Notch signaling pathway mediates anti-inflammatory effects of vagus nerve stimulation during lipopolysaccharide-induced acute kidney injury

Abstract

The cholinergic anti-inflammatory pathway (CAP) plays a central role in neuroimmunomodulation, and its activation is a potential strategy for ameliorating sepsis-associated acute kidney injury (SA-AKI). However, further investigations are necessary to understand the molecular mechanisms of CAP activation and develop therapies for SA-AKI. Here, we tested whether the Notch signaling pathway, which regulates cell-cell interactions, mediates the anti-inflammatory effects of CAP. Using a mouse model of lipopolysaccharide (LPS)-induced AKI, we found that CAP activation by vagus nerve stimulation (VNS) enhanced Notch2 signaling in macrophages, mitigating inflammation in the spleen and tissue damage in the kidneys. Consistently, macrophage-specific knockout of Notch2 resulted in an attenuation of these anti-inflammatory effects of VNS. We also demonstrated that VNS and macrophage-specific Notch2 signaling might upregulate transferrin, which maintains iron homeostasis, thereby protecting the kidneys. Taken together, our findings suggest the involvement of Notch signaling in the mechanisms of VNS-mediated CAP activation during LPS-induced AKI.

Sepsis-associated acute kidney injury has limited treatments. This study shows that Notch2 signaling in macrophages mediates the anti-inflammatory and kidney-protective effects of vagus nerve stimulation.

Introduction

Sepsis-associated acute kidney injury (SA-AKI) is a severe life-threatening complication that affects critically ill patients with sepsis or septic shock^1–6. It is characterized by a rapid decline of glomerular filtration rate, manifesting as decreased urine output and elevated serum creatinine levels. The risk of mortality is increased in SA-AKI and the risk of developing chronic kidney disease is also increased in surviving patients with SA-AKI. Furthermore, the pathophysiology of SA-AKI is highly complex and involves an early exaggerated inflammatory responses, microcirculatory dysfunction, metabolic reprogramming, and late immune suppression. Despite advances in understanding the mechanisms of SA-AKI, effective treatments remain limited.

The cholinergic anti-inflammatory pathway (CAP), a neuroimmune circuit discovered by Tracey et al^7,8., has gained attention as a potential therapeutic target for inflammation-related diseases including sepsis, peritonitis, central nervous system disorders, cardiovascular disease, and lung injury^9–13. This pathway involves the vagus nerve that detects peripheral inflammatory signals and initiates an anti-inflammatory reflex^1,14. Vagus nerve stimulation (VNS) promotes the noradrenaline release in the spleen by activating the splenic nerve¹⁵. This neurotransmitter interacts with β2-adrenergic receptors on choline acetyltransferase-positive CD4 T cells, which then release acetylcholine¹⁶. Acetylcholine binds to α7 nicotinic acetylcholine receptors (α7nAChR) on macrophages¹⁷, suppressing the production of pro-inflammatory cytokines^15–17. The anti-inflammatory and tissue-protective effects of VNS-mediated CAP activation have been shown in not only experimental studies but also clinical trials in rheumatoid arthritis and Crohn’s disease by using electrical stimulators of the vagus nerve^12,18,19.

The efficacy of CAP activation has also been demonstrated in acute kidney injury (AKI)^1,14,20. We have previously found that VNS, through electrical or optogenetic stimulation, protects the kidneys against ischemia-reperfusion injury (IRI)-induced AKI in the mouse model^21,22. In addition, a recent study suggested that CAP activation by the administration of an α7nAChR-specific agonist (GTS-21) lessened lipopolysaccharide (LPS)-induced AKI that partially mimics SA-AKI²³. However, there are relatively few drugs targeting CAP approved by the US Food and Drug Administration, and further understanding of the CAP activation mechanisms at the molecular level is required to develop CAP-targeted therapies for AKI including SA-AKI.

To achieve this goal, we recently performed RNA sequencing (RNA-seq) analysis and identified hairy and enhancer of split 1 (Hes1) as a downstream molecule of α7nAChR in macrophages²⁴. The mRNA expression of Hes1 was upregulated by nicotine-mediated stimulation of α7nAChR, and this induction of Hes1 was attenuated in macrophages from α7nAChR global knockout mice, indicating that Hes1 is a potential mediator of CAP activation²⁴. Hes1 is an inhibitory transcription factor, and its gene expression is regulated by the Notch signaling pathway, which plays an essential role in cell-cell interactions^25–27. Notch signaling controls cell fate decisions, tissue homeostasis, and inflammatory responses, and there are five Notch ligands (Jagged1 (Jag1), Jag2, Delta-like1 (Dll1), Dll3, Dll4) and four Notch receptors (Notch1-4) in mammals. Consistent with an increase in Hes1 by nicotine-mediated α7nAChR stimulation²⁴, a reanalysis of our RNA-seq datasets of macrophages²⁴ showed that nicotine treatment enhanced the mRNA expression of Jag1, Notch2, and Rbp-j (a downstream effector of Notch signaling) among Notch-associated genes (Supplementary Fig. 1). For these reasons, we hypothesized that the Notch signaling in macrophages is involved in the mechanisms of CAP activation. In this study, we investigated whether and how Notch signaling in macrophages mediates the anti-inflammatory and kidney-protective effects of CAP during LPS-induced AKI.

Results

Notch signaling pathway mediates anti-inflammatory effects of α7nAChR in peritoneal macrophages

To test our hypothesis that the anti-inflammatory effects of the CAP are mediated by the Notch signaling, we examined the relationship between Notch signaling and α7nAChR during an LPS-induced inflammatory state, because α7nAChR on macrophages is a central player in mitigating inflammatory responses during CAP activation¹⁷. From the analysis of previous RNA-seq datasets from murine peritoneal macrophages²⁴, we found that α7nAChR stimulation by nicotine upregulates Notch-associated genes besides Hes1 expression (Supplementary Fig. 1). However, nicotine is a nonselective agonist for nicotinic acetylcholine receptors, so we used GTS-21, which is a selective agonist of α7nAChR²³, to reduce the possibility that nicotinic acetylcholine receptors other than α7nAChR activate Notch signaling. Primary macrophages isolated from the peritoneal cavity of wild-type (WT) mice were cultured and subjected to GTS-21 and LPS (Fig. 1A). We first performed the Luminex assay (bead-based immunoassay) using culture supernatants from peritoneal macrophages to determine which cytokine/chemokine expression levels are relevantly affected by GTS-21 treatment. Among 32 cytokines tested, GTS-21 tended to decrease multiple pro-inflammatory cytokines/chemokines induced by LPS, however, GTS-21 only significantly reduced the LPS-induced increase in supernatant levels of the chemokine Ccl2 (also known as Mcp-1) compared to vehicle controls (Fig. 1B, C, Supplementary Table 1). We then studied whether Notch signaling mediates the decrease in Ccl2 by α7nAChR activation. Peritoneal macrophages were preincubated with DAPT, a chemical inhibitor of Notch signaling, prior to GTS-21 and LPS (Fig. 1A). ELISA experiments revealed that about 90% of LPS-induced Ccl2 levels were reduced by GTS-21, but this effect of GTS-21 on Ccl2 reduction was significantly attenuated by Notch blockade via the DAPT treatment (Fig. 1D). Furthermore, GTS-21 produced a robust increase in Hes1 mRNA expression that was suppressed by DAPT during LPS-triggered inflammation (Fig. 1E). These in vitro studies indicate that Notch signaling in peritoneal macrophages is activated downstream of α7nAChR, leading to reduced Ccl2 levels.

Fig. 1. Stimulation of α7nAChR, a component of CAP, enhances Notch signaling in peritoneal macrophages to dampen inflammation.

A Schematic showing the timeline of in vitro cell culture experiments (for B–E). Primary macrophages isolated from the peritoneal cavity of WT mice were cultured with combinations of DAPT (a chemical inhibitor of Notch signaling, 10 μM), GTS-21 (a selective agonist of α7nAChR, 50 μM), and LPS (100 ng/ml). The total treatment time of each compound is as follows: DAPT 48 h, GTS-21 15 min + 24 h, LPS 24 h. B Cell culture supernatants from peritoneal macrophages treated with GTS-21 or water and LPS or saline were collected, and cytokine and chemokine levels in those samples were then quantified by the Luminex assay. Every well in the heatmap indicates an average concentration of 4 samples obtained from independent experiments. The cross marks indicate that concentration values were outside the detection range. C Ccl2 concentrations in cell culture supernatants determined by the Luminex assay (independent biological replicates, N = 4). D ELISA experiments were performed to quantify Ccl2 concentrations in each sample. The ratios of Ccl2 levels in the GTS-21-LPS-treated group relative to those in the LPS-treated alone group were calculated from the results of ELISA. Each Ccl2 ratio (GTS-21 + LPS/Vehicle + LPS) in the presence or absence of DAPT was compared (independent biological replicates; N = 5). E The mRNA expression levels of Hes1 in peritoneal macrophages that were exposed to combinations of DAPT, GTS-21, and LPS relative to those in controls (in the absence of these three chemicals) (independent biological replicates; N = 6). Data are represented as means ± SEM. ns, not significant. *P < 0.05, ***P < 0.001, and ****P < 0.0001 (one-way ANOVA followed by post hoc Tukey’s multiple comparison test (C, E) and unpaired Student’s t test (D)).

CAP activation by VNS activates the Notch signaling in macrophages, leading to attenuation of LPS-induced inflammation in the spleen and kidneys

We previously showed that VNS protects the kidneys against IRI-induced AKI^21,22, yet the effectiveness of VNS has not been well evaluated in SA-AKI. To investigate if VNS attenuates inflammation in the spleen and kidneys during sepsis, a murine model of LPS-induced AKI was used in this study. The left cervical vagus nerve of WT mice was electrically stimulated 34 hours (h) before LPS injection, and these mice were euthanized 14 h after LPS (Fig. 2A). LPS-induced systemic inflammation causes splenomegaly²⁸, and as expected, spleens were enlarged by LPS in the current study. However, mice exposed to VNS prior to LPS had spleen weights that did not differ from those in sham-operated saline-treated mice (Fig. 2B). An LPS-induced increase in cytokine levels in the spleen was also suppressed by VNS (Fig. 2C, Supplementary Table 2). Among a panel of 32 cytokines/chemokines, VNS mainly affected chemokine levels rather than cytokines at this timepoint (14 h post-LPS injection), and LPS-induced increases in Ccl2, Ccl5, M-csf, G-csf, and Il-15 were significantly reduced by VNS in the spleen (Fig. 2C, D, Supplementary Table 2). Next, to examine the influence of VNS on the kidneys, renal injury due to LPS was assessed by mRNA levels of kidney injury molecule-1 (Kim-1, also known as Havcr1) as an early biomarker of kidney injury and histological analysis. As plasma creatinine levels were not elevated by LPS administration, suggesting that renal injury was mild in our LPS-induced AKI model (Supplementary Fig. 2), these two parameters were evaluated. Consistent with the anti-inflammatory responses in the spleen, mice that underwent VNS prior to LPS administration showed lower expression levels of Kim-1 in the kidneys compared to sham-stimulated mice (Fig. 2E). Periodic acid-Schiff (PAS) staining also revealed that VNS alleviated LPS-induced kidney tubular injury such as vacuolization (Fig. 2F). These data indicate that electrical stimulation of cervical vagus nerve provided anti-inflammatory effects to the spleen as well as tissue-protective effects to the kidneys against inflammation-induced damage in the context of endotoxin exposure. On the other hand, VNS application 1 h post-LPS injection did not suppress splenomegaly and an increase in renal Kim-1 expression induced by LPS (Supplementary Fig. 3).

Fig. 2. VNS leads to amelioration of LPS-induced inflammation in the spleen and kidneys, accompanied by Notch2 signaling activation in macrophages.

A Time course of mouse experiments (for B-H). Electrical stimulation (or sham stimulation) was applied to the left cervical vagus nerve of WT mice 34 h or 24 h before LPS i.p. injection (5 mg/kg). After 14 h or 24 h, these mice were euthanized, and the spleen and kidneys were harvested (34 + 14 h for B-G and 24 + 24 h for H). B Spleen size was measured and normalized to body weight (n = 8 to 12 per group). Scale bar = 1 cm. C A heatmap generated from the Luminex assay represents cytokine and chemokine levels in the spleen. Every well in the heatmap indicated an average concentration of 3 (sham + saline) or 5 samples (sham + LPS and VNS + LPS). The cross marks indicate that concentration values were outside the detection range. D Concentrations of Ccl2, Ccl5, M-csf, G-csf, and Il-15 in the spleen normalized to cell numbers (n = 3 to 5 per group). E Transcript levels of Kim-1 in the kidneys of mice that underwent VNS or sham stimulation followed by LPS administration. The values are relative to those in baseline controls (sham + saline) (n = 12 per group). F Representative PAS staining in kidney cortex with injured tubular areas (n = 6 to 9 per group). Arrowheads indicate kidney tubular vacuolization. Scale bar = 50 μm. G Representative immunofluorescence images of Notch2 (red) and F4/80⁺ macrophages (green) in the red pulp of the spleen (n = 8 to 13 per group). Scale bar = 10 μm. H Gene expression levels of Notch2 and Hes1 in splenic Cx3cr1⁺ macrophages isolated 24 h post-LPS injection and expressed as values relative to those in controls (sham + saline) (n = 4 to 7 per group). Data are means ± SEM. ns, not significant. *P < 0.05, **P < 0.01, and ****P < 0.0001 (one-way ANOVA followed by post hoc Tukey’s multiple comparison test (B, D, F, H) and unpaired Student’s t test (E)).

We next assessed the expression of Notch2 in the spleen of mice subjected to VNS and subsequent LPS challenge. LPS reduced Notch2 protein immunoreactivity to barely detectable levels in F4/80-positive macrophages; this suppression was prevented by VNS pretreatment (Fig. 2G). At 24 h after LPS injection, Notch2 mRNA expression in isolated Cx3cr1-positive splenic macrophages was comparable between VNS- and sham-operated mice, whereas the reduction in Hes1 expression during LPS-induced inflammation was prevented by VNS (Fig. 2H). We also investigated the influence of VNS application alone on Notch signaling in splenic macrophages. There was no difference in Notch2 and Hes1 levels between VNS- and sham-treated groups (Supplementary Fig. 4). These results suggest that VNS activates the Notch signaling via Notch2 receptor in splenic macrophages in the context of systemic inflammation, which attenuates inflammation-induced Notch suppression.

Notch2 deficiency in macrophages exaggerates pro-inflammatory responses in the spleen and kidneys

To further confirm the role of Notch2 in macrophages in VNS-mediated anti-inflammation, macrophage (MΦ)-specific Notch2 knockout (KO) mice were established by crossbreeding Cx3cr1-Cre mice and Notch2^flox/flox mice (Fig. 3A). There was no difference in body weight between Cx3cr1-Cre; Notch2^flox/flox (Notch2^MΦKO) and Notch2^flox/flox (Notch2^MΦWT) mice, while the spleen size of Notch2^MΦKO was bigger than that of Notch2^MΦWT (Supplementary Fig. 5A, B). Gene expression of Notch2 was decreased and Hes1 expression tended to be reduced in the splenic Cx3cr1-positive macrophages isolated from Notch2^MΦKO (Supplementary Fig. 5C, D). We also confirmed reduced Notch2 protein in splenic F4/80-positive macrophages by immunofluorescence (Supplementary Fig. 5E). In the kidneys, macrophage numbers were comparable between Notch2^MΦWT and Notch2^MΦKO mice, indicating that Notch2 deficiency does not affect the numbers of resident renal macrophages under physiological conditions (Supplementary Fig. 5F).

Fig. 3. The anti-inflammatory and kidney-protective effects of VNS-mediated CAP activation are diminished by macrophage-specific deletion of Notch2.

A Design of experiments using macrophage-specific Notch2 knockout (Notch2^MΦKO) mice (for B–F). WT (Notch2^MΦWT) or Notch2^MΦKO mice were subjected to VNS and LPS administration (5 mg/kg) 24 h after VNS. At 24 h post-injection, these mice were anesthetized and euthanized to collect blood, spleen, and kidneys. B Spleen weight was measured immediately after euthanasia and normalized to body weight (n = 6 to 10 per group). C Transcript levels of Ccl2 in splenic Cx3cr1⁺ macrophages of Notch2^MΦWT or Notch2^MΦKO mice that underwent VNS and subsequent LPS challenge. Expressed as values relative to those in the control group (WT + sham + saline) (n = 8 to 9 per group). D mRNA expression levels of Kim-1 in the kidneys of Notch2^MΦWT or Notch2^MΦKO mice subjected to VNS and LPS injection. Expressed as values relative to those in the control group (WT + sham + saline) (n = 11 to 14 per group). E Representative images of PAS staining of kidney sections and injured tubular areas (n = 4 to 6 per group). Arrowheads indicate intracellular vacuoles in renal tubules. Scale bar = 50 μm. (F) F4/80⁺ macrophages in the kidneys were visualized by immunohistochemical labeling and counted under a light microscope (n = 3 to 4 per group). Scale bar = 50 μm. Data are represented as means ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001 (one-way ANOVA followed by post hoc Tukey’s multiple comparison test (B, E, F) and unpaired Student’s t test (C, D)).

These mice were then subjected to VNS and LPS treatments to test if the suppressive effects of CAP on inflammatory responses are abrogated by Notch2 deletion in macrophages (Fig. 3A). VNS application before LPS administration mitigated splenomegaly in Notch2^MΦWT, but not in Notch2^MΦKO mice (Fig. 3B). Elevated Ccl2 expression was also observed in splenic Cx3cr1-positive macrophages from Notch2^MΦKO that underwent VNS and LPS treatments relative to those from Notch2^MΦWT (Fig. 3C). Renal injury induced by LPS, as indicated by higher Kim-1 expression in kidneys and injured tubular areas determined by histology, was mitigated by VNS in Notch2^MΦWT but the protective effects were lost in Notch2^MΦKO mice (Fig. 3D, E).

We next examined the effect of macrophage-specific Notch2 deficiency on macrophage infiltration into the kidneys since macrophages play a role in the pathophysiology of LPS-induced AKI^1,29. In addition, we showed that VNS and GTS-21-mediated activation of α7nAChR significantly decreased Ccl2 (a chemoattractant for macrophages/monocytes) levels in the spleen and culture supernatants of peritoneal macrophages, respectively (Fig. 2D and Fig. 1C). Consistent with these observations, LPS-induced infiltration of macrophages into the kidneys was significantly suppressed by VNS in Notch2^MΦWT, however, these effects of VNS were abolished in Notch2^MΦKO (Fig. 3F). Together, these findings suggest that macrophage-specific Notch2 provides anti-inflammatory and renoprotective effects during VNS-mediated CAP activation.

Candidate biological processes in kidneys altered by VNS and/or macrophage-specific Notch2 deficiency

Critical factors in the spleen associated with the anti-inflammatory effects of VNS include T cells expressing choline acetyltransferase¹⁶ and α7nAChR-positive macrophages¹⁷, and our current results now suggest the involvement of Notch signaling on macrophages in the CAP activation. Nonetheless, other factors in the kidneys remain elusive, thus we performed RNA-seq to identify key players in tissue protection by VNS in the kidneys (Fig. 4A, Supplementary Fig. 6). Whole kidney tissues from Notch2^MΦWT or Notch2^MΦKO mice that underwent VNS and LPS administration were used for these experiments (Fig. 4A). To broadly investigate the biological processes changed by VNS and/or macrophage-specific Notch2 deficiency, we initially focused on gene ontology (GO) analysis and made four paired comparisons (Fig. 4B–E, Supplementary Table 3). Significantly enriched GO terms were extracted from each comparison up to the top 15 terms. In Notch2^MΦWT mice, VNS altered multiple metabolic pathways under physiological conditions (Fig. 4B), while the processes involved in the immune and inflammatory systems were mainly affected by VNS under pathological conditions (Fig. 4C). Among altered pathways in the Notch2^MΦWT + VNS + LPS group (Fig. 4C), six biological processes were common in those enriched in Notch2^MΦKO groups with or without VNS: chemotaxis, chemokine-mediated signaling pathway, immune system process, inflammatory response, cellular response to interferon-gamma, and immune response (Fig. 4D, E). In particular, the involvement of chemotaxis and chemokine-mediated signaling pathway in VNS and/or macrophage-specific Notch2 knockout was consistent with our observations that VNS reduced various chemokines in the spleen and suppressed immune cell infiltration into the kidneys, and those effects of VNS were abolished in Notch2^MΦKO mice (Fig. 2C, D and Fig. 3C, F). Although there was no common biological process among the four paired comparisons (Fig. 4B–E), these data support our hypothesis that Notch signaling mediates the anti-inflammatory effects of VNS during LPS-induced AKI. In addition, these findings indicate that Notch activation by VNS potentially dampens inflammation via the regulation of chemokine function in kidneys.

Fig. 4. RNA-seq analysis identified biological processes in kidneys responding to VNS and/or macrophage-specific deletion of Notch2.

A Schematic of RNA-seq analysis of mouse kidneys (for B–E). Kidneys were isolated from Notch2^MΦWT or Notch2^MΦKO mice that underwent VNS (or sham) and LPS (or saline) injection (5 mg/kg), and total RNA extracted from those samples was used for the analysis (n = 3 per group and a total of 6 groups). B–E GO biological processes that were significantly enriched in the gene sets identified from DEG analysis (adjusted P values < 0.05, up to the top 15 GO terms). Two independent comparisons were made between VNS- and sham-treated groups in Notch2^MΦWT mice (B, Notch2^MΦWT + VNS + saline vs Notch2^MΦWT + sham + saline; C, Notch2^MΦWT + VNS + LPS vs Notch2^MΦWT + sham + LPS). Enriched GO terms in Notch2^MΦKO mice are also shown (D, Notch2^MΦKO + sham + LPS vs Notch2^MΦWT + sham + LPS; E, Notch2^MΦKO + VNS + LPS vs Notch2^MΦWT + VNS + LPS). Red squares indicate common biological processes among C, D, and E.

Kidney-protective effects of VNS are mediated by transferrin

To further elucidate the kidney-specific mechanisms underlying anti-inflammation and tissue protection by VNS, we next focused on candidate genes identified from the differentially expressed gene (DEG) analysis using RNA-seq datasets. A comparison of DEG profiles of VNS-treated kidneys under conditions with and without LPS revealed that VNS significantly increased the expression of 45 genes and 151 genes in each profile, respectively, and 11 genes were commonly upregulated by VNS between the two DEG profiles (Fig. 5A). We then examined whether these 11 genes were downregulated in the kidneys of Notch2^MΦKO mice because Notch2 deficiency attenuates anti-inflammatory effects of VNS (Fig. 3). Although none of these genes were identified as DEGs in transcriptomic profiles of Notch2^MΦKO mice, two and three of 11 genes tended to be downregulated in the profiles from Notch2^MΦKO with LPS and with VNS + LPS, respectively (Fig. 5A). Furthermore, we found one gene that overlapped between these two and three genes identified from each profile of Notch2^MΦKO mice: transferrin (Trf) (Fig. 5A). Validation by qPCR analysis also revealed that during LPS-induced AKI, 1) VNS upregulated gene expression of Trf, and 2) Notch2 deficiency in macrophages abrogated the VNS-mediated increase in Trf (Fig. 5B).

Fig. 5. Transferrin in the kidneys mediates the renoprotective effects of CAP activation by VNS.

A Strategy to identify candidate genes in the kidneys that play a central role in the anti-inflammatory effects of VNS. Eleven common upregulated genes in the kidneys of Notch2^MΦWT mice subjected to VNS were identified between two DEG profiles (log2 fold changes > 0.585 ( > 1.5-fold increase), adjusted P values < 0.10). Further selection revealed that transferrin (Trf) was a unique gene commonly downregulated in two transcriptomic profiles from Notch2^MΦKO mice among those 11 genes (log2 fold changes < -0.585 ( < 0.67-fold decrease)). B Validation of Trf expression in kidneys was performed by qPCR. Relative Trf expression levels to those in controls (WT + sham + saline) are shown (n = 4 to 14 per group). C Schematic of mouse experiments using ferristatin II (for D–G). WT mice were administered ferristatin II (10 mg/kg/day) or saline once every 24 h for 4 doses. At 48 h and 72 h after the first dose of ferristatin II, these mice underwent VNS and LPS injection (5 mg/kg), respectively. D Spleen size in saline + VNS + LPS group and ferristatin II + VNS + LPS group (n = 9 to 10 per group). E Plasma BUN concentrations (a marker of renal function) (n = 7 to 8 per group). F mRNA levels of Kim-1 in the kidneys of ferristatin II-treated or untreated mice subjected to electrical VNS and LPS challenge. The values are relative to those in baseline controls (saline + sham + saline) (n = 9 per group). G PAS staining in the cortex of kidney tissues and injured tubular areas calculated from those samples (n = 7 to 8 per group). Arrowheads indicate tubular vacuolization. Scale bar = 50 μm. Data are represented as means ± SEM. ns, not significant. *P < 0.05 and **P < 0.01 (one-way ANOVA followed by post hoc Tukey’s multiple comparison test (B) and unpaired Student’s t test (D–G)).

Transferrin (Trf), an iron-transporting protein, maintains iron homeostasis and is involved in multiple biological processes including inflammatory responses^30–32. Previous studies have demonstrated that Trf has therapeutic potential for diseases associated with inflammation such as bacterial infection and acute renal failure^33,34, whereas the involvement of Trf in the renoprotective effects of the CAP activation has not been reported. Hence, we aimed to investigate whether the blockade of Trf utilization attenuates kidney-protective effects of VNS against LPS-induced injury. WT mice were subjected to ferristatin II³⁵, an iron transport inhibitor targeting transferrin receptor 1 (TfR1), before VNS and LPS injection (Fig. 5C). VNS was applied 48 h after the first administration of ferristatin II since TfR1 internalization from the apical membrane of renal tubules or subsequent degradation of TfR1 was observed, as in previous reports^35,36, at that timepoint (Supplementary Fig. 7A, B). In addition, plasma BUN and renal Kim-1 levels were not altered by ferristatin II treatment alone, suggesting ferristatin II itself does not induce the kidney injury (Supplementary Fig. 7C, D). The spleen weight was comparable between ferristatin II- and vehicle-treated groups (Fig. 5D), while ferristatin II treatment increased plasma BUN and renal Kim-1 levels and exacerbated histological tubular injury in the kidneys (Fig. 5E–G). These results indicate that Trf utilization via TfR1 plays a crucial role in kidney protection against LPS-induced AKI in the context of CAP activation.

Discussion

In this study, we demonstrated that the Notch signaling in macrophages is responsible for the anti-inflammatory effects of CAP during LPS-induced AKI (Fig. 6). CAP triggered by VNS has been traditionally considered to suppress inflammatory responses via paracrine signaling between choline acetyltransferase-positive CD4 T cells and macrophages expressing α7nAChR^1,14,16,17. In contrast, Notch signaling is a contact-dependent signal transduction system^25–27. Therefore, in addition to the classical paracrine mechanism of CAP, our findings may illustrate a non-classical concept that physical contact between cells mediated by Notch signaling plays a critical role in CAP (Fig. 6). This idea is supported by a recent study showing that α7nAChR stimulation via GTS-21 treatment enhanced macrophage-macrophage contacts in the spleen to protect the kidneys against LPS-induced AKI, yet the influence of GTS-21 on T cell-macrophage interactions is still unclear²³. Our study also provided evidence that macrophage-specific Notch2 is central for reduced inflammation in the context of VNS-mediated CAP activation (Fig. 3). However, we could not identify which Notch ligands activate Notch2 receptor on macrophages during VNS (Fig. 6). From reanalysis of public single-cell RNA-seq data from the mouse spleen²³, we found that GTS-21-mediated stimulation of α7nAChR broadly upregulated Dll1 expression across multiple immune cell populations under LPS-induced inflammatory conditions (Supplementary Fig. 8). Previous research has also shown that Dll1 has a high affinity for Notch2 over Notch1³⁷. Based on these observations, direct contact between Dll1-expressing cells and Notch2-positive macrophages might be important for CAP activation and subsequent anti-inflammation.

Fig. 6. Hypothetical mechanism of Notch signaling in mediating the anti-inflammatory effects of CAP activation.

CAP activation by VNS enhances Notch signaling in macrophages to exert anti-inflammatory and tissue-protective effects against LPS-induced AKI. This means that the physical interaction between macrophages and neighboring immune cells via Notch signaling might contribute to CAP activation, not just the classical paracrine signaling mediated by α7nAChR on macrophages. VNS-triggered Notch2 activation in macrophages leads to reduction of cytokines and chemokines such as Ccl2 and Ccl5, and suppression of macrophage infiltration into the kidneys. Increased expression and utilization of transferrin within the kidneys play a key role in renoprotection by VNS, and Notch2-expressing macrophages may indirectly regulate renal transferrin expression. Dotted lines indicate unknown mechanisms. DMV dorsal motor nucleus of the vagus, CAP cholinergic anti-inflammatory pathway, NA noradrenaline, β2AR β2-adrenergic receptor, ACh acetylcholine, α7nAChR α7 nicotinic acetylcholine receptor, Hes1 hairy and enhancer of split 1.

Our experiments using peritoneal macrophages also revealed that the stimulation of α7nAChR activates Notch signaling to exert anti-inflammatory effects (Fig. 1). During LPS treatment, Notch inhibition by DAPT attenuated GTS-21 efficacy on the reduction of Ccl2 (Fig. 1D), indicating Notch signaling potentially mitigates inflammatory responses in an α7nAChR-dependent manner. These results are consistent with a previous study suggesting that Notch signaling dampens LPS-induced inflammation by suppressing NF-κB transcription activity in murine peritoneal macrophages³⁸, though Notch signaling in macrophages has both pro- and anti-inflammatory roles^26,27,39,40. However, the molecular mechanisms by which α7nAChR activates Notch signaling are poorly characterized (Fig. 6). One potential mediator of the Notch regulation by α7nAChR is the JAK2/STAT3 pathway, a downstream target of α7nAChR essential for the anti-inflammatory effects of VNS⁴¹. Furthermore, recent research has implicated that increased phosphorylation of STAT3 promotes the transcription of Notch ligands, Jag1 and Dll1, in breast cancer cells and primary monocytes, respectively^42,43. These findings suggest that JAK2/STAT3 upregulate Notch signaling. Conversely, Notch signaling has also been reported to activate JAK2/STAT3 signaling^43,44. The bidirectional communication between JAK2/STAT3 and Notch signaling^42–45 might explain why the Notch blockade by DAPT incompletely abrogated the suppressive effects of GTS-21-mediated α7nAChR stimulation on Ccl2 release (Fig. 1D). Other factors that may mediate the α7nAChR-Notch interaction are adenosine signaling and a neuronal guidance molecule netrin-1, which is also known as a ligand for adenosine A2B receptor^46–49. Previous studies have demonstrated that adenosine A2A receptor enhances α7nAChR upregulation by nicotine and VNS promotes netrin-1 expression in an α7nAChR-dependent manner^50,51. In addition, netrin-1 has been shown to activate Notch signaling by modulating the retention of Notch2 at the cell surface⁵². Further investigations are needed to elucidate the regulatory relationships between α7nAChR and Notch signaling in macrophages.

Accumulating evidence indicates that the spleen is indispensable for kidney protection against AKI by VNS^21–23,53. The removal of the spleen (splenectomy) prior to CAP activation abolished tissue-protective effects on the kidneys in various rodent AKI models including IRI-, LPS-, and cisplatin-induced injury^21–23,53. However, which factors mediate the spleen-kidney axis and how those factors protect the kidneys in the context of CAP activation remain open questions (Fig. 6). We believe that our findings offered two possibilities to answer these questions. First, splenic macrophages or other immune cells from the spleen might directly accumulate in the kidneys during inflammation, which is suppressed by VNS. We did not identify the origin of infiltrating macrophages into the kidneys, but there seemed to be a correlation between the spleen size and macrophage numbers in the kidneys in our model of LPS-induced AKI (Fig. 3B, F). Second, our study revealed that VNS prevented the production of various kinds of cytokines and chemokines in the spleen (Fig. 2C, D), and hence soluble factors released by the spleen potentially mediate kidney protection via the circulating blood or lymph. Based on our data, Ccl2, a potent chemoattractant for macrophages and monocytes, may play a role in the development of LPS-induced AKI by trafficking to the kidneys and activating resident renal cells. On the other hand, VNS could inhibit LPS-induced Ccl2 release from splenic macrophages, which dampens inflammation in the kidneys. In our experiments, after VNS and LPS treatments, Notch2^MΦKO mice showed increased numbers of macrophages in the kidneys accompanied by a larger spleen size, compared to Notch2^MΦWT mice (Fig. 3B, F). Higher levels of Ccl2 were observed in splenic macrophages of LPS-treated Notch2^MΦKO mice even after VNS (Fig. 3C). These results indicate that Notch2 in macrophages is crucial for the spleen-kidney axis to exert renoprotective effects during CAP activation and is associated with both the first and second possibilities above.

Although molecular mechanisms in the spleen underlying the anti-inflammatory and tissue-protective effects of the CAP are emerging^{12,23,24,54,55}, mechanisms in the kidneys have not been well investigated. To address this, we performed RNA-seq analysis using whole kidney sections and found that VNS mainly affected key metabolic pathways under physiological conditions (Fig. 4B). The role of VNS in renal metabolic regulation remains largely unexplored, whereas accumulating evidence demonstrates its critical functions in metabolic homeostasis in other organs⁵⁶. In the heart, VNS has been reported to promote mitochondrial biogenesis and energy metabolism by activating AMPK signaling pathways⁵⁷. A clinical study targeting patients with drug-resistant epilepsy also revealed that VNS upregulates SIRT1/PPARα expression in leucocytes and lowers de novo lipogenesis markers in erythrocytes, modulating systemic lipid metabolism⁵⁸. These metabolic factors should be considered as important targets for the kidney-protective effects of VNS in the future. Furthermore, we identified Trf as a principal VNS-responsive gene (Fig. 5A). Trf is a glycoprotein that regulates iron transport and homeostasis. It delivers iron to cells by binding to TfR1 on the plasma membrane. Low levels of Trf in the serum correlate with the severity of inflammatory diseases, indicating its involvement in host defense^30–32. Trf plays a role in the pathophysiology of AKI and protects kidneys against IRI^34,59,60. Consistent with these observations, we demonstrated that VNS enhanced Trf expression and VNS-mediated CAP activation might promote the utilization of Trf through TfR1 to attenuate kidney injury (Fig. 5). Our data also suggested that Notch2 in macrophages is possibly associated with Trf upregulation by CAP activation since Trf levels in Notch2^MΦKO mice were lower than in Notch2^MΦWT mice after the applications of VNS and LPS (Fig. 5B). To further examine the link between Notch2 and Trf, we attempted to investigate whether Trf supplementation attenuates pronounced inflammation observed in Notch2^MΦKO mice. However, a preliminary experiment using WT mice revealed that the administration of human apo-transferrin (hApo-Trf) prior to the LPS insult did not suppress splenomegaly and an increase in renal Kim-1 expression induced by LPS (Supplementary Fig. 9A–C). The mRNA expression of Trf in the kidneys was also not altered by hApo-Trf (Supplementary Fig. 9D). These results indicate that hApo-Trf treatment does not mimic the kidney-protective effects of VNS in our model. Considering that renal Trf mRNA levels were upregulated by VNS during LPS-induced inflammation (Fig. 5B), upregulated production of endogenous Trf within the kidneys, rather than circulating (exogenous) Trf, might be crucial for the kidney protection against LPS-induced AKI. Recent research has demonstrated that Trf translocated from the cytosol into the nucleus, where it bound to PPARα and promoted the transcription of fatty acid oxidation-related genes in liver tumor cells⁶¹. Since our RNA-seq analysis also revealed that VNS modulates Trf expression and lipid metabolism (Fig. 5B and Fig. 4B), endogenous Trf within the kidneys may accelerate fatty acid oxidation for renoprotection. Further assessment of whether kidney-specific overexpression of Trf counteracts the loss of VNS effects in Notch2^MΦKO mice is needed to expand our knowledge of the mechanisms by which macrophage-specific Notch2 enhances Trf expression to protect the kidneys against LPS-induced AKI.

This study has two major limitations. First, we employed a murine model of LPS-induced AKI that is widely utilized to investigate the early pathophysiology of SA-AKI⁶². LPS administration typically results in a rapid, transient spike in cytokine levels, whereas clinical sepsis is characterized by sustained infection or viable bacteremia, which leads to more prolonged, lower-magnitude systemic cytokine production⁶³. Alternative sepsis models such as cecal ligation and puncture (CLP) may more closely recapitulate these features of sepsis, but they introduce substantial confounding variables that increase experimental complexity⁶⁴. For example, CLP can result in variable bacterial exposure depending on the size and number of cecal punctures, the length of bowel ligated, and the amount of stool expressed. In addition, CLP introduces variability related to bacterial burden, peritonitis, systemic hemodynamic instability, fluid shifts, anesthetic exposure, and differences in host immune responses to live bacterial infection. These factors can independently influence inflammatory signaling pathways and organ injury, thereby complicating mechanistic interpretation. For the current mechanistic studies, we therefore used LPS-induced sepsis as a more controlled experimental model to establish proof of concept and determine whether Notch signaling is required for VNS-mediated CAP activation and protection. Future studies may incorporate more biologically relevant models such as CLP or cecal slurry injection. Second, our experiments using Notch2^MΦKO mice did not eliminate the developmental effects of Notch2 deficiency on inflammatory responses. The Notch signaling plays a role in cell fate determination during embryonic development, and global knockout of Notch2 in mice is embryonic lethal^25,65. In addition, although the body weight and the numbers of resident renal macrophages were comparable between Notch2^MΦKO mice and Notch2^MΦWT mice, Notch2^MΦKO mice showed enlarged spleens compared to controls (Supplementary Fig. 5A, B, F), indicating that germline knockout of Notch2 in macrophages may promote inflammation from birth to adulthood even under basal conditions. Hence, conditional gene knockout studies are needed to clarify the precise functions of macrophage-specific Notch2 in anti-inflammation during adulthood.

In conclusion, our findings indicated that Notch2 signaling in macrophages mediates the anti-inflammatory effects of VNS during LPS-induced AKI (Fig. 6). We also identified transferrin as a potential target for protecting kidneys against endotoxin-induced inflammation in the context of CAP activation, whose expression in the kidneys may be regulated by macrophage-specific Notch2 (Fig. 6). We believe that this study provides a comprehensive insight into molecular mechanisms underlying the anti-inflammatory and tissue-protective effects of CAP.

Methods

Animals

We have complied with all relevant ethical regulations for animal use. All animal procedures were performed in accordance with the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals or Basic Guidelines for the Proper Conduct of Animal Experiments in Research Institutions (Notice No. 71, MEXT, Japan). All animal protocols were approved by the University of Virginia (UVA) Institutional Animal Care and Use Committee or the Animal Care Committee of Chiba University (A7-174 and A8-087).

Male mice (aged 7-13 weeks) were used for all experiments. Mice were maintained in standard vivarium housing (up to five mice per cage, 12-hour light/dark cycle) on a chow diet, and water was freely available. Macrophage (MΦ)-specific Notch2 knockout (KO) mice (Notch2^MΦKO) were generated by crossing heterozygous CX3C motif chemokine receptor 1 (Cx3cr1)-Cre mice (B6J.B6N(Cg)-Cx3cr1^{tm1.1(cre)Jung}/J, Jackson Laboratory (#025524), provided by Dr. Kodi S. Ravichandran (UVA))⁶⁶ with Notch2^flox/flox mice (B6.129S-Notch2^tm3Grid/J, Jackson Laboratory (#010525), provided by Dr. James Zimring (UVA))⁶⁷. These two strains were on a mixed background, and Cre-negative Notch2^flox/flox mice were used as wild-type (WT) controls (Notch2^MΦWT). WT C57BL/6J mice were purchased from Jackson Laboratory (#000664) or Japan SLC, Inc. and bred in our facilities. The sample size was based on similar previous studies. Mice were anesthetized with ketamine/xylazine (120/12 mg/kg, intraperitoneal (i.p.) injections) or 1.5-3% isoflurane in O₂ (vaporizer) for surgeries and euthanasia.

Electrical vagus nerve stimulation (VNS)

All mice were anesthetized, and body temperature was maintained at 37.0 ± 0.5 °C with a servo-controlled temperature pad (TC-1000; CWE). Electrical stimulation of the left cervical vagus nerve was performed as previously described^21,22. Briefly, the left cervical vagus nerve was isolated via a midline cervical incision, and the nerve was put on a bipolar silver wire electrode (Cooner Wire, Cat#AS633). Electrical stimulation of the vagus nerve (square wave, 5 Hz, 50 or 100 μA intensity, 1 ms pulses) was applied for 10 minutes (min) using an Isostim Stimulator (A320; World Precision Instruments) or Constant Current Stimulator USE-210 (Unique Medical, Japan). In parallel, the vagus nerve was isolated without electrical stimulation for the sham operation.

Lipopolysaccharide (LPS)-induced AKI model

LPS-induced AKI was induced by i.p. injection of lipopolysaccharide (LPS) from Escherichia coli O111:B4 (5 mg/kg in a vehicle of saline, Sigma-Aldrich, Cat#L4391, Lot: 0000140462 or 0000185317 or 0000297998). VNS or sham treatments were applied 34 or 24 hours (h) before LPS administration, and mice were euthanized 14 h or 24 h after the LPS injection. For the analysis of therapeutic effects of VNS, mice were subjected to VNS or sham applications at 1 h post-LPS injection and were euthanized 23 h after VNS (or sham VNS).

Ferristatin II administration

Ferristatin II (Cayman Chemical, Cat#36621) was dissolved in saline (1 mg/ml). Male Notch2^MΦWT mice were administered ferristatin II (10 mg/kg/day) or saline by i.p. injection once a day for 4 days. On days 2 and 3 of ferristatin II treatment, mice were subjected to VNS (or sham VNS) and treatment with LPS (or saline), respectively. VNS or LPS treatment occurred immediately after the injection of ferristatin II on days 2 and 3, respectively, and mice were euthanized 24 h after LPS. For the assessment of drug efficacy (TfR1 internalization or degradation)^35,36, ferristatin II (10 mg/kg/day) was injected intraperitoneally once a day for 2 days. 24 h after the second dose, the spleen and kidneys were harvested from euthanized mice.

Collection of blood, spleen, and kidneys

Blood was collected under anesthesia from the retro-orbital sinus or by cardiac puncture, and plasma was prepared by centrifuging at 8500 rpm for 10 min at room temperature. Spleens were harvested immediately after euthanasia followed by weighing. The spleen size was normalized to body weight on the day of euthanasia. The kidney capsules were removed prior to kidney dissection for immunostaining and RNA extraction.

Preparation of splenic single-cell suspensions and Cx3cr1⁺ macrophages

Whole spleens were mechanically disrupted using the rubber end of a 10 ml syringe plunger and subsequently passing through a 40 μm cell strainer (Fisher Scientific, Cat#22363547) with phosphate-buffered saline (PBS) to a single-cell suspensions. The suspensions were centrifuged at 400 g for 5 min at 4 °C. The cell pellets were suspended in 2 ml of red blood cell (RBC) lysis buffer (BioLegend, Cat#420301 or #420302) per sample, and RBCs were lysed for 4 min at room temperature. The cells were then washed with 8 ml of PBS, centrifuged, and resuspended in PBS. After counting cell numbers, the cells were used for further experiments. Splenic Cx3cr1⁺ macrophages were isolated from these suspensions using the magnetic cell separation MojoSort^TM Mouse CX3CR1 Selection Kit (BioLegend, Cat#480056) and PBS supplemented with 0.5% bovine serum albumin (BSA, Sigma-Aldrich, Cat#A9418) and 2 mM ethylenediaminetetraacetic acid (EDTA) as an isolation buffer.

Periodic Acid-Schiff (PAS) staining and blood urea nitrogen (BUN) measurement

Each one-quarter of the right and left kidneys was fixed with 10% neutral-buffered formalin for 24 h and then stored in 70% EtOH. Kidney tissues were embedded in paraffin, sectioned (5 μm), and stained with Periodic acid-Schiff (PAS) (UVA Research Histology Core). Injured tubular areas were assessed by using a light microscope (AxioImager Z.1, Carl Zeiss), Stereo Investigator software (MBF Bioscience), and the area fraction fractionator (AFF) probe (a stereological method that provides unbiased estimation of the area fraction of injured tubules with intracellular vacuoles in the cortex within the kidney sections). The AFF was applied using a systematic random sampling scheme with a 400 μm x 400 μm counting frame, an 800 μm x 800 μm sampling grid size, and an 85 μm grid spacing. The sampling fraction was 25% of a total average area ( ± SEM) of 7.96 × 10⁶ ± 0.144 × 10⁶ μm² for each kidney section. Plasma blood urea nitrogen (BUN) was measured using the Urea Nitrogen (BUN) Colorimetric Detection Kit (Arbor Assays).

Kidney immunohistochemistry

Paraffin sections of the kidneys were deparaffinized in xylene and rehydrated through a graded ethanol series. Antigen retrieval was performed by treating the sections with proteinase K (Bio Basic, Cat#PB0451) for 15 min at 37 °C. Endogenous peroxidase activity was quenched by incubation with 3% H₂O₂ for 30 min at room temperature. The sections were then incubated with a blocking solution (Protein Block Serum-Free, Agilent, Cat#X0909) for 1 h at room temperature. Primary antibodies for F4/80 (listed in Supplementary Table 4) diluted in the blocking solution were applied to the sections and incubated overnight at 4 °C. Immunodetection was performed using VECTASTATIN Elite ABC-HRP kit (Vector Laboratories, Cat#PK-6104), and the reaction products were visualized by treating the sections with Metal Enhanced DAB Substrate Kit (Thermo Fisher Scientific, Inc., Cat#34065). The sections were counterstained with methyl green, dehydrated through graded ethanol, cleared in xylene, and mounted with a permanent mounting medium. The quantification of F4/80-positive cell numbers in the kidney was performed using a light microscope (AxioImager Z.1, Carl Zeiss) and Stereo Investigator software (MBF Bioscience). Five random fields were selected per kidney (ten random fields per mouse), and F4/80-positive cells were counted manually at x630 magnification. The average number of positive cells per field was calculated for each sample.

Immunofluorescence staining of the spleen and kidneys

Spleen tissues were immersed in 4% paraformaldehyde and kidney tissues were in periodate-lysine-paraformaldehyde fixative (2.0% paraformaldehyde/1.4% L-lysine/0.2% sodium periodate in 0.1 M phosphate buffer, pH 7.4) overnight. Following fixation, the tissues were washed three times with PBS and cryoprotected with sucrose at 4 °C. Spleen tissues were immersed in 10% sucrose solution for 2 h, followed by 30% sucrose solution for 3 h. Kidney tissues were treated with 18% sucrose solution for 5 h. The tissues were then embedded in optimal cutting temperature compound, and frozen sections were cut at 5 μm thickness by UVA Research Histology Core. The sections were fixed in 4% paraformaldehyde for 10 min at room temperature. Antigen retrieval was performed by treating the sections with proteinase K (Bio Basic, Cat#PB0451) for 15 min at 37 °C. Subsequently, the sections were permeabilized with 0.25% Triton X-100 diluted in PBS for 10 min at room temperature then incubated with a blocking solution (Protein Block Serum-Free, Agilent, Cat#X0909) for 1 h at room temperature. Primary antibodies for F4/80, Notch2, and TfR1 (listed in Supplementary Table 4) diluted in the blocking solution were applied to the sections and incubated overnight at 4 °C. Samples were then incubated in fluorophore-tagged secondary antibodies (listed in Supplementary Table 4) for 1 h at room temperature. The slides were mounted using a medium containing 4’,6-diamidino-2-phenylindole (DAPI) for nuclear counterstaining. Imaging was performed using a confocal microscope (Leica Stellaris 8 with DIVE, Leica).

Preparation of peritoneal macrophages and cell culture experiments

After euthanasia, the parietal peritoneum covering the peritoneal cavity of WT mice was gently exposed. 8 mL of ice-cold Dulbecco’s Modified Eagle Medium (DMEM, gibco, Cat#11965-092) supplemented with 10% fetal bovine serum (FBS, GeminiBio, Cat#100-106) and 1% penicillin-streptomycin (PS, gibco, Cat# 10378016) was injected into the peritoneal cavity using a 26-gauge needle and a 10 ml syringe. The peritoneum was gently massaged, and the injected fluid was collected by a 21-gauge needle with the same syringe. The cell suspension was filtered through a 70 μm cell strainer (Fisher Scientific, Cat#22363548) and then centrifuged (400 g for 5 min at 4 °C). The resulting peritoneal cells were resuspended in culture medium (DMEM + 10% FBS + 1% PS), manually counted with Tuerk’s Solution (Sigma-Aldrich, Cat#1092770100), plated in untreated 24-well cell culture plates (Thermo Fisher Scientific, Inc., Cat#144530) at a density of 2.0 ×10⁶ cells/well, and cultured for 22–24 h at 37 °C. After washing cells with culture medium once, DAPT (N-[N-(3,5-difluorophenacetyl-L-alanyl)]-S-phenylglycine tert-butyl ester, abcam, Cat#ab120633), dissolved in dimethyl sulfoxide (DMSO, 100 mM stock), was added at a final concentration of 10 μM (0.01% final concentration of DMSO). After 24 h, the culture medium was replaced with fresh DAPT solution, and cells were treated with GTS-21 (3-(2,4-dimethoxybenzylidene)-anabaseine dihydrochloride) (50 μM final concentration, 100 mM stock solution in water, abcam, Cat#ab120560) for 15 min followed by LPS stimulation (100 ng/ml final concentration, 1 mg/ml stock solution in PBS, Sigma-Aldrich, Cat#L4391, Lot: 0000140462) for 24 h (total treatment time: DAPT 48 h, GTS-21 15 min + 24 h, LPS 24 h). Cell culture supernatants and RNA were collected for further experiments.

Cytokine and chemokine measurement

A bead-based immunoassay (Luminex^® multiplex assay) was performed to comprehensively quantify cytokine/chemokine levels in cell culture supernatants of peritoneal macrophages (described in the previous section) and tissue lysates of the spleen. To obtain spleen lysates, single-cell suspensions of the mouse spleen in which RBCs were removed were centrifuged and the supernatants were discarded. The cells were then lysed using Pierce™ IP Lysis Buffer (Thermo Fisher Scientific, Inc., Cat#87787) supplemented with Halt™ Protease and Phosphatase Inhibitor Cocktail (Thermo Fisher Scientific, Inc., Cat#78440) and EDTA (at a final concentration of 5 mM). After centrifugation at 13,000 rpm for 15 min at 4 °C, the supernatants were used for the assay. The samples (macrophage culture supernatants and spleen lysates) were analyzed using the MILLIPLEX^® Mouse Cytokine/Chemokine Magnetic Bead Panel (MilliporeSigma, Cat#MCYTOMAG-70K-PX32), xMAP INTELLIFLEX^® system, and MILLIPLEX^® Analyst software (UVA Flow Cytometry Core). The panel included a total of 32 cytokines and chemokines: Tnf-α, Il-1α, Il-1β, Il-2, Il-3, Il-4, Il-5, Il-6, Il-7, Il-9, Il-10, Il-12 (p40), Il-12 (p70), Il-13, Il-15, Il-17, Ifn-γ, Lif, Ccl2 (Mcp-1), Ccl3 (Mip-1α), Ccl4 (Mip-1β), Ccl5 (Rantes), Ccl11 (Eotaxin), Cxcl1 (Kc), Cxcl2 (Mip-2), Cxcl5 (Lix), Cxcl9 (Mig), Cxcl10 (Ip-10), G-csf, M-csf, Gm-csf, Vegf. Values for cytokine/chemokine concentrations in the spleen were normalized to numbers of splenocytes without RBCs. Every well in the heatmap indicates an average concentration of each experimental group. In addition, enzyme-linked immunosorbent assays (ELISA) were performed for the measurement of Ccl2 concentrations in the culture supernatants of peritoneal macrophages. Ccl2 levels were determined using the Mouse CCL2/JE/MCP-1 DuoSet ELISA (Cat#DY479) and DuoSet ELISA Ancillary Reagent Kit 2 (R&D Systems). Synergy HT Microplate Reader (Biotek) was used as an ELISA plate reader.

Quantitative real-time PCR (qPCR)

Total RNA from the frozen kidney was extracted and then purified with TRI Reagent® (Molecular Research Center, Inc., Cat#TR118) according to the manufacturer’s instructions. RNeasy Mini kit (QIAGEN) or Quick-RNA MiniPrep Kit (Zymo Research) was used for RNA isolation from the splenic Cx3cr1⁺ macrophages and peritoneal macrophages. cDNA synthesis was performed with iScript^TM cDNA synthesis kit (BIO-RAD). The mRNA expression levels of each gene were quantified by real-time RT-PCR with iTaq^TM Universal SYBR^® Green Supermix (BIO-RAD) on a CFX Connect Real-Time System Bio-Rad instrument or a CFX Opus 96 Real-Time PCR System (BIO-RAD). Expression levels were normalized against those of murine β-actin, and the relative values of each data point were calculated using the ΔΔCt method. The primer sequences are listed in Supplementary Table 5.

RNA sequencing (RNA-seq)

Total RNA was extracted from mouse kidneys as described in the qPCR section above. This analysis included three factors (1, Notch2^MΦWT or Notch2^MΦKO; 2, Sham or VNS; 3, Saline or LPS) and a total of six groups (triplicate biological samples per group). The quality of total RNA was evaluated by Bioanalyzer (Agilent) at the UVA Genome Analysis and Technology Core. RNA-seq library preparation and sequencing were performed at GENEWIZ^® from Azenta Life Sciences (South Plainfield, New Jersey, USA). ERCC RNA Spike-In Mix (Thermo Fisher Scientific, Inc., Cat#4456740) was added to normalized total RNA prior to library preparation following the manufacturer’s protocol. RNA sequencing libraries were prepared using the NEBNext Ultra II RNA Library Prep for Illumina (New England Biolabs) using the manufacturer’s instructions. Briefly, mRNAs were initially enriched with Oligo d(T) beads. Enriched mRNAs were fragmented for 15 min at 94 °C. First-strand and second-strand cDNA were subsequently synthesized using primers which included random priming sites and Unique Molecular Identifiers (UMIs). UMI sequences were incorporated in the resultant cDNA. cDNA fragments were end-repaired and adenylated at 3’ ends, and universal adapters were ligated to cDNA fragments, followed by index addition and library enrichment by PCR with limited cycles. The sequencing libraries were multiplexed and clustered onto a flowcell on the Illumina NovaSeq X Plus instrument according to the manufacturer’s instructions. The samples were sequenced using a 2x150bp Paired-End (PE) configuration.

Data analysis of RNA-seq

Data processing and analysis were performed by GENEWIZ^® from Azenta Life Sciences (South Plainfield, New Jersey, USA). Sample barcode and UMI information were extracted from the FASTQ files and saved in the header of the transformed FASTQ files with umis 1.0.7. The transformed FASTQ files were then aligned to mouse (mus musculus) GRCm38 transcriptome using STAR aligner v.2.5.2b and tag count for genes were generated by umis 1.0.7. The STAR aligner is a splice aligner that detects splice junctions and incorporates them to help align the entire read sequences. Unique gene hit counts were calculated by using feature Counts from the Subread package v.1.5.2 after removing PCR duplicates by UMI sequence. Only unique reads that fell within exon regions were counted. After the extraction of gene hit counts, differentially expressed gene (DEG) analysis was performed using DESeq2. The Wald test was used to generate P values and Log2 fold changes. Genes with adjusted P values < 0.10 and absolute log2 fold changes > 0.585 ( > 1.5-fold increase or < 0.67-fold decrease) were determined as DEGs for each comparison. Gene ontology (GO) analysis was performed on the statistically significant set of DEGs by implementing the software GeneSCF. The GO list was used to cluster the set of genes based on their biological process and determine their statistical significance. Significant DEGs were clustered by their GO and the enrichment of GO biological process terms was tested using Fisher exact test (GeneSCF v1.1-p2). Significantly enriched GO terms with adjusted P values less than 0.05 in the DEG sets were extracted up to the top 15 terms.

Statistics and reproducibility

Data are represented as means ± standard error of the mean (SEM). Statistical analysis was performed with GraphPad Prism software ver. 9 or 10 (GraphPad Software Inc.). We applied the unpaired two-tailed Student’s t test for comparisons between two groups. One-way ANOVA followed by post hoc Tukey’s multiple comparison test was performed to determine the significance of differences among more than three groups. No data were excluded from the analysis unless significant outliers were detected using a statistical outlier test (ROUT Test; GraphPad Prism). P < 0.05 was considered statistically significant. The experiments in this study were independently repeated at least three times with consistent results.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Author contributions

S.K. and M.O. designed research; S.K., H.I., W.N., J.Y., and S.Z. performed research; S.K., H.I., W.N., V.S., and R.H. analyzed data; and S.K., H.I., W.N., R.S., T.M., and M.O. wrote the manuscript.

Peer review

Peer review information

Communications Biology thanks Hernando Gómez and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. Primary Handling Editors: Connie Wong and Joao Valente. A peer review file is available.

Funding

This work is supported by the National Institutes of Health (R01 DK123248 and R01 DK085259 to M.O.), the Uehara Memorial Foundation Research Fellowship (202030018 to S.K.), the Japan Society for the Promotion of Science (JSPS) Overseas Research Fellowships (202360504 to S.K.), and the JSPS KAKENHI (26K18552 to S.K. and 25K02439 to T.M.). V.S. was supported by the NIH training grant (1U2CDK129500/DK132771).

Data availability

The data that support the findings of this study are available from the corresponding authors upon reasonable request. The source data behind the graphs in the main and Supplementary Figs. are provided in Supplementary Data. The RNA-seq dataset reported in this study has been deposited in the Gene Expression Omnibus (GEO) under the accession number GSE284008.

Competing interests

The authors declare no competing interests.

Supplementary information

The online version contains supplementary material available at https://doi.org/10.1038/s42003-026-10620-2.