Mechanisms by which macrophage Ltb4r1 mediates tubular epithelial cell injury to facilitate the advancement of diabetic kidney disease

Abstract

Objective

To investigate the role of leukotriene B4 receptor 1 (LTB4R1) in high glucose-induced renal tubular epithelial cell injury and its regulatory association with macrophage polarization, and to validate the protective effect of LTB4R1 silencing in diabetic renal injury in vivo.

Methods

THP-1 cells were differentiated and polarized into M0, M1 (LPS + IFN-γ-induced) and M2 (IL-4 + IL-13-induced) macrophages, with phenotypic validation by immunofluorescence. HK-2 renal tubular epithelial cells were treated with gradient glucose concentrations (5.5, 15, 30, 45 mM) to assess dose-dependent effects on LTB4R1 expression and apoptosis. Indirect co-culture of HK-2 cells with polarized macrophages under high glucose (HG) conditions was performed using a Transwell system, with an osmotic control group to exclude osmotic artifacts. For in vivo studies, STZ-induced diabetic C57BL/6 mice were treated with AAV-mediated LTB4R1 shRNA. qPCR, Western blot, ELISA and flow cytometry were used to detect molecular expression, inflammatory cytokines and apoptosis.

Results

THP-1 cells were successfully differentiated into M0, M1 and M2 macrophages with distinct phenotypic marker expression. High glucose dose-dependently upregulated LTB4R1 mRNA and protein levels (all P < 0.01) and promoted HK-2 cell apoptosis (P < 0.01). HG specifically induced HK-2 cell injury, which was exacerbated by M1 macrophages and alleviated by M2 macrophages; concurrent changes included HG-induced upregulation of LTB4R1/CD68 and downregulation of CD163/CD206, which were reversed by M2 co-culture (all P < 0.01). In diabetic mice, LTB4R1 silencing significantly reduced renal Ltb4r1, cleaved caspase-3 and NF-κB p65 transcripts, suppressed pro-inflammatory cytokines (TNF-α, IL-1β) and elevated anti-inflammatory IL-10, restored CD163/CD206 expression, and inhibited renal apoptosis (all P < 0.01).

Conclusion

LTB4R1 mediates high glucose-induced renal tubular epithelial cell injury and diabetic renal injury by regulating macrophage polarization and NF-κB signaling. Silencing LTB4R1 mitigates diabetic renal inflammation and apoptosis, suggesting LTB4R1 as a potential therapeutic target for diabetic nephropathy that warrants further investigation.

Supplementary Information

The online version contains supplementary material available at https://doi.org/10.1186/s12882-026-05158-8.

Introduction

Diabetic kidney disease (DKD) is a leading cause of end-stage renal disease (ESRD) worldwide, and its incidence continues to climb alongside the global diabetes epidemic [1, 2]. Approximately 30–40% of individuals with diabetes ultimately develop DKD, conferring substantial morbidity, mortality, and socioeconomic burden [3, 4]. Although prior work has centered on glomerular injury, accumulating evidence now establishes that early tubulointerstitial damage—particularly involving proximal tubular epithelial cells (PTECs)—is a decisive driver of DKD progression. PTECs are highly susceptible to metabolic stress, oxidative insult, and inflammatory stimuli; their maladaptive repair following injury represents a major force behind renal fibrosis in DKD. Accordingly, targeting tubulointerstitial inflammation has emerged as a promising therapeutic strategy [5, 6]. Nevertheless, the upstream molecular mechanisms that govern tubulointerstitial inflammation under hyperglycemic conditions remain incompletely defined.

Macrophages are central immune cells that link metabolic dysregulation to chronic renal inflammation [7]. In the diabetic kidney, they display remarkable plasticity, dynamically shifting along the M1 (pro-inflammatory)–M2 (repair/pro-fibrotic) polarization spectrum. Through secretion of pro-inflammatory cytokines such as TNF-α and IL-1β, or anti-inflammatory mediators including IL-10, macrophages critically shape the extent of tubular injury and interstitial fibrosis [8–10]. In DKD, M1-dominant polarization drives early pro-inflammatory tubular damage, whereas M2 polarization can either resolve inflammation or, in later stages, contribute to fibrosis [11, 12]. Notably, the degree of M1 macrophage infiltration in kidney biopsies from DKD patients correlates positively with the severity of tubular injury; however, the upstream signals that direct this polarization imbalance remain poorly characterized [13].

Leukotriene B₄ (LTB₄) and its high-affinity receptor LTB₄R1 (also termed BLT1) constitute a key signaling axis that regulates innate immune cell chemotaxis and activation [14, 15]. LTB₄R1 is a G protein–coupled receptor that transduces the effects of LTB₄, a potent lipid mediator derived from arachidonic acid metabolism [16]. Recent evidence indicates that LTB₄R1 activation promotes macrophage polarization toward a pro-inflammatory M1 phenotype via the NF-κB pathway, thereby exacerbating tissue injury in autoimmune and chronic inflammatory diseases [17–19]. However, existing studies have largely focused on extra-renal contexts, and the role of LTB₄R1 in DKD has not been systematically investigated. In particular, whether LTB₄R1 regulates macrophage polarization and macrophage–tubular epithelial crosstalk in the diabetic kidney remains unknown. Preliminary data suggest that LTB₄/LTB₄R1 expression is upregulated in diabetic nephropathy, pointing to a potential pro-inflammatory function [20]. Yet it is unresolved whether hyperglycemia-driven LTB₄R1 activation contributes to tubular injury through macrophage polarization, and whether downstream pathways such as NF-κB are involved.

Current DKD management relies on glycemic control, blood pressure regulation, and inhibition of the renin–angiotensin–aldosterone system (RAAS) [21]. Despite these interventions, a substantial proportion of patients still progress to ESRD, underscoring the need to identify novel immuno-inflammatory targets beyond metabolic control [22]. On the basis of these knowledge gaps, we propose a testable hypothesis: hyperglycemia directly upregulates LTB₄R1 in PTECs to initiate tubular injury; injured PTECs recruit monocytes and promote M1 macrophage polarization; the hyperglycemic microenvironment concurrently upregulates LTB₄R1 in M1 macrophages, which activates NF-κB signaling to propagate pro-inflammatory responses; these responses in turn amplify LTB₄R1 expression in PTECs, thereby establishing a pathogenic positive-feedback loop between tubular injury and macrophage activation. By contrast, M2 macrophages may interrupt this loop and confer renal protection by suppressing LTB₄R1 expression in both cell types.

This study aims to validate the above hypothesis and to define the role and mechanisms of LTB₄R1 in macrophage–tubular epithelial crosstalk during DKD. We first examined the dose-dependent effects of hyperglycemia on LTB₄R1 expression and tubular injury in vitro, then established co-culture systems to assess the phenotype-specific effects of macrophages. In parallel, we employed a streptozotocin (STZ)-induced diabetic mouse model to evaluate the therapeutic potential of LTB₄R1 silencing in vivo. By delineating the LTB₄R1–NF-κB–macrophage polarization axis in DKD, this work fills a critical mechanistic gap and identifies LTB₄R1 as a promising therapeutic target.

Materials and methods

Cell culture

Human renal proximal tubular epithelial cells (HK-2; ATCC, Manassas, Virginia, USA) and human monocyte THP-1 cells (ATCC) were used in this study. All cell lines were authenticated by short tandem repeat (STR) profiling and routinely tested for mycoplasma contamination using a PCR-based detection method, with only mycoplasma-negative cells used for experiments. HK-2 cells were cultured in DMEM/F-12 medium (Thermo Fisher Scientific, Waltham, MA, USA) containing 10% fetal bovine serum (FBS; PS-FB5-SA, Peak Inc., USA) and 1% penicillin-streptomycin (15140122, Thermo Fisher Scientific, Waltham, MA, USA). THP-1 cells were cultured in RPMI-1640 medium (L211013, Basalmedia Biotechnology, Shanghai, China) containing 10% FBS and 1% penicillin-streptomycin. Cells were incubated at 37 °C in a humidified atmosphere containing 5% CO₂. Upon reaching 80–90% confluence, HK-2 cells were subcultured following detachment with 0.25% trypsin-EDTA (C0201, Beyotime, Shanghai, China). Then cells were treated with trypsin at 37 °C for 2–5 min, neutralized with complete medium, centrifuged, and reseeded at a ratio of 1:3. All cell culture experiments were performed with biological replicates (n = 3, independent cell passages) and technical duplicates per biological replicate.

In vivo animal model establishment and treatment

Male C57BL/6 mice (6–8 weeks old, 20–22 g) were purchased from Beijing Vital River Laboratory Animal Technology Co., Ltd. (Beijing, China; SCXK (Jing) 2021-0011). Mice were housed in a specific pathogen-free (SPF) environment with a 12-hour light/dark cycle, constant temperature (22 ± 2 °C) and humidity (50 ± 5%), and free access to food and water. After 1 week of acclimatization, the diabetic model was induced by a single intraperitoneal injection of streptozotocin (STZ; Sigma-Aldrich, St. Louis, MO, USA; Cat. No. S0130) dissolved in citrate buffer (pH 4.5) at a dose of 150 mg/kg body weight. Mice in the non-diabetic control (NC) group were injected with an equal volume of citrate buffer. Fasting blood glucose (FBG) levels were measured 72 h after STZ injection using a glucose meter (Roche, Basel, Switzerland; Accu-Chek Active). Mice with FBG ≥ 16.7 mmol/L were considered diabetic and included in the study.

Diabetic mice were randomly divided into three groups: diabetic model (DM) group, DM + shRNA-negative control (shRNA-NC) group, and DM + AAV-shRNA-Ltb4r1 group (n = 6 per group). The AAV-shRNA-Ltb4r1 and AAV-shRNA-NC vectors (serotype: AAV9, titer: 1 × 10¹² vg/mL) were constructed by Hanbio Biotechnology (Shanghai, China). Mice in the DM + shRNA-NC and DM + AAV-shRNA-Ltb4r1 groups were injected with 100 µL of the corresponding AAV vector via the tail vein, while mice in the NC and DM groups were injected with an equal volume of normal saline. All mice were continuously fed for 8 weeks after injection. During the experimental period, body weight and FBG levels were monitored weekly.

Renal tissue sample preparation

At the end of the 8-week treatment period, all mice were anesthetized with pentobarbital sodium (50 mg/kg, intraperitoneal injection; Sigma-Aldrich, St. Louis, MO, USA; Cat. No. P3761). Blood samples were collected from the orbital venous plexus, centrifuged at 3000 rpm for 15 min to obtain serum, and stored at -80 °C for subsequent analysis. Mice were then sacrificed by cervical dislocation, and the kidneys were quickly excised. After removing the renal capsule, one kidney was fixed in 4% paraformaldehyde for histological analysis, and the other was snap-frozen in liquid nitrogen and stored at -80 °C for RNA extraction, protein extraction, and ELISA detection. For tissue homogenate preparation, frozen renal tissue was weighed, homogenized in cold PBS (1:10, w/v) using a tissue homogenizer (Servicebio, Wuhan, China; Cat. No. SB-5900D), and centrifuged at 12,000 rpm for 20 min at 4 °C. The supernatant was collected for ELISA analysis.

Macrophage differentiation and polarization

THP-1 monocytes were differentiated into M0 macrophages by a 12-hour exposure to 320 nmol/L phorbol 12-myristate 13-acetate (PMA; Sigma-Aldrich, St. Louis, MO, USA; Cat. No. P8139), succeeded by a 24-hour incubation in fresh culture medium. Subsequent polarization towards the M1 phenotype was induced by stimulation with 100 ng/mL lipopolysaccharide (LPS; LPS25, Sigma-Aldrich, St. Louis, MO, USA) and 20 ng/mL interferon-γ (IFN-γ; 300-02-500UG, PeproTech, Cranbury, NJ, USA) for 36 h. M2 macrophages were generated by treating with 20 ng/mL interleukin-4 (IL-4; 200-04-20UG, PeproTech, Cranbury, NJ, USA) and 20 ng/mL interleukin-13 (IL-13; 200-13-10UG, PeproTech, Cranbury, NJ, USA) for 36 h. Cell culture was carried out in a CO₂ incubator (BC-J160S, Shanghai Boxun). Morphological evaluation was performed using an inverted phase contrast microscope (Nikon TS100). All cell culture procedures were conducted in a biosafety cabinet (SW-CJ-2FD, Shanghai Boxun).

Immunofluorescence staining

Following 24 h of culture, cells were subjected to fixation using 4% paraformaldehyde (Leagene, Beijing, China; Cat. No. DF0135) for 20 min. Subsequently, cells were permeabilized utilizing 0.5% Triton X-100 (Beyotime, Shanghai, China; Cat. No. ST795) and blocked with 5% bovine serum albumin (BSA; Beyotime, Shanghai, China; Cat. No. ST023) prior to incubation with primary antibodies against CD14 (ab182422, Abcam, Shanghai, China, 1:200), CD86(ab228485, Abcam, Shanghai, China, 1:200), CD163 (ab182422, Abcam, 1:200), and CD206 (ab64693, Abcam, 1:200) for 2 h. For renal tissue immunofluorescence, proximal tubular epithelial cells were identified using aquaporin-1 (AQP1) antibody (ab9566, Abcam, 1:200), and M1 macrophages were identified using CD86 antibody (ab228485, Abcam, 1:200). After the blocking procedure, cells were exposed to an Alexa Fluor™ 488-labeled secondary antibody (Thermo Fisher Scientific, Waltham, MA, USA; Prod. No. A-11008) for 60 min. For nuclear visualization, DAPI (Beyotime Biotechnology, Shanghai, China; Catalog No. C1005) was employed as a counterstain for 10 min. An inverted fluorescence microscope (DMI3000B, Leica, Germany) was used to acquire the resulting pictures.

Renal tubular epithelial cell model and high-glucose induction

To investigate the dose-dependent effect of glucose, HK-2 cells were first treated with a gradient of D-glucose concentrations: 5.5, 15, 30, and 45 mmol/L (Sigma-Aldrich, St. Louis, MO, USA; Prod. No. G7021). An osmotic control group supplemented with mannitol (Sigma-Aldrich, St. Louis, MO, USA; Cat. No. M9625) to match the osmolarity of 30 mmol/L D-glucose was additionally set up to exclude osmotic effects. Five experimental groups of HK-2 cells were established: a control group (Con) maintained in a 5.5 mmol/L D-glucose-containing medium (Sigma-Aldrich, St. Louis, MO, USA; Prod. No. G7021), and a high-glucose group (HG) sustained in a 30 mmol/L D-glucose-supplemented medium; HG + M0 group co-cultured with M0 macrophages; HG + M1 group co-cultured with M1 macrophages; and HG + M2 group co-cultured with M2 macrophages. The Transwell system (3421, Corning, Corning, NY, USA) was used for indirect co-culture. HK-2 cells (5 × 10⁵ cells per well) were introduced into the lower compartment of a 6-well plate, subsequently cultured with either 5.5 mmol/L or 30 mmol/L D-glucose-based medium. THP-1 cells (1.5 × 10⁶ cells/insert) were induced to M0, M1, or M2 phenotypes and seeded in the Transwell inserts. The upper and lower chambers were not in direct contact but allowed for cytokine exchange. After 24 h of co-culture, cells were collected for subsequent experiments.

Cell proliferation assay

After 24 h of treatment, HK-2 cells from each group were collected and seeded at a density of 2,000 cells/well in a 96-well plate. Following the addition of 10 µL of CCK-8 reagent (C0038, Beyotime Biotechnology, Shanghai, China) to each well, the cells were maintained at 37 °C for 2 h. Absorbance values were obtained at 450 nm. The relative growth rate (RGR) was calculated by normalizing the experimental group absorbance against the control group, using the formula: (experimental group absorbance - blank absorbance) / (control group absorbance - blank absorbance).

Cell apoptosis

Cellular apoptosis was quantified via Annexin V-FITC/PI apoptosis detection assay (Yeasen, Shanghai, China; Cat. No. 40302ES50), adhering to the supplier’s guidelines. Analysis of the stained samples was performed utilizing a CytoFLEX flow cytometer (Beckman Coulter, Brea, CA, USA).

Quantitative real-time PCR (qPCR) analysis

Total RNA was extracted from cells or renal tissues using TRIzol reagent (Invitrogen, Carlsbad, CA, USA; Cat. No. 15596026) according to the manufacturer’s instructions. RNA purity and concentration were determined using a NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific). Reverse transcription of 1 µg total RNA into cDNA was performed using a PrimeScript™ RT reagent kit with gDNA Eraser (Takara, Dalian, China; Cat. No. RR047A) to eliminate genomic DNA contamination. qPCR was conducted on a StepOnePlus Real-Time PCR System (Thermo Fisher Scientific) using TB Green^® Premix Ex Taq™ II (Takara, Dalian, China; Cat. No. RR820A). The reaction conditions were: 95 °C for 30 s, followed by 40 cycles of 95 °C for 5 s and 60 °C for 30 s. Relative gene expression levels were calculated using the 2⁻ΔΔCt method, with GAPDH as the internal reference gene. The primer sequences used are listed in Table 1.

Table 1.

Primer sequences

Species	Gene	Direction	Sequence (5’→3’)
Human	LTB4R1	F	CCTGTGTCACTATGTCTGCGGA
Human	LTB4R1	R	ATCGCCTTGGTGCGTAGCTTCT
Human	GAPDH	F	GTCTCCTCTGACTTCAACAGCG
Human	GAPDH	R	ACCACCCTGTTGCTGTAGCCAA
Mouse	LTB4R1	F	CTGCTGGTGCTGAACTTGG
Mouse	LTB4R1	R	CCGTGATGGCTTCGAAGAG
Mouse	cleaved caspase-3	F	GAGCTTGGAACGCGAAGAAA
Mouse	cleaved caspase-3	R	TTGCGAGCTGACATTCCAGT
Mouse	NF-κB p65	F	TCCTGTTCGAGTCTCCATGCAG
Mouse	NF-κB p65	R	GGTCTCATAGGTCCTTTTGCGC
Mouse	GAPDH	F	TGTGATGGGTGTGAACCACGAGA
Mouse	GAPDH	R	TTTTTGTGGGTGCAGCGAA

Western blot analysis

Protein lysates were generated for Western blotting via extraction with RIPA buffer (Solarbio, Beijing, China; Cat. No. R0010). rotein samples from the same batch were separated by SDS-PAGE gel electrophoresis (Servicebio, Wuhan, China; Cat. No. G2018) and subsequently transferred onto the same polyvinylidene difluoride (PVDF) membrane (Amersham, Freiburg, Germany; Cat. No. 10600023). T Based on differences in molecular weight between target proteins and the internal control GAPDH, the membrane was cut into separate strips and sequentially incubated with the corresponding primary antibodies, followed by incubation with horseradish peroxidase (HRP)-conjugated secondary antibodies. The secondary antibodies used were HRP-conjugated goat anti-rabbit IgG (1:5000 dilution; Abcam, UK; Cat. No. ab6721) or HRP-conjugated rabbit anti-mouse IgG (H + L) (1:5000 dilution; Abcam, Cat. No. ab6728). Protein bands were visualized using an enhanced chemiluminescence (ECL) detection system (Thermo Fisher Scientific, Waltham, MA, USA; Cat. No. 34580), and band intensities were quantified using ImageJ software (National Institutes of Health, Bethesda, MD, USA). All target proteins and the loading control were derived from the same gel and transferred onto the same membrane, ensuring accurate normalization using GAPDH. Primary antibodies used were as follows: for human samples, CD86 (ab 220188, Abcam, Shanghai, China, 1:1000), CD163 (1:1000, Abcam, ab182422), CD206 (1:1000, Abcam, ab64693), LTB4R1 (1:1000, Cell Signaling Technology, 74190), CD14 (ab221678, Abcam, 1:1000), CD86 (ab 119857, Abcam, 1:1000), cleaved caspase-3 (1:1000, CST, 9661), phosphorylated NF-κB p65 (p-p65; 1:1000, CST, 3031), and GAPDH (1:5000, Abcam, ab9485) as an internal reference.

Enzyme-Linked Immunosorbent Assay (ELISA)

The concentrations of TNF-α, IL-1β, and IL-10 were measured using commercial ELISA kits, following the manufacturers’ protocols. The ELISA kits used were: TNF-α (ab208348 for mouse and ab181421 for human, Abcam, Shanghai, China), IL-1β (ab197742 for mouse and ab214025 for human, Abcam, Shanghai, China), and IL-10 (ab255729 for mouse and ab185986 for human, Abcam, Shanghai, China).

Statistical analysis

Prior to performing parametric statistical tests, all experimental data were first subjected to normality test (Shapiro-Wilk test) using SPSS 26.0 software (IBM Corp., Armonk, NY, USA) to verify whether the data conformed to a normal distribution, and variance homogeneity test (Levene’s test) also using SPSS 26.0 software to verify the homogeneity of variance among groups. Only data that passed both normality and variance homogeneity tests were subjected to parametric tests. GraphPad Prism 8 software (GraphPad Software, San Diego, CA, USA) was employed for the statistical examination of the data, which are expressed as the average ± SD. One-way ANOVA was utilized for multiple group comparisons, whereas Student’s t-test was utilized for comparisons between two groups. We defined statistical significance as a P-value less than 0.05. We performed each experiment independently in triplicate.

Results

Differentiation and phenotypic identification of THP-1 cells into macrophages

Microscopic imaging showed that PMA-treated THP-1 cells underwent typical macrophage morphological transformation, shifting from a suspended monocytic state to adherent, irregularly shaped macrophages (Fig. 1). Immunofluorescence staining was performed to verify the phenotypic characteristics of undifferentiated and polarized macrophages. The M0 lineage marker CD14 (Fig. 2A) was abundantly expressed in unpolarized M0 macrophages, while its fluorescence intensity was markedly attenuated in M1 macrophages polarized with LPS and IFN-γ and moderately recovered (lower than M0 levels) in IL-4/IL-13-induced M2 macrophages. The M1-specific costimulatory marker CD86 (Fig. 2B) was barely detectable in M0 cells, strongly upregulated in M1 macrophages, and substantially diminished in the M2 phenotype. The canonical M2 marker CD163 (Fig. 2C) exhibited weak basal expression in M0 cells, was nearly absent in M1 macrophages, and showed robust fluorescence signals in M2 macrophages. Similarly, the M2-specific mannose receptor CD206 (Fig. 2D) presented low basal levels in M0 cells, was downregulated in M1 macrophages, and was profoundly enriched in M2 macrophages. Collectively, these immunofluorescence results confirm the successful differentiation of THP-1 monocytes into macrophages and the efficient generation of distinct M0, M1, and M2 macrophage phenotypes with unique marker expression profiles.

Fig. 1.

Morphological changes of THP-1 cells upon differentiation into macrophages. n = 3 per group

Fig. 2.

Immunofluorescence staining of macrophage phenotypes. A: Staining of CD14 (a marker associated with M0 macrophages). B: Staining of CD86 (a M1-specific co-stimulatory marker). C: Staining of CD163 (a M2-specific marker). D: Staining of CD206 (a M2-specific mannose receptor). Top row: Fluorescence signal of the target marker (green); Middle row: DAPI-stained nuclei (blue); Bottom row: Merged images of the marker and DAPI. n = 3 per group

Dose-dependent regulation of LTB4R1 expression and cell apoptosis by glucose

To determine the regulatory effect of glucose on LTB4R1 expression, HK-2 cells were exposed to a gradient of glucose concentrations (5.5, 15, 30, and 45 mM). Quantitative analysis demonstrated that LTB4R1 mRNA levels increased progressively with elevated glucose concentrations (Fig. 3A; all P < 0.01). LTB4R1 protein expression exhibited a consistent dose-dependent upregulation across the glucose gradient (Fig. 3B; all P < 0.001). Flow cytometry-based apoptotic detection using Annexin V-FITC/PI staining further revealed a gradual increase in cellular apoptosis rates as glucose concentrations rose (Fig. 3C; P < 0.01). These data demonstrate a strict dose-dependent relationship, whereby hyperglycemia drives progressive LTB4R1 upregulation at both transcriptional and translational levels, accompanied by aggravated tubular epithelial cell apoptosis.

Fig. 3.

Dose-dependent effects of glucose on LTB4R1 expression and cell apoptosis. A: Quantitative analysis of relative LTB4R1 mRNA expression in cells treated with 5.5, 15, 30, or 45 mM glucose. B: Representative Western blot images showing LTB4R1 protein expression and GAPDH as the internal loading control, together with densitometric quantification of LTB4R1 normalized to GAPDH. C: Representative Annexin V-FITC/PI flow cytometry scatter plots showing apoptosis and quantitative analysis of the apoptotic rate (Annexin V-positive cells). n = 3 per group. Statistical significance: ns (not significant), **P < 0.01, ***P < 0.001

High glucose-specific injury to renal tubular epithelial cells: modulation by macrophage polarization

A mannitol-matched osmotic control group was included to exclude osmotic interference. No significant differences in cell function or molecular indicators were observed between the normal control and osmotic control groups, verifying that the cellular alterations observed in this study were specifically induced by hyperglycemia rather than osmotic stress. High glucose (HG) treatment significantly reduced the viability of HK-2 cells compared with control and osmotic control groups (P < 0.01). Compared with HG treatment alone, co-culture with M1 macrophages further suppressed cell viability (P < 0.01), whereas M2 macrophage co-culture effectively restored cell viability (P < 0.01); M0 macrophage co-culture exerted no notable impact on HG-induced cell viability changes.

Western blot analysis showed that HG exposure significantly upregulated LTB4R1 and CD68 expression (P < 0.001) and downregulated M2 macrophage markers CD163 and CD206 (P < 0.01). Relative to the single HG group, co-culture with M1 macrophages further elevated LTB4R1 and CD68 levels (P < 0.01), while M2 macrophage co-culture reversed these upregulations (P < 0.001). Consistently, M2 co-culture fully restored the HG-suppressed expression of CD163 and CD206 (P < 0.001). Cytokine quantification revealed that HG significantly increased the secretion of pro-inflammatory TNF-α and IL-1β (P < 0.001) and decreased the level of anti-inflammatory IL-10 (P < 0.001). M1 macrophage co-culture further amplified HG-triggered pro-inflammatory cytokine production and IL-10 reduction, whereas M2 macrophage co-culture normalized TNF-α and IL-1β to basal levels and markedly elevated IL-10 secretion (all P < 0.001). Furthermore, HG significantly increased HK-2 cell apoptosis (P < 0.001), an effect aggravated by M1 macrophage co-culture and prominently alleviated by M2 macrophage co-culture (all P < 0.001).

Fig. 4.

High glucose-specific injury to HK-2 cells and its modulation by macrophage polarization. A: Cell viability (CCK-8 assay). B: Western blot (WB) analysis of LTB4R1, CD68 (M1 marker), CD163 and CD206 (M2 markers); GAPDH served as a loading control. C: Cytokine secretion (TNF-α, IL-1β: pro-inflammatory; IL-10: anti-inflammatory) measured by ELISA. D: Representative Annexin V-FITC/PI flow cytometry plots for apoptosis and quantitative apoptotic rate. Control (normal glucose), HG (high glucose), Osmotic control (mannitol-matched osmolarity control), HG + M0/M1/M2 (HG-treated HK-2 cells co-cultured with M0/M1/M2 macrophages). n = 3 per group. Statistical significance: ns (not significant), *P < 0.05, **P < 0.01, ***P < 0.001

Effect of LTB4R1 silencing on renal injury in STZ-induced diabetic C57BL/6 mice

To interrogate the in vivo role of LTB4R1 in diabetic kidney disease, we established an STZ-induced diabetic model in C57BL/6 mice (n = 6 per group) and silenced renal Ltb4r1 via AAV-mediated shRNA. Compared with non-diabetic controls, DM exhibited marked elevations in serum creatinine, blood urea nitrogen, uric acid and 24-h urinary protein (all P < 0.001; Fig. 5A–D). Kidney hematoxylin and eosin staining revealed tubular epithelial cell swelling, vacuolar degeneration, luminal dilation, interstitial edema and inflammatory cell infiltration (Fig. 5E). Treatment with a non-targeting control shRNA (DM + shRNA‑NC) did not significantly alter any renal function or histopathological parameter (all P > 0.05). In striking contrast, Ltb4r1-targeted shRNA significantly reduced all renal injury markers (all P < 0.001) and markedly ameliorated the pathological damage. Dual immunofluorescence colocalization analysis demonstrated that LTB4R1 was significantly colocalized with proximal tubular marker AQP1 and M1 macrophage marker CD86 in diabetic kidneys (Fig. 5F–I). Quantitative analysis showed that fluorescence intensities of AQP1-, CD86-, and LTB4R1-positive signals were significantly higher in the diabetic group compared with the normal control group (all P < 0.001). After Ltb4r1 silencing, LTB4R1 fluorescence intensity was markedly reduced, accompanied by concurrent decreases in AQP1 and CD86 signal intensities (all P < 0.001). The highly consistent expression patterns, together with morphological colocalization results, indicate that proximal tubular epithelial cells and M1 macrophages are the major cellular sources of LTB4R1 in diabetic kidneys.

Fig. 5.

Ltb4r1 silencing improves renal function, alleviates renal pathological damage and defines the cellular localization of LTB4R1 in STZ-induced diabetic mice. A: Quantitative analysis of serum creatinine; B: Quantitative analysis of blood urea nitrogen; C: Quantitative analysis of serum uric acid; D: Quantitative analysis of 24-h urinary protein; E: Representative H&E staining images of renal tissues; F: Representative immunofluorescence colocalization images of LTB4R1 (green) and tubular epithelial cell marker AQP1 (red); G: Quantitative analysis of relative fluorescence intensity of AQP1 and LTB4R1; H: Representative immunofluorescence colocalization images of LTB4R1 (green) and M1 macrophage marker CD86 (red); I: Quantitative analysis of relative fluorescence intensity of CD86 and LTB4R1. NC, non-diabetic control; DM, STZ-induced diabetic model; DM + shRNA-NC, diabetic mice treated with negative-control shRNA; DM + AAV-shRNA-Ltb4r1, diabetic mice with AAV-mediated Ltb4r1 knockdown. n = 6 per group. ns, no significant difference; ***P < 0.001

Molecular analyses revealed that in DM kidneys, transcript levels of Ltb4r1, cleaved caspase‑3 and NF‑κB p65 were significantly elevated (all P < 0.001; Fig. 6A). At the protein level, LTB4R1, the M1 markers CD14 and CD86, cleaved caspase‑3 and phosphorylated NF‑κB p65 were all upregulated, whereas the M2 markers CD163 and CD206 were downregulated (all P < 0.001; Fig. 6B). shRNA‑NC treatment had no effect on any of these proteins (all P > 0.05), whereas Ltb4r1 knockdown significantly reduced pro-inflammatory and pro-apoptotic proteins, suppressed NF‑κB pathway activity (all P < 0.05), and restored M2 marker expression (all P < 0.001), thereby reversing the aberrant molecular profile induced by diabetes. Concordantly, DM kidneys contained elevated levels of the pro-inflammatory cytokines TNF‑α and IL‑1β and reduced levels of the anti-inflammatory cytokine IL‑10 (all P < 0.001; Fig. 6C), and exhibited a significantly higher apoptotic rate (P < 0.001; Fig. 6D). Ltb4r1 silencing markedly suppressed the pro-inflammatory cytokines, increased IL‑10 production (all P < 0.001), and substantially reduced renal apoptosis (P < 0.001). Collectively, these results demonstrate that in the diabetic kidney, LTB4R1 is predominantly expressed in tubular epithelial cells and M1 macrophages; its silencing alleviates renal dysfunction and pathological injury by promoting M2 macrophage polarization and restraining excessive NF‑κB activation, thereby attenuating renal inflammation and apoptosis.

Fig. 6.

Ltb4r1 silencing modulates renal molecular expression, inflammatory response and cell apoptosis in diabetic mice. A: Transcript levels of Ltb4r1, cleaved caspase-3 and NF-κB p65 detected by qPCR; B: Protein expression of LTB4R1, CD14, CD86 (M1 markers), CD163/CD206 (M2 markers), cleaved caspase-3 and p-NF-κB p65 measured by Western blot. GAPDH was used as the internal reference; C: Secretion levels of pro-inflammatory cytokines TNF-α, IL-1β and anti-inflammatory cytokine IL-10 detected by ELISA; D: Representative Annexin V-FITC/PI flow cytometry plots and quantitative analysis of renal cell apoptotic rate. n = 6 per group. ns, no significant difference; *P < 0.05; **P < 0.01; ***P < 0.001

Discussion

This study systematically investigated the roles of distinct macrophage phenotypes and LTB4R1 in DKD, with a particular focus on their contributions to renal tubular epithelial cell injury. By establishing M0, M1, and M2 macrophage models derived from THP-1 cells and exposing them to a high-glucose microenvironment, we examined their effects on tubular epithelial cell proliferation, apoptosis, and LTB4R1 expression. These in vitro findings were further corroborated by in vivo animal experiments, providing new insights into macrophage-mediated renal injury in DKD.

PMA-induced differentiation of THP-1 monocytes into macrophages was fully validated in our system. Immunofluorescence profiling of canonical lineage and polarization markers—CD14 for M0, CD86 for M1, and CD163/CD206 for M2—confirmed successful differentiation and stable polarization plasticity. As expected, LPS/IFN-γ stimulation efficiently drove M1 polarization with elevated CD86 expression, whereas IL-4/IL-13 treatment skewed cells toward an M2 phenotype marked by robust CD163 and CD206 upregulation. These results are consistent with a prior report by Li et al., which verified the specific induction effects of these cytokine regimens on macrophage polarization phenotypes [23].

Hyperglycemic stress rendered renal tubular epithelial cells more susceptible to apoptotic injury, an effect further exacerbated by M1 macrophage co-culture to accelerate DKD progression. Glucose-induced tubular damage exhibited clear dose dependence across physiological to pathological concentrations (5.5, 15, 30 and 45 mmol/L). Notably, mannitol osmotic control experiments confirmed that tubular injury was triggered specifically by hyperglycemia, rather than non-specific osmotic perturbation. Our findings align with in vivo evidence showing excessive M1 macrophage infiltration exacerbates renal damage in diabetic mouse models [24]. Of note, Wang et al. reported that moderate M1 macrophage recruitment can initiate early inflammation and clear cellular debris to facilitate tissue repair and regeneration [25]. Such apparent discrepancies likely stem from differences in glucose dosage, intervention duration and cellular experimental systems used across studies. In contrast, our results demonstrate that M2 macrophages effectively protect tubular epithelial cells against high glucose-induced damage via inherent anti-inflammatory and reparative functions. As reported in previous mechanistic studies, M2 macrophages alleviate HK-2 cell injury and apoptosis induced by calcium oxalate and other stressors, largely through suppressing NADPH oxidase/ROS signaling and modulating the p38 MAPK and PI3K/Akt pathways [26]. Collectively, these findings highlight the decisive role of macrophage phenotypic switching in DKD pathogenesis, and suggest that targeting macrophage polarization could preserve tubular epithelial cell viability and modify disease trajectories. Consistent with established evidence from renal fibrosis research, this regulatory axis also participates in fibroblast activation and epithelial–mesenchymal transition (EMT): M1 macrophages promote EMT primarily via pro-inflammatory cytokine secretion, whereas M2 macrophages attenuate renal fibrotic progression by restraining this pathological transition [27, 28].

Tubular epithelial cell dysfunction and injury are early and irreversible events driving DKD progression. Accumulating evidence indicates that hyperglycemia disrupts tubular structural integrity and cellular homeostasis, concurrently suppressing proliferation and promoting apoptotic cell death [29]. Our in vitro data recapitulate these findings, showing that high glucose markedly inhibits HK-2 cell proliferation and induces apoptosis. In vivo, renal apoptotic cell abundance was markedly elevated in diabetic mice, and this phenotype was substantially rescued by LTB4R1 silencing. These observations are in line with established mechanisms whereby hyperglycemia triggers tubular apoptosis through oxidative stress, advanced glycation end product accumulation and hyperactivated inflammatory signaling cascades [30, 31]. Mechanistically, M1 macrophages amplify hyperglycemic tubular injury by enhancing the secretion of pro-inflammatory TNF-α and IL-1β while repressing anti-inflammatory IL-10 production, thereby exacerbating epithelial apoptosis. Conversely, M2 macrophages exert cytoprotective effects by reshaping the cytokine profile toward an anti-inflammatory and reparative signature. Consistent with published study, M1 macrophages promote tubular apoptosis via TNF-α/IL-1β release [32, 33], whereas M2 macrophages confer renoprotection by secreting anti-inflammatory mediators including IL-10 and TGF-β, supporting their dual roles in tissue repair and fibrosis restraint [34].

Leukotriene B4 (LTB4), a bioactive lipid metabolite generated via the arachidonic acid cascade, is a key regulator of innate inflammation and immune cell activation [35]. LTB4 drives macrophage recruitment and functional activation primarily through its high-affinity receptor LTB4R1 [36]. The LTB4–LTB4R1 signaling axis is hyperactivated in metabolic disorders such as obesity and diabetes, perpetuating chronic inflammatory stress [37]. In preclinical renal injury models, LTB4R1 is markedly upregulated in damaged kidney tissues, and pharmacological or genetic LTB4R1 blockade alleviates renal inflammation and parenchymal injury, implying its indispensable role in renal interstitial inflammation and tissue deterioration [38]. In our study, LTB4R1 expression was elevated by high glucose in a dose-dependent manner in vitro, and this upregulation was further augmented upon co-culture with M1 macrophages, accompanied by increased tubular epithelial apoptosis. Importantly, our in vitro data only establish a strong correlative relationship between LTB4R1 upregulation and hyperglycemic apoptosis; direct causal validation via pharmacological LTB4R1 inhibition is absent in cellular experiments. Future in vitro loss-of-function assays using receptor antagonists or targeted siRNA will be required to delineate whether elevated LTB4R1 is a direct driver of tubular apoptosis or a secondary stress-responsive event. In vivo, diabetic kidneys exhibited robust LTB4R1 induction, and AAV-mediated Ltb4r1 knockdown suppressed LTB4R1 expression, reduced NF-κB p65 phosphorylation, and restored inflammatory cytokine homeostasis. These in vivo genetic interference results provide functional evidence supporting LTB4R1 as an important mediator of inflammation and injury in the diabetic kidney, consistent with its previously documented pro-inflammatory and pro-injury roles in other types of renal disease. Integrating in vitro and in vivo observations, LTB4R1 upregulation is tightly coupled with M1 macrophage-elicited tubular damage, with NF-κB signaling serving as a plausible downstream effector. Nevertheless, the precise hierarchical regulatory relationship remains incompletely resolved, warranting further rigorous loss- and gain-of-function assays to dissect the causal ordering of LTB4R1 induction, NF-κB activation and tubular apoptosis.

Mechanistically, prior studies have demonstrated that LTB4R1 ligation activates Gα_i-dependent signaling, which relieves SOCS1-mediated repression of MyD88. This de-repression elevates MyD88 abundance and subsequently amplifies canonical NF-κB signaling. The activated cascade further induces pro-inflammatory cytokine production and oxidative stress mediator release, ultimately triggering apoptosis and tissue damage in target cells [19]. Emerging evidence also reveals preferential BLT1 (LTB4R1) enrichment in pro-inflammatory macrophages, with M1 subsets expressing inherently higher BLT1 levels than M2 counterparts [15]. This molecular characteristic explains the exaggerated NF-κB-driven inflammatory response observed in our M1 macrophage co-culture system, and aligns well with our in vivo results showing that Ltb4r1 inhibition blunts NF-κB activation and renal apoptosis. Collectively, our findings support a conserved pro-inflammatory LTB4R1–NF-κB signaling axis that contributes to DKD pathogenesis.

Despite providing novel immunological insights into DKD pathogenesis, this study has several limitations. First, PMA-induced THP-1-derived macrophages may exhibit non-physiological signaling and do not fully recapitulate primary or kidney-resident macrophages, warranting validation in primary models. Second, metabolic stress parameters (e.g., ROS, mitochondrial function, ATP) were not systematically assessed, leaving the interplay between LTB4R1 and metabolic injury undefined. Third, our in vitro studies lacked direct pharmacological inhibition or genetic knockdown of LTB4R1, limiting causal inference between high-glucose–induced LTB4R1 upregulation and tubular apoptosis; confirmatory in vitro intervention experiments are required. Fourth, endpoint macrophage phenotyping was not performed after co-culture; while the maintained M2-characteristic cytokine and marker profiles suggest functional retention, high glucose may induce partial repolarization, warranting independent endpoint verification in future studies. Fifth, the in vivo LTB4R1 silencing experiments were conducted only in diabetic mice without parallel shRNA intervention groups (NC + shRNA-NC / NC + shRNA-Ltb4r1) in non-diabetic controls. This limits the ability to exclude potential off-target effects of the viral vector and to evaluate the impact of LTB4R1 knockdown on normal kidney homeostasis. Sixth, short-term in vitro experiments and the STZ-induced acute diabetic model do not fully capture the chronic, progressive nature of DKD, including sustained inflammation and fibrosis; future studies using spontaneous diabetic models (e.g., db/db mice) are needed. Finally, the absence of renal biopsy validation from DKD patients precludes definitive clinicopathological correlations, and the therapeutic potential of M2 macrophage-targeted strategies and M2-derived exosomes warrants further investigation.

Conclusion

This study demonstrates that under high-glucose conditions, activation of LTB4R1 promotes M1 macrophage polarization, activates the NF-κB signaling pathway, and exacerbates inflammatory cytokine imbalance, thereby aggravating renal tubular epithelial cell injury. Conversely, LTB4R1 silencing confers renal protection by reversing these pathogenic processes. These findings refine the regulatory network linking lipid signaling, immune cell activation, and tubular injury in DKD and identify LTB4R1 as a potential target deserving further preclinical evaluation. Future studies addressing the aforementioned limitations—particularly those incorporating patient-derived samples—are essential to facilitate the translation of LTB4R1-targeted strategies from bench to bedside.

Supplementary Information

Below is the link to the electronic supplementary material.

Author contributions

W.C. and Q.L. wrote the main manuscript. C.N. and S.L. prepared the data collection. J.C. and G.L. prepared tables. S.L. and F.D. analyse and interpret of results. All authors reviewed the results and approved the final version of the manuscript. All authors would be informed each step of manuscript processing including submission, revision, revision reminder, etc.

Funding

This study was supported by Shandong Provincial Natural Science Foundation (Grant No.ZR2022MH232).

Data availability

The datasets used and analysed during the current study available from the corresponding author on reasonable request.

Declarations

Ethics approval and consent to participate

This study was approved by the Ethics Committee. Informed consent was obtained from all the participants. All methods were carried out in accordance with Declaration of Helsinki.

Consent for publication

Verbal informed consent was obtained from all individual participants included in the study. All authors have read and approved the final version of the manuscript submitted for publication.

Declaration of generative AI in scientific writing

The authors did not use generative AI or AI-assisted technologies in the development of this manuscript.”

Competing interests

The authors declare no competing interests.