M1 macrophage‐derived exosomes aggravate diabetic nephropathy by regulating the WTAP/S1PR2 axis

ABSTRACT

Background

Diabetic nephropathy (DN) is a severe microvascular complication of diabetes mellitus. The specific role of M1 macrophage‐derived exosomes in DN progression remains largely unexplored.

Methods

THP‐1 monocytes were differentiated into M0 macrophages and polarized into M1 macrophages for exosome extraction. WTAP was silenced using shRNA to generate WTAP‐deficient exosomes. Glomerular endothelial cells (GECs) were exposed to high glucose (HG) and co‐incubated with modified exosomes. Cell viability, oxidative stress, apoptosis, barrier function, and angiogenic capacity were assessed. The WTAP‐S1PR2 interaction was validated by RIP, MeRIP, and dual‐luciferase assays. In vivo, db/db mice received tail vein injections of respective exosomes for 8 weeks, followed by assessments of renal function, histopathology, and inflammation.

Results

M1 exosomes were internalized by GECs. WTAP delivered by shNC/M1‐Exo bound S1PR2 mRNA, enhancing its m6A modification and stability, thereby activating the RhoA/ROCK1 axis and aggravating HG‐induced GEC injury, oxidative stress, apoptosis, and endothelial permeability. Conversely, shWTAP/M1‐Exo attenuated these effects, and S1PR2 overexpression reversed the protective effects. In vivo, shWTAP/M1‐Exo improved renal function, ameliorated histopathological damage and fibrosis, and reduced systemic inflammation in db/db mice.

Conclusion

M1 macrophage‐derived exosomes promote DN progression by delivering WTAP to stabilize S1PR2 mRNA in an m6A‐dependent manner. Engineered exosomes lacking WTAP represent a promising targeted nanomedicine strategy for DN treatment.

INTRODUCTION

Diabetic nephropathy (DN) serves as the primary cause of end‐stage renal disease and represents a microvascular complication in populations with glucose metabolism disorders. ¹ , ² DN is frequently accompanied by profound risks of cardiovascular and renal deterioration, as well as elevated mortality rates. ³ , ⁴ , ⁵ The organic renal damage induced by this condition predominantly targets the glomeruli. Excessive glucose concentrations lead to the abnormal thickening of the glomerular basement membrane and a marked reduction in the effective filtration area, eventually culminating in localized tissue sclerosis and extensive fibrosis. ⁶ Glomerular endothelial cells (GECs), distributed along the capillary inner walls, constitute a crucial element in maintaining the structural integrity of the vascular barrier. ⁷ Being directly exposed to the circulatory environment, these cells are highly susceptible to multiple insults triggered by high glucose (HG), including oxidative stress, inflammatory responses, and lipotoxicity. ⁷ , ⁸ Consequently, a comprehensive elucidation of the underlying mechanisms by which HG induces GEC injury holds promise for paving novel pathways for the intervention of such renal pathologies.

As a crucial component of the body's defense system, macrophages are widely distributed across the glomeruli and interstitial regions. Literature has confirmed that these cells are deeply involved in the pathogenesis of DN. ⁹ Typically, this cell population is classified into the proinflammatory M1 phenotype and the reparative M2 phenotype. Under hyperglycemic conditions, the body tends to drive the massive generation of the former while exerting negative regulation on the differentiation of the latter; such phenotypic polarization is a core driver in triggering renal inflammation and fibrotic alterations. ¹⁰ Exosomes are nanoscale membrane‐bound structures secreted by various cells in the body, primarily serving as communication mediators in bodily fluid circulation. ¹¹ Recent studies have revealed that vesicles derived from the M1 phenotype accelerate the progression of DN. For instance, these vesicles could bind to sirtuin 2 via miR‐212‐5p and block the Akt/glycogen synthase kinase‐3 beta (GSK‐3β)/β‐catenin signaling axis, thereby impairing normal insulin secretion from pancreatic β‐cells in obese type 2 diabetic mouse models and ultimately leading to glucose intolerance. ¹² Furthermore, the methyltransferase‐like 14 (METTL14) contained within them can rely on the m6A methylation modification of progestin and adipoQ receptor family member 3 (PAQR3) to exacerbate the apoptotic process, oxidative stress, and inflammatory responses of GECs under high‐glucose conditions. ¹³ Nevertheless, the specific regulatory network through which M1 phenotype‐derived exosomes mediate GEC injury still warrants more in‐depth investigation.

Within eukaryotes, N6‐methyladenosine (m6A) represents a highly prevalent class of chemical modifications on messenger RNA. ¹⁴ , ¹⁵ Regarding this methylation process, wilms tumor 1‐associated protein (WTAP) plays an indispensable role. This molecule is capable of precisely intervening in RNA splicing and is deeply involved in regulating numerous physiological activities, including cell expansion, cell cycle progression, and even embryonic growth. ¹⁶ , ¹⁷ Existing academic findings revealed that WTAP could facilitate m6A‐level alterations in tripartite motif containing 22 (TRIM22). This action disrupted normal mitochondrial functioning by relying on the ubiquitination and degradation mechanism of OPA1 mitochondrial dynamin‐like GTPase, ultimately propelling the onset and progression of DN. ¹⁸ WTAP‐induced m6A methylation of atonal homolog 8 (Atoh8) promoted cell proliferation and fibrosis in DN. ¹⁹ Additionally, sphingosine‐1‐phosphate receptors (S1PRs) constitute a superfamily of G protein‐coupled transmembrane receptors specifically dedicated to mediating various cellular cascades triggered by sphingosine‐1‐phosphate. ²⁰ As a member of the S1PRs family, S1PR2 can exacerbate the deterioration of this renal complication by stimulating the endothelial‐to‐mesenchymal transition while concurrently compromising the integrity of the endothelial barrier. ²¹

Preliminary bioinformatics analyses (based on the SRAMP and RBPsuit databases) indicate that the S1PR2 mRNA sequence harbors both potential m6A modification sites and WTAP protein‐binding motifs, suggesting that the posttranscriptional regulation of S1PR2 may be highly dependent on WTAP. Synthesizing the aforementioned literature, the present study proposes the following scientific hypothesis: M1‐Exo may accelerate the pathological progression of DN by mediating the aberrant expression of WTAP. The underlying molecular mechanism may involve WTAP catalyzing the m6A modification of S1PR2 mRNA to maintain or upregulate its expression level, thereby exacerbating HG‐induced injury in GECs. This study aims to conduct an in‐depth validation of this hypothesis, with the goal of providing novel insights into the pathogenesis of DN and identifying potential intervention targets for the clinical targeted therapy of this disease.

MATERIALS AND METHODS

Cell culture and HG treatment

Human glomerular endothelial cells (HRGECs, GECs, Xuanke Biotech, Shanghai, China) were cultured in human glomerular endothelial cell complete medium (Procell, Wuhan, China). Human monocytes THP‐1 (Procell) were cultured in Roswell Park Memorial Institute‐1,640 medium (RPMI‐1640, Procell) supplemented with 10% fetal bovine serum (FBS, Procell), 1% penicillin/streptomycin (Procell), and 0.05 mM β‐mercaptoethanol (Maokang Biotech, Shanghai, China). All cell cultures were maintained in an incubator at 37°C with 5% CO₂. After being propagated to the logarithmic growth phase, the GECs were randomly allocated into two distinct experimental sets: a normal glucose (NG) control cohort and a HG treatment group. Specifically, the cells in the NG control cohort were maintained in endothelial cell complete medium supplemented with 5.5 mM glucose (Beyotime, Shanghai, China) for a duration of 48 h. In contrast, the HG treatment group was cultured using complete endothelial cell medium containing an elevated concentration of 30 mM glucose.

Differentiation induction and identification of THP‐1

THP‐1 cells were stimulated with 100 ng/mL phorbol‐12‐myristate‐13‐acetate (PMA, Thermo Fisher, Waltham, MA, USA) for 24 h to induce their differentiation into M0 phenotype macrophages. Subsequently, 100 ng/mL lipopolysaccharide (LPS, Maokang Biotech) and 20 ng/mL interferon‐gamma (IFN‐γ, Beyotime) were added to the system, and the cells were cultured for an additional 48 h to drive the polarization of M0 macrophages into the M1 phenotype. Following the aforementioned induction, both types of macrophages were collected, digested with trypsin (Beyotime), and counted. The cells were centrifuged at 1000 rpm for 1 min. An appropriate volume of sterile phosphate buffer solution (PBS) was added to resuspend the cells for a single wash. After discarding the supernatant and performing a second centrifugation, an anti‐CD80 antibody (Elabscience, Wuhan, China) was added to each tube, and the cells were incubated at 4°C for 3 min. Finally, the samples were analyzed on a flow cytometer to determine the expression level of CD80 protein on the cell surface.

Exosome isolation and identification

Well‐growing M1 macrophages were incubated under standard conditions for 48 h, after which the medium was replaced with RPMI‐1640 medium exosome‐depleted FBS (System Biosciences, Beijing, China) for a further 24‐h incubation. Subsequently, 50 mL of the culture supernatant was collected, and exosomes were isolated and purified according to the instructions of the ExoQuick kit (System Biosciences) for subsequent characterization. A 5 μL aliquot of the exosome suspension was dropped onto a transmission electron microscopy copper grid and left to stand for 10 min to allow the sample to settle naturally. Excess liquid was then absorbed using filter paper, followed by the addition of phosphotungstic acid solution to the grid for a 1‐min negative staining. The residual liquid was carefully blotted away with filter paper again. After the grid was completely dried under a heat lamp, it was mounted on a sample holder and placed under a transmission electron microscope (TEM) to observe the particle morphology and capture images. Additionally, exosomes were subjected to western blotting analysis to detect the specific markers tumor susceptibility gene 101 (TSG101), cluster of differentiation 81 (CD81), and CD9 in isolated exosomes, while simultaneously evaluating the abundance of the negative control protein Calnexin in cells. GECs were treated with 20 μg/mL exosomes for further study.

Fluorescent labeling and tracking of exosomes

The isolated exosomes were thoroughly mixed with 10 μM Dil dye (Beyotime) and incubated at 37°C in the dark for 30 min. Subsequently, the treated exosomes were co‐cultured with M1 macrophages at 37°C for 24 h. Following the co‐incubation, the cells were fixed at room temperature for 1 h. Finally, the cells were incubated with 4′,6‐Diamidino‐2‐Phenylindole (Beyotime) for 10 min and observed under a fluorescence microscope to evaluate and record the specific uptake of exosomes by the GECs.

Cell transfection

GenePharma (Shanghai, China) provided the small hairpin RNA targeting WTAP (shWTAP), S1PR2 overexpression plasmid (oe‐S1PR2), and the matched controls (shNC and oe‐NC). shWTAP and its negative control shNC were transfected into M1 macrophages and GECs using Lipofectamine 3,000 (Thermo Fisher), while oe‐S1PR2 and its negative control oe‐NC were transfected into GECs using Lipofectamine 3,000 (Thermo Fisher).

Quantitative real‐time polymerase chain reaction (qRT‐PCR)

Total RNA from cells was obtained using TsingZol (Tsingke, Shanghai, China). Subsequently, the extracted RNA samples were reverse transcribed into cDNA templates utilizing the AMV first‐strand cDNA synthesis kit (Baiao Leibo Technology Co., Ltd., Beijing, China). During the quantitative PCR amplification, β‐actin was selected as the internal reference gene, and SYBR Green (Thermo Fisher) was employed to perform the quantitative detection. Finally, the data were calculated using the 2^−ΔΔCT method to determine the abundance of the target gene mRNA in the cells. WTAP 5′‐TTGTAATGCGACTAGCAACCAA‐3′ and 5′‐GCTGGGTCTACCATTGTTGATCT‐3′; S1PR2 5′‐ATCGTGCTAGGCGTCTTTATCG‐3′ and 5′‐AGTGGGCTTTGTAGAGGATCG‐3′; β‐actin 5′‐CTTCGCGGGCGACGAT‐3′ and 5′‐CCACATAGGAATCCTTCTGACC‐3′.

Western blotting assay

Following the collection of samples from each experimental group, RIPA lysis buffer (Beyotime) was added, and the samples were placed on ice for 30 min to lyse the cells and tissues. Subsequently, the mixtures were centrifuged for 20 min, and the supernatants were carefully collected. The obtained protein extracts were then subjected to routine procedures, including separation by SDS polyacrylamide gel, membrane transfer, and blocking using 5% skimmed milk. Next, primary antibodies against the target proteins, including WTAP (1:8000, Proteintech, Rosemount, MN, USA), TSG101 (1:2000, Proteintech), CD81 (1:1000, ab79559, Abcam, Cambridge, MA, USA), CD9 (1:1000, ab236630, Abcam), Calnexin (1:5000, 10,427‐2‐AP, Proteintech), S1PR2 (1:1000, 21,180‐1‐AP, Proteintech), GTP‐bound RhoA protein (GTP‐RhoA, 1:1000, AF6352, Affinity, Nanjing, China), Rho‐associated, coiled‐coil containing protein kinase 1 (ROCK1, 1:8000, 21,850‐1‐AP, Proteintech), VE‐Cadherin (1:1000, ab33168, Abcam), and the internal reference β‐actin (1:5000, AF7018, Affinity) were added, followed by slow incubation in a shaker overnight. After repeated washes with TBST buffer, secondary antibodies (S0001/S0002, 1:5000, Affinity) were added. Finally, an ECL chemiluminescence system and a gel imaging device were utilized to analyze and record the specific expression abundance of the respective protein bands.

Cell counting kit‐8 (CCK‐8)

GECs were seeded into 96‐well plates. Following the corresponding experimental interventions, each well was washed twice with PBS. Subsequently, a mixture of human glomerular endothelial cell complete medium and CCK‐8 reagent (Beyotime) was added to the wells, and the culture plates were returned to the incubator for a 1‐h reaction protected from light. Finally, the plates were transferred to a microplate reader, and the absorbance values of each well were measured.

Reactive oxygen species (ROS) level analysis

Following the careful collection of GECs samples from each group, the cells were rinsed with PBS buffer two to three times. Subsequently, a working solution of the 2′,7′‐dichlorodi‐hydrofluorescein diacetate (DCFH‐DA) probe (Elabscience) at a concentration of 10 mmol/L was added to the wells to resuspend the cells. The cell suspensions were then transferred to a dark environment at 37°C and incubated for 30 min. Upon completion of the incubation period, the original liquid was discarded. Finally, a flow cytometer was utilized to detect and acquire the fluorescence signal intensities of the samples.

Colorimetric assays

Malondialdehyde (MDA) and superoxide dismutase (SOD) levels in GECs were analyzed using a microplate reader, employing the MDA Content Assay Kit (Solarbio, Beijing, China) and SOD Activity Assay Kit (Solarbio), respectively.

Cell apoptosis analysis

GECs from each experimental group were, respectively, collected and subjected to two to three washes with pre‐chilled PBS. Following the washing steps, the cell pellets were resuspended in an appropriate amount of incubation buffer, after which Annexin V‐FITC reagent (Solarbio) was added to the suspension and thoroughly mixed. Subsequently, propidium iodide (PI) dye (Solarbio) was supplemented, and the mixture was completely blended as well. The samples were then placed in a light‐protected environment at 4°C to react for 10 min. Within a strict 30‐min timeframe following the completion of the aforementioned staining procedures, the samples had to be immediately transferred to a flow cytometer to accurately evaluate and detect the apoptotic status of the cells.

Transendothelial electrical resistance (TER) assay

GECs were seeded into 24‐well Transwell inserts equipped with 0.4 μm polycarbonate membranes (Corning, Madison, New York, USA). Subsequently, the GECs were suspended in serum‐free human glomerular endothelial cell complete medium (Procell) and added to the apical chambers, while the basolateral chambers were supplemented with complete medium containing fetal bovine serum. The cells were incubated in a cell culture incubator and continuously monitored under an inverted microscope until confluent and a steady state was achieved. Following the respective experimental treatments, the resistance of the transendothelial cell monolayer was measured using a cell electrical resistance meter. The cell‐specific resistance was calculated by subtracting the resistance of the cell‐free membrane from the resistance measured for the cell monolayer. Subsequently, the TER was determined by multiplying this value by the effective bottom area of the Transwell insert.

FITC‐dextran permeability assay

A FITC‐dextran solution (500 μg/mL, Abmole, Shanghai, China) prepared in serum‐free medium was added to the apical chambers of the Transwell inserts, while 600 μL of serum‐free medium was added to the basolateral chambers. Subsequently, the inserts were incubated in an atmosphere protected from light for 1 h. Upon completion of the incubation, the liquid in the basolateral chambers was gently pipetted to ensure thorough mixing, and a 100 μL aliquot was transferred to black 96‐well plates. The fluorescence intensity was then measured using a microplate reader to evaluate the permeability of macromolecules across the endothelial monolayer.

Tube formation assay

An appropriate amount of Matrigel (Corning) was mixed with serum‐free medium, and 200 μL of the mixture was added to each well of 24‐well plates, followed by incubation at 37°C for 30 min to allow solidification. Subsequently, GECs subjected to different treatments and reaching 80% confluency were detached using trypsin, and the cell pellets were collected via centrifugation. The harvested cells were then seeded into the wells pre‐coated with Matrigel for continuous incubation. After 6 h of culture, the tube formation status of the cells was evaluated under an inverted microscope, and images were captured.

GTP‐RhoA activity analysis

GTP‐RhoA activity was analyzed using the RhoA G‐LISA Activation Assay Kit (Amyjet Scientific, Wuhan, China) according to the manufacturer's instructions. A microplate luminometer was used to analyze the results.

RNA immunoprecipitation (RIP) assay

RIP was performed using a commercial RIP kit (Millipore, Bradford, MA, USA), with anti‐IgG (1:100, ab109489, Abcam) and anti‐WTAP antibodies (RAB01195, 1:100, Thermo Fisher). Briefly, GECs were lysed in lysis buffer supplemented with RNase inhibitors (Thermo Fisher). The remaining lysates were incubated overnight at 4°C with antibody‐conjugated magnetic beads to specifically capture RNA‐protein complexes. After washing to remove non‐specifically bound substances, the purified RNA was subjected to qRT‐PCR to detect the specific enrichment of S1PR2 mRNA in the WTAP protein complexes.

m6A RNA immunoprecipitation (MeRIP) assay

MeRIP was conducted in accordance with the standard protocols of a MeRIP m6A kit (BersinBio, Guangzhou, China). Following the extraction of total RNA from each group of cells, the RNA was fragmented into lengths of approximately 100 nucleotides. Initially, the magnetic beads were incubated with the antibodies at room temperature for 30 min. After thoroughly washing away the residual uncoupled antibody components, the aforementioned fragmented RNA was added to the beads and incubated at 4°C with gentle shaking for 2 h to facilitate binding. Upon completion of the reaction, the supernatant was discarded, and the beads were gathered using a magnetic separation rack. Finally, the target nucleic acids were recovered and concentrated utilizing the RNeasy MinElute Kit (Qiagen, Valencia, CA, USA), enabling the subsequent determination of the m6A modification enrichment level on the S1PR2 transcript.

Dual‐luciferase reporter assay

DNA fragments derived from the S1PR2 gene, which contained either the wild‐type (WT) or mutant (MUT) sequences interacting with WTAP, were, respectively, inserted into the pmirGLO vector (Promega, Madison, WI, USA). Following this cloning procedure, the resulting recombinant constructs were designated as WT‐S1PR2 as well as MUT‐S1PR2. Subsequently, GECs underwent simultaneous transfection procedures utilizing either shWTAP or shNC, alongside the aforementioned reporter constructs. Ultimately, the relative luminescence levels were accurately measured through the application of a commercial dual‐luciferase reporter assay system (Promega).

Actinomycin D assay

GECs in each group were treated with culture medium supplemented with 5 μg/mL actinomycin D (Abcam) for varying durations (0, 4, 8, and 12 h). Cell samples were harvested at each of the aforementioned time points, followed by the application of qRT‐PCR to evaluate changes in S1PR2 mRNA expression within the cells.

Establishment of DN mouse model

The study was approved by the Animal Care Committee of Zhengzhou Central Hospital Affiliated to Zhengzhou University. To establish an animal model of DN, male spontaneous db/db mice and age‐ and background‐matched male db/m mice were purchased from SPF (Beijing) Biotechnology Co., Ltd. All animals were procured at six weeks of age, with body weights ranging from 40 to 50 g. The mice in the model group were transitioned to a specialized high‐sugar and high‐fat diet, whereas the control group was continuously maintained on a standard chow. Throughout the 10‐week feeding period, fasting blood glucose levels were routinely measured on a weekly basis, and urine samples were periodically collected using metabolic cages for the quantitative analysis of microalbumin. Successful establishment of the DN model was confirmed when the continuously monitored fasting blood glucose levels stably remained at or above 16.7 mmol/L, accompanied by significantly elevated 24‐h urinary albumin levels. Subsequently, the mice that met the aforementioned criteria were allocated into four experimental cohorts: the db/m group, the db/db group, the db/db + shNC/M1‐Exo group, and the db/db + shWTAP/M1‐Exo group, with each cohort comprising five animal subjects. Over the ensuing 8‐week period, the two exosome intervention groups received targeted delivery via the tail vein at a dosage of 1 × 10⁹ particles per mouse once a week. Concurrently, the remaining two groups were administered an equal volume of PBS via tail vein injections at the same frequency. Upon completion of the intervention cycle, all animals were subjected to a 12‐h fast with free access to water, followed by routine blood collection to determine biochemical parameters, including blood glucose, blood urea nitrogen (BUN), serum creatinine (Scr), and urinary albumin. Ultimately, pentobarbital sodium (40 mg/kg, Sigma, St Louis, MO, USA) was administered intraperitoneally to induce deep sedation. Following the confirmation of complete anesthesia, the renal target tissues and peripheral blood samples were meticulously dissected and harvested for subsequent analyses. Scr concentration and BUN were tested using the Quantichrom Creatinine Assay Kit (Amyjet Scientific, Wuhan, China) and Quantichrom Urea Assay Kit (Amyjet Scientific), respectively. Urinary albumin detection kit (Sigma) was used for urinary albumin analysis.

Histological analysis

Histological analysis was performed using Hematoxylin and Eosin (H&E) staining and Masson's Trichrome staining. For H&E staining assay, the renal tissues of the experimental animals were immersed in a 4% paraformaldehyde solution (Solarbio) for fixation, followed by routine embedding to prepare paraffin pathology sections. The sections were stained with hematoxylin (Solarbio), followed by bluing and differentiation using a 1% hydrochloric acid in ethanol solution, and were subsequently counterstained with eosin dye (Solarbio). Thereafter, the tissues were thoroughly dehydrated through an ethanol gradient, cleared with xylene, and ultimately mounted using neutral mounting medium and a coverslip. The prepared slides were placed under a light microscope for image acquisition.

For Masson's Trichrome staining analysis, the renal tissues were fixed in 4% paraformaldehyde (Solarbio) before undergoing paraffin embedding and sectioning. Following deparaffinization and hydration, the slides were first incubated in Gill's modified hematoxylin reagent (Sigma), followed by a gentle rinse with running tap water for 5 min. Subsequently, the sections were immersed in a 1% hydrochloric acid‐ethanol differentiation solution for approximately 10 s, rinsed again with running water for 5 min, and then transferred to Masson's trichrome composite stain (Jinpan, Shanghai, China) to react for 5 to 10 min. The sections were sequentially dehydrated in absolute ethanol, cleared in xylene, and mounted with neutral balsam. Images of the microscopic fields were then captured and recorded.

A semi‐quantitative scoring system ranging from 0 to 5 was performed by an investigator who was blinded to the study hypothesis and experimental groups to assess tubular injury, interstitial injury (H&E staining), and the degree of fibrosis (Masson's trichrome staining). The scoring criteria were as follows: 0 points: No injury observed; 1 point: Lesion area involving 1%–10%; 2 points: Lesion area involving 11%–25%; 3 points: Lesion area involving 26%–50%; 4 points: Lesion area involving 51%–75%; 5 points: Lesion area involving >75%. The tubular injury score was based on the presence and distribution of exfoliated epithelial cells, cellular debris, and hyaline casts within the tubules, as well as the severity of tubular degeneration and dilation. The interstitial injury score was based on the degree of interstitial expansion and the infiltration of inflammatory cells and tissue debris. The fibrosis score was based on the distribution of blue collagen staining in the renal interstitial area.

Enzyme‐linked immunosorbent assay (ELISA)

The levels of interleukin‐1β (IL‐1β) and tumor necrosis factor‐α (TNF‐α) in serum samples of mice were analyzed using commercial ELISA kits, including Mouse IL‐1β ELISA Kit (Beyotime) and Mouse TNF‐α ELISA Kit (Beyotime), according to the guidebooks.

Statistical analysis

All of the statistical evaluations were carried out utilizing GraphPad Prism version 7.0. The presented results are expressed as the means ± standard deviations (SD), derived from a total of three independent replicates. To assess differences between exactly two sets of data, an unpaired Student's t‐test was applied. Conversely, when comparing three or more distinct groups, a one‐way analysis of variance was conducted followed by Tukey's multiple comparisons test. P < 0.05 was considered to denote a statistically significant difference.

RESULTS

Treatment with exosomes from WTAP‐deficient M1 macrophages relieved HG‐induced increases of WTAP expression

THP‐1 cells were treated with PMA to differentiate into M0 macrophages, followed by treatment with LPS and IFN‐γ to polarize them into M1 macrophages. The study then detected CD80 and WTAP expression in M0 macrophages and M1 macrophages. As shown in Figure 1a, CD80 was mainly expressed in M1 macrophages. The results also showed that WTAP expression at the mRNA and protein levels was higher in M1 macrophages than in M0 macrophages (Figure 1b,c). The study isolated exosomes from M1 macrophages, and TEM revealed that the exosomes exhibited a cup‐shaped morphology with a diameter of approximately 100 nm (Figure 1d). Moreover, Western blotting assay revealed that the exosomal markers (TSG101, CD81, and CD9) were enriched in the M1 macrophage‐derived exosomes, while the cellular contaminant marker calnexin was exclusively detected in the parent M1 macrophages (Figure 1e). Subsequently, M1 macrophages were transfected with WTAP‐specific shRNA or a negative control (shNC) to silence WTAP expression, with knockdown efficiency confirmed in Figure 1f and g. Fluorescence microscopy revealed that Dil‐labeled exosomes from M1 macrophages were internalized by GECs following co‐incubation (Figure 1h). Further, the results showed that HG treatment increased WTAP mRNA and protein levels in GECs. Notably, this upregulation was enhanced by exosomes from shNC‐transfected M1 macrophages (shNC/M1‐Exo), but this promoting effect was reversed when using exosomes from shWTAP‐transfected M1 macrophages (shWTAP/M1‐Exo) (Figure 1i and j).

Figure 1.

Treatment with exosomes from WTAP‐deficient M1 macrophages relieved HG‐induced increases of WTAP expression. (a–c) THP‐1 cells were treated with PMA to differentiate into M0 macrophages, followed by treatment with LPS and IFN‐γ to polarize them into M1 macrophages. (a) Flow cytometry was performed to detect CD80⁺ macrophages. (b) WTAP mRNA expression was analyzed by qRT‐PCR in M0 macrophages and M1 macrophages. (c) Western blotting assay was performed to analyze WTAP protein expression in M0 macrophages and M1 macrophages. (d) TEM was applied to analyze the morphology of exosomes from M1 macrophages. (e) Western blotting was performed to detect the protein expression of TSG101, CD81, CD9, and calnexin in M1 macrophages and M1 macrophage‐derived exosomes. (f and g) The efficiency of WTAP knockdown in M1 macrophages was analyzed by qRT‐PCR and Western blotting. (h) The exosomes from M1 macrophages were isolated and labeled with Dil, followed by incubation with GECs. Red fluorescence in GECs was observed using fluorescence microscopy. (i and j) GECs were divided into the NG + PBS group, the HG + PBS group, the HG + shNC/M1‐Exo group, and the HG + shWTAP/M1‐Exo group. (i) WTAP mRNA expression was analyzed by qRT‐PCR in GECs. (j) Western blotting assay was performed to analyze WTAP protein expression in GECs. *P < 0.05, **P < 0.01 and ***P < 0.001.

Exosomes derived from M1 macrophages aggravated HG‐induced GEC injury in a WTAP‐dependent manner

GECs were exposed to HG and then co‐incubated with shNC/M1‐Exo or shWTAP/M1‐Exo. Subsequent analysis showed that HG treatment inhibited cell viability, and the effect was exacerbated by shNC/M1‐Exo (Figure 2a). The results also showed that HG treatment increased ROS and MDA levels and decreased SOD levels; these effects were augmented after treatment with shNC/M1‐Exo (Figure 2b–d). Additionally, shNC/M1‐Exo treatment enhanced the HG‐mediated elevation in apoptosis and cell permeability, as well as the reduction in normalized TER (Figure 2e–g). Further, treatment with shNC/M1‐Exo enhanced the inhibitory effect of HG treatment on tube formation (Figure 2h). However, these detrimental effects were ameliorated when the cells were treated with shWTAP/M1‐Exo instead (Figure 2a–h). Thus, these results showed that exosomes derived from M1 macrophages aggravated HG‐induced GEC injury in a WTAP‐dependent manner.

Figure 2.

Exosomes derived from M1 macrophages aggravated HG‐induced GEC injury in a WTAP‐dependent manner. GECs were divided into the NG + PBS group, the HG + PBS group, the HG + shNC/M1‐Exo group, and the HG + shWTAP/M1‐Exo group. (a) Cell viability was analyzed by CCK‐8 assay. (b) Flow cytometry was performed to quantify ROS levels. (c and d) Colorimetric assays were performed to analyze MDA and SOD levels. (e) Cell apoptosis was analyzed by flow cytometry. (f and g) Transwell assays were used to analyze cell permeability and normalized TER. (h) Tube formation was analyzed by tube formation assay. *P < 0.05, **P < 0.01 and ***P < 0.001.

WTAP silencing destabilized S1PR2 mRNA in a m6A‐dependent manner

The SRAMP database predicted the presence of m6A modification sites with “very high confidence” in the S1PR2 mRNA (Figure 3a). Additionally, the RBPsuit database predicted potential binding sites between the WTAP protein and S1PR2 mRNA (Figure 3b). The study then transfected WTAP shRNA and the matched control shNC into GECs to determine the association of WTAP and S1PR2. The efficiency of WTAP knockdown was analyzed by qRT‐PCR and Western blotting, and the results are shown in Figure 3c,d. As shown in Figure 3e,f, WTAP knockdown inhibited the mRNA and protein levels of S1PR2 in GECs. The RIP assay revealed that the S1PR2 mRNA could be enriched by the WTAP antibody (Figure 3g). Moreover, the decreased expression of WTAP reduced the m6A levels of S1PR2 in GECs (Figure 3h). The dual‐luciferase reporter assay also revealed that WTAP knockdown inhibited the luciferase activity of the wild‐type S1PR2 reporter plasmid, whereas it did not affect the luciferase activity of the mutant (Figure 3i). Further, the decreased WTAP expression accelerated the degradation of the S1PR2 mRNA in GECs (Figure 3j). Thus, WTAP knockdown destabilized S1PR2 mRNA in a m6A‐dependent manner.

Figure 3.

WTAP knockdown destabilized S1PR2 mRNA in a m6A‐dependent manner. (a) The SRAMP database was used to predict m6A modification sites in the S1PR2 mRNA. (b) The RBPsuit database was applied to predict potential binding sites between the WTAP protein and S1PR2 mRNA. (c and d) The efficiency of WTAP knockdown was analyzed by qRT‐PCR and Western blotting in GECs. (e and f) The effects of WTAP silencing on the mRNA and protein levels of S1PR2 were analyzed by qRT‐PCR and Western blotting, respectively, in GECs. (g–i) The association of WTAP and S1PR2 was analyzed by RIP, MeRIP, and dual‐luciferase reporter assays in GECs. (j) The effect of WTAP silencing on the stabilization of S1PR2 mRNA was analyzed by actinomycin D assay in GECs. Ns: not significant, *P < 0.05, **P < 0.01 and ***P < 0.001.

S1PR2 overexpression counteracted the effects of shWTAP/M1‐Exo in HG‐exposed GECs

Next, the team investigated whether the effects of shWTAP/M1‐Exo on HG‐induced GEC injury involved S1PR2 regulation. As shown in Figure 4a, oe‐S1PR2 was effective in increasing S1PR2 in GECs. Moreover, shWTAP/M1‐Exo treatment decreased S1PR2 protein expression, whereas the effect was relieved after transfection with oe‐S1PR2. Subsequent analysis revealed that S1PR2 overexpression inhibited cell viability, increased ROS and MDA levels, and decreased SOD levels (Figure 4b–e). Ectopic S1PR2 expression also induced cell apoptosis, enhanced cell permeability, reduced normalized TER, and inhibited tube formation (Figure 4f–i). Moreover, the data showed that shWTAP/M1‐Exo treatment‐induced effects on cell viability, oxidative stress, cell apoptosis, cell permeability, normalized TER, and tube formation were attenuated after S1PR2 overexpression (Figure 4b–i). Thus, S1PR2 was involved in the regulation of shWTAP/M1‐Exo in HG‐induced GEC injury.

Figure 4.

S1PR2 was involved in the regulation of shWTAP/M1‐Exo in HG‐induced GEC injury. GECs were divided into the HG + oe‐NC group, the HG + oe‐S1PR2 group, the HG + shNC/M1‐Exo group, the HG + shWTAP/M1‐Exo group, and the HG + shWTAP/M1‐Exo + oe‐S1PR2 group. (a) S1PR2 protein expression was detected by Western blotting. (b) Cell viability was analyzed by CCK‐8 assay. (c) Flow cytometry was performed to quantify ROS levels. (d and e) Colorimetric assays were performed to analyze MDA and SOD levels. (f) Cell apoptosis was analyzed by flow cytometry. (g and h) Transwell assays were used to analyze cell permeability and normalized TER. (i) Tube formation was analyzed by tube formation assay. *P < 0.05, **P < 0.01 and ***P < 0.001.

Exosomes derived from WTAP‐deficient M1 macrophages inactivated the RhoA/ROCK1 pathway and increased VE‐cadherin expression by regulating S1PR2 expression

As shown in Figure 5a,b, S1PR2 overexpression increased the protein expression of GTP‐RhoA and ROCK1, decreased VE‐Cadherin protein expression, and enhanced GTP‐RhoA activity in GECs. The data also showed that in HG‐induced GECs, shWTAP/M1‐Exo treatment decreased the protein expression of GTP‐RhoA and ROCK1, increased VE‐Cadherin protein expression, and weakened GTP‐RhoA activity when compared with the shNC/M1‐Exo; however, these effects were relieved after S1PR2 overexpression (Figure 5a,b).

Figure 5.

Exosomes derived from WTAP‐deficient M1 macrophages inactivated the RhoA/ROCK1 pathway and increased VE‐Cadherin expression by regulating S1PR2 expression. (a and b) GECs were divided into the HG + oe‐NC group, the HG + oe‐S1PR2 group, the HG + shNC/M1‐Exo group, the HG + shWTAP/M1‐Exo group, and the HG + shWTAP/M1‐Exo + oe‐S1PR2 group. (a) The protein expression of GTP‐RhoA, ROCK1, and VE‐Cadherin was detected by Western blotting. (b) GTP‐RhoA activity analysis by a commercial kit. *P < 0.05, **P < 0.01 and ***P < 0.001.

Exosomes derived from M1 macrophages aggravated renal injury in mice with DN in a WTAP‐dependent manner

A DN model was established using spontaneous diabetic db/db mice. Subsequently, the DN mice were administered exosomes (shNC/M1‐Exo or shWTAP/M1‐Exo) via tail vein injection. The results showed that the levels of blood glucose, BUN, Scr, and urinary albumin were higher in db/db mice than in db/m mice, and the effects on these latter three renal injury markers were enhanced after treatment with shNC/M1‐Exo (Figure 6a–d). Similarly, db/db mice showed higher kidney damage scores and larger Masson's trichrome‐stained areas than db/m controls, and these pathological changes were further aggravated by shNC/M1‐Exo administration (Figure 6e,f). Moreover, the serum levels of IL‐1β and TNF‐α were higher in db/db mice than in db/m mice, and these effects were enhanced after treatment with shNC/M1‐Exo (Figure 7a,b). As shown in Figure 7c,d, the protein expression of WTAP, S1PR2, ROCK1, and GTP‐RhoA was higher in renal tissues from the db/db mice in comparison with the db/m control, but VE‐Cadherin protein expression was lower; however, shNC/M1‐Exo administration aggravated these effects in db/db mice. Conversely, treatment with shWTAP/M1‐Exo significantly attenuated all these abnormalities compared to the shNC/M1‐Exo group (Figure 6b–f and Figure 7a–d).

Figure 6.

Exosomes derived from M1 macrophages aggravated renal injury in mice with DN in a WTAP‐dependent manner. A DN model was established using spontaneous diabetic db/db mice, with db/m mice as controls. Subsequently, the DN mice were administered exosomes (shNC/M1‐Exo or shWTAP/M1‐Exo) via tail vein injection. After 8 weeks, all mice were euthanized, and blood and renal tissues were harvested for the following analysis. (a) Blood glucose levels were measured using a portable blood glucose meter. (b–d) BUN, Scr, and urinary albumin were measured using commercial kits. (e) HE staining of renal tissues, and kidney damage scores were determined. (f) Masson staining of renal tissues, and Masson's trichrome‐stained areas were analyzed. Ns: not significant, **P < 0.01 and ***P < 0.001.

Figure 7.

Exosomes derived from M1 macrophages aggravated renal inflammation in mice with DN in a WTAP‐dependent manner. A DN model was established using spontaneous diabetic db/db mice, with db/m mice as controls. Subsequently, the DN mice were administered exosomes (shNC/M1‐Exo or shWTAP/M1‐Exo) via tail vein injection. After 8 weeks, all mice were euthanized, and blood and renal tissues were harvested for the following analysis. (a and b) ELISAs were performed to analyze serum levels of IL‐1β and TNF‐α. (c and d) Western blotting was used to detect the protein expression of WTAP, S1PR2, ROCK1, GTP‐RhoA, and VE‐Cadherin protein in renal tissues. *P < 0.05, **P < 0.01 and ***P < 0.001.

DISCUSSION

Approximately 30% of diabetic patients develop DN, a complication that not only poses a remarkably high risk for cardiovascular events but also incurs substantial public health expenditures. ²² Unfortunately, despite extensive efforts within the medical community, effective pharmacological interventions capable of halting the progression of this disease remain elusive, highlighting an urgent need for novel therapeutic approaches. Exosomes are deeply involved in the pathophysiological alterations associated with this renal condition, demonstrating tremendous potential for clinical translation. ²³ , ²⁴ , ²⁵ Notably, the lesion tissues of DN patients are highly enriched with M1‐polarized macrophages, and the extent of their infiltration correlates positively with the rate of glomerular filtration rate decline. ²⁶ Given this context, elucidating the specific molecular pathways by which exosomes derived from M1 macrophages drive disease exacerbation is crucial for the development of innovative targeted therapies. To this end, a series of validation experiments were designed in the present study. The data revealed that these specific exosomes served as carriers to deliver WTAP protein into GECs; subsequently, through an m6A‐dependent RNA modification mechanism, they stabilized S1PR2 transcripts, ultimately accelerating the deterioration of DN.

Our data showed that WTAP was highly expressed in M1 macrophages, and their derived exosomes were successfully internalized by GECs. HG‐induced WTAP upregulation in GECs, which was further amplified by M1 exosomes in a WTAP‐dependent manner. WTAP directly bound to S1PR2 mRNA, enhancing its m6A modification and stability. M1 macrophage‐derived exosomes significantly aggravated HG‐induced suppression of GEC viability and tube formation, while concurrently promoting oxidative stress, apoptosis, and endothelial permeability through the WTAP‐mediated upregulation of S1PR2. In vivo, these exosomes also significantly exacerbated renal injury in db/db mice, an effect that was notably attenuated when utilizing exosomes from WTAP‐deficient M1 macrophages. A previous study indicated that exosomal METTL14 from M1 macrophages promoted HG‐induced GEC injury via PAQR3 m6A modification. ¹³ However, that research solely focused on the m6A reader/writer METTL14 without exploring the dynamic alterations of other critical m6A regulators within the exosomes. The current findings extend this biological concept by identifying WTAP as a novel and functional m6A modulator transferred from M1 macrophages to GECs, revealing a distinct m6A‐dependent mechanism centered on S1PR2 rather than PAQR3. Another separate investigation reported that macrophage‐derived exosomes under HG conditions aggravate podocyte injury by regulating miR‐25‐3p. ²⁷ This previous study was limited to a non‐epigenetic miRNA mechanism and primarily focused on podocytes. The present results surpassed this mechanistic scope by establishing a previously unrecognized exosomal m6A RNA modification pathway targeting GECs. Together, these comparisons highlight a novel paradigm wherein exosomal WTAP acts as a crucial mediator of M1 macrophage‐to‐GEC communication under diabetic conditions, providing new mechanistic insights into m6A‐modulated renal endothelial dysfunction.

Mechanistically, the upregulation of S1PR2 may act as a central pivot that robustly activates the downstream RhoA/ROCK1 signaling axis to drive DN progression. Once activated by exosomal WTAP, this cascade may trigger cytoskeletal rearrangement and the degradation of VE‐Cadherin, directly leading to endothelial barrier dysfunction and increased vascular permeability. Furthermore, consistent with previous reports, the sustained activation of the S1PR2/RhoA/ROCK1 axis may synergistically induce mitochondrial fission via dynamin‐related protein 1 (Drp1) and promote endothelial‐to‐mesenchymal transition (EndMT) through β‐catenin nuclear translocation, ²¹ , ²⁸ which collectively shifts GECs toward a pro‐fibrotic and dysfunctional phenotype. Importantly, our functional rescue experiments demonstrated that exosomes derived from WTAP‐deficient M1 macrophages effectively inactivated this S1PR2/RhoA/ROCK1 signaling axis and successfully restored VE‐Cadherin expression, thereby reversing endothelial damage.

Despite the compelling evidence demonstrating the pathogenic role of the exosomal WTAP/S1PR2 axis in DN, certain limitations warrant acknowledgment. First, the current investigation primarily relies on a single animal model (db/db mice), which predominantly reflects type 2 diabetes; thus, the generalizability of these findings to type 1 diabetic nephropathy or other chronic kidney disease models remains undetermined. To address this issue, future studies should validate this molecular axis in alternative murine models, such as streptozotocin‐induced diabetic mice. Second, while the study establishes that S1PR2 activates the RhoA/ROCK1 cascade to degrade VE‐Cadherin, the precise crosstalk between this signaling pathway and other downstream pathological processes—such as Drp1‐mediated mitochondrial fission and EndMT described in previous literature—was not exhaustively elucidated. Incorporating comprehensive mitochondrial dynamics assessments and specific EndMT marker profiling in subsequent research would provide a more holistic understanding of how the WTAP/S1PR2 axis orchestrates comprehensive endothelial dysfunction. Third, it is important to note that tail vein injection results in systemic distribution, and shWTAP/M1‐Exo may potentially benefit other diabetic vascular complications. However, our in vitro evidence of direct exosome uptake by GECs and the consistent regulation of the renal WTAP/S1PR2 axis support a direct mechanism of renal protection, rather than an indirect effect mediated by other organs. Fourth, the use of whole kidney tissue lysates for Western blotting is a limitation of this study, as the abundance of tubular proteins may mask specific changes in the glomerular compartment. Future studies utilizing isolated glomeruli or laser capture microdissection are warranted to precisely delineate the cell‐specific regulatory mechanisms. Further, this study primarily relied on H&E and Masson staining combined with renal function indicators to assess renal injury, lacking TEM and immunofluorescence localization of cell‐specific markers. These experiments will be prioritized in future research.

In conclusion, the present study elucidated a novel macrophage‐endothelial communication mechanism wherein M1 macrophage‐derived exosomes transferred WTAP to GECs. Through m6A‐dependent posttranscriptional modification, WTAP stabilized S1PR2 mRNA, thereby activating the RhoA/ROCK1 signaling axis, disrupting VE‐Cadherin‐mediated endothelial barrier integrity, and ultimately accelerating the progression of DN. Targeting the WTAP/S1PR2/RhoA/ROCK1 regulatory network presents a novel and precise therapeutic strategy. Interventions designed to intercept pathogenic exosome cargo delivery or inhibit S1PR2 could offer tailored approaches to restore glomerular endothelial integrity and halt the deterioration of renal function in diabetic patients.

AUTHOR CONTRIBUTIONS

Lei Li designed and performed the research; Hongmei Liu, Yu Mao, Huanhuan Wang, Lige Song, Zhiqiang Kang analyzed the data; Lei Li wrote the manuscript. All authors read and approved the final manuscript.

DISCLOSURE

The authors declare no conflicts of interest.

Approval of the research protocol: The study was approved by the Animal Care Committee of Zhengzhou Central Hospital Affiliated to Zhengzhou University.

Informed consent: N/A.

DATA AVAILABILITY STATEMENT

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