Five-week-old Swiss female mice were obtained from the Multidisciplinary Center for Biological Research at the University of Campinas (Campinas, Brazil). The mice were kept in a temperature-controlled environment with a 12-h photoperiod. Experiments were performed in accordance with the ethical guidelines and regulations for use of laboratory animals. Ethics approval for this study, including the design for sepsis development and mortality analysis presented in this paper, was obtained from the State University of Campinas Ethics Committee (Protocol 5733-1). Importantly animals did not experience any form of suffering throughout the study. The female mice were randomly separated into two groups (25 dams per group), fed with either a HFD or a standard diet (SC) (NUVILAB^® Cr-1, Nuvital, PR, Brazil) ( Table 1 ) for 4 weeks ad libitum before mating. Dams continued with the diet during pregnancy and lactation. After birth, the litter size was culled to eight mice per litter. Male offspring were weaned on postnatal day 18 (P18) and fed with standard chow until P28 ( Figure 1 ). Each experimental protocol, such as surgery and survival tests, was conducted using one pup from each dam to constitute the respective group. Offspring at different ages were utilized for the receptor expression experiment, specifically at birth (P0), P56, and P82. The HFD was prepared in our laboratory according to the AIN-93G but modified for high-fat content (45%) as described previously (34).

Mice were anaesthetized with a mixture containing ketamine (139.2 mg kg^-1 body weight [bw]), diazepam (4 mg kg^-1 bw), and xylazine (18.4 mg kg^-1bw) and subsequently euthanized by decapitation for tissue collection. Tissue samples were frozen in liquid nitrogen and stored at -80°C until processing. Isoflurane, an inhalational anaesthetic, was used for induction and maintenance of general anaesthesia during stereotaxic surgery. It was used at 3%–4% for induction and reduced to 2% during surgery.

The offspring were separated as described below to evaluate the inflammatory response.

Design 1

An LPS-induced sepsis mouse model was used. Mice were treated with a lethal dose of LPS diluted in sterile saline and administered intraperitoneally (IP) at 30 mg kg^-1 bw. The offspring were observed for 72 h. The survival rate was recorded every 1 h. In the survival study, mice were allowed ad libitum access to food and water. The experiment was replicated twice to validate the obtained results.

In a separate study, mice were treated intracerebroventricularly (ICV) with benztropine (mesylate) 20 min before the LPS challenge. Benztropine, an m1mAChR antagonist, or phosphate-buffered saline (PBS) was injected at 40 µg kg^-1 bw. The mice were sacrificed 10 h after LPS administration; the serum was collected for analysis. The time was defined based on the survival rate.

Design 2

To explore the underlying mechanisms of central m1mAChR-mediated anti-inflammatory effects in mice, McN-A-343, an m1mAChR agonist, was administered ICV at 5 ng kg^-1 bw. The mice were euthanized 2 h after injection. In a separate study, the mice were treated with an agonist (ICV) 20 min before LPS challenge (1 mg kg^-1 bw IP). The mice were euthanized 2 h after LPS challenge.

At P28, offspring of both groups (SC-O and HFD-O) were perfused with 4% paraformaldehyde (PFA). Then, the brains were extracted and fixed in 4% PFA. Subsequently, the brains were embedded in Tissue-Tek (Sakura, Torrance, CA, USA), frozen, and cut into 15-µm thick coronal sections, following The Rat Brain in Stereotaxic Coordinates by Charles Watson and George Paxinos. The slides were incubated in blocking solution (3% bovine albumin; Sigma-Aldrich, St. Louis, MO, USA) for 90 min, followed by incubation with specific primary antibodies overnight at 4°C. The primary antibodies used were anti-Chrm1 diluted 1:500 (sc-365966, Santa Cruz Biotechnology, Inc, California) and anti-F480 diluted 1:500 (ab6640, Abcam, Inc, Boston). The slides were washed and incubated with appropriate secondary antibodies for 120 min. The secondary antibodies used were Donkey anti-rabbit conjugated to Alexa 488 diluted 1:500 (A-21206, Thermo Fisher Scientific, Inc, Waltham, MA) and goat anti-Rat conjugated to Cy3 diluted 1:1000 (ab6953, Abcam). TO-PRO-3 Iodide was used for nuclear labelling (1:1000; Life Technologies Inc, Carlsbad, CA). The slides were visualized and images were captured using a TCS SP5II Leica confocal microscope (Leica Microsystems, Wetzlar, Germany).

The offspring were sacrificed by decapitation on P28 after overnight fasting, and blood was collected. The samples were centrifuged at 1300 rpm for 15 min at room temperature. The serum was collected and stored at -80°C until processing. The cytokine levels were measured using DuoSet enzyme-linked immunosorbent assay kits, DY410-05 mouse TNF; DY401-05 mouse IL1-β/IL1-F2; and DY417-05 mouse IL-10 (R&D Systems, Minneapolis, MN, USA). CD14 levels were measured using Mouse CD14 Quantikine ELISA Kit, #MC140 (R&D Systems, Minneapolis, MN, USA. The C-reactive protein (CRP- K059-8.1) and albumin levels (K040-1) were obtained from biochemical analysis provided by BIOCLIN (Quimica Basica LTDA, Belo Horizonte, Brazil).

The offspring from control and HFD dams received 3%–4% isoflurane inhalational anaesthesia and were placed in a stereotaxic instrument (Stoelting Co. Wood Dale, Illinois). Isoflurane was reduced to 2% during surgery for cannula implantation. Afterwards, a 26G needle was used for the cannula and inserted into the lateral ventricle through a cranial incision. The following coordinates relative to the bregma were used to access the lateral ventricle: anterior/posterior axis, 0.34 mm from bregma to the rear; lateral, 1 mm from the midline; dorsoventral, 2.2 mm from the surface of the skull. Dental acrylic glue was added to secure the cannula following correct positioning. After surgery, the animals were allowed to recover from anaesthesia on a warm pad. Carprofen, an analgesic, was administered for postoperative pain (5 mg kg^-1 bw, IP). The cannula placement was tested 6 days after the surgery by measuring the dipsogenic response to angiotensin II injection (2 µL of a 1 × 10^-6M solution, ICV) (Sigma-Aldrich Inc, MERK, St Louis, MO). The time and dose of McN-A-343 (agonist) and benztropine (antagonist) treatment were standardized from time-course and dose-response experiments (data not shown).

Bone marrow cells were isolated as described previously (21). The long bones (femur and tibia) were removed from the offspring and placed in 0.5 mL perforated tubes inside 1.5 mL tubes, which were then centrifuged at 1200 g for 15 s at 4°C. The cell pellet was resuspended in Roswell Park Memorial Institute (RPMI) 1640 culture medium (Invitrogen, Carlsbad, CA, USA) supplemented with 10% foetal bovine serum (FBS; Invitrogen) and 1% penicillin (100 U/mL)/streptomycin (100 μg/mL) (Invitrogen). The cells were counted with a Neubauer chamber and placed on 60 mm culture dishes. The cells were cultivated for 7 days at 37°C in an atmosphere containing 5% CO₂ and 95% humidity. After this period, photos were taken of the culture, and cells were trypsinized and collected for western blotting, RT-PCR, and flow cytometry analysis.

Tissues were homogenized in freshly prepared ice-cold buffer (1% v/v Triton X-100, 0.1 M Tris, pH 7.4, 0.1 M sodium pyrophosphate, 0.1 M sodium fluoride, 0.01 M EDTA, 0.01 M sodium vanadate, 0.002 M PMSF (Phenylmethylsufonyl fluoride), and 0.01 mg mL^-1 aprotinin). The samples were centrifuged at 12,000 rpm for 30 min at 4°C. The supernatant was removed and the protein concentration was determined using the Bradford dye-bleeding method. The samples were resuspended in Laemmli sample buffer and boiled for 5 min before separation by SDS-PAGE using a miniature slab gel apparatus (Bio-Rad, Richmond, CA, USA). The separated proteins were electrotransferred from the gel to a nitrocellulose membrane for 30 min in a transfer buffer that contained methanol and SDS. These membranes were incubated overnight at 4°C with specific antibodies: α7nAChR (bs-1049R, Bioss Antibodies Inc, Woburn, MA), phosphorylated JNK (#9255; Cell Signaling Technology Inc, Danvers, MA), phosphorylated STAT3 (#9145, Cell Signaling), GAPDH (sc-32233, Santa Cruz Biotechnology, Inc., California), and phosphorylated NF-κB (#30335, Cell Signaling Technology Inc, Danvers, MA). Then, after washing with Tris-buffered saline (TBS)-Tween 20 (TTBS; 10 mM Tris, 150 mM NaCl, 0.5% Tween 20), the nitrocellulose membranes were probed with peroxidase-conjugated secondary antibodies (KPL, Gaithersburg, MD, USA) for 90 min at room temperature. Proteins were detected by a chemiluminescence kit (SuperSignal West Pico Chemiluminescent Substrate, Thermo Fisher Scientific Inc) and bands were evaluated by densitometry using Scion Image software (ScionCorp, MD, USA). The intensities of the bands were normalized to the loading control (GAPDH).

Frozen tissues were homogenized in TRIzol reagent (Life Technologies) for RNA extraction according to the manufacturer’s instructions. After incubation for 5 min at room temperature for complete dissociation, chloroform was added to the homogenate. Following centrifugation, the RNA phase was precipitated with isopropyl alcohol and the pellet was washed with 75% and 100% ethanol. After drying, the pellet was resuspended in ultra-pure water and stored at -80°C. RNA was quantified with a Nanodrop ND-2000 (Thermo Fisher Scientific). Reverse transcription was performed with 3 µg of total RNA using the High-Capacity cDNA Reverse Transcription kit (Life Technologies). Relative expression was determined using TaqMan Gene Expression Assays (Thermo Fisher Scientific) and SYBR Green Master Mix (Bio-Rad). The following TaqMan Gene Expression Assays were used: Chrna7 (Mm01312230_m1), Il6 (Mm01312230_m1), Tnf (Mm00443258_m1), Il1b (Mm00434228_m1), Socs3 (Mm00545913_s1), and Il10 (Mm01288386_m1). Gapdh (4351309; Applied Biosystems, USA) was used as endogenous control.

Quantitative PCR was performed with the SYBR Green Master Mix (Bio-Rad). The primers used are listed in Table 1 . Real-time PCR was performed on an AB/Prism 7500 fast platform. The data were analysed using the Sequence Detection System 2.0.5 software.

Cells isolated from the spleen and bone marrow were submitted to flow cytometry analysis. The spleen and bone marrow cells were evaluated with a macrophage panel, which determined the number of CD45⁺, F480⁺, CD11c⁺, and CD206⁺ cells. Spleen cells were also evaluated with a lymphocyte panel, which determined the number of CD3⁺, CD4⁺, CD8⁺, and Ly6 G⁺ cells.

Spleens were collected, and then gently dissociated by a needle and rinsed with PBS. Single-cell suspensions (1 × 10⁶) were then suspended in DMEM/FBS). The cells were treated with specific antibodies conjugated to FITC (CD45; CD206), PECy7 (CD45), APC (F480; CD4), PE (CD11c; Ly6G; CD8), and Alexa 488 (CD3). The incubation was at room temperature for 15 min protected from light. The analysis was performed in BD-FACS Accuri Cytometry (Becton Dickinson, MD, USA) and 10,000 events were acquired. The data were analysed using the FlowJo 7.6 software.

Bone marrow cells were collected after 7 days of spontaneous differentiation. The cells were treated with specific antibodies conjugated to FITC (CD206), PECy7 (CD45), APC (F480), and PE (CD11c). The incubation was at 4°C for 15 min protected from light. Analysis was performed in BD-FACS Accuri Cytometry (Becton Dickinson) and 10,000 events were acquired. The data were analysed using the FlowJo 7.6 software.

The results are presented as the mean ± standard error. The data were evaluated with the Kolmogorov–Smirnov test to determine whether they were normally distributed. After confirming a normal distribution, Student’s t-test for unpaired samples or analysis of variance (ANOVA) was used. ANOVA was followed by the Bonferroni post hoc test to determine differences between more than two groups. The log-rank test was used to analyse the survival rate. Statistical significance for all analyses was set at p < 0.05. All statistical comparisons were performed using GraphPad Prism 9.5.3 (GraphPad Software, San Diego, CA, USA).

In the study, we administered a lethal dose of LPS to SC-O and HFD-O to induce sepsis and then recorded their survival rates. First, we assessed whether the LPS challenge was effective in inducing sepsis in both groups. We measured biomarkers of sepsis and the inflammatory response in the serum of the offspring 10 h after the LPS challenge.

The CRP levels were higher after LPS treatment, with no significant difference between SC-O and HFD-O. Albumin levels did not show any significant differences between the groups. CD14 levels were elevated in all offspring treated with LPS and exceeded the highest point of the standard curve ( Table 2 ). TNF, IL1-β, and IL-10 were undetectable in SC-O and HFD-O that were not challenged with LPS. TNF and IL-10 levels were lower in HFD-O compared with SC-O after LPS injection. However, this decrease was prevented by treatment with an m1mAChR antagonist. IL1-β was only detected in SC-O after LPS injection and in HFD-O treated with the m1mAChR antagonist ( Table 2 ). We also evaluated the spleen weight. LPS significantly increased the spleen weight in SC-O compared with SC-O that were not treated with LPS ( Table 2 ). However, there was no difference in spleen weight in HFD-O treated with LPS.

Considering that LPS was sufficient to activate the inflammatory response, we assessed the SC-O and HFD-O survival curves. The mortality rate was significantly higher in SC-O compared with HFD-O ( Figure 2 ). However, prior ICV treatment with benztropine (mesylate), a pharmacological antagonist of m1mAChR, increased the mortality of HFD-O to levels comparable to SC-O ( Figure 2 ). This finding suggests that central m1mAChR is involved in protecting HFD-O against sepsis.

We evaluated the distribution and expression of cholinergic receptors (α7nAChR and m1mAChR) in the hypothalamus ( Figure 3 ). First, we evaluated the mRNA levels of both receptors at P0, P28, P56, and P82. While m1mAChR mRNA expression was higher in HFD-O at P0, P28, and P82 compared with SC-O ( Figure 3A ), α7nAChR mRNA expression in HFD-O was reduced at P0 and increased at P82 compared with SC-O ( Figure 3B ). Additionally, we evaluated m1mAChR protein expression by western blot at P28, P56, and P82. We noted increased hypothalamic m1mAChR protein expression in HFD-O at P28 and P56 compared with SC-O ( Figures 3C, D ). Considering the higher hypothalamic expression of m1mAChR, we evaluated he distribution of m1mAChR expression with immunofluorescence at P28 ( Figure 3E ). There were more m1mAChR⁺ cells in the median eminence of HFD-O compared with SC-O ( Figure 3E ). Nevertheless, m1mAChR expression seems to be higher in the arcuate nucleus compared with the median eminence of SC-O ( Figure 3E ).

To assess the impact of central m1mAChR activation on liver signalling pathways and α7nAChR expression in offspring at P28, we administered McN-A-343 (ICV), a pharmacological agonist of m1mAChR ( Figure 4A ). Figures 4B, C indicate that hypothalamic activation of m1mAChR increased α7nAChR and phosphorylated STAT3 (pSTAT3) protein expression in the liver of SC-O and HFD-O. Notably, pSTAT3 expression was higher in the liver of HFD-O than SC-O. Additionally, m1mAChR activation was accompanied by a reduction in phosphorylated JNK (pJNK) expression in the liver ( Figures 4B, C ).

We also evaluated liver cytokine mRNA levels. Il1b expression was not different between the groups, indicating similar levels of this cytokine. Interestingly, Tnf expression in HFD-O liver was lower compared with SC-O liver. However, treatment with the m1mAChR agonist (McN-A-343) via ICV administration reduced Tnf mRNA expression in SC-O liver, but there was no additional effect in HFD-O liver ( Figure 4D ). Il10 mRNA expression increased in HFD-O liver compared with SC-O liver, but ICV administration of McN-A-343 significantly increased Il10 mRNA expression in SC-O liver compared with HFD-O liver. Furthermore, McN-A-343 delivered via ICV administration demonstrated a greater inhibitory effect on liver Il6 mRNA expression in HFD-O compared with SC-O. Similarly, the mRNA levels of Socs3, an important regulatory molecule in inflammation, were higher in HFD-O liver ( Figure 4D ). This indicates that maternal HFD consumption may lead to increased SOCS3 expression in HFD-O liver, potentially impacting the regulation of inflammatory responses.

As shown in the Figure 4E , we investigated the hepatic inflammatory response in SC-O and HFD-O after LPS treatment. We simultaneously administered SC-O and HFD-O LPS (1 mg kg^-1, IP) and an m1mAChR agonist. Liver pSTAT3, a component of the anti-inflammatory pathway, was increased in all LPS-challenged mice. This suggests activation of the anti-inflammatory response in both SC-O and HFD-O following LPS treatment. Moreover, phosphorylated NF-κB (pNF-κB) and pJNK expression was decreased in the liver of LPS-treated HFD-O compared with LPS-treated SC-O. This indicates that maternal HFD consumption may have modulated the hepatic inflammatory response in the offspring, resulting in reduced activation of these pro-inflammatory signalling pathways. Interestingly, when we administered LPS-treated HFD-O with an m1mAChR agonist, there was a further decrease in pNF-κB and pJNK expression in the liver ( Figures 4F, G ). This suggests that activation of m1mAChR can enhance the anti-inflammatory response and attenuate the activation of pro-inflammatory signaling pathways in the liver of HFD-O following LPS challenge, as depicted in Figure 4H . These findings indicate that maternal HFD consumption and m1mAChR activation can influence the hepatic inflammatory response in offspring, potentially leading to a more pronounced anti-inflammatory state and reduced activation of pro-inflammatory pathways in HFD-O following LPS treatment.

The inflammatory and immune response to pathogens depends on the spleen. We investigated the levels of inflammatory cytokines in the spleen of SC-O and HFD-O following ICV treatment with the m1mAChR agonist. However, there were no significant differences in the inflammatory markers between the groups (data not shown).

Next, we performed flow cytometry using macrophage and lymphocyte panels to evaluate the inflammatory response of the spleen. In the macrophage panel, there was a decrease in CD45⁺CD11c⁺ cells in HFD-O compared with SC-O, and an increase in CD45⁺CD206⁺ cells in HFD-O ( Figure 5A ). These findings were supported by the differential expression of macrophage markers in the spleen. Similarly, Nos2 mRNA expression was decreased in HFD-O compared with SC-O ( Figure 5C ), whereas Arg1 mRNA expression appeared to be increased in HFD-O ( Figure 5C ). Furthermore, treatment with the m1mAChR agonist decreased Nos2 mRNA expression and increased Arg1 expression only in SC-O after LPS challenge ( Figure 5D ). Taken together, these observations confirm that the macrophages present in the spleen exhibit an anti-inflammatory profile.

The lymphocyte panel revealed an increase in CD3⁺CD4⁺ cells in the spleen of HFD-O compared with SC-O, and a reduction in CD3⁺CD8⁺ cells in HFD-O ( Figure 5B ). These findings were accompanied by a decrease in Il17 and Tgfb mRNA expression in the spleen of HFD-O compared with SC-O ( Figure 5C ). To investigate the increase in T-helper lymphocytes, we determined the specific type of T-helper lymphocytes present in the spleen of HFD-O after LPS challenge and treatment with an m1mAChR agonist. Interestingly, we observed a reduction in Il17 mRNA expression in the presence of LPS ( Figure 5D ), while Il22 mRNA expression appeared to be increased in HFD-O ( Figure 5C ). These observations highlight the presence of distinct lymphocyte profiles in HFD-O. Additionally, Ifng mRNA expression was decreased in HFD-O following LPS challenge, and the m1mAChR agonist reduced Ifng mRNA expression in LPS-treated SC-O ( Figure 5D ). Overall, these findings suggest reduced differentiation of lymphocytes towards the Th17 profile in HFD-O in the basal state and after LPS challenge.

To investigate the relationship between central muscarinic receptor and macrophage activation, we isolated bone marrow cells from the offspring. We cultured these cells and after 7 days of differentiation, we submitted them for further analysis. We counted the number of cells with a Neubauer chamber immediately after isolation ( Figure 6A ). There were fewer cells isolated from HFD-O compared with SC-O ( Figure 6B ). Additionally, a representative image shows that after 7 days of differentiation, the number of macrophages appears to be reduced in HFD-O compared with SC-O ( Figure 6C ). However, when evaluating the macrophage panel with flow cytometry, there were no significant differences between the groups in terms of the number of positive cells for either inflammatory or anti-inflammatory markers ( Figure 6D ).

Next, we evaluated the impact of central m1mAChR activation with McN-A-343 on the macrophage profile of isolated bone marrow cells ( Figure 7 ). We noted a decrease in IL1-β protein expression in the culture medium in both groups following treatment with the m1mAChR agonist ( Figure 7A ). We also assessed Tnf, Il1b, Il10, Il6, Nos2, and Arg1 mRNA expression ( Figure 7B ). Specifically, Il1b, Il6, and Nos2 mRNA expression was significantly reduced upon administration of the agonist McN-A-343. Conversely, McN-A-343 administration increased Il10 mRNA expression. Interestingly, Il10 mRNA expression increased in both groups upon agonist treatment. Furthermore, there was decreased Tnf and Il6 mRNA in HFD-O compared with SC-O. Nos2 mRNA expression, indicative of the inflammatory profile, was decreased in HFD-O, and the agonist treatment further decreased Nos2 mRNA expression in both groups. Conversely, Arg1 mRNA expression, a marker of the anti-inflammatory profile, was increased in HFD-O ( Figure 7B ). Notably, phosphorylation of STAT3, a protein downstream in the cholinergic anti-inflammatory pathway, was increased in both groups following treatment with the m1mAChR agonist ( Figures 7C, D ).