This study downloaded the uniformly standardized pan-cancer dataset from the University of California, Santa Cruz Genomic (UCSC) database and extracted ITPR2 expression data for each sample. Acute myeloid leukemia-M3 (AML-M3) samples were excluded, and all expression data were transformed to log2(x + 1) form. Differences in ITPR2 expression between samples were analyzed using R software (version 4.2.1, R Foundation for Statistical Computing, Vienna, Austria) [10]. The association between ITPR2 level and OS was investigated using data from the Gene Expression Omnibus (GEO) database (accession No: GSE12417; n = 242), with the median ITPR2 level used as the threshold for grouping.

Bone marrow (BM) samples were obtained from 82 newly diagnosed AML patients and 30 normal donors at the First Affiliated Hospital of Guangxi Medical University, China, from 2013 to 2021. The diagnosis of AML was confirmed according to the World Health Organization (WHO) classification system. Ethical approval for this research was granted by the Ethical Review Commission of the First Affiliated Hospital of Guangxi Medical University, Nanning, China (Approval code: 2025-E0668).

Bone marrow mononuclear cells (BM-MNCs) were isolated by centrifugation, from which total RNA was extracted. RNA purity was assessed via spectrophotometer, with a 260/280 nm ratio greater than 1.8 indicating adequate purity. The total RNA was then converted into complementary DNA by reverse transcription. Human $\beta$-actin serves as a reference gene to normalize. The relative levels of expression were calculated using the 2^{-Δ⁢Δ⁢Ct} method. The primer sequences were as follows: $\beta$-actin forward 5^′-GTGGCCGAGGACTTTGATTG-3^′ and reverse 5^′-CCTGTAACAACGCATCTCATATT-3^′, ITPR2 forward 5^′-TGCGCCAATCAGCTACTTCT-3^′ and reverse 5^′-TCAGGATTAAGCTCTGCAGCTA-3^′.

ITPR2 expression data were extracted from The Cancer Genome Atlas–Acute Myeloid Leukemia (TCGA-LAML) (non-M3) database. The samples were divided into high and low expression groups based on the median ITPR2 level. Differential expression analysis between the two groups was performed using the limma package, with differentially expressed genes (DEGs) judgement criteria of $|$log2FC$|$ $>$1 and p.adj 0.05. The enrichment analyses were subsequently performed on the R clusterProfiler package [11]. DEGs with p.adj 0.05 were selected for differential expression analysis. The c2.all.v2022.1.Hs.symbols.gmt gene set (a standardized, curated gene set file from the molecular signatures (MSigDB), specifically designed for functional enrichment analysis in human [Homo sapiens] molecular biology and biomedical research) were obtained from the MSigDB database for Gene Set Enrichment Analysis (GSEA) [12]. The enriched pathway results met the thresholds of p.adj 0.05 and a q 0.25. The Estimation of Stromal and Immune cells in Malignant Tumor tissues using Expression data (ESTIMATE) algorithm was applied to calculate the immune infiltration score of the AML samples. The single sample gene set enrichment analysis (ssGSEA) algorithm was applied to determine the association between ITPR2 level and the immune infiltration levels [13]. Finally, Pearson correlation analysis was conduct to clear the association between ITPR2 level and 47 immune checkpoint-related genes [14]. The analysis results were visualized with R ggplot2 package [15].

Tohoku Hospital Pediatrics-1 (THP-1) cells were acquired from the Shanghai Cell Bank (https://www.cellbank.org.cn). THP-1 cells were cultured in RPMI-1640 (Gibco, NY, USA) with 10% fetal bovine serum (FBS) (Biological Industries, Cromwell, CT, USA) and 1% penicillin-streptomycin at 37 °C. All cell lines were validated by STR profiling and tested negative for mycoplasma.

siRNA was used to silence the ITPR2 gene. The plasmid construct was synthesized and generated by Guangzhou Ribobio Co., Ltd. (a commercial biotechnology company specializing in nucleic acid synthesis and vector construction). The interfering sequences targeting the ITPR2 gene were as follows: siRNA-ITPR2-1: 5^′-CAACGAAATTAGCGAGAGA-3^′, siRNA-ITPR2-2: 5^′-CCCTTAGCCTACCACATCA-3^′, siRNA-ITPR2-3: 5^′-GTGGCGCTTTCATGTCGAA-3^′. THP-1 cells were seeded in 6-well plates and cultured until 50% confluency. Subsequently, transfection of 20 nM negative control siRNA (si-NC) with a scrambled sequence that does not target any human genes or ITPR2-specific siRNA into THP-1 cells was performed using the CALNP™ RNAi in vitro (Beijing D-Nano Therapeutics, Beijing, China) reagent at 37 °C for 24 h. Subsequently, at other indicated time points, the cells were harvested and stored.

Western Blot (WB) cellular proteins were lysed using radio-immunoprecipitation assay (RIPA) buffer and centrifuged for 5 minutes at 4 °C. Equal amounts of protein (40 µg) were loaded into each lane and separated by 10% SDS-polyacrylamide gel electrophoresis (SDS/PAGE). The proteins were then electrotransferred to a polyvinylidene fluoride (PVDF) membrane. The membranes were blocked via 8% nonfat milk for 1 h and overnight incubated with primary antibodies at 4 °C. The specific primary antibodies used in this study were: anti-ITPR2 (Affinity, CAT#DF13336, dilution ratio 1:1000), anti-Cyclin D1 (Proteintech, CAT#26939-1-AP, dilution ratio 1:2000), anti-Cyclin E1 (Proteintech, CAT#11554-1-AP, dilution ratio 1:2000), anti-CDK6 (Proteintech, CAT#14052-1-AP, dilution ratio 1:1000), anti-Bcl-2 (Proteintech, CAT#12789-1-AP, dilution ratio 1:4000), anti-Bax (Proteintech, CAT#50599-2-Ig, dilution ratio 1:5000), and anti-actin (Abcam, CAT#ab8226, dilution ratio 1:2000). After washing via phosphate-buffered saline (PBS), the membranes were incubated for 1 h with corresponding secondary antibodies (Anti-Rabbit IgG (Zenbio, CAT#511203, dilution ratio 1:5000) Anti-Mouse IgG (Zenbio, CAT#511103, dilution ratio 1:4000)) in a blocking solution. Chemiluminescence was detected using a kit from chemiluminescence detection kit (Bio-Rad) [16] with $\beta$-actin serves as a loading control. ImageJ software (v1.8, National Institutes of Health, Bethesda, MD, USA) was applied to determine the band intensity.

CCK-8 (NCM Biotech, Suzhou, Jiangsu, China) was utilized to evaluate cell viability. Ten microliters of CCK-8 were given each well and cultured for 2 h at 37 °C. The cells absorbance were detected with a spectrophotometer (Molecular Devices, San Jose, CA, USA) at 0, 24, 48, 72 h post-seeding.

Cell Cycle Analysis Kit (Beyotime, Shanghai, China) was applied to conduct the cell cycle assessment. Cells were subjected to two rounds of washing with PBS. After that fixed them in ethanol at 4 °C for 12 hours. Subsequently, centrifuged at 800 rpm for 5 minutes, then stained with propidium iodide (PI) and RNase A, and incubated at 37 °C for 30 minutes without light. The stained cells were detected via the CytoFLEX flow cytometer (Beckman Coulter, Brea, CA, USA). G0/G1, S, and G2/M phase distribution were measured using FlowJo software (v10.8.1, FlowJo LLC, Ashland, OR, USA).

The Annexin V-Fluorescein Isothiocyanate (FITC) Apoptosis Detection Kit (Beyotime) was utilized to measure cellular apoptosis following the relevant interpretion. Cells from various groups were plated in 6-well plates, then collected and washed with PBS. Next, 195 µL of Annexin V-FITC binding solution was used to resuspend them. For each tube, 5 µL of Annexin V-FITC and 10 µL of PI were added, then cultured for 20–25 minutes without light. Apoptosis rates of cells from each group were evaluated by flow cytometry. Additionally, late apoptosis was evaluated via labeling DNA fragments of apoptotic cells with a Terminal Deoxynucleotidyl Transferase dUTP Nick-End Labeling (TUNEL) Assay Kit (Elabscience, Wuhan, China). Following fixation, permeabilization, equilibration, labeling, and washing per the instructions, detected with flow cytometer. TUNEL-positive cells were quantified with FlowJo software (v10.8.1).

Fluorescent indicators Fluo-4 AM (Beyotime) and Rhod-2 AM (Yesen, Shanghai, China) were employed to determine the levels of cytoplasmic Ca²⁺ and mitochondrial Ca²⁺, respectively. THP-1 cells were prepared as single-cell suspensions and loaded with 5 µM Fluo-4 AM and 5 µM Rhod-2 AM by incubation at 37 °C for 30 minutes. Then, the cells were washed with PBS to eliminate any extracellular Fluo-4 AM and Rhod-2 AM. The cytoplasmic and mitochondrial Ca²⁺ concentrations were then simultaneously measured by CytoFLEX flow cytometry (Beckman Coulter).

The MMP was assessed via flow cytometry with JC-1 staining, following the manufacturer’s instructions (C2006, Beyotime) [17]. Cells were collected and resuspended in 0.5 mL of cell culture medium and then incubated with 0.5 mL of a JC-1 staining solution in the dark at 37 °C for 20 minutes. Then, the cells were harvested, washed three times with JC-1 staining buffer, and resuspended in JC-1 staining buffer. The red/green fluorescence of JC-1-labelled cells was detected using a CytoFLEX flow cytometry (Beckman Coulter). The difference in MMP were determined by the color transformation of the fluorescence.

Intracellular ROS levels were detected with a DCFH-DA probe (Beyotime). The cells were treated with serum-free media containing 10 µM DCFH-DA probe and incubated at 37 °C in the dark for 30 minutes, with gentle agitation every 5 minutes. After incubation, the cell pellets were collected, washed with PBS, and resuspended in PBS for CytoFLEX flow cytometry (Beckman Coulter) detection. The resulting green fluorescence was measured and the mean fluorescence intensity (MFI) serves as an indicator of the relative intracellular ROS levels.

A Genomic DNA Mini Preparation Kit (Beyotime) was used to isolate total DNA from treated cells. Reactions for RT-qPCR were performed with Real Star Fast SYBR qPCR (Genstar, Shenzhen, China). Human $\beta$-actin was used as a reference gene to normalize. The primer sequences were as follows: $\beta$-actin forward 5^′-GTGGCCGAGGACTTTGATTG-3^′ and reverse 5^′-CCTGTAACAACGCATCTCATATT-3^′, mtND1 forward 5^′-CCCTAAAACCCGCCACATCT-3^′ and reverse 5^′-GAGCGATGGTGAGAGCTAAGGT-3^′.

Cells were seeded in the medium containing varying concentrations of 2-Aminoethyl diphenylborinate (2-APB) or dimethyl sulfoxide (DMSO) control. After a specified incubation interval, 10 µL of CCK-8 reagent was added, and the plates were cultured for viability of the cells was determined according to this formula: cell viability rate = [(As–Ab)/(Ac–Ab)] $\times$ 100. The half maximal inhibitory concentration (IC50) was calculated using regression analysis with GraphPad Prism 8.0 software (GraphPad Software, Inc., San Diego, CA, USA) based on the cell viability data.

Statistical analysis were performed on SPSS 25.0 (IBM Corp., Armonk, NY, USA) and GraphPad Prism 8.0 software. Each in vitro experiment was replicated a minimum of three times for every sample, repeat, group, and condition. The t-test was used to judge statistical variances in quantitative datasets. For nonparametric data, the Mann-Whitney U test was implemented. Correlations between two parameters were determined via the Pearson’s analysis. The criteria of p 0.05 was set to define statistical significance. Survival analyses were depicted via Kaplan-Meier plots, with log-rank tests for the significance in survival distributions.

Given the potential critical role of ITPR2 in AML, we first investigated the expression profiles of ITPR2 across malignancies. This research collected a harmonized pan-cancer dataset from the UCSC database, which revealed that ITPR2 expression exhibits significant variations among multiple tumor types (Fig. 1A). To explore the clinical significance of ITPR2 in AML prognosis, the gene expression and clinical data of 242 non-M3 AML patients from the GSE12417 dataset were downloaded. Kaplan–Meier analysis revealed that high ITPR2 level was associated to significantly shorter OS (Fig. 1B)

Fig. 1.

ITPR2 expression was increased in public databases vs. normal group and associated with prognosis. (A) Expression of ITPR2 in pan-cancer. (B) OS of GSE12417 patients (n = 242) stratified by high vs. low ITPR2 expression. * p 0.05, *** p 0.001, **** p 0.0001; ·, indicates 0.5 $\le$ p 0.1, and -, indicates p $\ge$ 0.1. ITPR2, Inositol 1,4,5-trisphosphate receptor type 2; OS, Overall survival.

To validate the results, BM samples and clinical data from 82 non-M3 AML patients and 30 healthy donors were collected. RT-qPCR analysis revealed significantly higher ITPR2 expression in AML patients than in controls (Fig. 2A). Receiver operating characteristic (ROC) curve analysis revealed an area under the curve (AUC) of 0.906 (95% CI: 0.851–0.961, p 0.001, Fig. 2B) for distinguishing AML patients from controls. To assess the clinical relevance of ITPR2 in AML, the included cases were divided into favourable, intermediate, and adverse risk groups on the basis of diagnostic cytogenetic and molecular features. ITPR2 expression increased with risk severity, with the lowest expression in the favourable group and the highest expression in the adverse risk group (Fig. 2C).

Fig. 2.

ITPR2 expression was increased in clinical samples vs. normal group and associated with clinical features. (A) The expression of ITPR2 in AML patients (n = 82) and normal group (n = 30). (B) ROC analysis of ITPR2 for diagnosing AML. (C) ITPR2 expression associated with increasing risk stratification. (D) Kaplan-Meier analysis of AML patients showed significantly lower survival in the ITPR2 high expression group compared with the low expression group. * p 0.05, *** p 0.001. AML, Acute myeloid leukemia; ROC, Receiver operating characteristic.

To further analysis the relationship between ITPR2 expression and OS in AML patients, subjects were divided into low and high ITPR2 expression groups with the median relative ITPR2 expression as the cutoff. The Kaplan-Meier analysis indicated a significantly shorter OS in AML cases with high ITPR2 level versus those with low ITPR2 level (Fig. 2D). Table 1 compares the clinical and biological characteristics of the two groups. Notably, high ITPR2 expression was significantly associated with a greater number of BM blasts (p = 0.01). The Cox regression analyses of the AML subgroups indicated that high ITPR2 level was a significant independent factor for short OS (p 0.05, Table 2).

To analysis the functional significance of ITPR2 in AML, TCGA samples were divided into two different expression groups based on the median ITPR2 level. Differential expression analysis was performed using the limma package. DEGs were screened with $|$log2FC$|$ $>$1 and adjusted p 0.05, followed by Gene Ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) analyses (Fig. 3A,B). GO analysis revealed that DEGs were significantly enriched in biological processes such as cellular immune responses and generation of superoxide anions; in cellular components such as cytoplasmic vesicle lumens; and molecular functions such as immune receptor activity, activation of nicotinamide adenine dinucleotide (Phosphate) reduced (NAD(P)H) oxidase activity, and Ca²⁺-dependent protein binding. KEGG analysis revealed significant enrichment of DEGs in the B-cell receptor signalling pathway and haematopoietic cell lineage. In addition, DEGs from the limma-based differential analysis, screened with adjusted p 0.05, were used for GSEA (Fig. 3C).

Fig. 3.

Functional enrichment and immune infiltration analysis of ITPR2 expression in TCGA-LAML patients. (A,B) GO and KEGG enrichment analysis of DEGs. (C) GSEA analysis of DEGs. (D–F) Correlation analysis of ESTIMATE score (D), immune score (E) and stromal score (F), respectively. (G) Correlation analysis of ITPR2 with different immune cells. (H–N) Correlation between ITPR2 and the immune checkpoint CD160 (H), CD244 (I), CD80 (J), TIGIT (K), CD200 (L), TNFSF4 (M) and TMIGD2 (N), respectively. ** p 0.01, *** p 0.001, ns: not significant (p $>$ 0.05). TCGA-LAML, The Cancer Genome Atlas-Acute Myeloid Leukemia; GO, Gene Ontology; KEGG, Kyoto Encyclopedia of Genes and Genomes; DEGs, Differentially expressed genes; GSEA, Gene Set Enrichment Analysis; ESTIMATE, Estimation of Stromal and Immune cells in Malignant Tumor tissues using Expression data; TIGIT, T cell immunoreceptor with Ig and ITIM domains; TNFSF4, Tumor necrosis factor superfamily member 4; TMIGD2, Transmembrane and immunoglobulin domain containing 2.

For the purpose of exploring ITPR2’s role in AML-related immune regulation, the ESTIMATE algorithm was utilized. Outcomes revealed an inverse association between ITPR2 level and both the ESTIMATE and immune score (Fig. 3D–F). Using the ssGSEA algorithm, the correlation of ITPR2 with the infiltration score was analysed (Fig. 3G). ITPR2 level was positively related to the T helper and T-cell infiltration but negatively related to the Tregs, Th17 cells, macrophages, and NK CD56dim cells. Furthermore, the associations between ITPR2 and immune checkpoint genes were evaluated through pearson correlation analysis. ITPR2 was positively correlated with the expression of CD160, CD244, CD80, etc. (Fig. 3H–N).

To clarify the role of ITPR2 in AML, siRNAs were used to knockdown ITPR2 in the THP-1 cell line. The knockdown efficiency was verified by RT-qPCR and WB (Fig. 4A,B). In functional validation experiments, CCK-8 assay data confirmed that ITPR2 knockdown obviously blocked the proliferation of THP-1 cells (Fig. 4C). According to the result of flow cytometry, it was found that the loss of ITPR2 caused an increase in the ratio of cells in the G0/G1 stage (Fig. 4D) and suggested a potential induction of apoptosis (Fig. 4E,F). WB analysis further verified that ITPR2 knockdown not only reduced the expression of G1/S transition regulatory factors (cyclin D1, cyclin E1, and cyclin-dependent kinase 6 (CDK6)) but also downregulated the levels of Bcl-2 and Bcl-2-associated X protein (Bax) (Fig. 4G). While both Bcl-2 and Bax decreased, the resulting Bcl-2/Bax ratio 1 was shown to promote apoptosis in previous study [18].

Fig. 4.

siRNA-mediated ITPR2 knockdown suppresses THP-1 cell proliferation and induces apoptosis. (A,B) RT-qPCR and WB analysis were employed to examine the efficiency of ITPR2 knockdown in THP-1 cells (n = 3). (C) CCK-8 assay demonstrated that ITPR2 knockdown inhibited THP-1 proliferation (n = 3). (D) Cell cycle distribution of ITPR2-knockdown THP-1 cells analyzed by flow cytometry (n = 3). (E,F) The apoptotic rate of ITPR2 knockdown THP-1 cells was increased by using the Annexin V-FITC Apoptosis Detection Kit (E) and the One-step TUNEL Apoptosis Kit (n = 3) (F) for cell staining separately through flow cytometry. (G) WB analysisdemonstrated the expression levels of cell cycle and apoptosis-related proteins in ITPR2 knockdown cells. Grayscale values were quantified using ImageJ. The quantitative results of the levels of cycle and apoptosis-related proteins were adjusted using the level of $\beta$-actin protein. The image represents one of the 3 independent parallel experiments. Data were normalized to the Control group, so error bars are not included for the Control group. si‑, small interfering; Control, THP-1 cells without any treatment; NC, negative control; Compared with si-NC, * p 0.05, ** p 0.01, *** p 0.001. RT-qPCR, Quantitative real-time PCR; WB, Western blot; THP-1, Tohoku Hospital Pediatrics-1; CCK-8, Cell counting kit-8; V-FITC, V-Fluorescein Isothiocyanate; TUNEL, Terminal Deoxynucleotidyl Transferase dUTP Nick-End Labeling; CDK6, Cyclin-dependent kinase 6; Bcl-2, B-cell lymphoma-2; Bax, Bcl-2-associated X protein.

To explore the treatment potential of targeting ITPR2, an in vitro drug sensitivity assay was performed. 2-APB is a specific inhibitor of ITPRs [19]. RT-qPCR and WB confirmed that 2-APB effectively inhibited the expression of ITPR2 (Fig. 5A,B). The viability of THP-1 cells exposed to 2-APB was assessed with CCK-8 assays. The results revealed that prolonged drug exposure significantly decreased cell viability, with an IC50 value of 202 µM after 48 h (Fig. 5C). Therefore, in the subsequent experiments, this concentration and treatment time were selected for the subsequent tests. Additionally, the results of the cell proliferation assay indicated that compared with the control group, 2-APB led to more pronounced inhibition of cell growth (Fig. 5D). Flow cytometry analysis revealed that 2-APB increased the proportion of cells arrested in the G0/G1 phase (Fig. 5E) and suggested a potential induction of apoptosis (Fig. 5F,G). WB analysis further demonstrated that 2-APB treatment reduced the levels of cyclin D1, cyclin E1, Bcl-2, and Bax (Fig. 5H).

Fig. 5.

Exposure of THP-1 cells to 200 µM 2-APB for 48 h resulted in impaired proliferation and enhanced apoptosis. (A,B) RT-qPCR and WB analysis of THP-1 cells treated with 2-APB revealed decreased ITPR2 expression (n = 3). (C) Effect of different concentrations of 2-APB on the cell viability of THP-1 cell line (n = 6). (D) CCK-8 assay demonstrated 2-APB treatment effectively suppressed THP-1 growth (n = 3). (E) Cell cycle distribution of 2-APB-treated THP-1 cells analyzed by flow cytometry (n = 3). (F,G) The apoptotic rate of 2-APB-treated THP-1 cells was increased by using the Annexin V-FITC Apoptosis Detection Kit (F) and the One-step TUNEL Apoptosis Kit (G) for cell staining separately through flow cytometry (n = 3). (H) WB examined cell cycle and apoptosisrelated protein expression in 2-APB-treated THP-1 cells. Grayscale values were quantified using ImageJ. The quantitative results of the levels of cycle and apoptosis-related proteins were adjusted using the level of $\beta$-actin protein. The image represents one of the 3 independent parallel experiments. Data were normalized to the DMSO group, so error bars are not included for the DMSO group. Compared with DMSO, * p 0.05, ** p 0.01, *** p 0.001. 2-APB, 2-Aminoethyl diphenylborinate; DMSO, Dimethyl sulfoxide; IC50, Half maximal inhibitory concentration.

ITPR2 mediates Ca²⁺ transport. To investigate whether the aberrant upregulation of ITPR2 in AML cells disrupts intracellular Ca²⁺ homeostasis, this research detected the cytosolic and mitochondrial Ca²⁺ levels via the Fluo-4 and Rhod-2 probes, respectively. Our results showed that compared with si-NC, si-ITPR2 significantly reduced Ca²⁺ concentrations in both compartments (Fig. 6A,B). Mitochondrial Ca²⁺ imbalance will result in elevated oxidative stress and mitochondrial dysfunction. With the MMP-sensitive fluorescent dye JC-1, changes in the MMP were evaluated by analyzing the red/green fluorescence ratio. Knockdown of ITPR2 significantly depolarized the MMP (Fig. 6C), a hallmark of mitochondrial dysfunction associated with early apoptosis. The intracellular ROS levels were determined using DCFH-DA. We found that ITPR2 knockdown increased the level of intracellular ROS (Fig. 6D). To evaluate the effect of ITPR2 on mtDNA-encoded respiratory chain gene expression, NADH dehydrogenase 1 (ND1) mRNA was quantified in THP-1 cells. Knockdown of ITPR2 significantly downregulated ND1 mRNA (Fig. 6E). These results are consistent with those in 2-APB-treated THP-1 cells (Fig. 6A–E).

Fig. 6.

ITPR2 Knockdown and 2-APB treatment reduce intracellular Ca²⁺ and induce mitochondrial dysfunction in THP-1 Cells. (A,B) Flow cytometric analysis revealed reduced intracellular Ca²⁺ (A) and mitochondrial Ca²⁺ (B) levels in THP-1 cells after ITPR2 knockdown or 2-APB treatment. (C) Flow cytometry showed reduced MMP in THP-1 cells following ITPR2 knockdown or 2-APB treatment. (D) Flow cytometry showed increased ROS levels in THP-1 cells following ITPR2 knockdown or 2-APB treatment. (E) RT-qPCR showed reduced mtND1 expression in THP-1 cells with ITPR2 knockdown or 2-APB treatment. The image represents one of the 3 independent parallel experiments. si‑, small interfering; NC, negative control. Compared with si-NC or DMSO * p 0.05, ** p 0.01, *** p 0.001. Ca²⁺, Calcium; MMP, Mitochondrial membrane potential.