Abstract

Background

   The role of periodontal ligament stem cells (PDLSCs) and macrophage
   polarization in periodontal tissue regeneration and bone remodeling
   during orthodontic tooth movement (OTM) has been well documented.
   Nevertheless, the interactions between macrophages and PDLSCs in OTM
   remain to be investigated. Consequently, the present study was proposed
   to explore the effect of different polarization states of macrophages
   on the osteogenic differentiation of PDLSCs.

Methods

   After M0, M1 and M2 macrophage-derived exosomes (M0-exo, M1-exo and
   M2-exo) treatment of primary cultured human PDLSCs, respectively,
   mineralized nodules were observed by Alizarin red S staining, and the
   expression of ALP and OCN mRNA and protein were detected by RT-qPCR and
   Western blotting, correspondingly. Identification of differentially
   expressed microRNAs (DE-miRNA) in M0-exo and M2-exo by miRNA
   microarray, and GO and KEGG enrichment analysis of DE-miRNA targets,
   and construction of protein–protein interaction networks.

Results

   M2-exo augmented mineralized nodule formation and upregulated ALP and
   OCN expression in PDLSCs, while M0-exo had no significant effect.
   Compared to M0-exo, a total of 52 DE-miRNAs were identified in M2-exo.
   The expression of hsa-miR-6507-3p, hsa-miR-4731-3p, hsa-miR-4728-3p,
   hsa-miR-3614-5p and hsa-miR-6785-3p was significantly down-regulated,
   and the expression of hsa-miR-6085, hsa-miR-4800-5p, hsa-miR-4778-5p,
   hsa-miR-6780b-5p and hsa-miR-1227-5p was significantly up-regulated in
   M2-exo compared to M0-exo. GO and KEGG enrichment analysis revealed
   that the downstream targets of DE-miRNAs were mainly involved in the
   differentiation and migration of multiple cells.

Conclusions

   In summary, we have indicated for the first time that M2-exo can
   promote osteogenic differentiation of human PDLSCs, and have revealed
   the functions and pathways involved in the DE-miRNAs of M0-exo and
   M2-exo and their downstream targets.

Supplementary Information

   The online version contains supplementary material available at
   10.1186/s12903-022-02682-5.

   Keywords: Orthodontic tooth movement; Periodontal ligament stem cells;
   Macrophage polarization; Exosome, microRNA

Introduction

   Orthodontic tooth movement (OTM) is a process of bone remodeling
   occurring in periodontal tissues through a series of physiological
   reactions, with bone remodeling on the tension side of the bone and
   bone resorption on the pressure side, ultimately achieving tooth
   movement [[37]1, [38]2]. The periodontium connects the cementum to the
   alveolar bone through the type I fibers, which maintain the nutritional
   supply of the orthodontic teeth, and transmit and absorb mechanical
   stresses [[39]3]. Periodontal ligament stem cells (PDLSCs), a critical
   cell in periodontal tissue engineering, were first identified by Seo et
   al. [[40]4] when isolated and cultured from periodontal tissue of third
   molars. PDLSCs have been shown to have a positive effect on OTM through
   their multipotential differentiation capacity towards osteogenic,
   lipogenic and chondrogenic lineages similar to other mesenchymal stem
   cells, forming new periodontal support tissues to reconstruct the
   attachment relationship between cementum and alveolar bone [[41]4,
   [42]5]. Consequently, maintaining and promoting the properties of
   PDLSCs is essential for OTM.

   Exosomes are nanoscale vesicles that can be secreted by most cells,
   with a diameter of about 30–150 nm, and are widely present in various
   body fluids in the human body. Exosomes carry a variety of biologically
   active substances, such as proteins, lipids, DNA and RNA, which play an
   important role in intercellular communication and signal transduction,
   and are involved in the regulation of biological processes, including
   the immune response, cell proliferation and differentiation. [[43]6,
   [44]7]. Previous studies have demonstrated that human exfoliated
   deciduous teeth (SHED)-derived exosomes can promote osteogenic
   differentiation of PDLSCs through activation of Wnt and BMP signaling
   pathways [[45]8]. Bone marrow mesenchymal stem cells (BMSCs)-derived
   exosomes enhance migration, proliferation and differentiation of PDLSCs
   to promote periodontal regeneration [[46]9]. Mechanical strain-induced
   osteocyte-derived exosomes inhibit PTEN/AKT signaling pathway via
   miR-181b-5p to promote proliferation and osteogenic differentiation of
   human PDLSCs (hPDLSCs) by BMP2/Runx2 [[47]10]. It is indicated that
   multiple sources of exosomes contribute to the osteogenic
   differentiation of PDLSCs, and it is positive to deeply explore the
   effect of exosomes on PDLSCs for OTM treatment.

   Macrophages, as one of the main immune cells in the periodontal
   microenvironment, are involved in OTM. Macrophage polarization exists
   mainly in the state of classically activated macrophages (M1
   macrophages) and alternatively activated macrophages (M2 macrophages)
   [[48]11, [49]12]. Osteoclasts, which play a major role on the pressure
   side of the OTM process, are derived from monocytes/macrophages.
   Earlier studies have shown the presence of M1 macrophages on the stress
   side of OTM and that the proportion is significantly increased [[50]13,
   [51]14]. Notably, the interaction between macrophages and PDLSCs via
   exosomes may be indispensable for repair and regeneration of
   periodontal tissue and OTM. Previous studies have revealed that PDLSCs
   induce macrophages towards the M2 phenotype, and contribute to the
   enhancement of periodontal regeneration during the early stages of
   tissue repair [[52]15]. Liu et al. [[53]16] also showed that PDLSCs
   promote M2 polarization in macrophages via the JNK pathway.
   LPS-stimulated PDLSCs induce M1 polarization of macrophages via
   extracellular vesicles, which is detrimental to osteogenic
   differentiation [[54]17]. He et al. [[55]18] indicated that PDLSCs
   produce H[2]S in response to mechanical stimulation to promote
   macrophage polarization to the M1 phenotype, which contributes to bone
   remodeling and OTM. It is suggested that the regulation of macrophage
   polarization by PDLSCs is an important pathway for periodontal tissue
   regeneration and OTM. However, it remains to be investigated whether
   exosomes derived from different polarization states of macrophages have
   an effect on the osteogenic differentiation of PDLSCs.

   Based on the above studies, we hypothesized that exosomes derived from
   different polarization states of macrophages are involved in regulating
   the osteogenic differentiation process of PDLSCs. Furthermore, we
   analyzed the differentially expressed microRNAs (DE-miRNA) in M0
   macrophage-derived exosomes (M0-exo) and M2 macrophage-derived exosomes
   (M2-exo) by miRNA microarray. It aimed to elucidate the interaction
   mechanism between macrophages and PDLSCs in OTM, and to disclose the
   key factors.

Materials and method

Induction and identification of macrophage polarization

   THP-1 cells and complete medium were purchased from National Collection
   of Authenticated Cell Cultures (China; Cat. No.: TCHu 57). THP-1 cells
   were inoculated in complete medium and cultured in incubator conditions
   of 37 °C, 5% CO[2], and 90% humidity. THP-1 cells (1 × 10^6 cells /mL)
   were inoculated in 6-well plates and M0 macrophages were induced by
   stimulation with 100 ng/ml PMA (Solarbio, China) for 6 h. M0
   macrophages were incubated with 20 ng/ml IFN-γ and 100 ng/ml LPS
   (Solarbio) for 48 h to induce M1 macrophages. M0 macrophages were
   incubated with 20 ng/ml IL-4 and 20 ng/ml IL-13 (Solarbio) for 48 h to
   induce M2 macrophages. Macrophages were collected, and CD68 and CD80,
   CD16 and CD86, CD163 and CD206 (BD Biosciences, San Jose, CA, USA) were
   detected using a BD-FACSAria™ Fusion flow cytometer (BD Biosciences,
   San Jose, CA, USA) to identify M0, M1 and M2 macrophages as described
   previously, respectively [[56]19, [57]20].

Isolation and identification of macrophage-derived exosomes

   Briefly, after induction of THP-1 cells (1 × 10^6 cells /mL) into M0,
   M1 and M2 macrophages, RPMI-1640 complete medium was continued for 24 h
   and then replaced with serum-free RPMI-1640 medium for 24 h. M0
   macrophage-derived exosomes (M0-exo), M1 macrophage-derived exosomes
   (M1-exo) and M2 macrophage-derived exosomes (M2-exo) were prepared by
   differential centrifugation. Briefly, the supernatants of M0, M1 and M2
   macrophages were collected and centrifuged at 300 g for 10 min to
   remove cells, at 2000 g for 10 min to remove dead cells and at 10,000 g
   for 30 min with 4℃ to remove cell debris. Subsequently, the supernatant
   was transferred to an ultracentrifuge tube and centrifuged at 100,000 g
   for 90 min with 4℃ to precipitate exosomes. Finally, the exosomes were
   resuspended in pre-cooled PBS and centrifuged again at 100,000 g for
   90 min with 4℃ to obtain exosomes. Moreover, M0-exo, M1-exo and M2-exo
   were quantified using a spectrophotometer at 405 nm by referring to
   instructions of EXOCET Exosome Quantitation Assay Kit (System
   Biosciences, San Francisco, CA, USA). The concentrations of M0-exo,
   M1-exo and M2-exo were 5.32 (OD[405nm] = 0.09821), 5.96
   (OD[405nm] = 0.1059) and 4.36 (OD[405nm] = 0.0867) μg/μl, respectively.
   According to the standard curve (y = 0.0012x + 0.0176) conversion,
   1 × 10^6 THP-1 cells can be separated approximately 6.718 × 10^8
   M0-exo, 7.358 × 10^8 M1-exo and 5.758 × 10^8 M2-exo, respectively. The
   morphological features of M0-exo, M1-exo and M2-exo were observed by
   Talos F200X G2 TEM transmission electron microscopy (TEM; Thermo Fisher
   Scientific, Waltham, MA, USA) as previously described [[58]21].
   Expression of exosome markers CD9, TSG101 and ALIX were detected by
   Western blotting for M0-exo, M1-exo and M2-exo and the corresponding
   supernatants. After identification, the lipid dyes PKH67
   (Sigma-Aldrich) were used to label M0-exo, M1-exo and M2-exo by
   referring to the instructions.

Isolation and identification of primary hPDLSCs

   The hPDLSCs were separated from premolar teeth (15–30 years of age),
   which required extraction for orthodontic treatment. Inclusion
   criteria: (1) no systemic disease; (2) no history of smoking; (3) no
   history of long-term medication use; (4) healthy periodontal tissues.
   Exclusion criteria: (1) dental tissues with dental disease or
   inflammation; (2) patients refused to sign the informed consent form.
   All patient or guardian has signed an informed consent form, and the
   study has been approved by the Ethics Committee of Hospital of
   Stomatology, Kunming Medical University/Yunnan Stomatology Hospital
   (Approval Number: KYKQ2022MEC005). Primary culture of hPDLSCs was
   performed as described previously [[59]22], and 3–5 generations of
   hPDLSCs were used for subsequent studies. CD4, CD90, CD34 and CD45 were
   detected by flow cytometry (BD Biosciences) as previously described
   [[60]23, [61]24] to identify hPDLSCs. Lipogenic and osteogenic
   differentiation of hPDLSCs were induced as previously described
   [[62]24], and were identified using oil red O staining and Alizarin red
   S staining, respectively.

   After identification, osteogenic inducers (10–8 mol/L dexamethasone,
   50 μg/ml vitamin C and 10 mmol/L sodium β-glycerophosphate) induced
   osteogenic differentiation of hPDLSCs, and hPDLSCs were treated with
   10, 25, 50 and 100 μg/ml of M0-exo, M1-exo and M2-exo, respectively,
   for subsequent assays.

Alizarin red S staining (ARS)

   On day 21 of osteogenesis induction and exosome treatment, each group
   of hPDLSCs was fixed with 4% paraformaldehyde at room temperature for
   30 min and incubated with 1% alizarin red staining solution (Solarbio)
   at 37 ℃ for 30 min. Mineralized nodules were observed and photographed
   under an inverted microscope. The calcified nodules were lysed in
   alizarin red-stained cells by adding 500 μl of 1% cetylpyridine
   chloride solution for 1 h. Subsequently, 200 μl of this solution was
   used to measure the absorbance of the calcified nodules by
   spectrophotometry at 562 nm.

Western blotting assay

   RIPA buffer (Beyotime, China) and ProteoPrep® Total Extraction Sample
   Kit (Sigma-Aldrich, St. Louis, MO, USA) were used to lyse hPDLSCs
   (osteogenesis induction and exosome treatment at d 7 and 21) and
   exosomes, respectively, to obtain total protein. The concentration of
   hPDLSCs and total exosomal proteins was measured using BCA Protein
   Assay Kit (Beyotime). Equal amounts of proteins were added on SDS-PAGE
   and electrophoresis was performed, followed by transfer to PVDF
   membrane. The membrane was blocked with 5% BSA for 2 h at room
   temperature and the exosome-loaded membranes were incubated overnight
   at 4 °C with rabbit monoclonal antibodies CD9 (1:1000; ab236630),
   TSG101 (1:3000; ab125011) and ALIX (1:1000; ab275377). The hPDLSCs
   samples were incubated overnight with rabbit polyclonal antibodies to
   ALP (1:1000; ab229126) and OCN (1:2000; ab93876). The membrane was
   washed 3 times with PBST and incubated with Goat Anti-Rabbit IgG H&L
   (HRP) (1:5000; abcam; ab205718) for 1 h at 37 °C. β-actin was used as
   an internal reference. The ECL kit (Beyotime) was used for development
   and the bands on the membrane were scanned and imaged by E-Gel Imager
   gel imaging system (Thermo Fisher Scientific). All the above antibodies
   were purchased from Abcam plc (Cambridge, MA, USA). The results were
   quantified using Image J software.

RNA extraction and RT-PCR assay

   Total RNA in hPDLSCs was extracted using Trizol reagent (Thermo Fisher
   Scientific). Extraction of miRNA from exosomes by Qiagen miRNeasy Mini
   Kit (QIAGEN, Duesseldorf, Germany). Total RNA was reverse-transcribed
   to cDNA according to the instructions of SuperScript First-Strand
   Synthesis System (Thermo Fisher Scientific), and RT-qPCR was performed
   by SYBR GreenMaster Mix (Thermo Fisher Scientific) to detect the
   expression of mRNA and miRNA. The primers for mRNA and miRNA are shown
   in Table [63]1. β-actin and U6 were used as internal references for