Designing Ionic-Complementary

DESIGNING IONIC-COMPLEMENTARY

HYDROGELS FOR BONE TISSUE REPAIR

A thesis submitted to the University of Manchester for the degree of Doctor of Philosophy (PhD) in the Faculty of Chemical Engineering and Analytical Science

2014

MSc. LUIS ALBERTO CASTILLO DIAZ

School of Chemical Engineering and Analytical Science

CONTENTS

List of figures………………………………………………………………...... 4

List of abbreviations…………………………………………………………….……..5

Declaration…………………………………………………………………………….8

Acknowledgements…………………………………………………………………..10

Thesis Structure……………………………………………………………….…..….11

Abstract………………………………………………………………………………12

Objectives of the thesis………………………………………………………………13

1 Chapter 1 - Introduction. Versatile peptide hydrogels for bone and dental tissue regeneration………………….…………………….………………………..…….….14

2 Chapter 2- Human osteoblasts within soft peptide hydrogels promote mineralisation in vitro…………………………………………………………………….….70

3 Chapter 3 - Functional peptide hydrogels for bone formation applications………..71

4 Chapter 4 - Osteogenic differentiation of human mesenchymal stem cells promotes mineralisation within an octa-peptide hydrogel……………………...…....99

5 Chapter 5- Materials and methods……………………….………………………129

5.1 Materials…………………………………………………………………….…..129

5.2 Methods…………………………………………………………………………130

5.3 References………………………………………………………………………139

6 Chapter 6 – Conclusions, outlook and recommendations for future work………..140

6.1 Conclusions……………..……………………………………………………....140

6.2 Outlook………………………………………………………………………….141

6.3 Recommendations for future work……………………………………………...141

Appendix……………………………………………………………………………143

LIST OF FIGURES

Figure 1. Representative of a peptide hydrogel containing cell culture media. Gel has been plated into a 12 well insert of 15 mm of inner diameter and 3 µm of pore diameter……………………………………………………………………………..132

Figure 2. Representative of the flow of media in between the cell culture insert and the well plate to aid the gelation of the FEFEFKFK gel. The media flows from the bottom to the top of the gel………………………………………………………....132

Figure 3. Oscillatory rheometer……………………………………………………135

LIST OF ABBREVIATIONS

2-D two-dimensional 3-D three-dimensional AB alveolar bone ALP or alkphos alkaline phosphatase AP-1 activator protein-1 ATR-FTIR attenuated total reflectance-Fourier transform infrared spectroscopy BGs bioactive glasses bFGF basic fibroblast growth factor BMP’s bone morphogenetic proteins BMUs basic multicellular units BSA bovine serum albumin BSP bone sialoprotein CD circular dichroism CDs clusters of differentiation COL-I1 type I collagen or collagen I COX-2 cyclooxygenase-2 CPC cetylpyridinium chloride DCs dendritic cells ddH2O double distilled water DMEM Dulbecco’s modified Eagle’s medium DPBS Dulbecco’s phosphate-buffered saline DPSCs dental pulp stem cells ECM extracellular matrix EDTA ethylenediaminetetraacetic acid EGF epidermal growth factor EthD-1 ethidium homodimer FBS fetal bovine serum FN fibronectin FSH follicle-stimulating hormone GAGs glycosaminoglycans GCs glucocorticoids GMSCs gingival mesenchymal stem cells GPa gigapascal HA hyaluronic acid HA hydroxyapatite HCA hydroxycarbonate apatite hMSCs human mesenchymal stem cells HOBs or hOBs human osteoblasts HPLC high-performance liquid chromatography HSCs hematopoietic stem cells ICC immunocytochemistry IEC ion-exchange chromatography IGF-1 insulin growth factor-1 IFN-γ interferon gamma IFR infrared spectroscopy

IL interleukin MCGS mesenchymal cell growth supplement MCP-1 monocyte chemoattractant protein-1 M-CSF macrophage colony-stimulator factor MIP-1α macrophage inflammatory protein-1 alpha MMPs metalloproteinases of matrix MPa megapascal MSCs mesenchymal stem cells NF-κB nuclear factor-κB NKs natural killer cells NMR nuclear magnetic resonance NOD nucleotide-binding oligomerisation domain OBs osteoblasts OCN osteocalcin OCs osteoclasts ODP osteopontin-derived peptide OPF oligo poly (ethylene glycol) fumarate OPG osteoprotegerin OPN osteopontin ONN osteonectin pAb primary antibody PBS phosphatase-buffered saline PCL poly (ε-caprolactone) PDFG platelet-derived growth factor PDL periodontal ligament PDLSCs periodontal ligament stem cells PEG polyethylene glycol PEG-2 prostaglandin-2 PEGDMA polyethylene glycol dimethacrylate PEO poly (ethylene oxide) PFA paraformaldehyde PGA polyglicolic acid PLA polylactic acid PLLA poly-L-lactide acid PLGA poly (lactic-coglycolide) PMNs polymorphonuclear cells PNIPAAm poly (N-iso-propylacrylamide) pNPP p-Nitrophenyl phosphate PPF poly (propylene fumarate) PTH parathormone PVA poly (vinyl alcohol) RANK receptor activator of NF-κB RANKL receptor activator of NF-κB ligand ROS reactive oxygen species

RT room temperature sAb secondary antibody SAXS small angle X-ray scattering SEC size exclusion chromatography SPPS solid phase peptide synthesis TCP tissue culture plastic TEOS tetraethylorthosilicate TEM transmission electron microscopy TFA trifluoroacetic acid TGF-β transforming factor-beta Ths helper T lymphocytes TIMPS tissue inhibitors of metalloproteinase TLRs toll-like receptors TNF-α tumor necrosis factor-alpha TRAP tartrate-resistant acid phosphatase Tregs regulator T cells VCAM-1 vascular cell adhesion molecule-1 VEGF vascular endothelial growth factor

DECLARATION

The University of Manchester PhD by published work candidate declaration

Candidate name: Luis Alberto Castillo Díaz

Faculty: Chemical Engineering and Analytical Science

Thesis title: Designing ionic-complementary hydrogels for bone tissue repair

Declaration to be completed by the candidate:

I declare that no portion of this work referred to this thesis has been submitted in support of an application for another degree or qualification of this or any other university or other institute of learning.

Signed: Date:

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1This excludes material already printed in academic journals, for which the copyright belongs to said journal and publisher. Pages for which the author does not own the copyright are numbered differently from the rest of the thesis.

ACKNOWLEDGEMENTS

First and foremost I would like to thank my supervisor Prof. Aline Fiona Miller, for giving me the opportunity to work on this fascinating project, and for her generous and unconditional support. Another thanks goes to my co-supervisor, Prof. Julie Elizabeth Gough and to Dr Alberto Saiani for their support and the opportunity to work in an extraordinary interdisciplinary network.

I am also proud and grateful to have been part of the Polymers and Peptides, and the Tissue Engineering research groups, where I had the opportunity to work in a great atmosphere meeting extraordinary people. I would like to thank my colleagues, Dr Ayeesha Mujeeb, Dr Claire Tang, Dr Jean Baptiste-Guilbaud, Dr Jonathan Gibbons, Dr Louise Carney, Dr Mohamed Elsawy, Dr Mi Zhou, Dr Kate Alexandra Meade, Dr Richard Balint, Dr Stephen Boothroyd, Dr Natasha Bhuiyan, Simon Wan, Shirley Bentley, Dr Nabilah Russlan, Dr Laura Szkolar, Dr Israa Sabree, Dr Deepak Kumar, Chistopher Hickling, and Dr Dave Roberts for their comradeship and kindly support.

A special thanks to CONACyT-Mexico, for the generous fellowship support.

Last but not least, another thanks goes to my wife Dr Juana Elizabeth Reyes Martinez, my parents and sisters, María Priscila del Castillo Frías, Dr Benjamín Raziel Jaramillo Ávila and Dr Elsa Aurora Calleja Quevedo, because of without their support, the culmination of this work would not have been possible.

THESIS STRUCTURE

This thesis describes the design of peptide hydrogels for bone tissue engineering applications. The body of this thesis includes an article that has been already published, and three manuscripts, which will be submitted for publication. The format of both the published paper and the three manuscripts has been modified to fit in the template of this thesis, however the main text was not modified. The references either for the article or manuscripts are at the end of each chapter, as in the submitted version. In this thesis, a review manuscript (Chapter 1) serves as a general introduction, which provides an overview of bone tissue, and the factors and diseases affecting its metabolism is provided. Likewise, this literature review includes a general introduction to the biomaterials, and the state of the art of peptide hydrogels for the regeneration of bone and dental tissue regeneration. The first published original article (Chapter 2) describes how the FEFEFKFK peptide hydrogel works as a three- dimensional (3-D) niche for the culture of human osteoblasts (bone-forming cells). This chapter also discusses how the gel fosters bone mineralisation of cells cultured within it, highlighting its possible use for bone tissue engineering. The second manuscript (Chapter 3) discusses the effect that the functionalisation of the FEFEFKFK peptide hydrogel with the short peptide sequence RGD have on the physiological response of human osteoblasts. The third manuscript (Chapter 4) deals with the suitability of the non-functionalised peptide FEFEFKFK hydrogel to induce the osteogenic differentiation of human mesenchymal stem cells (hMSCs). The chapters in this thesis follow a logical sequence. First, we studied the response of osteoblasts within the non-functionalised FEFEFKFK hydrogel, and then the cellular response within the functionalised gel (RGDFEFEFKFK). Subsequently, taking advantage of the inherent plasticity of stem cells, we also explored the suitability of such octa-peptide gel to induce the osteogenic differentiation of hMSCs and mineralisation by the differentiated cells under osteogenic stimulation. The methods used in this project are described in detail in Chapter 5. Finally, conclusions, outlook and recommendations for future work are discussed at the end of this thesis (Chapter 6). In summary, this work has yielded a promising peptide gel-based approach, which might serve for inducing the regeneration of bone and dental tissue.

ABSTRACT

In recent years, the degradation and subsequent loss of tissues is an issue that has affected people worldwide. Although there are treatments addressing the degradation of tissues, such treatments involve complicated and expensive procedures, where full tissue regeneration is not achieved. For these reasons, in recent years, tissue engineering has developed cutting-edge biomaterials capable of inducing effective tissue regeneration both under cellular or acellular conditions. Peptide hydrogels are versatile biomaterials composed of the basic components of life amino acids, which act as building blocks to form hierarchical structures, which subsequently go on to form well-defined scaffolds. Biomaterials have been widely used for the culture of mammalian cells, tissue engineering, regenerative medicine, drug delivery, etc. This is thanks to their capability of providing a three-dimensional architecture to cells, which mimics the natural architecture of the extracellular matrix (ECM). Peptide- based hydrogels can be easily functionalised with active biological cues, which can direct the cellular response. It has been shown that the ionic-complementary FEFEFKFK hydrogel, succeeded to support the culture of mammalian cells such as bovine chondrocytes. In this work, we used the same FEFEFKFK hydrogel to investigate the capability of this hydrogel to support the three-dimensional culture of both human osteoblasts (hOBs), and human mesenchymal stem cells (hMSCs) for bone regeneration applications. To achieve this goal, hOBs were cultured within both FEFEFKFK (non-functionalised) and RGD-FEFEFKFK (functionalised) gels. Then the suitability of the FEFEFKFK gels to induce cellular proliferation, synthesis of bone ECM and mineralisation was explored. In addition, taking advantage of the inherent plasticity of hMSCs, we also investigated the capability of the FEFEFKFK gel to foster the osteogenic differentiation of hMSCs, and subsequently to induce bone mineralisation in 3-D under osteogenic stimulation. Based on the results obtained in this work, the FEFEFKFK gel arises as a promising biomaterial for both bone and dental tissue regeneration applications.

OBJECTIVES OF THE THESIS

Developing cutting-edge biomaterials capable of regenerating damaged tissues is important to provide alternative routes to treat people suffering of tissue degradation caused by degenerative disease or trauma. Using peptide hydrogels to provide a 3-D environment to cells is an interesting approach for the study of the cellular response with tissue engineering applications. In addition, hydrogels (alone or combined with other biomaterials) can be used to heal/regenerate damaged tissues such as bone, and dental tissue. The main goal of this work was designing an ionic-complementary peptide hydrogel capable of providing a 3-D niche for the culture hOBs and hMSCs, and induce the formation of bone-like tissue. The hydrogel used in this work has the peptide sequence FEFEFKFK, where F, E and K are phenylalanine, glutamic acid and lysine respectively. To achieve this goal we used three different approaches. (1) We studied the suitability of the non-functionalised FEFEFKFK gel to support the 3-D culture of hOBs. This involved evaluating several aspects of cell physiology such as capability to survive, proliferate, synthesise ECM and mineralise within the gel. (2) We modified the FEFEFKFK peptide sequence by incorporating the adhesive short peptide RGD (RGDFEFEFKFK). Then, we studied the effect of the peptide functionalisation on the physiological performance of hOBs to form bone-like tissue. Finally, (3) we studied the capability of the non-functionalised FEFEFKFK gel to foster both the osteogenic differentiation of hMSCs, and to induce mineralisation by differentiated cells. We expect that this work will provide fundamental insights related to the possible application of a peptide gel for inducing bone and dental tissue regeneration in the near future.

CHAPTER ONE

Versatile peptide hydrogels for bone and dental tissue regeneration

Luis A. Castillo Díaz, Alberto Saiani, Julie E. Gough, and Aline F. Miller

All authors contributed to the realisation of this manuscript.

14 Versatile peptide hydrogels for bone and dental tissue regeneration

Luis A Castillo Díaz,1,2 Alberto Saiani,2,3 Julie E Gough3* and Aline F Miller1,2* .

1 School of Chemical Engineering and Analytical Science, The University of Manchester (UK).

2 Manchester Institute of Biotechnology, The University of Manchester (UK).

2 School of Materials, The University of Manchester (UK)

Corresponding authors:

Aline F Miller, Manchester Institute of Biotechnology, School of Chemical Engineering and Analytical Science, The University of Manchester, 131 Princess Street, Manchester, M1 7DN, UK. Email: [email protected].

Julie E Gough, School of Materials, The University of Manchester, Grosvenor Street, Manchester M13 9PL, UK. Email: [email protected]

15 Abstract

For many years the regeneration of bone and oral tissues has been a challenging problem to overcome for medical applications. Nowadays, the treatments used to address small or large quantities of bone degradation have not yet been able to achieve an effective tissue restoration. Whilst bone grafting incorporates biological elements, such as growth factors to improve the bone healing, concerns related to risk of infection during surgery, morbidity, undesirable immunological responses, and inappropriate bone regeneration, encouraged bioengineers to search for better therapies. Tissue engineers have recently taken advantage of peptide-based hydrogels, which offer three-dimensional (3-D) matrices. These matrices allow the culture of distinct mammalian cells to create platforms to regenerate tissues. In this perspective, we focus this review on the recent progress that biomaterials, including peptide hydrogels, have achieved regarding the regeneration of bone and dental tissue. The scope of this review consists of a brief study of the bone physiology and pathophysiology. Thereafter, the basis of a bottom-up approach to create synthetic peptide-hydrogels with tissue engineering will be presented. Finally, the recent progress that hydrogels have achieved in tissue engineering and regenerative medicine, particularly in the regeneration of bone and teeth, will be discussed.

16 Contents

1. Introduction………………………………………………………………………..18 2. Bone……………………………………………………………………………….20 2.1 Bone structure and ECM composition…………………………………………...20 2.2 Bone metabolism…………………………………………………………………22 2.3 Biology of oral tissues...... 23 3. Pathophysiology of bone and oral tissues…………………………………………25 3.1 Osteoporosis……………………………………………………………………...25 3.2 Other bone pathologies…………………………………………………………..27 3.3 Periodontal disease (Periodontitis)… ………………………………...………...27 3.4 Caries/dental pulp infection……………………………………………………...29 4. Biomaterials for bone and dental tissues regeneration……..………..…………….30 4.1 Metallics………………………………………………………………………….31 4.2 Hydrogels……………………………………………………………………..….32 4.2.1 Natural scaffolds……………………………………………………………….33 4.2.2 Synthetic scaffolds……………………………………………………………...37 4.2.3 Other synthetic scaffolds……………………………………………………….39 5. Peptide-based hydrogels…………………………………………………………...43 5.1 Synthesis of peptides……………………………………………………………...44 5.2 The basis of a bottom up strategy/peptides self-assembly………………………..45 5.3 Mimicking the ECM……………………………………………………………...46 5.4 Peptide-based hydrogels for bone regeneration both in vitro and in vivo…….…47 5.5 Peptide-based hydrogels applied in dental tissue regeneration in vitro and in vivo…………………………………………………………………………….50 6. Conclusions and future perspectives………………………………………………52 Acknowledgments……………………………………………………………………53 References……………………………………………………………………………58

17 1. Introduction

Bone is a type of highly organised connective tissue with a complex homeostasis (1). Bone metabolism can be altered by diseases involving distinct factors, which include genetic disorders, hormonal imbalance, and/or infections. Also, age or traumatic wounds can affect bone turnover (2-4). An imbalance between bone formation and bone degradation can lead to the loss of either small or large mass of bone. Nowadays, the degradation of alveolar bone is a case that cannot be completely cured by the current treatments. Autogenous or autologous bone graft is still considered the gold standard to treat most of bone affections, including alveolar bone resorption (5). Whilst the bone graft approaches involve the incorporation of growth factors such as autologous platelet-rich plasma (6) or bone morphogenetic proteins (BMPs) (7) to stimulate bone formation, issues related to growth factors dosage, and graft degradation may lead either to a poor bone regeneration or bone overgrowth (6, 8). Other additional complications include high cost and risk of infections, post-operation pain, the need of additional surgeries and special equipment (9). Despite several biomaterials such as collagen sponge, bioactive glasses, ceramics or, metallics have been used to regenerate bone; none of these materials meet all the needed features to promote “an ideal bone regeneration”. It is believed that biomaterials intended for bone regeneration applications should accomplish at least the following features: osteoinductivity, osteoconductivity, biocompatibility, good mechanical properties and degradability, among others (10, 11). Recently, several strategies have been exploited in tissue engineering to create biomaterials for tissue regeneration applications (12-14). The strategy that proved to be most successful is the bio-inspired on a bottom-up approach, where amino acids acting as building blocks are engineered to form complex structures. This approach was inspired from nature, where the physicochemical interactions between amino acids or nucleotides are capable of forming complex macromolecules such as proteins and nucleic acids, respectively. These macromolecules form the genome as the genetic information storage database and the proteome as the molecular toolbox, for the execution of the various biochemical process essential for life in both prokaryotic and eukaryotic organisms as described by Ricardo, A., and Szostak J. W., 2009 (15), and Shapiro, R., 2007 (16).

18 Amino acids provide a chemical toolbox for designing a wide range of possible peptide sequences that can adopt bio-inspired protein secondary structures such as β- sheets (17), β-hairpins (18), α-helices (19), amphiphilic structures (20). These secondary structures can be engineered to self-assemble forming higher supramolecular structures through non-covalent interactions such as electrostatic, hydrophobic, van der Waals, and hydrophobic interactions. For example, the so-called ionic self-complementary peptides have been designed to have alternating hydrophobic and charged amino acids, which assemble into β-sheet structures that can grow into nanofibres (20). On the other hand, amphiphilic peptide sequences have been designed to have 2 blocks (20, 21). Systems built from α-helix/coiled coil peptides (19) can form supramolecular structures with different architectures such as fibrils, tapes, ribbons, tubes, spheres, cylinders, and rood-micelles (12, 20, 22). The above presented designs have been used to develop various peptide-based hydrogel systems, in which the gelation process can be triggered by external stimuli such as pH, temperature, ionic strength, light and enzymes (20). These gels are soft viscoelastic materials with tunable mechanical properties, where the peptide design, concentrations and some external factors above mentioned play a key role in giving them versatile features (20). Peptide hydrogels are highly hydrated materials (> 95%) and posses inherent biocompatibility thanks to their natural amino acid composition (23). In addition, the inner nano-architecture of hydrogels relatively resembles the architecture of the extracellular matrix (ECM). Moreover, the peptide sequences can be decorated with biological entities, which allow the formed gels to direct the cellular fate according to the need. This is useful for tissue engineering and regenerative medicine applications (24-26). Thus, in this review we will introduce the recent advances in using peptide- based biomaterials related to the regeneration of bone and dental tissue. We will also report on examples for strategies used to direct the cellular response to regenerate the above-mentioned tissues. These strategies include the incorporation of ECM biomolecules, growth factors, drugs, and the creation of hybrid systems. Finally, we will discuss the scope and outlook of peptide hydrogels as biomaterials with bone and dental tissue engineering applications.

19 2. Bone

Bone is a highly specialised tissue with a variety of functions within the body such as serving as a reservoir of minerals, supporting organs and tissues as well as acting as a framework for locomotive functions. Bone has a dynamic metabolism, which is susceptible to changes induced by local (i. e. cytokines, growth factors) and systemic factors (hormones, genetics, age). Other external factors such as fractures or wounds can also significantly alter the bone homeostasis (27, 28). Macroscopically, there are two modalities of bone: cortical bone and trabecular bone (1, 27). Alveolar bone, which supports teeth, is another type of bone that share characteristics of both cortical and trabecular bone (27, 29) (Figure 1). Cortical bone is constituted of Haversian systems, which are built from concentric layers of compact bone (lamellae). Lamellae is highly vascularized and innervated by Haversian channels (1, 27, 30). Peripherally, cortical bone is coated by periosteum, which is a fine and irrigated membrane with significant functions, which include appositional growth and bone repairing, since it contains both hematopoietic (HSCs) and mesenchymal stem cells (MSCs). On the other side, the inner endosteal bone surface also has a key role in bone turnover, as it is also harboring MSCs (31). Cortical bone has ∼ 5-10 % porosity with pore diameters ranging from ∼ 10 - 50 µm, moreover this type of bone has a compressive strength (σ) ca. 110 - 150 MPa, and a Young’s modulus (E) ca. 18 - 22 GPa (32). Trabecular bone is formed by semicircular trabecular osteons, and as cortical bone, it acquires a lamellar pattern, where collagen fibrils rearrange in alternating pattern orientations (1, 27). This type of bone is highly porous (∼ 75-85 % porosity), possess bigger pore sizes ranging from ∼ 300 to 600 µm in diameter. Trabecular bone has a compressive strength of ca. 2.6 MPa and Young’s modulus of ca. 0.1 - 0.3 Gpa (32). Overall, mineralised bone possess a compressive modulus range from ca. 100 kPa to 1000 kPa (33), as well as a Young’s modulus of ca. 109 – 1010 Pa according to literature (34).

2.1 Bone structure and ECM composition

Bone is composed by an inorganic phase (∼ 70 %), and an organic phase (∼ 20 %), where overall the cells and water represents ∼ 10 % of the whole tissue content (1, 2,

20 27, 30). The inorganic phase, is basically constituted by minerals such as calcium phosphate (CaP), which subsequently becomes hydroxyapatite (HA) with the chemical composition Ca10(PO4)6(OH)2. Other minor elements found in the inorganic phase include carbonate, acid phosphate, sodium (Na) magnesium (Mg), potassium (P), zinc (Zn), and fluoride (F-). Overall, the inorganic phase provides both stiffness and rigidity to bone tissue (1, 27, 30). On the other hand, the organic matrix, is mainly made of type I collagen or collagen I (COL-I) (ca. 88 %), and a little of other type of collagens such as collagen III, V, and X (1, 35). The organic phase also contains various non-collagen proteins such as alkaline phosphatase (ALP), osteocalcin (OCN), osteopontin (OPN), osteonectin (ONN), bone sialoproteins (BSPs) and other adhesive proteins such and fibronectin (FN) and vitronectin (1, 35). Additional non- collagen proteins include serum albumin, aggrecan, versican and decorin, which promote the adhesion of calcium and facilitate bone mineralisation. Finally, growth factors such as transforming growth factor beta (TGF-β), insulin growth factor (IGF-1), platelet-derived growth factor (PDFG), basic fibroblast growth factor (bFGF), epidermal growth factor (EGF), vascular endothelial growth factor

(VEGF), vascular cell adhesion molecule-1 (VCAM-1), and bone morphogenetic proteins (BMPs) are also part of the bone ECM (1, 35).

Bone cells are found either on bone surface or in bone marrow, which is a soft tissue having a stiffness of ca. 0.2 kPa (storage modulus) (36). Precursors of bone cells are also found within the vascular tissue in Haversian systems. The two major bone cells are osteoblasts (OBs), and osteoclasts (OCs) (27, 35). Osteoblasts have a mesenchymal origin, which means that mature osteoblasts come from bone marrow mesenchymal progenitor cells. The main function of osteoblasts is to synthesise osteoid (non-mineralised bone) via the production of ALP. These cells also synthesise other enzymes such as matrix metalloproteinases (MMPs), which have a key role in the subsequent mineralisation of bone (37). Osteoblasts are also active orchestrators of bone metabolism triggering the activation of osteoclasts (OCs), which are bone- resorbing cells. OBs synthesise and express the osteocytokines, such as the macrophage colony-stimulator factor (M-CSF), the receptor activator of NF-κB ligand (RANKL). These osteocytokines promote the differentiation and activation of pre-osteoclasts and OCs. Nonetheless, OBs inhibits the activation of OCs via the

21 expression of osteoprotegerin (OPG), which is a decoy receptor for RANKL (1, 35, 38). Osteoclasts are bone-resorbing cells formed by the fusion of mononuclear progenitors, which belong to the monocyte/macrophage lineage. Progenitors of OCs can be found either in circulating blood or within bone marrow (hematopoietic stem cells (HSCs). OCs are responsible of the regulation of bone mass through the degradation of the tissue using enzymes such as tartrate-resistant acid phosphatase (TRAP) and MMP-9 (39, 40). Osteoclasts express the RANKL receptor called RANK, and other receptors for other biomolecules involved in bone metabolism such as calcitonin, parathormone (PTH), glucocorticoids (GCs), and estrogens. OCs are also important source of pro-inflammatory cytokines such as interleukin-1 (IL-1), IL- 6, and tumor necrosis factor-alpha (TNF-α), which act as autocrine stimulators to orchestrate the degradation of bone (1, 35, 40). Other important cells present in bone include mesenchymal stem cells (MSCs), and hematopoietic stem cells (HSCs), which can be found in several areas of bone such as Haversian system, bone marrow, periosteum and endosteum (31). Stem cells renew the cell populations as required, allowing and facilitating the bone turnover (31). Table 1 summarises the characteristics and functions of the major cells of bone.

2.2 Bone metabolism

As a highly dynamic system, bone itself sculpts and renews through two fascinating processes, modeling and remodeling, which provide of shape and maintain an adequate density to bone through life. Modeling is held during childhood and adolescence, where bone is formed by endochondral ossification. During this process the bone extremities (epiphyseal and metaphyseal areas) are predominantly formed of cartilage (1, 38, 41). Subsequently, the soft tissue becomes vascularised, and cartilage plates are replaced with stiffer tissue, where both OBs and OCs are predominant, thus regulating the mineralisation of the new osteoid formed. It is believed that the modeling process is triggered by the physiological demands during growth; likewise this process also can be initiated by mechanical stress, where bone growing is tightly regulated according to needs. During the bone modeling, both apposition and resorption process are not entirely coupled. Apposition occurs in one side of bone, whilst resorption normally occurs in the opposite side (1, 38, 41).

22 On the other hand, bone remodeling is a process that occurs throughout life, and its major function is renewing the tissue. Under physiological conditions, from 5 to 10 % of bone volume is renewed every year (1, 41). During the bone remodeling process, old tissue is replaced by new osteoid, which is subsequently mineralised. Remodeling is carried out in the so-called Basic Multicellular Units (BMUs), which are formed by mature OBs and OCs. Unlike modeling, remodeling is a process tightly coupled, where bone resorption is immediately followed by apposition in the same site of bone “bone coupling” (Figure 2). Remodeling is held where physiologically necessary, either in old tissue areas or in sites where self-repair is a priority; nonetheless it is also believed that this process may occur randomly. Overall the process initiates with the recruitment and activation of mononuclear precursors, which following several differentiation states become mature osteoclasts. As previously mentioned, OCs activation is interestingly regulated by RANKL and OPG expressed by OBs. Other biomolecules involved in bone metabolism as well as in the activation of OCs include M-CSF, IL-1, IL-6 and TNF-α. It is worth mentioning here that prostaglandin-2 (PEG-2) has also a significant role in bone metabolism, since its expression is enhanced during bone resorption, stimulating the expression of other pro- inflammatory cytokines. It is believed that immediately after bone resorption occurs, cells and cytokines such as TGF-β, IGF-1, and BMPs among others, are exposed and released respectively, triggering the activation, differentiation and migration of OBs precursors to the resorbed site in order to start the synthesis of new osteoid. Thus, a remodeling cycle maintains the bone integrity (1, 41).

2.3 Biology of oral tissues

Alveolar bone (AB) and teeth constitute the major hard tissues in the oral cavity. Alveolar bone forms both maxilla and jaw, and although it is considered as having a small anatomy, its major role in mouth is to support teeth and the mastication forces (27, 29). As AB is composed of a combination of cortical and trabecular bone, its ECM composition is very similar to both types of bone mentioned earlier, having layers of lamella with parallel orientation (27, 29). The alveolar bone process is formed by trabecular bone coated with cortical bone plates in both lingual and labial faces. Along the trabecular area of the tissue, there are numerous bone marrow areas

23 harbouring a high content of HSCs, MSCs, and other cells, which nourish and support the bone metabolism. Covering the cortical bone surface, periosteum also nourishes, innervates and serves as a reservoir of cells, such as HSCs, MSCs, OBs, and OCs, which regulate the AB turnover. The endosteum covers the inner surface of lamellar and trabecular bone, and has similar functions to periosteum. AB remodelling is constantly induced either by tooth eruption, or mechanical forces from mastication (27, 29).

Teeth are also highly specialised organs; their ECM is very similar to bone. Teeth are structured in four different tissues, which include enamel, dentin, cementum and pulp. Enamel is formed by both an inner and outer enamel epithelium, which harbour ameloblasts (enamel-forming cells). Enamel is mainly constituted by hydroxyapatite (∼ 90 %), nonetheless it also contains tiny amounts of proteins such as amelogenins, ameloblastins, and enamelins. Just below enamel, dentin is a tissue formed by apatite crystals well oriented on an organic matrix made of collagen. Collagen and minerals provide to teeth their high load-bearing characteristic. Dentin also contains odontoblasts, which produce ECM and secondary dentin when it is needed. Teeth attaches to alveolar bone through the periodontal ligament (PDL), which is attached to the roots of teeth. PDL is mainly formed by collagen, which is produced by a highly elevated population of fibroblasts. The periodontal ligament is linked to the cementum of teeth, which is a layer of tissue that covers the roots of teeth, and where cementoblasts initiates the formation of PDL. The main function of PDL is acting as a cushion to support the mastication forces. Alveolar bone, PDL, and cementum are considered like a single unit called “periodontium”, which supports teeth and the mastication forces (42, 43). The most inner tissue within teeth is the dental pulp, which is a highly innervated and vascularized tissue, where dental pulp stem cells (DPSCs) are capable to differentiate into odontoblasts. It is also known that DPSCs are also capable to differentiate into other type of cells such as adipocytes, chondrocytes, osteoblasts, and neurons in vitro, which is an interesting feature that makes them promising to be used in tissue engineering applications (44).

24 3. Pathophysiology of bone and oral tissues

Most bone pathologies induce the degradation of tissue, however other such as osteoporosis, induce an abnormal and excessive bone formation. In this section the etiology and physiology of some bone and oral diseases will be described.

3.1 Osteoporosis

Osteoporosis affects more than 200 million people worldwide, including both women and men. This disease is characterised by loss of bone mass accompanied with a deterioration of tissue. Consequently, the integrity of the tissue is compromised and bone becomes fragile, which increases the risk of fractures (2, 45). There are two modalities of osteoporosis: primary and secondary. Primary osteoporosis can be divided into type I (postmenopausal osteoporosis) and type II (senile osteoporosis). In type I osteoporosis, the estrogen levels dramatically decrease, mainly during the menopause period. In addition, calcium levels also decrease because of estrogen deficiency. Estrogen deficiency increases bone resorption rather than impairs any bone apposition, as this hormone can induce osteoclast’s apoptosis (2, 45). During osteoporosis, both processes formation and resorption occur, however bone resorption is clearly predominant. Estrogens inhibit the expression of pro-inflammatory cytokines such as IL-1, IL-6 and TNF-α by monocyte/macrophage cells and stromal cells, cytokines whose function is to induce osteoclastogenesis through the system RANK/RANKL/OPG (2, 45). It has been reported that follicle-stimulating hormone (FSH), which stimulates the production of estrogens, paradoxically also promotes osteoclastogenesis and bone resorption at later stages of postmenopausal osteoporosis (2, 28). Thus, the low estrogen levels of estrogens in postmenopausal osteoporosis, facilitates the OCs’ job, making it more comfortable. The current therapeutic protocols to tackle osteoporosis include using osteoclastogenesis inhibitors, such as estrogens, bisphosphonates, and calcitonin. Moreover, parathormone (PTH), vitamin D, and calcium supplements may also improve the bone mineral density (2, 45).

Like type I osteoporosis, type II involves the participation of estrogens and additional age-related hormones such as PTH or androgens in both women and men

25 respectively. Type II osteoporosis is characterized by a substantial decrease in women bone mass during menopause, which continue at a comparable rate to elderly males. PTH is a systemic regulator of calcium that promotes bone degradation, conversely when administrated at intermittent doses it enhances bone formation. With age, PTH levels increase promoting a faster bone turnover activity, which subsequently induces loss of bone mass, particularly of cortical bone (45). Moreover, the progressive increase in reactive oxygen species (ROS) with age may also impair osteoblasts function. Thus the presence of ROS along with a drop in the antioxidants levels can also promote osteoporosis. To summarise, the hormonal imbalance together with the stressful high levels of ROS accompanied with senility, contribute to the development of female osteoporosis. In a similar way, the lower levels of sex hormones such as androgens and estrogens favour the development of male osteoporosis (2, 45).

Secondary osteoporosis has several etiological factors, which include the administration of glucocorticoids (GCs), sedentary life, complications derived from therapeutic interventions, etc. The administration of GCs to treat inflammatory diseases can directly affect the physiology of OBs, OCs and osteocytes, promoting an imbalanced remodeling (2). It has been shown that glucocorticoids can induce the differentiation of MSCs into an adipocyte lineage instead of an osteoblastic lineage (46). Additionally, GCs can also induce the apoptosis of mature OBs and osteocytes. Secondly, GSs indirectly enhance the activity and lifespan of OCs, hence decreasing the expression levels of OPG and increasing that of RANKL, respectively. GCs can also impair the intestinal absorption of calcium and increase its renal excretion. Nowadays, bisphosphonates and well-controlled doses of GCs along with calcium and vitamin D supplements are considered as efficient therapeutic regime for treatment of secondary osteoporosis. Sedentary life triggers the development of osteoporosis since bone remodeling is impaired when mechanical loading decreases. It is believed that osteocytes are able to sense the mechanical loadings applied on bone, and subsequently they initiate the tissue remodeling accordingly. Nevertheless, bone mechanical stimulation is poor in a sedentary person, which diminishes the formation of tissue and subsequently bone remodeling process. Therefore, physical activity is important to stimulate bone remodeling and the formation of bone (2).

26 3.2 Other bone pathologies

Piaget’s disease is a bone pathology characterised by an abnormal bone formation derived from a disorganised bone remodeling. Both formation and degradation of bone can be excessive. Piaget’s disease is very often accompanied by pain, arthritis and neurological alterations. Viral infections (paramyxovirus, for instance) may trigger the development of this disease (2, 45). Moreover, when large quantities of bone are lost, either by trauma or cancer, the treatment of choice in this case consist of autologous or allogenous bone grafting (5). However, this treatment strategy involves invasive surgery, which is very often accompanied by undesirable side effects such as high risk of infection, morbidity (which may last a few weeks or years), the need of additional surgeries, etc. (9).

Osteopetrosis is a bone disease that is generally related to genetic disorders, where bone formation is predominant and subsequently becomes excessive. For example, mutations in both carbonic anhydrase and cathepsin K genes can impair the physiological function of osteoclasts proton pump (H+ transport, chloride channel, RANK expression, and carbonic anhydrase II activity). This causes the development of brittle bones and increases the risk of fractures. There are lethal forms of osteopetrosis and the current treatment for this disease is based on hematopoietic stem cell transplantation (2, 45).

3.3 Periodontal disease (Periodontitis)

Periodontitis is an oral pathology that has negative effects on alveolar bone and the surrounding tissues. This pathology is characterised by a severe and chronic inflammatory process generally caused by bacteria. At latter stages, periodontitis may induce the destruction of PDL, and eventually induce the loss of teeth (47). It is believed that approximately 700 species of bacteria colonizes the oral cavity. Some of the most common species of bacteria related to the development of periodontitis include: Aggregatibacter actynomicetemcomitans is associated to juvenile periodontitis whilst Porphyromonas gingivalis, Treponema denticola, Tannerella forsythia, Fusobacterium, Fusobacterium nucleatum, Prevotella, Prevotella intermedia, Campylobacter species, Parvimonas micra and Eubacterium nodatum are

27 linked to adult periodontitis. Normally, bacteria tend to attach to teeth surfaces, nevertheless gram-negative anaerobic bacteria have preference to live in deeper areas with low oxygen concentration such as the gingival sulcus, which is the area between the cervical teeth crown/root surface, and the adjacent gingival epithelium, forming thus the dental plaque (3, 48). During the development of the periodontal disease, the destruction of both connective tissue and alveolar bone is induced by the immune response triggered by infection. During periodontitis, there is an intermittent inflammatory process, which follows phases of high cytokine production accompanied with periods of remission. This process progressively culminates with the destruction of soft tissues and finally with the loss of alveolar bone (3).

When bacteria infect periodontium, the first immune cells facing bacteria invasion are polymorphonuclear cells (PMNs), which have the mission to eliminate the invader agent through out the production of superoxides, hydrogen peroxide, hydroxyl radicals, hypochlorous acid and chloramines. Thereafter, the second group of defensive cells arrives to support the PMNs defenses. This second group of cells includes lymphocytes, monocytes and macrophages. Cells can recognise virulence factors such as peptidoglycans, lipopolysaccharide (LPS), fimbriae and DNA via toll- like receptors (TLRs) recognition (i. e. TLR-2 and TLR-4) (3, 27). Downstream the activation of TLRs culminates with the activation of transcription factors such as nuclear factor-κΒ (NF-κΒ) and the activator protein-1 (AP-1), both involved in the expression of pro inflammatory cytokines such as IL-1, IL-6, TNF-α, and chemokines such as CXCL8, etc. (3, 49, 50). Subsequently, if the infection persist, these pro- inflammatory cytokines upregulate the expression of other factors such as cyclooxygenase-2 (COX-2), prostaglandin E2 (PGE2), leukotriene B-4 and MMPs, which in turn enhance the production of RANKL by OBs, B lymphocytes, T lymphocytes and mononuclear cells (3, 27). Thus, once inflammatory cells infiltrate and accumulate in the infected area, great quantities of collagen are lost since PMNs and monocytes produce collagenases and elastases that can degrade ECM. The toxic environment produced “in the battle field”, not only kills bacteria, but also native cells such as fibroblasts and osteoblasts, which subsequently will disappear from the crestal bone. At the same time, a large number of activated osteoclasts degrade the bone crest promoting the detachment and degradation of periodontal fibres (27). Interestingly,

28 other immune cells such as dendritic cells (DCs), also contributes to the inflammatory process through the production of IL-1β, IL-12, TNF-α and TNF-β. Helper T cells-17 (Ths-17) also produce IL-17. Likewise, the interferon gamma (IFN-γ) production is also triggered from Th-1 and natural killers (NKs) (3). As mentioned earlier the inflammatory cytokines released enhance the production of RANKL, activating OCs that subsequently will degrade bone.

During periodontitis only a few cells are capable of producing anti-inflammatory cytokines. Cells like regulator T cells (Tregs) are able to produce IL-10, TGF-β, and CTLA-4 in order to downregulate the inflammatory effect created, and thus, reducing the destruction of bone and connective tissue. Nevertheless, their activity is lower compared to the cytokine storm previously triggered. Other proteins such as tissue inhibitors of metalloproteinase (TIMPs) as well as OPG may contribute to modulating the inflammatory process (3). Nowadays, guided tissue regeneration along with autologous and allogenous bone graft is the gold standard to repair periodontium (5, 43). Growth factors such as BMPs or platelet-derived growth factor (PDGF) (6, 43), drugs (bisphosphonates) (51, 52) and vitamin supplements can also be incorporated, in order to improve bone healing (52, 53).

3.4 Caries/dental pulp infection

Caries is a dental pathology that promotes the destruction of the dental tissue including dental pulp, PDL and AB. Caries results from a demineralization process in which, enamel is degraded due to the presence of organic acids produced by bacteria during the fermentation of dietary carbohydrates. Bacteria such as Streptococcus mutans, Streptococcus sobrinus, Actinomyces and lactobacilli among other, initiate teeth degradation creating cavities on the enamel surface through the production of lactic acid, which leads to acidify the pH in the mouth. Lactic acid is the end product of the carbohydrates fermentation such as sucrose. Once enamel demineralisation is undertaken, bacteria can reach inner tissues including dentin and pulp (54). Thus, when dental hard tissues are degraded, dental pulp is exposed to a wide myriad of obligate anaerobic bacteria such as Porphyromonas endodontalis, Fusobacterium nucleatum, Bacteroides species, Eubacterium species and Peptostreptococcus micros.

29 The colonization of pulp by bacteria initiates an inflammatory process, which generally culminates with tissue necrosis, impairing the physiological function of defensive cells such as leukocytes, thus bacteria remain out of reach the immune cells. However, via TLRs and nucleotide-binding oligomerisation domain (NOD) several native cells, which include hematopoietic cells, mesenchymal cells, PMNs and pulp fibroblasts can be alerted by virulence factors (3). If infection persists, periapical and AB degradation may become significant (3, 54). The basis of alveolar bone destruction induced by caries is similar as observed in bone resorption induced by periodontitis, where after an exacerbated inflammatory process is triggered, bone destruction is inevitable (3). Nonetheless, some lipid mediators derived from polyunsatured fatty acids such as resolvins, protectins, lipoxins and maresins, may help to reduce the inflammatory process triggered during infection. These lipid-based mediators are able to switch the macrophages pro-inflammatory profile into a “proresolutive state”. Thus, it is believed that macrophages might help to diminish the pulp inflammation via the expression of anti-inflammatory cytokines and growth factors such as IL-1ra, IL-10, TGF-β, IGF-1, VEGF, among others (3, 55). On the other hand, AB loss also can be promoted by other causes such as iatrogenic, dental extractions, defective occlusion, poor fit of dentures and unfavourable loadings (3, 56), where like in the case of periodontitis, the gold standard treatment involves bone grafting (5, 43).

4. Biomaterials for bone and dental tissues regeneration

Over the last few decades, cell-based bioassays have been routinely done on two- dimensional (2-D) cell cultures, which do not allow for physiologically relevant cellular responses due to the lack of proper cell-to-cell (only lateral) and cell-to- matrix (only basal) interactions. Recently, three-dimensional (3-D) systems have arisen as an attractive approach to mimic the natural 3-D cell niche. The cellular responses obtained from 3-D cell cultures might help to better understand the cell behaviour in nature, making it easier to find more reliable and new alternative routes to regenerate functional tissues (57). Nevertheless, are indeed the current biomaterials available capable to achieve what nowadays still has not been possible: create an “ideal” biomaterial for tissue

30 regeneration? Since biomaterials are systems created in vitro, which subsequently will be transferred in situ or in vivo, they have to be biocompatible and biodegradable. In addition, biomaterials for bone tissue regeneration have to promote the osteogenic differentiation of stem cells (osteoinductivity). Likewise, these materials must induce the deposition of ECM (osteoconductivity). The mechanical properties of biomaterials with bone regeneration applications also should allow for cell survival and form new tissue in a balanced manner. Metallics, for instance, are biomaterials designed for supporting high load-bearings and can function as substitutes of bone. Unlike metallics, the mechanical properties of soft materials such as hydrogels are significantly lower, though can provide a cell a niche that mimics the architecture of ECM. These soft materials allow cells to produce the needed proteins to form new osteoid, which is softer than mineralised bone (58). Although tissue engineering has achieved a significant progress in tissue regeneration, still there is a need to develop biomaterials that individually possess all the features mentioned earlier in order to achieve effective bone regeneration. In the following section, it will briefly discuss the progress that different biomaterials, such as metallics, natural and synthetic scaffolds (58, 59).

4.1 Metallics

Metallics have been widely used as implants to restore both large and small bone defects. In dentistry for example, the use of titanium implants is common in order to support prosthetics. Titanium possesses excellent features, which include chemical inertia, good mechanical properties, low density, biocompatibility and resistance to corrosion (60, 61). There are metallic porous implants, which not only improve osteointegration and retention, but also possess pores that can be coated with bioactive membranes, where both cells and new tissue formed interconnect each other (62). Metallics can also be coated with calcium, which have shown good biocompatibility when human osteoblasts are cultured in vitro (60). In addition, titanium alloys functionalised with the peptide “P”, showed to be effective to stimulate the growth of bone tissue in vivo. Peptide P includes the sequence DIWA (where D is aspartate, I isoleucine, W tryptophan, and A alanine) from the BMP-2 receptor, which is capable to stimulate the growth of bone when transplanted into femurs of rabbits. Likewise, when DIWA is incorporated into metallics having rough

31 surfaces, bone formation is enhanced (63). Aparicio et al., have reported that the roughness of materials provides both retention and osteoconductivity, where the roughness of titanium surfaces had positive impact on the formation of alveolar bone in vivo (64). The drawbacks of metallic biomaterials include changes of pH and temperature fluctuations after grafting. These changes disrupt the composition of implants leading to corrosion and wearing of materials, which subsequently may promote the release of toxic metallic debris (61, 65, 66). Likewise, the high mechanical properties of metallics can lead to the reduction of bone mass, fractures and subsequently fail of implants, as the strength that metallics can reach very often are far higher than the strength of the bone (59).

4.2 Hydrogels

Natural and synthetic-derived polymers are attractive materials to regenerate bone and dental tissues, thanks to their biocompatibility and biodegradability as well as the ability to mimic the architecture of the ECM. Both natural and polymer-based biomaterials are generally formulated with each other or with other elements in order to improve their functionality, thus forming composites. There is a wide variety of hydrogels made from natural sources, for example type I collagen, hyaluronic acid, chitosan, alginate, and fibrin hydrogels (67). The availability of these materials, as well as the inherent functional biomolecules that they possess, make them suitable to be used as scaffolds for tissue engineering. On the other hand, concerns regarding disease transmission and batch-to-batch variability are the major drawbacks for using these natural hydrogel scaffolds when any cellular response is studied (58, 59, 67).

Polymers and peptide-based hydrogels are biomaterials made of well-identified components, which diminish the batch-to-batch variability observed in natural gels. Likewise, the components used to create synthetic scaffolds are of low toxicity to cells and thus possess good biodegradability. For example, amino acids forming peptide gels, self-assemble via covalent and non-covalent interactions to form fibril or fibrils, which entangle to form self-supported hydrogels (67, 68). The nanofibres formed in peptide hydrogels resemble the nano-architecture present in ECM, and when biological moieties are incorporated in these systems, the resultant matrix “mimics” the ECM in which cells naturally live. Moreover, the tunable mechanical

32 properties of polymer gels allow to control and direct the cellular response. Synthetic polymers used with cell culture and tissue engineering applications include polylactic acid (PLA), polyglicolic acid (PGA), poly (ε-caprolactone) (PCL), polyethylene glycol (PEG), and poly (lactic-coglycolide) (PLGA), etc.(57, 58). Peptide hydrogels include Fmoc amino acids, Fmoc dipeptides, RADA-16 (Ac-

RADARADARADARADA-CONH2), P11 (CH3-CO-QQRQQQQQQQQ-NH2)

KLD12 (AcN-KLDLKLDLKLDL-CONH2), FEFEFKFK, MAX-1 D (VKVKVKVKV PPTKVKVKVKV-NH2), etc (20, 23). In RADA peptide, R is arginine, A stands for alanine, and D for aspartic acid. For P11 peptide, Q is glutamine, and for KLD12 peptide, V is valine, P is proline, T is threonine and K is lysine. In the case of FEFEFKFK peptide, F stands for phenylalanine and E for glutamic acid.

4.2.1 Natural scaffolds

Collagen hydrogels are scaffolds used in tissue engineering because of their good biocompatibility, biodegradability, and availability. Furthermore, collagen hydrogels can degrade via body enzymes such as collagenases, serine proteases, and MMPs (67). In 2013, Uchihashi, et al. reported that murine osteoblasts cultured on type I collagen hydrogels were able to migrate through the gel, produce ECM and subsequently differentiate into osteocyte-like cells (69). Fibril type I collagen-based scaffolds are one of the most class of the natural scaffolds used for tissue engineering applications. However, there are concerns regarding risk of disease transmission as they are obtained from animal sources, which include bovine skin, tendons, porcine skin, rat tail, among other. Moreover, concerns regarding batch variability and sterilisation are also drawbacks to be overcome. Nonetheless, nowadays recombinant human collagen production is a promising tool to overcome most of these drawbacks (70).

Alginate hydrogels are another type of natural scaffolds widely used in tissue engineering. Alginate-based gels are constituted by linear polysaccharides containing (1-4)-linked β-D-mannuronic acid (M) and α-L-guluronic acid (G) monomers. Alginate is a hydrophilic material, whose gelation is triggered by the incorporation of ions such as Ca2+, Mg2+, Bar2+ or Sr2+, which interact with G monomers and form

33 ionic interactions (67). Alginate-based hydrogels possesses good biocompatibility and injectability, and their tunable degradability make this material attractive to be used for 3-D cell culture, tissue engineering and drug delivery applications. Nonetheless, since cells lack of receptors to recognize alginate polymers, cells cannot naturally attach to alginate gels. Therefore, it is necessary to incorporate functional cues to alginate hydrogels via covalent links or microcarriers. The major additives used to this aim include the incorporation of RGD moieties, as well as other adhesion proteins from ECM such as laminin, fibronectin, and collagen (71-73).

It has been shown that RGD-alginate hydrogels can effectively release BMP-2 in order to induce bone regeneration in vivo (74). Moreover, RGD-alginate microspheres have shown to support the viability of human periodontal ligament stem cells (PDLSCs) and human gingival mesenchymal stem cells (GMSCs) in vitro, as well as for the osteogenic differentiation GMSCs and subsequent mineralisation was observed in vivo (75). On the other hand, it has been reported that the functionalization of alginate hydrogels using biological cues may be difficult to control, leading to non-specific cell interactions. Likewise, the degradation rate of alginate is often poorly regulated. Alginate does not degradate enzymatically; therefore it is necessary to incorporate ionic exchanging systems via the loss of divalent ions into the surrounding medium. Finally, it is also known that the mechanical properties of alginate gels are low (67, 72).

Chitosan is a material derived from the natural polymer chitin, which can be found in invertebrates, crustacean shells, mushrooms, green algae, yeasts, etc. Chitin is chemically structured by a linear polysaccharide that resembles the natural glycosaminoglycans (GAGs), as it contains both glucosamine and N- acetylglucosamine. (67, 76). The chemical structure of chitosan contains positively charged amino groups, which facilitates that this material adheres to mucosa. Likewise, the positively charged amino groups in chitosan, provides to this material of antibacterial and antifungal activity. The protonable amino groups along D- glucosamine residues of chitosan, can form stable covalent bonds with natural and synthetic species such as DNA and some negatively charged polymers such as poly (acrylic acid). The gelation of chitosan hydrogels is based on non-covalent interactions such as electrostatic, hydrophobic or hydrogen bonds, which are

34 undertaken between polymer chains. These interactions depend on several factors such as temperature, concentration or pH. Other available presentations of Chitosan include biofilms, sponges or porous membranes (76). Recently chitosan hydrogels containing amelogenin were used to induce the regeneration of enamel. Ruan and coworkers were onto show that such chitosan gel induced the formation of CaP clusters, and that this gel fused with tooth enamel crystals in situ to form an enamel-like tissue. Likewise, saliva cultured in bacteria medium where the Chitosan gel was present, showed lower optical densities than cultures without gel, suggesting antimicrobial properties of this system (77). Costa-Pinto and co-coworkers reported that hMSCs, adhered, proliferated and differentiated on chitosan fibre scaffolds under osteogenic stimulation in vitro. Then, when this construct was implanted into a murine bone defect, cells were capable of forming bone after 8 weeks (78). In a similar approach murine stem cells 3-D cultured within chitosan porous gelatin induced the regeneration of alveolar bone in vivo (79).

It seems the inherent properties of Chitosan make this material suitable for bone tissue engineering, however it is also known that Chitosan does not degrades easily. It has been reported that unmodified chitosan only dissolves in acidic environments, due to the strong hydrogen bonds formed during its gelation, which limits its use as an injectable biomaterial. Likewise, the low degradation rate of this material makes it difficult to analyse (67, 76). Other drawbacks related to Chitosan include its low mechanical properties, nevertheless when combined with other systems such as poly (ethylene oxide) PEO, both its mechanics and biocompatibility improve. Contamination with organic and inorganic during its production is another drawback to overcome (76).

With similar characteristics, hyaluronic (HA) acid is a non-sulfated GAG found through out the ECM. Multiple disaccharide units of N-acetyl-D-glucosamine and D- glucoronic acid constitute this glycosaminoglycan, which can degrade enzymatically via hyaluronidase. In ECM, hyaluronic acid is involved in the diffusion of nutrients, tissue hydration, as well as cellular differentiation, therefore, HA gels provide of these advantages when used in cell culture (67). HA hydrogels possess an interconnected porous structure, are biocompatible and biodegradable, and it is possible to produce this biomaterial in large via microbial fermentation. Thus HA gels can be used in drug

35 delivery, cell culture and tissue engineering applications. Nonetheless, HA gels frequently are combined with other polymers and adhesion peptides, due to cells and proteins not adhering to the surfaces of this polymer structure (67, 80, 81). Recently, Kim et al. showed that a biomineralised HA hydrogel crosslinked with vinyl phosphonic acid (VPAc), could act as an efficient drug deliver system, promoting bone mineralisation in vitro (82). Likewise HA hydrogels has shown to be effective to release growth factors such as BMPs and induce the subsequent osteogenic commitment of MSCs into bone-forming cells. For example, composites such as HA-bisphosphonates or HA-PEG serving as a carrier of BMPs, showed to be effective to release the BMPs, which subsequently enhanced the physiological functions of rat MSCs, and OBs in vitro (83). In a similar approach, the composite HA/MSCs/BMP-2 induced bone formation in vivo due to the MSCs stimulation by BMP-2. The osteogenically differentiated stem cells were also able to produce angiogenesis-related proteins such as VEGF (84).

The last natural-derived hydrogel described here is fibrin. Fibrin is a protein that participates in the formation of clots during bleeding. This biomaterial has intrinsic biological activity and biocompatibility, containing integrin and growth factor binding sites. Fibrin can be obtained from autologous and allogenous samples. It is known that both MMPs and plasmin can induce the degradation of fibrin. Fibrin hydrogels have been widely used in tissue engineering to regenerate different types of tissues including bone (85, 86). Recently, Murphy and coworkers showed that the survival of human MSCs spheroids as well as the production of VEGF, were enhanced within fibrin hydrogels, making this system suitable to be used for bone regeneration purposes and also to induce angiogenesis (87). In 2013 Sasaki et al., used fibrin hydrogels containing 3-D patterned biomimetic matrices (type I collagen and HA), in order to regulate murine osteoblast function under tensile stress. Fibril gel/matrices promoted the production of calcium and OPN by murine osteoblasts in vitro, as well as the formation of bone in vivo (88). Other applications of fibrin hydrogels include drug delivery and cell culture. Despite its intrinsic biological activity, one of the main limitations of using fibrin gels includes its very fast degradation in vivo. Nonetheless fibrin gels can be combined with synthetic polymers such as PEG, in order to improve their mechanical properties.

36 The use of protease inhibitors is a promising alternative to overcome the fast degradation of this type of gels (85).

4.2.2 Synthetic scaffolds

Synthetic polymers have been widely used to elaborate hydrogels with tissue engineering purposes, since they are made of well-characterised components. The well-controlled chemical processes to elaborate these scaffolds provide good batch-to- batch reproducibility, and well-controlled mechanical properties. Polymers are biodegradable and relatively non-cytotoxic, moreover, it is possible to control the architecture, and geometry of the matrix structures formed. Unlike natural polymers, synthetic hydrogels generally lack of natural biomolecules, therefore, it is necessary to incorporate naturally occurring biomolecules or combine them with natural polymers such as hyaluronic acid, fibrinogen or chitosan (67, 89, 90). Some of the most popular synthetic hydrogels used in bone tissue engineering, include poly (ethylene glycol) (PEG), poly (vinyl alcohol) (PVA), poly (propylene fumarate) (PPF), poly (N-iso-propylacrylamide) (PNIPAAm), as well as poly (lactic- co-glycolc) acid (PLGA), poly (glycolic acid) (PGA), poly (lactic acid) (PLA) and poly (ε-caprolactone) (PCL) copolymers. Peptide-based hydrogels are other synthetic scaffolds widely used in the regeneration of bone tissue, and composites made of peptides and polymers have been also widely used in tissue engineering and regenerative medicine (57, 67). Other synthetic biomaterials include ceramics, synthetic glasses, composites, and glass ionomers (10). In this section, recent advances in synthetic hydrogels and other composites will be described with focus on bone and dental tissue regeneration applications both in vitro and in vivo. PEG hydrogels have been widely applied on bone formation, including the regeneration of alveolar bone. As mentioned, the combination of polymers improves both the mechanical and chemical properties of materials, which gives significant advantages when used for hard tissues regeneration such as bone. The work by Ni et al. is a recent example of polymer conjugation to regenerate bone, where authors combined a brittle acellular bone matrix (ABM) with the polymer system PEG-PCL- PEG called PECE in order to overcome the brittleness of such a matrix and form a stable hydrogel. In PECE, the acellular matrix provides biofunctionality to the

37 polymer, making the system suitable to induce bone formation. Thus, PECE/ABM enhanced bone formation in a bony defect after 20 weeks of treatment with the composite (91). PEG hydrogels functionalised with the moiety c(RRETAWA), induced the osteogenic differentiation of hMSCs in vitro. According to authors, the cell differentiation is attributed to the interaction of c(RRETAWA) with α5β1 integrin of cells. The osteogenic differentiation of stem cells resulted in the production of ALP. On the other hand, authors claimed that the mechanical properties of the system ca. 25 kPa) induced the osteogenic differentiation of hMSCs, even without the use of osteogenic stimuli, cells were capable to differentiate and produce ALP (92). In another interesting approach, rat MSCs were cultured inside an oligo (poly (ethylene glycol) fumarate) (OPF) hydrogels in vitro. OPF gels were decorated with RGD, and an osteopontin-derived peptide (ODP). Stem cells were able to proliferate, and differentiate into osteoblasts, producing bone proteins such as ALP and osteopontin without the addition of osteogenic media. Authors hypothesised that only when the biofunctional peptides were present cell differentiation is induced within the hydrogel (93). In 2014, Killion et al. combined maleic polyvinyl alcohol (PVA) and polyethylene glycol dimethacrylate (PEGDMA) to create a gel able to release dexamethasone, in order to induce the osteogenic differentiation of MSCs (94). The releasing rate of dexamethasone was swelling and pore size-dependent within gels, factors that can be easily tuned by changing the concentration of polymers (94). Thoma and coworkers induced the regeneration of alveolar bone using a PEG hydrogel in vivo. The biodegradable PEG-based gel, which acted as a barrier membrane, was used to host a bony graft. When the composite was transferred into an alveolar bone defect, the alveolar bone formation was increased after two weeks of implantation (95). Lutolf et al. reported that a PEG-RGD hydrogel, which is MMPs- dependent biodegradable, was successful to induce the regeneration of bone through the release of BMP-2 in situ (96).

Shah et al. went onto show that a PLGA hydrogel coated with the growth factors BMP-2 and PDGF-BB was capable of releasing such growth factors, and induce bone formation in vivo. The formation of bone was attributed to the effective rate of growth factors release over two weeks of implantation (97). With a similar approach, a

38 PLGA/PEG hybrid hydrogel was capable of releasing HA to induce bone regeneration in vitro (98). An example of polymer and peptide hybrids for bone regeneration applications is the composite RADA16-P24-PLGA. In this composite, RADA16 was coupled with a BMP-2 peptide, the so called P24. The authors showed that PLGA acted as a good carrier to host RADA16-P24, and that the release of BMP-2 was effective to promote the adhesion and proliferation of bone marrow stem cells in vitro and induce the formation of bone in vivo (99). Other polymer hybrids such as PNIPAM/Chitosan, nano HA (nHA) (CS-gPNIPAM/nHA), as well as PLA/HA, have shown to be highly compatible and effective to induce the proliferation of murine osteoblasts in vitro (100) and the regeneration of bone in vivo (101), respectively. With dentistry applications, PVA was mixed with a formulation of calcium phosphate cement (H-cem), which possesses mineralisation properties (102). After PVA/Hcem was grafted into a bone defect, new bone was successfully formed over 3 months. Due to the physical properties of the cement used, this system could have potential to be used as a graft biomaterial to induce the regeneration of alveolar bone, where relatively a small quantity of tissue has to be restored (102). There are a plethora of publications showing the role of synthetic polymers in the regeneration of bone. It is clear that the design of composites optimises and complements the individual properties and capabilities of individual systems. In addition, the inclusion of biological cues into the scaffolds brings closer both a more physiological niche to cells and subsequently a more physiologically relevant cellular response.

4.2.3 Other synthetic scaffolds

In addition to metallics and peptide gels, tissue engineers have developed other interesting scaffolds, including nano and microparticles, bioactive composites, ceramics, etc. Polymer-based microparticles are scaffolds, which provide to cells a solid substrate to adhere, proliferate and synthesise ECM. The advantages of using nano and microparticles include their suitability to act as carriers of cells and bioactive molecules such as drugs, growth factors, cell, etc. Likewise, this type of scaffolds can be injected into the defects. Microparticles can be made of synthetic polymers such as PLGA, PLLA, PLG, as for natural polymers such as Gelatin, Chitosan, etc. Moreover, there is a wide rate of sizes of microparticles, which range from 10 µm to 1,000 µm size(103).

39 In 2013, Rogers at al. showed that microparticles made of poly (D,L-lactic-co-glycolic acid) (PGLA) succeeded to support the adherence, proliferation and osteogenic differentiation of human mesenchymal stem cells. Under osteogenic stimulation, cells adhere on the surface of PGLA microparticles and expressed several bone markers, such as CBF-A, ALP, osteopontin, etc. Using alizarin red staining, authors showed that differentiated cells mineralised on the surface of the microparticles in time- dependent fashion (104). Recently, Rahman and coworkers induced the formation of bone in a cranial defect using microparticles made of (D,L-lactic-co-glycolic acid) (PGLA) and PEG. The microparticles, having 100-200 µm size, previously showed to be effective to support the proliferation of human mesenchymal stem cells, and inducing the osteogenic differentiation of murine myoblasts through the well- controlled release of BMP-2. When microparticles (PLGA/PEG/BMP-2) were transferred into the bone defect, a significant formation of bone tissue could be induced after 6 weeks of treatment as showed by micro-computed tomography and histological techniques (105). Seeking for alternatives to regenerate bone, one interesting approach is the creation of porous scaffolds, which can provide an inner architecture similar to that observed in bone, thus foam polymer scaffolds arise as an interesting option to achieve this. Poly

(DL-lactide) (PDLLA) and Poly (DL-lactide-co-glycolide) (PDLLGA) were created via melting process and supercritical CO2 foaming to form porous scaffolds having 50- 200 µm pores size. The authors showed that both scaffolds supported the viability of human supermatogonial stem cells (SSCs) over two weeks of culture. In addition, cells were capable of infiltration into the porous architecture of scaffolds and differentiate into bone-forming cells as evidenced by the production of collagen I and ALP. The authors claimed that such cell differentiation observed might be due to the three-dimensional architecture provided by the scaffold porosity, where several adhesion sites are provided to cells, which triggered signal transductions related to cell differentiation (106).

Bioactive composites made of synthetic polymers and non-degradable materials such as tricalcium phosphate (TCP) and HA are another interesting option for the regeneration of bone. For example hydroxyapatite-based materials provide biological functions to polymers since ions can mimic the inorganic phase of bone and at the

40 same time they reinforce the mechanical properties of polymers (57). Thus for example, an effective strategy to diminish the progressive degradation of polymers is the incorporation of ceramic into the system. The ions from ceramics stabilise the structure of polymers, delaying their degradation. Inversely, the acidic degradation of polymers favours the solubility of ceramics, enhancing the availability of calcium phosphates, which subsequently will stimulate cells to synthesise an osteoid (107). Ceramics have been also used in dentistry; nevertheless their high brittleness, and concerns regarding unstable oxygen flow and nutrients when used, limits their use (10, 57). Recently ceramics can be found as injectable materials, which facilitates their administration where needed. Likewise, there is a wide range of porosity of ceramics, and depending on the application they can be designed to have pore sizes ranging from a few micrometers to 100 µm or more. Likewise, it has been reported that their functionality and degradation improves when combined with autogenous grafts or synthetic polymers (10, 59, 108). In 2014, Zhang and coworkers designed a spiral ceramic composed by HA and PCL (nano-HA/PCL). This composite showed to support the adherence, proliferation and osteogenic differentiation of human osteoblasts on 2-D culture. Cells were able to produce key bone proteins, such as COL-I, ALP, BSP, OC and ONN, and subsequent mineralisation was observed (108). Bicoral porous scaffold made of calcium carbonate (pore size from ∼150 to 600 µm in diameter) was capable of inducing the osteogenic differentiation of human dental pulp stem cells (DPSCs) when 3-D co- cultured with human osteoblasts. Differentiated DPSCs were synthesised OCN, OPN and BSPs, and mineralisation activity was reported too (109). On the other hand, in 2010 Bernhardt, et al. reported that a phosphatase β-tricalcium phosphate ceramic supported the proliferation of MSCs, as well as the production of ALP under osteogenic stimulation. This type of ceramic has interconnected channels with pore diameters ranging from 750 to 1400 µm (110). In a similar approach, Manjubala and coworkers also reported good osteoinductivity and the formation of bone in a diphasic calcium phosphate ceramic made of HA and TCP using an in vivo model (111). In order to induce osteogenesis Itoh et al. used hydroxyapatite discs coated with the peptide EEEEEEEPRGDT (E7PRGT), which incorporates a consecutive sequence of glutamic acid and RGD both present in osteopontin. HA is a material with inert osteoconductive properties, and the repetitive amino acids Q and D

41 provides to the composite good affinity to HA and adhesion sites for cells. Authors reported an increase in the attachment and proliferation of murine osteoblasts in vitro, which was RGD-dependent. In addition, osteoblats were capable to express collagen I, OCN and ALP, among other proteins. Moreover, the composite induced the deposition of mineralisation nodules (112).

Bioactive glasses (BGs) are another group of biomaterials widely used for bone tissue engineering applications. These materials are made of amorphous silica, which contains Ca, providing to ceramics the capability to interact with the ECM of bone. After BGs contact with body fluids, an intermediate layer of hydroxycarbonate apatite (HCA) is formed on its surface. BGs are capable of releasing soluble silica and calcium ions, which can stimulate bone cell function (113, 114). Glasses are commercially available as powder such as Bioglass 45S5, which is composed of SiO2-

CaO-Na2O-P2O5, nonetheless there are other different versions with various compositions (114). The degradability of glasses depends on the presence of HA and calcium, and that sol-gel derived glasses generally are more bioactive and resorb more efficiently than melted derived glasses with similar composition (58, 114). Silica glasses have shown to stimulate the adhesion and proliferation of human osteoblasts, as well as to foster the formation of calcium phosphate, apatite, and enhance bone mineralisation (115-117). Additionally, bioactive glasses have been used as particulate bone grafts, however its brittleness makes it difficult to design porous versions except when either sucrose, gelatin or PMMA microbeads are incorporated via crystallisation, which limits their clinical use (58, 114, 118). New design methods to produce bioactive glasses have been developed in order to overcome its fragility. BGs have been combined with materials such as graphene, strontium and synthetic polymers (PCL and PLA), in order to form porous fibres with bone regeneration purposes (119-121). Likewise when electrospinning and sol-gel methods are combined to create BGs, nano fibres with different porosity can be obtained, avoiding the need to incorporate other synthetic polymers. One example of this approach is the silica precursor tetraethylorthosilicate (TEOS), which forms a porous and flexible cotton-wool-like bioactive glass. This material has shown to support the adhesion and spreading of murine bone-forming cells. The mechanics and flexibility of this scaffold make it a promising route for the regeneration of alveolar bone (122).

Finally, glass polyalkenoate cements better known as “glass ionomers”, are biomaterials that form a paste, which is commonly used by dentists like a base cement. This type of material is made of calcium or a strontium aluminum/fluorosilicate glass powder, which once combined with a soluble acid polymer, it forms a porous paste that may reach high strength and elastic modulus similar to that observed for cortical bone. Glass ionomer is effective to release fluoride ions, but interestingly it is also able to take fluorides from the adjacent dental tissue. Thereafter, fluoride is released from the ionomer, which helps to diminish the solubility and degradation of teeth when exposed to the acidic environment formed by bacteria. Glass ionomer has the capability to release fluoride over at least 5 years (10, 123). In table 2, we have summarised the characteristics and functions of the biomaterials mentioned in this section.

5. Peptide-based hydrogels

Peptide hydrogels are highly hydrated systems formed by the non-covalent interactions between peptide chains that self-assemble into higher fibrillar/supramolecular structures. Peptide hydrogels are dynamic systems that have the capability to absorb and retain water, swell and progressively degrade (124, 125). Interestingly, the 3-D nano-architecture of the peptide hydrogel fibrillar structures “mimic” the nanoarchitecture of the ECM, simulating the natural niche of cells. Peptide hydrogels are also biocompatible, biodegradable and soft injectable materials. Thus, these materials are suitable 3-D scaffolds for the encapsulation of mammalian cells for tissue engineering and regenerative medicine applications (124, 125). The 20 different natural α-acids provide a chemical tool-box of versatile building blocks that allow for designing various peptide hydrogel structures (22, 126). Amino acids can be combined with a wide variety of different arrangements, leading to the formation of peptide sequences that can adopt various secondary structures. The characteristics of each secondary structure depend on the properties, order and composition of the building blocks (22, 126).

43 Peptide-based hydrogels formed from aminoacids can be used for multiple applications, ranging from 3-D cell culture and tissue engineering to regenerative medicine and drug delivery (125, 127). In this section we will briefly described the process peptide synthesis, thereafter the bottom-up approach for designing peptide hydrogels with bone and oral tissue regenerations is described. Finally, we will discuss the recent progress that peptide hydrogels have achieved regarding the regeneration of bone and dental tissues.

5.1 Synthesis of peptides

There are several methods to synthesise peptides, which include solid phase peptide synthesis (SPPS), native chemical ligation (NCL), Staudinger ligation, N- Carboxyanhydride (NCA) polymerisation and genetic biotechnology. Overall, the most commonly used method is SPPS (128, 129). In SSPS method, peptide sequences are grown on solid support through a stepwise synthesis, where alternating cycles of N-terminus deprotection and coupling cycles are executed. In the deprotection cycles, the amino terminal protecting group, either a Boc- or a Fmoc- group, can be removed using trifluoroacetic acid (TFA) in dichloromethane or 20% piperidine in dimethylformamide respectively. The number of deprotections depends on the length of the peptide chain to be synthesised. Once the final amino acid has been coupled to the peptide chain, the whole peptide can be cleaved from the insoluble resin, using. TFA. This technique allows synthesise both linear and cyclic peptides. For a more extensive description of the different types of peptide synthesis the reader is referred to Ramakers and van Hest, 2014 (128, 129). At the end of the synthesis, the crude peptide product has to be routinely purified. There are several methods to purify peptides, which include Reversed-Phase High-Performance Liquid chromatography (RHPLC), Ion-Exchange Chromatography (IEC) and Size-Exclusion Chromatography (SEC), among others (126, 130). The structural characterisation and analysis of peptides include a wide variety of methods such as Circular Dichroism (CD), Infrared Spectroscopy (IR), Attenuated Total reflectance-Fourier Transform Infrared Spectroscopy (ATR-FTIR), Nuclear Magnetic Resonance (NMR) Spectroscopy, X-ray crystallography, Small Angle X- ray Scattering (SAXS), Transmission Electron Microscopy (TEM), Mass Spectrometry (MS), Ultraviolet (UV) Fluorescence Spectroscopy, etc. (126, 130).

44 5.2 The basis of a bottom up strategy/peptides self-assembly

To understand why peptide hydrogels have become a promising route for the regeneration of tissues, it is important to define the basis of the peptide self-assembly, where a molecular state known as sol-gel transition is held. As briefly described earlier, amino acids act as building blocks to form hierarchical structures. Amino acids are bind by amide bonds between the carboxyl group of one building block and the amino group of the next block to form peptides. During the self-assemble process, several intrinsic forces and external factors drive the interaction of oligopeptides. It is believed that these factors are in direct relation with the inert characteristics of the amino acids forming the peptides. The chemical interactions involved in such process includes the participation of covalent and non-covalent forces such as hydrogen bonds, ionic and hydrophobic interactions, as well as van der Waals and π-π stacking forces, depending on the type of amino acids composing the peptide (22, 23). The chemical conformation that peptides acquire in solution “sol form” include secondary structures such as such as β-sheets, β-hairpins, α-helixes, amphiphilic structures, etc. (22, 23). The self-assembly of these secondary structures depends on the design of the peptide sequence and is thermodynamically driven by the non-covalent interactions earlier described in response to external factors such as pH (130, 131), peptide concentration (130, 132), ionic strength (133, 134), temperature (130, 135), enzymes, etc. (133, 136, 137). The interaction of these secondary structures leads to the formation of several hierarchical structures such as fibrils, fibres, tapes, ribbons, micelles, etc. Thus, the network formed provides the characteristic 3-D architecture of self-supported hydrogels (20, 23) (Figure 3). Peptide hydrogels are classified according to the peptide secondary structure into amphiphilic peptides, α-helix and β-sheet forming peptides (ionic self- complementary, β-hairpin and short peptides) (20). Since peptide hydrogels are formed of natural α-amino acid building blocks, these materials are expected thus to possess excellent biocompatibility and biodegradability, characteristics that make peptide gels suitable for cell culture and tissue engineering. The mechanical properties of peptide hydrogels are easily tunable, by controlling various factors such as pH, temperature, etc. (20, 23). However, the mechanical properties of peptide hydrogels are relatively weak if compared to metallic for

45 instance. This could be due to the heterogeneous structure formed and the insufficient entanglements among fibres, as for the dynamic association-dissociation of non- covalent interactions between the peptide chains (138, 139). The relatively low mechanical properties of these hydrogels could be a drawback when used for regenerative medicine applications, where it is required to stay in situ for relatively long time periods to allow for cell functions in a physiological manner. Nevertheless, several strategies are recently being developed to overcome this concern. For instance, the stability of peptide hydrogels is peptide-concentration dependent. By increasing the peptide concentration, the storage modulus (G′) of the hydrogels can increase from ca. 10 to 1,000 Pa (139), to reach far higher mechanical properties with G′ modulus ranging from ca. 500 to 20,000 Pa under specific conditions (24). Other parameters, which include the incorporation of food additives, inorganic compounds, natural polysaccharides as well as enzymatic crosslinking, among others, may significantly improve the mechanical properties of peptide gels (139). It is important to highlight that hydrogels are biomaterials primordially designed to encapsulate cells and allows the subsequent formation of tissue, rather than serve as a load-bearing scaffold.

5.3 Mimicking the ECM

Over last decades tissue engineers have developed biomaterials to study the cell biology within 3-D scaffolds, which are relatively capable “to mimic” the nano- architecture found in natural tissues. This strategy has allowed to study the cell behaviour in a more physiologically relevant manner in order to apply the acquired observations to regenerate different tissues (140). Nonetheless, it is important to understand that the human body is meticulously engineered and as such, its physiology is something extremely hard to mimic. Current biomaterials are far away from laboratory reproduction of what occurs in the human body. Thus, it is not enough to create biomaterials that “mimic the architecture and biofunctionality of the ECM” and subsequently encapsulate cells inside these materials (24). To regenerate human tissue, it is necessary to find a balance among components of materials as well as, cells and biomolecules used. Whilst, the in vitro 2-D cell culture has been the starting technology for the study of the cell biology, the transition into 3-D culture is currently a watershed, where the 3-

46 D scaffolds, like hydrogels, relatively provide a nano-architecture that resembles the natural niche of cells in the body. Additionally, decorating peptide hydrogels with active biomolecules make these materials more suitable to encapsulate cells with the ability to trigger/direct biological events (141, 142). Tissue engineering has taken advantage of the properties of various biological systems and has developed sophisticated scaffolds to regenerate tissues (143). Bone ECM is a complex niche, where a myriad of functional biomolecules interact with each other to keep the homeostasis of the tissue. In this environment, the cellular physiology is tightly orchestrated and regulated. However, cells are not the main component of the ECM, therefore they have to work hard to make suitable site for they live and remodeling their environment when needed (144). Therefore, directing the function of cells is one of the approaches of tissue engineering to induce the regeneration of tissues, including bone. To achieve this, both synthetic and natural hydrogels can be decorated with natural ECM proteins, such as adhesion moieties, growth factors, hormones, etc. These proteins direct the cellular activity, which is closer to that observed physiologically in the body. Some of the cell functions influenced by these factors include cell viability, adhesion, proliferation and ECM synthesis. Thus, the idea that hydrogels are passive 3-D scaffolds where cells acquire a random behaviour has fallen behind (26, 61, 141, 145). The common bone ECM cues incorporated into peptide hydrogels used for bone tissue engineering include RGD, BMPs, HA, VEGF, and TGF-β, etc. (125, 146). Hydrogels used to promote the formation of new osteoid and induce bone mineralisation in vitro (147-149) and in vivo have shown excellent progress, as they have proven to support the culture of cells and promote the formation of bone ECM (99, 150). Nonetheless, still is necessary to find an ideal balance in the components used to design the systems used for tissue engineering applications. This is important in order to overcome concerns regarding the modulation of 3-D structures, rearrangement of scaffolds, signal availability and disposition, as well as rate of cellular induction (140).

5.4 Peptide-based hydrogels for bone regeneration both in vitro and in vivo

There are extensive studies for the use of synthetic polymers for bone and dental regeneration. The use of peptide hydrogels for this application is less studied (148).

47 Nevertheless, peptide hydrogels have showed to be effective scaffolds for regenerate tissues including bone and dental tissues. As discussed earlier, hydrogels are not the ideal materials to support high-load bearings in bone tissue, however hydrogels may act as effective bone tissue inducers, fostering the production of new osteoid by cells within the gels or local cells in situ. Using peptide hydrogels as scaffolds for the regeneration of relatively small quantities of bone such as alveolar bone, could be an promising strategy to overcome the degradation of this type of bone (148). One of the most common systems used in tissue engineering and regenerative medicine is the β-sheet-based forming peptide hydrogels. This system is basically constituted by peptide chains having both negatively and/or positively charged amino acids, which are alternatively placed along the peptide chain with hydrophobic residues. The properties of amino acids can be controlled by changing the pH and the ionic strength of the system. The interaction forces driving the molecular self- assembly of this class of peptides include electrostatic, hydrogen bonds, hydrophobic interactions, van der Waals, etc. (17, 20, 151). The system RADA16-1 has been widely used for bone tissue formation purposes. RADA has showed to support the 3- D culture of murine osteoblasts in vitro when functionalised with ECM moieties (152) such as the fibronectin-derived integrins recognition sequence RGD, where the attachment, spreading and proliferation of bone murine cells is enhanced (153). Composites where β-sheet-based gels are used also have shown to promote bone regeneration. The composite formed by RADA16 and the polymer PolyHIPE (PHP) induced the growth, adhesion and differentiation of rat osteoblasts. Additionally, the in vitro production of ALP and other bone markers as well as the synthesis of new osteoid was observed. According to the authors, the porosity and the 3-D fibre environment provided by both biomaterials polymer and peptide fibres, respectively, provided a suitable environment to encourage cells to synthesise bone (154). The osteogenic differentiation of MSCs within peptide hydrogels is also an approach that provides significant advantages in regenerative medicine. In order to improve the mechanical properties and the osteoconductivity properties of RADA16, hydroxyapatite was incorporated. The system induced the osteogenic differentiation of rat MSCs (rMSCs), which produced ALP, and subsequently mineralised under osteogenic conditions in vitro (148).

48 In an attempt to simplify long 16-mer peptide sequences RDADA16, we have developed in our laboratory the β-sheet-forming octapeptide FEFEFKFK. The FEFEFKFK hydrogel showed to be suitable for supporting both the viability and proliferation of human osteoblasts when 3-D cultured within the gel. In addition, osteoblasts were capable to produce key bone proteins such as collagen I and ALP, and subsequently mineralised in vitro (147). The significance of such work relies on the fact that a shorter peptide sequence was used to create a suitable 3-D niche for the culture of human osteoblast cells (147). A different system created by Dettin and coworkers, is the ionic-complementary peptide EAbuK (Ac-Abu-E-Abu-E-Abu-K-Abu-K-Abu-E-Abu-E-Abu-K-Abu-K- OH), where Abu is 2-aminobutyric acid. This peptide was decorated with RGD and used to coat titanium surfaces in order to induce the release of IGF-1. This system effectively enhanced human osteoblasts functions such as adherence and proliferation in vitro (149). Beniash et al. developed a panel of amphiphile peptides such as the P Alkyl-C4G3S RGD-COOH peptide, where C is cysteine, G glycine and S serine. The amphiphile peptides were functionalised with distinct ECM moieties such as RGD, IKVAV, etc. The authors reported that osteoblast viability and proliferation were enhanced within these peptide hydrogels (155). On the other hand, Anderson and coworkers functionalised the amphiphile peptides CH3(CH2)14CONH-GTAGLIGQ- RGDS (PA-RGDS), (G stands for glycine, T threonine, L leucine and I isoleucine) and PA-DGEA. The short adhesion moieties RGD and DGEA derived from fibronectin and collagen, respectively. According to the authors, the osteogenic differentiation of hMSCs was significantly induced within PA-RGDS, even without the use of osteogenic stimulation. However, both functionalised systems were able to induce the cell differentiation with or without osteogenic stimuli. Authors suggested that such cell differentiation could be triggered via the recognition of ECM moieties by α5β1 and α2β1 (156).

Studies have reported the use of peptide hydrogels with bone formation applications in vivo. Some of these studies include that recently reported by Pan et al. in 2013, where RADA16 was coupled with a BMP-2 related peptide with the peptide sequence S(PO4)KIPKASSVPTELSAISTLYLDDD-CONH2, where S and Y are serine and tyrosine, respectively. Both RADA16-P24 and a PLGA (RADA16-P24-PLGA

49 construct) were combined in order to culture rat bone marrow stromal cells, which successfully adhered to the system in vitro. Stem cells subsequently differentiated and were capable to form new osteoid over 8 weeks in vivo (99). A composite combining RADA16 and HA increased the production of collagen 1, ALP, and other early osteogenic markers by rat bone marrow mesenchymal stem cells over two weeks in vivo (157). Mata et al. have previously evaluated the capability of two functionalised PAs: GS(P)EELLLAAA-C16 and SDGRKKLLLAAA-C16. In order to promote bone formation in vivo, each PA was functionalised with a phosphoserine residue S(P)-PA, and RGDS (PA-RGDS), respectively. The scaffolds showed to enhance bone regeneration more efficiently when combined and transferred into the bone defect than when transferred individually. The authors claimed that the highest bone formation observed under the scaffold combination, might be due to the phosphorylated site of the construct, which could stimulates both calcium and phosphate ion deposits, while RGD provided adhesion sites to cells (150). An alternative approach in bone regeneration is the growth factors release. A collagen sponge reinforced with poly (glycolic acid) PGA was combined with a PA hydrogel to form a hybrid scaffold capable of carrying and releasing bFGF. Authors reported extended periods (up to 750h) of sustained bFGF release from this system in vitro, where 90% of bFGF was released. Likewise, the production of both ALP and osteocalcin increased along with the bone mineral density in vivo over 4 weeks (158). It is clear that peptide hydrogels either individually or conjugated with other synthetic polymers or additives are capable to induce bone formation. Thus, peptide gels once functionalised; are capable to provide both good osteoinduction and osteoconduction conditions, which are necessary to induce cells to form new bone tissue in vitro and in vivo.

5.5 Peptide-based hydrogels applied in dental tissue regeneration in vitro and in vivo

The regeneration of alveolar bone and teeth has acquired significant interest in tissue engineering and regenerative medicine. The current therapeutic strategies for oral diseases such as caries or periodontal disease, require expensive and traumatic procedures, which frequently are accompanied by post-interventions and morbidity (159). For example, the clinical therapeutics “operative dentistry” used to treat caries, implies the removal not only of the infected tissue but also the removal of healthy

50 tissue, leading to teeth fragility and the need of more complicated and expensive port- treatments (159). Several researches such as Amalia Aggeli (160, 161), Jeffrey Hartgerink, and Rena D’Souza (162), have developed peptide gels in order to regenerate dental tissue.

Aggeli’s group developed the aforementioned P11-4, which self-assembles under specific pH and ionic strength conditions in situ. Thus, a “mineralising solution” composed of NaCl, NaHCO3, CaCl2 and Na2HPO4, which is introduced into the system, facilitates the nucleation of HA by the peptide gelation. The capability of the system to promote HA nucleation was assessed using a caries-like model in human permanent premolar teeth. Authors reported that when the peptide was applied into the dental defect, enamel mineralisation increased over 5 days. Using electron microscopy, authors claimed to show the formation of dense deposits associated to gel fibrils, which might represent hydroxyapatite nucleations. Likewise, using X ray spectroscopy, the authors showed the presence of calcium deposits (160). Recently,

Brunton et al. transferred the P11-4 in situ, where enamel regeneration in teeth of patients with caries-like lesions was induced without the need of drilling (161). On the other hand, a PA with the peptide sequence KKGGGAAAK functionalised with RGD (bRGDS PA) promoted the formation of enamel nodules when injected in situ (mice incisors in formation). Using different microscopy techniques, micro computed tomography and histology; the authors showed that the transference of teeth containing the PA into a mice kidney capsule for 8-10 weeks, induced to ameloblasts forming enamel nodules (163). Amphiphile peptides combined with RGD have also shown to be effective to form enamel in vitro as reported by Galler et al. in 2008. The RGD-PA having the peptide sequence GTAGLIGQERGDS, was used to 3-D culture stem cells from human exfoliated deciduous teeth (SHED) and DPSCs under osteogenic conditions (164). SHED cells were capable of proliferation, produced collagen and ALP, while DPSCs differentiated into ameloblasts and also produced ALP. In addition, DPSCs were capable of mineralising within the scaffold (164). The regeneration of dental pulp is another interesting field, in which peptide hydrogels have had promising advances. The multidomain peptide (MDP) having the sequence K(SL)3RG(SL)3KGRGDS has been used to harbour and release growth factors involved in the regeneration of dental pulp such as FGF- 2, TGF-β1 and VEGF. The MDP peptide efficiently released such growth

51 factors during relatively long time periods (two weeks). The release of growth factors induced morphological changes in cells, and cell proliferation was enhanced by FGF-2 in vitro. Subsequently, the MDP peptide, cells and growth factors were incorporated within dentin canals, which were subsequently transferred into mice. Using immunohistochemistry and histology analysis, the formation of a vascularised-like connective tissue in vivo was observed (162). Thus, the peptide systems here described have shown interesting properties, which make them promising tools to induce the regeneration of dental tissues.

6. Conclusions and future perspectives

It is clear that in the recent years, peptide-based biomaterials have achieved an interesting progress in tissue regeneration. There is no doubt about that peptide hydrogels provide a suitable environment for cells to grow and proliferate. When functionalised with ECM moieties or combined with other biomaterials, peptide hydrogels significantly improve the cellular functions. Therefore, peptide hydrogels represent a promising class of scaffold that proved to be useful for bone and dental tissues regeneration. Nevertheless, so far, most of the studies in vitro and in vivo have been carried out using murine cells, which is not an ideal physiologically relevant model. It is widely known that humans and rodents share similar genome and physiological functions; nonetheless it is evident that there are clear differences in the physiological cell response between both species. Thus, the findings obtained from studies where murine cells are used, might be confusion and subjective, since the main goal of tissue engineering is the regeneration of human tissues. In addition, it is important to consider that there is still a necessity to find an adequate balance in the composition of the “ingredients” (i. e.: amino acids, biological molecules, cells) used to create biomaterials in order to induce the formation of healthy and functional tissues. Likewise, it is necessary to understand more about the immune response triggered by professional and non-professional immune cells, which can reveal important insights to generate more effective and efficient biomaterials. However, the advances that tissue engineering have achieved in biomaterials field, later or sooner will allow us to develop the optimal materials that possess most of the desired characteristics discussed in this review. This will allow us not only regenerating

52 tissues, but also preventing many of the pandemic diseases affecting people worldwide.

Acknowledgements

The authors would like to thank CONACyT-Mexico for financial support, and members of the Polymers and Peptides Research Group, Biomaterials and Tissue Engineering Group, as well as Professor Sarah Cartmell’s Group for the helpful discussions and support.

53 List of figures

A B

Figure 1. Schematic representation of bone micrographs. A. Cortical bone (taken and modified from web). B. Trabecular bone (taken and modified from web). C. Mouse alveolar bone, taken and modified from FESI, UNAM-Mexico.

HematopoieCc5stem5cell5 Stromal5stem5cell5

AcCvaCon5of5osteoclast5precursor5 Osteoblasts5 by5stromal5stem5cell5 Osteoclast5 5coming5out5

BMPs" VCAM" Mononuclear5precursors5fusion5 TGF$β" IGF$1"

Growth5factors5released5aAer55 Osteoclast5resorbing5bone5 Bone5 bone5resorpCon5by5osteoclast5

Figure 2. Representative of bone coupling during bone remodeling. Osteoblast precursors and mature osteoblasts differentiate, and active respectively to synthesise new osteoid after bone resorption by osteoclasts.

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55 Tables

Table 1. Features and physiological functions of the distinct bone cells.

Cells Features Functions Round/cuboidal shape (ca. 20-30 m), life span ca. 1-10 weeks (35, µ Bone-forming cells. 165). Synthesise collagen, ALP, MMP-13, proteoglycans, RANKL Calcium storage. HA Osteoblasts OPG, M-CSF, IL-1, IL-6, IGF-I, PDGF, TGF-b, bFGF, and BMPs. formation. Transport of Receptors for PTH, estrogens, androgens, vitamin D3, and GCs (1, ions (1, 35, 38). 37, 38). Ring shape, large multinucleated (ca. 100 m), have brush border to µ Bone-resorbing cells. resorb bone surfaces. Lifespan ca. 12 days (165). Secrete TRAP, Regulation of bone Osteoclasts hydrogen ions, acidic collagenase, and cathepsin K. Express RANK, metabolism (35, 39, TRAP, MMPs. Receptors for calcitonin, androgens, insulin, PTH, 40). IGF-I, IL-1, CSF-1, and PDGF (1, 35, 39, 40). Maintenance of bone structure. Mechanical Quiescent, star-shaped (ca. 5-20 m), multiple cytoplasmic stress sensing. Osteocytes µ processes. Produce MMP-13 (30, 35, 166). Transport of CaP and inorganic phosphate (Pi) (27, 30, 35). Bone formation. Quiescent, elongated shape, capability to return into its previous Transport of ions. Bone lining cells activates condition. Express bone sialoprotein, OPN, OCN, ALP. Regulation of bone Receptor for PTH (35, 37, 167). metabolism (35, 37, 167). Quiescent, with reticular or pericyte-like morphology (in vivo). Elongated/fibroblast-like morphology (in vitro), subsequently wider Maintenance and Mesenchymal morphology (at later subcultures). CD34+, CD44+, Stro 1+, CD90+ regeneration of tissues stem cells (Thy-1), CD45-, CD19- (168-170). Express TGF-a, TGF-b, (168, 169). hepatocyte growth factor (HGF), (FGF-2), IGF-1, VEGF, and EGF (169). Quiescent, with reticular morphology (in vivo) (170). CD34+, Maintenance and Hematopoietic CD150+, CD90-/+, Lin-, CD38-, CD48-, CD41-, CD45RA-, CD49f+, regeneration of tissues stem cells and Rholo (171). (168, 169).

Table 2. Comparison of biomaterials according to their characteristics, highlighting their advantages and disadvantages when used for tissue engineering applications.

Biomaterial Advantages Disadvantages Need of invasive surgery for implantation. Thermo-conductors. Lacking of bioactivity. Releasing of Good mechanical properties. Metallics toxic metallic ions/or particles from Biocompatibility. Bioinert (62). corrosion or wearing. Mechanical properties higher than natural bone. Stress-shielding process (62). Potential immunogenicity. Risk of Contain natural biological infections and disease transmission. components. ECM biomolecular Variability between batches. Poor Natural scaffolds properties. Natural degradation. handling, cell adhesion and mechanical Good availability (67). properties. Easy degradability.

Sterilisation issues (58, 59). Reproducibility. Control of chemical Limited control over nanoscale components. Easy handling. structure. Biologically inactive (67, Biocompatibility. Scaffold structure 172). Resistance to enzymatic Synthetic mimics ECM. Control of degradation. Sterilisation issues. Low hydroxyesteres mechanical properties. Easy bio- biocompatibility, and formation of polymers functionalisation and formation of acidic degradation products. Stealth hybrids. Drug delivery and effect. Enhanced permeability and Injectability properties (58, 67). retention effect (68, 172, 173). Hierarchical structure formation. Reproducibility. Control of chemical components and matrix structured formed, which mimics ECM. Low- cost production, immunogenicity Synthetic Lacking of bio-functionality. and toxicity. Easy handling. peptide-based Sterilisation issues. Low mechanical Enzymatic and controllable hydrogels properties (67). degradability (172). Tunable mechanical properties. Easy functionalisation and formation of hybrids. Drug delivery and injectability properties (58, 67, 172). Control of chemical composition Other synthetic and rate of degradation. Release of Low toughness, brittle (174). biomaterials ions involved in bone formation and angiogenesis (174).

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69 CHAPTER TWO

Human osteoblasts within soft peptide hydrogels promote mineralisation in vitro

Luis A. Castillo Díaz, Alberto Saiani, Julie E. Gough, and Aline F. Miller

All authors contributed to the realisation of this original article.

70 TEJ0010.1177/2041731414539344Journal of Tissue EngineeringCastillo et al. 539344research-article2014

Original Article

Journal of Tissue Engineering Volume 5: 1–12 Human osteoblasts within soft peptide © The Author(s) 2014 DOI: 10.1177/2041731414539344 hydrogels promote mineralisation in vitro tej.sagepub.com

Luis A Castillo Diaz1,2, Alberto Saiani2,3, Julie E Gough3 and Aline F Miller1,2

Abstract Biomaterials that provide three-dimensional support networks for the culture of cells are being developed for a wide range of tissue engineering applications including the regeneration of bone. This study explores the potential of the versatile ionic-complementary peptide, FEFEFKFK, for such a purpose as this peptide spontaneously self-assembles into -sheet-rich fibres that subsequently self-associate to form self-supporting hydrogels. Via simple live/dead cell assays, we demonstrated that 3 wt% hydrogels were optimal for the support of osteoblast cells. We went on to show that these cells are not only viable within the three-dimensional hydrogel but they also proliferate and produce osteogenic key proteins, that is, they behave like in vivo bone cells, over the 14-day period explored here. The gel elasticity increased over time when cells were present – in comparison to a decrease in control samples – indicating the deposition of matrix throughout the peptide scaffold. Moreover, significant quantities of calcium phosphate were deposited. Collectively, these data demonstrate that ionic-complementary octapeptides offer a suitable three-dimensional environment for osteoblastic cell function.

Keywords Scaffold, peptide hydrogel, osteoblast cells, bone formation, mechanical properties Received: 25 February 2014; accepted: 2 May 2014

Introduction The majority of work within cell biology over the past These peptide-based hydrogels are typically fabricated few decades has been carried out in vitro on two-dimen- from short amino acid sequences that spontaneously self- sional (2D) systems, such as tissue culture plastic (TCP).1 assemble into -sheet- or α-helix-rich fibres. These go on There has been a move more recently to develop and use to self-associate to form a self-supporting matrix that three-dimensional (3D) scaffolds for the study of cells as resembles natural extracellular matrix (ECM) when above these materials better reflect their in vivo environment and hence give more reliable results.2–4 For example, the adhesion and migration behaviour of mammalian cancer 1 School of Chemical Engineering and Analytical Science, The University cells displayed different behavioural patterns within 3D of Manchester, Manchester, UK materials in comparison with cells cultured in 2D on flat- 2 Manchester Institute of Biotechnology, The University of Manchester, tened substrates.5–7 Several classes of materials have been Manchester, UK 3School of Materials, The University of Manchester, Manchester, UK tested and used for the fabrication of 3D matrices for host- ing and culturing cells, including polymers, protein and Corresponding authors: peptide-based materials. The latter is currently attracting Julie E Gough, School of Materials, The University of Manchester, significant attention as they inherently offer many of the Grosvenor Street, Manchester M13 9PL, UK. required properties of a biomaterial, including immuno- Email: [email protected] compatibility, biodegradability, easy handling, tunable Aline F Miller, Manchester Institute of Biotechnology, The University of structure and mechanical properties and ease of introduc- Manchester, 131 Princess Street, Manchester M1 7DN, UK. ing functionality.3,4,8 Email: [email protected]

Creative Commons CC-BY-NC: This article is distributed under the terms of the Creative Commons Attribution-NonCommercial 3.0 License (http://www.creativecommons.org/licenses/by-nc/3.0/) which permits non-commercial use, reproduction and distribution of the work without further permission provided the original work is attributed as specified on the SAGE and Open Access page (http://www.uk.sagepub.com/aboutus/openaccess.htm).Downloaded from tej.sagepub.com at The University of Manchester Library on July 7, 2014 2 Journal of Tissue Engineering

Table 1. FEFEFKFK gel concentrations and buffer conditions used for the preparation of hydrogels ready for the 3D culture of human osteoblast cells.

Gel (wt%) Peptide weight (mg) Water volume (µL) 1 M NaOH (µL) PBS (µL) 2 24 800 70 100 3 36 800 105 100 4 48 800 140 100 5 60 800 175 100

3D: three-dimensional; PBS: phosphate-buffered saline. a critical concentration.3,4,9 Consequently, many studies hydrophobic, and E and K are negatively and positively have focussed on the development of these scaffolds for charged, respectively, at physiological pH.20 FEFEFKFK the culture of several cell types, including bone-forming self-assembles into an antiparallel -sheet secondary struc- cells for bone tissue regeneration. For example, Zhang et ture, where all Fs are placed on the same side of the -sheet al. developed the ionic-complementary peptide sequence, while the charged amino acids are located on the other side. RADARADARADARADA (RADA-16), where R, A and This peptide subsequently forms a network of -sheet-rich D are arginine, alanine and aspartic acid, respectively. nanofibres, which branch to form a self-supporting hydro- They showed that this sequence forms extended and gel depending on the pH, temperature and ionic strength of ordered -structured nanofibres that form self-supporting the media.20,21 Recently, we demonstrated that hydrogels hydrogels. This system has since been used to support the prepared using FEFEFKFK were able to encapsulate 3D culture of mammalian cells including human osteo- bovine chondrocyte cells within 3D and the matrix sup- blasts (HOBs).10,11 Pochan and Schneider studied ported their viability and proliferation.21 -sheet hairpin peptides, such as VKVKVKVKVDPLP Here, we extend this work and explore the suitability of

TKVKVKVKV-NH2 (MAX 1), where V is valine, K is our FEFEFKFK-based hydrogel to act as a scaffold for the lysine and P is proline. This amphiphilic peptide self- 3D culture of primary HOB cells and the subsequent deposi- assembles into -hairpin secondary structures in response tion of minerals key for the formation of bone. First, we to a change in environmental conditions, such as salt con- explore the optimal mechanical strength of hydrogel centration. This leads to the formation of stable fibrous required for cell viability. Second, we explore cell viability hydrogels capable of supporting the culture of various over longer culture times and proliferation, as well as the cells, such as mesenchymal stem cells, osteoblasts and production of ECM proteins within the gel. Finally, we murine fibroblasts.4,12,13 Beniash et al.14 developed sur- monitor the capability of the gel to facilitate the bone miner- factant-based amphiphilic peptide hydrogels, which alisation process. encapsulated and supported the proliferation of HOBs, making this system suitable for bone tissue engineering applications. One other -sheet forming peptide – Ace- Materials and methods QQRFEWEFEQQ-NH (P-11) (Q, F, E and W are glu- 2 Hydrogel preparation tamine, phenylalanine, glutamic acid and tryptophan, respectively) – developed by Aggeli et al.15 has been FEFEFKFK peptide (>95% purity) was purchased from shown to facilitate teeth enamel re-mineralisation via Biomatik Corporation, Canada. Peptide solutions were pre- hydroxyapatite nucleation. pared initially at 2, 3, 4 or 5 wt% by dissolving the peptide

It is widely known that some bone diseases such as powder in 800 µL of doubly distilled water (ddH2O) (see osteoporosis and periodontal disease induce a progressive Table 1). They were subsequently vortexed and centrifuged and significant loss of bone. Periodontitis, in particular, (4000 r/min), before placing in an oven at 90°C for 2 h to can cause the loss of teeth at late stages.16,17 Currently, ensure complete dissolution. To neutralise the sample, 1 M deep scaling and root planning (debridement) is the stand- NaOH and Dulbecco’s phosphate-buffered saline (DPBS) ard method to treat this pathology.18 Nonetheless, when a were added, respectively (Table 1). This induced gelation. significant quantity of bone is lost, bone grafting is needed, The resulting transparent gel was again vortexed and cen- but this unfortunately comes with several drawbacks, such trifuged (5000 r/min) and placed back into the oven at 90°C as surgical complications, residual pain and increased risk for 12–24 h before being cooled at room temperature (RT) of infection. In addition, some osteogenic factors are not to ensure formation of a homogeneous hydrogel. completely viable after transplantation, and this has a neg- 19 ative effect on any bone regeneration. Oscillatory rheology Here, we wish to build on our groups’ previous work on the self-assembling behaviour and application of the pep- The viscoelastic behaviour of the FEFEFKFK hydrogels at tide FEFEFKFK. In this octapeptide, F is non-charged and days 0, 1, 3, 7 and 14 of cell culture was determined using

Downloaded from tej.sagepub.com at The University of Manchester Library on July 7, 2014 Castillo et al. 3 an ARG2 rheometer with a 20-mm parallel plate geometry. were viewed under a Leica TCS SP5 confocal microscope. Samples with and without cells were run in parallel, where For quantitative cell viability, gels containing cells were the latter acted as the control. In each case, ~150 µL of gel rinse twice in PBS, diluted using trypsin was pipetted onto the bottom plate and left for 10 min to (0.05%)–EDTA·4Na (0.53 mM). Subsequently, fresh cell equilibrate before recording the elastic (G) and viscous culture media were added and mixed before transferring (G″) moduli as function strain (0.01%–100%) at 1 Hz and the cell suspension into a micro-centrifuge tube. Here, it oscillatory frequency (0.1–100 Hz) at 1% strain. All sam- was diluted 1:1 in trypan blue dye to enable counting the ples were maintained at physiological temperature (37°C) cells using a haemocytometer. Cell counts and optical using a Peltier stage and solvent evaporation minimised images were obtained using an inverted microscope (Leica using a solvent trap. DM IL) and a SPOT insight camera (model 3.2.0; Diagnostic Instruments Inc., Michigan, USA). Cell culture Osteoblast proliferation HOBs (PromoCell, Heidelberg, Germany) were grown and maintained under standard cell culture conditions in Osteoblast proliferation was assessed using PicoGreen® Dulbecco’s modified Eagle’s medium (DMEM) supple- dsDNA assay (Life Technologies, Carslbad, CA, USA). mented with 10% foetal bovine serum (FBS), 1% penicillin, After gels were rinsed twice with DPBS, they were resus- 1% streptomycin and 50 µg/mL ascorbic acid. At 80%–90% pended in 500 µL of cold lysis buffer (200 mM Tris–HCl, of confluence, cells were subcultured using 0.05% 20 mM EDTA/ddH2O/1% Triton-X100) for 25 min. To trypsin–0.53 mM ethylenediaminetetraacetic acid (EDTA) ensure complete lysis, samples were vortexed vigorously 4Na solution (Gibco–Invitrogen, UK) and transferred to a and subject to three freeze–thaw cycles. Thereafter, sam- Falcon tube to be centrifuged to remove the trypsin solution. ples were vortexed again and 100 µL gel–cell suspension The cell pellet was then resuspended in fresh medium and was plated into black 96-well plates, where 100 µL of a adjusted to the required cell concentration. working solution of Quant-iT PicoGreen reagent was added. 3D cell culture within FEFEFKFK hydrogel The samples were incubated for 2–5 min at RT. Fluorescence readings were obtained using a plate reader Gels were sterilised by exposure to ultraviolet (UV) radia- (FLUOstar OPTIMA; BMG LABTECH) at wavelengths tion for 30 min, and 200 µL of cell suspension was then of 435 nm (excitation) and 529 nm (emission). A standard pipetted on top and gently mixed by stirring with the pipette curve was determined using calf thymus DNA in serial tip before gently pipetting up and down to create a homoge- dilutions in 1% Triton X. A blank gel was used to correct neous gel/cell suspension. A volume of 250 µL of gel con- the background absorbance and the assay was performed taining 1.5 × 105 cells was transferred into each well in a in triplicate. 12-well cell culture insert (ThinCert™ Greiner Bio-One) for all experiments except rheology, where 500 µL of gel con- Immunocytochemistry taining 2.5 × 105 cells was used. In addition, 1 × 104 cells/ cm2 were pipetted directly onto a tissue culture plate. The The presence of collagen type I (col-I) inside the gel was left to set for 10 min at 37°C. Gels were then placed FEFEFKFK hydrogels was determined visually using a in an incubator at 37°C in a 95% humidified atmosphere standard immunocytochemistry method after 7 and 14 days

(20% O2) with 5% CO2 and fresh media changes were of culture. After 24 h of culture, each sample was prepared repeated five times at 20-min intervals to stabilise the pH at by culture in osteogenic media containing 10−5 mM dexa- 7.2. Thereafter, the media were changed every 2 days to aid methasone (D4902-16; Sigma–Aldrich Co.), 10 mM cell growth within the gel. -glycerophosphate (G9422-100G; Sigma–Aldrich Co.) and 2.83 × 10−7 mM ascorbic acid (A8960-5G; Sigma- Cell viability Aldrich Co., St. Louis, MO, USA). At each time-point, the gels were fixed with 3% paraformaldehyde (PFA) for 30 Cell viability was tested qualitatively using a live/dead min at RT. Thereafter, samples were permeabilised using assay (Invitrogen) and quantitatively using standard cell Triton X-100 at 0.05% in DPBS for 15 min at RT. Samples counting (haemocytometer). For the live/dead assay, 1.5 were subsequently blocked with 1% bovine serum albumin mL of PBS containing 2.5 µL of 4 µM ethidium homodi- (BSA) for 40–50 min at RT and incubated for 1 h at RT mer-1 (EthD-1) assay solution and 1.5 µL of 2 µM calcein with a primary antibody (pAb) (diluted in 1% BSA) rabbit AM assay solution was prepared. The live/dead assay polyclonal to col-I (Abcam, UK), at a ratio stipulated by the solution was pipetted on top of each hydrogel and then manufacturer (1:250). Subsequently, the samples were incubated under standard cell culture conditions for 20 incubated for 1 h at RT in the dark with a goat anti-rabbit min. The staining solution was then removed and samples IgG-Alexa Fluor 594 (Abcam) as secondary antibody (sAb)

Downloaded from tej.sagepub.com at The University of Manchester Library on July 7, 2014 4 Journal of Tissue Engineering along with Alexa fluor 488 phalloidin to target col-I and vortexed and pipetted into a 96-well plate. Subsequently, a F-actin, respectively. The samples were rinsed in cycles of working anti-OST-HRP solution (100 µL) was added to 5 × 5–10 min, with DPBS washings between each step. each well, the plate covered and incubated for 2 h at RT. Samples were mounted on glass slides using ProLong Each well was then rinsed three times with a washing solu- Antifade Reagent (Invitrogen), and images were obtained tion before adding 100 µL of a chromogen solution (tetra- using a Leica TCS SP5 confocal microscope. methylbenzidine) and incubating for 30 min at RT in the dark. The reaction was stopped by adding 100 µL of a stop Quantification of bone mineralisation proteins solution (1 N HCl). Absorbance measurements were obtained within 1 h at 450 nm, using a plate reader (Tecan Bone formation was monitored by quantifying mineralisa- Infinite M200). Two independent assays were performed tion proteins that were present within each gel at days 7 in triplicate. and 14. At each time-point, gels were rinsed twice with

DPBS and resuspended in cold distilled water to ensure gel Mineralisation activity dissolution. Cells within these samples were subjected to three freeze–thaw cycles which led to cell lysis. Three Mineralisation activity was evaluated using Alizarin Red markers were selected and procedures are outlined below. staining. At days 7 and 14, the cells were fixed with 70%

In each case, a blank gel was used to correct the back- ethanol for 30 min, rinsed three times with ddH2O and then ground absorbance. stained with 40 nM Alizarin Red at RT in the dark for 45 min. Samples were then rinsed eight times with PBS. For Quantification of collagen optical density measurements, 10% cetylpyridinium chloride (CPC) was dissolved in 10 mM sodium phosphate and Collagen production was determined using a total collagen 1 mL of this was added to each gel before incubating at RT assay (QuickZyme Biosciences, Park, Leiden, The for 30 min. A volume of 200 µL of each solution was plated Netherlands). After resuspending and lysing, samples were into a clear, flat-bottomed 96-well plate (Nunc, UK) and vortexed and hydrolysed over 20 h at 95°C with 100 µL 12 the absorbance at 562 nm determined using a Labsystems M HCl. Samples were subsequently diluted 1:1 in 4 M HCl Multiskan Ascent plate reader. The assay was performed in and pipetted into a 96-well plate, before adding 75 µL of triplicate. an assay buffer and incubating for 20 min at RT. Following this, 75 µL of a detection reagent was added and the sam- Statistical analysis ple transferred into the oven (60°C) for 1 h. Absorbance measurements were obtained at 570 nm, using a plate Statistical significance in rheology, the PicoGreen, col-I, reader (Tecan Infinite M200). Three independent assays alkphos activity and OCN quantification assays, was were undertaken in triplicate. determined using one-way analysis of variance (ANOVA) followed by post hoc comparisons (Tukey’s method). Quantification of alkaline phosphatase activity Results and discussion Alkaline phosphatase (alkphos) activity was monitored using a colorimetric alkphos and peroxidase substrate Initial experiments were undertaken with the aim of opti- detection system (Sigma–Aldrich Co.). A volume of 20 µL mising the concentration of peptide hydrogel needed for of the cell lysis was added to transparent 96-well plates the encapsulation, maintenance and proliferation of HOBs. together with 200 µL of p-nitrophenyl phosphate (pNPP) It is known that varying the peptide concentration influ- solution (1 mg/mL pNPP, 0.2 M Tris buffer in 5 mL ences the mesh size of the network formed and also the ddH2O) (SIGMAFAST™ pNPP Tablets, N1891-50SET; mechanical strength of the hydrogel, both of which influ- Sigma–Aldrich Co.), and the following reaction was ence the flow of nutrients and waste products and also cell stopped with 3 M NaOH. The absorbance of samples was behaviour. To this end, four different concentrations of measured using a plate reader (Labsystems Multiskan FEFEFKFK peptide hydrogel were prepared (2, 3, 4 and 5 Ascent; Thermo Scientific, UK) at 405 nm every 30 s for wt%) in cell culture media and their elastic properties 30 min. Two independent assays were performed in determined. Each concentration formed a stable, self- triplicate. supporting hydrogel at pH 7 where the elastic modulus, G, increased with increasing concentration: 6.2 ± 1.2, 7.8 ± Quantification of osteocalcin 0.5, 13.3 ± 3.5 and 24.6 ± 3.5 kPa for the 2, 3, 4 and 5 wt% samples, respectively. This is in agreement with our previ- Osteocalcin (OCN) production was determined using a ous work, and others, where the density of the fibre net- Human Osteocalcin ELISA Kit (Invitrogen–Life work increases with concentration.20–22 The magnitude of Technologies). After the cells were lysed, samples were the G values is higher than those obtained when

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Castillo et al. 5

Gel (3wt%) Gel (4wt%) Gel (5wt%) Day 1 Day 1 Day 1

Day 7 Day 7 Day 7

Figure 1. Live/dead imaging representing living cells (green) and dead cells (red) within FEFEFKFK hydrogel prepared at distinct peptides concentrations (magnification = 10×, scale bar = 500 µm). the peptide was dissolved in pure water (in the order of with only a very few dead cells (Figure 2(a)–(c)). 10–100 s Pa for the concentration range studied here). This Importantly, cells were also incorporated throughout the is due to the salt within the cell culture media and PBS peptide hydrogel, as evidenced by the even cell distribu- screening the charges of the peptide, leading to an enhance- tion in the Z-direction shown in the confocal image in ment of the hydrophobic forces and hydrogen bonding. Figure 2(d) (~250 µm thickness). Such entrapment and This increases fibre aggregation and association, which viability is similar to our previous work with bovine chon- increases the strength of the 3D network.21 drocytes21 and demonstrates the hydrogels can support The effect of the changing mechanical properties on cells within 3D. The optical micrographs in Figure 2(e)– cell viability was subsequently explored by seeding each (g) reveal that the HOBs typically acquired a rounded mor- gel with 1.5 × 105 cells and monitoring cell fate via a live/ phology during their 2 weeks in culture within the peptide dead assay over 7 days (Figure 1). When cells were incor- hydrogel. When HOBs are present within natural bone, porated within the 2 wt% sample, the resulting gels were and their synthesis activity is induced, they tend to acquire rather weak and not stable, hence it was decided this sys- a cuboidal/plump 3D morphology. This is in contrast to the tem was not suitable for further study. It was clear that rather flattened morphology commonly seen for bone lin- osteoblast viability was favoured when cells were cultured ing cells, and also when the cells are grown on many 2D within 3 wt% gel where the majority of cells were alive substrates.23,24 This suggests that the HOBs within our (green) with only a few dead cells (red), in comparison to peptide hydrogel adopt an in vivo–like morphology, sug- a majority of dead cells in both 4 and 5 wt% samples gesting the peptide hydrogel is mimicking the 3D in vivo (Figure 1). It is postulated that this is due to the higher matrix environment. Such behaviour has been observed density of the fibre network which leads to smaller gel previously when HOBs have been cultured within syn- porosity, which perhaps is limiting the diffusion of nutri- thetic poly(ethylene glycol) (PEG)22 and natural alginate ents, the exchange of gases and the output of cellular scaffolds.25 It was also observed that HOBs formed some waste, or simply because the cells prefer to interact with clusters throughout the FEFEFKFK hydrogel. These are the softer network. Consequently, 3 wt% gels were selected also naturally present during osteoid synthesis.24 To quan- for further cell studies. tify the viability of HOBs within the peptide gels over their Cell viability within 3 wt% FEFEFKFK gel samples 2 weeks in culture, the percentage of viable cells was was extended up to 14 days, and fluorescent micrographs determined using the trypan blue exclusion quantification of the live/dead assay and optical micrographs monitoring method. It is clear from these data presented in Figure 3 cell morphology over time are given in Figure 2. It is evi- that the percentage viability of HOBs increases over the dent from live/dead screening that the majority of cells whole 14 days. This increase is most significant over the within each gel after 1, 7 and 14 days in culture were alive, first 7 days.

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6 Journal of Tissue Engineering

Day 1 Day 1

(a) (e)

Day 7 Day 7

(b) (f)

Day 14 Day 14

G (c) G(g)

Day 14

(d)

Figure 2. (a-c) Live/dead imaging of HOB viability within 3 wt.% FEFEFKFK hydrogels during 14 days of culture. Living and dead cells are represented by green and red fluorescence respectively. Magnification=10×, scale bars 500 mm). (d) Axial (Z-axis) imaging show homogeneous HOB dispersion through the gel. (e-g) Optical micrographs (grey) show cell morphology within the gels. Magnification=10×, scale bars represent 100 mm.d. Axial (Z-axis) imaging show homogeneous HOB dispersion through the gel. HOB: human osteoblast.

To identify the rate of proliferation of HOBs within the alginate gels.25 A decrease in DNA content over longer time peptide hydrogel, the DNA content was quantified over periods has also been reported for HOBs when cultured time using a PicoGreen assay. It is clear from the data pre- within, for example, PEG hydrogels. These possess higher sented in Figure 4(a) that the DNA content within the gel mechanical properties (10–300 kPa) in comparison to the increased significantly over the first 3 days of culture, and FEFEFKFK hydrogel studied here.22,26 It is known that cal- then remained steady, within experimental error, up to day cified bone possesses high stiffness,22,27 which is superior 14. Such an increase in cell proliferation over the first few to the strength of a typical hydrogel system,21,22,25 therefore days has been seen for HOBs encapsulated within different there is a possibility that the relatively low mechanical types of hydrogels such as peptide amphiphile14 and strength provided by our peptide hydrogel might influence

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as hormones and other local factors, which include FEFEFKFK-gel cytokines and growth factors released after bone resorp- 100 tion occurs. The role of osteocytes sensing micro- * alterations or mechanical loads on bone tissue is also very 75 important to activate HOBs. Finally, the autocrine activation via bone-specific ECM proteins is fundamental to 24,28,29 50 induce the proliferation of osteoblasts. The rate of proliferation of HOBs within 3D culture systems is, there- 25 fore, complex and clearly dependent on cell culture conditions used, such as type of scaffold, type of cells and/or the cell densities used. Here, we demonstrated that our system 0 1 7 14 is able to support the viability and proliferation of HOBs, Time (Days) without any additional cell proliferative factor. Consequently, the next step is to test the ability of cells to Figure 3. Quantification of cell viability as a function of produce key proteins for bone formation, including colla- culture time. Data are reported as mean values ± standard gen-I (col-I). Figure 5 shows the fluorescence and immu- deviation for n = 3. nocytochemistry images of stained gels for the *p < 0.05 when comparing cell numbers between time points. identification of F-actin that defines the cell architecture, and col-I, at days 7 and 14. It is clear that there is signifi- the rate of proliferation of HOBs (i.e. decrease) over pro- cant production of col-I inside the cell cytoplasm over the longed periods of time. This is unlikely in this case, how- first 7 days of culture. There is also some evidence of ever, given the cells were not viable within the stiffer 4 and extracellular col-I staining, albeit with less intensity. After 5 wt% gels. Furthermore, gels are dynamic systems and as 14 days in culture, however, a high concentration of col-I such are prone to expose less surface area to cells over time. was detected, not only inside the cells but also surrounding This might also impact on the extent and rate of cell prolif- the cells, particularly in areas of the gel where cells formed eration. The DNA content quantified for HOBs cultured in clusters. The high concentration of col-I suggests that cells 2D on TCP is given in Figure 4(b). A similar pattern is evi- are functional within the gel and consequently synthesise dent here; a significant increase over the first few days proteins to remodel their niche. This confirms that HOBs which then remains constant over longer times. This behav- can survive in the in vitro hydrogel and moreover produce iour is typical for 2D cell culture and is likely to be due to ECM proteins that are important for their survival and over-confluence at later times (Figure 4(b)). bone formation. It is also important to consider that HOBs are special- To corroborate the visual trend observed for the produc- ised cells, and as such, their predominant role is the secre- tion of col-I from the immunocytochemistry staining tion of proteins. It is believed that their activation and experiments, the quantity of protein present was deter- proliferation are triggered mainly by systemic factors such mined (Figure 6(a)) using a quantitative colorimetric

(a) (b) FEFEFKFK-Gel TCP

1.0x10-2 1.0x10-2 ns *** 7.5x10-3 7.5x10-3 ns

5.0x10-3 * 5.0x10-3

2.5x10-3 2.5x10-3

0 0 3 7 14 3 7 14 Time (days) Time (Days)

Figure 4. Quantification of DNA content from HOBs cultured (a) in 3D within FEFEFKFK peptide hydrogel and (b) in 2D on TCP. Data are reported as mean ± standard deviation for n = 3. HOB: human osteoblast. *p < 0.05; ***p < 0.001; ns denotes no significant difference when comparing cell number between time points.

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F-actin Type I Collagen Merge Day 7 Day 7 Day 7

Day 14 Day 14 Day 14

Figure 5. Fluorescence micrographs of actin cytoskeleton staining (green) and collagen I production (red) by HOBs cultured in 3D within 3 wt% FEFEFKFK hydrogel over 14 days (magnification = 10×, scale bar = 500 µm). HOB: human osteoblast.

method. Total collagen was found to increase between instead we are exploring our gels’ ability to host HOBs days 7 and 14, which fits well with qualitative results and the ability of the cells in this environment to synthe- observed in Figure 5. Such an increase in collagen produc- sise and release in situ bone-forming proteins, and conse- tion suggests that the cells might also be producing other quently deposit calcium phosphate, for non-load bearing ECM proteins involved in the process of bone formation, applications. The effect of HOBs and the production of such as alkphos and OCN. These have, therefore, also been matrix protein on the stiffness of the peptide hydrogel quantified and data reveal a similar pattern as collagen was monitored using oscillatory rheology. As a control, (Figure 6(b) and (c), respectively). The data for the pro- we also studied the stiffness of a gel without cells that duction of all three ECM proteins for the 2D culture of had identical media changes over time. The results are cells on TCP cultures were also determined and showed given in Figure 7 and indicate that the elasticity of the gel similar increasing production over time, as expected. Such network without cells decreased from 8 to 3.2 ± 0.6 kPa ECM protein production within the FEFEFKFK hydrogel over 14 days in culture media. This drop is likely due to is a significant finding, as it highlights it as a promising the natural dissolution of gel that inevitably occurs dur- scaffold for the culture of bone-forming cells. It also dem- ing media changes, given our peptide hydrogel is a physi- onstrates the gels’ potential for the culture of other types of cal gel. In contrast, the elasticity of the FEFEFKFK mammalian cells. The pattern of ECM production exhib- hydrogel with cells increased from ~5.4 to 22.6 ± 1.2 kPa ited by the HOBs is similar to that observed by other over the 14 days of culture. This differing behaviour is groups using different types of scaffolds, including poly(L- most likely to be due to the cells remodelling the gel and lactide) (PLLA) copolymers, RADA-16, PEG and gelatin the deposition of ECM proteins. A similar trend has been gels.10,22,30,31 reported by other authors for FEFEFKFK hydrogel21 and It is known that cell responses can be influenced by also the commercial PuraMatrix32 systems for the culture the characteristics of the niche, including stiffness, topog- of chondrocytes. raphy and roughness.27 The natural ECM matrix that Bone mineralisation is the last phase in the bone forma- hosts bone in vivo has a high content of non-mineralised tion process, where several proteins such as OCN, osteo- collagen and has a compressive modulus stiffness of pontin and bone sialoprotein are expressed.33 It is known ~100 kPa, which increases to ~1000 kPa once osteoid that during this process, osteoblasts release vesicles con- mineralises.22,27 Both these values are significantly higher taining minerals such as calcium and phosphate, which are than the elastic modulus reported for peptide hydro- essential for healthy bone formation.23,33,34 The increasing gels.11,14,21 Nevertheless, we are not aiming to match the presence of OCN during culture within the hydrogels has in vivo mechanical strength in our FEFEFKFK hydrogel, been monitored and the results are shown in Figure 6. We

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Castillo et al. 9

(a) FEFEFKFK-gel (b) *** TCP 2.0x10-3 1.0x10-3 ns

1.5x10-3 7.5x10-4

1.0x10-3 5.0x10-4

5.0x10-4 2.5x10-4

0 0 7 14 7 14 Time (Days) Time (Days) (c) (d) TCP FEFEFKFK-gel 1.0x10-2 2.5x10-3 ***

2.0x10-3 7.5x10-3 *** 1.5x10-3 5.0x10-3 1.0x10-3

-3 2.5x10 5.0x10-4

0 0 7 14 7 14 Time (Days) Time (Days) (f) (e) FEFEFKFK-gel TCP 2.0x10-3 *** 1.0x10-3

8.0x10-4 1.5x10-3 ns 6.0x10-4 1.0x10-3 4.0x10-4 5.0x10-4 2.0x10-4

0 0 7 14 7 14 Time (Days) Time (Days)

Figure 6. Total collagen production within (a) gel and (b) TCP; alkaline phosphatase activity within (c) gel and (d) TCP and osteocalcin production inside (e) gel and (f) TCP over 14 days in culture. Data are reported as mean ± standard deviation for n = 3 (col-I) and n = 2 (alkphos and OCN). Alkphos: alkaline phosphatase; OCN: osteocalcin. ***p < 0.001; ns denotes no significant difference for comparison in the protein production between days 7 and 14. went on to monitor the presence of any calcium deposits presence of calcium and consequently confirms calcium using Alizarin Red staining, both qualitatively from imag- phosphate formation. It is clear that discrete calcium ing and quantitatively using absorbance at 562 nm. The deposits are present within the gels after the first 7 days of photographs of the stained samples are shown in Figure culture, with even stronger staining intensity observed 8(a) where the intensity of the red stain indicates the after 14 days (Figure 8(a)). Staining of 2D cell culture on

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10 Journal of Tissue Engineering

FEFEFKFK-gel (-Cells) FEFEFKFK-gel (+Cells)

** ** ** 100000 ** ** ** ** 10000

1000

100 10

1 1 3 7 14 1 3 7 14 Time (Days) Figure 7. Elastic modulus, G, of FEFEFKFK hydrogels over 14 days in cell culture media conditions, both with and without the presence of HOBs. Data are reported as mean ± standard deviation for n = 3. HOB: human osteoblast. **p < 0.05, for comparison of the G values.

(a) FEFEFKFK-gel 2-D culture (TCP)

Day 7 Day 7

Day 14 Day 14

(b) FEFEFKFK-gel (c) TCP -6 1.0x10-5 5.0x10 ** 4.0x10-6 7.5x10-6 3.0x10-6 * 5.0x10-6 2.0x10-6 2.5x10-6 1.0x10-6 0 0 7 14 7 14 Time (Days) Time (Days )

Figure 8. (a) Alizarin red staining images showing the presence of calcium phosphate deposits in red both in FEFEFKFK hydrogel, and on 2D culture (TCP). Quantification assay of calcium deposits within (b) gel and (c) TCP, using Alizarin Red staining via extraction with 10% CPC. Data are reported as mean ± standard deviation for n = 3. 2D: two-dimensional; CPC: cetylpyridinium chloride. *p < 0.01 and **p < 0.05 for comparison of the calcium ions’ detection between time points.

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Castillo et al. 11

TCP was also observed, but the signal was not as intense as 4. Kretsinger JK, Haines LA, Ozbas B, et al. Cytocompatibility in the 3D studies. This was expected given there is only a of self-assembled beta-hairpin peptide hydrogel surfaces. thin film present for the 2D experiments. Quantitative Biomaterials 2005; 26: 5177–5186. results are given in Figure 8(b) where the absorbance of 5. Weaver VM, Petersen OW, Wang F, et al. Reversion of the specific dye was recorded at 562 nm over time. It is the malignant phenotype of human breast cells in three- dimensional culture and in vivo by integrin blocking anti- clear that in both 3D and 2D cases, the mineralisation bodies. J Cell Biol 1997; 137: 231–245. assay confirms a significant increase in mineralisation 6. Webb DJ and Horwitz AF. New dimensions in cell migra- over time within the gels (Figure 8(b) and (c), respec- tion. Nat Cell Biol 2003; 5: 690–692. tively). The intensity of absorbance is an order of magni- 7. Cukierman E, Pankov R, Stevens DR, et al. Taking cell- tude higher for the sample where cells were incorporated matrix adhesions to the third dimension. Science 2001; 294: throughout the hydrogel. This indicates that the 1708–1712. FEFEFKFK hydrogel not only supports, maintains and 8. Maude S, Ingham E and Aggeli A. Biomimetic self- allows the proliferation of primary HOBs but also allows assembling peptides as scaffolds for soft tissue engineering. the production of ECM proteins and the deposition of cal- Nanomedicine (Lond) 2013; 8: 823–847. cium phosphate and hence shows bone formation. Other 9. Cavalli S, Albericio F and Kros A. Amphiphilic peptides work on different systems report similar increases in the and their cross-disciplinary role as building blocks for nano- science. Chem Soc Rev 2010; 39: 241–263. deposition of calcium over time that ranges from 7 to 21 10. Zhang S, Holmes TC, DiPersio CM, et al. Self- 22,30,35,36 days in culture. complementary oligopeptide matrices support mammalian cell attachment. Biomaterials 1995; 16: 1385–93. Conclusion 11. Cunha C, Panseri S, Villa O, et al. 3D culture of adult mouse neural stem cells within functionalized self-assembling pep- Here, we have demonstrated that a 3 wt% FEFEFKFK tide scaffolds. Int J Nanomedicine 2011; 6: 943–955. fibrillar hydrogel is an excellent host for the 3D culture of 12. Haines-Butterick L, Rajagopal K, Branco M, et al. primary HOB cells. Cell viability and proliferation are Controlling hydrogelation kinetics by peptide design for maintained over 14 days within the gel (maximum length three-dimensional encapsulation and injectable delivery of of time tested), as is the production of bone matrix pro- cells. P Natl Acad Sci USA 2007; 104: 7791–7796. teins. Mineralisation within the hydrogel has been shown 13. Yan C, Altunbas A, Yucel T, et al. Injectable solid hydrogel: mechanism of shear-thinning and immediate recovery and correlates nicely with an increase in the mechanical of injectable -hairpin peptide hydrogels. Soft Matter 2010; properties of the gel over the 14 days in culture. These 6: 5143–5156. ionic-complementary peptide gels have potential for use as 14. Beniash E, Hartgerink JD, Stendahl JC, et al. Self- 3D bone regeneration scaffolds and as in vitro osteoblast assembling peptide amphiphile nanofiber matrices for cell culture systems. entrapment. Acta Biomater 2005; 1: 387–397. 15. Kirkham J, Firth A, Vernals D, et al. Self-assembling pep- Acknowledgements tide scaffolds promote enamel remineralization. J Dent Res 2007; 86: 426–430. The authors would like to thank the members of the Polymers 16. Silva N, Dutzan N, Hernandez M, et al. Characterization and Peptides, and Biomaterials and Tissue Engineering groups of progressive periodontal lesions in chronic periodontitis for helpful discussions. patients: levels of chemokines, cytokines, matrix metalloproteinase-13, periodontal pathogens and inflammatory Declaration of conflicting interest cells. J Clin Periodontol 2008; 35: 206–214. The authors declare that there is no conflict of interest. 17. Graves DT, Oates T and Garlet GP. Review of osteoimmunology and the host response in endodontic and periodontal Funding lesions. J Oral Microbiol 2011;3. 18. Ryan ME. Nonsurgical approaches for the treatment of peri- LCD acknowledges a PhD scholarship grant from CONACyT, odontal diseases. Dent Clin North Am 2005; 49: 611–636, vii. Mexico 19. Giannoudis PV, Dinopoulos H and Tsiridis E. Bone substitutes: an update. Injury 2005; 36(Suppl. 3): S20–S27. References 20. Saiani A, Mohammed A, Frielinghaus H, et al. Self- 1. Alberts B, Johnson A, Lewis J, et al. 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12 Journal of Tissue Engineering

23. Clarke B. Normal bone anatomy and physiology. Clin J Am 31. Idris SB, Arvidson K, Plikk P, et al. Polyester copoly- Soc Nephrol 2008; 3: S131–S139. mer scaffolds enhance expression of bone markers in 24. Nakamura H. Morphology, function, and differentiation of osteoblast-like cells. J Biomed Mater Res A 2010; 94: bone cells. J Hard Tissue Biol 2007; 16: 15–22. 631–639. 25. Place ES, Rojo L, Gentleman E, et al. Strontium- and zinc- 32. Maher SA, Mauck RL, Rackwitz L, et al. A nanofibrous alginate hydrogels for bone tissue engineering. Tissue Eng cell-seeded hydrogel promotes integration in a cartilage gap Part A 2011; 17: 2713–2722. model. J Tissue Eng Regen Med 2010; 4: 25–29. 26. Benoit DS, Durney AR and Anseth KS. Manipulations in 33. Neve A, Corrado A and Cantatore FP. Osteoblast physiol- hydrogel degradation behavior enhance osteoblast func- ogy in normal and pathological conditions. Cell Tissue Res tion and mineralized tissue formation. Tissue Eng 2006; 12: 2011; 343: 289–302. 1663–1673. 34. Boonrungsiman S, Gentleman E, Carzaniga R, et al. The role 27. Engler AJ, Sen S, Sweeney HL, et al. Matrix elasticity of intracellular calcium phosphate in osteoblast-mediated bone directs stem cell lineage specification. Cell 2006; 126: apatite formation. P Natl Acad Sci USA 2012; 109: 14170–14175. 677–689. 35. Clarke MS, Sundaresan A, Vanderburg CR, et al. A three- 28. Datta HK, Ng WF, Walker JA, et al. The cell biology of dimensional tissue culture model of bone formation utiliz- bone metabolism. J Clin Pathol 2008; 61: 577–587. ing rotational co-culture of human adult osteoblasts and 29. Hadjidakis DJ and Androulakis II. Bone remodeling. Ann N osteoclasts. Acta Biomater 2013; 9: 7908–7916. Y Acad Sci 2006; 1092: 385–396. 36. Guo XY, Gough JE, Xiao P, et al. Fabrication of nano- 30. Sachar A, Strom TA, Serrano MJ, et al. Osteoblasts structured hydroxyapatite and analysis of human osteo- responses to three-dimensional nanofibrous gelatin scaf- blastic cellular response. J Biomed Mater Res A 2007; 82: folds. J Biomed Mater Res A 2012; 100: 3029–3041. 1022–1032.

Downloaded from tej.sagepub.com at The University of Manchester Library on July 7, 2014 CHAPTER THREE

Functional peptide hydrogels for bone formation applications

Luis A. Castillo Díaz, Alberto Saiani, Julie E. Gough, and Aline F. Miller

All authors contributed to the realisation of this manuscript.

71 Functional peptide hydrogels for bone formation applications

Luis A Castillo Díaz,1,2 Alberto Saiani,2,3 Julie E Gough3* and Aline F Miller1,2* .

1 School of Chemical Engineering and Analytical Science, The University of Manchester (UK).

2 Manchester Institute of Biotechnology, The University of Manchester (UK).

2 School of Materials, The University of Manchester (UK)

Corresponding authors:

Julie E Gough, School of Materials, The University of Manchester, Grosvenor Street, Manchester M13 9PL, UK. Email: [email protected]

72 Abstract

Tissue engineering is still seeking to develop a biomaterial capable of effectively regenerating several tissues including bone. In dentistry for example, the resorption of alveolar bone caused by dental infection or periodontitis, is very often accompanied by tooth loss. Bone grafting, which is the gold standard to treat the resorption of alveolar bone, has not overcome concerns related to defective bone formation, morbidity, etc. Peptide hydrogels made of nano-fibre networks, mimic the three- dimensional (3-D) architecture to that displayed by the extracellular matrix (ECM), which provides a significant advantage in the understanding of the cellular response in vitro. Directing the cellular response within a 3-D scaffold is an interesting approach to promote the formation of tissues such as bone. This can be achieved through the incorporation of ECM ligands, such as the RGD peptide found in fibronectin. This study investigates whether FEFEFKFK peptide gel decorated with RGD improves the physiological response of human osteoblasts when cultured in 3- D. It is expected that peptide functionalisation to induce cell spreading thanks the adhesion sites provided through the fibre network of the gel. Cellular viability, proliferation, synthesis of ECM and mineralisation also might be enhanced under osteogenic stimulation. To assess whether the functionalisation of peptide gels improve cellular response, we used three different peptide gels, RGD-FEFEFKFK (active gel), RDG-FEFEFKFK and FEFEFKFK (the latter two as inactive control gels). The findings of this work revealed that the functionalised gel induced cellular spreading, and significantly increased cell proliferation under osteogenic stimulation. Moreover, in all three gels the synthesis of ECM and mineralisation were observed, however both processes were undertaken in a lower rate in active gel compared to controls. Thus, due to the functionalisation of the FEFEFKFK peptide gel increased cell proliferation and the allowed synthesis of ECM and mineralisation, we believe that the RGD-FEFEFKFK gel might function as an effective scaffold to promote the formation of alveolar bone for dentistry applications.

73 Introduction

Bone loss affects people worldwide, and the resorption of alveolar bone, which supports teeth, is still an issue that dentists face with poor results. Bone grafting, which is the gold standard to address alveolar bone resorption, does not achieve effective bone regeneration. This procedure has other disadvantages, such as low availability, high cost, morbidity and surgery complications. This encourages tissue engineers to seek alternative routes to regenerate alveolar bone (1). The use of biomaterials used for tissue regeneration has increased considerably. Some biomaterials used for tissue regeneration include synthetic and natural hydrogels, bioactive glasses, metallics, ceramics, composites, etc. (2). It is considered that materials intended for bone regeneration must meet with certain characteristics to successfully achieve adequate bone formation. These parameters include osteoinductivity (capability to induce the differentiation of progenitor cells into bone- forming cells), osteoconductivity (capability to promote in mature cells the synthesis of ECM), osteointegration (suitability of the material to be retained and subsequently integrate into the existing tissue), degradability, and not triggering any immunological response (3). Individually, none of the current biomaterials is able to fulfill the “ideal” requirements, hence, materials are frequently combined to complement and improve their individual features (1, 4). Both synthetic and natural hydrogels function as effective scaffolds for the culture of cells in tissue engineering applications (5-7). Peptide hydrogels are highly hydrated and dynamic systems composed by amino acids. (8) The latter act as building blocks to form hierarchical structures, which go on to form fibrils or fibres, that subsequently entangles to form self-supported structures (8-10). A significant advantage of peptide hydrogels in tissue engineering is that they provide a three-dimensional (3-D) niche to cells, which resembles the architecture of the natural extracellular matrix (ECM). This brings in vitro studies closer to the natural cell behaviour. Moreover, cells within hydrogels are protected while they perform their physiological functions. The inner gel structure allows the homogenous flow (in and out) of nutrients, oxygen and biomolecules secreted by cells. On the other hand, gels as viscoelastic materials, can be injected into the damaged tissue avoiding invasive therapeutics (11-13). The inert biodegradability of hydrogels facilitate its integration into the body, meanwhile cells synthesise new tissue (1). The success of peptide hydrogels in the regeneration of bone, relies on their osteoinductive

74 and osteoconductive properties, and not on acting as tissue substitutes or supporting high load bearings (14). Thus, the idea that hydrogels are not suitable materials for hard tissues regeneration is probably far from the truth. Recently, it has been reported that peptide-based hydrogels foster the growth osteoblasts as well as the osteogenic differentiation of stem cells (15-17). Biomaterials including different hydrogel systems (7, 18, 19) such as peptide hydrogels (15, 17, 20), can be easily decorated with ECM ligands, drugs, and other biomolecules to encourage bone formation (21, 22). Some of the most common ECM moieties that are incorporated into peptide hydrogels include the adhesive peptide sequence RGD, REDV, IKVAV and YIGSR. In RGD, R stands for arginine, G for glycine and D for aspartic acid. In REDV, R represents arginine, E glutamic acid, D aspartic acid and V valine. In IKVAV, the I is isoleucine, K lysine, V valine, A alanine and V valine. Finally, in YIGSR Y is tyrosine, I isoleucine, G glycine, S serine, and R arginine. The first two peptides are displayed in distinct domains of fibronectin, and the latter two belong to laminin (22- 24). These moieties serve as adhesion sites for cells and trigger transcription factors involved in cell spreading, proliferation, and synthesis of ECM (25). RGD is also found in other bone proteins such as type I collagen, vitronectin, osteopontin, bone sialoproteins, etc (26). Cells can recognise ECM through integrins such as α3β1, α5β1, α8β1, αIIbβ3, αvβ1, αvβ3, αvβ5, among others (25). This directs the cellular response with bone formation purposes as reported in the literature (27, 28).

The present article is a continuation of previous work, where we showed that the FEFEFKFK hydrogel successfully supports the production of ECM and that human osteoblasts encapsulated inside the gel were capable of producing mineralised tissue- like. Here, we functionalise incorporate the peptide RGD into the FEFEFKFK peptide sequence, in order to provide adhesion sites for osteoblasts within gel. Based on this strategy, it is expected to enhance the production of ECM and mineralisation under osteogenic stimulation (16). Our findings evidence that cells spread within the RGD- FEFEFKFK gel. In addition cell proliferation is significantly increased inside functionalised gel. Despite this, the highest synthesis of ECM and mineralisation were observed within non-functionalised gels. Finally, the mechanical properties of all gels containing cells decreased over time, this effect resulted more evident in control gels.

75 Experimental

Hydrogel preparation

Three different hydrogels were prepared: GRGDS-FEFEFKFK (active peptide sequence), GRDGS-FEFEFKFK (scrambled peptide sequence) and FEFEFKFK (unfunctionalised). All three peptides (> 95% purity) were purchased from Biomatik Corporation, Canada. To prepare the GRGDS-FEFEFKFK and GRGDS-FEFEFKFK gels, 10.8 mg of each peptide powder were weighted, and combined with 25.2 mg of FEFEFKFK peptide respectively to obtain a 36 mg mixture of each peptide (30% functionalised peptide and 70% unfunctionalised peptide). Thereafter each peptide mixture was dissolved in 800 µl of ddH2O. To prepare the unfunctionalised gel, 36 mg of FEFEFKFK peptide powder were weighted and also dissolved in 800 µl ddH20 (see Table 1). Peptide solutions were subsequently vortexed and centrifuged (4000 rpm min−1), before placing into an oven at 90°C for 2 h to ensure complete dissolution. To induce gelation, 1 M NaOH (90-95 µl) and DPBS (100 µl) were added. A further vortex/centrifugation cycle (5000 rpm min−1) was carried out to finally place the gel back into the oven at 90ºC for 2 h before being cooled at room temperature (RT) and ensure formation of a homogeneous hydrogel. The final concentration of all gels after the addition of media was 3 w/v %.

Oscillatory rheology

The viscoelastic properties of RGD-FEFEFKFK and control gels were determined using an ARG2 rheometer with 20 mm parallel plate geometry at days 1, 7 and 14 under cell culture conditions. Rheology measurements were undertaken following the protocol previously described by Castillo et al. 2014 (16).

Cell culture

Primary human osteoblasts (hOBs) were purchased from PromoCell, Heidelberg, Germany. The cells were grown and maintained following the cell culture conditions

76 indicated by the supplier’s protocol. For more details the reader is referred to Castillo et al. 2014 (16).

Three-dimensional culture within gel

A total volume of 200 µl of cell suspension (2×106 cells), were pipetted on top of the gel. Subsequently the cell suspension was gently mixed by stirring with the pipette tip before gently pipetting up and down until create a homogeneous gel/cell suspension. Thereafter 250 µl of gel containing 5×105 cells was transferred into 12-well cell culture inserts (ThinCertTM Greiner bio-one). Each gel was left to set for 10 min at

37°C in a 95% humidified atmosphere (20% O2) with 5% CO2 to aid gelation. Fresh media changes (× 5) at 20 minutes intervals were carried out to stabilize the pH of gels to physiological. After 24 h, an additional change of media was carried out and subsequent media changes were carried out every 4 days to aid cell growth within gel. In the case of two-dimensional cultures (2-D) on TCP 2×104 cells/cm2 were pipetted directly onto a 48-well cell culture plate. For ICC staining experiments, cells (4×104 cells cm2) were cultured on glass using 12-well cell culture plates.

Cell viability

Cell viability was explored using live/dead assay (Invitrogen, UK). For confocal imaging of live/dead assay, samples were processed according to the manufacturer’s instructions. For more details the reader is referred to Castillo et al. 2014 (16). To obtain the percentage of cell viability, we manually counted both live and dead cells over 14 days of culture. For this, we used live/dead assay imaging with three different confocal Z-scan series per sample (two samples per time-point).

Cell proliferation

Osteoblast proliferation was quantified using Picogreen dsDNA assay purchased from Life technologies, UK. Samples were processed according to the manufacturer’s instructions. For more details the reader is referred to Castillo et al. 2014 (16).

77 Immunofluorescence imaging for type I collagen production

The production of type I collagen within gels was carried out using immunofluorescence imaging. For more details of the immunocytochemistry staining method used in this work, the reader is referred to Castillo et al. 2014 (16).

ECM quantification

The production of ECM was assessed using spectrophotometric methods. All samples were subject to osteogenic stimuli over two weeks of culture. After each time-point gels were subjected to three freeze/thaw cycles, to ensure cellular lysis. We quantified three osteogenic markers: total collagen, osteocalcin (OCN) and alkaline phosphatase (ALP). In each case a blank gel without cells was used to correct background absorbance. All experiments to quantify bone proteins were normalised to cell numbers from the Picogreen assay. Two independent assays were performed in triplicate.

Total collagen synthesis

The total collagen production was determined using a Total Collagen Assay (QuickZyme Biosciences). Samples were processed according to the manufacturer’s instructions. For more details the reader is referred to Castillo et al. 2014 (16).

Alkaline phosphatase (ALP) activity

The alkaline phosphatase activity was monitored using a colorimetric peroxidase substrate detection system (Sigma-Aldrich). All samples were processed according to the manufacturer’s instructions. For more details the reader is referred to Castillo et al. 2014 (16).

Osteocalcin (OCN) synthesis

The osteocalcin production was determined using a Human Osteocalcin ELISA Kit (Invitrogen-Life technologies). All samples were processed according to the

78 manufacturer’s instructions. For more details the reader is referred to Castillo et al. 2014 (16).

Mineralisation

The deposition of bone-like nodules to determinate bone mineralisation within gels was explored using Alizarin red staining. For more details on the protocol used in this work the reader is referred to Castillo et al. 2014 (16).

Statistical analysis

For statistical analysis of picogreen, percentage of viability, oscillatory rheology and ECM quantification, results were displayed as mean ± SD using one-way ANOVA followed by appropriate post-hoc comparisons (Tukey’s method).

Results and discussion

Cell viability, morphology and proliferation

The first objective of this work was to determine the effect of the RGD moiety on the viability and morphology of human osteoblasts (hOBs) when cultured within the RGD-FEFEFKFK gel. We expected that through integrin recognition, the ligand RGD induces cells to display a rearrangement in their cytoskeleton. This would subsequently lead to a spread cell conformation, similar to that observed when cells are on bone surfaces in vivo. Live/dead assay was used to investigate the viability of hOBs cultured within active RGD-FEFEFKFK and RDG-FEFEFKFK and FEFEFKFK gel controls. Confocal imaging revealed that most osteoblasts were viable within all gels and only a few dead cells were observed (Figure 1 A-I), suggesting that the three different gels offered a suitable environment to cells over 12 days of culture. This result is consistent with what we previously reported, when hOBs were cultured inside the FEFEFKFK gel (16). It is believed that the RGD peptide sequence is capable of inducing physical (cell-cell), and chemical (paracrine) cell interactions via integrin-mediated signaling. RGD can also trigger focal adhesions, which might increase both cell viability and proliferation, while

79 simultaneously inhibiting apoptosis (25). The relative numbers of cell viability showed no significant difference among the three different gels compared (Figure 2). This result is in good agreement to a previous report where human mesenchymal stem cells were cultured in alginate microspheres with and without RGD (29). Nevertheless, notably cells spread within the active gel (Figure 1 A-C), which is consistent with previous reports (18, 20, 30, 31). On the other hand, in control samples, most hOBs show a round morphology (Figure 1 D-I). This is probably due to the inactive gels do not provide of adhesion ligands to the cells. The round cellular morphology found in control gels is consistent with what we previously reported for the FEFEFKFK peptide gel (16). Interestingly, a few cells within RDG-FEFEFKFK control (scrambled peptide sequence), spread at day 14 of culture (Figure 1 F, white circles). This suggests that cells might recognise the inactive peptide sequence. There is little evidence regarding the cell recognition of scrambled peptide sequences such as RDGS, GRGESP or GRADSP (P is proline) when used as controls (32-34), fact that is not fully understood. It is believed that the expression pattern of integrins (αvβ3, α5β1 and αIIbβ3, etc.), is not undertaken statically, opening the possibility that cells can recognise such scrambled peptide sequences presenting an aberrant conformation state (33). The next aim in this work was to investigate the effect of RGD on cell proliferation. Interestingly, the rate of hOBs proliferation was significantly higher within the RGD- FEFEFKFK gel than in the FEFEFKFK control (Figure 3). This is in accordance to the reported by others (20, 28, 35). There is evidence showing that RGD-dependent cell spreading, significantly enhances cell proliferation, which eventually leads to the down regulation of pro-apoptotic signal cascades (25). Further experiments involving the participation of pro-apoptotic molecules (p53, caspases, cytochrome c releasing, Apaf-1), are needed to determine the influence of RGD on both cell viability and proliferation within gels. Likewise, further experiments exploring the participation of death cell inhibitors (Bcl-2 or Bcl-XL) during RGD-cell interaction are also necessary (25, 36). On the other hand, no significant difference on cell proliferation between controls was observed. This was expected and is good agreement with reported by Benoit et al. 2005 and Olbrich et al. 1996 (19, 32). Human osteoblast proliferation was also tracked over 14 days on two-dimensional (2-D) culture, without comparison intentions. As expected, cells presented a time-dependent increase of the proliferation

80 rate (Figure S 1), which is typical for 2-D cultures. The results described above suggest that the functionalisation of the FEFEFKFK gel with RGD has a positive effect on the functions of human osteoblasts, in particular cell spreading and proliferation. This might aid the osteogenic activity of osteoblast cells.

Gel mechanical properties

Cells and the matrix harboring them may influence each other depending on both the cells functions (37-39), and on the stiffness and topography of the matrix (40, 41). The mechanical properties of hydrogels including the FEFEFKFK gel rely on the chemistry, physics and type of amino acids forming the peptide gel. Other factors such as peptide concentration, density of secondary structures formed, ionic strength, pH, etc., can influence the mechanical properties of peptide gels, making them tunable (8, 40). We previously showed that 3 w/v % FEFEFKFK gel, which has ca. 10 kPa storage/G prime (G′) moduli, successfully supports the 3-D culture of human osteoblasts. There, cells are capable of undertaking physiological functions such as synthesising ECM and mineralisation (16). In this work we investigate whether the incorporation of the RGD short peptide sequence has an impact on the mechanical properties of the gel. We also explore whether a change on the gel mechanics affects the osteoblats functions within the gel. We used oscillatory rheology to evaluate the mechanical properties of all three 3 w/v % gels (RGD, RDG, and unmodified sequence). The storage moduli of the RGD gel containing cells recorded ca. 10 kPa G′ moduli values (day 1), which subsequently decreased to ca. 7 kPa over time in day 14 (Figure 4). Whilst the mechanics of control gels shows a similar trend, (the G′ moduli values also decrease over time), in RGD gels this phenomenon was less pronounced (Table 2, days 1 to 14). High cell densities increase the fibre network-cell interactions in gels, which leads to an increase in the mesh size, accelerating the degradation of gels (40, 42). Moreover, higher cell densities produce higher cellular metabolic activity (hydrolysis and/or enzymatic degradation), leading to the rupture of the fibre linkages (43). In this work, to promote cell-cell interactions, we increased the cell numbers twice and three fold, in comparison to cell densities used to culture bovine chondrocytes and human osteoblasts within the FEFEFKFK gel, respectively (11, 16). We believe that a higher

81 cell density along with the expected gel wearing produced by the cell culture media exchange, might explain the progressive decreasing of the mechanical properties of all three gels over time. Some works report an increase (11, 16, 44) in the mechanical properties of synthetic and natural hydrogels, while other report a decrease (42, 45, 46). Further experiments using different cell densities are necessary to show the direct influence of cells on the mechanical properties of gels. Several variables in these systems, such as cell type, cell density, properties and chemical ratio of components, make gel mechanics a controversial phenomenon yet to be fully investigated. Evaluating the mechanical properties of gels is particularly important when cells are present. The G′ modulus of the inactive control FEFEFKFK gel were higher than both RGD and RDG gels at the first day of culture. However, the G′ moduli of the former gel becomes lower than the values recorded for the latter two gels over time (Figure 5). Despite this, we only observed cell spreading within the RGD-FEFEFKFK gel, suggesting that cell spreading is RGD-dependent. For control gels without cells, we observed a similar time-dependent decrease of the G′ modulus in all gels (Figures 4 and 5), which is consistent with what is reported in the literature (11, 16, 44).

ECM synthesis and mineralisation

The final aim of this work was to investigate the impact of the RGD ligand on the hOBs capability to synthesise ECM and mineralise within the gel. Type I collagen is one of the major components of bone ECM, hence we consider that testing its expression within gels is a key parameter to explore the ability of cells to synthesise ECM. Using immunocytochemistry (ICC) staining, we observed that hOBs are capable of synthesising and depositing type I collagen within the RGD-FEFEFKFK and control gels during the first 7 days of culture (Figure S 2). At day 14 of culture, the deposition of this protein seems to be lower within the functionalised gel in comparison with controls (Figure 6). This was quantitatively corroborated using a quantitative assay, where cells produced higher amount of type I collagen within control gels (Figure 7A) unlike reported when alginate microspheres were functionalised with RGD to induces osteogenic differentiation of hMSCs (29). Likewise, the production of osteocalcin showed a similar trend (Figure 7B). On the other hand, we could not detect a significant difference in the production of ALP between the RGD-FEFEFKFK gel and controls (Figure 7 C). Overall, these findings

82 strongly suggest that the incorporation of RGD into the peptide gel increases the proliferation of cells than the synthesis of ECM. This is consistent with some previous reports. For example, bone marrow stem cells showed spread morphology when agarose gels were functionalised with several ligands such as the RGD peptide, a recombinant fibronectin fragment (FnIII17-10) and a collagen mimetic peptide with the sequence GFOGER. However the cell-ligands interaction, inhibited the expression of factors related the expression of collage type II, aggrecan (mRNA), as well as the deposition of glycosaminoglycans. Additionally, the expression of OCN was also inhibited (47). Thus the biochemical stimuli of ligands may control the differentiation of cells, where the expression and synthesis of ECM proteins is not always upregulated through ligand stimuli. On the other hand, in order to mimic fibronectin adhesion sequences, a PEG gel was functionalised with the peptide sequences RGD and PHSRN (P is proline, H histidine, S serine, R arginine and N asparagine. This gel was capable of enhancing the metabolic activity of osteoblasts, however the expression of total ECM decreased in comparison to controls (unmodified and scrambled peptide sequences) gels (19). Thus, the incorporation of ligands directing the cellular response, not always enhances the production of ECM. It is believed that adhesion sites sensed by cells within a functionalised system can downregulate the signal transductions related to the expression of ECM (19, 47, 48), and upregulate those related to cell proliferation (49). This is consistent with the fact that high concentrations of immobilized adhesion sites tend to decrease the synthesis and deposition of ECM (25). Different studies suggest that the cellular mechanical tension induced by the RGD ligand might lead to both the formation of stress fibres and changes on the genetic program of cells. There, the synthesis of ECM generally is downregulated (25, 37, 50). Thus a spindle cell morphology induced by adhesive ligands is associated with a proliferative program, which derive from the cell-cell interaction/stimulation. This is unlike what is observed when cells acquire a rounded morphology, where the lack of cell contact is associated with cell differentiation accompanied with synthesis of ECM (50). Further studies are needed to elucidate whether RGD is capable of triggering cytoskeletal rearrangement in osteoblasts within the gel. This triggering occurs via the activation of proteins such as Rho (actomyosin filament promoter), microtubules, stress actin microfilaments, as well as gap junctions. It is also necessary to explore whether this process has any correlation with both the synthesis of ECM and cell proliferation. Finally, we also investigated

83 the capability of human osteoblasts of mineralising within gels using alizarin red staining. Consistent with what we observed in ECM production, we visually found that the mineralisation displayed by human osteoblasts within the RGD modified gel is lower in comparison to controls (Figure 8 A). This fact that was corroborated quantitatively (Figure 8 B). Likewise, the time-points tested here do not show a significant difference in the calcium deposits inside the RGD gel containing cells. This is in good agreement with the ALP activity reported here previously. So far these findings suggest that the ECM production by hOBs is not proportional to the rate of proliferation induced under the influence of RGD, which might also have a direct impact on the mineralisation rate. Although in this work 2-D cultures were only used as a reference, a similar discrepancy between the rate of proliferation and mineralisation was observed, when the mineralisation was normalised to cell numbers from picogreen (S. 3).

Conclusions

In this work, it was show that incorporating the RGD ligand into the FEFEFKFK peptide gel, enhances cell spreading and significantly increases the proliferation of human osteoblasts under osteogenic stimuli in vitro. This ligand might contribute to the formation of bone in situ, once such gel is transferred into bone defects. On the other hand, the lower ECM expression, and mineralisation observed within the functionalised RGDFEFEFKFK gel does not correlate with the high rate of cell proliferation. This suggests that cells commit into a proliferative state, which might have a positive impact of bone formation at subsequent time periods. The mechanical properties of the hydrogel allow cells to proliferate and undertake physiological functions. However, further investigation is necessary to show whether cell densities have a direct effect in the gel mechanical properties. Thus, the functionalisation of the FEFEFKFK hydrogel with the RGD peptide sequence might be an interesting approach to induce the proliferation and synthesis of bone-like tissue for bone regeneration applications.

84 Acknowledgements

The authors would like to thank CONACyT-Mexico for funding support as well as the Polymers and Peptides and Biomaterials and Tissue Engineering research groups for helpful discussions and kind support.

85 List of figures.

Day 1 Day 7 Day 14

A Viable/Dead B Viable/Dead C Viable/Dead

FEFEFKFK -

RGD

D Viable/Dead E Viable/Dead F Viable/Dead

FEFEFKFK -

RDG

G Viable/Dead H Viable/Dead I Viable/Dead

FEFEFKFK

Figure 1. Live/dead assay confocal imaging. Viable (green) and dead (red) cells inside RGD-FEFEFKFK, RDG -FEFEFKFK and FEFEFKFK gels over 14 days of culture (n=2). White circles enclose spread cell morphology inside the control RDG-FEFEFKFK gel. Magnification = 20×. Scale bars = 250 µm.

RGD-FEFEFKFK gel 100 RDG-FEFEFKFK gel FEFEFKFK gel 80

60 40

viability(%) Percent 0 1 7 14 Time (Days) Figure 2. Viability percentage from live/dead assay confocal imaging. Three different confocal Z-scan series from each sample were used to quantify both live and dead cells over 14 days of culture (n = 2).

2.5×10-1 RGD-FEFEFKFK gel p = 0.0064 RDG-FEFEFKFK gel -1 2.0×10 FEFEFKFK gel g/ml) µ 1.5×10-1

1.0×10-1 5.0×10-2 DNA content ( 0.0 3 7 3 7 3 7 14 14 14 Time (Days)

F ig 3. DNA content representing cell proliferation within RGD-FEFEFKFK, RDG-FEFEFKFK, and FEFEFKFK gels over 14 days of culture (n = 3).

A p = 0.0003 100000 RGD-FEFEFKFK gel (+ cells) p < 0.0001 p < 0.0001 RGD-FEFEFKFK gel (- cells) 10000

1000 100

(Pa) modulus G' 10 1 1 7 1 7 14 14 Time (Days)

p < 0.0001 B p < 0.0001 100000 RDG-FEFEFKFK gel (+ cells) p < 0.0001 p < 0.0001 RDG-FEFEFKFK gel (- cells) 10000

1000 100

(Pa) modulus G' 10 1 1 7 1 7 14 14 Time (Days)

C p < 0.0001 100000 FEFEFKFK gel (+ cells) p < 0.0001 FEFEFKFK gel (- cells) p < 0.0001 10000

1000

100

(Pa) modulus G' 10

1 1 7 1 7 14 14 Time (Days)

Figure 4. Storage modulus (G′ modulus) of (a) RGD-FEFEFKFK, (B) RDG-FEFEFKFK, and (C) FEFEFKFK gels with and without cells (grey and white bars respectively) over two weeks of culture (n = 3).

RGD-FEFEFKFK gel (+ cells) RGD-FEFEFKFK gel (- cells) RDG-FEFEFKFK gel (+ cells) RDG-FEFEFKFK gel (- cells) FEFEFKFK gel (+ cells) FEFEFKFK gel (- cells)

100000 p < 0.0001 p < 0.0001 p < 0.0001 10000

1000

100

(Pa) modulus G' 10

1 1 7 14 Time (Days) Figure 5. Storage modulus (G′ modulus) comparison between RGD-FEFEFKFK, RDG- FEFEFKFK, and FEFEFKFK gels with cells (patterned bars) and the same gels without cells (plain graphs) over two weeks of culture (n = 3).

DAY 14

F-actin Type I collagen Merge

FEFEFKFK FEFEFKFK

RGD

F-actin Type I collagen Merge

FEFEFKFK FEFEFKFK -

RDG

F-actin Type I collagen Merge

FEFEFKFK

Figure 6. ICC staining for type I collagen deposition within RGD-FEFEFKFK (A-C), RDG- FEFEFKFK (D-F), and FEFEFKFK (G-I) gels over two weeks of culture (n = 2). Magnification = 10×. Scale bars = 500 µm.

4.5×10-3 RGD-FEFEFKFK gel p = 0.0002 RDG-FEFEFKFK gel p = 0.0176 FEFEFKFK-gel -3 g/ml/cell) 3.0×10 p = 0.0024 μ

1.5×10-3

Total collagen ( 0.0 7 7 7 14 14 14 Time (Days)

B p < 0.0001 RGD-FEFEFKFK gel 4.0×10-3 p < 0.0001 RDG-FEFEFKFK gel FEFEFKFK gel 3.0×10-3 p = 0.0117

2.0×10-3

1.0×10-3

Osteocalcin (ng/ml/cell) Osteocalcin 0.0 7 7 7 14 14 14 Time (Days)

1.0×10-3 RGD-FEFEFKFK gel RDG-FEFEFKFK gel p = 0.0120 -4 FEFEFKFK gel 8.0×10 p = 0.0409 p < 0.0001 6.0×10-4

-4 4.0×10 2.0×10-4

0.0

activiy/nmol/ml/30min/cell pNPP 7 7 7 14 14 14 Time (Days)

Figure 7. Normalised production of (A) total collagen, (B) osteocalcin, and (C) alkaline phosphatase within RGD-FEFEFKFK and control gels over 14 days of culture (n = 2).

FEFEFKFK RGD-FEEFKFK RDG-FEFEFKFK FEFEFKFK

(- cells) (+ cells) (+ cells) (+ cells)

Day 7

Day 14

B RGD-FEFEFKFK gel 3×10-5 p = 0.0113 RDG-FEFEFKFK gel FEFEFKFK gel 2×10-5 p = 0.0069

1×10-5

Absorbance/cell (562 nm) 0 7 7 7 14 14 14 Time

Figure 8. Calcium deposition (red) within gels determined by Alizarin red staining (A). Normalised absorbance from calcium phosphate ions (CPC extraction) over two weeks of culture (B) (n = 2).

Supplementary images

0 2-D Culture (TCP) 1.0×10 p = 0.0007

7.5×10-1 g/ml)

5.0×10-1 p = 0.0131

p = < 0.001 2.5×10-1

DNA content ( 0.0 1 3 7 14 Time (Days)

Supplementary figure 1. DNA content representing cell proliferation on TCP culture over 14 days (n = 2).

DAY 7

F-actin Type I collagen Merge

FEFEFKFK -

RGD

F-actin Type I collagen Merge

FEFEFKFK -

F-actin Type I collagen Merge

EFEFKFK

Supplementary figure 2. ICC staining for type I collagen deposition within RGD-FEFEFKFK (A-C), RDG -FEFEFKFK (D-F), and FEFEFKFK (G-I) gels over 7 days of culture (n = 2). Magnification = 10×. Scale bars = 500 µm.

A TCP B

2-D Culture (TCP) 3×10-4

-4 p = 0.0016 Day 7 2×10

1×10-4

Absorbance/cell (562 nm) 0

Day 14 7 14 Time (Days)

Supplementary figure 3. Alizarin red staining of visual (A), and quantitative mineralisation (normalised to cell numbers) (B) on TCP over two weeks of culture (n = 3).

Tables

Table 1. FEFEFKFK gel concentrations, and buffer volume conditions used for the preparation of hydrogels for the 3-D culture of human osteoblast cells.

GRGDS GRDGS FEFEFKFK Water Total gel 1M NaOH 10× PBS Gel peptide peptide peptide weight volume w/v % /µl /µl weight /mg weight /mg /mg /µl RGD- 10.8 - 25.2 3 800 90 100 FEFEFKFK RDG- - 10.8 25.2 3 800 90 100 FEFEFKFK FEFEFKFK - - 36 3 800 90 100

Table 2. Storage moduli values recorded from RGD-FEFEFKFK, and control gels under cell culture conditions over 14 days.

RGD-FEFEFKFK gel RDG-FEFEFKFK gel FEFEFKFK gel Time G ′ modulus G ′ modulus G ′ modulus G ′ modulus G ′ modulus G ′ modulus (Days) Gel + cells Gel - cells Gel + cells Gel - cells Gel + cells Gel - cells

1 10300 ± 1100 10760 ± 830 10300 ± 1100 10300 ± 1800 10530 ± 900 15700 ± 1580

7 11830 ± 1000 9500 ± 1050 7900 ± 500 10100 ± 1000 4320 ± 670 15250 ± 800

14 7300 ± 1500 7900 ± 830 5800 ± 800 9000 ± 2800 2640 ± 260 11370 ± 1280

94 References

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98 CHAPTERFOUR

Osteogenic differentiation of human mesenchymal stem cells promotes mineralisation within an octa- peptide hydrogel

Luis A. Castillo Díaz, Alberto Saiani, Julie E. Gough, and Aline F. Miller

All authors contributed to the realisation of this manuscript.

99 Osteogenic differentiation of human mesenchymal stem cells promotes mineralisation within an octa-peptide hydrogel

Luis A Castillo Díaz,1,2 Alberto Saiani,2,3 Julie E Gough3* and Aline F Miller1,2* .

1 School of Chemical Engineering and Analytical Science, The University of Manchester (UK).

2 Manchester Institute of Biotechnology, The University of Manchester (UK).

2 School of Materials, The University of Manchester (UK)

Corresponding authors Aline F. Miller, Manchester Institute of Biotechnology, School of Chemical Engineering and Analytical Science, The University of Manchester, 131 Princess Street, Manchester, M1 7DN, UK. Email: [email protected]

Julie E. Gough, Materials Science Centre, School of Materials, The University of Manchester, Grosvenor Street, Manchester M13 PL, UK. Email: [email protected]

100 Abstract

An attractive strategy for the regeneration of tissues has been the use of extracellular matrix (ECM) analogous biomaterials. Peptide based fibrillar hydrogels have been shown to mimic the structure and properties of ECM. In this study, the capability of a simple ionic-complementary peptide FEFEFKFK (F, E and K are phenylalanine, glutamic acid and lysine) hydrogel to host human mesenchymal stem cells (hMSCs) in 3-dimensions (3D) and induce their osteogenic differentiation is demonstrated. Assays showed a sustained cell viability and proliferation throughout the hydrogel over 12 days of culture, and these hMSCs differentiated into osteoblasts simply upon addition of osteogenic stimulation. Differentiated cells synthesized key bone proteins, including collagen-1 (Col-1), osteocalcin (OCN) and alkaline phosphatase (ALP). Moreover, mineralization occurred within the hydrogel. These findings demonstrate that a simple octapeptide can host and induce the differentiation of stem cells, and as such it has the potential for the regeneration of hard tissues.

101 Introduction Bone is the major structural and supportive tissue in the body, but can be compromised by degenerative diseases or trauma (1, 2). It is understandable, therefore, that research into developing and optimizing the process of bone regeneration is intense and remains of great interest. It is known that such regeneration involves a complex series of biological events of bone induction and conduction where a number of different healthy cells or tissues lend themselves to restore lost or damaged osseous tissues. This becomes a serious challenge within the field of regenerative medicine where there are either large or small quantities of missing tissue (1-3). One example of this is in periodontitis, which is an oral pathology that induces the degradation of alveolar bone (2, 4). Currently, bone grafting is the ‘gold standard’ method used to tackle the resorption of alveolar bone (5); nevertheless, it does not achieve effective bone regeneration (6). Additional concerns with this methodology include high cost and the high risks associated with a surgical procedure (7). Over the past few years, the potential of using human mesenchymal stem cells (hMSCs) to regenerate different tissue types has been highlighted due to the cells inherent capability to commit into different type of mature cells such as osteoblasts, chondrocytes, among others (8, 9). The differentiation of hMSCs into bone-forming cells has also been reported where 3-dimensional (3D) scaffolds have been used to host the cells and subsequently induce and control differentiation via several different approaches, including tuning the matrix stiffness (10), incorporating growth factors (11), combining growth factors with low-power laser photo activation (12), heat shock stimuli (13), or using strontium(14). Several different types of 3D hydrogels have been reported in the literature, including both natural and synthetic systems. Examples of natural gels include collagen, alginate, hyaluronic acid or Matrigel (15, 16). These materials inherently have active biomolecules and offer good biocompatibility, but control of their components (batch to batch variability) makes it difficult to define the cause of any cellular response (16, 17). On the other hand, synthetic biomaterials such as poly (ethylene glycol) (PEG), and peptide-based systems overcome these issues, since these materials are made of well-known components providing a minimalistic approach to the culture of cells (18, 19). In addition, the mechanical properties of synthetic gels are easily tunable offering an attractive route to direct the cellular response. One limitation of these synthetic

102 materials is that they lack of bioactive molecules, however these can be easily incorporated successively (16). Peptide gels are highly versatile, their self-assembly can be controlled from the bottom-up to form secondary structures such as α-helixes or β-sheets for example, which self-assemble to form fibrils or fibres that subsequently entangle to form a self-supporting structure that mimics the extracellular matrix (ECM) (20, 21). With the modular peptide based systems, the solution to gel transition, the fibre and gel morphology, and consequently the resulting mechanical properties of the 3D hydrogel can be tuned easily by peptide design, or varying peptide concentration, pH, ionic strength, and/or temperature (20, 21). Moreover such peptide hydrogels are inherently biocompatible, and biodegradable and as a consequence, they have found a wide variety of applications, including drug delivery, cell culture, tissue engineering, biosensors and supports for biocatalysts (19). Despite their numerous advantages, these peptide hydrogels have only been used in a few studies for the culture and controlled differentiation of MSCs for bone regeneration (22-24). One example is from Anderson et al. where they incorporated the ECM moieties RGDS (arginine-glycine-aspartic acid-serine), and DGEA (aspartic acid-glycine-glutamic acid-alanine) to the end of a self-assembling peptide amphiphile (CH3(CH2)14CONH-GTALIGQ - where G, T, A, L, I, G, Q, are glycine, threonine, alanine, leucine, isoleucine, glycine and glutamine). They used the nano- fibrillar hydrogels to induce differentiation of hMSCs into osteoblasts by culturing under osteogenic stimulation. They went onto show that the differentiated cells were able to synthesize bone in vitro (22). Self-assembling peptides have also been used to create composite materials in an attempt to control the differentiation of hMSCs. For example the RADA16 peptide

(AcN-RADAADARADARADA-CONH2), has been combined with a collagen sponge, and the hybrid construct induced the differentiation of hMSCs. Once the composite containing undifferentiated cells was transferred in vivo, alveolar bone mass was significantly increased, and at the same time the population of osteoclasts (bone-resorbing cells) was down regulated. Authors hypothesised that hMSCs might have the ability to shift the pro-inflammatory macrophage program into an anti- inflammatory state, which subsequently aided the formation of bone (23). The same peptide was also mixed with hydroxyapatite (HA), and Matsumoto et al. demonstrated that this composite provided a 3D niche with good stiffness, and

103 osteoinductive properties, that led to the osteogenic differentiation of murine MSCs in vivo (24). Thermal stimulation has also proved successful in the differentiation of stem cells, where hMSCs within the RADA16-I hydrogel deposited calcium and expressed several key bone markers only after being induced by a change in temperature (13).

In the aforementioned approaches generally long peptide sequences that require decoration with additional ECM signals are needed to induce MSC differentiation. In this paper we present a simplified approach where we use a cost-effective ionic- complementary octa-peptide based hydrogel, FEFEFKFK where F, E and K are phenylalanine, glutamic acid and lysine. This builds on our previous work where we have shown that this ionic-complementary peptide self-assembles into β-sheet rich fibres that entangle to form a 3D self-supporting matrix with controllable mechanical properties (Figure 1-C) (25-27). We have also shown that this hydrogel can support the culture of osteoblast cells and promote mineralization under osteogenic stimulation (28). Here, we explore the capability of the FEFEFKFK peptide hydrogel to support the 3D culture of hMSCs, and induce the hMSCs commitment into bone- forming cells. In doing this we are demonstrating its potential for bone tissue engineering applications.

Materials and reagents

FEFEFKFK-peptide > 95% purity was purchased from Biomatik Corporation, Canada. Primary human mesenchymal stem cells (hMSCs), Osteoimage™ Mineralisation Assay, growth and osteogenic cell culture medium were purchased from Lonza Walkersville Inc., Basel, Switzerland. Trypsin/EDTA, DAPI-ProLong Antifade Reagent, Alexa fluor 488-phalloidin, Rhodamine-phalloidin, Live/dead assay, Picogreen assay, and Human Osteocalcin ELISA Kit were purchased from Life technologies, UK. Cell culture inserts were purchased from ThinCertTM Greiner Bio- One Limited, UK. Human mesenchymal stem cells (hMSCs) characterization kit was purchased from Merck Millipore, UK. Secondary antibody (sAb) anti-mouse IgG Atto-594, and SIGMAFAST™ p-NPP Tablets, N1891-50SET were purchased from Sigma-Aldrich Co., UK. Rabbit polyclonal antibody for type I collagen and goat anti- rabit IgG-Alexa Fluor 594 were purchased from Abcam, UK. Total Collagen Assay was purchased from QuickZyme Biosciences, UK.

104 Methods

Hydrogel preparation

FEFEFKFK-peptide (> 95% purity), was used to prepare a peptide solution at 2.5 wt.%. The peptide powder was dissolved in 800 µL of doubly distilled water

(ddH2O). The peptide solution was subsequently vortexed and centrifuged (4000 rpm min−1), before placing in an oven at 90ºC for 1h to ensure complete dissolution. To induce gelation, 1 M NaOH (90-95 µl) and DPBS (100 µl) were added. A further vortex/centrifuge (5000 rpm min−1) cycle was applied and the hydrogel placed in the oven at 90ºC for 12-24h before finally being cooled at room temperature (RT) to ensure formation of a homogeneous hydrogel.

Cell culture

Primary human mesenchymal stem cells (hMSCs) obtained from a 38 year old Caucasian male donor Cells were grown and maintained following standard cell culture conditions in hMSCs growth medium. At 50-60% of confluence, cells were subcultured using Trypsin/EDTA. Cell culture medium was changed every 4 days.

3D Cell culture within the FEFEFKFK hydrogel

Gels were sterilized by UV radiation for 35 mins before 1×106 cells were seeded in the hydrogels. This was done by pipetting 200 µl of cell suspension containing hMSCs in growth/osteogenic medium on top of the hydrogel. The cell suspension was gently mixed by stirring with the pipette tip before gently pipetting up and down to until create a homogeneous gel/cell suspension. 300 µl of gel containing 3×105 cells was transferred into each 12-well cell culture insert. The hydrogel was left to set for

10 min at 37°C in a 95% humidified atmosphere (20% O2) with 5% CO2. Fresh media changes were repeated 5 times at 20 mins intervals to stabilize the pH of gels to physiological pH. After 24 h, a fresh media change was carried out, and subsequent media changes were undertaken every 4 days to aid cell growth within the hydrogel. For all stimulated samples, cells were treated with osteogenic media after two days of culture.

105 Cell viability Cell viability was explored using standard live/dead assay. Live/dead assay solution was prepared using 1.5 ml−1 of PBS containing 2.5 µl of 4 µM ethidium homodimer-1 (EthD-1) assay solution and 1.5 µl of 2 µM calcein AM assay solution. The live-dead assay solution was pipetted on top of each hydrogel and then incubated under standard cell culture conditions for 20 mins. The staining solution was then removed and samples were viewed under a LEICA TCS SP5 confocal microscope. To determinate the numbers of viable cells, 5 different confocal microphotographs (representing 5 different areas of gel) were used to manually quantify viable cells. Cell clusters were considered as a single cell.

Cell proliferation

Human mesenchymal stem cell proliferation was quantified using Picogreen dsDNA assay. Gels were rinsed twice using DPBS, and then they were resuspended in 700 µl of cold lysis buffer (200 mM Tris-HCl, 20 mM EDTA/ddH2O/1% Triton-X100) for 25 mins. To ensure complete cell lysis, samples were vortexed vigorously and subject to 3 freeze-thaw cycles. Thereafter, samples were resuspended, and 100 µl gel/cell suspension was plated into a dark 96-wells, where 100 µl of a working solution of Quant-iT PicoGreen reagent was added. The samples were incubated for 2-5 mins at RT. Fluorescence readings were obtained in a plate-reader (FLUOstar OPTIMA BMG LABTECH) at 435 nm (excitation) and 529 nm (emission). A standard curve was determined using calf thymus DNA in serial dilutions in 1% (v/v) Triton X. A blank gel was used to correct the background absorbance and the assay was performed in triplicate.

Human mesenchymal stem cells (hMSCs) characterization

Stem cell characterization was carried out using immunocytochemistry (ICC). A hMSC characterization kit from which Stro-1, Thy-1 (CD90), and CD19 markers were used to characterize the undifferentiated state of cells either with or without osteogenic stimulation. After 2 days, stimulated samples were treated with osteogenic media. At each time-point, gels were rinsed twice (5 mins each) with 1 × PBS. Thereafter, samples were fixed in 4% (w/v) paraformaldehyde (PFA) for 30 mins at

106 RT. Subsequently samples were washed three times (5 mins each) with 1 × PBS, and permeabilized using Triton X-100 at 0.005% (v/v) in DPBs for 15 mins at RT. Samples were then blocked with 1% (v/v) bovine serum albumin (BSA) for at least 2 h at RT, and incubated over night at 4°C with mouse primary antibodies (pAb) anti- STRO-1, anti-THY-1 (CD90) and anti-CD19 respectively, at the ratio stipulated by the manufacturer (1:500 dilution). The next day, samples were washed twice (5 mins each) with 1 × PBS, and twice with blocking solution. After the last wash, samples were left in the blocking solution for at least 30 mins. Thereafter, samples were incubated for 2 h at RT in the dark with a secondary antibody (sAb) anti-mouse IgG Atto-594, along with Alexa fluor 488-phalloidin to target F-actin filaments. Both sAb and phalloidin, were used at ratios stipulated by the manufacturer (1:250). Samples were finally washed 3 times (5 mins each) with 1 × PBS before being mounted on glass slides using DAPI-ProLong Antifade Reagent, to stain cell nuclei. Images were obtained using a LEICA TCS SP5 confocal microscope. Two independent assays were performed, each in duplicate.

Immunocytochemistry (ICC) for type I collagen

Type 1 collagen production within the gel sample was examined visually using the same ICC staining procedure as for hMSCs characterization, except here a rabbit polyclonal antibody for type I collagen was used as the stain at a ratio stipulated by the manufacturer (1:250). After 24 h, samples were incubated for 2 h at RT in the dark with a goat anti-rabbit IgG-Alexa Fluor 594 along with Alexa fluor 488- phalloidin to target F-actin filaments. Both sAb and phalloidin, were used at ratios stipulated by the manufacturer (1:250). The samples were mounted and confocal images obtained as before. Two independent assays were again performed in duplicate.

Quantification of proteins

The production of bone proteins by hMSCs was quantified using spectrophotometric assays. After 6 and 12 days in culture, cell loaded gels were subjected to 3 freeze/thaw cycles, to ensure cellular lysis. Osteogenic markers quantified included total collagen, osteocalcin (OCN) and alkaline phosphatase (ALP). For all protein quantification experiments cells without osteogenic stimulation were used as control

107 and a blank gel was used to correct for background absorbance. All experiments were normalised to cell numbers obtained from the Picogreen assay. Details for each assay are given below.

Total collagen

Total collagen production was determined using a Total Collagen Assay. After resuspending and lysing, samples were vortexed and hydrolysed over 20 h at 95°C with 100 µl 12 M HCl. Samples were subsequently diluted 1:1 in 4M HCl, pipetted into a 96-well plate, before adding 75 µl of an assay buffer and leaving to incubate for 20 mins at RT. Afterwards, 75 µl of a detection reagent was added and the sample transferred into the oven (60°C) for 1 h. Absorbance measurements were obtained at 570 nm, using a plate-reader (TECAN Infinite M200). Two independent assays were undertaken in triplicate.

Osteocalcin

Osteocalcin (OCN) expression was determined using a Human Osteocalcin ELISA Kit. After cellular lysis using cold distilled water, samples were vortexed and pipetted into a 96-well plate. Subsequently, a working Anti-OST-HRP solution (100 µl) was added to each well, the plate covered and incubated for 2 h at RT. Each well was then rinsed 3 times with a washing solution before adding 100 µl of a chromogen solution (tetramethylbenzidine) and incubating for 30 mins at RT in the dark. The reaction was stopped by adding 100 µl of a stop solution (1N HCl). Absorbance measurements were obtained within 1 h at 450 nm using a plate-reader (TECAN Infinite M200). Two independent assays were performed in triplicate.

Alkaline phosphatase

Alkaline phosphatase (ALP) activity was monitored using a colorimetric ALP and peroxidase substrate detection system. The cell lysate (20 µl) was added to transparent 96-well plates together with 200 µl of p-Nitrophenyl phosphate (pNPP) solution (1 mg ml−1 pNPP, 0.2 M Tris buffer in 5 ml ddH2O) SIGMAFAST™ p-NPP Tablets, N1891-50SET. The reaction was stopped with 3 M NaOH. The absorbance of samples was measured using a plate-reader (Lab systems Multiskan Ascent) at

108 405 nm every 30 secs for 30 mins. Two independent assays were performed in triplicate.

Mineralisation (hydroxyapatite deposition)

The extent of mineralization was determined using an Osteoimage™ Mineralisation Assay. After each culture time-point, the gels were washed with 1 × PBS before being fixed with 4% (w/v) paraformaldehyde (PFA) for 20 mins at RT. Samples were subsequently washed a further twice (5-10 mins each) with Osteoimage™ wash buffer and then incubated with 1.5 ml staining reagent, along with rhodamine- phalloidin (1:250 dilution) to target F-actin filaments at RT in the dark for 30 mins. After incubation, gels were washed three times (5 mins each) with wash buffer before mounting the samples on glass slides using a DAPI-ProLong Antifade Reagent to stain the cell nucleus. Images were obtained using a LEICA TCS SP5 confocal microscope. To quantify the extent of mineralization, washed samples were resuspended in 700 µl wash buffer and their fluorescence determined in a fluorescent plate reader at a 492/520 nm ratio. Two independent assays were performed, again in triplicate.

Statistical analysis

Statistical significance in picogreen, total collagen, ALP activity, and OCN quantification assays, was determined using one-way ANOVA followed by post-hoc comparisons (Tukey’s method).

Results and discussion

Cell viability, morphology and proliferation

The first step in this work was to confirm the capability of the FEFEFKFK peptide hydrogel to support the viability and proliferation of human mesenchymal stem cells (hMSCs) when incorporated within the peptide hydrogel in 3-dimensions (3D). Cell viability was therefore assessed initially using a live/dead assay over a period of 12 days of culture. It is clear from the confocal images shown in Figure 2 that the majority of hMSCs remained viable over the 12 days of culture within the gel, with

109 only a few dead cells being present. This suggests that cells sensed an adequate niche to survive. Cell viability was subsequently quantified (Figure 3) and it was found that the number of living cells increased significantly during the first 6 days and then remained steady over the remaining days in culture. Such viability data, is consistent with what we have previously reported for human osteoblasts (28) and bovine chondrocytes (29) when encapsulated throughout our FEFEFKFK hydrogel. Additionally, the cell viability observed within the FEFEFKFK gel is similar to what Bidarra et al. reported where alginate microspheres were functionalised with RGD to induce the osteogenic differentiation of hMSCs (30). Other peptide and polymer- based hydrogels such as RADA and PEG gels where rat neural stem cells and human mesenchymal stem cells were used, showed a steady viability within such gels at similar cell culture time respectively (31, 32). It is clear from examining cell morphology using confocal microscopy (Figure 2) that most cells acquired a round/ovoid morphology during the first day of culture. As time progressed further (days 6 and 12), cells acquired a definite round shape. This was expected as cells typically remain rounded when embedded within a 3D niche that contains no adhesion sites. Optical microscopy was also used to confirm cell morphology (Figure 4A) and the data obtained corroborated the conclusions from confocal imaging. Interestingly, hMSCs were also cultured in 2D on top of tissue culture plastic (TCP) where they acquired a typical spindle morphology and these became wider under osteogenic stimulation (Figure 4B), as is typical for flattened surfaces (9). The extent of cell proliferation within the FEFEFKFK hydrogels was subsequently quantified using the Picogreen assay and the results are given in Figure 5. It is clear that cell numbers were sustained over the 12 days of culture. There was a slight drop in number, which matches with the viability tend at the same time point, however, moving from days 6 to 12, the rate of overall cell proliferation observed is likely to be sufficient to lead to bone formation if and when osteogenic differentiation is induced within the gel. Such behaviour is also commensurate with similar systems in the literature (33, 34). It should be noted that the stiffness of the gels might impact cell number. It is known that stiff hydrogels with elastic moduli of ca. 25 kPa can induce osteogenic differentiation of MSCs even without the use of any osteogenic supplements (18). Once differentiated, the rate of hMSC proliferation can be impaired (22). Previously we have reported that a 3 wt. % FEFEFKFK gel has a relatively high

110 stiffness (ca. 10 kPa) (28) in comparison with other peptide hydrogel systems used for this purpose (35, 36). The differentiation of hMSCs might potentially, therefore, be induced by the gel stiffness here thus leading to the observed decrease in cell proliferation. Thus, the low rate of stem cell proliferation after a week of culture may have a multifactorial explanation.

MSCs characterisation

To demonstrate that the hMSCs seeded into the peptide hydrogels were fully undifferentiated initially and remained so over time, the plasticity of the cells within the hydrogels was determined using immunocytochemistry (ICC) staining for two well-known positive hMSCs markers; stro-1 and CD90 (thy-1), as well as staining for CD19 as the negative control. This was done for cells embedded within the hydrogel, both with and without osteogenic stimulation. Confocal images shown in Figures 6A and B revealed the presence of both Stro-1 and Thy-1 respectively over the 12 days of culture in growth medium only, i.e. non-stimulated conditions. This confirmed the stem cell characteristic of cells within the gel. Interestingly, the expression of the positive stem cell markers was decreased slightly at day 12. This stem cell marker behaviour has been reported in the literature, where either with or without osteogenic stimuli, sub-cultured stem cells lose the expression of their stemness markers over time (37, 38). Likewise, it has been reported the loss of the expression of Thy-1 in hMSCs cultured under chondrogenic stimuli and using different substrates including alginate gels (39). When stem cells within our hydrogels were stimulated to differentiate using osteogenic media, the positive ‘stem cell’ markers were found to be expressed at a much reduced level in comparison to observed cells cultured with growth media, a result that matches with other studies reported in the literature (37, 38). In support of the characterization of the hMSCs, there was no evidence of the presence of the negative control CD19 within the hydrogel (Figure 7) both with and without osteogenic stimulation. This was expected as it is a marker exclusively expressed by B lymphocytes. These findings suggest that stem cells may shift from their undifferentiated state into a differentiated state in the presence of the osteogenic medium.

111 Extracellular matrix (ECM) production

To confirm the possible differentiation of hMSCs within the hydrogel matrix, the expression of type I collagen (Col-1) was monitored over time in culture by staining for the prototypical marker using ICC. The expression of Col-1 is a good parameter of cell differentiation in this case, as it is the major component of organic bone matrix (~90%) (1). As a first step samples were imaged without the presence of the Col-1 antibody, and images taken did not exhibit any fluorescent signal, thus confirming the hydrogel and cells did not display any unspecific or auto-fluorescence. Confocal imaging of samples with the Col-1 antibody revealed some production of Col-1 inside the hydrogel at days 6 and 12 under non-stimulating conditions as evidenced by the red staining (Figure 8). This corroborates our earlier hypothesis that some of the cells had indeed differentiated over extended time in culture, but not necessarily toward an osteoblastic lineage. Under osteogenic stimulating conditions Col-1 was also observed at day 6 of culture, and this increased significantly at 12 days of culture confirming the cells committed themselves to an increasing differentiated profile over time. Furthermore, it is clear from the confocal images in Figure 8 that the stimulated cells embedded within the FEFEFKFK matrix were also able to deposit the synthesized collagen in the surrounding gel, not just close to the cells. This behavior is also reminiscent of our previous work where human osteoblasts were cultured in 3D within the same hydrogel (28). The total collagen synthesized by the differentiated cells was subsequently quantified using a spectrophotometric assay. The normalized total collagen production observed during these assays is shown in Figure 9A for both the non-stimulated and stimulated conditions. The data corroborates the trends observed in the ICC staining experiments, where more collagen was produced in the stimulated conditions and the quantity of collagen increased over prolonged time in culture. This pattern in collagen synthesis is in good agreement with data reported for the differentiation of MSCs into osteoblast cells in other matrices such as PEG (31) and honeycomb collagen scaffold (40). The production of Col-1 is, however, not exclusive to osteoblast cells, therefore, we also quantified the production and activity of two key bone proteins involved in bone formation and mineralisation; osteocalcin (OCN) and alkaline phosphatase (ALP). Data normalized to cell numbers for each of these are given in Figure 9B and 9C respectively. It is clear that the data for the presence of OCN mirror those observed

112 for the production of collagen (Figure 9A), where the highest production of OCN was observed at the latest time-point for the osteogenic stimulated sample. This is in good agreement with similar studies reported in the literature (22, 40) and again points to osteogenic differentiation of the hMSCs. Similar trends were observed for ALP activity (Figure 9C), although there was less of an increase between the non- stimulated and stimulated cells in this case. In the literature the majority of studies report an increase in the presence of ALP once the osteogenic differentiation of hMSCs is induced in alginate microspheres, RADA-16 and Polycaprolactone (PCL) scaffolds (13, 30, 41). It is known that the differentiation of hMSCs into osteoblasts in vitro can take between ~14 and 30 days, where cells generally proliferate during the first 7 to 14 days. Only at this point does the differentiation process start where the transcription and the expression of ALP is carried out (13, 42). Therefore, in this study we believe the differentiation of hMSCs and subsequent expression of ALP could have been accelerated and reached its peak over the first 12 days. This has been reported by others (13, 41), and could explain the lack of significant increase in ALP activity between days 6 and 12 under osteogenic stimulation within the hydrogel. Based on these studies, any cell differentiation observed during culture could be concluded to be induced either by the environment surrounding cells (i.e. the hydrogel) or the osteogenic stimulation applied (i.e. presence of the osteogenic media). Certainly further studies are needed to determinate whether the mechanical properties of gel or the 3D environment induced this acceleration of the ALP production rate. As we mentioned earlier, it is well known that the mechanical properties of the ECM may have a significant influence either on cell the proliferation (43, 44), or on the differentiation of MSCs even in the absence of any osteogenic stimuli (18, 43-45). In addition, it is known that stiffest matrices can trigger the osteogenic differentiation of hMSCs, which is reasonable if we consider that bone cells are surrounded by a relatively stiff matrix (~100 kPa compressive moduli), which becomes stiffer (~1000 kPa) once mineralization occurs (46). We consider that 2.5% wt FEFEFKFK gel (~9 kPa) stiffness (26, 29), has an intermediate range of stiffness, which is far below bone stiffness but above bone marrow stiffness (~0.2 kPa storage modulus), which is the natural niche of these MSCs (13). Thus, at this stage we do not rule out that both the osteogenic stimulation and the mechanical properties of the gel might impact the plasticity of hMSCs. The influence of gel mechanical strength on hMSC

113 differentiation profile can be explored by varying the concentration of peptide, and/or the ionic strength of its media as well as the type of peptide used to self-assemble (28, 29). Thus far we have shown that the FEFEFKFK peptide hydrogel can act as a 3D support for the proliferation and differentiation of hMSCs into osteoblast cells that go onto produce key bone proteins involved in bone formation. This is greatly facilitated when the cells are stimulated by osteogenic media. We further needed to demonstrate that the new osteoblast cells promote mineralization within the hydrogel under the same cell culture conditions. This was therefore monitored by introducing a staining reagent into the hydrogel that specifically binds to hydroxyapatite. Any binding was then detected using a fluorescent assay. Confocal images of such stained samples at days 6 and 12 of culture, both with and with-out the addition of osteoblast stimuli, are given in Figure 10A. It is clear from these images that green staining indicating the presence of hydroxyapatite is present after 6 days of culture with osteogenic stimulation and the quantity of staining increases significantly after 12 days of culture with osteogenic stimulation. Our findings correlates with that reported in the literature where differentiated hMSCs under osteogenic stimulation mineralized within alginate microspheres (30) and PEGDA hydrogels functionalised with the fibronectin short peptide sequence RGD (47, 48). Although bone mineralisation is a process not very well understood yet, it is believed that apatite binds to type I collagen fibrils, which serve as a spatial framework for crystal deposition. Non-collagen proteins including osteocalcin are needed to complete this process (49-51). Thus, we believe that hydroxyapatite deposits observed within gel derive from the interaction of amorphous calcium phosphate with the bone proteins deposited by differentiated cells in the fibril network of gels. Only a faint hint of green fluorescence is observed after 12 days in culture with-out the addition of osteogenic stimulation, which may be explained due to the lower production of bone proteins observed in the protein quantification. The amount of mineralization occurring in each case was also quantified, and it is clear from the data in Figure 10B that significant mineralization only occurred after prolonged time in culture under stimulated conditions. Such behavior agrees well with the previous protein quantification work presented in Figure 9 indicating bone ECM production. This finding clearly demonstrates the suitability of the FEFEFKFK hydrogel to host hMSCs and promote their differentiation into osteoblast cells, in particular under osteogenic stimuli. In addition, this finding is in good agreement with

114 previous studies where the osteogenic differentiation of murine and human stem cells have been induced in the longer RADA16 peptide sequence (52) and in a phosphate- containing PEG (53) gel respectively over a similar time period of 1 to 4 weeks of culture. The advantage of our octa-peptide based system is its short length (hence reduced cost), it doesn’t require the addition of ECM functionalized peptides groups (such as RGDS and DGEA) and also its ease of handling and injectability (41).

Conclusions

Results from this study demonstrate human mesenchymal stem cells can be incorporated homogeneously and proliferate within a 3D simple ionic-complementary peptide FEFEFKFK hydrogel. These cells were subsequently induced to differentiate into osteoblast cells simply via the addition of osteogenic culture medium. This was significant as it eliminates the prerequisite of adding any ECM biological cues. Once differentiated these osteoblast cells were able to synthesize proteins such as Col-1, OCN and ALP, which are all typically involved in the formation of bone. Moreover, mineralization occurred within the hydrogel. This work, therefore, exemplifies a simple and cost effective biomaterial that can be employed for the regeneration of hard tissues, such as alveolar bone. It requires no additional functionalization with for example, ECM isolated ligand sequences. Furthermore it does not require any complex synthesis step, which is associated with arginine rich peptides such as the RADA systems (41). This octa-peptide system has the added potential of being injected in vivo or in situ, avoiding most of the issues related to the complex surgical procedures currently employed during bone grafting today.

Acknowledgements

The authors would like to thank CONACyT-Mexico for financial support, and members of the Polymers and Peptides, and the Biomaterials and Tissue Engineering Research Groups at UoM for helpful discussions.

115 List of figures

Figure 1. Schematic representation of the FEFEFKFK peptide self-assembly where hydrophobic

phenylalanine (F) is orange, hydrophilic and negatively charged glutamic acid (E) is red, and hydrophilic and positively charged lysine (K) is blue, (A) as building blocks, (B) as amino acids in the octa-peptide interact through electrostatic interactions to form β-sheet secondary structures, and (C) form elongated fibres through non-covalent interactions, which in turn are capable of entangling into a nano-fibrous network.

Day 0 Day 6 Day 12

Figure 2 Live/Dead assay showing viable (green) and non-viable (red) hMSCs within the FEFEFKFK gel over 12 days of culture. Magnification = 20× Scale bars = 250 µm.

116

200 p = 0.0083 150

100

50 Number of viable cells 0 0 6 12 Time (Days)

Figure 3. Quantification of viable cells encapsulated within the FEFEFKFK gel over 12 days of culture (n = 3).

117

A Non-stimulated cells Stimulated cells

Day 6

Day Day 12

B Non-stimulated cells Stimulated cells

Day 6

Day 12

Figure. 4. Optical microscopy imaging showing the morphology of hMSCs when (A) encapsulated within the FEFEFKFK hydrogels and (B) cultured on TCP, with and without osteogenic induction over 12 days of culture Magnification = 10×. Scale bars = 100 µm.

118

6×10-1 p = 0.0243

g/ml)

µ 4×10-1

-1 content (

2×10

DNA 0 1 6 12 Time (Days)

Figure 5. Quantified DNA content within the FEFEFKFK gels over 12 days of culture (n=3).

119

A B

Non-stimulated cells Stimulated cells Non-stimulated cells Stimulated cells

Cell nuclei/F-actin/Stro-1 Cell nuclei/F-actin/Stro-1 Cell nuclei/F-actin/Thy-1 Cell nuclei/F-actin/Thy-1

Cell nuclei/F-actin/Stro-1

Day

Cell nuclei/F-actin/Stro-1 Cell nuclei/F-actin/Stro-1 Cell nuclei/F-actin/Thy-1 Cell nuclei/F-actin/Thy-1

Day Day 6

Cell nuclei/F-actin/Stro-1 Cell nuclei/F-actin/Stro-1 Cell nuclei/F-actin/Thy-1 Cell nuclei/F-actin/Thy-1

Day Day 12

Figure 6. (A) ICC staining for Stro-1 (red) and (B) Thy-1 (red) within the FEFEFKFK hydrogel with and without osteogenic stimuli over 12 days of culture (n=2). Cell nucleus and F-actin are stained in blue and green respectively to characterize the cell architecture. Magnification = 40×. Scale bars = 100 µm.

120

Non-stimulated cells Stimulated cells Cell nuclei/F-actin/CD19 Cell nuclei/F-actin/CD19

Day 2

Cell nuclei/F-actin/CD19 Cell nuclei/F-actin/CD19

Day Day 6

Cell nuclei/F-actin/CD19 Cell nuclei/F-actin/CD19

Day 12

Figure 7. ICC staining for CD19 (red) within the FEFEFKFK gel with and without osteogenic . stimuli over 12 days of culture (n=2). Cell nucleus and F-actin are stained in blue and green respectively to determine the cell architecture. Magnification = 40×. Scale bars = 100 µm.

121

Non-stimulated cells Stimulated cells

Cell nuclei /F-actin/ Collagen 1 Cell nuclei /F-actin/ Collagen1

Day

Cell nuclei /F-actin/ Collagen 1 Cell nuclei /F-actin/ Collagen 1

Day 12

Figure 8. ICC staining for type 9 collagen (red) within the FEFEFKFK gel with and without osteogenic stimuli over 12 days of culture (n=2). Cell nucleus and F-actin are stained in blue and green respectively to determine the cell architecture. Magnification = 20×. Scale bars = 250 µm.

122

5.0×10-4 Non-stimulated MSCs p = 0.0008 -4 Stimulated MSCs 3.8×10 g/ml/cell) µ p = 0.0445 2.5×10-4

1.3×10-4

Total collagen ( collagen Total 0.0 6 6 12 12 Time (Days)

B -2 1.0×10 Non-stimulated MSCs 7.5×10-3 p = 0.0005 Stimulated MSCs

5.0×10-3 p = 0.0020

2.5×10-3

Osteocalcin (ng/ml/cell) Osteocalcin 0.0 6 6 12 12 Time (Days) C

1.5×10-2 Non-stimulated MSCs Stimulated MSCs 1.0×10-2

5.0×10-3

0.0 6 6

pNPP activity (nmol/ml/30min/cell) activity pNPP 12 12 Time (Days)

Figure 9. Quantification of bone proteins by hMSCs normalised to cell numbers from the Picogreen assay. Bone proteins quantification was undertaken with and without osteogenic stimuli within the hydrogel over two weeks of culture. (A) Total collagen production, (B) Osteocalcin production and (C) Alkaline phosphatase activity. Data are reported as mean ± standard deviation for n=2.

123

A Non-stimulated cells Stimulated cells

Cell nuclei/F-actin/Hydroxyapatite Cell nuclei/F-actin/Hydroxyapatite

Day Day 6

Cell nuclei/F-actin/Hydroxyapatite Cell nuclei/F-actin/Hydroxyapatite

Day Day

1.0 Non-stimulated cells Stimulated cells 0.8 p = 0.0255 0.6

0.4

0.2 RFU/Cell (492/520) 0.0 6 6 12 12 Time (Days)

Figure 10. (A) Confocal imaging of mineralization/hydroxyapatite deposits (green) within the FEFEFKFK hydrogel with and without osteogenic stimuli over 12 days of culture (n=2). Cell nuclei, and F-actin are stained in blue and red respectively. Magnification = 40×. Scale bars = 100 µm. (B) Quantification of relative fluorescence units normalised to cell numbers from the Picogreen assay to determine mineralisation within the hydrogel with and without stimuli over 12 days of culture (n=2).

124 References

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128 CHAPTER FIVE

Materials and methods

5.1 Materials The materials used in this thesis are classified and enlisted into two groups: chemical (Table 1) and biological materials (Table 2). A detailed use of all materials will be described in the methods section.

Table 1. List of chemical materials.

CHEMICAL MATERIALS SUPPLIER

FEFEFKFK-acid peptide powder. Biomatik Corporation, Canada. GRGDS-FEFEFKFK-acid peptide powder. Biomatik Corporation, Canada. GRDGS-FEFEFKFK-acid peptide powder. Biomatik Corporation, Canada. Sodium hydroxide (NaOH). Sigma-Aldrich, UK. Triypan Blue. Sigma-Aldrich, UK. Live/Dead assay (Ethidium homodimer/ Calcein Invitrogen, UK. AM). Triton-X100. Sigma-Aldrich, UK. Alexa Fluor 488-Phalloidin. Life technologies Ltd., UK. Rhodamine-Phalloidin. Life technologies Ltd., UK. DAPI-ProLong Antifade Reagent. Invitrogen, UK. Alizarin red. Sigma-Aldrich, UK. Cetylpyridinium chloride (CPC). Sigma-Aldrich, UK.

129

Table 2. List of biological materials

BIOLOGICAL MATERIALS SUPPLIER

Primary Human Osteoblasts (hOBs) obtained from Promo Cell, Heidelberg, Germany hip/knee of donor. Primary Human Mesenchymal Stem Cells (hMSCs) Lonza Walkersville Inc., Basel, Switzerland obtained from a 38 years old Caucasian male donor.

Dulbecco’s Modified Eagle’s Medium (DMEM) Sigma-Aldrich, UK supplemented with 10% fetal bovine serum (FBS). hMSCs growth medium. Lonza Walkersville Inc., Basel, Switzerland

Differentiation basal osteogenic medium. Lonza Walkersville Inc., Basel, Switzerland

Dulbecco’s Phosphate Buffered Saline (PBS) with Sigma-Aldrich, UK MgCl2 and CaCl2.

0.05% Trypsin / 0.53mm EDTA. Gibco-Invitrogen, UK

Polyclonal anti-human collagen 1 produced in rabbit. Abcam, UK

Anti-rabbit IgG-Alexa Fluor 594 produced in goat. Abcam, UK

5.2 Methods Hydrogel preparation FEFEFKFK-peptide (> 95% purity) was purchased from Biomatik Corporation, Canada. The concentrations of the peptide used to prepare the gels were 2, 3, 4, 5 w/v % FEFEFKFK (chapter 2), 3 w/v % RGD-FEFEFEFKFK, 3 w/v % RDG- FEFEFKFFK (chapter 3), and 2.5 w/v % FEFEFKFK (chapter 4). All peptide solutions were initially prepared at the peptide concentrations above mentioned by dissolving the peptide powder in 800 µL of distilled water (ddH2O), vortexing and centrifuging (4000 rpm min-1) the solution before placing in an oven at 90ºC for 2h to ensure complete dissolution. To neutralise the sample and to induce gelation 1M NaOH and DPBS were added. The resulting transparent gel was again vortexed and centrifuged (5000 rpm min-1), and placed back into the oven at 90ºC for 2-24h before

130 being cooled at room temperature (RT) to ensure formation of a homogeneous hydrogel.

Cell culture Human osteoblasts (PromoCell, Heidelberg, Germany) and human mesenchymal stem cells (Lonza Walkersville Inc., Basel, Switzerland) were routinely grown and maintained under standard cell culture conditions in a 95% humidified atmosphere

(20% O2) with 5% CO2. Osteoblasts were grown in Dulbecco’s modified Eagle’s (DMEM) cell culture medium, supplemented with 15% FBS, 1% penicillin and streptomycin and 50 µg ml-1 ascorbic acid. Mesenchymal stem cells were grown in growth media (Lonza Walkersville Inc., Basel, Switzerland). At 80-90% of confluence, cells were subcultured using 0.05% trypsin-0.53 mM EDTA 4Na solution (Gibco-Invitrogen, UK).

Viable cell count using a C-chip haemocytometer Cell counting was carried out using a C-chip haemocytometer. Cells in cell culture flasks were trypsinized using 2.5 ml-1 trypsin/EDTA for 5 minutes, then the cell suspension was centrifuged at 1300 rpm for 5 min. Subsequently the supernatant (media and trypsin) was removed and the cell pellet resuspended in 5 ml-1 fresh culture media. Thereafter a dilution (1:2) of the cell suspension was done by taking out 20 µl of the cell suspension and adding 20 µl of trypan blue. This diluted cell suspension was mixed and transferred (10 µl) into each edge of the haemocytometer. Cells in the haemocytomeer were examined under the microscope using a 10× objective. The cell counting was carried out in 5 of the 9 major squares. The cells overlapping the top and left border of squares were counted but no those overlapping bottom and right borders. Cells stained with blue colour were considered as non- viable cells. The cell counting was repeated using the second chamber to give a total of 10 squares. To calculate the cell numbers, the average number of cells in all 10 squares was consider to obtain the number of cells in 1 x 10-4/mL. As the original cell suspension was diluted 1:2, the number of cells was multiplied by 2 and then by 10,000 to obtain the number of cells per ml-1.

131 Cell culture within gel Gels were sterilised under UV ray for 45 minutes and then 250 µl-1 of cell suspension (please refer to chapters 2, 3, and 4 for details related to cell numbers) were added on top of the gel and gently mixed by stirring the media with the pipette tip. The gel/cell suspension was homogenously mixed by pipetting gently up and down. Then the required volume of gel/cell mixture was transferred into a cell culture insert, which was previously placed into a cell culture well plate (Figure 1).

Cell culture insert

Gel with media Cell culture plate

Figure 1. Represents a peptide hydrogel containing cell culture media, which has been plated into a 12 well cell culture insert of 15 mm inner diameter and 1 µm pore diameter.

Thereafter, the gel was leave to set into the oven for 10 minutes at 37°C to aid gelation, and then very gently 1.5 ml-1 of fresh media were added in between the cell culture insert and the well plate (Figure 2). The gel was replaced into the oven for 20 minutes to complete gelation. The cell culture media flows through the gel within ca. 5 minutes once the gel is in the oven. Finally, 5 media changes (20 minutes intervals each) were repeated to stabilize the pH at physiological. Finally, the media was changed every 3 days within gel to aid cell growth.

Gel with media Media

F igure 2. Represents the flow of media in between the cell culture insert and the well plate to aid the gelation of the FEFEFKFK gel. The media flows from the bottom to the top of the gel.

132 Live/Dead assay Live/Dead assay is a simple fluorescent technique that allows differentiating between live and dead cells in a determined system. The principle of this assay relies on the reaction of fluorescent dyes with cellular components. As it is known dyes cannot penetrate through living cells membrane, thus the non-fluorescent cell-permeant polyanionic calcein AM dye is enzymatically converted (by intracellular esterase activity) into an intense fluorescent calcein, which is retained within living cells, resulting in an intense uniform green fluorescence when cells are alive. On the other hand, the high-affinity nucleic acid Ethidium homodimer-1 (EthD1) stain has a weak fluorescence until it pass through disrupted membranes of dying cells, and subsequently binds to DNA of these cells, where emits a intense red fluorescence (http://tools.lifetechnologies.com/content/sfs/manuals/mp03224.pdf). For the Live/Dead assay used in this work to determine the viability of cells within gels, 1.5 mL of PBS containing 2.5 µL of 4mM ethidium homodimer-1 (EthD-1) assay solution and 1.5 µL of 2µM calcein AM assay solution was prepared. The live/dead assay solution was pipetted on top of each hydrogel and then incubated under standard cell culture conditions for 20 minutes. The staining solution was then removed and the samples analysed under a Leica TCS SP5 confocal microscope.

Picogreen assay Picogreen assay is a fluorescent method based on a fluorescent nucleic acid stain that allows to quantify double stranded DNA (dsDNA) from cells. This ultrasensitive technique, allows to detect as little as 25 pg/ml-1 of dsDNA using a standard spectrofluorometer (http://tools.lifetechnologies.com/content/sfs/manuals/mp07581.pdf.) Osteoblast and mesenchymal stem cells proliferation was assessed using Picogreen dsDNA assay (Life Technologies, Carslbad, CA, USA). After gels were rinsed twice with DPBS, they were resuspended in 500 µL of cold lysis buffer (200 mM Tris-HCl, 20 mM

EDTA/ddH2O/1% (v/v) Triton-X-100) for 25 minutes. To ensure complete lysis, samples were vortexed vigorously and subjected to three freeze-thaw cycles. Thereafter, samples were vortexed again and 100 µL gel-cell suspension was plated into black 96-well plates, where 100 µL of a working solution of QuantiT PicoGreen reagent was added. The samples were incubated for 2-5 mins at RT. Fluorescence

133 readings were obtained in a plate-reader (FLUOstar OPTIMA BMG LABTECH) at 435 nm (excitation) and 529 nm (emission). A standard curve was determined using calf thymus DNA in serial dilutions in 1% (v/v) Triton X. A blank gel was used to correct the background absorbance and the assay was performed in triplicate.

Oscillatory rheology Oscillatory rheology is a method to determinate the mechanical properties of a viscoelastic material. The application of a force known as “shear stress” induces the deformation of a determinate sample at controlled conditions. Such deformation can be recoverable and the energy applied is stored in the solid phase of the sample (elastic), meanwhile in the liquid phase (viscous), the deformation recovery is not possible and the energy applied is dissipated as heat. The viscoelastic behaviour is described by two parameters: Gꞌ (elastic or storage modulus) and Gꞌꞌ (viscous or loss modulus). To determinate the linear visco-elasticity of a determined material, it is necessary to apply both a stress amplitude sweep and a frequency sweep stress on the sample. During amplitude sweeps stress, a shear stress at constant frequency is applied on the sample, and the resulting shear strain is monitored as the shear amplitude stress increases. This test allows the identification of the linear viscoelastic region (LVR) of sample, were both Gꞌ and Gꞌꞌ remain constant as the amplitude of the shear stress increases. During the application of the shear stress, samples may break down or flow [1, 2]. A typical rheometer is composed of a stationary bottom plate connected to a Peltier element to control the temperature, as well as of a parallel rotating plate, which applies the shear stress (Figure 4).

134

Parallel plate

Sensor

Bottom plate

Figure 3. Oscillatory rheometer.

The sample is placed in a small gap i. e. 20 mm high separation between bottom and parallel plates. Once the sample is placed, the parallel plate is lowered until it contacts with the sample, where a sinusoidal force is applied. The bottom plate sensor is able to measure the induced deformation on the sample. The instrument software converts the applied sinusoidal force to sinusoidal shear strain and the sample deformation to sinusoidal shear stress. In addition, it also can be obtained the phase angle delta (δ), which determinates the differential phase between both sinusoidal shear strain and sinusoidal shear stress. Overall the phase angle delta provides information regarding how solid-like or liquid-like is a determined sample [1, 2]. The viscoelastic properties of the gels in chapters 2, 3 and 4, were studied using an ARG2 rheometer with a 20 mm/2° parallel plate geometry. At each-time point, ∼150 µL gel was loaded onto a horizontal plate and then the elastic (G') and viscous (G'') modulus were recorded as function of frequency sweeps between 0.1 to 100 Hz at 0.01-100.00 % strain at 37C. Gel viscoelastic properties measurements for both gels, gel containing cells (for cell densities the reader is referred to chapters 2, 3 and 4), and gel without cells, were studied keeping all variables constant.

Immunocytochemistry (ICC) Immunostaining is a method to study the expression of proteins by cells. Its principle relies on the use of antibodies to specifically target the protein of interest. Within immunostaining assays, one of the most widely used methods is immunocytochemistry (ICC), where cells are stained to be visualised under the

135 microscope, unlike immunohistochemistry (IHC), where tissue sections are studied [3].

Osteoimage mineralisation assay Osteoimage mineralisation assay is a fluorescent in vitro assay, to investigate bone cell mineralisation. This assay, allows the visual determination of mineralisation and its quantification by using fluorescent microscopy and spectrophotometric measurements respectively. The assay is based on the specific binding of the fluorescent Osteoimage staining reagent to the hydroxyapatite portions of bone-like nodules deposited by osteogenic stem cells, primary osteoblasts or bone-like cell lines (http://bio.lonza.com/uploads/tx_mwaxmarketingmaterial/Lonza_ManualsProductInst ructions_Instructions_-_OsteoImage_Mineralization_Assay.pdf).

Mineralisation and collagen type I staining The presence of type I collagen (col-I), and mineralisation deposits was visually determined using an ICC staining. Human osteoblasts were stimulated with osteogenic media containing 10-5 mM dexamethasone (D4902-16; Sigma-Aldrich Co.), 10 mM β-glycerophosphate (G9422-100G; Sigma-Aldrich Co., St. Louis, MO, USA) and 2.83 × 10-7 mM ascorbic acid (A8960-5G; Sigma-Aldrich CO., St. Louis, MO, USA) and human mesenchymal stem cells with Osteogenic Media (Lonza Walkersville Inc., Basel, Switzerland). At each time-point, the gels were fixed with 3% w/v paraformaldehyde (PFA) for 30 minutes at RT. Thereafter, samples were permeabilised using Triton X-100 at 0.05% (v/v) in DBPS for 15 minutes at RT. Samples were subsequently blocked with 1% (w/v) bovine serum albumin (BSA) for 40-50 minutes at RT and incubated for 1h at RT with a primary antibody (pAb) (diluted in 1% BSA) rabbit polyclonal to col-I (Abcam, UK), at the ratio stipulated by the manufacturer (1:250). The samples were incubated for 1h at RT in the dark with a goat anti-rabbit IgG-Alexa Fluor 594 (Abcam) as secondary antibody (sAb) along with Alexa fluor 488 phalloidin to target the protein of interest and F-actin, respectively. For mineralisation assay antibodies were not used. The samples were rinsed in cycles of 5 × 5-10 minutes, with DPBS washing between each step. Samples were mounted on glass slides using ProLong Antifade Reagent (Invitrogen), and images were obtained using a Leica TCS SP5 confocal microscope.

136 Total collagen assay Total collagen assay is based on the colorimetric quantification of hydroxyproline residues obtained via the acid hydrolysis of collagen http://www.quickzyme.com/wp- content/uploads/2012/07/Manual-QuickZyme-total-collagen-assayjuly-2012.pdf. The quantitative assay to monitor the production of total collagen by human osteoblasts and mesenchymal stem cells within gels was determined using a total collagen assay (QuickZyme Biosciences, Park, Leiden, The Netherlands). After resuspending and lysing, samples were vortexed and hydrolysed over 20 h at 95°C with 100 µL 12M HCl. Samples were subsequently diluted 1:1 in 4M HCl and pipetted into a 96-well plate, before adding 75 µL of an assay buffer and incubating for 20 minutes at RT. Following this, 75 µL of a detection reagent was added and the sample transferred into the oven (60°C for 1h). Absorbance measurements were obtained at 570 nm, using a plate reader (Tecan Infinite M200).

Human Osteocalcin Enzyme Amplified Sensitivity Immunoassay (EASIA) The human Osteocalcin EASIA, is an immunoenzymatic assay for the quantitative measurement of human osteocalcin. The principle of the assay relies on the use of monoclonal antibodies directed against distinct epitopes of human osteocalcin. Samples react with monoclonal antibodies covering microtiter wells; subsequently a second monoclonal antibody labeled with horseradish peroxidase reacts with samples (osteocalcin) captured. Bound enzyme labeled antibody is measured through a chromogenic reaction, after the incorporation of a chromogenic solution. The amount of substrate turnover is proportional to the human osteocalcin concentration (http://tools.lifetechnologies.com/content/sfs/manuals/KAQ1381.pdf). Osteocalcin (OCN) production by human osteoblasts and mesenchymal stem cells was determined using a Human Osteocalcin ELISA Kit (Invitrogen–Life Technologies). After the cells were lysed, samples were vortexed and pipetted into a 96-well plate. Subsequently, a working anti-OST-HRP solution (100 µL) was added to each well, the plate covered and incubated for 2 h at RT. Each well was then rinsed three times with a washing solution before adding 100 µL of a chromogen solution (tetramethylbenzidine) and incubating for 30 min at RT in the dark. The reaction was stopped by adding 100 µL of a stop solution (1 N HCl). Absorbance

137 measurements were obtained within 1 h at 450 nm, using a plate reader (Tecan Infinite M200).

Alkaline phosphatase assay The alkaline phosphatase assay relies on the principle that pNPP substrates develop the yellow end-product, p-nitrophenol, when hydrolysed by alkaline phosphatase. This system forms a soluble end product, which is suitable for ELISA. The products of this assay are supplied as ready-to-use buffered alkaline phosphatase substrate solutions containing p-NPP. Alkaline phosphatase activity by human osteoblasts and human mesenchymal stem cells within gels was monitored using a colorimetric alkphos and peroxidase substrate detection system (Sigma–Aldrich Co.). A volume of 20 µL of the cell lysis was added to transparent 96-well plates together with 200 µL of p-nitrophenyl phosphate (pNPP) solution (1 mg/mL pNPP, 0.2 M Tris buffer in 5 mL ddH2O) (SIGMAFASTTM pNPP Tablets, N1891-50SET; Sigma–Aldrich Co.), and the following reaction was stopped with 3 M NaOH. The absorbance of samples was measured using a plate reader (Labsystems Multiskan Ascent; Thermo Scientific, UK) at 405 nm every 30 seconds for 30 minutes.

Alizarin red assay Alizarin red staining is a method that has been used for many years to explore calcium deposits in both histology and cell cultures in vitro. Once osteoblasts or differentiated stem cells produce extracellular calcium deposits, alizarin red binds to the calcium portion of calcium phosphates to form chelates, where a bright orange-red staining is formed, indicating the presence of mineralised tissue. Alizarin red can be chemically extracted form the stained sample via cetylpyridinium chloride (CPC), to be semi quantified using a spectrophotometer [4, 5]. Mineralisation activity by human osteoblasts within gels was evaluated using Alizarin Red staining. At each time-point, the cells were fixed with 70% ethanol for 30 minutes, rinsed three times with ddH2O and then stained with 40 nM Alizarin Red at RT in the dark for 45 min. Samples were then rinsed eight times with PBS. For optical density measurements, 10% cetylpyridinium chloride (CPC) was dissolved in 10 mM sodium phosphate and 1 mL of this was added to each gel before incubating at RT for 30 minutes. A volume of 200 µL of each solution was plated into a clear flat-bottomed 96-well plate (Nunc,

138 UK) and the absorbance at 562 nm was determined using a Labsystems Multiskan Ascent plate reader.

5.3 References [1] Yan C, Pochan DJ. Rheological properties of peptide-based hydrogels for biomedical and other applications. Chem Soc Rev 2010;39:3528-40. [2] Jobling A. An introduction to rheology H. A. Barnes, J. F. Hutton and K. Walters, Elsevier Science Publishers, Amsterdam, 1989. pp. v + 199, Price $6050/Dfl 115.00. ISBN 0-444-87469-0. Polymer International 1991;25:61-. [3] Lerner KLLBW. World of microbiology and immunology. Detroit: Gale; 2003. [4] Puchtler H, Meloan SN, Terry MS. On the history and mechanism of alizarin and alizarin red S stains for calcium. J Histochem Cytochem 1969;17:110-24. [5] Gregory CA, Gunn WG, Peister A, Prockop DJ. An Alizarin red-based assay of mineralization by adherent cells in culture: comparison with cetylpyridinium chloride extraction. Anal Biochem 2004;329:77-84.

139 CHAPTER SIX

Conclusions, outlook and recommendations for future work 6.1 Conclusions

Peptide hydrogels have shown to play a significant role in tissue engineering and more recently in regenerative medicine. Along with the use of mamalian cells including human cells, peptide-based scaffolds are not only capable of inducing phenotypical changes in either mature or progenitor cells, but also directing and improving their physiological functions. Using both celluar and acelluar approaches, peptide hydrogels have showed to induce the regeneration of different tissues, incliding bone, potencially reducing costs, time and complicated treatments that are currently being used to restore damaged tissues. The bottom-up strategy, where amino acids are used as building blocks to create peptide gels is an promising route that can overcome concerns related to cytocompatiblity, osteoconductivity and osteoinductivity. In this work, a hydrogel made of eigth ionic-complementary amino acids, was designed to create a three-dimensional scaffold in an attempt to simplify long peptide sequences, and provide a suitable niche for the culture of human cells. The survival and proliferation of human osteoblasts and human mesenchymal stem cells within the FEFEFKFK gel, were sustained, and the physiological functions of such cells, were also fostered under specific conditions. Cells showed to be able of synthesising bone proteins and mineralising, leading to the formation of a bone-like tissue within hydrogels. In addition, the incorporation of biofunctional ligands/cues from adhesive proteins found in the extracelluluar matrix such as RGD, allows peptide gels mimicking the extracelluar matrix, and directing the cell behaviour as needed, bringing closer a physiological cellular response in vitro. Unlike other systems such as RADA-16, which is composed of 16 amino acids, the FEFEFKFK peptide gel is only made of 8 amino acids. The latter peptide gel is also capable of providing an adequate three-dimensional environment for cells, at the same time, this peptide hydrogel offers higher mechanical properties, which can be easily tuned as needed. A short peptide sequence system is cheaper and easier to design. Lower

140 mechanical properites of other systems such as RADA or MAX1 might limit the study of the cellular response for long time periods. In addition, unlike natural scaffolds such as Chitosan, Alginate Matrigel, etc., when using the FEFEFKFK hydrogel for tissue regeneration puposses, concerns related to cell attachment, gel degradation or batch-batch variability can be easily overcome. Thus, the FEFEFKFK gel arises as an promising route for the regeneration of bone tissue, which might be an interesting alternative to be used in the the regeneration of other tissues in vitro, in vivo or in situ, and subsequently for clinical applications.

6.2 Outlook

In this work it was shown that by simply using a peptide hydrogel, it is possible to provide a suitable environment for cells and induce the production of key components involved in the formation of bone. This opens the possiblity that through tight regulation and balance in the components composing such scaffold, cells can behave more in vivo-like, with promising bone tissue regeneration purposes. Thereafter it would be possible transferring such peptide gel either into an animal model system or in situ, to induce the regeneration of bone (i. e. alveolar bone). Considering that here it was shown that the FEFEFKFK peptide gel succeeded to induce the formation of bone-like tissue by human osteoblasts and differentiated human mesenchymal stem cells, either a celluar or an acelluar approach might be used, where the gel can acts as a three-dimensional scaffold capable of directing the formation of bone and other tissues.

6.3 Recommendations for future work As mentioned earlier, the FEFEFKFK gel showed to support the growth and physiological functions of bone-forming cells. Nonetheless, it is important to consider further aspects regarding the research carried out during this project. The FEFEFKFK hydrogel showed to be osteoinductive and osteoconductive, however studies comparing the effectiveness of the FEFEFKFK hydrogel against the effectiveness of other peptide and natural hydrogels are necessary. Overall, the peptide sequence design of the FEFEFKFK hydrogel provides a neutral matrix, which lacks of charge at physiological pH. This fact can influence the cellular performance. The design of

141 peptide sequences providing either positive or negative charges through the fibril- network of the hydrogel could direct some cellular responses such as adhesion, osteoinductivity and osteoconductivity. Additionally, further experiments are needed to investigate the potential of the FEFEFKFK hydrogel to induce the commitment of mesenchymal stem into other type of cells such as chondrocytes, neurons or muscle cells. Likewise, the culture and study of the response of mature cells cultured within the FEFEFKFK hydrogel will provide significant insights regarding the capability of this material to induce the regeneration of different tissues. Although in this work it was shown that the RGD ligand had positive effects on cell proliferation and spreading, further experiments are needed for testing the concentration effect of RGD on the cell behaviour. For instance, the osteogenic differentiation of hMSCs and the production of ECM by cells might be differently affected in different ways, depending on the concentration of the RGD functionalising the FEFEFKFK peptide fibres. However, this needs to be experimentally tested. In addition, studies using different cell densities are needed to compare the effect of cellular density on the hydrogel mechanical properties. Further investigations for exploring the progressive degradation of hydrogels are required, mainly when cells are encapsulated inside hydrogels. Likewise, exploring the effect of gel degradation on the cell performance (proliferation, production of ECM, etc.) is necessary. This information can help to complement the work presented here, opening the possibility of using this peptide hydrogel not only for the regeneration of bone tissue, but also for the regeneration of various types of body tissues.

142 APPENDIX Standard curves of the protein quantification assays from the Chapters 2, 3, and 4 of this thesis.

0.9" 1.2" y"="0.0026x"+"0.0813" y"="0.0032x"+"0.0236" 0.8" R²"="0.9951" R²"="0.99557" 1" 0.7" 0.6" 0.8" 0.5" 0.6" 0.4" 0.3" 0.4" Absorbance*(570nm)* Absorbance*(570nm)* 0.2" 0.2" 0.1" 0" 0" 0" 50" 100" 150" 200" 250" 300" 350" 0" 50" 100" 150" 200" 250" 300" 350" Total*collagen*(µg/mL)* Total*collagen*(µg/mL)*

Figure 1. Standard curves of the Total Collagen Assay (two-independent experiments) from Chapter 2.

3.5" y"="0.0322x"+"0.1691" 3" R²"="0.97108" y"="0.0301x"+"0.1425" 3" 2.5" R²"="0.98076"

2.5" 2" 2" 1.5" 1.5" 1" Absorbance*(450nm)*

1" Absorbance*(450nm)* 0.5" 0.5" 0" 0" 0" 10" 20" 30" 40" 50" 60" 70" 80" 90" 100" 0" 10" 20" 30" 40" 50" 60" 70" 80" 90" 100" Osteocalcin*(ng/mL)* Osteocalcin*(ng/mL)*

Figure 2. Standard curves of the Human Osteocalcin Assay (two-independent experiments) from Chapter 2.

0.12" 0.12"

0.1" 0.1"

0.08" 0.08" y"="0.0002x"+"0.0934" y"="0.0003x"+"0.0894" R²"="0.99296" R²"="0.98911" 0.06" 0.06" 0.04" 0.04" Absorbance*(405nm)* Abserbance)(405nm)) 0.02" 0.02"

0" 0" 10" 20" 30" 40" 50" 60" 70" 0" 0" 10" 20" 30" 40" 50" 60" 70" Measurement)count) Measurement*count*

Figure 3. Standard curves of the Alkaline Phosphatase Assay (two-independent experiments) from Chapter 2.

143

1.4" 1.2" y"="0.004x"+"0.1071" y"="0.0033x"+"0.0725" R²"="0.98249" R²"="0.99571" 1.2" 1" 1" 0.8" 0.8" 0.6" 0.6" 0.4" 0.4" Absorbance*(570nm)* Absorbance*(570nm)* 0.2" 0.2" 0" 0" 0" 50" 100" 150" 200" 250" 300" 350" 0" 50" 100" 150" 200" 250" 300" 350" Total*collagen*(µg/mL)* Total*collagen*(µg/mL)*

Figure 4. Standard curves of the Total Collagen Assay (two-independent experiments) from Chapter 3.

3" 1.8" y"="0.0144x"+"0.0675" y"="0.0293x"+"0.1777" R²"="0.99082" R²"="0.97406" 1.6" 2.5" 1.4" 2" 1.2" 1" 1.5" 0.8" 1" 0.6" Absorbance*(450nm)* Absorbance*(450nm)* 0.4" 0.5" 0.2" 0" 0" 0" 10" 20" 30" 40" 50" 60" 70" 80" 90" 100" 0" 10" 20" 30" 40" 50" 60" 70" 80" 90" 100" 110" Osteocalcin*(ng/mL)* Osteocalcin*(ng/mL)*

Figure 5. Standard curves of the Human Osteocalcin Assay (two-independent experiments) from Chapter 3.

0.45" 0.14"

0.4" y"="0.0042x"+"0.1317" 0.12" R²"="0.99805" 0.35" 0.1" 0.3" y"="0.0003x"+"0.104" R²"="0.99251" 0.25" 0.08"

0.2" 0.06" 0.15" 0.04" Absorbance*(405nm)* Absorbance*(405nm)* 0.1" 0.02" 0.05" 0" 0" 0" 10" 20" 30" 40" 50" 60" 70" 0" 10" 20" 30" 40" 50" 60" 70" Measurement*count* Measurement*count*

Figure 6. Standard curves of the Alkaline Phosphatase Assay (two-independent experiments) from Chapter 3.

144

1.4" y"="0.0039x"+"0.1025" 1.6" y"="0.0048x"+"0.0665" R²"="0.97762" 1.2" 1.4" R²"="0.9978" 1" 1.2" 1" 0.8" 0.8" 0.6" 0.6" 0.4" Absorbance*(570nm)* Absorbance*(570nm)* 0.4" 0.2" 0.2" 0" 0" 0" 50" 100" 150" 200" 250" 300" 350" 0" 50" 100" 150" 200" 250" 300" 350" Total*collagen*(µg/mL)* Total*collagen*(µg/mL)*

Figure 7. Standard curves of the Total Collagen Assay (two-independent experiments) from Chapter 4.

2.5" 3" y"="0.0223x"+"0.1463" y"="0.0266x"+"0.1535" R²"="0.98065" R²"="0.98983" 2.5" 2"

2" 1.5"

1.5" 1"

1" Absorbance*(450nm)* Absorbance*(450nm)* 0.5" 0.5" 0" 0" 0" 10" 20" 30" 40" 50" 60" 70" 80" 90" 100" 0" 10" 20" 30" 40" 50" 60" 70" 80" 90" 100" Osteocalcin*(ng/mL)* Osteocalcin*(ng/mL)*

Figure 8. Standard curves of the Human Osteocalcin Assay (two-independent experiments) from Chapter 4.

0.14" 0.12"

0.12" 0.1"

0.1" y"="0.0006x"+"0.094" 0.08" y"="0.0002x"+"0.0989" 0.08" R²"="0.99854" R²"="0.96663" 0.06" 0.06"

0.04" 0.04" Absorbance*(405nm)* Absorbance*(405nm)*

0.02" 0.02"

0" 0" 0" 10" 20" 30" 40" 50" 60" 70" 0" 10" 20" 30" 40" 50" 60" 70" Measurement*count* Measurement*count*

Figure 9. Standard curves of the Alkaline Phosphatase Assay (two-independent experiments) from Chapter 4.

145