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HDACi combination therapy with IDO1i remodels the tumor microenvironment and boosts antitumor efficacy in colorectal cancer with microsatellite stability

Journal of Nanobiotechnology volume 22, Article number: 753 (2024) Cite this article

Abstract

Background

Immunotherapy for colorectal cancer (CRC) with microsatellite stability (MSS) and mismatch repair proficiency (pMMR) has shown limited success in clinical trials. The combination of immunomodulators and immune checkpoint inhibitors (ICIs) is a potential strategy for treating CRC.

Methods

Histone deacetylase (HDAC) and indoleamine 2,3-dioxygenase 1 (IDO1) expression in CRC tissues and adjacent normal tissues was analyzed via database analysis, immunohistochemistry, and western blotting. A nanodrug designated as NP-I/P was subsequently formulated, encapsulating an IDO1 inhibitor (IDO1i; namely, epacadostat) and an immunomodulatory HDAC inhibitor (HDACi; namely, panobinostat). The antitumor efficacy of the nanoparticles and their effects on tumor microenvironment features were evaluated via in vitro and in vivo experiments.

Results

In the present study, we found that HDAC overexpression and IDO1 expression were attenuated in MSS/pMMR CRC. Thus, a nanodrug designated as NP-I/P was formulated to encapsulate epacadostat and panobinostat. In vitro, NP-I/P treatment promoted the apoptosis of tumor cells and induced the release of damage-associated molecular patterns, thereby leading to cell death–associated immune activation. The in vivo results revealed that NP-I/P treatment reversed the immunosuppressive phenotype of the microenvironment by inducing tumor immunogenic cell death (ICD), promoting CD8+ T cell infiltration, and reducing the numbers of Tregs, tumor-associated macrophages, and myeloid-derived suppressor cells. Finally, the results of the patient-derived xenograft and patient-derived organoid models demonstrated that NP-I/P treatment triggered tumor cell death and modulated the immune microenvironment in human CRC.

Conclusion

The combination of IDO1 and HDAC inhibitors represents a promising strategy for CRC treatment, and NP-I/P is a candidate for clinical trials.

Introduction

Colorectal cancer (CRC) is the third most frequently diagnosed cancer and second leading cause of cancer-related death worldwide [1]. Although advances in surgery, chemotherapy, and targeted therapy have markedly improved CRC prognosis [2, 3], effective treatments for patients with advanced CRC and microsatellite stability (MSS)/ mismatch repair-proficiency (pMMR) are lacking. Recently, immunotherapy with PD-1/PD-L1 inhibitors has been shown to be beneficial for patients in multiple cancers, such as melanoma and lung cancer [4, 5]. Unfortunately, immunotherapy has almost no effect on MSS/pMMR CRC [6, 7]. The immunosuppressive microenvironment is the major factor contributing to the poor response of MSS/pMMR CRC to immunotherapy or combined therapy [8, 9].

Various immune checkpoints have been investigated, and immune checkpoint inhibitors (ICIs) have been developed to improve cancer prognosis [10]. Indoleamine 2,3-dioxygenase 1 (IDO1) is an immune checkpoint that has been targeted for cancer treatments in recent years [11]. IDO1 is a critical enzyme in tryptophan-kynurenine metabolism, and catalyzes the first rate-limiting step in this process [12]. IDO1 is overexpressed in various cancers and cancer-associated cells, such as tumor-associated macrophages (TAMs), tumor-associated fibroblasts, mesenchymal stromal cells (MSCs), and myeloid-derived suppressor cells (MDSCs) [13,14,15]. Kynurenine that has accumulated in the tumor microenvironment (TME) may create an immunosuppressive environment and be correlated with poor patient survival and cancer prognosis. IDO1 expression and kynurenine prevent the activation of CD8+ T cells and NK cells [16, 17]; however, they can also promote the expansion of Tregs, MDSCs, and TAMs, which further inhibits the function of effector cells [18].

IDO1 is a promising immunotherapy target. Numerous IDO1 inhibitors, such as epacadostat, navoximod, and PF-06840003 have been tested in preclinical studies and clinical trials [19]. However, poor therapeutic responses to IDO1 inhibitors have been reported in these trials, especially in those on CRC [11]. The expression of IDO1 and lack of immune effector cells may have restricted the response to the IDO1 inhibitors. The expression level of IDO1 and the number of immune cells is lower in MSS-CRC than in MSI-H CRC, and these features lead to poor immunotherapeutic efficacy [20]. Therefore, boosting IDO1 expression and increasing the number of immune cells are potential methods for improving the efficacy of IDO1 inhibitors in CRC.

Histone deacetylases (HDACs) regulate gene expression via chromatin modification [21] and participate in angiogenesis, apoptosis, autophagy, the cell cycle, transcriptional regulation, and DNA damage repair [22]. Under normal physiological conditions, HDACs regulate histone acetylation in a balanced manner. In contrast, when cells undergo abnormal transformation, HDACs activity increase and disrupts normal gene expression, leading to uncontrolled cell proliferation and malignant cellular transformation [23]. Excessive HDACs expression and activity have been observed in multiple cancers, and HDACs are considered targets for tumor therapy [24]. Therefore, HDACs inhibitors (HDACis), such as panobinostat, tucidinostat, and vorinostat, have been synthesized and shown antitumor effects in clinical trials [25,26,27].

Recent studies have shown that the HDACi panobinostat can promote apoptosis, arrest the cell cycle, and suppress angiogenesis in various cancers [28, 29]. In addition, panobinostat contributes to the immunogenic cell death (ICD) of cancer cells [30] and facilitates dendritic cells (DCs) maturation, leading to the activation of the immune response in the TME. The accumulation of immune cells in the TME may provide conditions for the application of ICIs. Moreover, the inhibition of HDACs can promote the expression and secretion of IFN-γ [31], which is a factor that promotes IDO1 expression. Therefore, it may be assumed that the combination of IDO1 and HDAC inhibitors can boost antitumor efficacy. However, the combination of drug administration and drug delivery efficiency in vivo is a major factor that restricts therapeutic effects and causes side effects. In our previous studies, we reported that nanodrugs increased drug accumulation in tumors and reduced drug exposure in normal tissues, thus increasing antitumor efficacy and reducing the incidence of side effects in colon cancer [32, 33].

In this study, we designed a nanomedicine to deliver IDO1 and HDAC inhibitors to remodel the TME and achieve antitumor effects in MSS CRC. An mPEG -PCL nanoparticle was used for encapsulation of an IDO1i (epacadostat) and an HDACi (panobinostat) to form the nanodrugs NP-I/P. NP-I/P treatment not only restored the sensitivity of tumor cells to death and diminished tryptophan-kynurenine metabolism but also increased the release of damage-associated molecular patterns (DAMPs) and induced ICD in tumor cells. Cell line-derived xenograft(CDX)model experiments revealed that NP-I/P elicited a robust immune response by promoting ICD in tumor cells and activating CD8+ T cells. In addition, NP-I/P treatment promoted remodelling of the immunosuppressive microenvironment by reducing the number of Tregs and MSDCs and reversing M2 macrophage polarization. Moreover, we evaluated the antitumor and immune activation effects of NP-I/P in patient-derived xenograft (PDX) and patient-derived organoid (PDO) models. The results demonstrated the therapeutic effects of this strategy and its ability to modulate immunity in humanized tumor models. Our findings indicate that NP-I/P mediated synergistic HDAC- and IDO1-targeting immunotherapies may achieve favorable therapeutic effects in MSS CRC, providing a potential combination treatment for MSS CRC (Fig. 1).

Fig. 1
figure 1

Schematic illustration showing that the nanodrug encapsulating panobinostat and epacadostat remodels the tumor microenvironment and boosts antitumor efficacy in CRC with microsatellite stability.

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Methods and materials

Human subjects

Twenty-three patients who underwent surgery at the Third Affiliated Hospital of Sun Yat-Sen University (Guangzhou, China) with proven CRC were recruited for this study. Patients who received preoperative antitumor treatments (neoadjuvant radiotherapy or chemotherapy) were excluded. Tumor tissues and adjacent normal tissues were obtained as soon as possible after resection for the corresponding experiments. Patient consent was obtained from each patient, and the Ethics Committee of the Third Affiliated Hospital of Sun Yat-sen University approved the study. The demographic and clinical information and histopathological data for each patient are shown in Supplementary Table 1.

Public database

R2, UALCAN [34], and TIMER are interactive web portals that allow in-depth analysis of TCGA gene expression data. The R2: Genomics Analysis and Visualization Platform is publicly available at http://r2.amc.nl. UALCAN is publicly available at http://ualcan.path.uab.edu. TIMER is publicly available at https://cistrome.shinyapps.io/timer/.

Western blotting

Treated tumor cells were lysed with RIPA lysis buffer(Beyotime Biotech, China)supplemented with PMSF and phosphatase inhibitor at 4℃. For CRC samples, 20 mg of tissues was lysed with 200 µL of RIPA buffer in a grinder and then sonicated for 1 min. The lysate was subsequently centrifuged at 12,000 rpm for 10 min at 4℃. After the protein concentration was determined with a bicinchoninic acid protein assay kit (Beyotime Biotech, China), 30 µg of each sample was separated via 10% SDS-PAGE and then transferred to PVDF membranes (Millipore, USA). Then, the membrane was blocked with 5% BSA for 1 h and incubated with the corresponding primary antibodies (shown in the Supplementary Table 2) overnight at 4℃. The membrane was washed with TBST three times and incubated with horseradish peroxidase–conjugated secondary antibodies at room temperature for 2 h. Finally, the bands were visualized via an automatic chemiluminescence image system (Tanon 5200, China) and enhanced chemiluminescence detection kits (Beyotime Biotech, China).

Immunohistochemistry and immunofluorescence analyses of tissue samples

The paraffin-embedded tissues were sectioned into slices at a thickness of 4 μm, and the sections were deparaffinized and rehydrated after heated in 65 °C for 1 h. Tris-EDTA antigen retrieval solution was subsequently used for heat-induced epitope retrieval with a microwave oven. After incubation with 5% BSA at room temperature for 1 h, the sections were incubated with primary antibodies (shown in Supplementary Table 2) overnight at 4℃. For immunohistochemistry, the sections were incubated with peroxidase-conjugated antibodies and detected with a DAB system (Servicebio Technology, China). For immunofluorescence staining, the samples were incubated with fluorescently labelled secondary (Servicebio Technology, China) antibodies for 60 min at room temperature and the nuclei were stained with DAPI. Finally, the sections were imaged via a confocal microscope (Leica, Leica STELLARIS STED, Germany) or fluorescence microscope (Nikon, ECLIPSE Ti2-U, Japan).

Enzyme linked immunosorbent assay (ELISA)

To evaluate the release of kynurenic acid, ATP, and HMGB1 from CT26 tumor cells, the cells were seeded in 6 well plates at a density of 5 × 105 per well and incubated in RPMI-1640 medium supplemented with 10% FBS for 24 h. Then, the medium was removed and replaced with fresh medium containing phosphate-buffered saline (PBS), NP-I, NP-P, or NP-I/P. After incubation for 48 h, the culture medium was collected and kynurenic acid, ATP, and HMGB1 levels were measured by using ELISA kits according to the manufacturer’s instructions (Meimian, China). To evaluate the release of kynurenic acid and TGF-β from MDSCs, purified MDSCs were seeded in 6 well plates at a density of 5 × 105 per well and incubated in 1640 medium supplemented with 10% FBS for 24 h and then the same procedure as above was performed.

In vitro cytotoxicity experiments

Cell Counting Kit-8 (CCK8) assays were performed to evaluate the cytotoxicity of the nanodrugs in vitro. CT26 colon cancer cells were seeded into 96 well plate at a density of 5 × 103/100 µL and cultured for 24 h. Then, the culture media were replaced and nanodrugs were added to the media at a specific concentration gradient. After 48 h, CCK-8 reagent (Beyotime Biotech, China) was added, and cell viability was assessed according to the manufacturer’s instructions.

Preparation and characterization of nanodrugs

mPEG-PCL was purchased from Tansh Technology (Guangzhou China) and its structure was confirmed by nuclear magnetic resonance (NMR, Supplementary Fig. 1). Panobinostat and epacadostat were obtained from MedChemExpress Co., Ltd (Shanghai, China). A self-assembly method was utilized to prepare the nanodrugs as described previously [35]. For the synthesis of a nanodrug loaded with epacadostat and panobinostat (NP-I/P), 20 mg of mPEG-PCL was first dissolved in tetrahydrofuran (THF) solution. Then, 0.3 mg of panobinostat and 0.12 mg of epacadostat were added to the above mPEG-PCL solution. The mixture was added dropwise to ultrapure water (6 ml) while stirring at 1000 rpm, after which evaporation of THF at room temperature was performed for 48 h. Finally, the formed NP-I/P was collected and filtered through a 0.22 μm filter. For the synthesis of nanodrugs loaded with epacadostat (NP-I), panobinostat (NP-P), 1,1’-dioctadecyl-3,3,3’,3’-tetramethylindotricarbocyanine iodide (DiR), or coumarin, only epacadostat, panobinostat, DiR, or coumarin was added to the mPEG-PCL solution, respectively. The size distribution and morphology of the nanodrugs were measured via dynamic light scattering (DLS) and transmission electron microscopy (TEM). To confirm the stability of NP-I/P, 0.1 mL of NP-I/P solution was diluted in 0.9 mL of PBS containing 10% FBS at 37℃, and nanoparticle size was measured via DLS at predetermined time points.

Apoptosis analysis

A total of 5 × 105 CT26 cells or MDSCs were seeded in six well plate and treated with PBS, NP-I, NP-P, or NP-I/P for 48 h. The treated cells were collected and washed with PBS. The cells were subsequently resuspended and stained with the Annexin V-EGFP/PI Apoptosis Detection Kit (KeyGEN Biotech, China) at room temperature and analyzed via flow cytometry.

Cell cycle analysis

A total of 5 × 105 CT26 cells were seeded in six-well plates and treated with PBS, NP-I, NP-P, or NP-I/P for 48 h. The treated cells were collected and fixed in cold ethanol overnight at 4 °C. The cells were subsequently resuspended and stained with Cell Cycle Detection Kit (KeyGEN Biotech, China) reagents at room temperature for 30 min and analyzed via flow cytometry.

Live/dead viability assay

CT26 cells were seeded in 35 mm dishes at a density of 2 × 105 cells per dish and incubated in RPMI-1640 medium for 24 h. Then, the medium was removed and replaced with fresh medium containing PBS, NP-I, NP-P, and NP-I/P. After incubation for 48 h, the cells were stained with calcein acetoxymethyl ester (calcein AM) and PI (KeyGEN Biotech, China) and observed via fluorescence microscopy.

Migration, invasion, and colony formation assays

Cell migration and invasion assays were performed in 24-well Transwell chambers with 8-µm pore size polycarbonate. Matrigel was coated on the chamber for the invasion assay. A total of 5 × 104 cells were counted, suspended in serum-free culture medium, and then plated in the upper chamber. A total of 600 µL of cell culture medium containing 10% foetal bovine serum was added to the bottom chamber. After incubation for 24 h at 37 °C, the cells that migrated or invaded through the pores of the membrane were fixed with 4% paraformaldehyde and then stained with 0.1% crystal violet staining solution. The stained cells were visualized and counted under a microscope. For cell colony formation, the collected cells were counted, seeded into six-well cell culture plates (1000 cells per well) and incubated at 37 °C for 1 week. Then, the colonies were fixed with 4% paraformaldehyde for 15–20 min and stained with 0.1% crystal violet staining solution for 20–30 min. The cell colonies were photographed and counted.

Flow cytometry

Cells from different experiments were harvested and stained with fluorochrome-conjugated Abs according to the manufacturer’s instructions. For surface staining, the cells were collected and resuspended in the corresponding staining buffer. For intracellular staining, the cells were first stained with surface markers, fixed and permeabilized with Cytofix/Cytoperm Soln Kit (Becton Dickinson, USA) reagents, and finally stained with intracellular markers. To detect foxp3 in the nucleus, mononuclear cells were fixed and permeabilized with specific reagents (eBioscience, USA) after surface staining. For some experiments, T cells were stimulated with Leukocyte Activation Cocktail (Becton Dickinson, USA) at 37 °C for 4 h before staining. Finally, the data were acquired with a Calibur flow cytometer (Becton Dickinson, USA) and analysed with FlowJo software. The antibodies used in this study are summarized in Supplementary Table S2.

Cell immunofluorescence analysis

For cell immunofluorescence analysis, CT26 cells (1 × 105 cells per dish) were seeded into 30 mm dishes with 2 ml of RPMI-1640 and incubated at 37 °C for 24 h. PBS, NP-I, NP-P, and NP-I/P were added to different dishes at the same concentration of panobinostat and epacadostat and cultured for 48 h. After fixation with 4% paraformaldehyde for 20 min and antigen blocking with 5% BSA, the cells were stained with anti-CRT (Abcam, England) or anti-HSP70 (Abcam, England) antibody overnight at 4 °C and subsequently labelled with a FITC-conjugated secondary antibody for 1 h at room temperature. The cells were subsequently stained with DAPI and observed via laser scanning confocal microscopy (LSCM).

Isolation and purification of MDSCs and CD8+ T cells

For the isolation of MDSCs, a CT26-bearing mouse model was first constructed as described above. When the tumor size reached 500 mm3, the mice were sacrificed. The spleens were harvested and mechanically digested, followed by filtration through a 70 mm filter to obtain a single-cell suspension. Mononuclear cells were subsequently isolated via a standard Ficoll procedure as described previously [36]. MDSCs were purified via an anti-mouse Gr-1-biotin antibody and anti-biotin microbeads (Miltenyi Biotech, Germany) according to the manufacturer’s instructions.

For CD8+ T cell purification, BALB/c mice (male, 4–6 weeks of age) were sacrificed, and the spleens were subjected to isolation of mononuclear cells. Then, the CD8+ T cells were purified with anti-CD8a (Ly-2) MicroBeads (Miltenyi Biotech, Germany). Finally, the purity of the MDSCs and CD8+ T cells was determined via flow cytometry.

Isolation and culture of bone marrow-derived mononuclear cells (BMMCs)

BALB/c mice (aged 4–6 weeks) were sacrificed, and the femurs and tibias were collected after immersion in alcohol for 30 min. The ethanol on the surface of the tibia and femur was washed off with PBS. After the two ends of the femur and tibia were cut with scissors, a 1 mL syringe of cold PBS was used to aspirate the bone marrow from the femur and tibia. The bone marrow was subsequently filtered through a 70 μm mesh, and the blood cells were removed with red blood cell lysate to obtain BMMCs. The BMMCs were washed with PBS and resuspended in 1640 medium for the next experiment.

For macrophage induction, 20 ng/ml M-CSF was added to the culture medium of BMMCs, which were subsequently incubated at 37 °C with 5% CO2 for 7 days to obtain bone marrow-derived macrophages (BMDMs).

For DC induction, 15 ng/ml GM-CSF and 10 ng/ml IL-4 were added to the culture medium of BMMCs and incubated at 37 °C with 5% CO2 for 7 days. In accordance with routine culture, half of the culture medium was replaced every other day, and GM-CSF/IL-4 was added. On day 7, the culture medium was discarded, and loosely adhered BMDCs were collected to obtain immature DCs (iDCs).

Coculture of immune cells with CT26 tumor cells

For CD8+ T cells cocultured with CT26 tumor cells, purified CD8+ T cells were seeded in 6-well plates precoated with a-CD3 and a-CD28 was added to the culture medium. CT26 cells were placed into the plate at a ratio of 10:1 (CD8+ T cells:CT26 cells ) after treatment with different nanodrugs for 48 h. Then, the cells were cocultured for another 48 h, and the CD8+ T cells were harvested for IFN-γ detection. In some experiments, purified CD8+ T cells were first labelled with CFSE and then cocultured with CT26 cells to investigate the impact on cell proliferation.

For coculture of macrophages and DCs with CT26 tumor cells, macrophages and DCs isolated from the bone marrow were seeded in a 6-well plate, and CT26 cells were placed into the plate at a ratio of 1:2 (macrophage/DC: CT26) after treatment with different nanodrugs for 48 h. Then, the cells were cocultured for another 48 h and harvested for flow cytometry.

Animals and cell lines

BALB/c male and BALB/c-nude mice (4–6 weeks of age) were purchased from JinWei Biotechnology Co., Ltd. (Guangzhou, China). All animals were maintained under specific pathogen-free conditions. The animal experimental procedures were approved by the Institutional Animal Care and Use Committee of Guangdong Pharmaceutical University and were performed in accordance with the guidelines. CT26 cells were purchased from ATCC via iCell Bioscience, Inc. (Shanghai. China).

CT26-bearing mouse model and treatment

To construct the CT26-bearing mouse model, CT26 cells (5 × 105 cells) were subcutaneously inoculated into the left flanks of male BALB/c mice. The diameters of the tumors were measured with caliper, and the tumor volume was calculated as 1/2a×b2.

For in vivo fluorescence imaging, DiR-labelled NP-I/P was injected into CT26-bearing mice at a DiR dose of 0.75 mg∙kg− 1 body weight via the tail vein when the tumor volume reached 200 mm3. A fluorescence imaging system (Carestream IS 4000, USA) was subsequently used to capture in vivo images at 1, 3, 6, 12, 24, and 48 h after DiR injection. Forty-eight hours later, the tumors and major organs (heart, liver, spleen, lung, and kidney) were harvested for in vitro imaging.

For the antitumor response study, the mice were randomly divided into five groups (n = 6 mice per group) when the tumor volume reached 100 mm3 and treated with 200 µL of saline, I/P, NP-I, NP-P, or NP-I/P containing panobinostat and/or epacadostat at doses of 5 mg∙kg and 2 mg∙kg− 1 body weight every three days. Tumor size and body weight were recorded every two days. After five cycles of treatment, the mice were sacrificed, and the tumors, major organs, and sera were collected for further experiments.

To evaluate the in vivo toxicity of the nanodrugs, the sera of CT26 tumor-bearing mice were utilized to assess liver function and renal function, and the tissues of major organs were stained with haematoxylin and eosin (H&E).

H&E staining

Formalin-fixed paraffin-embedded tissues were sectioned into slices at a thickness of 4 μm. The sections were deparaffinized and rehydrated after heated in 65 °C for 1 h. Then, the sections were stained with haematoxylin solution for 5 min and rinsed with water for 5 min. Next, the tissues were stained with eosin solution. After being sealed with neutral gum, the sections were observed under Nikon light microscope (Nikon, ECLIPSE Ti2-U, Japan).

Sirius red staining

Formalin-fixed paraffin-embedded tissues were sectioned into slices at a thickness of 4 μm. The sections were deparaffinized and rehydrated after heating in 65 °C for 1 h. Then, the sections were stained with Sirius red for 8 min and dehydrated with anhydrous ethanol. After being sealed with neutral gum, the sections were observed under a Nikon light microscope (Nikon, ECLIPSE Ti2-U, Japan).

PDX model and treatment

To establish the PDX mouse model, fresh tumor tissues from CRC patients were sliced into 3–5 mm-diameter pieces, and then, the tissues (defined as G0) were subcutaneously implanted in the left flank of BALB/c-nude mice (defined as G1). When the tumors in G1 mice reached 500 mm3, the tumors were harvested, sliced into 3–5 mm pieces and subcutaneously implanted in the left flank of another BALB/c-nude mouse to generate G2 mice, which were suitable for in vivo experiments.

For the antitumor efficacy study in PDX model, the mice were randomly divided into five groups (n = 6 mice per group) when the tumor volumes reached 100 mm3 and treated with 200 µL of saline, I/P, NP-I, NP-P, or NP-I/P containing panobinostat and/or epacadostat at doses 5 mg∙kg− 1 and 2 mg∙kg− 1 body weight every four days. Tumor size and body weight were recorded every two days. After five cycles of treatment, the mice were sacrificed, and the tumors were harvested for further experiments.

Human PDO culture

Human PDOs were cultured as described previously. Type I collagen gel solution was prepared according to the manufacturer’s instructions, added to 30 mm Transwells inserts containing membranes with 0.4 μm pores and solidified at 37 °C for 30 min. Fresh tumor tissues were minced into 125–500 mm3 fragments, washed twice with ADMEM/F12 (Invitrogen) containing 1X Normocin (InvivoGen), resuspended in 1 mL of type I collagen gel and placed into inner Transwells precoated with gel. The Transwells inserts containing the tumor tissues were subsequently placed into other culture dishes containing 1 mL of ADMEM/F12 culture medium supplemented with Wnt3a, RSPO1, or Noggin-conditioned media (L-WRN, ATCC) supplemented with HEPES (1 mM, Invitrogen), glutamine (1X, Invitrogen), gastrin (10 mM, Sigma), N-acetylcysteine (1 mM, Sigma), epidermal growth factor (50 ng/mL, PeproTech), IL-2 (6000 IU/mL, PeproTech), G-CSF (50 ng/mL, PeproTech) or GM-CSF (50 ng/mL, PeproTech). Nanodrugs were added to the culture medium and incubated for 5 days, after which the samples were collected for further experiments.

Drug phagocytosis in PDOs

For the drug phagocytosis experiment with the PDO model, NP-encapsulated coumarin was added to the PDO culture medium as described above. After incubation for 2, 4, 8, or 24 h, the samples were collected and made into frozen slices. The frozen slices were stained with DAPI and observed under a fluorescence microscope to assess whether the nanodrugs could be phagocytosed by PDO samples.

Statistical analysis

The GraphPad Prism 7 software program was used for statistical analysis of the experimental data, and the results are presented as the means ± standard deviations (SDs). Significant differences between two unpaired groups were determined by the log-rank test or Student’s t test. P < 0.05 was considered to indicate statistical significance. The significance level was indicated as follows: *P < 0.05, **P < 0.01, ***P < 0.001 and ****P < 0.0001.

Results

HDACs and IDO1 are promising targets for synergistic immunotherapy in colorectal cancer

According to The Cancer Genome Atlas (TCGA), HDACs (HDAC2, HDAC4, HDAC6, and HDAC8) display different expression patterns in different tumors and corresponding normal tissues (S-Fig. 2). We compared the expression of HDACs (HDAC2, HDAC4, HDAC6, and HDAC8) between MSS and MSI-H CRC via the R2 platform. The expression levels of these HDACs in MSS CRC were greater than those in MSI-H CRC (Fig. 2A). The microsatellite stability states of three tumor tissues from MSI-H and MSS CRC patients were confirmed by detecting MLH1, PMS2, MSH2, and MSH6 expression (S-Fig. 3). MSS CRC samples presented higher protein levels of HDAC2, HDAC4, HDAC6, and HDAC8 (Fig. 2B). Moreover, compared with normal tissues, the tumors contain more HDAC2, HDAC4, HDAC6, and HDAC8 (S-Fig. 4), indicating that HDACs may be therapeutic targets for CRC. We subsequently utilized panobinostat, an HDACi, to treat CT26 cells. Interestingly, IDO1 expression in CT26 cells was upregulated following panobinostat treatment (Fig. 2E). IDO1 is an immune checkpoint that is expressed at a much lower level in MSS CRC than in MSI-H CRC (Fig. 2F, G). IDO1 deficiency in CRC may contribute to the poor therapeutic effects of IDO1 inhibitors. Therefore, HDAC and IDO1 are promising targets for the synergistic immunotherapy of CRC.

Fig. 2
figure 2

Expression of HDAC and IDO1 in colorectal cancer. (A) TCGA data analysis of HDAC (HDAC2, HDAC4, HDAC6, and HDAC8) expression in MSS and MSI-H CRC samples (n = 77) (from the R2 platform). (B) Western blotting analysis of HDAC (HDAC2, HDAC4, HDAC6, and HDAC8) expression in MSS and MSI-H CRC samples (n = 3). (C, D) Western blotting analysis of HDACs (HDAC2, HDAC4, HDAC6, and HDAC8) expression in CRC tissues and normal colorectal tissues (n = 8). (E) Western blotting analysis of IDO1 in CT26 cells after treatment with panobinostat (n = 3). (F) TCGA data analysis of IDO1 expression in MSS and MSI-H CRC (n = 77) (from the R2 platform). (G) Immunohistochemical staining of IDO1 in MSS and MSI-H CRC samples (n = 3). Scale bar: 20 μm

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Preparation and characterization of polymeric nanoparticles

We used mPEG-PLC, which is capable of self-assembling into nanoparticles (NPs), as a carrier to prepare polymeric NPs (Fig. 3A). The half-maximal inhibitory concentration (IC50) values of the IDO1 inhibitor, epacadostat, and the HDACi, panobinostat, were determined via ELISA (S-Fig. 5A) and a CCK-8 assay (S-Fig. 5B). Panobinostat and epacadostat were coloaded into the NPs to obtain the nanodrug NP-I/P, as described in the experimental section. NP-encapsulated panobinostat (NP-P) and epacadostat (NP-I) were prepared as experimental controls. The hydrodynamic sizes of the NPs were measured via dynamic light scattering, and the diameter of NP-I/P was found to be 101.5 ± 0.3 nm (Fig. 3B). In addition, the stability of NP-I/P in a physiological environment was verified by monitoring the particle size in 10% foetal bovine serum containing PBS at 37℃ for a long duration (Fig. 3C). The spherical structures of the NPs were observed via TEM (Fig. 3D). NP-P and NP-I exhibited similar morphologies and hydrodynamic diameters (S-Fig. 6A-D).

NP-I/P inhibits tumor growth in vitro

To investigate the cytotoxicity of the nanodrugs, a CCK-8 assay was performed to assess the viability of CT26 cells treated with different concentrations of the nanodrugs. NP-I/P exhibited great cytotoxicity against CT26 cells. The half-maximal inhibitory concentrations of NP-I/P were 200 nM (in terms of the panobinostat dose) and 100 nM (in terms of the epacadostat dose), compared with 300 nM for NP-P (Fig. 3E). Both NP-I and NP-I/P restrained the production of kynurenine, whereas NP-P only slightly inhibited the production of kynurenine for the induction of IDO1 (Fig. 3F). We further investigated the effects of the NPs on CT26 cell apoptosis. After incubation with NP-I, NP-P or NP-I/P for 48 h in CT26 cells, cleaved caspase-3 expression increased markedly, and total caspase-3 expression remained unchanged (Fig. 3G). The apoptosis rates of the CT26 cells incubated with PBS, NP-I, NP-P, and NP-I/P were 6.0%, 15.8%, 38.2%, and 47.5%, respectively (Fig. 3H). The effects of the nanodrugs on the cell cycle were further evaluated. The percentage of the cells in the G2/M and S phases clearly decreased, whereas that of the cells in the G0/G1 phase increased after incubation with NP-P and NP-I/P, indicating that a portion of the cells were arrested at the G0/G1 phase by the HDACi rather than the IDO1i (Fig. 3I). According to live/dead cell fluorescence analysis, the group treated with NP-P or NP-I/P had more dead cells (red fluorescence) and fewer live cells (green fluorescence) than the groups treated with PBS or NP-I (Fig. 3J). Moreover, after a pretreatment with different nanodrugs, the migration, invasion, and colony formation abilities of the CT26 cells decreased to varying degrees (S-Fig. 7). NP-I/P exhibited the most robust inhibition of tumor growth and tumor-killing effects. These results indicate that the nanodrugs possess an anti-tumor cytotoxic effect and that a synergistic effect exists between panobinostat and epacadostat.

Fig. 3
figure 3

Nano-I/P characterization and enhanced antitumor efficacy in vitro. (A) Schematic diagram of nanodrug preparation. (B) Size distribution of NP-I/P detected by dynamic light scattering (n = 3). (C) Stability of NP-I/P in PBS containing 10% FBS (n = 3). (D) Transmission electron microscopy (TEM) imaging of NP-I/P. Scale bar, 100 nm. (E) Cytotoxicity of NP-I, NP-P, and NP-I/P in vitro (n = 3). (F) Concentration of kynurenic acid in the medium after treated with NP-I, NP-P, or NP-I/P detected by ELISA (n = 3). (G) Western blotting analysis of caspase-3 and cleaved caspase-3 in CT26 cells after treated with NP-I, NP-P, or NP-I/P (n = 3). (H) Apoptosis of CT26 cells in different groups detected by flow cytometry (n = 3). (I) Cell cycle distribution of CT26 cells in different groups (n = 3); (J) Representative immunofluorescence staining images of live (green) and dead (red) CT26 cells after various treatments (n = 3). Scale bar: 100 μm

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Codelivery of an HDACi and an IDO1i induces ICD and regulates immune cell function in vitro

ICD is a type of programmed cell death triggered by multiple antitumor treatments. In addition to releasing DAMPs, ICD can facilitate the maturation of DCs and the activation of innate and adaptive immune responses. We hypothesized that nanodrugs can induce the release of DAMPs and tumor antigens to elicit ICD. Therefore, we investigated whether these nanodrugs can induce the release of CRT, HSP70, HMGB1, and ATP. The surface expression of CRT and HSP70 on CT26 cells was assessed via immunofluorescence and flow cytometry. NP-I/P treatment induced CRT expression on the cell membrane (Fig. 4A, S-Fig. 8A, B). Similarly, HSP70 expression on the CT26 cell membrane was markedly increased according to LSCM and flow cytometry (Fig. 4B, S-Fig. 8C). The release of HMGB1 into the culture media was measured by ELISA, which revealed that HMGB1 secretion significantly increased after NP-I/P treatment (Fig. 4C). Moreover, the HMGB1 level in the cell lysate significantly decreased, as detected by western blotting (S-Fig. 8D). Compared with that of the controls, the ATP concentration in the culture supernatant of the CT26 cells treated with NP-I/P significantly increased (Fig. 4D). We further investigated the mechanism by which NP-I/P induces ICD. The PKC/MEK pathway contributes to various biological processes, including cell proliferation, migration, and motility [37]. The activation of the PKC/MEK pathway and upregulation of CRT were observed when CT26 cells were treated with NP-P or NP-I/P (Fig. 4E), and the CRT level and activation of the PKC/MEK pathway were restored after the addition of the PKC inhibitor, sotrastaurin (Fig. 4F). NP-P and NP-I/P may therefore trigger ICD, at least partially, via the PKC/MEK pathway. Next, we investigated whether the nanodrugs could regulate immune cell function in vitro. After pretreatment with the nanodrugs, CT26 cells were cocultured with DCs, CD8+ T cells isolated with magnetic beads (S-Fig. 9), and macrophages for 72 h. In the NP-I/P group, mature DCs (labelled with CD80+CD86+) accounted for 31.8% of the cells, and this percentage was greater than that in the control groups (Fig. 4G). In addition, the proliferation and ability of CD8+ T cells to secrete IFN-γ were significantly greater in the NP-I/P group than in the control group (Fig. 4H, I). In contrast, the percentage of M2-type macrophages noticeably decreased after a coculture of CT26 cells pretreated with NP-I/P (Fig. 4J). Moreover, IDO1 expression was positively correlated with the expression of MDSC markers, such as ANPEP, CD33, and CSF1R, in CRC (S-Fig. 10A). The MDSCs had higher IDO1 protein levels than the CT26 and RAW264.7 cells did (S-Fig. 10B). After treatment with the nanodrugs, the percentage of apoptotic MDSCs increased significantly (Fig. 3K), and Arg-1 was downregulated (Fig. 4L, S-Fig. 11), especially in the NP-I/P group. Furthermore, NP-I/P inhibited TGF-β secretion and tryptophan-kynurenine metabolism (Fig. 4M, N) and thus inhibited the function of MDSCs. These results indicate that the codelivery of an HDACi and an IDO1i can induce ICD, enhance the immune response, and relieve immunosuppression in vitro.

Fig. 4
figure 4

NP-I/P treatment induces ICD in cancer cells and regulates immune cell function in vitro. (A) Representative immunofluorescence images of CRT after various treatments (n = 3). Scale bar: 40 μm. (B) Representative flow cytometry analysis and quantification of HSP70 in CT26 tumors after treatment with nanodrugs (n = 3). (C) The release of ATP, as evaluated via ELISA (n = 3). D (D) The release of HMGB1, as evaluated via ELISA (n = 3). (E) Western blot analysis of CRT, PKC/MEK, and caspase-3 expression in response to treatment with NPs (n = 3). (F) Western blot analysis of CRT, PKC/MEK, and caspase-3 expression in response to treatment with NPs with or without PKCis (n = 3). (G) Representative flow cytometry analysis and quantification of the percentage of mature DCs (CD80+CD86+ cells) after coculture with CT26 cells pretreated with nanodrugs (n = 3). (H) Representative flow cytometry analysis and quantification of CSFE for CD8+ T cells after coculture with CT26 cells pretreated with nanodrugs (n = 3). (I) Representative flow cytometry analysis and quantification of the percentage of IFN-γ-positive CD8+ T cells after coculture with CT26 cells pretreated with nanodrugs (n = 3). (J) Representative flow cytometry analysis and quantification of the percentage of M2 TAMs after coculture with CT26 cells pretreated with nanodrugs (n = 3). (K) Representative flow cytometry analysis and quantification of the percentage of apoptotic MDSCs after treatment with nanodrugs (n = 3). (L) Representative flow cytometry analysis of Arg-1-positive MDSCs after treatment with nanodrugs (n = 3). (M) ELISA evaluation of TGF-β in MDSCs treated with nanodrugs (n = 3). N. ELISA evaluation of kynurenic acid released by MDSCs treated with nanodrugs (n = 3)

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IDO1i enhances the antitumor efficacy of the HDACi in subcutaneous CT26 tumor-bearing mouse models of MSS CRC

Fluorescence imaging was performed to evaluate the tumor-targeting capacity and distribution of NP-I/P in tumor-bearing mice. NP-I/P was labelled with DiR and injected into the mice via the tail vein. The fluorescence intensity gradually increased at the tumor site (Fig. 5A), and after 48 h, the tumor and major organs (heart, liver, spleen, kidneys, and lungs) were harvested and imaged. The fluorescence intensity of the tumor was much greater than that of the organs (Fig. 5B), indicating that NP-I/P effectively accumulated in the CT26 tumors in vivo and favour the regulation of the TME by the drugs. We then investigated the antitumor efficacy of the nanodrugs in a subcutaneous mouse model following the treatment schedule shown in Fig. 5C. The tumor volumes were assessed every 2 days (Fig. 5D), and the tumors were harvested at the end of the treatment (Fig. 5E). According to the changes of tumor volume during treatment (S-Fig. 12) and tumor weights (Fig. 5F), NP-I/P suppressed tumor growth. To evaluate the in vivo side effects of the nanodrugs, we analyzed the body weights of the mice (Fig. 5G) via H&E staining of the major organs (S-Fig. 13A). The results revealed no difference among the five groups. The serum alanine aminotransferase, aspartate aminotransferase, and serum creatinine (S-Fig. 13B) levels were also within their normal ranges after treatment in all groups. Western blot analysis revealed that the expression of IDO1 increased in the I/P, NP-P, and NP-I/P groups, which further confirmed that the HDACi can upregulate IDO1 in tumors (Fig. 5H). Furthermore, an in vitro study revealed that the HDACi may upregulate IDO1 expression via the IFN-JAK2/STAT1-IRF1 pathway (S-Fig. 14), suggesting that the addition of the IDO1i can enhance the effects of the HADCi. To assess the effects of the nanodrugs on tumor proliferation, apoptosis, and angiogenesis, tumor sections were assessed for Ki67, cleaved caspase-3, and CD31expression. The tumor tissues from the NP-I/P group presented the lowest proliferation rate, highest apoptosis ratio, and lowest angiogenesis levels (Fig. 5I). Owing to the ability of the nanodrug to accumulate in tumors, NP-I and NP-P had stronger antitumor effects than did I/P. Treatment with NP-I/P, which contains both the IDO1i and the HDACi, further enhanced the therapeutic effects, revealing the synergistic antitumor effects of the IDO1i and HDACi.

Fig. 5
figure 5

Antitumor effects of nanodrugs in a CT26 tumor– bearing mouse model. (A) In vivo time-dependent fluorescence images after intravenous injection of NP-I/P and quantification of fluorescence intensity in tumors (n = 6). (B) In vitro fluorescence images and quantification of tumor and major organs after intravenous injection of NP-I/P for 48 h (n = 6). (C) Timeline for the treatment of CT26 tumor– bearing mice. (D) Changes in tumor volume were measured for 22 days after tumor inoculation(n = 6). (E) Photographs of tumors after different treatments(n = 6). (F) Ex vivo tumor weights of tumors after different treatments (n = 6). (G) Body weights of the CT26 tumor-bearing mice after different treatments (n = 6). (H) Western blot analysis of CRT, IDO1, and caspase-3 expression in tumor tissues after different treatments (n = 6). (I) Immunohistochemical staining of ki67, caspase-3, and CD31 in tumor tissues (n = 6). Scale bar: 40 μm

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Inhibition of HDAC and IDO1 boosts the antitumor immune response in vivo

We evaluated antitumor immune responses in CT26 tumor–bearing mice after nanodrug treatment. First, we examined DAMPs release in vivo via immunofluorescence to detect CRT and HMGB1 expression in tumors. CRT expression on the tumor cell membrane was greater in the NP-I/P group than in the other groups (Fig. 6A). In parallel, HMGB1 accumulated in the tumor stroma after treatment with NP-I/P but was localized in the tumor cells in the control group. These findings indicated that NP-I/P can effectively induce the release of DAMPs in vivo (Fig. 6B). We then assessed the phenotype and function of immune cells via flow cytometry (S-Figs. 15 and 16). More initial DCs were converted into mature DCs (CD80+CD86+ DCs) in the NP-I/P group (Fig. 6C). Mature DCs present tumor antigens to T cells, thereby enhancing T-cell antitumor immunity. The percentage of CD3+CD8+ T cells within the tumors increased (Fig. 6D, E), whereas that of CD3+CD4+ T cells decreased. Thus, the CD8/CD4 ratio increased in the NP-I/P group (Fig. 6F). In addition, the number of Treg cells (Foxp3+CD25+CD4+ T cells) significantly decreased (Fig. 6G). We then used immunofluorescence to confirm the distribution of CD8+ and CD4+ T cells in the tumors after the treatments (Fig. 6H). Compared with those in the control groups, the number of CD8+ effector T cells (IFN-γ+CD8+T and GZMB+CD8+T cells) in the tumors also increased (Fig. 6I, J). Immune memory plays a vital role in antitumor immunity regarding tumor-specific antigens. To assess the immune memory effects of the nanodrugs, we assessed the changes in the number of effector memory T (Tem) cells in the spleen after therapy via flow cytometry and found that the NP-I/P group contained a much greater percentage of Tem cells in the spleen (Fig. 6K). MDSCs and TAMs are central immunosuppressive cells in the tumor environment and promote tumor invasion. The flow cytometry results revealed that NP-I/P decreased the abundance of MDSCs (Fig. 6L) and M2 TAMs(F4/80+CD206+) in tumors (Fig. 6M), indicating that NP-I/P can reverse immunosuppression, which was confirmed by immunofluorescence (Fig. 6N). These results collectively demonstrate that the inhibition of HDAC and IDO1 boosts the antitumor immune response in vivo.

Fig. 6
figure 6

Synergetistic antitumor immune response of NP-I/P in vivo. (A) Representative immunofluorescence staining images of CRT expression in tumor tissues after various treatments (n = 6). Scale bar: 40 μm. (B) Representative immunofluorescence staining images of HMGB1 expression in tumor tissues after various treatments (n = 6). Scale bar: 40 μm. (C) Quantification of matured DCs (CD80+CD86+ cells) in tumor tissues subjected to various treatments (n = 6). D, E. Quantification of CD8+ T (D) and CD4+ T (E) cells in tumor tissues after various treatments (n = 6). (F) The ratio of CD8/CD4 in tumor tissues after various treatments (n = 6). (G) Quantification of Tregs in tumor tissues after various treatments (n = 6). (H) Representative immunofluorescence staining images of CD4 (green) and CD8 (red) in tumor tissues (n = 6). Scale bar: 40 μm. (I, J) Flow cytometry analysis results of the percentage of IFN-γ +CD8+ T cells (I) and GMZB+CD8+T cells isolated from the tumors (n = 6); (K) Flow cytometry analysis results of the percentage of effector memory T (TEM) cells in spleen after different treatments (n = 6). (L, M) Flow cytometry analysis results of the percentage of MDSCs (L) and M2 TAMs (M) isolated from tumors (n = 6). (N) Representative immunofluorescence staining images of CD206 (green) and Gr-1(red) expression in tumor tissues (n = 6). Scale bar: 40 μm

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The HDACi and IDO1i induces synergistic antitumor effects in PDX models of MSS CRC

To better simulate the tumor characteristics of CRC, we constructed a PDX model using BALB/c nude mice, as described in our previous study [33]. The PDX model was established by subcutaneously implanting CRC tumor tissues (generation 0; G0) in the left flank of BALB/c nude mice (generation 1; G1). When passaged to G2 (generation 2) (Fig. 7A), the tumors were harvested for H&E staining, Sirius red staining, and an evaluation of the microsatellite state via immunofluorescence. The G1 and G2 tumors maintained similar tumor characteristics and collagen contents as those in G0 tumors did (S-Fig. 17). In addition, the G1 and G2 tumors maintained the same MSS state as the G0 tumors did (S-Fig. 18); thus, the model was used for subsequent experiments. Fluorescence imaging was used to evaluate the tumor-targeting capacity and distribution of NP-I/P in the PDX model as in the subcutaneous tumor model. The fluorescence intensity gradually increased at the tumor site within 48 h (Fig. 7B), after which, the tumor and major organs were harvested. The fluorescence intensity of the tumor was greater than that of the heart, spleen, lungs, and kidneys but lower than that of the liver (S-Fig. 19A). The fluorescence imaging results suggested that the nanodrugs could also accumulate in tumors in the PDX model. The mice were intravenously treated every 4 days for five rounds after the tumor reached 100mm3 with saline, control NPs, or NP-I/P (S-Fig. 19B). Combination therapy with IDO1i and HDACi in NPs achieved excellent antitumor efficacy in a CRC PDX model after five treatment cycles. During the duration of therapy, the increase in tumor volumes gradually slowed in the NP-I/P group, and in the control group, the tumor sustained rapid growth (S-Fig. 19C). At the end of the treatment period, the tumors were harvested (Fig. 7D). The final tumor weight in the NP-I/P group was markedly lower than that in the control groups (Fig. 7E), but the body weight was not significantly different among the groups (S-Fig. 20A). The tumor tissues were subsequently subjected to pathological evaluation. H&E staining and Sirius red staining revealed that the tumor and collagen levels decreased after treatment with the nanodrugs (S-Fig. 20B). Immunohistochemistry and immunofluorescence staining revealed that the density of cleaved caspase-3 staining increased in the NP-I/P group (Fig. 7F). In contrast, the Ki67 and CD31 levels were attenuated in the NP-I/P group (Fig. 7G), indicating that NP-I/P inhibited tumor growth and angiogenesis in the PDX model. Finally, we focused on the effects of NP-I/P on ICD and immune regulation in the PDX model. NP-I/P induced CRT expression on the tumor cell surfaces (Fig. 6H), and HMGB1 was released into the stroma (Fig. 7I). NP-I/P also triggered ICD via the PKC-MEK pathway (Fig. 7J), which was consistent with our previous in vitro results (Fig. 4E, F). Moreover, NP-I/P suppressed the expression of TGF-β on tumor cells (Fig. 7K), and NP-P and NP-I/P encapsulated panobinostat induced IDO1 expression (Fig. 7L), which was also consistent with our previous results in vitro and in the CT26 mouse model. However, we found that the PDX model retained few humanized immune cells; therefore, we investigated whether the nanodrugs can regulate mouse immune cells. Interestingly, the percentages of MDSCs and TAMs in the mice were markedly reduced after treatment with NP-I/P (S-Fig. 21). These results indicate that the HDACi and IDO1i induced synergistic antitumor effects in the PDX model of MSS CRC. Other models are needed to further evaluate the antitumor immune efficacy of these nanodrugs.

Fig. 7
figure 7

The HDACi and IDO1i induced synergetic antitumor effects in PDX models. (A) Patient-derived xenograft (PDX) models were generated by implanting CRC tissue in mice and transplanting G1 tumor tissue into a larger number of mice with G2 tumors. (B) In vivo time-dependent fluorescence images after intravenous injection of NP-I/P and quantification of fluorescence intensity in tumors(n = 3). (C) Changes in tumor volume were measured for 32 days after tumor inoculation(n = 6). (D) Photographs of tumors after different treatments(n = 6). (E) Ex vivo tumor weight of tumors after different treatments (n = 6). (F) Immunohistochemical staining of cleaved caspase-3 and ki67 in tumor tissues (n = 6). Scale bar: 40 μm. (G) Representative immunofluorescence staining images of CD31 in tumor tissues after various treatments (n = 6). Scale bar: 40 μm. (H) Representative immunofluorescence staining images of CRT in tumor tissues after various treatments (n = 6). Scale bar: 40 μm. (I) Representative immunofluorescence staining images of HMGB1 in tumor tissues after various treatments (n = 6). Scale bar: 40 μm. (J) Western blot analysis of CRT and PKC/MEK expression in tumor tissues after various treatments (n = 6). (K) Immunohistochemical staining of TGF-β in tumor tissues (n = 6). Scale bar: 40 μm. (L) Western blot analysis of IDO1 expression in tumor tissues after various treatments (n = 6)

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Enhanced synergistic antitumor immune response with the combination of the HDACi and the IDO1i in PDO models of MSS CRC

In the absence of humanized immune cells, we could not clarify the effects of the nanodrugs on immune regulation in the PDX model. Thus, we constructed a PDO model of MSS CRC to further investigate the antitumor immune efficacy of the nanodrugs (Fig. 8A). After being cultured for 5 days, the PDO tissues were collected and stained for α-SMA, E-cadherin, β-catenin, CDX2, CK20, and CK7. The architecture, epithelial, and stromal compartments of the original tumor were preserved in the PDOs, and the histopathological characteristics of the PDOs were similar to those of the tumor tissues (Fig. 8B). We further investigated whether PDO tissues can phagocytose nanodrugs in vitro. Coumarin accumulated in the tumor in a time-dependent manner and was phagocytosed by tumor cells (Fig. 8C) and monocytes (S-Fig. 22A) in 24 h rather than T cells (S-Fig. 22B), which indicated that nanodrugs can be phagocytosed by tumor tissues in PDOs. Various nanodrugs were then added to the culture system and incubated for 5 days. Tumor cell proliferation was notably suppressed in the NP-I/P group (Fig. 8D), whereas NP-I/P facilitated tumor cell death (Fig. 8E). These results suggest that NP-I/P treatment induced synergistic antitumor effects in PDO models.

We then assessed the status of immune cells in these organoids. The tumor organoids retained the stability of immune cells when cultured in vitro (S-Fig. 23). Significantly increased CD8 expression was observed in the NP-I/P-treated tumor organoids, which produced large amounts of GZMB (Fig. 8F). Furthermore, the number of TAMs in the tumor organoids decreased after treatment with NP-I/P, and M1-type macrophages expansion was observed (Fig. 8G). Treatment with NP-I/P markedly decreased the in situ expression of arginase 1 (ARG-1), a suppressive molecule of MDSCs, in CD11b+ myeloid cells (Fig. 8H). These data indicate that the combination of the HDACi and IDO1i can synergistically enhance the antitumor immune response in PDO models of MSS CRC.

Fig. 8
figure 8

NP-I/P induced a synergetic antitumor immune response in PDO models. (A) Schematic diagram of PDO culture and treatment in vitro. (B) Immunofluorescence analysis of marker expression (α-SMA, E-cadherin, β-catenin, CDX2, CK20, and CK7) in tumors and patient-derived organoids. Scale bar, 40 μm. (C) Phagocytosis of nanodrug of PDOs in vitro. Scale bar, 40 μm. (D) Representative immunofluorescence images of ki67 in PDOs after various treatments (n = 6). Scale bar, 40 μm. (E) Representative immunofluorescence images of TUNEL staining in PDOs after various treatments (n = 6). Scale bar, 40 μm. (F) Representative immunofluorescence staining images of CD8+ T cells in PDOs after various treatments (n = 6). Scale bar, 40 μm. (G) Representative immunofluorescence images of CD86/CD206 in PDOs after various treatments (n = 6). Scale bar, 40 μm. (H) Representative immunofluorescence images of CD11b/Arg-1 in PDOs after various treatments (n = 6). Scale bar, 40 μm

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Discussion

Immune checkpoints such as PD-1/PD-L1, CTLA-4, TIGIT, and IDO1, have been identified and investigated in various cancers [10]. Immune checkpoint blockade (ICB) has been used as a first-line therapy for the treatment of cancers, including non-small cell lung cancer and melanoma. Immunotherapy has been shown to be a promising and powerful therapy for cancers [4, 5]. However, because of the immunosuppressive microenvironment and lack of effector immune cells, ICB results in poor antitumor responses in some cancers, especially MSS/pMMR CRC [7]. MSS/pMMR CRC is considered a “cold tumor” and possesses an “immune desert” TME, resulting in a poor response to immunotherapy [38]. Chemotherapy, radiotherapy, and photodynamic therapy have been regarded as potential strategies to improve the response of ICB in CRC [39,40,41], owing to the induction of tumor cell ICD and the triggering of antitumor immunity, severe side effects, and harsh service conditions that limit the application of these treatments. Thus, developing a novel strategy that induces an effective antitumor immune response and has a good safety profile is crucial for the treatment of MSS/pMMR CRC.

In this study, we first assessed the expression of HDACs and IDO1 in CRC, which was not previously confirmed, especially in MSS and MSI-H CRC. HDACs are targets for antitumor treatment in multiple cancers [24], and we found that HDACs are overexpressed in MSS/pMMR CRC. Thus, HDACs may be therapeutic targets for CRC. The pan- HDACi, panobinostat, which has been shown to exert antitumor effects [42], was selected as one of the two drugs for the composite nanomedicine in this study. Consistent with the finding of a previous study [43], we found that IDO1 was expressed at low levels in MSS CRC, but that panobinostat treatment facilitated the expression of IDO1 in tumor cells via the JAK2/STAT1/IRF1 pathway. We hypothesized that combining panobinostat with an IDO1 inhibitor (epacadostat) may enhance the antitumor effect and immune response to these two drugs in MSS CRC.

The short half-life, quick-forming tolerance, and pronounced side effects of cytotoxic drugs and ICBs are major factors that limit their application in cancer therapy. These limitations also exist for panobinostat and epacadostat. Nanomedicines were the first treatment type to enable the administration of antitumor drugs to solid tumors by passively targeting and reducing access to healthy tissues [44]. The slow and controlled release of nanomedicines improves the effects of drugs and reduces the required drug dose [45]. Therefore, we developed an NP platform for delivering panobinostat and epacadostat to tumor and immunosuppressive cells in CRC models. Our results demonstrated that NP-I/P successfully accumulated in tumor tissues and caused fewer side effects in vivo than the naked drugs, indicating that NP-I/P is safe for treating tumors.

Panobinostat suppresses tumors by inducing apoptosis and autophagy, arresting the cell cycle, and inhibiting angiogenesis [29]. In addition, the HDACi induces the release of DAMPs, thereby triggering tumor ICD [30]. Epacadostat decreased kynurenine production, relieved the immunosuppression of microenvironment, and boosted the antitumor immunity of immune cells such as CD8+ T cells. Our results demonstrate that NP-I/P successfully triggers antitumor immune responses by inducing the release of DAMPs and ICD via the PKC‒MEK pathway. Moreover, the inhibition of IDO1 by NP-I/P suppressed the immunosuppressive phenotype of the tumor microenvironment and facilitated effector cell activation. NP-I/P directly induced tumor cell death and suppressed tumor neovascularization, but also triggered immune response, enhanced CD8+ T cell function, and decreased the number of immunosuppressive cells, including Tregs, MDSCs, and TAMs. MDSCs and TAMs are the principal immunosuppressive cells that form an immunosuppressive microenvironment in tumors [46]. A previous study and the present study demonstrated that the expression of IDO1 in MDSCs and TAMs is much greater. NP-I/P can be phagocytosed by MDSCs and TAMs to further inhibit their immunosuppressive function. In addition, we compared the antitumor efficacy of naked panobinostat/ epacadostat, NP-I, NP-P, and NP-I/P in mouse models of CRC and validated the potent antitumor efficacy of the combination strategy and the efficiency of the nanomedicine. NP-I/P successfully remodelled the TME and shifted the tumor from a “cold tumor” phenotype to a “hot tumor” phenotype.

Mouse tumor models have been used to simulate human systems and preliminarily validate the effects of drugs in vivo [47]. This study confirmed the antitumor effect and immune modulation of NP-I/P in mouse tumor models. However, further studies are needed to verify the effects of these drugs on mouse tumors, and these should be further validated in human tumor models. PDX models are suitable for exploring the antitumor efficacy in human tumors, and can help in the evaluation of the physiological and pathological processes of human tumor cells [48]. By using PDX models, we confirmed that NP-I/P accumulates in CRC tissue, thereby inducing CRC cell apoptosis and triggering ICD in CRC cells. PDOs can be cultured efficiently using human tumor tissues and accurately simulate human tumors [49]. The behaviour of malignant cells and their interactions with the microenvironment can also be investigated in PDOs [50]. In the present study, we used human MSS/pMMR CRC PDO models to explore the effects of NP-I/P on immune modulation. NP-I/P was shown to accumulate in PDO tissues and regulate the quantity and function of immune cells. These results support the potential of treating MSS/pMMR CRC with NP-I/P.

Notably, this study focused on panobinostat and epacadostat, so it is not clear whether the codelivery of other HDACis or IDO1is might have similar effects. Therefore, investigating other combinatorial strategies for treating MSS/pMMR CRC is of interest. Although NP-I/P altered the immune microenvironment in CDX, PDX, and PDO models, further investigation is needed to elucidate the molecular mechanisms underlying its immune regulation. Additionally, the PDX model used in this study did not retain the original human immune cells, and a humanized mouse PDX model should be constructed for further investigation.

In summary, our study provides a potent strategy for eliciting an antitumor immune response in MSS/pMMR CRC by codelivering HDACs and IDO1 inhibitors via self-assembled NPs. This effect was validated in patient-derived tumor models, highlighting that NP-I/P may improve the clinical treatment of patients with MSS/pMMR CRC.

Data availability

No datasets were generated or analysed during the current study.

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Acknowledgements

Not applicable.

Funding

This work was supported in part by grants from Guangdong Basic and Applied Basic Research Foundation (2022A1515111195), City-College-Institute allied Projects of Guangzhou City (SL2023A03J00413), Five Five Engineering Projects of the Third Affiliated Hospital of Sun Yat-sen University (2023WW502), the Science and Technology Planning Project of Guangdong Province (2021A0505030020).

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Author notes

  1. Rongpu Liang and Dongbing Ding contributed equally to this work.

Authors and Affiliations

  1. Department of Gastrointestinal Surgery, The Third Affiliated Hospital of Sun Yat-sen University, Guangzhou, Guangdong, 510630, P.R. China

    Rongpu Liang, Dongbing Ding, Yiquan Li, Shengxin Huang, Jiannan Ren & Bo Wei

  2. Department of Gastrointestinal Surgery, Lingnan Hospital, The Third Affiliated Hospital of Sun Yat-sen University, Guangzhou, P.R. China

    Rongpu Liang, Dongbing Ding, He Huang & Bo Wei

  3. Central Laboratory, The Third Affiliated Hospital of Sun Yat-sen University, Guangzhou, P.R. China

    Tianyun Lan

  4. Center of Electronic and Light Microscopy, Institute of Physiology of the National Academy of Sciences of Belarus, Minsk, Belarus

    Svetlana Ryabtseva

  5. Department of Gastrointestinal Surgery, Lingnan Hospital, The Third Affiliated Hospital of Sun Yat-sen University, Guangzhou, Guangdong, 510630, P.R. China

    He Huang

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  1. Rongpu Liang
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Contributions

R.L., D.D., H.H, and B.W. contributed to conceived and designed the experiments; R.L., D.D., H.H, and B.W. were involved in analyzed the data; R.L., D.D., H.H, T.L., Y.L., S. R., S.H., and J.R. contributed to performed the experiments and writing — original draft; B.W. was involved in writing — review and editing and funding acquisition; all authors finally approved the manuscript.

Corresponding authors

Correspondence to He Huang or Bo Wei.

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Liang, R., Ding, D., Li, Y. et al. HDACi combination therapy with IDO1i remodels the tumor microenvironment and boosts antitumor efficacy in colorectal cancer with microsatellite stability. J Nanobiotechnol 22, 753 (2024). https://doi.org/10.1186/s12951-024-02936-0

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Journal of Nanobiotechnology

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