NIPA1 depletion in tumor-associated macrophages via IGFBP2/EGFR attenuates acute myeloid leukemia progression and chemoresistance

Abstract

Acute myeloid leukemia (AML) is an aggressive and heterogenous malignancy, with unsatisfactory clinical outcomes even with very intensive treatment strategies. Previous research has revealed that Prader-Willi/Angelman syndrome 1 (NIPA1) is highly expressed in patients with AML and is negatively correlated with overall survival. However, studies on the molecular mechanism of NIPA1 in AML remain limited. To explore the role of NIPA1and its molecular mechanism in AML, we discovered through cellular experiments that NIPA1 was upregulated in M2 macrophages. NIPA1 knockdown inhibited M2 macrophage polarization and the survival of AML cells. In animal experiments, we also found that NIPA1 depletion restricted tumor growth in mice with AML. Further studies indicated that NIPA1 knockdown inhibited IGFBP2/epidermal growth factor receptor (EGFR) signaling in M2 macrophages, thereby weakening leukemia cell survival and reducing anthracycline resistance. Using both parental HL-60 and an adriamycin-resistant (HL-60/ADR) model, we show that TAM-targeted NIPA1 depletion enhances chemosensitivity and attenuates AML progression. Thus, NIPA1 could be used as a potential therapeutic strategy for immunotherapy in acute myeloid leukemia (AML).

Supplementary Information

The online version contains supplementary material available at 10.1007/s00277-025-06688-1.

Introduction

Acute myeloid leukemia (AML) is a genetically, epigenetically, and clinically heterogeneous malignancy characterized by the accumulation of somatic mutations in immature myeloid progenitors and rapid disease progression if untreated [1]. Although mutation-guided targeted therapies have improved outcomes, most patients still face high relapse rates, drug resistance, and suboptimal complete remission, with a 5-year survival below 40% [2]. Hence, there is an urgent need for more effective therapeutic strategies and a more comprehensive understanding of the mechanisms of AML. Chemoresistance remains a principal barrier to durable remission in AML, particularly to anthracyclines such as doxorubicin/adriamycin (ADR), which are core components of induction therapy. Both cell-intrinsic mechanisms (drug efflux, apoptosis evasion, metabolic rewiring) and extrinsic cues from the tumor microenvironment, especially tumor-associated macrophages (TAMs), can confer or maintain resistance. TAM-derived cytokines and growth factors support leukemic cell survival under chemotoxic stress and raise the effective IC50 to standard agents. To directly assess how macrophage NIPA1 signaling impacts therapy response, we employed an established ADR-resistant AML cell line (HL-60/ADR) alongside parental HL-60 cells, enabling us to test whether modulating TAMs via NIPA1 alters both baseline survival and anthracycline sensitivity.

Tumor-associated macrophages (TAMs) are abundant and functionally plastic components of the AML tumor microenvironment (TME). In AML, TAMs are skewed toward an immunosuppressive, pro-tumor phenotype that supports leukemic blast survival and proliferation, nurtures leukemic stem/progenitor cells, promotes angiogenesis, and contributes to therapy resistance through cytokines, chemokines, metabolic cues, and immune checkpoint signaling [3–5]. Therapeutically reprogramming TAMs toward antitumor states is therefore an attractive strategy in AML.

NIPA1 (SPG6) is a conserved transmembrane magnesium transporter best known for pathogenic variants causing hereditary spastic paraplegia and has also been implicated in non-oncologic conditions, underscoring pleiotropic biological roles beyond hematologic malignancies [6, 7]. Coincidentally, Chen et al. revealed that NIPA1 was highly expressed in AML patients and was negatively correlated with overall survival. NIPA1 has been suggested to be involved in growth and survival of AML, and targeting NIPA1 may be a promising strategy for AML treatment [8]. However, whether NIPA1 within cells of the TME—particularly macrophages—regulates AML progression remains unknown.

Interestingly, the STRING website (https://cn.string-db.org/) predicted a protein-protein interaction between NIPA1 and insulin-like growth factor binding protein 2 (IGFBP2). IGFBP2 is a crucial cell-autonomous factor that promotes the survival and migration of AML cells [9]. Notably, IGFBP2 has been reported to promote tumor progression by activating epidermal growth factor receptor (EGFR) signaling and downstream STAT3 in several cancers, including melanoma, where IGFBP2 upregulates PD-L1 via an EGFR–STAT3 axis [10]. These observations suggest that IGFBP2 can interface with EGFR pathway components, potentially via complex formation at the cell membrane. Based on these data and our STRING prediction, we hypothesized that NIPA1 may be linked to an IGFBP2–EGFR signaling module in tumor-associated macrophages (TAMs). We therefore examined IGFBP2–EGFR association and their relationship to NIPA1 using co-immunoprecipitation and co-localization assays.

We hypothesized that macrophage-intrinsic NIPA1 promotes AML progression by skewing TAM polarization toward an immunosuppressive, pro-leukemic state, potentially through an axis involving IGFBP2. To test this, a series of biological experiments was conducted to explore the potential molecular role of NIPA1 in AML development, which might be correlated with IGFBP2 and TAMs. This study provides a novel immunological explanation of this phenomenon.

Methods and materials

Cell culture, treatment and transfection

The human monocytic leukemia cell line (THP-1) and human leukemia cell line (HL-60) were obtained from Procell Life Science and Technology Co. Ltd. (Wuhan, China). ADR-resistant HL-60 cells (HL-60/ADR) were obtained from the Institute of Hematology, Chinese Academy of Medical Sciences (Tianjin, China). THP-1 cells were incubated in RPMI-1640 medium (Hyclone, Logan, UT, USA) supplemented with 10% fetal bovine serum (FBS; Gibco, Grand Island, NY, USA), 0.05 mM β-mercaptoethanol, and 1% penicillin/streptomycin mixture, while HL-60 cells were incubated with IMDM containing 20% FBS and 1% penicillin/streptomycin. HL-60/ADR cells were cultured in RPMI-1640 medium with 10% FBS and 0.2–0.4 µg/mL ADR to maintain the drug-resistant phenotype, and were cultured in ADR-free medium for 1 weeks prior to experiments. All cells were cultured in a humidified incubator with 5% CO₂ at 37℃.

THP-1 cells were induced with 100 ng/mL phorbol myristate acetate (PMA; Sigma-Aldrich, Carlsbad, CA, USA) to differentiate into M0 macrophages. Subsequently, the macrophages were stimulated towards M2 polarization with IL-4 (50 ng/ml, R&D Systems)/IL-13 (20 ng/ml, R&D Systems) for 48 h. M2 macrophages were transfected with two shRNAs of NIPA1 (sh-NIPA1-1 and sh-NIPA1-2) and a negative control (sh-NC), or IGFBP2-overexpression vector (Ov-IGFBP2) along with its negative control (Ov-NC), using Lipofectamine 3000 reagent (Invitrogen) for 48 h according to the manufacturer’s instructions. Subsequently, the conditioned medium (CM) from the macrophages was used to culture HL-60 cells to investigate the effect of NIPA1-mediated macrophages on HL-60 cellular behavior. For EGFR inhibition in macrophages, cetuximabwas added only to THP-1–derived macrophages for the final 24 h of the 48 h transfection/overexpression period (sh-NIPA1 ± Ov-IGFBP2). CM was then collected, centrifuged to remove cells/debris, and applied to HL-60 or HL-60/ADR cells without cetuximab. Leukemia cells were not exposed directly to cetuximab in these assays.

Quantitative real-time PCR

RNA was extracted from cells using TRIzol reagent (Invitrogen). After reverse transcription using a ReverTra Ace qPCR RT Kit (Toyobo, Osaka, Japan), real-time PCR was performed using the LightCycler 96 Instrument (Roche, Mannheim, Germany) and the SYBR Master Mix kit (OriGene Technology, Rockville, MD, USA) according to the manufacturer’s instructions. The mRNA levels were calculated using the 2^−△△Ct method and normalized to β-actin levels.

Western blot

Cells were collected, washed with PBS, and lysed with RIPA buffer to extract the total protein. The protein concentration was determined using a bicinchoninic acid Protein Assay Kit (Beyotime, Shanghai, China). Protein lysates for Bax/Bcl-2 analyses were collected after 48 h exposure to CM. The proteins (30 µg) were loaded onto 12% SDS-PAGE gels and electro-transferred to polyvinylidene fluoride (PVDF) membranes (Bio-Rad, Hemel Hempstead, UK). Subsequently, the membranes were blocked with skimmed milk for 2 h and then incubated overnight at 4℃ with primary antibodies, followed by an incubation with HRP-conjugated secondary antibodies for 2 h at room temperature. The protein bands were visualized using an enhanced chemiluminescence kit (GE Healthcare).

Flow cytometry analysis

To assess apoptosis in HL-60 cells, the cells were exposed to macrophage-CM for 48 h before Annexin V/PI staining (Beyotime, Shanghai, China) for 15 min in the dark. To assess the proportion of macrophages, FITC-conjugated anti-CD68, anti-CD86, and anti-CD163 antibodies were used to analyze the surface antigen expression of macrophages. Finally, the stained cells were analyzed using a FACS Calibur flow cytometer (BD Biosciences, San Jose, USA).

Cell counting kit-8 assay

HL-60 or HL-60/ADR cells were cultured in CM from NIPA1-knockdown M2 macrophages for 24, 48, and 72 h, respectively. CCK-8 solution (Dongren Chemical Technology, Shanghai, China) was added to each well, and the cells were incubated for an additional 3 h. Cell viability was assessed based on the absorbance of each well at 450 nm using a microplate reader (Biotek, Winooski, VT, USA).

In addition, HL-60 and HL-60/ADR cells were treated with increasing concentrations of ADR (0, 1, 2, 4, 8, 16, 32, 64, 128, and 256 µM) for 24 h. Cell viability was determined using the CCK-8 assay, and the half-maximal inhibitory concentration (IC50) value was calculated.

5-Ethynyl-2-deoxyuridine (EdU) incorporation assay

After HL-60 cells were labelled with EdU, the Click reaction mixture from the Click-iT EdU Alexa Fluor 488 kit (Invitrogen) was added and incubated for 30 min at room temperature in the dark. EdU was added for the final 2 h of a 24 h incubation with CM prior to fixation and Click-iT labeling. Nuclei were counterstained with DAPI. Images were acquired on an Olympus fluorescence microscope using identical exposure settings across groups. For quantification, five non-overlapping fields per well were imaged at 20× (three technical replicate wells per condition; n = independent experiments as indicated in figure). EdU-positive nuclei (Alexa Fluor 488 channel) and total nuclei (DAPI channel) were segmented using ImageJ/Fiji with a fixed intensity threshold and watershed separation applied uniformly to all images. The percentage EdU incorporation was calculated as (EdU + nuclei/total DAPI + nuclei) × 100% for each field, and field-level values were averaged per well, then per experiment. Analyses were performed blinded to condition.

Co-immunoprecipitation (CoIP) assay

Total protein was extracted from cells as previously described. A portion of the total protein served as the input control, whereas other proteins were incubated with IP-indicated antibodies at 4℃ overnight, followed by incubation with Protein A/G agarose beads (Calbiochem, Temecula, CA, USA) at 4℃ for 4 h. After washing and elution, immunoprecipitation was performed using western blotting.

Immunofluorescent assay

Cells seeded on coverslips were fixed with 4% paraformaldehyde and permeabilized with 0.1% Triton X-100 at room temperature. After blocking with 3% BSA at room temperature for 30 min, the cells were incubated with primary antibodies overnight at 4℃, followed by incubation with FITC-conjugated goat anti-rabbit IgG antibody for 1 h at room temperature. After counterstaining with DAPI, fluorescence images were obtained using a fluorescence microscope (Olympus, Tokyo, Japan).

Animal study

A total of 24 male BALB/c nude mice (4–6 weeks old, 18–22 g) were purchased from Charles River Laboratory Animal Technology Co., Ltd. (Beijing, China). All the mice were maintained under controlled temperature, humidity, and illumination conditions. The mice were randomly divided into three groups (n = 8 mice per group). For the AML mouse model, HL-60 cells were co-transplanted subcutaneously with THP-1–derived M2 macrophages in which NIPA1 was transiently knocked down prior to injection. Briefly, THP-1 cells were differentiated to M0 and polarized to M2 as described above, then transfected with sh-NIPA1-2 or sh-NC for 48 h. After confirming NIPA1 knockdown (qRT-PCR/western blot), HL-60 cells were mixed with the indicated M2 macrophages at a 1:1 ratio (5 × 10^6 HL-60 + 5 × 10^6 M2 macrophages in 100 µL PBS/Matrigel 1:1) and injected into the right flank. Three groups were used: (i) HL-60 + M2(sh-NC), (ii) HL-60 + M2(sh-NIPA1), and (iii) HL-60 only. Tumor size was measured every three days using a Vernier caliper, and tumor volume was recorded by measuring the perpendicular diameters using the formula: (min² × max)/2. After sacrifice, single cells were isolated from the fresh tumor tissues using Collagenase IV, followed by staining with CD163 antibody at 4 °C for 30 min. The stained cells were then subjected to flow cytometry. All animal procedures were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals and Animal Care, and were approved by the Ethics Committee of the First Affiliated Hospital of Ningbo University (Approval number: 12491).

Immunohistochemical staining

The resected tumors were fixed in 4% paraformaldehyde, embedded in paraffin, and mounted on slides. After deparaffinization and rehydration, tumor sections were incubated in 3% hydrogen peroxide to inhibit endogenous peroxidase activity. Following antigen retrieval with citrate buffer (pH6.0), the sections were blocked with 10% goat serum and then probed overnight at 4℃ with primary antibodies. Detection was performed using a Dako IHC kit (Dako EnVision + System, HRP, Agilent Technologies). Images were obtained using a light microscope (Olympus).

Terminal-deoxynucleoitidyl transferase mediated Nick end labeling (TUNEL) assay

After deparaffinization and rehydration, the tumor sections were washed with PBS and incubated with 20 µg/ml Proteinase K at 37℃ for 30 min. Subsequently, the tissues were incubated with TUNEL detection mixture (Beyotime) at 37℃ for 60 min in the dark. Images were obtained using a light microscope (Olympus).

Statistical analysis

All results are presented as the mean ± standard deviation (SD). Statistical analyses were performed using GraphPad Prism software. The means of the variables were examined using one-way ANOVA with Tukey’s post-hoc test. Differences were considered statistically significant when the p-value was less than 0.05.

Results

NIPA1 is involved in the regulation of M2 macrophages polarization

To investigate the role of NIPA1 in macrophage polarization, THP-1 cells were differentiated into M0 macrophages using PMA, as evidenced by the high expression of CD68 (Fig. 1A), a well-known marker for M0 macrophages. After stimulation with IL-4/IL-13, the mRNA levels of CD163 and the percentage of CD163⁺ cells were significantly increased compared to the control (Fig. 1B-C), confirming macrophage polarization towards M2 phenotype. Additionally, the expression level of NIPA1 was also upregulated in IL-4/IL-13-treated macrophages compared to the control (Fig. 1D), suggesting that NIPA1 was upregulated in M2 macrophages.

Fig. 1.

NIPA1 is involved in the regulation of M2 macrophages polarization (A) THP-1 cells were differentiated into M0 macrophages by PMA, and the mRNA level of CD68 was examined by qRT-PCR. (B) M0 macrophages were stimulated with IL-4/IL-13, and the mRNA level of CD163 was examined by qRT-PCR. (C) Flow cytometry analysis was performed to detect CD163⁺ cells. (D) The mRNA levels of NIPA1 were examined by qRT-PCR. **p < 0.01, ***p < 0.001

NIPA1 knockdown inhibits M2 macrophage polarization

To clarify the specific function of NIPA1 in macrophage polarization, two shRNAs targeting NIPA1 (sh-NIPA1-1 and sh-NIPA1-2) were designed. As exhibited in Fig. 2A-B, NIPA1 expression was significantly decreased at both the mRNA and protein levels in the sh-NIPA1-1 and sh-NIPA1-2 groups compared to sh-NC group. Because of its higher transfection efficacy, sh-NIPA1-2 was selected for the further experiments. Figure 2C-F illustrate that in IL-4/IL-13-induced M2 macrophages, NIPA1 knockdown significantly promoted the mRNA level of CD86, a classical marker of M1 macrophages, as well as the percentage of CD86⁺ cells. In contrast, the mRNA levels of CD163 and the percentage of CD163⁺ cells were significantly decreased following NIPA1 knockdown. These findings suggested that silencing NIPA1 could reduce macrophage polarization towards the M2 phenotype.

Fig. 2.

NIPA1 knockdown inhibits M2 macrophage polarization (A) Two shRNAs of NIPA1 (sh-NIPA1-1 and sh-NIPA1-2) were transfected into macrophages, and the mRNA level of NIPA1 was examined. (B) The protein expression of NIPA1 was detected using western blot. (C) The mRNA levels of CD86 were examined by qRT-PCR. (D) CD86⁺ cells were detected using flow cytometry. (E) The mRNA levels of CD163 were examined by qRT-PCR. (F) CD163⁺ cells were detected using flow cytometry. ***p < 0.001

NIPA1 depletion in M2 macrophages contributes to decreased proliferation and increased apoptosis of HL-60 and HL-60/ADR cells.

Subsequently, Subsequently, HL-60 and HL-60/ADR cells were cultured in CM for 24–72 h. Cell viability was measured at 24/48/72 h (Fig. 3A), EdU incorporation after 24 h CM exposure with EdU added for the final 2 h (Fig. 3B), and apoptosis after 48 h CM exposure (Fig. 3C). Bax/Bcl-2 were assessed after 48 h CM exposure (Fig. 3D-E). HL-60 and HL-60/ADR cells were cultured in CM from M2 macrophages to determine whether NIPA1 knockdown influenced the function of macrophages in tumor development and drug resistance. As shown in Fig. 3A, the OD value in the sh-NIPA1 group was significantly lower than those in the sh-NC group for both HL-60 and HL-60/ADR cells, indicating that the supernatant from NIPA1 knockdown-mediated M2 macrophages could restrict the viability of AML cells. Similarly, the number of EDU-positive cells in the sh-NIPA1 group was significantly reduced compared to the sh-NC group (Fig. 3B), revealing that NIPA1 knockdown-mediated M2 macrophages also suppressed the proliferation of AML cells. In addition, flow cytometry analysis revealed that NIPA1 depletion in M2 macrophages significantly promoted the apoptosis of HL-60 and HL-60/ADR cells, which was further confirmed by the decreased protein expression of Bcl-2 and increased protein expression of Bax in the sh-NIPA1 group compared to that in the sh-NC group (Fig. 3D-E).

Fig. 3.

NIPA1 depletion in M2 macrophages contributes to decreased proliferation and increased apoptosis of HL-60 and HL-60/ADR cells (A) HL-60 cells or HL-60/ADR cells were cultured with condition medium from M2 macrophages. Cell viability was assessed using CCK-8 assay. CCK-8 viability at 24, 48, and 72 h after exposure to CM . (B) EDU staining was performed to assess cell proliferation ability. EdU incorporation after 24 h CM exposure; EdU was pulsed for the final 2 h. (C) Flow cytometry analysis was conducted to detect cell apoptosis. Annexin V/PI apoptosis measured after 48 h CM exposure. (D-E) The protein expression of Bax and Bcl-2 was detected using western blot. Bax/Bcl-2 by western blot after 48 h CM exposure. **p < 0.01, ***p < 0.001

NIPA1 knockdown inhibits IGFBP2/epidermal growth factor receptor (EGFR) in M2 macrophages

Next, we explored the potential regulatory mechanisms of NIPA1 in macrophages, as shown in Fig. 4A. NIPA1 knockdown in M2 macrophages significantly inhibited IGFBP2 and EGFR protein expression. According to the STRING website (https://cn.string-db.org/), there may be a protein-protein interaction between NIPA1 and IGFBP2. To validate these interactions, co-immunoprecipitation (Co-IP) assays were conducted. As exhibited in Fig. 4B-C, not only did NIPA1 interact with IGFBP2, but IGFBP2 also interacted with EGFR. Moreover, two-color immunofluorescence showed overlapping signals for IGFBP2 (red) with NIPA1 (green) and, in a separate experiment, IGFBP2 (red) with EGFR (green) (Fig. 4D), indicating spatial proximity of these protein pairs (Fig. 4D), consistent with co-distribution but not proof of direct molecular interaction. Protein–protein association was supported by Co-IP (Fig. 4B–C). These data suggested that NIPA1 positively regulated IGFBP2/EGFR in M2 macrophages via protein interactions.

Fig. 4.

NIPA1 knockdown inhibits IGFBP2/EGFR in M2 macrophages (A) In NIPA1 knockdown-mediated M2 macrophages, the protein expression levels of IGFBP2 and EGFR were examined using western blot. (B) CoIP assays were performed to examine the protein-protein interaction between IGFBP2 and NIPA1. (B) CoIP assays were performed to examine the protein-protein interaction between IGFBP2 and EGFR. (D) Two separate two-color immunofluorescence assays: top row, IGFBP2 (red) with NIPA1 (green); bottom row, IGFBP2 (red) with EGFR (green). Nuclei were counterstained with DAPI. Images illustrate overlapping signals for each protein pair. ***p < 0.001

CM from NIPA1-knockdown M2 macrophages modulates AML cell survival and chemosensitivity via the IGFBP2/EGFR axis

CM collected from M2 macrophages transfected with sh-NIPA1, with or without IGFBP2 overexpression and/or EGFR inhibition, was applied to HL-60 and HL-60/ADR cells to assess effects on survival and chemosensitivity. Following co-transfection, IGFBP2 mRNA levels were significantly upregulated in the sh-NIPA1 + Ov-IGFBP2 group compared with those in the sh-NIPA1 + Ov-NC group (Fig. 5A). Cetuximab was applied only to the macrophage cultures during the final 24 h before harvest to probe the role of macrophage EGFR signaling in polarization; no inhibitor was added to leukemia cells. The results showed that NIPA1 knockdown-mediated CD86 mRNA upregulation and CD163 mRNA downregulation were significantly reversed by IGFBP2 overexpression. This reversal was partially inhibited by the additional treatment with an EGFR inhibitor (Fig. 5B). Consistently, flow cytometry analysis also revealed that, compared to the sh-NIPA1 + Ov-NC group, the proportion of CD86⁺ cells was significantly reduced, whereas the proportion of CD163⁺ cells was significantly increased in the sh-NIPA1 + Ov-IGFBP2 group, which was partly restored by the EGFR inhibitor (Fig. 5C). These results suggested that NIPA1 knockdown-mediated inhibition of macrophage polarization towards the M2 phenotype is regulated by the IGFBP2/EGFR axis.

Fig. 5.

The impact of NIPA1 knockdown on macrophages polarization was modulated by IGFBP2/EGFR (A) M2 macrophages were transfected with sh-NIPA1 alone or co-transfected with sh-NIPA1 and Ov-NC/Ov-IGFBP2. The mRNA levels of IGFBP2 were examined using qRT-PCR. (B) The mRNA levels of CD86 and CD163 were examined using qRT-PCR. (C) CD86⁺ cells and CD163⁺ cells were detected using flow cytometry. *p < 0.05, **p < 0.01, ***p < 0.001

The impacts of NIPA1 knockdown on AML cell survival and chemosensitivity were modulated by IGFBP2/EGFR

CM from M2 macrophages transfected with sh-NIPA1, with or without IGFBP2 overexpression, and treated ± cetuximab during the final 24 h of macrophage culture, was applied to HL-60 or HL-60/ADR cells. Cetuximab was not added to leukemia cells, ensuring that observed effects reflect macrophage-secreted factors rather than direct EGFR blockade on AML cells. As shown in Fig. 6A, compared to the sh-NIPA1 + Ov-NC group, the OD value in the sh-NIPA1 + Ov-IGFBP2 group was significantly increased in both HL-60 and HL-60/ADR cells, which was partly reversed by additional treatment with an EGFR inhibitor. Compared to the sh-NIPA1 + Ov-NC group, the number of EDU-positive cells in the sh-NIPA1 + Ov-IGFBP2 group was significantly elevated, which was partly reversed in the sh-NIPA1 + Ov-IGFBP2 + EGFR inhibitor group (Fig. 6B). Moreover, flow cytometry analysis revealed that NIPA1 depletion in M2 macrophages caused an increased rate of apoptosis in both HL-60 and HL-60/ADR cells, which was significantly reduced by IGFBP2 overexpression, accompanied by downregulation of Bax protein expression and upregulation of Bcl-2 protein expression. Compared to the sh-NIPA1 + Ov-IGFBP2 group, the cell apoptosis rate was greatly increased in the sh-NIPA1 + Ov-IGFBP2 + EGFR inhibitor group (Fig. 6C-E). The results were shown in Fig. 6F revealed that NIPA1 knockdown in macrophages significantly reduced the IC50 value for both HL-60 and HL-60/ADR cells in response to ADR treatment. In addition, the reduced IC50 value due to NIPA1 knockdown increased significantly following IGFBP2 overexpression, and the EGFR inhibitor could partly inhibit this elevation.

Fig. 6.

The impacts of NIPA1 knockdown on AML cell survival and chemosensitivity were modulated by IGFBP2/EGFR (A) HL-60 cells or HL-60/ADR cells were cultured with CM from M2 macrophages. Cell viability was assessed using CCK-8 assay. Viability at 48 h. (B) EDU staining was performed to assess cell proliferation ability. EdU after 24 h CM with 2 h EdU pulse. (C) Flow cytometry analysis was carried out to detect cell apoptosis. (D-E) The protein expression of Bax and Bcl-2 was detected using western blot. (F) HL-60 cells or HL-60/ADR cells were treated with increasing concentrations of ADR, and the IC50 value was calculated. *p < 0.05, **p < 0.01, ***p < 0.001

NIPA1 depletion restricts tumor growth in AML mice

We established an AML xenograft model by co-injecting HL-60 cells with M2 macrophages transfected with either sh-NIPA1 or sh-NC, thereby specifically manipulating NIPA1 in the macrophage compartment at the time of engraftment. As shown in Fig. 7A, NIPA1 depletion significantly slowed tumor growth and reduced both tumor weight and volume. The results of immunohistochemical staining and the TUNEL assay presented in Fig. 7B demonstrated that NIPA1 knockdown significantly reduced the expression of NIPA1 and Ki67 in tumor sections and increased the number of TUNEL-positive cells in tumor tissues. This indicated that NIPA1 depletion effectively inhibited cell proliferation and promoted cell apoptosis in vivo. This was also confirmed by the downregulated protein expression of Bcl-2 and upregulated protein expression of Bax following NIPA1 knockdown (Fig. 7C). Furthermore, flow cytometry analysis confirmed that NIPA1 knockdown significantly increased the infiltration of CD86⁺ cells and decreased CD163⁺ cell infiltration in tumor tissues, confirming that NIPA1 knockdown promoted macrophage polarization towards the M1 phenotype (Fig. 7D). These in vivo experiments were conducted without ADR treatment; thus, the data demonstrate the impact of macrophage NIPA1 depletion on tumor growth and immune phenotype at baseline.

Fig. 7.

NIPA1 depletion in co-transplanted M2 macrophages restricts tumor growth in AML mice (A) BALB/C nude mice were injected subcutaneously with HL-60 cells mixed 1:1 with THP-1–derived M2 macrophages transfected for 48 h with sh-NIPA1 (M2 sh-NIPA1) or control shRNA (M2 sh-NC). Tumor volume and tumor weight were measured. (B) Representative images of immunohistochemical staining (NIPA1 and Ki67) and TUNEL assay. (C) Protein expression of Bcl-2 and Bax in tumor tissue was detected using western blot. (D) Flow cytometry analysis was used to examine CD86⁺cell and CD163⁺cells infiltration in tumor tissues. ***p < 0.001

Discussion

Acute myeloid leukemia (AML) is a highly heterogeneous malignancy. Currently, a considerable number of therapeutic strategies have been developed to improve the outcomes of AML patients, yet the complete remission rate remains low [2]. Consequently, there is an urgent need to develop novel diagnostic and therapeutic approaches. To the best of our knowledge, this is the first functional demonstration that NIPA1 is essential for macrophage polarization. The NIPA1 depletion in M2 macrophages hindered the survival of HL-60 cells and enhanced the chemosensitivity of HL-60/ADR cells by regulating the IGFBP2/EGFR axis. In general, NIPA1 could be used as a therapeutic agent for AML.

A key objective of this study was to determine whether TAM NIPA1 signaling contributes to chemoresistance, a major clinical failure mode in AML. We therefore included an adriamycin-resistant line (HL-60/ADR) to mirror anthracycline-refractory disease. NIPA1 depletion in M2 macrophages reduced leukemia cell viability and increased apoptosis in both HL-60 and HL-60/ADR cells, and significantly decreased ADR IC50 values. Rescue by IGFBP2 overexpression and partial reversal by EGFR inhibition indicate that a NIPA1–IGFBP2–EGFR axis in TAMs sustains survival signaling under chemotoxic stress. These data support the translational relevance of targeting TAM NIPA1 to overcome microenvironment-mediated anthracycline resistance.

Inspired by research on the TME in solid tumors, the study of TAMs in hematologic malignancies has gained significant attention. Macrophages are highly adaptable cells that can change their phenotype and function in response to various factors, including epigenetic changes, the surrounding environment, and external stimuli. They can exhibit characteristics of both the pro-inflammatory M1 and anti-inflammatory M2 phenotypes. TAMs mainly represent an M2-like phenotype with potent immunosuppressive activity and play a tumor-promoting role in carcinogenesis, metastasis, and chemoresistance [11–14]. Numerous studies have indicated that a high proportion of M2 TAMs was correlated with poor prognosis in patients with AML [15]. M2 macrophages derived aggressive leukemia development in vivo by enhancing resistance to phagocytosis, improving mitochondrial metabolism, and altering macrophage polarization. This presented a promising strategy for cancer treatment [16]. For instance, chenodeoxycholic acid had been reported to inhibit AML progression by promoting lipid peroxidation by reducing M2 macrophage polarization [17]. On the other hand, Tian et al. revealed that let-7b was a potentially aberrant gene in leukemia-associated macrophages that drives the M2 phenotype. The knockdown of let-7b could repolarize leukemia-associated macrophages towards the M1 phenotype, which might help inhibit the development of AML [15]. In this study, we found that NIPA1 was consistently upregulated when macrophages were polarized to the M2 phenotype. Additionally, NIPA1 knockdown decreased the proportion of M2 macrophages, suggesting that NIPA1 might be a macrophage M2 co-expression factor. In addition, the CM from NIPA1-depleted macrophages caused decreased proliferation and increased apoptosis of HL-60 and HL-60/ADR cells in vitro, which was also confirmed in an in vivo animal study. These results indicated that NIPA1 depletion could hinder macrophage polarization towards the M2 phenotype, which in turn reduced leukemia cell survival and chemoresistance, thereby blocking AML development.

To date, research on the regulatory role of NIPA1 in AML and its molecular mechanisms is still limited. This study revealed for the first time that NIPA1 might be an aberrant gene that influences macrophages to adopt the M2 phenotype. Nevertheless, the mechanism by which NIPA1 regulates macrophage polarization remains unclear. It was discovered from the STRING website (https://cn.string-db.org/) that NIPA1 might interact with IGFBP2, which was verified by the CoIP assay. IGFBP2 has been identified as a gene associated with cancer-associated fibroblasts that can induce M2 macrophage polarization in the TME, contributing to tumor development [18]. Accordingly, our finding suggested that IGFBP2 overexpression counteracted the impact of NIPA1 knockdown on macrophage polarization, indicating that NIPA1 knockdown might reduce M2 macrophage polarization, in part by inhibiting IGFFBP2. Our IF data indicate overlapping subcellular distribution of NIPA1, IGFBP2, and EGFR, whereas Co-IP supports protein–protein association. We therefore interpret the microscopy findings as co-distribution consistent with, but not demonstrating, complex formation. This aligned with a previous report that IGFBP2 facilitated melanoma progression by activating EGFR [10]. EGFR belongs to the human epidermal growth receptor family and plays a critical role in tumorigenesis [19]. Research has been shown that it influences the growth and differentiation of AML cells, leading to anticancer effects [20, 21]. EGFR-mediated activation of macrophages towards the M2 phenotype plays an important role in tumorigenesis and metastasis [22, 23]. Cannabinoid Receptor-1, serving as a targeted antitumor therapy, has been demonstrated to suppress M2 macrophage polarization by downregulating EGFR [24]. In the present study, we observed an inhibitory effect of NIPA1 knockdown on EGFR expression. NIPA1 might positively regulate IGFBP2/EGFR in M2 macrophages via protein interactions. Therefore, NIPA1 depletion might attenuate M2 macrophage polarization by downregulating the IGFBP2/EGFR axis. While our in vitro studies show that macrophage NIPA1 depletion reduces ADR IC50 and enhances apoptosis in both parental and ADR-resistant AML cells, the current in vivo xenograft experiments were performed without anthracycline treatment. Therefore, we cannot directly conclude in vivo chemosensitization from these data. Future studies will test whether targeting NIPA1 in TAMs augments the efficacy of ADR in vivo, including dosing, scheduling, and pharmacodynamic readouts of the IGFBP2/EGFR axis.

In conclusion, these findings revealed that NIPA1 might act as a co-expression factor for M2 macrophages. NIPA1 depletion could hinder macrophages polarization towards the M2 phenotype, which in turn reduced the leukemia cell survival and chemoresistance, thereby blocking AML development. Thus, targeting NIPA1 could serve as a promising therapeutic approach for immunomodulation in AML, with the potential to mitigate anthracycline resistance, which warrants evaluation in ADR-treated in vivo models.

Supplementary Information

Below is the link to the electronic supplementary material.

Author contributions

GO designed the study; ST, LS, QM, YW, LC, KX, MZ, ZX, and AW collected the data; LS, QM, YW, LC and KX analyzed the data; ST and ZX interpreted the data; and ST drafted and GO revised the manuscript. ST and GO confirm the authenticity of all the raw data. All authors have read and reviewed the final manuscript.

Funding

This work was supported by the Zhejiang Province Leading Geese Plan under Grant 2025C02247(SD2).

Data availability

No datasets were generated or analysed during the current study.

Declarations

Ethic statement

All animal experiments were conducted in accordance with the Guide for the Care and Use of Laboratory Animals and approved by the Animal Care and Use Committee of the First Affiliated Hospital of Ningbo University.

Consent for publication

Its publication has been approved by all coauthors.

Consent to participate

Not applicable.

Competing interests

The authors declare no competing interests.