α-Ketoglutaric acid sodium salt (Cat# K1875), ACh chloride (Cat# A6625), apyrase (Cat# A6410), adenosine 5′-triphosphate disodium salt hydrate (Cat# A2383), bovine serum albumin (BSA, Cat# A9418), choline chloride (Cat# C7017), CRP (Cat# AG723-M), dimethyl sulfoxide (DMSO, Cat# D2650), glutamate-pyruvate transaminase (GPT, Cat# 10737127001), glutamic-oxalacetic transaminase (GOT, Cat# 10105554001), glutaminase grade V (Cat# G880), hydroxylamin hydrochloride (Cat# 159417), L-glutamic dehydrogenase (GLDH, Cat# G2626), L-lactate dehydrogenase (LDH, Cat# 10107085001), L-malate dehydrogenase (L-MDH, Cat# LMDH-RO), LPS (E. coli O111:B4, Cat# L2630; E. coli O26:B6, Cat# L2654), recombinant human macrophage colony-stimulating factor (M-CSF, Cat# SRP3110), mecamylamine hydrochloride (Cat# M9020), (meta)periodate (Cat# S1878), nicotine hydrogen tartrate salt (Cat# N5260), nigericin sodium salt (Cat# N7143), phorbol 12-myristate 13-acetate (PMA, Cat# P1585), PC chloride calcium salt tetrahydrate (Cat# P0378), potassium hydroxide (KOH, Cat# 1.05033), and recombinant mouse interferon-γ (INF-γ, Cat# IF005) were purchased from Merck (Darmstadt, Germany). Gibco penicillin-streptomycin solution (Cat# 151401222) and L-glutamine solution were purchased from Thermo Fisher Scientific (Dreieich, Germany). Glycine (Cat# 3908.2), glycerol (Cat# 3783.2), hydrazinium sulphate (Cat# HN96.2), hydrochloric acid (HCl, Cat# P074.4), L-alanine (Cat# 3076.1), L-aspartic acid (Cat# 1690.1), L-glutamic acid (Cat# 1743.1), L-glutamine (Cat# HN08.1), ortho-phosphoric acid (H₃PO₄, Cat# 6366.1), pyruvic acid sodium salt (Cat# 8793.1), sodium hydroxide (NaOH, Cat# 6771.2), sodium L-serine (Cat# 1714.1) and TRIS (Cat# 4855.2) were from Carl Roth, Karlsruhe, Germany. Glucose hexokinase FS (Cat# 125119910920), TruCal U (Cat# 591009910064), TruLab N (Cat# 590009910061), TruLab P (Cat# 590509910061), Cleaner (Cat# 188309910885), Cleaner A (Cat# 186109910923), Cleaner B (Cat# 186509910923) were purchased from Diasys, Flacht, Germany. Acetic acid (Cat# 0820), β-nicotine amide adenine dinucleotide (NAD, Cat# A1124), ß-nicotine amide adenine dinucleotide disodium salt (NADH, Cat# A1393), di-potassium hydrogen phosphate anhydrous (Cat# A1042), potassium di-hydrogen phosphate (Cat# A1043), and sodium acetate anhydrous (Cat# A1522), were purchased from AppliChem GmbH, Darmstadt, Germany. Adenosine 5′-diphosphate (ADP, Cat# BD18698) was obtained from BLDPharm, Reinbeck, Germany. BzATP (2’(3’)-O-(4-benzoyl-benzoyl)ATP trieethylammonium salt) was purchased from Jena Bioscience (Jena, Germany), 0.5 M pyrogen-free ethylenediaminetetraacetic acid (EDTA) solution from bioWORLD (Dublin, OH, United States, Cat# 40520000), recombinant human INF-γ from R&D Systems (Minneapolis, MN, United States; Cat# 285-IF-100), and PNU-282987 (N-(3R)-1-azabicyclo[2.2.2]oct-3-yl-4-chlorobenzamide, Cat# 2303) from Tocris (Bio-Techne GmbH, Wiesbaden-Nordenstadt, Germany). IL-4 (Cat#200-04) and IL-13 (Cat#200-13) were purchased from Preprotec, Thermo Fisher Scientific, Waltham, MA USA. PMA and PNU-282987 were dissolved in DMSO. Nigericin was dissolved in ethanol (EtOH). When appropriate, control experiments were performed with the corresponding concentrations of DMSO/EtOH without drugs. The conopeptides [V11L;V16D]ArIB (specifically antagonizing α7 nACh) (40–42) and RgIA4 (antagonizing nAChRs composed of subunits α9 and α10) (36, 43, 44) were produced and characterized as described previously (44, 45).

THP-1 cells (German Collection of Microorganisms and Cell Cultures, Cat# ACC16) were cultured for a maximum of 20 passages under 5% CO₂ atmosphere at 37°C using RPMI 1640 medium from Sigma (Merck, Cat# R8758) supplemented with 10% fetal bovine serum (FBS) from Capricorn (Ebsdorfergrund, Germany, Cat# FBS-16A). For IL-1β release experiments, monocytic THP-1 cells were spun down (500 g, 8 min, room temperature (RT)) and the cell pellet was resuspended in FBS-free RPMI medium. 0.5 × 10⁶ cells/0.5 ml/well were seeded into 48-well plates (Greiner Bio-One Frickenhausen, Germany). Cells were left untreated or primed for 5 h with 1 µg/ml LPS (E. coli O26:B6) (46–48). After priming, BzATP (100 µM) (49, 50) or nigericin (50 µM; added together with 0.5 U/ml apyrase) (51) was added for 40 min in the presence or absence of cholinergic agonists and antagonists. At the end of the experiments, cells were spun down (500 g, 8 min, 4°C) and the cell-free supernatants were collected and stored at −20°C for later cytokine measurements.

For generating THP-1 cell-derived macrophages, either 0.3 × 10⁶ cells/ml and per well were seeded in 12-well plates (Greiner) or 6 x 10⁶ cells were seeded in T75 cell culture flasks (Sarstedt, Nümbrecht, Germany) and incubated with 50 nM PMA for 24 h. Thereafter, a 24 h resting phase in fresh complete medium without PMA followed. To induce cell differentiation into M1-like macrophages, cells were cultured in fresh complete medium supplemented with 10 ng/ml human IFN-γ and 10 ng/ml LPS (E. coli O111:B4) for 48 h. Differentiation into M2-like macrophages was induced by culturing the cells in fresh complete medium supplemented with 20 ng/ml IL-4 and 20 ng/ml IL-13 for 48 h. M0-like macrophages were differentiated with PMA in parallel to M1-like and M2-like macrophages followed by a 72 h resting phase in fresh complete medium without PMA.

To investigate IL-1β release on day 5, the medium was replaced by fresh RPMI 1640 medium (Sigma) supplemented with 10% FBS. The cells were stimulated with 1 µg/ml LPS (E. coli O26:B6) for 5 h and handled as described for monocytic THP-1 cells. The identity of M0-like, M1-like and M2-like macrophages on day 5 was evaluated by flow cytometry (see below). Cells were washed once with Dulbecco´s phosphate buffered saline without calcium and magnesium (PBS; Merck Cat# D8537) and detached using TrypLE™ Express (Thermo Fisher Scientific, Cat# 12605010) according to the manufacturer’s protocol. The cell number was determined, and cells were stored on ice until flow cytometry.

Human PBMC were obtained from self-reported healthy non-smoking adult volunteers (male and female). All procedures were approved by the local ethics committee at the University of Giessen (approval No. 90/18 and No. 98/23). Blood was drawn into sterile syringes containing EDTA (final concentration 1 mM). PBMCs were separated on Leucosep gradients (Greiner Bio-One, Cat# 227261). Isolated PBMCs were washed twice with PBS + 1% BSA and finally resuspended in monocyte attachment medium (PromoCell, Heidelberg, Germany, Cat# C-28051). 0.5 x 10⁶ cells per well were seeded in a 48-well plate and cultured for 2 h. Non-adherent cells were removed, and cell culture medium was replaced by RPMI 1640-medium (Sigma) + 10% FBS (Capricorn) + 50 U penicillin/ml + 50 μg streptomycin/ml + 10 ng/ml M-CSF. The cells were cultured at 37°C, 5% CO₂. After 72 h, the medium was replaced by fresh RPMI 1640-Medium (Sigma) + 10% FBS (Capricorn) + 50 U penicillin/ml + 50 μg streptomycin/ml + 10 ng/ml M-CSF to obtain a M0-like phenotype. To obtain a M1-like phenotype, 50 pg/ml LPS (E. coli 026:B6) + 20 ng/ml IFN-γ were added, or 40 ng/ml IL-4 for a M2-like phenotype and cultured for a further incubation time of 72 h. On day 6 of differentiation, cells were washed with and cultured in fresh RPMI 1640 medium + 10% FBS and used for IL-1β release experiments as described above. For flow cytometry (see below), differentiated cells were washed once with PBS (Merck Cat# D8537) and detached using TrypLE™ Express (Thermo Fisher Scientific, Cat# 12605010) according to the manufacturer’s protocol. The cell number was determined, and cells were stored on ice until flow cytometry.

For the isolation of peritoneal macrophages, peritoneal dialysate from female and male patients (above the age of 18 years) admitted to the University hospital Giessen was used. All experiments were approved by the local ethics committee at the University of Giessen, Germany (approval No. AZ251/20). The privacy rights of the patients have been observed and written informed consent was given by each patient. Peritoneal dialysates were collected during planned peritoneal dialysis training 14 days after implantation of the peritoneal dialysis catheter required due to chronic renal insufficiency requiring dialysis. Exclusion criteria were chronic or acute infectious diseases, increased inflammatory parameters (leukocytosis, CRP above 10 mg/l, procalcitonin levels above 0.5 ng/ml, fever), ongoing antibiotic therapy, immunosuppressive therapy, chemotherapy within the last 3 months, pregnancy, or lack of consent.

Dialysates were stored on ice and all isolation procedures were performed on ice or at 4°C. Cells were isolated by centrifugation at 200 x g, 20 min. The cell pellets were washed with cold PBS/0.1% BSA.

Contamination with erythrocytes were removed by using RBC lysis buffer (BioLegend, San Diego CA USA, Cat# 420301) according to the manufacturer’s protocol. Isolated cells were resuspended in RPMI medium (Merck) plus 10% FBS (Capricorn) and 50 U penicillin/ml, 50 µg streptomycin/ml.

For cytokine release experiments, 1 x 10⁶ cells per ml were seeded per well in a 12-well plate and cultured overnight at 37°C and 5% CO₂. On the next day, macrophages were enriched by adherence selection. Peritoneal macrophages were left untreated or primed for 5 h with 0.1 or 1 µg/ml LPS (E. coli O26:B6). After priming, BzATP (100 µM) was added for 40 min in the presence or absence of cholinergic agonists and antagonists. At the end of the experiments, cells were spun down (500 x g, 8 min, 4°C) and the cell-free supernatants were collected and stored at −20°C for later cytokine measurements.

To analyze the cell types present in the peritoneal dialysates before and after adherence selection, cytospots were generated using the Cellspin I centrifuge (28 x g, 4 min, RT). Cytospots were air dried and fixed using BD Cytoperm™ for 20 min on ice. Moreover, 0.1 x 10⁶ cells were seeded in 10-well chamber-view slides (Greiner, Cat# 543079) and cultured overnight. On the next day, the RPMI medium was discarded, and cells were fixed using BD Cytoperm™ as described above. For immunocytochemical staining, cytospots and chamber-view slides were incubated with PBS plus 1% hydrogen peroxide on ice for 30 min, washed three times with PBS, and incubated with BD Perm Wash™ plus 1% BSA for 30 min at RT. Samples were stained with murine monoclonal antibodies to human CD14 (BioLegend, Cat# 325602) and CD68 (Agilent Dako, Cat# GA613) for 1 h followed by 1 h incubation with horseradish-peroxidase (HRP)-conjugated anti-mouse Ig antibodies (from Agilent Dako, Cat.# P0260 and Cat.# K400311-2). Thereafter, all samples were washed three times with TBS-buffer (0.5 M TRIS, 1.54 mM NaCl, 5% (v/v) HCl; pH 7.6) followed by incubation with 3,3’-diaminobenzidine tetrahydrochloride (DAB; Merck, Cat# D5905) in PBS/0.015% hydrogen peroxide (0.5 mg/ml) for 10 min at RT. At the end, samples were stained with Mayer’s Haematoxylin for 6 min and then cover-slipped with Glycergel mounting medium (Agilent Dako, Cat# C0563). Stained cells of 6–9 randomly selected areas per sample were counted using the microscope Leica DMLS (20 x objective) and the ZEN (blue edition) software (Zeiss). Cells were categorized in CD14⁺, CD14^-, CD68⁺, and CD68^- cells. Granulocytes were identified by their endogenous myeloperoxidase activity resulting in dark-brown stained granules and their typical polymorphic nuclei. After counting, percentages of CD14⁺ and CD68⁺ macrophages before and after adherence selection were calculated and compared.

Total RNA was isolated from monocytic THP-1 cells, THP-1 cell-derived M0- and M1-like macrophages, and peritoneal macrophages using the RNeasy Plus Mini Kit (Qiagen, Hilden, Germany Cat#74134) in combination with the RNase-Free DNase Set (Qiagen; Cat#79256) according to the manufacturer’s instructions. The quantity and quality of the RNA were assessed using the NanoDrop2000 (Thermo Fisher Scientific). Ratios of absorbance at 260/280 (acceptable value range: 1.8 – 2.0) and 260/230 (acceptable value range: 2.0 – 2.2) were determined to confirm that all samples were suitable for further analysis. The obtained RNA (1 µg) was reversely transcribed using QuantiTect^® Reverse Transcription Kit Qiagen; Cat# 205311) according to the manufacturer’s instructions. Universal Human Reference RNA (Thermo Fisher Scientific; Cat# QS0639) was included as a positive control. Thereafter, real-time RT-PCR was performed (n = 5–6 biological replicates per experimental group, each sample was assessed in technical duplicates) using the Real-Time PCR Cycler StepOne Plus^® (Applied Biosystems, Carlsbad, CA, USA) and SsoAdvanced Universal SYBR Green Supermix (Bio Rad, Hercules, CA, USA). Primers were obtained from Thermo Fisher Scientific ( Table 1 ). As a negative control, samples were run under the same conditions, using nuclease-free water instead of cDNA. For each cDNA sample and each primer pair, duplicate reactions were performed. The PCR protocol consisted of an initial activation of the polymerase at 95°C for 5 min, followed by 40 cycles of a denaturation step at 95°C for 5 s and an annealing step at 61°C for 10 s and extension step at 72°C for 30 s, followed by a melting curve analysis. The mRNA expression is given as cycle threshold (Ct) values. For mRNAs that were undetectable in these experiments, the Ct value was artificially set to the cut of value Ct = 40 and this value was included in further analyses. The mean Ct value of technical duplicates was further used to calculate the fold change gene expression level based on the 2^-ΔΔCt method (52) to compare the expression of the genes of interest (CHRNA7, CHRNA9, CHRNA10, CHRFAM7A; see Table 1 ) after normalization by geometric averaging of the three reference genes glyceraldehyde-3-phosphate dehydrogenase (GAPDH), peptidylprolyl isomerase A (PPIA), and ribosomal protein L37a (RPL37A) (53–57).

The specificity of the RT-PCR was confirmed by separation of the PCR products in a 1.5% agarose gel and sequencing (Eurofins Genomics, Ebersberg, Germany). The sequencing results were then analyzed using the public interface of BLAST, http://www.ncbi.nlm.nih.gov/blast, at the NCBI website at the NCBI website.

To assess the morphology of THP-1 cell-derived macrophages, 0.1 x 10⁶ monocytic cells/0.5 ml RPMI 1640 medium (Merck) supplemented with 10% FBS (Capricorn) were cultured in 8-well cell culture chambers (Sarstedt, Cat# 94.6170.802). THP-1 cell-derived M0-like, M1-like and M2-like macrophages were differentiated as described above. On day 5, cells were washed once with PBS, followed by fixation with BD Cytoperm™ (BD, Franklin Lakes, NJ, USA, Cat# 554722) for 20 min on ice. Thereafter, cells were washed with PBS and chambers were air-dried. Macrophages were stained with May-Grünwald’s dye for 1–2 min, then incubated in double distilled water (ddH₂O) for 5 min, followed by Giemsa’s staining for 4–10 min. After the staining, the chamber frames were removed, slides cleaned thoroughly with ddH₂O, dried and cover-slipped with Neo-Mount^® (Merck, Cat# 1.09016) before microscopy (Leica DMLS, Leica Microsystems).

Flow cytometry was performed to evaluate the presence of cell surface markers of THP-1 cell-derived macrophages (M0-, M1- and M2-like) and hPBMC-derived macrophages using a panel of antibodies against cluster of differentiation (CD)38, CD80, CD83, CD86, CD163, CD206, CD209 and HLA-DR (see Supplementary Table S1 ). Macrophages were detached using TrypLE (see above) and resuspended in flow cytometry buffer (PBS, 2 mM EDTA, 5% FBS). Thereafter, 1 - 2 × 10⁵ cells were incubated with appropriate antibodies for 30 min on ice. All antibodies ( Supplementary Table S1 ) were used at the manufacturer’s recommended dilution or titrated to determine the optimal staining concentrations (between 1:20 and 1:100). Cells were also incubated with the control isotype antibody corresponding to each primary antibody. Thereafter, cells were washed twice in flow cytometry buffer. Then, the cells were fixed with 1% freshly dissociated paraformaldehyde (PFA) before flow cytometry analysis. Cells were analyzed on a BD FACSVerse™ System (BD Biosciences) using the BD FACSuite™ software, recording at least 10,000 events for each sample. Forward scatter (FSC) files were exported and analyzed in FlowJo software version 10.8.1 (FlowJo, LLC). First, macrophages were gated by FSC–area (FSC-A) and side scatter–area (SSC-A), followed by single cell gating by SSC-A and side scatter-width (SSC-W). Surface marker-positive cells were gated based on Fluorescence Minus One (FMO) or isotype controls and percentage of surface markers positive cells was measured and exported.

5 x 10⁵ monocytic THP-1 cells were seeded in 12-well plates and divided into 4 groups. The monocytic THP-1 cell group was cultivated for 5 days without differentiation supplements. The three other cell groups were treated with differentiation factors as described above to yield M0-, M1- and M2-like macrophages. On day 5 of cultivation, the monocytic THP-1 cells were centrifuged and resuspended in fresh medium while in the wells of the adherent growing M0-, M1- and M2-like macrophages the medium was exchanged to fresh medium without supplementation of differentiation factors. After 6 h the supernatants of the individual wells were collected and stored at -80°C. In parallel, the number of cells were counted in the corresponding wells. The frozen medium supernatants were heated for 15 min at 95°C and subsequently centrifuged at 8000 x g for 5 min. Glucose, lactate, glutamine, glutamate, pyruvate, aspartate, alanine and serine concentrations were determined photometrically using a Respons^® 920 bench top analyzer (DiaSys Diagnostic Systems GmbH, Holzheim, Germany), as previously described (58). The conversion rates of the metabolites were calculated in [nmol/(h*10⁵ cells)] relative to the respective metabolite concentrations in control medium samples without cells which were cultivated for 6 hours in parallel to the wells with cells. Medium used for the metabolic conversion rates: Dulbecco’s modified Eagle’s medium (DMEM) without glucose, glutamine and phenol red (Gibco™, Thermo Fisher Scientific, Cat# A1443001) supplemented with 10% FBS Xtra (Capricorn, Cat# FBS-16A), 11 mM D-glucose (Carl Roth, Karlsruhe, Germany, Cat# X997.1), 2 mM L-glutamine (Merck, Cat# G3126), 50 U/ml penicillin, and 50 μg/ml streptomycin (Gibco™, Thermo Fisher Scientific, Cat# 151401222).

Cytokine concentrations of IL-1β, IL-1α and IL-18 were measured in the above-described cell-free supernatants using the human IL-1 beta/IL-1F2 DuoSet enzyme-linked immunosorbent assay (ELISA) from R&D Systems (Cat# DY201), human IL-1 alpha/IL-1F1 DuoSet ELISA (Cat# DY200-05), and human Total IL-18 DuoSet ELISA (Cat# DY318-05) according to the supplier’s instructions.

To measure LDH activity, the CytoTox 96^® Non-Radioactive Cytotoxicity Assay (Promega, Madison, WI, United States; Cat# G1780) was performed according to the manufacturer’s instructions. LDH activities determined in cell-free supernatants are given as percentage of 100% cell lysis control.

With exception of the data obtained in metabolic conversion rate measurements, all data were analyzed using SPSS^® (Version 29, IBM^®, Armonk, NY, USA). Multiple independent data sets were first analyzed by the non-parametric Kruskal–Wallis test. In case of a p ≤ 0.05, the non-parametric Mann–Whitney U test was performed and again, a p ≤ 0.05 was considered as evidence for statistical significance. Dependent data sets were analyzed first by the Friedman test followed by the Wilcoxon signed-rank test.

All numbers (n) of the individual experiments (biological replicates) are indicated in the Results section and in the Figures. No outliers were excluded from the analyses. The free and open-source software Inkscape version 0.48.5 r10040 (licensed under the GPL) was used for visualization of the data.

Data obtained in metabolic conversion rate measurements are presented as mean ± standard deviation (SD) using GraphPad Prism^® (version 10.2.3, GraphPad Software, Boston, Massachusetts, USA). Data analysis was performed using SAS^® statistical software (version 9.4, Statistical Analysis System Institute, Cary, North Carolina, USA). A two-way analysis of variance (ANOVA) was conducted to quantify metabolic conversion rates of the mean values of 5 biological replicates of glucose, pyruvate, lactate, glutamine, glutamate, aspartate, alanine and serine, including interaction effects between monocytic THP-1 cells, THP-1 cell-derived M0-, M1- and M2-like macrophages. The normality of residuals was assessed to validate the model assumptions. Post-hoc pairwise comparisons were performed and adjusted for multiple testing using the Bonferroni correction.

THP-1 cells develop a macrophage-like phenotype resembling primary human macrophages when exposed to PMA (59). A PubMed search in August 2025 using the search terms “THP-1” “macrophage” “differentiation” resulted in 1,211 hits. The published differentiation and characterization protocols of THP-1 cell-derived macrophages, however, vary largely between these publications and thus, may unknowingly affect the relevance of research output across research groups (60). In this study, THP-1 cell-derived macrophages were used to investigate the cholinergic control of the BzATP-mediated release of IL-1β. To test the efficiency of the differentiation protocols used, first the morphology of stained macrophages was assessed ( Figure 1A ). M0-like macrophages depicted a rounded or a spindle-shaped morphology, whereas a fibroblast-like, spindle-shaped morphology dominated in M1- and M2-like macrophages ( Figure 1A ). Moreover, M0-like macrophages appeared to be smaller compared to M1-like macrophages and THP-1 cell-derived M1-like macrophages possessed amoeboid protrusions.

Flow cytometry was performed on differentiated THP-1 cells to assess cell surface markers typical for polarized macrophages (59, 61–63). A panel of antibodies was used against the following markers: the pro-inflammatory phenotype markers CD38 (64, 65), CD80, CD83 (66, 67) and CD86 (68, 69), the anti-inflammatory phenotype markers CD163 (61, 70) CD206 and CD209 (71, 72), and the human leukocyte antigen DR (HLA-DR) which is known to be abundantly expressed on human monocytes and macrophages (73). In comparison to THP-1 cell-derived M0-like macrophages, the M1-like differentiation protocol ( Figure 1 ) resulted in an upregulation of CD38, CD80, and CD86 (p ≤ 0.05; n = 5 each; Figure 1B,C ). The typical cell surface markers of M2-like macrophages CD206 and CD209 were increased in the THP-1 cell-derived M2-like macrophages when compared to the M0-like (p ≤ 0.05; n = 5 each; Figures 1B, C ). Of note, CD83 was weakly expressed by all three macrophage populations and, thus, not useful to distinguish between the M0-, M1- and M2-like macrophages (n = 5; data not shown). In addition, the known M2-like macrophage surface marker CD163 was not upregulated by the used differentiation protocol for M2-like macrophages (n = 5; data not shown).

To further characterize the THP-1 cell-derived macrophages, metabolic conversion rates were analyzed ( Figure 2 ). Even if cell passage-dependent minimal variations were detected within the 5 different cell batches used ( Supplementary Figure S1 ), a direct comparison of the general metabolic conversion rates revealed phenotype-specific differences. In M1-like macrophages glucose consumption was 4-fold higher in comparison to monocytic THP-1 cells and 2-fold higher in comparison to M0- and M2-like macrophages. Lactate production was 3-fold higher in M1-like macrophages in comparison to monocytic THP-1 cells, M0-like as well as M2-like macrophages (p ≤ 0.05; n = 5; Figures 2A-C ). The correlation analysis of glucose consumption and lactate production showed that in M1-like macrophages at least 25% of the glucose consumed was converted to lactate, while in monocytic THP-1 cells, M0-like and M2-like macrophages the conversion of glucose to lactate was not of great importance ( Supplementary Figure S2 ; Supplementary Table S2 ). Aspartate conversion changed from conversion rates close to the detection limit in monocytic THP-1 cells and spread conversion between consumption and production in M2-like macrophages to pronounced consumption in M1-like macrophages which was in addition 5-fold higher in comparison to M0-like macrophages (p ≤ 0.05; n = 5; Figures 2A, G ). In terms of serine conversion, M1- and M2-like macrophages were characterized by 5-fold higher serine consumption rates compared to monocytic THP-1 cells (p ≤ 0.05; n = 5; Figures 2A, I ). Glutamine consumption and glutamate production were higher in all THP-1 cell-derived macrophages compared to monocytic THP-1 cells. However, only the difference in glutamate production between M2-like macrophages and monocytic THP-1 cells reached a significant level (p ≤ 0.05; n = 5; Figures 2A, D, E ). In M0- and M2-like macrophages, strong positive correlations between glutamine consumption and both lactate and glutamate production indicate a high glutaminolytic capacity with glutamate being released as part of the glutamine, whereas M1-like macrophages show inverse correlations ( Supplementary Table S2 ). Alanine was consumed in M0- and M2-like macrophages while in monocytic THP-1 cells and M1-like macrophages alanine conversion spread between consumption and production ( Figures 2A, H ).

Next, the release of IL-1β by LPS-primed THP-1 cell-derived M0-, M1- and M2-like macrophages in response to BzATP (100 µM), ATP (2 mM) and nigericin (50 µM) were compared ( Supplementary Figure S3 ). Phenotype-specific responses were revealed. LPS-primed M0-like macrophages were found to release IL-1β in response to BzATP from 71 to 282 pg/ml, ATP from 188 to 492 pg/ml, and nigericin from 20 to 1081 pg/ml (n = 6; p ≤ 0.05; Supplementary Figure S3A ). LPS-primed M1-like macrophages were found to release the highest amount of IL-1β in response to BzATP (478 to 2351 pg/ml), ATP (577 to 1936 pg/ml) and nigericin (96 to 482 pg/ml; n = 6; p ≤ 0.05; Supplementary Figure S3B ). No BzATP- and ATP-mediated IL-1β release was detected by LPS-primed THP-1 cell-derived M2-like macrophages (n = 7 – 11; Supplementary Figure S3C ). In response to nigericin, however, LPS-primed THP-1 cell-derived M2-like macrophages released significant amounts of IL-1β ranging from 4 to 181 pg/ml (n = 10; p ≤ 0.05; Supplementary Figure S3C ). Without LPS-priming, the cells did not release IL-1β in response to nigericin (n = 4; Supplementary Figure S3C ).

First, transcript levels of human CHRNA7, CHRNA9 and CHRNA10 were determined in monocytic THP-1 cells (n = 6), THP-1 cell-derived M0- and M1-like macrophages (n = 6 each) and human peritoneal macrophages (n = 5). Moreover, the expression of the Dupα7 nAChR (CHRFAM7A), known as a dominant negative regulator of α7 nAChR functions, was analyzed. The cells were left untreated or primed with LPS for 5 h. The specificity of all primers used was validated by agarose gel electrophoresis ( Supplementary Figure S4 ) and sequencing the PCR products. Transcripts of all genes of interest (CHRNA7, CHRNA9, CHRNA10 and CHRFAM7A) were obtained ( Supplementary Figure S4). The Ct values of all nAChR subunit genes of interest were above 30 ( Supplementary Table S3 ; Supplementary Table S4 ). CHRNA9 was often undetected (Ct ≥ 40), for example in three out of five biological human peritoneal macrophages samples ( Supplementary Table S4 ).

The relative expression of the genes of interest in LPS-primed THP-1 cells (monocytic, M0-like and M1-like) were further calculated by normalization to the three reference genes (GAPDH, PPIA, RPL37A) via the 2^-ΔΔCt method ( Figure 3 ). Apart from CHRFAM7A in THP-1 cell-derived M0-like macrophages (n = 6; p ≤ 0.05), no differences between the genes of interest in untreated and LPS-primed cells were found ( Figure 3 ).

To test if classical and unconventional nAChR agonists inhibit the ATP-induced IL-1β release, human monocytic THP-1 cells were primed for 5 h with LPS (1 µg/ml) ( Figure 4 ). Thereafter, 100 µM BzATP, a P2RX7 agonist (74), was applied for 40 min in presence and absence of the cholinergic agonists (ACh, choline, nicotine, PC, CRP). To test for the involvement of nAChRs, the antagonist conopeptides [V11L;V16D]ArIB (antagonizing α7 nAChRs (40–42)) and RgIA4 (antagonizing nAChRs containing subunits α9 and/or α10 (36, 43, 44)) were used. IL-1β concentrations were measured in cell culture supernatants. As expected (46, 47), priming with LPS alone did not induce the release of relevant amounts of IL-1β by monocytic THP-1 cells (untreated 1.8 ± 3.4 pg/ml; LPS 5.4 ± 10.2 pg/ml; n = 34). Stimulation of LPS-primed cells with BzATP resulted in elevated IL-1β levels in supernatants of monocytic THP-1 cells in the range of 15 to 156 pg/ml (n = 34). The classical nAChR agonists ACh ( Figure 4A ; n = 6), choline and nicotine ( Figure 4B ; n = 5) significantly (p ≤ 0.05) inhibited the BzATP-induced release of IL-1β. Similar results were found for the unconventional endogenous agonists PC ( Figure 4C ; n = 7) and CRP in concentrations of 10 and 20 µg/ml ( Figure 4D ; n = 6 – 9). The viability of the cells was not affected by the agonists as measured by LDH release ( Supplementary Table S5 ). The inhibitory effect of all tested nAChR agonists was reverted by the conopeptides [V11L; V16D]ArIB and RgIA4, suggesting that nAChR subunits α7 and α9* are involved in the cholinergic signaling ( Figure 4 ).

Next, we tested if the BzATP-induced release of IL-1β by LPS-primed THP-1 cell-derived M0-like and M1-like macrophages is controlled by nAChR agonists ( Figure 5 ). Stimulation of LPS-primed THP-1 M0-like macrophages with BzATP resulted in elevated IL-1β levels ranging from 75 to 397 pg/ml ( Figure 5A ; n = 6), whereas the BzATP-induced release of IL-1β by M1-like macrophages was in the range of 110 to 1732.5 pg/ml ( Figures 5B–D ; n = 17). Similar to experiments with monocytic THP-1 cells ( Figure 4 ), the classical nAChR agonists ACh, choline and nicotine as well as CRP significantly (p ≤ 0.05) inhibited the BzATP-induced release of IL-1β by both M0-like and M1-like macrophages ( Figure 5 ). The inhibitory effect of PC on the BzATP-induced release of IL-1β by THP-1 cell-derived M1-like macrophages has been shown before (46). Again, the inhibitory effect of all tested nAChR agonists was reverted by mecamylamine ( Figure 5B ; n =5) and the conopeptides [V11L; V16D]ArIB and RgIA4 ( Figures 5C, D ; n = 5 – 6), suggesting the involvement of nAChR subunits α7 and α9* ( Figure 5 ). When using the selective α7 nAChR agonist PNU-282987 (75, 76), the same results were observed. PNU-282987 inhibited the BzATP-induced release of IL-1β by monocytic THP-1 cells and THP-1 cell-derived M1-like macrophages, and the inhibitory effect was reverted by the conopeptides [V11L; V16D]ArIB and RgIA4 (p ≤ 0.05; n = 6; Supplementary Figure S5 ). The viability of the cells was not affected by the agonists, conopeptides, and mecamylamine as measured by LDH activity in cell culture supernatants ( Supplementary Table S4 ). While LDH values of LPS-primed M0-like macrophages treated with BzATP were 8.2 ± 3.7% (n = 6), LPS-primed M1-like macrophages showed higher LDH values of 28.7 ± 13.7% in response to BzATP (n = 18; Supplementary Table S6 ).

To investigate if this cholinergic mechanism inhibits other pathways of NLRP3 inflammasome activation, we next tested for an ATP-independent mechanisms of inflammasome activation and IL-1β secretion. For this purpose, LPS-primed monocytic THP-1 cells (n = 5; Figure 6A ) and THP-1 cell-derived M1-like macrophages (n = 5; Figure 6B ) were treated with the pore-forming bacterial toxin nigericin (50 µM) instead of BzATP. In these experiments, the ATP-degrading enzyme apyrase (0.5 U/ml), was added together with nigericin to ensure that only ATP-independent mechanisms are investigated. When IL-1β secretion was triggered by nigericin, the nAChR agonists ACh, choline, PC and nicotine had no impact on the release of IL-1β (p ≥ 0.05; Figure 6 ).

To confirm that the cholinergic mechanism inhibiting the BzATP-mediated release of IL-1β is active in primary human macrophages, freshly isolated PBMCs from blood samples of healthy human volunteers were used. The PBMCs were differentiated into hPBMC-derived M0-like and M1-like macrophages. Differences in the levels of surface markers on day 6 of differentiation were detected when comparing the hPBMC-derived M0-like and M1-like macrophages by flow cytometry (n = 3; Supplementary Figure S6 ). On day 6 of differentiation, the cells were primed with LPS (1 µg/ml). Thereafter, BzATP (100 µM), was applied in presence and absence of the nAChR agonists ( Figure 7 ). Priming with LPS alone did not induce the release of relevant amounts of IL-1β by hPBMC-derived M0-like (1 to 8 pg/ml) and M1-like macrophages (2 to 15 pg/ml). Stimulation of LPS-primed cells with BzATP resulted in elevated IL-1β levels in cell culture supernatants of hPBMC-derived M0-like macrophages in the range of 10 to 48 pg/ml and by hPBMC-derived M1-like in the range of 21 to 90 pg/ml. While hPBMC-derived M0-like macrophages showed mixed responses to the nAChR agonists PC, ACh and nicotine ( Figure 7A ), the agonists significantly inhibited the BzATP-induced release of IL-1β by hPBMC-derived M1-like macrophages ( Figure 7B ). In control experiments, PC, ACh and nicotine showed no impact on the LPS-induced release of IL-1β in the absence of BzATP ( Figure 7B ). The viability of the cells was not affected by the agonists as measured by LDH activity in cell culture supernatants ( Supplementary Table S7 ).

We further investigated human peritoneal macrophages ( Figure 8 ) isolated from peritoneal dialysates. Macrophages were enriched via adherence selection and characterized by anti-CD14 and anti-CD68 staining ( Supplementary Figure S7 ). Quantification of the CD14⁺ and CD68⁺ cells revealed a significant increase in the proportion of macrophages by adherence selection (n = 7 each, p ≤ 0.05; Supplementary Figure S7 ). The cells were primed with either 0.1 µg/ml ( Figure 8A ) or 1 µg/ml ( Figure 8B ) LPS for 5 h. Thereafter, BzATP (100 µM), was applied for 40 min in presence and absence of the nAChR agonists ( Figure 8 ). Priming with 0.1 µg/ml LPS alone did not induce the release of relevant amounts of IL-1β by peritoneal macrophages (17 to 78 pg/ml), while additional stimulation with BzATP resulted in IL-1β levels in cell culture supernatants ranging from 768 to 3390 pg/ml (n = 7). Cell priming with 1 µg/ml LPS resulted in IL-1β levels ranging from 16 to 321 pg/ml, that were increased by BzATP to the range of 133 to 2898 pg/ml (n = 7). Interestingly, we detected nAChR agonist-specific responses of human peritoneal macrophages. The BzATP-induced release of IL-1β by peritoneal macrophages was insensitive to ACh (10 µM and 100 µM) and only slightly sensitive to CRP, whereas nicotine was effective in both, cells primed with 0.1 µg/ml or 1 µg/ml LPS ( Figure 8 ). PC inhibited the BzATP-induced release of IL-1β in peritoneal macrophages primed with 0.1 µg/ml LPS (p ≤ 0.05; Figure 8A ) while a tendency of inhibition was seen in peritoneal macrophages primed with 1 µg/ml (p = 0.063; Figure 8B ). The LDH activity remained below 5% independent of the treatment ( Supplementary Table S8 ).

In addition to IL-1β, the cytokines IL-1α and IL-18 were studied. In these experiments the cytokine concentrations of supernatants from the same samples were analyzed in parallel for all three cytokines by ELISA, with IL-1β levels serving as internal controls for the efficiency of the cholinergic agonists (PC, ACh, nicotine, choline) to control the BzATP-mediated IL-1β release. In LPS-primed monocytic THP-1 cells stimulation with BzATP resulted in elevated levels of IL-1β (ranging from 72 to 468 pg/ml; n = 6) and IL-18 (ranging from 67 to 199 pg/ml; n = 6), whereas the concentrations of IL-1α were below the detection levels of the ELISA (7.8 pg/ml). All tested nAChR ligands significantly (p ≤ 0.05; n = 6) reduced the BzATP-induced release of IL-1β and IL-18 ( Figures 9A, B ). LPS-primed THP-1 cell-derived M1-like macrophages responded to BzATP with elevated levels of IL-1β (ranging from 882 to 1621pg/ml), IL-1α (ranging from 14 to 25 pg/ml), and IL-18 (ranging from 35 to 67 pg/ml; n = 6 each). The BzATP-induced release of IL-1β, IL-1α and IL-18 by THP-1 cell-derived M1-like macrophages was significantly reduced by all tested nAChR ligands (p ≤ 0.05; n = 6; Figures 9C–E ).