Immunomodulation activity of alkali extract polysaccharide from Plantago asiatic L. seeds

Abstract

Numerous polysaccharides have been isolated from plants and used as a source of biological response modifiers in therapeutic agents. We previously found a novel acidic polysaccharide from Plantago asiatic L. seeds, named PLP, having strong antioxidant abilities in vitro. To further elucidate the role of PLP as a biological response modifier, the immunomodulating activities of PLP in macrophage cells were explored in the current study. We confirmed that stimulation of murine macrophage cells with PLP resulted in cell proliferation. PLP could directly bind to macrophage with saturable and reversible character and required TLR2 and TLR4 participation. PLP interaction with TLR2 and TLR4 led to the activation of intracellular p38, ERK and JNK. Furthermore, specific inhibitors of p38, ERK and JNK could weaken the ability of PLP to induce macrophage cell proliferation. Overall, this study indicated for the first time the immunostimulating properties of PLP on macrophage cells through a receptor-mediated mechanism, which involves TLR2 and TLR4 MAPK signaling pathways, and highlighted the role of PLP as an efficacious biological modifier in oncologic immunotherapy.

---

1. Introduction

In recent decades, polysaccharide such as Plantago L. from herbaceous plants have garnered considerable attention as a rich source of novel bioactive compounds, which has been proven to be a healthy food and has advantages, including relaxing bowel, reducing blood fat and blood pressure.^1–4 Psyllium can also be used as a substitute for gluten in bread.⁵ In China, Plantago major, Plantago asiatic, and Plantago depressa have been used in functional foods, and P. asiatic is most widely distributed.^6–8

In our previous studies, a novel homogenous polysaccharide, referred to as PLP, was purified from P. asiatica L. seeds that inhabit in Jiangxi province, China. It has a backbone of β-1,4-linked Xylp with three α-GlcAp-(1 → 3)-Araf attached to the O-3 position and one α-T-linked-GlcAp and one α-Araf-(1 → 5)-Araf attached to the O-2 position every eight monosaccharide residues.⁹ In vitro experimental PLP showed scavenging abilities against hydroxyl, peroxyl anion, and DPPH radicals and displayed significant binding capacities against cholic and chenodeoxycholic acids.^10–12 The antioxidant activity and bile acid-binding capacities of PLP were closely connected to the number of chemical and molecular polysaccharide structure.^13–15

Although the mechanisms underlying the immunomodulating activity of polysaccharides need to be further searched, one of the primary mechanisms involves toll-like receptors (TLRs). The mammalian TLR family is a group of germ-line encoded receptors that trigger immune responses via recognition of structures conserved among microbial species known as pathogen-associated molecular patterns (PAMPs) such as LPS, peptidoglycan, and lipoprotein.^16,17 The family comprises at least 11 members, among which TLR2 and TLR4 are well characterized as the transmembrane receptors involved in the recognition of ligands containing carbohydrate moieties, e.g. LPS¹⁸ and various natural polysaccharides.^19–21 Upon sensing the presence of these ligands, TLRs trigger the downstream signaling cascade of MyD88/TIRAP-IRAK1-TRAF6-TAK1, which in turn results in the activation of mitogen-activated protein kinases (MAPKs), and further leads to the regulation of genes that orchestrate the proliferation, survival and immune responses.^22,23

In this study, we aimed to investigate the immunomodulating property of PLP in murine macrophage cells,^24–27 including cells proliferation, cytotoxicology assay, and discharge of NO, especially focusing on the involvement of TLR signaling in PLP-mediated macrophage cells responses. We find that PLP is capable of inducing proliferation in macrophage cells, the mechanism of which is direct, saturable and reversible binding of PLP to TLR2 and TLR4 with subsequent activation of MAPK signaling pathways. Collectively, these data indicate that PLP is an efficacious stimulant of macrophage cells, which may serve as a scientific foundation for developing their possible application in antitumor foods.

2. Experimental

2.1. Materials

DEME medium and FCS were obtained from Life Technologies (USA). Abs specific to mouse TLR2, TLR4 and the isotype controls were from Novus Biologicals (USA). ECL western blotting substrate was from BIORAD (USA). LPS, ConA, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), o-phenylenediamine, fluoresceinamine (FLA) and 1-cyano-4-dimethylaminopyridine tetrafluoroborate (CDAP) were from Sigma-Aldrich (USA). RAW 264.7 mouse macrophage cells were obtained from the Chinese Academy of Sciences (Shanghai, China).

2.2. Purification of polysaccharide PLP

The polysaccharide PLP was prepared from the dried seeds of P. asiatica L. following the protocols described previously. The purity of PLP was determined using a high-performance size exclusion chromatograph (Agilent, Santa Clara, CA, USA) equipped with a gel filtration column (Shodex SUGAR KS-805, 8 mm i.d. × 300 mm, Showa Denko, Japan) and a refractive index detector. Deionized water as the mobile phase was eluted at a flow rate of 1.0 ml min⁻¹.

2.3. Proliferation assay

Macrophage cells were cultured in 96-well microplates at a density of 2 × 10⁶ cells per ml in RPMI-1640 medium containing 10% FCS, supplemented with 60 mg l⁻¹ penicillin, 100 mg l⁻¹ streptomycin. The cells were stimulated with PLP (25–400 nM) and LPS (20 nM) for 48 h in a CO₂ incubator, followed by incubation with MTT (5 mg ml⁻¹) for another 4 h. The formazan crystals formed from MTT by living cells were fully dissolved in DMSO for 10 min. The absorbance was determined at 570 nm in a multiskan spectrum (Thermo Fisher Scientific, Vantaa, Finland) and induction of cell proliferation was expressed as the proliferation index, calculated by dividing A570 of stimulated cells with A570 of control cells.

2.4. Neutral red cell proliferation and cytotoxicology assay

Macrophage cells were cultured in 96-well microplates at a density of 2 × 10⁶ cells per ml in RPMI-1640 medium containing 10% FCS, supplemented with 60 mg l⁻¹ penicillin and 100 mg l⁻¹ streptomycin. The cells were stimulated with PLP (25–400 nM) and LPS (20 nM) for 24 h in a CO₂ incubator, followed by incubation with neutral red (20 μl) for another 2 h (Beyotime, Shanghai, China). Neutral red was dissolved in lysis buffer for 10 min. The absorbance was determined at 540 nm in a multiskan spectrum.

2.5. Discharge of NO

Macrophage cells were cultured in 96-well microplates at a density of 2 × 10⁶ cells per ml in RPMI-1640 medium containing 10% FCS, supplemented with 60 mg l⁻¹ penicillin and 100 mg l⁻¹ streptomycin. The cells were stimulated with PLP (25–400 nM) and LPS (20 nM) for 24 h in a CO₂ incubator, followed by the addition of 50 μl cell culture and 50 μl standards to a new 96-well microplates, and 50 μl Griess reagent I and Griess reagent II (Beyotime, Shanghai, China). The absorbance was determined at 540 nm in a multiskan spectrum.

2.6. Fluoresceinamine labeling of PLP

The polysaccharide PLP was conjugated to FLA using the CDAP-activation method, as previously described, with slight modifications. In brief, 10 mg of CDAP was added into an aqueous solution containing 30 mg of PLP with gentle stirring and maintained at pH 9.0 for 2.5 min. The CDAP-activated PLP was then mixed with 2 mg of FLA (pH adjusted to 8.0) and incubated at room temperature overnight. Fluoresceinamine-labeled PLP (fl-PLP) was separated from the excess free FLA with an Amicon Ultra-15 centrifugal filter unit (Millipore, Billerica, MA, USA). The FLA and PLP amounts in fl-PLP were quantified by measuring absorbance at 440 nm and phenol–sulfuric acid assay. Results for FLA labeling, expressed as mole of FLA per mole of PLP, were 62.0 ± 3.4.

2.7. Specific binding and competition assay

Macrophage cells were spotted on slides and stained with fl-PLP or FLA for 1 h on ice. Cells were then washed thrice with ice-cold PBS containing 1% BSA and photographed under a confocal laser scanning microscope (Leica, Solms, Germany). In another experiment, aliquots of macrophage cell suspensions at a density of 1 × 10⁶ cells per ml in PBS containing 1% BSA were incubated with fl-PLP at serial concentrations (25–200 nM) for 1 h. After three washes, cells were examined on a FACS Calibur flow cytometer (BD Biosciences, San Jose, CA, USA) with a 488 nm laser excitation and a 530 nm emission filter. Data were acquired from a minimum of 10 000 cells and analyzed using the FlowJo program (BD Biosciences, USA). Macrophage cells were incubated with a mixture of 100 nM fl-PLP and unlabeled PLP (200–3200 nM) for 1 h. The binding was then analyzed by flow cytometry as aforementioned.

2.8. Antibody blocking and signaling inhibition assay

Macrophage cells were pretreated with anti-TLR2, anti-TLR4 or respective isotype controls (20 μg ml⁻¹) at 37 °C for 2 h or with SB203580, U0126 and SP600125 (20 μM) at 37 °C for 30 min prior to addition of PLP (80 nM). Cell proliferation in the culture supernatants was determined 48 h later using an MTT assay.

2.9. Affinity adsorption precipitation and western blot analysis

Macrophage cells were collected for extraction of cytomembrane proteins using a membrane protein reagent extraction box. The polysaccharide PLP was conjugated to FLA using the CDAP-activation method, as previously described with slight modifications. Briefly, 5 mg CDAP was added into an aqueous solution containing 10 mg of PLP with gentle stirring and maintained at pH 9.0 for 5 min, and then it was added into 200 μl EAH-Sepharose 4B (GE Healthcare, USA). The PLP-Se4B were incubated and kept shaking at room temperature over 72 hours. The PLP-Se4B were then mixed with 1 ml cytomembrane proteins (2 mg ml⁻¹) and incubated at 4 °C over 2 hours. The polymer was washed four times with ice-cold PBS, then added into 50 μl 1× SDS loading buffer and heated for 3 min in boiling water, and finally the supernate was collected. The supernate was resolved on 12% SDS–polyacrylamide gel, electro-transferred onto a BioTrace NT nitrocellulose membrane (Pall, Ann Arbor, MI, USA) and then incubated in TBST buffer (50 mM Tris, pH 7.6, 150 mM NaCl, 0.05% Tween-20) containing 3% BSA at 37 °C for 2 h. The membrane was subsequently incubated with Abs against TLR2 and TLR4 overnight, followed by incubation with the corresponding secondary antibodies conjugated with horseradish peroxidase at 37 °C for 1 h. The protein bands were finally visualized using an enhanced chemiluminescence (ECL) system.

2.10. Statistical analysis

The Prism 6.0 program (GraphPad Software, La Jolla, CA, USA) was used for statistical analysis. Mann–Whitney U test or Kruskal–Wallis test followed by Dunn's post hoc test was performed to determine significant differences where needed.

3. Results and discussion

3.1. PLP promotes macrophage cells proliferation

Macrophage cells were cultured with PLP or LPS, a well-known cell stimulant, as the positive control. Cell proliferation was measured after 48 h by MTT assay. The results indicated that PLP significantly stimulated the macrophage cell to proliferate in a dose-dependent manner (Fig. 1A). The stimulation obtained with PLP was weaker than that mediated by equimolar amounts of LPS, suggesting that PLP may be a safe and effective immunostimulant, which does not induce acute and robust inflammation as LPS.

Fig. 1 PLP promotes macrophage cell proliferation, induces NO response and accelerates macrophage cell phagocytosis. (A) Effect of PLP on macrophage cell proliferation. Macrophage cells were treated with PLP or LPS at the indicated concentrations, and cell proliferation was measured by MTT assay at 48 h (*p ≤ 0.05 vs. medium, **p ≤ 0.01 vs. medium). (B) Effect of PLP on discharge of NO by NO text box (**p ≤ 0.01 vs. medium). (C) Effect of PLP on phagocytosis of macrophages by neutral red pinocyfic test (**p ≤ 0.01 vs. medium).

3.2. PLP stimulates macrophage cell phagocytosis and discharge of NO

Neutral red, on its intake by living cells, accumulates in the lysosomes. In cell proliferation and populations, this neutral red quantity increases. However, when a cell damage occurs, the intake of neutral red decreases. Therefore, depending on the ability of neutral red intake, the cell proliferation or toxicity can be determined. Macrophage cells were cultured with PLP and LPS, and the phagocytic extent was measured by a neutral red box. The results showed that PLP can accelerate macrophage cell phagocytosis (Fig. 1B).

NO is a type of endogenous vascular contraction factor, which is mainly caused due to activating macrophages and killing pathogenic microorganisms. Macrophage cells were cultured with PLP and LPS, and the discharge ability was measured by a NO box. The results indicated that PLP can promote the discharge of NO (Fig. 1C).

3.3. PLP binds to macrophage cells directly, saturably and reversibly

It is well-known that numerous polysaccharides can utilize their immunomodulating activities via direct binding to specific receptors or partners on immunocytes. To investigate whether PLP stimulated the macrophage cells through the same mechanisms, we prepared a fluoresceinamine-labeled probe of PLP (fl-PLP) and examined its direct binding capacity to macrophage cells by confocal microscopy analysis and flow cytometry. It was observed clearly that macrophage, cells incubated with fl-PLP, showed bright fluorescence under the confocal microscope, whereas the control group exhibited no fluorescence, suggesting the positive binding of fl-PLP to macrophage cells. The FLA moiety-dependent binding of fl-PLP was excluded by the faint fluorescence found in cells treated with FLA at an equimolar dose (Fig. 2).

Fig. 2 Fluoresceinamine-labeled PLP binds to macrophage cells positively. Macrophage cells were incubated with media alone, fl-PLP, or FLA at the indicated concentrations for 1 h at 4 °C and then spotted on a slide and subjected to confocal laser scanning. Experiments were performed in triplicate. The representative cells with fluorescence and normal light observation and a merged image for each group are presented.

Next, both saturation and competition assay were performed to study the binding characteristics of fl-PLP in macrophage cells. In a saturation assay, macrophage cells were stained with fl-PLP at serial concentrations for 1 h, and then the binding ability of fl-PLP was determined using flow cytometry. As shown in Fig. 3A, the mean fluorescent intensity of macrophage cells was gradually elevated with the incremental increase in fl-PLP concentration ranging from 25 to 400 nM and reached the saturation state at higher doses. The K_d and B_max values for PLP binding to macrophage cells were ∼490 nM and 248, respectively. In a competition assay, macrophage cells were incubated with fl-PLP and excess unlabeled PLP together for 1 h and then subjected to flow cytometric analysis. As expected, additional unlabeled PLP from 200 to 3200 nM (from 2- to 16-fold of fl-PLP) resulted in a dose-dependent decrease in the mean fluorescent intensity of macrophage cells, suggesting that fl-PLP binding to macrophage cells can be competitively inhibited by unlabeled PLP. Curve-fitting analysis by 3-parameter logistic model showed that PLP had an IC₅₀ of ∼1200 nM (Fig. 3B). All these observations indicated that PLP binding to macrophage cells was a direct, dose-dependent, saturable and reversible process and implied that the stimulation of macrophage cells by PLP may be a receptor-mediated event.

Fig. 3 PLP binds to macrophage cells in a saturable and reversible manner. (A) Saturable binding kinetics of PLP in macrophage cells. Macrophage cells were incubated with fl-PLP at the indicated concentrations for 1 h at 4 °C. The mean fluorescence intensity of each group was examined by flow cytometry. The combined results are presented in panel A. (B) Competitive inhibition of fl-PLP binding to macrophage cells by unlabeled PLP. Cells were incubated with 100 nM of fl-PLP for 1 h in the absence or presence of unlabeled PLP with concentrations from 200 to 3200 nM and then harvested for flow cytometric analysis. The combined results expressed as a percentage mean fluorescence intensity of fl-PLP with unlabeled PLP are presented in panel B.

3.4. TLR2 and TLR4 are required for PLP activities in macrophage cells

TLRs have been demonstrated to play a direct role in the control of macrophage cells responses, including proliferation, up-regulation of activation markers, cytokine secretion, terminal differentiation and antibody production.^28–30 Based on these findings and taking into consideration that TLR2 and TLR4 can recognize carbohydrate-containing molecules, we focused our study on the functional relevance of TLR2 and TLR4 in PLP-mediated macrophage cell responses.

The requirement of TLR2 and TLR4 for PLP function was first investigated in macrophage cells following receptor blocking with specific antibodies. Macrophage cells were stimulated with PLP in the presence of antibodies to TLR2 or TLR4 and then subjected to proliferation assay. Antibodies to TLR2 and TLR4 significantly reduced the proliferative effect of PLP by 49.45% and 39.4%, respectively, when compared with Ab-free control (Fig. 4A). The reduction was found to be synergetic upon the combined treatment with anti-TLR2 and anti-TLR4. In contrast to the substantial suppression observed in anti-TLR2 and/or anti-TLR4 treatment group, proliferation was significantly altered in PLP-stimulated macrophage cells in the presence of the isotype control (Fig. 4A). Therefore, we propose that TLR2 and TLR4 might be functionally correlated to the stimulation of macrophage cells by PLP. To further confirm this statement, affinity adsorption precipitation and western blot analysis were used to separate TLR2 and TLR4. These findings, together with those from antibody blocking assay and WB assay results (Fig. 4B), provided convincing evidence that both TLR2 and TLR4 were responsible for the immunostimulating property of PLP in macrophage cells.

Fig. 4 TLR2 and TLR4 are required for PLP activities in macrophage cells. (A) Antibodies to TLR2 and TLR4 attenuate PLP functions in macrophage cells. Macrophage cells were pretreated with anti-TLR2, anti-TLR4 or respective isotype controls (20 μg ml⁻¹) for 2 h before the addition of PLP. After 48 h incubation, cell proliferation was measured using MTT assay (*p ≤ 0.05, **p ≤ 0.01). (B) Precipitation of TLR2 and TLR4 from macrophage cell membrane extracted by PLP-coupled Sepharose 4B. Membrane extracts from macrophage cells were incubated with PLP-Sepharose 4B and EAH-Sepharose 4B for 2 h at 4 °C. The bound proteins were harvested by precipitation and then subjected to western blot analysis probing with anti-TLR2 and anti-TLR4 antibodies.

3.5. MAPK signaling is greatly involved in PLP-induced macrophage cell response

To value the consequential downstream events upon PLP binding to TLR2 and TLR4 in macrophage cells, the responses of three MAPK proteins, including p38, ERK and JNK, which have been well-characterized as the signal transducers for TLR2 and TLR4, were investigated.²² First, the macrophage cells were treated with different concentrations of PLP (20, 100, 200 nM) for 30 min. Then, the cellular extracts from each group were subjected to the detection of MAPK phosphorylation by western blot analysis. As shown in Fig. 5A and B, PLP increased the phosphorylation of p38 (at Tyr182), ERK (at Tyr204) and JNK (at Thr183 and Tyr185) in a dose-dependent manner, without affecting the total expression of p38, ERK or JNK significantly. These findings suggested that PLP can trigger all the three MAPK signaling pathways in macrophage cells.

Fig. 5 PLP-induced macrophage cell activation involves MAPK signaling. (A and B) PLP-mediated MAPK activation in macrophage cells. Macrophage cells were treated with PLP at the indicated concentrations for 30 min, and then cellular extracts obtained from each group were subjected to western blot analysis probing with anti-p38, anti-phospho-p38, anti-ERK, anti-phospho-ERK, anti-JNK, anti-phospho-JNK, or β-actin antibodies. Images shown in panel A are representative of triplicates, and the band intensities are presented in panel B. (C) Abrogation of PLP-mediated macrophage cell proliferation by specific MAPK inhibitors. Cells were stimulated with PLP for 48 h along with a pretreatment with either SB203580, U0126 or SP600125 for 30 min. Cell proliferation were measured using MTT assay (n = 6, **p ≤ 0.01 vs. inhibitor-free by Mann–Whitney U test).

To further elucidate the relevance of the MAPK pathways to PLP activities in macrophage cells, specific inhibitors of p38 (SB203580), ERK (U0126) and JNK (SP600125) were used to examine if they could interfere with PLP-mediated macrophage cell responses. We found that 20 μM of p38 inhibitor SB203580 decreased PLP-dependent cell proliferation by 15.95%, when compared with inhibitor-free control, suggesting that p38 MAPK signaling partly played a pivotal role during the macrophage cell activation process (Fig. 5C). A similar effect can also be observed from the experiment performed with the ERK inhibitor U0126, in which additional usage of U0126 at 20 μM resulted in 34.3% suppression in PLP-mediated cell proliferation (Fig. 5C). Unlike SB203580 and U0126, the JNK inhibitor SP600125 at an equimolar dose reduced the proliferative activity of PLP in macrophage cells by 7.1% (Fig. 5C). Combining the data from the signaling inhibition assay and western blot analysis, we found that a broader profile of signaling pathways, including p38, ERK and JNK, was utilized by PLP to promote macrophage cell proliferation.

4. Conclusions

To summarize, we demonstrated the immunomodulating potential of a Plantago asiatic L. seed polysaccharide PLP in macrophage cells. The almost certain mechanism accounting for this involves TLR2- and TLR4-mediated MAPK signaling pathways. Given that immunotherapy has become an important component of cancer treatment during the recent decades, our findings clearly support the role of PLP as an efficacious and safe biological response modifier, which might be applied in the future clinical practice and antitumor therapy.

Acknowledgements

We thank the Funding Scheme for Training Young Teachers in Shanghai Colleges (Grant No. yyy11020), the Shanghai Committee of Science and Technology (No. 13430503400), the Science Foundation of Shanghai Institute of Technology (No. YJ2011-75) and the Scientific Research Foundation for the Returned Overseas Chinese Scholars, State Education Ministry (No. ZX2012-05).

Notes and references

1. I. N. Beara, M. M. Lesjak, D. Z. Orčić, N. Đ. Simin, D. D. Četojević-Simin, B. N. Božin and N. M. Mimica-Dukić, LWT--Food Sci. Technol., 2012, 47, 64–70.
2. I. N. Beara, D. Z. Orčić, M. M. Lesjak, N. M. Mimica-Dukić, B. A. Peković and M. R. Popović, J. Pharm. Biomed. Anal., 2010, 52, 701–706.
3. J. H. Cummings, CRC Handbook of Dietary Fiber in Human Nutrition, 2001, vol. 3, pp. 183–252.
4. M. Rodríguez-Morán, F. Guerrero-Romero and G. Lazcano-Burciaga, J. Diabetes Complications, 1998, 12, 273–278.
5. R. P. Zandonadi, R. B. A. Botelho and W. M. C. Araújo, J. Am. Diet. Assoc., 2009, 109, 1781–1784.
6. C. Fang, Nutr. Res., 2000, 20, 695–705.
7. J. Ma and S. Clemants, Taxon, 2006, 55, 451–460.
8. A. L. Romero, K. L. West, T. Zern and M. L. Fernandez, J. Nutr., 2002, 132, 1194–1198.
9. L. Gong, H. Zhang, Y. Niu, L. Chen, J. Liu, S. Alaxi, P. Shang, W. Yu and L. Yu, J. Agric. Food Chem., 2015, 63, 569–577.
10. J.-L. Hu, S.-P. Nie, C. Li, Z.-H. Fu and M.-Y. Xie, J. Agric. Food Chem., 2013, 61, 6092–6101.
11. J.-L. Hu, S.-P. Nie, F.-F. Min and M.-Y. Xie, J. Agric. Food Chem., 2012, 60, 11525–11532.
12. J. Y. Yin, S. P. Nie, C. Zhou, Y. Wan and M. Y. Xie, J. Sci. Food Agric., 2010, 90, 210–217.
13. G.-Y. Kim, G.-S. Choi, S.-H. Lee and Y.-M. Park, J. Ethnopharmacol., 2004, 95, 69–76.
14. Y. Niu, Z. Xie, H. Zhang, Y. Sheng and L. Yu, J. Agric. Food Chem., 2013, 61, 596–601.
15. A. Prieto, M. Bernabé and J. Leal, Mycol. Res., 1995, 99, 69–75.
16. A. Roeder, C. J. Kirschning, R. A. Rupec, M. Schaller and H. C. Korting, Trends Microbiol., 2004, 12, 44–49.
17. O. Takeuchi and S. Akira, Cell, 2010, 140, 805–820.
18. K. Hoshino, O. Takeuchi, T. Kawai, H. Sanjo, T. Ogawa, Y. Takeda, K. Takeda and S. Akira, J. Immunol., 1999, 162, 3749–3752.
19. S. Han, Y. Yoon, H. Ahn, H. Lee, C. Lee, W. Yoon, S. Park and H. Kim, Int. Immunopharmacol., 2003, 3, 1301–1312.
20. X. Li and W. Xu, J. Ethnopharmacol., 2011, 135, 1–6.
21. K.-I. Lin, Y.-Y. Kao, H.-K. Kuo, W.-B. Yang, A. Chou, H.-H. Lin, L. Y. Alice and C.-H. Wong, J. Biol. Chem., 2006, 281, 24111–24123.
22. X. Li, S. Jiang and R. I. Tapping, Cytokines, 2010, 49, 1–9.
23. S. L. Peng, Curr. Opin. Immunol., 2005, 17, 230–236.
24. C. J. Hackett, J. Allergy Clin. Immunol., 2003, 112, 686–694.
25. H.-t. Li, B.-L. Zhao, J.-W. Hou and W.-J. Xin, Biochem. Biophys. Res. Commun., 1996, 223, 311–314.
26. C. Nathan and Q.-w. Xie, Cell, 1994, 78, 915–918.
27. I. A. Schepetkin, G. Xie, L. N. Kirpotina, R. A. Klein, M. A. Jutila and M. T. Quinn, Int. Immunopharmacol., 2008, 8, 1455–1466.
28. I. Bekeredjian-Ding and G. Jego, Immunology, 2009, 128, 311–323.
29. L. M. Ganley-Leal, X. Liu and L. M. Wetzler, Clin. Immunol., 2006, 120, 272–284.
30. Y. Nagai, T. Kobayashi, Y. Motoi, K. Ishiguro, S. Akashi, S.-i. Saitoh, Y. Kusumoto, T. Kaisho, S. Akira and M. Matsumoto, J. Immunol., 2005, 174, 7043–7049.

---

Footnote

† These authors contributed equally to this work.

---