The current study was approved by the Ethics Committee of Zhoupu Hospital (approval no. ZPYYLL-2018-02; Shanghai, China) and was carried out in accordance with The Declaration of Helsinki. Three healthy male donors aged 54±14 years and three patients with UC (two male patients and one female patient) aged 34±19 years were selected for the present study. Peripheral colonic tissue from pediatric (age, <16 years) and adult subjects were collected after obtaining written informed consent from the patients or the pediatric patient's parent or guardian. The samples were collected during colonoscopy. The healthy donors underwent colonoscopy as part of a routine health check-up at the hospital. The patients with UC were diagnosed by a gastroenterologist and a pathologist. The puncture performed during colonoscopy yielded colon tissue, which underwent hematoxylin and eosin (H&E) staining and immunofluorescence detection using the following antibodies: Anti-CD62P (1:200; cat. no. 60322-1-Ig; Proteintech Group, Inc.), anti-inducible nitric oxide synthase (iNOS; 1:200; cat. no. 18985-1-AP; Proteintech Group, Inc.), anti-PF4 (1:100; cat. no. 21157-1-AP; Proteintech Group, Inc.) and anti-CXCR3 (1:100; cat. no. 26756-1-AP; Proteintech Group, Inc.).

THP-1 cells (cat. no. SCSP-567) were obtained from The Cell Bank of Type Culture Collection of The Chinese Academy of Sciences and were cultured in RPMI 1640 culture medium (cat. no. L210KJ; Shanghai Basalmedia Technologies Co., Ltd.) supplemented with 10% fetal bovine serum (cat. no. 098-150; Wisent, Inc.) and 1% penicillin-streptomycin (cat. no. 15140122; Gibco; Thermo Fisher Scientific, Inc.). The cells were maintained at 37°C with 95% O₂ and 5% CO₂ in a humidified atmosphere. Phorbol 12-myristate 13-acetate (PMA), PF4, AMG487 (CXCR3 inhibitor), PD98059 (ERK inhibitor) and SC75741 (p65 inhibitor) were purchased from MedChemExpress (cat. nos. HY-18739, HY-P70618, HY-15319, HY-12028 and HY-10496, respectively). Lipopolysaccharide (LPS), which was used to induce the polarization of THP-1 cells to a proinflammatory phenotype, was purchased from Beijing Solarbio Science & Technology Co., Ltd. (cat. no. L8880). LPS (1 μg/ml) was added to THP-1 cells for 6 h at 37°C and the samples were collected. PMA (100 ng/ml) was added to THP-1 cells for 24 h at 37°C to induce the differentiation of monocytes into macrophages (24). After adding PMA to THP-1 cells to transform them into macrophages, the cells were treated as required. Subsequently, the samples were treated with AMG487 (2.5 μM) (25), PD98059 (10 μM) (26) or SC75741 (5 μM) (27) for 2 h at 37°C, after which PF4 (100 ng/ml) was added for a further 6 h at 37°C and the samples were collected.

Human venous blood (5 ml) was collected from three of the aforementioned healthy volunteers at the Zhoupu Hospital Affiliated to Shanghai University of Medicine and Health Sciences (Shanghai, China). A portion of the blood was placed in a sodium citrate anticoagulant tube (BD Biosciences) and was subjected to centrifugation at 200 × g for 15 min at room temperature, thereby obtaining platelet-rich plasma (PRP). Prostaglandin 2 (500 ng/ml; Sigma-Aldrich; Merck KGaA) was added to PRP to prevent platelet activation. Platelet precipitation was carried out by centrifugation at 800 × g for 3 min at room temperature. The platelets were resuspended and washed in Tyrode's Salts buffer (140 mM NaCl, 10 mM NaHCO₃, 2.5 mM KCl, 0.5 mM Na₂HPO₄, 1 mM MgCl₂, 22 mM sodium citrate and 0.55 mM glucose; pH 6.5; Sigma-Aldrich; Merck KGaA), counted, and added to RPMI 1640 complete culture medium in an appropriate proportion for use (6). THP-1 cells were co-cultured with platelets at a ratio of 1:100 for 6 h. Light-field microscopic observations were performed and images were captured using an inverted microscope (DMi1; Leica Biosystems).

A total of 20 healthy female Balb/c mice (age, 6-8 weeks; weight, 20-22 g) were provided by Hangzhou Qizhen Experimental Animal Technology Co., Ltd. The experimental procedure was conducted in accordance with the guidelines set by the National Institutes of Health (28), and all animal experiments were approved by the Animal Care and Use Professional Committee of Zhoupu Hospital (approval no. ZPYYLL-2018-02). The mice were maintained under the following controlled conditions: Temperature, 23±1°C; humidity, 40-50%; 12-h light/dark cycle; ad libitum access to food and water. A 7-day acclimation period was allowed for the mice prior to the commencement of the study. The animals were euthanized if any case-predefined humane endpoints were observed, including: i) Weight loss, rapid loss of 15-20% of original body weight; ii) weakness, unable to eat and drink on their own, unable to stand for up to 24 h or unable to stand with extreme reluctance; and iii) signs of depression and hypothermia (<37°C) without anesthesia or sedation. No humane endpoints were met throughout the experiments.

A murine model of acute colitis was established by administering 2.5% DSS (Shanghai Yeasen Biotechnology Co., Ltd.) as previously described (29). The mice were randomly divided into the following four groups (n=5/group): Control, DSS, clopidogrel and AMG487 groups. Clopidogrel is a commonly used drug to inhibit platelet activation, and inhibition of platelet activation can reduce the production of platelet factors, including PF4. Following a 1-week period of acclimation, all experimental groups with the exception of the control group were administered a DSS solution, which was freshly prepared and replaced every other day, and the mice freely drank water mixed with DSS; mice were treated with AMG487 or clopidogrel (or control) for 3 days before DSS after the 1 week acclimation period. The clopidogrel group was administered an oral gavage of 20 mg/kg/day clopidogrel (MedChemExpress) (30,31), whereas the AMG487 group was administered an intraperitoneal injection of 5 mg/kg/day AMG487 (32). The control and DSS model groups were administered an intraperitoneal injection of a sterile PBS solution containing 20% β-cyclodextrin (MedChemExpress). The disease activity index (DAI) and histological examination (H&E staining) were used to assess the colitis model. DAI was scored as follows: Weight loss: 0, no weight loss; 1, 0-5%; 2, 5-10%; 3, 10-15%; 4, 15-20%. The degree of fecal softness: 0, no change; 1, soft stool; 2, loose stools. Fecal occult blood condition: 0, the color did not change within 2 min; 1, 1-2 min discoloration; 2, color change within 1 min; 3, discoloration within 10 sec; 4, immediate discoloration or hematochezia. The sum of the three scores is the DAI score. The fecal occult blood test was performed with the use of a fecal occult blood kit (cat. no. BA2020B; Baso Diagnostic, Inc.). Upon completion of the animal experiments, anesthesia was initiated with 3% isoflurane and was maintained with 1.5% isoflurane, and blood (0.5 ml) was taken from the apex of the heart, after which, the mice were euthanized with CO₂ at a volume replacement rate of 40% vol/min according to merican Veterinary Medical Association Guidelines for the Euthanasia of Animals: 2020 Edition (33). The colon was then removed for subsequent analysis. The blood samples were allowed to stand at room temperature for 60 min prior to centrifugation. After centrifugation at 2,000 × g for 15 min at room temperature, serum was obtained to assess the levels of inflammatory cytokines. The colon tissues were measured and washed to remove non-tissue debris. Subsequently, additional colon samples were stored at −80°C for future analysis.

Human and murine colon samples were sectioned after fixation with 4% paraformaldehyde for 24 h at room temperature and embedded in paraffin (Biosharp Life Sciences). The samples were then cut into 5-μm sections and mounted on slides. The sections were stained with H&E and then observed under a light microscope. Briefly, paraffin-embedded sections were oven-dried at 60°C for 2 h, deparaffinized and stained with hematoxylin for 10 min, rinsed and stained with eosin for 30 sec, washed and sealed; the entire procedure was operated at room temperature. Two pathologists scored the extent of intestinal mucosal damage and inflammatory cell infiltration in a blinded manner. Briefly, in the case of damage to the intestinal mucosa, the following criteria were applied: 0, no damage; 1, discrete mucosal epithelial damage; 2, superficial mucosal erosion; 3, extensive mucosal damage. For crypt abnormalities, the following criteria were applied: 0, normal crypts; 1, a few crypts showed structural changes or atrophy; 2, structural changes or atrophy of numerous crypts; 3, extensive crypt abnormalities and loss. For inflammatory cell infiltration, the following criteria were applied: 0, none or a small amount of inflammatory cells in the lamina propria; 1, increased numbers of inflammatory cells in the lamina propria; 2, inflammatory cells spread to the submucosa; 3, the whole layer has inflammatory cell infiltration. The sum of the three scores was calculated as the colon histological score.

A total of 1×10⁶ THP-1 cells/well were seeded into 6-well plates and divided into the following five experimental groups: i) Control group; ii) LPS group; iii) PLT group; iv) LPS + PLT group and v) LPS + PLT + CaCl₂ group. In the PLT groups, the THP-1 cells were co-cultured with platelets at a ratio of 1:100 for 6 h. In the LPS group, cells were collected 6 h after stimulation with LPS. In the LPS + PLT group, LPS was added during co-culture. In the LPS + PLT + CaCl₂ group, 10%w/v CaCl₂ was added to platelet conditioned medium for 5 min before co-culture with THP-1, and LPS was added at the same time. CaCl₂ is a commonly used platelet activator. All experiments were performed at 37°C. In addition, PMA (100 ng/ml) was added to THP-1 cells for 24 h at 37°C to induce the differentiation of monocytes into macrophages; these treatments are separate to the Control, LPS, PLT and LPS + PLT groups. After adding PMA to THP-1 cells to transform them into macrophages, experiments related to PF4/CXCR3 were performed. Subsequently, the samples were treated with AMG487 (2.5 μM/ml), PD98059 (10 μM) or SC75741 (5 μM) for 2 h at 37°C, after which PF4 (100 ng/ml) (25) was added for a further 6 h at 37°C and the samples were collected.

Total RNA was isolated from treated THP-1 cells and mouse colon tissue samples using TRIzol^® (Invitrogen; Thermo Fisher Scientific, Inc.); briefly, the cells were washed with PBS, the supernatant was discarded and TRIzol was added for 10 min. Subsequently, chloroform was added and mixed vigorously for 30 sec, the supernatant was left for 10 min, and was then centrifuged at 12,000 × g for 15 min at 4°C, and an equal volume of isopropanol was added for 10 min. After centrifugation at 12,000 × g for 15 min at 4°C, the supernatant was discarded and then pre-cooled 75% ethanol was added to the remaining pellet. The RNA precipitate was obtained by centrifugation at 7,500 × g for 5 min at 4°C and was resuspended in DEPC water. RNA at a final concentration of 1 μg was reverse transcribed into cDNA using a reverse transcriptase kit (Vazyme Biotech Co., Ltd). After mixing the relevant reagents of the kit, the samples were incubated at 50°C for 15 min and 85°C for 5 sec. qPCR was conducted on a Rotor-Gene Q-cycler (Qiagen, Inc.) using the Rotor-Gene SYBR Green qPCR Kit (Vazyme Biotech Co., Ltd.), according to the manufacturer's protocol. The thermocycling conditions were as follows: 95°C for 30 sec, followed by 40 cycles at 95°C for 10 sec and 60°C for 30 sec; and final steps at 95°C for 15 sec, 60°C for 1 min and 95°C for 15 sec. Human and mouse GAPDH were used as internal controls for parallel amplification. Relative changes in gene expression were quantified using the 2^−ΔΔCq method (34,35). All experiments were performed in triplicate. The specific primers (Sangon Biotech Co., Ltd.) used are listed in Table I. Among the specific primers, SRC was found in the 'Chemokine Signaling Pathway' and 'Cytokine-Cytokine Receptor Interaction' pathways when searching the downstream targets of PF4/CXCR3 in the Kyoto Encyclopedia of Genes and Genomes (KEGG) pathways (KEGG PATHWAY: map04062; https://www.genome.jp/entry/map04062 and KEGG PATHWAY: map04060; https://www.genome.jp/entry/map04060).

Total protein was isolated from THP-1 cells using RIPA buffer (Biosharp Life Sciences) containing protease and phosphatase inhibitors (Biosharp Life Sciences) on ice for 30 min. Subsequently, the cell fragments were centrifuged at 12,000 × g for 15 min at 4°C. The soluble protein component was mixed with 5X loading buffer (Biosharp Life Sciences), and the concentration of proteins was determined using the BCA protein assay kit (Beyotime Institute of Biotechnology). Equal amounts of protein (30 μg/lane) were subjected to SDS-PAGE on 10 or 12.5% gels, and were electroblotted onto PVDF membranes (MilliporeSigma). The membranes were blocked using 5% fat-free milk (Beyotime Institute of Biotechnology) for 2 h at room temperature and were then incubated at 4°C overnight with the following primary antibodies: Anti-CD62P (1:2,000; cat. no. 60322-1-Ig; Proteintech Group, Inc.), anti-inducible nitric oxide synthase (iNOS; 1:500; cat. no. 18985-1-AP; Proteintech Group, Inc.), anti-CD86 (1:500; cat. no. 26903-1-AP; Proteintech Group, Inc.), anti-β-actin (1:20,000; cat. no. 66009-1-Ig; Proteintech Group, Inc.), anti-Bcl-2 (1:500; cat. no. 66799-1-Ig; Proteintech Group, Inc.), anti-Bax (1:1,000; cat. no. ET1603-34; HUABIO), anti-caspase-3 (1:1,000; cat. no. 14220; Cell Signaling Technology, Inc.), anti-ERK1/2 (1:1,000; cat. no. 4695; Cell Signaling Technology, Inc.), anti-phosphorylated (p)-ERK1/2 (1:2,000; cat. no. 4370; Cell Signaling Technology, Inc.), anti-p65 (1:1,000; cat. no. 6956; Cell Signaling Technology, Inc.) and anti-p-p65 (1:1,000; cat. no. 3033; Cell Signaling Technology, Inc.). After incubation with horseradish peroxidase-conjugated secondary antibodies (1:5,000; cat. nos. SA00001-1 and SA00001-2; Proteintech Group, Inc.) for 1 h at room temperature, the membranes were incubated with enhanced chemiluminescence (ECL) FemtoLight Substrate (cat. no. SQ201L; Epizyme; Ipsen Pharma). Protein detection was performed using the automatic chemiluminescence image analysis system (Tanon 5200 Multi; Tanon Science and Technology Co., Ltd.). Protein expression was semi-quantified using ImageJ (version 1.5.3; National Institutes of Health).

Apoptosis was detected by flow cytometry. After the cells were treated with 2.5 μM AMG487 for 2 h, 100 ng/ml PF4 was added for 24 h and the harvested cells were washed three times with PBS. Apoptosis was detected by flow cytometry after staining with Annexin V-FITC and PI (Beyotime Institute of Biotechnology) for 30 min at room temperature. The percentage of apoptotic cells was calculated, and the data were analyzed using Novo Express Immunofluorescence Analysis(version 1.4.1; Agilent Technologies, Inc.). The apoptotic rate was determined using a NovoCyte 2000 Flow Cytometer (Agilent Technologies, Inc.) (36).

Mouse serum IL-1β and TNF-α levels were measured using an Automatic Biochemistry Analyzer (XPT; Siemens AG).

ROS levels in THP-1 cells treated with PF4 and AMG487 were measured according to the instructions of the ROS kit (Beyotime Institute of Biotechnology). PMA (100 ng/ml) was added to THP-1 cells for 24 h at 37°C to induce the differentiation of monocytes into macrophages. Subsequently, the samples were treated with AMG487 (2.5 μM) for 2 h at 37°C, after which PF4 (100 ng/ml) was added for a further 24 h at 37°C and the samples were collected. After washing with PBS, proportionally diluted DCFH-DA probes (in serum-free medium at a ratio of 1:1,000) were added to the cells. THP-1 cells were washed three times with PBS to completely remove the unbound DCFH-DA probe and were then fixed with 4% paraformaldehyde for 10 min at room temperature. After nuclear staining with DAPI, the intracellular ROS levels were assessed using ImageJ (version 1.5.3; National Institutes of Health) by detecting the fluorescence brightness of the DCFH-DA probe under FITC parameters. Imaging was performed using an inverted fluorescence microscope (DMi8; Leica Biosystems).

The levels of JC-1 in THP-1 cells were determined in accordance with the instructions provided in the JC-1 kit (Beyotime Institute of Biotechnology). PMA (100 ng/ml) was added to THP-1 cells for 24 h at 37°C to induce the differentiation of monocytes into macrophages. Subsequently, the samples were treated with AMG487 (2.5 μM) for 2 h at 37°C, after which PF4 (100 ng/ml) was added for a further 6 h at 37°C and the samples were collected. After washing with PBS, the JC-1 dye was added to the cells at a 1:200 dilution in JC-1 staining buffer and incubated at 37°C for 20 min. Following two washes of the THP-1 cells with JC-1 staining buffer to ensure the complete elimination of unbound JC-1 dye, the intracellular mitochondrial membrane potential levels were evaluated using a fluorescence microscope. This involved the detection of the fluorescence intensities of both red and green fluorescence. Imaging was performed using an inverted fluorescence microscope (DMi8; Leica Biosystems).

For human and mouse colon tissue sample sections, immunostaining was performed using standard procedures. Briefly, human and murine colon samples were sectioned after fixation with 4% paraformaldehyde for 24 h at room temperature and embedded in paraffin (Biosharp Life Sciences). Paraffin-embedded 5-μm sections were dewaxed using xylene and rehydrated using gradient alcohol, and antigen repair was performed by microwave heating with Tris-EDTA (pH 9.0) at 95°C for 20 min. The tissue was then blocked with 5% bovine serum albumin (cat. no. ST023; Beyotime Institute of Biotechnology) for 30 min at room temperature, after which the solution was removed and the sections were incubated overnight at 4°C with the corresponding primary antibodies. The sections were then stained with TSA fluorescent labeling reagent (cat. no. bry-0023-100; Shanghai Ruiyu Biotechnology Co., Ltd.) for 30 min at room temperature and DAPI working solution (cat. no. P0131; Beyotime Institute of Biotechnology) was added dropwise. Imaging was carried out using a laser scanning confocal microscope (TCS SP8; Leica Biosystems). The antibodies used in this experiment included: Anti-CD62P (1:200; cat. no. 60322-1-Ig; Proteintech Group, Inc.), iNOS (1:100; cat. no. 18985-1-AP; Proteintech Group, Inc.), CD206 (1:200; cat. no. 18704-1-AP; Proteintech Group, Inc.), PF4 (1:100; cat. no. 21157-1-AP; Proteintech Group, Inc.) and CXCR3 (1:100; cat. no. 26756-1-AP; Proteintech Group, Inc.).

Briefly, human and murine colon samples were sectioned after fixation with 4% paraformaldehyde for 24 h at room temperature and embedded in paraffin. Paraffin-embedded mouse colon sections (5 μm) were dewaxed using xylene and rehydrated using gradient alcohol. Sample permeabilization was performed through incubation with 20 μg/ml proteinase K for 20 min at room temperature. Endogenous peroxidase was inactivated via incubation with 3% H₂O₂ for 5 min at room temperature. The TUNEL reaction mixture (cat. no. Roche-11684795910; Sigma-Aldrich; Merck KGaA) was added to the sample and incubated for 1 h at 37°C. The DAPI working solution was added dropwise and imaging was carried out using a laser scanning confocal microscope (TCS SP8; Leica Biosystems).

Data are presented as the mean ± standard deviation (n≥3). Statistical analysis was performed using either unpaired, two-tailed Student's t-test, or one-way ANOVA followed by Bonferroni post hoc test using GraphPad Prism (version 9.0; Dotmatics). Categorical data, such as DAI and pathological scores in animal experiments, are presented as the median (range) and were analyzed using the non-parametric Kruskal-Wallis test followed by Dunn's post hoc test. P<0.05 was considered to indicate a statistically significant difference.

In present study, it was observed that, compared with in the normal control group, in the colon of patients with UC, bleeding at the colonic mucosa was shown to occur during active UC (Fig. 1A). This was also observed in the colon of DSS-induced mice, with more neovascularization and microvessels, suggesting that platelets enter the colon from blood vessels and exert their effects (Fig. 1A and B). The co-localization of platelets and macrophages was investigated by staining for a platelet activation marker (CD62P) and a macrophage M1 marker (iNOS); the findings revealed that the co-localization of platelets with macrophages was more pronounced in the colon of patients with UC compared with in healthy controls (Fig. 1C). This finding was confirmed by staining for PF4 and iNOS in the mouse colon to detect the colocalization of platelets with proinflammatory macrophages (Fig. 1D).

In vitro, platelets were extracted from healthy individuals and co-cultured with THP-1 human monocytes. Observations at the microscopic level revealed that platelets contributed to the transformation of monocytes into macrophages, THP-1 cells were transformed from suspended monocytes into adherent macrophages (Fig. 2A). PMA, a common drug that induces mononuclear-to-macrophage transformation, was used as a reference group to verify THP-1 adherence. To examine the effects of platelets on macrophages in an inflammatory environment, an in vitro model of an inflammatory environment induced by LPS was developed. RT-qPCR confirmed that platelets facilitated the release of the proinflammatory cytokines IL-1β, IL-6 and TNF-α from monocyte-derived macrophages. Furthermore, in an LPS-induced proinflammatory environment, the addition of CaCl₂ to platelets resulted in a more pronounced inflammatory response. By contrast, platelets had little impact on the anti-inflammatory cytokine IL-10 (Fig. 2B). After co-culturing THP-1 cells with platelets, RT-qPCR demonstrated a notable increase in the expression of PF4 as well as CXCR3 compared with THP-1 cells only (Fig. 2C). According to these results, platelets may serve a key role in facilitating the differentiation of monocytes into macrophages and exerting proinflammatory effects.

It was hypothesized that platelet activation could influence macrophages through the PF4/CXCR3 pathway, thereby affecting the progression of UC. Therefore, the CXCR3 inhibitor AMG487 was used during co-culture to validate its efficacy. As a result of the co-culture, the expression levels of CD62P, a platelet activation marker, and iNOS, a macrophage M1-type marker, were increased. In the presence of AMG487, iNOS expression was reduced, whereas the expression of CD62P was not affected (Fig. 3A). To better understand the role of this pathway, PF4 protein was added to THP-1 cells and the levels of inflammatory markers were measured. The results of RT-qPCR demonstrated that, as per the observations shown in Fig. 2B, the expression levels of inflammatory indicators, such as IL-1β, IL-6 and TNF-α, were elevated after the addition of PF4, whereas these expression levels were reduced following the addition of AMG487 (Fig. 3B).

According to the aforementioned findings, it was indicated that the addition of PF4 enhanced the M1 proinflammatory phenotype, as demonstrated by the M1 phenotypic markers CD86 and iNOS. Both CD86 and iNOS were upregulated after the addition of PF4 to THP-1 cells, whereas the inhibition of CXCR3 suppressed inflammation in macrophages (Fig. 3C). These results suggested that platelets interact with macrophages through the PF4/CXCR3 pathway, thereby promoting macrophage polarization towards a proinflammatory state.

ROS have been demonstrated to be associated with the development of inflammation. The levels of ROS were significantly elevated following the addition of PF4, whereas they were decreased following the addition of a CXCR3 inhibitor (Fig. 4A). Notably, the function of mitochondria can be impaired by excessive ROS; therefore, changes in the mitochondrial membrane potential were observed in macrophages following the addition of PF4 by JC-1 staining. When PF4 was added, the green fluorescence of the monomer was markedly increased, indicating a decrease in mitochondrial membrane potential, which AMG487 reversed (Fig. 4B). Reduced mitochondrial membrane potential can initiate the apoptosis signaling pathway; therefore, apoptosis was detected by Annexin-V FITC. FITC can be used to detect cells entering early apoptosis, PI detects cell necrosis, whereas cells positively stained for both FITC and PI are cells entering late apoptosis. FITC/PI staining was significantly increased following the addition of PF4, whereas this was reversed after the addition of AMG487 (Fig. 4C). To further clarify the regulatory factors of apoptosis, the key proteins involved in this process were detected. Compared with in the control group, the PF4 group exhibited significantly higher expression levels of the pro-apoptotic protein Bax, and lower expression levels of the anti-apoptotic protein Bcl-2, suggesting that PF4 increased apoptosis. In addition, caspase-3, which is a marker of irreversible apoptosis, was cleaved in response to PF4, whereas AMG487 reversed this effect (Fig. 4D).

The current study examined the regulatory function of PF4/CXCR3 by focusing on two established pathways associated with inflammatory processes: i) NF-κB and ii) MAPK. The protein expression levels of p-p65 and p-ERK were increased in response to PF4 and were decreased when CXCR3 was inhibited (Fig. 5A). To confirm this finding, the ERK inhibitor PD98059 and the p65 inhibitor SC75741 were used to treat the cells, and the inflammatory factors and macrophage M1 marker iNOS were subsequently detected, as well as another key gene downstream of CXCR3 and upstream of the two pathways, SRC. In the KEGG pathways 'Chemokine Signaling Pathway' and 'Cytokine-Cytokine Receptor Interaction', PF4 acts as a chemokine and activates downstream signals by binding to CXCR3. SRC may be involved in signal amplification and regulation downstream of CXCR3, and affect the activation and migration of immune cells. The results showed that the expression levels of iNOS, the M1 proinflammatory phenotypic marker, and ERK, p65 and SRC, the key genes of the two pathways, were decreased after the addition of the inhibitors, and the effect was equally efficacious when the two inhibitors were added together (Fig. 5B and C). Therefore, it was hypothesized that PF4/CXCR3 may regulate macrophage inflammation through the NF-κB and MAPK pathways.

The mice were treated with clopidogrel or AMG487 for 3 days prior to UC induction, followed by 7 days of free drinking of 2.5% DSS (Fig. 6A). During DSS administration, body weight was measured and fecal occult blood testing was performed daily, and the DAI score was calculated based on body weight change, fecal softness and occult blood conditions. Following the addition of DSS, the weight of mice decreased significantly, whereas the DAI score was significantly increased. Both the body weight and DAI of mice were significantly improved after administration of clopidogrel or AMG487 (Fig. 6B and C). The length of the colon is also a factor in determining UC. In the DSS group, the colon was significantly shortened, which was reversed in the clopidogrel and AMG487 groups (Fig. 6D). As a result of H&E staining, colon pathology was assessed and scored according to relevant inflammatory factors. The results indicated that the DSS group had the highest pathological score and the most severe disease, whereas clopidogrel and AMG487 effectively alleviated the condition (Fig. 6E).

As a result of fluorescent double labeling of colon macrophage M1 and M2 markers (iNOS/CD206), an increase in M1 proinflammatory expression and a decrease in M2 expression in the mouse colon was observed after DSS stimulation, which was ameliorated by clopidogrel and AMG487 (Fig. 7A). As demonstrated by peripheral blood measurement of inflammatory cytokines and colon RT-qPCR, inflammation was elevated in the DSS group, which was ameliorated in the clopidogrel and AMG487 groups (Fig. 7B and C). Additionally, double labeling of PF4 and CXCR3 was performed. In the DSS group, confocal staining of PF4 and CXCR3 was detected, indicating that UC was aggravated by PF4/CXCR3 signaling in mice. Among the treatments administered, clopidogrel could inhibit the activation of platelets and reduce the content of PF4, whereas AMG487 inhibited CXCR3 expression (Fig. 7D). This finding was also confirmed by PF4/CXCR3 double labeling on colon samples from patients with UC (Fig. 7E).

TUNEL assay was then performed to detect apoptosis in mouse colons, and it was observed that apoptosis was markedly enhanced in the DSS group, whereas it was reduced in the clopidogrel and AMG487 groups (Fig. 8A). A similar pattern of results was observed when tight junction proteins were used to measure the function of the colonic barrier. Zonula occludens 1, mucin 2 and occludin, which serve important roles in intestinal homeostasis, were stably expressed in the colon of the control group. Mice in the DSS group had severe colon tissue injury, and the expression levels of these genes were decreased; however, clopidogrel and AMG487 could alleviate colonic tissue damage and reverse these changes in expression levels (Fig. 8B). Thus, it was indicated that the PF4/CXCR3 pathway may be one of the key pathways in the progression of UC, which not only serves a key role in inflammatory progression, but also has a certain role in colonic apoptosis and intestinal barrier function.

The molecular mechanism through which platelets regulate macrophages through PF4/CXCR3 to enhance inflammation in UC was identified in the present study (Fig. 9). When UC develops, platelets bleed through vascular injury and enter the colon to activate platelets and release cytokines. PF4 released by platelets binds to the macrophage receptor CXCR3 in the colon, activating macrophages and polarizing them towards the proinflammatory phenotype through signaling pathways such as the MAPK and NF-κB pathways, serving a proinflammatory role and releasing proinflammatory factors. A variety of proinflammatory cells may be recruited into the colon by these inflammatory factors, causing inflammatory edema and affecting colon cell apoptosis. Under the influence of the PF4/CXCR3 pathway, macrophages are fully activated, ROS are released, mitochondrial function is disrupted and macrophage apoptosis is induced.