Myeloid cells, including proinflammatory monocytes and neutrophils, have important roles in the pathology of multiple sclerosis and its animal model, experimental autoimmune encephalomyelitis (EAE). These cells infiltrate the CNS in the early stages of disease development and contribute to the inflammatory response that is associated with symptom severity. It is thus crucial to identify and understand new mechanisms that can regulate the CNS infiltration of proinflammatory myeloid cells. Nicotinic acetylcholine receptors (nAChRs) have been increasingly studied for their immune-regulatory properties. In this study, we assessed the ability of nicotine, an nAChR ligand, to modulate proinflammatory myeloid cell numbers within the bone marrow, spleen, blood, and CNS of EAE mice. We found that nicotine significantly inhibits the infiltration of proinflammatory monocytes and neutrophils into the CNS at time points where these cells are known to play critical roles in disease pathology. In contrast, nicotine does not affect the expansion of other monocytes. We also show that nicotine exerts these effects by acting on α7 and α9 nAChR subtypes. Finally, mRNA transcript levels for CCL2 and CXCL2, chemokines involved in the chemotaxis of proinflammatory monocytes and neutrophils, respectively, are reduced in the brain of nicotine-treated EAE mice before the massive infiltration of these cells. Taken together, our data provide evidence that nAChRs can regulate proinflammatory cell infiltration into the CNS, which could be of significant value for the treatment of neuroinflammatory disorders.

Inflammation is one of the hallmark features of multiple sclerosis (MS) (1), a debilitating CNS disease that afflicts ∼2.5 million individuals worldwide and lacks effective treatments. Although research in the pathology of MS and its animal model, experimental autoimmune encephalomyelitis (EAE), has focused mostly on the roles of the adaptive immune response, the importance of myeloid cells (monocytes, macrophages, microglia, myeloid dendritic cells, and neutrophils) is becoming increasingly clear. Their critical role is underscored by the fact that myeloid cells are the predominant immune cells found in active MS brain lesions (24). It is well-known that myeloid cells can promote inflammation and thus contribute to disease progression. However, it is increasingly clear that some myeloid cells, in particular monocytic cells (monocytes, macrophages, and microglia), also play important roles in repair mechanisms, which are crucial for disease recovery (5).

The paradoxical roles of monocytic cells in disease pathology are thought to be explained by the presence of at least two subsets of monocytes/macrophages, which are the “classically activated” M1 cells that display mostly proinflammatory functions, and the “alternatively activated” M2 cells, which play anti-inflammatory or regulatory roles (6, 7). M1 and M2 cells can also be distinguished based on surface marker expression, because M1 cells express the chemokine receptor CCR2, as well as high levels of Ly6C (herein called CCR2+Ly6Chigh cells), whereas M2 cells are positive for CX3CR1, another chemokine receptor, and express low levels of Ly6C (7). More importantly, the balance between M1 and M2 cells has been shown to influence the inflammatory outcome, where high proportions of M1 cells appear to promote myelin damage and aggravate symptoms in EAE (5). The distribution dynamics of CCR2+Ly6Chigh cells in the blood and CNS of EAE mice has been the object of a recent study, which found that, although the proportion of myeloid cells in the blood that are CCR2+Ly6Chigh increases before disease onset, disease severity is rather correlated to the ratio of CCR2+Ly6Chigh to total myeloid cells in the CNS (8). This finding underlines the importance of CCR2+Ly6Chigh cell infiltration into the CNS as a pathological mechanism for EAE.

Neutrophils are another type of myeloid cell that have increasingly been the focus of EAE studies. Indeed, neutrophils migrate from the blood to the spinal cord parenchyma within a day before disease onset, and their numbers remain high for a few days before returning to normal levels during the recovery stage (3, 4). Their presence within the CNS is a contributor to disease initiation, because data show that neutrophil depletion significantly ameliorates clinical scores (3, 9). New modalities to control neutrophil infiltration into the brain and spinal cord could thus be highly beneficial toward the treatment of inflammatory disorders of the CNS.

The balance between beneficial and detrimental consequences of myeloid cell activity may well depend on endogenous mechanisms that regulate their numbers and functions. Evidence supports the notion that inflammation is modulated by cholinergic signaling (10), and cholinergic ligands such as nicotine have been shown to reduce the severity of EAE symptoms and ameliorate recovery (1113). Some of these previous studies have reported reduced myeloid cell numbers in the CNS of nicotine-treated EAE mice (11, 13). However, it is unknown whether nicotine specifically regulates CCR2+Ly6Chigh cell numbers within the CNS. In addition, although it is clear that the α7 nicotinic acetylcholine receptor (nAChR) subunit, 1 of the 16 identified nAChR subunits, is a key player in the beneficial effects of nicotine, recent evidence suggests that other nAChRs may also be involved in immune regulation (1315).

In this study, we thus investigated whether nicotine specifically regulates proinflammatory CCR2+Ly6Chigh monocyte and neutrophil cell numbers in immunologically important organs, specifically the bone marrow (BM), spleen, and blood, as well as the brain and spinal cord. We provide evidence that nicotine significantly reduces the amount of CCR2+Ly6Chigh monocytes and neutrophils that enter the brain and spinal cord, and also decreases the numbers of CCR2+Ly6Chigh monocytes in the blood of EAE mice. This effect is most prominent during the peak of disease stages, where the highest level of proinflammatory cell infiltration normally occurs. Nicotine appears to modulate cell infiltration by inhibiting the production of CCL2 and CXCL2 chemokines in the CNS. Finally, our data demonstrate that both α7 and α9 nAChRs are important for nicotinic regulation of myeloid cell distribution in the blood and CNS. This study therefore identifies nAChRs as new molecular targets to control proinflammatory myeloid cell infiltration into the CNS.

C57BL/6J wild type (WT) and α7 nAChR knockout (KO) mice (B6.129S7-Chrnatm1Bay, stock number 003232) were purchased from The Jackson Laboratory (Bar Harbor, ME). The α7 nAChR KO (α7KO) was prepared by the deletion of the last three exons (810) of the Chrna7 gene, encoding the second through the fourth transmembrane domains and the cytoplasmic loop. The α9 nAChR KO (α9KO) heterozygous breeder mice, generated by the deletion of exons 1 and 2 containing the translation/transcription initiation sites, were generously provided by Barbara J. Morley (Boys Town National Research Hospital, Omaha, NE). The α7 and α9 KO lines were maintained in a C57BL/6J background. The genotype of all lines was confirmed by PCR with the suppliers’ protocol and primer sequences: mouse nAChR α7 common forward primer 5′-TTC CTG GTC CTG CTG TGT TA-3′, α7 WT reverse primer 5′-ATC AGA TGT TGC TGG CAT GA-3′ and α7 KO reverse primer 5′-TAG CCG AAT AGC CTC TCC AC-3′ and mouse nAChR α9 WT forward primer 5′-GCC CCA TCC CTG CAT CT-3′, α9 WT reverse primer 5′-GTA GCT TTG GAA TGA GTG GAT GAG C-3′, nAChR α9 KO forward primer 5′-CGG ACC AAC TAA TGA TAC ACT GGA G-3′, and α9 KO reverse primer 5′-GAC CCA CAG AAT GAA CTG AGT TGA C-3′. All animals were housed in individually microventilated cages, up to five mice per cage. Mice used were at least 7–8 wk of age at the experiment’s inception. The experiments were reviewed, approved, and conducted in accordance with the policies outlined by the Canadian Council for Animal Care and were approved by the Université de Moncton’s animal care committee.

To induce acute EAE, we injected mice s.c. in the hind flank with 200 μg myelin oligodendrocyte glycoprotein (MOG)35–55 peptide (single-letter amino acid sequence; M-E-V-G-W-Y-R-S-P-F-S-R-V-V-H-L-Y-R-N-G-K; Synpeptide, Shanghai, China) in IFA (Difco, Detroit, MI) with 500 μg nonviable, desiccated Mycobacterium tuberculosis (Difco). On the day of and 2 d after immunization, the mice were also inoculated with 200 ng pertussis toxin i.p. (List Biologic, Campbell, CA). Mice were monitored daily for symptoms scored on an arbitrary scale of 0 to 5 with 0.5 increments: 0, no symptoms; 1, flaccid tail; 2, hind-limb weakness or abnormal gait; 3, complete hind-limb paralysis; 4, complete hind-limb paralysis with forelimb weakness or paralysis; 5, moribund or deceased.

As described previously (11), nicotine bitartrate (Sigma-Aldrich, St. Louis, MO) in PBS (100 mg/ml) or a solution containing PBS alone was freshly prepared and loaded into Alzet osmotic minipumps (model 1007D and model 2002; DurectCorporation, Cupertino, CA) 12 h before pump implantation. The pumps were implanted s.c. on the back of the mice and continuously delivered either PBS or nicotine salt at 0.39 mg nicotine free base per mouse per day until the indicated days for sacrifice (7, 12, 16, or 22 d). The pump implantation surgeries were done 2 d before MOG immunization.

On the day of sacrifice (3, 7, 12, 16, or 22 d postimmunization), mice were anesthetized with 2% isoflurane (Partenaires Pharmaceutiques du Canada, Richmond Hill, ON). Once the animals were under complete anesthesia, the thoracic cage was opened to expose the heart and the spleen. Blood was collected (see the following subsection Blood Cells), the spleen was harvested (see later Spleen Cells subsection), and animals were then perfused by intracardiac puncture with cold PBS for 2 min. Afterward, BM cells (BMCs; see BMCs subsection later in this article), the brain, and spinal cord (see later CNS Cells subsection) were collected. Tissues were then further processed to generate single-cell suspensions, as described in the following subsections.

Blood cells.

A total of 0.5–1 ml blood was withdrawn by intracardiac puncture using 0.5M EDTA-coated syringes and then transferred into tubes containing a 10% vol 0.5M EDTA. A total of 300 μl 3% dextran and 300 μl PBS were added to the blood to sediment RBCs under room temperature for 45 min. The supernatant was then transferred to a fresh tube, and the remaining RBCs in the supernatant were removed using the RBC lysis buffer (BioLegend, San Diego, CA) as per the manufacturer’s protocol. Cells were then spun (5 min at 450 × g) and the supernatant discarded. An additional wash step with PBS was then completed.

Spleen cells.

Collected spleens were gently dissociated by grinding and sieving through 70-μm cell strainers (Corning, Durham, NC) with PBS. RBC lysis buffer was applied to remove the RBCs. The total spleen cells were passed through a 40-μm cell strainer to remove cell debris. Cells were then spun (5 min at 450 × g) and the supernatant was discarded. An additional wash step with PBS was then completed.

BMCs.

BMCs were flushed out of the femur and tibia with PBS, and RBCs were lysed with RBC lysis buffer. Cells were then passed through a 40-μm cell strainer (Corning) to obtain single cells. Cells were then spun (5 min at 450 × g) and the supernatant was discarded, followed by an additional wash step with PBS.

CNS cells.

Cold PBS was delivered intracardially to eliminate the contamination of blood cells in the CNS. Brain and spinal cord were removed, finely dissected, and then subjected to enzymatic digestion using the Neural Tissue Dissociation Kit (P) (Miltenyi Biotec, Auburn, CA), and dissociated automatically using the gentleMACS Octo Dissociator with Heaters (Miltenyi Biotec), using the default program for neural tissues. Myelin debris were then removed by first diluting the cell suspension 2-fold with 9% sucrose in PBS, followed by centrifugation (700 × g for 15 min). Pelleted cells were washed with PBS and centrifuged (450 × g for 5 min), after which the supernatant was discarded.

Single-cells suspensions from the various tissues were resuspended at the concentration of 105–106/100 μl in PBS. Cells were incubated with anti-mouse TruStain fcX (BioLegend, San Diego, CA) at room temperature for 10 min to block the Fc receptors. Monocyte phenotype was analyzed by staining for one or more of the following mouse Ags (targeted by the indicated Ab fluorescently tagged with either Alexa Fluor 488, PE, allophycocyanin, Pacific blue, Alexa Fluor 700, PE/Cy7, or allophycocyanin-Cy7): CD11b(M1/70), Ly6G(1A8), CD80(16-10A1), CD86(PO3), MHC-II(M5/114.15.2), CCR2(475301), Ly6C(HK1.4), CD45(30-F11), CD49d/VLA-4(R1-2), and LFA-1(H155-78). All cytometric Abs were purchased from BioLegend except for CCR2-PE, which was purchased from R&D Systems (Burlington, ON, Canada). Appropriate isotype controls were included. Cell viability was measured with 7-aminoactinomycin D (7-AAD) (eBioscience), as per the supplied protocol. All data were acquisitioned using a FC500 or MoFlo XDP flow cytometer (Beckman Coulter), and results were analyzed using the Kaluza software (Beckman Coulter).

Clean femoral and tibial bones from hind limbs were obtained by removing muscle tissues and sterilizing bones in EtOH 70%. BMCs were flushed from the bones with PBS using a 10-ml syringe and a 21-gauge needle for the femur, or a 25-gauge needle for the tibia, and collected in a 50-ml tube. Cell aggregates were dislodged by gentle pipetting. RBCs where lysed by RBC lysis buffer, as per the supplier’s protocol. Debris was removed by passaging the suspension through a 40-μm BD Falcon Cell Strainer. Cells were harvested by centrifugation at 300 × g for 10 min, and the supernatant was removed. BMCs were solubilized in complete medium (RPMI 1640 supplemented with 10% heat-inactivated FBS, 1% penicillin/streptomycin, and 2 mM l-glutamine) at a concentration of 2 × 106 cells/ml, supplemented with M-CSF (10 ng/ml) and 2.5 ng/ml recombinant mouse IFN-γ (Cell Guidance Systems, Cambridge, U.K.). Cells were cultured in a humidified incubator at 37°C and 5% CO2 for 3 d. Suspended cells were harvested by gentle pipetting of the medium, and adherent cells were washed with PBS and detached using a 10 mM EDTA solution at a pH of 7.4. Suspension and adherent cells were combined and spun at ∼450 × g for 5 min. The supernatant was removed and cells were washed once by adding 10 ml PBS followed by a 5-min spin at 450 × g. After the wash, cells were counted and resuspended in PBS at a concentration of ≤1 × 106 cells/100 μl. Cells were then incubated with 7-AAD, anti-CD11b, and anti-Ly6G Abs, as described in the Flow cytometric analysis section. Viable monocytes (CD11b+Ly6G) and neutrophils (CD11b+Ly6G+) were then sorted into separate tubes containing complete media with a MoFlo XDP cell sorter (Beckman Coulter). 7-AAD+ and low forward scatter events were always discriminated against while sorting to eliminate nonviable cells, cell debris, and microparticles. A small portion of sorted cells was analyzed using the MoFlo XDP to ensure the efficiency of the sorting. Cells were then used for transwell migration assays.

Migration assays were conducted in Boyden chambers (CytoSelect 96-well cell migration assay; Cell Biolabs, San Diego, CA), as per the supplied protocol. Membrane pore sizes were 5 μm for monocytes and 3 μm for neutrophils. In brief, 100,000 cells were seeded in the upper chamber in RPMI 1640 medium containing 5% FBS. The lower chamber contained 100 ng/ml CCL2 (MCP-1) for monocytes or 10 ng/ml IL-8 for neutrophils, also in RPMI 1640 medium containing 5% FBS. Cells were allowed to migrate for 2 h in a humidified incubator at 37°C and 5% CO2. At the end of the assay, migratory cells were dissociated from the membrane with the supplied cell detachment buffer. Migratory cells were then counted with the MoxiZ cell counter.

Cells were lysed in RiboZol (Amresco, Solon, OH) and kept in −80°C until use. Total RNA was purified using the Qiagen RNeasy Mini Kit (Valencia, CA), as per the supplied protocol. Total RNA yield and concentration were quantified using NanoDrop 1000 (Thermo Scientific, Waltham, MA). cDNA was then generated using the qScript XLT cDNA SuperMix (Quanta Biosciences, Gaithersburg, MD), as per the supplied protocol, followed by RNA digestion with RNase H (2 U/μl) at 37°C for 20 min (Ambion, Burlington, ON). Quantitative real-time PCR was then performed on the CFX Connect Real-Time PCR Detection System (Bio-Rad), using SsoAdvancedSYBR Green Supermix (Bio-Rad) and the following primers: mouse CCL2 forward primer 5′-CTGCTGTTCACAGTTGCCG-3′ and reverse primer 5′-GCACAGACCTCTCTCTTGAGC-3′; mouse CX3CL1 forward primer 5′-CACGAATCCCAGTGGCTTTG-3′ and reverse primer 5′-GGCGTCTTGGACCCATTTCT-3′; mouse CXCL2 forward primer 5′-AGGGCGGTCAAAAAGTTTGC-3′ and reverse primer 5′-CAGGTACGATCCAGGCTTCC-3′; mouse hypoxanthine phosphoribosyltransferase 1 forward primer 5′-TGCTGACCTGCTGGATTACA-3′ and reverse primer 5′-TTTATGTCCCCCGTTGACTGA-3′; mouse matrix metalloproteinase-9 (MMP-9) forward primer 5′-TAGATCATTCCAGCGTGCCG-3′ and reverse primer 5′-GCCTTGGGTCAGGCTTAGAG-3′; mouse ICAM-1 forward primer 5′-CAA TTT CTC ATG CCG CAC AG-3′ and reverse primer 5′-AGC TGG AAG ATC GAA AGT CCG-3′; mouse VCAM-1 forward primer 5′-TGA ACC CAA ACA GAG GCA GAG T-3′ and reverse primer 5′-GGT ATC CCA TCA CTT GAG CAG G-3′. Cycle conditions were 95°C for 30 s, followed by 40 cycles of 95°C for 5 s and 60°C for 30 s, and finally a melt curve 65–95°C (0.5°C increment) for 5 s/step. Results were analyzed using CFX manager software.

The Mann–Whitney U tests were used to compare differences among each group. Data are presented as mean ± SEM. For all statistical analysis, the level of significance was set at p < 0.05.

Clinical score of the disease was recorded from the day after immunization until the peak phase of EAE in WT, α7KO, and α9KO mice. As shown in Fig. 1A, there was a significant decrease in the clinical scores of nicotine-treated WT EAE (WT-EAE-Nic) animals from day 8 until day 15 postimmunization. However, nicotine did not significantly alter clinical scores in α7KO animals (Fig. 1B). In contrast, there was a significant decrease in clinical scores from 10 to 16 d postimmunization in α9KO mice, independent of treatment. However, nicotine treatment itself had no additional effect in α9KO mice (Fig. 1C). As depicted in Fig. 1D, the average clinical score from 13 to 16 d postimmunization in WT-EAE-PBS mice was significantly different from in WT-EAE-Nic animals, and in α9KO animals regardless of nicotine treatment [3.00 ± 0.18, 2.45 ± 0.19, 2.56 ± 0.25, 2.59 ± 0.17, 2.45 ± 0.16, and 2.07 ± 0.23 for PBS-treated WT EAE (WT-EAE-PBS), WT-EAE-Nic, PBS-treated α7KO EAE (α7KO-EAE-PBS), α7KO-EAE-Nic, PBS-treated α9KO EAE (α9KO-EAE-PBS), and α9KO-EAE-Nic, respectively; p < 0.05 when comparing WT-EAE-PBS versus WT-EAE-Nic or WT-EAE-PBS versus α9KO-EAE-PBS]. Similarly, the cumulative clinical score (Fig. 1E) also decreased in WT-EAE-Nic animals and all α9KO animals (19.2 ± 1.7, 12.6 ± 1.0, 16.0 ± 1.7, 15.5 ± 1.3, 14.3 ± 1.1, and 12.0 ± 1.4 for WT-EAE-PBS, WT-EAE-Nic, α7KO-EAE-PBS, α7KO-EAE-Nic, α9KO-EAE-PBS, and α9KO-EAE-Nic, respectively; p < 0.05 when comparing WT-EAE-PBS versus WT-EAE-Nic or WT-EAE-PBS versus α9KO-EAE-PBS). Nicotine treatment also delayed disease onset in WT mice, without much effect in α7KO or α9KO animals (7.8 ± 0.2, 9.7 ± 0.3, 8.1 ± 0.5, 8.8 ± 0.3, 8.7 ± 0.2, and 9.0 ± 0.2 d postimmunization for WT-EAE-PBS, WT-EAE-Nic, α7KO-EAE-PBS, α7KO-EAE-Nic, α9KO-EAE-PBS, and α9KO-EAE-Nic, respectively; p < 0.05 when comparing WT-EAE-PBS versus WT-EAE-Nic). These data confirm previous findings that nicotine protects against EAE via α7 nAChRs (12, 13) while genetic deletion of α9 nAChRs also attenuates disease symptoms (14).

FIGURE 1.

Clinical features of EAE in WT, α7KO, or α9KO mice after nicotine treatment. Mice were immunized with MOG35–55 and treated with nicotine for up to 16 d via osmotic pumps, and disease severity was scored each day as described in 2Materials and Methods. (A) Disease progression was significantly altered in WT-EAE-Nic (black lines with squares, n = 14) compared with WT-EAE-PBS (black lines with circles, n = 17) from days 8 to 15. (B) Clinical scores were not different between α7KO-EAE-PBS (green line with black circles, n = 11) and nicotine-treated α7KO (α7KO-EAE-Nic, green line with green diamond, n = 11) mice, nor between α7KO EAE (PBS-treated and nicotine-treated combined) and WT-EAE-PBS mice (black line). (C) Although there were no differences between α9KO-EAE-PBS (red line with black circles, n = 20) and nicotine-treated α9KO (α9KO-EAE-Nic, red line with red diamond, n = 20) mice, disease progression was significantly lower in α9KO mice, irrespective of drug treatment, compared with WT-EAE-PBS mice. (D and E) Average clinical scores from days 13 to 16 (D) and cumulative clinical scores from disease onset to day 16 (E) were reduced by nicotine treatment (patterned bars) in WT mice (black and white bars), but not in α7KO (green bars) or α9KO (red bars) mice. Again, both these parameters were significantly lower in α9KO mice compared with their WT counterparts, regardless of drug treatment. (F) Disease onset was also delayed in WT-EAE-Nic mice, whereas no drug or genetic effects were observed in α7KO or α9KO mice. These data confirm previous findings that nicotine is protective against EAE, that both α7 and α9 nAChRs appear to be involved in these protective effects. In addition, our data support the hypothesis that the α9 nAChR may play a role in endogenous immune-regulatory mechanisms. Asterisk denotes a statistically significant difference between groups (Mann–Whitney U test, *p < 0.05).

FIGURE 1.

Clinical features of EAE in WT, α7KO, or α9KO mice after nicotine treatment. Mice were immunized with MOG35–55 and treated with nicotine for up to 16 d via osmotic pumps, and disease severity was scored each day as described in 2Materials and Methods. (A) Disease progression was significantly altered in WT-EAE-Nic (black lines with squares, n = 14) compared with WT-EAE-PBS (black lines with circles, n = 17) from days 8 to 15. (B) Clinical scores were not different between α7KO-EAE-PBS (green line with black circles, n = 11) and nicotine-treated α7KO (α7KO-EAE-Nic, green line with green diamond, n = 11) mice, nor between α7KO EAE (PBS-treated and nicotine-treated combined) and WT-EAE-PBS mice (black line). (C) Although there were no differences between α9KO-EAE-PBS (red line with black circles, n = 20) and nicotine-treated α9KO (α9KO-EAE-Nic, red line with red diamond, n = 20) mice, disease progression was significantly lower in α9KO mice, irrespective of drug treatment, compared with WT-EAE-PBS mice. (D and E) Average clinical scores from days 13 to 16 (D) and cumulative clinical scores from disease onset to day 16 (E) were reduced by nicotine treatment (patterned bars) in WT mice (black and white bars), but not in α7KO (green bars) or α9KO (red bars) mice. Again, both these parameters were significantly lower in α9KO mice compared with their WT counterparts, regardless of drug treatment. (F) Disease onset was also delayed in WT-EAE-Nic mice, whereas no drug or genetic effects were observed in α7KO or α9KO mice. These data confirm previous findings that nicotine is protective against EAE, that both α7 and α9 nAChRs appear to be involved in these protective effects. In addition, our data support the hypothesis that the α9 nAChR may play a role in endogenous immune-regulatory mechanisms. Asterisk denotes a statistically significant difference between groups (Mann–Whitney U test, *p < 0.05).

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To assess the effect of nicotine treatment on monocyte infiltration and subpopulation distribution in the CNS during the course of EAE, we sacrificed nicotine-treated and PBS-treated animals on the 7th, 12th, 16th, or 22nd day postimmunization. We measured the infiltration of monocytes and its subpopulations into the CNS by analyzing cells isolated from the brain and spinal cord by flow cytometry. The strategy for the analysis was to first use the combination of the markers CD11b and Ly6G to distinguish between monocytes/macrophages/microglia (CD11b+Ly6G) and neutrophils (CD11b+Ly6G+) (Fig. 2A, 2B). Resident microglia are CD11b+CD45med and can be distinguished from CD11b+CD45high cells, which include infiltrating monocytes/macrophages and activated microglia (Fig. 2C) (16, 17). Among the CD11b+CD45high cells, proinflammatory monocytes/macrophages are CCR2+Ly6Chigh (Fig. 2D). See Table I for the averages ± SEM of the percentages of myeloid cells in the CNS for each group and time point.

FIGURE 2.

Nicotine inhibits the recruitment of proinflammatory CCR2+Ly6Chigh monocytes into the CNS of EAE mice. The brain and spinal cord from WT naive (white background), WT-EAE-PBS (dark gray background), or WT-EAE-Nic (light gray background) mice were harvested on the 7th, 12th, 16th, and 22nd day postimmunization, and brain monocytes/activated macrophages and microglia were analyzed by flow cytometry (n = 6–8 per group per time point). (AD) The gating strategy used is as follows: total monocytic cells were first identified based on their expression of CD11b [(A), CD11b+ gate], followed by their lack of Ly6G expression [(B), Ly6G gate]. While gating on CD11b+Ly6G cells, microglia (CD11b+Ly6GCD45med, upper left gate) and infiltrating monocytes/activated macrophages (CD11b+Ly6GCD45high, upper right gate) were then identified (C). Infiltrating monocytes/activated macrophages were then further analyzed based on the expression of CCR2 and Ly6C, of which proinflammatory cells are CCR2+Ly6Chigh [(D), upper right quadrant]. (E) The percentages of resident microglia (patterned portion of bars), infiltrating monocytes/macrophages (solid portion of bars), and total monocytic cells (infiltrating monocytes/macrophages plus microglia) with respect to total brain cells were analyzed in the brain (left panel) and spinal cord (right panel) at various time points postimmunization. Nicotine significantly inhibits the percentages of infiltrating monocytes/macrophages found in the brain and spinal cord at day 16 postimmunization, whereas not affecting the proportions of microglia. (F) Infiltrating monocytes/macrophages were further subdivided into the proinflammatory CCR2+Ly6Chigh population (solid portion of bars), whereas all other monocytic cells (patterned portion of bars) were grouped together. Nicotine significantly inhibited the recruitment of CCR2+Ly6Chigh cells only. These data show that nicotine prevents the overall EAE-induced increase in monocytic cell numbers in the CNS specifically by preventing the recruitment of proinflammatory monocytes/macrophages. Data shown are means ± SEM, whereas an asterisk denotes a significant difference between indicated groups (Mann–Whitney U test, *p < 0.05).

FIGURE 2.

Nicotine inhibits the recruitment of proinflammatory CCR2+Ly6Chigh monocytes into the CNS of EAE mice. The brain and spinal cord from WT naive (white background), WT-EAE-PBS (dark gray background), or WT-EAE-Nic (light gray background) mice were harvested on the 7th, 12th, 16th, and 22nd day postimmunization, and brain monocytes/activated macrophages and microglia were analyzed by flow cytometry (n = 6–8 per group per time point). (AD) The gating strategy used is as follows: total monocytic cells were first identified based on their expression of CD11b [(A), CD11b+ gate], followed by their lack of Ly6G expression [(B), Ly6G gate]. While gating on CD11b+Ly6G cells, microglia (CD11b+Ly6GCD45med, upper left gate) and infiltrating monocytes/activated macrophages (CD11b+Ly6GCD45high, upper right gate) were then identified (C). Infiltrating monocytes/activated macrophages were then further analyzed based on the expression of CCR2 and Ly6C, of which proinflammatory cells are CCR2+Ly6Chigh [(D), upper right quadrant]. (E) The percentages of resident microglia (patterned portion of bars), infiltrating monocytes/macrophages (solid portion of bars), and total monocytic cells (infiltrating monocytes/macrophages plus microglia) with respect to total brain cells were analyzed in the brain (left panel) and spinal cord (right panel) at various time points postimmunization. Nicotine significantly inhibits the percentages of infiltrating monocytes/macrophages found in the brain and spinal cord at day 16 postimmunization, whereas not affecting the proportions of microglia. (F) Infiltrating monocytes/macrophages were further subdivided into the proinflammatory CCR2+Ly6Chigh population (solid portion of bars), whereas all other monocytic cells (patterned portion of bars) were grouped together. Nicotine significantly inhibited the recruitment of CCR2+Ly6Chigh cells only. These data show that nicotine prevents the overall EAE-induced increase in monocytic cell numbers in the CNS specifically by preventing the recruitment of proinflammatory monocytes/macrophages. Data shown are means ± SEM, whereas an asterisk denotes a significant difference between indicated groups (Mann–Whitney U test, *p < 0.05).

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Table I.
Myeloid cell percentages in the CNS
7 d12 d16 d22 d
BrainSpinal CordBrainSpinal CordBrainSpinal CordBrainSpinal Cord
% CD11b+CD45+ cells in CNS 
 Naive 8.79 ± 0.79 8.34 ± 1.34 8.79 ± 0.79 8.34 ± 1.34 8.79 ± 0.79 8.34 ± 1.34 8.79 ± 0.79 8.34 ± 1.34 
 WT-EAE-PBS 7.87 ± 2.18 9.49 ± 2.79 12.83 ± 2.14 12.59 ± 3.65 18.67 ± 2.05 17.08 ± 2.75 11.99 ± 1.67 11.80 ± 2.30 
 WT-EAE-Nic 7.96 ± 1.50 11.12 ± 1.55 10.04 ± 1.08 9.2 ± 1.56 12.69 ± 1.93 11.80 ± 2.35 10.5 ± 1.38 10.93 ± 1.03 
% CD11b+CD45med cells in CNS 
 Naive 7.49 ± 0.77 7.75 ± 1.31 7.49 ± 0.77 7.75 ± 1.31 7.49 ± 0.77 7.75 ± 1.31 7.49 ± 0.77 7.75 ± 1.31 
 WT-EAE-PBS 6.69 ± 2.09 7.14 ± 1.54 8.30 ± 1.42 8.04 ± 3.37 8.98 ± 2.18 9.60 ± 2.44 7.65 ± 1.35 7.05 ± 1.97 
 WT-EAE-Nic 6.66 ± 0.87 9.90 ± 1.42 7.36 ± 0.76 6.67 ± 2.23 7.84 ± 2.07 6.29 ± 1.78 6.72 ± 1.04 7.29 ± 1.37 
% CD11b+CD45high cells in CNS 
 Naive 1.47 ± 0.22 0.90 ± 0.32 1.47 ± 0.22 0.90 ± 0.32 1.47 ± 0.22 0.90 ± 0.32 1.47 ± 0.22 0.90 ± 0.32 
 WT-EAE-PBS 2.32 ± 0.37 2.10 ± 0.95 4.55 ± 1.53 5.22 ± 1.44 9.69 ± 1.27 9.27 ± 1.86 4.34 ± 0.32 4.75 ± 0.57 
 WT-EAE-Nic 2.25 ± 0.39 1.11 ± 0.15 3.24 ± 0.20 3.46 ± 1.37 4.85 ± 1.17 5.51 ± 0.64 3.78 ± 0.80 3.65 ± 0.69 
 α7KO-EAE-PBS — — — — 6.11 ± 3.27 5.77 ± 4.34 — — 
 α7KO-EAE-Nic — — — — 10.98 ± 4.02 12.94 ± 2.82 — — 
 α9KO-EAE-PBS — — — — 8.32 ± 1.90 10.86 ± 1.39 — — 
 α9KO-EAE-Nic — — — — 9.56 ± 2.46 10.73 ± 2.02 — — 
CCR2+Ly6Chigh in CNS 
 Naive 0.42 ± 0.13 0.21 ± 0.13 0.42 ± 0.13 0.21 ± 0.13 0.42 ± 0.13 0.21 ± 0.13 0.42 ± 0.13 0.21 ± 0.13 
 WT-EAE-PBS 1.32 ± 0.36 0.51 ± 0.29 2.42 ± 0.72 2.78 ± 0.77 4.96 ± 1.21 4.83 ± 1.14 0.86 ± 0.21 0.79 ± 0.39 
 WT-EAE-Nic 0.80 ± 0.28 0.29 ± 0.10 1.89 ± 0.32 2.21 ± 1.07 2.11 ± 1.01 1.86 ± 0.40 0.66 ± 0.15 0.77 ± 0.49 
 α7KO-EAE-PBS — — — — 2.62 ± 1.46 4.15 ± 1.45 — — 
 α7KO-EAE-Nic — — — — 5.21 ± 2.32 4.69 ± 1.81 — — 
 α9KO-EAE-PBS — — — — 3.59 ± 0.92 5.76 ± 0.99 — — 
 α9KO-EAE-Nic — — — — 4.38 ± 1.52 4.45 ± 1.31 — — 
Non-CCR2+Ly6Chigh in CNS 
 Naive 1.05 ± 0.17 0.72 ± 0.24 1.05 ± 0.17 0.72 ± 0.24 1.05 ± 0.17 0.72 ± 0.24 1.05 ± 0.17 0.72 ± 0.24 
 WT-EAE-PBS 1.00 ± 0.25 0.87 ± 0.15 5.107 ± 0.67 2.45 ± 0.88 5.11 ± 0.67 4.44 ± 1.36 3.48 ± 0.27 3.96 ± 0.26 
 WT-EAE-Nic 1.44 ± 0.33 0.86 ± 0.10 1.36 ± 0.28 1.25 ± 1.60 4.21 ± 0.94 4.03 ± 0.23 3.12 ± 0.66 2.88 ± 0.21 
% Neutrophils in CNS 
 Naive 0.74 ± 0.12 1.17 ± 0.79 0.74 ± 0.12 1.17 ± 0.79 0.74 ± 0.12 1.17 ± 0.79 0.74 ± 0.12 1.17 ± 0.79 
 WT-EAE-PBS 1.46 ± 0.36 0.93 ± 0.40 3.04 ± 0.73 5.68 ± 3.18 3.90 ± 0.59 10.54 ± 4.13 2.46 ± 0.27 2.17 ± 0.78 
 WT-EAE-Nic 1.27 ± 0.43 1.93 ± 1.67 3.27 ± 0.79 5.52 ± 2.92 2.24 ± 0.43 4.06 ± 1.61 2.82 ± 0.50 2.45 ± 1.43 
 α7KO-EAE-PBS — — — — 3.48 ± 0.49 6.04 ± 2.18 — — 
 α7KO-EAE-Nic — — — — 3.11 ± 1.25 6.95 ± 2.80 — — 
 α9KO-EAE-PBS — — — — 3.93 ± 0.80 9.52 ± 2.72 — — 
 α9KO-EAE-Nic — — — — 4.07 ± 0.54 8.43 ± 2.74 — — 
7 d12 d16 d22 d
BrainSpinal CordBrainSpinal CordBrainSpinal CordBrainSpinal Cord
% CD11b+CD45+ cells in CNS 
 Naive 8.79 ± 0.79 8.34 ± 1.34 8.79 ± 0.79 8.34 ± 1.34 8.79 ± 0.79 8.34 ± 1.34 8.79 ± 0.79 8.34 ± 1.34 
 WT-EAE-PBS 7.87 ± 2.18 9.49 ± 2.79 12.83 ± 2.14 12.59 ± 3.65 18.67 ± 2.05 17.08 ± 2.75 11.99 ± 1.67 11.80 ± 2.30 
 WT-EAE-Nic 7.96 ± 1.50 11.12 ± 1.55 10.04 ± 1.08 9.2 ± 1.56 12.69 ± 1.93 11.80 ± 2.35 10.5 ± 1.38 10.93 ± 1.03 
% CD11b+CD45med cells in CNS 
 Naive 7.49 ± 0.77 7.75 ± 1.31 7.49 ± 0.77 7.75 ± 1.31 7.49 ± 0.77 7.75 ± 1.31 7.49 ± 0.77 7.75 ± 1.31 
 WT-EAE-PBS 6.69 ± 2.09 7.14 ± 1.54 8.30 ± 1.42 8.04 ± 3.37 8.98 ± 2.18 9.60 ± 2.44 7.65 ± 1.35 7.05 ± 1.97 
 WT-EAE-Nic 6.66 ± 0.87 9.90 ± 1.42 7.36 ± 0.76 6.67 ± 2.23 7.84 ± 2.07 6.29 ± 1.78 6.72 ± 1.04 7.29 ± 1.37 
% CD11b+CD45high cells in CNS 
 Naive 1.47 ± 0.22 0.90 ± 0.32 1.47 ± 0.22 0.90 ± 0.32 1.47 ± 0.22 0.90 ± 0.32 1.47 ± 0.22 0.90 ± 0.32 
 WT-EAE-PBS 2.32 ± 0.37 2.10 ± 0.95 4.55 ± 1.53 5.22 ± 1.44 9.69 ± 1.27 9.27 ± 1.86 4.34 ± 0.32 4.75 ± 0.57 
 WT-EAE-Nic 2.25 ± 0.39 1.11 ± 0.15 3.24 ± 0.20 3.46 ± 1.37 4.85 ± 1.17 5.51 ± 0.64 3.78 ± 0.80 3.65 ± 0.69 
 α7KO-EAE-PBS — — — — 6.11 ± 3.27 5.77 ± 4.34 — — 
 α7KO-EAE-Nic — — — — 10.98 ± 4.02 12.94 ± 2.82 — — 
 α9KO-EAE-PBS — — — — 8.32 ± 1.90 10.86 ± 1.39 — — 
 α9KO-EAE-Nic — — — — 9.56 ± 2.46 10.73 ± 2.02 — — 
CCR2+Ly6Chigh in CNS 
 Naive 0.42 ± 0.13 0.21 ± 0.13 0.42 ± 0.13 0.21 ± 0.13 0.42 ± 0.13 0.21 ± 0.13 0.42 ± 0.13 0.21 ± 0.13 
 WT-EAE-PBS 1.32 ± 0.36 0.51 ± 0.29 2.42 ± 0.72 2.78 ± 0.77 4.96 ± 1.21 4.83 ± 1.14 0.86 ± 0.21 0.79 ± 0.39 
 WT-EAE-Nic 0.80 ± 0.28 0.29 ± 0.10 1.89 ± 0.32 2.21 ± 1.07 2.11 ± 1.01 1.86 ± 0.40 0.66 ± 0.15 0.77 ± 0.49 
 α7KO-EAE-PBS — — — — 2.62 ± 1.46 4.15 ± 1.45 — — 
 α7KO-EAE-Nic — — — — 5.21 ± 2.32 4.69 ± 1.81 — — 
 α9KO-EAE-PBS — — — — 3.59 ± 0.92 5.76 ± 0.99 — — 
 α9KO-EAE-Nic — — — — 4.38 ± 1.52 4.45 ± 1.31 — — 
Non-CCR2+Ly6Chigh in CNS 
 Naive 1.05 ± 0.17 0.72 ± 0.24 1.05 ± 0.17 0.72 ± 0.24 1.05 ± 0.17 0.72 ± 0.24 1.05 ± 0.17 0.72 ± 0.24 
 WT-EAE-PBS 1.00 ± 0.25 0.87 ± 0.15 5.107 ± 0.67 2.45 ± 0.88 5.11 ± 0.67 4.44 ± 1.36 3.48 ± 0.27 3.96 ± 0.26 
 WT-EAE-Nic 1.44 ± 0.33 0.86 ± 0.10 1.36 ± 0.28 1.25 ± 1.60 4.21 ± 0.94 4.03 ± 0.23 3.12 ± 0.66 2.88 ± 0.21 
% Neutrophils in CNS 
 Naive 0.74 ± 0.12 1.17 ± 0.79 0.74 ± 0.12 1.17 ± 0.79 0.74 ± 0.12 1.17 ± 0.79 0.74 ± 0.12 1.17 ± 0.79 
 WT-EAE-PBS 1.46 ± 0.36 0.93 ± 0.40 3.04 ± 0.73 5.68 ± 3.18 3.90 ± 0.59 10.54 ± 4.13 2.46 ± 0.27 2.17 ± 0.78 
 WT-EAE-Nic 1.27 ± 0.43 1.93 ± 1.67 3.27 ± 0.79 5.52 ± 2.92 2.24 ± 0.43 4.06 ± 1.61 2.82 ± 0.50 2.45 ± 1.43 
 α7KO-EAE-PBS — — — — 3.48 ± 0.49 6.04 ± 2.18 — — 
 α7KO-EAE-Nic — — — — 3.11 ± 1.25 6.95 ± 2.80 — — 
 α9KO-EAE-PBS — — — — 3.93 ± 0.80 9.52 ± 2.72 — — 
 α9KO-EAE-Nic — — — — 4.07 ± 0.54 8.43 ± 2.74 — — 

The average percentages of each subpopulation of myeloid cells with respect to the total number of cells in the CNS in each group and time point are indicated ± SEM. Dashes indicate that no data were collected at these time points.

As shown in Fig. 2E, we observed an increase in total CD11b+CD45+ cell (infiltrating monocytes/macrophages and microglia) percentage in the brain and spinal cord of PBS-treated mice from the 12th day onward; this increase peaked at 16 d and remained greater than normal until 22 d after immunization. The increase in total CD11b+CD45+ cells at 16 d was inhibited by nicotine treatment. To determine which CD11b+CD45+ subpopulation contributed to the increase in total cell numbers, we then quantified the proportions of resident microglia (CD11b+CD45med) and infiltrating monocytes/activated macrophages (CD11b+CD45high). We found that the proportion of resident microglia relative to total cells in the brain or spinal cord remained constant throughout the different time points, and these numbers were unchanged after nicotine treatment. In contrast, there was an increase in the infiltrating monocyte/activated macrophage percentage relative to total brain or spinal cord cells, starting from 12 d postimmunization, peaking at 16 d, and diminishing but remaining higher than naive mice until 22 d. Interestingly, the increase in infiltrating monocyte/activated macrophage cells was inhibited by nicotine treatment, an effect that was most pronounced at 16 d postimmunization.

To further identify the subpopulation distribution among the infiltrating monocytes/activated macrophages, we also analyzed the proportions of proinflammatory CCR2+Ly6Chigh cells within the CNS. As shown in Fig. 2F, the number of cells in the CCR2+Ly6Chigh subpopulation within the brain and spinal cord started increasing at 7 d postimmunization, peaked at 16 d, followed by full recovery at 22 d. Intriguingly, CCR2+Ly6Chigh cell numbers were significantly reduced at 16 d postimmunization, but not significantly affected at other time points. In parallel, the numbers of all other infiltrating monocytes/activated macrophages (non-CCR2+Ly6Chigh) only increased significantly from 16 d onward and remained greater than normal until the 22nd day. Nicotine treatment did not alter the numbers of other monocyte phenotypes at any of the time points. These data show that nicotine specifically inhibits the recruitment of proinflammatory CCR2+Ly6Chigh, rather than all monocytes, into the CNS of EAE mice, especially at 16 d postimmunization, when their infiltration into the CNS and disease severity reached their maximum.

Infiltrating monocytes/macrophages of the CNS are derived from peripheral blood, whereas blood cells originate from BM. We thus also analyzed the monocyte cell numbers and the subpopulation variations in BM, blood, and spleen, after PBS or nicotine treatment (see Table II for specific values). As shown in Fig. 3A, the monocyte percentages in total BMCs began increasing at 3 d, peaked at 7 d, and remained greater than normal until 16 d postimmunization. Nicotine treatment had no effect on monocyte cell numbers compared with PBS-treated animals in BM. Similarly, Fig. 3B shows that the production of CCR2+Ly6Chigh from BM started increasing at 3 d and remained high until 12 d postimmunization, and was not altered by nicotine treatment. The total monocyte percentages in spleen (Fig. 3C) was increased at 7 and 12 d, and returned to normal at 16 d postimmunization. Nicotine treatment further increased monocyte percentages at 12 d postimmunization, but had no effect at other time points. Among monocytes in the spleen, CCR2+Ly6Chigh proportions (Fig. 3D) were increased from 3 until 16 d postimmunization, without significant differences in nicotine-treated groups.

Table II.
Myeloid cell percentages in BM, spleen, and blood
3 d7 d12 d16 d
% Monocytes in total BMCs 
 Naive 10.14 ± 0.51 10.14 ± 0.51 10.14 ± 0.51 10.14 ± 0.51 
 WT-EAE-PBS 15.10 ± 0.96 16.39 ± 1.19 12.84 ± 1.15 12.88 ± 0.39 
 WT-EAE-Nic 14.05 ± 0.72 16.00 ± 1.69 14.13 ± 1.34 12.01 ± 0.55 
% Monocytes in total spleen cells 
 Naive 2.03 ± 0.18 2.03 ± 0.18 2.03 ± 0.18 2.03 ± 0.18 
 WT-EAE-PBS 2.45 ± 0.21 2.91 ± 0.24 2.87 ± 0.30 2.48 ± 0.46 
 WT-EAE-Nic 3.02 ± 0.45 3.11 ± 0.43 4.50 ± 0.48 2.44 ± 0.39 
% Monocytes in total blood cells 
 Naive 8.16 ± 0.74 8.16 ± 0.74 8.16 ± 0.74 8.16 ± 0.74 
 WT-EAE-PBS 5.41 ± 1.61 8.49 ± 1.82 11.28 ± 2.10 13.48 ± 1.85 
 WT-EAE-Nic 4.52 ± 1.04 8.28 ± 5.91 9.23 ± 5.31 7.65 ± 1.69 
 α7KO-EAE-PBS — — — 18.39 ± 5.07 
 α7KO-EAE-Nic — — — 13.74 ± 3.77 
 α9KO-EAE-PBS — — — 12.62 ± 2.54 
 α9KO-EAE-Nic — — — 13.06 ± 3.35 
% CCR2+Ly6Chigh in total BMCs 
 Naive 4.75 ± 0.31 4.75 ± 0.31 4.75 ± 0.31 4.75 ± 0.31 
 WT-EAE-PBS 6.19 ± 0.78 8.60 ± 1.47 6.94 ± 0.70 5.30 ± 0.10 
 WT-EAE-Nic 6.65 ± 0.53 8.65 ± 1.42 7.11 ± 0.63 6.24 ± 0.29 
% CCR2+Ly6Chigh in total spleen cells 
 Naive 0.19 ± 0.04 0.19 ± 0.04 0.19 ± 0.04 0.19 ± 0.04 
 WT-EAE-PBS 0.54 ± 0.07 0.81 ± 0.15 0.75 ± 0.11 0.61 ± 0.19 
 WT-EAE-Nic 0.59 ± 0.14 0.96 ± 0.21 1.17 ± 0.22 0.47 ± 0.10 
% CCR2+Ly6Chigh in total blood cells 
 Naive 2.20 ± 0.50 2.20 ± 0.50 2.20 ± 0.50 2.20 ± 0.50 
 WT-EAE-PBS 1.20 ± 0.46 2.04 ± 0.65 2.80 ± 1.20 3.68 ± 0.67 
 WT-EAE-Nic 1.27 ± 0.52 2.33 ± 1.19 2.58 ± 0.90 1.83 ± 0.43 
 α7KO-EAE-PBS — — — 3.64 ± 0.89 
 α7KO-EAE-Nic — — — 3.56 ± 0.93 
 α9KO-EAE-PBS — — — 3.04 ± 1.00 
 α9KO-EAE-Nic — — — 2.44 ± 0.44 
% Neutrophils in total BMCs 
 Naive 34.12 ± 2.87 34.12 ± 2.87 34.12 ± 2.87 34.12 ± 2.87 
 WT-EAE-PBS 41.55 ± 2.83 52.83 ± 2.40 58.40 ± 2.43 58.62 ± 2.38 
 WT-EAE-Nic 47.96 ± 2.99 53.12 ± 1.69 60.02 ± 2.19 63.21 ± 2.42 
% Neutrophils in total spleen cells 
 Naive 0.76 ± 0.10 0.76 ± 0.10 0.76 ± 0.10 0.76 ± 0.10 
 WT-EAE-PBS 5.76 ± 1.46 6.84 ± 1.58 7.31 ± 1.06 4.48 ± 0.77 
 WT-EAE-Nic 4.45 ± 1.52 5.73 ± 0.88 9.33 ± 1.10 6.80 ± 1.28 
% Neutrophils in total blood cells 
 Naive 19.04 ± 2.92 19.04 ± 2.92 19.04 ± 2.92 19.04 ± 2.92 
 WT-EAE-PBS 36.74 ± 9.79 57.87 ± 6.30 76.10 ± 4.04 46.70 ± 10.12 
 WT-EAE-Nic 24.51 ± 8.45 47.60 ± 5.03 72.46 ± 7.48 68.45 ± 9.01 
3 d7 d12 d16 d
% Monocytes in total BMCs 
 Naive 10.14 ± 0.51 10.14 ± 0.51 10.14 ± 0.51 10.14 ± 0.51 
 WT-EAE-PBS 15.10 ± 0.96 16.39 ± 1.19 12.84 ± 1.15 12.88 ± 0.39 
 WT-EAE-Nic 14.05 ± 0.72 16.00 ± 1.69 14.13 ± 1.34 12.01 ± 0.55 
% Monocytes in total spleen cells 
 Naive 2.03 ± 0.18 2.03 ± 0.18 2.03 ± 0.18 2.03 ± 0.18 
 WT-EAE-PBS 2.45 ± 0.21 2.91 ± 0.24 2.87 ± 0.30 2.48 ± 0.46 
 WT-EAE-Nic 3.02 ± 0.45 3.11 ± 0.43 4.50 ± 0.48 2.44 ± 0.39 
% Monocytes in total blood cells 
 Naive 8.16 ± 0.74 8.16 ± 0.74 8.16 ± 0.74 8.16 ± 0.74 
 WT-EAE-PBS 5.41 ± 1.61 8.49 ± 1.82 11.28 ± 2.10 13.48 ± 1.85 
 WT-EAE-Nic 4.52 ± 1.04 8.28 ± 5.91 9.23 ± 5.31 7.65 ± 1.69 
 α7KO-EAE-PBS — — — 18.39 ± 5.07 
 α7KO-EAE-Nic — — — 13.74 ± 3.77 
 α9KO-EAE-PBS — — — 12.62 ± 2.54 
 α9KO-EAE-Nic — — — 13.06 ± 3.35 
% CCR2+Ly6Chigh in total BMCs 
 Naive 4.75 ± 0.31 4.75 ± 0.31 4.75 ± 0.31 4.75 ± 0.31 
 WT-EAE-PBS 6.19 ± 0.78 8.60 ± 1.47 6.94 ± 0.70 5.30 ± 0.10 
 WT-EAE-Nic 6.65 ± 0.53 8.65 ± 1.42 7.11 ± 0.63 6.24 ± 0.29 
% CCR2+Ly6Chigh in total spleen cells 
 Naive 0.19 ± 0.04 0.19 ± 0.04 0.19 ± 0.04 0.19 ± 0.04 
 WT-EAE-PBS 0.54 ± 0.07 0.81 ± 0.15 0.75 ± 0.11 0.61 ± 0.19 
 WT-EAE-Nic 0.59 ± 0.14 0.96 ± 0.21 1.17 ± 0.22 0.47 ± 0.10 
% CCR2+Ly6Chigh in total blood cells 
 Naive 2.20 ± 0.50 2.20 ± 0.50 2.20 ± 0.50 2.20 ± 0.50 
 WT-EAE-PBS 1.20 ± 0.46 2.04 ± 0.65 2.80 ± 1.20 3.68 ± 0.67 
 WT-EAE-Nic 1.27 ± 0.52 2.33 ± 1.19 2.58 ± 0.90 1.83 ± 0.43 
 α7KO-EAE-PBS — — — 3.64 ± 0.89 
 α7KO-EAE-Nic — — — 3.56 ± 0.93 
 α9KO-EAE-PBS — — — 3.04 ± 1.00 
 α9KO-EAE-Nic — — — 2.44 ± 0.44 
% Neutrophils in total BMCs 
 Naive 34.12 ± 2.87 34.12 ± 2.87 34.12 ± 2.87 34.12 ± 2.87 
 WT-EAE-PBS 41.55 ± 2.83 52.83 ± 2.40 58.40 ± 2.43 58.62 ± 2.38 
 WT-EAE-Nic 47.96 ± 2.99 53.12 ± 1.69 60.02 ± 2.19 63.21 ± 2.42 
% Neutrophils in total spleen cells 
 Naive 0.76 ± 0.10 0.76 ± 0.10 0.76 ± 0.10 0.76 ± 0.10 
 WT-EAE-PBS 5.76 ± 1.46 6.84 ± 1.58 7.31 ± 1.06 4.48 ± 0.77 
 WT-EAE-Nic 4.45 ± 1.52 5.73 ± 0.88 9.33 ± 1.10 6.80 ± 1.28 
% Neutrophils in total blood cells 
 Naive 19.04 ± 2.92 19.04 ± 2.92 19.04 ± 2.92 19.04 ± 2.92 
 WT-EAE-PBS 36.74 ± 9.79 57.87 ± 6.30 76.10 ± 4.04 46.70 ± 10.12 
 WT-EAE-Nic 24.51 ± 8.45 47.60 ± 5.03 72.46 ± 7.48 68.45 ± 9.01 

The average percentages of each subpopulation of myeloid cells with respect to the total number of cells in the BM, spleen, or blood in each group and time point are indicated ± SEM. Dashes indicate that no data were collected at these time points.

FIGURE 3.

Nicotine diminishes the numbers of proinflammatory monocytes in blood. On the indicated days postimmunization, BMCs, blood, and spleen were harvested from WT naive (white bars), WT-EAE-PBS (black bars), and WT-EAE-Nic (patterned bars) mice and analyzed by flow cytometry (n = 6–8 per group and time point). The same gating strategies were used to identify total monocytes (CD11b+Ly6G) and CCR2+Ly6Chigh monocytes, as described in Fig. 2. The percentage of total monocytes (A, C, and E) and CCR2+Ly6Chigh monocytes (B, D, and F) in BMCs (A and B), spleen (C and D), and blood (E and F) are represented in graphical format (means ± SEM). Notably, nicotine completely inhibits the EAE-induced increase in total monocytes and CCR2+Ly6Chigh monocytes in the blood on day 16 postimmunization, whereas the drug increases monocyte percentages in the spleen on day 12 postimmunization. Data shown are means ± SEM, whereas an asterisk denotes a significant difference between indicated groups (Mann–Whitney U test, *p < 0.05).

FIGURE 3.

Nicotine diminishes the numbers of proinflammatory monocytes in blood. On the indicated days postimmunization, BMCs, blood, and spleen were harvested from WT naive (white bars), WT-EAE-PBS (black bars), and WT-EAE-Nic (patterned bars) mice and analyzed by flow cytometry (n = 6–8 per group and time point). The same gating strategies were used to identify total monocytes (CD11b+Ly6G) and CCR2+Ly6Chigh monocytes, as described in Fig. 2. The percentage of total monocytes (A, C, and E) and CCR2+Ly6Chigh monocytes (B, D, and F) in BMCs (A and B), spleen (C and D), and blood (E and F) are represented in graphical format (means ± SEM). Notably, nicotine completely inhibits the EAE-induced increase in total monocytes and CCR2+Ly6Chigh monocytes in the blood on day 16 postimmunization, whereas the drug increases monocyte percentages in the spleen on day 12 postimmunization. Data shown are means ± SEM, whereas an asterisk denotes a significant difference between indicated groups (Mann–Whitney U test, *p < 0.05).

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Monocytes must circulate via the peripheral blood before infiltrating into the inflamed CNS. We found an increase in total monocytes in peripheral blood (Fig. 3E) of WT-EAE-PBS mice at 16 d, an effect that was prevented by nicotine treatment. There was no increase at other time points when comparing WT-EAE-PBS mice with naive animals, and nicotine did not have any effect on blood monocytes at these other time points. Similarly, the blood CCR2+Ly6Chigh subpopulation (Fig. 3F) significantly increased at 16 d postimmunization, whereas nicotine treatment completely reversed this effect. The CCR2+Ly6Chigh subpopulation did not change significantly at other time points, and again was not altered by nicotine treatment. Overall, these results suggest that nicotine can significantly alter the distribution of monocyte subpopulations in the blood of EAE mice.

The disease-protective and CCR2+Ly6Chigh monocyte distribution-altering effects of nicotine in EAE mice may be caused by the ligand acting on various nAChRs expressed by monocytes/macrophages (13, 14). To investigate the role of α7 nAChRs and α9 nAChRs in this cholinergic anti-inflammatory pathway, we induced EAE in α7KO and α9KO mice, and monocyte/macrophage numbers in the blood and CNS were analyzed 16 d postimmunization in the same manner as done in WT mice. As shown in Fig. 4A, the nicotine-dependent inhibition of total blood monocyte percentages was completely abrogated in both α7KO and α9KO mice. Similarly, Fig. 4B demonstrates that the inhibitory effect of nicotine on blood CCR2+Ly6Chigh monocyte percentages was also reversed in α7KO and α9KO mice. As depicted in Fig. 4C and 4D, the inhibitory effect of nicotine on total monocyte infiltration into the CNS was again no longer observed in α7KO or α9KO mice. More specifically, CCR2+Ly6Chigh monocyte infiltration was not affected by nicotine treatment in brain (Fig. 4E) or spinal cord (Fig. 4F) of α7KO and α9KO mice. The inhibitory effect of nicotine treatment on monocytes and CCR2+Ly6Chigh subpopulation infiltration thus appear to be mediated by both of these nAChR subtypes.

FIGURE 4.

Nicotine no longer inhibits the percentages of CCR2+Ly6Chigh monocytes in the blood or CNS of α7KO or α9KO mice. Naive (solid white bars), WT (black or white backgrounds), α7KO (dark gray backgrounds), or α9KO (light gray backgrounds) mice were immunized and treated with either PBS (solid bars) or nicotine (patterned bars) and sacrificed at day 16 postimmunization (n = 6–8 per group). The percentages of total monocytes (A, C, and D) and CCR2+Ly6Chigh monocytes (B, E, and F) in the blood (A and B), brain (C and E), and spinal cord (D and F) were significantly decreased in WT nicotine-treated EAE mice compared with PBS-treated EAE mice. These effects of nicotine were absent in α7KO and α9KO mice. Data shown are means ± SEM, whereas an asterisk denotes a significant difference between indicated groups (Mann–Whitney U test, *p < 0.05).

FIGURE 4.

Nicotine no longer inhibits the percentages of CCR2+Ly6Chigh monocytes in the blood or CNS of α7KO or α9KO mice. Naive (solid white bars), WT (black or white backgrounds), α7KO (dark gray backgrounds), or α9KO (light gray backgrounds) mice were immunized and treated with either PBS (solid bars) or nicotine (patterned bars) and sacrificed at day 16 postimmunization (n = 6–8 per group). The percentages of total monocytes (A, C, and D) and CCR2+Ly6Chigh monocytes (B, E, and F) in the blood (A and B), brain (C and E), and spinal cord (D and F) were significantly decreased in WT nicotine-treated EAE mice compared with PBS-treated EAE mice. These effects of nicotine were absent in α7KO and α9KO mice. Data shown are means ± SEM, whereas an asterisk denotes a significant difference between indicated groups (Mann–Whitney U test, *p < 0.05).

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Apart from monocytes, the infiltration of neutrophils into the CNS also contributes to disease progression during EAE (3, 4). As shown in Fig. 5A, flow cytometric analysis was performed on the CD11b+Ly6G+ (neutrophil) population to measure the proportions of circulating and infiltrating neutrophils in different tissues. The proportions of neutrophils (Fig. 5B) in the BM began increasing at 3 d and remained greater than normal until 16 d postimmunization, whereas this effect was not affected by nicotine treatment. The accumulation of neutrophils in the spleen (Fig. 5C) and peripheral blood (Fig. 5D) started from 3 d and remained at higher levels until 16 d. Nicotine treatment did not alter neutrophil levels in the spleen or blood of EAE mice at any given time point. Comparatively, neutrophil infiltration into the brain (Fig. 5E) and spinal cord (Fig. 5F) increased at 12 d, remained high at 16 d, and slightly declined at 22 d. Nicotine treatment inhibited the infiltration of neutrophils into the CNS at 16 d only. Once again, the effects of nicotine on neutrophil percentages in the brain (Fig. 5G) and spinal cord (Fig. 5H) at 16 d postimmunization were reversed in α7KO and α9KO mice. These data demonstrate that nicotine can also significantly reduce the infiltration of neutrophils into the CNS of EAE mice in a α7- and α9-dependent fashion.

FIGURE 5.

Nicotine inhibits the recruitment of neutrophils into the CNS of WT but not α7KO or α9KO mice 16 d postimmunization. (A) Neutrophils were identified based on their expression of CD11b and Ly6G (CD11b+Ly6G+, top right quadrant). The percentage of BM (B), spleen (C), blood (D), brain (E and G), and spinal cord (F and H) cells that were neutrophils in WT (black or white backgrounds), α7KO (dark gray backgrounds), or α9KO (light gray backgrounds) treated with either PBS (solid bars) or nicotine (patterned bars) are depicted in graphical format (n = 6–8 per group and time point). The percentages of neutrophils increase in all tissues at nearly all time points postimmunization in all PBS-treated animals. Although nicotine prevents the increase in the brain and CNS of WT mice at the 16-d time point, it fails to do so in α7KO or α9KO mice. Data shown are means ± SEM, whereas an asterisk denotes a significant difference between indicated groups (Mann–Whitney U test, *p < 0.05).

FIGURE 5.

Nicotine inhibits the recruitment of neutrophils into the CNS of WT but not α7KO or α9KO mice 16 d postimmunization. (A) Neutrophils were identified based on their expression of CD11b and Ly6G (CD11b+Ly6G+, top right quadrant). The percentage of BM (B), spleen (C), blood (D), brain (E and G), and spinal cord (F and H) cells that were neutrophils in WT (black or white backgrounds), α7KO (dark gray backgrounds), or α9KO (light gray backgrounds) treated with either PBS (solid bars) or nicotine (patterned bars) are depicted in graphical format (n = 6–8 per group and time point). The percentages of neutrophils increase in all tissues at nearly all time points postimmunization in all PBS-treated animals. Although nicotine prevents the increase in the brain and CNS of WT mice at the 16-d time point, it fails to do so in α7KO or α9KO mice. Data shown are means ± SEM, whereas an asterisk denotes a significant difference between indicated groups (Mann–Whitney U test, *p < 0.05).

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We next assessed whether the lower numbers of blood-derived proinflammatory monocytes and neutrophils in the CNS of nicotine-treated EAE mice 16 d postimmunization could be caused by the death of these cells before their infiltration. We found that the number of 7-AAD+ monocytes at 14 d postimmunization (Fig. 6A) was similar in PBS-treated and nicotine-treated EAE mice (10.5 ± 2.3 versus 11.1 ± 2.3% in WT-EAE-PBS versus WT-EAE-Nic, p > 0.05). Nicotine also did not affect the viability of CCR2+Ly6Chigh monocytes (21.1 ± 5.7 versus 15.38 ± 2.9% in WT-EAE-PBS versus WT-EAE-Nic, p > 0.05; Fig. 6B) or neutrophils (3.4 ± 1.1 versus 2.6 ± 0.4% in WT-EAE-PBS versus WT-EAE-Nic, p > 0.05; Fig. 6C).

FIGURE 6.

Migratory capabilities of BM-derived monocytes are inhibited by nicotine. (AC) 7-AAD+ cells in the total monocyte population (A), CCR2+Ly6Chigh monocytes (B), and neutrophils (C) were quantified by flow cytometry. (D and E) The number of migratory M-CSF and IFN-γ–induced BM-derived monocytes (D) and neutrophils (E) were assessed by transwell migration assays. (F and G) VLA-4high expression in total monocytes (F) and CCR2+Ly6Chigh monocytes (G), and (H and I) LFA-1high expression in total monocytes (H) and CCR2+Ly6Chigh monocytes (I) were assessed by flow cytometry. Although nicotine was able to directly inhibit monocyte migratory capabilities in vitro, these data suggest that nicotine does not reduce CNS monocyte or neutrophil numbers by inducing the death of blood monocytes or neutrophils before their infiltration or by reducing their expression of key integrins. Data shown are means ± SEM, whereas an asterisk denotes a significant difference between indicated groups (Mann–Whitney U test, *p < 0.05).

FIGURE 6.

Migratory capabilities of BM-derived monocytes are inhibited by nicotine. (AC) 7-AAD+ cells in the total monocyte population (A), CCR2+Ly6Chigh monocytes (B), and neutrophils (C) were quantified by flow cytometry. (D and E) The number of migratory M-CSF and IFN-γ–induced BM-derived monocytes (D) and neutrophils (E) were assessed by transwell migration assays. (F and G) VLA-4high expression in total monocytes (F) and CCR2+Ly6Chigh monocytes (G), and (H and I) LFA-1high expression in total monocytes (H) and CCR2+Ly6Chigh monocytes (I) were assessed by flow cytometry. Although nicotine was able to directly inhibit monocyte migratory capabilities in vitro, these data suggest that nicotine does not reduce CNS monocyte or neutrophil numbers by inducing the death of blood monocytes or neutrophils before their infiltration or by reducing their expression of key integrins. Data shown are means ± SEM, whereas an asterisk denotes a significant difference between indicated groups (Mann–Whitney U test, *p < 0.05).

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The recruitment of monocytes and neutrophils from the periphery into the CNS relies on multiple cellular mechanisms, such as the expression of adhesion molecules, metalloproteinases, and the production of chemokines from the sites of inflammation. For instance, the expression of the integrins VLA-4 and LFA-1 by leukocytes, and their binding partners VCAM-1 and ICAM-1 on endothelial cells, respectively, are integral for leukocyte migration across the blood vessel wall. The chemokine CCL2 is involved in the recruitment of the proinflammatory CCR2+Ly6Chigh monocyte subpopulation (18), whereas CX3CL1 plays a role in the recruitment of CX3CR1+ monocyte subpopulations (19). In parallel, the chemokine CXCL2 is one of the chemokines responsible for the recruitment of neutrophils into the CNS (20). MMP-9 is an extracellular matrix-degrading enzyme also known to be involved in diapedesis and important for EAE pathology (21).

We thus investigated whether the migratory ability of monocytes and neutrophils was affected by nicotine using transwell migration assays (Fig. 6D, 6E), and found that nicotine significantly reduced CCL2-induced monocyte migration by 32.7 ± 8.2% (p < 0.05). In contrast, neutrophil migration in response to IL-8 was unaffected by nicotine (p > 0.05). We then assessed the expression of leukocyte integrins VLA-4 and LFA-1 on the total population of blood monocytes, as well as specifically in CCR2+Ly6Chigh monocytes 14 d postimmunization (Fig. 6F–I). We found that nicotine failed to alter VLA-4 expression in total (6.1 ± 1.4 versus 5.8 ± 1.9% in WT-EAE-PBS versus WT-EAE-Nic; Fig. 6F) and CCR2+Ly6Chigh (11.7 ± 3.2 versus 10.2 ± 3.0% in WT-EAE-PBS versus WT-EAE-Nic; Fig. 6G) monocytes. Similarly, LFA-1 expression was also unchanged in total (23.2 ± 2.0 versus 25.7 ± 3.4% in WT-EAE-PBS versus WT-EAE-Nic; Fig. 6H) or CCR2+Ly6Chigh (6.9 ± 0.6 versus 6.7 ± 1.7% in WT-EAE-PBS versus WT-EAE-Nic; Fig. 6I) monocytes. This suggests that nicotine affects monocyte and neutrophil infiltration into the CNS by mechanisms other than the modulation of leukocyte integrin expression.

We next determined whether nicotine could alter monocyte and neutrophil infiltration into the CNS by regulating the expression of VCAM-1 and ICAM-1, the receptors for VLA-4 and LFA-1, in the brain of WT mice 7, 12, or 16 d postimmunization. Again, we found that nicotine did not inhibit the relative mRNA expression of VCAM-1 (3.5 ± 0.3 versus 3.8 ± 1.3, 1.4 ± 0.2 versus 1.8 ± 0.2, 1.7 ± 0.3 versus 1.7 ± 0.4 relative expression for WT-EAE-PBS versus WT-EAE-Nic at 7, 12, and 16 d postimmunization, respectively; p > 0.05 at each time point; Fig. 7A) or ICAM-1 (4.4 ± 1.2 versus 4.0 ± 0.3, 3.3 ± 0.4 versus 3.1 ± 0.3, and 5.3 ± 1.9 versus 5.3 ± 1.9 relative expression for WT-EAE-PBS versus WT-EAE-Nic at 7, 12, and 16 d postimmunization, respectively; p > 0.05 at each time point; Fig. 7B) at any of the time points studied for either transcript. Likewise, we measured the effects of nicotine on the mRNA expression of MMP-9 within the CNS and found that transcript levels of the metalloproteinase (Fig. 7C) were increased at 7 d (0.1 ± 0.4 versus 1.0 ± 0.6 relative expression, for naive versus PBS-treated EAE, p < 0.05), 12 d (0.1 ± 0.04 versus 1.5 ± 0.3 relative expression, for naive versus PBS-treated EAE, p < 0.05), and 16 d (0.1 ± 0.04 versus 1.2 ± 0.6 relative expression, for naive versus PBS-treated EAE, p < 0.05) postimmunization; however, nicotine was unable to inhibit the expression of this enzyme at any time point (0.8 ± 0.3, 1.4 ± 0.5, and 1.6 ± 0.3 relative expression at 7, 12, and 16 d postimmunization, respective; p > 0.05). In contrast, the transcript levels of CCL2 (Fig. 7D) were increased at 12 d (1.6 ± 0.9 versus 9.0 ± 1.6 relative expression, for naive versus PBS-treated EAE, p < 0.05) and remained greater than normal at 16 d postimmunization (1.6 ± 0.9 versus 8.8 ± 3.5 relative expression, for naive versus PBS-treated EAE, p < 0.05). Nicotine treatment inhibited the increase in transcript levels of CCL2 in the brain at 12 d postimmunization (9.0 ± 1.6 versus 3.5 ± 0.5 relative expression, for PBS- versus nicotine-treated EAE, p < 0.05), without effect on any other time points (1.7 ± 0.6 and 6.9 ± 3.4 relative expression for 7 and 16 d, respectively; p > 0.05). For CX3CL1 (Fig. 7E), there was no significant change in PBS-treated EAE mice compared with naive animals at 7, 12, or 16 d (1.3 ± 1.0 relative expression for naive mice, compared with 1.6 ± 0.2, 0.6 ± 0.1, or 0.9 ± 0.1 relative expression for PBS-treated EAE mice at 7, 12, or 16 d, respectively; p > 0.05). However, nicotine treatment increased the transcription of CX3CL1 at 7 d postimmunization (1.6 ± 0.2 versus 2.2 ± 0.3 relative expression for PBS- versus nicotine-treated EAE, p < 0.05), without any effect at other time points (0.7 ± 0.2 and 0.8 ± 0.1 relative expression for 12 and 16 d, respectively; p > 0.05 when compared with PBS-treated animals, respectively). As depicted in Fig. 7F, there was also an increase in mRNA levels for CXCL2 at 12 d (0.4 ± 0.2 versus 6.2 ± 1.7 relative expression for naive versus PBS-treated EAE, p < 0.05) and 16 d (0.4 ± 0.2 versus 2.4 ± 1.07 relative expression for naive versus PBS-treated EAE, p < 0.05) postimmunization, whereas nicotine treatment inhibited the increased transcription levels at 12 d (6.2 ± 1.7 versus 2.3 ± 0.6 relative expression for PBS-treated versus nicotine-treated EAE, p < 0.05), without effect at other time points (1.2 ± 0.7 versus 0.4 ± 0.1 and 2.4 ± 1.1 versus 2.9 ± 0.7 relative expression for PBS-treated versus nicotine-treated EAE on 7 and 16 d; p > 0.05). Overall, these data indicate that nicotine influences CCL2, CX3CL1, and CXCL2 mRNA expression in the brain, whereas the molecule has no effect on the expression of integrins and metalloproteinases, all of which are crucial for monocyte and neutrophil recruitment during EAE.

FIGURE 7.

Nicotine modulates the mRNA expression levels of chemokines involved in monocyte and neutrophil chemotaxis. Relative mRNA expression levels for the leukocyte integrin receptors VCAM-1 (A) and ICAM-1 (B), MMP-9 (C), monocyte-attracting chemokines CCL2 (D) and CX3CL1 (E), and the neutrophil-attracting chemokine CXCL2 (F) in the brain were analyzed by RT-qPCR. Nicotine inhibited the increase in CCL2 and CXCL2 expression levels at 12 d postimmunization, whereas the drug increased the expression of CX3CL1 and had no effect on VCAM-1, ICAM-1, or MMP-9 transcript levels. Data shown are means ± SEM, whereas an asterisk denotes a significant difference between indicated groups (Mann–Whitney U test, *p < 0.05).

FIGURE 7.

Nicotine modulates the mRNA expression levels of chemokines involved in monocyte and neutrophil chemotaxis. Relative mRNA expression levels for the leukocyte integrin receptors VCAM-1 (A) and ICAM-1 (B), MMP-9 (C), monocyte-attracting chemokines CCL2 (D) and CX3CL1 (E), and the neutrophil-attracting chemokine CXCL2 (F) in the brain were analyzed by RT-qPCR. Nicotine inhibited the increase in CCL2 and CXCL2 expression levels at 12 d postimmunization, whereas the drug increased the expression of CX3CL1 and had no effect on VCAM-1, ICAM-1, or MMP-9 transcript levels. Data shown are means ± SEM, whereas an asterisk denotes a significant difference between indicated groups (Mann–Whitney U test, *p < 0.05).

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Although smoking is a risk factor, nicotine without the many other chemicals found in tobacco smoke has been shown to have an anti-inflammatory effect and to protect against inflammatory diseases, such as EAE (1113). Nicotine can inhibit the proliferation of lymphocytes in peripheral immune compartments (1113), as well as the overall infiltration of leukocytes into the CNS (11, 13). Among leukocytes, myeloid cells play an important role in EAE initiation and progression (22). However, the effect of nicotine treatment on the development, migration, and distribution of myeloid cell subpopulations, especially proinflammatory M1 monocytes and neutrophils, during the course of EAE is not clear. The main goal of this study was to evaluate the impact of nicotine on the CNS infiltration of CD11b+CD45+ cell subsets, including infiltrating monocytes/macrophages (Ly6GCD11b+CD45high), with a particular focus on proinflammatory M1 monocytes (CCR2+Ly6Chigh), as well as microglia (Ly6GCD11b+CD45med) and neutrophils (Ly6G+CD11b+CD45+), which all play a prominent role in EAE pathology. After confirming our hypothesis that nicotine selectively inhibits proinflammatory cell recruitment to the CNS of EAE mice, our secondary goals were then to assess the role of α7 and α9 nAChR subtypes, and to determine whether nicotine could achieve this by modulating cell viability, migration capability, or chemokine expression.

We first assessed the clinical scores in EAE mice of WT, α7KO, or α9KO background pretreated with nicotine or PBS. In support of previous studies (1113), the average clinical scores during the peak stages of disease and the cumulative scores throughout the time course were reduced, whereas disease onset was delayed, in WT-EAE-Nic mice. In addition, nicotine no longer affected disease course in α7KO mice, thus confirming once again that nicotine has protective properties against EAE, and that the α7 nAChR subtype is an important target for this ligand. However, recent evidence suggested that additional nAChR subtypes may be involved (13), and it was later shown that α9 nAChRs are involved in endogenous mechanisms that regulate EAE pathology, presumably via acetylcholine or choline (14). Our data support this hypothesis, because disease severity (average and cumulative scores) were significantly reduced in α9KO mice, regardless of nicotine or PBS treatment.

We then proceeded to analyze the proportions of myeloid cells in the brain and spinal cord of nicotine- or PBS-treated EAE mice at key stages of disease development. We first found that, similar to a previous study (8), total monocytic cell numbers in the brain and spinal cord (CD11b+CD45+Ly6G) began to increase 12 d postimmunization and peaked at 16 d, coinciding with disease onset and peak severity, respectively. The overall increase in monocytic cells in the CNS of EAE mice was significantly reduced by nicotine treatment, as expected (11, 13). We then determined which subpopulation of monocytic cells was affected by nicotine, because a reduction in total monocytic cell numbers could be explained by a decrease in microglia or blood-derived monocyte/macrophages. Interestingly, we found that blood-derived monocyte/macrophages were the predominant cell type that contributed to the higher total monocytic cell numbers in EAE, and that nicotine prevented their increase, while not affecting the presence of microglia. In turn, CCR2+Ly6Chigh cell proportions in the CNS are known to be positively correlated with EAE clinical scores (8). We thus assessed their numbers and confirmed that CCR2+Ly6Chigh cells are major contributors to the increased infiltrating monocytes/macrophages observed in the CNS during disease development and, more importantly, that nicotine specifically inhibited the recruitment of these proinflammatory cells while not affecting the numbers of other monocytic subpopulations. Overall, our data demonstrate that nicotine specifically inhibits the recruitment of proinflammatory monocytes/macrophages into the CNS of EAE mice. Because these cells are major players in EAE pathology, our data thus provide a concrete explanation for the disease-modifying properties of this nAChR ligand.

The determinant role of neutrophils in the early stages of EAE pathology is becoming increasingly clear. Neutrophils expand in the BM, spleen, and blood within a few days after immunization (23). These cells also begin to infiltrate the CNS before the onset of EAE clinical symptoms, and evidence suggest that they play a crucial role in the disruption of the blood–brain and blood–spinal cord barriers (3). In this study, we confirm that neutrophil numbers increase rapidly in the preclinical stages of EAE in all tissues examined, and that their maximal numbers in the brain coincide with peak disease severity at the 16-d time point. Of greater significance, however, is the novel finding that nicotine treatment reduces the numbers of neutrophils at 16 d, both in the brain and in the spinal cord. It is thus possible that nicotine treatment prevents the disruption of the blood–brain and blood–spinal cord barriers by limiting neutrophil numbers and functions; however, further studies will be required to assess this hypothesis.

Multiple mechanisms could be involved in the reduction in the recruitment of CCR2+Ly6Chigh monocytes and neutrophils into the CNS. First, nicotine could be lowering the numbers of these cells in the blood, thereby diminishing the pool of cells that are able to enter the CNS. This could occur by a diminished production of these cells in the BM, or by their sequestration in the BM or spleen. Second, nicotine could be modulating the expression of CCL2 and CXCL2, the chemokines responsible for the chemotaxis of CCR2+Ly6Chigh monocytes and neutrophils, respectively (18, 20). Third, nicotine could be directly compromising the ability of these cells to migrate across the blood–brain barrier, perhaps by inhibiting the expression of key proteins such as the metalloproteinase MMP-9 or adhesion molecules such as VLA-4, LFA-1, ICAM-1, and VCAM-1. We thus assessed all of these possibilities.

First, our data show that nicotine inhibits the increase in the proportions of CCR2+Ly6Chigh cells, but not neutrophils, normally observed in the blood of EAE mice (23). This finding supports the idea that nicotine decreases CCR2+Ly6Chigh cell numbers in the CNS by diminishing the pool of these cells in the blood; however, this does not appear to be the mechanism responsible for the lower numbers of neutrophils in the CNS. In addition, nicotine thus does not appear to reduce proinflammatory monocyte or neutrophil numbers in the blood by inducing the death of these cells, because their viability was unchanged in the blood of EAE mice.

We thus assessed whether nicotine could directly inhibit the migratory capabilities of monocytes and neutrophils. Indeed, treatment with the molecule inhibited the migration of proinflammatory BM-derived monocytes, induced by M-CSF and IFN-γ (24), in transwell migration assays. Surprisingly, however, in vivo nicotine treatment did not alter the expression of VLA-4 and LFA-1 on blood monocytes, two integrins that are necessary for leukocyte migration across the blood vessel cell wall. It is therefore possible that nicotine-treated proinflammatory monocytes are less able to respond to CCL2, or that other mechanisms are responsible for their reduced migratory capability. In contrast, nicotine had no effect on neutrophil migration in vitro, thus suggesting that other indirect mechanisms must explain the reduction in CNS neutrophils.

We therefore investigated whether changes occurred in the CNS, such as altered expression of the integrin receptors VCAM-1 and ICAM-1, the metalloproteinase MMP-9, or the expression of monocyte (CCL2 and CX3CL1) and neutrophil (CXCL2)-attracting chemokines. Our results demonstrate that nicotine inhibits the expression of CCL2 and CXCL2 in the brain of EAE mice at 12 d postimmunization, coinciding with the time at which proinflammatory monocytes and neutrophils begin their massive infiltration into the CNS. Interestingly, we also found that nicotine increased the expression of CX3CL1, a chemokine that can attract M2 monocytes; however, it is not clear whether these cells were increased in the CNS of our mice. Finally, nicotine did not affect the expression levels of VCAM-1, ICAM-1, or MMP-9 in the brain at any time point. Overall, our data suggest that nicotine mainly affects leukocyte migration into the CNS by inhibiting the expression of chemokines responsible for proinflammatory monocyte and neutrophil chemotaxis. Although more studies are required to determine whether other mechanisms may also be involved, our data provide important insight into the mechanisms by which nicotine modulates proinflammatory cell recruitment into the CNS of EAE mice, thus altering the course of disease.

A previous study demonstrated that nicotine can affect EAE outcome to various extents depending on the timing of drug administration (11). EAE time of onset and disease severity are reduced, whereas recovery is ameliorated when nicotine is administered before or concomitantly with MOG immunization. In contrast, time of onset is not affected when nicotine treatment begins 7 d postimmunization. However, all three treatment paradigms are equally effective in reducing disease severity and increasing recovery. Shi et al. (11) also showed that nicotine treatment during the first 7 d postimmunization reduced leukocyte numbers in the CNS during peak disease. In this study, animals were pretreated 2 d before MOG immunization, and EAE clinical scores and CNS leukocyte numbers are in accordance with the previous data. Because a pretreatment was used, we cannot exclude the possibility that our results are due to the general inhibition of disease induction mechanisms in a pathologically important target tissue. Nonetheless, the EAE-induced expansion of monocytes and neutrophils in the BM and spleen was not affected by nicotine. In contrast, nicotine pretreatment specifically affected these cells in the blood and CNS precisely at the time points where they reach their maximum numbers. Overall, the previous study mentioned (11) and our present findings suggest that the observed effects of nicotine on disease course and pathology are due to specific pathological mechanisms, such as those involved in the regulation of cell migration into the CNS, and are not due to the general reduction of disease initiation.

Finally, we also assessed the role of α7 and α9 nAChRs in the modulation of CCR2+Ly6Chigh monocyte and neutrophil numbers in the blood and CNS of EAE mice. Our data show that nicotine was no longer able to reduce cell numbers in either α7KO or α9KO mice, confirming a role for both of these nAChR subtypes in cholinergic modulation of proinflammatory cell recruitment into the CNS. This finding supports the established notion that α7 nAChRs are indeed a major target of nicotine in mediating inflammation (25), but also reinforces previous findings that other nAChRs, including the α9 subtype, may also play a determinant role in the cholinergic anti-inflammatory pathway (13, 14). Surprisingly, however, we did not observe any changes in myeloid cell numbers in the blood or CNS of α9KO mice compared with WT mice at the 16-d time point. This is contrary to previous findings, which showed that total myeloid cell numbers were increased in the brain of α9KO mice 7 d postimmunization (14). This may be explained by the difference in the time points between the two studies; in this study, we only assessed myeloid cell numbers 16 d postimmunization because this was the only time point where a nicotinic effect was observed in these tissues. Moreover, the previous study did not assess CCR2+Ly6Chigh cells or neutrophils, whereas these cells were the focus of this study. Nonetheless, our findings add important information regarding the role of nAChRs in the mechanisms of nicotinic protection against disease and suggest that the reduced clinical symptoms observed in α9KO mice are not due to changes in proinflammatory monocyte or neutrophil infiltration into the CNS, and instead are explained by other α9 nAChR-dependent immune-modifying mechanisms. More studies will be required to shed light on this issue.

In summary, this study provides evidence that nicotine alters the infiltration of proinflammatory monocytes and neutrophils into the CNS of EAE mice via multiple nAChRs, including the α7 and α9 subtypes. Nicotine appears to achieve these effects by inhibiting the expression of CCL2 and CXCL2, two cytokines involved in the chemotaxis of proinflammatory monocytes and neutrophils, respectively. The use of ligands that are selective for one or both of these nAChR subtypes may offer a beneficial clinical outcome, and thus provide a valuable therapeutic strategy for neuroinflammatory disorders such as MS.

We acknowledge Marc Surette (Université de Moncton) for kindly sharing laboratory equipment, as well as Anick Beaulieu (Université de Moncton) and Crystal Morrison (Université de Moncton) for technical expertise.

This work was supported by grants from the Multiple Sclerosis Society of Canada (to A.R.S.), the New Brunswick Health Research Foundation (to A.R.S.), the New Brunswick Innovation Foundation (to A.R.S.), the Nebraska Tobacco Settlement Biomedical Research Fund (to B.J.M.), and the National Institutes of Health (Grant R01DC006907 to B.J.M.). Salary support was provided by the Centre de Formation Médicale du Nouveau-Brunswick (to W.J.) and the New Brunswick Innovation Foundation (to S.S-P. and P.R.).

Abbreviations used in this article:

7-AAD

7-aminoactinomycin D

BM

bone marrow

BMC

BM cell

EAE

experimental autoimmune encephalomyelitis

KO

knockout

α7KO

α7 nAChR KO

α9KO

α9 nAChR KO

α7KO-EAE-PBS

PBS-treated α7KO EAE

α9KO-EAE-PBS

PBS-treated α9KO EAE

MMP-9

matrix metalloproteinase-9

MOG

myelin oligodendrocyte glycoprotein

MS

multiple sclerosis

nAChR

nicotinic acetylcholine receptor

WT

wild type

WT-EAE-Nic

nicotine-treated WT EAE

WT-EAE-PBS

PBS-treated WT EAE.

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The authors have no financial conflicts of interest.

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