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Adipose Tissue–derived Microvascular Fragments as Vascularization Units for Dental Pulp Regeneration

  • Xun Xu
    Affiliations
    National Engineering Laboratory for Oral Regenerative Medicine, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    Department of Oral and Maxillofacial Surgery, West China Hospital of Stomatology, Sichuan University, Chengdu, China
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  • Cheng Liang
    Affiliations
    National Engineering Laboratory for Oral Regenerative Medicine, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    Department of Oral and Maxillofacial Surgery, West China Hospital of Stomatology, Sichuan University, Chengdu, China
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  • Xin Gao
    Affiliations
    National Engineering Laboratory for Oral Regenerative Medicine, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    Department of Oral and Maxillofacial Surgery, West China Hospital of Stomatology, Sichuan University, Chengdu, China
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  • Haisen Huang
    Affiliations
    National Engineering Laboratory for Oral Regenerative Medicine, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    Department of Oral and Maxillofacial Surgery, West China Hospital of Stomatology, Sichuan University, Chengdu, China
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  • Xiaotao Xing
    Affiliations
    National Engineering Laboratory for Oral Regenerative Medicine, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    Department of Oral and Maxillofacial Surgery, West China Hospital of Stomatology, Sichuan University, Chengdu, China
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  • Qi Tang
    Affiliations
    West China School of Public Health and West China Fourth Hospital, Sichuan University, Chengdu, China
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  • Jian Yang
    Affiliations
    National Engineering Laboratory for Oral Regenerative Medicine, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    Department of Oral and Maxillofacial Surgery, West China Hospital of Stomatology, Sichuan University, Chengdu, China
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  • Yutao Wu
    Affiliations
    National Engineering Laboratory for Oral Regenerative Medicine, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    Department of Oral and Maxillofacial Surgery, West China Hospital of Stomatology, Sichuan University, Chengdu, China
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  • Maojiao Li
    Affiliations
    National Engineering Laboratory for Oral Regenerative Medicine, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    Department of Oral and Maxillofacial Surgery, West China Hospital of Stomatology, Sichuan University, Chengdu, China
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  • Huanian Li
    Affiliations
    National Engineering Laboratory for Oral Regenerative Medicine, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    Department of Oral and Maxillofacial Surgery, West China Hospital of Stomatology, Sichuan University, Chengdu, China
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  • Li Liao
    Correspondence
    Address requests for reprints to Dr Li Liao, State Key Laboratory of Oral Disease, West China School of Stomatology, Sichuan University, 14# South Renmin Road Chengdu, China 610018, or Dr Weidong Tian, Department of Oral and Maxillofacial Surgery, West China Hospital of Stomatology, Sichuan University, No. 14, 3rd Section, Renmin South Road, Chengdu 610041, China.
    Affiliations
    National Engineering Laboratory for Oral Regenerative Medicine, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    Department of Oral and Maxillofacial Surgery, West China Hospital of Stomatology, Sichuan University, Chengdu, China
    Search for articles by this author
  • Weidong Tian
    Correspondence
    Address requests for reprints to Dr Li Liao, State Key Laboratory of Oral Disease, West China School of Stomatology, Sichuan University, 14# South Renmin Road Chengdu, China 610018, or Dr Weidong Tian, Department of Oral and Maxillofacial Surgery, West China Hospital of Stomatology, Sichuan University, No. 14, 3rd Section, Renmin South Road, Chengdu 610041, China.
    Affiliations
    National Engineering Laboratory for Oral Regenerative Medicine, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    State Key Laboratory of Oral Diseases, West China Hospital of Stomatology, Sichuan University, Chengdu, China

    Department of Oral and Maxillofacial Surgery, West China Hospital of Stomatology, Sichuan University, Chengdu, China
    Search for articles by this author
Published:April 19, 2021DOI:https://doi.org/10.1016/j.joen.2021.04.012

      Abstract

      Introduction

      The transplantation of dental pulp stem cells (DPSCs) has emerged as a novel strategy for the regeneration of lost dental pulp after pulpitis and trauma. Dental pulp regeneration of the young permanent tooth with a wide tooth apical foramen has achieved significant progress in the clinical trials. However, because of the narrow apical foramen, dental pulp regeneration in adult teeth using stem cells remains difficult in the clinic. Finding out how to promote vascular reconstitution is essential for the survival of stem cells and the regeneration of dental pulp after transplantation into the adult tooth.

      Methods

      Adipose tissue–derived microvascular fragments (ad-MVFs) were isolated from human adipose tissues. The apoptosis and senescence of DPSCs cultured in conditioned media were evaluated to explore the effects of ad-MVFs on DPSCs. DPSCs combined with ad-MVFs were inserted into the human tooth root segments and implanted subcutaneously into immunodeficient mice. Regenerated pulplike tissues were analyzed by hematoxylin and eosin and immunohistochemistry. The vessels in regenerated tissues were analyzed by Micro-CT and immunofluorescence.

      Results

      The isolated ad-MVFs contained endothelial cells and pericytes. ad-MVFs effectively prevented the apoptosis and senescence of the transplanted DPSCs both in vivo and in vitro. Combined with DPSCs, ad-MVFs obviously facilitated the formation of vascular networks in the transplants. DPSCs combined with ad-MVFs formed dental pulp–like tissues with abundant cells and matrix after 4 weeks of implantation. The supplementation of ad-MVFs led to more odontoblastlike cells and increased the formation of mineralized substance around the root canal.

      Conclusions

      Cotransplantation with ad-MVFs promotes the angiogenesis and revascularization of transplanted DPSC aggregates, leading to robust regeneration of dental pulp.

      Key Words

      ad-MVFs enhance the vascularization of grafts and the regeneration of dental pulp. Considering the availability of autologous ad-MVFs in the clinic, ad-MVFs might be used to solve the challenge of dental pulp regeneration in adult permanent teeth.
      Dental pulpitis caused by trauma and infection is a common disease. Because of the structure of the root canal and the low capacity for self-repair, the pulpitis usually results in necrosis of the dental pulp. Root canal therapy, the conventional treatment for pulpitis, needs to remove the infected dental pulp totally and fill the pulp cavity and root canal with inorganic materials, but the tooth becomes devitalized, brittle, and vulnerable to be broken after pulp loss
      • Soares C.J.
      • Rodrigues M.P.
      • Faria E.S.
      • et al.
      How biomechanics can affect the endodontic treated teeth and their restorative procedures?.
      . At present, no practical treatment is available to regenerate the lost dental pulp in the clinic.
      The transplantation of dental pulp stem cells (DPSCs) has emerged as a promising strategy to regenerate dental pulp
      • Miran S.
      • Mitsiadis T.A.
      • Pagella P.
      Innovative dental stem cell-based research approaches: the future of dentistry.
      . A recent clinical trial demonstrated that the transplantation of aggregates of DPSCs successfully regenerates the dental pulp of injured young permanent teeth
      • Xuan K.
      • Li B.
      • Guo H.
      • et al.
      Deciduous autologous tooth stem cells regenerate dental pulp after implantation into injured teeth.
      , but dental pulp regeneration in permanent adult teeth has not achieved a breakthrough. The main obstacle is the narrow apical foramen, which limits the supply of oxygen and nutrition to maintain the survival and function of the transplanted stem cells
      • Yang J.
      • Yuan G.
      • Chen Z.
      Pulp regeneration: current approaches and future challenges.
      ,
      • Rombouts C.
      • Giraud T.
      • Jeanneau C.
      • About I.
      Pulp vascularization during tooth development, regeneration, and therapy.
      . The chronic hypoxic environment after transplantation leads to cell senescence and dysfunction. Moreover, because microvasculatures are essential niches for stem cells, the lack of microvasculature would affect the self-renewal and differentiation capacity of the transplanted stem cells
      • Rouwkema J.
      • Rivron N.C.
      • van Blitterswijk C.A.
      Vascularization in tissue engineering.
      . Physiologically, a vascular system crossing the apical foramen is essential for transplanting oxygen and nutrients, but the growth rate of microvessels is about 5 μm/h, which could not satisfy the survival of the transplants
      • Utzinger U.
      • Baggett B.
      • Weiss J.A.
      • et al.
      Large-scale time series microscopy of neovessel growth during angiogenesis.
      . Therefore, how to promote the revascularization of grafts is a critical issue to protect the function of seeded cells and promote the regeneration of dental pulp.
      Prevascularization of the transplants is a practical strategy to promote the angiogenesis of the transplants. Several studies have confirmed that combining DPSCs with endothelial cells (ECs) facilitates vascularization after transplantation in animals
      • Dissanayaka W.L.
      • Hargreaves K.M.
      • Jin L.
      • et al.
      The interplay of dental pulp stem cells and endothelial cells in an injectable peptide hydrogel on angiogenesis and pulp regeneration in vivo.
      ,
      • Khayat A.
      • Monteiro N.
      • Smith E.E.
      • et al.
      GelMA-encapsulated hDPSCs and HUVECs for dental pulp regeneration.
      . In these studies, ECs were derived from the umbilical cord of the newborn. However, autologous ECs are hard to obtain in adults. Because of the immune rejection and ethical issues of heterogeneous allogeneic ECs, this strategy is limited in the clinic.
      Adipose tissue–derived microvascular fragments (ad-MVFs) are functional vessel segments enzymatically isolated from adipose tissue. ad-MVFs contain several essential cell types (ECs, pericytes, and mesenchymal stem cells [MSCs]) for angiogenesis and can rapidly assemble into vascular networks after transplantation
      • Frueh F.S.
      • Spater T.
      • Lindenblatt N.
      • et al.
      Adipose tissue-derived microvascular fragments improve vascularization, lymphangiogenesis, and integration of dermal skin substitutes.
      . Until now, researchers have used ad-MVFs to vascularize an epicardial patch
      • Shepherd B.R.
      • Hoying J.B.
      • Williams S.K.
      Microvascular transplantation after acute myocardial infarction.
      , a pancreatic encapsulating device
      • Hiscox A.M.
      • Stone A.L.
      • Limesand S.
      • et al.
      An islet-stabilizing implant constructed using a preformed vasculature.
      , skin grafts
      • Frueh F.S.
      • Spater T.
      • Korbel C.
      • et al.
      Prevascularization of dermal substitutes with adipose tissue-derived microvascular fragments enhances early skin grafting.
      , and porous scaffolds
      • Laschke M.W.
      • Kleer S.
      • Scheuer C.
      • et al.
      Vascularisation of porous scaffolds is improved by incorporation of adipose tissue-derived microvascular fragments.
      . Because of the extensive resource of autologous adipose tissue in liposuction, ad-MVFs seem to be a promising source of autologous ECs for vascularizing the implants of DPSCs.
      In this study, we constructed a complex combining ad-MVFs with DPSC aggregates and evaluated the vascularization and dental pulp regeneration after transplantation in vivo.

      Materials and Methods

       Isolation and Culturing of DPSCs

      The primary human DPSCs used in this study were isolated from freshly extracted intact third maxillary molars of healthy patients aged 13–20 years who received orthodontic treatment at West China Hospital of Stomatology, Sichuan University, Chengdu, China, as described previously
      • Gronthos S.
      • Mankani M.
      • Brahim J.
      • et al.
      Postnatal human dental pulp stem cells (DPSCs) in vitro and in vivo.
      . The collection of human third molars was performed after receiving approval from the Ethics Committee of West China Hospital of Stomatology, Sichuan University and written informed consent from the volunteers. Briefly, the extracted teeth were cleaned with phosphate-buffered saline (PBS; OriGene, Rockville, MD) containing 10% penicillin-streptomycin (Solarbio, Beijing, China). We ground longitudinal grooves on the tooth with TF-11 diamond burs (SBT, Ningbo, China) and split the tooth along the grooves by elevator to harvest the pulp. Then, the separated pulp tissues were washed several times with 2% penicillin-streptomycin, shredded, and digested with 3 mg/mL collagenase type I (Sigma-Aldrich, St Louis, MO) for 30 minutes at 37°C. After the suspension was centrifuged at 1200 revolutions per minute (rpm) for 5 minutes, the supernatant was aspirated. The remaining tissues and cells were seeded in 25-cm2 culture flasks (Corning Incorporated, Corning, NY) containing alpha-minimum essential medium (α-MEM; HyClone, Marlborough, MA) supplemented with 10% fetal bovine serum (FBS; Gibco, Grand Island, NY) and 1% penicillin-streptomycin and cultured under 5% CO2 at 37°C. Three human pulp samples from diverse individuals were blended to decrease individual differences. DPSCs from passages 3 to 5 were used in all experiments.

       Formation of DPSC Aggregates

      DPSCs were detached using 0.25% Trypsin-EDTA (Gibco) and seeded on 100-mm culture dishes (Thermo Fisher Scientific, Waltham, MA) at a concentration of 1 × 106 cells per dish in 10% FBS/α-MEM. The cells were maintained at 37°C in a 5% CO2 humidified atmosphere and maintained until 80% confluence was achieved. Then, DPSCs were cultured in 10% FBS/α-MEM containing 50 μg/mL ascorbic acid (Sigma-Aldrich). The culture medium was refreshed every 3 days. After they were detached from the culture plates, the DPSCs could aggregate together into a cell pellet and then be used in transplantation.

       Characterization of DPSCs

      Flow cytometry analysis was conducted to detect the expression of MSC-related markers. DPSCs (>1 × 105 cells) were washed and resuspended in 2% FBS/PBS containing suggested concentrations of the following conjugated antibodies according to the manufacturer’s instructions for 30 minutes at 4°C: CD31/FITC (BD, Franklin Lakes, NJ), CD34/FITC (BD), CD45/FITC (BD), CD73/PE (BD), CD90/PE (BD), and CD105/PE (BD). Then, cells were washed 3 times and resuspended in 2% FBS/PBS for analysis using an Accuri C6 flow cytometer (BD). The exported data were analyzed using FlowJo V10 software (BD). The results of the flow cytometry analysis suggested the DPSCs used in this study highly expressed MSC markers, including CD73, CD90, and CD105 (Supplemental Fig. S1A is available online at www.jendodon.com).
      Additionally, the osteo-/odontogenic and adipogenic capacity of DPSCs was confirmed by alizarin red (Sigma-Aldrich) staining and oil red O (Sigma-Aldrich) staining as previously described
      • Li R.
      • Guo W.
      • Yang B.
      • et al.
      Human treated dentin matrix as a natural scaffold for complete human dentin tissue regeneration.
      (Supplemental Fig. S1B and C is available online at www.jendodon.com). The expression of tubulin β III (Abcam, Cambridge, UK) in DPSCs after neurogenic induction was detected under a fluorescence microscope (Olympus, Tokyo, Japan) to evaluate the ability of neurogenic differentiation (Supplemental Fig. S1D is available online at www.jendodon.com). To verify the self-renew of DPSCs, we seeded 1 × 103 on a 100-mm culture dish and cultured DPSCs for 10 days with 10% FBS/α-MEM. The colonies were stained with crystal violet (Solarbio) after being fixed in 4% formaldehyde (KESHI, Chengdu, China) (Supplemental Fig. S1E is available online at www.jendodon.com), and the ratio of clone formation was counted (18.7%).

       Isolation and Characterization of ad-MVFs

      After obtaining informed consent, ad-MVFs were isolated from adipose tissues from patients conducting cosmetic liposuction at Sichuan Huamei Zixin Medical Aesthetic Hospital, as described previously
      • Frueh F.S.
      • Spater T.
      • Scheuer C.
      • et al.
      Isolation of murine adipose tissue-derived microvascular fragments as vascularization units for tissue engineering.
      . Briefly, the adipose tissues were washed with α-MEM containing 2% penicillin-streptomycin. Then, fat tissues were cut up with scissors and digested for 15 minutes with collagenase I under vigorous stirring at 37°C. The digestion was neutralized with α-MEM supplemented with 20% FBS, and the suspension was incubated at 37°C for 5 minutes. Next, the fat supernatant was aspirated. The remaining suspension was filtered using a 500-μm strainer (pluriSelect, Leipzig, Germany) to remove undigested adipose tissue and a 20-μm strainer (pluriSelect) to harvest ad-MVFs. Finally, ad-MVFs were washed from the 20-μm strainers and enriched by a 5 minutes of centrifugation at 600g. Vascular-associated antibodies involving rabbit CD31 (Abcam), mouse CD34 (Abcam), and rabbit α-SMA (Abcam) were used to identify the isolated microvessels.

       Culture of ad-MVFs

      ad-MVFs were cultured on the 1-mm-thick Matrigel (Corning) with the endothelial medium containing DMEM:F12 medium (Gibco), 15% FBS, 20% Knockout Serum Replacement (KOSR), GlutaMAX (Gibco, Grand Island, NY), non-essential amino acids (NEAA, Gibco), 100 ng/mL vascular endothelial growth factor A, and 100 ng/mL fibroblast growth factor 2 (Peprotech, Cranbury, NJ). Pictures were taken with a microscope every day to observe the morphology of the vessels. After 14 days, the vascular networks were identified by immunofluorescence labeled by CD31 antibody.

       Influence of ad-MVFs to Senescent DPSCs In Vitro

      DPSCs were induced to senescence with 30 g/L D(+)-Galactose (Solarbio) for 48 hours and 10 μmol/L H2O2 (KESHI) for 24 hours, respectively. The conditioned media (media culturing ad-MVFs for 24 hours) was used to culture the senescent DPSCs. The Cell Counting Kit 8 (KeyGEN BioTECH, Nanjing, China) was used to detect cell proliferation following the recommended instructions. To detect the influence of ad-MVFs on the osteogenesis of DPSCs, we cultured senescent DPSCs with the conditioned media for 4 days and then the osteogenic medium for the next 12 days.

       Effects of ad-MVFs on Senescent DPSCs In Vivo

      We collected samples of DPSC aggregates inserted into human tooth root segments (hRTSs) and implanted subcutaneously for 3 and 7 days to explore the senescence and apoptosis of cells. Frozen sections were made to detect senescence-associated β-galactosidase (SA-β-Gal) with the β-Galactosidase Staining Kit (Solarbio) and apoptosis with the One Step TUNEL Apoptosis Assay Kit (Beyotime, Shanghai, China) following the manufacturers’ instructions. Then, the quantification of stained cells was performed with ImageJ software (National Institutes of Health, Bethesda, MD)
      • Schneider C.A.
      • Rasband W.S.
      • Eliceiri K.W.
      NIH Image to ImageJ: 25 years of image analysis.
      .

       Quantitative Real-Time Polymerase Chain Reaction

      Total RNAs were extracted from DPSCs using quick-RNA McroPrep (ZYMO, Irvine, CA); 1 μg total RNAs was used for reverse transcription into first-strand complementary DNA using HiScript (Vazyme, Nanjing, China). Quantification polymerase chain reaction (qPCR) analysis was conducted using SYBR Green qPCR Master Mix (Bimake, Houston, TX). The primer sequences used are listed in Supplemental Table S1 (available online at www.jendodon.com).

       Preparation of hTRSs

      Mandibular and maxillary premolars were harvested from patients younger than 25 years old who received tooth extraction at West China Hospital of Stomatology, Sichuan University. After removing the cementum, dental pulp tissues, and predentin, the roots of these teeth were horizontally sectioned into 10 mm-long segments, and the root canal was enlarged to 5 mm in diameter of both ends using sterilized FG-701L fissure burs (SBT). Then, the root segments soaked into deionized water were cleaned using an ultrasonic cleaner for 10 minutes. After that, the root canal was treated with 17%, 10%, and 5% EDTA (KESHI), respectively, for 30 minutes to demineralize the dentin matrixes on the root canal surface. The treated root canal was finally soaked into penicillin-streptomycin at 4°C over 3 days.

       Ectopic Regeneration of Dental Pulp in Immunodeficient Mice

      The animal experiments in this study were approved by the Experimental Animal Ethics Committee of West China Hospital of Stomatology, Sichuan University. Six-week-old male Balb/c nude mice were purchased from GemPharmatech Co, Ltd (Nanjing, China). All experimental procedures were performed under relevant guidelines and regulations.
      The subcutaneous implantation of hRTSs containing DPSC aggregates was performed as previously reported
      • Li X.
      • Ma C.
      • Xie X.
      • et al.
      Pulp regeneration in a full-length human tooth root using a hierarchical nanofibrous microsphere system.
      . In brief, thermoplasticized gutta-percha was heated to a molten state to seal 1 end of the hRTSs. To prevent leakage, we extended gutta-percha outward to the edge of the segments when blocking with pluggers. After that, we injected saline into the chambers of the sealed segments to check the liquid leakage. Freshly isolated ad-MVFs were resuspended in 20 μL saline and combined with cell sheets of DPSCs. Then, the grafts were rolled up and mildly inserted into the hTRS inner chamber. hTRSs were then bilaterally implanted in the subcutaneous space of the dorsum of immunodeficient mice. The implants were harvested after 4 weeks. In a preliminary experiment, we selected the dosage of 30,000 U and 60,000 U ad-MVFs pretransplant according to the studies of angiogenesis in other tissues
      • Spater T.
      • Korbel C.
      • Frueh F.S.
      • et al.
      Seeding density is a crucial determinant for the in vivo vascularisation capacity of adipose tissue-derived microvascular fragments.
      . The blood vessels formed in the implants of DPSC aggregates combined with 60,000 U ad-MVFs were more abundant than those combined with 30,000 U ad-MVFs (Supplemental Fig. S2 is available online at www.jendodon.com). As a result, we selected the dosage of 60,000 U ad-MVFs for transplant in the following experiments.

       Labeling Sign of Calcein

      To study the de novo mineral deposits, we injected 10 μg/mL calcein (Sigma-Aldrich)/PBS into mice 3 and 10 days before being sacrificed. The hard tissue sections of the hRTSs were made, and images were taken using a fluorescence microscope (Olympus).

       Perfusion of Microfil

      Angiography was performed to evaluate blood vessel formation as previously described
      • Bolland B.
      • Kanczler J.M.
      • Dunlop D.G.
      • et al.
      Development of in vivo muCT evaluation of neovascularisation in tissue engineered bone constructs.
      . After subcutaneous transplantation for 4 weeks, the mice were anesthetized with pentobarbital. An extended midline incision was made across the thorax and abdomen. The left sternum was cut to expose the heart, and the inferior vena cava was ligated. The left ventricle was cannulated with a 26-G intravenous catheter after the inferior vena cava was cut off, and 10 mL heparinized saline was injected. Next, Microfil MV-120 Blue (Flow-Tech, Carver, MA) and MV-Diluent solutions (2:3, Flow-Tech) along with MV Curing agent (Flow-Tech, 6% of total volume) were infused at 2 mL/min until the subcutaneous vessels were visualized to be filled with blue contrast agent. Finally, the mice were stored at 4°C for about 3 hours until the silicon compound polymerized. The samples were collected and fixed by 4% paraformaldehyde and scanned with µCT 50 (SCANCO Medical, Brüttisellen, Switzerland) using settings of 70 kV and 200 μA with a 10-mm voxel size. The 3-dimensional images were reconstructed to show the new blood vessels formed in the hTRSs. The density of the blood vessels was analyzed with EVALUATE software (SCANCO Medical).

       Histologic and Immunohistochemical Analysis

      After 4 weeks of transplantation, the implants were retrieved, fixed in 4% formaldehyde (KESHI) overnight at 4°C, and then demineralized with 10% EDTA until the dentin was softened. The demineralized samples were embedded in paraffin and sectioned at 5-μm thickness. Paraffin tissue sections were deparaffinized in dewaxing liquid (Solarbio) and rehydrated through graded ethanol solutions. Then, these sections were used for hematoxylin and eosin (H&E), immunofluorescence, and immunohistochemistry staining according to the manufacturers’ recommended protocols. Regarding immunofluorescence staining, primary antibodies of rabbit CD31 (Abcam), mouse p16 (Abcam), rabbit p21(Abcam), and mouse p53 (Abcam) and secondary antibodies of Alexa Fluor 488–conjugated goat antimouse (Invitrogen, Waltham, MA) and Alexa Fluor 555–conjugated goat antirabbit (Invitrogen) were used. Stained sections were finally inspected under a fluorescence confocal microscope (Olympus FV1000). For immunochemical staining, primary antibodies of dentin matrix protein 1 (DMP-1) (Biovision, Milpitas, CA) and dentin sialoprotein (DSP) (Santa Cruz, Dallas, TX) and secondary antibodies of goat antirabbit immunoglobulin G–horseradish peroxidase (Zen, Chengdu, China) and goat antimouse immunoglobulin G–horseradish peroxidase (Zen) were used. Images were scratched by a light microscope (Olympus BX43F).

       Semiquantitative Analysis of Blood Vessels and ECs

      For the analysis of the blood vessels, the sections of demineralized transplants were stained with H&E. Three sections of each hTRS were selected. The 2 × 4 mm region at the center of each section was selected for quantification. The images were scanned using Aperio AT (Leica, Buffalo Grove, IL) at 40× magnification. We quantified the vessels and the vessel areas with ImageJ software. For the analysis of ECs, red fluorescence–labeled CD31-positive cells were recognized as ECs, and the area of the lumen was calculated. Also, the number of cells adjacent to the dentin was counted, and the percentage of the matrix in H&E images was measured.

       Statistical Analysis

      Data were presented as mean ± standard deviation using GraphPad Prism 8 (GraphPad Software, La Jolla, CA). All statistical analyses were performed using 1-way analysis of variance or the Student t test. Values were compared using multiple comparisons where P < .05 was considered significant.

      Results

       ad-MVFs Are Potent in Regenerating Microvessel Networks

      Adipose tissue obtained from liposuction contained an abundant network of microvessels (Supplemental Fig. S3A is available online at www.jendodon.com). ad-MVFs in human adipose tissues were obtained by enzyme digestion and filtration as previously reported
      • Frueh F.S.
      • Spater T.
      • Scheuer C.
      • et al.
      Isolation of murine adipose tissue-derived microvascular fragments as vascularization units for tissue engineering.
      . Microscopic examination revealed that isolated ad-MVFs maintained integrated capillary networks (Supplemental Fig. S3B is available online at www.jendodon.com). Immunofluorescence showed ad-MVFs were surrounded by ECs (CD31+ and CD34+) and pericytes (α-SMA+) (Supplemental Fig. S3C is available online at www.jendodon.com).
      To confirm the capacity of ad-MVFs to regenerate microvessels, we cultured ad-MVFs on Matrigel for 2 weeks. ad-MVFs seeded on Matrigel formed multiple colonylike cell clusters and started to sprout vascularlike structures by 3 days. The neovessels extended and connected into continuous networks after culture for 14 days (Supplemental Fig. S3D is available online at www.jendodon.com). The newly formed microvessel networks expressed the red fluorescence–labeled CD31 marker, indicating the vessels were surrounded by ECs (Supplemental Fig. S3E is available online at www.jendodon.com).

       Angiocrine Factors of ad-MVFs Preserve the Function of Senescent DPSCs In Vitro

      The loss of physiologic niche is a factor of cellular senescence after transplantation. Because MSCs are usually located in the perivascular niche, ECs could support the functionality of MSCs through paracrine factors. To verify the influence of ad-MVFs on the senescence of DPSCs, we compared the changes in cell proliferation and mineralization abilities in cellular senescence models of DPSCs cultured with or without the conditioned media of ad-MVFs. As shown in Figure 1A, the conditioned media of ad-MVFs promoted the proliferation of senescent DPSCs. Furthermore, qPCR revealed that alkaline phosphatase (ALP), DMP, bone morphogenetic protein (BMP), and dentin sialophosphoprotein (DSPP) were markedly up-regulated in senescent DPSCs treated with the conditioned media of ad-MVFs relative to senescent DPSCs (Fig. 1B). These results suggest that paracrine factors from ad-MVFs could alleviate the senescence of DPSCs.
      Figure thumbnail gr1
      Figure 1Effects of the conditioned media of ad-MVFs on senescent DPSCs in vitro. (A) The proliferation curves of normal DPSCs, senescent DPSCs, and senescent DPSCs were treated with the conditioned media of ad-MVFs (DPSCs + ad-MVFs). (B) The expression of markers of odontogenesis and osteogenesis of DPSCs after osteogenic inducing for 12 days.

       ad-MVFs Promote the Vascularization of the Regenerated Dental Pulp

      We made 10-mm-long hTRSs with 1 end sealed to imitate the environment of tooth regeneration in the clinic. DPSC aggregates combined with ad-MVFs were inserted into the hRTSs and implanted subcutaneously into immunodeficient mice for dental pulp regeneration (Fig. 2A). These transplantations were harvested for analysis 4 weeks later.
      Figure thumbnail gr2
      Figure 2valuation of angiogenesis in the transplants of DPSC aggregates combined with or without ad-MVFs. (A) The immunodeficient mice model for the subcutaneous transplantation. ad-MVFs were combined with DPSC aggregates and then the rolled-up grafts were inserted into hTRSs to transplant subcutaneously into mice. (B) The mice were injected with Microfil 4 weeks after transplantation. Three-dimensional reconstruction of vessel networks filled with Microfil by micro–computed tomographic imaging. (C) The quantification of the vessel density in pulplike tissues of DPSCs + ad-MVFs compared with DPSCs. (D and E) H&E staining of the selected regions of the transplants of the 2 groups. F and G are an enlargement of the framed area in D and E, respectively. The black arrowheads show the vessel structures in F and G. (H) Semiquantitative measurement of the proportion of lumen areas. (I) Immunofluorescence labeling of ECs in DPSCs and DPSCs + ad-MVFs. The white arrowheads in I point to CD31-positive ECs. (J) Semiquantification of the ratio of CD31-positive areas. Scale bar: (B) 1 mm, (D and E) 500 μm, (F and G) 200 μm, and (I) 50 μm. ∗∗P < .01. ∗∗∗∗P < .0001.
      To assess vascularization in the DPSC and DPSC + ad-MVF groups, the vascular networks were filled with Microfil. Micro–computed tomographic images revealed that vasculatures in the DPSC + ad-MVF group were more evident and abundant than those in the DPSC group (Fig. 2B and C). Then, we analyzed angiogenesis in the transplant by histologic analysis. H&E staining showed integrated vessels were perfused with blood cells in both groups (Fig. 2D-G). The numbers and diameters of vessels in the DPSC + ad-MVF group were higher than that of the DPSC group. The semiquantitative analysis showed combination with ad-MVFs resulted in a 25% increase in the vessel area compared with the group without ad-MVFs (Fig. 2H). Immunofluorescence assay confirmed CD31+ EC-lined vascular networks in the pulplike tissues (Fig. 2I). Semiquantification of the ratio of vessels confirmed an increase in the vessels in the DPSC + ad-MVF group compared with those in the DPSC group (Fig. 2J).

       ad-MVFs Alleviate the Senescence and Apoptosis of DPSCs after In Vivo Implantation

      Insufficient blood supply leads to the apoptosis and senescence of the transplanted cells. We examined whether ad-MVFs could alleviate the apoptosis and senescence of DPSCs in vivo. TUNEL (Terminal Deoxynucleotidyl Transferase [TdT] -mediated dUTP Nick-End Labeling) assays showed the ratio of apoptotic cells in the DPSC + ad-MVF group was significantly lower than that in the DPSC group on the third and seventh days (Fig. 3A and B). According to SA-β-Gal staining, the DPSC + ad-MVF group showed a significant reduction in senescent cells after implantation compared with the DPSC group. Importantly, senescence in the DPSC + ad-MVF group was almost undetectable on the seventh day (Fig. 3C and D), indicating ad-MVFs can prevent cell senescence in the transplanted tissues. Accordingly, immunofluorescence of cellular senescence markers, such as p16, p21, and p53, confirmed the results of SA-β-Gal staining (Fig. 3E).
      Figure thumbnail gr3
      Figure 3Apoptosis and senescence in grafts after implantation for 3 and 7 days. Samples were collected after subcutaneous implantation for 3 and 7 days, respectively. Frozen sections were made to detect senescence-associated β-galactosidase and apoptosis. (A) The detection of apoptotic cells by TUNEL staining. (B) The ratio of apoptotic cells in different samples. (C) The visualization of senescent cells by SA-β-Gal staining. (D) The percentage of senescent cells in samples. (E) Immunofluorescence for p16, p21, and p53 in transplanted grafts at both time points. Scale bar: (A) 50 μm, (B) 50 μm, and (C) 100 μm.

       ad-MVFs Promote the Regeneration of Dental Pulp after Transplantation

      Histologic analysis confirmed that the DPSC + ad-MVF group showed enhanced cellularity compared with the DPSC group (Fig. 4AF). Moreover, the extracellular matrix increased in the DPSC + ad-MVF group (Fig. 4G). Immunohistochemical staining revealed that the cells adjacent to the dentin expressed odontoblast markers DMP-1 and DSP (Fig. 4H), suggesting the transplanted DPSCs differentiated into the odontoblastic lineage. According to the quantification, there were significantly more odontoblastlike cells in the DPSC + ad-MVF group than in the DPSC group (Fig. 4I). We also label the newly formed minerals in the root canal using calcein (Fig. 4J). The distance between the 2 fluorescence lines of the DPSC + ad-MVF group is broader than that of the DPSC group, indicating that ad-MVFs accelerated the mineral deposition in the DPSC + ad-MVF group (Fig. 4K).
      Figure thumbnail gr4
      Figure 4The regeneration of dental pulp. DPSCs with or without ad-MVFs were inserted into the hRTSs and subcutaneously implanted into immunodeficient mice. Then, the regenerated tissues were evaluated after transplantation of 4 weeks. (A and B) An overview of regenerated pulplike tissues of the DPSC and DPSC + ad-MVF groups. C and D are magnified images of the black rectangle framed region in A and B. E and F present enlarged views of the red rectangle framed region in A and B. (G) Semiquantitatively measuring the ratio of the red matrix in H&E staining. (H) Immunohistochemistry for DMP-1 and DSP in grafts. (I) The quantification of DPSCs adjacent to dentin. (J) The observation of odontogenic deposits by calcein with green fluorescence. (H) The measurement of the distance between 2 green fluorescent lines in the 2 groups. Scale bar: (A and B) 1 mm, (C and D) 100 μm, (E, F, and H) 50 μm, and (J) 20 μm. ∗P < .05. ∗∗P < .01. ∗∗∗∗P < .0001.

      Discussion

      Recent studies have applied 3-dimensional structures or microspheres made of poly-D, L-lactide/glycolide materials
      • Huang G.T.
      • Yamaza T.
      • Shea L.D.
      • et al.
      Stem/progenitor cell-mediated de novo regeneration of dental pulp with newly deposited continuous layer of dentin in an in vivo model.
      , hydrogels
      • Alqahtani Q.
      • Zaky S.H.
      • Patil A.
      • et al.
      Decellularized swine dental pulp tissue for regenerative root canal therapy.
      , and nanofibers
      • Li X.
      • Ma C.
      • Xie X.
      • et al.
      Pulp regeneration in a full-length human tooth root using a hierarchical nanofibrous microsphere system.
      combined with DPSCs to regenerate the dentin-pulp complex. Although the stem cell–based strategies seem to be practical in clinic trials, an essential challenge is to rapidly vascularize regenerated tissues and obtain blood perfusion to ensure the survival of implanted cells
      • Dissanayaka W.L.
      • Zhu L.
      • Hargreaves K.M.
      • et al.
      Scaffold-free prevascularized microtissue spheroids for pulp regeneration.
      . In the present study, we explored an effective and practical approach to rebuild the blood circulation of the dental pulp. To our knowledge, this is the first study to apply ad-MVFs as a vascularized component in dental pulp regeneration. Compared with previous studies using ECs such as human umbilical vein ECs, our strategy using ad-MVFs has several advantages. First, ad-MVFs contain 3 essential cell types of small vessels: ECs, pericytes, and stromal cells
      • McDaniel J.S.
      • Pilia M.
      • Ward C.L.
      • et al.
      Characterization and multilineage potential of cells derived from isolated microvascular fragments.
      . Second, ad-MVFs maintain the morphologic and functional characteristics of small vessels, which facilitates the regeneration of the vascular network
      • Hoying J.B.
      • Boswell C.A.
      • Williams S.K.
      Angiogenic potential of microvessel fragments established in three- dimensional collagen gels.
      . Third, autologous ad-MVFs are available for the majority of patients in the clinic. These advantages would be beneficial to the later clinical application of ad-MVFs in dental pulp regeneration.
      Transplanted stem cells are faced with an environment of ischemia and hypoxia due to the lack of blood perfusion. In such a situation, mitochondrial respiration is inhibited, and reactive oxygen species generated by cells leads to oxidative stress, resulting in stem cells apoptosis
      • Samakova A.
      • Gazova A.
      • Sabova N.
      • et al.
      The PI3k/Akt pathway is associated with angiogenesis, oxidative stress and survival of mesenchymal stem cells in pathophysiologic condition in ischemia.
      and senescence
      • Vono R.
      • Jover Garcia E.
      • Spinetti G.
      • Madeddu P.
      Oxidative stress in mesenchymal stem cell senescence: regulation by coding and noncoding RNAs.
      . Uncontrollable cell apoptosis and senescence are among the major reasons for the failure of stem cell transplantation
      • Shafiq M.
      • Jung Y.
      • Kim S.H.
      Insight on stem cell preconditioning and instructive biomaterials to enhance cell adhesion, retention, and engraftment for tissue repair.
      . In this study, we confirmed that the ratio of apoptotic cells gradually increased transplantation
      • Nunes S.S.
      • Greer K.A.
      • Stiening C.M.
      • et al.
      Implanted microvessels progress through distinct neovascularization phenotypes.
      . Importantly, we found that the apoptotic cells in the group containing ad-MVFs were significantly less than that in the DPSC group, and ad-MVFs significantly alleviated cell senescence after implantation for 7 days, suggesting that the presence of ad-MVFs prevents cell apoptosis and senescence in the transplant of DPSCs. Because the functional vessels have not been formed in the short-term, the protective effect of ad-MVFs might be due to angiocrine factors secreted by the cells residing in ad-MVFs
      • Laschke M.W.
      • Kleer S.
      • Scheuer C.
      • et al.
      Vascularisation of porous scaffolds is improved by incorporation of adipose tissue-derived microvascular fragments.
      . The hypothesis was supported by our in vitro assay using the conditional media of ad-MVFs. However, the mechanism of ad-MVFs preventing cellular senescence and apoptosis remains unknown.
      Our results found that the regeneration of dental pulp tissue in groups with ad-MVFs is improved compared with transplantation with DPSCs only. Furthermore, regularly arranged odontoblastlike cells expressing DMP and DSP can be observed almost vertically adjacent to the dentin, which indicates that DPSCs could differentiate into odontoblasts and acquired the ability to secrete dentin matrix
      • Suzuki S.
      • Sreenath T.
      • Haruyama N.
      • et al.
      Dentin sialoprotein and dentin phosphoprotein have distinct roles in dentin mineralization.
      ,
      • Chen Y.
      • Yu Y.
      • Chen L.
      • et al.
      Human umbilical cord mesenchymal stem cells: a new therapeutic option for tooth regeneration.
      . Although we observe no evident mineral deposits on H&E images, calcein showed the growth activity of new mineralized tissues. These effects might be majorly due to the fact that ad-MVFs facilitate the revascularization of transplants. Moreover, ad-MVFs are a rich source of proangiogenic factors that stimulate not only angiogenesis
      • Laschke M.W.
      • Menger M.D.
      Adipose tissue-derived microvascular fragments: natural vascularization units for regenerative medicine.
      but also odontogenesis. Further investigations are necessary to confirm the direct and indirect manner of ad-MVFs to promote dental pulp regeneration.
      In this study, we tested the effect of ad-MVFs on angiogenesis using a model of ectoptic dental pulp regeneration. hTRSs of human premolars deprived of dental pulp were used to simulate dental pulp loss in the clinic
      • Li X.
      • Ma C.
      • Xie X.
      • et al.
      Pulp regeneration in a full-length human tooth root using a hierarchical nanofibrous microsphere system.
      . We sealed 1 end of hTRSs with thermoplasticized gutta-percha to imitate the environment of clinical treatment. It was reported that thermoplasticized gutta-percha provides an apical seal superior to mineral trioxide aggregate in the clinic
      • Vizgirda P.J.
      • Liewehr F.R.
      • Patton W.R.
      • et al.
      A comparison of laterally condensed gutta-percha, thermoplasticized gutta-percha, and mineral trioxide aggregate as root canal filling materials.
      . Compared with thermoplasticized gutta-percha, MTA still has several drawbacks, including the possible release of hazardous substances, delayed setting time, and the handling inconvenience, which probably brings negative effects on the survival of implants and difficulties in operation
      • Parirokh M.
      • Torabinejad M.
      Mineral trioxide aggregate: a comprehensive literature review--part III: clinical applications, drawbacks, and mechanism of action.
      . Therefore, we chose thermoplasticized gutta-percha in this study. Although an ectopic dental pulp regeneration model is widely used in the research of dental pulp regeneration, we understand that the process of revascularization on the dental pulp in the subcutaneous site might be different from that in the alveolar bone. To overcome the limits of the ectopic model, an in situ dental pulp regeneration model using large animals might be valuable to confirm the potential of ad-MVFs.

      Conclusion

      In summary, we successfully improved the vascularization of DPSC aggregates with ad-MVFs for dental pulp regeneration. ad-MVFs present a powerful vascularization ability and enhance the odontogenesis of DPSCs. The senescence of DPSCs induced by hypoxia is alleviated in this process. Hence, ad-MVFs are a promising source of vascularization units used for dental pulp regeneration.

      Acknowledgments

      Supported by the National Key Research and Development Program of China (grant no. 2017YFA0104800 ), Key Project of Sichuan Province (grant nos. 2019YFS0515 and 2019YFS0311), the Fundamental Research Funds for the Central Universities (grant no. YJ201878 ), and the Nature Science Foundation of China (grant nos. 81600912 and 31601113 ).
      The authors deny any conflicts of interest related to this study.

      Supplementary Material

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