EDITED BY: Giovanna Orsini, Victor E. Arana-Chavez and Thimios Mitsiadis PUBLISHED IN: Frontiers in Physiology

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ISSN 1664-8714 ISBN 978-2-88919-984-6 DOI 10.3389/978-2-88919-984-6

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# **DENTAL AND PERIODONTAL TISSUES FORMATION AND REGENERATION: CURRENT APPROACHES AND FUTURE CHALLENGES**

Topic Editors:

**Giovanna Orsini,** Polytechnic University of Marche, Italy **Victor E. Arana-Chavez,** University of São Paulo, Brazil **Thimios Mitsiadis,** University of Zurich, Switzerland

Figure composition inspired by a scanned human molar (microCT image, Institute of Oral Biology, University of Zurich), multiplied in various scales and pseudocolors. Production and copyright by Thimios Mitsiadis.

Sequential and reciprocal interactions between oral epithelial and cranial neural crest-derived mesenchymal cells give rise to the teeth and periodontium. Teeth are vital organs containing a rich number of blood vessels and nerve fibers within the dental pulp and periodontium. Teeth are composed by unique and specific collagenous (dentin, fibrillar cementum) and non-collagenous (enamel) highly mineralized extracellular matrices. Alveolar bone is another collagenous hard tissue that supports tooth stability and function through its close interaction with the periodontal ligament. Dental hard tissues are often damaged after infection or traumatic injuries that lead to the partial or complete destruction of the functional dental and supportive tissues.

Well-established protocols are routinely used in dental clinics for the restoration or replacement of the damaged tooth and alveolar bone areas. Recent progress in the fields of cell biology, tissue engineering, and nanotechnology offers promising opportunities to repair damaged or missing dental tissues. Indeed, pulp and periodontal tissue regeneration is progressing rapidly with the application of stem cells, biodegradable scaffolds, and growth factors. Furthermore, methods that enable partial dental hard tissue repair and regeneration are being evaluated with variable degrees of success. However, these cell-based therapies are still incipient and many issues need to be addressed before any clinical application. The understanding of tooth and periodontal tissues formation would be beneficial for improving regenerative attempts in dental clinics.

In the present e-book we have covered the various aspects dealing with dental and periodontal tissues physiology and regeneration in 6 chapters:

**Citation:** Orsini, G., Arana-Chavez, V. E., Mitsiadis, T., eds. (2016). Dental and Periodontal Tissues Formation and Regeneration: Current Approaches and Future Challenges. Lausanne: Frontiers Media. doi: 10.3389/978-2-88919-984-6

# Table of Contents

*07 Editorial: A New Era in Dentistry: Stem Cell-Based Approaches for Tooth and Periodontal Tissue Regeneration*

Thimios A. Mitsiadis and Giovanna Orsini

#### **Chapter one: General principles on the use of stem cells for regenerating craniofacial and dental tissues**

*10 Dental pulp stem cells as a multifaceted tool for bioengineering and the regeneration of craniomaxillofacial tissues*

Maitane Aurrekoetxea, Patricia Garcia-Gallastegui, Igor Irastorza, Jon Luzuriaga, Verónica Uribe-Etxebarria, Fernando Unda and Gaskon Ibarretxe

*20 Manufacturing of dental pulp cell-based products from human third molars: current strategies and future investigations*

Maxime Ducret, Hugo Fabre, Olivier Degoul, Gianluigi Atzeni, Colin McGuckin, Nico Forraz, Brigitte Alliot-Licht, Frédéric Mallein-Gerin, Emeline Perrier-Groult and Jean-Christophe Farges

*28 Use of Rat Mature Adipocyte-Derived Dedifferentiated Fat Cells as a Cell Source for Periodontal Tissue Regeneration*

Daisuke Akita, Koichiro Kano, Yoko Saito-Tamura, Takayuki Mashimo, Momoko Sato-Shionome, Niina Tsurumachi, Katsuyuki Yamanaka, Tadashi Kaneko, Taku Toriumi, Yoshinori Arai, Naoki Tsukimura, Taro Matsumoto, Tomohiko Ishigami, Keitaro Isokawa and Masaki Honda

*40 Changing Paradigms in Cranio-Facial Regeneration: Current and New Strategies for the Activation of Endogenous Stem Cells*

Luigi Mele, Pietro Paolo Vitiello, Virginia Tirino, Francesca Paino, Alfredo De Rosa, Davide Liccardo, Gianpaolo Papaccio and Vincenzo Desiderio

## **Chapter two: The roles of nerves, vessels and stem cell niches in tissue regeneration**

#### *53 The Nervous System Orchestrates and Integrates Craniofacial Development: A Review*

Igor Adameyko and Kaj Fried

Anna Woloszyk, Davide Liccardo and Thimios A. Mitsiadis

# *82 The Perivascular Niche and Self-Renewal of Stem Cells*

Min Oh and Jacques E. Nör

# **Chapter three: Dental pulp regeneration and mechanisms of various odontoblast functions**

and Dimitrios Tziafas

*106 Human odontoblast-like cells produce nitric oxide with antibacterial activity upon TLR2 activation*

Jean-Christophe Farges, Aurélie Bellanger, Maxime Ducret, Elisabeth Aubert-Foucher, Béatrice Richard, Brigitte Alliot-Licht, Françoise Bleicher and Florence Carrouel

Masaki Sato, Tadashi Furuya, Maki Kimura, Yuki Kojima, Masakazu Tazaki, Toru Sato and Yoshiyuki Shibukawa

# **Chapter four: Dental root and periodontal physiology, pathology and regeneration**

Min Liu, David A. Reed, Giancarlo M. Cecchini, Xuanyu Lu, Karan Ganjawalla, Carol S. Gonzales, Richard Monahan, Xianghong Luan and Thomas G. H. Diekwisch

*159 Orthodontic Forces Induce the Cytoprotective Enzyme Heme Oxygenase-1 in Rats*

Christiaan M. Suttorp, Rui Xie, Ditte M. S. Lundvig, Anne Marie Kuijpers-Jagtman, Jasper Tom Uijttenboogaart, René Van Rheden, Jaap C. Maltha and Frank A. D. T. G. Wagener

# **Chapter five: Physiology and regeneration of the bone using various scaffolds and stem cell populations**

*184 Bone resorption: an actor of dental and periodontal development?* Andrea Gama, Benjamin Navet, Jorge William Vargas, Beatriz Castaneda and Frédéric Lézot

*191 Biomimetically enhanced demineralized bone matrix for bone regenerative applications*

Sriram Ravindran, Chun-Chieh Huang, Praveen Gajendrareddy and Raghuvaran Narayanan

Anna Spina, Roberta Montella, Davide Liccardo, Alfredo De Rosa, Luigi Laino, Thimios A. Mitsiadis and Marcella La Noce

## **Chapter six: Physiology, pathology and regeneration of enamel using dental epithelial stem cells**

*234 Distribution of the amelogenin protein in developing, injured and carious human teeth*

Thimios A. Mitsiadis, Anna Filatova, Gianpaolo Papaccio, Michel Goldberg, Imad About and Petros Papagerakis

*242* **In vivo** *administration of dental epithelial stem cells at the apical end of the mouse incisor*

Giovanna Orsini, Lucia Jimenez-Rojo, Despoina Natsiou, Angelo Putignano and Thimios A. Mitsiadis

# Editorial: A New Era in Dentistry: Stem Cell-Based Approaches for Tooth and Periodontal Tissue Regeneration

Thimios A. Mitsiadis 1 \* and Giovanna Orsini 2

1 Orofacial Development and Regeneration, Centre for Dental Medicine, Institute of Oral Biology, University of Zurich, Zurich, Switzerland, 2 Department of Clinical Sciences and Stomatology, Polytechnic University of Marche, Ancona, Italy

Keywords: tooth, dentin, enamel, stem cells, scaffolds, vasculature, craniofacial, regenerative dentistry

#### **The Editorial on the Research Topic**

#### **Dental and Periodontal Tissues Formation and Regeneration: Current Approaches and Future Challenges**

#### Edited by:

Gianpaolo Papaccio, Seconda Università degli Studi di Napoli, Italy

#### Reviewed by:

Vincenzo Desiderio, Seconda Università degli Studi di Napoli, Italy

> \*Correspondence: Thimios A. Mitsiadis<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Craniofacial Biology, a section of the journal Frontiers in Physiology

Received: 15 July 2016 Accepted: 04 August 2016 Published: 19 August 2016

#### Citation:

Mitsiadis TA and Orsini G (2016) Editorial: A New Era in Dentistry: Stem Cell-Based Approaches for Tooth and Periodontal Tissue Regeneration. Front. Physiol. 7:357. doi: 10.3389/fphys.2016.00357 The continuously increasing progress in stem cell biology has expanded our horizons into the principles of health and disease. This new acquired knowledge and the recent advances in the field of biotechnology have formed the substrate for translation into novel therapeutic solutions (Mitsiadis et al., 2015). Indeed, traditional dental therapies have successfully incorporated a good number of new materials that are more compatible and friendly to dental and periodontal tissues. Although these treatments are suitable and satisfactory for long-lasting dental restorations, the idea of stem cell-based tissue regeneration is tempting and represents the decisive objective of introducing new therapeutic approaches in the clinics. Teeth develop through continuous and reciprocal interactions between cranial neural crest-derived mesenchymal stem cells and oralderived epithelial stem cells during early embryogenesis (Mitsiadis and Graf, 2009; Smith et al., 2009; La Noce et al., 2014). These highly mineralized organs are composed by collagenous (such as the mesenchyme-derived dentin and cementum structures) and non-collagenous (such as the epithelium-derived enamel structure) extracellular matrices. The alveolar bone is another collagenous hard tissue, which through its close interaction with the periodontal ligament supports tooth stability and function. Tooth vitality is ensured by blood vessels and nerve fibers within the dental pulp and periodontium (Carmeliet and Jain, 2011; Pagella et al., 2015). Regenerative approaches include the formation of new enamel, dentin, pulp, periodontal ligament, and alveolar bone after tooth damage due to genetic pathology, traumatic injuries, caries, and periodontal lesions (Mitsiadis and Harada, 2015). Therefore, the recent progress made in the fields of stem cell biology, tissue engineering, and nanotechnology offers promising opportunities to repair damaged or missing dental tissues (Mitsiadis et al., 2012). Several considerable efforts have been made recently in the dental field and new concepts concerning dental treatment strategies have arisen with the hope to translate basic research findings into the clinics. Repairing teeth by autologous cell grafting is highly desired, but some ethical, economic, and legislative issues should be first resolved.

In this research topic, prominent researchers within the dental field provided important information, gave responses, and generated stimulating hypotheses concerning the development and regeneration of dental and periodontal tissues. A balanced mosaic of original and review articles point out different facets of the interplay between stem cell biology and dental tissue formation, homeostasis, and regeneration.

The different sources of stem cells, their differentiation potential, and the current state of stem cell-based strategies for dental or craniofacial tissue regeneration are discussed in several articles. The cranial neural crest-derived dental stem cells or stem cells from other tissues can be used with success for dental, periodontal, and craniofacial repair purposes (Akita et al.; Aurrekoetxea et al.; Ducret et al.). The appropriate selection of stem cells in conjunction with the creation of an adequate and respective microenvironment is essential for a successful clinical outcome. Therefore, the identification and characterization of stem cell niches within dental tissues, and the analysis of niche-derived signals during tooth homoeostasis and repair is of prime importance (Oh and Nor). The exploitation of various adult stem cell populations localized in the different tooth components is challenging. Furthermore, success depends on well-controlled and coordinated cytodifferentiation and mineralization processes during tissue repair and regeneration (Eapen and George).

Small-scale engineered biomaterials and tailor made empty scaffolds that will attract specific endogenous stem cell populations can be designed for the regeneration the periodontium (Green et al.) as well as of various tissues within the craniofacial complex (Mele et al.). Innervation of craniofacial tissues, including teeth, is essential for ensuring their morphology, physiological function, and regeneration (Fried and Adameyeko). Equally, blood supply of craniofacial structures and teeth are important for their function, vitality, and longevity, and therefore it is important to develop methods for analyzing angiogenesis in regenerating tissues (Woloszyk et al.). One of these methods allowed us to evaluate and compare the ability of human dental pulp stem cells and gingival fibroblasts in attracting vessels into silk fibroin scaffolds (Woloszyk et al.). The tissue-engineering concept for dental pulp regeneration implicates stem cells, scaffolds, and signaling molecules (Passos et al.; Yang et al.). Pulp-derived stem cells have the potential to differentiate into odontoblasts and other cell types and thus could be used for clinical applications. Odontoblasts produce nitric oxide with antibacterial activity, which is an important issue for successful tissue regeneration purposes (Farges et al.). During tooth homeostasis and pathology, communication between odontoblasts is activated via specific channels (Sato et al.). Hydrogels may be used as stem cell-seeded matrices for pulp and periodontal tissue repair purposes in the clinics (Teti et al.). Polymerized resin-based materials could affect cellular physiology and induce cell apoptosis (Teti et al.).

#### REFERENCES

Several pathologies exist that lead to root malformations and consequently affect tooth eruption or/and physiology (Luder). Tooth eruption is a complicated event that can differ from species to species (Liu et al.). Resorption of the alveolar bone that surrounds the tooth root could be an actor of periodontal development and remodeling (Gama et al.). Orthodontic forces affect alveolar bone remodeling and periodontal ligament regeneration through defined molecular mechanisms (Suttorp et al.). The use of biomimetically enhanced demineralized bone matrices could represent an interesting approach for bone regenerative applications (Ravindran et al.). However, the outcome of tissue engineering strategies for bone formation depends on the origin and the culture conditions of the applied stem cells (Mattioli-Belmonte et al.; Spina et al.).

Amelogenin, one of the most important organic elements of enamel, is re-expressed during dental pathology and repair, indicating that it can act as a signaling molecule for dental tissue repair events (Mitsiadis et al.). Furthermore, injection of stem cells at dental pathological sites may enhance the possibility of de novo enamel formation. For testing this hypothesis a mouse model was used for the injection of epithelial stem cells derived from the continuously growing incisors (Orsini et al.).

Besides many open scientific questions, strategic concerns also need to be addressed. Controversies in scientific data, opinions, and formulated hypotheses set the starting point for additional research that will benefit to the dental field. Repair and regeneration of dental tissues using stem cell-based approaches combined with new biotechnological tools is an exciting and highly relevant area of research. Further evaluation of existing tools is needed in order to optimize future therapeutic strategies in dentistry. The time for applying novel regenerative therapies in dental clinics has come, thus designing a new era, full of excitement and expectations.

#### AUTHOR CONTRIBUTIONS

The authors contributed to the writing, reading, and editing of the present editorial.

#### ACKNOWLEDGMENTS

This work was supported by institutional funds from University of Zurich (TM) and by funds from the Polytechnic University of Marche (GO). The authors contributed to the writing, reading, and editing of the present editorial.

craniomaxillofacial development and tissue repair. Eur. Cell. Mater. 28, 348–357.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

The reviewer VD and handling Editor declared their shared affiliation, and the handling Editor states that the process nevertheless met the standards of a fair and objective review.

Copyright © 2016 Mitsiadis and Orsini. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Dental pulp stem cells as a multifaceted tool for bioengineering and the regeneration of craniomaxillofacial tissues

Maitane Aurrekoetxea † , Patricia Garcia-Gallastegui † , Igor Irastorza, Jon Luzuriaga, Verónica Uribe-Etxebarria, Fernando Unda\* and Gaskon Ibarretxe\*

*Department of Cell Biology and Histology, Faculty of Medicine and Dentistry, University of the Basque Country, Leioa, Spain*

#### Edited by:

*Thimios Mitsiadis, University of Zurich, Switzerland*

#### Reviewed by:

*Gianpaolo Papaccio, Second University of Naples, Italy Zhi Chen, Wuhan University, China*

#### \*Correspondence:

*Fernando Unda<EMAIL_ADDRESS>Gaskon Ibarretxe<EMAIL_ADDRESS>
*These authors have contributed equally to this work.*

*†*

#### Specialty section:

*This article was submitted to Craniofacial Biology, a section of the journal Frontiers in Physiology*

Received: *31 July 2015* Accepted: *01 October 2015* Published: *16 October 2015*

#### Citation:

*Aurrekoetxea M, Garcia-Gallastegui P, Irastorza I, Luzuriaga J, Uribe-Etxebarria V, Unda F and Ibarretxe G (2015) Dental pulp stem cells as a multifaceted tool for bioengineering and the regeneration of craniomaxillofacial tissues. Front. Physiol. 6:289. doi: 10.3389/fphys.2015.00289* Dental pulp stem cells, or DPSC, are neural crest-derived cells with an outstanding capacity to differentiate along multiple cell lineages of interest for cell therapy. In particular, highly efficient osteo/dentinogenic differentiation of DPSC can be achieved using simple *in vitro* protocols, making these cells a very attractive and promising tool for the future treatment of dental and periodontal diseases. Among craniomaxillofacial organs, the tooth and salivary gland are two such cases in which complete regeneration by tissue engineering using DPSC appears to be possible, as research over the last decade has made substantial progress in experimental models of partial or total regeneration of both organs, by cell recombination technology. Moreover, DPSC seem to be a particularly good choice for the regeneration of nerve tissues, including injured or transected cranial nerves. In this context, the oral cavity appears to be an excellent testing ground for new regenerative therapies using DPSC. However, many issues and challenges need yet to be addressed before these cells can be employed in clinical therapy. In this review, we point out some important aspects on the biology of DPSC with regard to their use for the reconstruction of different craniomaxillofacial tissues and organs, with special emphasis on cranial bones, nerves, teeth, and salivary glands. We suggest new ideas and strategies to fully exploit the capacities of DPSC for bioengineering of the aforementioned tissues.

Keywords: DPSC, differentiation, tooth, bone, salivary gland, nerve, cell therapy

# INTRODUCTION: DPSC AND TISSUE ENGINEERING OF THE ORAL CAVITY

The oral cavity is a complex multi-organic structure. Because oral tissues and organs are functionally connected at many levels, irreversible damage to any of them is likely to eventually affect the others, causing extensive malfunction. Tooth decay, periodontal disease, alveolar bone resorption, orthodontic problems, orofacial neuropathic pain, and impaired salivary gland function are conditions that seriously affect oral health of a large part of the world population. Owing to their functional connectivity, once damage is diagnosed to one organ of the oral cavity it is important to intervene rapidly and efficiently, to repair or replace the injured or lost tissues, to avoid severe degradation of oral health.

Synthetic replacement materials and prostheses (fillings, bridges, implants, etc.) have traditionally been the treatment of choice to treat dental decay. However, all functions of the original biological tooth are not fully restored by this kind of replacement therapies. Other organs of the oral cavity (e.g., nerves, salivary glands) are simply not amenable to mechanical substitution approaches. Thus, tissue engineering represents a new collection of treatment options for the complete biological regeneration of craniomaxillofacial tissues and organs. The development of this field requires three essential components: (i) stem cells, (ii) biomaterial scaffolds, and (iii) stimulating factors or inductive signals. Tissue engineering is now fully considered as an alternative to the conventional treatments for dental injury and disease, offering substantial advantages over traditional dental restoration techniques (Nör, 2006; Wang et al., 2012).

Stem cells are the cornerstone of regenerative cell therapy. An enormous variety of multipotent stem cells have been isolated and studied from different human tissues, such as the bone marrow (Ding and Morrison, 2013), adipose tissue (Kapur et al., 2015), skin (Blanpain and Fuchs, 2009), and the umbilical cord (Yan et al., 2013; Kalaszczynska and Ferdin, 2015). Among them, mesenchymal stem cells (MSC) are the most promising for clinical purposes (Rastegar et al., 2010; Ménard and Tarte, 2013). In the oral cavity, adult tooth tissues also contain different active populations of stem cells with mesenchymal phenotype (Huang et al., 2009). Unlike other types of MSC, dental stem cells originate from the neural crest (Janebodin et al., 2011) and are lineage-related with peripheral nerve glial progenitor cells (Kaukua et al., 2014), which places them in a privileged position to mediate regeneration of both connective and nerve tissues (Ibarretxe et al., 2012; Martens et al., 2013). Several types of human dental stem cells have been identified. Among them; DFPC, Dental Follicle progenitor cells; PDLSC, Periodontal ligament stem cell; SCAP, Stem cells from apical papilla; SHED, Stem cells from primary exfoliated deciduous teeth; DPSC, Dental pulp stem cells, which are without a doubt the most used for research purposes (**Figure 1**). Although it still remains a considerable challenge to obtain all the different types of neuronal and glial cells from DPSC, their differentiation to specialized connective tissue cells has proven to be relatively simple. Efficient protocols for obtaining adipocytes, chondroblasts and osteo/odontoblasts are firmly established in the literature (Gronthos et al., 2002; Huang et al., 2009; Kawashima, 2012).

In this review, we summarize the current understanding of DPSC and their capacity to regenerate injured orofacial tissues, with special focus on the teeth and associated periodontal tissues, peripheral cranial nerves, and the salivary glands. We present the different strategies used in experimental restorative models with potential applications in dentistry and oral medicine, together with the different scaffold biomaterials and stimulating factor combinations employed to elicit an optimal cellular response required to regenerate damaged tissues. We critically comment on some crucial aspects to be considered for bioengineering and the regeneration of each of these tissues, based on our own research experience.

# DPSC FOR BONE REGENERATION

cell lineages.

Regeneration of maxillary and mandibular bone is fundamental in the field of implantology (Lee et al., 2014). Osteogenic differentiation of DPSC is easily induced in vitro by adding ascorbic acid (Asc), dexamethasone (Dexa), and βglycerophosphate (β-gly) to the culture medium, along with fetal bovine serum (Laino et al., 2005; Langenbach and Handschel, 2013). Asc is an essential enzyme cofactor to generate procollagen (Vater et al., 2011), which is necessary for the correct synthesis of Collagen type I, the main organic component of the bone matrix. Dexa is required for osteoblast differentiation, working as Runx2 inductor by activating WNT/β-catenin signaling in MSC (Hamidouche et al., 2008). Runx2 transcription factor expression is fundamental in driving cellular commitment to osteogenic lineages. Finally, β-gly is a donor of inorganic phosphate, which is essential for creating hydroxyapatite mineral, and as a signaling molecule to induce DPSC osteodifferentiation (Foster et al., 2006; Fatherazi et al., 2009; Tada et al., 2011).

The application of this differentiation cocktail is highly effective in inducing DPSC to generate a mineralized bonelike extracellular matrix in vitro. When DPSC are exposed to this differentiation cocktail for 7–21 days, they start to strongly express different osteoblast differentiation markers, such as Osterix, Runx2, osteocalcin, and bone sialoprotein. Alkaline phosphatase (ALP), and Alizarin red staining are positive under these conditions, and are used to demonstrate effective differentiation of DPSC to functional osteoblasts, and the generation of a calcified hydroxyapatite (HA)-containing extracellular matrix (Mangano et al., 2010; Yu et al., 2010; Li et al., 2011). DPSC-derived osteoblasts are phenotypically and functionally similar to normal primary osteoblasts, although some differences remain in terms of their gene expression profiles (Carinci et al., 2008). Naïve undifferentiated DPSC are negative for the adult osteo/odontoblast differentiation marker Osterix, but nevertheless exhibit detectable levels of Runx2, suggesting they are primed or pre-committed to differentiate to osteo/odontoblasts (Yu et al., 2010). When the osteogenic differentiation cocktail is added to DPSC, Runx2, and Osterix expression increase and these cells generate extracellular calcified bone-like matrix nodules, that stain distinctively and positively with Alizarin red (Laino et al., 2005; **Figure 2**).

In the last few years the use of DPSC for in vivo bone regeneration by tissue engineering has been extensively studied by relying on these protocols and using different experimental models and types of scaffolds with different results (Morad et al., 2013). Woven bone chips obtained by human DPSC (hDPSC) osteoinduction are able to induce the generation of adult bone tissue with an integral blood supply, when they are heterotopically transplanted in immune-compromised

FIGURE 2 | Osteogenic differentiation of DPSC. In the presence of a three-ingredient cocktail containing dexamethasone, β-glycerophosphate and ascorbate added to the culture medium for 7 days, DPSC differentiate to osteoblast-like cells that generate a mineralized bone tissue matrix that stains positively for Alizarin red.

(IC) animals (d'Aquino et al., 2007). However, the use of scaffolding materials is often necessary to optimize the 3D structure of the formed bone tissue and enhance the osteoblastic differentiation of hDPSC. To this end, different materials have been successfully employed as a vehicle of hDPSC, including 3D gelatin scaffolds (Li et al., 2011), self-assembling biodegradable peptide nanofiber hydrogels (Chan et al., 2011), CaP/PLGA (calcium phosphate/polylactic-co-glycolic acid) scaffolds (Graziano et al., 2008a,b), particulate hydroxyapatite/tricalcium phosphate (HA/TCP) ceramic scaffolds (Zhang et al., 2008), and natural scaffolds like biocoral (Mangano et al., 2011) and chitosan/collagen (Yang et al., 2011). Finally, alginate and PuraMatrix™ hydrogels also provide a three dimensional (3D) biodegradable carrier of stem cells for bone tissue engineering purposes, with the additional large advantage that these formulations are injectable, forming 3D hydrogels upon contact with the physiological environment and thus completely filling the tissue lesion, with minimally invasive dental and orthopedic surgeries. Dental stem cells encapsulated in either alginate or PuraMatrix™ exhibited high capacities for osteo-differentiation both in vitro and in vivo (Misawa et al., 2006; Cavalcanti et al., 2013; Moshaverinia et al., 2013).

Inductive signals also play a critical role to stimulate development of new bone tissue. The development of a welldefined, safe, and controlled technique to obtain and locally deliver growth factors derived from autologous plasma has provided a powerful tool to enhance bone tissue regeneration, which has been successfully implemented in the clinical practice (Anitua et al., 2008, 2012). Formulations such as Platelet-rich plasma (PRP), Platelet-rich fibrin (PRF), and Plasma rich in growth factors (PRGF) provide a biologically enriched culture medium for the stimulation and functional differentiation of primary cells with latent regenerative potential. Several studies have shown that PRGF in particular provides additional benefits for bone regeneration (Paknejad et al., 2012; Rivera et al., 2013). It has been reported that PRP and PRGF induce proliferation and enhance osteogenic differentiation in SHEDs, DPSCs, and PDLSCs (Lee et al., 2011). Other studies have used recombinant human Bone morphogenetic protein 2 (rhBMP-2) to enhance osteogenic differentiation of DPSC in animal models of alveolar bone reconstruction (Liu et al., 2011). The use of rhBMP-2 is approved and widely extended to perform maxillary sinus floor augmentation in dental practice (Nazirkar et al., 2014; Freitas et al., 2015).

Despite a large body of preclinical evidence, very few clinical trials have still been performed to evaluate new advanced therapeutic medicinal products (ATMP) for bone regeneration based on DPSC (La Noce et al., 2014). A notable exception is one clinical trial performed with seven patients to repair mandibular bone defects by transplant of DPSC in a collagen scaffold (d'Aquino et al., 2009). After 3 years of follow-up, the clinical outcomes were positive (Giuliani et al., 2013). However, several obstacles remain, including a changing regulatory framework, a shortage of technology for automation of cell production by Good manufacturing practice (GMP) standards, a non-alignment between academic and industrial research, and a need for longterm product follow-up (La Noce et al., 2014; Leijten et al., 2015). These limitations have hampered the emergence of DPSC as a tool to enhance bone regeneration at clinical level.

#### DPSC FOR DENTAL PULP REGENERATION

Caries and root fracture are the cause of approximately fifty percent of all tooth extractions. Tooth injuries such as pulp exposure, deep caries and irreversible pulpitis are currently treated by endodontics and dental refilling/reconstruction. This procedure usually entails the amputation or entire removal of the dental pulp and replacement of the tissue with an inert material. Root fracture and tooth loss are closely related to this loss of pulp vitality because innervation and vasculature affect pulp homeostasis and root dentin regeneration. The regeneration of pulp injuries has become a goal for functional tooth restoration in dentistry (Nakashima and Akamine, 2005; Nakashima et al., 2009).

The first research reports on DPSC showed that these cells present optimal characteristics to attain functional regeneration of the dental-pulp chamber. Xenogenic transplant of HA/TCP scaffolds with hDPSC into the dermis of IC mice resulted in the formation of pulp-like tissue and polarized odontoblasts that produced tubular dentin, showing that dentinogenic differentiation was one of the prime or default programs established in the DPSC phenotype (Gronthos et al., 2000, 2002). Recently, more refined studies combining SHED with injectable scaffolds (Puramatrix™ and rhCollagen type I) have demonstrated that dental stem cells are not only kept alive when injected into human premolar root canals in vivo, but are also able to fully reconstruct vascularized pulp tissue throughout the root canal, and to differentiate to DSPP and DMP-expressing odontoblasts that generate new dentin (Rosa et al., 2013). In a similar experimental design, other authors employed combinations of DPSC and umbilical vein endothelial cells embedded in Puramatrix™ scaffold, to improve vascularization and angiogenesis of de novo formed dental pulp-like tissue (Dissanayaka et al., 2015).

Other studies have focused on the regenerative potential of different cytokines and growth factors, with the goal to improve cellular chemotaxis and cell homing into the emptied dental pulp space in vivo. Different combinations of cytokines and growth factors which included basic Fibroblast growth factor (bFGF), Platelet derived growth factor (PDGF), or Vascular endothelial growth factor (VEGF) in the presence of basal levels of Nerve growth factor (NGF) and BMP-7 were effective in regenerating a revascularized and recellularized dental pulp-like tissue integrated into endodontically-treated root canal dentinal walls in vivo (Kim et al., 2010). In addition, it has been shown that bFGF and Stromal-derived factor (SDF1) also exert a potent chemotactic recruitment on DPSC, while BMP-7 induces their differentiation (Suzuki et al., 2011). In a recent study, SDF1 loaded onto silk-fibroin scaffolds promoted the complete regeneration and revascularization of the dental pulp of mature dog teeth that had been subjected to endodontic treatment in situ (Yang et al., 2015), suggesting a promising future for the functional reconstruction of dental pulp tissue in next generation dentistry.

# DPSC FOR COMPLETE DENTAL REGENERATION

In 2009, a ground-breaking report was published by the group of T. Tsuji on the generation of complete and fully functional teeth by combining isolated mouse dental epithelial and MSC (Ikeda et al., 2009). More recently, in 2011, the same research group developed a complete tooth unit consisting of a mature tooth, periodontal ligament and alveolar bone, using a similar recombination technique (Oshima et al., 2011). These tooth units were transplanted into toothless mouse jaw regions in vivo, and erupted correctly and in occlusion. They also presented an adequate dental structure, an adequate hardness of enamel and dentin, a proper function of the periodontal ligament, a positive alveolar bone remodeling response to orthodontic forces, and a positive response to noxious stimuli such as mechanical stress and pain.

Although precise replication of these results has not yet been reported on the literature, the aforementioned work represents a substantial advance in stem cell combination technology, with the potential to develop completely new bioengineered replacement organs for next generation regenerative therapy. One of the main limitations to date is probably the lack of consistent sources of epithelial cells that can be combined with dental mesenchymal cells (DPSC or others) to generate a complete bioengineered tooth germ. The team of T. Tsuji employed embryonic mouse tooth germ epithelia as precursors of the enamel organ, a structure that fully disappears in adult teeth. In the search for dental-competent sources of epithelial stem cells, some authors have reported success using adult gingival epithelial cells, which upon recombination with mesenchymal cells generate mature teeth and form enamel and dentin (Angelova Volponi et al., 2013). Other authors have employed iPS-derived cells to generate mature bioengineered teeth by similar recombination methods (Wen et al., 2012; Cai et al., 2013; Otsu et al., 2014). The substantial progress made in the field of dental bioengineering warrants further research on this line for the next years.

# DPSC FOR REGENERATION OF NERVE TISSUE AND DAMAGED CRANIAL NERVES

DSPC originate from the neural crest (Janebodin et al., 2011) and share a common origin with peripheral nerve glial progenitor cells (Kaukua et al., 2014). These features make DPSC a very interesting choice for regeneration of the peripheral nervous system, including nerves of the oral cavity. Some reports claim that DPSC can even differentiate to functionally active adult neurons (Arthur et al., 2008; Kiraly et al., 2009; Gervois et al., 2015). These conclusions are based on the acquisition of neuronspecific gene and protein markers by DPSC, and also on their capacity to generate neuronal-like voltage-dependent sodium and calcium currents, and action potential-like membrane voltage oscillations. However, it should be noted that to date there has been no definite proof of differentiation of DPSC to genuine neurons that can exhibit repetitive firing of action potentials upon electrical stimulation, and establish synapses showing functional plasticity as identified by transmission electron microscopy (TEM).

However, it is undeniable that DPSC present some striking similarities with neural stem cells. When DPSC are grown in culture conditions lacking fetal serum, they reorganize morphologically and switch from a uniform cell monolayer to a more quiescent state characterized by the appearance of prominent spheroid structures that resemble CNS-derived neurospheres which stain positively for the neural stem cell marker Nestin (Ibarretxe et al., 2012; Bonnamain et al., 2013; Xiao and Tsutsui, 2013; Gervois et al., 2015; **Figure 3**). Migratory cells are occasionally observed to leave the DPSC spheroids and virtually all these cells express the neuron-lineage marker β3-tubulin. Moreover, their morphology is in some cases surprisingly similar to migratory neuroblasts, with a long and thin leading process terminated by a prominent growth cone, and a trailing cell body displaying a few short cytoplasmic processes (Marín et al., 2006). However, a note of caution should be added: another cell type of the dental pulp, the odontoblast, is also characterized by the expression of Nestin, the generation of voltage-dependent Na+ currents, and a similar morphology (Fujita et al., 2006; Ichikawa et al., 2012). The DPSC population grown in these conditions is fairly heterogeneous, both morphologically and physiologically, with a range of different morphologies from fibroblast-like to neuroblast-like cells, and also a variable rate of response to stimulation with neurotransmitter receptor agonists.

Regardless of the debate on the true capacity of DPSC to differentiate to neurons and other neural cell lineages, there is little doubt that these cells could be a very interesting choice to repair injured nerve tissues as a result of their active secretion of neurotrophic and immune-modulatory factors (Nosrat et al., 2004; Pierdomenico et al., 2005). The high expression of certain neural markers and neurotransmitter receptors by DPSC also suggests that these cells may actively respond to signals of the neural environment and effectively integrate into injured nerve tissues, promoting the reestablishment of functional nerve connectivity (Arthur et al., 2009; Kadar et al., 2009; Kiraly et al., 2009). For instance, some recent reports show that transplanted DPSC are able to differentiate in vivo to myelinating cells in the spinal cord, replacing lost cells and promoting regeneration of transected axons in acute models of spinal cord injury (Sakai et al., 2012).

In the case of the peripheral nerve system, treatment of nerve injuries represents a clinical challenge because of the difficulties of regenerating transected nerves. Though numerous surgical successes have been reported with a short nerve gap, there is still no satisfactory technique for long nerve defects, which often require a complex clinical reconstruction. Autologous nerve grafting has been considered the gold standard for repairing peripheral nerve gaps caused by traffic accidents or tumor resectioning (Kumar and Hassan, 2002; Hayashi and Maruyama, 2005; Bae et al., 2006). However, this technique has inevitable disadvantages, such as a limited supply of available nerve grafts and permanent loss of the sacrificed donor nerve function. Brainderived neural progenitor cells also promote regeneration of

transected nerves (Murakami et al., 2003). However, the use of cells from other neural tissues involves potentially serious clinical complications along with ethical considerations.

Taking all these arguments into account, there is an active search for new sources of cells to be used in craniofacial nerve bridging and regeneration. Considerable advances have been made in this field using DPSC for the treatment of facial nerve injuries. Particularly, patients with facial paralysis, especially younger ones, may experience tremendous psychosocial distress about their condition (Chan and Byrne, 2011). Recent studies have used DPSC transplanted in PLGA tube scaffolds to achieve a complete functional regeneration of the facial nerve in rats to recovery levels comparable to those obtained with peripheral nerve autografts (Sasaki et al., 2011, 2014). Interestingly, recent research also indicates that hDPSC can be differentiated to Schwann-like cells that efficiently myelinate DRG neuron axons in vitro, a finding confirmed by ultrastructural TEM analysis (Martens et al., 2014). Considering the important role that Schwann cells play in axonal protection and regeneration of peripheral nerves (Walsh and Midha, 2009), and the difficulty of their harvesting and maintenance, the generation of DPSCderived autologous Schwann cells may represent a milestone in the design of new treatments for conditions of peripheral nerve injury, including facial paralysis.

Finally, another important property of DPSC is their active secretion of neurotrophic factors (Nosrat et al., 2004; Bray et al., 2014), which may be exploited to treat neuropathic pain states associated with peripheral nerve injury. In the case of orofacial pain, some of the most distressing and painful conditions that can be experienced by a human being are neuralgias affecting the trigeminal nerve, or CN V. These are characterized by intense stabbing pain and spasms, usually associated with a mechanical injury, compression, demyelination and inflammation of trigeminal sensory fibers (Love and Coakham, 2001; Sabalys et al., 2013). It is known that local application of Glial derived neurotrophic factor (GDNF) exerts a potent analgesic effect and reverses the symptoms associated with neuropathic pain (Boucher et al., 2000). Because DPSC secrete important amounts of GDNF, it is conceivable that autologous DPSC transplantation to the injured trigeminal nerve may also potentially provide important benefits to treat severe orofacial pain disorders, alone or in combination with pharmacological enhancers of GDNF signaling (Hedstrom et al., 2014), some of which are even formulated to permit topical application.

# DPSC FOR SALIVARY GLAND REGENERATION

Salivary gland regeneration is an area of intense research because the most common treatment for head and neck cancer (i.e., radiotherapy) irreversibly affects salivary gland function, causing hyposalivation and xerostomia. In consequence, difficulty in mastication, swallowing, taste, speech, rampant dental caries and mucosa infections appear, leading to high discomfort in patients and a reduced quality of life (Vissink et al., 2010). Palliative treatments such as mechanical stimulation of salivary gland activity (chewing gum), or pharmacological agents such as pilocarpine (Salagen™), as well as saliva substitutes, show a limited efficacy in relieving the symptoms (Taylor, 2003).

Tissue engineering of a salivary gland replacement organ would be a conclusive way to treat these patients. To accomplish this complex task, there are some issues that must be taken into account: (i) optimization of the 3D scaffold material properties (including non-toxicity, permeability and biodegradability, among others), (ii) identification of an epithelial stem cell population capable of appropriate salivary differentiation (iii) definition of ideal culture conditions and extracellular matrix components (ECM) with laminin, integrins, MMPs to mimic the microenvironment of the gland, and (iv) selection of a population of MSC to be combined with salivary epithelial cells to generate a full bioengineered salivary gland germ. Finally, in comparison with dental bioengineering, salivary gland bioengineering presents an important additional requirement: apart from the communication between the epithelium and the mesenchyme, epithelial-axonal interactions are also required from the very beginning of salivary gland morphogenesis, because innervation plays an important role in regulating the activity of epithelial progenitor cells (Knox et al., 2010) and salivary gland branching and tubulogenesis (Nedvetsky et al., 2014). Therefore, salivary gland bioengineering strategies must also take into account that this is an organ that heavily relies on the establishment of a functional innervation to properly complete its development (Lombaert et al., 2011; Knosp et al., 2012).

Some studies indicate that isolated mouse salivary gland stem cells (cKit+, CK14+, Msi-1+, and Sca-1+) can grow and form salispheres that produce amylase (Lombaert et al., 2008). Moreover, injection of a liquid suspension containing only 300–400 cKit+ cells into the mouse SMG was shown to partially rescue the loss of morphology and function induced by irradiation insults (Lombaert et al., 2008; Coppes and Stokman, 2011; Nanduri et al., 2013). In recent years, the technology to generate salivary gland organoids in vitro has progressed remarkably. Recent major advances in the field include the advent of bioengineered glands constructed from dissociated cells from adult and fetal salivary glands (Ogawa et al., 2013; Nanduri et al., 2014). An important study by the group of T. Tsuji showed that the bioengineered salivary glands generated by a cell recombination protocol contained epithelium, mesenchyme, endothelium, and nerve cells. Most importantly, the entire organ could be transplanted into adult mice, reconnect with the existing ductal system and function properly (Ogawa et al., 2013; Ogawa and Tsuji, 2015). Thus, the salivary gland joins the tooth in the list of organs that can be fully generated ex vivo by tissue engineering. As in the case of the tooth, the main challenge to translate these findings to clinical practice is still the need for sources of stem cells that are sufficiently abundant and competent for salivary epithelial differentiation, and that do not involve destruction of donor salivary gland tissue.

In the search for stem cell sources to be employed in salivary gland bioengineering, some progress has been recently made using DPSC. It has been shown that DPSC contribute to the generation of salisphere-like structures when combined with Human salivary gland (HSG) cell lines in either Matrigel™ or hyaluronic acid hydrogel scaffolds (Janebodin and Reyes, 2012). These salispheres expressed markers of terminally differentiated acinar cells, and generated a well-irrigated salivary glandlike tissue after subcutaneous transplantation in IC mice. The principal role of DPSC in salivary gland bioengineering is to generate the salivary stroma or mesenchymal-derived compartment, and these cells are in theory one of the best choices for that purpose because the tooth mesenchyme and the salivary gland mesenchyme share a common embryonic origin (i.e., neural crest).

HSG are stable cell lines derived from neoplastic tissues (Shirasuna et al., 1981) and definitely not the best choice to bioengineer a physiologically functional salivary gland. In order to evaluate whether trophic support provided by DPSC sustains proliferation and ramification of primary salivary gland epithelia in developmental stages, in vitro recombination experiments between isolated embryonic mouse salivary epithelia and dissociated hDPSC may be performed in 3D laminin or other extracellular matrix scaffolds (Sequeira et al., 2013). In

these conditions, DPSC bind and interact with isolated salivary epithelia, improving the in vitro viability of the tissue (**Figure 4**). However, recombination with DPSC alone is clearly insufficient to promote long-term growth and ramification of developing salivary epithelia. One missing but important potential factor is that this system cannot reproduce the extensive innervation that occurs within normal salivary gland tissues and plays a crucial role in salivary gland growth and morphogenesis (Knox et al., 2010; Knosp et al., 2012). Next steps in this direction

#### REFERENCES

should involve the addition of specific growth factors that sustain epithelial cell growth and commitment to salivary gland cell differentiation, as well as the use of coculture systems between bioengineered salivary glands and nerve cells to attempt to mimic the growth stimulatory effect of natural salivary innervation.

#### CONCLUDING REMARKS

DPSC have emerged as a very promising tool with a great potential to be used in tissue engineering models aimed at the functional reconstruction of different craniomaxillofacial organs. Some of the main advantages of these cells are their multifaceted differentiation capacity, along with their nontumorigenic phenotype, a high proliferation rate, a relative simplicity of extraction and culture, and their possibility of cryopreservation, which ultimately make possible to obtain patient-specific cell lines for their use in autologous cell therapy. However, many issues and challenges still need to be addressed before these cells can be employed in clinical practice. The full control of differentiation of DPSC to specific fates is still one important issue, and although DPSC derivation to certain connective tissue-lineage cells appears to be relatively simple, their differentiation into nerve-tissue lineages still poses important unanswered questions. Moreover, the development of the cell recombination technology required to create next generation bioengineered replacement organs will necessitate extensive research for the coming years. In this case, DPSC may be an ideal choice to be used in experimental models of tissue-engineered reconstruction of organs of the oral cavity, and the paradigmatic cases of the tooth and the salivary gland may constitute the leading edge of a massive research effort on next generation organ replacement therapies.

#### FUNDING

This work was financed with project grants from the University of the Basque Country UPV/EHU (UFI11/44) and the Basque Government (IT831-13). MA, PG, and JL were sponsored by the University of the Basque Country (UPV/EHU) and VU by Jesus Gangoiti Barrera Foundation.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2015 Aurrekoetxea, Garcia-Gallastegui, Irastorza, Luzuriaga, Uribe-Etxebarria, Unda and Ibarretxe. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Manufacturing of dental pulp cell-based products from human third molars: current strategies and future investigations

Maxime Ducret 1, 2, 3, Hugo Fabre1 , Olivier Degoul 4 , Gianluigi Atzeni 4 , Colin McGuckin4 , Nico Forraz 4 , Brigitte Alliot-Licht 5 , Frédéric Mallein-Gerin1 , Emeline Perrier-Groult 1 and Jean-Christophe Farges 1, 2, 3 \*

#### Edited by:

Thimios Mitsiadis, University of Zurich, Switzerland

#### Reviewed by:

Gianpaolo Papaccio, Second University of Naples, Italy Lucia Jimenez-Rojo, University of Zurich, Switzerland Giovanna Orsini, Marche Polytechnic University, Italy

#### \*Correspondence:

Jean-Christophe Farges, Laboratoire de Biologie Tissulaire et Ingénierie thérapeutique, Institut de Biologie et Chimie des Protéines, UMR5305 Centre National de la Recherche Scientifique/Université Claude Bernard Lyon 1, 7 passage du Vercors, 69367 Lyon, France<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Craniofacial Biology, a section of the journal Frontiers in Physiology

Received: 03 June 2015 Accepted: 16 July 2015 Published: 06 August 2015

#### Citation:

Ducret M, Fabre H, Degoul O, Atzeni G, McGuckin C, Forraz N, Alliot-Licht B, Mallein-Gerin F, Perrier-Groult E and Farges J-C (2015) Manufacturing of dental pulp cell-based products from human third molars: current strategies and future investigations. Front. Physiol. 6:213. doi: 10.3389/fphys.2015.00213

Frontiers in Physiology | www.frontiersin.org August 2015 | Volume 6 | Article 213 |

1 Laboratoire de Biologie Tissulaire et Ingénierie thérapeutique, UMR5305 Centre National de la Recherche Scientifique/Université Claude Bernard Lyon 1, Lyon, France, 2 Faculté d'Odontologie, Université de Lyon, Université Claude Bernard Lyon 1, Lyon, France, 3 Hospices Civils de Lyon, Service de Consultations et Traitements Dentaires, Lyon, France, 4 CTI-BIOTECH, Cell Therapy Research Institute, Meyzieu, France, 5 Faculté d'Odontologie, Institut National de la Santé et de la Recherche Médicale UMR1064, Centre de Recherche en Transplantation et Immunologie, Université de Nantes, Nantes, France

In recent years, mesenchymal cell-based products have been developed to improve surgical therapies aimed at repairing human tissues. In this context, the tooth has recently emerged as a valuable source of stem/progenitor cells for regenerating orofacial tissues, with easy access to pulp tissue and high differentiation potential of dental pulp mesenchymal cells. International guidelines now recommend the use of standardized procedures for cell isolation, storage and expansion in culture to ensure optimal reproducibility, efficacy and safety when cells are used for clinical application. However, most dental pulp cell-based medicinal products manufacturing procedures may not be fully satisfactory since they could alter the cells biological properties and the quality of derived products. Cell isolation, enrichment and cryopreservation procedures combined to long-term expansion in culture media containing xeno- and allogeneic components are known to affect cell phenotype, viability, proliferation and differentiation capacities. This article focuses on current manufacturing strategies of dental pulp cell-based medicinal products and proposes a new protocol to improve efficiency, reproducibility and safety of these strategies.

Keywords: human dental pulp, stem cells, tissue engineering, immunophenotyping, expansion, cryopreservation, good manufacturing practices, cell-based medicinal products

## Introduction

Over the two last decades, mesenchymal stromal cells (MSC) have been intensely studied due to their potential clinical applicability to treat tissue and organ defects resulting from diseases, trauma or aging (Caplan, 1991). Their use has been proposed to repair and regenerate human mesenchymal tissues, alone or combined to scaffolds and/or morphogenic molecules (Langer and Vacanti, 1993). Bone marrow and adipose tissue are conventional sources of MSC, but invasive cell collection protocols, frequent use of general anesthesia and risk of morbidity at the collection site have stimulated the search for alternative tissues (Huang et al., 2009; Zuk, 2010; Davies et al., 2015). Third molars are frequently removed for therapeutic reasons and the connective tissue it contains, the dental pulp, can be easily recovered. They are now considered a valuable source of MSC for tissue repair and regeneration (Mayo et al., 2014). In this context, numerous investigators have attempted to obtain clinical-grade dental pulp stem/progenitor cells (DPSC) from these teeth. However, most manufacturing procedures reported so far may not be totally satisfactory, since they may alter the biological properties of the cells and the quality of the derived cell-based products (Ménard and Tarte, 2013). If such procedures are currently permitted by European and American regulation authorities, further studies are necessary to develop more efficient, reproducible, safe and standardized manufacturing processes of dental pulp cell-based medicinal products (Tirino and Papaccio, 2012; Albuquerque et al., 2014; Eubanks et al., 2014; Huang and Garcia-Godoy, 2014; La Noce et al., 2014; Nakashima and Iohara, 2014).

Dental pulp mesenchymal cells have been successfully used to regenerate human craniofacial bone (d'Aquino et al., 2009; Giuliani et al., 2013). However, these studies were performed in the absence of defined, universally accepted protocols for large-scale, clinical-grade production of DPSC (Fekete et al., 2012). This point is important in the light of recent reports indicating moderate, irreproducible and non-suitable benefits of therapies performed with various sources of MSC (Allison, 2009; Tyndall, 2011; Daley, 2012). These results were explained in part by the fact that cell performances are affected by cell isolation and expansion conditions and indicate the need for optimized and standardized procedures for MSC-based products' manufacturing (Allison, 2009; Pacini, 2014). The European Union (EU) and United States (US) have established classifications and recommended guidelines for manufacturing MSC-based products. In Europe, MSC are defined as "cell therapy products" and referred to as Advanced Therapy Medicinal Products (ATMP) (European Regulation 1394/2007). ATMP are considered Cell-Based Medicinal Products (CBMP) when containing living cells or tissues. CBMP are "medicinal products presented as having properties for, or used in or administered to, human beings with a view to treating, preventing or diagnosing a disease in which the pharmacological, immunological or metabolic actions are carried out by cells or tissues" (Schneider et al., 2010; Pacini, 2014). DPSC belong to this category and can be referred to as Dental Pulp (DP)-CBMP. In the US, DPSC are considered as Human Cells, Tissues or cellular and tissue-based Products (HCT/Ps) (Code of Federal Regulation (CFR) Title 21 CFR 1271). They are classified in two categories: (1) products that are "minimally manipulated" and used clinically in a homologous manner, and (2) products that are either "more than minimally manipulated" or used in a non-homologous manner. A cellbased product is considered as being "more than minimally manipulated" when the inherent biological characteristics of the cells have been significantly altered (Pacini, 2014).

Production and delivery of MSCs should be made in accordance with European Good Manufacturing Practices (GMP), whereas, in the US, it must comply with Current Good Tissue Practice requirements (GTP) (Fekete et al., 2012; Kellathur and Lou, 2012; Sensebé et al., 2013). GMP/GTP require many quality controls regarding donor eligibility, sample recovery, label, transport and receipt, process and storage, laboratory equipment, supplies and reagents, cell-based product distribution to recipient patients and documentation that must be maintained by the handler (Alici and Blomberg, 2010; Abou-El-Enein et al., 2013; Sensebé et al., 2013; Wuchter et al., 2015). These controls make GMP/GTP procedures long and costly, and further studies are encouraged to develop shorter, less expensive and more standardized procedures for DP-CBMP manufacturing (Albuquerque et al., 2014; Eubanks et al., 2014; Huang and Garcia-Godoy, 2014; La Noce et al., 2014; Nakashima and Iohara, 2014; Hilkens et al., 2015). In the present paper, we will firstly review current international guidelines regarding the five manufacturing steps of DP-CBMP (**Figure 1**), and then we will highlight the drawbacks and potential risks of actual strategies. Finally we will propose modifications of the protocols intended to increase the efficiency, reproducibility and safety of these strategies, from tooth extraction to the harvest of clinical-grade DP-CMBP.

# Teeth Collection and Pulp Tissue Recovery

Since the discovery of DPSC by Gronthos et al. (2000), numerous papers have reported the isolation of stem/progenitor cells from the dental pulp of human third molars. However, since there are no rules specifying the best tooth development stages for pulp cell collection, teeth were collected in patients of various ages and therefore at various developmental stages. It also greatly impairs the interpretation and comparison of the experimental results.

Transport from the operating block to the laboratory notably requires a medium that does not affect cell viability. It was previously shown that DPSC remain viable up for 5 days when extracted teeth are maintained in phosphate-buffered saline (PBS) (Perry et al., 2008; Woods et al., 2009) and this time is more than enough for the transport of samples to the laboratory and pulp recovery.

## Dental Pulp Cell Isolation and Enrichment

After dental pulp recovery, two options are possible for isolating dental pulp cells: enzymatic dissociation and explant culture. Enzymatic dissociation consists of digesting the pulp tissue with collagenase and dispase enzymes to liberate the cells that are then plated on culture dishes. However, a growing number of authors consider that enzymatic dissociation is not adapted to medicinal manufacturing, owing to its putative consequences on cell phenotype and properties (Shah et al., 2013; Busser et al., 2014; Ohnuma et al., 2014). In addition, tissues and cells exposed to collagenase are considered "more than minimally manipulated" by FDA [Code of Federal Regulation (CFR) Title 21 CFR 1271] and potentially require the use of pharmaceutical grade manufactured enzymes, which significantly increases the scale-up costs. By contrast, cell isolation by explant culture increasingly appears easier, faster, safer, less expensive and more in line with GMP guidelines to obtain clinical-grade amounts of MSC (Hilkens et al., 2013). It is based on the growth of cells out of tissue fragments (explants) that are plated on culture dishes. It recently allowed for efficient recovery of human adipose or Wharton jelly stem/progenitor cells in serum-free,

xeno-free medium conditions (Busser et al., 2014; Swamynathan et al., 2014). Additionally, explant-derived DPSC display similar or enhanced differentiation abilities compared with cells from dissociated tissue (Spath et al., 2010; Hilkens et al., 2013).

safe and less expensive procedures (Step 2), cell expansion in defined,

Cell selection by sorting methods has been proposed to enrich the cultured cell population in stem/progenitor cells based on their expression of specific surface markers such as CD34, CD184, Stro-1, CD146, CD271, and MSCA-1 (Shi and Gronthos, 2003; d'Aquino et al., 2009; Waddington et al., 2009; Yu et al., 2010; Jiang et al., 2012; Tomlinson et al., 2015). However, the use of such a procedure is today limited by the complexity of the technique and the prohibitive cost (Kawashima, 2012; Nakashima and Iohara, 2014). Besides, multiplying steps and using additional reagents increase the risk of microbial contamination and the difficulty to obtain CBMP in GMP conditions. The same reservation can be made regarding the use of biophysical markers that have been found relevant to isolate MSC in an easier and more predictable way than biochemical markers (Lee et al., 2014).

# Dental Pulp Cell Culture and Expansion

Among other factors, the composition of the culture medium and the presence of a coating material on the culture dish may influence the nature and the quality of the final CBMP and therefore the clinical results (Lopez-Cazaux et al., 2006; Majd et al., 2009; Jung et al., 2012; Pisciotta et al., 2012; Pacini, 2014). Currently, CBMP manufacturing under GMP procedures recommends the use of xeno-free materials and reagents to prevent the risk of viral, bacterial, fungal and prion contamination, and the possible induction of immunizing effects in the final recipient. Additionally, industrial production is responsible for frequent batch-to-batch serum variability and the serum itself can promote early cell differentiation (Mannello and Tonti, 2007; Jung et al., 2012). For these reasons, the supplementation of the cell culture medium with xeno- or allogeneic products should be limited to "cases for which a valid alternative cannot be found" (European Regulation 1394/2007). Today, the development of xeno-free, serum-free, defined media, able to rapidly expand stem/progenitor cells without impairing their differentiation capabilities, represents a major objective for the standardization of DP-CBMP production (Tekkatte et al., 2011; Jung et al., 2012; Bonnamain et al., 2013; Carvalho et al., 2013). Multiple passages are often necessary to obtain a clinicalscale amount of cells, but they may lead to a slow-down of the proliferation rate, progressive cell senescence and loss of multipotentiality that prevent future cell differentiation (Baxter

development of a bioassay (Step 5).

et al., 2004; Bork et al., 2010; Yu et al., 2010; Sensebé et al., 2013). In our culture conditions, cell doubling times remained constant from P1 to P4 (≈40 h) and we calculated that more than 25.107 cells could be theoretically obtained after four passages with one dental pulp, which is likely to be a sufficient cell number for one pulp regeneration, bone socket filling, or for localized periodontal treatment (Kaigler et al., 2013; Albuquerque et al., 2014).

# DP-CBMP Manufacturing Control

DP-CBMP manufacturing requires advanced quality controls of the safety, identity and efficacy of the final product (Wang et al., 2005; Sensebé et al., 2013). Since CBMP cannot undergo sterilization before implantation, the absence of bacteria, virus, fungi and prion contamination has to be checked. The presence of endotoxin must also be tested to prevent immune reactions in the recipient patient. Long-term ex vivo expansion of cells increases the risk of genetic instability and the occurrence of potential chromosomal abnormalities, since there exists a close relation between cell senescence and risk of transformation (Baxter et al., 2004; Rubio et al., 2005; Campisi, 2007). To limit this risk, the number of population doublings should be kept to a minimum. In addition, conventional karyotyping must be combined with fluorescence in situ hybridization (FISH) or comparative genomic hybridization (CGH array) to assess the genomic stability of scaled-up cell populations (Barkholt et al., 2013).

The control of the population identity into expanding cell cultures is generally realized by flow cytometry analysis of surface antigens. During the past decade, most of these controls have been realized in compliance with the recommendations of the International Society of Cellular Therapy (ISCT) (Dominici et al., 2006). However, it is today acknowledged that several markers initially proposed by ISCT for the positive characterization of MSC (for instance CD73, CD90, and CD105) are shared by several populations of cells including progenitor cells, mature fibroblasts or perivascular cells (Russell et al., 2010; Alt et al., 2011; Halfon et al., 2011; Al-Nbaheen et al., 2013; Lv et al., 2014).

# DP-CBMP Uses

Over recent years, DP-CBMP were clinically tested with the aim to regenerate human craniofacial bone. DP-CBMP were implanted, in association with a collagen I-based sponge scaffold, in mandibular bone sockets in a phase I clinical trial (d'Aquino et al., 2009). Three years after DP-CBMP grafting, the tissue regenerated in the graft site was compact bone (Giuliani et al., 2013). Case reports of osteoradionecrosis treatment using DP-CBMP were also reported (Manimaran et al., 2014). The angiogenic, neurogenic and odontogenic potential of DP-CBMP was also successfully tested in preclinical studies (Gandia et al., 2008; Iohara et al., 2009; Sakai et al., 2012; Ishizaka et al., 2013). In addition, a phase I clinical trial is currently under progress to evaluate the DP-CBMP potential to regenerate the human dental pulp (Nakashima and Iohara, 2014). Despite these successes, potential applicability of DP-CBMP will be closely dependent on their final production cost and their large-scale clinical outcomes. In particular, a high cost-efficacy ratio would constitute a serious impediment for their routine use. Hence, it is necessary to have a clear overview and understanding of the complete value chain to try to reduce costs (Abou-El-Enein et al., 2013, 2014; Leijten et al., 2015).

Storage of cryopreserved cell-based products (cryobanking) over long periods of time offers unique opportunities to increase DP-CBMP applicability. However, similar to cell culture and expansion, cryopreservation is associated with infective, prion, toxicological and immunological risks owing to the presence of human or animal components and additives such as DMSO in the storage medium (Papaccio et al., 2006; Perry et al., 2008; Woods et al., 2009; Lee et al., 2012). Accordingly, xeno-free, defined cryopreservation media must be privileged.

DP-CBMP could also be used in biomedical research as components of bioassay kits to investigate the effects of drugs on dental pulp cells in a reproducible "humanized" system (Jurga et al., 2010; Leeb et al., 2011; Forraz et al., 2013). Such kits are reliable preclinical alternatives to animal models in the actual regulatory context. Assessment of the risks related to chemical products' use and screening or testing new therapeutic molecules are indeed extremely complicated and costly. The average costs to take a blockbuster drug to clinical trials are estimated to be around 1 billion euros. Furthermore, the accuracy of toxicological and preclinical studies greatly depends on the experimental animal models used for such evaluations. In particular, rodent species, widely use, are known to only partially mimic the human biological system. Development of DP-CBMP bioassay kits would offer a prime platform to successfully induce dentinogenesis, osteogenesis or neurogenesis in vitro (Zhang et al., 2006; Woloszyk et al., 2014; Jensen et al., 2015; Leijten et al., 2015).

# Proposals for a Protocol with a More GMP Compliant Approach (Figure 2)

We recently proposed the use of impacted third molars between Nolla's developmental stages 5 (crown almost completed) and 7 (one third root completed). The presence of large, open apices in teeth without roots or with roots partially developed allows for an easy access to the pulp tissue and its gentle, atraumatic extirpation from the enamel/dentin shell with fine tweezers. It avoids the cell stress resulting from the crown-root mechanical separation with a drill or a clamp that is necessary for recovering pulps from teeth with more developed or complete roots (Perry et al., 2008; Takeda et al., 2008; Ducret et al., in press). Additionally, human dental pulp cells (HDPC) isolated at around the crowncompleted stage displayed short cell doubling times and high growth rate (Takeda et al., 2008). We found similar results in our study. We also selected impacted teeth to minimize the risk of pulp tissue contamination and disease transmission by oral microorganisms (Nolla, 1960; Ducret et al., in press). This choice may enable to skip the step of sample disinfection performed

with chemicals such as chlorhexidine or povidone-iodine/sodium thiosulfate (Perry et al., 2008; d'Aquino et al., 2009). When using PBS as a transport medium, we failed to detect any contamination in cultures of HDPC (n > 50 patients) during the isolation and expansion steps, contrary to others (Perry et al., 2008; Ducret et al., in press). This might be related to our selection of impacted teeth from young patients (13–17 year-old) that have never been in contact with the septic oral cavity, versus the selection of erupted ones from older patients (18–30 year-old) by those authors.

Regarding cell isolation, we used explant culture for recovering human dental pulp cells for clinical application. Each pulp sample was cut into about 20 explants that allowed for the harvest of a total of one million dental pulp cells after 14 days of culture (Ducret et al., in press). This result is in agreement with other studies reporting that, whatever the technique used (tissue dissociation or explant culture), 2 weeks of culture allow for the recovery of about 106 cells from one third molar pulp (Eubanks et al., 2014).

We pre-coated the culture dish surface for cell isolation and culture with an equal mixture of human placental collagens I and III. This composition was chosen because they are the two most abundant collagens in the dental pulp extracellular matrix. Xenofree dissociating reagents (such as TrypLe R or Accutase R ) and xeno-free defined culture medium (such as SPE-IV R [ABCell-Bio, France], containing clinical grade human albumin, α-MEM, rhIGF-1 and rhFGF-2) are recommended for cell culture and passaging instead of the products commonly used (Carvalho et al., 2013; Ducret et al., in press). Moreover, cryopreservation of dental pulp cells in serum-free medium had no negative impact on cell doubling times and cumulative cell numbers (Ducret et al., in press). Although the viability of cells cryopreserved in serumfree medium was decreased compared to fresh cells, it is similar to that previously reported (Lee et al., 2012).

Future investigations are required for identifying more specific membrane markers for these cells. In our study, immunophenotypic analysis of 17 surface markers revealed that our dental pulp cell expanding population was made of mesenchymal cells, a percentage of whom expressed the mesenchymal stem cell/progenitor markers CD146 and MSCA-1. The number of cells expressing these markers remained similar from P1 to P4, suggesting that the cell fate was not significantly affected by our culture conditions. In addition, cell karyotyping by G-band analysis showed that this rapid expansion did not lead to genomic instability that would be potentially harmful for the recipient patient (Ducret et al., in press).

#### Conclusion and Perspectives

Recent successes in bone and dental pulp regeneration therapies carry the promise to use dental pulp-cell-based medicinal products in the near future. However, current strategies to manufacture DP-CBMP are not totally satisfactory since they do not comply with current international guidelines. New manufacturing standardized protocols, intended to increase efficiency, reproducibility and safety of these strategies, are

#### References

urgently needed. Further investigations are also warranted to estimate the real benefit of DP-CBMP use compared to current therapeutic options and precisely determine the cost-efficacy ratio that risks being a major block for the large-scale clinical use of these cell-based products.

#### Acknowledgments

The authors thank the CNRS, the French Ministry of Higher Education and Research, the French Institute for Odontological Research (IFRO), the Société Française de Rhumatologie and the Société Française d'Endodontie for their financial support. HF holds a doctoral fellowship from the Région Rhône-Alpes.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2015 Ducret, Fabre, Degoul, Atzeni, McGuckin, Forraz, Alliot-Licht, Mallein-Gerin, Perrier-Groult and Farges. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Use of Rat Mature Adipocyte-Derived Dedifferentiated Fat Cells as a Cell Source for Periodontal Tissue Regeneration

Daisuke Akita1 , Koichiro Kano2 , Yoko Saito-Tamura3 , Takayuki Mashimo4 , Momoko Sato-Shionome5 , Niina Tsurumachi 3 , Katsuyuki Yamanaka6 , Tadashi Kaneko6 , Taku Toriumi 7 , Yoshinori Arai 8 , Naoki Tsukimura1 , Taro Matsumoto9 , Tomohiko Ishigami 1 , Keitaro Isokawa7 and Masaki Honda10 \*

*1 Department of Partial Denture Prosthodontics, School of Dentistry, Nihon University, Tokyo, Japan, 2 Laboratory of Cell and Tissue Biology, College of Bioresource Science, Nihon University, Fujisawa, Japan, 3 Department of Orthodontics, School of Dentistry, Nihon University, Tokyo, Japan, 4 Department of Oral and Maxillofacial Surgery, Faculty of Medicine, Juntendo University, Tokyo, Japan, 5 Department of Pediatric Dentistry, School of Dentistry, Nihon University, Tokyo, Japan, 6 GC Corp. R&D, Tokyo, Japan, 7 Department of Anatomy, School of Dentistry, Nihon University, Tokyo, Japan, 8 School of Dentistry, Nihon University, Tokyo, Japan, 9 Division of Cell Regeneration and Transplantation, Department of Functional Morphology, School of Medicine, Nihon University, Tokyo, Japan, 10 Department of Oral Anatomy, School of Dentistry, Aichi-Gakuin University, Nagoya, Japan*

#### Edited by:

*Thimios Mitsiadis, University of Zurich, Switzerland*

#### Reviewed by:

*Thomas G. H. Diekwisch, University of Illinois at Chicago, USA Petros Papagerakis, University of Michigan, USA Giovanna Orsini, Marche Polytechnic University, Italy*

> \*Correspondence: *Masaki Honda<EMAIL_ADDRESS>
#### Specialty section:

*This article was submitted to Craniofacial Biology, a section of the journal Frontiers in Physiology*

Received: *16 November 2015* Accepted: *02 February 2016* Published: *23 February 2016*

#### Citation:

*Akita D, Kano K, Saito-Tamura Y, Mashimo T, Sato-Shionome M, Tsurumachi N, Yamanaka K, Kaneko T, Toriumi T, Arai Y, Tsukimura N, Matsumoto T, Ishigami T, Isokawa K and Honda M (2016) Use of Rat Mature Adipocyte-Derived Dedifferentiated Fat Cells as a Cell Source for Periodontal Tissue Regeneration. Front. Physiol. 7:50. doi: 10.3389/fphys.2016.00050* Lipid-free fibroblast-like cells, known as dedifferentiated fat (DFAT) cells, can be generated from mature adipocytes with a large single lipid droplet. DFAT cells can re-establish their active proliferation ability and can transdifferentiate into various cell types under appropriate culture conditions. The first objective of this study was to compare the multilineage differentiation potential of DFAT cells with that of adipose-derived stem cells (ASCs) on mesenchymal stem cells. We obtained DFAT cells and ASCs from inbred rats and found that rat DFAT cells possess higher osteogenic differentiation potential than rat ASCs. On the other hand, DFAT cells show similar adipogenic differentiation, and chondrogenic differentiation potential in comparison with ASCs. The second objective of this study was to assess the regenerative potential of DFAT cells combined with novel solid scaffolds composed of PLGA (Poly *d, l*-lactic-co-glycolic acid) on periodontal tissue, and to compare this with the regenerative potential of ASCs combined with PLGA scaffolds. Cultured DFAT cells and ASCs were seeded onto PLGA scaffolds (DFAT/PLGA and ASCs/PLGA) and transplanted into periodontal fenestration defects in rat mandible. Micro computed tomography analysis revealed a significantly higher amount of bone regeneration in the DFAT/PLGA group compared with that of ASCs/PLGA and PLGA-alone groups at 2, 3, and 5 weeks after transplantation. Similarly, histomorphometric analysis showed that DFAT/PLGA groups had significantly greater width of cementum, periodontal ligament and alveolar bone than ASCs/PLGA and PLGA-alone groups. In addition, transplanted fluorescent-labeled DFAT cells were observed in the periodontal ligament beside the newly formed bone and cementum. These findings suggest that DFAT cells have a greater potential for enhancing periodontal tissue regeneration than ASCs. Therefore, DFAT cells are a promising cell source for periodontium regeneration.

Keywords: periodontal tissue regeneration, dedifferentiated fat cells (DFAT cells), PLGA scaffold, cell transplantation, periodontal fenestration defect

#### Akita et al. DFAT Cells for Periodontal Tissue Regeneration

# INTRODUCTION

The use of adult mesenchymal stem cells (MSCs) seems to be ideal for practical periodontal regenerative medicine, because they are not subject to the restrictions such as embryonic stem cells (ESCs) or induced pluripotent stem cells (Prockop, 1997; Pittenger et al., 1999; Zuk et al., 2001; Reyes et al., 2002; Safford et al., 2002; Akita et al., 2014). ASCs may be a promising cell source for periodontal-tissue regeneration (Tobita et al., 2008; Tobita and Mizuno, 2013; Akita et al., 2014). The use of ASCs has several advantages over bone marrow-derived or dental tissuederived MSCs such as dental follicle (Honda et al., 2007, 2010, 2011), dental pulp (Aurrekoetxea et al., 2015; Sato et al., 2015), and periodontal ligament (Saito et al., 2013). Adipose tissue contains large numbers of stromal cells and is available in larger quantities than bone marrow and dental tissue (Fraser et al., 2006). Furthermore, the method for obtaining adipose tissue is relatively easy and is less invasive than that for bone marrow, and adipose tissue can be obtained from patients without teeth. The utility of ASCs for periodontal tissue regeneration has been demonstrated in rat and canine models in several studies, but successful use of human ASCs for such purpose has not been described yet (Tobita et al., 2008; Tobita and Mizuno, 2013; Akita et al., 2014).

Adipose tissue is connective tissue composed mostly of adipocytes surrounded by fibroblasts. Mature adipocytes with a large single lipid droplet are generally considered to be terminally differentiated cells which have lost their proliferative ability. However, by using a ceiling culture methods based on buoyancy, lipid-free fibroblast-like cells, known as dedifferentiated fat (DFAT) cells, can be separated from mature adipocytes in our previous studies (Yagi et al., 2004; Nobusue et al., 2008; Nobusue and Kano, 2010). DFAT cells can re-establish their active proliferation ability and can differentiate into various cell types under appropriate culture conditions (Matsumoto et al., 2008; Kono et al., 2014). DFAT cells may have the potential to contribute to dental tissue regeneration (Kaku et al., 2015).

A three-dimensional (3D) biodegradable scaffold is likely to be necessary for the delivery of stem cells or progenitor cells to periodontal defect, both to preserve space for the formation of new periodontal tissue and to provide initial support for the growing transplanted cells (Wang et al., 2005). Since the most extensively used synthetic polymers in tissue engineering studies are poly(glycolic acid), poly(lactic acid), and their copolymers (e.g., Poly d, l-lactic-co-glycolic acid; PLGA), we previously examined the in vivo performance of solid PLGA scaffolds seeded with ASCs (Akita et al., 2014). Solid PLGA scaffolds have large fully interconnected pores and substantially higher compressive strength than sponge-like PLGA-based scaffolds. Recently, the possibility of using DFAT cells to promote periodontal tissue regeneration was raised by researchers who seeded an atelocollagen sponge-like scaffold with DFAT cells (Sugawara and Sato, 2014). An advantage of the higher compressive strength of solid PLGA scaffolds is that they typically offers higher primary stability than natural scaffolds such as those composed of atelocollagen. Our results showed that the PLGA scaffolds maintained their structural integrity for 5 weeks when used for in vivo transplants (Akita et al., 2014). We concluded that these solid PLGA scaffolds are useful for regeneration of periodontium.

To date, no studies evaluating DFAT cells combined with solid PLGA scaffolds for periodontal tissue regeneration have been published. We first compared the characteristics of rat DFAT cells with those of rat ASCs—including proliferative and multipotent differentiation potential. We then evaluated the in vivo potential for periodontal tissue regeneration of rat DFAT cells combined with solid PLGA scaffolds in periodontal fenestration defects created in mandibular alveolar bone, and compared the performance of rat DFAT cells in this context with that of ASCs.

#### MATERIALS AND METHODS

All animal experiments were reviewed and approved by the Animal Research and Care Committee at the Nihon University School of Dentistry (AP10D014 and AP15D006).

#### Isolation of Rat DFAT Cells and ASCs

To isolate DFAT cells and ASCs, 9-week-old male F344 rats (n = 5, body weight 190 ± 10 g) were purchased from CLEA Japan, Inc. (Tokyo, Japan). Isolation of DFAT cells from mature adipocytes was done with a modified version of a method that has been described previously (Matsumoto et al., 2008). Briefly, ∼1 g of inguinal subcutaneous fat tissue was washed extensively with phosphate-buffered saline (PBS; Wako, Osaka, Japan) and minced and digested in 0.1% (w/v) collagenase solution (C6885; Sigma-Aldrich, St. Louis, MO) at 37◦C for 60 min with gentle agitation. After filtration and centrifugation at 135 g for 3 min, the floating primary mature adipocytes in the top layer were collected. After three washes with PBS, cells (5 × 104 ) were placed in 12.5 cm2 culture flasks (BD Falcon, England) filled completely with Dulbecco's modified Eagle's medium (DMEM; Sigma-Aldrich, UK) and supplemented with 20% fetal bovine serum (FBS; Nichirei Bioscience Inc., Tokyo, Japan), and were incubated at 37◦C in 5% CO2. Mature adipocytes floated up and adhered to the top inner surface (ceiling surface) of the flasks. After about a week, the medium was removed and changed into DMEM supplemented with 20% FBS, and the flasks were inverted so that the cells were on the bottom (**Figure 1**). The medium was changed every 4 days until the cells reached confluence.

Cultured ASCs were prepared as described previously (Tobita et al., 2008; Tobita and Mizuno, 2013; Akita et al., 2014). Briefly, the stromal vascular fraction (SVF) was isolated as the pellet fraction from collagenase-digested adipose tissue by centrifugation at 180 g for 5 min after collecting of the floating uppermost layer as described above. The remaining cells were plated in DMEM supplemented with 20% FBS and 1% antibiotics as a growth medium. The cells were referred to as SVF cells, and were cultured at 37◦C in a humidified atmosphere containing 5% CO2 (**Figure 1**). In the comparison experiments, second and third passage ASCs and DFAT cells from the same samples were used. All animal experiments were reviewed and approved by the Animal Research and Care Committee at the Nihon University School of Dentistry (AP10D014 and AP15D006).

# Adipogenic, Osteoblastic, and Chondrocytic Differentiation In vitro

Differentiation assays for adipocytes, osteoblasts, and chondrocytes were performed as previously described (Matsumoto et al., 2008; Mikami et al., 2011; Saito et al., 2013; Akita et al., 2014; Sato et al., 2015). The three cell lines obtained from three rats were used in those studies.

For adipogenic differentiation, the cells were plated in 6-well plates at a density of 5 × 104 cells and grown to confluency with growth medium. The cells were then incubated for 21 days in DMEM containing 10% FBS, 1 mM dexamethasone (Sigma-Aldrich), 0.5 mM isobutylmethylxanthine (Sigma-Aldrich), and 1× insulin-transferrin-selenium-A (ITS; GIBCO, Grand Island, NY) as an adipogenic induction medium, and lipid vacuole formation was examined by Oil red O staining (Wako Pure Chemical Industries, Osaka, Japan). To evaluate the adipogenic differentiation ability, Oil red O-positive cells were count under the phase contrast microscope.

For osteogenic differentiation, cells were plated in 6-well plates at a density of 5 × 104 cells and grown to confluency with growth medium. The growth medium was replaced with osteogenic induction medium including DMEM containing 10% FBS, 100 nM dexamethasone, 10 mM glycerol 2-phosphate disodium salt hydrate (Sigma-Aldrich), and 50 mM L-ascorbic acid phosphate magnesium salt n-hydrate (Wako Pure Chemical Industries). The cells were cultured with the osteogenic induction medium for 21 days and stained with alkaline phosphatase (ALP) activity using nitro-blue tetrazolium plus 5-bromo-4 chloro-3′ -indolyphosphate (NBT/BCIP) ready-to-use tablets, pH 9.5 (Roche Diagnostics, Pentzberg, Germany). To detect calcium deposition, cells were incubated in 1% Alizarin red S (Sigma-Aldrich). Ca2+ evaluated was also determined by measuring Ca2+ uptake using the Calcium E-test (Wako Pure Chemical Industries).

For chondrogenic differentiation, cell pellets were collected in centrifuge tube (2.5 × 104 cells/tube) with growth medium. After 2 days in culture, the growth medium was replaced with chondrogenic induction medium including DMEM containing 1% FBS, 50µM L-ascorbic acid phosphate magnesium salt n-hydrate, 40µg/mL proline (Sigma-Aldrich), 100µg/mL pyruvate (Sigma-Aldrich), 10 ng/mL recombinant human TGFβ3 (R&D Systems, Inc., Minneapolis, MN, USA), and 1 × ITS for 21 days. The microscopic structure of cells was examined after cell pellets were fixed for 10 min with 10% neutral buffered paraformaldehyde, dehydrated in a graded series of ethanol solutions, embedded in paraffin, and cut into 7µm sections. After deparaffinization, sections were stained with Alcian blue (0.1 N, pH 1.0). Each test was conducted three times and results are presented as the mean ± standard deviation.

#### Preparation of Cell-Scaffold Complex

The PLGA-based solid scaffolds (LA:GA = 75:25, Mw. 25 kDa; PLGA scaffold, GC Dental Product Co. Ltd.), with a porosity of 80%, were prepared as previously described (Akita et al., 2014; Yamanaka et al., 2015). Briefly, PLGA scaffolds were resized to ∼2 × 3 × 1 mm3 and soaked in 70% ethanol for 30 min to improve wetting, and then a vacuum pump was used to remove air bubbles. The scaffolds were then washed three times (30 min each) in PBS to remove residual ethanol, and 100µL aliquots of the cell suspension (1 × 107 cells/mL, 1 × 106 cells/scaffold) were seeded onto the tops of prewetted PLGA scaffolds and left undisturbed in an incubator for 1 h. As a negative control, empty PLGA scaffolds were prepared.

#### Cell Transplantation

Fifteen healthy male F344 rats (10-weeks-old; 200 ± 10 g) were used for in vivo experiment. Under inhalational device of Isoflurane (KN-1071 Marcobit-E; Natsume Seisakusho Co. Ltd., Tokyo, Japan), the rat periodontal fenestration defect model was prepared as previously described (King et al., 1997; Yang et al., 2010; Han et al., 2013; Akita et al., 2014). A skin incision was made along the inferior border of the left mandible, then the masseter muscle and the periosteum covering the buccal surface of the mandible were elevated as a flap. The alveolar bone overlying the mandibular first and second molar roots was removed with a dental inverted bur. The size of the periodontal defect was ∼2 × 3 × 1 mm in height, width, and depth, respectively, with the anterior margin mesial to the distal root of the first molar and the posterior margin just distal to the second molar. The coronal margin was ∼1.5 mm apical to the crest of the alveolar bone and the inferior margin was ∼2.5 mm apical to the alveolar crest (**Figures 2A,B**). The exposed roots of the first and second molars were denuded and cementum was completely removed together with the supporting bone, which occasionally resulted in penetration into the root dentin (**Figures 2C,D**). PLGA scaffolds with DFAT (DFAT/PLGA) or ASCs (ASCs/PLGA) were placed in the defects and covered with membrane (GC membrane; GC Dental Product Co. Ltd.). The unilateral mandibular first molar in each of the 15 rats were selected for analysis and the defects were randomly assigned to the following three treatment groups: (1) PLGA scaffolds covered with GC membrane (GC membrane; GC Dental Product Co. Ltd.) without cells (PLGA-alone group) (n = 5), (2) ASCs-loaded PLGA scaffolds covered with membrane (ASCs/PLGA group) (n = 5), and (3) DFAT cells-loaded PLGA scaffolds covered with membrane (DFAT/PLGA group) (n = 5). The right side of the

# In vivo Micro Computed Tomography (CT) Imaging and Analysis

ensure healing by primary intention.

mandibular was not operated to maintain oral function. After the masseter muscle was repositioned, the skin incision was closed to

In vivo x-ray micro computed tomography (R\_mCT; Rigaku Corporation, Tokyo, Japan) was used as previously described (Akita et al., 2014). The exposure parameters were 17 s, 90 kV, and 100µA. The isotropic voxel size was 30µm.

Briefly, hard tissue regeneration images were obtained from each rats immediately after surgery and each week until 5 weeks after surgery. The images were constructed into 3D images using i-View (J. Morita Co., Kyoto, Japan). Bone volume was measured in the regions of interest (ROIs) from voxel images using bone volume-measuring software 3 by 4 viewer 2011 (Kitasenjyu Radist Dental Clinic I-View Image Center, Tokyo, Japan). The ROI size was 4 × 3 × 1.5 mm, which covered the surgically created periodontal tissue defect. The bone volume in the ROI was measured immediately after surgery and each week until 5 weeks after surgery. The increase in bone volume in individual rats was then calculated by subtracting the bone volume on day 0 from each of the subsequent values. The increase in bone was considered to be defect re-ossification.

# Histological and Histometric Analysis

Five weeks after surgery, the harvested specimens were fixed in 10% neutral buffered paraformaldehyde for 24 h, decalcified in 10% ethylenediaminetetraacetic acid (EDTA) for 5 weeks, dehydrated through a graded series of ethanol solutions, and then embedded in paraffin. For the rat specimens, frontal plane sections (7µm thick) were prepared with a microtome (Leica RM2165, Nussloch, Germany) and the paraffin sections of the first molar central and distal roots were stained with hematoxylin and eosin (H&E). To observe the periodontal ligament fibers and Sharpey's fibers, the paraffin sections were stained with Azan and Picrosirius-red.

For the quantitative analysis, five H&E-stained sections per specimen were selected and the thickness of newly formed cementum and periodontal ligament in the first molar central and distal roots were measured by light microscopy (ECLIPSE LV100POL, Nikon, Tokyo, Japan).

After decalcification, the sections were stained as follows: dewaxing and rehydration; immersion in 0.1% Picrosirius-red

FIGURE 2 | Creation of the periodontal fenestration defect. (A) Rat mandibular frame specimen. The black flame indicates the site of the surgically created periodontal fenestration defect site. (B) Macroscopic view of a surgically created periodontal defect on the buccal surface of the mandibular first molar. The arrow indicates the first molar mesial root and the arrowhead indicates the first molar distal root. (C) H&E staining showing the surgically exposed central root of the first mandibular molar. The defect includes alveolar bone, periodontal ligament, and cementum. The arrows indicate the defect site and the asterisk indicates the central root of the first molar on the buccal side. Scale bar = 1000 µm. (D) H&E staining showing surgically exposed distal root of the first mandibular molar. The arrows indicate the defect site and the asterisk indicates the distal root of the first molar on the buccal side. Scale bar = 1000 µm.

solution (Sirius red 0.1 g dissolved into 100 mL of saturated picric acid solution) for 1 h of staining; rinsing with water for 5 min; re-staining with Harris hematoxylin for 5 min, dehydration with a gradient series of ethanol solutions; treatment with a xylene solution; and sealing with a rubber mount. The amount, distribution, and morphology of each different type of collagen were assessed by polarized light microscopy.

# Localization of Fluorescent-Labeled DFAT Cells

Four healthy male F344 rats (10-weeks-old; 200 ± 10 g) were used for this experiment. Cells harvested with 0.25% trypsin-EDTA, resuspended at 1 × 106 cells/ml in DMEM, and labeled with a fluorescent dye—chloromethylbenzamido 1,1′ -dioctadecyl-3,3,3′ ,3′ tetramethylindocarbocyanine (DiI; Vybrant R , V22885, Life Technologies, Eugene, OR, USA). Fluorescent lipophilic tracer was added at 5µL/mL in DMEM. After incubation for 20 min at 37◦C with 5% humidified CO2, the cells were centrifuged at 180 g for 5 min and washed twice with PBS. To prepare cryosections, fixed, and dehydrated specimens after 5 weeks fluorescent-labeled cell transplantation were embedded immediately in OCT compound (Sakura Finetechnical Co. Ltd., Tokyo) and frozen in liquid nitrogen. After cryosections were incubated for 30 min with a fluorescent 4′ ,6-diamidino-2-phenylindole solution (DAPI; Sigma-Aldrich), fluorescence microscopy (Biozero BZ-8000, Keyence, Osaka, Japan) was used to investigate the survival and localization of transplanted cells.

#### Statistical Analysis

Data was expressed as the mean and standard deviation (SD) for each group. Statistical analysis was performed using ExcelStatistical File software (ystat2008.xls; Igakutosyosyuppan, Tokyo, Japan). The Mann–Whitney U-test was used for intergroup comparisons. P < 0.05 was considered statistically significant.

## RESULTS

#### Characterization of Rat DFAT Cells

After ceiling culture, the adhered mature adipocytes divided asymmetrically and generated cuboidal cells with scanty cytoplasm as described previously (Matsumoto et al., 2008). The cuboidal cells formed colonies on day 7 (**Figure 3A**). We also successfully prepared ASCs from SVF fractions; these cells also expanded and formed colonies on day 7 as described in our previous reports (**Figure 3B**) (Akita et al., 2014). The morphology of the ASCs was slightly different from that of the cells derived from mature adipocytes. The ASCs have more cytoplasm than the DFAT cells.

Next, differentiation potential was analyzed. Patterns of increase in ALP activity were detected in both cell populations with or without differentiation-inducing medium. Since the extent of the formation of Alizarin Red-positive mineralized nodules in DFAT cells was slightly higher than that observed in ASCs on day 14 (**Figure 4A**), calcium accumulation was measured. Calcium accumulation in osteogenic induction medium was significantly higher in DFAT cells than in ASCs at day 14 and 21 (**Figure 4B**). Newly synthesized glycosaminoglycan, as determined by Alcian-blue positive staining, was observed in both cell populations when subjected to conditions that favor chondrogenic differentiation (**Figure 4C**). DFAT cells showed Alcian-blue positive areas in the cell pellets in chondrogenic induction medium compared with that of ASCs. The accumulation of lipid vacuoles as determined by positive Oil red O staining was observed in the cytoplasm in both cell populations when subjected to conditions that favor adipogenic differentiation (**Figure 4D**). The number of Oil red O-positive cells in the DFAT cell population was similar to that in the ASC population (**Figure 4E**).

# Hard Tissue Formation

Furcation periodontal tissue defects were artificially prepared and transplanted with PLGA-alone, ASCs/PLGA, or DFAT/PLGA scaffolds. No severe inflammation or swelling was observed in any examined sites throughout the experimental period. To evaluate hard tissue formation in the alveolar bone areas, reconstituted micro computed tomography (micro-CT) images and horizontal sections were prepared by micro-CT until week 5 because the artificially created defect area was almost filled with newly formed hard tissue by week 5 in the DFAT/PLGA scaffold group (**Figure 5**).

Newly formed hard tissue was clearly visible at the alveolar bone defect sites in all groups at week 2. Interestingly, cortical bone-like tissue was observed at the outer layer of the defect sites at 4 weeks in the DFAT/PLGA scaffold, but not the ASCs/PLGA and PLGA-alone scaffolds by the horizontal images. At 5 weeks, the newly formed hard tissue was completely connected with the native cortical bone at both mesial and distal side in the DFAT/PLGA group. The translucent image between dental root and the newly formed hard tissue was clearly visible in all the samples by week 5. Since the DFAT/PLGA scaffolds showed large amounts of newly formed hard tissue compared with the ASCs/PLGA scaffolds, we calculated the quantity of new hard tissue formation. The hard tissue volume in DFAT/PLGA scaffolds was significantly higher than that in ASCs/PLGA scaffolds at 2, 3, and 5 weeks (**Figure 6**) (P < 0.05).

FIGURE 3 | Morphology of primary cultured DFAT cells and ASCs. (A) When the flasks had been inverted for 7 days, DFAT cells grew to form a colony. Some DFAT cells in the colony still have a lipid droplet (arrowheads). Scale bar = 500 µm. (B) Cultured ASCs are more spindle-shaped than DFAT cells. Scale bar = 500 µm.

# Transplantation of DFAT/PLGA Enhanced Periodontal Tissue Regeneration

By week 5, the created defect sites were easily identifiable by H&E in the PLGA-alone group because the remnant of the PLGA scaffold was clearly visible (**Figures 7A,D**, **8A,D**). At low magnification, formation of mineralized tissues such as bone and cementum was evident in the DFAT/PLGA and ASCs/DFAT groups at 5 weeks (**Figures 7B,C**), but not in the PLGA-alone group (**Figure 7A**). The newly formed bone structures within the defect were coalesced with surrounding native alveolar bone (**Figures 7B,C**, **8A–C**). The presence of osteoblasts lining the organic matrix and osteocytes trapped in the lacunae of newly formed bone structures was detected in the DFAT/PLGA and ASCs/PLGA groups (**Figures 7E,F**, **8E,F**). Vascularization was observed in all samples (**Figures 7G–I**, **8G–I**).

Periodontal ligament organization was recognized in the cementum-ligament-alveolar bone complex in the DFAT/PLGA and ASCs/PLGA groups (**Figures 7H,I**, **8G–I**). The orientation of the ligament fibers, visualized by Picrosirius-red staining, resembled the oblique fibers in the functional native periodontal ligament tissue (**Figures 7K,L**, **8K,L**). In the PLGA-alone group, sparse, and disorganized fibers were observed between dentin and scaffold because there was no cementum or nearby bone formation in the alveolar bone defect (**Figures 7J**, **8J**).

The results of the histometric analysis are summarized in **Figure 9**. The newly formed cementum on the central root in the DFAT/PLGA group was significantly thicker than that in the ASCs/PLGA and PLGA-alone groups (P < 0.05) (**Figure 9A**). Furthermore, the thickness of periodontal ligament regenerated in the tissue defects at the part of the central root in the DFAT/PLGA group was also significantly larger than in the

the cell number stained by Oil red O.

February 2016 | Volume 7 | Article 50 |

ASCs/PLGA and PLGA-alone groups (P < 0.05) (**Figure 9B**). Finally, the newly formed alveolar bone at the part of central root in the DFAT/PLGA group was significantly thicker than that in the ASCs/PLGA and PLGA-alone groups.

## Localization of Transplanted DFAT Cells in the Periodontal Tissue Defects

Fluorescent-labeled DFAT cells and ASCs were visualized in the newly formed alveolar bone and cementum, as well as in the newly formed periodontal tissues (**Figures 10A–H**).

# DISCUSSION

The aim of this study was to assess the periodontal regenerative potential of DFAT cells derived from rat mature adipocytes in combination with solid PLGA scaffolds transplanted into surgically created periodontal tissue defects (King et al., 1997; Akita et al., 2014). In addition, we compared the proliferation and differentiation potential in vitro and periodontal tissue-forming capability in vivo of DFAT cells and ASCs.

A successful periodontal tissue-engineering strategy requires a suitable scaffold that maintains space for cell growth and differentiation. A suitable scaffold for periodontal tissue regeneration should be biodegradable, so that it can eventually be replaced by regenerated tissue (Rungsiyanont et al., 2012). We previously demonstrated the in vivo performance of a PLGA scaffold seeded with ASCs (Akita et al., 2014). When a PLAGalone scaffold was transplanted into a periodontal defect in the present study, the defect space was occupied by connective tissue. In addition, newly formed connective tissue was observed

the area equivalent to (G–I), stained with Picrosirius-red to show Sharpey's fibers. Arrowheads indicate newly formed collagen fibers. Scale bar = 100 µm. NB, newly

around the remaining scaffold 5 weeks after transplantation and no inflammation was observed during the 5-week experimental period. These results suggest that the PLGA scaffold contribute to maintaining space for tissue regeneration in the periodontal defect because the structural integrity of the PLGA scaffold is higher than that of soft scaffolds such as those made from hydro gels and collagen sponge, which have been used in many other studies (Dogan et al., 2003; Kawaguchi et al., 2004; Zhao et al., ˘ 2004; Tobita et al., 2008; Li et al., 2009; Nuñez et al., 2012).

formed bone; Ce, newly formed cementum; De, root dentin.

Interestingly, based on ALP activity and mineralization activity in vitro, DFAT cells showed higher osteoblastic differentiation capacity than ASCs. Furthermore, DFAT cells showed greater potential for hard tissue formation in vivo than ASCs by micro-CT analysis. These results indicate that the osteogenic potential of DFAT cells is higher than that of ASCs, which is consistent with previous findings (Kishimoto et al., 2013). In addition, based on Oil red O and Alcian blue analysis, DFAT cells and ASCs had similar adipogenic and chondrogenic differentiation potential. Our group has studied DFAT cells in term of tissue-specific functions and genes that regulate of differentiation, such as those encoding SFRP2, PRRX1, HEY2, AEBP1, PEG10, PRRX2, RUNX1, FZD7, and IGFBP5. The results indicate DFAT cells have a multilineage differentiation capacity and a feature of progenitor cells committed to various cell lineages rather than the stem cell fate (Ono et al., 2011). Furthermore, our previous study showed that when a single cell-derived clonal population of DFAT cells expressed the specific genes of various cell lineages, it was capable of

periodontal ligament fibers. Arrowheads indicate newly formed collagen fibers. Scale bar = 100 µm. NB, newly formed bone; Ce, newly formed cementum; De, root

differentiating into multiple lineages (Kazama et al., 2008; Matsumoto et al., 2008). Taken together, these findings suggest that DFAT cells could have the ability to differentiate into cementoblasts, fibroblasts, and osteoblasts.

In our study, we observed in vivo periodontal tissue regeneration including cementum, periodontal ligament, and alveolar bone following both transplantation of DFAT cells and transplantation of ASCs. For the functional repair of the cementum-ligament-bone complex, fibrous connective tissues must insert into the cementum, and bone to achieve optimal biomechanical integration. In our study, the PLGA-alone group showed no cementum or alveolar bone formation, This is because the damaged tissue does not naturally repair owing to the lack of MSCs in periodontal tissues. In contrast, when DFAT cells or ASCs were transplanted into the periodontal tissue defect, newly formed cementum, and alveolar bone formation was observed in the periodontal fenestration defect. Interestingly, the regenerated cementum and bone volume in the DFAT/PLGA group was significant higher than in the ASCs/PLGA group. This is the first comparative study on periodontal tissue regeneration to show that rat DFAT cells have highly potential for cementum and alveolar bone formation than ASCs. However, the rat is not a reliable animal model because rats have a very high capability of regenerating bone. Therefore, we need to examine the periodontal tissue regeneration potential of DFAT cells in a large animal model before clinical use is attempted.

In contrast to our methods, Sugawara, and Sato used DFAT cells that were cultured in an osteogenic differentiation induction

dentin.

first molar, the cementum in the DFAT/PLGA group was significantly thicker than that in the ASCs/PLGA and PLGA-alone groups. In the distal root of the first molar, there were no significant differences between DFAT/PLGA and ASCs/PLGA. (B) In the central root of first molar, the periodontal ligament of the DFAT/PLGA group was significantly wider than that of the ASCs/PLGA and PLGA-alone groups. In the distal root of first molar, there were no significant differences between DFAT/PLGA and ASCs/PLGA groups. (C) In the central root and distal root of the first molar, the alveolar bone in the DFAT/PLGA group was significantly thicker than in the ASCs/PLGA and PLGA-alone groups (*n* = 12, \**P* < 0.05).

medium before transplantation. They observed a small amount of a cementum on the exposed root surface (Sugawara and Sato, 2014). In our study, we found thicker cementum and alveolar bone formation. The discrepancy could be because of difference in the number of transplanted cells, the scaffold, and the treatment before transplantation. We seeded 1 × 106 DFAT cells in the PLGA scaffold. In addition, DFAT cells in our study had not been induced to undergo differentiation into specialized cells in vitro. Further research is needed to clarify suitable cell number and suitable pre-transplantation treatment.

We specifically observed fibrous connective tissue in the newly formed cementum and alveolar bone in the DFAT cell group and the ASC group by using Picrosirius-red staining and Azan staining. Interestingly, the periodontal ligament space in the DFAT/PLGA was wider than that in the ASCs/PLGA. The results suggest that the transplanted DFAT cells participated in the repair of periodontal tissue via secretion of trophic factors or that they could directly participate into the various cell types constituting the periodontium (Chen et al., 2008; Gnecchi et al., 2008; Mooney and Vandenburgh, 2008; Chen and Jin, 2010). Our group has reported that DFAT cells secrete several cytokines associated with bone formation and angiogenesis (Kikuta et al., 2013). In addition, some GFP-positive DFAT cells clearly remained in the regenerated periodontal tissue area, as also reported in a previous study on periodontal tissue regeneration (Sugawara and Sato, 2014). Our analysis also revealed that the transplanted DFAT cells were detected at the injection site 5 weeks after transplantation. Both of these results suggest that at least some of the DFAT cells engrafted onto injected periodontal tissue and contributed to local periodontal tissue regeneration. Further research is required to determine specifically whether the labeled DFAT cells generated the differentiated cementoblasts, osteoblasts, and periodontal ligament fibroblasts. Previous studies have demonstrated that MSCs, at least in part, can differentiated into cementoblasts, osteoblasts, and fibroblasts in regenerated periodontal tissue (Hasegawa et al., 2006; Tobita et al., 2008).

In conclusion, we found a significantly greater increase in crucial tissues, cementum, and alveolar bone in the DFAT/PLGA group compared with the ASCs/PLGA group. The regenerated periodontium restored the architecture to an arrangement that was similar to the original architecture, collagen fibers inserted into the cementum, and alveolar bone layers. DFAT cells could be a promising cell source for periodontal tissue regeneration. However, the impact of the

#### REFERENCES

local differentiation microenvironment on DFAT cells and the mechanisms controlling the differentiation into cementoblasts, periodontal ligament fibroblasts, and osteoblasts remain to be examined further.

## AUTHOR CONTRIBUTIONS

DA: Performed most of work, in particular in-vivo study. KK: Developed the technique to isolate DFAT cells. YS: Supported the experiment performed by DA. T. Mashimo: Performed in vivo experiments. N. Tsurumachi: Isolated DFAT cells from rat. KY: Developed the scaffolds in our study. TK: Developed the scaffold. TT: Performed Cell culture and differentiation assay. N. Tsukimura: Made the experimental design. T. Matsumoto: Established the differentiation assay and cell culture technique. TI: Prepared the manuscript. KI: Managed the overall experimental design. MH: Controlled the overall paper until completion.

# FUNDING

This work was supported in part by Grant-in-Aid for Scientific Research (B and Houga) (21390528, 15H05037, and 15K15724 to MH, and 15H04607 to KK), research grants from the Promotion Project of Medical Clustering of Okinawa Prefecture (KK), Grants from the Dental Research Center, the Nihon University School of Dentistry (MH), Nihon University Multidisciplinary Research Grant for 2012 and 2013 (MH), and the Nihon University Graduate School of Dentistry research fund (10010105 to DA). This work was also supported by the MEXT-Supported Program for the Strategic Research Foundation at Private Universities (S1411018).

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2016 Akita, Kano, Saito-Tamura, Mashimo, Sato-Shionome, Tsurumachi, Yamanaka, Kaneko, Toriumi, Arai, Tsukimura, Matsumoto, Ishigami, Isokawa and Honda. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Changing Paradigms in Cranio-Facial Regeneration: Current and New Strategies for the Activation of Endogenous Stem Cells

Luigi Mele1 , Pietro Paolo Vitiello2 , Virginia Tirino1 , Francesca Paino1 , Alfredo De Rosa3 , Davide Liccardo1 , Gianpaolo Papaccio1 \* and Vincenzo Desiderio1

1 Department of Experimental Medicine, Section of Biotechnology and Medical Histology and Embryology, Second University of Naples, Naples, Italy, 2 Medical Oncology, Dipartimento Medico-Chirurgico di Internistica Clinica e Sperimentale "F. Magrassi e A. Lanzara," Second University of Naples, Naples, Italy, 3 Department of Odontology and Surgery, Second University of Naples, Naples, Italy

#### Edited by:

Thimios Mitsiadis, University of Zurich, Switzerland

#### Reviewed by:

Ariane Berdal, UMRS 1138 Institut National de la Santé et de la Recherche Médicale, Université Paris Diderot - Paris 7, France Catherine Chaussain, Université Paris Descartes, France Michel Goldberg, UMR-S 1124, Institut National de la Santé et de la Recherche Médicale and University Paris Descartes, France

> \*Correspondence: Papaccio Gianpaolo<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Craniofacial Biology, a section of the journal Frontiers in Physiology

Received: 04 January 2016 Accepted: 09 February 2016 Published: 24 February 2016

#### Citation:

Mele L, Vitiello PP, Tirino V, Paino F, De Rosa A, Liccardo D, Papaccio G and Desiderio V (2016) Changing Paradigms in Cranio-Facial Regeneration: Current and New Strategies for the Activation of Endogenous Stem Cells. Front. Physiol. 7:62. doi: 10.3389/fphys.2016.00062 Craniofacial area represent a unique district of human body characterized by a very high complexity of tissues, innervation and vascularization, and being deputed to many fundamental function such as eating, speech, expression of emotions, delivery of sensations such as taste, sight, and earing. For this reasons, tissue loss in this area following trauma or for example oncologic resection, have a tremendous impact on patients' quality of life. In the last 20 years regenerative medicine has emerged as one of the most promising approach to solve problem related to trauma, tissue loss, organ failure etc. One of the most powerful tools to be used for tissue regeneration is represented by stem cells, which have been successfully implanted in different tissue/organs with exciting results. Nevertheless, both autologous and allogeneic stem cell transplantation raise many practical and ethical concerns that make this approach very difficult to apply in clinical practice. For this reason different cell free approaches have been developed aiming to the mobilization, recruitment, and activation of endogenous stem cells into the injury site avoiding exogenous cells implant but instead stimulating patients' own stem cells to repair the lesion. To this aim many strategies have been used including functionalized bioscaffold, controlled release of stem cell chemoattractants, growth factors, BMPs, Platelet–Rich-Plasma, and other new strategies such as ultrasound wave and laser are just being proposed. Here we review all the current and new strategies used for activation and mobilization of endogenous stem cells in the regeneration of craniofacial tissue.

Keywords: craniofacial abnormalities, regenerative medicine, stem cell transplantation, stem cells and regenerative medicine, stem cells recruitment, SDF1, bioscaffold, BMP signaling

# INTRODUCTION

Regenerative medicine is the field of translational research that aims to replace and repair cells, tissues and organs to restore their normal functions (Mason and Dunnill, 2008). In the last 20 years, the possibility to use endogenous and exogenous stem cells for tissue repair has emerged producing enthusiasm in the scientific and medical community, but on the other side, raising a series of ethical and practical issues. The craniomaxillofacial complex is of fundamental importance for many different functions, such as breathing and eating, others non-vital but still important for social relationships, such as aesthetics and the delivery of senses such as sight, smell, and sound. For these reasons, craniomaxillofacial tissue damages have serious physiological and psychological consequences, and it is of paramount importance to seek solutions to optimize the life of people with maxillofacial trauma (Teo and Vallier, 2010). Many disciplines, such as biology, medicine, chemistry, and engineering, are involved in the study and development of techniques or products that regenerate the native conditions of such a complicated body region as the craniofacial tissues such as bone, muscle, cartilage and nervous tissue. The research moved toward two main different directions: one that involves the study of stem cell biology, including the development of novel techniques to isolate, characterize and transplant stem cells, the other related to the material science, which aims to the creation of new biomaterials and technologies suitable for the use with stem cells. As for stem cell biology, the research focused on stem and progenitor cells isolation, characterization, expansion, and transplantation for replacing damaged tissues (Giuliani et al., 2013) and on the study of growth-factors and genetic technology to control proliferation, migration and ability in tissue repair and regeneration (Alvarez et al., 2012; Han et al., 2014).