Skip to main content
NCBI home page
Food Science & Nutrition logoLink to Food Science & Nutrition
. 2020 Jul 23;8(9):5063–5070. doi: 10.1002/fsn3.1805

Anti‐inflammatory effect of Antirrhinum majus extract in lipopolysaccharide‐stimulated RAW 264.7 macrophages

Mi Jang 1, Inguk Hwang 1, Byungsoon Hwang 1, Gichang Kim 1,✉
PMCID: PMC7500786  PMID: 32994966

Abstract

Antirrhinum majus (AM) has attracted attention as a rich source of phytochemicals, which are beneficial for human health. However, the anti‐inflammatory effects of AM have not been studied scientifically. Therefore, we investigated the antioxidative properties and anti‐inflammatory effects of AM extract (AME) in lipopolysaccharide (LPS)‐stimulated RAW 264.7 macrophages. AME showed high radical‐scavenging ability. Viability of RAW 264.7 cells was not significantly altered by AME at the concentrations of 0–300 µg/ml. LPS‐induced nitric oxide (NO) production was decreased by treatment with 0–300 µg/ml AME in a concentration‐dependent manner. AME pretreatment significantly inhibited the protein expression of inducible nitric oxide synthase (iNOS) and cyclooxygenase‐2 (COX‐2) in a concentration‐dependent manner. AME also considerably inhibited the mRNA and protein expression of inflammatory cytokines, such as tumor necrosis factor‐a (TNF‐α), interleukin‐1 β (IL‐1β), and interleukin‐6 (IL‐6). These findings provide a foundation for further studies and use of AM in nutraceuticals.

Keywords: anti‐inflammatory, antioxidative, Antirrhinum majus, cytokines, ethanol extract

Antirrhinum majus extract showed strong inhibition of inflammatory cytokines (TNF‐alpha, IL) and an protein expression of iNOS and COX‐2. Also, it show high radical‐scavenging ability. These findings provide a foundation for further studied and use as nutraceuticals.

graphic file with name FSN3-8-5063-g006.jpg

1. INTRODUCTION

Inflammatory response refers to a response in which the immune cells secrete various inflammatory mediators to protect the body in the event of trauma caused by an external physical or chemical stimulus or introduction of foreign substances such as bacteria or virus (Dinarello, 2010; Olefsky & Glass, 2010). Sustained inflammatory response can lead to an excessive secretion of inflammatory mediators, which cause various adult diseases, including obesity, diabetes, and hypertension (Charo & Taub, 2011; Deans & Sattar, 2006; Ohashi, Shibata, Murohara, & Ouchi, 2014; Wojdasiewicz, Poniatowski, & Szukiewicz, 2014; Wong, Spence, Lamki, Freeman, & McDonald, 1986). Macrophages, which are distributed throughout the body tissues, play a key role in this inflammatory response. They not only possess the ability to defend against pathogens and cancer cells but are also activated by various stimuli or cytokines secreted by immune cells; this enables macrophages to secrete various physiologically active substances and pro‐inflammatory cytokines, such as nitric oxide (NO), tumor necrosis factor (TNF‐α), interleukin (IL)‐1β, and IL‐6, to maximize the immune response (Scheller, Chalaris, Schmidt‐Arras, & Rose‐John, 2011; Valledor, Comalada, Santamaría‐Babi, Lloberas, & Celada, 2010). Also, MCP‐1, a potent chemical factor in the CC‐chemotactic family, is caused by the stimulation of LPS in macrophages. It is one of the key chemokines the regulate infiltration and migration of monocytes and macrophages to sites of active inflammation (Jamie et al., 2012; Satish, Sergey, Shohreh, & Bassel, 2009). Inflammation within the body causes the production of a large amount of inflammatory factors, such as NO and prostaglandin E2 (PEG2), by inducible nitric oxide synthase (iNOS) and cyclooxygenase‐2 (COX‐2). In the inflamed state, excessive amount of NO produced by iNOS not only facilitates inflammatory response but also exacerbates the inflammation by facilitating the biosynthesis of inflammatory mediators (Choi, Kim, & Lee, 2014; Fiorucci et al., 2004). In other words, the expression of iNOS and COX‐2 and the production of NO and PEG2 serve as typical inflammatory factors of immune cells.

Moreover, another factor that can induce an inflammatory response is the oxidative stress generated inside the body. The biochemical reactions of normal cells, along with various environmental pollutants and chemicals, increase oxidative stress in the body, which causes cellular damage, including cell membrane damage and protein degradation. Furthermore, because it is linked to inflammatory response, oxidative stress can also induce chronic inflammatory response that leads to DNA damage and various diseases such as cancer, heart disease, arteriosclerosis, Alzheimer's disease, and Parkinson's disease (Halliwell & Gutteridge, 1984, 2015; Hybertson, Gao, Bose, & McCord, 2011; Reuter, Gupta, Chaturvedi, & Aggarwal, 2010; Uttara, Singh, Zamboni, & Mahajan, 2009; Valko et al., 2007). Accordingly, chronic inflammatory response can be reduced by decreasing oxidative stress via consumption of ingredients with excellent antioxidative effect (Di Penta et al., 2013; Elmarakby & Sullivan, 2012).

Antirrhinum majus (AM), which belongs to the Scrophulariaceae family, is a perennial plant that originates from Africa and Southern Europe. AM was so named because its flower resembles the mouth of a goldfish. Its seeds, leaves, and flowers are used as herbal medicines, and AM is known to have anti‐inflammatory, analgesic, and diuretic effects; however, scientific studies on the physiological activity of AM are still lacking. Therefore, the present study aimed to identify the functionality of AM extract (AME) to explore whether it can be used as a basic ingredient in the development of agents for preventing and treating inflammatory diseases. We investigated the in vitro antioxidative activities of AME, along with its inhibitory effects on the production of NO, iNOS, COX‐2, and inflammatory cytokines in lipopolysaccharide (LPS)‐induced RAW 264.7 cells.

2. MATERIALS AND METHODS

2.1. AM sample and extract preparation

Antirrhinum majus used in the present study was purchased from Five Mountain Herb Farm in 2016. The purchased AM was washed twice with water, and the moisture was removed. Subsequently, 100 g AM was submersed in 95% ethanol and ground with a homogenizer (IKA) and then incubated in the dark for 24 hr at 25°C. AM‐ethanol extract was filtered using a filter paper (NO. 2; Whatman), followed by decompression concentration with a rotary vacuum evaporator (R‐205; Buchi) at 37°C. The concentrated AM extract (AME) was freeze‐dried, powdered, and stored at −20°C for use in further experiments.

2.2. Measurement of total polyphenol content

Total polyphenol content from the AME was measured via a method modified from the Folin–Denis method (Singleton & Rossi, 1965). Here, 500 μl of the extract was reacted with 500 μl of 0.5 N Folin–Ciocalteu's phenol reagent in the dark for 3 min, followed by addition of 1.5 ml of 2% Na2CO3 and incubation for 30 min in the dark at room temperature. The samples were dispensed into a 96‐well plate, and the absorbance at 760 nm was measured using a microplate reader (Infinite M200 PRO; Tecan). TPC was calculated by obtaining the slope and intercept values from the standard calibration curve constructed using the absorbance values based on gallic acid solutions at four different concentrations (1, 10, 100, and 1,000 μg/ml) and then substituting the absorbance value with that of each sample.

2.3. Measurement of total flavonoid content

Total flavonoid content was measured via the method described by Moreno, Isla, Sampietro, and Vattuone (2000). Briefly, 1 ml of the extract was mixed with 1 ml of 2% AlCl3 and incubated in the dark for 30 min, followed by measuring the absorbance at 430 nm using a microplate reader. TFC was calculated by obtaining the slope and intercept values from the standard calibration curve constructed using the absorbance values based on quercetin solutions at four different concentrations (1, 10, 100, and 1,000 μg/ml) and then substituting the absorbance value with that of each sample.

2.4. Measurement of DPPH and ABTS radical‐scavenging ability

The Blois method was used to measure the electron donating ability of AME samples with respect to 1,1‐diphenyl‐2‐picrylhydrazyl (DPPH; Sigma‐Aldrich Co.) by using (Blois, 1958). The AME samples were diluted to concentrations of 1, 2, 5, and 10 μg/ml. Then, 1 ml of 0.15 mM DPPH solution was added to 1 ml of each diluted sample, the mixtures were incubated for 30 min in the dark, and the absorbance at 517 nm was measured. The 2,2'‐azinobis(3‐ethylbenzothiazoline‐6‐sulfonate) (ABTS) radical‐scavenging ability was measured using the method described by Pellegrini et al (Re et al., 1999). After thoroughly mixing 7 mM ABTS (Sigma‐Aldrich Co.) and 2.45 mM K2S2O8 (potassium persulfate; Sigma‐Aldrich, Co.) solutions, the mixture was stored for 12–16 hr in the dark. The ABTS solution was used by precalibrating its OD value at 734 nm to be 0.7–1.1. After mixing 100 μl of ABTS solution with 100 μl of samples diluted to varying concentrations, the mixtures were dispensed to a 96‐well plate and allowed to react for 30 min in the dark, after which, absorbance at 732 nm was measured. Ascorbic acid was used as the positive control for DPPH and ABTS radical‐scavenging ability.

2.5. Cell culture

RAW 264.7 macrophages were obtained from the American Type Culture Collection (ATCC). RAW 264.7 macrophages were cultured in 37°C, 5% CO2 incubator with Dulbecco's modified Eagle's medium (DMEM; Sigma‐Aldrich Co.) containing 10% fetal bovine serum (FBS; Gibco), and 1% Penicillin‐Streptomycin solution (Sigma‐Aldrich Co.).

2.6. Cytotoxicity

Cytotoxicity in RAW 264.7 macrophages was tested via 3‐(4,5‐dimethylthiazole‐2‐yl)‐2,5‐diphenyl‐tetrazolium bromide (MTT; Sigma‐Aldrich Co.) assay. Cells were seeded into a 96‐well plate, with 1 × 103 cells/well, and cultured in 37°C, 5% CO2 incubator for 24 hr. After treating the cells with varying concentrations of AME, the cells were cultured again for 24 hr under the same conditions. Next, 50 μl of 5 mg/ml MTT solution was added to each well, and the cells were cultured for 4 hr. Then, the medium was removed, the cells were washed twice with phosphate‐buffered saline (PBS), and 200 μl of dimethyl sulfoxide (DMSO; Sigma‐Aldrich Co.) was added to each well. Absorbance at 540 nm was measured using a microplate reader.

2.7. Measurement of NO concentration

RAW 264.7 macrophages were cultured in 96‐well plates, under the conditions mentioned above. After pretreating with varying concentrations of AME for 4 hr, the cells were treated with LPS (10 ng/ml) and cultured for 24 hr. The cell culture broth was retrieved, mixed with Griess reagent at a 1:1 ratio, and incubated in the dark at room temperature for 10 min. Absorbance at 540 nm was measured using a microplate reader.

2.8. Western blotting

Raw 264.7 macrophages were seeded in a 6‐well plate (2 × 105 cells/well) and cultured for 24 hr. After pretreating with varying concentrations of AME for 4 hr, the cells were treated with LPS (10 ng/ml) and cultured for another 24 hr. The medium was removed; the cells were washed with PBS and then centrifuged at 5,000 g for 3 min. The pellet thus obtained was resuspended in lysis buffer (GenDEPOT) and incubated for 30 min on ice, followed by sonication for 60 s. After confirming that the lysis buffer had lost its viscosity, the mixture was centrifuged at 10,000 g for 10 min to collect the supernatant. Proteins in the supernatant were quantified using BSA assay kit (Thermo Fisher Scientific Inc.). Quantified protein was mixed in 4× sample buffer and inactivated for 10 min at 100°C, followed by electrophoresis with 12% SDS poly‐acrylamide gel. The isolated proteins were transblotted onto polyvinylidene difluoride (PVDF) membrane, which was then blocked by 2‐hr incubation with 5% skim milk. The membrane was incubated with primary antibodies against iNOS (1:3,000, sc7271; Santa Cruz Biotechnology), COX‐2 (1:5,000, ab15191; Abcam), IL‐1β (1:3,000, ab9722; Abcam), TNF‐α (1:3,000, ab9739; Abcam), and GAPDH (1:3,000, ab9485; Abcam) for 24 hr at 4°C, and washed 3 times for 10 min with Tris‐buffered saline Tween‐20 (TBST). Secondary antibodies were diluted to a ratio of 1:5,000 and allowed bind for 2 hr at room temperature. After washing 3 times for 10 min with TBST buffer, the samples were treated with enhanced chemiluminescence (ECL) solution and imaged using the ChemiDoc system (Bio‐Rad).

2.9. Total RNA extraction and RT‐qPCR

RAW 264.7 macrophages, cultured as described in the earlier sections, were washed twice with PBS. Total RNA was isolated using the NucleoSpin® RNA Plus kit (Macherey‐Nagel) and quantified using NanoDrop One spectrophotometer (Thermo Fisher Scientific Inc.).Then, 1 μg of total RNA was reverse‐transcribed into cDNA using the High Capacity cDNA Reverse Transcription Kit (Thermo Fisher Scientific Inc.). Real‐time PCR was performed using 1 μl of the synthesized cDNA, together with 1 μl of primer, 10 μl of SYBR master mix, and 8 μl of RNA‐free water. The mRNA expression of all genes tested is normalized to the RPS3 expression. Gene‐specific information of the genes analyzed is provided in Table 1.

TABLE 1.

Sequences of primers used for RT‐qPCR

Primer name Sequence
iNOS
Forward AATCTTGGAGCGAGTTGTGG
Reverse CAGGAAGTAGGTGAGGGCTTG
COX‐2
Forward ACTCACTCAGTTTGTTGAGTCATT
Reverse TTTGATTAGTACTGTAGGGTTAATG
TNF‐α
Forward GCCACCACGCTCTTCTGCCT
Reverse GGCTGATGGTGTGGGTGAGG
MCP‐1
Forward TCTGGACCCATTCCTTCTTG
Reverse AGGTCCCTGTCATGCTTCTG
IL‐6
Forward CCAGAGATACAAAGAAATGATGG
Reverse ACTCCAGAAGACCAGAGGAAAT
IL‐1β
Forward AAATACCTGTGGCCTTGGGC
Reverse CTTGGGATCCACACTCTCCAG
RPS3
Forward ATCAGAGAGTTGACCGCAGTTG
Reverse AATGAACCGAAGCACACCATAG
Open in a new tab

2.10. Statistical analysis

Significance between groups was tested using GraphPad Prism version 5.02 (GraphPad Software Inc.), and the results were expressed as mean ± SEM. Significance of differences between the mean values for two groups was assessed using t test. For all statistical significance, differences with p‐values <.05 were considered significant.

3. RESULTS AND DISCUSSION

3.1. High TPC and TFC in AME

Phenols are the most abundant of the various physiologically active substances present in plants (Manach, Scalbert, Morand, Rémésy, & Jiménez, 2004). Polyphenolic compounds are largely divided into flavonoids and tannins. Of the two, flavonoids belong to the largest subgroup of polyphenols synthesized by plants. The total polyphenols present in AM were measured in terms of gallic acid equivalents, whereas the total flavonoids were measured in terms of quercetin equivalents. The AME showed a high gallic acid and quercetin equivalent contents of 31.65 and 0.28 mg/g, respectively (Table 2). The high content of polyphenols, which are antioxidative substances that convert active oxygen in the body into harmless substances, in AM confirms that AM would have excellent antioxidative effects.

TABLE 2.

Total polyphenol and flavonoid contents of Antirrhinum majus extracts

Sample Total polyphenol content Total flavonoid content
(mg GAE/g) (mg QE/g)
95% ethanol extract of Antirrhinum majus 31.65 ± 0.63 0.28 ± 0.02
Open in a new tab

Abbreviations: GAE, gallic acid equivalent; QE, quercetin equivalent.

3.2. Radical‐scavenging activities of AME

DPPH and ABTS assays were performed to analyze the antioxidative properties of AME in terms of radical‐scavenging ability. DPPH is a free radical with a unique color. However, when free radical DPPH reacts with an antioxidant, the free radicals are scavenged by the antioxidative action of the sample, thereby resulting in a color change from purple to yellow. The method based on this principle is widely used for measuring antioxidative activities (Bondet, Brand‐Williams, & Berset, 1997). The DPPH radical‐scavenging ability of AME increased as the concentration of AME increased (Figure 1). ABTS radical‐scavenging ability is measured on the basis of the principle that antioxidants within the sample scavenge ABTS free radicals produced by the reaction between the ABTS reagent and potassium persulfate, thereby resulting in the loss of cyan color. Similar to the results of the DPPH assay, the ABTS radical‐scavenging ability of AME increased as the concentration of AME increased. Both DPPH and ABTS radical‐scavenging abilities were the highest at the concentration of 10 μg/ml. This confirms that AME has high antioxidative activities.

FIGURE 1.

FIGURE 1

Open in a new tab

Antioxidant activity of Antirrhinum majus extract (AME). DPPH radical‐scavenging activity was measured for a mixture containing 0.15 mM DPPH and different concentrations of AME. For the ABTS assay, 7 mM ABTS solution was incubated with 2.45 mM potassium persulfate; and the ABTS radical‐scavenging ability of a mixture of 100 μl of this solution and different concentrations of AME was measured

3.3. Cytotoxicity assessment in RAW 264.7 cells

MTT assay was used to assess the cytotoxicity of AME in RAW 264.7 macrophages (Figure 2). Sample concentrations with cell viability of ≥90% relative to the control group were considered to be safe, non‐toxic concentrations. Cell viability measurement after treatment with 50, 100, 150, 300, and 500 μg/ml of AME showed cell viability of ≥90% up to the concentration of 300 μg/ml, revealing no cytotoxicity, whereas 500 μg/ml AME showed cell viability of <90%. On the basis of these results, the 50, 100, 150, and 300 μg/ml AME samples were selected for further experiments.

FIGURE 2.

FIGURE 2

Open in a new tab

Cytotoxicity of Antirrhinum majus extract (AME) in RAW 264.7 cells. RAW 264.7 cells were incubated for 24 hr in the presence or absence of AME at the indicated concentration. Cell viability was evaluated by the MTT assay. Data represent the mean ± SEM of triplicate determinations from three separate experiments. *p < .05 versus CON

3.4. Inhibitory effects of AME on NO production

Lipopolysaccharide are extracellular components of gram‐negative bacteria and they act as powerful stimuli for various cells such as monocytes and macrophages. In particular, when macrophages are activated after stimulation by LPS, they produce and release inflammatory mediators such as NO via regulation of pro‐inflammatory factors (Lin, Juan, Shen, Hsu, & Chen, 2003; Xu et al., 2014). Therefore, in the present study, we examined the effects of AME on the production of NO, which is one of the active oxygen species and is known to play a key role in inflammation induction. The amount of NO produced was measured in terms of NO2 − content of the cell culture broth by using the Griess reagent. Based on the results obtained, the inhibitory effects of AME against NO production by LPS‐stimulated RAW 264.7 cells were measured. NO production after LPS stimulation increased to 35.8 μM in the untreated cells. This LPS‐induced increase in NO was significantly inhibited by AME treatment in a concentration‐dependent manner (Figure 3). In particular, excellent inhibitory effect of ≥50% on NO production was found at the AME concentration of 300 μg/ml.

FIGURE 3.

FIGURE 3

Open in a new tab

Effect of Antirrhinum majus extract (AME) on nitrite production in RAW 264.7 cells. After pretreatment with the indicated concentration of AME for 4 hr, RAW 264.7 cells were treated with 10 ng/ml lipopolysaccharide (LPS) for 24 hr. Nitrite levels in the cell media were measured spectrophotometrically using Griess reagent. Data are provided as mean ± SEM. ***p < .01 versus CON; ### p < .05 versus LPS treatment only

3.5. Anti‐inflammatory effects of AME in RAW 264.7 cells

Nitric oxide, a mediator of inflammatory response, is biosynthesized by the enzymes NOS and COX. Therefore, we next performed a western blot analysis to determine whether AME also inhibits the protein expression of iNOS and COX‐2. After pretreating Raw 264.7 cells with LPS (10 ng/ml), the cells were treated with AME and cultured for 24 hr to observe expression of iNOS and COX‐2. The group that was not treated with LPS showed no iNOS and COX‐2 expression, whereas the LPS‐treated group treated showed increased expression of iNOS and COX‐2, compared to that observed in the control group. However, AME treatment showed strong inhibition of iNOS and COX‐2 expression in a concentration‐dependent manner (Figure 4a). Next, the inhibition of iNOS and COX‐2 mRNA expression was studied in AME‐treated RAW 264.7 cells. The results were similar to those obtained for experiments studying the expression of iNOS and COX‐2 proteins; the AME‐treated group showed inhibitory effects on mRNA expression, compared to the mRNA expression observed in the control group that was treated with LPS only (Figure 4b). These findings show that AME has strong inhibitory effects on NO production and early‐inflammatory factor production and that NO production is inhibited via inhibition of iNOS expression.

FIGURE 4.

FIGURE 4

Open in a new tab

Inhibitory effects of Antirrhinum majus extract (AME) on lipopolysaccharide (LPS)‐induced iNOS and COX‐2 expression. Cells were treated with AME for 4 hr followed by LPS treatment (10 ng/ml) for 24 hr. (a) Concentration‐dependent effect of AME on the expression of iNOS and COX‐2. Representative Western blot images and quantification data are shown. (b) mRNA expression of inflammatory genes. Cells were treated with agents for 24 hr, and RT‐qPCR was performed. Data are provided as mean ± SEM. **p < .01, ***p < .01 versus CON; # p < .05, ## p < .05, ### p < .05 versus LPS treatment only

3.6. Inhibitory effects of AME on pro‐inflammatory cytokine production

Macrophages secrete proteins called cytokines, which are signaling molecules that react with receptors on the cell surface to trigger certain reactions. LPS stimulates macrophages to secrete cytokines that typically play a central role in an inflammatory response, such as TNF‐α, IL‐6, IL‐1β, and MCP‐1. Therefore, Western blot analysis was performed to examine the inhibitory effects of AME on the expression of cytokines in RAW 264.7 cells activated by LPS (Figure 5a). The expression of TNF‐α and IL‐1β increased considerably in the group with RAW 264.7 cells treated with LPS, compared to that in the control group. However, the expression of TNF‐α and IL‐1β was significantly inhibited in the AME‐treated and LPS‐stimulated group. Moreover, by treating RAW 264.7 cells with AME, the expression of mRNA of pro‐inflammatory cytokines, TNF‐α, IL‐6, IL‐1β, and MCP‐1, was also found to be inhibited. Thus, compared to the control group that was treated with LPS only, the AME‐treated group showed similar inhibitory pattern for protein and mRNA expression of the studied inflammatory cytokines.

FIGURE 5.

FIGURE 5

Open in a new tab

Inhibitory effects of Antirrhinum majus extract (AME) on lipopolysaccharide (LPS)‐stimulated pro‐inflammatory cytokine expression in RAW 264.7 cells. RAW 264.7 cells were treated with AME for 4 hr followed by LPS treatment (10 ng/ml) for 24 hr. (a) The pro‐inflammatory cytokine protein level was measured by Western blotting. The experiments were repeated at least three times, and representative Western blot images and quantification data are shown. (b) mRNA levels for inflammatory genes, as measured by RT‐qPCR in RAW 264.7 cells. **p < .01, ***p < .01 versus CON; # p < .05, ## p < .05, ### p < .05 versus LPS treatment only

Antirrhinum majus, a plant with the edible flowers used as a food item in Korea, has attracted attention as a rich source of phytochemicals, which are beneficial for human health. It has been used as an alternative therapy and traditional medicine to treat various diseases. However, its biological properties have not been fully elucidated. Therefore, in the present study, we investigated the effects of flavonoids involved in Korean AM. AME prevented the induction of iNOS and COX‐2 by LPS in RAW 264.7 macrophages. Furthermore, AME suppressed both the LPS‐induced production of pro‐inflammatory cytokines, at the mRNA and protein level, in macrophages. Although the anti‐inflammatory effects of AME are known, this is the first report demonstrating that AME inhibits inflammation in macrophages.

Flavonoids are widely existed in the plants and various biological functions in previous studies (Kumar & Pandey, 2013; Lopes, Schulman, & Hermes‐Lima, 1999). These studies suggest that flavonoids have various pharmacological properties such as anti‐cancer, antioxidant, antihypertensive, antiatherosclerotic, and anti‐inflammatory properties (Lee et al., 2012; Ray et al., 1999; Wang et al., 1999). Of these biological properties, the anti‐inflammatory effects of flavonoids have used for a long time. Inflammation is the normal physiological response to a variety of tissue injuries, chemical agents, and bacterial infection (Ariel, Maridonneau‐Parini, Rovere‐Querini, Levine, & Mühl, 2012; Khan et al., 2011; Liu, Zou, Chai, & Yao, 2014). Macrophages play a significant role in inflammation by overproducing pro‐inflammatory cytokines and inflammatory mediators (Chawla, Nguyen, & Goh, 2011; Laskin, 2009; Wynn, Chawla, & Pollard, 2013). Chronic inflammation causes the increase of pro‐inflammatory mediators, including iNOS and COX‐2, and various cytokines, including MCP‐1, TNF‐α, IL‐6, and IL‐1β (Gabay, 2006; Kushner, 1993). NO, synthesized by iNOS, has been recognized as important inflammatory mediator and a mediator of cell communication in macrophages (Wang et al., 1999; Wojdasiewicz et al., 2014). iNOS is expressed after exposure to specific stimulants such as LPS and cytokines (Wong et al., 1986). Thus, the regulation of iNOS and COX‐2 is important in the inflammation response.

4. CONCLUSIONS

In this study, an ethanol extract of AM was prepared and its antioxidative and anti‐inflammatory properties were investigated. AME showed high polyphenolic and flavonoid contents as well as high DPPH and ABTS radical‐scavenging ability. To test the cytotoxicity of AME, the viability of RAW 264.7 macrophages was assessed. The anti‐inflammatory effects of AME were investigated via inhibition of NO production and changes in expression of inflammation‐related proteins and mRNA in RAW 264.7 cells activated by LPS stimulation. Exposure of RAW 264.7 macrophages to LPS was associated with an accumulation of NO in the culture medium, suggesting an enhanced NO production. This LPS‐induced NO production by Raw 264.7 macrophages was inhibited by AME, in a concentration‐dependent manner. Furthermore, AME inhibited iNOS and COX‐2 protein expression in a concentration‐dependent manner in LPS‐stimulated Raw 264.7 macrophages. Investigation of inhibitory effects on expression of cytokines that play a central role in inflammatory stage showed that AME treatment inhibited the LPS‐induced expression of TNF‐α and IL‐1β protein expression as well as the mRNA levels of TNF‐α, IL‐6, IL‐1β, and MCP‐1 in Raw 264.7 macrophages in a concentration‐dependent manner. Collectively, these findings indicate that AME showed anti‐inflammatory effects in macrophages. Although more detailed research is needed to obtain further insights into the exact anti‐inflammatory mechanism of AME, the findings of our study may serve as important basic data for studies on anti‐inflammatory substances via extraction of active ingredients from AM, as well as studies on isolation and mechanism of action of inflammation‐inhibiting components for prevention or treatment purposes.

CONFLICT OF INTEREST

The authors declare that they do not have any conflict of interest.

ETHICAL APPROVAL

This study does not involve any human or animal testing.

Jang M, Hwang I, Hwang B, Kim G. Anti‐inflammatory effect of Antirrhinum majus extract in lipopolysaccharide‐stimulated RAW 264.7 macrophages. Food Sci Nutr. 2020;8:5063–5070. 10.1002/fsn3.1805

Funding information

This research is supported by National Institute of Agricultural Sciences project of South of Korea (PJ0135352020).

REFERENCES

  1. Ariel, A. , Maridonneau‐Parini, I. , Rovere‐Querini, P. , Levine, J. S. , & Mühl, H. (2012). Macrophages in inflLPSmation and its resolution. Frontiers in Immunology, 3, 324 10.3389/fimmu.2012.003240 [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Blois, M. S. (1958). Antioxidant determinations by the use of a stable free radical. Nature, 181, 1199–1200. 10.1038/1811199a0 [DOI] [Google Scholar]
  3. Bondet, V. , Brand‐Williams, W. , & Berset, C. (1997). Kinetics and mechanisms of antioxidant activity using the DPPH free radical method. LWT ‐ Food Science and Technology, 30(6), 609–615. 10.1006/fstl.1997.0240 [DOI] [Google Scholar]
  4. Charo, I. F. , & Taub, R. (2011). Anti‐inflammatory therapeutics for the treatment of atherosclerosis. Nature Reviews Drug Discovery, 10(5), 365–376. 10.1038/nrd3444 [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Chawla, A. , Nguyen, K. D. , & Goh, Y. P. (2011). Macrophage‐mediated inflammation in metabolic disease. Nature Reviews Immunology, 11(11), 738–749. 10.1038/nri3071 [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Choi, Y. H. , Kim, G. Y. , & Lee, H. H. (2014). Anti‐inflammatory effects of cordycepin in lipopolysaccharide‐stimulated RAW 264.7 macrophages through Toll‐like receptor 4‐mediated suppression of mitogen‐activated protein kinases and NF‐κB signaling pathways. Drug Design, Development and Therapy, 8, 1941–1953. 10.2147/DDDT.S71957 [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Deans, K. A. , & Sattar, N. (2006). “Anti‐inflammatory” drugs and their effects on type 2 diabetes. Diabetes Technology & Therapeutics, 8(1), 18–27. 10.1089/dia.2006.8.18 [DOI] [PubMed] [Google Scholar]
  8. Di Penta, A. , Moreno, B. , Reix, S. , Fernandez‐Diez, B. , Villanueva, M. , Errea, O. , … Villoslada, P. (2013). Oxidative stress and proinflammatory cytokines contribute to demyelination and axonal damage in a cerebellar culture model of neuroinflammation. PLoS One, 8(2), e54722 10.1371/journal.pone.0054722 [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Dinarello, C. A. (2010). Anti‐inflammatory agents: Present and future. Cell, 140(6), 935–950. 10.1016/j.cell.2010.02.043 [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Elmarakby, A. A. , & Sullivan, J. C. (2012). Relationship between oxidative stress and inflammatory cytokines in diabetic nephropathy. Cardiovascular Therapeutics, 30(1), 49–59. 10.1111/j.1755-5922.2010.00218.x [DOI] [PubMed] [Google Scholar]
  11. Fiorucci, S. , Mencarelli, A. , Distrutti, E. , Baldoni, M. , delSoldato, P. , & Morelli, A. (2004). Nitric oxide regulates immune cell bioenergetic: A mechanism to understand immunomodulatory functions of nitric oxide‐releasing anti‐inflammatory drugs. The Journal of Immunology, 173, 874–882. 10.4049/jimmunol.173.2.874 [DOI] [PubMed] [Google Scholar]
  12. Gabay, C. (2006). Interleukin‐6 and chronic inflammation. Arthritis Research & Therapy, 8(Suppl2), S3 10.1186/ar1917 [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Halliwell, B. , & Gutteridge, J. M. (1984). Oxygen toxicity, oxygen radicals, transition metals and disease. The Biochemical Journal, 219(1), 1–14. 10.1042/bj2190001 [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Halliwell, B. , & Gutteridge, J. M. (2015). Free radicals in biology and medicine (p. 968). New York, NY: Oxford University Press Co. [Google Scholar]
  15. Hybertson, B. M. , Gao, B. , Bose, S. K. , & McCord, J. M. (2011). Oxidative stress in health and disease: The therapeutic potential of Nrf2 activation. Molecular Aspects of Medicine, 32(4–6), 234–246. 10.1016/j.mam.2011.10.006 [DOI] [PubMed] [Google Scholar]
  16. Jamie, L. M. , Davis, J. M. , Jennifer, L. S. , Reilly, T. E. , Jung, S. H. , James, A. C. , … Murphy, E. A. (2012). Linking tumor‐associated macrophages, inflammation, and intestinal tumorigenesis: Role of MCP‐1. American Journal of Physiology‐Gastrointestinal and Liver Physiology, 303(10), G1087–G1095. 10.1152/ajpgi.00252.2012 [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Khan, S. , Shin, E. M. , Choi, R. J. , Jung, Y. H. , Kim, J. , Tosun, A. , & Kim, Y. S. (2011). Suppression of LPS‐induced inflammatory and NF‐κB responses by anomalin in RAW 264.7 macrophages. Journal of Cellular Biochemistry, 112(8), 2179–2188. 10.1002/jcb.23137 [DOI] [PubMed] [Google Scholar]
  18. Kumar, S. , & Pandey, A. K. (2013). Chemistry and biological activities of flavonoids: An overview. The Scientific World Journal, 29(2013), 162750 10.1155/2013/162750 [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Kushner, I. (1993). Regulation of the acute phase response by cytokines. Perspectives in Biology and Medicine, 36(4), 611–612. 10.1353/pbm.1993.0004 [DOI] [PubMed] [Google Scholar]
  20. Laskin, D. L. (2009). Macrophages and inflammatory mediators in chemical toxicity: A battle of forces. Chemical Research in Toxicology, 22(8), 1376–1385. 10.1021/tx900086v [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Lee, E. J. , Kim, C. , Kim, J. Y. , Kim, S. M. , Nam, D. , Jang, H. J. , … Ahn, K. (2012). Inhibition of LPS‐induced inflammatory biomarkers by ethyl acetate fraction of Patrinia scabiosaefolia through suppression of NF‐κB activation in RAW 264.7 cells. Immunopharmacology and Immunotoxicology, 34(2), 282–291. 10.3109/08923973.2011.602412 [DOI] [PubMed] [Google Scholar]
  22. Lin, H. Y. , Juan, S. H. , Shen, S. C. , Hsu, F. L. , & Chen, Y. C. (2003). Inhibition of lipopolysaccharide‐induced nitric oxide production by flavonoids in RAW264.7 macrophages involves heme oxygenase‐1. Biochemical Pharmacology, 66(9), 1821–1832. 10.1016/s0006-2952(03)00422-2 [DOI] [PubMed] [Google Scholar]
  23. Liu, Y. C. , Zou, X. B. , Chai, Y. F. , & Yao, Y. M. (2014). Macrophage polarization in inflammatory diseases. International Journal of Biological Sciences, 10(5), 520–529. 10.7150/ijbs.8879 [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Lopes, G. K. , Schulman, H. M. , & Hermes‐Lima, M. (1999). Polyphenol tannic acid inhibits hydroxyl radical formation from Fenton reaction by complexing ferrous ions. Biochimica et Biophysica Acta, 1472(1–2), 142–152. 10.1016/S0304-4165(99)00117-8 [DOI] [PubMed] [Google Scholar]
  25. Manach, C. , Scalbert, A. , Morand, C. , Rémésy, C. , & Jiménez, L. (2004). Polyphenols: Food sources and bioavailability. American Journal of Clinical Nutrition, 79(5), 727–747. 10.1093/ajcn/79.5.727 [DOI] [PubMed] [Google Scholar]
  26. Moreno, M. I. , Isla, M. I. , Sampietro, A. R. , & Vattuone, M. A. (2000). Comparison of the free radical‐scavenging activity of propolis from several regions of Argentina. Journal of Ethnopharmacology, 71(1–2), 109–114. 10.1016/s0378-8741(99)00189-0 [DOI] [PubMed] [Google Scholar]
  27. Ohashi, K. , Shibata, R. , Murohara, T. , & Ouchi, N. (2014). Role of anti‐inflammatory adipokines in obesity‐related diseases. Trends in Endocrinology and Metabolism, 25(7), 348–355. 10.1016/j.tem.2014.03.009 [DOI] [PubMed] [Google Scholar]
  28. Olefsky, J. M. , & Glass, C. K. (2010). Macrophages, inflammation, and insulin resistance. Annual Review of Physiology, 72, 219–246. 10.1146/annurev-physiol-021909-135846 [DOI] [PubMed] [Google Scholar]
  29. Ray, P. S. , Maulik, G. , Cordis, G. A. , Bertelli, A. A. , Bertelli, A. , & Das, D. K. (1999). The red wine antioxidant resveratrol protects isolated rat hearts from ischemia reperfusion injury. Free Radical Biology and Medicine, 27(1–2), 160–169. 10.1016/s0891-5849(99)00063-5 [DOI] [PubMed] [Google Scholar]
  30. Re, R. , Pellegrini, N. , Proteggente, A. , Pannala, A. , Yang, M. , & Rice‐Evans, C. (1999). Antioxidant activity applying an improved ABTS radical cation decolorization assay. Free Radical Biology and Medicine, 26(9–10), 1231–1237. 10.1016/s0891-5849(98)00315-3 [DOI] [PubMed] [Google Scholar]
  31. Reuter, S. , Gupta, S. C. , Chaturvedi, M. M. , & Aggarwal, B. B. (2010). Oxidative stress, inflammation, and cancer: How are they linked? Free Radical Biology and Medicine, 49(11), 1603–1616. 10.1016/j.freeradbiomed.2010.09.006 [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Satish, L. D. , Sergey, K. , Shohreh, A. , & Bassel, E. S. (2009). Monocyte chemoattractant protein‐1 (MCP‐1): An overview. Journal of Interferon and Cytokine Research, 29(6), 313–326. 10.1089/jir.2008.0027 [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Scheller, J. , Chalaris, A. , Schmidt‐Arras, D. , & Rose‐John, S. (2011). The pro‐ and anti‐inflammatory properties of the cytokine interleukin‐6. Biochimica et Biophysica Acta, 1813(5), 878–888. 10.1016/j.bbamcr.2011.01.034 [DOI] [PubMed] [Google Scholar]
  34. Singleton, V. L. , & Rossi, J. A. (1965). Colorimetry of total phenolics with phosphomolybdic‐phosphotungstic acid reagents. American Journal of Enology and Viticulture, 16, 144–158. [Google Scholar]
  35. Uttara, B. , Singh, A. V. , Zamboni, P. , & Mahajan, R. T. (2009). Oxidative stress and neurodegenerative diseases: A review of upstream and downstream antioxidant therapeutic options. Current Neuropharmacology, 7(1), 65–74. 10.2174/157015909787602823 [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Valko, M. , Leibfritz, D. , Moncol, J. , Cronin, M. T. , Mazur, M. , & Telser, J. (2007). Free radicals and antioxidants in normal physiological functions and human disease. International Journal of Biochemistry & Cell Biology, 39(1), 44–84. 10.1016/j.biocel.2006.07.001 [DOI] [PubMed] [Google Scholar]
  37. Valledor, A. F. , Comalada, M. , Santamaría‐Babi, L. F. , Lloberas, J. , & Celada, A. (2010). Macrophage proinflammatory activation and deactivation: A question of balance. Advances in Immunology, 108, 1–20. 10.1016/B978-0-12-380995-7.00001-X [DOI] [PubMed] [Google Scholar]
  38. Wang, M. , Shao, Y. , Li, J. , Zhu, N. , Rangarajan, M. , LaVoie, E. J. , & Ho, C. T. (1999). Antioxidative phenolic glycosides from sage (Salvia officinalis). Journal of Natural Products, 62(3), 454–456. 10.1021/np980436g [DOI] [PubMed] [Google Scholar]
  39. Wojdasiewicz, P. , Poniatowski, Ł. A. , & Szukiewicz, D. (2014). The role of inflammatory and anti‐inflammatory cytokines in the pathogenesis of osteoarthritis. Mediators of Inflammation, 2014, 561459 10.1155/2014/561459 [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Wong, D. G. , Spence, J. D. , Lamki, L. , Freeman, D. , & McDonald, J. W. (1986). Effect of non‐steroidal anti‐inflammatory drugs on control of hypertension by beta‐blockers and diuretics. Lancet, 1(8488), 997–1001. 10.1016/s0140-6736(86)91271-7 [DOI] [PubMed] [Google Scholar]
  41. Wynn, T. A. , Chawla, A. , & Pollard, J. W. (2013). Macrophage biology in development, homeostasis and disease. Nature, 496(7446), 445–455. 10.1038/nature12034 [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Xu, X. , Yin, P. , Wan, C. , Chong, X. , Liu, M. , Cheng, P. , … Xu, J. (2014). Punicalagin inhibits inflammation in LPS‐induced RAW264.7 macrophages via the suppression of TLR4‐mediated MAPKs and NF‐κB activation. Inflammation, 37(3), 956–965. 10.1007/s10753-014-9816-2 [DOI] [PubMed] [Google Scholar]

Articles from Food Science & Nutrition are provided here courtesy of Wiley

RESOURCES