regeneration. FASEB Journal, 24(10), 3625-3632.

Huang, D. M., Hsiao, J. K., Chen, Y. C., Chien, L. Y., Yao, M., Chen, Y. K., Ko, B. S., Hsu, S. C., Tai, L. A., Cheng, H. Y., Wang, S. W., Yang, C. S., & Chen, Y. C. (2009). The promotion of human mesenchymal stem cell proliferation by superparamagnetic iron oxide nanoparticles. Biomaterials, 30(22), 3645-3651.

Hughes, S., El Haj, A. J., & Dobson, J. (2005). Magnetic micro- and nanoparticle- mediated activation of mechanosensitive ion channels. Medical Engineering and Physics, 27(9), 754-762.

Jayakumar, P., & Di Silvio, L. (2010). Osteoblasts in bone tissue engineering. Proceedings of the Institution of Mechanical Engineers. Part H, 224(12), 1415- 1440.

Jegal, S. H., Park, J. H., Kim, J. H., Kim, T. H., Shin, U. S., Kim, T. I., & Kim, H. W. (2011). Functional composite nanofibers of poly(lactide-co-caprolactone) containing gelatin-apatite bone mimetic precipitate for bone regeneration. Acta Biomaterialia, 7, 1609-1617.

Meng, J., Xiao, B., Zhang, Y., Liu, J., Xue, H., Lei, J., Kong, H., Huang, Y., Jin, Z., Gu, N., & Xu, H. (2013). Super-paramagnetic responsive nanofibrous scaffolds under static magnetic field enhance osteogenesis for bone repair in vivo. Scientific Reports, 3(1), 2655.

Dobson, J. (2008). Remote control of cellular behaviour with magnetic nanoparticles. Nature Nanotechnology, 3(3), 139.

Jun, Y., Seo, J., & Cheon, J. (2008). Nanoscaling laws of magnetic nanoparticles and their applicabilities in biomedical sciences. Accounts of Chemical Research, 41(2), 179-189.

Kanczler, J., Sura, H., Magnay, J., Green, D., Oreffo, R., Dobson, J., & El Haj, A. (2010). Controlled differentiation of human bone marrow stromal cells using magnetic nanoparticle technology. Tissue Engineering. Part A, 16(10), 3241-3250.

Kannarkat, J. T., Battogtokh, J., Philip, J., Wilson, O. C., & Mehl, P. M. (2010). Embedding of magnetic nanoparticles in polycaprolactone nanofiber scaffolds to facilitate bone healing and regeneration. Journal of Applied Physics, 107(9), 09B307.

Kantipuly, C., Katragadda, S., Chow, A., & Gesser, H. D. (1990). Chelating Polymers and Related Supports for Separation and Preconcentration of Trace Metals. Talanta, 37, 491-517.

Karp, J. M., Dalton, P. D., & Shoichet, M. S. (2003). Scaffolds for Tissue Engineering. MRS Bulletin, 28(04), 301-306.

Khojasteh, A., Behnia, H., Naghdi, N., Esmaeelinejad, M., Alikhassy, Z., & Stevens, M. (2013). Effects of different growth factors and carriers on bone regeneration: A systematic review. Oral Surgery, Oral Medicine, Oral Pathology and Oral Radiology, 116(6), E405-E423.

Kigami, R., Sato, S., Tsuchiya, N., Sato, N., Suzuki, D., Arai, Y., Ito, K., & Ogiso, B. (2014). Effect of basic fibroblast growth factor on angiogenesis and bone regeneration in non-critical-size bone defects in rat calvaria. Journal of Oral Science, 56(1), 17-22.

Kovacic, P., & Somanathan, R. (2010). Electromagnetic fields: Mechanism, cell signaling, other bioprocesses, toxicity, radicals, antioxidants and beneficial effects. Journal of Receptors and Signal Transduction, 30(4), 214-226.

Landis, W. (1995). The strength of a calcified tissue depends in part on the molecular structure and organization of its constituent mineral crystals in their organic matrix. Bone, 16(5), 533-544.

Lanyon, L. E. (1974). Experimental support for the trajectorial theory of bone structure. The Journal of Bone and Joint Surgery. British Volume, 56, 160–166.

Lanyon, L. E, & Baggott D. G. (1976). Mechanical function as an influence on the structure and form of bone. The Journal of Bone and Joint Surgery. British Volume, 58-B, 436–443.

Last, J., & Reiser, K. (1984). Collagen biosynthesis. Environmental Health Perspectives, 55, 169-177.

Lee, K., Silva, E. A., & Mooney, D. J. (2011). Growth factor delivery-based tissue engineering: general approaches and a review of recent developments. Journal of the Royal Society Interface, 8(55), 153-170.

Levi, B., & Longaker, M. T. (2011). Concise review: adipose‐derived stromal cells for skeletal regenerative medicine. Stem cells, 29(4), 576-582

Li, X., Wang, L., Fan, Y., Feng, Q., Cui, F. Z., & Watari, F. (2013). Nanostructured scaffolds for bone tissue engineering. Journal of Biomedical Materials Research Part A, 101(8), 2424-2435.

Lima, J., Gonçalves, A. I., Rodrigues, M. T., Reis, R. L., & Gomes, M. E.. (2015). The effect of magnetic stimulation on the osteogenic and chondrogenic differentiation of human stem cells derived from the adipose tissue (hASCs). Journal of Magnetism and Magnetic Materials, 393, 526-536.

Liu, J., Zhang, Y., Yang, T., Ge, Y., Zhang, S., Chen, Z., & Gu, N. (2009). Synthesis, characterization, and application of composite alginate microspheres with magnetic and fluorescent functionalities. Journal of Applied Polymer Science, 113(6), 4042-4051.

Liu, X., Wang, P., Chen, W., Weir, M. D., Bao, C., & Xu, H. H. (2014). Human embryonic stem cells and macroporous calcium phosphate construct for bone regeneration in cranial defects in rats. Acta Biomaterialia, 10(10), 4484-4493.

Liu, T. Y., Hu, S. H., Liu, T. Y., Liu, D. M., & Chen, S. Y. (2006). Magnetic-sensitive behavior of intelligent ferrogels for controlled release of drug. Langmuir, 22, 5974-5978

Logeart-Avramoglou, D., Anagnostou, F., Bizios, R., & Petite, H. (2005). Engineering bone: challenges and obstacles. Journal of Cellular and Molecular Medicine, 9 (1), 72-84.

Lohmann, C. H., Schwartz, Z., Liu, Y., Guerkov, H., Dean, D. D., Simon, B., & Boyan, B. D. (2000) Pulsed electromagnetic field stimulation of MG63 osteoblast-like cells affects differentiation and local factor production. Journal of Orthopedic Research 18, 637-646.

Luu, Y. K., Pessin, J. E., Judex, S., Rubin, J., & Rubin, C. T. (2009). Mechanical signals as a non-invasive means to influence mesenchymal stem cell fate, promoting bone and suppressing the fat phenotype. IBMS BoneKEy, 6(4), 132-149.

Macotela Y., Emanuelli B., Mori M.A., Gesta S., Schulz T.J., Tseng Y.H., Kahn C.R. (2012) Intrinsic differences in adipocyte precursor cells from different white fat depots. Diabetes 61, 1691–1699.

Markov, M. S. (2007) Pulsed electromagnetic field therapy history, state of the art and future. Environmentalist, 27, 465-475.

Marks, S., & Popoff, S. (1988). Bone cell biology: The regulation of development, structure, and function in the skeleton. American Journal of Anatomy, 183(1), 1- 44.

Marolt, D., Campos, I. M., Bhumiratana, S., Koren, A., Petridis, P., Zhang, G., Spitalnik, P. F., Grayson, W. L., & Vunjak-Novakovic, G. (2012). Engineering bone tissue from human embryonic stem cells. Proceedings of the National Academy of Sciences, 109(22), 8705-8709.

Mauney, J. R., Sjostorm, S., Blumberg, J., Horan, R., O'Leary, J. P., Vunjak-Novakovic, G., Volloch, V., & Kaplan, D. L. (2004). Mechanical stimulation promotes osteogenic differentiation of human bone marrow stromal cells on 3-D partially demineralized bone scaffolds in vitro. Calcified Tissue International, 74(5), 458- 468.

McKibbin B. The biology of fracture healing in long bones. (1978) The Journal of Bone and Joint Surgery. British Volume 60-B, 150–162.

Meng, J., Zhang, Y., Qi, X., Kong, H., Wang, C., Xu, Z., Xie, S., Gu, N., & Xu, H. (2010). Paramagnetic nanofibrous composite films enhance the osteogenic responses of pre-osteoblast cells. Nanoscale, 2(12), 2565-2569.

Misawa, H., Kobayashi, N., Soto-Gutierrez, A., Chen, Y., Yoshida, A., Rivas-Carrillo, J. D., Navarro-Alvarez, N., Tanaka, K., Miki, A., Takei, J., Ueda, T., Tanaka, M., Endo, H., Tanaka, N., & Ozaki, T. (2006). PuraMatrix facilitates bone regeneration in bone defects of calvaria in mice. Cell transplantation, 15(10), 903-910.

Mizuno, H. (2009). Adipose-derived stem cells for tissue repair and regeneration: ten years of research and a literature review. Journal of Nippon Medical School, 76(2), 56-66.

Morgan, E. F., Gleason, R. E., Hayward, L. N. M., Leong, P. L., & Salisbury Palomares, K. T. (2008). Mechanotransduction and fracture repair. The Journal of Bone & Joint Surgery, 90(Suppl 1): 25–30.

Myster, D. L., & Duronio, R. J. (2000). Cell cycle: To differentiate or not to differentiate?. Current Biology, 10(8), R302-R304.

Nagai, Y., Unsworth, L. D., Koutsopoulos, S., & Zhang, S. (2006). Slow release of molecules in self-assembling peptide nanofiber scaffold. Journal of Controlled Release, 115(1), 18-25.

Nagatomi, J., Arulanandam, B. P., Metzger, D. W., Meunier, A., & Bizios, R. (2003). Cyclic pressure affects osteoblast functions pertinent to osteogenesis. Annals of biomedical engineering, 31(8), 917-923.

Nauth, A., Ristevski, B., Li, R., & Schemitsch, E. H. (2011). Growth factors and bone regeneration: How much bone can we expect? Injury, 42(6), 574-579.

Neves, M. I., Wechsler, M. E., Gomes, M. E., Reis, R. L., Granja, P. L., & Peppas, N. A. (2017). Molecularly imprinted intelligent scaffolds for tissue engineering applications. Tissue Engineering Part B: Reviews, 23(1), 27-43.

Orr, A. W., Helmke, B. P., Blackman, B. R., & Schwartz, M. A. (2006). Mechanisms of mechanotransduction. Developmental Cell, 10(1), 11–20.

O’Toole, G. C., & Boland, P. (2006). Metastatic bone cancer pain: etiology and treatment options. Current pain and headache reports, 10(4), 288-292.

Panagopoulos, D. J., Messini, N., Karabarbounis, A., Philippetis, A. L., & Margaritis, L. H. (2000). A Mechanism for Action of Oscillating Electric Fields on Cells. Biochemical and Biophysical Research Communications, 272(3), 634-640.

Panagopoulos, D. J., Karabarbounis, A., & Margaritis, L. H. (2002). Mechanism for action of electromagnetic fields on cells. Biochemical and Biophysical Research Communications, 298(1), 95-102.

Panagopoulos, D. J., Johansson, O., & Carlo, G. L. (2015). Polarization: A Key Difference between Man-made and Natural Electromagnetic Fields, in regard to Biological Activity. Scientific Reports, 5(1), 14914.

Panek, M., Marijanović, I., & Ivković, A. (2015). Stem cells in bone regeneration. Periodicum biologorum, 117(1), 177-184.

Panseri, S., Cunha, C., D'Alessandro, T., Sandri, M., Russo, A., Giavaresi, G., Marcacci, M., Hung, C. T., & Tampieri, A. (2012). Magnetic Hydroxyapatite Bone Substitutes to Enhance Tissue Regeneration: Evaluation In Vitro Using Osteoblast-Like Cells and In Vivo in a Bone Defect. PLoS ONE, 7(6), E38710.

Panseri, S., Russo, A., Sartori, M., Giavaresi, G., Sandri, M., Fini, M., Maltarello, M. C., Shelyakova, T., Ortolani, A., Visani, A., Dediu, V., Tampieri, A., & Marcacci, M. (2013). Modifying bone scaffold architecture in vivo with permanent magnets to facilitate fixation of magnetic scaffolds. Bone, 56(2), 432-439.

Parfitt, A. M. (1994). Osteonal and hemi-osteonal remodeling: the spatial and temporal framework for signal traffic in adult human bone. Journal of Cellular Biochemistry, 55, 273-286.

Parikh, S. N. (2002). Bone graft substitutes: past, present, future. Journal of Postgraduate Medicine, 48(2), 142-148.

Pavalko, F., Chen, N., Turner, C., & Burr, D. (1998). Fluid shear-induced mechanical signaling in MC3T3-E1 osteoblasts requires cytoskeleton-integrin interactions. American Journal of Physiology, 44(6), C1591-C1601.

Pavalko, F. M., Norvell, S. M., Burr, D. B., Turner, C. H., Duncan, R. L., & Bidwell, J. P. (2003). A model for mechanotransduction in bone cells: The load‐bearing mechanosomes. Journal of Cellular Biochemistry, 88(1), 104-112.

Pilla, A. (2002). Low-intensity electromagnetic and mechanical modulation of bone growth and repair: Are they equivalent? Journal of Orthopaedic Science, 7(3), 420-428.

Polo-Corrales, L., Latorre-Esteves, M., & Ramirez-Vick, J. E. (2014). Scaffold design for bone regeneration. Journal of Nanoscience and Nanotechnology 14, 15-56.

Polo-Corrales, L., Ramirez-Vick, J., & Feria-Diaz, J. J. (2018). Recent Advances in Biophysical Stimulation of MSC for Bone Regeneration. Indian Journal of Science and Technology 11, 1-41.

Rauch, F., & Glorieux, F. H. (2004). Osteogenesis imperfecta. The Lancet, 363(9418), 1377-1385.

Rauh, J., Milan, F., Guenther, K. P., & Stiehler, M. (2011). Bioreactor systems for bone tissue engineering. Tissue Engineering Part B: Reviews, 17(4), 263-280.

Rho, J. Y., Kuhn-Spearing, L., & Zioupos, P. (1998). Mechanical properties and the hierarchical structure of bone. Medical Engineering and Physics, 20(2), 92-102.

Rogel, M., Qiu, H., & Ameer, G. (2008). The role of nanocomposites in bone regeneration. Journal of Materials Chemistry, 18(36), 4233-4241.

Rose, F. R., & Oreffo, R. O. (2002). Bone tissue engineering: hope vs hype. Biochemical and biophysical research communications, 292(1), 1-7.

Ross, C. L., Siriwardane, M., Almeida-Porada, G., Porada, C. D., Brink, P., Christ, G. J., & Harrison, B. S. (2015). The effect of low-frequency electromagnetic field on human bone marrow stem/progenitor cell differentiation. Stem Cell Research, 15, 96-108.

Rubin, J., Rubin, C., & Jacobs, C. R. (2006). Molecular pathways mediating mechanical signaling in bone. Gene, 367, 1-16.

Rubio Ayala, M., Syrovets, T., Hafner, S., Zablotskii, V., Dejneka, A., & Simmet, T. (2018). Spatiotemporal magnetic fields enhance cytosolic Ca2+ levels and induce actin polymerization via activation of voltage-gated sodium channels in skeletal muscle cells. Biomaterials, 163, 174-184.

Russo, A., Bianchi, M., Sartori, M., Parrilli, A., Panseri, S., Ortolani, M., Sandri, M., Boi, M., Salter, D. M., Maltarello, M. C., Giavaresi, G., Fini, M., Dediu, V., Tampieri, A., & Marcacci, M. (2016). Magnetic forces and magnetized biomaterials provide dynamic flux information during bone regeneration. Journal of Materials Science: Materials in Medicine, 27(3), 1-13.

Salgado, A. J. and O. P. Coutinho, & Reis, R. L. (2004). Bone tissue engineering: state of the art and future trends. Macromolecular Bioscience, 4(8): 743-765.

Samba Sivudu, K., & Rhee, K. Y. (2009). Preparation and characterization of pH- responsive hydrogel magnetite nanocomposite. Colloids and Surfaces A: Physicochemical and Engineering Aspects, 349, 29-34.

Sapir-Lekhovitser, Y., Rotenberg, M., Jopp, J., Friedman, G., Polyak, B., & Cohen, S. (2016). Magnetically actuated tissue engineered scaffold: Insights into mechanism of physical stimulation. Nanoscale, 8(6), 3386-3399.

Sasaki, N., Matsushima, N., Ikawa, T., Yamamura, H., & Fukuda, A. (1989). Orientation of bone mineral and its role in the anisotropic mechanical properties of bone— Transverse anisotropy. Journal of Biomechanics, 22(2), 157-164.

Schmidt, C., Pommerenke, H., Dürr, F., Nebe, B., & Rychly, J. (1998). Mechanical stressing of integrin receptors induces enhanced tyrosine phosphorylation of cytoskeletally anchored proteins. The Journal of Biological Chemistry, 273(9), 5081-5085.

Sela, J., & Bab, I. (2012). Principles of bone regeneration. New York: Springer.

Semino, C. E. (2008). Self-assembling peptides: from bio-inspired materials to bone regeneration. Journal of dental research, 87(7), 606-616.

Sharir, A., Barak, M.M., & Shahar, R. (2008). Whole bone mechanics and mechanical testing. Veterinary Journal, 177, 8-17.

Stevens, M. M. (2008). Biomaterials for bone tissue engineering. Materials Today, 11(5), 18-25.

Taichman, R. (2005). Blood and bone: Two tissues whose fates are intertwined to create the hematopoietic stem-cell niche. Blood,105(7), 2631-2639.

Tchkonia T., Tchoukalova Y.D., Giorgadze N., Pirtskhalava T., Karagiannides I., Forse R.A., Koo A., Stevenson M., Chinnappan D., & Cartwright A. (2005). Abundance of two human preadipocyte subtypes with distinct capacities for replication, adipogenesis, and apoptosis varies among fat depots. American Journal of Physiology, Endocrinology and Metabolism 288, E267–E277.

Tchkonia T., Giorgadze N., Pirtskhalava T., Thomou T., DePonte M., Koo A., Forse R.A., Chinnappan D., Martin-Ruiz C., von Zglinicki T., Kirkland J.L. (2006). Fat depot-specific characteristics are retained in strains derived from single human preadipocytes. Diabetes 55, 2571–2578.

Teng, S., Liu, C., Krettek, C., & Jagodzinski, M. (2014). The application of induced pluripotent stem cells for bone regeneration: Current progress and prospects. Tissue Engineering. Part B, Reviews, 20(4), 328-339.

Velasco, M. A., Narváez-Tovar, C. A., & Garzón-Alvarado, D. A. (2015). Design, materials, and mechanobiology of biodegradable scaffolds for bone tissue engineering. BioMed research international, 2015, 729076.

Vo, T. N., Kasper, F. K., & Mikos, A. G. (2012). Strategies for controlled delivery of growth factors and cells for bone regeneration. Advanced Drug Delivery Reviews, 64(12), 1292-1309.

Voog, J., & Jones, D. L. (2010). Stem cells and the niche: a dynamic duo. Cell Stem Cell 6, 103-115.

Walleczek, J. (1991). Electromagnetic Field Effects on Cells of the Immune System: The Role of Calcium Signalling.

Wan, D. C., Shi, Y. Y., Nacamuli, R. P., Quarto, N., Lyons, K. M., & Longaker, M. T. (2006). Osteogenic differentiation of mouse adipose-derived adult stromal cells requires retinoic acid and bone morphogenetic protein receptor type IB signaling. Proceedings of the National Academy of Sciences, 103(33), 12335-12340.

Wang, H., Li, Y., Zuo, Y., Li, J., Ma, S., & Cheng, L. (2007). Biocompatibility and osteogenesis of biomimetic nano-hydroxyapatite/polyamide composite scaffolds for bone tissue engineering. Biomaterials, 28(22), 3338-3348.

Wang, X., Horii, A., & Zhang, S. (2008). Designer functionalized self-assembling peptide nanofiber scaffolds for growth, migration, and tubulogenesis of human umbilical vein endothelial cells. Soft Matter, 4(12), 2388-2395.

Wei, Y., Zhang, X., Song, Y., Han, B., Hu, X., Wang, X., Lin, Y., & Deng, X. (2011). Magnetic biodegradable Fe3O4/CS/PVA nanofibrous membranes for bone regeneration. Biomedical Materials, 6(5), 055008.

Weiner, S., & Wagner, H. (1998). THE MATERIAL BONE: Structure-Mechanical Function Relations. Annual Review of Materials Science, 28(1), 271-298.

Woo, S. L., Kuei, S. C., Amiel, D., Gomez, M. A., Hayes, W. C., White, F. C., & Akeson, W. H. (1981) The effect of prolonged physical training on the properties of long bone: a study of Wolff’s Law. The Journal of Bone and Joint Surgery. American Volume 63, 780–787.

Wozney, J. M., Rosen, V., Celeste, A. J., Mitsock, L. M., Whitters, M. J., Kriz, R. W., Hewick, R. M., & Wang, E. A. (1988) Novel regulators of bone formation: molecular clones and activities. Science 242, 1528–1534.

Wu, Y., Jiang, W., Wen, X., He, B., Zeng, X., Wang, G., & Gu, Z. (2010). A novel calcium phosphate ceramic–magnetic nanoparticle composite as a potential bone substitute. Biomedical Materials, 5(1), 015001.

Xu, H. Y., & Gu, N. (2014). Magnetic responsive scaffolds and magnetic fields in bone repair and regeneration. Frontiers of Materials Science, 8(1), 20-31.

Yokoi, H., Kinoshita, T., & Zhang, S. (2005). Dynamic reassembly of peptide RADA16 nanofiber scaffold. Proceedings of the National Academy of Sciences of the United States, 102(24), 8414-8419.

Yoshimura, K., & Sokabe, M. (2010). Mechanosensitivity of ion channels based on protein–lipid interactions. Journal of The Royal Society Interface, 7(Suppl 3), S307-S320.

Yousefi, A. M., James, P. F., Akbarzadeh, R., Subramanian, A., Flavin, C., & Oudadesse, H. (2016). Prospect of stem cells in bone tissue engineering: a review. Stem cells international, 2016, 6180487.

Yun, H. M., Lee, E. S., Kim, M. J., Kim, J. J., Lee, J. H., Lee, H. H., Park, K. R., Yi, J. K., Kim, H. W., & Kim, E. C. (2015). Magnetic Nanocomposite Scaffold-Induced Stimulation of Migration and Odontogenesis of Human Dental Pulp Cells through Integrin Signaling Pathways. PLoS One.10, e0138614

Zeng, X., Hu, H., Xie, L., Lan, F., Jiang, W., Wu, Y., & Gu, Z. (2012). Magnetic responsive hydroxyapatite composite scaffolds construction for bone defect reparation. International Journal of Nanomedicine, 7, 3365-3378

Zhao, X., & Zhang, S. (2006). Molecular designer self-assembling peptides. Chemical Society Reviews, 35(11), 1105-1110.

Zhang, S., Holmes, T., Lockshin, C., & Rich, A. (1993). Spontaneous assembly of a self- complementary oligopeptide to form a stable macroscopic membrane. Proceedings of the National Academy of Sciences, 90(8), 3334-3338.

Zhang, S., Lockshin, C., Cook, R., & Rich, A. (1994). Unusually stable beta-sheet formation in an ionic self-complementary oligopeptide. Biopolymers, 34(5), 663- 672.

Zhang, S., Holmes, T. C., DiPersio, C. M., Hynes, R. O., Su, X., & Rich, A. (1995). Self- complementary oligopeptide matrices support mammalian cell attachment. Biomaterials, 16(18), 1385-1393

Zhang, S. (2004). Beyond the Petri dish. Nature biotechnology, 22(2), 151-152.

Zhang, S. (2002). Emerging biological materials through molecular self- assembly. Biotechnology Advances, 20(5/6), 321-339.

Zhao, X., & Zhang, S. (2006). Molecular designer self-assembling peptides. Chemical Society Reviews 35, 1105-1110.

Zhao, X., Kim, J., Cezar, C. A., Huebsch, N., Lee, K., Bouhadir, K., Mooney, D. J. (2011). Active scaffolds for on-demand drug and cell delivery. Proceeding of the National Academy of Sciences U S A 108, 67-72.

Zhu, Y., Yang, Q., Yang, M., Zhan, X., Lan, F., He, J., Gu, Z., & Wu, Y. (2017). Protein Corona of Magnetic Hydroxyapatite Scaffold Improves Cell Proliferation via Activation of Mitogen-Activated Protein Kinase Signaling Pathway. ACS Nano.11:3690-3704.

Zimmerman, D., Jin, F., Leboy, P., Hardy, S., & Damsky, C. (2000). Impaired Bone Formation in Transgenic Mice Resulting from Altered Integrin Function in Osteoblasts. Developmental Biology, 220(1), 2-15

Zioupos, P., Currey, J., & Hamer, A. (1999). The role of collagen in the declining mechanical properties of aging human cortical bone. Journal of Biomedical Materials Research, 45(2), 108-16.

Appendix I

To estimate the magnetic force generated on each cell to activate the mechanotransduction process we decided to calculate the total force and divide it by a number of cells in each well plate. The magnetic force (F) acting on magnetic NPs inside a magnetic field which is defined as:

푉∆휒 퐹 = (퐵. 훻)퐵 µₒ where 푉 is the NP volume (in m3), ∆χ is the difference in magnetic susceptibilities between the NPs and the surrounding medium (dimensionless), µₒ is the permeability of vacuum, which is a constant equal to 4π×10-7 T·m/A, B is the applied magnetic field (in T), (퐵. 훻) is the gradient of the magnetic field (in T/m).

The magnitude of magnetic flux density (B) generated from the Helmholtz coils was measured from coil featured and the current intensity, can be measured according to the following formula:

3 5 2 푁 퐵 = ( ) 휇 퐼 4 0 푅 where N is the number of turns in each coil (N = 124), R is mean coil radius (R = 150 mm),

I is the electric current passing through the Helmholtz coil to generate magnetic fields (I =

1.345 A); thus, the magnetic field intensity is

3 5 124 B = ( )2 4휋 × 10−7 × 1.345 × = 1 mT = 1 × 10−3 T 4 0.15

The expression for the gradient of the magnetic field at the center of coils carrying current in the reverse direction, this gradient is given by:

5 ⅆ퐵 3 4 2 푁휇0퐼 (퐵. ∇) = | = ( ) ⋅ 2 ⅆ푥 푥=0 2 5 푅 where N is the number of turns in each coil (N = 124), R is mean coil radius (R = 150 mm),

I is the current (I = 1.345 A).

5 −7 ⅆ퐵 3 4 2 124×4휋×10 × 1.345 (퐵∇) = | = ( ) 2 = 0.007998 T/m ⅆ푥 푥=0 2 5 (0.15)

To measure superparamagnetic iron oxide NP volume (V), depending on the manufacturer information, each nanoparticle has an average diameter of d = 10 nm. So,

4 4 V = 휋푟3 = 휋(5 × 10−9)3 m3/NP 3 3

To quantify the difference in magnetic susceptibilities ∆χ we need to have the NPs and the surrounding medium (dimensionless) susceptibilities. The susceptibility of NPs (is dependent on the frequency of the magnetic field) χNP = 0.115 (Grüttner, et al., 2007), and susceptibility of media χ0 = 0.

Thus, according to the previously calculated information, the magnetic force generated from a single NP is given by

4휋 −9 3 −3 푉∆푥 (5×10 ) ×0.115×0.007998×10 퐹 = (퐵훻)퐵 = 3 µₒ 4휋×10−7

F = 37.36× 10−25 N/NP = 37.36 × 10−13 pN/NP

To estimate the number of NP per well, we will use the manufacturer information about

NP density (1 g/ml), NP concentration (5 mg/mL). The NPs concertation added during magnetic scaffold synthesis was 60 µg/ml, so the NP mass per well is calculated as shown below:

60×0.5 Mass of NPs in each well = = 3 × 10−3 g/well 100×103

mass 3×10−3 푔 The total volume of NPs in each well = = = 3 × 10−3 ml = ⅆ푒푛푠푖푡푦 1 푔/푚푙

3 × 10−9 푚ᶟ/well

Thus, the

−9 3×10 푚ᶟ 15 Total number of NPs per well = 4 = 6 × 10 NPs/well 휋(5×10−9 )3 푚ᶟ 3

Using the number of cells seeded at day 0 as 15×104 cells, and from the total number of

NPs per well, we can estimate the total magnetic force generated from the NPs around each cell considering that NPs are not affecting other cells:

total number of NPs per well 6×1015 Number of NPs around each cell = = = 푡표푡푎푙 푛푢푚푏푒푟 표푓 푐푒푙푙푠 푝푒푟 푤푒푙푙 15×104

0.4 × 1011 NPs/cell

Therefore, the total force generated from the NPs around each cell will be

F = 0.4 × 1011 × 37.36 × 10−13 pN/cell = 0.148 pN/cell

This is a conservative calculation simply because we cannot calculate the exact resultant force vector on each cell.