SA microgels were prepared following established protocols [16, 17]. A calcium-EDTA (Ca-EDTA) complex was formulated by combining calcium chloride and disodium EDTA in a 1:1 molar ratio, and adjusting the pH to 7.4 using sodium hydroxide. The aqueous phase consisted of 1% SA (MACKLIN, Shanghai, China) and 50 mM Ca-EDTA dissolved in distilled water, followed by filtration through a 0.22 µm syringe filter (FMC201030, JETBIOFIL, Guangzhou, China). For cell-laden microgels, primary human PMCs (PHPMCs) at a density of 5 $\times$ 10⁶ cells/mL were suspended in the aqueous phase. Two milliliters of droplet generation oil (FluidicLab, China) were filtered using a 0.22 µm syringe filter. All microfluidic encapsulation devices were sterilized with 75% ethanol and exposed to ultraviolet light overnight prior to use. The aqueous and oil phases were loaded into separate syringes and mounted on a laboratory syringe pump (LSP01-3A, LONGER, Tucson, AZ, USA), connected to a microfluidic chip (PDMS-FF-100B, FluidicLab, Shanghai, China) to ensure uniformity. Flow rates were maintained at 7 µL/min for the aqueous phase and 35 µL/min for the oil phase. The resultant gel was collected in culture medium for subsequent experiments. The encapsulation rate and release curve of sodium alginate microgel were determined by cell counting kit 8 (CCK-8) method.

The study protocol was approved by the Human Ethics Committee of Wenzhou Medical University (Acceptance number: KY2024-R179). The research adhered to Good Clinical Practice guidelines and followed the principles of the Declaration of Helsinki. PHPMCs were isolated from PD effluent of patients undergoing dialysis at the First Affiliated Hospital of Wenzhou Medical University. Inclusion criteria were: (1) absence of peritonitis during PD; (2) regular PD follow-up; (3) age between 40 and 60 years; (4) no malignant tumors; (5) no ascites; and (6) signed informed consent. No statistically significant disparities in age, gender, or educational background were observed across the participant cohorts (p $>$ 0.05). PD effluent samples were collected after durations of 1 week (1W), 1 year (1Y), and 3 years (3Y) of PD (n = 30 per group). The surface of the PD fluid specimen bag was disinfected with 75% ethanol before sterile collection. Samples were aliquoted into 50 mL centrifuge tubes and centrifuged at 1000 $\times$g for 5 minutes at 4 °C to pellet the PHPMCs, with the supernatant discarded. Cells were resuspended in Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F12) (Gibco, CA, USA) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin, and cultured in dishes at 37 °C with 5% CO₂. The medium was refreshed every 2–3 days to remove non-adherent cells. PHPMCs were characterized by immunofluorescence staining for cytokeratin 18 (CK18) and vimentin to confirm mesothelial origin. Upon reaching 80–90% confluence, cells were trypsinized with 0.25% trypsin-EDTA and reseeded for expansion.

In vitro models were established as follows: (1) the control (Con) group: PHPMCs cultured in DMEM/F12 medium for 96 h; (2) the high glucose (G) group: HPMCs cultured in DMEM/F12 medium supplemented with 60 mM glucose for 48 h to induce fibrosis, followed by replacement with normal DMEM/F12 medium for an additional 48 h; (3) the high glucose+SA (G+SA) group: PHPMCs cultured in high-glucose medium (60 mM) for 48 h, followed by co-culture with freshly prepared blank SA microgels for an additional 48 h (total 96 h); (4) the high glucose+SA/PMC (G+SA/PMC) group: PHPMCs cultured in high-glucose medium (60 mM) for 48 h, followed by co-culture with freshly prepared SA microgels encapsulating additional PHPMCs for another 48 h (total 96 h). Since PHPMCs are non-immortalized cells, they were used after 5–7 passages in both the in vivo and in vitro experiments.

Cell proliferation was assessed using the CCK-8 assay kit (Beyotime, Shanghai, China). PMCs were seeded in 96-well plates at a density of 5 $\times$ 10³ cells per well and incubated at 37 °C with 5% CO₂ for 24 h to allow cell attachment. Subsequently, PMCs were co-incubated with SA and SA/PMC for 0, 24, 48, and 72 h. Following the designated incubation period, CCK-8 reagent was introduced into each well, subjected to gentle agitation, and further incubated for 1 h. Absorbance measurements (optical density, OD) were then conducted at 450 nm with a microplate reader (Thermo Fisher Scientific, Waltham, MA, USA). Each treatment group included at least three replicates, and the experiment was repeated three times. Cell proliferation was evaluated by comparing OD values across treatment groups at each time point. The PMCs cell lines were validated by STR profiling and tested negative for mycoplasma.

BALB/c nude mice (male, 8 weeks old) were provided by Charles River Laboratory Animal Technology Co., Ltd (Zhejiang, China). The mice were housed under standard laboratory conditions with a 12-h light/dark cycle, a temperature of 22 $\pm$ 1 °C, and ad libitum access to food and water. After one-week of acclimatization, the mice were randomly divided into five experimental groups (n = 3–5 per group): (1) the Con group received daily intraperitoneal (IP) administration of 10 mL/100 g of physiological saline for four consecutive weeks; (2) the PD group received IP injections of 10 mL/100 g of 4.25% glucose PD solution for four consecutive weeks; (3) the PD+SA group received IP injections of 10 mL/100 g of 4.25% glucose PD solution for four consecutive weeks, and SA microgels at the beginning of the second week; (4) the PD+PMC group received IP injections of 10 mL/100 g of 4.25% glucose PD solution for four consecutive weeks, and 5 $\times$ 10⁶ PHPMCs at the beginning of the second week; (5) the PD+SA/PMC group received IP injections of 10 mL/100 g of 4.25% glucose PD solution for four consecutive weeks, and SA microspheres encapsulating 5 $\times$ 10⁶ PHPMCs at the beginning of the second week. The body weights of the mice were systematically monitored and documented both at baseline and at regular intervals during the experimental timeline.

To begin the terminal sampling procedure, mice were anesthetized with isoflurane vapor. Induction used 3–5% isoflurane in 100% oxygen (via nose cone or chamber), with a 1–3% concentration to maintain anesthesia. The depth of anesthesia was monitored every 2–3 minutes by checking foot-pinch reflexes and breathing rate. Throughout the process, the mice rested on a heating pad for thermoregulation, and eye lubricant was applied to prevent dryness. Since this was a terminal procedure, no post-operative pain management was needed. Once a surgical plane of anesthesia was achieved, we immediately performed cardiac blood collection, perfusion, or organ collection. For euthanasia, we used CO₂ inhalation ($\ge$99% purity) at a fill rate of 30–70% of the chamber volume per minute without pre-charging. After the animals lost consciousness, CO₂ flow was continued for at least one minute. We then applied a secondary method, such as bilateral thoracotomy, exsanguination, or cervical dislocation, to ensure death. The blood, liver, heart, lungs, kidneys, and peritoneum of the mice were collected and preserved. Some portions of the tissues were immediately stored in a –80 °C freezer, while other portions were fixed in 4% paraformaldehyde.

Harvested tissue specimens, including those from the liver, heart, lung, kidney, and peritoneum, underwent fixation in 4% paraformaldehyde for a duration of 48 h. Fixed tissues were dehydrated through a graded ethanol series, cleared in xylene, and embedded in paraffin. Paraffin-embedded tissues were sectioned at a thickness of 5 µm using a microtome. Deparaffinization of tissue sections was carried out using xylene, followed by rehydration via a graded ethanol series, and culminating in immersion in distilled water. H&E staining was performed according to the instructions provided in the stain kit (Solarbio, Beijing, China). Sections were stained with hematoxylin for 5 minutes to visualize nuclei, then rinsed in running tap water. Eosin staining was performed for 2 minutes to stain the cytoplasm and extracellular matrix. Stained sections were dehydrated through increasing ethanol concentrations, cleared in xylene, and mounted with a coverslip using a synthetic resin. Images were captured using a Leica microscope (VT1000S, Shanghai, China). All experiments were repeated at least in triplicate.

MASSON staining was performed using a stain kit (Solarbio, Beijing, China) and following the manufacturer’s protocol. Sections were immersed in Bouin solution overnight at 37 °C, then rinsed with running water until the yellow color disappeared. The sections were subsequently stained with aniline blue for 2 minutes and briefly rinsed with water. Mayer’s hematoxylin was applied for 2 minutes, followed by a brief water rinse. The sections were differentiated in acidic ethanol for a few seconds and washed in running water for 10 minutes. Biebrich scarlet-acid fuchsin was applied for 10 minutes, the sections were then briefly rinsed with distilled water, followed by treatment with phosphomolybdic acid solution for about 10 minutes. Without washing, the aniline blue solution was directly applied for 5 minutes. The sections were subsequently treated with a weak acid solution for 2 minutes and quickly dehydrated in 95% ethanol. They were then dehydrated in absolute ethanol three times, each for 5–10 seconds, followed by clearing in xylene three times, each for 1–2 minutes. Finally, the sections were mounted with neutral resin, and images were captured using a Leica microscope. All experiments were repeated at least in triplicate.

Immunohistochemical analysis was conducted on tissue sections prepared from paraffin-embedded samples. The 4 µm slices were dewaxed in xylene and hydrated with graded ethanol solutions. Antigen retrieval was performed with citrate buffer, followed by treatment with 3% hydrogen peroxide. Subsequently, sections were blocked with goat serum for 1 h. The tissue sections were then incubated overnight at 4 °C with primary antibodies, including anti-periostin (POSTN) (1:100, DF6746, Affinity, Jiangsu, China), anti-nuclear factor kappa-B (NF-$\kappa$B) P65 (1:100, ab32536, Abcam, Cambridge, MA, USA), anti-phospho-NF-$\kappa$B P65 (p-P65, 1:100, AF2006, Affinity, China), anti-CXCL8 (1:100, DF6998, Affinity, China), anti-$\alpha$-smooth muscle actin ($\alpha$-SMA, 1:100, AF1032, Affinity, China), anti-collagen 1 (Col-1, 1:100, AF7001, Affinity, China), anti-transforming growth factor-$\beta$ (TGF-$\beta$, 1:100, BF8012, Affinity, China), and anti-FN (1:100, AF5335, Affinity, China). After washing with phosphate buffered saline (PBS), HRP-labeled secondary antibodies were applied, followed by 3,3^′-diaminobenzidine (DAB) chromogenic staining to visualize the bound antibodies, and hematoxylin counterstaining to highlight the nuclei. Images were captured using a Leica microscope, and all experiments were repeated in triplicate.

Paraffin-embedded tissue sections (4 µm) were dewaxed in xylene and rehydrated through graded ethanol solutions. Antigen retrieval was performed using citrate buffer followed by 4% paraformaldehyde. Cell samples were washed three times with PBS and the cells then fixed with 4% paraformaldehyde. Both tissue and cell samples were permeabilized with 0.1% Triton X-100, and blocked with goat serum for 1 h. Samples were then incubated overnight at 4 °C with one or two primary antibodies from different sources. After washing with PBS, samples were incubated at 37 °C for 1 h with corresponding secondary antibodies, such as Alexa Fluor 488 goat anti-rabbit (A-11008, Invitrogen, Carlsbad, CA, USA) or Alexa Fluor 594 goat anti-mouse (A-11005, Invitrogen, Carlsbad, CA, USA). 4^′,6-diamidino-2-phenylindole (DAPI) (Solarbio) was used for nuclear visualization. Images were captured using a KFBIO digital pathology slide scanner (KF-PRO-005, KFBIO, Ningbo, Zhejiang, China). All experiments were repeated at least in triplicate.

Animal tissue samples were homogenized using a tissue homogenizer in radio immunoprecipitation assay (RIPA) lysis buffer (Beyotime, Shanghai, China) containing 0.1% phenylmethylsulfonyl fluoride (PMSF). Cells were lysed using RIPA lysis buffer with 0.1% PMSF. The homogenized cell lysates were sonicated at 30 Hz for 3 seconds, repeated three times. Both cell and animal tissue samples were then centrifuged at 15,000 rpm for 20 minutes at 4 °C, and the supernatant collected for protein analysis. Protein concentrations were determined using a protein quantification kit. Protein samples were prepared with a 5$\times$ loading buffer and adjusted with PBS to ensure equal loading. Equivalent quantities of protein were resolved via sodium dodecyl sulfate-polyacrylamide gel electrophoresis and then electrophoretically transferred onto polyvinylidene fluoride membranes. After blocking for 2 h with 5% skim milk, the membrane was washed with 1$\times$ tris buffered saline with tween-20 (TBST) and incubated with primary antibodies overnight at 4 °C. The next day, the membrane was washed with TBST and incubated with the corresponding Horseradish Peroxidase-labeled (HRP-labeled) secondary antibody for 1 h at room temperature. Protein bands were visualized using an electrochemiluminescence detection kit (4AW011, 4A Biotech, Suzhou, China), with the signal captured using a chemiluminescence imaging system. Band intensities were quantified by analyzing grayscale values using ImageJ software (National Institutes of Health, Bethesda, MD, USA). All experiments were repeated at least in triplicate.

Total RNA was extracted from tissue or cell samples using a commercially available RNA extraction kit (Beyotime, Shanghai, China) according to the manufacturer’s instructions. RNA concentration and purity were assessed using a spectrophotometer (DS-11 FX/FX+, Denovix, Wilmington, DE, USA). Complementary DNA (cDNA) was synthesized from 1 µg of total RNA with a reverse transcription kit (Vazyme, Nanjing, China). Quantitative real-time PCR was conducted on a QuantStudio 5 instrument (Thermo Fisher Scientific, USA) using SYBR Green chemistry. Relative gene expression levels were determined with the comparative threshold cycle (Ct) method, and normalized to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (internal reference gene). All experiments were repeated at least in triplicate.

Prism 9 software (GraphPad Software, San Diego, CA, USA) was utilized to generate bar charts and conduct statistical analyses. Data conforming to a normal distribution are expressed as the mean $\pm$ standard deviation, while non-normally distributed data are reported as the median with interquartile range. For the comparison of data between groups, the two independent samples t-test was applied when the variance between groups was homogeneous. A p-value 0.05 was considered statistically significant.

The production process for SA microgels is illustrated in Fig. 1a. Optical microscopy revealed the diameter of SA microgels ranged from approximately 60 to 100 µm (Fig. 1b). Encapsulation of PHPMCs within SA (SA/PMC) did not significantly alter the microgel diameter. CCK-8 assays demonstrated a significant increase in PMC proliferation after 48 h of co-culture with both SA and SA/PMC. Moreover, SA/PMC exhibited accelerated cell proliferation compared to the SA-only group (Fig. 1c). As the ratio of the aqueous phase to the oil reservoir flow rate (Va/Vo) increases, the entrapment efficiency shows a downward trend (Fig. 1d). Meanwhile, with the extension of time, the cumulative release rate gradually rises, reflecting that this hydrogel has a certain time dependence in substance release (Fig. 1e). These findings suggest that SA microgels effectively encapsulate PHPMCs and facilitate their release, thereby enhancing cell proliferation.

Fig. 1.

Characteristic of sodium alginate (SA) microgel. (a) Production of SA microgel. (b) Microstructure of the SA and SA microgels encapsulating peritoneal mesothelial cells (SA/PMCs). Scale bar = 200 µm. (c) Cell counting kit 8 (CCK-8) OD values at 450 nm for primary human peritoneal PMCs co-cultured with SA (PMC+SA), and primary human PMCs co-cultured with SA/PMC (PMC+SA/PMC). (d) Cell encapsulation efficiency under different aqueous and oil phase flow rates (Va/Vo). (e) Cumulative cell release rate. All data are presented as the mean $\pm$ SD. *p 0.05, ***p 0.001.

Co-staining with cytokeratin 18 (CK18) and vimentin confirmed the identity of cells isolated from PD fluid as PHPMCs (Fig. 2a). PD fluid was collected from patients with PD durations of 1W, 1Y, and 3Y. A high number of PHPMCs was collected in the 1W-PD group, with these cells displayed a cobblestone-like morphology when cultured for 11 days. In contrast, fewer PHPMCs were found in the 1Y-PD group, with the cells undergoing EMT characterized by transition from a cobblestone-like to a spindle-shaped morphology. The 3Y-PD group showed even fewer PHPMCs, with the cells losing their proliferative ability almost entirely (Fig. 2b). These observations indicate that prolonged PD may lead to cell fibrosis and induce EMT.

Fig. 2.

Reduced proliferative capacity of PHPMCs was associated epithelial-mesenchymal transition in long-term peritoneal dialysis (PD) patients. (a) Representative immunofluorescence images of markers vimentin and cytokeratin 18 (CK18) in PHPMCs. Scale bar = 50 µm. (b) The growing images of PHPMCs of PD patients for 1 week (1W-PD), 1 year (1Y-PD), and 3 year (3Y-PD). Scale bar = 200 µm. D, day.

To investigate the mechanisms underlying peritoneal fibrosis in long-term PD patients, PHPMCs were isolated from PD fluid of patients undergoing dialysis for 1W and 1Y. According to the sequencing results, POSTN and CXCL 8 were increased in patients undergoing peritoneal dialysis for 1 year (Fig. 3a). Immunofluorescence staining indicated a marked upregulation of POSTN expression in the 1Y-PD group relative to the 1W-PD cohort. NF-$\kappa$B p65, a downstream target of POSTN, also exhibited increased expression in the 1Y-PD group, as confirmed by immunofluorescence. The activation and nuclear translocation of p-p65 exhibited pronounced nuclear accumulation in the 1Y-PD group. Furthermore, CXCL8, an inflammatory marker associated with peritoneal fibrosis, showed increased protein expression in the 1Y-PD group (Fig. 3d). Importantly, fibrosis-related markers, such as $\alpha$-SMA, Col-1, TGF-$\beta$, and FN also showed elevated protein expression in the 1Y-PD group (Fig. 3b). The same results were comfirmed by qRT-PCR and western blots (Fig. 3c–o). Collectively, these findings suggest that in patients undergoing peritoneal dialysis for one year, peritoneal mesothelial cells demonstrate upregulated expression of POSTN, NF-$\kappa$B, and CXCL8, along with elevated fibrosis markers.

Fig. 3.

Increased expression of periostin/nuclear factor kappa-B/C-X-C motif chemokine ligand 8 (POSTN/NF-$\kappa$B/CXCL8) in long-term PD patients with peritoneal fibrosis. (a) Heatmap showing differential gene expression in PMCs from 1W-PD and 1Y-PD patients. (b) Representative immunofluorescence images of staining for POSTN, NF-$\kappa$B P65, p-P65, CXCL8, $\alpha$-smooth muscle actin ($\alpha$-SMA), collagen I (Col-1), transforming growth factor-$\beta$ (TGF-$\beta$), and FN in PMCs from 1W-PD and 1Y-PD patients. Scale bar = 50 µm. (c–i) mRNA expression levels in the two groups. (j–o) Representative Western blot images and quantitative analysis of POSTN, p-P65/P65, CXCL8, $\alpha$-SMA, and Col-1 expression in the two groups. All data are presented as the mean $\pm$ SD. **p 0.01, ***p 0.001.

Primary human PMCs were cultured in a 60 mM high glucose medium to examine the role of elevated POSTN expression in fibrosis under high glucose conditions. Immunofluorescence staining revealed increased POSTN expression in the G and G+SA groups, whereas the G+SA/PMC group showed reduced POSTN expression. Similarly, NF-$\kappa$B P65 expression was elevated in the G and G+SA groups. High glucose also promoted the phosphorylation and nuclear translocation of P65, thereby increasing CXCL8 expression, but this was inhibited by SA/PMC. Furthermore, elevated expression of fibrosis-related markers ($\alpha$-SMA, Col-1) was seen in the G and G+SA groups, whereas the G+SA/PMC group exhibited reduced levels (Fig. 4a). mRNA analysis confirmed that high glucose increased POSTN/NF-$\kappa$B/CXCL8 expression, while SA/PMC treatment reduced the expression of these markers (Fig. 4b–h). Western blot results also demonstrated increased expression levels of POSTN, NF-$\kappa$B P65, and CXCL8 in the G group, with reduced levels in the G+SA/PMC group. The fibrosis marker $\alpha$-SMA was similarly increased in the G group and decreased in the G+SA/PMC group (Fig. 4i–m). These results demonstrate that high glucose induces POSTN/NF-$\kappa$B/CXCL8 expression and EMT in PMCs, which can be mitigated by SA/PMC treatment.

Fig. 4.

SA/PMC reduces the high glucose-induced increase in POSTN/NF-$\kappa$B/CXCL8 expression and fibrosis in PHPMCs. (a) Representative immunofluorescence images of POSTN, NF-$\kappa$B P65, p-P65, CXCL8, $\alpha$-SMA and Col-1 staining in the Control (Con), high glucose (G), high glucose+SA (G+SA), and high glucose+SA/PMC (G+SA/PMC) groups. Scale bar = 50 µm. (b–h) mRNA expression level of POSTN, NF-$\kappa$B P65, CXCL8, $\alpha$-SMA, Col-1, TGF-$\beta$, and FN in the four groups. (i–m) Representative Western blot images and quantitative analysis of POSTN, NF-$\kappa$B P65, CXCL8, and $\alpha$-SMA in the four groups. All data are presented as the mean $\pm$ SD. *p 0.05, **p 0.01, ***p 0.001.

A murine model of PD was used to further investigate the involvement of POSTN in peritoneal fibrogenesis. MASSON staining indicated significant peritoneal fibrosis in the PD, PD+SA, and PD+PMC groups, whereas the PD+SA/PMC group showed reduced fibrosis (Fig. 5a). Additionally, the peritoneal thickness was significantly greater in the PD and PD+SA groups compared to the control group, but significantly smaller in the PD+SA/PMC group compared to the PD and PD+PMC groups (Fig. 5b). Immunohistochemical assessment indicated elevated POSTN expression in the PD group, which was substantially attenuated in the PD+SA/PMC treatment group. Importantly, POSTN expression in the PD+SA/PMC group was lower than in the PD+PMC group. Similarly, the levels of NF-$\kappa$B P65, p-P65, and CXCL8 were elevated in the PD group and reduced in the PD+SA/PMC group (Fig. 5c–g). Western blot analysis corroborated the findings that POSTN, NF-$\kappa$B, and CXCL8 protein expression levels were elevated in the PD group, but significantly reduced following PD+SA/PMC treatment, with lower levels in the PD+SA/PMC group compared to the PD+PMC group (Fig. 5h–k). Additionally, immunofluorescence showed increased fluorescence intensity for POSTN, NF-$\kappa$B P65, p-P65, CXCL8, $\alpha$-SMA, Col-1, and TGF-$\beta$ in the PD group, which was reduced in the PD+SA/PMC group (Fig. 5l,m). These findings indicate that high glucose PD induces POSTN/NF-$\kappa$B/CXCL8 expression and peritoneal fibrosis in mice, with SA/PMC treatment showing superior antifibrotic effects compared to SA.

Fig. 5.

Increased expression of POSTN/NF-$\kappa$B/CXCL8 and marked peritoneal fibrosis in the mouse PD model. (a) Hematoxylin and eosin (H&E) and MASSON staining of parietal peritoneum in the mouse PD model. Scale bar = 50 µm. (b) Thickness of the parietal peritoneum. (c) Representative immunohistochemical staining for POSTN, NF-$\kappa$B P65, p-P65, and CXCL8. Scale bar = 10 µm. (d–g) Quantitative analysis of the expression of POSTN, NF-$\kappa$B P65, p-P65, and CXCL8 shown in (c). (h–k) Representative Western blot images and quantitative analysis of POSTN, NF-$\kappa$B P65, and CXCL8 proteins shown in (h). (l) Representative immunofluorescence images of staining for POSTN, NF-$\kappa$B P65, p-P65 and CXCL8. Scale bar = 10 µm. (m) Representative immunofluorescence images of staining for $\alpha$-SMA, Col-1, and TGF-$\beta$. All data are presented as the mean $\pm$ SD. Scale bar = 10 µm. *p 0.05, **p 0.01, ***p 0.001.

To evaluate the biosafety of SA microgels, blood and tissue samples were collected from the various groups of PD mice. Hemolysis tests indicated that adding SA to mouse blood resulted in a hemolysis rate of 2%, suggesting good blood compatibility (Fig. 6a,b). Moreover, no significant changes in body weight were observed during the PD period (Fig. 6c). To assess the impact of SA microgel on liver and kidney function, we measured the levels of alanine aminotransferase (ALT), aspartate aminotransferase (AST), blood urea nitrogen (BUN), serum creatinine (Scr), and glomerular filtration rate (eGFR). None of these parameters showed any significant abnormalities (Fig. 6d–g). Furthermore, histological analyses using H&E and Masson staining revealed no significant abnormalities in liver, heart, lung, and kidney tissues (Fig. 6h–k). Collectively, these findings suggest that SA microgels exert minimal adverse effects on the physiological and histological parameters assessed, indicating their potential safety as a therapeutic modality.

Fig. 6.

Biosafety of SA microgels. (a) Images of hemolysis test and hemolysis rate. (b) Body weight of mice before and after PD. (c) Alanine aminotransferase (ALT) level of PD mice. (d) Aspartate aminotransferase (AST) level. (e) Blood urea nitrogen (BUN) level. (f) Serum creatinine (Scr) level. (g) Glomerular filtration rate (eGFR). (h–k) Representative H&E and MASSON images of liver, heart, lung, and kidney. All data are presented as the mean $\pm$ SD. Scale bar = 50 µm. ns, no significant difference.