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Introduction

The spleen is the largest secondary lymphoid organ, and an extensive meshwork of nerve fibers is distributed throughout the splenic compartments including the splenic nodules, periarteriolar lymphoid sheath, marginal zones, trabeculae, white pulp and red pulp.^1,2 Even more striking, the nerve fibers are co-localized with various subsets of dendritic cells (DCs), macrophages (Mφ), and lymphocytes (ie B cells and T cells).^2,3 As early as 30 years ago, some studies demonstrated that sympathetic nerve terminals in the spleen secrete norepinephrine (NE) interacting with T cells in response to stimulation, while the content of splenic norepinephrine significantly decreases following chemical or surgical sympathectomy.^4,5 It was not until around 2000 that the cholinergic anti-inflammatory pathway (CAP) concept was formally proposed and extensively studied because of its vital function in modulating the mammalian immune response.^6–10 The CAP mechanism relies on the parasympathetic vagus nerve transmitting signals from the brain to the adrenergic splenic nerve that interacts with immune cells (Supplementary Figure 1). Munyaka et al¹¹ revealed that CAP is activated after central cholinergic activation by intracerebroventricular infusion of the M1 muscarinic acetylcholine (ACh) receptor agonist in mice but is suppressed by vagotomy or splenic neurectomy. Moreover, Ji et al¹² proved that central cholinergic activation induced by the acetylcholinesterase inhibitor galantamine or the muscarinic ACh receptor agonist relieves colitis in mice, which is inhibited after vagotomy, splenic neurectomy or splenectomy. These reports indicate that spleen is a vital target of the CAP.

In the past decade, many researchers applied physical methods to trigger the splenic neural-immune reflex for regulating the immune response to chronic inflammatory diseases.¹³ Inoue et al¹⁴ utilized a bipolar silver wire electrode stimulating the isolated left cervical vagus nerve to attenuate kidney ischemia-reperfusion injury (IRI), suggesting that vagus nerve stimulation (VNS)-mediated IRI treatment and systemic inflammation depend on α7 nicotinic acetylcholine receptors (α7nAChR)-positive splenocytes. Koopman et al^15,16 reported that stimulation of the CAP by VNS to treat rheumatoid arthritis (RA) in experimental models and clinical trials exerts a significant mitigation effect, whilst the symptoms aggravated following unilateral cervical vagotomy or α7nAChR knockout in mice. Besides, Meregnani et al¹⁷ demonstrated that the symptoms of colonic colitis in rats induced by trinitrobenzene sulphonic acid was significantly reduced after VNS performed with electrode cuffs for 5 days. Moreover, electroacupuncture intervention eliciting splenic efferent vagus nerve activity not only reduced tumor proliferation in breast tumor-bearing mice by alleviating inflammation and enhancing antitumor immunity,¹⁸ but also could alleviate the severity of chemotherapy-induced nausea and vomiting in patients with advanced cancer.^19,20 In addition, bioelectronic devices for VNS in clinical application have already been approved by the European Medicines Agency and the US Food and Drug Administration (FDA) for the treatment of drug-refractory epilepsy and depression.^13,21,22

Unsatisfactorily, nerve stimulation using permanently implanted electrodes, transcutaneous bioelectronic devices, or slightly injurious electroacupuncture was limited to stimulate large nerves or superficial nerves.²³ Peripheral organs, not just the spleen, are extensively traversed by peripheral nervous system, which makes it difficult to selectively stimulate subsets of axons that terminate in spleen and uniquely modulate the immune response to inflammatory diseases.²³ For example, cervical nerve stimulation usually activates the extensive efferent and afferent neural pathways that elicit neural responses from non-target organs.^{13,16,24–26} Although scientists are committed to developing more advanced, miniaturized and sophisticated electrode designs for implantation on specific nerves near the target organ,^27,28 invasive implantation easily causes damage to the delicate nerve fibers of the target organ and it is difficult to stimulate nerves throughout the organ. Thus, new nonintrusive strategies are imperative to trigger specific nerves, especially the splenic nerve, and mobilize the splenic immune function against diseases.

Notably, recent studies showed that noninvasive ultrasound successfully elicits peripheral nerve activation in the spleen to treat chronic inflammatory diseases (Supplementary Figure 1). Gigliotti JC and co-workers²⁹ used an ultrasound imaging probe to successfully irradiate the neural innervation in the spleen mitigating IRI in mice, whose mechanism is mainly attributable to the CAP activation.^6,9,16,24 Later, Cotero et al²³ reported that ultrasound modulating splenic nerve reduces endotoxin-induced cytokine production at a level comparable with traditional VNS performed with electrodes. Zachs et al³⁰ applied ultrasound stimulation targeting the spleen, which significantly reduces inflammation severity in RA mouse models. These studies provide ample evidence on the potential of precise splenic ultrasound stimulation to replace bioelectronic devices for the translation of peripheral neuromodulation-based therapies for inflammatory disease. However, the potential roles and underlying mechanisms of splenic ultrasound stimulation in cancer management have been rarely reported and superficially defined. Upon validation of its significant antitumor efficacy, splenic ultrasound stimulation would represent a groundbreaking clinical advancement by circumventing multiple limitations of conventional cancer therapies: surgical resection-associated recurrence and visceral injury compromising quality of life, chemotherapy-induced systemic toxicity, off-target risks of immunotherapy, and the technical complexities of personalized gene therapy.³¹

Further investigation is needed to establish whether ultrasound directly regulates the immunomodulatory function of the spleen by interfering with splenic immune cells and microenvironment. Ultrasound targeting the spleen is crucial in achieving conspicuously therapeutic effects, since the stimulation of other body locations or the lack of immune cells in mice is noneffective, which suggests the indispensability and importance of immune cells.^29,30 Cotero et al²³ and Zachs et al³⁰ concluded that ultrasound activating CAP indirectly exerts an influence on splenic immune cells or cytokines to reduce the severity of lipopolysaccharide acute inflammation and RA, but they mainly focused on the analysis of neuro-immune mechanism and did not perform a systematic research on the changes of splenic immune cells in response to ultrasonic stimulation. Generally, ultrasonic irradiation on cells directly irritates the expression of multiple key genes/signaling pathways or interferes with the secretion of cytokines to regulate cell proliferation, differentiation and migration, such as stimulating the expression of vascular endothelial growth factor in several cells including endothelial cells, neural cells and ischemic cells.^32–35 Therefore, we hypothesized that ultrasound could effectively immunomodulated on the splenic immune cells responding to cancer cell antigens in splenic microenvironment, and enhance the proliferation, activation, migration, and information dissemination of immune cells during cancer immunotherapy.

Accordingly, this study was designed to explore whether focused ultrasound precisely stimulating spleen (FUS sti. spleen) was effective in suppressing tumor proliferation, and the underlying mechanisms regulating splenic ultrasound stimulation in cancer management, including the activation of splenic nerve-related CAP, and splenic immune cells responding directly to ultrasonic capabilities. The therapeutic efficacy of FUS sti. spleen was first assessed on various tumors under specific parameters screened before. Furthermore, the splenic immunomodulation through FUS sti. spleen was evaluated based on the changes of splenic immune cell population and cytokine levels. Subsequently, nerve blockade, immune cell clearance and RNA sequencing were performed to identify the underlying mechanisms of splenic ultrasound stimulation in cancer management, such as splenic nerve-related CAP activation and FUS directly modulating immune cells, and the related molecular mechanism.

Methods

Cell Lines and Animal Models

Hepatocellular carcinoma (HCC) cell lines (H22 and hepa1-6) and 4T1 breast cancer cell line were purchased from the American Type Culture Collection (ATCC) and used for subsequent analysis. H22 cells were reproduced by ascites after intraperitoneal injection in C57black/6 mice (0.5–1.5*10^7 cells/mL; 200 µL per mouse). Hepa1-6 cells and 4T1 cells were cultured in Dulbecco’s modified Eagle’s medium (Cat. No. 11320033, Gibco, Invitrogen, Carlsbag, USA) supplemented with 10% fetal bovine serum (FBS; Cat. No. A5669701, Gibco, Invitrogen, Carlsbag, USA) and 1% penicillin-streptomycin solution (Cat. No. 15140122, Gibco, Invitrogen, Carlsbag, USA), and incubated at 37 °C under 5% CO₂ and 100% humidity.

Seven-week (weighted ~17 g) C57black/6 mice, purchased from the Beijing Animal Experiment Center (Chinese Academy of Sciences, Beijing, China), were housed at 24±2 °C under a 12-h light/dark cycle and acclimatized for at least 1 week before the experiments. All animals had access to sterilized food and water ad libitum. An amount of 0.2 mL H22 cancer cells (approximately 1–3*10^6 cells/mL) or 4T1 cancer cells (1–3*10^6 cells/mL) was hypodermically injected into the dorsal hindlimb region of the mouse to establish a subcutaneous tumor model. Regarding the in situ xenograft HCC models, an incision of approximately 1 cm in length was made at the upper end of the midabdominal line after the mouse was anesthetized with isohalothane and fixed on the anatomic stage. A total of 25 µL H22 cancer cells (approximately 0.5–1.5*10^7 cells/mL) or 50 µL hepa1-6 cancer cells (approximately 4–6*10^6 cells/mL) was injected into left hepatic lobe with an insulin syringe, and the incision was closed after applying pressure on the pinhole with a medical cotton swab for 3–4 minutes.

The tumor volume was calculated as follows: volume = 0.5*L*W^2, where L and W were the length and width of the tumor, respectively, measured by a caliper. The weight of spleen, tumor and mice was assessed by electronic scales. The splenic index was calculated by the ratio of spleen weight to mouse weight.

Ultrasonic Platform Set-up and Application for Spleen Stimulation

A diagram of the FUS system is shown in Supplementary Figure 2A. A function generator (Cat. No. 33120A, Agilent, Santa Clara, USA) produced a pulsed sinusoidal waveform triggering the power amplifier (Cat. No. AG1019, California, USA; or RPR-4000, RITEC Inc., Warwick, RI, USA) to drive a 1.04 MHz FUS transducer with a 100 mm aperture and 65 mm focus, whose focal region was ~1.4*1.4*8.6 mm³ (Supplementary Figure 2B and C). The acoustic pressure and spatial beam profile of the FUS transducer were measured using a hydrophone (Cat. No. HNR-1000, ONDA, Videlles, France). The FUS transducer was mounted on a XYZ motorized positioning stage to control ultrasonic duration, and the position of the FUS focus relative to the mice was adjusted under a B-mode imaging guidance with a 3.2 MHz phased array positioned at the center of the FUS transducer.

The mice were anesthetized with 2–3% inhaled isoflurane and placed on a manual translation stage equipped with heating pad. Then, a centrifuged coupling gel was immediately applied to the shaved skin, and the manual translation stage was adjusted to allow the spleen (or tumor) site to be in tight contact with the bottom of the water tank under the B-mode imaging guidance (Supplementary Figure 2D). The FUS transducer was moved across the whole spleen (or tumor) through the XYZ programmable logical controller (Supplementary Figure 2E).

Flow Cytometry

Single-cell suspensions from the spleen, peripheral blood, tumor, and para-carcinoma tissue were obtained after red blood cells were lysed using lysis buffer (Cat. No. 555899, BD Bioscience, USA) according to the manufacturer’s instructions. Cell suspensions were washed with sterilized PBS, and then incubated with the antibodies listed in Supplementary Table 1, and the scheme of antibody labeling for each immune cell is shown in Supplementary Table 2. Next, the immune cells were counted by flow cytometry (FCM) using CytoFLEX LX (Beckman Coulter Life Sciences, USA) after filtration through a 70-μm nylon cell strainer (Cat. No. 352350, Corning, USA). Data were analyzed using the FlowJo software (FlowJo 10, LLC, Ashland, OR).

Cytokine Measurement

Luminex Assay

Cytokine/chemokine quantification in plasma was performed by Luminex xMAP technology using a magnetic Luminex assay (R&D Systems, Minneapolis, MN, USA). The quantification was carried out using a Luminex^® 200 Flow Cytometry System (Cat. No. x-200, Thermo Fisher Scientific, MA, USA) and Milliplex Analyst software (Version 5.1, Merck Millipore, MA, USA).

ELISA

The concentration of NE, ACh, granzyme and perforin in plasma, respectively, detected by enzyme-linked immunosorbent assay (ELISA). The operation processes were carried out in strict accordance with the instruction manual from Mouse Perforin ELISA Kit (Cat. No. F30718-A, FANKEW, Shanghai, China), Mouse Granzyme-B ELISA Kit (Cat. No. F3214-A, FANKEW, Shanghai, China), Mouse ACh ELISA Kit (Cat. No. MU30072, Bioswamp, Wuhan, China) and Mouse NE ELISA Kit (Cat. No. MU30372, Bioswamp, Wuhan, China), respectively.

Histological and Immunohistochemical Staining

Hematoxylin eosin (HE) staining was applied to evaluate the pathological changes of the spleen after FUS irradiation. Transferase-mediated deoxyuridine triphosphatebiotin nick end labeling (TUNEL) staining was performed to observe splenic cell apoptosis through the colorimetric TUNEL Apoptosis Assay Kit (Cat. No. C1091, Beyotime, Shanghai, China). The primary antibodies of anti-Ki67 (Cat. No. RM9106S1, Thermo Fisher Scientific, MA, USA; dilution 1:200), anti-NK1.1 (Cat. No. 108759, BioLegend, California, USA), anti-CD8a (Cat. No. ab4055, Abcam, Cambridge, UK; dilution 1:1000), and anti-c-Fos (Cat. No. ab222699, Abcam, Cambridge, UK; dilution 1:2000) were used for immunohistochemical staining to detect the protein expression of cyclin-D1, Ki67, NK1.1, CD8a, and c-Fos respectively. At least 5–10 different regions in each section were randomly selected for image acquisition using a fluorescence microscopy (observer3, Carl Zeiss, Jena, Germany), and the positive area ratio was quantified using the Image Pro Plus 6.0 software (Media Cybernetics, CA, United States).

RNA Sequencing

Approximately 1*10⁷ CD8 T cells (CD45⁺ CD3⁺ CD8a⁺) and 5*10⁶ NK cells (CD45⁺ CD3⁻ NK1.1⁺) were sorted from the spleen using the Beckman Kurt MoFlo Astrios ultra high-speed flow cytometry sorting system (MoFlo Astrios EQ, Beckman Coulter, lnc, USA). Subsequently, general transcriptome sequencing was performed when the positive cell rate was greater than 90% detected by flow cytometry (CytoFLEX LX, Beckman Coulter Life Sciences, USA). The cDNA library construction and sequencing of all RNA samples were performed by Shanghai OE Biotech Co., LTD. Transcriptome sequencing was performed to screen differentially expressed genes (DEGs). Then, gene ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analysis were applied to annotate the function of the DEGs. The DEG screening criteria were as follows: P < 0.05 and differential fold change |logFC| ≥ 1.

Nerve Signal Blockade

Propranolol Hydrochloride Blocking CAP

Propranolol hydrochloride (Cat. No. abs44129405, Absin, Shanghai, China), a kind of adrenergic receptor antagonist, was intraperitoneally administered at a dose of 3 mg/kg to block CAP by directly binding to the adrenergic receptor at 1.5 hours before FUS exposure to spleen according to previous reports,^36–38 which ensured FUS sti. spleen at the optimal time point of CAP blockade.

Absolute Ethanol-Induced Splenic Nerve Block

Absolute ethanol was used for splenic nerve block according to previous studies.^39–44 Briefly, the mice were anesthetized with the isoflurane, and disinfection with iodophor was performed on the shaved skin. A 7–10 mm incision was made next to the left side of the spleen. The spleen was isolated with tweezers to keep it away from the peritoneal cavity for a clear exposure of the three main blood supply vascular trees. A wet cotton was used to protect the peritoneal cavity and other organs. Subsequently, absolute ethanol was repeatedly applied to those vascular trees with a cotton tip for 5–10 seconds each time, with an interval of five seconds, for a total of seven times to block the splenic nerve fibers. Excessive ethanol dripping from the cotton tip was carefully avoided, to avoid significant vasospasm, which could lead to permanent damage to blood vessels, consequently leading to spleen necrosis and organ complete absorption. As regards the sham-operated mice, the same entire surgical procedure was performed except for the repeated application of anhydrous ethanol that was replaced by normal saline (pH = 7.4).

In vivo Immune Cell Depletion

The experimental C57BL/6 mice were treated with an intravenous injection of Ultra-LEAF™ purified anti-mouse NK1.1 antibody (Cat. No. 108759, BioLegend, California, USA) or Ultra-LEAF™ purified anti-mouse CD8a antibody (Cat. No. 100763, BioLegend, California, USA) at a dose of 50 μg/mouse on day 0, 3, 7, 12 and 17 to deplete NK cell or CD8 T cell. As regards MDSC depletion, the Ultra-LEAF™ Purified anti-mouse Ly-6G antibody (Cat. No. 127650; BioLegend, California, USA) was intraperitoneally administered to C57BL/6 mice at a dose of 0.25 mg/mouse on day 0, 6, 12, and 18. The cell clearance efficacy was assessed by FCM.

Calcium Detection and in vitro Anticancer Experiment of FUS Stimulating Splenic CD8 T Cells or NK Cells Enhanced by Calcium

Von Kossa Staining

The splenic specimens were cut into 5 μm-thick sections and subjected to VON KOSSA Calcium Staining Kit (Cat. No. JM1519; HPBIO, Shanghai, China) for the histological visualization of calcium deposits (mineralization). Calcium deposition in splenic cells was assessed by fluorescence microscopy (Zeiss observer3), where mass deposits appeared black, while dispersed deposits appeared gray. The positive area ratio was quantified using the Image Pro Plus 6.0 software.

FCM Detection of Fluo-4 AM Labeled Cells

The splenic cell suspensions were washed twice with sterilized PBS, and then incubated with Fluo-4 AM dye (1 mL dye/10⁶ cells; Cat. No. S1061M, Beyotime, Shanghai, China) for 30 min at 37 °C according to the manufacturer’s instructions. Next, the fluorescent cells were monitored by FCM using the CytoFLEX LX (Ex/Em = 490/525 nm).

In vitro Experiments Verifying Calcium-Strengthened FUS Stimulating Splenic NK Cells Against Tumor

Approximately 1.2*10⁶ NK cells were sorted from the spleen using the Beckman Kurt MoFlo Astrios ultra high-speed flow cytometry sorting system, and then subjected to FUS stimulation with or without high calcium concentration (300 nM). Then, NK cells were co-cultured with pEGFP-C1 plasmid-transfected Hepa1-6 cancer cells at a ratio of 1:1, where GFP fluorescent protein was used to distinguish cancer cells from splenic cells. Transwell devices (Corning Incorporated, NY, USA) with 3-µm diameter holes were used to distinguish immune cells in the superstratum and cancer cells in the substratum. The cells were co-cultured for 48 h before GFP fluorescence observation with the Zeiss observer3, CCK-8 assay (Cat. No. ABS50003-500T, Univ, China), and crystal violet (CV) staining to assess cancer cell suppression and immune cell proliferation. The tumoral background was mimicked by adding ultrasound shattered cancer cell suspension to the NK cells solution during FUS stimulation.

Statistical Analysis

Statistical analysis was performed using the statistical product and service solutions software (SPSS, USA). Statistical significance was analyzed using Student’s t-test for parametric data and Mann–Whitney U-test for nonparametric comparisons, with Bonferroni correction (specially for multiple comparisons), and the results were expressed as mean ± SEM. A value of p < 0.05 was considered statistically significant (*p < 0.05, **p < 0.01, and ***p < 0.001).

Results

Screening and Optimization of Ultrasonic Parameters for Spleen Stimulation

In view of the different ultrasonic parameters and platforms applied in previous studies of splenic ultrasound stimulation, it is pseudoscientific to indiscriminately imitate their ultrasonic parameters or methods to this study for regulating splenic immunotherapy to anti-cancer. Based on the pre-experiment results, 11 processing groups were set-up: G1-G11 group (detailed in Table 1), for the screening and optimization of ultrasonic parameters, and FUS sti. spleen in subsequent experiments was performed once every other day starting from the 2nd day after cancer cell implantation, with a total experimental duration of 28 days (Figure 1A). At the experimental endpoint (day 29), animals underwent terminal anesthesia followed by comprehensive tissue harvesting for other experimental analyses (eg, HE and TUNEL staining). The degree of subcutaneous H22 tumor suppression and spleen injury (assessed by HE and TUNEL staining) was used as the decisive criterion to establish the appropriate ultrasonic parameters.

Table 1 Scheme of Different Ultrasonic Parameters for FUS Sti. Spleen

Figure 1 Inhibitory effect of FUS sti. spleen on subcutaneous H22 tumor under various ultrasonic parameters. (A) experimental flow diagram. (B) images of spleen and tumor in G1-G11 groups. (C) tumor growth curves. (D) tumor weight. (E) mice weight. (F) spleen weight. (G) spleen index. (H) HE and TUNEL staining of the spleen irradiated with FUS under different ultrasonic parameters; the red arrow indicates the area of the magnified image, and the green arrow indicates the TUNEL-positive cells. (I) analysis of TUNEL positive area. (n = 5; * p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001; mean ± SEM). G1-G11 group details are shown in Table 1.

The phenotypic results showed that FUS sti. spleen significantly suppressed the proliferation of subcutaneous H22 tumor, especially in the G4, G5, G8 and G11 group (Figure 1B–D). The weight of the mice subjected to FUS sti. spleen was lower than that of the non-FUS mice (Figure 1E), and FUS sti. spleen showed a tendency to decrease splenomegaly (Figure 1F and G). By comparison, the increase of the duty cycle or the FUS exposure time at 1.8 MPa did not improve the anti-tumor effect (G3, G6, G9 and G10 group; Figure 1B–D). However, the high peak negative pressure of 2.8 MPa induced a severe spleen injury, easily leading to the death of mice (G5 group; Figure 1B, H and I). In contrast, FUS sti. spleen effectively suppressed tumor proliferation without damaging the splenic cells at 2.3 MPa (Figure 1B–D, H and I). However, the antitumor effect was significantly reduced when the FUS exposure time of 20s was shortened to 10s (G4 and G7 group; Figure 1B–D). FUS damaged a few splenic cells once increasing the exposure time of 20s to 40s or the duty cycle of 1% to 10% (G4, G8 and G11 group; Figure 1B–D, H–I). The above splenic stimulation results demonstrated a clear “dose response” to ultrasound with a distinct power level required for the specific regulation of biophysical effects on splenic immune cells, which was similar to previous studies.^23,29,30 In conclusion, the ultrasonic parameters of the G4 group (2.3 MPa, 1%, 20s) were selected for subsequent experiments of splenic immunomodulation for cancer therapy.

FUS Universally Activated Spleen Immunological Function to Suppress the Proliferation of Various Tumors

FUS Activated Splenic Immune Cells to Suppress the Proliferation of the Xenograft HCC in situ

The mouse model of HCC in situ, which was better to resemble the situation of primary liver cancer, was used to assess the anticancer efficiency of FUS sti. spleen. The orthotopic HCC mice were subjected to FUS sti. spleen from day 0 after cancer cell implantation, and once every two days for a total experimental duration of 20 days (Figure 2A). At the experimental endpoint (day 21), animals underwent terminal anesthesia followed by comprehensive tissue harvesting for other experimental analyses (eg, FCM).

Figure 2 FUS sti. spleen suppressed the proliferation of the xenograft HCC in situ. (A) experimental flow diagram of FUS sti. spleen after HCC cancer cell implantation. (B–I) results of FUS sti. spleen suppressing the proliferation of the xenograft H22 HCC in situ. (B) images of spleen and xenograft H22 HCC in situ. (C and D) tumor volume and tumor weight, respectively. (E–G) liver weight, mice weight and spleen weight, respectively. (H) spleen index calculated by the ratio of spleen weight to mouse weight. (I) survival curves. S1, control group of xenograft H22 HCC in situ; S2, FUS sti. spleen started on day 0 after H22 cancer cell implantation. (J–Q) results of FUS sti. spleen suppressing the proliferation of the xenograft Hepa1-6 HCC in situ. (J) images of spleen and xenograft Hepa1-6 HCC in situ. (K and L), tumor volume and tumor weight, respectively. (M–O, liver weight, mice weight and spleen weight, respectively. (P) spleen index calculated by the ratio of spleen weight to mouse weight. (Q) survival curves. (n = 10; * p ≤ 0.05; ***p ≤ 0.001; mean ± SEM). L1, control group of xenograft Hepa1-6 HCC in situ; L2, FUS sti. spleen started on day 0 after Hepa1-6 cancer cell implantation.

Twenty orthotopic H22 HCC mice were randomly divided into two groups: the S1 group was used as the control group, and the S2 group was subjected to FUS sti. spleen. The results of visual observation, tumor volume and tumor weight demonstrated that FUS sti. spleen significantly suppressed the proliferation of xenograft H22 carcinoma in situ, with a tumor inhibition rate of up to ~70% as compared with the control group (S2 vs S1, p<0.001; Figure 2B–D). Although the mean value of liver weight (total weight of the tumor and liver parenchyma) showed a decreasing trend in the S2 group compared to the S1 group, no significant difference was observed between the two groups (Figure 2E). In addition, the weight of the mice in the S2 group was lower than that of the mice in the S1 group (p<0.001; Figure 2F). The spleen weight and spleen index showed a decreasing trend of splenomegaly in the S2 group as compared with the S1 group, but without statistically significant difference (Figure 2G and H). More importantly, the survival statistics proved that the orthotopic H22 HCC mice treated with FUS sti. spleen had a longer survival time than the control group (Figure 2I).

Sixteen orthotopic Hepa1-6 HCC mice were randomly divided into two groups: L1 group was performed as the control group, and L2 group was administered to FUS sti. spleen. The results of visual observation, tumor volume and tumor weight showed that FUS sti. spleen remarkablely inhibited the proliferation of xenograft Hepa1-6 carcinoma in situ, with an antitumor ratio of up to ~80% (L2 vs L1, p<0.001; Figure 2J and L). Additionally, the weight of the liver and mice in the L2 group was lower than that of the mice in the L1 group (p<0.05; Figure 2M and N). No significant difference in spleen weight and spleen index was observed, although there was a decreasing trend shown in the L2 group as compared with the L1 group (Figure 2O and P). It was noteworthy that the survival time of the mice in the L2 group was significantly longer than that of the mice in the L1 group (Figure 2Q).

Thereafter, FCM results revealed that the proportion of tumor suppressor related immune cells in the spleen, blood, tumor and para-carcinoma tissue was significantly increased in the S2 group as compared with the S1 group, such as Th2 cells, NK cells, CD8 T cells, Mφ (Mφ2) and DC1 in the spleen; B cells, NK cells, CD8 T cells, Mφ1 and DC1 in the blood; CD4 T cells, NK cells, CD8 T cells, Mφ (Mφ1) and DC1 in the tumors; Th1 cells, NK cells, CD8 T cells and Mφ (Mφ1) in the para-carcinoma tissue (Table 2, Supplementary Figures 3–6). Moreover, the proportion of negative immune cells was significantly reduced, such as Treg cells and MDSCs (M-MDSCs and PMN-MDSCs) in both spleen and tumor; Th17, PMN-MDSCs in the blood; and PMN-MDSCs in the para-carcinoma tissue (Table 2, Supplementary Figures 3–6).

Table 2. Statistical Significance of FCM Results of Immune Cells in the Spleen, Blood, Tumor and Para-Carcinoma Tissue From Orthotopic HCC Mice Subjected to FUS Sti. Spleen or Non-Treated Mice

As regards to orthotopic Hepa1-6 HCC mice, FCM results showed that the number of positive anticancer immune cells in the spleen, blood, tumor and para-carcinoma tissue was significantly increased in the L2 group than in the L1 group, such as Th2 cells, NK cells, CD8 T cells, Mφ and DCs in the spleen; CD4 T cells, Th2 cells, NK cells, CD8 T cells, Mφ1, Mφ2 and DCs in the blood; NK cells, CD8 T cells, Mφ and DCs in the tumor; and NK cells, CD8 T cells and Mφ (Mφ1 and Mφ2) in the para-carcinoma tissue (Table 2, Supplementary Figures 7–10). The number of negative immune cells was significantly reduced, such as Treg cells and PMN-MDSCs in the spleen; Th17 and MDSCs in the blood; Th17 and PMN-MDSCs in the tumor; and PMN-MDSC in the para-carcinoma tissue (Table 2, Supplementary Figures 7–10).

These results fully demonstrated the significant effect of FUS on splenic immunomodulation for cancer immunotherapy. Particularly, FUS sti. spleen remarkably increased the proportion of NK cells and CD8 T cells in the spleen, blood, tumor and para-carcinoma tissue, followed by Mφ and DCs, whilst it reduced the number of PMN-MDSCs in all tissues with one accord, and then Treg cells.

Additionally, we assessed the therapeutic efficacy of FUS sti. spleen on subcutaneous 4T1 breast tumor, and further quantitatively evaluated the alterations in immune cell proportions following FUS intervention. Obviously, FUS sti. spleen demonstrated potent antitumor efficacy, achieving 70% tumor growth inhibition and significantly prolonging survival. FCM revealed that FUS elicited significant immunomodulatory effects, particularly enhancing NK cell and CD8+ T cell populations while suppressing immunosuppressive cells. The complete results are available in Supplementary File 1.

FUS Sti. Spleen Altered the Cytokine Levels

In addition to the modulation of cellular immunity by FUS sti. spleen, humoral immunity may also be significantly regulated, which represents one dominant factor in the anticancer process during FUS irradiation. Herein, we selectively detected the cytokine levels of GM-CSF, TNF-alpha, IL-12, CCL2, IL-1beta, IL-2, IL-4, IL-6, IL-10, IL-13, IL-17, IFN-gamma, CXCL10, M-CSF, IL-1alpha, CCL4, CXCL12, IL-27, perforin and granzyme in the peripheral blood using the Luminex xMAP technology. Interestingly, the concentration levels of TNF-alpha, IFN-γ, perforin and granzyme were significantly increased in the FUS sti. spleen group as compared with the control group in both H22 and Hepa1-6 HCC in situ models (Figure 3A–C), which indicated the activation of cytotoxic CD8 T cells and NK cells against tumor. Usually, the naive CD8 T cells in the immune process undergo activation and clone expansion, which in turn produce the effector cytokine TNF-alpha, IFN-gamma, perforin and granzyme.^45,46 In addition, the secretion level of IFN-gamma, perforin and granzyme specifically indicated the activation of NK cells to fight cancer.⁴⁷

Figure 3 Cytokine detection and bioinformatics analysis. (A and B), heatmap of cytokine levels in orthotopic H22 and Hepa1-6 HCC mice subjected to FUS sti. spleen. (n = 5; * p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001; mean ± SEM). (C) Venn diagram of up- and down-regulated cytokines in both orthotopic H22 and Hepa1-6 HCC mice subjected to FUS sti. spleen. (D) protein-protein network analysis of cytokines and target genes based on STRING database. (E) KEGG pathway enrichment analysis of cytokines and target genes.

The concentration levels of the pro-inflammatory chemokines CCL2, CCL4 and CXCL10 were significantly elevated in the FUS sti. spleen group of both orthotopic H22 and Hepa1-6 HCC models compared with those in the control group (Figure 3A–C). These pro-inflammatory chemokines promote immune cells especially effector T cells successfully migrating into metastatic tumor sites to perform an anti-tumor activity.^48–50 CXCL12 is a homeostatic chemokine involved in physiological processes like embryogenesis, neurogenesis, cardiogenesis, hematopoiesis, leukocyte homing, and angiogenesis, by inducing the migration and activation of hematopoietic progenitor and stem cells, endothelial cells, and most leukocytes.^51,52 In this study, the CXCL12 level was significantly increased in the FUS sti. spleen group (Figure 3A–C). As a final level of activity regulation, CXCL12 synergizes with CXCL8 but also with CXCL9, CXCL10, CXCL11, and multiple CC chemokines to attract B- and T-lymphocytes, DCs, monocytes, and CD34⁺ progenitor cells.⁵³ Moreover, CXCL12 binding to its receptor CXCR4 mediates the biological behavior of tumor cells such as movement, chemotaxis, adhesion, secretion (such as MMPs and VEGF), angiogenesis, growth, and proliferation by activating various signaling pathways.^54,55

Besides, GM-CSF and M-CSF are 2 important cytokines that regulate macrophage population and function. Many experimental loads of evidence suggest that the aberrant expression of GM-CSF and its receptors are present in many cancers such as glioblastoma, small cell carcinoma, skin carcinoma, meningiomas, colon cancer, lung cancer, and particularly HCC, promoting cancer progression by regulating the tumor microenvironment involving Mφ and MDSCs, and promoting epithelial to mesenchymal transition, angiogenesis, and expression of immune checkpoint molecules.^56–59 The concentration level of GM-CSF in this study was significantly decreased as both orthotopic H22 and Hepa1-6 HCC mouse models underwent FUS sti. spleen (Figure 3A–C). In addition, the M-CSF level was increased after orthotopic HCC mice were subjected to FUS sti. spleen (Figure 3A–C), which regulated the survival, proliferation and differentiation of the monocyte-macrophage lineage from progenitors to mature cells and activated several important functions of mature tissular Mφ.^60,61

The proinflammatory cytokine IL-1beta showed an association with tumor progression,⁶² but interestingly, although the inhibition of IL-1beta signaling has potent anti-metastatic effects, the inhibition of the activity of this cytokine has contradictory effects on primary tumors, sometimes reducing but often promoting their growth.⁶³ Moreover, anti-cancer treatments are able to promote IL-1beta production by cancer or immune cells, with opposite effects on cancer progression,⁶⁴ which resembles the tumor-suppressing effect of FUS sti. spleen. However, beyond that, the cytokine levels of IL-1alpha, IL-17 and IL-27 showed no significant differences in consistency as orthotopic H22 and Hepa1-6 HCC mice subjected to FUS sti. spleen (Figure 3A–C). In addition, IL-2 is a pleiotropic cytokine required for both effector lymphocyte proliferation/differentiation and regulatory T cell expansion/survival,⁶⁵ which was significantly secreted as spleen responded to FUS stimulation (Figure 3A–C). Additionally, IL-4 mainly secreted by T cells and NK cells induces Mφ2 activation, and regulates the expression of IL-6, IL-10, IL-12 and IL-13.⁶⁶ Some studies also reported that polarized Th1 cells produced IL-2, IL-12 and IFN-gamma, and polarized Th2 cells secreted IL-4, IL-6, IL-10 and IL-13 that promoted tumor proliferation⁶⁷ (Figure 3A–C). In this study, IL-6 level was increased and IL-13 level was decreased when the orthotopic HCC mice were subjected to FUS sti. spleen (Figure 3A–C). Nevertheless, the cytokine levels of IL-4, IL-10 and IL-12 were altered between orthotopic H22 and Hepa1-6 HCC mice models subjected to FUS sti. spleen (Figure 3A–C).

Protein–protein networks of these cytokines and target genes, and KEGG pathway enrichment analysis showed that FUS stimulating splenic immune cells to regulate cytokines against tumor was associated with several signaling pathways, especially JAK-STAT signaling pathway and PI3K-AKT signaling pathway (Figure 3D and E). It was evident that these cytokines characterized the tumor cytotoxicity of NK cells and CD8 T cells, mainly through cytokine-cytokine receptor interaction pathway to obtain an anticancer effect (Figure 3D and E).

FUS Directly Activated Splenic Immune Cells for Cancer Immunotherapy

FUS Promoted Splenic Positive Immune Cell Proliferation and Activation to Combat Tumor

According to previous studies, splenic ultrasound stimulation activated CAP to inhibit arthritis, colitis, hyperglycemia and other inflammatory diseases, but they paid more attention to the modulation of inflammatory factors such as TNF-α, IL-6 and IL-10, as well as CAP validation, but not to the changes of immune cells.^23,68,69 In this study, FUS sti spleen significantly altered the proportion of immune cells in the spleen, blood, and tumor, especially NK cells and CD8 T cells that increased uniformly in various tumor models (Table 2, Supplementary File 1), thus arousing our special attention. Inexplicably, the previous studies based on inflammatory disease models (eg, pneumonia, myocarditis, and colitis) proved that splenic ultrasound stimulation activates CAP to reduce Mφ and inhibits the secretion of pro-inflammatory cytokines (eg TNF-α), but in this study based on tumor models, FUS sti. spleen significantly increased Mφ and promoted TNF-α secretion. Therefore, our speculation was that FUS directly stimulated splenic immune cells stressing tumor signals, which promoted the proliferation and activation of positive immune cells (ie NK cells and CD8⁺ T cells) to suppress cancer cell proliferation. To further confirm this hypothesis, splenic nerve blockade was performed with propranolol hydrochloride targeting CAP to explore the impact of CAP deficiency on tumor suppression during FUS sti. spleen. Subsequently, NK cells and CD8 T cells were depleted with Ultra-LEAF™ purified anti-mouse NK-1.1 antibody and Ultra-LEAF™ purified anti-mouse CD8a antibody, respectively, to further explore their role and importance in the tumor-suppressing process of FUS sti. spleen. Forty-eight orthotopic H22 HCC mice were randomly divided into eight groups: the Q1 group was used as the control group; the Q2 group was subjected to FUS sti. spleen; the Q3 group was treated with an intraperitoneal injection of propranolol hydrochloride to block CAP; the Q4 group was subjected to FUS sti. spleen after an intraperitoneal injection of propranolol hydrochloride; the Q5 group was subjected to NK cell clearance; the Q6 group was subjected to FUS sti. spleen after NK cell depletion; the Q7 group was subjected to CD8 T cell clearance; the Q8 group was subjected to FUS sti. spleen after CD8 T cell depletion. The operation of FUS sti. spleen and experimental timeline were identical to those described in Figure 2A.

The therapeutic results once again demonstrated the significant tumor suppression of FUS sti. spleen (Q2 vs Q1, p < 0.001; Figure 4A–C). It is noteworthy that specifically CAP blockage did not significantly attenuate the therapeutic efficacy of FUS sti. spleen on the tumor (Q4 vs Q3, p < 0.001; Figure 4A–C), although the anti-tumor rate was slightly reduced in the Q4 group as compared to the Q2 group. It confirmed our previous speculation that CAP did not play a dominant role in the process of FUS sti. spleen for tumor suppression. We supposed that cancer is different from inflammatory diseases such as arthritis, colitis, and pneumonia, and FUS sti spleen to inhibit inflammatory diseases mainly relied on CAP affecting humoral immunity, while it suppressed tumor proliferation mainly depending on the direct modulation of cellular immunity.

Figure 4 Therapeutic effect of FUS sti. spleen on xenograft H22 carcinoma in situ after CAP blockage, NK cells depletion and CD8 T cell clearance. (A) images of spleen and orthotopic H22 tumor. (B and C) tumor volume and tumor weight, respectively. (D–F) liver weight, mice weight and spleen weight, respectively. G, splenic index calculated by the ratio of spleen weight to mouse weight. (H–K) concentration of NE, ACh, granzyme and perforin, respectively, detected by ELISA. (n = 6; * p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001; mean ± SEM). Q1, control group of xenograft H22 HCC in situ; Q2, FUS sti. spleen on day 2 after H22 cancer cell implantation; Q3, injection of propranolol hydrochloride to block CAP; Q4, FUS sti. spleen after injection of propranolol hydrochloride; Q5, NK cell clearance; Q6, FUS sti. spleen after NK cell depletion; Q7, CD8 T cell clearance; Q8, FUS sti. spleen after CD8 T cell depletion.

Furthermore, the tumor proliferation was significantly promoted after NK cell depletion (Q5 vs Q1, p < 0.001; Figure 4A–C). Interestingly, FUS sti. spleen did not show any tumor suppressive effect after depleting NK cells (Q6 vs Q5; Figure 4A–C). Moreover, CD8 T cell clearance promoted tumor proliferation (Q7 vs Q1, p<0.05; Figure 4A–C), but as opposite as NK cell clearance, did not reduce the therapeutic efficacy of FUS sti. spleen on tumor (Q8 vs Q2; Figure 4A–C). These results proved that NK cells were much more indispensable and important in cancer suppression in response to FUS stimulation than CD8 T cells. The statistical results of liver weight, mice weight, spleen weight and spleen index also showed that NK cell depletion had a significant effect on tumor proliferation and splenomegaly, and elucidated the attenuation of FUS sti. spleen on tumor suppression and splenomegaly (Figure 4D–G).

The concentration of NE and ACh in the plasma was significantly increased after FUS sti. spleen (Q2 vs Q1, Q6 vs Q5, and Q8 vs Q7, p < 0.05; Figure 4H and I), which indicated that FUS sti. spleen indeed activated splenic CAP. In addition, propranolol hydrochloride significantly blocked the binding of NE to adrenergic receptors, that is, effectively blocking CAP, thus increasing the NE content (Q3 vs Q1, p<0.01; Figure 4H) but no significant changes in ACh (Q3 vs Q1; Figure 4I). Therefore, NE content increased after FUS sti. spleen (Q4 vs Q3; Figure 4H), while ACh did not significantly change (Q4 vs Q3; Figure 4I). Nevertheless, it is important to note that the results of granzyme and perforin detection by ELISA further confirmed the significance of splenic NK cells activation and their secreting cytotoxic molecules in response to FUS stimulation for an anti-cancer effect, but not affected by CAP (Figure 4J and K).

FCM results proved that Ultra-LEAF™ purified anti-mouse NK-1.1 antibody and Ultra-LEAF™ purified anti-mouse CD8a antibody effectively eliminated NK cells and CD8 T cells in the body, respectively (Q5/6 and Q7/8 vs Q1/2; Tables 3–6, Supplementary Figures 11–14). In the case of CAP blockage and NK or CD8 T cell clearance, the proportion of Mφ, DCs and MDSCs in the body was also affected, especially the number of Mφ and DCs that was generally reduced after depleting NK cells and CD8 T cells (Q3/4, Q5/6 and Q7/8 vs Q1/2; Tables 3–6, Supplementary Figures 11–14). However, FUS sti. spleen consistently increased the number of NK cells and CD8 T cells in the body regardless of CAP blockade and NK or CD8 T cell clearance, followed by Mφ1 and DC1 (Q4 vs Q3, Q6 vs Q5, and Q8 vs Q7; Tables 3–6, Supplementary Figures 11–14), while the variation tendency of other immune cells (including PMN-MDSCs) was not consistent in the spleen, blood, tumor and para-carcinoma tissue (Tables 3–6, Supplementary Figures 11–14). Clearly, changes in the proportion of immune cells (especially NK cells) with or without modulation by splenic ultrasound stimulation were closely related to the anti-tumor effect.

Table 3. FCM Results of Immune Cells in the Spleen

Table 4. FCM Results of Immune Cells in the Blood

Table 5. FCM Results of Immune Cells in the Tumor

Table 6. FCM Results of Immune Cells in Para-Carcinoma Tissue

The results of the above CAP blockage experiments demonstrated that CAP did not play a dominant role in tumor suppression by FUS sti. spleen; however, it could not exclude the impact of splenic nerve or other nerve signals. Thereafter, we applied absolute ethanol to denervate the splenic nerve to explore the role of the splenic nerve on spleen immunomodulation and anti-cancer effect by FUS sti. spleen. Moreover, the spleen was removed to exclude the possibility of immune regulation and tumor suppression by FUS irradiation on non-splenic organs or tissues. Thus, forty-two orthotopic H22 HCC mice were randomly divided into six groups: H1 group was used as the control group; H2 group was subjected to FUS sti. spleen; H3 group was subjected to splenic nerve denervation by absolute ethanol; H4 group was subjected to FUS sti. spleen after splenic nerve denervation; H5 group was subjected to the removal of the spleen; and H6 group was exposed to FUS on spleen position after splenectomy.

The results showed that splenic nerve denervation promoted tumor proliferation to some extent, although not significant (H3 vs H1, p = 0.3430 in tumor volume and p = 0.1590 in tumor weight), suggesting that the splenic nerve had a certain role in the anti-tumor effect. However, more importantly, splenic nerve denervation did not attenuate the tumor suppression efficacy of FUS sti. spleen, which was up to ~75% (H4 vs H3, p<0.001; Figure 5A–C) that was superior to or consistent with the tumor inhibition rate of FUS sti. spleen without splenic nerve denervation (~70%, H2 vs H1, p<0.001; Figure 5A–C). It further ruled out the indispensable role of splenic nerve on tumor suppression by FUS sti. spleen, which was also direct evidence that FUS directly stimulated splenic immune cells to regulate immunity against cancer.

Figure 5 Therapeutic effect of FUS sti. spleen on tumor suppression after splenic nerve denervation and splenectomy. (A) images of spleen and orthotopic H22 tumor. (B and C) tumor volume and tumor weight, respectively. (D–F) liver weight, mice weight and spleen weight respectively. (G) splenic index calculated by the ratio of spleen weight to mouse weight. (n = 7; * p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001; mean ± SEM). (H and I) concentration of NE and ACh, respectively, detected by ELISA. (J) c-Fos immunohistochemical staining and statistical result of positive area in section; the red arrows indicate the c-Fos-positive cells. (K and L) concentration of granzyme and perforin, respectively, detected by ELISA. (n = 5; * p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001; mean ± SEM). H1, control group of xenograft H22 HCC in situ; H2, FUS sti. spleen on day 2 after H22 cancer cell implantation; H3, splenic nerve denervation by absolute ethanol; H4, FUS sti. spleen after splenic nerve denervation; H5, removal of the spleen; H6, FUS stimulated the splenic position after splenectomy.

Liver weight and mice weight also indirectly reflected the anti-cancer effect of FUS sti. spleen after splenic nerve denervation (Figure 5D and E). However, it must be noted that spleen weight and spleen index clearly indicated that splenic nerve denervation dramatically promoted tumor-induced splenomegaly (H3 vs H1, p < 0.01; Figure 5F and G), while FUS sti. spleen significantly reduced splenomegaly (H4 vs H3, p < 0.01; Figure 5F and G). The comprehensive analysis of the changes in splenic characteristics and tumor proliferation showed that the splenic nerve played an important role in the anti-cancer effect of the spleen under the natural state but not the splenic ultrasound stimulation.

ELISA results of NE and ACh showed that splenic neurons were not activated by FUS under splenic nerve denervation; thus, there was no significant change in the concentration of CAP-related NE and Ach (H4 vs H3; Figure 5H and I). The c-Fos immunohistochemical staining also proved that few splenic neurons responded to FUS stimulation after splenic nerve denervation (Figure 5J). However, splenic nerve denervation did not affect the response of splenic immune cells to FUS stimulation, especially the NK cells that secreted significantly higher levels of perforin and granzyme (H4 vs H3, p<0.01; Figure 5K and L). Particularly, after splenectomy, FUS irradiation in splenic position did not show any anti-tumor effect (H6 vs H5; Figure 5A–E, K-L), which demonstrated that the spleen irradiated with FUS was the only cause of the anti-cancer effect, and directly showed that the integrity of spleen was critical during FUS sti. spleen for tumor suppression. In summary, comprehensive analysis of splenectomy and splenic nerve denervation experiments clearly confirmed that FUS activated splenic anticancer immune function through the direct FUS stimulation of immune cells, thus further suppressing tumor proliferation.

FCM results proved that splenic nerve denervation did not affect cell proportion (H3 vs H1; Supplementary Figures 15–18), particularly not weaken the modulation properties of FUS sti. spleen on the increase of the positive immune cells (especially NK cells, CD8 T cells, Mφ1 and DC1) in the body, and decrease of the negative immune cells PMN-MDSCs (H4 vs H2/3; Supplementary Figures 15–18). It fully demonstrated that FUS directly stimulated the splenic immune cells to promote their proliferation and activated the anti-cancer capability. After the removal of the spleen, the proportion of most immune cells in the peripheral blood was significantly decreased (H5/6 vs H1/2; Supplementary Figure 16), which indicated the importance of spleen for the body’s immune system. However, after splenectomy, the spleen location irradiated by FUS did not show any impact on the number of immune cells (H6 vs H5; Supplementary Figures 16–18), which demonstrated that the spleen integrity or the presence of immune cells was essential for responding to FUS stimulation in tumor immunotherapy.

Association of Splenic Positive and Negative Immune Cells Responding to FUS Irradiation

Our previous study demonstrated that the proportion of PMN-MDSCs in the spleen and blood from tumor-bearing mice was significantly increased, which suppressed NK cells and CD8 T cells.⁷⁰ After FUS sti. spleen, the proportion of positive immune cells such as NK cells and CD8 T cells in the body was significantly increased, while the proportion of negative immune cells such as PMN-MDSCs was significantly decreased (Table 2, Supplementary File 1). Why did FUS, as a non-specific physical method, cause the number of positive and negative immune cells to change in two directions? The depletion of NK cells and CD8 T cells did not show consistent changes of PMN-MDSCs in the spleen and blood (Q5/6 and Q7/8 vs Q1/2; Tables 3 and 4); thus we proposed to reversely demonstrate the association of NK cells and CD8 T cells to PMN-MDSCs by depleting PMN-MDSCs. Forty orthotopic H22 HCC mice were randomly divided into four groups: R1 group was used as the control group; R2 group was subjected to FUS sti. spleen; R3 group was subjected to PMN-MDSCs depletion; and R4 group was subjected to FUS sti. spleen after PMN-MDSCs clearance.

PMN-MDSCs depletion inhibited tumor proliferation to a certain extent (R3 vs R1; Supplementary Figure 19A–D) and the subsequent FUS sti. spleen further enhanced tumor suppression (R4 vs R3; Supplementary Figure 19A–D). However, the tumor suppressive effect of FUS sti. spleen after PMN-MDSCs clearance was slightly weaker than that without PMN-MDSCs depletion (R4 vs R2; Supplementary Figure 19A–C). Mice weight did not clearly show a significant difference among the four groups (Supplementary Figure 19E), but spleen weight and spleen index showed a decreased trend in the R4 group as compared to the R3 group (Supplementary Figure 19Fand G).

FCM results showed that Ultra-LEAF™ purified anti-mouse Ly-6G antibody effectively eliminated PMN-MDSCs from the spleen, blood, tumor, and para-carcinoma tissue (R3 vs R1; Supplementary Figures 19H–K and 20–23). Interestingly, it significantly increased the proportion of Mφ, especially Mφ2, in the spleen and blood (R3 vs R1; Supplementary Figures 19H and I, 20 and 21). Moreover, PMN-MDSC depletion significantly increased NK cells and CD8 T cells only in the blood (R3 vs R1; Supplementary Figures 19I and 21). These results explained the reason that PMN-MDSCs clearance suppressed tumor proliferation to a certain extent (Supplementary Figure 19A–D). Even more important, PMN-MDSC depletion in spleen did not promote the proliferation of splenic NK cells and CD8 T cells (R3 vs R1; Supplementary Figures 19H and 20). Combined with the observation that the depletion of NK cells and CD8 T cells in the spleen did not increase the proportion of splenic PMN-MDSCs (Q5 and Q7 vs Q1; Table 3), it proved that there was no direct interaction between PMN-MDSCs and NK cells or CD8 T cells. Accordingly, it is reasonable to deduce that FUS sti. spleen increased the number of NK cells and CD8 T cells while the decrease of PMN-MDSCs was not directly correlated. PMN-MDSC decrease may be attributed to the increase of other immune cells in the spleen such as Mφ, especially Mφ2 mentioned above, or even to the rise of total tumor suppressor related immune cells in the spleen and blood. Of course, this mechanism has yet to be explored in the light of more direct and conclusive evidence.

FUS Regulating Calcium-Related Signaling Pathways to Modulate Splenic Immunity

The above results revealed that splenic NK cells and CD8 T cells were critical for suppressing tumor growth in response to FUS stimulation. Thereafter, RNA sequencing was performed to verify their changes at the molecular level and investigate the biological mechanism underlying this phenomenon.

According to the differential gene expression clustering heatmap and volcano plot, 655 upregulated genes and 344 downregulated genes were found in NK cells (Supplementary Figure 24A–C), as well as 1869 upregulated genes and 877 downregulated genes in CD8 T cells (Supplementary Figure 24A–C) in the FUS sti. spleen group compared with the control group (Supplementary Figure 24D–F). Both GO and KEGG enrichment analysis showed that the DEGs were significantly enriched in pathways of cell adhesion, cell migration, immune process (ie ECM-receptor interaction, antigen processing and presentation, and intestinal immune network for IgA production) and signal transduction (ie PI3K-AKT signaling pathway, Rap1 signaling pathway, Hippo signaling pathway and calcium signaling pathway) in the FUS sti. spleen group compared with the control group (Supplementary Figure 24G–N).

Further analysis revealed that genes related to NK cell immune function, especially activation, proliferation, and motility, were significantly up-regulated after FUS sti. spleen (Supplementary Figure 25A–C). Genes involved in calcium-related signaling pathways were also significantly highly expressed (Supplementary Figure 25D). According to GO functional annotation analysis, these upregulated genes were not only involved in the positive regulation of cell proliferation, migration and kinase activity, but also mainly involved in integrin-mediated signaling pathway and epidermal growth factor receptor signaling pathway (Supplementary Figure 26A). Moreover, MHC class II protein complex was suppressed (Supplementary Figure 26B), which promoted NK cell activation in mice as reported by Li et al.⁷¹ Particularly, KEGG enrichment analysis of these upregulated genes showed the upregulation of PI3K-Akt signaling pathway, Hippo signaling pathway, ErbB signaling pathway, and Ras signaling pathway (Supplementary Figure 26C), which were closely associated with cell proliferation, differentiation and activation, especially under ultrasound stimulation.^72–74 Downregulated genes were significantly enriched in the intestinal immune network for IgA production (Supplementary Figure 26D), which might be beneficial to tumor suppression as previous studies reported that IgA inhibits NK activity of NK cell-enriched lymphoid cells and gamma-interferon-treated effector cells.^75,76

More importantly, protein-protein networks revealed that among these significantly different signaling pathways, it is likely that the calcium signaling pathway played a central role and affected the expression of other signals to promote NK cell proliferation, migration and activation (Supplementary Figure 26E). Furthermore, GSEA analysis confirmed that FUS stimulation significantly regulated various signaling pathways of NK cells, such as up-regulating ECM-receptor interaction, PI3K-AKT signaling pathway, Hippo signaling pathway and calcium signaling pathways (Supplementary Figure 26F).

Additionally, the genes related to CD8 T cell activation, proliferation, and motility were significantly up-regulated in the FUS sti. spleen group compared to the control group (Supplementary Figure 27E–G). Genes involved in calcium-related signaling pathways were also highly expressed (Supplementary Figure 27H). GO and KEGG enrichment and network analysis of the differential genes showed significant enrichment of pathways in signal transduction (ie Rap1 signaling pathway, MAPK/ERK signaling pathway, and calcium-related signaling pathways) and immune process (ie IL-6 production, IgA production, Fc epsilon RI signaling pathway and Fc gamma R-mediated phagocytosis) (Supplementary Figure 28A–D). Particularly, the calcium signaling pathway was involved in regulating most of the others and it was probably the initial response to FUS stimulation (Supplementary Figure 28E). GSEA analysis proved that these signaling pathways involving CD8 T cells proliferation, activation and migration, such as ECM-receptor interaction, PI3K-AKT signaling pathway, Rap1 signaling pathway and calcium signaling pathways were significantly upregulated (Supplementary Figure 28F). Although the GO and KEGG enrichment analysis of the downregulated genes showed the downregulation of MAPK signaling pathway, TNF signaling pathway and others (Supplementary Figure 28D), GSEA analysis revealed that these signaling pathways were highly expressed when the tumor-bearing mice were subjected to FUS sti. spleen (Supplementary Figure 28G).

These functional annotations suggested that FUS first activated calcium-related signaling pathways and then regulated other signaling pathways to promote the proliferation, activation, and migration of NK cells and CD8 T cells. Many previous studies confirmed that ultrasound affects the proliferation, activation, and differentiation of various cells by altering intracellular calcium-related signaling pathways to achieve specific biological effects.^77–79 The further exploration of splenic calcium changes in response to FUS stimulation by the Von Kossa staining and FCM detecting Fluo-4 AM labeled cells revealed that the splenic cell population of Von Kossa stained and Fluo-4 AM labeled were observably increased in the FUS sti. spleen group as compared with the control group (Figure 6A and B). Calcium deposits in spleen can promote cytokine production, such as Mφ secreting IL-1β and TNF-α.^80,81 Particularly, the results of in vitro experiments (Figure 6C) fully demonstrated that FUS stimulation promoted NK cell proliferation and activation to suppress cancer cell deterioration (note: H22 cancer cells were unable to adhere to the bottom and difficult to separate from NK cells that migrated from the upper layer of the transwell to the bottom layer, which led to a significant error in the subsequent detection results. Thus, Hepa1-6 cancer cells were chosen for the in vitro study). Moreover, calcium strengthened the impact of FUS stimulation on splenic NK cells to suppress the proliferation of cancer cells in vitro (Figure 6C). Besides, ultrasound shattered cancer cell suspension further enhanced the anticancer effect of calcium-involved FUS simulation on splenic NK cells. Additionally, we also verified that calcium reinforced the effect of FUS stimulation on splenic CD8 T cells to suppress cancer cell deterioration in vitro (not supplying the data). Overall, the above experimental results succinctly proved that FUS altered calcium-related signaling pathways in NK cells and CD8 T cells to reinforce the antitumor effect. However, in the future, it is worthwhile comprehensively and systematically analyzing the specific mechanism of splenic immune cells in response to FUS stimulation by spatial region-resolved proteome and spatial single-cell sequencing.

Figure 6 Calcium strengthened FUS-stimulated splenic NK cells against Hepa1-6 cancer cells. (A) Von Kossa staining of calcium deposition in the spleen with or without FUS stimulation. (B) FCM detected Fluo-4 AM labeled cells in the spleen with or without FUS stimulation. (n = 4; * p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001; mean ± SEM). (C) results of calcium strengthened impact of FUS stimulation on splenic NK cells to suppress cancer cells in vitro, including GFP fluorescence imaging, CV staining and CCK-8 assay to analyze the proliferation and activity of Hepa1-6 cancer cells, and CCK-8 assay to characterize the proliferation and cytoactivity of NK cells. (n = 3; * p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001; mean ± SEM). T1, Hepa1-6 cancer cells; T2, NK cells; T3, NK cells cocultured with Hepa1-6 cancer cells in the transwell; T4, FUS stimulated NK cells; T5, NK cells subjected to FUS irradiation, and then cocultured with Hepa1-6 cancer cells in the transwell; T6, NK cells mixed with Hepa1-6 lysate and cocultured with Hepa1-6 cancer cells in the transwell; T7, NK cells mixed with Hepa1-6 lysate, subjected to FUS irradiation, and then cocultured with Hepa1-6 cancer cells in the transwell; T8, NK cells mixed with Hepa1-6 lysate in 300 nM calcium culture medium and cocultured with Hepa1-6 cancer cells in the transwell; T9, NK cells in 300 nM calcium culture medium subjected to FUS irradiation, and then cocultured with Hepa1-6 cancer cells in the transwell; T10, NK cells mixed with Hepa1-6 lysate in 300 nM calcium culture medium, subjected to FUS irradiation, and then cocultured with Hepa1-6 cancer cells in the transwell.

Therapeutic Application of FUS Sti. Spleen on Intermediate and Advanced Tumors

The above studies in which FUS sti. spleen was performed on the second day after cancer cells implanted into the mice were equivalent to the start of the treatment in the early stage of cancer (before tumor formation). However, the anticancer effect of FUS sti. spleen after tumor formation is one of the worthiest explorations in clinical transformation and application. Therefore, a comparative study was performed to investigate the therapeutic application of FUS sti. spleen on 5^th and 9^th day after H22 cancer cell transplantation into the liver deemed to be tumorigenesis (Figure 7A). Four experimental groups were set as follows: K1 group was used as the control group of xenograft H22 HCC in situ; K2-K4 groups were subjected to FUS sti. spleen on days 0, 4 and 8 after H22 cancer cell implantation, and once every two days (Figure 7A). At the experimental endpoint (day 21), animals underwent terminal anesthesia followed by comprehensive tissue harvesting for other experimental analyses (eg, FCM).

Figure 7 FUS sti. spleen suppressed the proliferation of the xenograft H22 HCC in situ. (A) experimental flow diagram of FUS sti. spleen on day 0, 4 and 8 after H22 cancer cell implantation. (B) images of spleen and xenograft H22 HCC in situ. (C) tumor volume. (D) tumor weight. (E) liver weight. (F) curve of mice weight. (G) spleen weight. (H) spleen index calculated by the ratio of spleen weight to mouse weight. (n = 7; * p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001; mean ± SEM). (I–L), FCM results of NK cells, CD8 T cells and PMN-MDSCs in the spleen, blood, tumor, and para-carcinoma tissues, respectively. (n = 4; * p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001; mean ± SEM). K1, control group of xenograft H22 HCC in situ; K2-K4, FUS sti. spleen on day 0, 4 and 8, respectively, after H22 cancer cell implantation. Representative FCM plots and statistical diagram of immune cells in K1-K4 groups are shown in Supplementary Figures 29–32. The results shown in Figures 7 and 5 were from the same batch of experiments. With animal welfare in mind, only one control group and FUS sti. spleen group were set up; thus, the K1 and K2 group were the same as the H1 and H2 group respectively.

The results showed that FUS sti. spleen on day 0, 4 and 8 significantly suppressed the proliferation of xenograft H22 HCC carcinoma in situ, with an anti-tumor rate of up to ~70%, ~67% and ~48% respectively (K2/3/4 vs K1; Figure 7B–D). The results tautologically demonstrated that FUS sti. spleen exerted an evident tumor suppression, regardless of being performed before or after tumorigenesis, but the later ultrasound irradiation on the spleen was performed after the onset of the tumor, with a smaller effect on tumor suppression. Additionally, the statistical results of liver weight and mice weight showed a decreasing trend in the K2-K4 groups compared to the K1 group (Figure 7E and F). The spleen weight and spleen index also demonstrated that FUS sti. spleen attenuated the symptoms of splenomegaly caused by cancer, as same as in Figure 7G–H.

Statistic FCM results showed that the proportion of NK cells and CD8 T cells was significantly increased in the spleen, blood, tumor, and para-carcinoma tissue of the K2-K4 groups compared with the K1 group, and the proportion of PMN-MDSCs was significantly reduced (Figure 7I–L, Supplementary Figures 29–32). However, no significant difference was found in the number of immune cells among the K2-K4 groups except that NK cells were significantly higher in the K2 group (Figure 7I–L, Supplementary Figures 29–32). In addition, the proportion of Mφ1 and DC1 in four tissues of the K2-K4 groups was significantly increased (Supplementary Figures 29–32). What needs to be highlighted is the inconsistency in the changes in the cell proportion of Mφ2, DC2 or other immune cells in the spleen, blood, tumor, and para-carcinoma tissue (Supplementary Figures 29–32). Our hypothesis was that this phenomenon was mainly attributed to the migration of splenic immune cells to tumor lesions after responding to ultrasonic stimulation and stressing the tumor microenvironment, which was fully demonstrated in our previous work that CD11b⁺ CD43^hi Ly6C^lo splenocyte-derived Mφ migrate into the lesion in a liver fibrosis model.⁸²

Special attention should be paid to the fact that the biological effect of ultrasound on spleen varies in different disease models. As reported by Nunes et al⁶⁸ and Liu et al,⁸³ FUS sti. spleen mainly affects Mφ to inhibit the secretion of inflammatory cytokines in colitis and myocarditis. Also related to RA, CD 4 T cell, CD8 T cell, B cell and myeloid cell populations were significantly changed in response to ultrasound, specifically the CD8 T cells show an evident correlation with FUS sti. spleen based therapy.⁸⁴ However, this study fully confirmed that FUS sti. spleen significantly promoted the proliferation of NK cells, CD8 T cells, Mφ1 and DC1 in various tumor models, and reduced PMN-MDSCs, but mainly suppressed cancer progression by stimulating NK cells. Additionally, the biological effects of ultrasound on different types of cells are also varied, including enhancing MC3T3-E1 osteoblasts and hematopoietic stem cell ingrowth, proliferation, and early differentiation,^85,86 inducing neural progenitor cell polarization,⁷² promoting proliferation and migration of HaCaT keratinocytes,⁷³ facilitating the extracellular matrix synthesis of degenerative human nucleus pulposus cells,⁸⁷ and motivating the expression of brain-derived neurotrophic factor in astrocyte.⁷⁷ However, the difference in biological effects of ultrasound irradiation on various immune cells enriched in the spleen is not yet known and needs to be detailedly studied in the future.

In conclusion, although FUS sti. spleen non-specifically activated spleen immunological function to universally suppress tumor proliferation, it was effective on the early stage of cancer or cancer prevention. Nevertheless, it might be necessary to combine FUS sti. spleen with other treatments in patients with advanced cancer, such as applying FUS qualitatively and quantitatively intervention on the tumor (FUS int. tumor).

FUS Sti. Spleen & FUS Int. Tumor Synergistically Suppress Tumor Proliferation

Previous studies reported that ultrasonic intervention on tumor lesions enhances tumor suppression by promoting the proliferation, activation, and infiltration of immune cells.^88,89 In this study, comparing and discussing FUS sti. spleen and FUS int. tumors – two strategies for immunomodulation to counteract the swelling by interfering from immune organs and tumor foci, respectively – are crucial. Five experimental groups were considered: #1 group, normal mice (mice without any treatment); #2 group, control group of subcutaneous H22 tumor mice; #3 group, FUS sti. spleen on day 0 after H22 cancer cell implantation, and once every two days; #4 group, FUS int. tumor; #5 group, FUS sti. spleen and FUS int. tumor synergistic treatment. At the experimental endpoint (day 29), animals underwent terminal anesthesia followed by comprehensive tissue harvesting for other experimental analyses (eg, FCM). According to pre-experiments and our previous results,⁹⁰ FUS int. tumor was performed every 4 days from day 9 after the subcutaneous inoculation of H22 cancer cells (Figure 8A), and the ultrasonic parameters were set to 3.3 MPa, 1%, 20s to achieve controlled disruption of tumor’s extracellular matrix barrier and tumor microenvironment, thereby enhancing immunomodulatory effects without excessively compromising tissue architecture or inducing cellular necrosis.

Figure 8 FUS sti. spleen and FUS int. tumor synergistically or individually suppressed the proliferation of the subcutaneous H22 HCC tumor. (A) experimental flow diagram of FUS int. tumor every four days started on day 8 after H22 cancer cell implantation. (B) image of spleen and tumor. (C) tumor volume. (D) tumor weight. (E) curve of mice weight. (F) spleen weight. (G) spleen index, the ratio of spleen weight to mouse weight. (n = 8; * p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001; mean ± SEM). (H–J) FCM results of NK cells, CD8 T cells and PMN-MDSCs in the spleen, blood, and tumor, respectively. (n = 4; * p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001; mean ± SEM). (K and L) schematic illustration of the mechanism during FUS int. tumor to enhance anticancer immune response of FUS sti. spleen. #1, normal mice group; #2, control group of subcutaneous H22 tumor mice; #3, FUS sti. spleen on the second day after H22 cancer cell implantation; #4, FUS int. tumor; #5, FUS sti. spleen and FUS int. tumor synergistic effect.

The results of #3 vs #2 group again confirmed the ability and application prospect of FUS sti. spleen in suppressing tumor proliferation (Figure 8B–D). Moreover, FUS int. tumor showed an effective tumor suppression (#4 vs #2 group; Figure 8B–D), but it did not show any significant difference as compared with the #3 group (Figure 8B–D). It was noteworthy that FUS sti. spleen and FUS int. tumor synergistically induced the best tumor suppression (Figure 8B–D). In addition, FUS sti. spleen and FUS int. tumor synergistically or either alone reduced the mice weight (Figure 8E), but the combined application significantly exacerbated splenomegaly (Figure 8F and G).

FCM results revealed that the proportion of NK cells and CD8 T cells in the spleen, blood and tumor was significantly and synergistically increased compared to the effect of each of the two taken individually (Figure 8H–J). The comparison of the #3-#5 groups revealed that the number of NK cells and CD8 T cells in the spleen and blood was significantly higher after H22 tumor-bearing mice were subjected to the synergistic effect of FUS sti. spleen and FUS int. tumor compared to the effect of each of the two taken individually (Figure 8H and I). However, such a trend in the number of immune cells was not present in tumor tissues, which was mainly attributed to the fact that FUS int. tumor not only disrupted the structure of tumor stromal barrier and cancer cells, but also killed immune cells. Additionally, FUS sti. spleen, FUS int. tumor or both synergistically significantly increased the proportion of Mφ1 or DC1 in the spleen, blood, and tumor (Supplementary Figures 33–35). However, consistent with the FCM results in Table 2, FUS sti. spleen had no significant consistent effect on CD4 T cells or their subtype in the spleen, blood, and tumor from subcutaneous H22 tumor-bearing mice.

FUS has been gradually developed into a clinical strategy approach for targeted tumor disruption through thermal mechanisms.^91–93 In essence, FUS at high intensity generated focal temperatures exceeding 60 °C that instantaneously caused coagulative necrosis of the target soft tissue (Figure 8K).^94,95 In contrast, non-thermomechanical disruption may have a more relevant role in low intensity FUS ablation, particularly in terms of immunomodulatory effects. As opposite to thermal ablation causing protein coagulation necrosis, the mechanical disruption of FUS int. tumor converts immunologically “cold” (poorly infiltrated) tumors into immunologically “hot” (well-infiltrated) ones by damaging cancer cells to release antigens, collapsing the tumor matrix barrier to enhance antigens captured by immune cells, and especially intensifying the infiltration of immune cells and cytokines into the tumor lesions, which strengthened the immunotherapeutic responses to cancer (Figure 8K and L).^88,96 Although FUS int. tumor alone exerted an anti-tumor effect, to a certain extent, the tumor matrix damage would increase the risk of cancer cell spread and metastasis.⁹⁷ However, in this study, the synergistic use of FUS sti. spleen and FUS int. tumor was much more effective, since tumor disruption by FUS provided an antigen source for activating splenic immune cells during the process of FUS sti. spleen to suppress tumor proliferation (Figure 8L).

Discussion

The comparison of tumor inhibition and spleen injury under different ultrasonic parameters of FUS sti. spleen revealed that the spleen (or splenic cells) had significant ultrasound dose dependence and tolerance. This was also reported in previous studies; for example, Liu et al⁸³ successfully screened an effective ultrasonic parameter of 0.35 MPa with a 1s on /5s off duty cycle to significantly alleviate autoimmune myocarditis, and regulate the proportion and function of Tregs and Mφ by activating CAP. Besides, Cotero et al²³ discovered that non-invasive splenic ultrasound stimulation at 0.83 MPa effectively alleviates the severity of arthritis and reduces cytokine response to endotoxin through CAP modulation based on CD4⁺ T cells. However, according to a summary of studies (Supplementary Table 3), although splenic ultrasound stimulation can effectively suppress various inflammatory diseases, the ultrasonic parameters applied for spleen stimulation are extremely variable. Some studies only referred to the ultrasonic parameters mentioned in previous reports but lacked screening and optimization; for example, Hu et al⁸⁴ and Morton et al⁹⁸ used the parameters of 0.35 MPa and 1-s on/5-s off reported by Zachs et al³⁰ for splenic immunomodulation to treat inflammatory diseases. However, in practical applications, different ultrasound platforms lead to significant differences in ultrasonic focal region. More importantly, although some studies proved that the mechanism of splenic ultrasound stimulation for inflammatory disease therapy is based on CAP activation,^23,29,69 some other studies demonstrated the direct action of ultrasound on splenic immune cells.^30,84 Therefore, the dependence, specificity, and mechanism of spleen response to ultrasonic stimulation is also different, which may be one of the reasons for the different ultrasonic parameters applied in the above studies. This study also demonstrated that FUS sti. spleen to modulate immune function for tumor suppression was not mainly by activating CAP but depended on the direct stimulation of the proliferation and activation of NK cells, as well as the biological effects of ultrasonic stimulation on CD8 T cells, Mφ, DCs and MDSCs.

At present, there is a lack of studies on splenic ultrasound stimulation to regulate immunotherapy for various diseases; thus, ultrasonic parameters are not objectively consistent, which seriously restricts the translational progress of clinical application. Even if the ultrasonic parameters were screened and optimized in this study, there are still some shortcomings. For example, when the ultrasonic intensity is less than 2.3 MPa, the effect of a longer duration of ultrasound irradiation on splenic immune modulation for tumor suppression needs to be further studied. Fortunately, there are two studies of splenic ultrasound stimulation for immunotherapy on inflammatory diseases registered in http://www.clinicaltrials.gov (NCT03690466, and NCT03548116), but their clinical results are not yet officially published. We look forward to their positive news, and expect that FUS sti. spleen can be widely employed in clinical treatment to alleviate patient suffering.

Conclusion

Combined under optimized ultrasound parameters (2.3 MPa, 1%, and 20s), FUS sti. spleen has been demonstrated to directly activate splenic immune cells, particularly NK cells and CD8⁺ T lymphocytes, through altering calcium-dependent signaling but without significant involvement of cholinergic neuroimmune modulation, thereby effectively suppressing early-to-intermediate stage malignancies. With perspective, concurrent FUS sti. spleen and FUS int. tumor significantly achieve superior therapeutic outcomes, indicating substantial clinical translation potential that warrants further validation through animal and clinical trials.

Abbreviations

IRI, ischemia-reperfusion injury; VNS, vagus nerve stimulation; α7nAChR, α7 nicotinic acetylcholine receptors; FUS, focused ultrasound; DC, dendritic cell; NK, natural killer; MDSC, myeloid-derived suppressor cell; M-MDSC, monocytic myeloid-derived suppressor cell; PMN-MDSC, polymorphonucler myeloid-derived suppressor cell; Mφ, macrophage; Treg, regulatory T cells; FCM, flow cytometry; HCC, hepatocellular carcinoma; TNF-α, tumor necrosis factor α; CAP, cholinergic anti-inflammatory pathway; NE, norepinephrine; ACh, acetylcholine; IgA, immunoglobulin A.

Data Sharing Statement

Raw and analyzed sequencing data in this study have been deposited in the NCBI’s Gene Expression Omnibus (under series accession code GSE267237 and GSE267445).

Ethical Approval

All animal experiments were performed in accordance with the institutional guidelines and approved by the Animal Experimentation Ethics Committee of Xi’an Jiaotong University. Additionally, all methodological protocols were designed and reported in accordance with the ARRIVE guidelines for the welfare of the laboratory animals.

Acknowledgments

We thank Hongwei Tian, Gaixia He and Haiyan Chen for providing technical support in preparation of animal models. We thank Pro. Pengfei Liu, Pro. Guangyao Kong, Pro. Yujin Zong, Pro. Shemin Lv and Pro. Tielin Yang for their careful guidance on the design of experimental scheme. In addition, Wei Dong thanks the care and support from Yinggang Zhang, Shoufei Qu, Yameng Wei, Shaoying Zhang, Pro. Jun Li and Pro. Fanpu Ji in the past 2 years.

We would like to thank MogoEdit (https://www.mogoedit.com) for its English editing during the preparation of this manuscript. And we sincerely thank Shanghai OE Biotech Co., LTD (https://www.oebiotech.com/) for the RNA sequencing and data analysis.

This paper has been uploaded to ResearchSquare and bioRxiv as a preprint: https://www.researchsquare.com/article/rs-4639146/v1; https://www.biorxiv.org/content/10.1101/2025.03.31.646454v1.

Funding

This work was supported by the National Natural Science Foundation of China (No. 12204370), the Innovation Ability Supporting Program of Shaanxi Province (No. 2023WGZJ-ZD-09), and the Basic-Clinical Integration Innovation Project of Xi ‘an Jiaotong University (No. YXJLRH2022092).

Disclosure

The authors declare that they have no competing interests in this work.

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Introduction

As a benign disease, keloids are featured by the overgrowth of dense fibrous tissue on the affected skin. Although benign, they often cause troublesome symptoms including itching, pain, and pruritus. Furthermore, their high recurrence rates make them particularly challenging in clinical practice.¹ Recommended treatments for keloids include compression therapy, intralesional injections, laser therapy, topical treatments, surgical excision, postoperative radiotherapy, and cryotherapy.² However, the nonspecific nature of these therapies increases the risk of damaging surrounding tissues. Intralesional corticosteroid injections, a standard first-line treatment, frequently result in side effects such as localized pain and changes in skin appearance.³ The combination of corticosteroids and 5-Fluorouracil has demonstrated potential in improving treatment outcomes, yet persistent side effects remain a concern.⁴ Furthermore, irregular post-operative radiotherapy, or intralesional injections, and high-risk sites have also been implicated in the recurrence of keloids.^5–7 Therefore, targeting the distinct genetic and protein expression profiles of keloids compared to normal skin necessitates the identification of novel therapeutic targets and corresponding ligands to advance keloid treatment.⁸

Previous studies on keloid therapy have primarily focused on the transforming growth factor-beta (TGF-β)/Smad signaling pathway,^9,10 which plays a crucial role in the prolonged stimulation of fibroblasts and myofibroblasts, leading to excessive collagen production in keloids.⁹ However, considering the involvement of TGF-β in multiple biological processes, therapies targeting this signaling may cause systemic cytotoxic effects, limiting the advancement of TGF-β-based treatments for fibrotic diseases.¹⁰ Therefore, further research is warranted to refine targeted therapies for keloids.

In our previous study, we identified several potential targets for keloid therapy through Mendelian randomization and single-cell sequencing analyses. However, these studies did not reveal any functional targets. Notably, proteins from the BAG family have attracted our interest due to their overexpression in both tumors and, to some extent, keloids, highlighting the tumor-like properties of these lesions.¹¹ Among them, BAG cochaperone 2 (BAG2) has emerged as a promising target for keloid therapy according to the single-cell sequencing analysis. Previous research has shown that the high BAG2 expression among tumor-associated fibroblasts correlated with the poor prognosis in breast cancer and its anti-apoptotic characteristics,^12,13 suggesting that BAG2 may play a similar role in keloid progression. Mechanistically, BAG2 negatively regulates the chaperone-associated ubiquitin ligase, C terminus of Hsc70-interacting protein (CHIP), which facilitates the ubiquitin-mediated degradation of misfolded proteins. This suggests that BAG2 may inhibit the degradation of specific proteins, including overexpressed collagen in keloids. Additionally, BAG2 interacts with the MAPK signaling pathway, influencing downstream cellular proliferation.^14,15 In keloids, BAG2 may inhibit collagen degradation and promote abnormal collagen accumulation, warranting further investigation.

This study aimed to validate the role of BAG2 in keloid progression and identify compounds targeting BAG2, employing high-throughput screening technology tailored for keloid patients. The experimental setup is illustrated in Figure S1. Additionally, we explored the intrinsic connection between BAG2 and the progression of keloids to provide insights for more precise keloid management.

Materials and Methods

Ethics Statement

Keloid biopsies (10 patients with 4 men and 6 women age ranging from 26 to 42 years old, with keloids from operative excision of previous abdominal incisionor chest incision, having the lesion of over one year without reduction, without previous surgical treatment or radiotherapy, diagnosed by 2 experienced clinical experts with lesions extending beyond the wound boundary into the normal skin and other clinical features) and normal skin samples (3 female patients, aged 38, 41 and 42, with skin excision from facial plastic surgery as brow lifting) were obtained from the Shanghai Jiao Tong University affiliated ninth people’s hospital in accordance with the institutional review board (SH9H-2024-TK561). All patients provided formal, informed and written consent to supply a biopsy for this study. The study complied with the Declaration of Helsinki.

Keloid Fibroblasts (KFs) Isolation and Culture

Ten keloid samples from four men and six women were obtained after surgical excision. The sterile samples were rinsed in PBS, cut into pieces, and digested with 0.2% collagenase IV for 4 hours at 37 °C. After centrifugation, the sedimentary cells were resuspended in DMEM supplemented with 10% fetal bovine serum (FBS), 100 U/mL penicillin, and 100 µg/mL streptomycin, and incubated at 37°C in a 5% CO₂ incubator. The spindle keloid fibroblasts (KFs) from at least three patients at passages 2 to 3 were mixed for usage in further experiments.¹⁶

Single-Cell Data Analysis

To explore BAG2 gene expression in different cell groups, we downloaded the single-cell dataset GSE181297 of keloid patients from the Gene Expression Omnibus (GEO) database.¹⁷ This dataset was generated via the Illumina NovaSeq 6000 platform for scRNA-seq including keloid and normal skin samples from human subjects. The data processing included normalizing, filtering low-quality cells, de-batching, selecting highly variable genes for dimensionality reduction, and analyzing BAG2 expression in different cell clusters.

Quantitative Real-Time Polymerase Chain Reaction (qRT-PCR)

qRT-PCR was performed as previously described.¹⁸ Briefly, total RNA was extracted from cells after treatment using an RNA isolation kit (Takara Bio, Shiga, Japan). RNA purity was assessed by measuring the A260/A280 ratio, with acceptable values ranging from 1.8 to 2.0. The primers used for gene amplification are listed in Table 1. The results from three independent reactions were used to determine relative gene expression, normalized against β-actin expression.

Table 1 Primer Sequence

Cell Transfection

Small interference RNA (si-RNA) targeting BAG2 for BAG2 knockdown and its negative control (si-NC) were synthesized by Sangon Biotech (Shanghai, China). The sequence of si-BAG2 was listed in Supplementary Table 1. The non-transfected KFs were set as controls. Cell transfection was conducted using RNATransMate transfection reagents (Sangon Biotech, Shanghai, China) according to the manufacturer’s protocol.

Lentiviral Production and Infection

DNA oligonucleotides encoding primers for obtaining the target gene BAG2 (sequences: BAG2-1, 5’-AGGTCGACTCTAGAGGATCCCGCCACCATGGCTCAGGCGAAGATCAACGCTAAAG-3’, BAG2-2, 5’-TCCTTGTAGTCCATGGATCCATTGAATCTGCTTTCAGCATTTTG-3’), or a negative control were inserted into the lentiviral vector GV703 (Genechem, Shanghai, China). Human mRNA-BAG2 cDNA (synthesized by Genechem, Shanghai, China) was inserted into the BamHI/Age I sites of the lentiviral vector GV703. For lentiviral production, 293T cells were transfected with the lentiviral vector along with packaging plasmids using Lipofectamine 2000 (Life Science, USA) according to the manufacturer’s instructions. At 48 h and 72 h after transfection, culture media was collected, pooled and filtered. Human Fibroblast cell line (HHF1) obtained from The Institute of Cell Biology, Chinese Academy of Sciences was infected with the indicated lentivirus, and BAG2 expression was determined by WB at 48 h after infection.

Ex-Vivo Explant Culture of Keloid Tissue

After harvesting aseptic keloid tissue, and removing epidermis, the remaining dermis of keloids was cut into about 3×2 × 2 mm pieces by a scalpel. The dermal fragments were divided into different groups, and cultured for 3 days in DMEM containing 10% FBS as previously described.¹⁹ After tissue attachment, the medium was replaced for the control, siRNA-treated, and compounds-treated groups, which were then incubated for an additional five and seven days. The explants were collected after treatment.

Cell Proliferation Assay

The KFs transfected with si-BAG2 and si-NC (2×10³ cells/well) were seeded into a 96-well plate. Following the manufacturer’s protocol, 10 μL of CCK-8 reagent (Beyotime, Shanghai, China) was added to each well and incubated for 2 hours at 37°C. Subsequently, cell proliferation was assessed daily from day 1 to day 5. The number of cells was quantified by measuring absorbance at 450 nm using a microplate reader. Each treatment group was evaluated in triplicate to ensure reliable results.

Cell Cycle Analysis

KFs were treated with si-BAG2, C16-PAF (Targetmol, #T21547, 10 nM), or si-BAG2 alone for 24 and 48 hours, along with the si-NC control. After treatment, the cells were rinsed once with PBS, and fixed with 70% ethanol overnight. The cell cycle analyses were performed according to the instructions of the Cell Cycle Kit (Qihai Biotechnology, Shanghai, China), and flow cytometric analyses were performed by a flow cytometer (Beckman Coulter) equipped with ModiFit LT v2.0 software.

Cell Migration Assay

To evaluate the effect of BAG2 knockdown on the KFs migration, a scratch assay was performed. KFs were seeded in multi-well plates and cultured to confluence. The culture medium was then removed, and the cell monolayers were scratched using a 200 µL pipette tip. The cells were rinsed with PBS and subsequently treated with either si-BAG2, si-NC, or control. Images of cell migration were captured at 0 and 24 hours using an inverted light microscope (TE2000 Nikon, Japan) at 40×magnification at 0 and 24 hours. These experiments were performed in triplicate, and migration areas were quantified using ImageJ analysis software (National Institutes of Health, Bethesda, MD).

Histological, Immunohistochemical and Immunofluorescence Analyses

Keloid tissue and explant samples were fixed overnight in 4% paraformaldehyde at 4°C, embedded in paraffin, and sectioned to 5 μm thickness. The sections were subsequently stained with hematoxylin and eosin (H&E) and Masson’s trichrome for histological examination. Furthermore, keloid tissue sections were also treated with antibodies specific to BAG2 and α-SMA at dilutions from 1:2000 to 1:100. Antibody binding was visualized using 3,3’-diaminobenzidine (DAB) chromogen (Dako, Glostrup, Denmark) and counterstained with hematoxylin for immunohistochemical analyses. For immunofluorescence staining, after overnight incubation with primary antibodies, specimens were incubated with secondary antibodies for 1 hour. Cell nuclei were stained with 4,6-diamidino-2-phenylindole (DAPI). Finally, digital images were obtained using a V10-ASW 4.2 computerized image analysis system (Olympus, Tokyo, Japan). The antibodies used were: BAG2 antibody (Affinity, #DF2650, 1:100), Goat Anti-Rabbit IgG (H+L) HRP (Jackson, #111-035-045, 1:200), α-SMA antibody (Proteintech, #67735-1-Ig, 1:100), Goat Anti-Mouse IgG (H+L) Red (Jackson, #115-295-003, 1:200), and Goat Anti-Rabbit IgG (H+L) FITC (Jackson, #111-095-003, 1:200). The positive rate of BAG2 was defined as the proportion of BAG2-positive cells among all the cells in the field.

Western Blot

Western blot was performed on tissue samples or cultured cells as indicated. Total protein was extracted using RIPA lysis buffer, as previously described.¹⁸ The protein concentration of each lysate was determined using a BCA protein assay kit. Protein samples were separated by SDS-PAGE electrophoresis and transferred onto polyvinylidene fluoride (PVDF) membranes. The membranes were then blocked with 5% skimmed milk and immunoblotted with specific primary antibodies, including COL1, COL3, α-SMA, MEK, p-MEK and β-actin, diluted between 1:2000 and 1:1000 in TBST. After overnight incubation with the primary antibodies and three subsequent TBST washes, the membranes were incubated with the appropriate secondary antibodies for 1 hour at room temperature. Protein bands were detected using an enhanced chemiluminescence (ECL) kit (Amersham Biosciences, Chalfont St. Giles, UK). The intensity of each protein band was normalized against the β-actin band for comparison. The primary antibodies used in this study were: COL3 (Abcam, #Ab184993, 1:1000), COL1 (Huabio, #HA722517, 1:1000), α-SMA (Affinity, #AF1032, 1:1000), BAG2 (Affinity, #DF2650, 1:1000), MEK (Affinity, #AF3385, 1:1000), p-MEK (Affinity, #AF6385, 1:1000), and β-actin (Affinity, #AF7018, 1:2000).

Surface Plasmon Resonance (SPR) for Affinity Screening and Affinity Determination

SPR experiments were performed in at 25 °C on a BIAcore T200 with CM5 sensor chips, and data were analyzed with BIAcore T200 Evaluation software (GE Healthcare), following the manufacturer’s instruction. BIAcore T200 optical biosensor was used to screen for BAG2 affinity and to measure equilibrium dissociation constant (K_D) values for protein–ligand interactions. A cell on the CM5 sensor chip was activated using a mixture of 200 μM 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide and 50 μM N-hydroxysuccinimide at a flow rate of 10 μL/min for 420 seconds. Subsequently, BAG2 protein (50 μL) was mixed with 180 μL of 10 mM sodium acetate solution at pH 5.0 and immobilized on the cell surface at the same flow rate and duration for two cycles as previously described.²⁰ The cell was then blocked using 1 M ethanolamine. A neighboring channel, used as a reference, underwent the same activation and blocking steps, except immobilization was carried out with PBS adjusted to pH 5.0. Both channels were equilibrated with PBS afterward. Grouped SPR single concentration screening: Fifty compounds were grouped, with each compound contributing 1 μL, combined in an EP tube to total 50 μL. EP tubes were labeled from 1 to 54, and 950 μL of PBS was added to achieve a screening concentration of 10 μM for each compound. The DMSO content was maintained at 5% within the correction range. Each run flowed at 10 μL/min for 150 seconds. After each run, the chip was regenerated for 5 minutes with 10 mM glycine-HCl (pH 2.0), repeating until all groups (54 in total) were tested. Sequential SPR single concentration screening: The two groups with the highest RU values from the first screening round (100 compounds total) were selected. Each compound was taken as 2 μL from the plate and mixed with 198 μL of PBS to achieve a screening concentration of 100 μM. Each run flowed at 10 μL/min for 150 seconds. At the end of each run, the chip was regenerated for 5 minutes with 10 mM glycine-HCl (pH 2.0) solution, repeating this process until all compounds (100 in total) were completed. Data were collected from the sample cell using BIAcore T200 Control software (v. 2.0, GE Healthcare) and normalized against the reference cell. Association and dissociation constants were calculated using global fitting to a 1:1 Langmuir binding model with BIAcore T200 Evaluation software (v. 2.0, GE Healthcare). Final figures were generated using Origin 7 software (v. 7.0552, OriginLab).

To determine the K_D value for protein-ligand interactions, each compound was diluted 6 times from 12 μM to 15.625 nM. The compounds were flowed over a chip with immobilized BAG2 protein from lowest to highest concentration at 10 μL/min for 150 seconds. After each concentration, the chip was regenerated with 10 mM glycine-HCl (pH 2.0) for 5 minutes at the same flow rate. Data were recorded in real time, with molecular weight adjustment and solvent correction applied to address non-specific binding and signal drift molecular effects. Data analysis was conducted using BIAcore T200 software (GE Healthcare) following the manufacturer’s guidelines.

Statistical Analysis

All data are presented as means ± SD and analyzed using GraphPad Prism version 8 software (GraphPad Software Incorporation, La Jolla, California, United States). The cell migration area data were analyzed using Image J software. Sample sizes were determined based on practical and experimental considerations, without predetermined statistical methods. Differences among multiple groups were analyzed using analysis of variance (ANOVA), followed by Tukey’s post hoc test for comparisons between groups (α = 0.05). Differences among two groups were analyzed using t-test. Statistically significant differences were indicated by asterisks, with a two-tailed p-value of less than 0.05 considered significant.

Results

Higher BAG2 Expression in KF Clusters Compared to Normal Skin and Higher BAG2 Protein Expression in Keloid Tissue

After quality control of the GSE181297 dataset, we conducted dimensionality reduction and clustering analysis. Unique molecular identifiers between 200 and 6000 were selected. Cell clusters were categorized based on original molecular markers. These clusters included endothelial cell (EC), fibroblast (FB), proliferating myofibroblast (MFB), keratinocyte (KC), immune cells (lymphocytes, macrophages, and mast cells), lymphatic endothelial cell (LEC), and neural cell (Figure S2A and B). Besides, elevated BAG2 gene expression in KFs was observed in differential expression analysis (Figure S2C), with no significant difference in BAG2 expression between ECs and LECs in keloid and normal skin samples. Immunohistochemical staining confirmed BAG2 expression around fibrous nodules (Figure 1A). Additionally, immunofluorescent staining showed colocalization of BAG2 with α-SMA-positive cells, while BAG2 expression was also observed in α-SMA-negative cells, consistent with single-cell sequencing results (Figure 1B). Western blot analysis further demonstrated that BAG2 protein levels were significantly higher in keloid tissue than in normal skin samples (Figure 1C). The images of individual channel of immunofluorescence staining of keloid samples were showed in Figure S3A. The negative controls of immunohistochemical staining for keloid and normal skin, and the Hematoxylin and Eosin Staining of the keloid tissue was showed in Figure S3B and Figure S4.

Figure 1 The expression of BAG2 in keloid and normal skin samples. (A) Immunohistochemical analysis reveals enhanced positive staining for BAG2 in and around the fibrous nodules of keloid tissues compared to normal skin samples. (Scale bar = 100 μm at 200× magnification; Scale bar = 50 μm at 400× magnification); (B) The statistical analysis of BAG2 protein expression in keloid tissues and normal skin samples, as detected by immunohistochemical staining; (C) Immunofluorescence staining of a keloid sample slide shows nuclear staining with DAPI (blue) and colocalization of BAG2 (green) with α-SMA (red) positive cells; BAG2 expression is also visible in α-SMA negative cells. (Scale bar = 100 μm); (D) BAG2 protein levels in keloid tissue were significantly elevated compared to normal skin samples. (K: Keloid; (N) Normal Skin). ***P<0.001.

Inhibition of BAG2 Reduced Collagen Synthesis and Deposition of KFs, and in an Ex-Vivo Model

The relative gene expression of collagen type I (COL1), which is associated with collagen synthesis and excessive deposition in keloids,²¹ and Tissue inhibitor of metalloproteinase 1 (TIMP1), which inhibits matrix metalloproteinases and subsequently promotes COL1 deposition in keloids,¹⁹ was significantly decreased following BAG2 inhibition (Figure 2A). Additionally, Western Blot analysis showed reduced protein expression of BAG2, COL1, collagen type III (COL3), and α-SMA in si-BAG2-treated KFs, highlighting the role of BAG2 in abnormal collagen deposition (Figure 2B). Consistent with these findings, collagen structures in si-BAG2-treated ex-vivo keloid explants appeared thinner and more degraded compared to the control group (Figure 2C). Furthermore, COL1 and COL3 protein levels were significantly reduced in ex-vivo explants treated with si-BAG2 (Figure 2D).

Figure 2 Inhibition of BAG2 reduced keloid collagen synthesis and deposition. (A) The relative gene expression of COL1A1 and TIMP1 in si-BAG2-treated keloid fibroblasts compared to control groups. Gene expression related to collagen synthesis and the inhibition of collagen degradation was significantly reduced following BAG2 inhibition; (B) Western blot analysis showing reduced protein expression of BAG2, COL1, COL3, and α-SMA in si-BAG2-treated keloid fibroblasts; (C) Masson’s trichrome staining of keloid explants from different groups. Collagen structures in the si-BAG2-treated groups appeared thinner compared to control groups; (D) Western blot analysis demonstrates decreasing protein expression of COL1 and COL3 in si-BAG2-treated ex-vivo keloid explants. The reduction in COL1 and COL3 protein expression was statistically significant. Statistical analysis of relative protein expression is provided (n=3). **P<0.01, **** P<0.0001.

Inhibition of BAG2 Reduced Migration and Proliferation of KFs, Correlating with the MEK Pathway

BAG2 inhibition significantly reduced the migration and proliferation of KFs (Figure 3A and B). Additionally, the relative gene expression of TGF-β, an indicator of the proliferation and migration of KFs, was significantly decreased in si-BAG2-treated KFs (Figure 3C).²² In addition, Western Blot further demonstrated decreased levels of phosphorylated MEK (p-MEK) following si-BAG2 treatment (Figure 3D). Furthermore, si-BAG2 treatment significantly increased KFs’ percentage within the G0/G1 phase, suggesting cell cycle arrest. This effect was reversed by C16-PAF, an MEK activator, which restored G0/G1 phase percentages in si-BAG2-treated KFs (Figure 3E).

Figure 3 Inhibition of BAG2 reduced KF migration and proliferation via the MEK signaling pathway. (A) Representative images and bar graph depict the migration of KFs treated with or without si-BAG2 at 0 and 24 hours after scratching (n=3). Scale bar = 500 μm; (B) CCK-8 assays were performed on control and si-BAG2-treated KFs, showing a significant reduction in the proliferation of si-BAG2-treated cells (n=3); (C) Relative gene expression of TGF-β in si-BAG2-treated and control KFs, with significantly decreased TGF-β expression observed following si-BAG2 treatment (n=3); (D) Western blot analysis revealing decreased p-MEK in si-BAG2-treated cells; (E) Representative cell cycle profiles of KFs from various groups, assessed by flow cytometry, accompanied by statistical analysis (n=3). *P<0.05, ** P<0.01, ***P<0.001, **** P<0.0001. KF, keloid fibroblast.

Screening of Compounds with High‑affinity to BAG2 via Surface Plasmon Resonance (SPR) and Their Cell Inhibition Rates

BAG2 protein was found to play a significant role in collagen deposition in keloid tissues and the proliferation of KFs, making it a potential target for keloid treatment. To explore compounds with high-affinity to BAG2, the structure of BAG2 predicted by AlphaFold3 is shown in Figure S5. SPR was employed as an affinity screening technology to identify potential high-affinity compounds for BAG2.

A total of 2,732 compounds from the FDA and anti-fibrosis libraries were screened (Supplementary Table 2). In the first round, 2 groups of compounds were identified, and 6 compounds were identified as putative BAG2 ligands in the second round (Figure 4A and B). The primary cell inhibition rates of these 6 compounds on KFs were determined via CCK8 assays on Day 3 (Supplementary Table 3). Three compounds (Saikosaponin B1, Bazedoxifene acetate, and Ponesimod) showed significant inhibition of KFs and were selected for further investigation. Subsequently, the affinity index (K_D values) of these compounds with BAG2 was determined using SPR, and their inhibitory effects on KFs were evaluated at varying concentrations. The results indicated that Saikosaponin B1, Bazedoxifene acetate, and Ponesimod had K_D values of 6.35E−7 M, 2.84E−6 M, and 5.87E−6 M, respectively. Additionally, the fitted curves demonstrating the effects of different compound concentrations on the relative cell viability of KFs revealed IC50 values of 9.32 μM for Bazedoxifene acetate and 24.96 μM for Ponesimod, indicating that both drugs exhibit strong binding affinity and inhibitory effects (Figure 4C and D). Although Saikosaponin B1 exhibited relatively high binding affinity to BAG2, its inhibitory effect on KFs was below 40% at a concentration of 50 μM (Figure S6), suggesting it is not an optimal candidate for BAG2 targeting. The association rate constants (Ka) and dissociation rate constants (Kd) of these compounds are listed in Supplementary Table 4. Overexpressing BAG2 in human fibroblasts, and adding Bazedoxifene (10μM) acetate or Ponesimod (25μM) for 48h, it was found that these two compounds could inhibit BAG2-mediated cell cycle transition and cell proliferation (Figure 4E). The overexpression of BAG2 was validated by Western blot (Figure S7).

Figure 4 SPR screening of high-affinity compounds with BAG2 and validation in vitro. (A) Initial screening of 54 compound groups via SPR identified two groups demonstrating high affinity for BAG2 (shown in blue), compared to the positive control, BAG2 rabbit monoclonal antibody (mAb) from ABclonal Technology (A8775) (shown in red); (B) A second screening via SPR of two groups comprising 100 compounds revealed six compounds with high affinity for BAG2 (shown in blue), using the same BAG2 rabbit mAb as a positive control (shown in red); (C-D) SPR assays were used to analyze the binding affinities of Bazedoxifene acetate and Ponesimod to human BAG2 protein. Additionally, dose-response curves displaying the response of KFs to Bazedoxifene acetate and Ponesimod treatment over 72 hours are presented (n=6); (E) Representative cell cycle profiles of human fibroblasts from control, BAG2 overexpressed (oe-BAG2), oe-BAG2 treated with Bazedoxifene acetate or Ponesimod groups, assessed by flow cytometry, accompanied by statistical analysis (n=3). **** P<0.0001. KF, keloid fibroblast.

Compounds with High-Affinity to BAG2 Exhibited Inhibition of Collagen Deposition in Keloid Tissue

To further evaluate the therapeutic effects of the identified compounds on keloids, the effects of Bazedoxifene acetate and Ponesimod were assessed through protein expression analysis of COL1 and COL3, as well as histological evaluation of ex-vivo keloid explants. The experiments revealed that treatment with Bazedoxifene acetate and Ponesimod for five days or longer significantly reduced both the protein expression of collagen and the histological deposition of collagen in the ex-vivo explants (Figure 5). Additionally, Bazedoxifene acetate showed a superior inhibitory capacity for the expression of COL1 and COL3. These findings confirmed that compounds with high-affinity to BAG2 could markedly inhibit collagen deposition in ex-vivo keloid explants, indicating their clinical potential as therapeutic agents for keloids.

Figure 5 Masson’s trichrome of keloid explants in different groups. (A) In ex-vivo keloid explants treated with Bazedoxifene acetate and Ponesimod, the collagen structures appeared thinner compared to control groups; (B) Western blot analysis showed reduced protein expression of COL1 and COL3 in ex-vivo keloid explants treated with Bazedoxifene acetate and Ponesimod. Notably, Bazedoxifene acetate demonstrated a superior inhibitory effect on collagen deposition; (C) Statistical analysis of the relative protein expression is included (n=3). *P<0.05, **P<0.01.

Discussion

Keloids, a benign fibrogenic skin disease, share numerous characteristics with tumors, including the absence of spontaneous regression, excessive proliferation, and high recurrence rates.²³ Consequently, it is critical to explore therapeutic agents that target specific mechanisms involved in keloid formation. In this study, we identified BAG2 as a novel therapeutic target and screened thousands of compounds to find high-affinity BAG2 ligands. Our findings include the identification of Ponesimod and Bazedoxifene acetate as high-affinity ligands to BAG2, whose inhibitory effects were evaluated on keloid explants.

The present study observed an upregulation of BAG2 in keloid tissues, meanwhile, decreasing BAG2 with siBAG2 inhibited the proliferation of KFs, which correlated with MEK signaling. Consistent with the alterations of TGF-β following BAG2 inhibition, TGF-β modulates cell proliferation through the MEK pathway via a Smad-independent signaling mechanism (Derynck et al, 2003). This is consistent with prior findings linking MEK pathway activation to the proliferation of KFs,²⁴ and to the proliferative and migratory behaviors of cancer cells.^25,26 As evidenced by our findings, si-BAG2 suppressed KFs’ progression at the G0/G1 phase, a process that could be reversed by MEK activation. This indicates that BAG2 facilitates KF proliferation, which is correlated with the MEK signaling pathway, as an early indicator of a long-term process involving ECM remodeling. In accordance with previous studies, inhibiting the MEK pathway could induce G0/G1 phase arrest.^27,28 Additionally, the initial phosphorylation site Ser20, located within the curled helical domain near the amino terminus of BAG2, is phosphorylated by MAPKAPK-2. The p38 MAPK-MAPKAPK-2-BAG2 phosphorylation cascade, activated in response to extracellular stress, influences cell proliferation activities,²⁹ and may explain BAG2’s role in activating the MEK pathway. The BAG2-Heat shock protein (HSP) complex maintains basement membrane integrity and regulates ECM and cell junctions in glial cells, suggesting a similar role in keloids.³⁰ BAG2 also has anti-apoptotic effects that promote cell survival and, in some cases, enhances TIMP synthesis and activity by boosting intracellular anti-apoptotic factors. Since TIMPs inhibit MMP activity to control ECM degradation, BAG2-mediated upregulation of TIMPs may reduce MMP-driven ECM breakdown, preventing excessive degradation.¹³

While the proliferation and migration of KFs are well-documented, accelerated collagen accumulation also plays a significant role in keloid pathology.¹⁹ Additionally, mechanical stress at keloid sites has been recognized as a risk factor for the initiation and progression of this condition.³¹ The stiffness of the extracellular matrix (ECM) in keloids, largely due to collagen accumulation, augments the effects of mechanical stress, which, in turn increases the production of ECM by KFs.³² Therefore, halting the collagen deposition cycle is vital for effective keloid therapy.³³ Traditional therapies for keloids fail to adequately target the essential pathological processes of collagen synthesis and degradation.³⁴ In this study, altering BAG2 activity reduced both collagen synthesis and deposition in vitro and ex vivo. Additionally, our findings demonstrate that drugs with high-affinity to BAG2 can significantly reduce collagen accumulation, highlighting their potential as novel targeted therapies for keloids that inhibit collagen deposition and reduce the viability of KFs. In pulmonary fibrosis, TGF-β treatment promotes α-SMA expression through acetylated CCAAT/enhancer binding protein β (C/EBP-β), and a similar mechanism may exist in keloids, which warrants further investigations.³⁵

Several skin diseases, including psoriasis, vitiligo, dermatitis, and skin cancers, have been successfully treated with targeted therapies.^36–39 Drugs such as vemurafenib and trametinib, which target the BRAF mutation and MEK respectively, have significantly advanced the field of targeted therapy, although many such drugs are still in the preclinical research stage. Specifically, drugs targeting keloids, due to their localized presence on the body surface, could offer more precise treatment options. Nevertheless, there are currently no specific inhibitors targeting BAG2, a keloid-associated marker identified through this study. In this study, several high-affinity BAG2 ligands with were identified, including Saikosaponin B1, Ponesimod, and Bazedoxifene acetate, and verified their capability to inhibit collagen expression and deposition in ex-vivo keloid tissue. Saikosaponin B1, a bioactive molecule found in Radix Bupleuri, has been shown to exhibit anticancer properties⁴⁰ and antifibrotic activity in liver fibrosis.⁴¹ Ponesimod, a Sphingosine 1-phosphate (S1P) modulator, has been investigated as a treatment for Multiple Sclerosis, focusing on its ability to regulate S1P activity in lymphocytes.⁴² Bazedoxifene acetate, a selective Estrogen Receptor modulator, was examined for its role as a BAG2 ligand and its capacity to inhibit the viability of KFs in this study.⁴³ Interestingly, tamoxifen, another Estrogen Receptor modulator, has also been shown to reduce keloid collagen fibers.⁴⁴ Like bazedoxifene acetate, tamoxifen’s impact on keloids appears to extend beyond simple Estrogen Receptor modulation. Bazedoxifene acetate appears to influence keloid pathology by correlating with the inhibition of BAG2. The mechanisms through which bazedoxifene acetate and ponesimod act on keloids necessitate further detailed exploration.

Despite the lack of prior reports on these drugs as keloid inhibitors, their strong affinity for BAG2 and effectiveness in inhibiting KFs suggest their potential as promising candidates for targeted keloid therapy. Bazedoxifene acetate, in particular, stands out for its lower K_D, lower IC₅₀, and higher binding affinity. As an FDA-approved medication, bazedoxifene acetate may manage antifibrotic activity by affecting both collagen deposition and the viability of KFs. Moreover, due to their relatively high SLogP values, as detailed in Supplementary Table 5, bazedoxifene acetate and ponesimod were more likely to facilitate transdermal absorption, potentially reducing side effects associated with systemic administration. Topical application of these compounds may provide a viable therapeutic option for keloid treatment.

Despite the promising potential of drugs targeting BAG2 in keloid therapy, treatment with these agents may induce unintended off-target effects on other proteins. Therefore, further functional analyses and the development of systems to predict potential side effects are warranted. In this study, we found that inhibiting BAG2 could reduce collagen deposition. Additionally, the system of MMPs and TIMPs was also affected, warranting further investigation. Furthermore, the direct interaction between BAG2 and MEK signaling requires further investigation to elucidate the role of BAG2 in keloid formation. In addition, proliferation is a confounding factor for the scratch assay, the effect of BAG2 in the migration of KFs should be further investigated. Further, in vivo models are warrant for validation of the role of BAG2 in keloids. Compared to the ex-vivo model, the in-vivo model provides more comprehensive information on physiological responses and systemic pathological changes. In addition, as indicated by SPR, the binding affinity of Bazedoxifene acetate and Ponesimod to BAG2 is relatively higher than that of the other tested drugs. To further validate the effect of Bazedoxifene acetate and Ponesimod on keloids via BAG2, silencing experiments should be conducted in future studies. In summary, this study highlighted BAG2’s impact on collagen deposition and the proliferation of KFs, which are critical to the pathogenesis of keloids, benign skin tumors with high recurrence rates. Additionally, we identified compounds with high-affinity ligands to BAG2 that could serve as potential therapeutic targets. Among the compounds tested, Bazedoxifene acetate demonstrated superior affinity for BAG2 and more effective inhibition of keloid fibroblasts and keloid tissue compared to other ligands, suggesting novel approaches for the precise management of keloid disease.

Conclusion

This study revealed the pathogenic role of BAG2 in keloid and identified its high-affinity ligands, Bazedoxifene acetate and Ponesimod. The therapeutic capabilities of these compounds demonstrated their potential to improve targeting therapy for keloids.

Resource Availability

The data underlying this article are available in the article and in its online supplementary material. Single cell sequencing data is available in [GEO database] at [https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE181297]. Further data underlying this article will be shared on reasonable request to the corresponding author.

Acknowledgments

We thank all the patients who provided samples for the experiment for their support. This work was supported by National Nature Youth Foundation of China (Grant No. 82402938 & 82102319) Science and Technology Commission of Shanghai Municipality (Grant No. 22MC1940300) and Wuxi Taihu Lake Talent Plan, Supports for Leading Talents in Medical and Health Profession. The funding body did not play a role in the study design, data collection, analyses, interpretation, manuscript preparation, and in the decision to submit the manuscript.

Author Contributions

All authors made a significant contribution to the work reported, whether that is in the conception, study design, execution, acquisition of data, analysis and interpretation, or in all these areas; took part in drafting, revising or critically reviewing the article; gave final approval of the version to be published; have agreed on the journal to which the article has been submitted; and agree to be accountable for all aspects of the work.

Disclosure

The authors have declared that no conflict of interest exists. The patents associated with this article include patent application number 202411342096.X, credited to Yinmin Wang, Lin Lu, and Jun Yang.

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