Isolation and Characterization of Dental Pulp Stem Cell-Derived Exosomes Across Cell Passages

Summary

This protocol outlines a standardized workflow for isolating and characterizing dental pulp stem cells (DPSC)-derived exosomes across passages, enabling reproducible assessment of exosome quality and functional activity for regenerative applications.

Abstract

Dental pulp stem cells (DPSCs) represent an accessible and clinically relevant source of mesenchymal stem cells, and their derived exosomes have emerged as promising bioactive vesicles for cell-free applications in tissue engineering and regenerative medicine. However, reproducible and standardized methods for isolating and functionally validating DPSC-derived exosomes remain limited, particularly with respect to variability introduced during cell expansion across passages. Here, we describe a comprehensive, standardized, kit-based workflow for the isolation and purification of exosomes from DPSC-conditioned medium that is compatible with routine laboratory practice. This method enables consistent recovery of intact vesicles suitable for morphological, molecular, and functional characterization using commonly available techniques. Exosome identity is assessed based on morphology, size distribution, and marker expression, while biological activity is evaluated using a defined in vitro anti-inflammatory assay. To examine passage-related effects, exosomes derived from different DPSC passages are compared using this functional readout, providing a practical framework for assessing consistency during cell expansion. By integrating isolation, purification, characterization, and functional validation into a single workflow, this protocol offers a reproducible and scalable approach for generating functionally validated DPSC-derived exosomes for biomaterials research and regenerative applications.

Introduction

Exosomes derived from mesenchymal stem cells serve as crucial mediators of intercellular communication and demonstrate significant therapeutic potential in the fields of tissue engineering and regenerative medicine. Exosomes are vesicles measuring 30–150 nm in diameter, capable of delivering bioactive molecules and regulating various functions in recipient cells, such as proliferation, differentiation, and immune response^1,2. Among the various mesenchymal stem cell (MSC) sources, dental pulp stem cells (DPSCs) are of particular interest because of their ready accessibility, robust proliferative capacity, low tumorigenic risk, and minimal ethical concerns^3,4,5. Exosomes derived from DPSCs (DPSC-Exos) contain a diverse repertoire of functional cargo, including proteins, mRNAs, and microRNAs, enabling them to retain many of the biological properties of their parental cells. Previous studies have demonstrated that DPSC-Exos can promote angiogenesis, neural repair, bone regeneration, and anti-inflammatory responses. Beyond their roles in tissue engineering, exosomes are increasingly recognized as important regulators in oncogenesis and cancer therapy, where they contribute to tumor microenvironment remodeling, drug resistance, and immune evasion⁶. MSC-derived exosomes, including those from DPSCs, possess inherent tumor-homing capabilities and immunomodulatory properties, positioning them as promising next-generation vehicles for targeted anti-cancer drug delivery or as cell-free immunotherapeutic agents. By circumventing several limitations associated with cell-based therapies, DPSC-Exos have emerged as a promising acellular bioactive component for regenerative applications^{7,8,9,10,11,12}.

Despite these advantages, the translation of DPSC-Exo research into reproducible experimental or clinical use remains challenging. A major limitation is the lack of unified and standardized protocols for exosome isolation and purification. Currently used approaches include ultracentrifugation¹³, ultrafiltration, precipitation, size-exclusion chromatography, and immunoaffinity capture^14,15. Among these methods, polymer-based precipitation offers practical advantages, including high throughput, compatibility with a wide range of sample volumes, and operational simplicity, making it suitable for studies involving large sample numbers or scale-up requirements^16,17,18. Comparative analyses of different isolation methods, particularly regarding protein yield and particle-to-protein ratio, have been widely documented in the literature¹⁹. However, the influence of DPSC passage number on the characteristics and biological activity of derived exosomes has not been sufficiently addressed, representing an important gap for the standardization and quality control of DPSC-Exo production.

To address these technical limitations, this study presents a standardized, kit-based protocol for isolating and purifying exosomes from DPSC-conditioned medium. In addition to physical and molecular characterization, the protocol incorporates a functional assay to evaluate the anti-inflammatory activity of exosomes derived from different DPSC passages, enabling a comparative assessment of passage-dependent consistency in exosome quality and function. The workflow is designed for straightforward implementation in laboratories equipped with standard cell culture and molecular biology facilities. It relies on commercially available kits to reduce inter-experimental variability and can be readily scaled to accommodate different experimental needs. To minimize donor-to-donor variability, DPSCs are isolated from individual teeth and expanded independently rather than pooled from multiple donors. This protocol is particularly useful for researchers seeking to generate functionally validated DPSC-derived exosomes for downstream applications, including wound healing, bone tissue engineering, and immunomodulation studies, where batch-to-batch consistency is essential. Overall, it provides a practical and standardized framework for producing DPSC-derived exosomes for use in tissue engineering and regenerative medicine research.

Protocol

This study was conducted in accordance with the principles of the Declaration of Helsinki and approved by the institutional ethics committees. Human dental pulp tissues were obtained from discarded third molars or orthodontic premolars collected at Ningbo Stomatology Hospital in accordance with institutional ethical guidelines (Protocol No. NBKQYY2025LS-04; approval date: May 30 2025). Human skin tissues were obtained from discarded foreskin samples collected during routine circumcision procedures in the Department of Urology at the Affiliated People's Hospital of Ningbo University, in accordance with institutional ethical guidelines (Protocol No. 2024104; approval date: Oct 30 2024). Written informed consent was obtained from all donors or their legal guardians prior to sample collection.

1. Isolation of dental pulp stem cell-derived exosomes

1. Isolation, culture, and identification of DPSCs
   1. Sample collection and transport
      1. Collect intact human third molars from donors aged 18–25 years or premolars from donors aged 12–16 years following orthodontic extraction.
      2. Immediately place the extracted teeth in pre-chilled phosphate buffered saline (PBS) supplemented with 100 U/mL penicillin-G and 100 µg/mL streptomycin.
      3. Transport samples to the laboratory at 4 °C and process within 6 h of extraction.
   2. Dental pulp exposure and tissue dissociation
      1. Immerse in 70% ethanol for 30 s; wash three times with 1× PBS containing 3% P/S (30 s each). Carefully remove residual gingival and periodontal tissues from the tooth surface using sterile scissors or a scalpel.
      2. Wrap the tooth in sterile gauze, then carefully crack the crown using a sterilized hammer to expose the pulp chamber.
      3. Gently separate the pulp tissue from the dentin, mince it into small fragments (<1 mm³) using sterile surgical blades.
      4. Transfer the tissue fragments into a 1.5 mL centrifuge tube containing a digestion solution of 2.5 mg/mL Dispase II and 2.5 mg/mL collagenase type I.
      5. Incubate the mixture at 37 °C incubator for 30–45 min with gentle agitation.
         NOTE: The duration of enzymatic digestion was determined based on the volume and degree of homogenization of the dental pulp tissue.
   3. Digestion termination and primary cell culture
      1. Stop the enzymatic digestion by adding an equal volume of Dulbecco's Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS).
      2. Centrifuge the cell suspension at 200 × g for 5 min to pellet the cells and tissue fragments.
      3. Discard the supernatant and resuspend the pellet in 1 mL of complete culture medium (α-MEM with 20% FBS, 100 U/mL penicillin, and 100 µg/mL streptomycin).
      4. Seed the resuspended cells and explants into a 60-mm culture dish and add complete culture medium to a final volume of 5 mL.
      5. Incubate the cells at 37 °C in a humidified atmosphere containing 5% CO₂.
   4. Cell maintenance and subculture
      1. Refresh the culture medium every 2–3 days.
      2. Monitor cells daily.
         NOTE: Primary DPSCs will typically migrate from tissue explants and reach 80–90% confluence within 7–14 days.
      3. For subculture, detach cells using 0.05% trypsin- ethylenediaminetetraacetic acid (EDTA) when they reach 80–90% confluence.
         CAUTION: Trypsin is an irritant. Wear appropriate personal protective equipment, including gloves and safety glasses.
      4. Passage cells at a ratio of 1:3 using the complete culture medium (with 20% FBS).
      5. Use DPSCs at passages 2, 4, and 6 for all downstream exosome isolation and functional assays.
   5. Characterization of DPSCs by flow cytometry
      1. Detach DPSCs at the desired passage using 0.05% trypsin-EDTA, neutralize with complete culture medium, and collect the cell suspension.
      2. Centrifuge at 300 × g for 5 min at room temperature and discard the supernatant. Resuspend the cell pellet in PBS and count the cells. Adjust the cell density to approximately 1 × 10⁶ cells/mL.
      3. Aliquot 100 µL of the cell suspension into each flow cytometry tube. Incubate the cells with fluorochrome-conjugated antibodies against CD44, CD90, CD105, CD45, CD19, and CD14 at a dilution of 1:100 in PBS for 30 min at 4 °C in the dark.
      4. Wash the cells by adding 1 mL of PBS to each tube, followed by centrifugation at 300 × g for 5 min. Carefully discard the supernatant and resuspend the cell pellet in 300–500 µL of PBS for analysis.
      5. Include appropriate isotype control antibodies at matched concentrations for each fluorochrome to enable background correction and gating.
      6. Analyze samples using a flow cytometer equipped with appropriate lasers and filters for the selected fluorochromes. Acquire at least 10,000 events per sample.
      7. Perform data analysis using flow cytometry software. First, gate viable cells based on forward scatter (FSC) and side scatter (SSC) to exclude debris. Then apply singlet gating to remove doublets.
      8. Determine marker expression by comparing fluorescence intensity with corresponding isotype controls to define positive and negative populations.
2. Isolation and purification of DPSC-Exos
   1. Preparation of conditioned medium
      1. Culture DPSCs in a 10-cm culture dish until they reach 70–80% confluence.
      2. Remove the spent medium and then gently wash the cell monolayer twice with pre-warmed PBS.
      3. Add 10 mL of complete medium supplemented with 10% exosome-depleted FBS.
      4. Further incubate the culture at 37 °C until cell confluence reaches 90–100% (a process usually exceeding 24 h).
   2. Collection of conditioned medium
      NOTE: Keep all subsequent steps on ice or at 4 °C.
      1. Transfer the conditioned medium to a centrifuge tube.
      2. Centrifuge the medium at 4 °C and 3,000 × g for 10 min to remove cells and large debris.
      3. Carefully transfer the supernatant to a new centrifuge tube without disturbing the pellet.
      4. Centrifuge the supernatant at 4 °C and 10,000 × g for 10 min to pellet smaller particles and apoptotic bodies.
      5. Filter the supernatant through a 0.22 µm syringe filter into a clean 50 mL centrifuge tube.
      6. Process the clarified medium immediately for exosome isolation, or aliquot and store it at -80 °C.
         NOTE: The clarified conditioned medium can be stored at -80 °C for several months. Avoid repeated freeze-thaw cycles.
   3. Exosome isolation by polymer-based precipitation
      1. Thaw the clarified conditioned medium on ice if it was frozen.
      2. In a sterile tube on ice, combine the medium with an exosome concentration solution at a 4:1 volume ratio (e.g., 20 mL medium: 5 mL solution). Vortex the mixture for 1 min to ensure homogeneity.
      3. Leave the mixture undisturbed at 4 °C for 16–18 h to allow exosome precipitation.
         NOTE: The incubation time can be appropriately extended to potentially increase yield, but should not exceed 24 h.
      4. Centrifuge the mixture at 4 °C and 10,000 × g for 75–80 min to pellet the exosomes. Carefully aspirate and discard the supernatant without disturbing the pellet.
      5. Centrifuge the tube again at 4 °C and 10,000 × g for 2 min. Remove any residual supernatant with a pipette.
      6. Resuspend the exosome pellet in 1x PBS (Use 200 µL of PBS per 20 mL conditioned medium).
      7. Transfer the resuspension to a new 1.5 mL centrifuge tube and centrifuge at 4 °C and 12,000 × g for 2 min, retaining the supernatant, which is enriched with exosome particles (repeat if necessary).
   4. Purification of exosomes
      1. Apply the exosome suspension to an exosome purification filter column placed in a collection tube.
      2. Centrifuge the column at 4 °C and 3,000 × g for 10 min.
      3. Collect the flow-through liquid from the bottom of the collection tube. This liquid contains the purified exosomes.
   5. Storage of purified exosomes
      1. Aliquot the purified exosome suspension into single-use volumes suitable for downstream experiments.
      2. Store the aliquots at -80 °C.
         ​NOTE: Aliquoting is critical to avoid repeated freeze-thaw cycles, which can degrade exosome integrity and biological activity.
   6. Characterization and identification of DPSCs-Exo
      1. Perform morphological analysis by transmission electron microscopy (TEM) as described in steps 1.2.6.2–1.2.6.6.
      2. Use a pipette, draw 10 µL of the prepared exosome suspension sample, and drop it onto pre-prepared Parafilm. Place the formvar/carbon-coated copper grid, film side facing down, onto the droplet, allowing it to adsorb naturally for 10–15 min.
      3. Use a filter paper strip to wick away any excess liquid and allow it to air dry briefly.
      4. Use a pipette, draw 10 µL of 2% phosphotungstic acid solution, and drop it onto the Parafilm. Invert the grid that has completed the adsorption step so its film side faces the staining solution, and let it sit for 3–5 min.
      5. Use a filter paper strip to wick away excess liquid and allow the grid to dry under an incandescent lamp.
      6. Observe and capture images using a transmission electron microscope.
      7. Perform size distribution and concentration analysis by nanoparticle tracking analysis (NTA) as described in steps 1.2.6.8–1.2.6.12.
      8. Rinse the sample chamber three times with ultrapure water.
      9. Calibrate the instrument using standard 100 nm polystyrene beads according to the manufacturer's instructions. Proceed with measurements after successful calibration.
      10. Rinse the sample chamber with 1× PBS.
      11. Dilute the exosome sample in 1× PBS (1:500 dilution) and load it into the sample chamber. Monitor particle movement in real time using the instrument software.
      12. Acquire data and generate the corresponding analysis report.
      13. Identify the protein markers by Western blot analysis as described in steps 1.2.6.14–1.2.6.24.
      14. Lyse exosome samples using an exosome-specific protein lysis buffer and quantify protein concentration using a bicinchoninic acid (BCA) assay.
      15. Prepare resolving and stacking gels. Load equal amounts of protein per lane and add 5 µL of protein marker to each gel.
      16. Perform electrophoresis at a constant voltage of 80 V until the dye front reaches the resolving gel, then increase to 120 V and continue until the dye front reaches the bottom of the gel.
      17. Cut a polyvinylidene fluoride (PVDF) membrane to the appropriate size and activate it by soaking it in methanol for 5 min.
      18. Prepare transfer buffer consisting of 25 mM Tris base, 192 mM glycine, and 20% (v/v) methanol (pH = 8.3). Equilibrate all transfer components in the transfer buffer.
      19. Assemble the transfer sandwich in the following order from cathode to anode: sponge – filter paper – gel – PVDF membrane – filter paper – sponge. Remove any air bubbles and perform protein transfer at a constant current of 300 mA.
      20. After transfer, rinse the PVDF membrane briefly with Tris-buffered saline with 0.1% Tween20 (TBST) and block with 5% non-fat milk for 1 h at room temperature with gentle shaking.
      21. Incubate the membrane with primary antibodies (e.g., anti-CD63 and anti-TSG101) diluted in 5% non-fat milk overnight at 4 °C with gentle shaking. Wash the membrane three times with TBST for 5 min each.
      22. Incubate with the appropriate secondary antibody at room temperature for 30 min, followed by three washes with TBST (5 min each).
      23. Prepare fresh electrochemiluminescence (ECL) working solution (mix Solutions A and B at a 1:1 ratio), apply to the membrane, and detect signals using a chemiluminescence imaging system.

2.​ Isolation and culture of human primary skin epidermal keratinocytes

1. Sample collection and transport
   1. Obtain foreskin tissue samples from healthy male donors at The Affiliated People's Hospital of Ningbo University.
   2. Immediately place the extracted foreskin tissue in 4 °C PBS supplemented with 100 U/mL penicillin-G and 100 µg/mL streptomycin.
   3. Transport samples to the laboratory at 4 °C and process within 6 h.
2. Aseptic preparation and tissue fragmentation
   1. Immerse the tissue in 70 % ethanol for 30 s to 3 min.
      NOTE: Adjust the immersion time based on tissue size for initial disinfection.
   2. Using sterile scissors or a scalpel, dissect and remove subcutaneous fat and connective tissue from the washed tissue sample.
   3. Mince it into small fragments (fragment size <1 mm³) using sterile surgical blades.
      NOTE: Mincing 3–4 tissue samples typically requires less than 15 min. Add a small amount of PBS during mincing if the tissue appears dry.
   4. Transfer the minced tissue slurry into a 50 mL centrifuge tube containing a digestion solution of 2.5 mg/mL Collagenase Type I and 2.5 mg/mL Dispase II. Add 10–20 mL of digestion solution per piece of minced tissue.
   5. Incubate the mixture at a 37 °C incubator for 60 min. Maintain gentle agitation at 80 × g, or agitate the tube vigorously by hand every 20 min.
3. Secondary digestion and DNase treatment
   1. Add 0.25% Trypsin-EDTA to the digestion mixture at a 1:5 volume ratio. Incubate the tube at 37 °C for 25 min with gentle agitation.
      CAUTION: Trypsin is an irritant. Wear appropriate personal protective equipment, including gloves and safety glasses.
   2. Add DNase I solution (10 mg/mL) to the mixture at a ratio of 10 µL per mL of total digestion volume. Incubate at 37 °C for 5 min with gentle agitation.
4. Digestion termination and primary cell culture
   1. Stop the enzymatic digestion by adding an equal volume of DMEM supplemented with 10% FBS.
   2. Triturate the mixture thoroughly by pipetting up and down approximately 20 times to dissociate the tissue fragments. Filter the cell suspension through a 100 µm cell strainer into a new 50 mL centrifuge tube.
   3. Centrifuge the cell suspension at 200 × g for 5 min to pellet the cells and tissue fragments.
   4. Discard the supernatant and resuspend the pellet in 1 mL of complete culture medium (α-MEM with 20% FBS, 100 U/mL penicillin, and 100 µg/mL streptomycin).
   5. Seed the resuspended cells and explants into a 10-cm culture dish and add complete culture medium to a final volume of 6 mL.
   6. Incubate the cells at 37 °C in a humidified atmosphere containing 5% CO₂.
      NOTE: Supplement keratinocyte cultures with Y-27632 at a final concentration of 5 µM during culture.
5. Cell maintenance and subculture
   1. Refresh the culture medium every 2–3 days.
   2. Monitor cells daily.
      NOTE: Primary skin epidermal keratinocytes will typically migrate from tissue explants and reach 80-90% confluence within 10-14 days.
   3. For subculture, detach cells using 0.05% trypsin-EDTA when they reach 80–90% confluence.
      ​CAUTION: Trypsin is an irritant. Wear appropriate personal protective equipment, including gloves and safety glasses.

3. Functional verification of anti-inflammatory function of exosomes from different passages

1. Cell seeding and preparation
   1. Seed epidermal keratinocytes at a density of 1.5 × 10⁵ cells per well into a 12-well plate. Add 1 mL of complete medium to each well and culture the cells for 24 hours.
   2. When cell confluence reaches approximately 70%, carefully aspirate the medium, gently wash the cell monolayer twice with pre-warmed 1× PBS.
2. Inflammatory stimulation and exosome treatment
   1. Induce a psoriasiform cellular model, stimulate the cells with the proinflammatory cytokine mixture "M5" (containing IL-1α, IL-17A, IL-22, Oncostatin M, and TNF-α, each at a final concentration of 10 ng/mL).
   2. For the treatment groups, co-incubate the stimulated cells with DPSC-Exos (50 µg per well).
   3. For the negative control (NC) group, add an equivalent volume of PBS instead of exosomes.
      ​NOTE: Perform all stimulations and treatments in a medium devoid of Y-27632 to preclude its potential interference with inflammatory signaling pathways.
3. Sample collection and qRT-PCR analysis (functional comparison across passages)
   1. Harvest the cells and extract RNA after 24 h of treatment.
   2. Analyze the expression of the key inflammatory marker IL-23A.
      1. Compare the inhibitory effects on IL-23A expression among exosome treatment groups derived from different passages (P2, P4, and P6) using qRT-PCR to assess passage-dependent changes in exosome function.
      2. Perform qRT-PCR using the following cycling conditions: initial denaturation at 95 °C for 3 min, followed by 40 cycles of denaturation at 95 °C for 10 s and annealing/extension at 60 °C for 30 s.
      3. Perform a melt curve analysis from 65 °C to 95 °C to confirm amplification specificity.
         ​NOTE: Primer sequences: Human-IL-23A (forward: 5'-GAAGAGGGAGATGAAGAGAC-3', reverse: 5'-TATCCGATCCTAGCAGCTTC-3'); Human-H36B4 (forward: 5'-GCAATGTTGCCAGTGTCTGT-3', reverse: 5'-GCCTTGACCTTTTCAGCAAG-3')

Results

Identification and Characterization of Dental Pulp Stem Cells
Primary dental pulp stem cells (DPSCs) isolated using the described protocol exhibited stable growth and typical mesenchymal morphology during in vitro expansion. As shown in Figure 1, DPSCs at passages 2, 4, and 6 displayed a homogeneous population of spindle-shaped, fibroblast-like cells, with no apparent morphological differences or signs of spontaneous differentiation across pa...

Discussion

The standardized protocol described in this study provides a reproducible framework for the isolation of dental pulp stem cells (DPSCs), the production of DPSC-derived exosomes (DPSC-Exos), and the functional validation of these vesicles across multiple cell passages. The discussion below highlights critical procedural considerations, optimization strategies, methodological limitations, and the broader relevance of this workflow for regenerative medicine research.

The reliability of this proto...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.05% Trypsin-EDTA Solution	Gibco	25300054	Due to its digestive strength, trypsin is widely used for cell dissociation, routine cell culture passaging, and primary tissue dissociation.
100 mm Cell and Tissue Culture Dish	Biofil	TCD010100	Surface Treated/Non-Treated Sterile&Non-Pyrogenic DNase/RNase-Free
100 nm PS beads	Thermo Fisher	3100A	Prior to conducting nanoparticle tracking analysis (NTA) experiments, the instrument must be calibrated using a standard with a precisely known particle size (100 nm).
15-mL Bulk Conical Centrifuge Tube	KIRGEN	KG2611	Sterile, RNase/Dnase and pyrogenic free
200-mesh Formvar-carbon-coated copper grid	Zhongjing Scientific Instruments	AZH200	Supports exosome samples for TEM imaging. The Formvar-carbon coating enhances sample adhesion and conductivity, while the 200-mesh grid provides adequate open area for even particle distribution and stable imaging.
37 °C Incubator	Thermo Scientific	51032877	The 37-degree CO2 incubator provides an ideal in vitro environment.
50-mL Bulk Conical Centrifuge Tube	KIRGEN	KG2811	Sterile, RNase/Dnase and pyrogenic free
Anti-CD63 antibody [MX-49.129.5]	Abcam	ab193349	Anti-CD63 antibody [MX-49.129.5] (ab193349) is a mouse monoclonal antibody detecting CD63 in Western Blot, Flow Cytometry (Intra), Flow Cytometry, IHC-P, ICC/IF. Suitable for Human.
Anti-TSG101 antibody [EPR7130(B)]	Abcam	ab125011	Anti-TSG101 antibody [EPR7130(B)] (ab125011) is a rabbit monoclonal antibody detecting TSG101 in Western Blot, Flow Cytometry (Intra), Flow Cytometry, IHC-P, ICC/IF. Suitable for Human, Mouse, Rat.
Automated cell counter	Countstar	IC1000	Automated cell counter for fast, precise cell counting
BCA Protein Assay Kit	Beyotime	P0012	Have good linear relationship within the concentration range of 50 - 2000 μg/ml.
Benchtop Centrifuge	BIORIDGE	TD5	Benchtop centrifuge for quick and efficient sample separation in laboratory settings
Biological safety cabinet	Thermo Scientific	1379	Biological Safety Cabinet is a sterile containment device that protects operators, samples, and environments from biohazards by filtering airborne pathogens and preventing cross-contamination during microbiological work.
Cell Culture Plate,12-Well	Eppendorf	0030721110	Tissue culture treated,with lid,flat bottom Sterile,free of detectable pyrogens,RNase & DNase,human & bact.DNA Non-cytotoxic
Cell Culture Plate,96-Well	NEST	701001 0714B	Tissue Culture Treated,Polystyrene,Non-Pyrogenic,Sterile
Cellsaving	NCM	C40100	Serum-free,animal protein-free Cell Freezing Medium
Collagenase, Type I, powder	Gibco	17100017	Collagenase is a protease that cleaves the bond between a neutral amino acid (X) and glycine in the sequence Pro-X-Gly-Pro, which is found with high frequency in collagen.
Dispase II, powder	Gibco	17105041	Dispase II (neutral protease) is an amino-endo peptidase that hydrolyzes the N-terminal peptide bonds of non-polar amino acid residues.
Dulbecco's Modified Eagle Medium	Gibco	C11995500BT	[+]4.5g/L D-Glucose [+]L-Glutamine [+]110mg/L Sodium Pyruvate
Evo M-MLV Reverse Transcription Kit	AG	AG11707	A dedicated reverse transcription reagent for Real Time RT-PCR. It utilizes the Evo M-MLV reverse transcriptase with strong extension ability, enabling efficient synthesis of cDNA within a short period of time.
Exosome Concentration Solution	Umibio	UR52111	Extract exosomes from the cell supernatant
Exosome Depleted Fetal Bovine Serum	Umibio	UR50202	The exosome-free fetal bovine serum was obtained by filtering the sterilely collected healthy fetal bovine serum, and ≥99% of the endogenous exosomes in the fetal bovine serum were removed. It had the same cell growth rate and morphology as the fetal bovine serum before processing.
Fetal Bovine Serum, qualified, Australia	Gibco	10099141	Hemoglobin level: ≤30 mg/dL (levels routinely ≤25 mg/dL)
Human-H36B4	Sangon Biotech	SB002	Forward: 5'-GCAATGTTGCCAGTGTCTGT-3'; Reverse: 5'-GCCTTGACCTTTTCAGCAAG-3'
Human-IL-23A	Sangon Biotech	SB002	Forward: 5'-GAAGAGGGAGATGAAGAGAC-3'; Reverse: 5'-TATCCGATCCTAGCAGCTTC-3'
Inverted Biological Microscope	Leica	DMI1	Directly observe the living cells or tissues that are adhering to the walls of containers such as culture flasks and petri dishes.
Minimum Essential Medium	Gibco	C12571500BT	[+]L-Glutamine [+]Ribonucleosides [+]Deoxyribonucleosides
NcmSpin Rapid RNA Extraction Kit	NCM	M5106	This kit can rapidly extract total RNA from up to 3,000,000 cells and up to 10mg of tissue.
Penicillin-Streptomycin	NCM	C100C5	Contains 10,000 units/ml of penicillin and 10,000 micrograms/ml of streptomycin. It has been filtered to remove bacteria and can be directly used for cell culture. It effectively prevents bacterial contamination of the cells.
PerfectStart Green qPCR SuperMix	TransGen	AQ601-02-V2	A fluorescent dye that binds to the minor groove of all double-stranded DNA and emits green light upon excitation. During PCR amplification, it incorporates into newly amplified DNA, resulting in increased fluorescence. During the denaturation step, the DNA strands separate, releasing the dye and causing a significant drop in the fluorescence signal.
Phosphate Buffered Saline (PBS)	Viva Cell Biosciences	C3582-0500	Its function is to keep the pH of the medium within physiological range and maintain the osmotic pressure balance inside and outside the cell.
Phosphotungstic acid	Solarbio	G1870	Used as a negative stain for TEM sample preparation.
SW Shaking Water Baths	Julabo	(DIN 12876-1)I	Working temperature range from +20 to +99.9 °C
Syringe Filter,0.22 μm/33 mm,PES	Beyotime	FF362	DNase/RNase Free Low Protein Binding Non-Pyrogenic Individually Wrapped
Transmission Electron Microscope (TEM)	HITACHI	HT7700	Used for high-resolution imaging of nanoparticle morphology, including exosomes. Operated at an accelerating voltage of 80–100 kV to visualize negatively stained samples and confirm particle size, shape, and structural integrity.
ZetaView_Particle Metrix	Particle Metrix	PMX-120	Used for measuring the particle size, concentration, Zeta potential, and fluorescence properties of nanoparticles.