Enzymatically Synthesized Inorganic Polymers as Morphogenetically Active Bone Scaffolds

CHAPTER TWO

Enzymatically Synthesized Inorganic Polymers as Morphogenetically Active Bone Scaffolds: Application in Regenerative Medicine Xiaohong Wang1,*, Heinz C. Schröder1, Werner E.G. Müller1,*

1ERC Advanced Investigator Grant Research Group at the Institute for Physiological Chemistry, University Medical Center of the Johannes Gutenberg University Mainz, Germany *Corresponding authors: E-mail: [email protected], [email protected]

Contents 1.  Introduction29 2.  Bone Scaffolds 30 2.1  Bioinert Materials 31 2.2  Bioactive Materials 33 2.3  Regenerative Functional and Custom-Made Tissue Units 35 3.  Bone Cells 39 3.1 MSCs/Osteoblasts 39 3.2  Osteoclast Cell Differentiation 41 4.  Biogenic, Morphogenetically Active Inorganic Polymers 43 4.1 Biocalcite 43 4.2 Bio-polyphosphate 43 4.3 Biosilica 47 5.  Enzymes Controlling the Synthesis of Morphogenetically Active Inorganic Polymers: A Paradigm Shift in Bioinorganic Chemistry 50 5.1  Carbonic Anhydrase 51 5.2  Alkaline Phosphatase 57 6.  Biocalcite as Bioseed during Mammalian HA Formation 60 7.  CA Activators as Potential Novel Drugs to Stimulate Bone Mineral Formation 62 8.  ALP Activators—Potential Novel Compounds to Stimulate Bone Mineral Formation? 63 9.  Applications for Bioprinting Organs 65 10.  Concluding Remarks 67 Acknowledgments68 References68 International Review of Cell and Molecular Biology, Volume 313 ISSN 1937-6448 http://dx.doi.org/10.1016/B978-0-12-800177-6.00002-5

© 2014 Elsevier Inc. All rights reserved.

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Abstract In recent years a paradigm shift in understanding of human bone formation has occurred that starts to change current concepts in tissue engineering of bone and cartilage. New discoveries revealed that fundamental steps in biomineralization are enzyme driven, not only during hydroxyapatite deposition, but also during initial bioseed formation, involving the transient deposition and subsequent transformation of calcium carbonate to calcium phosphate mineral. The principal enzymes mediating these reactions, carbonic anhydrase and alkaline phosphatase, open novel targets for pharmacological intervention of bone diseases like osteoporosis, by applying compounds acting as potential activators of these enzymes. It is expected that these new findings will give an innovation boost for the development of scaffolds for bone repair and reconstruction, which began with the use of bioinert materials, followed by bioactive materials and now leading to functional regenerative tissue units. These new developments have become possible with the discovery of the morphogenic activity of bioinorganic polymers, biocalcit, bio-polyphosphate and biosilica that are formed by a biogenic, enzymatic mechanism, a driving force along with the development of novel rapid-prototyping three-dimensional (3D) printing methods and bioprinting (3D cell printing) techniques that may allow a fabrication of customized implants for patients suffering in bone diseases in the future.

ABBREVIATIONS 1,25(OH)2D3  1,25-dihydroxy-vitamin D3 3-D Three-dimensional ALP  Alkaline phosphatase ASP asialoprotein b-ALP  bone-specific alkaline phosphatase BMPs  bone morphogenetic proteins BSP  bone sialoprotein CA  carbonic anhydrase Ca2+ calcium CaCO3 Ca-carbonate CaP  calcium phosphate COLI  collagen type I CTR  calcitonin receptor ECM  extracellular matrix GGCX  γ-glutamyl carboxylase HA hydroxyapatite HCO3− bicarbonate IκBα  nuclear factor of kappa light polypeptide gene enhancer in B-cells inhibitor, alpha M-CSF  macrophage–colony-stimulating factor MAPK/ERK  mitogen-activated protein kinase/extracellular-signal-regulated kinases MSC  mesenchymal stem cells OCAL osteocalcin OPN osteopontin OPG osteoprotegerin

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Pi  inorganic phosphate polyP polyphosphates PPi pyrophosphate PPK  polyphosphate kinase QA  quinolinic acid RANK  receptor activator of nuclear factor κB RANKL  receptor activator of NF-κB ligand R-Smads  receptor-regulated Smads TCP  tricalcium phosphate TNAP  tissue-nonspecific type alkaline phosphatase TRAP  tartrate-resistant acid phosphatase TRIS 2-amino-2-hydroxymethyl-propane-1,3-diol β-GP  β-glycerophosphate WNT/SHH WNT/hedgehog

1.  INTRODUCTION The ultimate goal for any kind of reconstructive surgery had been tissue/ organ repair from ancient times until present. Repair of tissue/organ defects traditionally involves tissue grafting and/or organ transplantation as well as alloplastic or synthetic material replacement. Since early history until the seventeenth-century gold was used as implant material for hard tissue defects (Sanan and Haines, 1997) but also organically based materials, e.g., the organic marine sponge skeletons/scaffolds, were occasionally applied (Camper, 1771). The manifested limitations of those grafts were tried to compensate later by the application of implants based on synthetic materials of inorganic or organic nature. However, those implants very often failed to integrate into the host tissue and showed inherent disadvantage not to be replaceable by the body’s own cells and tissues. In the 1980s, tissue engineering emerged to overcome those limitations by tissue grafting and/or alloplastic tissue repair (Langer and Vacanti, 1993). In the last two decades, the concept of transplanting of compensatory porous and degradable materials, enriched with biofactors (cells, genes, and/or proteins) has been developed (­Hollister, 2005). Attempts that include stem cell approaches and gene therapy approaches followed (­Cutroneo, 2003; Audet, 2004; Anam and Davis, 2013).Very recently morphogenetically active scaffolds, suitable for the three-dimensional (3D) growth of mesenchymal stem cells (MSC) and likewise suitable for bioprinting (3D cell printing), have been developed.This chapter outlines strategies to fabricate tissue units by 3D bioprinting technology, as well as a novel approach to apply activators for the recently elucidated major enzymes involved in the biosynthesis of the bone biomineral, the carbonic anhydrase (CA) and the alkaline phosphatase (ALP).

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2.  BONE SCAFFOLDS It is the task of tissue engineering to develop biologically active organ/tissue substitutes that have the property to restore lost morphological and functional features of impaired or diseased organs. As outlined by Langer and Vacanti (1993) in tissue engineering new developments of functional substitutes for damaged tissue can only be successfully translated into practice if the basic principles of biology and engineering can be amalgamated to the invention. One discipline alone cannot achieve this goal. This interdisciplinary field should provide solutions for tissue creation and repair. From cell biology it is known that cells composing tissues, from the basis of the metazoan kingdom the sponges (phylum: Porifera) to the crown Metazoa, the mammals, and the insects, are not loosely embedded in the tissue but integrated to functional units by controlled and directed cell–cell interactions as well as cell–matrix interactions (Müller et al., 2004). This biological basic construction network for Metazoa allows intracellular as well as extracellular signaling information transmission networks but also via extracellular matrix (ECM) elements, e.g., collagen and fibronectin, concerted circuits that are regulated by biological, physical, and chemical cues of the microenvironment. Those tuned interactions provide the critical platform for integrated cell functions and behaviors. In turn, scaffolding materials to be designed and to be intended for tissue engineering applications must mimic those physiologic environments. Even more, these circuits not only integrate the cells between soft tissue and hard tissue but also, and there especially, determine the 3D geometrical, topographical, as well as physical units. Physiological matrices as well as fabricated 3D scaffolds are the crucial prerequisites to elicit, induce, and trigger the cells with the physiologically relevant stimuli in order to establish, maintain, and further develop their functionalities to associate and to build tissue. In turn, a shift of paradigm is presently proceeding which builds on (1) the collected experiences from the established bioinert scaffold materials as well as (2) the knowledge on bioactive materials and is culminating in (3) the development of regenerative and custom-made biosynthetic implants and tissue grafts. It is the aim to fabricate scaffolds for engineered tissue units using biodegradable soft and simultaneously porous materials that allow the embedding and integration of biological cells with growth and differentiation factors, exogenously added or synthesized by the cells themselves.

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2.1  Bioinert Materials The application of bioinert materials for tissue and bone tissue replacement is old. Especially the application of biomaterials from marine animals as plaster and in bone/tissue replacement has been a tradition since the Greek times. Camper (1771) described that the organic matrix of sponges can be successfully used in plastic surgery of the palate of the skull (Figure 2.1(a)). He was fabricating a nose using lime wood, covered it with a sponge and fixed it in the roof of the mouth via a silk thread which had been waded by small sponge slices. It might also be noted that the application of the siliceous skeleton of sponges as suitable scaffold onto which human stem cells can be seeded has recently been reported (Green et al., 2003). Basically two basis materials have been used to fabricate implants for bone reconstitution/reconstruction; (1) metals and (2) ceramics. In the first phase of those replacement supports, the intention for the use of those implants had been to stabilize the body at the position of the damaged bone (Chang et al., 1996). For decades metal implants have been used in orthopedics for mechanical skeletal repair. Those supports had to meet the challenge to strengthen the implant–bone interface and to prevent stressshielding effects. Those implants can be fabricated in customized processes, e.g., by 3D printing (Figure 2.1(b)). The key issue for a durable and successful implant is the establishment of a strong bone–implant interface. It emerged that smooth implant surfaces can result in the formation of encapsulation with the consequence of loosening of the implant (Greco, 1994). In one approach to extend and to promote long-term interface strength, porous materials and porous coatings have been developed (Engh and Bobyn, 1985). Those porous materials and coatings induce a partial to complete bone ingrowth, which has the advantageous property to enhance the strength of the interface bonding under simultaneous reduced tendency to cause capsule formation around the implant. A further challenging issue for most of those metalbased implants is the appearance of “stress shielding” (Jacobs et al., 1993; Amstutz, 1991). Even though it is well established that bone regeneration and repair processes are promoted by mechanical loads (Van Lenthe et al., 1997) metal materials such as titanium, still widely used today for bone implants, is much stiffer than native bone. Consequently, an implant of solid titanium can carry a disproportionate amount of the biological loads (see: Thelen et al., 2004). In turn, the surrounding bone undergoes a process of “stress shielding” and suffers from abnormally low levels of

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A

B

C

D

Figure 2.1 The development of bone scaffolds. (a) Bioinert materials, e.g., application of sponges in tissue replacement (Camper 1771). The damaged nose/nostrils (A, B, D, C) was modeled by a piece of lime wood covered with a sponge (T, U, V), and fixed in the roof of the mouth (W) via a silk thread (S) that had been surrounded with small sponge slices. (b) Computer-aided rapid prototyping/3D printing. (A) Data are generated for an organ or tissue unit using the computing process. Algorithms for the automated design and fabrication of a scaffold/bone part are developed based on an assembly-free process. (B to D) The bone unit is printed, like in an ink-jet printer, and comprises a tightfitting customized morphology. If it is formed of ceramic or titanium, the implant has usually only an osteoconductive property. (c) Fabrication of an osteoinductive scaffold made of glass, metal, or ceramic that allows the cells to migrate into its pores. The scaffold is coated with bioactive factors or polymers, e.g., bone morphogenetic proteins-2 or polyphosphate, that direct its associated cells, stem cells, to terminally differentiated cells that build tissue units like blood vessels.

stress, which finally results again in bone resorption, followed by loosening of the implant (Black, 1999). A further achievement toward a more advanced material to be used for orthopedics is the use of bioinert ceramics (Boutin, 1981). Ceramic, an

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inorganic, nonmetallic solid that is prepared by the action of heat and subsequent cooling, has been used since the 1970s in orthopedics (Smith, 1963; Eyring and Campbell, 1969). As material, ceramics could have a crystalline or partly crystalline structure, or be amorphous (e.g., a glass). Ceramics are increasingly used in the orthopedic surgery, especially for joint prostheses. It is used not only for joint bearings, but also for the bone–implant interface of prostheses (see: Hayashi et al., 1993), especially for implants as knee prosthesis, total ankle prosthesis, or total elbow prosthesis. The bioinert ceramics have been found to have the property of excellent resistance during carrying (Kumar et al., 1991). Since the beginning of the development of ceramic implants it is hoped that this material is more biocompatible than metal alloys since it is provided with the property of resistance to corrosion, to be less cytotoxic and to be hydrophilic. At present synthetic biodegradable polymers, interconnected with porous calcium hydroxyapatite (HA) ceramics have been found to be very suitable composite materials for implants, since they can be combined with growth and development factors in a carrier/scaffold system, e.g., with recombinant human bone morphogenetic protein-2 (rhBMP-2), that strongly promotes the clinical effects of rhBMP-2 in bone tissue regeneration (Bianco and Gehron, 2000). This study, like others that appeared during this time period (Hollister, 2005), predetermines the development to a second level of regenerative restorative implantology the “bioactive materials.”

2.2  Bioactive Materials The strategy to design more effective bioactive tissue engineering scaffolds is to implement in a complex manner three essential elements, first, a porous matrix (scaffold), second, to elicit osteoconductive signals, and third, to implant, adjacent to the matrix, osteogenic cells that can attach to the matrix and respond to their signals via an adequate blood supply (Finkemeier, 2002); Figure 2.1(c). In turn, the implants must be designed in a hierarchical way; first hierarchical porous structures must be prepared that are provided with a suitable mechanical stability and flexibility and allow the transfer and diffusion of growth factors and differentiation factors. The complex 3D anatomical shape of the bone substitution material must try to imitate from the nanometer to the millimeter level the functional properties of the natural bone.The scaffold formed must ideally meet the requirements of the cells to be provided with nutrients. The porous channels must allow cell migration, and their surface features must be suitable for cell attachment (Cukierman et al., 2001). The morphology/topography as well as the

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roughness of the scaffold surface should fall between the theoretical maximum given by the material and the theoretical minimum of zero predicted by composite theories (Torquato, 2002; Hollister, 2005). Consequently, the critical issue for the design of the surface texture is based on computer calculations that match the requirements to allow intercellular and transcellular signal transmission as well as to leave space for the development of a vascular system that allows mass transport. An effective permeability is determined by the 3D pore arrangement and the adhesion receptors associated with the plasma membrane, e.g., the integrins. The biological properties of the surfaces of the, basically inert, matrices are crucially important.The material must be bioactive along an increasing complexity and inducibility, from being osteoconductive to osteoinductive and allowing processes that cause osseointegration. According to the definition by Albrektsson and Johansson (2001) the term osteoconduction means that bone grows on a surface, e.g., bone surface, that supports the ingrowth of the osteoblasts into pores, channels, or pipes (Wilson-Hench, 1987). However, more often the surface used is not the bone but another biogenic growth platform (Glantz, 1987). The next higher level of bioactivity is osteoinduction. This term stands for the activity of a contact or soluble material that displays the potency to induce the undifferentiated and pluripotent stem cells to enter the differentiation pathway toward the bone-forming cell lineage, more specific to induce osteogenesis. Finally, osseointegration refers to the process by which the implant is stably anchored into the bone. At present and strictly speaking, only the bone morphogenetic proteins (BMPs) being members of the transforming growth factor-β (TGF-β) superfamily of growth factors and well established physiological regulators of osteoblastic differentiation (Lavery et al., 2008) can be considered as osteoinductive. Among them BMP-7 and BMP-2 display the highest osteoinductivity (e.g., Urist, 1965; Lavery et al., 2009; Sampath et al., 1990). Already in limited clinical use is BMP-2 that has been shown to induce new bone formation in spine fusions and long bone nonunion fractures (Gautschi et al., 2007). After binding of BMP to the integrated cell surface receptor, a tetramer of serine/threonine kinase transmembrane receptors consisting of two type I and two type II receptors, intracellular signaling occurs via intracellular signaling proteins to the receptor-regulated Smads (R-Smads). In turn R-Smads form heteromeric complexes with the common mediator Smad, Smad-4, and subsequently translocate to the nucleus where they act as transcription factors to induce the BMP responsive genes (Sebald et al., 2004). This implies that any kind of scaffold supplemented/coated with BMPs must be qualified to be

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an osteoinductively acting implant. However, recently it becomes overt that in osteoblasts also BMP-independent anabolically acting routes exist. Examples are polyphosphate (polyP) and biosilica-mediated pathways (Wang et al., 2014a,b), natural inorganic polymers that have the capacity to induce BMP-2 in osteoblasts. In addition, a BMP-independent route has been proposed that induces the differentiation of osteoblast precursor cells to mature functionally active osteoblasts (Müller et al., 2013a). It also appears likely that the phytoestrogen isoquercitrin acts synergistically with polyP, on the transcription factor RUNX2 (Wang et al., 2014c). In line with a broader interpretation of osteoinductivity (Amini et al., 2012) biomaterials, including natural and synthetic ceramics (i.e., HA and various calcium phosphate (CaP) compositions, and their composites), have been qualified as osteoinductive materials. Besides of synthetic CaP-based biomaterials, also in the form of sintered ceramics (Yamasaki and Sakai, 1992; Klein et al., 1994), cements (Habibovic et al., 2008), and coatings (Habibovic et al., 2004), natural and coral-derived ceramics (Ripamonti, 1991; Ripamonti et al., 2009) have been attributed to be osteoinductive. Especially highlighted should be porous bioglass that has been widely introduced into clinics (Jones, 2013). This bioactive ceramics, a biodegradable glass of a general formula “Na2O–CaO–SiO2–P2O5,” contains high levels of calcium (Ca2+); its most generally used formulation contains 46.1 mol% SiO2, 24.4 mol% Na2O, 26.9 mol% CaO, and 2.6 mol% P2O5 and has been termed 45S5 Bioglass (Hench et al., 1971). The bioactive glasses are reported to stimulate bone regeneration to a larger extent than other bioactive ceramics. This 45S5 Bioglass forms a semichemical bond with bone and, in vivo, bonds to other bioceramics. In comparison, the CaP-based materials are more widely used in the clinics.

2.3  Regenerative Functional and Custom-Made Tissue Units Already a decade ago it has been prognosticated that tissue engineering technology, based on computer-aided jet-based 3D organ/tissuelike printing, could be a solution of the organ transplantation crisis (Mironov et al., 2003). However, it is needed to distinguish between inert implants that cannot be replaced by cells/tissue of the recipient and functional graft substitutes that are disintegrated and subsequently replaced by cells and ECM filaments ingrowth from the surrounding tissue of the recipient. Many types of CaP biomaterials have been developed that comprise a similar composition like native bone mineral and its precursors, e.g., HA and α- and β-tricalcium phosphate in the form of ceramics, cements, and thin

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coatings (LeGeros, 2002). A few of these insoluble CaP materials are even osteoconductive and in some cases provided with the ability to induce bone formation, implying that those materials are osteoinductive (Damien and Parsons, 1991; Habibovic and de Groot, 2007). The implant, even though acting to a limited extent osteoinductively, remains as a major core in the previously damaged region.The process of biodegradation around the bone graft substitutes is favored since they prevent the disadvantageous resorption of the neighboring bone due to stress-shielding effects. The technique of free-form fabrication applicable for producing of 3D synthetic bone graft substitutes allows a precise control of the overall geometry and in turn also of the porous structure of the scaffold/implant. The 3D printing of CaP-based structures at room temperature has been successfully demonstrated (Gbureck et al., 2007). By modification of the surface of the implant, either with respect to the morphological, ultrastructural, or chemical properties, the capacity of the material to induce ectopic bone formation can be improved. The fabrication of 3D implants by direct cell printing based on computer-aided design files offers a sophisticated and challenging direction to engineer 3D tissuelike units, to be placed into living human organs (Figure 2.2). However, those approaches require three sequential hierarchical steps of increasing complexity. First, compilation of preprocessing followed by the development of computational “blueprints” of a given organ; second, processing or the program file for the printing of the actual organlike unit; and finally, postprocessing or organ conditioning and ultimately organ maturation. In recent years several types of cell printers have been developed (Tasoglu and Demirci, 2013). The basic principle is that the cells, either in suspension or as aggregates, are embedded into a printable matrix which is then sequentially layered under formation of predesigned blocks. It is surely feasible to fabricate implantable, by rapid-prototyping, 3D organlike units in the future; but it remains open when this concept of tissue engineering can be exploited and integrated into the constraints emerging from the biological, genetic rules of developmental biology. It is just extraordinarily difficult to tailor a suitable matrix into which the cells can be embedded to become provided with the required physiological solute and fibrous extracellular molecules. Only after deciphering the genetic blueprint of the cells and its time- and space-specific expression it will become possible to lay the ground with the chemical and physical cues for the stem/precursor cells to direct them toward an integrative assembly in an organ.

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Figure 2.2  Regenerative functional and customized tissue units bioprinted by the 3D cell printing approach. Sketch of the bioprinting of cells, embedded into a matrix, e.g., alginate/gelatin, using a 3D bioplotter. A cell suspension is filled into a cartridge hooked to the printing head (a). This control element is connected with the computer-guided printing apparatus; the alginate/gelatin/cells are passed through a needle into a CaCl2 bath, which hardens the scaffold (b, c). This scaffold with the bioprinted cells is submersed into medium/serum. Then the 3D scaffold is overlayed with an agarose layer containing the morphogenetically active factors or polymers, e.g., polyP (d). In such an environment the cells proliferate and differentiate.

For 3D bioprinting a critical size of the aggregates has to be intended. A cell density within the organic matrix of >106 cells/ml, according to our experiences, is preferable (Neufurth et al., 2014). The size of the aggregates formed within the scaffold matrix is determined by the supply of nutrients and growth factors and/or morphogenetically active polymers. Usually the aggregates are spherical and can reach sizes of a diameter of 500 μm. Incubation conditions must be developed that are favorable for the cells to spread onto a cell substrate, e.g., fibronectin or collagen. In turn, cell–cell and cell-substrate adhesion must be finely tuned; the cells must express the property to decrease substratum adhesivity while simultaneously allow increasing cell–cell cohesivity, and vice versa (Ryan et al., 2001). In addition, the cell environment should be tailored in a way that one cell type is directed towards cell–cell cohesion while the other cell type undergoes intensive cell spreading. The cell aggregates formed after bioprinting can be composed homocellularly, comprising only one type of cell or heterocellularly, being a hybrid composed of more than one cell type (Figure 2.3(a) and (b)). If homocellular aggregates are formed in a bioprinted matrix the subsequently formed

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Figure 2.3  Bioprinting of cells, embedded in an organic matrix. (a) Either homocellular aggregates are allowed to be formed that have the potency to form basic but simple building units. Alternatively, (b), heterocellular aggregates are allowed to be formed that can differentiate to functional units, e.g., comprising vascular structures (v), within cells derived from a different stem cell lineage. (c) It is still an ambition to bioprint cells of different stem cell origin that can functionally differentiate to different cell types that build an organ.

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tissue structures can show different morphologies but hardly form functional organ units (Figure 2.3(a)). In contrast, if heterocellular aggregates are allowed to be formed within the matrix the formation of internal structures can be expected, e.g., blood vessel within endothelial and smooth muscle cells. Alternatively, peripheral blood stem cells can be used that can differentiate either into endothelial cells or smooth muscle cells; endothelial progenitor cells are present in peripheral blood stem that can be induced to cells supporting neoendothelialization (Sugaya et al., 2012). At present it seems to be manageable to sequentially bioprint cell types with a different phenotype (Figure 2.3(b)). The solution of choice in the future will be to bioprint cells that, based on their cell adhesion properties and origin from a given stem cell lineage, proliferate and in parallel differentiate to a complex organ (Figure 2.3(c)).

3.  BONE CELLS Bone appears to be a solid, rigid organ; however, it is highly flexible and dynamic allowing bone anabolic and bone catabolic processes to proceed in a tuned interacting manner. The bone is under continuous remodeling that takes place throughout the life span of an individual. It should be highlighted that the morphology and the shape or the size of bone is genetically determined. Until now it remains unclear which genetic blueprint controls the form-giving processes in bone. Under physiological conditions the net balance between osteoblastic bone formation, mediated by osteoblasts, and osteoclastic bone resorption, driven by osteoclasts, is very much tuned. The bone anabolic cells, the osteoblasts, originate from MSCs having the potential to proliferate and the capacity to differentiate into several connective tissue/cell types. In contrast, the bone catabolic cells, the osteoclasts arise from hematopoietic stem cells (Teitelbaum, 2006).

3.1 MSCs/Osteoblasts The pluripotent MSCs have the potency to differentiate into osteoblasts, chondroblasts, bone marrow stromal cells, fibroblasts, muscle cells, or adipocytes depending on the presence of the growth and differentiation factors in their microenvironment (Wang et al., 2014a); Figure 2.4. Osteoblasts having a cuboidal or columnar shape are lining the bone surfaces at those sites that undergo active bone formation during bone development or fracture repair. Osteoblasts express high levels of type I collagen (COLI) and proteoglycans (glycosaminoglycans), the two main components of the bone matrix, also

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Figure 2.4  Multipotent differentiation of multipotent human mesenchymal stem cells (hMSC). Specific signaling molecules and growth factors as well as differentiation factors induce/activate transcription factors and by that determine both the commitment and the differentiation of hMSCs toward the osteogenic, chondrogenic, adipogenic, or myogenic lineage. The osteogenic and the chondrogenic lineages are involved in the restorative repair of bone and cartilage tissue (osteochondral tissue reconstitution). Biosilica and polyphosphate (polyP) display anabolic, morphogenetic effects on those two differentiation lines. BMP-2, bone morphogenetic proteins-2; ALP, alkaline phosphatase.

termed osteoid. Osteoblasts are also involved in mineralization of osteoid, very likely via the liberation of matrix vesicles, and by the deposition of calcium, carbonate, and phosphate (Landis et al., 1993; Hohling et al., 1978; Müller et al., 2013b). Osteoblasts are aligned by adherens-type junctions, including desmosomes and tight junctions (Safadi et al., 2009). Osteoblasts synthesize and secrete a variety of cytokines and colony-stimulating factors controlling myelopoiesis, e.g., interleukin-6, interleukin-11, granulocyte–macrophage colony-stimulating factor and macrophage–colony-stimulating factor

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(M-CSF). In addition, osteoblasts synthesize a series of growth factors, including TGF-β, BMPs, platelet-derived growth factors, and insulin-like growth factors. Finally, terminally differentiated osteoblasts possess receptors for the parathyroid hormone as well as for 1,25-dihydroxy-vitamin D (1,25(OH)2D3), the major hormones regulating bone metabolism and mineral deposition (Figure 2.5).

3.2  Osteoclast Cell Differentiation Osteoclasts, originating from the hematopoietic lineage (Boyle et al., 2003), undergo differentiation and maturation in the presence of the M-CSF and of the receptor activator of NF-κB ligand (RANKL). Markers for the multinucleated osteoclasts are the highly expressed tartrate-resistant acid phosphatase (TRAP), as well as calcitonin receptor and integrin avb3 (Cerri et al., 2003); Figure 2.5. The cytokine/receptor triad, RANKL with its receptor (RANK) and the endogenous decoy receptor osteoprotegerin (OPG) crucially control bone formation and bone remodeling (Boyce and Xing, 2008; Santini et al., 2011). While RANKL is synthesized by the osteoblastic lineage cells this signaling molecule is essential for the differentiation of those cells that are involved in bone resorption, the osteoclasts. RANKL is expressed on osteoblasts, T cells, dendritic cells, as well as their precursors from where it can be released by specific proteases (Zhang et al., 2009).This ligand (RANKL) binds to the cell surface receptor RANK, located on precursor and mature osteoclasts and by that promotes osteoclastogenesis. After binding of RANKL to RANK the osteoclasts become activated and resorb bone mineral; during this process the cells have close contact to the bone surface (Fuller et al., 2010). At this interphase, osteoclasts to bone, vesicles are formed, via integrin (avb3), that contain proton pumps and acid hydrolases, e.g., cathepsin K. Those enzymes and vesicles are present in those cells that are bone-apposed. “Resorptive hemivacuoles” are formed between osteoclasts and bone, allowing the protons to dissolve the HA scaffold of the bone (Figure 2.5). The intracellular pH is kept close to neutral via the chloride/bicarbonate exchanger.The function of RANKL is under control of OPG, a decoy receptor that is secreted by stromal cells and also by osteoblasts (Kearns et al., 2008). OPG scavenges RANKL by binding to it and neutralizes its function. In turn it has to be concluded that any deregulation of the tuned expression of the RANKL/RANK/OPG system causes a dysregulation of the differentiation pathways of the osteoblasts and the osteoclasts and in turn promotes catabolic bone remodeling (Boyce and Xing, 2008). By that, OPG prevents bone matrix from excessive

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Figure 2.5 Differentiation of the progenitor cells of the bone-forming osteoblasts (osteoblastogenesis) and the bone-resorbing osteoclasts (osteoclastogenesis). Upper panel: Osteoblast differentiation starts from the mesenchymal stem cells and ends with the osteocytes. The major transcription factor Runx2, which is under the control of bone morphogenetic proteins-2, is synthesized in chondrocytes and causes a stage-­ dependent increase in the structural and functional proteins in osteoblasts, for example, b-ALP (bone-specific alkaline phosphatase), COLI (collagen type I), OP (osteopontin), ASP (asialoprotein), BSP (bone sialoprotein), and OCAL (osteocalcin), as well as RANKL (receptor activator of NF-κB ligand). Lower panel: Principle differentiation stages from the hematopoietic stem cells via preosteoclasts to functionally active, bone-resorbing osteoclasts. The osteoblasts direct the preosteoclasts to the osteoclast through the interaction of RANKL with RANK (receptor activator of nuclear factor κB), an interaction that is blocked by OPG (osteoprotegerin). Differentiation from hematopoietic stem cells starts via activation of the PU.1 transcription factor and inflammatory signals. The CD34+ osteoclast precursor cells, after entering the circulating system and in the presence of M-CSF (macrophage–colony-stimulating factor) and 1,25-dihydroxy-vitamin D3 (vitamin D3), become recruited onto the surface of bone. The preosteoclasts, after the stimulation of the DAP12 adapter protein/receptor undergo multinucleation to the osteoclasts. Those cells express in the presence of 1,25-dihydroxy-vitamin D3 the receptor RANK. After binding of RANKL to RANK the osteoclasts dissolve HA by lowering the pH. Markers for the activated osteoclasts are TRAP (tartrate-resistant acid phosphatase) and CTR (calcitonin receptor). HA, hydroxyapatite.

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resorption by binding to RANKL and in turn abolishes the activation of the osteoclasts via inhibition of the RANK pathway. In conclusion, the relative concentrations of RANKL and OPG in bone are the major morphogenetic determinants of bone mass and strength.

4.  BIOGENIC, MORPHOGENETICALLY ACTIVE INORGANIC POLYMERS 4.1 Biocalcite Exemplarily the beneficial function of the calcareous corals in bone reconstruction has been demonstrated (Cooper et al., 2014). Especially the effects of secondary metabolites from soft corals acting against inflammation and tumor growth have been highlighted (Chen et al., 2013). Well understood is the inductive osteogenic differentiation effect of coral scaffold on MSCs (Puvaneswary et al., 2013). Even, with respect to some biological markers, calcareous scaffolds, derived from corals, have been found to be superior in comparison to bone grafts. A significantly higher level of expression of the osteogenic differentiation markers, ALP, osteocalcin (OCAL), and osteonectin, as well as of the transcription factor Runx2 has been described. Even more, the extent of mineralization within coral grafts has been found to be more extensive compared to bone grafts. Further studies revealed that coral products have a curative potential on bone deficits as well. The naturally occurring calcium, within the calcareous scaffold, in the form of aragonite found in the scleractinian hard corals and in the form of calcite deposits within the soft octocorals, contribute to anabolic bone restoration. This effect is especially pronounced if these minerals are administered together with zeolite, a microporous mineral, in mice induced to a menopausal state (Banu et al., 2012); this effect has been confirmed in rabbits as well (Parizi et al., 2012). Parallel with the effect of coral minerals on bone formation, their effect on dental deformities has been studied (Figueiredo et al., 2010). The data indicated that the coral skeleton in its unrefined form cannot be applied due to its necrotic potential. However, if purified coral minerals are applied a beneficial osteogenic effect on bone marrow stromal cells is seen; an effective repair of mandibular defects in canines has been reported (Yuan et al., 2010).

4.2 Bio-polyphosphate Like the chemically synthesized inorganic polymeric phosphate, polyP, the biogenic polyphosphate (bio-polyP) has an amorphous state (Kulaev et al.,

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2004; Rao et al., 2009; Omelon and Grynpas, 2008). In contrast to the chemically synthesized polyP, which is synthesized at high temperature, the biogenically produced bio-polyP is synthesized at ambient physiological conditions (Rao et al., 2009) via polyP kinases; Figure 2.6(a). The biopolymer bio-polyP is found in a wide range of organisms, including bacteria, fungi, algae, plants, and animals (see: Rao et al., 2009); it is readily water soluble in millimolar concentrations at chain lengths <100 phosphate units (Van Wazer, 1958; Rao et al., 2009). The natural bio-polyP is synthesized as a linear polymer of phosphate residues linked together via anhydride linkages from tens to hundreds of units. Even though in the polymer the phosphate groups are linked via phosphoanhydride bonds, the polymer is stable over wide temperature and pH ranges (Kulaev et al., 2004). PolyP is

Figure 2.6  The proposed polyP/bio-polyP metabolism in mammalian/bone cells. (a) Outline of the intracellular polyP metabolism, storage, and subsequent release to the extracellular space where ortho-phosphate serves with Ca2+ as substrates for HA synthesis. (b) Tuned interaction between osteoblasts (HA anabolic pathway) and osteoclasts (HA catabolic pathway). It is outlined that bio-polyP supports the progression of precursor osteoblasts to mature osteoblasts by induction of the genes encoding BMP-2 and b-ALP, followed by the increased release of b-ALP; furthermore OPG synthesis is stimulated. It is assumed that polyP activates a hypothetical TF (transcription factor). ALP hydrolyzes both polyP (Ca2+ salt) and β-GP (β-glycerophosphate). The CBE (chloride-bicarbonate exchanger) in concert with the CA (carbonic anhydrase) is involved in the homeostasis of the intracellular CO2 concentration and pH level. BMP-2, bone morphogenetic proteins-2; b-ALP, bone-specific alkaline phosphatase; OPG, osteoprotegerin; HA, hydroxyapatite; RANKL, receptor activator of NF-κB ligand.

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successfully used as food additive and a base material for cosmetic products (Omoto et al., 1997). The nutritional benefit of this polymer has been substantiated in animal experiments (Lee et al., 2008, 2009). Since polyP is a multivalent anion, it binds to the biologically essential cations Ca2+, Mg2+, Mn2+, Fe2+, and Co2+ (see: Rao et al., 2009); even more, polyP acts as a strong Ca2+ chelator and as an antioxidant. In turn, for in vitro studies polyP should be added as a salt, by mixing polyP together with CaCl2 in a stoichiometric ratio of 2:1 (polyP:CaCl2); this salt is designated “polyP·Ca2+complex” (Müller et al., 2011). The biological function of polyP has been studied in microorganisms and more recently also in animals (Kulaev, 1979; Wood and Clark, 1988; Schröder and Müller, 1999). It has been proposed that polyP acts as a storage substance of energy, as a chelator for metal cations, as a donor for sugar and adenylate kinase, and as an inducer of apoptosis; in addition, it is involved in mineralization processes of bone tissue (Schröder et al., 2000). Recently, some data have been presented that polyP acts as a modulator of gene expression (Usui et al., 2010). The studies indicate that in the osteoblastlike cell line, MC3T3-E1, polyP causes an increased gene expression of OCAL, osterix, bone sialoprotein, and tissue-nonspecific alkaline phosphatase (TNAP), all proteins known to be crucial for bone formation (Sinha et al., 2010; Sun et al., 2005). It remained unclear whether polyP causes the increased skeletal mineralization in its polymeric form or as monomeric phosphates that are formed from polyP through hydrolysis by phosphatases (Omelon et al., 2009). The susceptibility of polyP for phosphatases is well established (Lorenz et al., 1994a,b, Lorenz et al., 1997). As one consequence of the enzymatic hydrolysis of polyP, a release of Ca2+ ions has been proposed; this cation is metabolically utilized during HA formation (Omelon and Grynpas, 2008). The possible modulating effect of phosphatases on the size of polyP is also notable with respect to a probable parallel effect of the enzyme on the amount of the extracellular β-glycerophosphate (β-GP), which is a well established component required for biomineralization in vitro, for HA formation in mammalian cells in vitro (Chung et al., 1992). However, under in vivo conditions, β-GP is rapidly and virtually completely degraded to phosphate prior to or during the initial phase of mineralization (Bellows et al., 1992). Very recently, we could demonstrate that bio-polyP displays morphogenetic activity on bone-forming osteoblasts, SaOS-2 cells, and inhibitory activity on RAW 264.7 cells acting as osteoclasts (Figure 2.6(b)). The osteoblast-like SaOS-2 cells form HA crystals, in response to exposure to

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bio-polyP, based on their potency to express key molecules known to control HA formation (see: Wiens et al., 2010a; Müller et al., 2011; Wang et al., 2013a), e.g., the BMP-2, an inducer of bone formation, OPG, a cytokine that is expressed in osteoblasts with a significant role in the maturation of osteoclasts as well as in the control of bone mineral density, and RANKL, a mediator that binds to RANK which is a receptor that mediates maturation of osteoclasts. In turn, the relative concentration ratio between OPG and RANKL is crucial for the differentiation and survival of osteoclasts, since, as outlined, OPG can bind to RANKL and by that inactivates its function. Hence, the OPG:RANKL ratio controls the osteoinductivity on the level of RANKL, a decisive ligand required for the differentiation of osteoclasts. Likewise, the osteoclast-like RAW 264.7 cells have the potency to readily differentiate into osteoclasts when they are exposed to recombinant RANKL and, by that, have been successfully used as a model for studies of osteoclastogenesis in vitro. It is important to mention that bio-polyP can induce ALP, an enzyme which provides inorganic phosphate required for HA synthesis (Müller et al., 2011). The bio-polyP·Ca2+ complex was found to be a strong inducer of HA formation in SaOS-2 cells and, in particular, to cause an enhanced expression of BMP-2. In parallel, bio-polyP•Ca2+ strongly inhibits the progression of RAW 264.7 cells into osteoclasts, which is reflected by the reduction of cells expressing TRAP, a well established marker protein for terminally differentiated osteoclasts. As an additional endpoint marker for osteoclast differentiation, the effect of bio-polyP•Ca2+ on the function of IκBα kinase (IκBα, nuclear factor of kappa light polypeptide gene enhancer in B-cells inhibitor, alpha) was determined. This kinase is one key molecule which causes the activation of NF-κB during RANKL-caused (pre)osteoclast differentiation (Wang et al., 2013a). The results revealed that bio-polyP•Ca2+ inhibits at low concentrations (10–100 μM) the phosphorylation, and by that, the signaling function of IκBα via the respective kinase in RAW 264.7 cells. These data show that bio-polyP affects the tuned balance between osteoblasts and osteoclasts in the anabolic direction, implying that HA synthesis is favored at the expense of HA degradation/dissolution. The biopolymer, polyP, is synthesized and degraded enzymatically via the polyphosphate kinase (PPK), polyphosphate:glucose-6-phosphotransferase, exopolyphosphatase, polyphosphate:adenosine monophosphate phosphotransferase, 1,3-diphosphoglycerate:polyphosphate phosphotransferase, tripolyphosphatase, polyphosphate glucokinase, and endopolyphosphatase (Rao et al., 2009). The PPKs had been considered as the key enzymes since they are capable of reversibly shifting both energy and phosphate, storage

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or consumption, of phosphate energy control. The key polyP-metabolizing enzymes have been discovered not only in prokaryotes but also in Metazoa. The Ca2+ salts of polyP are less soluble compared to the Na+ or K+ salts; in turn the Ca2+ salts of polyP are stored in vesicles, especially in vesicles of bone cells (Boonrungsiman et al., 2012). Released from the storage vesicles within bone or their neighboring cells, polyP is transported into the extracellular space, where it undergoes degradation to orthophosphate via the bonespecific alkaline phosphatase (b-ALP). In this compartment, ortho-phosphate together with Ca2+ will serve as substrate for HA formation (Müller et al., 2011). Since the initial sites of HA formation might be present within cells (Mahamid et al., 2011), similar mobilization and deposition reactions can be proposed here also. A schematic outline of the polyP metabolism and the connection with HA deposition onto bone cells is given in Figure 2.6.

4.3 Biosilica Based on the experimentally established finding that all metazoans originate from one ancestor, to which the siliceous sponges are the most closely related to (Müller et al., 2004), it has been postulated that the siliceous skeleton of sponges shares functional relationship to the Ca-based skeletons of vertebrates (Müller, 2005; Müller et al. 2009). In addition, evidence has been presented indicating that silicate/silicon is an essential trace element in vertebrate nutrition (Schwarz and Milne, 1972; Van Dyck et al., 2000; Carlisle, 1986; Jugdaohsingh, 2007). Even more suggestive has been the result that silicon deprivation results in severe skeletal malformations (Carlisle, 1972). Those experimental studies in chicken showed that it is the connective tissue, which shows the highest silicon concentrations, in contrast to heart or muscle tissue, where the silicon concentrations are much lower. Moreover, a spatial correlation could be established between the areas of bone formation within animal tissue and the accumulation of silicon (Müller et al., 2009). In the lowest metazoan phylum the siliceous sponges (Hexactinellida and Demospongiae) silica, or more distinctly termed biosilica since it is biogenically synthesized (Morse, 1999), is formed enzymatically using the enzyme silicatein (Wang et al., 2012; Müller et al., 2013c). Considering the common body plan from the sponges (Porifera) to the higher Metazoa, it became overt that biosilica might display a common morphogenetic potential during skeleton formation in the animal kingdom. Based on these data, it was concluded that a burst of silicon accumulation occurs around the osteoid and osteoid–bone interfaces, implying that this

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inorganic component is essential for bone formation. Consequently, our group studied the effect of biosilica on the activity of osteoblasts and osteoclasts in vitro. SaOS-2 cells were grown in the mineralization activation cocktail (composed of 5 mM β-GP, 50 mM ascorbic acid, and 10 nM dexamethasone) on a support, coated either with HA or with biosilica. The cell layers that were grown on HA did not form HA crystals on their surfaces, while the cells that were cultivated for 5 days on biosilica well formed HA crystals that were often fusing to clusters (Wang et al. 2013b). This observation which had been supported by alizarin red S staining assays underscores that biosilica displays an inductive effect on SaOS-2 (Wiens et al., 2010a,b). Moreover, biosilica as a nontoxic polymer, causes a significant shift of the OPG:RANKL ratio toward the steadystate level of OPG in vitro (Wiens et al., 2010b). The data gathered revealed that in SaOS-2 cells, biosilica causes an increased gene expression of OPG, while the steady-state level of RANKL remains unchanged (Figure 2.7(b)). This increased OPG expression was verified on protein level, by application of an ELISA system and also by direct staining of the SaOS-2 with Alizarin Red S. In continuation we found that biosilica also displays a stimulatory effect on BMP-2 expression (Wiens et al., 2010b). This finding had an important impact on a more comprehensive understanding of the differentiation/proliferation property of biosilica. Again, by application of quantitative polymerase chain reaction analysis, it became overt that biosilica induces the expression of this important cytokine by twofold (Wiens et al., 2010a). In continuation, we analyzed if biosilica displays an effect on the proliferation propensity of the cells in vitro.This question is important in order to assess the osteogenic potential of this inorganic polymer. Incorporation studies revealed (Wiens et al., 2010a) that the ratio between [3H]dT incorporation into DNA and biomineral (HA) formation was significantly higher in cells that grew on silicatein/ biosilica-modified substrates. This latter finding is the first strong indication that the osteogenic potential of biosilica is more than merely an osteoconductive one. Following the above mentioned definition (­Albrektsson and Johansson, 2001), biosilica can be considered to have an osteoinductive capability since it combines, firstly, an increased expression of genes (proteins) required for the differentiation and secondly, an increased potential for proliferation of the osteogenic cells. In further ongoing attempts in our group, we also studied the effect of biosilica on COLI expression ­(Müller et al. 2013a).The PCR data disclosed that in SaOS-2 cells the gene expression of this collagen subtype I of the collagen family is upregulated almost in parallel with BMP-2. These two aspects of the biosilica effect on SaOS-2 cells in vitro suggest, or perhaps even demonstrate, that this polymer, biosilica, has not only a potential to accelerate

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Figure 2.7  Proposed mitogenic effect caused by silica/biosilica on bone cells. (a) After taken up by the organism, silica is proposed to accumulate extracellularly around the collagen/mucopolysaccharides fibrous network as Ca-salt. After hydrolysis to monomeric units, silica acts on the cellular level to the mitogen-activated protein kinase (MAPK)/­ extracellular-signal-regulated kinases (ERK) pathway, and TGF–TGF receptor signaling route and finally via the WNT/hedgehog (WNT/SHH) differentiation/polarity way to enhance the expression of the anabolic bone-forming mediators osteopontin (OPN), OCAL, and b-ALP. (b) Proposed effects of biosilica on osteoblasts, osteoclasts, and their progenitor cells. (Bio)silica causes an increased expression of OPG and BMP-2 in osteoblasts. In turn OPG counteracts RANKL, and by that inhibits the differentiation of osteoclasts. Finally, it is sketched that after exposure to biosilica the osteoblasts release a factor, the osteoblasts-derived inhibitory factor, that strongly inhibits the proliferation of osteoclasts. BMP-2, bone morphogenetic proteins-2; b-ALP, bone-specific alkaline phosphatase; OPG, osteoprotegerin; RANKL, receptor activator of NF-κB ligand; OPG, osteoprotegerin; RANK, receptor activator of nuclear factor κB; OCAL, osteocalcin; SHH, sonic hedgehog homolog.

cell proliferation and differentiation of osteogenic cells in vitro but also the capacity to provide and synthesize the matrix, the scaffold for the osteogenic cells (COLI), on which the differentiating cells can find their functional destination within a growing bone structure. In addition, functional evidence has been presented revealing that the osteoblast-related SaOS-2 cells release a factor that kills osteoclasts, e.g., RAW 264.7 cells (Schröder et al., 2012). The mode of action of biosilica on mammalian cells is partially understood.The above mentioned data indicating that biosilica comprises a positive osteogenic activity in vitro (Wiens et al., 2010a, 2010b) has been comprehensively confirmed recently (Han et al., 2013). These authors further substantiated that biosilica/ortho-silicate causes beneficial and promising effects for future potential applications in therapy of bone diseases, e.g., osteoporosis. These confirmatory data have been elaborated with bone marrow stromal cells and showed that silica causes an upregulation of the bone-related

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anabolic genes osteopontin (OPN), OCAL, and b-ALP, as well as an increased expression of genes involved in the WNT and sonic hedgehog homolog (SHH)-signaling pathways (Han et al., 2013); Figure 2.7(a). Based on these data together with the findings and conclusion published by Jugdaohsingh et al. (2002) a first interpretation of the mode of action of biosilica on cellular and subcellular level can be given (Figure 2.7(a)). Silica can be taken up by the oral route, then it can be transported to the bulky extracellular space of the connective tissue and very likely also into cells. In the extracellular space, ortho-silicate can be supposed to bind to Ca2+ ions that are abundant in this compartment which is rich in mucopolysaccharides (Schmidt et al., 1969). Since Ca-silicates have a lower solubility compared to the respective Na+ or K+ salts (Manzano et al., 2012), it can be postulated that Ca-silicates accumulate in the collagen-mucopolysaccharide fibrous scaffold around the bone tissue, where they can be mobilized again by changing the solubility product or by changing the pH milieu. In the extracellular space, polymeric silica can be hydrolytically degraded to monomeric silica perhaps by cathepsins or by lowering the concentration of the monomeric reactant, with respect to polysilica. The presumed uptake of ortho-silicate via an ion channel might be facilitated through the consumption of ATP (Schröder et al., 2004). Intracellularly, silica is supposed to activate the MAPK–ERK pathway in osteoblast-like cells (Shie et al., 2011); via this central signaling pathway, the silica-caused signal is translocated to the nucleus. Flanked is this pathway by activation of the transforming growth factor/tumor growth factor (TGF) pathway in response to which an enhanced cellular differentiation and collagen production occurs by binding of TGF-1 to its cognate receptors (Li et al., 2013a). Finally, data led to suggest that silica activates the WNT and SHH-signaling pathways with Axin2 and β-catenin as the central molecules (Han et al., 2013).The proposed mechanism favors the idea that silica acts on the different levels or pathways, primarily at the transcriptional level, of those genes that encode for proteins, stimulating bone anabolism, e.g., OCAL, OPN, and b-ALP.

5.  ENZYMES CONTROLLING THE SYNTHESIS OF MORPHOGENETICALLY ACTIVE INORGANIC POLYMERS: A PARADIGM SHIFT IN BIOINORGANIC CHEMISTRY Multicellular organisms larger than a few millimeters need to develop a hard skeleton in order to stabilize, harden, or stiffen their body form and shape (Lowenstam and Weiner, 1989; Knoll et al., 2012). The construction of

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such hard skeletons is based on biomineral deposits that have a species-specific morphology. The reactions resulting in the formation of these bioinorganic structural elements are controlled by the same principles as reactions that are involved in the synthesis of bioorganic macromolecules. The principles are: (1) to run at ambient, physiological conditions, where (2) the reactants are present at nonsaturating concentrations, and (3) the reaction velocities are accelerated by enzymes (Figure 2.8). Furthermore, the biomineralization processes are (4) integrated in the anabolic and catabolic pathways of the organisms, and, finally, (5) they are genetically controlled.The genetically or molecular biologically controlled biomineralization processes (Wang et al., 2014d) are initiated by bioseeds that act as nucleation sites; the subsequent growth of the biomineral is equally and effectively controlled by genetically based organic template(s). The paradigm shift in bioinorganic chemistry is illustrated in Figure 2.8. Mineralic, abiotically formed calcite crystals are found very abundantly in nature (Figure 2.8(a) and (b)). Their synthesis as well as degradation/­ dissolution occurs in nature under nonphysiological physical and chemical conditions (Rodriguez-Navarro et al., 2009), while both the synthesis and the dissolution of those calcite crystals in Metazoa proceeds enzymatically. It has only very recently been recognized that the formation of both silica and Cacarbonate (CaCO3) deposits in animals is driven enzymatically. The basic studies have been performed with the evolutionary oldest metazoans, the sponges (phylum: Porifera) (Müller et al., 2004). The skeletons, spicules, of these first animals on Earth are formed either of inorganic polymeric biosilica, as in the sponges grouped to the classes of Hexactinellida and Demospongiae, or of mineralic biocalcite, CaCO3, as in the sponge class of Calcarea. Biosilica is synthesized by the enzyme silicatein (Morse, 1999; Müller, 2003), while biocalcite deposits are formed by the rate-­determining enzyme CA; (Wang et al., 2014e).The calcareous skeletal elements of the sponges, the spicules, are illustrated here with the example from the calcareous sponge species Sycon raphanus (Figure 2.8(c) and (d)). These spicules are surrounded within the animals by an organic sheet (Müller et al., 2012, 2014a)

5.1  Carbonic Anhydrase The deposition of CaCO3 is an exergonic and thus a thermodynamically favored, possible process (Li et al., 2013b).The initial reaction, the formation of bicarbonate (HCO3−) from CO2 and H2O, is the rate-limiting step in CaCO3 formation (Reddy, 1981).The activation energy barrier, required to initiate the HCO3− formation can be overcome by the enzyme CA (Wang et al., 2014c, 2014d).

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Figure 2.8  Biochemical versus chemical mineralization. Center: Almost all exclusively biochemically driven biomineralization reactions (green filled segment in the reaction velocity versus substrate concentration curve) run between the substrate (S) concentration range close to zero up to slightly above the K m value (Michaelis–Menten constant). In contrast, chemical reactions (red segment) are usually performed at substrate saturation conditions. While biologically occurring biomineralization processes are physiologically controllable, and their products formed are usually soft and by that

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It should be stressed on this point that the modulation of the energy barriers posed by the activation energy of a given reaction enables an organism fine-tunely to control under which physiological conditions a thermodynamically possible reaction can be initiated or prevented (Meldrum and Cölfen, 2008). Almost exclusively, alterations of the heights of the activation energy barriers are adjusted by enzymes or by the surface architecture of membranes separating two phases. There are the CAes, a family of enzymes that accelerate the HCO3− formation from CO2 and H2O during biocalcite formation, especially in sponge spicules (Henry, 1996; Müller et al., 2013d). These highly prevalent enzymes that allow bicarbonate to be formed in an organism (Lindskog, 1997) are characterized by an extremely high-turnover number, allowing the catalysis of the reversible hydration of carbon dioxide (CO2) to HCO3−. If this reaction occurs in the presence of Ca ions (Sanyal and Maren, 1981) the mostly water-insoluble (under physiological conditions) CaCO3 is formed. Four of the seven metazoan CA isoenzymes are cytosolic, CA-I, -II, -III, and -VII; among them the CA-II is a widely studied one (Sly and Hu, 1995). Our group could present solid evidence that the CAes are involved in bone formation (Müller et al., 2013b). The mammalian CA-II, usually a cytosolic enzyme, is targeted in intact cell systems under certain physiological conditions to the cell membrane (Alvarez et al., 2001; Chang et al., 2012).The complete cDNA of the CA in the phylogenetically oldest animals that have a skeleton based on CaCO3, the calcareous sponges with S. raphanus as an example, has been cloned, expressed, and functionally tested (Müller et al., 2012, 2014a). The complete 1476 nts cDNA encodes, within its open reading frame from nt68–70 moldable, biodegradable, and in turn biocompatible, purely chemically produced materials are more permanent, usually hard, not degradable and are often bioincompatible. (a and b) Examples for chemically formed CaCO3 minerals. (a) An optical calcite (calcite) produces two refracted rays that are transmitted by this material, instead of one ray, producing a double image. One of the refracted rays penetrates independent of the orientation of the crystal, while the other rotates its position with the crystal. The crystal is laid out of the title page of the fundamental monograph on mineralization by Bütschli (1898). The two arrow point to the double refraction area with the authors’ name. (b) Calcite crystals are important rock-forming minerals in sedimentary environments. (c and d) Enzymatically formed CaCO3 biominerals, the calcareous spicules from the sponge Sycon raphanus (c). The spicules are surrounded by an organic sheet (os). In (d), the surfaces of the spicules are stained positive for carbonic anhydrase (ca) by using specific fluorescently labeled antibodies as a tool. (For interpretation of the references to color in this figure legend, the reader is referred to the online version of this book.)

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to nt1001–1003, the 312 aa putative CA, with a Mr of 33,251 and a pI of 5.81. The closest human related CA to the Sycon enzyme are the human CA-X (gene bank accession number Q9NS85) and the human CA- XI (O75493) that have been grouped to the “acatalytic” CAes (Lovejoy et al., 1998;Wang et al., 2014e). The Zn-binding sites that are involved in the catalytic reaction (hydration of CO2) are present in the CA-alpha (vertebrate-like) group stretch of the S. raphanus protein. Within the CA molecule the Zn ions are bound to the enzyme through the three His residues in the catalytic center of the enzyme (Tripp et al., 2001). The cDNA of the Sycon CA was prepared in a recombinant way and used to raise antibodies. Immunostructural studies revealed that the Sycon CA is localized on the surface of the mature, developed spicules, the ca. 300 μm-long diactines and the ca. 300 μm-large triactines and tetractines (Figure 2.8(c) and (d)). It is assumed that the membranous, organic sheaths described to cover the spicules (­Figure 2.8(c)) are composed predominantly of this enzyme (Müller et al., 2012); Figure 2.8(d). Subsequently, the recombinant enzyme was used to determine the in vitro calcium carbonate formation by applying the in vitro diffusion assay (­Müller et al., 2013d). Even though the present-day oceans are supersaturated with respect to calcium carbonate, only very rarely spontaneous abiotic CaCO3 precipitations are seen (Tyrrell, 2008). Like in other metazoans, e.g., mollusks and echinoderms, sponges take up CO2 from the aqueous environment as HCO3− via specific membrane transporters that are characterized by a Michaelis–Menten constant which has a value of around 50 mM (Chow et al., 1976), or it is produced metabolically. At this concentration, CaCO3 precipitates slowly to an extent of about 50% during an incubation period of only 20 h in an ammonium carbonate diffusion/“dessicator assay” at pH 7–8 (Li et al., 2013c). This purely chemically driven reaction is too slow to account for the rates of CaCO3 deposition occurring in vivo, e.g., during spicule formation in Sycon sp. (Ilan et al., 1996). In these animals the 100 μm long and about 4 μm-thick calcite spicules grow with a very fast growth rate of 65 μm/h. Consequently it must be postulated that an acceleration of the velocity of the exergonic reaction at ambient, physiological conditions has to meet the physiological prerequisites. The most obvious mechanism to achieve this demand is either to lower the activation energy by an enzyme, or to allow the CaCO3 process to proceed on a functionalized organic surface. In our previous studies we tested the first possibility (Müller et al., 2014a, 2013d;Wang et al., 2014f).We used as the substrate for the enzymatic reactions in the “dessicator” assay 50 mM CaCl2 over which

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CO2 vapor, generated from NH4HCO3 solution, was passed. The reaction was performed at a pH of 7.5. The progress of the mineralization process was recorded, based on the decrease of free Ca2+ concentration. In the absence of the enzyme CA the process started after an initial lag phase of 5 h. However, after addition of the recombinant CA (35 Wilbur–Anderson (W–A) units/500 μl) a significantly increase in the reaction velocity and an accelerated mineralization was determined; after an incubation period of 50 min already 25% of the CaCl2 was precipitated, under the conditions used (Figure 2.9(a) and (b)). An extent of 36% of precipitated CaCO3 was measured after 4 h; after an incubation period of 10 h, the extent of precipitated CaCO3 amounted to 80% (not shown in Figure 2.9). Two morphologically different CaCO3 deposits are formed in the in vitro assay in the presence of the CA (35 W-A CA units per assay): first prisms with an average size of 80–120 μm and second round-shaped deposits having similar dimensions (Müller et al., 2014a; Müller et al., 2013d; Wang et al., 2014f ). It is remarkable and likewise indicative for a biologically controlled biomineralization process that the enzymatic/biogenic CA-driven CaCO3 crystal formation can be frozen at the vaterite state, even though the overall thermodynamically possible “end-point” transition formation to calcite could be reached. In the biomimetic approach we could show that the CA-driven CaCO3 deposition process can be blocked at the vaterite crystal stage in the presence of silintaphin-2, a sponge-specific protein that is rich in aspartic acid (Asp, D) and glutamic acid (Glu, E) (Müller et al., 2014b). The hardness, elastic modulus, and creep of the two forms of the CaCO3 deposits formed in the CA-driven reaction, the round-shaped vaterite deposits and the rhombohedral calcite were determined by nanoindentation. The load–displacement curves obtained for the two CaCO3 forms revealed the following values: for the rhombohedral calcite 1.98 ± 0.31 GPa and for the round-shaped vaterite deposits only 1.38 ± 0.39 GPa. Parallel, a distinct decrease of the elastic modulus was measured for the vaterite deposits with 39.13 ± 8.04 GPa, in comparison to the rhombohedral calcite prisms with 72.83 ± 11.68 GPa. This significant difference in the mechanical properties between the two morphologies is also reflected from the creep behavior. While the creep characteristics for the rhombohedral calcitic prisms was found to be 5.44 ± 1.15 (per maximal depth (%)), the corresponding value for the round-shaped vaterite deposits is 9.95 ± 1.60 (Müller et al., 2014b). As expected the enzyme (CA)-mediated deposition of CaCO3 is temperature dependent (Müller et al., 2013d). While at 10 °C the reaction velocity of CaCO3 deposition is almost identical in the enzyme-containing

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Figure 2.9  Abiotic (absence of CA) as well as biotic (CA-driven) CaCO3 deposition in the ammonium carbonate diffusion assay. (a) The reactions were performed in the absence or (b) presence of 35 W-A units CA per 500 μl crystallization assay. The reaction assays either remained free of an additional compound (filled square) or were supplemented with 10 μM quinolinic acid (QA, open triangle). At the indicated incubation time points, samples were taken and the free Ca2+ concentration was determined. The decrease in the concentration of free Ca2+ indicates an increase in deposited CaCO3. Samples of six parallel determinations were quantitated; means ± SD are given. *p < 0.05. (c) Postulated interaction of QA (in red) with the active center of the human CA. Three His residues within the active center are interacting with the central Zn2+ ion. The three-hydrogen bond network, formed by Wat-150, Wat-129, and Wat-130, is indicated; modified after Ilies et al. (2002). It is outlined that QA interacts with its N-heteroatom to Wat-130 and one of its carboxylic acid groups to Zn2+. CA, carbonic anhydrase. (For interpretation of the references to color in this figure legend, the reader is referred to the online version of this book.)

and enzyme-lacking assays, at higher, physiologically more relevant incubation temperatures (e.g., 22 °C; Müller et al., 2014a), the reaction velocity of the CA-driven CaCO3 formation is significantly higher than that in the absence of CA. Likewise indicative is the pH dependence of the CaCO3 deposition reaction in the presence/absence of CA. While in the absence of CA the precipitation of CaCO3 increases only slightly from pH 6.0 to pH 8.1, the CA-driven reaction velocity increases markedly (by over

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5-fold) from pH 6.0 to pH 8.0. Importantly, the increased rate in the reaction velocities seen in the CA-containing assays can be inhibited almost completely by the CA-specific inhibitor acetazolamide at 3 μM (Müller et al., 2013d). In order to underscore the enzymatic contribution to the calcium carbonate deposition in vitro, the characteristic kinetic constant, the Michaelis–Menten constant, for CA, was determined (Müller et al., 2013d). Like any other enzymatic reaction also the CA-mediated CaCO3 deposition reaction follows a substrate saturation kinetics. Under the assay conditions used (50 mM CaCl2, pH 7.5, 25 °C), a linear increase of the reaction velocity is seen between 0 and 20 mM CaCl2. At higher concentrations a saturation level is approached. It is well established that the CAs function both as hydratases, in the formation of CaCO3, and also as esterases (­Kirley and Day, 1985). The Michaelis–Menten constants (Km) for both reactions are almost identical at around 5 mM for the hydratase (using CO2 as substrate) and for the esterase (with the substrate 4-nitrophenylacetate).

5.2  Alkaline Phosphatase In 1923, the importance of phosphate biochemistry for bone formation was strengthened by Robinson (Robinson, 1923) with his finding that hexosephosphoric esters are modulating ossification. Consequently the enzyme ALP has been implicated in phosphate metabolism in bone, due to the high levels and regional accumulation of these ions in areas of highest ossification (Lorch, 1949). The ALPs are a family of enzymes and exist in humans as four isoenzymes. Three of them are expressed in a tissue-specific pattern in the intestine, the placenta, and germ cells (Price, 1993), while the fourth ALP is abundant in bone and liver (Henthorn, 1996). Since then, the latter isoform has been used as a biochemical marker in serum and urine to assess bone formation and bone resorption (DeLaurier et al., 2002), especially in patients with metabolic diseases, such as osteoporosis and hyperparathyroidism (Christgau et al., 2000). In the following, this enzyme, the b-ALP was used as a target to successfully modulate phosphatase activity and, in turn, ossification, e.g., by bisphosphonates (Vaisman et al., 2005). The physiological role of bone ALP is partly understood. It has been proposed that b-ALP generates inorganic phosphate (Pi), which is needed for HA crystallization in the bone matrix (Millan, 2006), while in an alternative view evidence has been presented that this enzyme hydrolyzes the mineralization inhibitor inorganic pyrophosphate (PPi) (Rezende et al., 1998) in order to facilitate mineral precipitation and growth (Hessle et al., 2002). More recent studies favor the assumption that the major function of

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b-ALP is to maintain the proper homeostasis of this mineralization ­inhibitor to ensure balanced bone mineralization (Millan, 2006). Korenchevsky and Carr proposed in 1925 (Korenchevsky and Carr, 1925) that β-GP is the physiological phosphate donor for HA deposition in vivo and in vitro. β-GP is a synthesized intracellularly and then released from the cells to the extracellular space (Graff et al., 2003). In turn, β-GP has traditionally been used as one component in the activation cocktail added to HA-mineralizing cells in vitro (Cheng et al., 1994). Besides being a proposed substrate for HA formation, β-GP has been demonstrated to be a suitable substrate for ALP (Ahlers, 1975), especially in tissues characterized by physiological and pathological mineralization (Fratzl-Zelman et al., 1998). However, already for sometime it had been concluded that, especially under in vitro conditions, β-GP is completely converted into Pi (Bellows et al., 1992). Even more, the enzyme ALP also hydrolyzes inorganic polyPs (Lorenz and Schröder, 2001), naturally occurring inorganic polymers that are found in pro- and eukaryotes (see above). In continuation our group showed that the two naturally occurring inorganic polymers, biosilica and polyP, are potent inducers of biomineralization of HA-forming bone cells in vitro (Müller et al., 2011). In our studies we could show that Pi can substitute for β-GP during the biomineralization process on the SaOS-2 cells. The polymer polyP is synthesized and degraded in the mitochondria of mammals; there polyP is stored as a polyP•Ca2+ complex in order to neutralize the negative charges of the polymer (Müller et al., 2014c). After the export to the cytoplasm and to the extracellular space the polyP•Ca2+ complex undergoes hydrolytic cleavage to Pi and Ca2+ where the two components serve as the inorganic components for HA formation (Figures 2.6 and 2.10). Data from our group (Müller et al., 2011) strongly suggest that polyP acts inductive on the expression level of the TNAP in SaOS-2 cells. Even more, the enzyme activity is significantly upregulated in the presence of polyP, indicating that polyP acts as an inducer of the TNAP. In turn, this enzyme has been correlated with the anabolic pathway of bone formation (Millán, 2006). TNAP plays a key role in bone mineralization, and both initiates and promotes the formation of HA crystals in the matrix vesicles of osteoblasts as well as, pathologically, in hypertrophic chondrocytes (Anderson et al., 2004). Subsequently, the crystals are transported into the ECM and deposited onto the cell membrane (Anderson et al., 2004). Those matrix vesicles might represent extracellular particles that serve as bioseeds for the subsequent biomineralization processes. TNAP also degrades PPi under formation of

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Figure 2.10  Schematic outline of the Ca-phosphate deposition onto SaOS-2 cells. Intracellular polyP is stored and metabolized by catabolic and anabolic enzymes in the mitochondrion and there complexed to Ca2+. If required, polyP•Ca2+-complex is released into the cytoplasm and then into the extracellular space where the complex is hydrolyzed to inorganic phosphate (Pi) and Ca2+, the two main components of HA formation. Subsequently Pi and Ca2+ become deposited onto the CaCO3 bioseeds formed in the vicinity of osteocalcin (OCAL). Orthophosphate, released from polyphosphate (polyP), downregulates the activity of the carbonic anhydrase. ALP, alkaline phosphatase.

free Pi, which is the main component together with Ca2+ for HA formation. We also provided evidence that polyP increases the intracellular Ca2+ level in SaOS-2 cells (Müller et al., 2011). An increased level of cytosolic Ca2+ has been measured after exposure of the cells to polyP•Ca2+-complex, an effect that was not seen if polyP was added as a Na+ salt. These data show that polyP•Ca2+-complex is a potent inducer of ALP in SaOS-2 cells and additionally contributes with these activities and functions to the HA crystallite formation. Furthermore, a significant amount of phosphate consumed during HA formation is supplied from polyP synthesized in SaOS-2 cells adjacent to/or identical with the osteoblasts (Leyhausen

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et al., 1998; Hacchou et al., 2007; Müller et al., 2011). Interestingly enough, the hydrolytically released orthophosphate, Pi, from polyP inhibits the CA in a feedback manner and by that prevents a further enzymatic synthesis of HCO3−/CaCO3 and in turn inhibits the synthesis of the bioseeds that are required for HA formation (Müller et al., 2013b); Figure 2.10.

6.  BIOCALCITE AS BIOSEED DURING MAMMALIAN HA FORMATION The contribution of CaCO3 during HA/bone formation has been described. Bone is a biomineral, composed of a mineral phase (70% w/w), an organic matrix (≈25% w/w), and ≈10% of water (v/w). The inorganic phase is predominantly HA (Ca5(PO4)3(OH)). Earlier it could be determined that carbonate is present in the apatite mineral phase of bone and enamel at concentrations exceeded only by calcium and phosphate (Rey et al., 1989). Even more, it has been proposed that bone formation in osteoblasts starts from collagen fibrils around which poorly crystalline carbonated apatite crystals, carbonate apatite aggregates are deposited (Boonrungsiman et al., 2012). Recently we have shown that newly formed mineralic deposits on SaOS-2 cells consists, as expected, of calcium and phosphorus, but also of high levels of carbon (Müller et al., 2013b). The conclusion that CaCO3 crystallites act as bioseeds for HA deposition has been underscored by inhibition studies using inhibitors of the CA. If the cells were exposed to those inhibitors they failed to produce HA in larger amounts.The substrate for the enzymatic CaCO3 deposition, HCO3−, can be provided to the extracellular space in close vicinity to the Cl−/HCO− exchanger (AE), or intracellularly in the plasma membrane in proximity to the Na+:HCO3− cotransporter (NBC) (see: Müller et al., 2013b); Figure 2.11. Since the CA-II inhibitor acetazolamide has been found to inhibit biomineralization in SaOS-2 cells the participation of this enzyme has been implicated in the Ca-deposit formation onto the cell membranes. The CA-II is ubiquitously present in the cytoplasm of almost all metazoan cells and, focusing on mammalian bone cells, is probably involved in bone resorption (Margolis et al., 2008). There, CA-II causes proton production, resulting in a drastic acidification of the resorption lacuna/bone regions. Furthermore, studies suggest that this enzyme is also involved in bone formation (Sly and Hu, 1995). These surprising Janus-faced catabolic/anabolic metabolic reactions, controlled by CAs might be explained on account of the reversibility of the CA-catalyzed

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Figure 2.11  Proposed bioseed formation for the mineralization process on bone-forming cells (SaOS-2). The transcripts of Ca2+-binding protein OCAL are synthesized in SaOS-2 cells. The primary translation product is processed to the mature OCAL. During this process the protein undergoes carboxylation via the γ-glutamyl carboxylase (GGCX). We propose that OCAL delivers Ca2+ (first component of biomineralization onto bone cells) as a substrate for the CA, which in turn is used for the formation of HCO3− (second component) and then for CaCO3, which functions as bioseed. The CA is activated by sponge extract(s), like QA and HCO3−. OCAL, osteocalcin; CA, carbonic anhydrase; QA, quinolinic acid.

reaction. The CA acts both as a CaCO3 anabolic enzyme, facilitating and accelerating bicarbonate formation, a precursor molecule required for calcium carbonate mineral formation, and also as a catabolic enzyme that promotes calcium carbonate dissolution, as shown, e.g., in corals (Müller et al., 1984). Published experimental data revealed that during the initial phase of the controlled bone-synthesizing process poorly crystalline carbonated apatite is deposited, which contains several percents (4–6 wt%) of carbonate in the apatite crystals (Termine et al., 1973; Biltz and Pellegrino, 1977; ­Matsuura et al., 2009). Even more, recent studies suggest that the increased carbonate content in apatite crystals has an anabolic effect on

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bone formation (Matsuura et al., 2009). Our EDX mapping studies (Müller et al., 2013b) indicate that the crystallites initially formed onto SaOS-2 cells are not only rich in the elements calcium and phosphorous but also in carbon. In turn, we concluded that carbonate and phosphate deposits are co- or sequentially synthesized onto SaOS-2 cells, during the initial phase of mineral formation. Furthermore, the CA-II has been proven to be under certain physiological conditions (pH regulation) localized at the plasma membranes of human pancreatic cells (Alvarez et al., 2001), where the enzyme is involved both in pH regulation and in the secretion of bicarbonate through the Cl−/HCO− exchanger and/or an additional HCO3− channel (Alvarez et al., 2001). Based on these data we conclude that the CaP/HA deposition reactions in bone tissue are preceded by CaCO3 precipitation, a process that is driven by an increased CA activity (Figure 2.11). The origin of the second major component of mineral formation onto bone cells, of Ca2+, is not sufficiently known. We postulated that the bulk of Ca2+ is delivered by the Ca2+-binding OCAL to the cell membrane at the sites where the bioseeds are formed (Figure 2.11). Again a patched/localized accumulation of both proteins (OCAL) at the membranes would support the CaCO3 bioseed view during mineralization of bone cells.The process of OCAL formation can be subdivided into two steps; first, the expression of the OCAL transcripts and, second, the carboxylation of OCAL by the γ-glutamyl carboxylase (GGCX). This process is vitamin K-dependent through influencing the enzyme GGCX (Wood and Suttie, 1988; Lian and Gundberg, 1988;Thrailkill et al., 2012).

7.  CA ACTIVATORS AS POTENTIAL NOVEL DRUGS TO STIMULATE BONE MINERAL FORMATION Only very little experimental evidence is available on the potential therapeutic effect of CA activators on bone anabolism (Supuran and ­Scozzafava, 2000). Until now only a few CA activators have been identified, but none of them have been tested for its potential in the treatment of bone disorders (Supuran 2008). Very recently we found that the CA-driven CaCO3 deposition process onto SaOS-2 cells can be stimulated by CA activators both in vitro and also in intact cell system (Müller et al., 2014a). As activator(s) we have chosen extracts from the sponge Suberites domuncula and one component, isolated from these extracts, quinolinic acid (QA). In the in vitro CA-driven CaCO3 deposition assay we could show that both the S. domuncula extract and QA stimulate mineral formation (Figure 2.9(a) and (b)). In the experiments shown here the enzyme-driven CaCO3 deposition

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results, under the conditions used (Müller et al., 2014a), a CaCO3 formation of 28% during a 240 min incubation period, while in the presence of 10 μM QA 48% of CaCO3 are formed under otherwise the same conditions. This effect is amplified by polyP (Müller et al., 2014a). Based on these data we propose that the two “regulator enzymes,” the ALP and the CA, are under feedback control, offering new targets for pharmacological intervention of bone diseases. While the ALP is activated by polyP, the CA is inhibited by PO43−. In a preliminary approach a molecular model for the interaction of QA with the active center of the CA is proposed (Müller et al., 2014a); Figure 2.9(c). QA comprises, with its N-heteroatom in the pyridine backbone as well as the dicarboxylic acid side chains, two potential interacting groups with the Zn2+containing CA. The zinc prosthetic group in the CA is coordinated in three positions by histidine side chains, by His94, His96, and His119 (­Lindskog, 1997). The fourth coordination position at Zn is occupied by one water molecule and causes there a polarization of the hydrogen–oxygen bond, resulting in an increased negativity of the oxygen and a weakening of the bond to the Zn atom. A fourth histidine, His64, is placed close to the substrate of water and accepts a proton from the water molecule through general acid–base catalysis under formation of a hydroxide bound to the zinc (Figure 2.9(c)).These studies have been performed with human-crystallized CA. We propose that the QA with its dicarboxylic acid side chains interacts with the enzyme-bound Zn ion, like in the bipyridyl Zn chelators (Ilies et al., 2002). The mineralization studies with alizarin red also revealed that QA causes a significant upregulation of the biomineralization process by SaOS-2 cells in vitro.This effect is especially pronounced during the initial phase of mineralization, during the first 3 days. At this time point the increase caused by QA is 210% compared to the 100% of the controls (Müller et al., 2014a). Studies using SEM coupled with EDX analysis confirmed our previous findings (Müller et al., 2013b) that carbon is strongly accumulated in the mineral nodules on the surface of the SaOS-2 cells, whether incubated with S. domuncula extracts or with QA.

8.  ALP ACTIVATORS—POTENTIAL NOVEL COMPOUNDS TO STIMULATE BONE MINERAL FORMATION? The ALPs can be activated and/or inhibited by Mg2+ and Zn2+ (­Brunel and Cathala, 1973), an effect that is due to a conformational change of the active site of the enzyme (Hung and Chang, 2001). While Zn2+ is

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bound at the catalytic site of ALP, Mg2+ acts as an allosteric activator of ALP, at a site on the enzyme, which is distinct from the Zn2+ site. Besides of divalent metal ions, also amino alcohols (e.g., 2-amino-2-methyl-1-propanol, diethanolamine), and TRIS (2-amino-2-hydroxymethyl-propane-1,3-diol) buffer (Lamb et al., 1998) were found to activate ALP. In a recent attempt we determined the effect of oligoP (oligophos­ phate)•Ca2+-complex, having a chain length of the oligoP of six phosphate units linked in a circle. For these studies the Na-hexametaphosphate was reacted with a stoichiometric concentration of CaCl2. A crude extract has been prepared from SaOS-2 cells and then reacted in an in vitro assay using p-nitrophenyl phosphate as a substrate. After an incubation period of 15 min (37 °C) the released product, p-nitrophenol, was determined spectrophotometrically at 405 nm (Komoda et al., 1982). An extract from 105cells had been used in the 200 μl assay. As summarized in Figure 2.12, oligoP•Ca2+complex stimulated the ALP from SaOS-2 cells in a concentration-dependent manner; between the range of 5 and 30 μM a significant increase in the activity is measured. In comparison, the polyP•Ca2+-complex is less active but still significantly affects the enzyme in a stimulating manner. At present we explain this stimulatory effect on a direct action of the oligo/polymer on the ALP, or on an intermediate product, released by the phosphatase on the enzyme protein, resulting in a conformational change.

Figure 2.12  Effect of oligoP (oligophosphate)•Ca2+-complex between the concentration range of 5–50 μM (black bars) and of 30 μM polyP•Ca2+-complex (gray bar) on the activity of the ALP, prepared form SaOS-2 cells; the control without any oligo/polymer phosphate is indicated (open bar). The enzyme reactions were run for 15 min (37 °C) in an assay containing the substrate p-nitrophenyl phosphate. The differences between the control group (no additional phosphate compound) were evaluated using unpaired t-test.*p < 0.05; the means of 10 replicates had been performed.

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9.  APPLICATIONS FOR BIOPRINTING ORGANS It is self-evident that the repair of critical size bone defects with autogenous bone grafts is the gold standard (Epple, 2007). Because of several limitations which cannot be solved by using allogenous bone grafts, such as immunogenicity and risk of infection, synthetic bone scaffolds can provide several advantages if they meet the following conditions. Those scaffolds need to show similar physiochemical characteristics as the natural bone, and should have the ability to attract the bone-forming cells (the progenitor cells or functionally active differentiated cells), two challenging tasks, limiting the routine application of synthetic materials in the treatment of bone defects. The artificial scaffold to be applied for bone tissue engineering should be accepted by the cells as a suitable 3D platform for their growth, differentiation, and mineralization (HA deposition). And, further on a suitable scaffold should possess the inorganic/organic 3D structure of bone with an appropriate porosity (Holzwarth and Ma, 2011), allowing the ingrowth of cells and an efficient transport of cytokines, growth factors, and nutrients. Recently, much hope and some progress have been achieved in rapidprototyping/3D printing techniques. This 3D printing technology involves a computer-controlled layer-by-layer technology (Figure 2.13(a) and (b)). The advantage of this method is that the implants display a customized 3D geometry of the bone defect, based on medical imaging data. However, preferentially the implant should allow an optimal integration and also the potential to be replaced by the adjacent tissue with functional host cells/tissue. Based on published data alginate/chitin, also together with silica (Madhumathi et al., 2009; Gimeno-Fabra et al., 2011), provides a suitable matrix for the encapsulation of mammalian cells. In our studies we have demonstrated that SaOS-2 cells embedded into Na-alginate that has been supplemented with silica (Schloßmacher et al., 2013; Müller et al., 2013a;Wang et al., 2013a) retain the proliferating activity. Even more, if embedded in biosilica together with an alginate/ gelatin matrix the cells display morphogenetic activity, especially if they have been overlayed with polyP•Ca2+-complex. Based on these findings we have successfully started to print 3D structures in order to apply this technology for bioprinting as well as for construction of bioartificial tissues or organs. In a first step we have encapsulated separately bone-forming (SaOS-2) and bonedegrading (RAW 264.7) cells and subsequently printed this biomimetic synthetic scaffold to tissue units. The mechanical properties, including surface roughness and hardness, of the hydrogel containing cells were determined

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A

B

C

d-A d-B d-C Figure 2.13  Fabrication of tissue units by 3D bioprinting and subsequent sheathing/coating of the soft bioprinted tissue units with morphogenetically active electrospun mats. (a) Computer-aided rapid prototyping bioprinting. (a–A and -B). The sketch outlines the computer-guided extrusion of Na-alginate hydrogel (supplemented with biosilica or bicarbonate) through a capillary of a needle (n) in a meander-like pattern. This matrix contained SaOS-2 cells (Sa-2). Those printed blocks are incubated in a surrounding matrix/ medium into which RAW 264.7 cells (RAW) have been embedded as well. (a-C) A fabricated bioprinted 3D tissue unit, with a diameter of 11.5 mm, showing the arrangement of the printed alginate/gelatin cylinders (cy). (b) Preparation of an overlay (ol) around the bioprinted cylinders (cy), containing the cells (SaOS-2 or RAW 264.7 cells). (c) Outline of the formation of packed 3D bioprinted, bioartificial tissue units (tu). The typical electrospinning setup is shown, comprising a syringe pump (sp), attached to a syringe needle (sn). During the electrospinning process, the polymer (p) solution is induced to form, at

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and found to be too soft for an immediate application as implant. Therefore, we developed electrospinned mats, fabricated from poly(ε-caprolactone) and provided with suitable mechanical properties, that allow a sheathing/coating of the more soft bioprinted tissue units (Müller et al., 2014d). Such “composite/hybrid” implants should have the potential to be implanted and to act in the cavities as regeneration-inducing and replaceable implants (Figure 2.13).

10.  CONCLUDING REMARKS The advantages to utilize synthetic bone scaffolds include: elimination of disease transmission risk, fewer surgical procedures, reduced risk of infection or immunogenicity, and especially abundant availability of synthetic scaffold materials. The basic challenge to develop a suitable synthetic scaffold is to mimic the complex physiological environment in which bone cells grow and differentiate. In a physiological framework, the bone cells find a suitable scaffold that allows them to ingrow into a scaffold with the matching porosity where they can differentiate and communicate by signaling with the neighboring cells. Moreover, these cavities must allow the substrates for the osteoblasts to enter and to be available for the osteoid deposition, allow vascularization, and finally bone ingrowth. Focusing on bone tissue engineering strategies, e.g., such as cell transplantation, acellular scaffolds, stem cell therapy, again the physiological regulatory network of cytokines and growth factors must be provided to the MSCs after the removal from the donor ex vivo. In the future 3D cell printed implants enriched with morphogenetically active polymers, e.g., biosilica of bio-polyP, should be developed which additionally contain activators for the biomineralization process based on the CaCO3 and CaP biominerals. Since it is now established that the key enzymes that are involved in the formation of the bioseeds, formed of CaCO3, and the subsequent CaP/HA formation initiated by the ALP, the screening for activators of those enzymes will be continued in a boosted manner.

high voltage, thin microfibers (nf). The fibers are collected at an appropriate collector (c). Onto the motor (mo)-driven collector an electrospun polymer mesh/mat (m) is formed. (c) A trimmed/cut mat (m) is layered onto a well (w) of a microtiter plate, filled with the applied medium (m) (d-A). The mat sample is submersed into the well (d-B) and the cells are bioprinted (bp) onto the mat. The cells, encapsulated into the matrix, are printed in a cylinder–meander pattern to a bioartificial tissue unit (tu). Those units become enveloped into the electrospun mat (m) (d-C). Insert: An electrospun mat.

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ACKNOWLEDGMENTS W.E.G. M. is a holder of an ERC Advanced Investigator Grant (No. 268476 BIOSILICA).The work in our laboratory was supported by grants from the Deutsche Forschungsgemeinschaft (Schr 277/10-3) and the European Commission (“Bio-Scaffolds - Customized Rapid Prototyping of Bioactive Scaffolds”, No. 604036; Industry-Academia Partnerships and Pathways “CoreShell”, No. 286059;“MarBioTec*EU-CN*”, No. 268476; and “BlueGenics”, No. 311848).

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