Periodontal and Dental Pulp Cell-Derived Small Extracellular Vesicles: A Review of the Current Status

Extracellular vesicles (EVs) are membrane-bound lipid particles that are secreted by all cell types and function as cell-to-cell communicators through their cargos of protein, nucleic acid, lipids, and metabolites, which are derived from their parent cells. There is limited information on the isolation and the emerging therapeutic role of periodontal and dental pulp cell-derived small EVs (sEVs, <200 nm, or exosome). In this review, we discuss the biogenesis of three EV subtypes (sEVs, microvesicles and apoptotic bodies) and the emerging role of sEVs from periodontal ligament (stem) cells, gingival fibroblasts (or gingival mesenchymal stem cells) and dental pulp cells, and their therapeutic potential in vitro and in vivo. A review of the relevant methodology found that precipitation-based kits and ultracentrifugation are the two most common methods to isolate periodontal (dental pulp) cell sEVs. Periodontal (and pulp) cell sEVs range in size, from 40 nm to 2 μm, due to a lack of standardized isolation protocols. Nevertheless, our review found that these EVs possess anti-inflammatory, osteo/odontogenic, angiogenic and immunomodulatory functions in vitro and in vivo, via reported EV cargos of EV–miRNAs, EV–circRNAs, EV–mRNAs and EV–lncRNAs. This review highlights the considerable therapeutic potential of periodontal and dental pulp cell-derived sEVs in various regenerative applications.


Introduction
Extracellular vesicles (EVs) are membrane-bound bilayered lipid particles that are secreted from both prokaryotic and eukaryotic cells, carrying a cargo of biological molecules (i.e., protein, nucleic acid, lipids and metabolites) from their parent cells [1]. Initially, EVs were considered 'cellular dust', generated by cellular metabolism, until their biological role in the mineralization of bone was recognized [2,3]. A principal role of EVs is as an intercellular communicator of biological information into a recipient cell. This interaction can trigger signaling cascades and modulate cell behavior [4]. The biological function of EVs is defined by the parent cells from which they originate. EVs are involved in almost all cellular interactions, especially tumor metastasis, tissue homeostasis, and inflammatory regulation [4,5]. Due to their constituent biological molecules, EVs hold great promise as a therapeutic delivery system in regenerative medicine.
The definition, terminology and subtypes of EVs are still being debated. The International Society of Extracellular Vesicles (ISEV) recommends a division of EV subtypes based on their size: medium/large EV (>150 nm) and small EV (<150 nm) [6]. However, considering the discrepancies in the published literature, for simplification purposes, this review will define EV subtypes based on both their size and biogenesis ( Figure 1a): small extracellular vesicles (also known as exosomes) (sEVs, <200 nm), microvesicles (MVs,

Small EV (Exosomes)
Small EVs (sEV), or exosomes, originate from endosomes, and are biological nanoparticles that are smaller than 200 nm [5]. Further, sEVs are produced through an endocytic pathway, and their particle size is partially overlaid with that of microvesicles and apoptotic bodies. The biogenesis process for sEV is unique, whereby the endosomal network is the source of sEV that produce, classify, distribute and define the proper destination of the secreted sEV [2,8]. Endosome production can be categorized into the following three subtypes, according to each stage of development: early endosomes, late endosomes, and recycling endosomes. Early endosomes are formed by inward budding of the cell membrane, before a second inward budding of the endosomal membrane that results in the formation of late endosomes-intraluminal vesicles (ILVs). Late endosomes containing IVLs are named multi-vesicular bodies (MVBs), and the MVBs either fuse with lysosomes to degrade or follow the endocytic pathway for sEV generation. Once fusion with the plasma membrane is completed, the small membrane-enclosed vesicles are released into the extracellular matrix.
The biogenesis of sEV is affected by the following two main pathways that can induce multi-vesicular bodies (MVBs) generation: the endosomal sorting complex, required for the transport (ESCRT)-dependent pathway and ESCRT-independent pathway [9]. For the ESCRT-dependent pathway, ESCRT I and ESCRT II mediate the invagination of the late endosomal membrane, and ESCRT III will be recruited to the invaginated membrane sites. The cargo proteins are then deubiquitinated, and this stimulates the departure of the vesicle and the formation of MVBs. In the ESCRT-independent pathway, the neutral sphigo-mylinase2 (nSMase2) takes sphingolipids as substrates and converts sphingolipids to ceramide at the endosomal membrane. Following this, the microdomain is prepared for merging into a larger structure, which accelerates the endosomal budding and biogenesis of MVBs [10]. Moreover, sEVs that are produced by these different pathways possess different biomarkers, except CD63, which is the most common biomarker for all sEVs [11,12]. With respect to the ESCRT-dependent pathway, if endocytosis is mediated by Ras-related protein 27A/B (RAB27A/B), TSG101 is a biomarker of sEV. If the endocytosis is mediated by phospholipase D2 (PLD2) and RAB7, through the ESCRT-independent pathway, the

Small EV (Exosomes)
Small EVs (sEV), or exosomes, originate from endosomes, and are biological nanoparticles that are smaller than 200 nm [5]. Further, sEVs are produced through an endocytic pathway, and their particle size is partially overlaid with that of microvesicles and apoptotic bodies. The biogenesis process for sEV is unique, whereby the endosomal network is the source of sEV that produce, classify, distribute and define the proper destination of the secreted sEV [2,8]. Endosome production can be categorized into the following three subtypes, according to each stage of development: early endosomes, late endosomes, and recycling endosomes. Early endosomes are formed by inward budding of the cell membrane, before a second inward budding of the endosomal membrane that results in the formation of late endosomes-intraluminal vesicles (ILVs). Late endosomes containing IVLs are named multi-vesicular bodies (MVBs), and the MVBs either fuse with lysosomes to degrade or follow the endocytic pathway for sEV generation. Once fusion with the plasma membrane is completed, the small membrane-enclosed vesicles are released into the extracellular matrix.
The biogenesis of sEV is affected by the following two main pathways that can induce multi-vesicular bodies (MVBs) generation: the endosomal sorting complex, required for the transport (ESCRT)-dependent pathway and ESCRT-independent pathway [9]. For the ESCRT-dependent pathway, ESCRT I and ESCRT II mediate the invagination of the late endosomal membrane, and ESCRT III will be recruited to the invaginated membrane sites. The cargo proteins are then deubiquitinated, and this stimulates the departure of the vesicle and the formation of MVBs. In the ESCRT-independent pathway, the neutral sphigomylinase2 (nSMase2) takes sphingolipids as substrates and converts sphingolipids to ceramide at the endosomal membrane. Following this, the microdomain is prepared for merging into a larger structure, which accelerates the endosomal budding and biogenesis of MVBs [10]. Moreover, sEVs that are produced by these different pathways possess different biomarkers, except CD63, which is the most common biomarker for all sEVs [11,12]. With respect to the ESCRT-dependent pathway, if endocytosis is mediated by Ras-related protein 27A/B (RAB27A/B), TSG101 is a biomarker of sEV. If the endocytosis is mediated by phospholipase D2 (PLD2) and RAB7, through the ESCRT-independent pathway, the biomarkers of sEV are alix, syntenin, and syndecan. As for the RAB11/35-mediated ESCRT-independent pathway, CD81, Wnt and proteolipid protein (PLP) are the preferred biomarkers.
The function of sEV in intercellular communication is determined by the interconnection between sEV surface proteins and receptors on the recipient cells that subsequently activates a variety of signaling pathways [5]. Further, sEVs arising from different cell types have different cargos that dictate and direct different biological effects. The sEVs are highly abundant in biofluids [13][14][15], and they have been demonstrated to be associated with immune response, viral pathogenicity, osteogenesis, odontogenesis, neuroprotection, angiogenesis, and anti-tumor functions [16]. For example, oral cancer cell-derived sEVs create a mechanism that can promote tumor progression by modifying vesicular contents and establishing a distant premetastatic niche with molecules that favor cancer cell proliferation, migration, invasion, metastasis, angiogenesis, and even drug resistance [17]. Evidence that sEVs play an important role in cell differentiation suggests that sEVs may have a potential role in tissue regeneration.

Microvesicles
Microvesicles (MVs) are membrane vesicles of different sizes, surrounded by a lipid layer of membrane, and they range in size from 50 nm to 1 µm. Microvesicles are generated by the outward budding of the plasma membrane, and are abundant in tissues/cells and biofluids [18]. The contents of MVs are similar to that of sEV. The MV components of note include CD40, selectins, integrins, cytoskeletal proteins, and cholesterol [19].
The biogenesis of MV involves the contraction of cytoskeletal proteins and phospholipid redistribution, contributing to a dynamic interplay in the plasma membrane and the resultant formation of microvesicles. Within the plasma membrane, the aminophospholipid translocase regulates phospholipid distribution, transferring phospholipids from one leaflet to another. Once phosphatidylserine (PS) is translocated to the leaflet of the outer membrane, the outward blebbing of the membrane and microvesicle formation is initiated. The interaction between actin and myosin causes the cytoskeletal structure contraction, which mediates membrane budding [20].
MVs have been reported to maintain tissue homeostasis during tissue regeneration, angiogenesis, anti-tumor effects, and in pathologies such as tumorigenesis, chronic inflammation, and atherosclerosis [19]. MVs that are produced by blood cells (e.g., neutrophils, macrophages, and platelets) are involved in the pro-coagulatory response [21]. MVs can be both pro-inflammatory and anti-inflammatory; this is determined by the induction or stimulation that is received by their parent cells. MVs that are produced by tumor cells enhance invasiveness and accelerate cancer progression, as well as strengthen the drug resistance of tumor cells [22]. This indicates that MVs are potential therapeutic agents for tissue regeneration; however, the function of MVs in periodontal tissue healing and regeneration requires further investigation.

Apoptotic Bodies
Apoptotic bodies (ApoBDs) are produced by cells undergoing apoptosis, and vary in size from 50 nm to 2 µm [23,24]. ApoBDs result from the formation of subcellular fragments when an apoptotic cell disassembles. They are comprised of molecular components from living cells and provide a rich molecular pool for recipient cells. However, ApoBDs are engulfed by macrophages and digested by phagolysosomes shortly after they are released [25]. ApoBDs and apoptosis are not related to an inflammatory reaction, the constituents in dying cells and ApoBDs are not released automatically to the environment, and anti-inflammatory cytokines are not generated during engulfing. ApoBDs have phosphatidylserine (PS) on their surfaces, to attract engulfing cells, and are considered to be specific biomarkers for ApoBDs [26]. Autoimmune diseases may be associated with defects in the clearance of ApoBDs. ApoBDs may stimulate the formation of thrombus and improve anti-cancer immunity.
Increasing evidence suggests that ApoBDs have important immune regulatory roles, in autoimmunity, cancer, and infection [24], as well as promoting osteogenesis [27]. For example, ApoBDs that are derived from mature osteoclasts can induce osteoblast differentiation by activating the protein kinase B/phosphoinositide 3-kinases (PI3K/AKT) pathway [27]. However, knowledge of their function and role is still limited and more studies are required in this field.

The Source and Characteristics of Periodontal (Dental Pulp) Cells
Dental tissue-derived (or stem) cells have remarkable characteristics for therapeutic application, being easily accessible and a rich source of stem cells with a well-known regenerative capacity. A great variety of multipotent adult or postnatal stem cells can be retrieved from dental tissues, especially from periodontal tissue and dental pulp from extracted permanent teeth (dental pulp stem cells-DPSCs) and exfoliated deciduous teeth (SHED). A healthy periodontium consists of soft (periodontal ligament-PDL and gingiva) and hard (alveolar bone and cementum) tissue, and cells residing within the healthy periodontal tissues include periodontal ligament (stem) cells (PDLSCs), PDL and gingival fibroblasts (PDLF, GFs), or gingival stem cells (GMSCs), osteoblasts (OBs), osteoclasts (OCs), and various immune cells ( Figure 2) [28,29]. Moreover, stem cells can be obtained from dental apical papilla tissues (SCAP) and dental follicles (DFSCs, or DFCs) of the developing tooth [28,29]. Importantly, EV that is derived from these cells can be detected within periodontal tissues and biofluid (i.e., gingival crevicular fluid) ( Figure 2). specific biomarkers for ApoBDs [26]. Autoimmune diseases may be associated with defects in the clearance of ApoBDs. ApoBDs may stimulate the formation of thrombus and improve anti-cancer immunity. Increasing evidence suggests that ApoBDs have important immune regulatory roles, in autoimmunity, cancer, and infection [24], as well as promoting osteogenesis [27]. For example, ApoBDs that are derived from mature osteoclasts can induce osteoblast differentiation by activating the protein kinase B/phosphoinositide 3-kinases (PI3K/AKT) pathway [27]. However, knowledge of their function and role is still limited and more studies are required in this field.

The Source and Characteristics of Periodontal (Dental Pulp) Cells
Dental tissue-derived (or stem) cells have remarkable characteristics for therapeutic application, being easily accessible and a rich source of stem cells with a well-known regenerative capacity. A great variety of multipotent adult or postnatal stem cells can be retrieved from dental tissues, especially from periodontal tissue and dental pulp from extracted permanent teeth (dental pulp stem cells-DPSCs) and exfoliated deciduous teeth (SHED). A healthy periodontium consists of soft (periodontal ligament-PDL and gingiva) and hard (alveolar bone and cementum) tissue, and cells residing within the healthy periodontal tissues include periodontal ligament (stem) cells (PDLSCs), PDL and gingival fibroblasts (PDLF, GFs), or gingival stem cells (GMSCs), osteoblasts (OBs), osteoclasts (OCs), and various immune cells ( Figure 2) [28,29]. Moreover, stem cells can be obtained from dental apical papilla tissues (SCAP) and dental follicles (DFSCs, or DFCs) of the developing tooth [28,29]. Importantly, EV that is derived from these cells can be detected within periodontal tissues and biofluid (i.e., gingival crevicular fluid) ( Figure 2). Schematic showing the main cell population and cell products (EVs) within a healthy periodontium. Various cells reside in the periodontium, such as periodontal ligament (stem) cells (PDLSCs), fibroblasts (GFs) and stem cells (GMSCs) from the gingiva, osteoblasts (OBs), osteoclasts (OCs), and various immune cells. AB: alveolar bone; C: cementum; D: dentin; DP: dental pulp; PDL: periodontal ligament; SHED: dental pulp cells from human exfoliated deciduous teeth (SHED); SCAP: cells from periodontal apical papilla tissues (SCAP). Schematic showing the main cell population and cell products (EVs) within a healthy periodontium. Various cells reside in the periodontium, such as periodontal ligament (stem) cells (PDLSCs), fibroblasts (GFs) and stem cells (GMSCs) from the gingiva, osteoblasts (OBs), osteoclasts (OCs), and various immune cells. AB: alveolar bone; C: cementum; D: dentin; DP: dental pulp; PDL: periodontal ligament; SHED: dental pulp cells from human exfoliated deciduous teeth (SHED); SCAP: cells from periodontal apical papilla tissues (SCAP).
Dental mesenchymal stem cells originate from the neural crest ectomesenchyme and reside in stromal niches (perivasculature and peripheral nerve-associated glia cells). The current consensus holds that both perivascular cells [30] and glia cells [31] are responsible for dental MSCs origin, as revealed in mouse experiments [31]. Much like bone-marrowderived MSCs that originate from mesoderm [32], dental stem cells express MSCs markers and exhibit multipotent linage regeneration (i.e., osteogenic, chondrogenic, neurogenic) and immunomodulatory capabilities. These properties make these cells suitable candidates for therapeutic application (reviewed by Chalisserry et al. in [33]) in neurological disorders, angiogenesis, dentin-pulp regeneration and periodontal regeneration. PDLSCs, GFs, DPSCs, SHED, and DFSCs have been demonstrated to promote multiple-tissue regeneration, both in vitro and in vivo [34][35][36][37][38][39][40]. However, cell therapy has several challenges, including high cost, insufficient cell number, and associated regulatory barriers. On the other hand, a cell-free approach, centered around cell products (i.e., EVs derived from these cells), has been proposed, and there is an emerging focus on cell-derived EVs as potential therapeutic agents to promote periodontal regeneration. The utilization of sEVs for dental tissue regeneration is emerging as a viable cell-free treatment option, with 'proof of concept' studies reported using bone marrow or adipose MSC-derived sEVs (reviewed in [41][42][43]); yet, periodontal or dental pulp cell sources are likely to uniquely reflect the functional complexity of the periodontium and oral cavity.
The following sections will summarize the current methods for cell-derived sEV isolation and characterization, with particular emphasis on sEVs from periodontal and dental pulp cells.

General Concepts
Although sEVs have been studied for decades, there is still no standardized protocol for their isolation. Despite the presence of recommended guidelines for EV isolation and characterization, such as the Minimal Information for Studies of Extracellular Vesicles 2014 (MISEV2014) and MISEV 2018, these guidelines are not always followed.
Prior to the isolation of sEV, sequential centrifugation is commonly used to remove cell debris and large EVs, as follows: i Step 1: the cell conditional media (CM) is harvested and centrifuged at 300-400× g to remove cells, and the supernatant (SN) is collected; ii Step 2: the SN collected in step 1 is centrifuged at 2000-3000× g to remove cells debris and apoptotic bodies. The SN is collected from this step; iii Step 3: SN from step 2 is centrifuged at 10,000-20,000× g to remove the aggregates of biopolymers, microvesicles, and the other structures with a buoyant density higher than sEVs. The SN is collected from this step; iv Step 4: then, the following isolation methods are used to enrich the sEVs: ultracentrifugation, sucrose gradient centrifugation, size exclusion chromatography, precipitationbased isolation, immunoaffinity chromatography, and ultrafiltration.
Given the growing interest in EVs, technical standardization is critical, as many different methodologies have been utilized for isolation and analysis. The influence of these various techniques on the downstream composition and functionality of EV cargos remains unclear; accordingly, the ISEV position papers [6,44] have raised the need to define 'good practices' and ultimately archive standardization. However, many researchers are not following these four steps, due to a lack of standardized protocols. Here, our review briefly introduces each isolation method, and discusses its merits and disadvantages (listed in Table 1).

Ultracentrifuge
Ultracentrifugation is the gold-standard method for isolating sEV, as the equipment is relatively easy to access and the methodology is technically straightforward. The method involves an ultracentrifugation step at 100,000×-200,000× g to pellet sEV [45]. However, ultracentrifugation has disadvantages, in that it leads to a low recovery rate of sEV, it is time consuming (1.5-10 h), contains non-vesicular macromolecule contamination, and results in EV aggregation.

Floatation-Related Methods (Sucrose Gradient Centrifugation)
Floatation-related methods distribute molecules based on the buoyant density, and the protein aggregates and sEV can be sufficiently separated. Differential gradient centrifugation (usually takes 250 min-1 day) takes advantage of buoyant density to fractionate EVs using sucrose or idoxanol gradients [45]. The sEVs can be separated by the discontinuous gradient sucrose solution, with each layer containing the desired size of EV. Other chemical reagents (i.e., iodixanol) can also be utilized instead of sucrose, for continuous EV harvest with no layers. Non-vesicular protein contaminants are distributed at a reduced level within this method, resulting in less protein contamination. However, sucrose gradient centrifugation cannot separate large particles that have a similar sedimentation rate.

Size Exclusion Chromatography (SEC)
SEC can be used to isolate small sEV, based on the size of the molecules, where large particles pass through the gel earlier than the small-sized molecules. The small-sized particles are trapped in the tiny pores on the surface of the gel, while the larger molecules can bypass the gel or receive less interference from the gel [46]. This technique has been well established with commercialized SEC columns, including qEV (iZON Science), Exo-spin™ SEC columns (Cell Guidance Systems Ltd.) and Pure-EVs SEC columns (HandaBioMed, Lonza). SEC has been proposed as an effective alternative method for pure sEV isolation, Nanomaterials 2021, 11, 1858 7 of 34 with a key advantage being its time efficiency (~30 min, including 10 mL of column washing with PBS). However, the similarly sized sEV and microvesicles cannot be separated by SEC.

Precipitation-Based Isolation (Sodium Acetate, PEG, Protamine)
Precipitation-based isolation has the following two mechanisms: polymeric precipitation and neutralizing charges [47]. In polymeric precipitation, a soluble polymer, usually polyethylene glycol (PEG), is mixed with EV samples and the mixture is incubated overnight, and EVs are sedimented by low-speed centrifugation at 1500 g. PEG precipitation enables a simple process for a large number of samples. Commercial kits, such as ExoQuick (System Biosciences), total exosome isolation reagent (Invitrogen), EXO-Prep (HansaBioMed), exosome purification kit (Norgen Biotek), and miRCURY exosome isolation kit (Exiqon), are based on this principle. For the other precipitation method, all EVs possess negative charges, so positively charged molecules (i.e., sodium acetate and protamine) are chosen for the precipitation. This method is popular due to its straightforward protocol; however, these precipitation methods lead to low sEV purity due to co-precipitation of the components from CM or biofluids, such as protein, DNA and RNA, and hence further purification is required.

Immunoaffinity Chromatography
The monoclonal antibodies (mAbs) against specific sEV surface proteins (i.e., CD 9) are fixed on the column, to capture a specific sEV population [48]. Once the CM passes through the column, the EVs, which express certain exosomal markers on their membrane, will be captured by the mAbs. This method leads to a very pure EV population, but low yield and scalability. This is attributed to the fact that this step needs to be repeated several times to ensure the mAbs can capture sufficient EVs (~240 min).

Ultrafiltration
Semi-permeable membranes (ranging from 3 kDa to 100 kDa) are adapted for sEV fractionation within filtration-based isolation; the membrane function is determined by its pore size. However, sEVs cannot be fractionated according to their biogenesis or biomarkers, but it is normally used to concentrate sEVs. It is still an efficient way to eliminate the minimal sample volume (~130 min) with a simple procedure, and has been proven to yield higher recovery of sEVs than ultracentrifugation [49]. However, ultrafiltration might lead to low EV protein, but a rather higher concentration of non-EV proteins (i.e., albumin).

Current Isolation Challenge
As mentioned previously, the current challenges of sEV isolation include time-consuming procedures, impurities, insufficient EV yield, and low scalability [50]. Although many researchers have investigated combinations of these isolation methods, an urgent demand has arisen to investigate high-yielding and time-effective isolation protocols. Currently, there is no optimal sEV isolation method; however, a combination of ultracentrifugation, SEC, and ultrafiltration has been used for the pure sEV population, which is a critical factor for downstream therapeutic applications.

sEV Isolation and Characterization Methods for Periodontal (and Dental Pulp) Cells
To date, there are no standardized protocols for sEV isolation and characterization. From the 33 studies that are reported in this review, we have summarized periodontal (dental pulp) cell-derived sEV isolation and characterisation methods . Various isolation methods have been used for periodontal (dental pulp) cells sEVs, including ultracentrifugation (UC), precipitation-based methods, and ultrafiltration ( Figure 3a). Regarding EV characterization, the latest MISEV 2018 guidelines [6] suggest that all EV researchers should characterize sEV from at least three different aspects, such as EV particle numbers, EV Nanomaterials 2021, 11, 1858 8 of 34 morphology, and EV-enriched protein markers. However, most of the current studies did not follow the MISEV guidelines, and this requires additional attention for all EV research.
(dental pulp) cell-derived sEV isolation and characterisation methods . Various isolation methods have been used for periodontal (dental pulp) cells sEVs, including ultracentrifugation (UC), precipitation-based methods, and ultrafiltration ( Figure 3a). Regarding EV characterization, the latest MISEV 2018 guidelines [6] suggest that all EV researchers should characterize sEV from at least three different aspects, such as EV particle numbers, EV morphology, and EV-enriched protein markers. However, most of the current studies did not follow the MISEV guidelines, and this requires additional attention for all EV research. Different sEV isolation methods have been utilized for various cells (Figure 3a), with precipitation and ultracentrifugation methods being the two most commonly used techniques. In PDL(S)C-derived sEV isolation (10 studies), the precipitation-based method (i.e., a commercial ExoQuick kit) is the most commonly used (n = 6, 60%), followed by ultracentrifugation (n = 4, 40%). Among six studies in GFs/GMSC-derived sEVs, the precipitation-based method (n = 4, 66.7%) and ultrafiltration (n = 2, 33.3%) were used for GFs/GMMSCs-sEV isolation. Regarding DPSC-derived sEV, most researchers selected ultracentrifugation (n = 7, 77.8%), with one study using the precipitation-based method (n = 2, 22.2%). For SHED-sEVs, all of the studies (n = 4, 100%) used the ultracentrifugation method to isolate sEVs from SHED.
In summary, ultracentrifugation and precipitation-based methods are the two most common methods used for periodontal (dental pulp) cells sEV isolation. WB, TEM and Different sEV isolation methods have been utilized for various cells (Figure 3a), with precipitation and ultracentrifugation methods being the two most commonly used techniques. In PDL(S)C-derived sEV isolation (10 studies), the precipitation-based method (i.e., a commercial ExoQuick kit) is the most commonly used (n = 6, 60%), followed by ultracentrifugation (n = 4, 40%). Among six studies in GFs/GMSC-derived sEVs, the precipitation-based method (n = 4, 66.7%) and ultrafiltration (n = 2, 33.3%) were used for GFs/GMMSCs-sEV isolation. Regarding DPSC-derived sEV, most researchers selected ultracentrifugation (n = 7, 77.8%), with one study using the precipitation-based method (n = 2, 22.2%). For SHED-sEVs, all of the studies (n = 4, 100%) used the ultracentrifugation method to isolate sEVs from SHED.
In summary, ultracentrifugation and precipitation-based methods are the two most common methods used for periodontal (dental pulp) cells sEV isolation. WB, TEM and NTA are the most common methods for periodontal (dental pulp) cell-derived sEVs characterisation.
According to the latest MISEV 2018 guidelines [6], it is critical to consider several factors influencing the collection of EV, including characteristics of primary cell source (donor health status, age, gender), primary cell passage number, confluence at harvest, culture volume, media change frequency, CM harvesting conditions, as well as all culture media composition and preparation details. Thus, our review includes detailed donor information for primary cells (if mentioned), the cell culture condition prior to CM collection, and detailed sEV isolation protocols. This will allow future EV researchers to select appropriate protocols for CM harvesting and sEV isolation.
The following three aspects of hDPL(S)Cs-sEV analysis were evaluated in the 10 studies that have been included in this review: (1) EV size, (2) EV content (protein, RNA, etc.), and (3) EV function in cell differentiation in vitro and in vivo. Regarding the size of hPDL(S)Cs-sEV, three studies did not characterize the sEV size [52,53,60]. There is a large deviation for the reported EV size: <200 nm in five studies [51,54,55,57,58], two populations (90 ± 20 nm and 1200 ± 400 nm) in one study [59], and 100-710 nm in one study [56]. It is noted that two studies engineered the hPDLCs-EV using polyethyleneimine (PEI, yielding PEI-EV) [56,59]. The following two factors may contribute to this deviation: the EV isolation method (UC or precipitation methods), and the EV size characterization methods (TEM, or DLS, or NTA). We will define EV size <200 nm as sEV, and unclear EV size as EV.
The size of sEV derived from microcarrier cell cultures of PDLSCs: 112-182 nm. • hPDLSCs-sEV suppressed basal and LPS-induced activity of NF-kB in PDLSCs.

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Combined treatment with EV and anti-TLR4 antibody attenuated the inhibitory effect on the NF-kB activity • Uptake of hPDLSCs-sEV in hPDLSC-activated phosphorylation of Akt and GSK3β (Ser 9) indicating that PI3K/Akt signaling pathway may act as a suppressor of NF-kB activity • EVs did not significantly affect osteogenic mineralization of hPDLSCs cultures, but EV significantly increased expression of ALP, OPN, BSP, and CP23 gene expression, but downregulated BMP2 expression on the 10 days of osteogenic differentiation.
There is increasing evidence demonstrating that hPDL(S)Cs-EV can modulate in vitro angiogenesis (in HUVECs [52]), osteogenesis (in MG-63 OBs [54] and hPDLSCs [55,56,59]), anti-inflammation (in LPS-treated hPDLSCs [55,60] and J774.1 macrophages [57]), and immunoregulation (induced M1 polarization in THP-1 cells [58]) via modulating the TGF-beta pathway, MAPK pathway, mTOR pathway and FoxO signaling pathways [51], and PI3K/Akt signaling [55] and NF-kB signaling pathways [57]. The in vivo function of hPDL(S)Cs-EV was explored, either in rat calvaria defect [56,59] or intravenous administration in mouse multiple sclerosis disease [60] models. Pizzicannel-la et al. [56] created a calvarial defect, with a diameter of 4 mm and a height of 0.25 mm in male Wistar rats (300-350 g; n = 4 for each group). The hPDLSCs-EV and hPDLSCs-PEI-EV were loaded on collagen membranes and transplanted into the rat calvaria defect for 6 weeks, leading to enhanced bone and vascularization compared to the no-EV groups, with the PEI-EV group inducing better osteogenesis and vascularization compared to the EV group. Diomede et al. [59] revealed similar results, showing that hPDLSCs-PEI-EV leads to increased blood vessel formation after 6 weeks of the transplantation of hPDLSCs-EV-and hPDLSCs-PEI-EV-loaded collagen membranes into a rat calvarial defect. Rajan et al. [60] established a mouse model of MS disease, and intravenous administration of hPDLSCs-EVs decreased apoptosis and inflammation in the diseased mice.
In their in vivo investigations, Nakao et al. [61] created a ligature-induced periodontitis mice model, and locally injected hGMSCs-sEV or TNF-α-preconditioned GMSC-derived exosomes (hGMSCs-sEV-TNF) into the palatal gingiva of the ligated second maxillary molar. One week post-injection, both the interventions significantly reduced periodontal bone loss compared to the PBS control group, while hGMSCs-sEV-TNF further reduced the distance from the cementoenamel junction to the alveolar bone crest (CEJ-ABC) and the number of tartrate-resistant acid phosphatase (TRAP)-positive osteoclasts, indicating an anti-osteoclastic property for hGMSCs-sEV [61]. Moreover, Mao et al. [66] transplanted hGMSCs-sEV-loaded gelfoam sheets into the crush-injury sites of sciatic nerves in C57BL/6J mice, and the EV group had comparable beneficial effects on the functional recovery of the injured sciatic nerves of mice compared to the hGMSCs group. Further, hGMSCs-sEV enhanced the expression of neuronal and Schwann cell markers (β-tubulin III and S100 calcium-binding protein B-S100B) at one-month post-injury, compared with hGMSCs controls, suggesting that hGMSCs-sEV can promote neuron regeneration in vivo. Diomede et al. [67] loaded hGMSCs-EV or hGMSCs-PEI-EV into 3D-printed PLA scaffolds with/without hGMSCs, and transplanted them into rat calvaria defects for 6 weeks. Both the hGMSCs-EV and hGMSCs-PEI-EV groups enhanced bone and blood vessel formation, yet hGMSCs-PEI-EV performed better than the EV group.

Human Dental Pulp Cells (hDPSCs)-sEV
Ivica et al., 2020 [70] • hDPSCs from healthy third molars (n = 3, 16 to 25 years old) were extracted • Human bone marrow-derived mesenchymal stem cells (HBMMSCs). Passage 3-6 • Exosome-free medium (Invitrogen) was used for hDPSCs culture for 48 h before CM collection. • CM centrifuged at 300× g for 10 min; 2000× g for 30 min; total exosome isolation agent (Invitrogen) was added; the mixture was centrifuged at 10,000× g for 30 min • hDPSC-sEV ranged from 45 to 156 nm by TEM, with a mean size of 90 nm. • hDPSC-sEV encapsulated in fibrin gel promoted hBMMSCs migration and proliferation.   • hPDSCs were isolated from healthy dental pulp tissues and (n = 8) from the caries-free teeth that need to be extracted due to orthodontics • hBMMSCs were isolated from bone marrow aspirates of healthy people (n = 8).
• At 80% confluence, the medium was replaced with a serum-free medium for 48 h before CM collection.

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The CM was centrifuged at 300× g for 10 min, centrifuged at 16,500× g for 30 min at 4 • C; passed through a 0.2 µm; ultracentrifuged at 4 • C at 100,000× g for 70 min; ultracentrifuged again at 4 • C at 100,000× g for 70 min  RNA sequencing analysis showed that 28 microRNAs significantly changed in OD-Exo isolated under odontogenic conditions, of which 7 miRNAs increased and 21 miRNAs decreased. The qRT-PCR analysis showed that miR-5100 and miR-1260a levels in OD-Exo increased, while miR-210-3p and miR-10b-5p decreased, which were consistent with the miRNA sequencing. GO analysis showed that the differentially expressed miRNAs are associated with the TGFβ1 pathway. • OD-exo activated the TGFβ1 pathway by upregulating TGFβ1, TGFR1, p-Smad2/3, and Smad4 in DPSCs, compared to the control group and UN-Exotreated group. • Compared with UN-Exo, miR-27a-5p was expressed 11 times higher in OD-Exo.
With respect to in vivo studies, Zhou et al. [69] created a full-thickness excisional skin wound-healing model in male C57BL/6 mice (8 weeks old), and then subcutaneously injected hDPSCs-sEV (200 µg in 100 µL) from healthy or periodontitis patients derived hDPSCs-sEV (200 µg in 100 µL PBS) for 4, 9, and 14 days. Both the sEV groups promoted the wound healing process and vascularization compared to the PBS control group, while hDPSCs-sEV from the periodontitis patients increased the wound closure rate and the number of newly formed microvessels, with more CD31-and VEGF-positive cells compared to the sEV from a healthy patient. Shen et al. [72] established a ligation-induced periodontitis model in 6-8-week-old male C57BL/6J mice, and a chitosan hydrogel (CS) loaded with 50 µg of hDPSCs-sEV (hPSDCs-sEV-CS group) was locally injected after ligature removal, with a local injection of PBS or hPSDCs-sEV used as the controls. The results showed that the hDPSCs-sEV-CS group led to increased bone formation, a thick layer of epithelial layers, less inflammatory cells, and a lower amount of TRAP-positive osteoclasts, at 10 days post-treatment. Furthermore, hPSDCs-sEV-CS treatment significantly reduced pro-inflammatory cytokines (IL-23, IL-1α, TNF-α, IL-12, IL-1β, IL-27, and IL-17), and NF-κB p65 and p38 MAPK signaling, in periodontal tissues compared with other groups. RNA sequencing analysis of the periodontium showed that 7351 differentially expressed genes (DEGs) were found between the hDPSCs-sEV-CS and CS groups. GO term enrichment analysis of the top 200 DEGs demonstrated that they are associated with chemotaxis pathways and the immune response, which were downregulated in the hDPSCs-sEV-CS group. Most importantly, hDPSCs-sEV-CS induced macrophages converting from a proinflammatory phenotype to an anti-inflammatory phenotype in the periodontium of periodontitis mice, with more CD206+ anti-inflammatory macrophages and significantly decreased CD86+ in pro-inflammatory macrophages [72]. This indicates that hDPSCs-sEV can promote bone formation, epithelium re-growth, and reduce inflammation in a periodontitis mice model. Furthermore, Huang et al. [76] loaded hDPSCs-EV into clinicalgrade type I collagen membranes, and then placed them on a human tooth root slice (3-4 mm in thickness), before subcutaneously transplanting into athymic nude mice for 2 weeks. They resulted in enhanced dental-pulp-like tissues, with increased odontogenic proteins (dentin matrix acidic phosphoprotein 1-DMP1, and dentin phosphophoryn-DSPP) and endothelial cell marker protein (von Willebrand factor-vWF).
The function of SCAP-sEVs was investigated in vivo on gingival soft tissue [77] and dentin-pulp regeneration [78]. Liu et al. [77] created full-thickness circular gingival wounds in C57BL/6J mice, using a biopsy punch (soft tissue defects with a diameter of 2.0 mm). Following this, 40 µg of SCAP-sEVs, SCAP-siCdc42-sEVs, or PBS, was injected submucosally into the palates of the wounds sites. Seven days post-injection, SCAP-sEVs promoted palatal gingival tissue regeneration by enhancing vascularization in the early phase [77]. Zhuang et al. [78] loaded 50 µg/mL SCAP-sEVs and 4 × 10 5 BMMSCs with gelatin sponge onto a dentin slice, before subcutaneously transplanting them into immunodeficient mice. Significant dentin-pulp regeneration was observed 12 weeks post-transplantation in the SCAP-sEVs group compared to the PBS control group.
The action of SHED-sEVs on periodontitis disease and periodontal defect in vivo has been investigated in a mouse [80] and rat model [82], respectively. Wei et al. [80] locally injected 20 µg of SHED-sEVs into buccal and lingual sides of the first molar once per week, over 2 weeks, in ligature-induced periodontitis mice. After 2 weeks, SHED-sEVs reduced bone loss, with a decreased CEJ-ABC distance compared to the controls. Moreover, Wu et al. [82] generated a periodontal defect (4 × 2 × 1.5 mm 3 ) in their rat model, at the buccal alveolar bone of the first-to-third mandibular molars. SHED-sEVs were loaded into a β-TCP scaffold before placing them into the periodontal defect for four weeks, resulting in enhanced neovascularization and new bone formation compared to the β-TCP/PBS scaffold.
In their study, Shi et al. [83] injected gelatin hydrogels (100 µL), loaded with LPS-DFCs-sEVs (sEVs derived from LPS-treated DFCs) or DFCs-sEVs, into the periodontal pocket of the right maxillary second molar in a ligature-induced periodontitis rat model. The intervention was once a week for up to 8 weeks, and resulted in significantly reduced alveolar bone loss and TRAP-positive osteoclasts, as well as enhanced well-oriented PDL fibers in the LPS-DFCs-sEVs group.

Summary and Discussion
Periodontal cells (PDLCs/SCAP and GFs/GMSCs) and dental pulp (DPSCs/SHED)derived EVs can play an important role in augmenting the function of recipient cells, such as proliferation and osteo/odontogenic differentiation, as well as anti-inflammation and anti-cancer properties . In particular, one study of GMSCs-sEVs [61] and DPSCs-sEVs [72], two studies of SHED-sEVs [80,82], and one study of DFCs-sEVs [83] can promote alveolar bone, vasculature and well-organized PDL fibers regeneration, and reduced inflammation in a periodontitis animal model or a periodontal defect model. As such, these EVs may serve as potential 'cell-free' therapeutics to facilitate periodontal regeneration; however, more in vivo studies are required to confirm this concept.
All of the above factors are critical for a successful therapeutic outcome; thus, it is of great importance to follow the relevant guidelines and consider the above-discussed variables with more comparisons between different parameters.
From the included studies, except for one common EV-miRNA (miR-1260b) between DPSCs/SHED and GFs/GMSCs, there appears to be no common EV-miRNA detected between these cell types (shown in Venn diagram, Figure 4a). We also listed reported EV-miRNAs and EV-circRNAs from PDL(S)Cs-EV and hDPSCs-EV (Figure 4b). However, this needs further confirmation with more studies. Furthermore, 38 EV-miRNAs, 69-557 EV-circRNAs, 254-15,380 EV-mRNAs and 2907-11,581 EV-lncRNAs were reported for EV from periodontal (dental pulp) cells by RNA sequencing analysis. We have outlined that these EVs possess anti-inflammation, osteo/odontogenesis, anti-osteoclastogenesis, angiogenesis and immunomodulatory functions in vitro and in vivo. Thus, we propose that periodontal cell-derived EVs can modulate the cell function via EV cargos (Figure 4c). However, more studies for periodontal cell-derived EVs are required to further confirm this concept.
Given that cell source, CM collection, and EV isolation and characterization are critical in obtaining pure EV populations, future studies should take these factors into account and follow the latest MISEV guidelines. Researchers should consider adding EV purity (EV particles per µg protein), DNase/RNase/proteinase treatment and EV engineering before in vivo therapeutic research. Although current research has not yet standardized these factors, data from all 33 studies in this review suggest that periodontal (dental pulp) cell-derived EVs can function as potential therapeutics to promote periodontal regeneration and impart anti-inflammatory properties. However, investigating the effect of periodontal cell-derived EV on in vivo periodontal regeneration models is required to understand their potential therapeutic role in periodontal regeneration.

Conclusions and Future Perspectives
This review demonstrates that sEV can be isolated from periodontal and pulp cells, with 11 studies investigated the EV cargos, including sEV-miRNA [52,53,61,63,64,72,75,81], EV-circRNA [51,71], EV-lncRNA [51] and EV-mRNA [67,72]. We summarize the common EV-miRNA and EV-circRNA within periodontal (or dental pulp) cells (Figure 4a,b). From the included studies, except for one common EV-miRNA (miR-1260b) between DPSCs/SHED and GFs/GMSCs, there appears to be no common EV-miRNA detected between these cell types (shown in Venn diagram, Figure 4a). We also listed reported EV-miRNAs and EV-circRNAs from PDL(S)Cs-EV and hDPSCs--EV ( Figure 4b). However, this needs further confirmation with more studies. Furthermore, 38 EV-miRNAs, 69-557 EV-circRNAs, 254-15,380 EV-mRNAs and 2907-11,581 EV-lncRNAs were reported for EV from periodontal (dental pulp) cells by RNA sequencing analysis. We have outlined that these EVs possess anti-inflammation, osteo/odontogenesis, anti-osteoclastogenesis, angiogenesis and immunomodulatory functions in vitro and in vivo. Thus, we propose that periodontal cell-derived EVs can modulate the cell function via EV cargos (Figure 4c). However, more studies for periodontal cell-derived EVs are required to further confirm this concept. Given that cell source, CM collection, and EV isolation and characterization are critical in obtaining pure EV populations, future studies should take these factors into account and follow the latest MISEV guidelines. Researchers should consider adding EV purity (EV particles per μg protein), DNase/RNase/proteinase treatment and EV engineering before in vivo therapeutic research. Although current research has not yet standardized these factors, data from all 33 studies in this review suggest that periodontal (dental pulp) cell-derived EVs can function as potential therapeutics to promote periodontal regeneration and impart anti-inflammatory properties. However, investigating the effect of periodontal cell-derived EV on in vivo periodontal regeneration models is required to understand their potential therapeutic role in periodontal regeneration.