Micron 67 (2014) 155–168

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Micron journal homepage: www.elsevier.com/locate/micron

Plasticity of human dental pulp stromal cells with bioengineering platforms: A versatile tool for regenerative medicine Serena Barachini a,∗ , Serena Danti b , Simone Pacini a , Delfo D’Alessandro b , Vittoria Carnicelli b , Luisa Trombi a , Stefania Moscato a , Claudio Mannari a , Silvia Cei b , Mario Petrini a a b

Department of Clinical and Experimental Medicine, University of Pisa, Italy Department of Surgical, Medical, Molecular Pathology and Emergency Medicine, University of Pisa, Italy

a r t i c l e

i n f o

Article history: Received 24 March 2014 Received in revised form 15 July 2014 Accepted 20 July 2014 Available online 27 July 2014 Keywords: Stem cells Dental pulp Differentiation Bioengineering Regenerative medicine

a b s t r a c t In recent years, human dental pulp stromal cells (DPSCs) have received growing attention due to their characteristics in common with other mesenchymal stem cells, in addition to the ease with which they can be harvested. In this study, we demonstrated that the isolation of DPSCs from third molar teeth of healthy individuals allowed the recovery of dental mesenchymal stem cells that showed self-renewal and multipotent differentiation capability. DPSCs resulted positive for CD73, CD90, CD105, STRO-1, negative for CD34, CD45, CD14 and were able to differentiate into osteogenic and chondrogenic cells. We also assayed the angiogenic potential of DPSCs, their capillary tube-like formation was assessed using an in vitro angiogenesis assay and the uptake of acetylated low-density lipoprotein was measured as a marker of endothelial function. Based on these results, DPSCs were capable of differentiating into cells with phenotypic and functional features of endothelial cells. Furthermore, this study investigated the growth and differentiation of human DPSCs under a variety of bioengineering platforms, such as low frequency ultrasounds, tissue engineering and nanomaterials. DPSCs showed an enhanced chondrogenic differentiation under ultrasound application. Moreover, DPSCs were tested on different scaffolds, poly(vinyl alcohol)/gelatin (PVA/G) sponges and human plasma clots. We showed that both PVA/G and human plasma clot are suitable scaffolds for adhesion, growth and differentiation of DPSCs toward osteoblastic lineages. Finally, we evaluated the interactions of DPSCs with a novel class of nanomaterials, namely boron nitride nanotubes (BNNTs). From our investigation, DPSCs have appeared as a highly versatile cellular tool to be employed in regenerative medicine. © 2014 Elsevier Ltd. All rights reserved.

1. Introduction

Abbreviations: DPSCs, dental pulp stromal cells; MSC, mesenchymal stromal cells; SHEDs, stromal cells from human exfoliated deciduous; BM-MSCs, bone marrow-derived mesenchymal stromal cells; NTFs, neurotrophic factors; NGF, nerve growth factor; BDNF, brain-derived neurotrophic factor; PPP, platelet poor plasma; US, ultrasounds; LIUS, low intensity US; LFUS, low frequency US; PVA, poly(vinyl alcohol); G, gelatin; BNNTs, boron nitride nanotubes; ␣-MEM, minimum essential medium alpha modification; FBS, fetal bovine serum; PDT, population doubling time; CPs, chondrogenic pellets; HUVECs, human umbilical vein endothelial cells; SEM, scanning electron microscopy; TEM, transmission electron microscopy. ∗ Corresponding author at: Hematology Division, Department of Clinical and Experimental Medicine, University of Pisa, Via Roma 56, 56100 Pisa, Italy. Tel.: +39 050 993484; fax: +39 050830162. E-mail address: [email protected] (S. Barachini). http://dx.doi.org/10.1016/j.micron.2014.07.003 0968-4328/© 2014 Elsevier Ltd. All rights reserved.

Among mesenchymal stromal cells (MSCs) of autologous origin, human dental pulp stromal cells (DPSCs) have received growing attention in recent years firstly due to the easy accessibility of the tissue in any adult individual. Dental pulp tissue is thought to be derived from migratory neural crest during development (Peters and Balling, 1999). Dental pulp is a soft connective tissue and its main functions are to produce dentin, and to maintain the biological and physiological vitality of the dentin. DPSCs are multipotent stromal cells derived from neural crest and mesenchyme and have the capacity to differentiate into multiple cell lineages. Firstly, Gronthos et al. (2000, 2002) have isolated post-natal stromal cells from the human dental pulp of permanent teeth. Other stromal cell populations from surrounding tissues of the tooth have been isolated from periodontal ligament, human exfoliated deciduous teeth, apical papilla and

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dental follicle precursor cells (Miura et al., 2003; Seo et al., 2004; Sonoyama et al., 2006). These post-natal populations have MSC-like features, namely the capacity for self-renewal and the potential to differentiate into multiple lineages including osteoblasts and chondroblasts (Huang et al., 2009). However, DPSCs show higher self-renewal ability, immunomodulatory capacity and proliferation in vitro than bone marrow-mesenchymal stromal cells (BM-MSCs); furthermore, they preferentially differentiate to osteoblasts rather than into adipocytes (Gronthos et al., 2000; Pierdomenico et al., 2005). Although the majority of studies have focused their attention on the ability of DPSCs to differentiate into odontoblast-like cells (Almushayt et al., 2006; Cordeiro et al., 2008; Paula-Silva et al., 2009) or osteoblasts (Laino et al., 2005, 2006; D’Aquino et al., 2007), it has also been discovered that they are capable of differentiating into other cell types, including smooth muscle cells (Kerkis et al., 2006; Gandia et al., 2008) and neurons (Arthur et al., 2008; Kadar et al., 2009). DPSCs express nestin and glial fibrillary acidic protein and, under appropriate stimuli, are capable of differentiating into functionally active neurons (Arthur et al., 2008), influencing endogenous recruitment of neural stem cells and generating neurospheres (Sasaki et al., 2008). Neural stem cell markers, such as nestin, expressed in DPSCs reflect the neural origin of dental pulp (Kerkis et al., 2006; Estrela et al., 2011). Recently, we have investigated specific molecular profiles of human stromal stem cell populations derivated from different tissues, particularly with regard to the global HOX gene family expression profile (HOX code) and their three amino acid loop extension co-factor subfamilies (Picchi et al., 2013). The different levels of HOX expression detected in stromal cells with different potency strongly suggest that HOX genes may not only reflect positional and embryological cell identity, but also indicate the cellular position within the stem cell hierarchy. Such considerations support the growing evidence that HOX code provides a “biological fingerprint” to distinguish stem cell populations (Chang et al., 2002; Moens and Selleri, 2006). Our previous study shows that DPSCs exhibit extremely low levels of expression of a few HOX genes, confirming previous findings, and in line with the neuroectodermal origin of DPSCs (Couly et al., 2002; D’Antò et al., 2006). It is known that neural crests are HOX negative and indeed the few active HOX genes in DPSCs are expressed at barely detectable levels. MSCs appear to exert paracrine trophic effects through the secretion of bioactive molecules (Caplan and Dennis, 2006; Caplan, 2007) as the neurotrophic factors (NTFs). In particular, brainderived neurotrophic factor (BDNF) and nerve growth factor (NGF) produced by DPSCs, have been shown to have a crucial influence over neurons in the central nervous system such as motor neurons and dopaminergic neurons of the substantia nigra (Nosrat et al., 2001, 2004). In our previous study, we found that high levels of BDNF, NGF transcripts were constitutively expressed by DPSCs and that the neuroprotective effect of DPSCs against two neurotoxins on an in vitro model of Parkinson’s could be due to soluble factors, such as BDNF and NGF, released by DPSCs (Nesti et al., 2010). DPSCs may thus be an alternative source in cell therapy for neurological diseases. Recent highlights have started considering stem cells not only as the final therapeutic product, but rather as part of complex bioengineered therapeutic strategies. In this view, MSCs can be fully appreciated as a versatile tool in combination with biomaterials and biomedical devices to guide their commitment in various human body tissues (Kshitiz et al., 2012; Kinney and McDevitt, 2013). Therefore, stem cell plasticity thus represents a master feature. The potential use of stem cells in bioengineering-based therapies includes tissue engineering and nanotechnology. As a proof of concept, we tested the biological response of DPSCs in three different bioengineering platforms. As an example of biomedical device-assisted platforms for tissue regeneration, ultrasounds (US)

represent a non-invasive and versatile tool that has been shown to promote tissue repair, such as cartilage healing in animal models of articular cartilaginous defects (Cook et al., 2001). Studies on the underlying mechanism leading to cell response to US stimuli are ongoing, however preliminary evidence in rat chondrocyte cultures have highlighted that US can enhance the synthesis of proteoglycans depending on their application cycles and intensity (Parvizi et al., 1999). In line with these observations, recent reports have pointed out that low intensity US (LIUS) can exert a favorable effect on the proliferation, extracellular matrix synthesis and chondrogenic differentiation of MSCs in vitro (Lee et al., 2006; Shah et al., 2013). Differently from LIUS, used for diagnostic purposes, low frequency US (LFUS) (range 20–100 kHz) is used in sonophoresis and dentistry since they transfer higher mechanical energy. In our study we tested the efficacy of LFUS on DPSC pellets under chondrogenic differentiation with respect to traditional pellet culture. Moreover, we investigated DPSCs for tissue engineering applications, using biocompatible three-dimensional spongy materials to assess their differentiation capability. Poly(vinyl alcohol) (PVA) has been among one of first synthetic macromolecules employed in both implantable and non-implantable medical devices (e.g., contact lenses, artificial meniscus) due to its characteristics including low protein adsorption, biocompatibility, high hydrophilicity, easy processability and chemical inertia (Alves et al., 2011). Additionally, PVA can be easily added to biologic molecules to obtain bioartificial matrices with specific architectural features (Cascone et al., 2004). Among which, gelatin (G) is a natural protein derived from collagen able to promote cell adhesion (Dubruel et al., 2007). Spongy scaffolds based on PVA and G have been successfully tested with gingival fibroblasts, being reported as very promising substrates for tissue engineering (Moscato et al., 2008). In the present study, PVA/G sponges were prepared and investigated in vitro for threedimensional growth and osteogenic differentiation of DPSCs. Finally, nanomedicine is emerging as a powerful source of innovative cell-targeted-therapies based on nanoengineered materials, whose toxicological risks have yet to be disclosed. The balance of these features has thus become an intriguing challenge for successful development of therapeutically effective nanosystems. In particular, due to their small size, nanoparticles can enter the cell membrane and be used as non-viral vectors. Boron nitride nanotubes (BNNTs) are a class of ceramic nanoparticles with a tubular shape that have attracted recent attention due to their excellent physical properties and putative biocompatibility (Ciofani and Danti, 2012). In this study, we report our preliminary findings on the interactions of BNNTs with DPSCs, aimed at supporting the future disclosement of novel nanomedicine-based therapies. In summary, the specific aims of the current study are to improve understanding of the biological properties of human DPSCs. Moreover, we have investigated DPSC growth and differentiation in different bioengineering applications, evaluating their multipotent potential in regenerative medicine.

2. Materials and methods 2.1. Extraction and isolation of human DPSCs Human dental pulps were obtained from molars of healthy subjects (n = 10) 18–35 years of age, after informed consent (Oral Surgery Department, Santa Chiara Hospital, Pisa, Italy) according to a protocol approved by the local University committee on Ethics in Medicine. Each subject, before extraction, was checked for systemic and oral diseases and pre-treated a week before with professional dental hygiene. Before extraction, the dental crown was rinsed with a 0.2% chlorexidin gel (Dentosan, Johnson & Johnson Medical S.p.A., Rome, Italy) for 2 min. Radicular dental pulps were obtained, using

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a Gracey curette, from healthy and non-carious teeth. The samples were placed inside sterile tube containing 20 ml of proliferation medium, transported to the laboratory and immediately processed. Proliferation medium consisted of minimum essential medium alpha modification (␣-MEM; Sigma–Aldrich, St. Louis, MO, USA) and 10% fetal bovine serum (FBS; Sigma–Aldrich) supplemented with 100 IU/ml penicillin (Pharmacia & Upjohn S.p.A., Milan, Italy), 100 IU/ml streptomycin (Bristol-Myers Squibb S.p.A., Sermoneta, Italy) and 2 mM l-glutamine (Lonza, Wolkersville, MD, USA). Pulp tissue explants were placed in tissue flasks with the proliferation medium and were cultured at 37 ◦ C in 5% CO2 concentration at 95% humidity. Medium changes were carried out twice a week. On the confluence, the cells were detached with 0.05% trypsin–0.02% ethylenediaminetetraacetic acid (Life Technologies, Carlsbad, CA, USA) for further expansions. Cell growth was analyzed after the first passage (P1) by direct cell counts to determine the cumulative population doublings (PDs) and the population doubling time (PDT). PD number was calculated using the formula log10 (N)/log10 (2), where N = cells harvested/cells seeded ratio (Kern et al., 2006). Results were expressed as cumulative PDs. PDTs were calculated dividing total culture time in hours (h) by cumulative PDS. 2.2. Characterization of DPSCs 2.2.1. Immunophenotypic analysis The phenotypical analysis was performed on cells detached from passages 2 through 6, after incubation of the DPSCs with monoclonal antibodies (mAbs). A total of 100 ␮l of cell suspension (5 × 105 cells) was aliquoted per tube and appropriately labeled mAbs were added for multicolor analysis and incubated for 30 min at 4 ◦ C; then, samples were washed twice in MACSQuant running buffer (Miltenyi Biotech, Bergisch Gladbach, Germany). The flow cytometer instrument was set using cells stained with isotypic controls. Cells were gated on a forward versus side scatter plot to eliminate debris. Acquisition was performed using 10,000 cells that were analyzed with an MACSQuant cytofluorimeter running the MACSQuantify software (Miltenyi Biotech). DPSCs were stained using mAbs specific for CD105 PE-Cy7-conjugated (BioLegend, San Diego, CA, USA), CD73 PE-Cy7-conjugated (Miltenyi Biotech), CD90 PerCP-Cy5.5 (BioLegend), CD31 PE-conjugated (Miltenyi Biotech), CD146 PE-conjugated (BioLegend), CD309 PE-conjugated (BioLegend), SSEA-4 AlexaFluor® 488 (BioLegend), STRO-1 AlexaFluor® 647 (BioLegend), CD45 APC-Vio770 (Miltenyi Biotech) and CD34 VioBlue (Miltenyi Biotech). If not indicated, further experiments were performed in triplicate. 2.2.2. Neurotrophin quantification In order to evaluate BDNF and NGF levels in the medium of the DPSC cultures, the whole supernatant medium from three distinct experiments was collected weekly in duplicate. After centrifugation to remove particulates, 2 ml aliquots were stored at −20 ◦ C. After thawing at room temperature, 0.5 ml were collected from the aliquots and analyzed with an enzyme-linked immunosorbent assay (ELISA). NGF was detected with an ELISA kit (Koma Biotech, Korea). Briefly, NGF pre-coated 96-well plate was incubated with cell supernatants and NGF standards for 2 h. After, it was washed and incubated with a biotinylated rabbit anti-human NGF polyclonal antibody, washed again, and then incubated with avidin conjugated to horseradish peroxidase. BDNF was detected with an ELISA kit (Promega, USA). According to manufacturing recommendations, 96-well plate was coated with anti-BDNF mAb and blocked with the blocking buffer provided, then samples and standards were added. After sample incubation and washing, plate was incubated with anti-human BDNF polyclonal antibody, washed, and finally incubated with anti-IgG antibody conjugated to horseradish

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peroxidase. For both neurotrophins, the plates were incubated with a tetramethylbenzidine (TMB) solution and absorbance measured at 450 nm using a microplate reader (Model 550, BioRad Laboratories, Hercules, CA, USA). 2.3. Differentiation of DPSCs 2.3.1. Endothelial differentiation When 80–90% of confluence, DPSCs were detached by trypsin digestion, re-plated at a density of 1 × 104 cells/cm2 on 5 ␮g/cm2 fibronectin BioCoatTM (Becton Dickinson, San Jose, CA, USA) in 6-well plates. The cells were cultured in EGM-2 media (Lonza, Wolkersville, MD, USA) for 15 days, replacing the medium twice a week. An undifferentiated control group was cultured in proliferation medium. To evaluate endothelial differentiation, cells were processed for immunophenotype analysis, as described in the previous section, and PECAM1 (CD31) gene expression, as described later. Assessment of in vitro capillary tube-like formation was carried out using Matrigel (Becton Dickinson Bioscience, San Jose, CA, USA): an in vitro assay that shows the angiogenetic potential of cells. The Matrigel was thawed overnight at 4 ◦ C and mixed to homogeneity using cooled pipette tips. Aliquots of Matrigel (50 ␮l) were distributed as a thin layer on the bottom of a 12-well cell culture plates and left for polymerization at 37 ◦ C for 30 min. Differentiated DPSCs were detached and resuspended in EGM-2 (Lonza) to give a final concentration of 5 × 105 cells/ml, plated onto the Matrigel-coated surface and incubated for 1 h in a 37 ◦ C humidified incubator. Undifferentiated cells were cultured in proliferation medium. Tube-like structure formation was examined via light microscopy and imaged after 12 h. The uptake of Ac-LDL was investigated as an endothelial marker. It was assessed incubating cell samples for 4 h at 37 ◦ C with 5 ␮g/ml AlexaFluor-488® -conjugated Ac-LDL (Life Technologies). Finally, specimens were observed under fluorescence microscopy. 2.3.2. Osteogenic differentiation To promote osteogenic differentiation, the cells were seeded at a density of 4 × 103 cells/cm2 in 6-well plates (CoStar Group, Bethesda, MD, USA) and cultured in basal medium, ␣-MEM supplemented with 10% FBS, antibiotics and l-glutamine, until they reached 60–70% confluence. As soon as subconfluence was reached, osteogenic differentiation of cells was induced culturing them for 2.5 weeks with osteogenic induction medium, consisting of 100 ␮M dexamethasone (Sigma–Aldrich), 10 mM ␤glycerophosphate (Sigma–Aldrich), 100 ␮M ascorbic acid (Vitamin C, Roche, Indianapolis, IN, USA) and 10% FBS in ␣-MEM. Medium was changed twice a week. Cells kept in basal medium were used as negative controls. Osteogenic differentiation was investigated using the von Kossa and Alizarin S (Sigma–Aldrich) staining, to demonstrate the deposition of a mineralized matrix. The cells were fixed with 1% formalin (Bio-Optica) for 10 min at 4 ◦ C and stained for 15 min with 1% silver nitrate (Fluka, Millwaukee, WI, USA, and Sigma–Aldrich). The stain was developed incubating the cells in 0.5% pyrogallol (Fluka) and then fixed with 5% sodium thiosulfate (Fluka) for 5 min. Finally, the cells were counterstained with 0.1% nuclear fast red (Fluka). The samples were then dehydrated and mounted with DPX mountant (Fluka). Mineral deposition was evaluated as black granules using optical light microscopy. 2.3.3. Chondrogenic differentiation under US stimulation DPSCs at passage 2 were trypsinized, counted and placed in 5 ml tubes at a density of 250,000 cells/tube, centrifuged at 1200 rpm for 7 min to obtain chondrogenic pellets (CPs), which were washed in sterile PBS. Chondrogenic culture medium was thus added, consisting of DMEM/F12, 1.25 ␮g/ml bovine serum albumin (BSA), 5.35 ␮g/ml linoleic acid, 50 ␮g/ml ascorbic acid, 100 ␮g/ml sodium

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Table 1 Primer pairs designed for detection of PECAM1 and ACTB mRNA. NCBI ref#

Gene name

Primer pair

Amplicon length (bp)

NM 000442

PECAM1

231

NM 001101.3

ACTB

Forward GAACCTGTCCTGCTCCATC Reverse TCAAACTGGGCATCATAAGAAAT Forward CGCCGCCAGCTCACCATG Reverse CACGATGGAGGGGAAGACGG

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pyruvate, insulin–transferrin–selenium (ITS premix), 10−7 M dexamethasone and 10 ng/ml transforming growth factor beta 1 (TGF-␤1; PeproTech, Rocky Hill, NJ). If not otherwise specified, all reagents were purchased from Sigma–Aldrich. CPs were cultured in standard cell culture conditions (i.e., 5/95% CO2 /air, >90% humidity). One set of CPs was daily treated (3 times for 5 s) with LFUS (at 40 kHz frequency and 20 W output power) using a sonicator bath (Bransonic 2510; Bransonic, Danbury, CT, USA). After 3 weeks, samples were processed for histologic procedures. 2.3.4. Quantitative RT-PCR DPSCs from three independent samples were cultured in proliferation or in EGM-2 medium for 15 days. Cells were then harvested by trypsin–EDTA digestion and processed for gene expression analysis. Total RNA was extracted using RNeasy Mini Kit (Qiagen GmbH, Hilden, Germany), as indicated by manufacturer’s protocol. DNaseI digestion was performed on column. One microgram of each RNA sample was retrotranscripted with QuantiTect Reverse Transcription Kit (Qiagen GmbH) and two-fold dilutions of cDNAs were analyzed by quantitative RT-PCR on iCycler-iQ5 Optical System (BioRad Laboratories, Hercules, CA), using SsoFast EvaGreen SuperMix (BioRad). All samples were run in duplicate. Primers from PECAM1 and ACTB genes (Table 1) were designed from coding sequences published on GenBank database with the help of Beacon Designer v.7 Software (Premier Biosoft International, Palo Alto, CA, USA). Relative quantitative analysis was performed following 2−Ct Livak’s method (Livak and Schmittgen, 2001). Normalization was made with ACTB housekeeping gene. 2.4. DPSC interaction with different biomaterial platforms 2.4.1. DPSCs cultured inside human plasma clots Plasmas for the clot preparation were obtained from peripheral blood of healthy adult donors. The blood was collected into tubes containing EDTA as anticoagulant and the platelet poor plasma (PPP) was obtained by centrifugation (1700 × g for 20 min). The DPSCs were resuspended in PPP at three different densities, 0.25 × 105 , 0.5 × 105 and 1.0 × 105 cells/sample, and seeded in 6well plates (CoStar Group). To produce DPSC/clot constructs, CaCl2 (7 mM; Dade Berhing, Marburg, Germany) was added to each well (Trombi et al., 2008; Barachini et al., 2009). After 30 min of incubation at 37 ◦ C in 5% humidified CO2 atmosphere, constructs were harvested and cultured for 7 days in osteogenic medium or proliferation medium as control. After 7 days the constructs were fixed in 4% neutral buffered formalin and processed for histological procedures. As an osteoblastic marker, calcium deposition was tested on paraffin sections by means of von Kossa staining. Cell viability after 48 h, was assessed using alamarBlue® (Life Technologies) assay. 2.4.2. DPSCs cultured on PVA/G scaffolds PVA/G scaffolds (weight composition ratio of 80/20) were obtained via emulsion and freeze-drying (Moscato et al., 2008). Briefly, an aqueous solution of PVA (Mw = 85,000–124,000, from

Sigma–Aldrich) and G (gelatin type B from bovine skin, 75 bloom, from Sigma–Aldrich) was obtained at 50 ◦ C under stirring and further added with sodium lauryl sulfate (SLS, from Sigma–Aldrich) to obtain a dense foam that was quenched in liquid nitrogen and lyophilized. Dried foams were stabilized by crosslinking with glutaraldehyde (GTA; grade II, from Sigma–Aldrich) vapors, cut into cylinders (5 mm diameter, 1.5 mm thickness) and subsequently treated with glycine (Sigma–Aldrich) solution to block GTA unreacted binding sites. The resulting sponge is a highly porous, biocompatible, hydrophilic and bio-stable material, suitable for tissue engineering applications. The scaffolds were sterilized with absolute ethanol, washed with sterile 2× PEN-Strep (Sigma–Aldrich)/Diflucan (Pfizer Italia, Latina, Italy) saline and finally rinsed with phosphate buffered saline (PBS, Sigma–Aldrich) prior to cell seeding. Twice passaged DPSCs were seeded on PVA/G scaffolds at a density of 500,000 cells/sample. After 24 h, sets of cell/scaffold constructs were committed either to the osteogenic or to the chondrogenic lineage replacing the proliferation culture medium with appropriate differentiative media. Differentiations were performed for 21 days using osteogenic and chondrogenic culture medium, as reported in the previous sections. Undifferentiated controls of DPSC/scaffold constructs were also carried out culturing the specimens in proliferation medium until the endpoint. Along the culture time, viability of the constructs was assayed, while at the endpoint samples were processed for qualitative (scanning electron microscopy, histology) and quantitative (viability test, DNA quantification, glycosaminoglycan content) analyses. 2.4.3. DPSCs cultured with BNNTs BNNTs, supplied by the Australian National University, Canberra, Australia, were produced by a ball-milling and annealing method (Yu et al., 2005). Details of sample purity and composition, as provided by the supplier, comprise yield > 80%, boron nitride > 97 wt%, residual metal catalysts (Fe and Cr) ∼1.5 wt% and absorbed O2 ∼1.5 wt%. Stable aqueous dispersions of BNNTs were obtained using 0.1% poly-l-lysine (PLL, Mw 70,000–150,000; Fluka, Buchs, Switzerland) as described in a previous study (Ciofani and Danti, 2012). Excess PLL was removed by repeated ultracentrifugation (Allegra 64R; Beckman Coulter, Fullerton, CA, USA). Stable PLL-BNNT dispersions were thus obtained by the non-covalent coating of the nanotube walls with PLL. The final concentration of BNNTs in the dispersion was quantified using a UV/Vis/NIR spectrophotometer (LIBRA S12, Biochrom, Cambridge, UK), while the residual PLL concentration was assessed using the bicinchoninic acid (BCA) method, as previously reported (Ciofani and Danti, 2012). For cytotoxicological investigations, DPSCs were seeded on 96-well plates (20,000 cells/well) in proliferation medium. After 24 h, the medium was replaced with a modified medium, containing the following PLL-BNNT concentrations: 0, 5, 10 and 15 ␮g/ml. Cultures were performed for 72 h to monitor cell viability. To evaluate cellular uptake of the nanotubes, the DPSCs were seeded on T75 flasks (800,000 cells/flask) and, after 24 h, they were incubated overnight with medium containing PLL-BNNTs at 10 ␮g/ml to be subsequently treated for transmission electron microscopy (TEM; Zeiss 902 microscope, Zeiss, Oberkochen, Germany). 2.4.4. DPSC viability Viability of DPSCs cultured with different biomaterial platforms was investigated using the alamarBlue® assay. Data were acquired according to manufacturer’s instructions, and expressed ad percentage of reduced alamarBlue® (%ABred). This bioassay incorporates a REDOX indicator resulting in color change of the culture medium according to cell metabolism and can be performed multiple times on the same samples due to its negligible toxicity. Samples (n = 3 and n = 6, for cell/scaffold construct and PLL-BNNT experiments, respectively) and controls, including blank

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controls (i.e., scaffolds with no cells, and empty well plates for cell/scaffold construct and PLL-BNNT experiments, respectively), were incubated for 4 h with the dye according to the manufacturer’s recommendations. Samples were assayed at different culture timepoints: 1, 11, 21 days to assess cell/scaffold construct viability, and 24, 48, 72 h to assess PLL-BNNT cytotoxicity. After each assay, supernatants were removed from the cultures and replaced with fresh culture medium. Samples were analyzed with a spectrophotometer (BioRad Laboratories, Hercules, CA, USA) under a double wavelength reading (570 nm and 600 nm). Finally, dye reduction percentage was calculated using dye molar extinction coefficients and appropriate absorbance equations as provided by the manufacturer. 2.4.5. Double stranded (ds)-DNA and glycosaminoglycan (GAG) contents in PVA/G constructs At the endpoint (21 days), un-, osteo-, and chondro-differentiated cell/scaffold constructs were either fixed for morphological and histologic evaluation, or stored in enzymatic solution at −80 ◦ C for quantitative assays. Digestive enzymes consisted of proteinase K, pepstatin A and iodoacetamide (Sigma–Aldrich) solution in phosphate-buffered EDTA (Park et al., 2005). Cell lysates were obtained using a freeze/thaw/vortex treatment of the samples. Contents of double ds-DNA and GAGs could be thus quantified in cascade on the same specimens (n = 3). ds-DNA content in cell lysates was measured using the PicoGreen kit (Molecular Probes, Eugene, OR, USA), as previously reported (Ciofani and Danti, 2012). Briefly, working buffer and PicoGreen dye solution were prepared according to the manufacturer’s instructions using reagents provided within the kit. After 10 min incubation in the dark at room temperature, the fluorescence intensity of the samples was measured on a plate reader (Victor3; PerkinElmer, Waltham, MA, USA), using an excitation wavelength of 485 nm and an emission wavelength of 535 nm. To determine GAG content, the constructs underwent digestion in a 60 ◦ C water bath for 16 h, then the dimethylmethylene blue dye (DMMB) assay was performed (Park et al., 2005). Absorbance was measured at 520 nm on a plate spectrophotometer (BioRad Laboratories). GAG contents were finally reported per microgram of ds-DNA, as obtained from the PicoGreen assay. 2.4.6. Histologic analysis Three-dimensional cultures of DPSCs (i.e., osteo-differentiated in plasma clots, chondro-differentiated in pellets, and un-, osteo, and chondro-differentiated on PVA/G scaffolds) were fixed in 4% neutral buffered formalin diluted in 1× PBS (0.1 M, pH 7.2) overnight at 4 ◦ C, then washed in 1× PBS and stored in 70% ethanol. Whole chondrogenic pellets, osteogenic plasma clots and a parts of cell/scaffold constructs were processed for histologic procedures. Briefly, the samples were dehydrated with a graded series of ethanol aqueous solutions up to absolute ethanol (the latter for 6 h), clarified twice in xylene for 2 h, rinsed in liquid paraffin for 4 h at 60 ◦ C and finally paraffin-embedded. Sections were deparaffined in xylene and rehydrated in ethanol before being stained or immunostained. Prior to observation, the sections were rehydrated in ethanol, clarified in xylene and mounted in DPX medium (Fluka, Buchs, Switzerland). Toluidine blue staining reveals sulfated GAGs in violet color and was performed on chondrogenic pellets. Sections were air dried and treated with 0.2% Toluidine blue (Sigma–Aldrich) for 10 s, washed and air dried. Alcian Blue staining detects generic and sulfated GAGs in cyan color (at pH 2.5 and pH 1, respectively) and was performed on chondrogenic pellets. Sections were incubated in Alcian Blue solutions kit (Bio-Optica, Milan, Italy) according to manufacturer’s instructions and counterstained with 0.1% nuclear fast red (Fluka). Von Kossa staining highlights carbonate and phosphate crystals and was performed on

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DPSCs osteo-induced on 6-well plates and on fibrin clots. Sections were treated with 1% silver nitrate (Fluka) for 15 min at artificial light, soaked in 0.5% pyrogallol (Fluka) for 2 min treated with 5% sodium thiosulfate (Sigma–Aldrich) for 2 min and counterstained with nuclear fast red. Immunohistochemical analysis was carried out on CPs and cell/scaffold constructs. Samples were permeabilized with 0.2% Triton X-100 (Sigma–Aldrich) in 1× PBS for 10 min. Peroxidasis quenching was performed incubating the samples in a 0.6% H2 O2 methanol solution for 15 min in the dark. To block the aspecific binding sites, the samples were incubated with goat serum (Vector Lab, Burlingame, CA, USA) diluted 1:20 in 1× PBS at 37 ◦ C for 20 min. After washing, the specimens were incubated in a moisted chamber overnight at 4 ◦ C with mouse monoclonal anti-osteopontin (OPN) diluted 1:2000 (sc-21742, Santa Cruz Biotechnology, Santa Cruz, CA, USA) and mouse monoclonal anti-Aggrecan diluted 1:50 (sc-33695, Santa Cruz Biotechnology). Negative controls were performed incubating some sections with 0.1% BSA/1× PBS only. After 24 h, the samples were incubated with goat anti-mouse biotinylated secondary antibodies (Vector Lab) diluted 1:200 in 1.5% goat or horse serum–1× PBS solution for 60 min and then with streptavidin (Vectastain Elite ABC Kit Standard, Vector Lab) for 30 min. The reactions were revealed incubating the specimens in the substrate-chromogen solution (0.5 mg/ml, 3,3-diaminobenzidine tetrahydrochloride containing 0.02% H2 O2 ; Amresco, Solon, OH, USA) for 5 min in the dark, counterstained with hematoxylin and observed by a microscope (Leica Microsystems, Wetzlar, Germany). After each passage, washing in 0.01% Triton/1× PBS was performed. 2.4.7. Scanning electron microscopy (SEM) analysis After fixation (4% neutral buffered formalin in PBS, overnight at 4 ◦ C), cell/scaffold constructs were dehydrated in a graded series of ethanol aqueous solutions up to anhydrous ethanol, dried using the critical point method (Balzers CPD030, Oerlikon Balzers, Balzers, Liechtenstein), cross-sectioned, mounted on aluminum stumps, sputter-coated with gold (Sputter Coater Emitech K550, Quorum Technologies Ltd., West Sussex, UK), and examined with a scanning electron microscope (JEOL JSM-5200, JEOL Ltd., Tokyo, Japan). 2.4.8. Transmission electron microscopy (TEM) analysis TEM analysis was performed on DPSC specimens incubated overnight with culture medium containing PLL-BNNTs at 10 ␮g/ml. After removal of the culture medium, cells were trypsinized, centrifuged and fixed in a solution constituted of 0.5%/4% (w/v) GTA/formaldehyde in PBS 0.1 M at pH 7.2 for 2 h at 4 ◦ C. After washing, the samples were post-fixed in 1% w/v OsO4 PBS 0.1 M at pH 7.2 for 1 h, washed and dehydrated with acidified acetondimethylacetal (Fluka). Finally, the samples were embedded in Epon/Durcupan resin in BEEM capsules #00 (Structure Probe, West Chester, PA, USA) at 56 ◦ C for 48 h. Ultra-thin sections (20–30 nm thick) were obtained with an ultramicrotome (Ultratome Nova, LKB, Bromma, Sweden) equipped with a diamond knife (Diatome, Biel/Bienne, Switzerland). The sections were placed on 200 square mesh nickel grids, counterstained with saturated aqueous uranyl acetate and lead citrate solutions and observed with a transmission electron microscope (Zeiss 902, Carl Zeiss, Oberkochen, Germany). Drops of diluted PLL-BNNT aqueous solution were poured on copper grids and observed via TEM. 2.4.9. Statistical analysis Statistical significance in quantitative analyses was evaluated using the two-tailed t test for paired (alamarBlue® assay) or unpaired (RT-PCR, ds-DNA and GAG) data, followed by Bonferroni’s correction. Data underwent both descriptive, i.e., mean ± standard deviation (SD), and inferential statistics (p values).

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3. Results 3.1. Isolation and characterization of DPSCs Dental pulp tissues were collected from 10 clinically extracted human teeth. In this study, we observed that culturing human dental pulp tissue in plastic dishes led to proliferating adherent colonies of fibroblast-like cells that presented a durable proliferative capability in culture lasting for 4 months without evident morphological changes. DPSCs are able to form colonies, similar to the colonies formed by BM-MSCs, in 10–14 days of culture. When colonies were detached and replated at defined cell density, DPSCs formed a homogenous monolayer of cells which keep the fibroblast-like morphology (Fig. 1A). After studies of 10 passages, over a period of 4 months, cell cultures reached a doublings number of approximately 25 PDs and showed an average doubling time of 115 h. DPSCs were characterized via flow cytometry for surface protein expression. We analyzed the phenotypes of all populations, each of which corresponds to a dental pulp donor, cultivated and expanded at different passages. As shown in Fig. 1B, almost all cells expressed typical mesenchymal stromal cell antigens such as CD90, CD105, CD73, CD146, STRO-1, but they lacked the CD45, CD34, CD31 and SSEA-4 antigens. ELISA assay were performed on culture media to detect the amounts of NGF and BDNF secreted by the DPSCs. The analyses confirmed that the expression of BDNF and NGF was detectable in three different samples of DPSCs with values ranging from 3 to 4 pg/cell (Fig. 1C). 3.2. Differentiation of DPSCs The DPSCs were investigated throughout a functional characterization to assess their multipotency. 3.2.1. Endothelial differentiation Endothelial differentiation was performed by culturing DPSCs in the presence of EGM-2 for 15 days. Immunophenotype analysis showed more than 70% of differentiated DPSCs to be positive for CD309, and 40% to be positive for CD31 (Fig. 2A). Same antigens were not expressed by cells cultured under proliferating conditions. The Ac-LDL uptake assay showed that the differentiated DPSCs were able to internalize Ac-LDL into their cytoplasm (green in Fig. 2B) similarly to human umbilical vein endothelial cells (HUVECs, data not shown) while undifferentiated cells failed to uptake Ac-LDL (Fig. 2C). Differentiated DPSCs formed a tube-like structure within 12 h incubation on Matrigel (Fig. 2D), in contrast to the undifferentiated cells that were unable to form capillary-like structures (data not shown). Gene expression analysis of PECAM1 (CD31) mRNAs corroborated the immunophenotypic results, thus confirming the attainment of endothelial differentiation (Fig. 2E). 3.2.2. Osteogenic differentiation The DPSCs were induced to differentiate toward the osteoblast phenotype adding the culture medium with standard osteogenic supplements. After 21 days of culture, the deposition of calcium matrix nodules was revealed by both von Kossa and Alizarin S stain (Fig. 3A and B). Conversely, undifferentiated DPSCs did not show any positive stain for calcium deposition (Fig. 3C and D). 3.2.3. Chondrogenic differentiation CPs cultured in standard conditions and CPs treated with low frequency US (namely, CPs-US) underwent histologic evaluation for cartilaginous molecule expression (i.e., GAGs and aggrecan; Fig. 4A and H). CPs-US showed higher compactness and homogeneity than those of untreated CPs. Specifically, in CPs-US, cartilaginous ECM molecules were uniformly distributed in the samples and present to a higher extent than in CPs. Differently, in untreated

CPs, chondrogenic markers were often detected with zonal distribution. However, the investigated molecules were expressed with a similar intensity in CPs and CPs-US. In particular, generic GAGs were expressed with high intensity (+++) and uniform distribution in all the samples, as shown by Alcian Blue staining at pH 2.5 (Fig. 4E and F). Sulfated GAGs, typical of mature cartilage, were revealed with strong intensity (+++) both in the CPs and CPs-US, as demonstrated by both Toluidine blue (metachromatic reaction) and Alcian Blue staining at pH 1. In the CPs-US, the positivity was uniformly located (Fig. 4B and D), while in the CPs some areas were negative (Fig. 4A and C). Finally, as displayed by IHC, the aggrecan marker was weakly (+) and uniformly expressed in both CPs and CPs-US samples, but in the latter with an enhanced protein synthesis (Fig. 4G and H). 3.3. Cell culture with biomaterial platforms 3.3.1. Human plasma clots Significantly reduced cell viability was detected on clots after osteogenic induction (27.5 ± 5.5%, p < 0.05), in comparison with clots cultured in proliferation medium (40.9 ± 8.7%). Osteogenic differentiation was demonstrated by enhancement of mineralized matrix deposition, clearly evidenced through the presence of black granules (Fig. 3E), with respect to undifferentiated controls (Fig. 3F). The clot with the highest cell density (100,000 cells) demonstrated the most intense mineralization. 3.3.2. PVA/G scaffolds As detected by alamarBlue assay, DPSCs cultured on PVA/G scaffolds under diverse differentiative regimens were viable up to 21 culture days (Fig. 5A). Data of dye reduction percentage along the culture times (1, 11, 21 days), as calculated following the manufacturer’s formula were: 50.12 ± 7.79%, 51.95 ± 13.99%, 43.12 ± 10.11% for undifferentiated DPSCs; 50.49 ± 9.07%, 40.05 ± 4.97%, 46.43 ± 5.13% for osteo-differentiated DPSCs; and 40.37 ± 9.40%, 25.61 ± 8.04%, 30.69 ± 5.62% for chondrodifferentiated DPSCs. Between day 1 and day 21, no statistically significant differences in cell viability were detected within undifferentiated and osteo-differentiated groups (p > 0.05), while chondro-differentiated samples showed a statistically significant decrease (p = 0.0002). Viability of chondro-induced samples showed statistically significant reductions compared to both undifferentiated (p < 0.0001 on day 11 and p < 0.001 on day 21) and osteo-differentiated samples (p < 0.001 on day 11 and p < 0.0001 on day 21). At the endpoint, ds-DNA and GAG contents were detected with quantitative assays (Fig. 5B and C). Results of ds-DNA contents were 0.31 ± 0.12, 0.26 ± 0.13 and 0.26 ± 0.64 ␮g/sample for un-, osteo- and chondro-differentiated constructs, respectively. No statistically significant difference was highlighted (p > 0.05) (Fig. 5B). Findings on GAGs showed a similar trend, showing contents of 3.35 ± 0.38 ␮g/sample, 3.27 ± 0.34 ␮g/sample and 3.71 ± 0.56 ␮g/sample for un-, osteo- and chondro-differentiated constructs, respectively. On average, the highest GAG amount was found in the chondro-induced samples; however, in any case, statistically significant differences were not detected (p > 0.05) (Fig. 5C). SEM analysis highlighted DPSCs being able to penetrate in the pore network and deeply colonize the PVA scaffold in all the types of constructs (Fig. 6A and C). Cells were well stretched out with elongated morphology in un- and osteo-differentiated samples, while cells showed mildly spindle and round morphology, typical of the mature chondrocytes, in chondro-differentiated constructs (Fig. 6C). In un- and osteo-differentiated specimens, immunohistochemistry revealed a widespread presence, at both intra- and extracellular level, of the osteogenic marker OPN, with higher intensity (+++) in the osteo-differentiated than in the un-differentiated constructs (+) (Fig. 6D and E). In a similar fashion,

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Fig. 1. Morphological, immunological and functional characterization of DPSCs. (A) DPSCs formed a homogenous monolayer of cells that revealed typical spindle-shaped morphology. (B) Flow cytometrical analysis of DPSC showed typical immunophenotype characterized by the expression of CD90, CD105, CD73, CD146, and STRO-1. (C) ELISA assay confirmed that DPSCs of three different samples were able to secrete BDNF and NGF neurotrophins.

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Fig. 2. Differentiation of DPSCs toward endothelial lineage. (A) After induction of endothelial differentiation DPSCs acquired positive stain for anti-CD309 and anti-CD31 antibodies. (B) Moreover, differentiated DPSCs were able to internalize Ac-LDL (green in B) respect to (C) untreated cells, as well as (D) to form capillary tube-like structures after 12 h incubation on Matrigel (arrows). (E) Levels of PECAM1 mRNA, reported as fold expression change, of differentiated DPSCs resulted higher than undifferentiated DPSCs. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

the chondro-differentiated constructs displayed the chondrogenic marker aggrecan at intracellular level with a higher immunopositivity (++) than that detected in the un-differentiated constructs (−/+) (Fig. 6F and G). 3.3.3. BNNTs PLL-coated BNNTs were administrated with different doses (0, 5 and 10 ␮g/ml) to cultures of DPSCs up to 72 h, and cell viability was monitored using the alamarBlue test (Fig. 7A). Dye reduction percentage ranged between 55.36 and 63.31% in control (0 ␮g/ml PLL-BNNTs), 58.83 and 68.87% in 5 ␮g/ml, and 59.21 and 66.78% in 10 ␮g/ml samples. Within each dose-group, and between different dose-groups at same times, cell viability did not show any

statistically significant difference up to the endpoint (p > 0.05). Finally, PLL-BNNTs were observed via TEM both before and after administration in the culture medium (Fig. 7B and C), resulting in strongly contrasted nano-objects, which were detected as clusters in cytoplasm vesicles of DPSCs (Fig. 7C). The presence of such nano-objects at nuclear level was never highlighted. 4. Discussion Stem cell biology has emerged as a key pillar for regenerative medicine. In order to effectively regenerate tissues via engineered technologies, understanding the biology of stem cells has become an essential basis for driving and controlling the regeneration

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Fig. 3. Osteogenic differentiation of DPSCs. (A) Under proper stimuli, DPSCs were able to differentiate toward osteogenic lineage forming mineralized nodules (arrows), detected by von Kossa staining, or (B) Alizarin S. (C and D) Conversely, untreated cells did not reveal any matrix deposition. Osteogenic potential of DPSCs were also demonstrated in plasma clot scaffolds by the presence of black granules (arrows in E), while in non-osteoinductive condition (F) mineralized nodules were not detected.

processes. The simple question is what are the properties of cells that make them potentially ideal candidates for tissue engineering. Serakinci and Keith (2006) indicated some of the desirable properties that stem cells for transplantation should ideally possess: (i) to be easily isolated and purified, (ii) to maintain their multipotential lineage capacity, (iii) to show directed differentiation, (iv) to be autologous to the patient, (v) to allow genetic modification, (vi) to be highly expanded in culture, and (vii) to be non-tumorigenic. Nonetheless, in most cases applying multipotent cells to a surgical setting is not possible in the form of cell suspension, and requires the use of a scaffold avoiding cell dispersion in the surgical site. Tissue engineering is an interdisciplinary field that applies the principles of engineering and life sciences to the development of three-dimensional scaffolds able to sustain or improve the function of an applied regenerative cell population. The scaffold’s mechanical and biochemical properties determine the efficiency and avidity in which the cells can form the composite. The ideal biomaterial should be biocompatible and bioresorbable to support the replacement of normal tissue

without inflammation. Incompatible materials are destined for an inflammatory or foreign-body response that eventually leads to rejection and/or necrosis. In addition, the degradation products, if produced, should be removed from the body at an adequate rate via metabolic pathways to keep the concentration of these degradation products in the tissues at a tolerable level. Furthermore, the scaffold should provide an environment in which the appropriate regulation of cell behavior (e.g., adhesion, proliferation, migration, and differentiation) can occur to allow the formation of a functional tissue. Depending on their origin, biomaterials can be divided into three classes: (i) naturally derived (e.g. plasma clot), (ii) completely synthetic, and (iii) biomimetic materials (e.g. PVA/G). Naturally derived materials have the potential advantage of biologic recognition while synthetic polymers can be reproduced on a large scale with controlled properties of strength, degradation rate, and microstructure. Bioartificial materials can be produced from blends of biologic and synthetic polymers and have been developed to account for the advantages of both types of biomaterials.

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Fig. 4. Histologic analysis of chondroinduced DPSC pellets without (A, C, E, G) and with LFUS irradiation (B, D, F, H). (A and B) Toluidine Blue staining: sulfated GAGs are stained in violet. (C and D) Alcian Blue staining at pH 1: sulfated GAGs are stained in cyan, cell nuclei are showed in red. (E and F) Alcian Blue pH 2.5 staining: generic GAGs are stained in cyan, cell nuclei are showed in red. (G and H) Aggrecan is detected in brown by immunohistochemistry; cell nuclei are shown in blue. (A–H) Original magnification ×400; scale bars: 50 ␮m. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

Nonetheless, stem cells should be able to proliferate, migrate, and differentiate inside the scaffold itself. Thus, maintaining the cell biological properties, described by Serakinci, inside the scaffold should be considered as a further requisite of cell candidates in order to be a feasible tool in reconstructive medicine. The ability of a cell population to interact and functionalize the engineered substrate should not be predicted by assaying the biological properties “on plate” only. Therefore, we believe that it is necessary to confirm that cells maintain their characteristics once applied to different bioengineered platforms. The fundamental requisites of

biomaterial for tissue engineering is usually only related by the ability to efficiently promote the adhesion of the cells from which they will be colonized. Possible influences on cell proliferation and differentiation should also be evaluated in order to define the ideal composite for tissue regeneration. In order to evaluate human DPSCs as a feasible progenitor source for regenerative medicine, we assayed DPSC growth and multi-lineage differentiation focusing on their possible interference with different bioengineering platforms (i.e., LFUS, plasma clot and PVA/G scaffolds, BNNTs). The DPSCs were a homogeneous

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Fig. 5. Quantitative analyses on DPSC/PVA constructs. (A) Cell viability with alamarBlue test at different culture times. (B) Construct cellularity (ds-DNA content per sample) at the endpoint (21 days). (C) GAG content at the endpoint (21 days). Asterisks indicate different statistical significance of comparisons: *p < 0.01, **p < 0.001, ***p < 0.00001.

Fig. 6. SEM and immunohistochemical analysis of DPSC/PVA constructs. SEM analysis of DPSC/PVA constructs cultured (A) in standard medium, (B) in osteogenic medium (arrows: osteo-differentiated cells), (C) in chondrogenic medium (arrows: mature chondrocytes). Immunohistochemical analysis of the osteogenic and chondrogenic markers, performed on the DPSC/PVA constructs. OPN and aggrecan are displayed in brown (arrows), cell nuclei are showed in blue. OPN on the construct cultured (D) in standard medium and (E) in osteogenic medium. Aggrecan on the construct cultured (F) in standard medium and (G) in chondrogenic medium. SC = scaffold. Original magnification ×400; scale bars: 50 ␮m.

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Fig. 7. Results of BNNT-treated DPSCs. (A) Cytotoxicity with alamarBlue test with different PLL-BNNT doses (0, 5 and 10 mg/L). (B and C) TEM micrographs; magnification ×12,000. (B) Plain PLL-BNNTs (arrows); and (C) PLL-BNNTs internalized inside cytoplasm vesicles of a single DPSC (arrows). Scale bars: 1 ␮m.

population that expressed antigens typical of mesenchymal stromal cells, such as CD90, CD105, CD73, STRO-1 and the absence of CD45, CD34 and CD31, suggesting a lack of cells of hematopoietic and angiogenic lineages. In addition, these cells were positive for CD146, suggesting an involvement in the perivascular cell niche of dental pulp tissue. Indeed, stem cell properties are maintained in the adult tissue by a special microenvironment known as the stem cell niche (Scadden, 2006) that has been identified with the perivascular area in the pulp tissue (Shi and Gronthos, 2003). We also analyzed the angiogenic potential of DPSCs, evaluating the differentiation process of these cells. We demonstrated that differentiated DPSCs cultured in Matrigel promoted 3D capillary-like structures after 12 h and the uptake of Ac-LDL after 4 h. In addition, DPSCs differentiated into endothelial cells, expressed the endothelial markers CD309 and CD31 and showed an increase of PECAM by RT-PCR. Our study demonstrated that DPSCs support the generation of endothelial cells as revealed by the formation of capillary-tube like structures and this might be an indication of the possible contribution of these cells to neo-angiogenesis. A recent study showed that when DPSCs were injected intramyocardially after myocardial infarct, there was a significant increase in the size of the anterior wall and a significant decrease in the size of the infarct (Gandia et al., 2008). Another study further demonstrated the potential of DPSCs in preventing the progression of liver fibrosis in rats treated with carbon tetrachloride, restoring liver function (Ikeda et al., 2008). These findings suggest that even if DPSCs are not contributing directly to new tissue formation, they could support regenerative mechanisms by inducing neo-angiogenesis in the graft site. Furthermore, in the present study, we showed that DPSCs secrete BDNF and NGF in the supernatants of DPSC cultures, suggesting that in addition to neoangiogenic induction, these cells might contribute to the regeneration by paracrine effect of secreted growth factors. All of these interesting biological properties of DPSCs should be improved or at least maintained once they are applied in the biological bioengineering material in order to describe these cells as a feasible tool for regenerative medicine.

Considering this, we tested the possible influences of different bioengineering platforms on DPSC biology, applying bioartificial and biological scaffolds (PVA/G and plasma clot), evaluating LFUS application and testing BNNT cytocompatibility. Our previous works have recently demonstrated that the plasma clot as a natural substrate could represent the ideal support for the delivery of MSCs from BM and human umbilical cord blood, and that its moldability facilitates the work of the surgeon to adapt the material to the damaged surface (Trombi et al., 2008; Barachini et al., 2009). In line with BM-MSC/plasma clot behavior, we could observe a favorable interaction between DPSCs and the 3D architecture of the fibrin-microfiber clot and most importantly we demonstrated that osteogenic potential of these cells is maintained in the composite confirming that DPSCs could be successfully applied in combination with a biological scaffold. Fibrin clot aggregates in vivo undergo short-time degradation, thanks to processes such as fibrinolysis and phagocytosis, making these scaffolds highly resorbable. Conversely, PVA-based scaffolds are biocompatible matrices showing long-lasting biostability in the human body and they can suite other tissue engineering applications (Baker et al., 2012). In this study, DPSCs were seeded on PVA/G sponges obtained via emulsion and freeze-drying, a scaffolding technique which allows pore formation of a wider size than that of hydrogels (Hutmacher, 2000). Gelatin acts as a natural porogen and increases cell affinity toward the pristine synthetic polymer PVA. It has to be pointed out that pore features in the scaffolds play a key role in cell migration, survival and ECM production. Previous investigations have highlighted that PVA/G sponges act as suitable supports for connective-like tissue synthesized by human gingival fibroblasts (Moscato et al., 2008). We showed that DPSCs could colonize the entire 3D structure filling the scaffold porosity, remaining viable and synthetically active also under differentiative culture conditions. Since cellularity was not statistically different in un-, osteo- and chondro-differentiated constructs at the endpoint, the metabolic decline shown by the differentiated samples could depend on specific cell conditions

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induced by differentiation. Compared to un-differentiated controls, we observed enhanced expression only of early stage bone or cartilage markers, such as OPN in the osteo-differentiated and aggrecan in the chondro-differentiated samples. This indicated that these specific substrates were somehow able to maintain the DPSC stemness delaying their differentiation timeline. These data support our hypothesis that some cell biological properties could be influenced by the different scaffolds. Nonetheless these influences could represent an appealing feature where a spatio-temporal control of stem cell differentiation is desirable (Kinney and McDevitt, 2013). Similarly to different materials, other bioengineering procedures could affect the biological properties of the composites, i.e. the application of US at low frequency during their in vitro culture under chondrogenic commitment. In our findings, a minimal DPSC irradiation with LFUS resulted in a spatially uniform distribution and enhanced synthesis of GAGs and aggrecan, thus confirming the favorable effects reported using other US sources and mesenchymal stromal cell types (Lee et al., 2006; Shah et al., 2013). Indeed, while cartilagineous pellets obtained by standard culture methods displayed non-homogenous cell morphology and marker distribution, CPs-US were histologically homogenous. USbased therapies are routinely applied in medical practice and might disclose new approaches also in the translational regenerative medicine of cartilage defects if combined with stem cell transplants. Moreover, we tested DPSCs with BNNTs, a novel class of nanomaterials that has been receiving growing attention by the scientific community (Golberg et al., 2010). Similarly to carbon nanotubes, BNNTs can be employed as intracellular vectors for drug and plasmid delivery, thus playing a key role in regenerative medicine. However, owing to their chemical inertia, BNNTs have shown a superior biocompatibility and, owing to their physical properties, they can also act as nanotransducers for cell stimulation therapies (Ciofani et al., 2013). Our assays showed that DPSCs are able to uptake BNNTs by endocytosis without suffering from important cytotoxical affects. In recent years, human DPSCs have received growing attention due to their characteristics in common with other MSCs, in addition to the ease of their harvesting. Teeth can be recovered during routine dental procedures at any time during a lifespan. As third molar teeth are removed during standard prophylactic interventions and do not require additional surgical procedures, the utilization of such tissues for stem cell isolation offers an attractive alternative. In addition, owing to their immaturity, DPSCs virtually possess privileged immunoregulatory properties and could be considered as good substitutes of BM-MSCs for many therapeutical uses. Moreover, in contrast to BM aspirates, human dental pulp offers several advantages: ease of harvesting, absence of risks to donors and absence of ethical problems. BM-derived MSCs are among the most promising adult stromal cell types for regenerative medicine. The advantages of DPSCs include not only their high proliferation capability, but also the painless nature of stem cell collection with minimal invasion. In fact, the third molars are the most common source of dental stromal cells, because extraction of wisdom teeth is widely performed and the teeth are usually considered to be medical waste. In conclusion, DPSCs appear as a feasible cellular tool to be employed in regenerative medicine. Indeed, DPSCs are obtainable with non-invasive procedures, are easy to isolate and preserve a high replication capability. Moreover, DPSCs show multi-lineage differentiation, including the endothelial lineage. Finally and most importantly, DPSCs were highly versatile in interacting with different bioengineering strategies, making the specific combination between cells and a biomedical platform exploitable to target distinct regenerative strategies.

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