<figure><img src="image_56.png" /><figcaption>FIGURE 4</figcaption></figure>

(A,B) CCK-8 assay indicated that proliferation of HUVECs and HFF-1 after culturing with PNs, FHE10, and FHE10@PNs, respectively; (C,D) DAPI (blue)/ FITC-phalloidin (green) staining assay of HUVECs and HFF-1 with PNs, FHE10, and FHE10@PNs after culturing, respectively; (E) Transwell assay showed that migration of HUVECs and HFF-1 were stained by crystal violet; (F) Tube formation assay; (G) Quantitative analysis of tube formation; (H–J) qRT-PCR analysis of VEGF, PDGF, and bFGF.

In summary, FHE10 hydrogel and PNs were selected for follow-up studies.

## 3.2 Effects of PNs, FHE10, and FHE10@PNs on proliferation, morphology, migration, the tube formation, and angiogenesis-related gene expression in vitro

HUVECs and HFF-1 in good growth state were cultured by using the material extract. The FHE10@PNs promoted cell proliferation more than PNs or FHE10 alone, and with the increase of culture time, cell proliferation gradually increased, indicating that cells also adapted to the surrounding environment and achieved the best growth state (Figures 4A, B).

HUVECs and HFF-1 were cultured for 3 days, then their morphology was observed by a light microscope (Figures 4C, D). HUVECs cells in experiment group have no significant change in cell morphology when compared to the control group. The cell body of HFF-1 was fusiform or irregular triangular, with an oval nucleus in the center and a cytoplasmic protrusion, which was radial when growing. The characteristics were consistent with the normal proliferation of HUVECs and HFF-1.

The migration of HUVECs and HFF-1 co-cultured with the material was observed by a transwell migration assay (Figure 4E). The groups added with materials all could promote cell migration, and the FHE10@PNs had the strongest ability to promote cell migration, so the combined action of the two had a synergistic effect on cell migration. By a quantitative analysis of the number of migrating cells (Supplementary Figure S3), the results were consistent with the

staining, which proved that the materials indeed had the chemotactic attraction to cells.

*In vitro* tube formation assay was performed by co-cultured PNs, FHE10, and FHE10@PNs with HUVECs to evaluate the tube formation ability (Figures 4F, G). HUVECs co-cultured with each group and formed a network of capillary-like structures on the Matrigel after 6 h, and each group had some differences. The tube's diameter is shown in the supporting information (Supplementary Figure S4). Compared with the control group, the PNs, FHE10, and FHE10@PNs groups all could promote angiogenesis, and the FHE10@PNs group had the best-promoting effects.

The effects of PNs, FHE10, and FHE10@PNs on the expression of angiogenesis-related genes after co-culture with HUVECs and HFF-1 for 7 days were evaluated (Figures 4H–J). The gene expression levels of VEGF, PDGF, bFGF in the PNs, FHE10, and FHE10@PNs groups were better than those in the control group, while the FHE10@PNs group had the best gene expression levels.

## 3.3 *In vivo* wound tissue-repair ability

Encouraged by the controllable injectability, favorable biocompatibility and antibacterial activity, outstanding pro-angiogenic properties of the prepared hydrogels revealed above, FHE10@PNs was chosen and further used as a potential wound dressing for skin repair to evaluate the *in vivo* wound healing performance. Figure 5A visually showed the wound healing progress treated by the saline, PNs, FHE10, and FHE10@PNs for 0, 3, 7, and 14 days, respectively. The wound healing rates of the control, PNs, FHE10, and FHE10@PNs groups were measured by
<figure><img src="image_57.png" /><figcaption>FIGURE 5</figcaption></figure>

(A) Wound images of control, PNs, FHE10, FHE10@PNs at 0, 3, 7 and 14 days post-operation; (B) Traces of wound repair for each treatment group *in vivo*. Blue, green, yellow, and red areas correspond to the wound area at n (n = 0, 3, 7 and 14) days, respectively; (C) Wound healing ratio of the defects treated with saline (control), PNs, FHE10, FHE10@PNs at 0, 3, 7 and 14 days post-operation; (D) H&E stained images; (E) The reepithelialization rate of wound defect at 7 and 14 days after treatment with saline (control), PNs, FHE10, and FHE10@PNs; (F) Masson's trichrome stained images; (G,H) The collagen deposition in the wound sites was evaluated by ImageJ.

using ImageJ at each observation time point, and the FHE10@PNs group showed the best performance in wound healing rate, as compared with the other three groups with a significant difference at Day 7 (p < 0.05). The average wound healing rates of control, PNs, FHE10, and FHE10@PNs were 44.8%, 71.3%, 80.0%,

and 87.1%, respectively. The continuous observation for up to 14 days after operation showed unhealed wounds in the control, PNs, and FHE10 groups, which almost completely healed in the FHE10@PNs group with a wound healing rate of 99.5% (Figures 5A-C).
<figure><img src="image_58.png" /><figcaption>**FIGURE 6**
IHC and IF analysis of neovascularization in wound of control, PNs, FHE10, and FHE10@PNs groups, respectively. (A) IHC staining for CD31, red arrows mean newly formed blood vessels; (B) IF co-staining for α-SMA and CD31, yellow arrows mean mature vessels; (C) Area analysis of mature blood vessels at 7 and 14 days post-operation; (D) Quantitative analysis of newly formed blood vessels at 7 and 14 days post-operation; (E) Quantitative analysis of mature vessels at 7 and 14 days post-operation.</figcaption></figure>

We further clarified the quality of regenerated wound tissue by using H&E staining and Masson trichrome staining at the observation time points of day 7 and day 14 (Figures 5D, F). FHE10@PNs exhibited better healing effects than the other three groups (Figures 5D, E). After 7 days of the skin defect, the epidermal regeneration rates of the control group, PNs group, FHE group, and FHE10@PNs group reached 39.5%, 46.3%, 55.8%, and 68.3%, respectively. After 14 days of treatment, the wounds treated with FHE10@PNs exhibited almost no open wounds with a healing rate of 92.3% and become smooth with new epidermal tissue. In contrast, most of the wounds in the other groups remained significantly more open and the scars were uneven.

Collagen fibers are a key component related to granulation tissue and the dermal extracellular matrix during the wound healing processes (Diller and Tabor, 2022). Thus, Masson staining was performed to evaluate the deposition and organization of collagen fibers in regenerative skin tissue at 7 and 14 days post-surgery (Figures 5F, G). As presented in Figure 5G, in terms of the area of collagen deposition, a significant difference was found across the groups at 7 days (15.05%, 16.71%, 19.61%, 25.89% for control, PNs, FHE10, FHE10@PNs, respectively). On day 14, mature collagen fibers are observed in the FHE10@PNs group and the collagen deposition is denser and more organized (Figure 5H). On the other hand, the collagen fibers presented in the other groups are still partially dysplastic.

## 3.4 FHE10@PNs hydrogel facilitates angiogenesis

The angiogenesis is essential for nutrient and oxygen supply in the process of skin wound repair. To better elucidate the properties of the

FHE10@PNs hydrogel on angiogenesis, CD31 and α-SMA were selected for immunohistochemical staining analysis. As plotted in Figures 6A, D, the FHE10@PNs group exhibited significantly increased neovascularization based on a high level of CD31 expression on both day 7 and day 14. In contrast, either PNs or FHE10 alone displayed less CD31 expression, confirming that the FHE10@PNs hydrogels provided a favorable 3D microenvironment scaffold for blood vessel formation.

Moreover, the colocalization of α-SMA and CD31 was seen as a biomarker of a normalized vessel (Wu et al., 2020). Positively stained endothelial cells and smooth muscle cells lining the abundant vessel networks were observed in the skin regenerated tissue treated with FHE10@PNs, but significantly, less were observed in the control or other material-treated groups. The mature vessel areas on day 7 and day 14 was significantly bigger than the other groups (Figures 6B,C,E). On the basis of the above results, FHE10@PNs hydrogels not only showed the ability to facilitate neovascularization but also to promote subsequent vascular maturation in the process of skin wound repair.

## 4 Conclusion

In this study, a hybrid hydrogel was prepared by preparing PGS acrylate derivative photocurable nanospheres and mixing them with injectable oxidized hyaluronic acid hydrogel FHE10. The *in vitro* results showed that the FHE10@PNs group could up-regulate the proliferation, migration, and expression of genes related to angiogenesis of HUVECs and HFF-1. *In vivo* studies further demonstrated that the prepared FHE10@PNs group significantly promoted wound reepithelialization, collagen deposition, and neovascularization, and finally promoted wound healing. Therefore,
the preparation of injectable FHE10@PNs has better effects on promoting reepithelialization, collagen deposition, angiogenesis and wound healing than using PNs or FHE10 hydrogel alone. In conclusion, the FHE@PNs may be a potential material to promote reepithelialization, collagen deposition, and angiogenesis in wound healing.

## Data availability statement

The original contributions presented in the study are included in the article/Supplementary Material, further inquiries can be directed to the corresponding authors.

## Ethics statement

The animal study was reviewed and approved by Animal Research Ethics Committee of the First Affiliated Hospital of Zhejiang University.

## Author contributions

JL: investigation, methodology, and writing original draft. FS, YL, JD, SC, XJ, and JJ: methodology, and conceptualization. PR, YC, TZ, FS, and JL: conceptualization, project administration, and writing-review and editing.

## Funding

This research was funded by the Shanghai Rising-Star Program (20QC1401300), National Natural Science Foundation of China (81902186), Technology Support Project of Science and Technology Commission of Shanghai Municipality (22S31904700),

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## Acknowledgments

We thank the excellent technical support (microscopy platform: Xingguang Liang, Fangjie Lu, Yahong Wu, flow cytometry platform: Yi Guo) by the corefacility, Central Laboratory, the First Affiliated Hospital, Zhejiang University School of Medicine with the confocal laser microscopy, and flow cytometry.

## Conflict of interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

## Publisher's note

All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher.

## Supplementary material

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fbioe.2022.1091122/full#supplementary-material

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<figure><img src="image_59.png" /></figure>

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OPEN ACCESS

EDITED BY

Tonghe Zhu,
Shanghai University of Engineering
Sciences, China

REVIEWED BY

Jianhai Chen,
Peking University People's Hospital, China
Lichen Wang,
Stevens Institute of Technology,
United States
Kaihu Li,
Xiangya Hospital, Central South University,
China

*CORRESPONDENCE

Chengyu Zhuang,

zhuangchengyu@msn.com

Lei Wang,

ray_wangs@hotmail.com

¹These authors have contributed equally to this work

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a section of the journal
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RECEIVED 12 November 2022

ACCEPTED 30 January 2023

PUBLISHED 09 February 2023

CITATION

Li G, Zhang Y, Wu J, Yang R, Sun Q, Xu Y, Wang B, Cai M, Xu Y, Zhuang C and Wang L (2023), Adipose stem cells-derived exosomes modified gelatin sponge promotes bone regeneration.
*Front. Bioeng. Biotechnol.* **11**:1096390.
doi: 10.3389/fbioe.2023.1096390

COPYRIGHT

© 2023 Li, Zhang, Wu, Yang, Sun, Xu, Wang, Cai, Xu, Zhuang and Wang. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

Adipose stem cells-derived
exosomes modified gelatin sponge
promotes bone regeneration

Gen Li¹†, Yin Zhang¹†, Jiezhou Wu²†, Renhao Yang¹, Qi Sun²,
Yidong Xu¹, Bo Wang², Ming Cai², Yang Xu¹, Chengyu Zhuang¹* and
Lei Wang¹*

¹Department of Orthopaedics, Ruijin Hospital, Shanghai Jiaotong University School of Medicine, Shanghai, China, ²Department of Orthopaedics, Shanghai Tenth People's Hospital, Tongji University School of Medicine, Shanghai, China

**Background:** Large bone defects resulting from trauma and diseases still a great challenge for the surgeons. Exosomes modified tissue engineering scaffolds are one of the promising cell-free approach for repairing the defects. Despite extensive knowledge of the variety kinds of exosomes promote tissue regeneration, little is known of the effect and mechanism for the adipose stem cells-derived exosomes (ADSCs-Exos) on bone defect repair. This study aimed to explore whether ADSCs-Exos and ADSCs-Exos modified tissue engineering scaffold promotes bone defects repair.

**Material/Methods:** ADSCs-Exos were isolated and identified by transmission electron microscopy nanoparticle tracking analysis, and western blot. Rat bone marrow mesenchymal stem cells (BMSCs) were exposed to ADSCs-Exos. The CCK-8 assay, scratch wound assay, alkaline phosphatase activity assay, and alizarin red staining were used to evaluate the proliferation, migration, and osteogenic differentiation of BMSCs. Subsequently, a bio-scaffold, ADSCs-Exos modified gelatin sponge/polydopamine scaffold (GS-PDA-Exos), were prepared. After characterized by scanning electron microscopy and exosomes release assay, the repair effect of the GS-PDA-Exos scaffold on BMSCs and bone defects was evaluated *in vitro* and *in vivo*.

**Results:** The diameter of ADSCs-exos is around 122.1 nm and high expressed exosome-specific markers CD9 and CD63. ADSCs-Exos promote the proliferation migration and osteogenic differentiation of BMSCs. ADSCs-Exos was combined with gelatin sponge by polydopamine (PDA)coating and released slowly. After exposed to the GS-PDA-Exos scaffold, BMSCs have more calcium nodules with osteoinductive medium and higher expression the mRNA of osteogenic related genes compared with other groups. The quantitative analysis of all micro-CT parameters showed that GS-PDA-Exos scaffold promote new bone formed in the femur defect model *in vivo* and confirmed by histological analysis.

**Conclusion:** This study demonstrates the repair efficacy of ADSCs-Exos in bone defects, ADSCs-Exos modified scaffold showing a huge potential in the treatment of large bone defects.

KEYWORDS

adipose stem cells, exosomes, tissue engineering, bone defect, Biomaterials
# Introduction

Despite the great advance of bone tissue engineering in the last few years, bone defect caused by various reasons such as trauma and infection is still a challenge for clinicians in orthopedic surgery (NIGUEZ SEVILLA et al., 2019; TOROS and OZAKSAR, 2019). Traditional methods to repair the bone defect include autografts, allografts, and xenografts, which exhibit excellent osteoconductive and osteoinductive properties, they still have several limits include repetitive surgery, immune rejection, and risk of disease transmission. Bioactive implants may provide an effective solution for this problem. To improve the bioactivity of the implants, many researchers use different substances, including polymer materials (DONG et al., 2015), ions (FERRANDEZ-MONTERO et al., 2019), growth factors (LIU et al., 2017; CABALLERO AGUILAR et al., 2019), cells (LIU et al., 2017), to modified different implant materials.

Among these, mesenchymal stem cells (MSCs) show strong osteogenesis activity in bone defect repair and have emerged as an alternative modified method to improve the implants (ANNAMALAI et al., 2019). However, MSCs have relative limitations in the maintenance of biological activity, and the logistics delivery in clinical therapies (AL-MORAISSI et al., 2019).

Recently, MSC paracrine effects are considered to be principally responsible for the tissue repair potential, and increasing interest has focussed on the exosomes (AGHAJANI NARGESI et al., 2017; PHINNEY and PITTENGER, 2017). MSCs derived exosomes show lower immunogenicity, and will not directly form tumors compared with MSCs. Using MSCs derived exosomes may get around MSCs' side effects (QI et al., 2016). Compared with stem cells, MSCs derived exosomes show a safe, low-cost and efficient manner without immune or ethical restrictions (LIU et al., 2021).

Exosomes, secreted by most cell types with a diameter of 30–180 nm, are released into the extracellular environment with the cell membrane. Exosomes can carry many bioactive molecules such as lipids, proteins, mRNAs, tRNA, lncRNAs, and miRNAs (SOKOLOVA et al., 2011; GURUNATHAN et al., 2019). Researchers found the source of MSCs influence the biological effects of MSC-derived exosomes. MSCs obtained from bone marrow (BMSCs) are the most frequently used stem cells in cell therapy and tissue engineering. BMSCs derived exosomes also promote bone regeneration (FURUTA et al., 2016). Although most research on MSC-based cell therapy focused on BMSCs, there is an increasing importance of adipose tissue as an alternative MSCs source. Compared with an equivalent amount of bone marrow, adipose tissue can provide up to 500-fold more MSCs (HASS et al., 2011). Thus, adipose tissue can be the most efficient source of MSCs (DUSCHER et al., 2017). Several studies show that exosomes derived from adipose-derived stem cells (ADSCs-Exos) promote angiogenesis, cutaneous healing, and nerve regeneration (YANG et al., 2018; HONG et al., 2019). However, the repair effect of ADSCs-Exos on bone defect remains unclear. Here, we aim to evaluate the impact of ADSCs-Exos on BMSCs osteogenic differentiation in vitro and repair effect of ADSCs-Exos after combined with materials on bone defect in vivo, to estimate its potential therapeutic value on bone defect.

# Materials and methods

## Ethics statement and animals

All the animal procedures and operations complied with the Guidelines of Shanghai Laboratory Animal Center and the Policies

on the Use of Humans and Animals in Research of the Shanghai Tenth People's Hospital (SYXK: 2014-0026) and demonstrated to the principles outlined in the Declaration of Helsinki. All the animals in this work were looked after humanely, and all efforts were made to remove their discomfort. 6-weeks and 8-weeks old male Sprague-Dawley (SD) rats were purchased from Bikai Experimental Animal Company (Shanghai, China). All the rats were housed in the standard cage with thermostatic room (25°C) in a 12 h light and 12 h dark cycle. The standard diet and water were supplied. The rats were adapted to the experimental environment at least 2 weeks before starting this study.

## Isolation and culture of ADSCs

ADSCs derived from the inguinal fat pad of 6-weeks SD rats. In brief, the capillaries were removed from the lipoaspirate, minced with phosphate-buffered solution (PBS, GenomSciences, Hangzhou, China) and digested with 1% collagenase A (Gibco, Carlsbad, United States) for 1 h at 37°C. After filtration, the mixture was centrifuged (1,000 rpm, 10 min) at room temperature (RT), and the supernatant was discarded. The collected cells were washed with PBS, centrifuged (1,000 rpm, 5 min) and then resuspended in DMEM/F12 medium (Gibco, Carlsbad, United States) with 10% fetal calf serum (FCS; Gibco, Carlsbad, United States) and 1% penicillin-streptomycin (PS) in a humidified 5% CO₂ at 37°C. When the cells passed 3 generations, they were identified and used for next experiments.

## Flow cytometry

This study used stem cell surface markers to evaluate the phenotype of cells, including CD29-PE, CD90-FITC, CD105-PE-cy5, CD45-APC (all the antibodies from Abcam, Cambridge, United Kingdom) after 1 week expansion. In brief, cells were cultured with antibodies for 30 min at room temperature, and the analysis was performed by a FACSCalibur flow cytometer (BD Biosciences, San Jose, United States). The nonspecific IgG-labeled cells were used as controls. The data were analyzed by FlowJo software (Tree Star, Ashland, OR).

## ADSCs differentiation

ADSCs ($2 \times 10^5$ per well in 6-well plates) were treated with adipogenic media to induce adipogenesis. Adipogenic medium consisted of 0.5 mM 3-isobutyl-1-methyl-xanthine, 1 µM dexamethasone, 5 µg/mL insulin, and 50 µM indomethacin. The culture medium was replaced every 3 days. Seven days later, Oil Red O staining (Beyotime Institute of Biotechnology, Haimen, China) was used to measure lipid vesicles. For osteogenic differentiation, Osteogenic medium consisted of 100 nM dexamethasone, 10 mM βglycerophosphate, and 0.05 mM L-ascorbic acid-2-phosphate, the media was changed with osteogenic induction media supplied with the differentiation media every 3 days. After 21 days, osteonectin was detected. For chondrogenic differentiation, ADSCs were cultured in a chondrogenic differentiation medium (Cyagen, Santa Clara, CA, United States).
and refreshed every 3 days. 21 days later, cartilage matrix protein was detected. All the pictures were captured by an Olympus IX51 light microscope (Olympus, Tokyo, Japan).

## Exosomes isolation and identification

The exosomes derived from ADSCs were purified from serum-free cell medium by series centrifugation and filtration steps. Briefly, collected supernatant centrifuged at 300 g for 10 min to remove dead cells, 5,000 g for 10 min to remove cellular debris. Then the supernatant was ultrafiltrated by 0.22 µm filter (Whatman, Maidstone, United Kingdom) and centrifuged at 120,000 g for 90 min. The final pellets were resuspended in 1 mL PBS and stored at -80°C. The collected pellets were distinguished by transmission electron microscopy (TEM, Zeiss, Axio, Germany) and nanoparticle tracking analysis (NTA, NanoSight LM10, Malvern Instruments, Westborough, MA).

## PKH67-labeled exosomes

ADSCs derived exosomes were labeled with PKH67 (Sigma-Aldrich, St. Louis, MO). In brief, 2 µL PKH67 was incubated with exosomes in a total of 1 mL of diluent for 15 min. 1 mL of 1% of BSA was added into the mixture to stop the labeling and was centrifuged at 120,000g for 2 h in 4°C. Then, the supernatant was discarded, resuspended the pellets in 5 mL of PBS, and centrifuged at 120,000g for 1 h in 4°C. Finally, PKH67-labeled-exosomes were resuspended in 1 mL of PBS and stored at -80°C.

## Western blot

ADSCs or exosomes were lysed in RIPA buffer (Beyotime, Shanghai, China). The samples were separated on a 10%-15% SDS polyacrylamide gel and transferred to polyvinylidene difluoride (PVDF, Millipore, Billerica, MA, US) membranes. Then the PVDF membranes were blotted with 5% BSA for 1 h and incubated with primary antibodies overnight at 4°C. The membranes were incubated with horseradish peroxidase-conjugated anti-rat IgG (1:5,000, Santa Cruz Biotechnology, US) and visualized with an enhanced chemiluminescence kit (Amersham, GE Healthcare, Waukesha, WI, US) and quantified by ImageJ software ver. 1.52. Primary polyclonal antibodies against CD9 and CD63 (1: 1,000, Cell Signaling Technology).

## Fabrication and characterization of adipose stem cells-derived exosomes modified gelatin sponge/polydopamine scaffolds (GS-PDA-Exos)

Gelatin sponges were purchased from Guangzhou Kuaikang Medical Apparatus Co. (Guangzhou, China). According to the previous study (DINH et al., 2018), the gelatin sponges soaked in dopamine (DA) solution (2 mg/mL in 10 mM Tris-HCl, pH 8.5, Sigma-Aldrich, St. Louis, US) were incubated with shaking at 37°C for 24 h to form the polydopamine (PDA) coating. To remove the non-adherent PDA, the gelatin sponge/polydopamine (GS-PDA)

scaffolds were gently shaken in an ultrasonic cleaner with distilled water for 5 times. The GS-PDA scaffolds were dried and sterilized by ethylene oxide before the next step. Then the GS-PDA scaffolds were immersed in PKH67-labeled or non-labeled ADSCs-Exos solution (10¹⁰particles/scaffold) with shaking at 37°C for 12 h. To remove the non-adherent exosomes, and the GS-PDA-Exos scaffolds were gently shaken in an ultrasonic cleaner with distilled water for 5 times. The distribution of PKH67-labeled exosomes on the scaffolds was observed with the confocal imaging system (Nikon, Japan). To measure the ADSCs-Exos release effect of GS-PDA-Exos scaffolds. The amount of ADSCs-Exos released was measured using CD63 ELISA (Beyotime, Shanghai, China) assay. Scanning electron microscopy (SEM, Hitachi, Tokyo, Japan) was used to observe the surface morphology of the scaffolds.

## Isolation and culture of BMSCs

BMSCs were isolated from the femurs and tibias of 6-week SD male rats. In brief, The bone marrow was separated from the femurs and tibias of rats by flushing with a serum-free culture medium (DMEM/F12; Gibco, USA). The red blood cells were lysed in the lysate and cultured in DMEM/F12 (containing 10% FBS) and 1% penicillin-streptomycin (PS) in a humidified 5% CO2 at 37°C. Change the medium after 24-48 h, then replace the medium once every 2-3 days later. When the cells passed 3 generations, they were identified and used for next experiments BMSCs used in this study were P3-P6 generation cells.

## Cell proliferation and migration assay

The regulation of ADSCs-Exos and GS-PDA-Exos on the proliferation of rat BMSCs was assessed by the Cell Counting Kit-8 (CCK-8) assay (Dojindo, Kyushu Island, Japan) (ZHU et al., 2019). Briefly, BMSCs were seeded into 96 well plates at a density of 4,000 cells per well, respectively. The cells were cultured with 100 µL of growth medium containing different concentrations of exosomes (0, EXO-L 1 × 10⁹ particles/mL, EXO-H 1 × 10¹⁰ particles/mL) or 100 µL of growth medium with scaffolds immersed. Each well was incubated with 10 µL CCK-8 solution and cell proliferation curves were constructed at a wavelength of 450 nm.

Scratch wound assays assessed the effect of ADSCs-Exos and GS-PDA-Exos on Cell migration. 1.5 × 10⁵ BMSCs were seeded into 6-well plates and cultured in growth medium for 12 h. Next, one scratch was made in each well using a 200 µL pipette tip. After washing with PBS, the medium was then replaced with serum-free medium supplemented with ADSCs-Exos (0, 1 × 10⁹, 1 × 10¹⁰ particles/mL), or serum-free medium with scaffolds immersed. Wound closure was measured by capturing images at different time points. The scratched areas were measured using Image-Pro Plus software.

## Alkaline phosphatase activity assay and alizarin red staining

As described above, BMSCs were seeded into 6 well plates and cultured with growth medium containing different concentrations of
exosomes or with scaffolds immersed. After 24 h, the medium was replaced with the normal or osteoinductive (OIC) medium for 10 days, At different time points (1, 4, 7, and 10 days), the alkaline phosphatase (ALP) activity assay of the supernatant was performed with an ALP Assay Kit (Beyotime, Shanghai, China). To assess mineralization, cells were induced for 3 weeks, fixed with 4% paraformaldehyde solution and washed with PBS for 3 times, then gently washed with distilled water and stained with 2% Alizarin red (ARS, Sigma-Aldrich, St. Louis, US). To quantify the coloration of ARS. 10% of acetic acid was added to each cell. After incubation for 12 h, the suspension transferred to tubes and centrifuged at 20,000 g for 15 min. The supernatant was transferred to another tube and neutralized with 10% ammonium hydroxide. Then 100 µL of each sample was measured at a wavelength of 405 nm.

RNA extraction and real-time quantitative-
PCR (RT-PCR)

RT-PCR assessed the regulation of ADSCs-Exos and GS-PDA-
Exos on the osteogenic differentiation related genes. Rat BMSCs were
grown in normal, and osteoinductive media in the presence of ADSCs-
Exos or scaffolds. The total RNA of the samples was isolated based on
the manufacturer's protocol using the RNAiso plus kit (TaKaRa,
Tokyo, Japan). The RNA was treated with DNase in a 10 µL
reaction and conducted at 42°C for 2 min. Then, the PrimeScript
RT reagent Kit (TaKaRa, Tokyo, Japan) was used in a total volume of
20 µL for the mRNAs in a 15 min incubation period at 37°C and
stopped after a 5 s enzyme denaturing step at 85°C.

Real-time amplification was performed by the SYBR Premix Ex Taq (TaKaRa, Tokyo, Japan) in an ABI 7900 thermocycler (ABI; Foster City, CA, United States). The PCR cycling was 95°C for 30 s, followed by 40 cycles of 95°C for 5 s and 60°C for 30 s. Then a dissociation curve was at 95°C for 15 s, 60°C for 15 s, and 95°C for 15 s. GAPDH was used as a relative control and analyzed using the 2 the number of $e^{-ΔΔCT}$ method. Primer sequences used in this work were described in Supplementary Table S1.

Animal experiment

The studies involving animals were reviewed and approved by
RuiJin Hospital Ethics Committee. Eight-week-old male Sprague
Dawley (SD) rats were obtained from the Bikai Experimental
Animal Company (Shanghai, China). In total, 24 animals were
randomly divided into four groups as follows: 1) empty defect
(control) 2) GS scaffold 3) GS-PDA scaffold and 4) GS-PDA-Exos
scaffold, each group has 6 animals. After adaptation for two weeks,
300~350 g SD rats were used for establishing the critical-sized rat
femoral bone defect model. Briefly, All animals were anesthetized by
intraperitoneal injecting 2.5% pentobarbital (40 mg/kg). The knee was
shaved and sterilized, after cut open one by one to expose the distal
femur, a diameter of 3.0 mm defect was made at the femoral condyle
with a slow-speed electric drill. The ice saline solution was locally used
to lower the temperature. Then the scaffolds were implanted into the
defects, and the incision was stitched.

Radiologic and histological analysis

At 2 and 4 weeks after implantation, the femurs of the animals
were harvested and evaluated the new bone formation within the bone
defect using micro-computed tomography (micro-CT; Skyscan 1,076,
Bruker, Belgium) with a spatial resolution of 12 µm. The center of the
bone tunnel of a 3 mm diameter was selected as the region of interest
(ROI). The Scanco software was used to measure bone mineral density
(BMD), bone tissue volume/total tissue volume (BV/TV), trabecular
thickness (Tb.Th) and trabecular separation/spacing (Tb.Sp). After the
micro-CT scan, the femurs were used for histology analysis. The
samples were fixed in 4% paraformaldehyde for 48 h and decalcified in
10% ethylenediaminetetraacetic acid for 4 weeks. After decalcification,
the samples were dehydrated and embedded in paraffin. Hematoxylin
and eosin (HE) staining was performed with the 5 µm thick sections.

Statistical analysis

The SPSS 17.0 software was used for statistical analyses. All data
were presented as mean ± standard deviation (SD). Differences among
groups were assessed with one-way ANOVA. The statistical
significance was *p* < 0.05. All experiments were repeated at 3 replicates.

Results

Acquisition and identification of ADSCs in SD
rats

We obtained the ADSCs by standard methods and then performed
the identification of surface markers. Flow cytometry results showed
that over 99% of cells (passage 1) were positive for the mesenchymal
stem cell markers CD29, CD90, or CD105 and negative for the
leukocyte marker CD45 (Figure 1A). The ADSCs underwent 21-
day chondrogenic, and adipogenic induction and experienced 14-
day osteogenic induction to evaluate their multilineage differentiation
potential. The ADSCs were positive for Oil Red O staining after
adipogenic induction (Figure 1B). Cell immunofluorescence showed
the ADSCs cultured in osteogenic induction positive expression of
osteonectin. After adipogenic induction, ADSCs showed the
expression of cartilage matrix protein as detected by
immunofluorescence. The results indicate that the cells we obtained
are ADSCs (Figure 1B).

Characterization of ADSCs-derived exosomes

TEM analysis showed that the exosomes isolated from the
supernatants of ADSCs is a round membranous vesicle, which
diameter of approximately 100 nm (Figure 1C). NTA analysis
indicated that the parameters of the exosomes ranged mainly from
80 to 180 nm (Figure 1D). Western blotting analysis (Figure 1E)
showed that the exosome-specific markers CD63 and CD9 were
detected in ADSCs-derived exosomes, and the expression of
abundance is high.
<figure><img src="image_60.png" /><figcaption>FIGURE 1</figcaption></figure>

Identification of ADSCs and ADSCs-Exos. (A). Characterization of ADSCs marker expression by flow cytometry analysis. ADSCs stained with CD29-PE, CD90-FITC, CD105-PE-cy5, and CD45-APC antibodies, respectively. (B). Undifferentiated and differentiated ADSCs. Phenotypic characterization of ADSCs at high confluency; ADSCs were induced to differentiate into adipogenic cells under adipogenic differentiation conditions for 21 days and stained positive for Oil Red O; Under osteogenic differentiation conditions, ADSCs formed Calcium crystals and expressed osteonectin by immunofluorescence stained; Under chondrogenic differentiation conditions, ADSCs differentiate into chondrogenic cells and immunohistochemical stained positive for cartilage matrix protein. (200x) (C). Morphology of exosomes derived from ADSCs (ADSCs-Exos) observed by transmission electron microscopy (TEM). (D). Particle size distribution of ADSCs-Exos measured by nanoparticle tracking analysis: the diameter of exosomes mainly distributed at 80–180 nm; the peak value is 122.1 nm. (E). Western blots of exosomal protein of ADSCs-Exos. Blots were probed using antibodies against CD9 and CD63.

## ADSCs-derived exosomes promote BMSCS proliferation, migration, and osteogenic differentiation

Recent research has found that exosomes are the key substance of the paracrine function of stem cells for participating in the repair process of various tissues. Therefore, exosomes may be an ideal material to improve the osteogenic differentiation ability of stem cells.

The CCK-8 assay was used to determine whether ADSCs-derived exosomes affects the proliferation activity of BMSCs. Compared with the Exos-L and CON group, the OD value of Exos-H increased significantly at 4 and 7 days, and the results showed that ADSCs-derived exosomes could promote the proliferation of BMSCs (Figure 2A). After BMSCs were treated with Exos and stimulated for 12 h, the scratch test results showed that the migration distance of BMSCs cells in the Exos-H group was longer than that in the Exos-L

group. The mobility of Exos-L was (28.33 ± 3.05) %. The migration rate of Exos-H was (45.67 ± 3.51) %, which was statistically significant compared with the migration rate of the CON group (15.00 ± 3.00) % (Figure 2B). The result suggests that ADSCs-derived exosomes can promote the migration of BMSCs cells. And the effect is related to the number of exosomes particles. ARS staining and quantitative analysis showed that compared with Exos-L and CON, the Exos-H group had the most significant number of mineralized nodules. And ADSCs-derived exosomes could promote the mineralization of BMSCs as the concentration increases (Figure 2C). The expression level of ALP in BMSCs treated by ADSCs-derived exosomes during osteogenesis induction was detected to evaluate whether the exosomes promoted osteogenic differentiation. The results showed that ADSCs-derived exosomes could promote BMSCs osteogenic differentiation, which is related to the exosomes concentration (Figure 2D). Further RT-PCR results showed that ADSCs-derived exosomes could increase mRNA
<figure><img src="image_61.png" /><figcaption>FIGURE 2</figcaption></figure>

ADSCs-Exos promote the proliferation, migration, and osteogenic differentiation of BMSCs. (A). CCK-8 assay showed that BMSCs treated by Exos-H (1 x 10¹⁰ particles/ml) cultured grows faster with culture days as compared to Exos-L (1 x 10⁹ particles/ml) or CON in conditioned-media. (B). The scratch wound assay of BMSCs treated with Exos-H, Exos-L, and CON, respectively. By 12 h, the scratch test results showed that compared with the CON group, BMSCs had longer migration distances in both Exos-H and Exos-L groups. (C). Microscopic images showing the formation of mineralized nodules. The number of BMSCs in the normal medium (NON-OIC) group increased gradually within 21 days, but no mineralized nodules observed. On day 21, mineralized nodules were found in CON, Exos-L, Exos-H groups by ARS staining. Among them, the number of mineralized nodules in Exos-L, Exos-H group is larger than that in CON, and the Exos-H group has the most mineralized nodules. (D). The activity of alkaline phosphatase (ALP) in BMSCs was detected at 1, 4, 7, and 10 days. The ALP activity of Exos-L and Exos-H group was significantly higher than that of the CON group at the time points of 7 and 10 days, and there was a significant difference between the groups of Exos-L and Exos-H. (E). The effect of ADSCs-Exos on osteogenic differentiation related genes, including RNUX2, OSX, OCN, OPN, and COL1a. BMSCs from ADSCs-Exos group cultured in OIC medium for 10 days high expressed mRNA of osteogenic differentiation related gene. (*, p < 0.05; **, p < 0.01; ***, p < 0.001).

expression of runt-related transcription factor 2 (Runx2), Osterix (OSX), osteocalcin (OCN), osteopontin (OPN), and Collagen type I (COL1) during osteogenesis induction in BMSCs (Figure 2E).

## Fabrication and characterization of GS-PDA-Exos scaffolds

As shown in Figure 3A, the GS turned brown or black after soaked in DA solution and formed the PDA coating, but the GS-PDA and GS-PDA-Exos scaffolds maintained their shape as naked GS. The form of the PDA coating did not cause damage to the structure of the fibrous. Shown in SEM pictures, the surface of the GS scaffold was smooth, but that of GS-PDA and GS-PDA-Exos scaffolds exhibited increased surface roughness. Similarly, A considerable number of PKH-67-labeled exosomes (green dots) could be found on the surface of the scaffolds with the confocal scanning microscopic imaging. The

cumulative release curve of the amount of ADSCs-Exos released from GS-PDA-Exos scaffolds is shown in Figure 3B; the ADSCs-Exos showed burst release within 3 days, followed by a relatively slow release until 7 days.

## GS-PDA-Exos scaffold promotes osteogenesis differentiation of rat BMSCs in Vitro

The proliferation of rat BMSCs were compared at days 1, 2, 3, 4, and 7. The cell proliferation of BMSCs increased with time within the 7 days monitoring span. Compared with GS and GS-PDA scaffolds and control groups, the optical density value in the GS-PDA-Exos scaffold group was significantly higher from the fourth to seventh day (Figure 4A). The migration ability of BMSCs with GS-PDA or GS-PDA-Exos was enhanced (Figure 4B). Alizarin Red S staining showed
<figure><img src="image_62.png" /><figcaption>**FIGURE 3**

Characterization of GS-PDA-Exos scaffolds (A). Three scaffolds including gelatin sponge (GS), GS with the polydopamine coating scaffold (GS-PDA), and ADSCs-Exos modified GS-PDA scaffold (GS-PDA-Exos) were prepared. Scanning electron micrographs showing the surface morphology of the scaffolds. The adhesion of PKH67 labeled ADSCs-exos on GS-PDA-Exos was observed by fluorescence microscopy. (B). The release curve of ADSCs-Exos from GS-PDA-Exos scaffold.</figcaption></figure>

GS-PDA-Exos significantly increased the calcified nodules and promoted BMSCs osteogenesis (Figure 4C). Compared with other groups, the ALP activity of the GS-PDA-Exos scaffold group was higher on day 7 and 10 days (Figure 4D). After cultured in OIC media for 10 days, all osteogenic differentiation-related genes of BMSCs, including RUNX2, OSX, OCN, OPN, and Col1 significantly upregulated in GS-PDA-Exos group than that in other groups (Figure 4D). These data demonstrated the GS-PDA-Exos scaffold could promote osteogenesis in vitro.

GS-PDA-Exos promotes bone repair in the
femur defect model *in vivo*

The critical-sized femoral bone defect model of rats were made to
analysis the effect of scaffolds on bone repair (Figure 5A). After 2 or
4 weeks of implantation, animals' femurs harvested for micro-CT and
histological tests. In the representative HE stained sections, the defect
areas were filled with newly formed tissues in GS, GS-PDA, and GS-
PDA-Exos groups. On the contrary, newly formed bone tissue was
observed only along the border of the defect in the control group

(Figure 5B). The bone defect area of the rat femoral defect was analyzed by micro-CT. As shown in Figure 5C, GS-PDA-Exos group had obvious new bone formation as compared with other groups at 2 and 4 weeks. The quantitative analysis of all micro-CT parameters, including BMD, BV/TV, Tb.Th and Tb. Sp confirmed that GS-PDA-Exos promotes bone repair in the femur defect model *in vivo* (Figure 5D).

Discussion

The exosomes are released by most cell types and involved in a
wide range of tissue regeneration, including the skin, heart, kidney,
and skeletal musculature (DONG et al., 2019; GRANGE et al., 2019;
MENDT et al., 2019). Taisuke Furuta et al. revealed that exosomes
derived by bone marrow-derived MSCs promote fracture healing in a
mouse model (FURUTA et al., 2016). In this study, we have
successfully synthesized PDA coated GS for the controlled delivery
of GS-PDA-Exos. We aimed here to elucidate whether ADSCs-derived
exosomes could promote bone repair and whether GS-PDA-Exos
could promote the new bone generation of a critical-sized defect in the
<figure><img src="image_63.png" /><figcaption>FIGURE 4</figcaption></figure>

GS-PDA-Exos scaffold promotes the proliferation, migration, and osteogenic differentiation of BMSCs (A). The CCK-8 assay detected the effects of GS, GS-PDA, and GS-PDA-Exos on BMSCs proliferation. (B). The scratch wound assay detected the effects of GS, GS-PDA, and GS-PDA-Exos on the migration of BMSCs. Compared with the CON, GS, and GS-PDA, the mobility of BMSCs in the GS-PDA-Exos group increased significantly. (C). The number of mineralized nodules of BMSCs was detected by ARS staining. The results showed a large number of mineralized nodules were observed in CON, GS, GS-PDA, and GS-PDA-Exos groups. The number of stained nodules in GS-PDA-Exos was significantly higher than that in other groups. (D). The ALP activity of BMSCs was detected after exposure to the CON, GS-PDA, and GS-PDA-Exos scaffolds. The expression level of ALP in GS-PDA-Exos increased continuously within 10 days. At 4 days, it was higher than that of the CON, GS and GS-PDA groups, but there was no significant difference, while at 7 and 10 days, ALP expression was significantly higher than that of the other groups. (E). Effects of materials on mRNA expression of BMSCs osteogenesis related genes (RUNX2, OSX, OCN, OPN, and Col1a) detected by RT-PCR. There was no significant difference between GS and GS-PDA and CON. Compared with other groups, GS-PDA-Exos significantly promoted the expression of RUNX2, OSX, OCN, OPN, and Col1a. (", no significant difference *, p < 0.05; **, p < 0.01).

SD rat femur. Based on the *in vitro* and *in vivo* experiments, ADSCs-derived exosomes contribute to bone repair *via* inducing BMSCs osteogenic differentiation *in vitro*, and the implant of GS-PDA-Exos exerted an excellent bone healing capability in the critical-sized bone defect of the SD rat femur.

Different kinds of MSCs have been used for tissue engineering and regenerative medicine. Among these, BMSCs serve as ideal seed cells for tissue engineering in numerous studies, ADSCs share many of the characteristics of BMSCs. Meanwhile, ADSCs can be obtained more easily *in vitro* and abundant storage *in vivo*. With varies advantages, ADSCs are more applicable than other MSCs to use for tissue engineering. As shown in Figures 1A, B, the characterization of ADSCs was identified by flow cytometry and adipogenic, osteogenic, and chondrogenic differentiation, as in previous study (CHEN et al., 2019). Emerging evidence has shown the crucial roles of

exosomes from MSCs promote tissue repair (CHEW et al., 2019). MSCs-derived exosomes, as a critical product of paracrine secretion, are a kind of sphere-shaped extracellular vesicle whose diameter is about 30–180 nm. Previous studies have demonstrated that exosomes secreted by human-induced pluripotent stem cells could promote bone regeneration in critical-sized calvarial defects (QI et al., 2016). Qin et al. reported that exosomes secreted by BMSCs could improve bone regeneration in a calvarial defects model (QIN et al., 2016). Thomas Thomou et al. reported that Adipose-derived exosomal miRNAs could regulate gene expression in other tissues and serve as a vital adipokine (THOMOU et al., 2017). Based on these findings, we hypothesized that ADSCs-derived exosomes could promote bone regeneration. Firstly, the exosomes were isolated from ADSCs serum-free cell medium and characterized by TEM, NTA, and western blot. (Figures 1C–E). Secondly, Our results displayed ADSCs-derived-exos
<figure><img src="image_64.png" /><figcaption>FIGURE 5</figcaption></figure>

Micro-CT and histological analysis of the effect of GS-PDA-Exos on bone repair in vivo. (A). Construction of the critical-sized femoral bone defect model of rats. (B). Representative images of hematoxylin-eosin (HE) staining of the decalcification bones slice, showing the newly formed tissue, including the fibrous tissue, newly mineralized bone tissue. (C). Representative two-dimensional micro-CT images showing the effect of different scaffolds (CON, GS, GS -PDA, and Gs-PDA-Exos) on the new bone tissue formation inside the defect site. The red dot line circled the region of interest (ROI). (D). Summarized data showing the micro-architectural parameters of the newly formed bone tissue at 2 and 4 weeks by analyzing the micro-CT images using image analysis software. Bone mineral density (BMD), bone tissue volume/total tissue volume (BV/TV), trabecular thickness (Tb.Th), and trabecular separation/spacing (Tb.Sp) were shown in the panel. (*, no significant difference *, *p* < 0.05; **, *p* < 0.01).

effectively accelerate the migration and proliferation of BMSCs (Figures 2A, B). More importantly, ADSCs-derived-exos could significantly increase the mRNA expression of osteogenesis-related genes such as RUNX2, OSX, OCN, OPN, COL1 (Figure 2E). Similarly, ALP activity was dramatically higher in ADSCs-derived-exos groups (Figure 2D). Compared with the control group, the ADSCs-derived-exos significantly enhanced the mineralized matrix production of BMSCs when culturing for osteogenesis differentiation (Figure 2C). Our results suggested ADSCs-derived-exos effectively promote BMSCs' osteogenic differentiation.

To deal with bone defects, various kinds of bone graft substitutes are being synthesized to promote bone repair (KOKUBO et al., 2004; CAO et al., 2014; KANDA et al., 2015). In this study, we use GS as a scaffold to combine ADSCs-derived-exos. GS is a partial hydrolysis product of collagen with excellent biocompatibility, biodegradability, and low immunogenicity. Besides this, high interconnected pores with macroporous morphology of the GS could enhance cell growth. These characteristics make GS an ideal biomaterial for tissue engineering. To better carry exosomes than usual physical adsorption methods, here we introduce the mussel-inspired PDA coating to adhere to and carry

exosomes. The PDA coating has strong adhesion to various substrates and could second modification by other substrates through chemical reaction (LAVOIE et al., 2005). By this method, we synthesized a new biomaterial GS-PDA-Exos. As shown in Figure 3, exosomes could be immobilized by PDA coating and slowly released from the material.

Further, we investigated the effect of GS-PDA-Exos *in vitro* and *in vivo*. *In vitro*, Compared with other groups, GS-PDA-Exos could accelerate the migration and proliferation of BMSCs (Figures 4A, B), promote the activity of ALP (Figure 4D), increase mineralized nodules formation (Figure 4C), and significantly upregulated osteogenic differentiation-related genes, including, RUNX2, OSX, OCN, OPN, COL1 (Figure 4E). Based on these findings, we further investigated the *in vivo* osteogenesis functionality of the GS-PDA-Exos with a femur defect model of rats (Figure 5A). Consistent with the observation *in vitro*, micro-CT and histological analyses showed that the GS-PDA-Exos significantly promote bone repair in the femur defect model than the control groups (Figures 5B–D).

In summary, the present study suggests that ADSCs-derived exosomes enhance the migration, proliferation, and osteogenic differentiation of BMSCs *in vitro*, and a new tissue engineering GS-
PDA scaffold combined with ADSCs-derived exosomes significantly enhanced bone repair in the femur defect model in vivo.

## Data availability statement

The original contributions presented in the study are included in the article/Supplementary Material, further inquiries can be directed to the corresponding authors.

## Ethics statement

The animal study was reviewed and approved by Ruijin Hospital Ethics Committee.

## Author contributions

GL, YZ, and JW contributed equally to this work. GL: Conceptualization, data curation, formal analysis, funding acquisition, project administration, software, supervision, validation, writing—original draft, writing—review and editing. JW: Conceptualization, data curation, formal analysis, funding acquisition, project administration, software, supervision, validation, writing—original draft, writing—review and editing. YZ: Data curation, formal analysis, funding acquisition, project administration, software. RY: Investigation, methodology, project administration, resources, software, supervision, validation, visualization. QS: Resources, software, supervision, validation, visualization. YiX: Data curation, formal analysis, investigation. BW: Data curation, formal analysis, investigation, methodology. MC: Conceptualization, data curation, formal analysis, funding acquisition, project administration,

supervision. YaX: Data curation, investigation, methodology. CZ: Conceptualization, Data curation, formal analysis, funding acquisition, project administration, supervision, validation, visualization, writing—review and editing. LW: Conceptualization, data curation, formal analysis, funding acquisition, project administration, resources, supervision, validation, visualization, writing—review and editing.

## Funding

This work was supported by the National Natural Science Foundation of China (82102618 and 32171317), Shanghai Municipal Health Commission Health Industry Clinical Research Project (20204Y0351), key project of “Star of Jiaotong University” Biomedical Engineering Foundation (YG2021ZD06) and Local projects guided by the central government (YDZX20213100003001).

## Conflict of interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

## Publisher's note

All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher.

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OPEN ACCESS

EDITED BY
Yawei Du,
Shanghai Jiao Tong University, China

REVIEWED BY
Lichen Wang,
Stevens Institute of Technology,
United States
Giovanna Desando,
Rizzoli Orthopedic Institute (IRCCS), Italy

*CORRESPONDENCE

Wei Lin,
✉ linwei@scu.edu.cn
Guanglin Wang,
✉ wglfrank@wchscu.cn

†These authors have contributed equally to this work

SPECIALTY SECTION

This article was submitted to Biomaterials, a section of the journal Frontiers in Bioengineering and Biotechnology

RECEIVED 29 December 2022

ACCEPTED 14 March 2023

PUBLISHED 29 March 2023

CITATION

Dong C, Tan G, Zhang G, Lin W and Wang G (2023). The function of immunomodulation and biomaterials for scaffold in the process of bone defect repair: A review.

Front. Bioeng. Biotechnol. 11:1133995.

doi: 10.3389/fbioe.2023.1133995

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© 2023 Dong, Tan, Zhang, Lin and Wang. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY).

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# The function of immunomodulation and biomaterials for scaffold in the process of bone defect repair: A review

Changchao Dong¹,², Gang Tan², Guangyan Zhang³†, Wei Lin⁴* and Guanglin Wang¹,⁵*

¹Trauma Medical Center, Department of Orthopedics Surgery, West China Hospital, Sichuan University, Chengdu, Sichuan, China, ²Department of Orthopedics, West China School of Public Health and West China Fourth Hospital, Sichuan University, Chengdu, Sichuan, China, ³Department of Respiratory Medicine, The 7th Hospital of Chengdu, Chengdu, Sichuan, China, ⁴Department of Gynecology, West China Second Hospital, Sichuan University, Chengdu, Sichuan, China, ⁵Department of Orthopedics, West China Hospital, Orthopedics Research Institute, Sichuan University, Chengdu, Sichuan, China

The process of bone regeneration involves the interaction of the skeletal, blood, and immune systems. Bone provides a solid barrier for the origin and development of immune cells in the bone marrow. At the same time, immune cells secrete related factors to feedback on the remodeling of the skeletal system. Pathological or traumatic injury of bone tissue involves changes in blood supply, cell behavior, and cytokine expression. Immune cells and their factors play an essential role in repairing foreign bodies in bone injury or implantation of biomaterials, the clearance of dead cells, and the regeneration of bone tissue. This article reviews the bone regeneration application of the bone tissue repair microenvironment in bone cells and immune cells in the bone marrow and the interaction of materials and immune cells.

KEYWORDS

bone regeneration, BMSCs, immune cells, biomaterials, interaction

## 1 Introduction

Bone is a dynamic, complex system composed of organic and inorganic matter. The inorganic mineral scaffold of bone is endowed with a high degree of plasticity and dynamism of the skeletal tissue through the mosaic of the living cells, the cellular matrix, blood, and the immune system.

The organic and inorganic components occupy approximately 30% and 70% of the bone in the biogenic hierarchical composite matrix (Behzadi et al., 2017). The inorganic component as a bone framework mainly comprises hydroxycarbonated apatite (HA) with a crystal structure (Zhang et al., 2007). Due to its high biocompatibility, engineered HA has been widely used in the biomedical research field of bone regeneration materials. Various cells [Bone marrow mesenchymal stem cells (BMSCs), osteoblasts, osteoclasts, immune cells] embedded in the lacunar steel pipe network of the inorganic framework secrete many functional proteins and extracellular matrix, such as type I collagen fibers, proteoglycan molecules, Osteopontin (OPN), Bone morphogenetic protein (BMP) (Behzadi et al., 2017), Phosphate regulating gene with homologies to endopeptidases on the X
<figure><img src="image_66.png" /><figcaption>**FIGURE 1**
Bone cells and immune cells in bones.</figcaption></figure>

chromosome (PHEX) enzyme. The extracellular matrix and signaling molecules constitute the organic components of the bone tissue framework. The delicate “sculpting” of the bone matrix and HA cavity framework by organic functional protein molecules is essential for bone growth and fracture repair.

Bone marrow is the principal place of organisms’ hematopoiesis. The multi-polarized bone marrow cavity mainly contains hematopoietic stem cells (HSC), bone marrow and lymphoid progenitor cells, and mature immune cells. The immune cell population includes B, neutrophils, Macrophages, and T cells (Okamoto et al., 2017). Bone and immune cells and other cells in the bone marrow share the same variable microenvironment (Figure 1). In the complex and changeable microenvironment, they interact with each other to perform the “bone immune system” function. The prosperity of osteoimmunology as a self-governed research subject has significantly contributed to the observation of increased bone resorption in every kind of inflammatory skeleton diseases, such as Rheumatoid Arthritis (RA), Osteoporosis (OP), Osteoarthritis (OA), and periodontitis (Tsukasaki and Takayanagi, 2019).

The causes of bone resorption in RA, OP, and OA are not fully elucidated. The scientific community has summarized several reasons. Some views suggest that bone loss occurs due to dysregulation of the microenvironment surrounding these lesions,

characterized by increased RANKL expression. Osteocytes, Immune cells (T cells and B cells), and Inflammatory synovial fibroblasts are thought to be important factors in the production of RANKL (Vis et al., 2013; Weber et al., 2019; Fischer and Haffner-Luntzer, 2022; Komatsu and Takayanagi, 2022). In addition, it has been reported that the increased expression of inflammatory factors (IL-6, TNF-α, etc.) is also an important cause of bone loss. On the one hand, these inflammatory factors assist RANKL in inducing osteoclast formation. On the other hand, it regulates Dickkopf-1 (Dkk-1) and sclerostin (SOST) to inhibit the Wnt signaling pathway for reducing osteoblast diffibration (Hickman and Pierson, 2016; Weber et al., 2019; Komatsu and Takayanagi, 2022). The researchers have designed a variety of excellent biomaterials to interfere with the inflammatory process in three diseases. Strontium-substituted bioactive glass (SR-BG) significantly inhibited bone loss by inhibiting p38 and NF-KB pathways downstream of RANKL (Huang et al., 2020). Chitosan, similar to the structure of biological tissue glycosaminoglycans, has made polymer and derivative forms that improve the oral utilization of the drugs of Risedronate sodium and Alendronate sodium to achieve good effects in inhibiting osteoclasts (Rahimi et al., 2022). Bioactive particles such as HAMA microspheres (Li et al., 2022), modified liposomes (Feng et al., 2020), and chitosan-modified nanoparticles (Shi et al., 2018) can be used as gene delivery vectors (IL-6-siRNA, TNF-α, etc.)
to treat arthritis and bone loss by decreasing the expression of inflammatory factors in RA and OA.

Unlike the self-repair of minor fractures, the ability to self-heal from bone damage with large defects may be limited. The migration and differentiation of osteoblast precursor cells is the key process of bone injury repair, but it is difficult for these cells to find the attachment site in the condition of large bone injury (Abbasi et al., 2020). Therefore, applying biomaterials provides a promising solution for this kind of damage. The application of Osteogenic glue and artificial periosteum has successfully repaired comminuted fracture and large bone defect models (Wu et al., 2020; Xin et al., 2020; Tang et al., 2021). These successful cases of preclinical bioactive materials will provide references for developing more clinical biomaterials applications.

The application of autologous bone, allogeneic bone, or bone substitute material has been a common strategy in the surgical treatment of bone defects. The generation of blood vessels and the invasion of osteoblast precursor cells are the keys to the successful treatment of bone defects. Insufficient vascularization and poorly cast bone matrix may lead to cell necrosis or detachment of bone substitutes (Gotz et al., 2012; Zheng et al., 2022). Thus, the coupling of angiogenesis and osteogenesis is the key to the successful repair of bone defects. Autocrine and paracrine factors determine the interaction between osteogenesis and angiogenesis. Factors such as Vascular endothelial growth factor (VEGF), Hypoxia-inducible factor (HIF), and Osteopontin/tin can act bidirectionally on endothelium and osteoblasts (Gotz et al., 2012). It is beneficial for angiogenesis and new bone formation. Materials biologists have combined these factors to design a variety of advanced biomaterials that are different from conventional implantation therapy. Cross-linked engineered VEGF (TG-VEGF) modified fibrin matrix significantly promoted early vascular invasion and osteogenic differentiation of critical-size skull defects (Burger et al., 2022). In addition, exosomes from BMSCs carrying HIF-1α load onto traditional β-TCP scaffolds contribute to new bone regeneration and Neovascularization in critical-size bone defects (Ying et al., 2020). With the further study of bone immunology, the combination of immune factors has been introduced into new bioactive scaffolds for osteogenesis and angiogenesis. 3D-printed PCL scaffolds of Plla electrospun microfiber (3D-M-EF) and nanofiber (3D-N-EF) composites have the ability to modulate macrophage M2 polarization, and VEGF and BMP-2 are secreted by M2 to promote angiogenesis and osteogenesis of rat skull defect (Liu et al., 2021). Various metal particles have been studied for the interaction between immune cells and bone regeneration (Zhao et al., 2021a; Wan et al., 2022; Wu et al., 2022; Li et al., 2023). Therefore, the intervention of immune microenvironment materials will open a new chapter for the repair strategy of bone regeneration.

Innovation in biomedical technology will provide a better basis for bone defect repair, 3D printing technology has been successfully applied to materials science, optics, Robotics and chemistry, and other scientific fields. With the application of 3D printing in biomedicine, it has been successfully applied to print living cells or specific tissues, but there are some challenges in the application of 3D printing technology. The choice of printing ink determines its Biocompatibility and cellular distribution. In addition, changes in forced shear and photothermal induction during 3D printing may have a greater impact on cell viability. The biggest problem with 3D

printing is that it cannot respond to the biological needs of the tissue. These reasons led to the development of 4D bio-printing technology. Materials scientists have studied 4D bio-printing techniques that are programmable, multi-material combinatorial, and bio-tissue responsive to adaptation. Although the 4D printing technology has great challenges in easy manufacturing, economy, and extensibility, its considerable biomedical application prospect is worth paying attention to (Osouli-Bostanabad et al., 2022).

Various immune cells (T, B, Dc) regulate bone homeostasis. Moreover, Various molecules (cytokines, chemokines, and signaling molecules) in the bone microenvironment are regulated and utilized by various cells in the bone marrow (Okamoto et al., 2017). In addition, A growing body of evidence suggests that bone and immune cells regulate each other to regulate hematopoietic function further. Regarding the evolutionary perspective, the simultaneous development of acquired immunity and the skeletal system maybe lead to shared mechanisms and close the effect of skeletal cells and immune cells. Further, various cells and molecules interact with each other to regulate their development.

The interaction between the immune system and the biological material relies on the relationship between the graft and the surrounding tissues, mediating processes such as cell-specific innate defense and adaptive immune response. It is increasingly evident that macrophages residing in tissues or macrophages recruited from other regions play different parts in the healing stage, and the same material implanted in different parts will cause different effects (Reid et al., 2015). However, a common feature of biomaterials is to induce unfavorable immune responses, leading to excessive inflammation, tissue destruction, healing disorders, fibrotic encapsulation, and even isolation and rejection of medical materials. Resolving these side effects requires further exploration of the interaction between the material and the surrounding tissue microenvironment, which is both a challenge and an innovation.

## 2 Effects of the immune system on bone marrow cells

### 2.1 Bone marrow cells

Continuous bone remodeling occurs in the bones throughout the organism's life cycle. Osteoblasts, osteoclasts, and osteocytes work together to complete the dynamic bone remodeling of bones (Figure 2) (Naik and Wala, 2013; Behzadi et al., 2017). Bone remodeling involves the Interplay of the time and space of osteoblast, which leads to bone formation, and osteoclast, which leads to bone resorption (Naik and Wala, 2013).

Osteoblasts are clusters of bone marrow mesenchymal stem cells covering the bone's surface. They are metabolically active and can synthesize and secret collagen and non-collagen bone matrix proteins deposited between bone cells and the surface. These uncalcified deposits are called osteoid, and the osteoid mineralization cycle is 10 days. BMSCs can differentiate into osteoblasts. The process involves the expression of Runx2 and Osterix transcription factors based on external stimuli such as Parathyroid hormone (PTH), Prostaglandin E 2 (PGE2), and Insulin-like Growth Factor (IGF) (Tsukasaki and Takayanagi,
<figure><img src="image_67.png" /></figure>

<figure><img src="image_68.png" /><figcaption>FIGURE 2</figcaption></figure>

Bone remodeling and its involved cells. (A): Four types of cells are found within bone tissue. (B): Interaction between Osteoblasts-Osteoclasts-Osteocytes during bone remodeling. The graph was reprinted with permission from (Behzadi et al., 2017) and (Dar et al., 2018).

2019). Moreover, BMP-2 and Wnt signal pathways are responsible for osteoblast or BMSCs differentiation.

Osteocytes connect through a small tube connected to the bone surface and form a network, the dense signal communication path in the bones. Osteocytes are derived from bone osteoblasts, then wrapped in the bone matrix. However, Osteocytes begin to express some specific genes, but osteoblasts do not express these genes. Sclerostin is a product of osteocytes, which can bind to low-density lipoprotein receptor-related protein (LRP) and inhibit the Wnt signal pathway to reduce bone formation (Poole et al., 2005). Function Sclerostin (SOST) gene encoding sclerostin mutations results in an increased bone mass loss in humans, called sclerosis.

Osteoclasts originate from HSCs. HSCs first develop into monocytes. Then monocytes develop into mature osteoclasts under the stimulation of related factors. Osteoclast production requires macrophage colony-stimulating factor (M-CSF) and receptor activator of nuclear factor-κB ligand (RANKL) as induction signals. Osteogenic precursor cells, stromal cells, and synovial endothelial cells express M-CSF and RANKL to induce osteoclastogenesis. Along with BMMs, they differentiate and form mature osteoclasts. At this time, osteoclasts also begin to express specific genes and fuse. Moreover, RANKL is expressed on preosteoblasts and activated T cells (Wada et al., 2006; McInnes and Schett, 2007). RANKL combines its receptor RANK with pro-osteoclast cells, which is grave for the differentiation of osteoclasts and their bone resorption capacity. The interaction between RANKL and RANK is regulated by the RANKL competitive receptor OPG, which inhibits osteoclast production in vitro and in vivo. In addition to the RANKL and RANK affecting each other, other key pro-osteoclast signaling pathways depend on trigger receptors (TREM) expressed on bone marrow cells. The OSCAR and the tyrosine kinase DAP12 interact with triggering receptors expressed on myeloid cells (TREM), which strongly promotes osteoclastogenesis (Barrow et al., 2011).

## 2.2 Immune system

The immune system is essential for organisms to defend against foreign objects. In addition, the invasion of pathogens can also stimulate the evolution of the immune system. The body's adaptability and innate immune response participate in this process. As mentioned above, both immune and bone cells are present in bone. Meanwhile, the two systems influence each other (Lee et al., 2019) (Figure 3A). Bone cells and their surrounding environment secrete chemokines [Macrophage chemoattractant protein-1 (MCP-1), Stromal cell-derived factor 1 (SDF-1), etc.] that attract the arrival of immune cells, which subsequently secrete cytokines (RANKL, IL-1, IL-6, IL-10, TNF-α, etc.) that affect the development of bone cells. Thus, basic and clinical researchers explore the interaction between immune cells and their factors with bone cells (Naik and Wala, 2013). Although immune cells are various, they originate from a common ancestor-HSCs. HSCs are finally differentiated into T cells, B cells, macrophages, NK cells, and neutrophils through pluripotent stem cells, myeloid stem cells, and lymphoid stem cells and enter different tissues or organs with the bloodstream to exert their physiological functions further.

### 2.2.1 Macrophages

Macrophages belong to the mononuclear phagocytic cell system. They also originate from HSCs. In the bone marrow, HSCs develop into monocytes through immature monocytes, enter connective tissues and other organs with the bloodstream, and then turn into macrophages. The body's homeostasis processes include inflammation, foreign body removal, and bone tissue repair (Figure 3B) based on macrophages' differentiation and function (Xie et al., 2020).

Macrophages are a class of highly plastic immune cells whose main function is to play the role of phagocytosis. It engulfs foreign
<figure><img src="image_69.png" /></figure>

bodies (bacteria, viruses, nanoparticles, etc.) that invade the
organism and engulfs the dead and apoptotic cells it produces.
Phagocytosis of different substances by macrophages produces
different body responses. Macrophages form the “classical
activated” M1 phenotype after phagocytosis of pathogens and
dead cells, which mainly plays a pro-inflammatory role, whereas,
when macrophages engulf apoptotic cells, they mainly form
“alternatively activated” M2 phenotype, which promotes tissue
repair by secreting repair factors such as IL-10 and TGF-β (Xie
et al., 2020; Schlundt et al., 2021). In bone defect repair, plasma
proteins and foreign bodies attract macrophages to the defect. In the
early phase of inflammation, macrophages transform into
M1 macrophages, which in the early phase play a clearing role of
foreign bodies and dead cells, and subsequently, the transformation
of macrophages into M2 type initiates the process of bone tissue
regeneration. Therefore, the ratio of M1/M2 macrophages is a key
process in the transition from inflammation to regeneration (Zhao
et al., 2021b).

Compared with monocytes, multi-nucleated cells formed by the
fusion of macrophages have a phagocytic affinity for significant
substances. The ability of macrophages puts them at the center of the

evolutionary and functional roles of the bones and immune system
because monocytes can form granulomas and other giant cells in
inflammation regions, and they can also fuse in bones to form large
multi-nucleated cells called osteoclasts. The decision to proceed with
osteoclast forming is only controlled by the local cells factor
environment of RANKL (Dar et al., 2018). Part of macrophages
participate in the fight against harmful damage at the site of tissue
inflammation; they can perform multiple functions. Inflammatory
macrophages can clear apoptotic cells, proliferate and resident
matrix, infiltrate white blood cells, and kill parenchymal cells
(Naik and Wala, 2013). In addition, heterotopic bone formation
in fibrodysplasia ossificans progressiva (FOP) patients is thought to
be triggered by inflammation such as macrophages and mast cells l
(Convente et al., 2018).

2.2.2 T cells

T cells are lymphoid stem cells derived from the bone marrow.
They are a group of cells that develop and mature after migrating
into the thymus. It mainly includes CD4 T cells and CD8 T cells.
CD4 T cells mainly develop into Th cells. Naïve Th cells can
transform into subgroups according to the antigens and cytokine
stimuli they receive during activation. And then, Th cells can be well divided into four subgroups, namely, Th1, Th2, Treg, and Th17 cells. Unactivated Th cells can inhibit the formation of osteoclasts *in vitro* (Toraldo et al., 2003) or *in vivo* (Li et al., 2007). Under steady-state conditions, Th cells cannot secrete RANKL (Dar et al., 2018). In contrast, the activation of T cells under inflammatory conditions can lead to increased production of RANKL and Tumor Necrosis Factor-α (TNF-α), which promote osteoclast production and bone loss under kinds of inflammatory and autoimmune conditions (Colucci et al., 2004), such as periodontitis (Brunetti et al., 2005), cancer (Colucci et al., 2004) and osteoporosis (Faienza et al., 2009). However, not all kinds of T cells can activate osteoclasts. It is reported that CD8⁺ T cells have osteoprotective effects. CD8⁺ T cells can inhibit osteoclastogenesis by secreting OPG (Dar et al., 2018).

Th1 cells can secret Interleukin-2 (IL-2), Interleukin-12 (IL-12), interferon-γ (IFN-γ), and TNF-α to participate in the clearance of pathogens. Moreover, Th2 cells can secret IL-4, IL-5, IL-6, IL-9, and IL-13 cytokines to participate in the clearance of parasitic infections, extracellular microorganisms, and allergic diseases. It is now known that Th1 and Th2 cells can inhibit the formation of osteoclasts in different populations by secreting their iconic cytokines IFN-γ and IL-4, respectively (Harrington et al., 2006). In the past decade, two new CD4⁺ Th cell subgroups have been reported, which were Th17 cells (Harrington et al., 2006) and Treg cells. Roryt transcription factor defines Th17 cells, and the Foxp3 transcription factor specifically defines Treg cells. They have been confirmed that these two kinds of Th cells are the primary T cells that regulate osteoclast production (Dar et al., 2018).

Transforming growth factor-β (TGF-β), IL-6, and inflammatory stimuli induce Naïve Th cells to transform into Th17 cells. Th17 cells produce IL-17, the hallmark factor of Th17 cells, as well as IL-22, IL-26, and IFN-γ (Harrington et al., 2006; Gao et al., 2007). Th17 cells induce osteoclastogenesis by producing IL-17, a common factor that promotes osteoclastogenesis by inducing RANKL (Adamopoulos et al., 2010). However, IL-17 can promote osteoclast production through direct and indirect action, mainly through two inflammatory factors, TNFα and IL-1 (Dar et al., 2018). These cytokines in cells enhance the expression of RANKL that support osteoclastogenesis, activating pro-osteoclast through the RANKL-rank signal pathway. In the occurrence of various bone diseases, Th17 cells are involved, such as RA (Figure 3C) (Tsukasaki and Takayanagi, 2019), periodontitis (Figure 3D) (Tsukasaki and Takayanagi, 2019), osteoporosis, and OA (Maddur et al., 2012).

It has been reported that a specific type of TH17 cells-exFoxp3TH17 cells, compared with TH17 cells, can activate osteoclasts. Treg cells express the specific molecule Forkhead Box Protein P3 (FOXP3). In arthritis animals, when CD25loFoxp3+T cells lose FOXP3 expression and start to transform into TH17 cells, inflammation stimulates IL-6 produced by synovial cells. Importantly, Foxp3+IL-17+T cells are particularly prominent in the synovial tissue of patients with active RA but not detected in the synovial fluid of inactive patients, which has been observed in membrane tissue. It indicates the pathogenic effect of exFoxp3TH17 cells in RA. Furthermore, the increase of IL-6 secreted by synovial fibroblasts contributes to the production of exFoxp3TH17 cells. In turn, the increase of IL-17 can stimulate the production of CCL20, IL-6, granulocyte-macrophage colony-

stimulating factor (GM-CSF), and RANKL in synovial tissue, thereby exacerbating local inflammation and joint destruction (Hirota et al., 2018). Therefore, Bone-destructive T cells and synovial tissue's malignant circulation mechanism is an effective research field for treating bone diseases.

CD4⁺ Treg cells and TH17 cells have opposite effects. CD4⁺ Treg cells can suppress osteoclast differentiation and function by secreting a kind of cytokines which include TGF-β and IL-4 cytokines, in a cytokine-dependent manner. Recently, In the ovariectomized osteoporosis model, by producing IL-10 and TGF-β1, Treg cells can reduce osteoclast differentiation and bone resorption. Another reported mechanism is that Treg cells can produce Cytotoxic T Lymphocyte-Associated Antigen-4 (CTLA-4) to control immune function, bind to CD80/CD86 on pre-osteoclast cells, and inhibit inflammation (Dar et al., 2018). According to reports, oral probiotic Lactobacillus rhamnosus GG can stimulate osteogenesis through the Wnt process of Treg cells acting on CD8+T cells. In addition, in the process of bone healing, it is shown that Vγ6+γδ T cells produce IL-17 and promote bone formation (Ono et al., 2016). These findings demonstrate the critical role of immune cells in calcification formation.

NK T cells can clear viral infections, transformations, or abnormal cells. Activating natural NK T (iNK T) can cause active osteoclasts. In the synovial fluid of a confirmed RA patient, NK T cells account for 20% of all lymphocytes. Moreover, CD56bright NK T cell subgroups upregulate many adhesion molecules and chemokine receptors, which can help preferentially recruit NK T cells to RA In the patient's inflamed synovium. NK T cells also produce M-CSF and RANKL, which are beneficial for inducing osteoclastogenesis (Dar et al., 2018).

### 2.2.3 B cells

The role of B cells with other immune cells and bone cells is complex and diverse (Dar et al., 2018). B cells are differentiated from HSCs in the skeleton. The osteoblast cells, which are in the skeleton niche, can support HSC and B cell differentiation. B cells play a vital function in the pathogenesis of RA. High serum RA factor and Anti-Citrullinated Protein Ab (ACPA) are related to the course of aggressive bone injury. Immune complexes, including ACPA, have been seen to enhance osteoclast differentiation by activating Fcγ receptors on Osteoclast progenitor cells (OPCs) (Harre et al., 2015). It is a rare immune complex function separated from the host's defense environment. ACPA also accelerates osteoclast production through its Fab fragments. The citrulline epitopes on OPCs and osteoclasts bind to ACPA to stimulate TNF and IL-8 production, thereby enhancing osteoclast production in an autocrine manner (Krishnamurthy et al., 2016).

It has been reported that both B cell and B cell-derived plasma cells can express RANKL, decoy receptor 3 (DcR3), or IL-7 secretion (a potent bone resorption stimulator), which indirectly regulates osteoclastogenesis. Plasma cells derived from the tumor in patients with multiple myeloma can reduce bone formation by producing cellular molecules (like dickkopf-related protein 1 and sclerostin) (Dar et al., 2018). The lack of estrogen can cause the increase of B lymphocytes, and the corresponding treatment can inhibit the increase of B lymphocytes. Interestingly, the immature B cell population that expresses B220 can even be transdifferentiated into the osteoclast differentiation pathway under *in vitro*
conditions, which provides a source of osteoclast precursors for the bone loss caused by ovariectomy. It has been shown that the RANKL expression of B220+ cells is enhanced in ovariectomized mice. In addition, it has been reported that postmenopausal mammalian-derived B cells from the bone can secrete RANKL, and B cells have a positive role in OP, like OPG, whose lack can lead to increased osteoclast production. In addition, the effect of B and T cells can regulate the forming of many bone cell factors because B cells inhibit osteoclast production by Th1 cell activation but promote osteoclasts when it comes to Th2 cell s activation (Li et al., 2007). The two types of cells can also influence the OPG from B lineage cells through the CD40/CD40L interaction pathway, which limits basal bone resorption in vivo (Yamamoto et al., 2019).

### 2.2.4 Dendritic cells (DCs)

DCs are highly effective APCs and are key in treating pathogens and tumors mediated by cellular immunity. Under the action of immune cell CD4 T cells, mouse CD11c⁺DCs can successfully transform into functional osteoclasts and induce osteoclast-mediated bone loss (Dar et al., 2018). These findings suggest the strong effect of CD11c⁺DC subpopulations in osteoclast forming. The RANK-RANKL signaling pathway also plays a role in the osteoclast transformation process of DCs (Tucci et al., 2011). In addition, mature DC can promote Th17 cells to express IL-17, enhancing osteoclastogenesis (Dhodapkar et al., 2008).

### 2.2.5 Neutrophils

Neutrophils are derived from hematopoietic stem cells of bone marrow. After differentiation and development in bone marrow, neutrophils enter the blood or tissues. In the start stages of inflammation caused by bacterial infections, environmental exposures, and certain cancers, neutrophils are essential to innate immunity. In addition, high concentrations of RANKL are also derived from the neutrophils of the mammalian immune system, thereby activating bone loss (Dar et al., 2018). It has been further reported that neutrophils can regulate osteoblast function, leading to increased bone resorption (Brunetti et al., 2013).

## 2.3 Bone marrow cells regulation of HSCs and immune cells

The balance of HSCs in the bone is inseparable from the role of cytokines like Stem cell factor and chemokine C-X-C ligand12 (CXCL12). Since CXCL12 is produced in the body by abundant reticulocytes and leptin receptor-positive osteoblasts or osteoprogenitor cells instead of mature osteocytes, it is an essential source of these niche factors regulated by HSCs (Zhou et al., 2014). Immune cells originate from the bone. Osteoblasts modulate the differentiation of immune cells beyond doubt. For example, it has been reported that consuming osteoblasts' CXCL12 reduces the number of B lymphoid progenitor cells in the bone marrow. Moreover, osteoblasts express Notch ligand delta-like 4, which helps support the growth of T cell progenitor cells (Yu et al., 2015). Under certain conditions, osteoblasts may also have different roles in the process of hematopoiesis. For example, bone cell-derived Dickkopf-1 (DKK1) directly promotes hematopoietic reconstitution after bone marrow suppression by inhibiting HSCs

senescence and indirectly promotes hematopoietic reconstitution by inducing the secretion of bone epidermal growth factor. It is reported that bone marrow endothelial cells (Himburg et al., 2017) and osteoblasts can regulate red blood cell production. Activated osteoblasts can weaken the development of leukemia (Krevvata et al., 2014). Activated CD4⁺ T cells reduce the bone formation of osteoblasts and lead to imperfect B-cell lymphocyte production, which proves an interaction between bone and immune cells (Shono et al., 2010).

The osteoclast bone resorption produces the bone marrow cavity where the secretion of osteoblasts, bone cells, and other cells are attached. Due to the abnormal balance of osteoclastic and osteogenic differentiation in osteosclerotic animals, there is insufficient space in the bone marrow to support the differentiation of hematopoietic cells, ultimately leading to abnormal extramedullary hematopoiesis in the spleen and liver. This abnormal process is detrimental to the formation of immune cell differentiation and function (Okamoto et al., 2017). Patients with bone sclerosis may appear with anemia and infections due to abnormal hematopoietic function (Sreehari et al., 2011). Therefore, the cavity microenvironment formed by osteoclast resorption is necessary for normal functional hematopoiesis. Moreover, osteoclasts play a certain role in the mobilization of HSCs.

# 3 Interaction of materials and immune cells

## 3.1 Biomaterials

With the development of biomaterials, their status in medical care is getting higher and higher. At the same time, many new disciplines have been produced, and huge development and applications have been achieved, such as medical implants, drug delivery, tissue engineering, and immunotherapy (Vishwakarma et al., 2016; Roseti et al., 2017). Biomaterials include a series of compounds with very different functions and structural characteristics, from macromolecules taken from nature to completely synthetic nanoparticles. Biomaterials should be mechanically resilient, Biocompatibility, or capable of degradation in the time required for bone cell colonization and mineralization. Biocompatibility is the primary target of choice for bone implant materials. Moreover, bone is a load-bearing organ with certain mechanical strength, and biomaterials with the same Young's modulus as bone are highly sought after. For repairable bone injuries, biomaterials are generally selected to be biodegradable at a rate comparable to the bone healing cycle (Borchers and Pieler, 2010; Gorejová et al., 2018; Zhu et al., 2022).

There are many classification methods for biomaterials, which are classified into natural polymers and synthetic polymers according to polymers. The most commonly used tissue regeneration materials, especially orthopedics materials, like synthetic polymers, ceramics, and natural polymers (Mariani et al., 2019). Ceramic materials (glass, alumina, zirconia, calcium phosphate) are mainly used in orthopedics, dental implants, and bone filling. They have the characteristics of low elasticity, high-temperature resistance/hardness, and high brittleness. The chemical structure and physical properties similar to natural bone tissue show
<table><caption>TABLE 1 The effects of material properties on cell behavior.</caption><thead><tr><td>Biomaterials</td><td>Features</td><td>Function</td><td>Ref</td></tr></thead><tbody><tr><td>PCL/P3ANA-RGD nanofibers</td><td>Higher surface area and better mechanical properties</td><td>Favored cell attachment, proliferation, and osteogenic activity</td><td>Guler et al. (2017)</td></tr><tr><td>Laminin-PHB fibrous</td><td>Simulation of surface morphology and chemical composition of extracellular matrix (ECM)</td><td>The adhesion and proliferation of nerve cells are promoted</td><td>Sangsanoh et al. (2017)</td></tr><tr><td>PEDOT/Cs/Gel scaffolds</td><td>The incorporation of PEDOT on the scaffold increased the electrical conductivity, hydrophilicity, mechanical properties, and thermal stability</td><td>The adhesion and proliferation of neuron-like cells are promoted</td><td>Wang et al. (2017)</td></tr><tr><td>PLGA-GNP fibers</td><td>Poly (lactide-co-glycolide) (PLGA) can effectively control the loading of gold nanoparticles (GNP)</td><td>Osteogenic differentiation of human adipose-derived stem cells is increased</td><td>Lee et al. (2018)</td></tr><tr><td>β-Tricalcium phosphate (TCP) scaffolds decorated with polydopamine nanoparticles PDA-NPs</td><td>Affinity for a variety of proteins and peptides</td><td>Excellent osteoinductivity and bone-regeneration</td><td>Wang et al. (2016)</td></tr><tr><td>Cylindrical PVA hydrogel</td><td>Stiffness gradient</td><td>Investigate the effects of substrate stiffness on stem cell differentiation into specific cell types</td><td>Kim et al. (2015)</td></tr><tr><td>Randomly-oriented fibrous scaffolds</td><td>Different topographic materials utilized as bio-scaffolds in vivo</td><td>Cytoskeletal tension release abrogated the divergent differentiation pathways on different substrate topography</td><td>Yin et al. (2015)</td></tr></tbody></table>

Biomaterial polymers include natural and synthetic biomaterial polymers (Wei et al., 2020) (Table 2).

<table><caption>TABLE 2 Classification of biological materials.</caption><thead><tr><td>Natural biomaterial polymers</td><td>Synthetic biomaterial polymers</td></tr></thead><tbody><tr><td>Collagen</td><td>PCL, PGA</td></tr><tr><td>Chitosan</td><td>PLGA, PHB</td></tr><tr><td>Fibrin</td><td>Biodegradable Ceramics</td></tr><tr><td>Silk Fibroin</td><td>Bioactive glasses and biodegradable metals</td></tr></tbody></table>

good biocompatibility (Mariani et al., 2019). Synthetic polymers as promising biomaterials for bone tissue engineering research and use. Moreover, natural polymers similar to natural ECM make them great biocompatible. They combine synthetic and natural polymers' advantages, including immunological properties. The morphology and physical properties of the materials have different effects on cell behavior (Table 1). With the development of material technology, biodegradable biomaterials are particularly popular in bone tissue engineering.

The most common natural biomaterial polymers are collagen, chitosan, fibrin, and silk fibroin, which have good histocompatibility and low immunogenicity due to their close similarity to animal tissue components. Decomposition products are also the best raw materials for the body's biosynthesis, so they are prevalent in tissue engineering. However, these single materials have poor mechanical strength and rapid degradation, and their application in bone tissue is limited. Therefore, these materials are often combined with bioceramics and synthetic polymers.

Synthetic biomaterial polymers such as poly(ε-caprolactone) (PCL), polylactic acid, PGA, copolymer PLGA, poly(3-hydroxybutyrate) PHB, biodegradable ceramics, bioactive glasses, and biodegradable metal materials, which are biocompatible and have a controllable rate of degradation, and their degradation

products have no toxic effects on tissues in vivo. Moreover, manual control of design and synthesis parameters can produce polymers with better mechanical properties.

## 3.2 Immune cell responses to biomaterial implants

Preparing the materials mentioned above into biodegradable composite materials is the focus of current research. These composites possess excellent biocompatibility, osteoconductivity, mechanical strength, and osteogenic properties. Meanwhile, these composite materials have become the most promising materials in bone defect repair with the help of new preparation technologies that have emerged in recent years. A systematic understanding of the specific processes of materials in bone regeneration in the emergency immune response is the practical basis for designing

<figure><img src="image_70.png" /><figcaption>FIGURE 4
Immune cascade after bone implant material, the graph was reprinted with permission from (Xie et al., 2020).</figcaption></figure>
composite materials for immune cell regulation (Xie et al., 2020) (Figure 4).

The implant is placed a second later, and blood from the damaged blood vessel surrounds the biomaterial and begins interacting with the graft. And then, Plasma contents of the body, including proteins (fibrinogen, albumin, vitronectin, fibronectin, gamma globulin), attach to the implant surface quickly and autonomously (Wilson et al., 2005). These are affected by the process, quantity, composition, and conformational changes of adsorbed molecules of biological materials.

The clot formed by blood exudate defines the temporary matrix surrounding the biological material (Ekdahl et al., 2011) and is cleaved into fibrin by thrombin. Moreover, the complement protein is activated when it comes into contact with biological materials to support platelet adhesion and activation. The recruitment and adhesion of sufficient immune cells are seduced by abundant pro-inflammatory cytokines, chemokines, and growth factors (Anderson et al., 2008), and with the formation of the temporary matrix, acute and chronic inflammation also follows.

The chemokine produced by the host cell or damaged tissue induces activated neutrophils to be recruited from the peripheral blood and adhere to the implantation site (via β2 integrin). It also attempts to produce engulfing proteolytic enzymes and ROS to destroy/degrade biological materials (Grandjean-Laquerriere et al., 2007).

After neutrophils are activated, they will synthesize a large number of immunomodulatory signals (Mariani et al., 2019): CXCL8 (the most significant chemokine, the main target of which is the neutrophil itself), CCL2 (C. Chemokine ligands2), and CCL4. CCL2 and CCL4 are both effective chemotactic and activating factors for immune cells (Yamashiro et al., 2001). The gradual increase of these chemokines promotes the infiltration of monocytes and inhibits the infiltration of neutrophils. The lack of this signal cannot further activate neutrophils, enter the apoptotic pathway, and then be cleared from the site by phagocytes (Anderson et al., 2008). At the same time, circulating monocytes respond to chemotactic agents and bind to the fibrinogen in the temporary matrix of biological materials, thereby being activated (Shen et al., 2004) and differentiated into classical activation or “M1” macrophages (Mariani et al., 2019). The classification of these cells is based on the secretion of IL-1β, IL-6, TNF-α, chemokines (Jones et al., 2007; Mesure et al., 2010), and enzymes.

Macrophages induce invasion of inflammatory cells via CCL2, CCL4, and CXCL8 (Jones et al., 2007), and before experiencing the “frustrated” phagocytosis (because the biological material is too large), they try to decompose the biological materials by producing ROS and releasing degrading enzymes (Mariani et al., 2019). This pathway eventually leads to increased cytokine release (Underhill and Goodridge, 2012). Like the wound healing stage (Mariani et al., 2019), the adherent macrophages may eventually transfer to the “M2” phenotype (Mariani et al., 2019), which secrete IL-10 and so on (various anti-inflammatory cytokines). These cytokines can reduce their degradation ability, complete body activity reproduction, induce fibroblast movement and proliferation, and then achieve bone regeneration. Conversion between M1 and M2 and the mechanism of blocked phagocytosis leads to the fusion of macrophage membranes and the formation of foreign

body giant cells (FBGCs), a sign of chronic inflammation on the coverage of biological materials. FBGCs formation is usually a landmark part of Foreign body reaction (FBR) induced by biological materials. It promotes the formation of FBR by activating mast cells, basophils, and Th cells. These cells can produce IL-4 and IL-13, enhancing the fusion of macrophages on biological materials (Brodbeck et al., 2005).

In the chronic inflammatory stage, some cytokines are secreted by Th cells. Their rich cytokines produce a wide range of regulation of pro-inflammatory or anti-inflammatory sequels (Mariani et al., 2019). The interactions between M1 and M2 macrophages and the changes of Th1 to Th2 cells express cell factors, indicating that T cells play a vital role in promoting the resolution and regeneration of inflammation.

The synergy of immune cells induces the production of pro-fibrotic factors, such as Platelet-derived growth factor (PDGF) (Shen et al., 2006), vascular endothelial growth factor (VEGF) (Chen et al., 2010), and TGF-β (Garg et al., 2009), which can recruit some fibroblasts. Experimental repair of damaged tissue, activated type I and type III collagen. Fibroblasts are responsible for accumulating new substrate, which is the process of fibrosis reaction.

Regarding the mechanism of promoting regeneration, M2 macrophages with anti-inflammatory/anti-fibrotic phenotypes promote regeneration through mutual interference with regulatory T cell subsets (Tregs) that play a vital part in the immune system. These cells regulate the tilt of the local immune response to inflammation, which further promotes the cascade of tissue regeneration and repair. Moreover, maintain anti-inflammatory and anti-fibrotic phenotypes by secreting IL-10. In addition, Tregs can enhance the ability to heal by inducing type 2 responses. After the reduction of T cells, the level of resident Tregs is still elevated, which may be because they lack epidermal growth factor receptor (EGFR) (Zaiss et al., 2013; Arpaia et al., 2015), whose expression can promote the cell factors secreting by mast cells to keep the Tregs in the damaged spot (Zaiss et al., 2013). Once Treg cells appear, they will proliferate and upregulate the secretion of bimodules, which is necessary for cell regeneration and may produce cell proliferation or induce cell differentiation. Treg cells may also enhance the regeneration ability of endogenous stem cells and progenitor cells through growth factors secret.

# 4 Immunomodulatory applications of biomaterials in skeletal disease

The exploration of immunomodulation in biomaterials in regenerative medicine has attracted much attention. An in-depth understanding of materials' immune cascade reaction principle (Xie et al., 2020) (Figure 5A) is critical in developing and innovating new materials.

## 4.1 The application of neutrophils in the design of biomaterials

Although neutrophils do not make up much in the bone marrow, they are the first cells to be mobilized in inflammatory
<figure><img src="image_71.png" /><figcaption>FIGURE 5</figcaption></figure>

Immune cascade after bone implant material. (A) The interaction of biomaterials, skeletal cells, and immune cells, the graph was reprinted with permission from (Xie et al., 2020). (B–D) Zn²⁺ and BMP-2 peptide-modified titanium screws promote macrophage M2 polarization and synergize with BMP-2 to promote bone formation. The graph was reprinted with permission from (Wang et al., 2022).

and traumatic fractures. Neutrophils, pioneer cells of the inflammatory response, play a dual role in bone regeneration and repair. On the one hand, neutrophils eliminate cell debris, infectious substances, and phagocytic material scaffolds at the injury site by secreting proteases (e.g., matrix metalloproteinases, collagenases) and provide sites for subsequent bone regeneration. Cytokines such as IL-17 and TNF-α synthesized by neutrophils activate osteoclast through the RANKL-RANK signaling pathway (Chang et al., 2013). On the other hand, neutrophils stimulate angiogenesis (Christoffersson et al., 2012) and bone regeneration through direct or indirect action. Neutrophils in liver injury achieve angiogenesis and maturation by secreting MMP-9 to promote the expression of VEGF (Christoffersson et al., 2012). In the early stage of inflammation, neutrophils alleviate acute inflammation and reduce the secretion of inflammatory cytokines, thereby reducing the activation of osteoclast precursor cells and promoting bone repair (Bastian et al., 2018). The application of biologically active neutrophil membrane materials is enthusiastically sought after by scientific researchers. After implantation, the membrane material simulates the post-inflammatory process, and the apoptosis of neutrophils induces the entrapment of macrophages, which further promotes the polarization of macrophages to M2 to play an anti-inflammatory role and promotes bone regeneration. During this process, TGF-β and IL-10 secreted by macrophages promote anti-inflammatory and bone regeneration repair. IL-8-induced polarization of neutrophils contributes to endochondral ossification (Cai et al., 2021). These will provide strong support for designing materials to regulate neutrophil behavior.

## 4.2 The application of macrophages in the design of biomaterials

The application of macrophage polarization in the regeneration of bone defect sites is favored. The changes in

macrophage behavior promote vascular and bone regeneration (Loi et al., 2016; ElHawary et al., 2021). Angiogenesis provides various nutrients and cells for bone repair. Therefore, coupling blood vessels and bone regeneration is another entry point for material design in fracture repair. The modified application of various bioactive molecules combined with biomaterials promotes vascular or bone regeneration by regulating macrophage polarization (Loi et al., 2016; Wang et al., 2022). Calcium silicate combined with β-tricalcium silicate stimulates the polarization of macrophages to M1/M2 by releasing IFN-γ and Si to secrete further vascular growth factors (VEGF, CXCL-12, PDGF-BB) to promote angiogenesis (Loi et al., 2016). Mesoporous silica (MNS)-loaded BMP bioactive molecules were incorporated into 3D printed scaffolds made of modified GelMA to release bioactive molecules and promote macrophage M2 polarization, BMP active molecules, and M2-secreted anti-inflammatory cells factors to promote the repair of diabetic bone defects. The release of BMP-2 and zinc particles regulates the increased proportion of M2 macrophages on the screw-coated surface to promote bone regeneration (Wang et al., 2022) (Figures 5B–D). Although macrophage polarization occurs in both angiogenesis and bone regeneration, there is a particular deviation in the macrophage's temporal and spatial control time. It is urgent to develop new biomaterials to simultaneously realize the classification of the fate of macrophages in angiogenesis and bone regeneration.

## 4.3 The application of T cells in the design of biomaterials

T cells play an indirect role in bone regeneration and repair, and factors secreted by various T cell subtypes indirectly regulate the process of bone regeneration. Like macrophages, T cells can be roughly divided into pro-inflammatory cell subtypes (Th1, Th17) and anti-inflammatory cell subtypes (Th2, Treg). Th1 cells can inhibit the expression of RUNX2 in BMSCs by secreting the IFN-γ
<figure><img src="image_72.png" /><figcaption>**FIGURE 6**
Immunomodulatory material for bone regeneration. (A,B) DOPA4@BMP2pPEEK material regulates the differentiation of CD4 T cells and synergizes with BMP-2 to promote bone regeneration. The graph was reprinted with permission from (Zhao et al., 2021c).</figcaption></figure>

factor to reduce bone repair ability (Liu et al., 2011). In addition, IFN-γ secreted by Th1 cells is associated with the polarization of M1 macrophages, and sustained M1 polarization leads to sustained inflammatory activation that prevents bone repair from occurring (Spiller et al., 2015). Th17 acts as the ability to secrete pro-inflammatory cytokine IL-17, but its role can promote bone regeneration and repair. IL-17A combined with BMP-2 can significantly enhance the osteogenic ability of BMSCs (Moon et al., 2016). Th2 and Treg cells promote bone homeostasis towards osteogenesis and bone regeneration by inducing the formation of M2 macrophages (Chen et al., 2012) and inhibiting osteoclast differentiation (Lu et al., 2017). And the surface of PEEK implants promotes bone regeneration by regulating T cell differentiation via likable mussel-inspired azide-DOPA4 and BMP2p coupling (Zhao et al., 2021c) (Figure 6). Although there are few studies on T cells in bone regeneration biomaterials, the regulation of macrophages and osteoclasts by T cells will be a practical starting point for biomaterials to regulate the immune microenvironment to achieve bone regeneration and bone disease treatment. This will provide new therapeutic strategies for clinical bone defect regeneration and disease treatment.

# 5 Conclusion and future perspectives

The research of biomaterials is reviewed in bone regeneration and repair. We summarize some defects and improvement schemes of biomaterials and analyze the promising prospect of bone immune materials. It can be used for reference in the design of immune materials for bone regeneration.

1. Tricalcium phosphate ceramics (β-TCP) have been used as bone-filling materials in clinical practice due to their high biocompatibility and bone conductivity, but their non-degradability limits their application in bone regeneration. Then, biodegradable biomaterials with the same mechanical strength as β-TCP will be gradually explored. And the high biocompatibility and plasticity of the extracellular matrix (ECM) components will provide a broad application prospect for the research of bone regeneration materials.

2. Electrospun fibers' tenacity, plasticity, and core-shell structure can carry hydrophilic and non-hydrophilic drugs, proteins, peptides, etc. They have been used as periosteal-like and filling delivery materials in bone defect models to promote
bone regeneration. Still, the application of cytotoxic solvents in their synthesis process has not been able to promote their clinical application. The development of biological fibers will provide a new direction for the wide application of nanofibers.

3) 3D bio-printing materials have been widely used in the study of bone regeneration because of their ability to simulate organisms' tissue structure accurately. Although it can print a material resembling bone, it can not respond to internal activity when implanted in organisms. The intelligent programming of 4D bio-printing materials and the function of responding to the activity of implants will provide broad application space for the research of bone regeneration.

4) M1 macrophages are important in clearing foreign bodies and dead cells at defect sites. They are indispensable in bone defect repair. Therefore, controlling the ratio of M1/M2 is the key to regeneration repair. In addition, how to transform macrophages from type M1 to type M2 after implantation is also the key to initiating the regenerative repair.

5) Numerous articles have focused on the effects of materials on macrophage behavior, with little focus on studies of other immune cells such as T cells, neutrophils, and DC cells. Different subtypes of T cells play multiple roles in the regeneration process, and neutrophils are also key cell populations that initiate the regenerative function of macrophages. The design of immunomodulatory materials targeting T cell subtypes and other immune cell behaviors may provide a new route for regenerative applications.

The success or failure of biomaterials in bone injury repair is determined by the interaction of the host immune system with biomaterials. Continued activation of the inflammatory response will lead to a delay in the repair process or tissue necrosis. Therefore, the rational regulation of the foreign body reaction effect is the key to bone injury regeneration and repair. From the selection of raw materials and the synthesis process, Biomaterials endowed with bioactive factors or their characteristics regulate immune cells, thus realizing the regulation of tissue regeneration. Competent biomaterials regulate the biological

behavior of immune cells such as neutrophils, macrophages, or T cells. It promotes the differentiation of these immune cells to an anti-inflammatory phenotype, further promoting bone tissue regeneration. The research of new biomaterials to promote the real-time regulation of immune cell behavior to adapt to the high degree of compatibility between the immune system and bone regeneration, the tacit cooperation of these new biomaterials in innate and adaptive immunity needs to be further explored. Therefore, the design of biomaterials must consider the activation of immune cells and the mutual interference between different innate and adaptive cellular components to meet the research needs of clinical medicine.

## Author contributions

Conceptualization, GW; investigation, CD and GT; resources, CD and G-YZ; writing original draft preparation, CD; writing-review and editing, WL; supervision, GW; project administration, GW. All authors have read and agreed to the published version of the manuscript.

## Conflict of interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

## Publisher's note

All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher.

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<figure><img src="image_73.png" /></figure>

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OPEN ACCESS

EDITED BY
Yawei Du,
Shanghai Jiao Tong University, China

REVIEWED BY
Gang Li,
Soochow University, China
Tuo Yang,
University of California, San Diego,
United States

*CORRESPONDENCE
Shanliang Han,
✉ hanshanliang523@163.com
Chongwei Ke,
✉ shchweiz@126.com

†These authors have contributed equally to this work and share first authorship

RECEIVED 07 January 2023

ACCEPTED 17 April 2023

PUBLISHED 03 May 2023

CITATION
Cao Z, Chen L, Niu G, Li Y, Hu Z, Hong R, Zhang X, Hong L, Han S and Ke C (2023), Preparation and characterization of a novel triple composite scaffold containing silk fibroin, chitosan, extracellular matrix and the mechanism of Akt/FoxO signaling pathway in colonic cancer cells cultured in 3D.

Front. Bioeng. Biotechnol. 11:1139649.

doi: 10.3389/fbioe.2023.1139649

COPYRIGHT
© 2023 Cao, Chen, Niu, Li, Hu, Hong, Zhang, Hong, Han and Ke. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Preparation and characterization of a novel triple composite scaffold containing silk fibroin, chitosan, extracellular matrix and the mechanism of Akt/FoxO signaling pathway in colonic cancer cells cultured in 3D

Zhipeng Cao†, Liang Chen†, Gengming Niu, Yan Li, Zhiqing Hu, Runqi Hong, Xiaotian Zhang, Liang Hong, Shanliang Han* and Chongwei Ke*

Department of General Surgery, Fifth People's Hospital of Shanghai, Fudan University, Shanghai, China

This work examined the physical and chemical properties and biocompatibility in vivo and in vitro of a unique triple composite scaffold incorporating silk fibroin, chitosan, and extracellular matrix. The materials were blended, cross-linked, and freeze-dried to create a composite scaffold of silk fibroin/chitosan/colon extracellular matrix (SF/CTS/CEM) with varying CEM contents. The SF/CTS/CEM (1:1:1) scaffold demonstrated the preferable shape, outstanding porosity, favorable connectivity, good moisture absorption, and acceptable and controlled swelling and degradation properties. Additionally, HCT-116 cells cultivated with SF/CTS/CEM (1:1:1) showed excellent proliferation capacity, cell malignancy, and delayed apoptosis, according to the in vitro cytocompatibility examination. We also examined the PI3K/PDK1/Akt/FoxO signaling pathway and discovered that cell culture using a SF/CTS/CEM (1:1:1) scaffold may prevent cell death by phosphorylating Akt and suppressing FoxO expression. Our findings demonstrate the potential of the SF/CTS/CEM (1:1:1) scaffold as an experimental model for colonic cancer cell culture and for replicating the three-dimensional in vivo cell growth environment.

KEYWORDS

extracellular matrix, silk fibroin, chitosan, scaffolds, Akt/FoxO signaling pathway

## 1 Introduction

With over 940,000 annual deaths, colonic carcinoma (CC) is the second most lethal malignant tumor around the world, followed by lung cancer (Sung et al., 2021). Due to their socioeconomic development, developed countries have the highest incidence of this disease (Dekker et al., 2019). The occurrence and development of tumors are multi-stage processes involving multiple biological pathways, and the mechanisms involved are relatively complex (Zhang et al., 2018). By combining the construction of the tumor environment with the study of the biological behavior of tumors and utilizing new research technologies and methods,
greater understanding and progress have been made in tumor research in recent years. Therefore, developing novel tumor models and advancing tumor research are quite important.

These studies suggest that tumor growth has its own internal microenvironment. Cells are encapsulated by the outer matrix, and there are dynamic interactions between cells and the matrix and between cells and signal molecules. With different methods of building a tumor microenvironment, the goal is to simulate a microenvironment closer to the growth of tumors *in vivo* and facilitate biological behavior that is closer to real tumors *in vivo*. At this stage, biological research on tumors is mainly carried out at the level of two-dimensional (2D) single-cell culture. However, 2D culture has some limitations in the study of biological behavior and drug sensitivity of colorectal cancer, such as the reduction of tumor malignancy and changes in cell-related properties, such as differentiation and interaction between polarity and extracellular matrix (ECM), which cannot represent the real tumor focus. It has been reported that monolayer cell culture differs from the *in vivo* cells in terms of tumor drug mechanism and resistance, whereas various physiological activities of colon cancer cells in a three-dimensional culture environment are closer to the real environment *in vivo* (Shin et al., 2018). Cells in a three-dimensional (3D) culture system may retain a healthy proliferative state for a longer amount of time compared to 2D culture conditions, and cell activity is greatly increased. Therefore, we established previously a 3D tumor model for tumor drug screening (Lovitt et al., 2018). Although 3D scaffold materials are widely used in tissue engineering, there are few reports in the field of tumors, especially in colon cancer (Liang et al., 2020). Tissue engineering materials are creatively used to seed specific tumor cells into certain biomaterials, form cell biomaterial composites after *in vitro* culture, and implant them into mice to investigate the similarities and differences between tumor characteristics and traditional culture methods (Xie et al., 2016). In a 3D model, cells continue to proliferate and secrete matrix. At the same time, 3D materials are gradually absorbed. The final tumor tissue demonstrates significant differences in morphology, function, and other aspects of the tumor tissue with simple subcutaneous tumorigenesis. Therefore, some scholars have proposed that a 3D structure constructed *in vitro* should be fully used to simulate the tumor microenvironment (Landberg et al., 2020).

In three-position scaffold materials, the use of high-molecular-weight substances such as silk fibroin (SF) and chitosan (CTS) has been widespread. SF has a wide range of sources and good biological properties (Algarrahi et al., 2018; Galvez Alegria et al., 2019); thus, its increasing use in tissue engineering and has good prospects. The fast rate of breakdown of SF, however, limits its use (Chomchalao et al., 2013). As a linear polysaccharide with naturally occurring positively charged bases, CTS has excellent biofunctionality, plasticity, biodegradability, and safety (Zhao et al., 2020). A CTS 3D scaffold can provide space for cells to grow, multiply, and finally approximate the characteristics of organs with certain functions, which can then be used to study tumor tissues (Patrulea et al., 2015). Other components, including elastin, collagen, glycoproteins, and proteoglycans, can be found in the ECM. According to previous studies, the ECM can have an impact on a cell's basic functions, including cell division, proliferation, adhesion, and phenotypic expression. In addition to help organizing tissues, it also provides

crucial biochemical and biomechanical cues for regulating vascular and immune development, cell proliferation, migration, and differentiation. Furthermore, the ECM can control how dynamically cancers develop.

Researchers have been exploring composite systems with diverse polymers to overcome the drawbacks of scaffolds made of a single material. Combinations of various polymers are expected to confer their individual properties and form scaffolds that may promote cell adhesion, proliferation, and differentiation (Ghosh et al., 2019). Moreover, composite material scaffolds have been used and studied for cartilage repair (Bhardwaj et al., 2011), sciatic space repair (Gu et al., 2014), and bone defect reconstruction (Rios et al., 2009).

This paper presents a new study on CC 3D culture using SF/CTS/colon extracellular matrix (CEM) composite scaffolds *in vitro*. For the first time, we investigated the creation of composite scaffolds based on the SF/CTS/CEM polymer system. We established a new 3D tumor cell culture system by seeding HCT-116 cells on SF/CTS/CEM scaffolds.

# 2 Materials and methods

## 2.1 Materials and animals

Genuine silkworm cocoons were obtained from farmers in Shiquan, Shanxi province. CTS powder (900,000 Da, 95% deacetylated), acetic acid, lithium bromide, dimethyl sulfoxide, absolute ethanol, cell counting kit (CCK)-8, dialysis bags, 4% paraformaldehyde, 1-Ethyl-3-(3-Dimethylaminopropyl) carbodiimide hydrochloride (EDC), and N-Hydroxysuccinimide (NHS) were purchased from Sangon Biotech (Shanghai) Co., Ltd., and sodium carbonate was acquired from the Sinopharm Chemical Reagent Co., Ltd. DMEM medium, 4,6-Diamidino-2-phenylindole dihydrochloride (DAPI), DY-554-Phalloidin staining, YF-488-Annexin V, and PI Apoptosis Kit were purchased from Share-bio Co., Ltd. Fetal bovine serum was purchased from Shrabio (Shanghai) Co., Ltd. Female naked mice aged 6 weeks were purchased from JSJ-lab (Shanghai) Co., Ltd. The mice were housed in a germ-free environment. The human CRC cell line HCT-116 was obtained from the Cell Bank of Shanghai Fifth People's Hospital and cultured in DMEM medium.

### 2.1.1 Extraction of silk fibroin and preparation of chitosan solution

The silkworm cocoon shell was cut into 1 cm² pieces, immersed in 0.5% sodium carbonate solution, and boiled three times for 1 h each. It was then fully dried in an oven at 65 C after being rinsed three times in distilled water. The processed silk fibers were placed in a 9 M lithium bromide solution to form another solution, which was then dialyzed against distilled water (renewed every 12 h) in a dialysis bag (3.5 KD) for 72 h to obtain a 3% SF solution. To create a 3% CTS solution, CTS was dissolved in a solution of 3% glacial acetic acid.

### 2.1.2 Preparation of colonic extracellular matrix

The colon tissue was soaked in 2% SDS+0.5% EDTA solution, which was changed every 3 h, shaken at room temperature for 12 h, and rinsed up with PBS buffer. The tissues were then immersed in
1% Triton X100 + 0.5% EDTA, which was changed every 3 h, shaken at room temperature for 12 h, and then rinsed up with PBS buffer to obtain the CEM. The CEM was ground into a powder by cold extraction with liquid nitrogen.

## 2.2 Scaffold synthesis and block design

As previously reported, chemical cross-linking techniques and freeze-drying technology were used to create the scaffolds used in the study (Li et al., 2017). SF scaffolds were made using only the 3% SF solution. SF and CTS were combined in a 1:1 (w/w) ratio to create SF/CTS (1:1) scaffolds. Three distinct mass ratios of the SF solution, CTS solution, and CEM powder were used to make the SF/CTS/CEM scaffolds with different ratios: 1:1:0.5, 1:1:1, and 1:1:2 (w/w). The mixture was then added to a solution containing 95% aqueous ethanol, 50 mmol/L EDC, and 18 mmol/L NHS, and the combination was agitated magnetically for 30 min to produce a homogenous solution. The solutions were then cast into well-sized 24-well and 96-well plates and crosslinked for 12 h at 4 C. The samples were then frozen at -20 C for 12 h and stored at -80 C for a further 12 h. To create the scaffolds, the samples were put into a freeze-dryer for 48 h. Before beginning cell culture, we used a low-temperature plasma sterilizer to decontaminate the scaffolds.

## 2.3 Characterization of the scaffolds

### 2.3.1 Macroscopic appearance

Front and lateral views photographs were used to compare the various scaffold groups after they had been removed from the 24-well plate.

### 2.3.2 Internal morphology

After removing the scaffolds from the 24-well plate, a layer of scaffold film with the scaffold's cross-sectional structure was left at the bottom of the well, and the structure of the scaffold film was examined under an optical microscope and captured on camera. The microstructures of the scaffolds were observed using a scanning electron microscope (SEM). The scaffolds were cut into small fragments with a surgical blade. We placed the prepared samples on the sample table and plated them with platinum. Then, the samples were put into the SEM and observed after vacuuming. We observed the microstructure and the microchannel of the scaffolds under the SEM and took photographs.

### 2.3.3 Porosity evaluation

The liquid replacement method is used to assess scaffold's porosity. Briefly, the scaffold is placed in anhydrous ethanol with a volume of $V_0$. The volume of pure ethanol and the submerged scaffold after full immersion was $V_1$. $V_2$ was the volume remaining after the impregnated scaffold was removed. The following formula was used to determine porosity: $(V_0-V_2)/(V_1-V_2) \times 100\%$.

### 2.3.4 Water uptake ratio

The mass of the scaffold in the dry state was recorded as $W_1$. PBS buffer was used to rehydrate the dried scaffolds for 24 h at 37°C. With the aid of filter paper, any remaining liquid on the surface was removed,

and the wet sample's weight was recorded as $W_2$. The following formula was used to compute water uptake ratio: $(W_2-W_1)/W_1 \times 100\%$.

### 2.3.5 Degradation property

The mass of the scaffold in a dry environment was $W_0$. The scaffolds were placed in 6-well cell plate medium in PBS solution at 37 C, and then dried for 12 h at 65 C before being weighed at 1 d (day), 3 d, 7 d and 14 d ($W_n$). The following formula was used to compute the degradation ratio: $(W_0-W_n)/W_0 \times 100\%$.

## 2.4 Cell incorporation into scaffolds

Scaffolds were sterilized at low temperature in advance. We seeded 50,000 cells into each scaffold in 24-well plates, then slowly shook these plates, and finally added 1 mL of complete medium to each well. Every other day, the culture medium was changed.

### 2.4.1 Cell adhesion in scaffolds

In a nutshell, HCT-116 105 cell ($A_0$) cell suspensions were seeded on pre-wetted scaffolds, and adhesion rates were assessed 1, 3, and 6 h later. The scaffolds were taken out of the wells, and cells were counted ($A_1$). Cells clinging to the well walls were digested and counted when the media was removed ($A_2$). The following formula was used to calculate the cell adhesion rate: $(A_0-A_1-A_2)/A_0 \times 100\%$. Three experiments were performed for each condition, and the average adhesion rate was calculated.

### 2.4.2 Cell proliferation in scaffold

CCK-8 was used to track cell proliferation in 2D plates, SF/CTS (1:1), and SF/CTS/CEM (1:1:1) scaffolds. Then, 100 µL of a cell suspension containing $10^3$ HCT-116 cells was seeded onto scaffolds in 96-well plates. Following the manufacturer's instructions, proliferation of cell was assessed on 1 d, 3 d, and 5 d. Briefly, 10 µL CCK-8 reagent and 90 µL DMEM mixture was added to each well and the plate was placed for 90 min in the dark. Subsequently, we shocked the plates for 15 min and then the scaffolds were removed. To test absorbance at 450 nm, the remaining liquid was transferred to a fresh 96-well plate.

### 2.4.3 Cell growth, micro-structure, and ultra-structure in scaffolds

#### 2.4.3.1 DAPI and DY-455-Phalloidin staining

HCT-116 cells ($5 \times 10^5$) were cultured in a pre-prepared scaffold. For cytoskeletal staining, 0.5% Triton X-100 in PBS was used to permeabilize the cells for 10 min at room temperature after cells being fixed with 4% paraformaldehyde for 15 min on ice. Using 10 g/mL, cytoskeletons were stained after incubation for 20 min at room temperature with a solution of DY-455-Phalloidin conjugates, then rinsed with PBS to get rid of any unbound DY-455-Phalloidin conjugates. Nuclear staining was performed with DAPI. A fluorescent microscope was used to record the images (Leica DM2500).

#### 2.4.3.2 Scanning electron microscope

HCT-116 cells were cultured on different groups of scaffolds. On days 1 and 3, the scaffold was washed with PBS for 3 times after the culture medium was removed. Then we used 4% paraformaldehyde to fix the cells. The plates were then placed in a refrigerator set at
<figure><img src="image_74.png" /><figcaption>FIGURE 1</figcaption></figure>

Colon of fresh nude mice and its extracellular matrix after acellular treatment (A, B); Flowchart of the preparation of composite scaffolds by freeze-drying technique (C); Front and side views of SF, SF/Cs (1: 1), SF/Cs/CEM (1: 1: 0.5), SF/Cs/CEM (1: 1: 1), and SF/Cs/CEM (1: 1: 2) scaffolds (D, E). SF, silk fibroin; CTS, chitosan; CEM, colon extracellular matrix.

80 °C for 12 h before being frozen at 20 C for another 12 h. The culture plates were then freeze-dried in a freeze-dryer for 48 h, and a SEM was used to examine the cell morphology on the scaffold.

### 2.4.3.3 Hematoxylin-eosin staining

Hematoxylin and eosin (HE) staining was performed to observe biocompatibility of scaffolds *in vivo* and *in vitro*. On days 1 and 3 of scaffold culture, the composites were fixed with 4% paraformaldehyde. Meanwhile, 1 month after subcutaneous implantation of the scaffold containing cells, the nude mice were treated and the tumor was removed. HCT-116 cells and scaffolds from the removed tumors were fixed with 4% paraformaldehyde. After embedding in paraffin, the tissues were sectioned into slices of 3 µm thickness. Slices where then stained for HE and photographed with a Leica DM2500 microscope and an oil-immersion lens.

## 2.5 DY-488-Annexin V staining and flow cytometry assay

HCT-116 cells were seeded into different scaffolds and cultured in a 1 g/L low-glucose medium for 24 h to induce

apoptosis. Cells were digested with trypsin without EDTA (ShareBio, Shanghai, China) and collected in Eppendorf tubes. Cells were stained using the Annexin V FITC Apoptosis Kit (ShareBio, Shanghai). A flow observation analyzer was used to assess the staining.

## 2.6 Protein extraction and Western blotting

A RIPA solution containing 1% PMSF was used to lyse the cells, and the entire protein extraction process was carried out on ice. Following that, protein samples were centrifuged for 15 min at 10,000 rpm at 4°C. Using the BCA reagent, protein concentrations were measured. Proteins were separated by electrophoresis on SDS gels and then transferred to PVDF membranes. Prior to being incubated with primary antibodies for 14 h at 4 C, the PDVF membranes were blocked with 5% nonfat milk for 1-2 h at room temperature. The secondary antibody was incubated for 1-2 h at room temperature after three membrane washes. After washing the secondary antibody solution from the membrane three times, protein expression was detected using an enhanced chemiluminescence reagent.
<figure><img src="image_75.png" /><figcaption>**FIGURE 2**
Images of optical microscope (A1-E1, a1-e1) and scanning electron microscope (A2-E2, a2-e2) of SF, SF/CTS (1: 1), SF/CTS/CEM (1: 1: 0.5), SF/CTS/CEM (1: 1: 1), and SF/CTS/CEM (1: 1: 2) scaffolds. SF, silk fibroin; CTS, chitosan; CEM, colon extracellular matrix.</figcaption></figure>

<figure><img src="image_76.png" /><figcaption>**FIGURE 3**
Results of porosity (A), water uptake rate (B) and degradation rate (C) of different groups of scaffolds. \*p < 0.05, \*\*p < 0.01, \*\*\*p < 0.001.</figcaption></figure>
<figure><img src="image_77.png" /></figure>

<figure><img src="image_78.png" /><figcaption>FIGURE 4</figcaption></figure>

DAPI staining, Phalloidin staining and merging diagram of cultured HCT-116 cells in 2D, SF/CS (1:1) and SF/CS/CEM (1:1:1) (A). Results of adhesion rate (B) and cell proliferation (C) of the different groups of scaffolds. *p* < 0.05, **p** < 0.01, ***p** < 0.001. SF, silk fibroin; CTS, chitosan; CEM, colon extracellular matrix.

## 2.7 Immunohistochemical

Six-week-old mice were brought up in a pathogen-free environment. HCT-116 cells that had been conditioned in 2D, SF/CTS (1:1), and SF/CTS/CEM (1:1:1) were implanted subcutaneously into mice flanks. Using antibodies for Ki-67, PCNA, TUNEL, and Bcl-2 staining, tumor samples from mice were fixed in formalin and embedded in paraffin, sectioned into 5 µm slices, and subjected to immunohistochemical (IHC) staining. This was done 2 weeks and 1 month following cell implantation.

## 2.8 Statistical analysis

For all statistical calculations, the Graph Pad Prism 8 software was utilized. Each experiment was performed in triplicate. The mean and standard deviation (SD) were used to present quantitative data. The statistical significance was established using the Student's t-test and analysis of variance (ANOVA). When differences were *p* < 0.05, **p** < 0.01, or ***p** < 0.001, they were deemed significant.

# 3 Results

## 3.1 Characterization of the composite scaffolds in 3D

### 3.1.1 Macroscopic appearance

Figure 1 depicts the macroscopic characteristics of the SF, SF/CTS (1:1), SF/CTS/CEM (1:1:0.5), SF/CTS/CEM (1:1:1), and SF/CTS/CEM (1:1:2) scaffolds. We noticed that the shapes of all the scaffolds were comparable. Most of the scaffolds were yellowish-white in color, whereas the SF scaffold was completely white.

### 3.1.2 Internal structure

To understand the internal structure of the scaffolds, we employed an optical microscope and SEM. As seen in Figure 2, every scaffold contained porous networks with architectures featuring various pore sizes, homogeneity, and strong pore connection. The addition of CTS and CEM changed the structure of pure SF scaffolds. The pore diameters and homogeneity of the SF/CTS/CEM (1:1:0.5), SF/CTS/CEM (1:1:1), and SF/CTS/CEM (1:1:2) scaffolds varied. As the proportion of CEM increased, the scaffold's pore size steadily grew.
<figure><img src="image_79.png" /><figcaption>FIGURE 5</figcaption></figure>

Scanning electron microscope images of HCT-116 cells on SF/CTS (1: 1), SF/CTS/CEM (1: 1: 0.5), SF/CTS/CEM (1: 1: 1), and SF/CTS/CEM (1: 1: 2) scaffolds. HCT-116 cells were cultured on different groups of scaffolds on day 1 (A1-D1, a1-d1) and day 3 (A2-D2, a2-d2). SF, silk fibroin; CTS, chitosan; CEM, colon extracellular matrix.

### 3.1.3 Physical and chemical properties

#### 3.1.3.1 Porosity

All scaffolds had porosities greater than 68.69% (Figure 3A). Of all the scaffolds, the SF/CTS/CEM (1:1:1) scaffold had the highest porosity (88.33% ± 3.58%), followed by the SF/CTS/CEM (1:1:2) scaffold (81.11% ± 4.78%). The porosities of the SF/CTS and SF/CTS/CEM (1:1:0.5) scaffold were 81.11% ± 4.78% and 73.81% ± 5.12%, respectively. Our findings demonstrated that there was a statistically significant difference in the porosity of the SF/CTS (1:1) and SF/CTS/CEM (1:1:1) scaffolds ($p < 0.05$).

#### 3.1.3.2 Water uptake ratio

The water uptake ratios of the scaffolds are shown in Figure 3B. All of the scaffolds had water absorption rates in deionized water that were more than 501.9%. The maximum water uptake ratio was observed in the SF/CTS/CEM (1:1:1) scaffold (1,072% ± 48%), followed by the SF/CTS/CEM (1:1:2) scaffold (933.5% ± 36%). In contrast, the scaffold with the lowest water uptake ratio was the SF/CTS (1:1) scaffold (745.92% ± 44%). The results showed that there was significant difference in water absorption between SF/CTS (1:1) and SF/CTS/CEM (1:1:1) scaffolds.

#### 3.1.3.3 Degradation rates

Figure 3C displays the scaffolds' degradation rates submerged in the PBS solution. The addition of CEM slowed the deterioration of the scaffolds. The SF/CTS/CEM (1:1:1) scaffold exhibited the lowest degradation rate, followed by the SF/CTS/CEM (1:1:2) scaffold. A

significant difference observed between the two groups ($p < 0.05$). All five groups of scaffolds degraded to varying degrees within 14 days; however, the overall trends were similar. Day 3 was the turning point, as in the first 3 days degradation occurs rapidly, slowing down during the subsequent 4 days. From the 7th to 14th days, degradation stabilizes.

## 3.2 Cell incorporation into scaffolds

### 3.2.1 Cell adhesion in scaffolds

Counting cells on a cytometry plate at 1, 3, and 6 h after seeding the cells on the pre-wetted scaffolds allowed assessing cell adherence to the scaffolds. Figure 4B depicts the cell adhesion rate, showing that the SF/CTS/CEM (1:1:1) and SF/CTS/CEM (1:1:2) scaffolds had the highest rates. The SF/CTS/CEM (1:1:1) scaffold showed higher cell adhesion.

### 3.2.2 Cell proliferation in scaffold

To gauge cell proliferation on the various scaffold groups, the CCK-8 kit was used (Figure 4C). According to our findings, the scaffold with the highest proportion of proliferating cells at each time point was the SF/CTS/CEM (1:1:1) scaffold. Based on the statistical findings, the cell proliferation of the two scaffolds was not significantly different on day 1, whereas the cell proliferation of the triple scaffold was significantly higher on days 3 and 5.
<figure><img src="image_80.png" /><figcaption>FIGURE 6</figcaption></figure>

(A) Images of hematoxylin and eosin staining of scaffolds. Control groups were untreated SF/CTS (1:1) and SF/CTS/CEM (1:1:1) scaffolds, HCT-116 cells were cultured on SF/CTS (1:1) and SF/CTS/CEM (1:1:1) scaffolds on day 1 and day 3. (B) Images of hematoxylin and eosin staining of tumors formed after subcutaneous implantation of SF/CTS (1:1) and SF/CTS/CEM (1:1:1) scaffolds for 2 weeks and 1 month in nude mice. SF, silk fibroin; CTS, chitosan; CEM, colon extracellular matrix.

### 3.2.3 Cell growth, micro-structure and ultra-structure in scaffolds

#### 3.2.3.1 DAPI and DY-488-Phalloidin staining

To visualize the cytoskeleton organization, we stained actin filaments with DY-488-Phalloidin-FITC and imaged the cells using a light microscope Leica DM2500 (Figure 4A). In 2D culture, the majority of the cells were long and fusiform, while the majority of the cells seeded in SF/CTS (1:1) and SF/CTS/CEM (1:1:1) scaffolds were spherical, which was more representative of the *in vivo* cell morphology. However, cell-cell attachments tended to be more pronounced in the SF/CTS/ECM (1:1:1) scaffold than in the SF/CTS (1:1) scaffold, with a spherical morphology. Additionally, we used DAPI to stain cell nuclei to confirm cell division, as shown in Figure 4A, which shows a statistically significant difference in the number of cells across the three scaffolds (2D, SF/CTS (1:1), and SF/CTS/CEM (1:1:1)) at all time points (p < 0.001). The proliferation ability of the SF/CTS/CEM (1:1:1) group was the highest.

#### 3.2.3.2 SEM

Figure 5 depicts the SEM images of the cells on the scaffolds, displaying their morphology and health state. There was more room for cell division and proliferation in the SF/CTS/CEM (1:1:1) scaffold because the channel diameter was larger than that of the SF/CTS group (1:1). More cells proliferated on CEM-containing scaffolds. SF/CTS/

CEM (1:1:1) scaffolds supported higher cell proliferation than SF/CTS (1:1) scaffolds under the same conditions. When compared to typical culture dishes, the cell morphologies in 3D scaffolds (SF/CTS/CEM (1:1:1) and SF/CTS (1:1)) were very different. In 3D scaffolds, the cells do not fully adhere to the scaffold and retain their body shape. The cells cultured in these scaffolds also had integrated spheres and, even though the number of cells grew over time, they did not spread across the inner surface of the scaffold. We also observed that the SF/CTS/CEM (1:1:1) scaffold supported a higher cell proliferation under the same conditions. The center portions of the scaffolds and the image are obtained (Figures 10 and 11). Figure 10 shows that internal situation of SF/CTS/CEM (1:1:1) scaffolds. Figure 11 shows the cell growth in the inner area of the scaffold with different material components. HCT-116 cells grow in the inner area of the different scaffolds. It was found that compared with the control group, the cells in SF/CTS/CEM (1:1:1) scaffolds group grew in lumps and had stronger proliferation ability.

#### 3.2.3.3 HE staining

Images of HE-stained scaffolds are shown in Figure 6. Under the same conditions, the SF/CTS/ECM (1:1:1) scaffolds supported higher cell proliferation than the SF/CTS (1:1) counterparts. Compared with cells cultured in traditional 2D culture, cells on the scaffold tend to aggregate and grow. Notably, scaffolds implanted subcutaneously
<figure><img src="image_81.png" /><figcaption>FIGURE 7</figcaption></figure>

(A) Images of flow cytometry apoptosis of HCT-116 cells cultured in 2D, SF/CTS (1:1), and SF/CTS/CEM (1:1:1) scaffolds after apoptosis induction. (B)

Analysis of protein expression of HCT-116 cells cultured on 2D, SF/CTS (1:1), and SF/CTS/CEM (1:1:1) scaffolds for 14 days. SF, silk fibroin; CTS, chitosan; CEM, colon extracellular matrix.

were gradually absorbed over time, and the tumor cells grew
gradually. We also observed angiogenesis in the tumor.

3.3 Flow cytometry assay

Flow cytometry analysis of apoptosis is shown in Figure 7.
Under the same induction conditions, the apoptosis of HCT-116
cells cultured with the SF/CTS/CEM scaffold was the lowest,
followed by the SF/CTS scaffold, with the highest apoptosis being
observed in the 2D group.

3.4 Protein extraction and Western blotting

To confirm the influence of SF/CTS/CEM scaffolds on
apoptosis, the PI3K/PDK1/Akt/FoxO signaling pathway was
investigated by gel electrophoresis using protein samples of cells
cultured for 14 days in the different conditions. SF/CTS/CEM (1:1:1)
scaffolds activated Akt phosphorylation and inhibited the
expression of pro-apoptotic FoxO compared with SF/CTS (1:1)
and 2D cultures, as shown by protein expression (Figure 7).

3.5 IHC staining

We subcutaneously injected HCT-116 cells grown on SF/CTS/
CEM scaffolds or controls into the flanks of naked mice to determine
whether tumor malignancy was reduced *in vivo* (n = 5 per scaffold).
Figure 8 and Figure 9 illustrate how tumor cells cultured on SF/CTS/
CEM scaffolds expressed Ki67 and PCNA at significantly higher
levels than cells cultured in 2D. Bcl-2 and TUNEL expression were
also downregulated.

4 Discussion

Conventional and classic 2D cell culture methods offer a
practical platform for *in vitro* cancer. Cells cultivated on the
surface of 2D flat Petri dishes, however, exhibit much less
malignant phenotypes and do not accurately represent the same
cell-ECM and cell-cell interactions as tumor formations *in vivo*
(Smalley et al., 2006). To create scaffolds with the ideal culture
properties, SF, CTS, and CEM have stimulated the development of
3D cell culture systems. Compared to SF scaffolds of pure
silkworm, the blended scaffolds have different morphologies,
porosities, elasticities, swelling behaviors, and biochemical
compositions. Previously, the SF/CTS (1:1) mixture has been
shown to be a potential biomaterial for the generation of
scaffolds for cancer treatment (Gupta et al., 2009). In contrast
to substrates made of pure biomaterials or synthetic polymers,
according to a recent study 3D cell culture utilizing SF/CTS (1:1)
scaffolds can promote cell proliferation in prostate cancer (Bäcker
et al., 2017). Our preliminary experimental findings, however,
indicated that SF/CTS (1:1) scaffolds had poor water
absorption, a slow rate of breakdown, and a slow rate of cell
adhesion, which restricts their use in 3D cell culture. Solid tumors
are made up of genetically-mutated cancer cells surrounded by
ECM and an additional group of genetically normal cells. The latter
two elements are part of the tumor microenvironment and are
important regulators of tumor biology, which has an impact on
patients' prognoses. The tumor ECM has been the focus of research
for the past 20 years, exposing the fundamental biochemical and
biological concepts and mechanisms underlying its function in
tumor cell survival and proliferation. The ECM, however, also has a
significant impact on immune cells in the microenvironment,
controlling their differentiation and infiltration into tumor cells,
as well as their proliferation and survival (Kolesnikoff et al., 2022).
<figure><img src="image_82.png" /><figcaption>**FIGURE 8**
Immunohistochemical staining images of tumors formed by subcutaneous implantation of scaffolds. Two weeks and 1 month after implantation, tumor samples were subjected to immunohistochemical staining against TUNEL, and Ki67.</figcaption></figure>

<figure><img src="image_83.png" /><figcaption>**FIGURE 9**
Immunohistochemical staining images of tumors formed by subcutaneous implantation of scaffolds. Two weeks and 1 month after implantation, tumor samples were subjected to immunohistochemical staining against PCNA and Bcl-2.</figcaption></figure>

Currently, it is believed that the ECM chemical cues are the primary forces behind cancer formation and progression. Although the ECM mechanical forces have previously received little attention, they are now believed to be crucial to the development of illness and malignant cellular activity (Walker et al., 2018). The ECM surrounds the tumor cells, which interact dynamically with cytokines and signal transduction in its highly complicated milieu. The tumor microenvironment is a fertile ground for cell proliferation and malignant transformation, which can deter immune attacks, allow cancer cells to escape immune surveillance, cause surrounding infiltration and metastasis, and affect prognosis (Lu et al., 2020). We examined the changes in the biological features of cells following *in vivo* and *in vitro* culture using a 3D scaffolding method in order to evaluate the interaction between cells and the microenvironment (Liu et al., 2020). A 3D culture system was constructed, and preliminary exploration of the tumor microenvironment was performed (Wei et al., 2018). The study looked at how cells interact with the matrix using the biological framework of the TME that was built (Kievit et al., 2010; Mao et al., 2012). Based on the abovementioned advantages, to enhance the SF/CTS (1:1) composite scaffolds' qualities and make them better suited for cell culture, we added CEM to them. Furthermore, the interior structure of the SF/CTS (1:1) scaffold is simpler than the tumor cell microenvironment *in vivo*, notably in terms of chemical composition, while being more complex than that of a 2D cell culture system. The scaffold's chemical complexity can be increased, improving its ability to mimic the tumor milieu. This

is accomplished by adding a CEM component. We predicted that the triple biomaterial composite scaffold would perform better than those consisting of just two biomaterials in terms of both features and performance. Our findings were intriguing in that HCT-116 cell spheres displayed tumor-like morphological characteristics seen *in vivo* in all four 3D scaffolds (SF/CTS (1:1), SF/CTS/CEM (1:1:0.5), SF/CTS/CEM (1:1:1), and SF/CTS/CEM (1:1:2)). The extremely aggressive activity typical of tumor cells *in vivo* was more likely to be displayed by cells growing in a 3D scaffold made of SF/CTS/CEM (1:1:1).

The Interaction between the OH- and COO- groups in SF, CTS, and CEM complexes is thought to produce interactions, enhancing the tensile strength of triple composite polymers. CEM might be added to the SF/CTS (1:1) composite scaffolds to increase their overall porosity and average pore diameter. Cells require physical room to function, which improves their ability to utilize nutrients and oxygen, as well as promoting effective removal of metabolic waste. These variables significantly affect the metabolic processes, cell attachment, proliferation, distribution, and differentiation (Nava et al., 2016). The optimal pore diameter depends on the specific cell type (Murphy et al., 2016). In our study, the SF/CTS/CEM (1:1:1) scaffold was the best for the culture and development of HCT-116 cells, due to its wide average pore width. High-porosity scaffolds improve mechanical interlocking and cell infiltration (Ionescu and Mauck, 2013). The SF/CTS/CEM (1:1:1) scaffold showed the best improvement of cell proliferation in our investigation and had the maximum porosity (88.33% ± 3.58%). For cell-infiltrating
<figure><img src="image_84.png" /><figcaption>**FIGURE 10**
The interior environment of the SF/CTS/CEM(1:1:1) scaffolds can be seen in SEM images. The indicated region demonstrates how cells are expanding within the pores.</figcaption></figure>

<figure><img src="image_85.png" /><figcaption>**FIGURE 11**
The figure shows the cell growth in the inner area of the scaffold with different material components. HCT-116 cells grow in the inner area of the SF/CS, SF/CTS/CEM(1:1:0.5), SF/CTS/CEM(1:1:1) and SF/CTS/CEM(1:1:2) scaffold (A-D, a-d). Compared with (A, a; B, b; D, d), the cells in SF/CTS/CEM (1:1:1) scaffolds group (C, c) grew in lumps and had stronger proliferation ability.</figcaption></figure>

scaffolds, higher water absorption is preferred. A lower expansion rate helps maintain the structural stability of the scaffold. Here, a rise in the CEM percentage was associated with an increase in swelling and water absorption. According to our findings, the SF/CTS/CEM (1:1:1) scaffold exhibited a high water uptake ratio (1,072% ± 48%), was porous, and degraded at an optimum rate (Figure 3). Further characterization of the biological features of these scaffolds showed that the cells on the SF/CTS/CEM (1:1:1) scaffold exhibited the highest cell adhesion and proliferation rates (Figure 4), which supported the cell growth and morphological characteristics shown in the SEM images (Figure 5). The complete analysis of the biological features suggest that the optimal scaffold for the *in vitro* investigation of CC cells may be the SF/CTS/CEM

(1:1:1) scaffold (Figure 10). Staining of the scaffold and cells revealed that the cells in the 3D scaffold kept their *in vivo* shape. Significant alterations in cell structure, protein expression, and mechanical properties occur during their migration through new tissues *in vivo* or in a new environment *in vitro*. It is well known that during movement, migration, adhesion, and proliferation, the cytoskeleton can undergo changes (Jin et al., 2012). In addition, cytoskeletal networks are essential for preserving cell shape (Ma et al., 2012). According to previous reports, a crucial condition for cell cycle progression is the cytoskeleton's proper configuration (Jiang et al., 2013). According to our findings, the 2D culture cells developed the slowest, followed by those on the SF/CTS/CEM (1:1:1) and the SF/CTS (1:1) scaffolds. The majority
of the cells in the 2D condition were long and fusiform, but the cells on the SF/CTS/CEM (1:1:1) and SF/CTS (1:1) scaffolds were round or nearly round (Figure 11). This indicated that the cells cultivated on scaffolds were more compatible with the growth status of cells in vivo. The HCT-116 cells transplanted on the 3D scaffold tended to assemble into multicellular spheres in contrast to the 2D monolayer model. These 3D multicellular aggregates serve as a useful alternative to traditional tumor models in cancer research (Hirschhaeuser et al., 2010). Our findings demonstrated that, compared to the SF/CTS (1:1) scaffold, the SF/CTS/CEM (1:1:1) scaffold supported sphere growth more significantly (Figure 5) and (Figure 11).

We also found that the cells cultured on SF/CTS/CEM (1:1:1) scaffolds showed significant proliferative ability, malignancy, and delayed tumor cell apoptosis compared with the those cultured in SF/CTS (1:1) scaffolds and 2D conditions. The PI3K/PDK/Akt/FoxO signaling pathway is a classical apoptosis regulator; therefore, we assessed the expression of this pathway. The findings demonstrated that p-Akt expression was upregulated and FoxO expression was downregulated in SF/CTS/CEM-cultured cells. These results suggest that the delay in apoptosis in scaffold culture is related to the PI3K/PDK/Akt/FoxO signal transduction pathway.

Despite the fact that our research sheds light on the development and possibility of fabricating composite scaffolds employing complimentary biopolymers, some issues still need to be addressed. As an example, the fundamental mechanisms observed in this study are still not well understood. In addition, in terms of porosity, water absorption, cell adhesion, proliferation, and sphere formation, our SF/CTS/CEM (1:1:1) scaffold performed well, although it came in second place in terms of disintegration rate. The association between the rate of degradation and cell proliferation cannot be inferred. Before these 3D models may be altered and widely used in medicine and basic science, further research is required to validate and clarify our findings and to better understand the potential mechanisms of cell growth and proliferation on these scaffolds.

## 5 Conclusion

The biological activity of CC cells differed depending on the type of material employed, as demonstrated by our analysis of the different HCT-116 cell culture conditions (2D, SF/CTS (1:1), and SF/CTS/CEM (1:1:1) scaffolds). According to our findings, the SF/CTS/CEM (1:1:1) scaffold has the potential to be a useful tumor model for CC cell culture studies and a realistic representation of cell growth in a 3D environment, as in living organisms.

## Data availability statement

The original contributions presented in the study are included in the article/Supplementary Material, further inquiries can be directed to the corresponding authors.

## Ethics statement

The animal study was reviewed and approved by Department of Laboratory Animal Science, Fudan University. Written informed consent was obtained from the owners for the participation of their animals in this study.

## Author contributions

ZC, CK, and LC conceived the project and supervised all experiments. ZC designed the experiments and wrote the manuscript. LC analyzed the data. GN, YL, ZH, RH, XZ, and LH provided support with experimental techniques. The manuscript was revised by LC, SH, GN, YL, LH, and CK. All authors reviewed the manuscript. All authors read and approved the final manuscript.

## Funding

This study was supported by the Medical System of Shanghai Minhang District (No.2020MWDXK02), the High-level professional physician training program of Minhang District (No.2020MZYS21), and the Health Profession Clinical Research Funds of Shanghai Municipal Health Commission (No.20204Y0390).

## Acknowledgments

We appreciate Editage's (www.editage.cn) assistance with English language editing.

## Conflict of interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

## Publisher's note

All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher.

## Supplementary material

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fbioe.2023.1139649/full#supplementary-material