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. 2021 Dec 31;31(1):101–110. doi: 10.1007/s10068-021-01017-4

Immune-modulation effect of Halocynthia aurantium tunic lipid on RAW264.7 cells

A-yeong Jang 1,#, Chaiwat Monmai 2,#, Weerawan Rod-In 2, Ji-Eun Kim 1, SangGuan You 2, Tae Ho Lee 3, Woo Jung Park 1,2,✉
PMCID: PMC8733137  PMID: 35059234

Abstract

The current study evaluated the immune-regulatory potential of lipid extract from Halocynthia aurantium tunic on macrophage cells. The results showed that H. aurantium lipid is composed of primarily SFA (68.32%), followed by MUFA and PUFA (17.61% and 14.07%, respectively). Halocynthia aurantium lipid dose-dependently modulated the NO and PGE2 production in RAW264.7 cells without any LPS stimulation. The lipid effectively up-regulated the cytokine expression, including IL-1β, IL-6, and TNF-α in RAW264.7 cells. The COX-2 expression as a key biomarker for inflammation was also significantly increased. Conversely, H. aurantium lipid down-regulated the expression of inflammatory cytokines in LPS-stimulated RAW264.7 cells. Halocynthia aurantium lipid modulated the phosphorylation of NF-κB p-65, p38, ERK, and JNK, indicating that this lipid activated through NF-κB and MAPK pathways. These results provide insight into the immune-regulatory activities of H. aurantium tunic lipid and suggest that H. aurantium tunic may a potential lipid source for immune-regulatory molecules.

Keywords: Halocynthia aurantium, Lipid, Immune-regulation, NF-κB, MAPK

Introduction

Immunomodulation is one of the most important physiological mechanisms and is closely related to the occurrence and prevention of various diseases including cancer, inflammatory diseases, and infectious diseases (Chen and Mellman, 2017; Latinne and Fiasse, 2006; Sam et al., 2018). Inflammation is one of the body's defenses to repair damaged areas and it occurs via regulation of releasing inflammatory mediators when external factors such as bacteria or viruses stimulate the immune system (Fujiwara and Kobayashi, 2005). Macrophages are the primary cells in the initiation of immune response (Monmai et al., 2018) and they lead to phagocytosis as a protective role to primarily remove infectious pathogens and cancer cells. Those cells also act as important mediators of immune response by secretion of various cytokines and physiological process such as antigen presentation (Nathan, 1987). When they are activated in inflammation, they produce NO, TNF-α, IL-1β, and IL-6 and especially NO is generated by inducible nitric oxide synthase (iNOS) and acts as an important mediator in getting rid of immune stimuli and microorganisms such as bacteria and viruses (Fujiwara and Kobayashi, 2005; Korhonen et al., 2005).

Lipids such as fat-soluble vitamins, cholesterol and essential fatty acids are important components for human health and development and play important roles in preventing disease (Roche, 1999). These have been known to give effects on immune system (Riccioni et al., 2003; Vitale and Broitman, 1981). Especially, polyunsaturated fatty acids (PUFAs) including omega-3 (n − 3) and omega-6 (n − 6) PUFAs contain two or more double bonds that controlled a wide range of functions in the body, including blood pressure, hematic clotting, correct development and function of the brain and nervous system (Gammone et al., 2018). PUFAs have reported that they decrease the production of TNF-α, IL-1β, and IL-6 in response to LPS (Kang and Weylandt, 2008). PUFAs have been reported a function in reducing the regulation of inflammation in macrophages by modulating the NF-κB and MAPK activity (Lo et al., 2000; Novak et al., 2003).

Sea squirts are known to have various substances that improve human health (Oh et al., 2019). They inhibit the reverse transcription of the immunodeficiency virus (Murakami et al., 2000), show an anti-proliferative effect on various tumor cells (Loya et al., 1997), perform inhibition of both peroxides and nitric oxide generation (Nathan, 2002). Bioactive compounds extracted from sea squirts were reported with antimicrobial, anti-oxidant, anti-lipase, immune-stimulation, and anti-cancer activities (Heu et al., 2013; Kwon et al., 2011; Monmai et al., 2018; Oh et al., 2019). One of the sea squirts, Halocynthia roretzi, has been reported to contain various lipid-associated functional ingredients such as EPA, DHA, and carotenoids (Mikami et al., 2010). It is also known that the tunic of H. roretzi includes high levels of carotenoids and other nutrients (Rohmah et al., 2016). Konishi et al. (2008) also reported acetylene carotenoids in H. roretzi lipids to reduce the production of LPS-induced pro-inflammatory cytokines (Konishi et al., 2008).

Halocynthia aurantium, which grows on the east coast of Korea, is a valuable organism but has not been studied as much as H. roretzi even though there is a report that H. aurantium tunic lipid containing 53.10% of PUFAs (Fomenko et al., 2013). In particular, studies associated with the relation between total lipids extracted from the tunic of H. aurantium and immune regulation have not been reported. Therefore, this study investigated the immunomodulatory effect of lipid extracted from H. aurantium tunic on RAW264.7 macrophages.

Materials and methods

Halocynthia aurantium sample and lipid extraction

The sea squirts, H. aurantium, were purchased from the local market of the East Sea near Gangwon Province, South Korea. Their tunic was separated and collected and the lipid was extracted according to the modified method of Bligh and Dyer (1959). Briefly, 4.5 g of dried sample was homogenized in a Waring Blender for 2 min with a mixture of chloroform and methanol in the ratio of 2:1 (30 mL of chloroform: 15 mL of methanol). After centrifuge, the homogenate was filtered and collected in a separator funnel. The filtration was centrifuged to make the solution into two layers and then the volume of the chloroform layer was collected. The collected layer was dried and dissolved in DMSO at the final concentration of 20 mg/mL (as 100%) for the immuno-modulatory activity assays.

Gas chromatography (GC) analysis

Fatty acids were extracted from H. aurantium tunic lipid following the methods of Garces and Mancha (1993) and the modified one-step lipid extraction (Park et al., 2009) to prepare fatty acid methyl esters (FAMEs). The FAMEs were analyzed by an Agilent 7890A gas chromatograph (GC), equipped with flame ionization detection (FID) (Perkin Elmer, Waltham, MA, USA), and an HP- FFAP capillary column (30 m × 0.32 mm, 0.25 µm of film thickness). The condition of column temperature was initially maintained at 150 °C for 2 min, increased to 235 °C at 3.5 °C/min and then maintained at 235 °C for 10 min. Quantification of the fatty acid compositions was identified from chromatographic peaks by comparing their retention times with internal standard C17:0 (Sigma-Aldrich, St. Louis, MO, USA).

Cell culture

RAW264.7 cells were obtained from Korean Cell Line Bank (KCLB, Korea). Cells were grown in RPMI-1640 medium (Gibco™, Waltham, MA, USA), supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin. Cells were incubated at 37 °C in a humidified 5% CO2 atmosphere and maintained through the weekly passage.

Macrophage viability assay

RAW264.7 cells at a number of 1 × 105 cells/well in a 96-well plate were treated with the various concentrations of H. aurantium tunic lipid (0.25%, 0.50%, 0.75%, or 1.00%) for 24 h. EZ-cytox cell viability assay kit (Daeil Labservice, Seoul, Korea) was used to determine the cytotoxicity of the tunic lipid. The ratio of cell viability (%) was calculated based on the following formula:

Macrophageviabilityratio%=A450ofthetestgroupA450ofthecontrolgroup×100

where A450 is the absorbance of the treatment at 450 nm.

NO production assay

The accumulation of nitrite in culture media was used as an indicator of NO production. RAW264.7 cells were pre-treated with various concentrations of H. aurantium tunic lipid for 1 h. For stimulation, cells were added with or without 1 μg/mL of LPS (from Escherichia coli O111:B4, Sigma-Aldrich). After 24 h incubation, the Griess reagent (Sigma-Aldrich, USA) were used for nitric concentration and calculated the NO production ratio (%) according to the following formula:

NOproductionratio%=A540ofthetestgroupA540ofthecontrolgroup×100

where A540 is the absorbance of the treatment at 540 nm.

The control groups were untreated and the LPS group for immune-enhancement and anti-inflammation activity, respectively.

RNA isolation and cDNA synthesis

RAW264.7 cells (5 × 105 cells/well) were treated with H. aurantium tunic lipid for 1 h, followed by stimulation with or without LPS. After 24 h incubation, total RNA was isolated by Tri reagent® (Molecular Research Center, Cincinnati, OH, USA). The concentration of extracted RNA was measured using the nanophotometer (Implen, Germany). The cDNA synthesis was reverse-transcribed from 500 ng of RNA sample using the high capacity cDNA reverse transcription kit (Applied Biosystems, Carlsbad, CA, USA).

Measurement of immune gene expression by quantitative real-time PCR

Immune-associated gene (IL-1β, IL-6, COX-2, and TNF-α) expressions were determined with SYBR® Premix Ex Taq™ II (Takara Bio Inc., Shiga, Japan) in a QuantStudio™ 7 FlexReal-Time PCR System (ThermoFisher Scientific, Waltham, MA, USA). The mixture of PCR reaction consisted of 0.4 µM of each specific primer and 10 ng of cDNA templates using the condition at pre-denaturation (95 °C, 30 s) and denaturation (95 °C, 5 s) and annealing (60 °C, 5 s) for 40 cycles. The results were determined by comparing the cycle threshold (CT) of the melting/dissociation curve stages and normalized to the β-actin (Table 1).

Table 1.

The sequences of oligonucleotide primers used for macrophage test of immune genes

Gene Accession no. Sequence
IL-1β NM_008361.4

Forward: GGGCCTCAAAGGAAAGAATC

Reverse: TACCAGTTGGGGAACTCTGC
IL-6 NM_031168.2

Forward: AGTTGCCTTCTTGGGACTGA

Reverse: CAGAATTGCCATTGCACAAC
COX-2 NM_011198.4

Forward: AGAAGGAAATGGCTGCAGAA

Reverse: GCTCGGCTTCCAGTATTGAG
TNF-α D84199.2

Forward: ATGAGCACAGAAAGCATGATC

Reverse: TACAGGCTTGTCACTCGAATT
β-actin NM_007393.5

Forward: CCACAGCTGAGAGGGAAATC

Reverse: AAGGAAGGCTGGAAAAGAGC
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Western blot assay

Cells were harvested with radioimmunoprecipitation assay buffer (RIPA buffer) (Tech & Innovation, China) supplemented with 0.1% of protease inhibitor (Thermo Scientific, USA) and Pierce™ BCA Protein Assay Kit (Thermo Scientific, USA) were used to determine the protein concentration. Thirty micrograms of protein from each sample were loaded on SDS-PAGE and transferred to a polyvinylidene fluoride (PVDF) membrane. The membranes were incubated with antibodies specific for p-NF-κB p65, p-p38, p-ERK1/2, and p-JNK (Cell Signaling Technology, USA), and α-tubulin (Abcam, United Kingdom). To detect protein signals, the membranes were used Pierce® ECL Plus Western Blotting Substrate (Thermo Scientific, USA). The blot was summarized by the ChemiDoc XRS + imaging system (Bio-Rad, USA) with ImageLab software (version 4.1; Bio-Rad, USA).

PGE2 production assay

RAW264.7 cells (1 × 105 cells/well) were treated with H. aurantium lipid and stimulation by with or without 1 μg/mL of LPS. The plate was further incubated for 24 h. The cultured medium was collected in a micro-centrifuge tube and the tube was centrifuged at 3000 × g for 10 min. The supernatant was transferred to a new flat-bottomed 96-well plate. The PGE2 production was performed using the PGE2 ELISA kit (Enzo Life Sciences, Farmingdale, NY) in accordance with the manufacturer's instructions. The results were calculated based on a standard curve.

Effect of NF-κB and MAPK activation-inhibitors on TNF-α expression

RAW264.7 cells at 5 × 105 cells per well were seeded in a 24-wells plate and incubated for 24 h. Cells were treated with 100 nM of NF-κB activation inhibitor (Calbiochem, 481406) for 3 h, or 20 µM of MAPK activation-inhibitors which including ERK Inhibitor (Calbiochem, 328006), JNK Inhibitor II (Calbiochem, 420119), and p38 MAP Kinase Inhibitor (Calbiochem, 559389) for 1 h (Bhattacharyya et al., 2010). Cells were treated with varying concentrations of H. aurantium tunic lipid or 1 µg/mL of LPS. Total RNA was extracted from cells after 24 h incubation. TNF-α expression was examined by the real-time PCR.

Statistical analysis

The statistical software (Statistix 8.1) was used for analyzed statistical differences. Data are presented as means ± standard deviation. A one-way ANOVA was then used with Tukey’s honest significance test to determine differences in all treatment groups with p < 0.05.

Results and discussion

Fatty acid profiles of H. aurantium tunic lipid

The fatty acid compositions of total lipid from H. aurantium tunic were presented in Fig. 1. In the total fatty acids, the results showed that the lipid contains SFAs (68.32 ± 0.98%), MUFAs (17.61 ± 1.74%), and PUFAs (14.07 ± 1.00%). Stearic acid (18:0), palmitic acid (16:0), and myristic acid 14:0) were the most abundant of SFAs. Among MUFAs, oleic acid (OA, 18:1n-9) was found to be the highest content at 6.78 ± 0.28% of the total fatty acids. The PUFAs were represented by four major components, containing dihomo α-linolenic acid (20:3n3), eicosapentaenoic acid (EPA, 20:5n3), docosahexaenoic acid (DHA, 22:6n3), and linoleic acid (LA, 18:2n6) at about 4.09 ± 0.36, 3.88 ± 0.31, 3.38 ± 0.34, and 2.72 ± 0.23% of the total fatty acids, respectively. Similar to previous research, the total lipid in aqueous-alcohol extract from H. aurantium tunic consisted mainly of fatty acid composition of palmitic acid (23.80 ± 2.10%), stearic acid (17.37 ± 2.00%), EPA (10.98 ± 0.55%), DHA (5.76 ± 0.54%), oleic acid (4.89 ± 0.40%), myristic (4.63 ± 0.99%), and linoleic (1.13 ± 0.10%), but different the percentage of amount contents (Fomenko et al., 2013).

Fig. 1.

Fig. 1

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Fatty acid composition of lipid extracted from H. aurantium tunic. A The representative chromatogram of FAME analysis using GC-FID. B The percentage of fatty acids from H. aurantium tunic lipid. Data are presented as mean ± SD, n = 5. Different letters (a, b, c, d, and e) indicate significant differences (p < 0.05) between the amounts of fatty acids, which were obtained from the same of H. aurantium tunic lipid

Effect of H. aurantium tunic lipid on viability in macrophages

The yield of H. aurantium tunic lipid was 53.1 ± 1.7 mg which was 1.18 ± 0.04%. To determine the cytotoxicity of H. aurantium lipid, RAW264.7 cells were evaluated with various lipid-concentrations. As shown in Fig. 2A, B, tunic lipid at low concentration (0.25% and 0.50%) did not provide any cytotoxicity to RAW264.7 cells, however, the cellular viability was gradually decreased when the concentration of tunic lipid was increased (0.75% and 1.00%).

Fig. 2.

Fig. 2

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Effect of lipid extract from H. aurantium tunic on cellular cytotoxicity and NO production in macrophage. A The cellular cytotoxicity effect on macrophage viability in RAW264.7 cells. B The cellular cytotoxicity effect on macrophage viability in LPS-stimulated RAW264.7 cells. C The NO production effect in RAW264.7 cells. D The NO production effect in LPS-stimulated RAW264.7 cells. Data are presented as mean ± SD, n = 3. Different letters (a, b, and c) indicate statistical differences between treatment groups at p < 0.05

Effect of H. aurantium tunic lipid on NO production in macrophages

NO is an inorganic low molecular radical produced by nitric oxide synthase and it was released from macrophages is toxic to cells and promotes pro-inflammatory cytokine secretion to induce immune responses (Coleman, 2001). The immune-stimulation and anti-inflammatory effects of lipid were investigated by measuring the NO production. Figure 2C showed that tunic lipid significantly and dose-dependently increased the production of NO in RAW264.7 cells. The anti-inflammatory effect of tunic lipid using LPS to stimulate inflammation was shown in Fig. 2D. The result showed that tunic lipid significantly suppressed LPS-stimulated NO production in a dose-dependent manner.

Effect of H. aurantium tunic lipid on immune-associated gene expressions in macrophages

Macrophages play an essential role in inflammation and led to the production of pro-inflammatory cytokines such as IL-1β, IL-6, and TNF-α, COX-2 (Fujiwara and Kobayashi, 2005; Kim et al., 2007). Gene expressions of pro-inflammatory cytokines have been known to regulate the immune system in macrophages. In RAW264.7 cells, immune-associated gene expressions were increased in a dose-dependent depending on the lipid concentration (Fig. 3A). Treatment with 1.00% tunic lipid gave the highest effect in all cytokine gene expression. However, mRNA expressions in LPS-stimulated RAW264.7 cells dose-dependently were suppressed (Fig. 3B). In addition, COX-2 expression also up-regulated and down-regulated in RAW264.7 cells according to the concentration (Fig. 3C, D). Similarly, the lipid extracts from the Nostoc commune showed that the gene expression of pro-inflammatory cytokines including TNF-α, COX-2, IL-1β, IL-6, and iNOS in macrophages (Park et al., 2008). These results suggest that the addition of H. aurantium tunic lipid inhibited and enhanced the gene expression in RAW264.7 cells.

Fig. 3.

Fig. 3

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Effect of lipid extract from H. aurantium tunic on the immune-associated gene expression and the production of PGE2 in macrophage. A The cytokine expression in RAW264.7 cells. B The cytokine expression in LPS-stimulated RAW264.7 cells. C The COX-2 expression in RAW264.7 cells. D The COX-2 expression in LPS-stimulated RAW264.7 cells. E The PGE2 production effect in RAW264.7 cells. F The PGE2 production effect in LPS-stimulated RAW264.7 cells. Data are presented as mean ± SD, n = 3. Different letters (a, b, c, d, and e) indicate statistical differences between treatment groups at p < 0.05

Effect of lipid from H. aurantium tunic on PGE2 production

PGE2 is produced from arachidonic acid which is produced during inflammation (Li et al., 2018). The production of PGE2 was measured for evaluation of the immune-regulatory effect of H. aurantium tunic lipid. As shown in Fig. 3E, lipid extracted from H. aurantium tunic significantly and dose-dependently enhanced the PGE2 production in RAW264.7 cells. Figure 3F showed that stimulation of RAW264.7 cells with LPS also increased the production of PGE2 and the treatment of different concentrations of H. aurantium lipid suppressed the LPS-induced PGE2 production.

Effect of H. aurantium tunic lipid on MAPK and NF-κB signaling pathways

NF-κB regulates the expression of pro-inflammatory cytokines and inflammatory mediators such as iNOS and COX-2 to control the immune system. The signaling molecule belonging to MAPK are p38, ERK, and JNK, which is important for cell growth and cell differentiation. The activation of MAPK and NF-κB signaling pathways by lipid extract from marine sources can regulate inflammatory cytokine expression and inflammatory processes (Li et al., 2012; Park et al., 2008; Rod-In et al., 2019). To deepen the understanding of the molecular mechanism of how H. aurantium lipid exerts its immune-regulation, immune signaling pathways were examined. The effect of H. aurantium lipid on the activation of NF-κB signaling was measured through the phosphorylation of NF-κB p-65. MAPK activation was also examined with the phosphorylation of JNK, p38 and ERK1/2. As shown in Fig. 4A and B, H. aurantium lipid dose-dependently increased the phosphorylation of NF-κB p-65, JNK, ERK, and p-38 levels. In contrast, H. aurantium lipid suppressed the protein expression levels under LPS-stimulation in RAW264.7 cells (Fig. 4C, D).

Fig. 4.

Fig. 4

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The effect of H. aurantium tunic lipid on proteins associated with NF-κB and MAPK pathways. A Western blot of proteins from RAW264.7 cells. B Relative band intensity of proteins from RAW264.7 cells. C Western blot of proteins from LPS-stimulated RAW264.7 cells. D Relative band intensity of proteins from LPS-stimulated RAW264.7 cells. Data are presented as mean ± SD, n = 3. Different letters (a, b, c, d, and e) indicate statistical differences between treatment groups at p < 0.05

Effect of lipid from H. aurantium tunic on MAPK and NF-κB inhibited-RAW264.7 cell

TNF-α expression was used to evaluate the effect of H. aurantium lipid on the mechanism of immune-regulation which is known to play a crucial role in immunological and inflammatory reactions (Nimal et al., 2006). RAW264.7 cells were incubated with NF-κB and MAPKs (ERK, JNK, p38) activation-inhibitors and TNF-α expression were measured compared with various concentration of lipid treated-RAW264.7 cells. As shown in Fig. 5, treatment of H. aurantium tunic lipid (0.25–1.00%) showed dose-dependently increased the TNF-α expression in the NF-κB and MAPKs signaling inhibited-macrophages. In NF-κB and ERK inhibited-macrophages, treatment of 1.00% lipid decreased the expression of TNF-α (32.35 ± 0.76% and 50.45 ± 1.49%) compared to the treatment in RAW264.7 cells (non-inhibitor). Among NF-κB and MAPKs signaling, 1% lipid-treated group showed the highest TNF-α expression in the JNK inhibited-RAW264.7 cells.

Fig. 5.

Fig. 5

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TNF-α expression in H. aurantium tunic lipid-treated NF-κB and MAPK activation inhibited RAW264.7 cells. Data are presented as mean ± SD, n = 3. Different letters (a, b, c, d, and e) indicate statistical differences between treatment groups at p < 0.05

The present study demonstrated that lipid from H. aurantium significantly regulates the key immune-associated biomarkers involving the expression of the immune-associated genes and the production of NO and PGE2. Those regulations led to further activation or inactivation of NF-κB p-65 and MAPKs (ERK1/2, JNK, and p38) depending on the inflammatory status, thus controlling the immune response. These results might be helpful to understand the immune-regulatory mechanism of H. aurantium tunic lipid on immune cells and moreover suggested that H. aurantium may be a potential marine source for immunomodulation in the human physiological system.

Acknowledgements

This study was supported by the University Emphasis Research Institute Support Program (No. 2018R1A61A03023584) from the National Research Foundation of Korea.

Declarations

Conflict of interest

The authors declare no conflict of interest.

Footnotes

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

A-yeong Jang and Chaiwat Monmai equally contributed to the study.

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