Vagal Afferent Fibers Contribute to the Anti-Inflammatory Reactions by Vagus Nerve Stimulation in Concanavalin A Model of Hepatitis in Rats

doi:10.21203/rs.3.rs-46092/v1

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Research article

Vagal Afferent Fibers Contribute to the Anti-Inflammatory Reactions by Vagus Nerve Stimulation in Concanavalin A Model of Hepatitis in Rats

https://doi.org/10.21203/rs.3.rs-46092/v1

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published 03 Dec, 2020

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Background: Increasing number of studies provide evidence that the vagus nerve stimulation (VNS) dampens inflammation in peripheral visceral organs. However, the effects of afferent fibers of the vagus nerve (AFVN) on anti-inflammation have not been clearly defined. Here, we investigate whether AFVN are involved in VNS-mediated regulation of hepatic production of proinflammatory cytokines.

Methods: An animal model of hepatitis was generated by intraperitoneal injection of concanavalin A (ConA) into rats, and electrical stimulation was given to the hepatic branch of the vagus nerve. AFVN was selectively eliminated by treating animals with capsaicin (CAP). mRNA and protein expression in target tissues was analyzed by RT-PCR, real-time PCR, western blotting and immunofluorescence staining.

Results: TNF-α, IL-1β, and IL-6 mRNAs and proteins that were induced by ConA in the liver macrophages were significantly reduced by the electrical stimulation of the hepatic branch of the vagus nerve (hVNS). hVNS also induced an increase in phospho-STAT3 in the liver macrophages of ConA-injected animals, suggesting the activation of STAT3 transcription factor associated with cholinergic anti-inflammation. Capsaicin (CAP) treatment deteriorated transient receptor potential vanilloid 1 (TRPV1)-positive neurons and also induced an increase in apoptotic caspase-3 signals in nodose ganglion (NG) neurons. Concomitantly, CAP suppressed choline acetyltransferase (ChAT) expression that was induced by hVNS in DMV neurons of ConA-injected animals. Furthermore, hVNS-mediated downregulation of TNF-α, IL-1β, and IL-6 expression was reduced by CAP administration.

Conclusions: Our data indicate that the activity of AFVN contributes to hepatic anti-inflammatory responses mediated by hVNS in ConA model of hepatitis in rats.

Internal Medicine

vagus nerve stimulation

anti-inflammation

choline acetyltransferase

capsaicin cholinergic anti-inflammation

liver

Vagus nerve is the major parasympathetic nerve connecting the brain to the cardiopulmonary and most of the visceral organs. In addition to its physiological regulatory function, accumulating evidence shows that the vagus nerve activity is involved in mediating neuroimmune interaction (Reardon et al. 2018). The vagus nerve transmits neural activity originated from the central nervous system into the innate and adaptive immune cells in the visceral organs and thus regulates their pathophysiological function (Tracey, 2007; Bonaz et al. 2019). Studies using experimental animals have revealed that the VNS attenuates inflammation in peripheral organs such as spleen and gastrointestinal tracts, and the principles of cholinergic anti-inflammatory pathway in macrophages are widely taken as a mechanistic basis explaining anti-inflammation. For instance, VNS induces JAK2-STAT3 pathway associated with the activation of nicotinic acetylcholine receptor α7 and inhibits the production of pro-inflammatory cytokines such as TNF-α (Wang et al. 2003; de Jonge et al. 2005).

ConA is a lectin that binds to sugar group of glycoproteins on T lymphocytes. In the liver, Con-A bound T cells interact with antigen-presenting cells such as Kupffer cells via MHC-II complex, and consequently, both T cells and Kupffer cells produce inflammatory cytokines and lead to inflammation or cell death in the liver tissue (Tiegs, 1997; Krenkel & Tacke, 2017). Previous studies have reported that vagus nerve activity is involved in regulating hepatic ischemia/reperfusion injury, hepatocyte apoptosis, portal hypertension, inflammatory responses of Kupffer cells, phagocytic activity of macrophages, and oxidative damages in the liver tissues in animal models of liver disease (Zhang et al. 2019a; Özdemir-Kumral et al. 2017; Bockx et al. 2012; Hiramoto et al. 2008; Li et al. 2014; Nishio et al. 2017; Fonseca et al. 2019; Metz & Pavlov, 2018). It was further shown that T lymphocytes play a role in modulating the inflammatory responses that are controlled by sympathetic and parasympathetic nerve activities (Wong et al. 2011; Rosas-Ballina et al. 2011), raising the possibility that T lymphocytes activated in ConA-induced hepatitis may be involved in regulating inflammatory responses by VNS. However, the effects of VNS on hepatic inflammation in ConA-injected animals are not known yet.

Given the abundance of afferent fibers accounting 70–80% of the vagus nerve (Prechtl & Powley, 1990), VNS applied to cervically exposed nerve may increase the activity of afferent fibers and affect anti-inflammatory responses via contralateral efferent vagus nerve (Inoue et al. 2016). According to cholinergic anti-inflammatory reflex theory, pro-inflammatory cytokines produced from peripheral organs stimulate afferent sensory nerve and consequently augmenting vagal efferent outputs may downregulate inflammatory response (Tracey, 2002). However, the contribution of AFVN to VNS-induced anti-inflammation has not been clearly demonstrated in peripheral organs with vagal innervation. To investigate the effects of AFVN on the regulation of inflammatory responses, here we selectively eliminated AFVN by treating with CAP in concanavalin A (ConA)-model of hepatitis in rats. Our study presents new evidence that hVNS-induced activation of AFVN contributes to attenuating the hepatic production of pro-inflammatory cytokines.

Experimental Animals and Ethical Approval

Sprague-Dawley rats (male, 7–8 weeks old, 200–250 g) were purchased (Samtako Inc. Seoul, Korea, RRID:RGD:737903) and maintained in ventilated animal room with regulated temperature (22-23°C), 60% humidity under a standard 12 hour light and 12 hour dark cycle (light on from7:00 am to 7:00 pm). Animals were freely accessed to food pellets (Samyang Co., Seoul, Korea) and water. Total number of animals used for individual experiments are listed in Table 1. No experimental animals died during the experiments and any animals were not excluded in this study from analyses. All animal care procedures were approved by the Committee on Use of Live Animals for Teaching and Research at Daejeon University (approval number: DJUARB2019-029). Animal care and all experimental procedures were in accordance with the NIH Guide for the Care and Use of Laboratory Animals and also approved by the Committee on Use of Live Animals for Teaching and Research at Daejeon University (Daejeon, Korea). The authors complied with the ethical principles as outlined in Grundy (2015)

Table 1

Number of animals and experimental groups
Experiments	Animal groups	Real-time PCR / RT PCR	Western blotting	Immunofluorescence staining
VNS experiment	Untreated CTL	5
	ConA + Sham	4
	ConA + hVNS	4
	hVNS	4
	2d ConA + Sham		5
	2d ConA + hVNS		5
	3d ConA + Sham		5
	3d ConA + hVNS		5
	4d ConA + Sham		5
	4d ConA + hVNS		5
	cVNS			4
	Sham (cVNS)			4
CAP experiment	Vehicle		6
	CAP-L & H		7
	Vehicle + hVNS		5
	Vehicle + Con A		5
	Vehicle + Con A + hVNS		5
	Cap + hVNS		5
	Cap + Con A		5
	Cap + Con A + hVNS		5
Retrograde tracing	DiI injection	2

Drug administration

Rats were anesthetized by inhalation of isoflurane (2%; Hana Pharm Co., Ltd., Seoul, Korea) during CAP or vehicle injection. In order to minimize pain caused by CAP, 6 ml of atropine sulfate (0.5 mg/ml, Jeil Pharmaceutical Co., Daegu, Korea) was injected i.p., 20-30 min prior to CAP injection. CAP (Cayman Chemical Co., Ann Arbor, MI, USA) was dissolved in 10% ethanol, 10% Tween-80 and 0.9% NaCl and was injected i.p. with 40 mg/kg and supplemented with 80 mg/kg twice at 6 h and 24 h after the initial injection as a high-dose regimen (CAP-H) or injected initial 20 mg and supplemented 40 mg/kg twice at 6 h and 24 h later as a low-dose regimen (CAP-L) (Czaja et al. 2008). Volumes of CAP injected to individual animals were evenly adjusted as 1 ml per kg body weight. Equivalent volume of vehicle solution (VEH, 10% ethanol, 10% Tween-80 and 0.9% NaCl) was injected into the control animals. Three days later, animals were subjected to experiments of ConA and hVNS administration. Efficacy of capsaicin administration was examined by eye wiping test. Capsaicin was injected into rats with high-dose regimen as described above, and eye wiping behavior was measured 3 days after the initial injection by counting the frequency of eye wiping for 5 min period. Concanavalin A (7.5 mg/ml in saline, Sigma-Aldrich, St. Louis, MO, USA) was intravenously injected into the tail with a dose of 15 mg/kg.

Vagus nerve stimulation

Rats were anesthetized with ketamine (80 mg kg^-1; Yuhan, Seoul, Korea, Cat #8806421050707) and zylazine (5 mg kg^-1; Bayer, Leverkusen, Germany, Cat. #KR02315). This combined injection induced a stable anesthetic state for about 1 hour, which is an optimal time period to perform the surgery experiment without causing animals’ pain and suffering. The use of ketamine, an antipsychotic drug, has been approved by the Korea Ministry of Food and Drug Administration (Cheongju, Korea, Approval number: DJURFDA-130). The abdomen was incised and the middle and the left and right lateral lobes of liver were lifted up to expose the hepatic branch of the vagus nerve which is closely associated with portal vein, hepatic artery, bile duct, and liver hilus (Jensen et al. 2013). Exposed nerve was placed in a bipolar hook electrode of nichrome wire, and the electrical current (10 mA, 5 Hz, 5 ms of pulse duration, 5 min) was applied by using the isolated pulse stimulator (model 2100, A-M Systems Inc., Sequim, WA, USA). We used this stimulation condition based on our previous study showing an efficient induction of AFVN leading to activation of hippocampal neurons (Shin et al. 2019). After hVNS, animals were returned to animal room and sacrificed 1 – 3 days later. Sham treatment for hVNS was done by exposing the hepatic branch of the vagus nerve in anesthetized rats and suturing the skin without applying electrical stimulation. Animals underwent the same recovery procedure as hVNS group animals. The animals were sacrificed with an overdose of ketamine (150 mg kg^-1, i.p.).

Retrograde tracing

In order to identify vagal sensory neurons in the nodose ganglion (NG) innervating the liver tissue in rats, we performed a retrograde tracing experiment. Animals were anesthetized with ketamine (80 mg kg^-1) and xylazine (5 mg kg^-1), and 1,1′-Dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate (DiI, 3 μl of 0.5% in DMSO, Sigma-Aldrich) was taken by using Hamilton Syringe (#80330, Reno, NV, USA) and slowly applied to the cut end of the hepatic branch of the vagus nerve for 2-3 min. After suturing incised abdomen skin tissue, animals were recovered from narcosis and maintained in animal room for 7 days until the dissection of NG for further analysis. To dissect NG ganglion, we cut the anterior surface of the neck and exposed the carotid artery. The vagus nerve was carefully exposed from the carotid artery by removing muscles and connective tissues along the rostral direction. The NG was isolated just above the position where the hypoglossal nerve crosses the vagus nerve.

Western blot analysis

Liver tissue was dissected from rats and sonicated in RIPA buffer (150 mM NaCl, 1.0% CA-630, 0.1% SDS, 50 mM Tris, pH 8.0, 0.5% sodium deoxycholate; Thermo Fisher Scientific, Waltham, MA, USA) supplemented with protease inhibitor and phosphatase inhibitor cocktails (Roche Diagnostics, Canton, Switzerland). The lysate was centrifuged at 12,000 rpm, 15 min, and 4°C and the supernatant was collected. Extracted proteins (20 mg) were separated by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to PVDF membrane. PVDF membrane was reacted with blocking solution (5% BSA, 1X TBST (0.1% tween 20 in Tris-buffered saline)) and followed by primary and secondary antibody reactions. For the quantitative analysis of ChAT protein in the NTS and DMV, coronal brain sections (60 mm thickness) were prepared by using a cryostat (CM1850, Leica, Wetzlar, Germany). Sections were placed on the slides and the area covering NTS and DMV were scrapped off under the stereoscopic microscope by using 3 ml syringe. Preparation of cell lysates and the remaining steps of western blotting experiment were essentially the same as described above. Immunoblotting was performed with primary antibodies against TNF-α (ab9739, Rabbit-polyclonal, 1:2,000, Abcam, Cambridge, UK, RRID:AB_308774), IL-1β (ab9722, Rabbit-polyclonal, 1:2,000; Abcam, RRID:AB_308765), IL-6 (ab9324, Mouse-monoclonal, 1:2,000; Abcam, RRID:AB_307175), ChAT (ab181023, Rabbit-monoclonal, 1:2000, Abcam, RRID:AB_2687983), phospho-STAT3 (Y705, #9145, Rabbit-polyclonal, 1:1,000; Cell Signaling Technology, Danvers, MA, USA, RRID:AB_2491009), and β-actin (A1978, Mouse-monoclonal, 1:50,000; Sigma-Aldrich, RRID:AB_476692). Secondary antibodies were anti-rabbit IgG HRP (#7074, 1:5,000; CST, RRID:AB_2099233) and anti-mouse IgG HRP (#7076, 1:5,000; CST, RRID:AB_330924) antibodies. Intensity of protein bands in the X-ray film was determined by densitometric measurement using the i-Solution software (http://www.imt-digital.com/English2.0/html/home.php, version 21.1, Image & Microscope Technology, Daejeon, Korea).

Real-time PCR and RT-PCR

Total RNA was extracted from the liver and brain stem tissues by using trizol reagent (Thermo Fisher Scientific). cDNA was synthesized by incubating isolated RNA in the reaction containing 50 mM Tris-HCl, 3 mM MgCl₂, 75 mM KCl, 10 mM DTT), 104 μM dNTP mixture, RNasin (30 U), random primers (16 μM, Promega, Madison, WI, USA), and MMLV reverse transcriptase (200 U, Promega) for 2 hours at 37°C. RT-PCR was performed by using Green Master Mix (Promega) as described previously (Chang et al. 2018). The primer sequences for RT-PCR are as follows; the forward primer (5’-TTCTTTGTCTTGGATGTTGTCAT-3’) and reverse primer (5’-AACATTTCAACCTCAACCTTCTGG-3’) for ChAT mRNA, the forward primer (5’-TGTAGGCCTGCTGGATCAAC-3’) and reverse primer (5’- GCAGGATATCAGCTCGGTGT-3’) for AChE mRNA, and the forward primer (5’-CACACTGTGCCCATCTATGA-3’) and the reverse primer (5’- GCAGGATATCAGCTCGGTGT) for actin mRNA. Amplified DNA was analyzed by 1% agarose gel electrophoresis. Real-time PCR was performed in a 20 μl reaction volume containing synthetic cDNA (3 μg), 1X Power SYBR Green PCR Master mix (Life technologies, Carlsbad, CA, USA), 0.15 μM forward and reverse TNF-α primers. PCR for rat GAPDH gene was carried out by incubating synthesized cDNA (3 μg), 1X TaqMan Gene Expression Master Mix (AmpliTaq Gold DNA Polymerase, Thermo Fisher Scientific) and 1X rat GAPDH primer (Rat GAPD, Applied Biosystems, Foster City, CA, USA), endogenous control (VIC1/MGB probe primer) in a 20 μl of reaction volume. Real-time PCR reactions were performed using 96 well plate (MicroAmp Optical 96-well reaction plate, Applied Biosystems). The plate was covered with a film (MicroAmp Optical Adhesive Film, Applied Biosystems) and centrifuged briefly to spin down the sample (GS-6R Centrifuge, Beckman Coulter Life Sciences, Brea, CA, USA). Real-time PCR was carried out using an 7500 Real Time PCR System (Applied Biosystems) by activating Taq polymerase at 50°C for 2 min and at 95°C for 10 min, followed by 40 cycles with 15 sec at 95°C for denaturation and 1 min at 60°C for annealing. The RNA levels in each group were represented as fold changes in levels of target mRNAs to GADPH reference mRNA. The primer sequences for real-time PCR were as follows; the forward primer (5’-ACAAGGCTGCCCCGACTAT-3’) and the reverse primer (5’-CTCCTGGTATGAAGTGGCAAATC-3’) for TNF-α mRNA, the forward primer (5’-GGGCGGTTCAAGGCATAACAG-3’) and the reverse primer (5’-CTCCACGGGCAAGACATAGG-3’) for IL-1b mRNA, the forward primer (5’-CTGGTCTTCTGGAGTTCCGT-3’) and the reverse primer (5’-TGGTCCTTAGCCACTCCTTCT-3’) for IL-6 mRNA, and the forward primer (5’-GACCCAGAAGCTTCCAAGCCA-3’) and the reverse primer (5’-TGGGCATTGTAGTGACTCTCG-3’) for ChAT mRNA. The relative quantification (RQ) value of TNF-α, IL-1b, IL-6 and ChAT mRNA expression on GAPDH mRNA was calculated by the threshold cycle (Ct) data.

Immunofluorescence staining

Rats were anesthetized with overdose of ketamine and xylazine and perfused by using 4% paraformaldehyde in phosphate buffered saline (PBS). Tissues such as brain stem, NG, and liver were dissected and immersed overnight in 20% sucrose in PBS solution. After rapid freezing with -80^oC of dimethylbutane (Sigma-Aldrich), tissues were cut using a cryostat (Leica) and thaw-mounted on the slide (16 mm thickness). Immunofluorescence staining was performed as described previously (Chang et al. 2012). Briefly, sections were fixed, permeabilized, treated with blocking solution (2.5% BSA and 2.5% horse serum, 0.1% Triton X-100 in 1X PBS), and incubated with primary antibodies for 24 h at 4℃, washed three times with 1X PBST and incubated with secondary antibodies at room temperature for 2 h in a dark room. The primary antibodies used were anti-TNF-α (ab9739, Rabbit-polyclonal, 1:400; Abcam), anti-IL-1β (ab9722, Rabbit-polyclonal, 1:400; Abcam), anti-IL-6 (ab9324, Rabbit-polyclonal, 1:400; Abcam), anti-P2X2 (PA1-24624, Rabbit-polyclonal, 1:400; Thermo Fisher Scientific, Waltham, MA, USA, RRID:AB_2157912), anti-c-Fos (sc-166940, Mouse-monoclonal, 1:400; Santa Cruz Biotech, Dallas, Texas, USA, RRID:AB_10609634), anti-cleaved Caspase-3 (#9661, Rabbit-polyclonal, 1:400; CST, RRID:AB_2341188), anti-CD11b (554980, Mouse-monoclonal, 1:200; BD Biosciences, Franklin Lakes, NJ, USA, RRID:AB_2129492), anti-pY-STAT3 (#9145, Rabbit-polyclonal, 1:400; CST), anti-NF-200 (N0142, Mouse-monoclonal, 1:400; Sigma-Aldrich, RRID:AB_477257), anti-ChAT (ab181023, Rabbit-monoclonal, 1:400, Abcam), anti-VR1 (M-1714-100, Mouse-monoclonal, 1:400, Abcam, RRID:AB_2492520), anti-Albumin (NBP1-32458, Rabbit-polyclonal, 1:200, Novus Biologicals, Centennial, CO, USA, RRID:AB_10003946) primary antibodies, and normal mouse IgG (sc-2025, Mouse-monoclonal, 1:400, Santa Cruz Biotechnology, RRID:AB_737182) and normal rabbit IgG (#2729, Rabbit-polyclonal, 1:400, CST, RRID:AB_1031062) antibodies. Rhodamine-goat anti-rabbit IgG (R-6394, 1:400; Molecular Probes, Eugene, OR, USA, RRID:AB_2556551) and fluorescein-goat anti-mouse IgG (F-2761, 1:400; Molecular Probes, RRID:AB_2536524) antibodies were used as secondary antibodies. When necessary for nuclear staining, sections were incubated with Hoechst 33258 (2.5 μg/ml, bis-benzimide, Sigma) for 10 minutes before the final washing with 1X PBST. Signal intensity of immunofluorescence images were measured by using the program Image J (ImageJ, NIH, Bethesda, MA, USA, RRID:SCR_003070). Quantification was represented as either the signal intensity or the number of cells displaying effective pixel values. Fluorescence images were converted into grayscale mode, and the pixel density above the threshold which has been set in the program was adapted as being effective for further quantification. Fluorescence intensity in the images was presented as the pixel density relative to that of the control images. Also, individual cells showing the pixel density above the threshold which has been set in the same way as above were counted. The number of labeled cells or the pixel density in the field of image were counted and averaged for 3-5 nonconsecutive sections. Observers were blind to the slides that were used for the analysis by fluorescence microscopy.

Experimental design and statistical analysis

For the studies examining the effects of hVNS on the production of inflammatory cytokines, rats were randomly assigned to untreated control, ConA plus Sham, and ConA plus hVNS. Sham or hVNS treatments were performed in ConA-injected animals. Liver and brainstem tissues were collected for real-time PCR, western blotting and immunofluorescence experiments. For CAP treatment studies, all rats were administered with CAP and randomly assigned into CAP, CAP plus hVNS, CAP plus ConA and Sham, and CAP plus ConA and hVNS. Administration of ConA and hVNS were essentially the same as above. Brainstem tissues were collected and used for RT-PCR and western blot analyses, and liver tissues for western blot analysis. Some of CAP-injected rats and untreated control animals were used to examine eye wiping test. The experimenter was blinded to the animal’s groups during experimentation and statistical analysis.

All data were presented as mean ± standard deviation (SD). The mean number data among experimental groups were compared by using Student’s t test (unpaired) or one-way ANOVA and Tukey’s multiple comparison test for multiple comparisons (GraphPad Prism 7.00, GraphPad Software Inc., San Diego, CA, USA). Statistically significant differences were set at *p<0.05, **p<0.01, ***p<0.001.

hVNS downregulates hepatic expression of inflammatory cytokine mRNAs and proteins in ConA-injected animals

To investigate whether hVNS has the effects on the inflammatory responses in the liver tissue, we delivered hVNS 24 h after ConA injection and analyzed the hepatic expression of TNF-α, IL-1β and IL-6 inflammatory cytokines 24 h later. mRNA levels of TNF-α, IL-1β, and IL-6 were significantly elevated by ConA treatment and then down-regulated by hVNS (Fig. 1A-C). Similarly, protein levels of TNF-α, IL-1β, and IL-6 were markedly increased in the liver tissue after ConA treatment and down-regulated by hVNS (Fig. 1D-F). Immunofluorescence analysis showed that the signals of TNF-α, IL-1β, and IL-6 were clearly induced in the liver tissue by ConA treatment, being mostly colocalized with macrophage marker protein CD11b that was also increased by ConA (Fig. 2A upper panel). Then, the signals of TNF-α, IL-1β and IL-6 were reduced by hVNS in ConA-treated animals. Quantitative measurement of protein signals showed significant increases by ConA and decreases by hVNS (Fig. 2A lower panel). The protein signals, colocalized with CD11b in ConA-treated animals, were detected in perinuclear area, as identified by merged view with Hoechst-stained nuclei (Fig. 2B).

To investigate the persistency of anti-inflammatory effects of hVNS in ConA-treated liver tissue, we analyzed levels of inflammatory cytokines 2–4 days after ConA injection (; individual groups labeled 2d, 3d, and 4d ConA groups in Fig. 3A). Levels of TNF-α, IL-1β, and IL-6 were maintained or partly reduced in 2d and 3d ConA groups of animals and further decreased in 4d ConA group (Fig. 3B, D, F). Vagal modulation of inflammatory cytokines, in terms of the decreases in cytokine levels in the presence of hVNS relative to the absence of hVNS, was obvious in 2d ConA groups and became less effective in 3d and 4d ConA groups (Fig. 3C, E, G). In 3d ConA groups, levels of three cytokines were all decreased after hVNS administration though the changes were not statistically significant (% decreases and p values; 71% and p = 0.74 for TNF-α, 63% and p = 0.58 for IL-1β, 52% and p = 0.35 for IL-6).

hVNS induces an increase in STAT3 phosphorylation at tyrosine 705 (pY-STAT3) in the liver

It was previously reported that JAK2-STAT3 pathway is involved in cholinergic anti-inflammation after VNS (de Jonge et al. 2005). Here in the liver tissue, pY-STAT3 was not detected in the liver tissue of untreated control, slightly induced after ConA treatment, and the level was further elevated by hVNS (Fig. 4A), showing a significant increase of band intensity in ConA + hVNS group compared to the control group (Fig. 4B). Immunofluorescence images of liver tissue showed that induced signals of pY-STAT3 were localized with CD11b-labeled cells and were mostly detected around Hoechst-stained nuclei in both ConA + Sham and ConA + hVNS groups (Fig. 4C). Percentage of pY-STAT3-positive cells among the whole CD11-positive cells was significantly increased in ConA + hVNS group compared to CTL or ConA + Sham group (Fig. 4D). To examine the reaction specificity of CD11 antibody, we co-immunolabeled CD11b with albumin protein that is expressed only in parenchymal hepatocytes (Gu et al. 2017). CD11-lebeled cells were clearly distinguished from the outnumbered albumin-labeled hepatocytes in experimental groups (Fig. 4E). No signals were detected by normal IgG antibody reactions in all experimental groups, supporting the specificity of immunolabeling reactions.

hVNS increases the expression of ChAT mRNA and protein in DMV neurons

ChAT catalyzes the synthesis of acetylcholine in the presynaptic terminal of cholinergic neurons and its expression is induced in glutamate-applied cholinergic neurons and splenic T cells after VNS (Rosas-Ballina et al. 2011; Zhou et al. 2008). Here, we examined the expression of ChAT mRNA and protein in the DMV neurons. hVNS induced significant increases in ChAT mRNA level in the tissue lysate of DMV area (Fig. 5A). ConA treatment in Sham control or in hVNS group did not significantly change ChAT mRNA levels. However, the levels of ChAT mRNA significantly elevated by hVNS administration in both Sham and ConA groups. Western blot analysis showed that ChAT proteins were similarly increased by VNS (Fig. 5B) and quantitative comparison revealed significant increases in band intensity by hVNS in Sham and ConA groups (Fig. 5C). ChAT proteins were detected as multiple bands between 67–72 kDa with higher molecular weights in Sham and ConA + Sham groups than those in corresponding hVNS-administered groups. We detected no protein band from the cell lysates of the macrophages that showed no expression of ChAT in the previous report (Fujii et al. 2017), supporting the reaction specificity of ChAT antibody. Immunofluorescence analysis showed that in animals given hVNS, the number of ChAT-positive cells were slightly higher in the ipsilateral side (left DMV) than the contralateral side (right DMV), yet failing to show statistical difference. (% difference and p value, 11.7% and 0.5 respectively; Fig. 5D, E). ChAT expression in the DMV by hVNS was lower in the number and more restricted in its distribution when compared with the pattern of ChAT expression after VNS at the cervical location (labeled cVNS in Fig. 5D, E), indicating the suborganization of ChAT-expressing DMV neurons that respond to hVNS. Moreover, ChAT protein, displaying stronger signals in ConA + hVNS group compared to Sham control and ConA groups, was mostly colocalized with c-Fos protein in the DMV neurons (Fig. 5F). Enlarged image revealed that c-Fos proteins showing intense signal in the nuclear zone were colocalized with cytoplasmic ChAT-labeled neurons in the DMV (Fig. 5G).

CAP selectively degenerates NG sensory neurons

AFVN make vagovagal connections and innervate peripheral target organs through efferent fibers. To examine whether hVNS-induced activation of afferent fibers affects on anti-inflammatory responses in the liver, we set out to study the production of inflammatory cytokines after eliminating afferent vagus nerve activity. First, in order to identify NG neurons connected to the liver, we applied DiI into the hepatic vagal branch and found that a certain portion of NG neurons, as identified by immunofluorescence staining of NF-200 proteins, were retrogradely labeled with DiI (Fig. 6A). To investigate whether NG neurons are required for hVNS-mediated anti-inflammatory reaction, we attempted to remove NG sensory neurons by low- or high-dose of CAP administration (CAP-L or CAP-H). NG sensory neurons responding to CAP was identified by immunolabeling of TRPV-1 and additional labeling of P2 × 2 purinergic receptors. TRPV1-positive sensory neurons were colocalized with P2 × 2-labeled NG neurons (marked arrows), and they were shrunken or weakly stained after CAP treatments (Fig. 6B). The number of TRPV1-positive neurons was slightly reduced by CAP-L and further decreased by CAP-H treatment (Fig. 6C). The number of cells expressing active cleaved form of caspase3 was are increased by CAP in NG neurons, showing significant increases by CAP-H treatment (Fig. 6D, E), indicating apoptotic death of NG neurons by CAP. Collectively, CAP-H was effective at eliminating CAP-responding NG neurons and thus was used for the rest of the present study. Systemic administration of CAP-H resulted in significant increases in eye-wiping behavior in CAP-H-treated groups compared to vehicle control (Fig. 6F), suggesting the behavioral effects of nociceptive TRPV signaling in the cornea by CAP-treated rats (Hegarty et al. 2014).

CAP downregulates ChAT expression and hepatic production of inflammatory cytokines

To further examine that CAP-induced removal of NG neurons had an effect on efferent vagus nerve activity, we analyzed ChAT protein in the DMV after CAP treatment. Levels of ChAT protein in hVNS and ConA + hVNS groups were markedly decreased after CAP treatment, showing the similar size in molecular weights of ChAT as the uppermost band in the control group (Fig. 7A). No band was detected for the protein lysates from macrophages cells, as was in Fig. 5B, again confirming the absence of non-specific reaction of ChAT antibody. In the presence of CAP, ChAT protein levels did not show any significant differences among Sham, hVNS, ConA + Sham, and ConA + hVNS groups though the band intensity in CAP + ConA + Sham group was relatively stronger (Fig. 7B, C). Having noted the variation in the molecular weights of ChAT protein in DMV area among experimental groups, we investigated the possible involvement of posttranscriptional modification of ChAT mRNA expression. We identified a single DNA band at 529 bp (Fig. 7D upper), as expected by amplification of common-type ChAT mRNA by RT-PCR (Saito et al. 2009), implicating that the variation of ChAT protein size may be derived from posttranslational, not posttranscriptional, modification. Band intensity was significantly higher in the VNS and ConA + VNS groups compared with their corresponding control groups (Fig. 7D lower). We also investigated the expression of acetylcholinesterase (AChE) mRNAs by RT-PCR in the DMV. PCR band for AChE mRNA, which was not detected from the control tissue, was clearly seen only in ConA + Sham group (Fig. 7E upper) and was significantly decreased by VNS (Fig. 7E lower).

We then examined the effects of CAP treatment on hVNS-modulated production of inflammatory cytokines in the liver. ConA treatments induced significant increases in TNF-α band intensity in both CAP and non-CAP vehicle (VEH) groups that were downregulated by hVNS, showing less decrease in CAP groups than VEH group in terms of the ratio of band intensity of hVNS group to Sham group (% decreases and p values; 97% and p = 0.01 for VEH group vs. 72% and p = 0.12 for CAP groups, One-way ANOVA) (Fig. 8A, B). Similarly, ConA treatments induced significant increases in IL-1β band intensity in both CAP and VEH groups that were downregulated by hVNS, showing less decrease in CAP groups than VEH group in terms of the ratio of band intensity of hVNS group relative to Sham group (% decreases and p values; 72% and p = 0.02 for VEH groups vs. 60% and p = 0.04 for CAP groups, One-way ANOVA) (Fig. 8C, D). Finally, ConA induced large increases in IL-6 band intensities in both CAP and VEH groups that were downregulated by hVNS, showing less decrease in CAP groups than VEH group in terms of the ratio of band intensity of hVNS group relative to Sham group (% decreases and p values; 91% and p < 0.0001 for VEH groups vs. 62% and p = 0.002 for CAP groups, One-way ANOVA) (Fig. 8E, F), thus representing less change in CAP group than VEH group. A comparison of band intensity showed higher values in ConA-VNS-CAP group than ConA-VNS-Vehicle group (; the arithmetic mean ratios and p values of the band intensities of ConA-VNS-CAP to ConA-VNS-Vehicle groups; 9.4 and 0.91 for TNF-α, 1.7 and 0.92 for IL-1β, 4.1 and 0.27 for IL-6, respectively), but the comparison did not show statistical significance for all three inflammatory cytokines, which might reflect the overall weak, but highly variable band intensities among VNS-administered animal groups. Taken together, these results suggest that the inhibitory effects of hVNS on the production of inflammatory cytokines by CAP animals are attenuated by CAP treatment.

In the present study, we found that hVNS regulates hepatic expression of inflammatory cytokines TNF-α, IL-1β, and IL-6 in Con-A model of hepatitis and also increases the expression of ChAT gene in the DMV neurons. Pharmacological blockade of AFVN decreased ChAT expression and suppressed anti-inflammatory effects mediated by hVNS. Thus, our data present new evidence that afferent vagus nerve activity is required for hVNS-induced anti-inflammatory response in the liver of ConA-injected animals.

T cells are activated by ConA binding and stimulate antigen-presenting cells such as Kupffer cells in the liver through the molecular interaction between T cell receptor and MHC complexes (Tiegs, 1997). Inflammatory cytokines are produced from activated T cells and antigen presenting cells, causing hepatic inflammation. Given that T cells such as acetylcholine-synthesizing T cells and hepatic invariant NKT cells can be activated by adrenergic stimulation of celiac ganglionic neurons that are innervated by parasympathetic vagus nerve or from spinal sympathetic nerve activity (Wong et al. 2011; Rosas-Ballina et al. 2011), we hypothesized that the similar neuroimmune communication might be involved in the regulation of hepatic inflammation by hVNS. Here, we found that hVNS attenuated hepatic production of inflammatory cytokines TNF-α, IL-1β, and IL-6 in ConA-injected animals. We also found that the level of phospho-STAT3 (Y705) was increased by hVNS in ConA-injected animals. pY-STAT3 signals were slightly induced from CD11-positive macrophages in the liver of ConA-injected animals, suggesting the possible involvement of pY-STAT3 activity in macrophage activation that mediates proinflammatory reactions (Wang et al. 2011). Then, VNS further elevated the level of pY-STAT3 in CD11-positive hepatic macrophages. In contrast to short-term activation of STAT3, prolonged increases of pY-STAT3 by VNS may activate anti-inflammatory pathway leading to the upregulated expression of target genes such as suppressor of cytokine signaling 3 (SOCS3) and transcriptional repression of inflammatory cytokines (de Jonge et al. 2005; Yasukawa et al. 2003). Vagal efferent fibers are known to have synaptic connection within the ganglia at the hepatic hilus and the portal space to postganglionic fibers that are extended into hepatic vasculature, bile duct, and hepatic parenchyma (McCuskey, 2004; Sutherland, 1964; Skaaring & Bierring, 1976). It was also reported that the activation of α7 nicotinic acetylcholine receptors is involved in VNS-induced anti-inflammation in hepatic cells such as Kupffer cells and hepatic resident macrophages (Wang et al. 2003; Hiramoto et al. 2008; Nishio et al. 2017; Fonseca et al. 2019). However, the sequence of events connecting between ACh-releasing vagal efferent fibers and AChR-expressing target cells in the liver remains unclear, and thus the characterization of mediators of intercellular cholinergic signaling, if any, should be of great importance to understand VNS-induced anti-inflammation in the liver.

We found that c-Fos-positive DMV neurons in hVNS animals also induced increased expression of ChAT, suggesting that ChAT activity may be linked to hVNS-induced cholinergic anti-inflammation. How can hVNS induce ChAT gene expression in DMV neurons? One possible pathway would be antidromic conduction of electrical stimulation of efferent fibers into neuronal cell body in the DMV. Electrical stimulation of peripheral nerve fibers after injury is known to retrogradely transport lesion signals and facilitate regenerative responses in the nucleus at gene expression level (Al-Majed et al. 2000; Mahar & Cavalli, 2018). Another pathway is that the increased activity of AFVN is transmitted into NTS neurons through glutamate receptors and subsequently into DMV neurons utilizing GABA, glutamate, and norepinephrine as neurotransmitters (Andresen & Yang, 1990; Travagli et al. 2006). When we administered CAP in rats in order to remove AFVN, levels of ChAT which had been induced by hVNS were diminished close to baseline values of untreated control group, suggesting that the enhanced activity of AFVN by hVNS may function to induce ChAT expression in DMV neurons. However, it is unclear how neuronal excitability transmitted into the soma of DMV neurons induces ChAT gene expression. ChAT gene expression was shown to be induced in DMV neurons after vagotomy (Wang et al. 2011). Further studies are important to examine whether there are common signaling mechanisms inducing gene expression from injured axons and electrically stimulated axons.

ChAT expression generates two splice variants, common-type ChAT (cChAT) and peripheral type ChAT (pChAT) with molecular weights of 68 kDa and 48.9 kDa, respectively, which are differentially induced in DMV neurons after vagotomy (Saito et al. 2009; Tooyama & Kimura, 2000). In addition, electrophoretic mobility analysis of cChAT protein purified from rat brain showed multiple bands around 67 kDa (Dietz & Salvaterra, 1980; Hersh et al. 1984). In the present study, we have identified ChAT mRNA in DMV neurons as a single amplified band by RT-PCR that corresponds to cChAT transcript. However, immunoblotting of ChAT protein in DMV lysates showed multiple bands between 67–72 kDa in Sham and ConA-treated groups and a single band at 72 kDA in CAP-H-treated groups; In contrast, only one intense band was detected at ~ 67 kDa after hVNS in non-CAP VNS groups. Immunoblotting for the protein lysate from cultured RAW 264.7 macrophage cells with anti-ChAT antibody did not generate any protein band, implying the absence of possible non-specific detection of ChAT antibody with unrelated protein species. Taken together, our data suggest that the process of posttranslational modification may differentially regulate so as to produce variant ChAT proteins in DMV neurons, reflecting the effects of CAP and/or hVNS administrations. For instance, hVNS may facilitate the reaction of proteolytic cleavage of higher molecular weight precursors of ChAT and increase the stability of processed ChAT protein, leading to the accumulation of 67 kDa of ChAT protein. Interestingly, expression of AChE mRNA in the dorsal vagal complex was increased by ConA and downregulated by VNS. Along with the increased production of ChAT in DMV neurons of ConA-treated animals, hVNS-induced inhibition of AChE may play a role in facilitating cholinergic neurotransmission in the peripheral target tissues such as liver and spleen and contribute to the regulation of inflammation. Further studies on the regulation of AChE production by VNS in DMV neurons will be useful to understand the functional link of AChE to cholinergic anti-inflammation.

Previous studies have shown that systematic or perivagal application of CAP results in degeneration of AFVN (Czaja et al. 2008; Li & Owyang, 1993). CAP binds to non-selective cationic channel TRPV1, depolarize the nociceptive sensory neurons (Caterina et al, 1997), which can cause neuronal toxicity and death (Czaja et al. 2008). Here, we confirmed that the systemic administration of CAP induced apoptosis in the NG neurons and reduced hVNS-induced production of ChAT in the DMV neurons. Moreover, CAP treatment deteriorated NG sensory neurons positive to immunolabeling of TRPV1 receptors that were colocalized with P2 × 2-positive sensory neurons known to be largely present in the c-fibers of NG neurons transmitting signals via CAP receptor TRPV1 (Yu et al. 2005; Kwong et al. 2008). However, it was reported that CAP treatment inhibits the activation of T lymphocytes and decrease the inflammation and apoptosis of hepatocytes in ConA-treated animal (Zhang et al. 2019b). Given the difference in a time period of CAP pretreatment and injection dose (1 mg/kg for 30 min in Zhang et al. 2019b vs 200 mg/kg for 3 days in the present study) before the administration of ConA, acute CAP treatment may act on TRPV1 receptors of immune cells that are activated by ConA whereas the prolonged treatment of higher dose of CAP prior to ConA may be effective, preferentially on TRPV1 receptors in sensory nerve fibers, leading to distinct pathophysiological consequences in Con-A model of hepatitis. While our data strongly suggest the down-regulation of ChAT expression due to decreased activity of AFVN in CAP-treated animals, a possibility of direct action of CAP on vagal motor neuron leading to the suppression of ChAT gene expression cannot be excluded (Browning et al. 2013). Further studies on selective blocking of vagal afferents, for instance by using surgical vagal deafferentation (Baptista et al. 2007), are required to clarify this issue.

We discovered that anti-inflammatory effects of hVNS in ConA-treated animals were largely diminished by CAP treatment. The extent of reduction of TNF-α, IL-1β, and IL-6 levels after hVNS was suppressed by CAP treatment. However, a direct comparison of changes in inflammatory cytokine levels in ConA + VNS group of animals before and after the CAP treatment does not show the statistical significance. Having noted high variation of band intensities among experimental groups of animals given VNS, more quantitative measurement of these inflammatory cytokines (e.g., ELISA) would be useful to clarify this issue. After the removal of afferent fibers by CAP, the remaining efferent vagal activity caused by hVNS might be reduced and thus would be less effective in attenuating inflammatory in the liver, supporting the notion that the activation of vagal afferents contributes to hVNS-mediated hepatic anti-inflammation. This interpretation is consistent with the reduction of ChAT level by CAP treatment in DMV neurons. Since hVNS elevates ChAT levels in DMV neurons, ChAT in DMV may be used as one of indicators of hVNS-induced anti-inflammation. While our results claim the contribution of vagal afferent activity, a possibility of sympathetic nerve activity cannot be ruled out (see Fig. 8G). The sympathetic nerve pathway could be activated indirectly through vago-spinal descending activity and function to regulate inflammation (Komegae et al. 2018) by virtue of immunosuppression by increased adrenergic inputs into liver cells (Wong et al. 2011).

In conclusion, we have found that acute type of hVNS has a dampening effect on hepatic inflammation and demonstrated that afferent nerve activity contributes to hVNS-induced anti-inflammation. Further technical development, for instance by using optogenetic method (Chang et al. 2015) to stimulate separately the afferent and efferent components of vagus nerve fibers, would be of great advantage to develop therapeutic strategies and also to gain insights into interaction between pathological visceral organs and the brain.

AFVN, afferent fibers of the vagus nerve; VNS, vagus nerve stimulation; hVNS, hepatic branch of the vagus nerve; CAP, Capsaicin; ConA, concanavalin A; ChAT, choline acetyltransferase; DMV, dorsal motor nucleus of vagus nerve; NTS, nucleus tractus solitaries; TRPV1, transient receptor potential vanilloid 1; NG, nodose ganglion; SAC, sacrifice; CTL, untreated control; pY-STAT3, STAT3 phosphorylation at Y705; cVNS, cervical vagus nerve stimulation; DVC, dorsal vagal complex; VEH, vehicle; CG, celiac ganglion; PPF, parasympathetic postganglionic fiber; SPF, sympathetic postganglionic fiber.

Ethical Approval

All animal care procedures were approved by the Committee on Use of Live Animals for Teaching and Research at Daejeon University (approval number: DJUARB2019-029).

Availability of data and materials

The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request.

Funding

This work was supported by the Basic Science Research Programs through the National Research Foundation of Korea (NRF) funded by the Ministry of Education (NRF-2016R1D1A1B01014190 and NRF-2018R1A6A1A03025221).

Competing interests

The authors declare that they have no competing interests.

Author Contributions

All experiments were performed in the Daejeon University. UN and BJ conceived and designed the experiments. BJ performed the experiments. UN and BJ analyzed the data. UN supervised the projects and wrote the manuscript. BJ and UN reviewed and approved the final version of the manuscript. All authors read and approved the final manuscript."

Acknowledgements

We are indebted to former and present members of the Neurophysiology Laboratory at Daejeon University for valuable discussions and comments.

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Vagal Afferent Fibers Contribute to the Anti-Inflammatory Reactions by Vagus Nerve Stimulation in Concanavalin A Model of Hepatitis in Rats

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Abstract

Figures

Introduction

Materials And Methods

Experimental Animals and Ethical Approval

Drug administration

Vagus nerve stimulation

Retrograde tracing

Western blot analysis

Real-time PCR and RT-PCR

Immunofluorescence staining

Experimental design and statistical analysis

Results

hVNS downregulates hepatic expression of inflammatory cytokine mRNAs and proteins in ConA-injected animals

hVNS induces an increase in STAT3 phosphorylation at tyrosine 705 (pY-STAT3) in the liver

hVNS increases the expression of ChAT mRNA and protein in DMV neurons

CAP selectively degenerates NG sensory neurons

CAP downregulates ChAT expression and hepatic production of inflammatory cytokines

Discussion

Abbreviations

Declarations

Ethical Approval

Availability of data and materials

Funding

Competing interests

Author Contributions

Acknowledgements

References

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