Background
Active research during the last 20 years has revealed the important role of the vagus nerve in the regulation of immunity and inflammation in a physiological mechanism termed
the inflammatory reflex [
1,
2]. In the inflammatory reflex, sensory (afferent) vagus nerve signaling is activated by cytokines and other inflammatory molecules in response to pathogens, injury, or other pathophysiological events [
1,
3]. This signaling is integrated in the brainstem with motor (efferent) vagus nerve cholinergic signaling, which controls pro-inflammatory cytokine levels and inflammation [
2,
4]. This efferent arm of the inflammatory reflex was termed
the cholinergic anti-inflammatory pathway [
2,
5]. The α7 nicotinic acetylcholine receptor (α7nAChR) expressed on macrophages and other immune cells has been identified as a key mediator of cholinergic anti-inflammatory signaling [
6,
7]. Stimulation of the α7nAChR on macrophages activates downstream intracellular mechanisms, including suppression of NF-κB activation and results in decreased production of TNF and other pro-inflammatory cytokines [
8‐
11]. These discoveries opened an avenue of preclinical research revealing the anti-inflammatory efficacy of vagus nerve stimulation (VNS) and α7nAChR agonists in endotoxemia, sepsis and many other inflammatory conditions [
12,
13]. This research paved the way to recent successful clinical trials with VNS in patients with inflammatory disorders [
14].
Murine endotoxemia and cecal ligation and puncture (CLP) have been widely used in studying the role of the α7nAChR in the cholinergic regulation of inflammation. Endotoxemia, associated with robust systemic cytokine release and inflammation is considered by some as a model of gram negative sepsis, while CLP is a clinically relevant model of polymicrobial sepsis [
15]. Sepsis is a life-threatening condition characterized by organ dysfunction resulting from an excessive inflammatory response to infection. This organ system dysfunction is correlated with higher long term mortality, even if patients recover from their illness in the hospital [
16]. In mice, VNS or pharmacological cholinergic α7nAChR activation suppresses pro-inflammatory cytokine levels and mitigate mortality in mice with endotoxemia and CLP [
13,
17‐
20]. The role of macrophages in endotoxemia and sepsis is complex; some reports characterize macrophages as protective due to their crucial role in efferocytosis of neutrophils, phagocytosis of bacteria, and tissue repair, while other reports indicate their detrimental effects [
21‐
23]. The role of α7nAChR on macrophages in mediating cholinergic suppression of pro-inflammatory cytokine production in the cholinergic anti-inflammatory pathway has been characterized as a major mechanism underlying the neural control of immune responses. However, other potential mechanisms, such as modulating the recruitment of monocytes/macrophages to damaged tissue, remain unclear.
In this study, we investigated the broader role of the α7nAChR in inflammation by examining the migration and accumulation of macrophages during endotoxemia. We showed that the protective role of α7nAChR in endotoxemia is positively correlated with monocyte/macrophage migration to the inflamed tissues. Moreover, we found that α7nAChR-mediated migratory properties depend on the expression of a major adhesive receptor integrin αMβ2 (CD11b/CD18), thus indicating an important molecular link between cholinergic signaling and macrophage motility. Therefore, these results reveal a novel protective mechanism of the cholinergic anti-inflammatory pathway.
Materials and methods
Reagents and antibodies
Reagents were purchased from Sigma-Aldrich (St. Louis, MO, USA), BioRad (Hercules, CA, USA), BioLegend (San Diego, CA, USA), and Thermo Fisher Scientific (Waltham, MA, USA). Lipopolysaccharide (LPS, endotoxin) derived from E. coli O55:B5, and PNU-282987 were purchased through Sigma-Aldrich. Antibodies against cell surface markers Ly6-G (clone 1AB), Ly6-C (clone HK1.4), αM (clone M1/70), αL (clone M17/4), F4/80 (clone BM8), αX (clone N418), and α4 (clone R1–2) are from eBioscience. Antibodies against Siglec F (clone 1RNM44N) and L-selectin (clone MEL-14) are from Invitrogen. Antibodies against CCR2 (clone SA203G11) and CCR5 (clone HM-CCR5) are from BioLegend.
Animals
Wild-type (WT; C57BL/6J, stock #000664) and α7nAChR-deficient (α7nAChR−/−; B6.129S7-Chrna7tm1Bay/J, stock #003232) mouse colonies were purchased from Jackson Laboratory (Bar Harbor, ME, USA). The α7nAChR deficient strain was backcrossed to C57BL/6 for eight generations. Mice aged between 8 and 12 weeks were used for the study. Similar age WT and α7nAChR−/− mice were employed for each experiment. No comparative analysis was conducted across mice of varying sexes and ages. All animal procedures were performed according to animal protocols approved by East Tennessee State University IACUC. Protocol number is P210903.
Endotoxemia
In survival experiments, male or female WT and α7nAChR
−/− mice were intraperitoneally injected with a sublethal dose of LPS calculated based on body weight. Since LPS activity is slightly variable from bath to bath, we evaluated the sublethal dose for each vial preparation using increasing doses of LPS (
E. coli O55:B5) in C57BL/6J wild-type mice. Female mice are more resistant to LPS treatment compared to male mice [
24,
25]. Therefore, a gender-specific dosing was applied to reach a similar percentage of lethality for male and female mice. Depending on the LPS bath preparation we used 7–8 mg/kg for males and 9–12 mg/kg for females. Notably, the same concentration of LPS was used for all groups in each experiment.
In all endotoxemia experiments, body temperature was monitored twice daily using a rectal probe connected to a ThermoWorks (American Fork, UT, USA) MicroTherma meter.
To examine macrophage accumulation in the lungs, male or female WT and α7nAChR
−/− mice were given an intraperitoneal injection of LPS as described above. After 48h, mice were euthanized using Isothesia (Henry Schein Animal Health, Dublin, OH) and perfused, and lungs were removed. Lungs were digested using collagenase II as described below ( "
Flow Cytometry and Imaging Flow Cytometry Analyses" section) and prepared for flow cytometry. In an additional experiment, male and female WT mice were treated intraperitoneally with 3mg/kg PNU-282987, 15 min before the injection of LPS, to examine the effect of α7nAChR stimulation on macrophage accumulation. Control mice received DMSO (vehicle) 15 min before LPS. Samples were incubated with anti-αM/PE-Cy7, anti-CCR2/APC, anti-CCR5/PE-Cy7, anti-Siglec F/FITC, anti-Ly6-G/PE, anti-F4/80/PE, anti-F4/80/APC, and anti-αX/APC across multiple samples.
Isolation of peritoneal macrophages
Thioglycollate-induced peritoneal macrophages are a well-established source to study macrophage function in inflammatory conditions. Peritoneal macrophages from 8- to 12-week-old WT and α7nAChR−/− mice were collected via peritoneal lavage with 5mL of sterile PBS 72 h after intraperitoneal injection of 1mL 4% thioglycolate. Mice were euthanized via CO2 asphyxiation before the collection procedure. The cells were counted and plated in petri dishes for 2 h in RPMI 1640 (Corning, Corning, NY) with 10% FBS and 1% penicillin/streptomycin, after which non-adherent cells were removed.
Isolation of monocytes from mouse bone marrow
The isolation of bone-marrow-derived monocytes provides a pure population of monocytes that exceed the number of monocytes obtained from the peripheral blood of mice by approximately 10 folds. Monocytes were isolated from the femoral and tibial bone marrow of WT and α7nAChR−/− mice by first flushing out bone marrow with RPMI 1640, followed by lysis of red blood cells. Magnetic-assisted cell sorting (MACS) was then used to purify monocytes via a negative separation kit, following the manufacturer’s protocol (Miltenyi Biotec, Gaithersburg, MD, USA). Purity of the isolated monocytes was analyzed by flow cytometry using antibodies to αM/PE-Cy7, Ly6-G/PE, and Ly6-C/FITC. In all experiments, the purity was between 87% and 92%.
Adoptive transfer of monocytes in the model of endotoxemia
Monocytes were isolated from the bone marrow of WT and α7nAChR
−/− mice, as above, and labeled with red PKH26 (WT), or green PKH67 (α7nAChR
−/−) fluorescent dyes. A total of 1X10
6 red and 1X10
6 green monocytes were mixed equally and injected into the tail veins of WT mice or α7nAChR
−/− mice. These mice received a sub-lethal dose of LPS intraperitoneally within 5 min after injection of cells. After 48 h, the mice were sacrificed using Isothesia and perfused with PBS. Lungs, liver, and spleen were isolated and digested with 2mg/mL collagenase II (Sigma Aldrich, St Louis, MO, USA) prepared in HBSS as previously described [
26]. Digested cell suspension was filtered through a 70μm cell strainer and any remaining red blood cells were lysed. Cell filtrate was incubated with a viability dye and analyzed using flow cytometry (Fortessa X-20, Becton Dickson, Franklin Lakes, NJ, USA) and imaging flow cytometry (ImageStream Mark II, Amnis, Seattle, WA, USA) for the detection of fluorescently labeled cells. The dye colors were used with only WT cells in a separate experiment to confirm that dye color does not influence the result.
Adoptive transfer rescue in the model of endotoxemia
WT or α7nAChR
−/− monocytes were isolated from bone marrow as described above in "
Isolation of Monocytes from Mouse Bone Marrow" section. Freshly isolated cells, either WT or α7nAChR
−/−, were injected into the tail veins of WT or α7nAChR
−/− recipient mice. Cell injection was immediately followed by a sub-lethal dose of LPS. In all adoptive transfer rescue experiments, body temperature and morbidity were monitored twice daily. Mortality rate was analyzed using the Kaplan–Meier method.
Flow cytometry and imaging flow cytometry analyses
Flow cytometry analysis was used to assess cell surface markers listed in "
Reagents and Antibodies" section as well as determine the number of PHK26 and PKH67 positive cells in the lungs, liver, and spleen during adoptive transfer. For the analysis of cell surface markers, harvested cells were first incubated with FcR-Blocking solution (eBioscience) for 15 min on ice. Next, samples of 2 × 10
6 cells were incubated with appropriate antibody panels for 30 min on ice. Cells were then washed and analyzed using the Fortessa X-20 (Becton Dickson).
To detect labeled macrophages in tissue, the lungs, liver, and spleen were digested using 2mg/mL collagenase II (Sigma-Aldrich, St Louis, MO, USA) as described above in "
Adoptive Transfer of Monocytes in the Model of Endotoxemia" section. Cell suspension was next pre-cleaned via filtering through a 70μm cell strainer. Cells were incubated with live/dead viability dye for 30 min on ice (Thermo Fisher, Waltham, MA, USA). PKH26 and PKH67 labeled macrophages within the digested organs were analyzed with flow cytometry (Fortessa X-20) and imaging flow cytometry (Image Stream Mark II, Amnis). For analysis of αM on labeled macrophages, preparation was carried out as above.
Imaging flow cytometry analysis results were analyzed using IDEAS 6.2 software. The PKH26 and PKH67 labeled cells were captured on channels 2 and 3, respectively.
Macrophage 3D migration assay
WT and α7nAChR−/− peritoneal macrophages were labeled with PKH26 red fluorescent dye or PKH67 green fluorescent dye. An equal number of WT and α7nAChR−/− were mixed and plated on the membranes of 6.5mm transwell inserts with 8μm pores (Costar, Corning, NY) pre-coated with 4μg/mL fibrinogen for 3 h. A 3-D fibrin gel was made by mixing 0.75mg/mL fibrinogen containing 1% FBS and 1% penicillin/streptomycin with 0.5 U/mL thrombin, at a total volume of 100μL per transwell. MCP-1 (30nM) or RANTES (12.8nM) were added to 100μL of HBSS containing 1% FBS and 1% penicillin/streptomycin and added to the top of the gel to initiate migration. Transwells were incubated for 48h at 37 C in 5% CO2 in a 24-well plate. In each well, 650μL of HBSS containing 1% FBS and 1% penicillin/streptomycin was added beneath the transwell insert to prevent drying of the gel during incubation. Experiment run in two independent replicates with wells of each respective cytokine plaved in triplicate. Migrating cells were detected using confocal microscopy (Leica TCS SP8), and results were analyzed with IMARIS 8.0 software. Wells showing migrated cells were used in statistical analysis, MCP-1 (n = 4) and RANTES (n = 3).
qRT-PCR
Prior to RNA isolation, peritoneal macrophages were incubated overnight with LPS (10ng/mL) and PNU-282897 (30μM). Total RNA was extracted from thioglycolate-induced mouse peritoneal macrophages using the PureLink RNA Mini Kit (Invitrogen, Carlsbad, CA, USA). Reverse transcription was performed using the iScript cDNA Synthesis Kit (Bio Rad, Hercules, CA, USA). Roughly 0.8–1.0μg of cDNA was synthesized in a 20μL reaction volume, per the kit instructions. Real-time PCR reactions were set up in a 96-well qPCR plate using IQ SYBR Green Supermix (Biorad, Hercules, CA, USA) and run using the CFX96 Real Time Thermal Cycler fitted with a C1000 lid (BioRad). Each sample was plated in duplicate. Specific primers for each target were designed and are listed in Table
1 (Integrated DNA Technologies, Coralville, IA). Primer sequences were derived from previously published studies and verified using NCBI Blast and IDT Oligo Analyzer [
27,
28]. Fold changes were normalized to GAPDH. Relative expression of each target was calculated using the Livak Method [
29].
Table 1.
Primers sequence used for qPCR.
αX forward | CTGGATAGCCTTTCTTCTGCT | |
αX reverse | GCACACTGTGTCCGAACTCA | |
αM forward | TCCGGTAGCATCAACAACAT | |
αM reverse | GGTGAAGTGAATCCGGAACT | |
αD forward | GGAACCGAATCAAGGTCAAGT | |
αD reverse | ATCCATTGAGAGAGCTGAGCTG | |
CCR2 forward | ACAGCTCAGGATTAACAGGGACTTG | |
CCR2 reverse | ACCACTTGCATGCACACATGAC | |
CCR5 forward | TCCGTTCCCCCTACAAGAGA | |
CCR5 reverse | TTGGCAGGGTGCTGACATAC | |
MCP-1 forward | TGGAGCATCCACGTGTTGGC | |
MCP-1 reverse | ACTACAGCTTCTTTGGGACA | |
RANTES forward | GCTTCCCTGTCATTGCTTGCTC | |
RANTES reverse | AGATGCCCATTTTCCCAGGACC | |
β1 forward | GTGACCCATTGCAAGGAGAAGGA | |
β1 reverse | GTCATGAATTATCATTAAAAGTTTCCA | |
GAPDH forward | AAGGTCATCCCAGAGCTGAA | |
GAPDH reverse | CTGCTTCACCACCTTCTTGA | |
Trans-endothelial migration assay
Endothelial cells (HUVECS) were labeled using CellVue Claret (Sigma-Aldrich, St-Louis, MO) and incubated overnight on the membranes of 6.5mm transwell inserts with 8µm pores (Costar, Corning, NY) to form a monolayer. Non-adhered endothelial cells were gently washed out. WT and α7nAChR
−/− monocytes were isolated from bone marrow using magnetic-assisted cell sorting as described above in methods "
Isolation of Monocytes from Mouse Bone Marrow" section. Isolated monocytes were labeled using either PKH67 (green) or PKH26 (red), with colors switched to confirm that dye color does not influence the result. Stained monocytes were added on top of the endothelial cells. MCP-1 (30nM) or RANTES (12.8nM) were added to the bottom chamber to start migration, along with media containing 650μL of HBSS with 1% FBS and 2% penicillin/streptomycin to prevent drying. Each respective cytokine was plated in triplicate and the transmigration had three independent replicates. After 3 h, the migration was evaluated by confocal microscopy (Leica TCS SP8). Results were analyzed using IMARIS 8.0. Transwells with well-demarcated HUVEC monolayers were used for analysis, MCP-1 (n = 6), and RANTES (n = 9).
Isolation of peripheral blood monocytes
To evaluate the potential changes in adhesion receptor expressions on circulation monocytes after the LPS challenge, monocytes were isolated from the peripheral blood of WT and α7nAChR−/− mice. Male mice were injected with 8μg LPS per gram of body weight. After 3 h, mice were euthanized using Isothesia, and blood was collected in EDTA (200mM) coated syringes through cardiac puncture. Each mouse yielded 500–700μL of blood, which was diluted using an equal volume of balanced salt solution, prepared as instructed in the Cytiva Ficoll–Paque protocol. The monocytes were separated from whole blood using Ficoll–Paque 1.084 (Cytiva) according to manufacturer instructions. Isolated monocytes were then prepared for flow cytometry using anti-Ly6-C/FITC, anti-αM/PE-Cy7, anti-αL/APC, anti-L-selectin/PE, and anti-α4/PE across multiple samples.
Discussion
Here we showed that genetic α7nAChR deficiency is associated with reduced macrophage migration to the lungs during murine endotoxemia and specific pharmacological activation of this receptor results in increased macrophage migration. These observations indicate a previously unrecognized role for the α7nAChR, a key peripheral component of the cholinergic anti-inflammatory pathway, in mediating macrophage migration during acute inflammation. In parallel, α7nAChR deficiency results in increased mortality of mice during endotoxemia, which indicates a tonic protective function of the α7nAChR in inflammation. These findings identify macrophage migration as an important mechanism contributing to the physiological cholinergic regulation of inflammation. Additional mechanistic insight substantiates this notion, revealing that the expression of integrin αMβ2 is reduced on α7nAChR-deficient monocyte-derived macrophages, indicating its potential role in α7nAChR-mediated macrophage migration.
Macrophages are essential players in innate immunity that may have a protective or pathological contribution to the development of inflammatory diseases [
23]. Macrophage phenotype, tissue distribution, molecular environment, and disease stage define the outcomes of macrophage function. Inhibition of pro-inflammatory cytokine secretion by macrophages was the major anti-inflammatory function reported for α7nAChR [
10,
43,
44]. Pioneering work from Kevin Tracey’s group revealed that α7nAChR activation blocks the nuclear translocation of NF-κB, a master transcription factor for multiple pro-inflammatory genes that generate inflammatory responses [
6,
12,
45‐
47]. However, other potential mechanisms may have a significant contribution to the α7nAChR-mediated macrophage response.
Despite a well-characterized protective role of α7nAChR in endotoxemia and CLP sepsis, the direct effect of α7nAChR-deficiency on survival during endotoxemia was not investigated previously. Our results of α7nAChR
−/− mice compared with WT clearly demonstrate the detrimental impact of α7nAChR deficiency on survival during murine endotoxemia and complement previous observations that administration of the α7nAChR agonists GTS-21 or choline improve the survival of mice during endotoxemia and CLP sepsis [
18,
19]. The significant drop of body temperature at 24 h after LPS-injection is an important pathophysiological effect of endotoxemia. Consequently, body temperature analysis was implemented as a reliable verification of endotoxemia severity and progression in our in vivo experiments.
Previous studies reported that the accumulation of macrophages in the lungs during sepsis can have a protective function [
21,
22]. In contrast, the accumulation of neutrophils is a characteristic feature of sepsis-induced acute lung injury and is associated with poor outcomes [
37,
48]. One of the mechanisms by which macrophages provide protection is through the control of inflammation via efferocytosis of activated neutrophils (Bailey et al. 2021). Here, we evaluated the effect of α7nAChR activation using a specific agonist on macrophage accumulation in lungs. Previous studies demonstrated the potent anti-inflammatory effects of α7nAChR activation by agonists, such as GTS-21 and PNU-282987 in murine models of systemic inflammation and sepsis [
18,
19,
38,
44,
49]. We observed that wild-type mice treated with the agonist PNU-282987 exhibited a significant increase in the number of monocyte-derived macrophages and body temperature, along with a decrease in neutrophil numbers when compared to untreated mice. This finding is consistent with data reported by Huston et al. where nicotine treatment decreased the number of neutrophils accumulated in carrageenan-filled air pouches, as compared to controls [
50].
Consistent with our α7nAChR activation approach, we observed a significant reduction in the number of macrophages in the lungs of α7nAChR-deficient mice during endotoxemia. These data were supported by the decrease in body temperature in α7nAChR-deficient mice which indicates the greater severity of systemic inflammation in these animals.
To provide additional insights in our study, we evaluated in vivo migration by monitoring fluorescently labeled, adoptively transferred monocytes/macrophages in the model of endotoxemia [
32,
40,
41,
51]. We employed an internal control within each recipient mouse by injecting an equal number of monocytes from both WT and α7nAChR
−/− donors, facilitating direct comparison between the two monocyte types. An additional adoptive transfer tracking experiment was performed using α7nAChR-deficient recipients. Both experiments revealed the same pattern: more WT monocytes were detected in the lungs, liver, and spleen when compared to α7nAChR-deficient monocytes. In addition, to confirming the outcome of the experiment with WT recipients, the repetition with α7nAChR-deficient recipients suggests that the enhanced migration of WT monocytes does not depend on the expression of α7nAChR on other cell types. In both setups, the quality of the isolated donor monocytes was validated using flow cytometry, where a purity of 87–92% was confirmed.
To address any potential influence of fluorescent dyes on macrophage migration in vivo, we conducted a separate experiment comparing the migration of equal numbers of WT monocytes labeled with either PKH26 (red) or PKH67 (green) and found that both red and green-labeled WT macrophages exhibited similar motility when migrating towards inflamed tissue. These results provided evidence that the fluorescent dyes themselves do not significantly affect macrophage migration.
To demonstrate the direct involvement of α7nAChR in macrophage migration, we conducted in vitro 3D migration assays, a well-developed technique that we have previously used [
32,
40,
41]. Namely, our experimental setup provided a comprehensive assessment of macrophage migration, wherein monocyte-derived WT and α7nAChR
−/− macrophages, labeled with different fluorescent dyes, migrated through a fibrin matrix against gravity in the presence of a chemokine gradient. By including two types of fluorescently labeled cells (WT and α7nAChR
−/−) within the same matrix, we reduced data variability and enabled accurate calculation of the migration ratio between control and knockout macrophages in each sample. Within the fibrin matrix, we observed that α7nAChR-deficient macrophages exhibited reduced effectiveness in migrating along both RANTES and MCP-1 gradients compared to WT macrophages.
In comparison with our experimental protocol, previous studies that attempted to evaluate the contribution of α7nAChR to macrophage migration utilized macrophage-like cell lines, wild-type cell phenotype (no α7nAChR-knockout), and the most importantly, a simplified 2D transmigration setup without chemokine gradients or protein coatings. For example, the ability of the α7nAChR agonists (PHA-543613 and varenicline) to decrease migration of RAW264.7 cells was demonstrated by testing cell transmigration through uncoated trans-well membranes (Boyden chambers) without a chemokine gradient [
52,
53]. Similar results were obtained by others who showed that acetylcholine can inhibit LPS-induced RAW264.7 cell migration. This study suggested that the inhibition of migration was attributed to the blocking of MMP-9 expression [
54]. MMPs play a role in 3D macrophage migration through the extracellular matrix (ECM) by degrading ECM proteins and creating space for cell movement. However, it should be noted that the presented experiments do not directly verify this hypothesis, as the proposed model using un-coated transwells does not involve MMP-mediated ECM degradation. In this model, macrophages transmigrate via an 8-µm pore-size membrane without immobilized ligands and chemokine gradients, where cell motility is mostly regulated by gravity and diffusion. Therefore, the evaluation of the role of α7nAChR in macrophage migration remained incomplete. In this study, we provided advanced characterization by implementing an improved experimental design and methodology.
In addition to migration through the extracellular matrix, trans-endothelial migration is another crucial step in the recruitment of leukocytes during inflammation. Our findings revealed no significant difference in the transmigration of WT and α7nAChR−/− monocytes across an endothelial monolayer in response to either MCP-1 or RANTES. These data were supported by the similar expression of integrins αL, αM, α4, and L-selectin on WT and α7nAChR−/− mouse peripheral blood monocytes. These molecules are key adhesion receptors in the process of adhering to, and migrating across, the endothelial wall. Based on these results, we concluded that trans-endothelial migration does not contribute to the differential migration observed between WT and α7nAChR−/− monocytes/macrophages.
Reduced expression of integrin αX and integrin αM at the transcriptional level in α7nAChR
−/− macrophages provides a potential mechanistic explanation for their reduced migration. Integrin αMβ2 is a crucial adhesive receptor for the recruitment of monocytes and migration of macrophages through the extracellular matrix. Integrin αXβ2 possesses multiple regulatory functions on macrophages [
55,
56], but has a limited effect on macrophage migration due to relatively low level of expression on macrophage subsets. Therefore, the decreased αM mRNA and protein levels in α7nAChR
−/− macrophages suggest that their impaired migration may be due to altered adhesion.
In contrast to these findings, we observed an increased expression of integrin αDβ2 in α7nAChR
−/− macrophages. Integrin αDβ2 is significantly upregulated on pro-inflammatory (M1-like) macrophages in vivo and in vitro and contributes to the development of various chronic inflammatory diseases, such as atherosclerosis and diabetes [
32,
41]. Importantly, previous studies have shown that activation of α7nAChR leads to macrophage polarization toward the M2 phenotype [
38,
39,
57]; therefore, α7nAChR deficiency should be associated with M1 phenotype, where integrin αDβ2 is upregulated. Based on the levels of αMβ2 and αDβ2 expression on different macrophage subsets, it was suggested that αMβ2 is involved in macrophage migration to and from the sites of inflammation, while αDβ2 plays a role in the retention of pro-inflammatory, M1-polarized macrophages at the sites of chronic inflammation [
40]. Therefore, the modest upregulation of the low-expressed αDβ2 on monocyte-derived macrophages may have a limited impact on macrophage migration but highlights the potential pathological role of integrin αDβ2 in α7nAChR-deficient mice during the development of atherosclerosis and diabetes [
58‐
60].
Interestingly, the injection of WT monocytes did not completely rescue the phenotype of α7nAChR-deficient mice, resulting in only partial improvement in survival. It could be that this incomplete rescue is attributed to the already overwhelming NF-κB activation present in α7nAChR−/− mice; thus, adding a population of WT monocytes may not be sufficient to reverse the deleterious effects caused by α7nAChR deficiency.
The recent discovery of CHRFAM7A, a human-specific dominant negative regulator of α7nAChR function expanded our understanding of how the cholinergic anti-inflammatory pathway is regulated in humans [
61,
62]. The translated protein CHRFAM7A (dupα7) lacks the acetylcholine binding site, leading to reduced α7 receptor activity. In a study using peripheral blood mononuclear cells from septic patients, Cedillo et al. reported that CHRFAM7A has an inverse relationship with disease severity as well as cholinergic anti-inflammatory pathway activity, where patients with higher expression of CHRFAM7A had poorer prognoses [
63]. THP-1 human monocytic cells transfected with dupα7 demonstrated a reduced migration, colony formation, and chemotaxis toward MCP-1 [
64]. Therefore, the reduced α7 receptor activity inhibits macrophage migration, aligning with our observation that α7nAChR deficiency has a negative impact on the migration of mouse macrophages. Despite the translational incongruency, basic research of α7nAChR and the cholinergic anti-inflammatory pathway thus far in rodents, primary human monocytes, and cell lines has provides invaluable insights and presented therapeutic opportunities for the treatment of sepsis.
Based on our results, we propose that α7nAChR deficiency leads to reduced migration of macrophages to the lungs and other inflamed organs, thereby impairing the clearance of recruited neutrophils through efferocytosis. Consequently, neutrophils present in the tissues of α7nAChR
−/− mice secrete pro-inflammatory cytokines. Furthermore, the failed recruitment of α7nAChR-deficint monocytes results in their accumulation in the bloodstream and the secretion of pro-inflammatory cytokines via NF-κB-related mechanisms [
65,
66]. Collectively, these processes contribute to an increased cytokine storm and higher mortality rate.
Publisher's Note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.