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. 2023 Jul 18;102(10):102946. doi: 10.1016/j.psj.2023.102946

The release of zearalenone-induced heterophil extracellular traps in chickens is associated with autophagy, glycolysis, PAD enzyme, and P2X1 receptor

Hanpeng Wu 1, Xuhai Li 1, Zhan Zhang 1, Yingrong Ye 1, Yichun Chen 1, Jingjing Wang 1, Zhengtao Yang 1, Ershun Zhou 1,1
PMCID: PMC10428124  PMID: 37542939

Abstract

Zearalenone (ZEA) is produced mainly by fungi belonging to genus Fusarium in foods and feeds. Heterophil extracellular traps (HETs) are a novel defense mechanism of chicken innate immunity involving activated heterophils. However, the conditions and requirements for ZEA-triggered HET release remain unknown. In this study, immunostaining analysis demonstrated that ZEA-triggered extracellular fibers were composed of histone and elastase assembled on DNA skeleton, showing that ZEA can induce the formation of HETs. Further experiments indicated that ZEA-induced HET release was concentration-dependent (ranging from 20 to 80 μM ZEA) and time-dependent (ranging from 30 to 180 min). Moreover, in 80 μM ZEA-exposed chicken heterophils, reactive oxygen species (ROS) level, catalase (CAT), superoxide dismutase (SOD) activity, malondialdehyde (MDA) content, and glutathione (GSH) content were increased. Simultaneously, ZEA at 80 μM activated ERK and p38 MAPK signaling pathways by increasing the phosphorylation level of ERK and p38 proteins. Pharmacological inhibition assays revealed that blocking nicotinamide adenine dinucleotide phosphate (NADPH) oxidase, ERK, and p38 mitogen-activated protein kinase (MAPK) reduced ZEA-induced ROS levels but had no impact on HET formation. Furthermore, immunostaining analysis indicated that the heterophil underwent the formation of autophagosome based on being stained with LC3B. The pharmacological inhibition assays demonstrated that rapamycin-, wortmannin-, and 3-methyladenine (3-MA)-treatments modulated ZEA-triggered HET formation, indicating that heterophil autophagy played a key role in ZEA-induced HET formation. Further studies on energy metabolism showed that inhibition of lactate/glucose transport, hexokinase-2 (HK-2), fructose-2,6-biphosphatase 3 (PFKFB3) in glycolysis abated ZEA-induced HETs, implying that glycolysis was one of the factors influencing the ZEA-induced HET formation. Besides, inhibition of the peptidylarginine deiminase (PAD) enzyme and P2X1 significantly reduced the ZEA-induced HET formation. In conclusion, we demonstrated that ZEA-triggered HET formation, which was associated with glycolysis, autophagy, PAD enzyme, and P2X1 receptor activation, providing valuable insight into the negative effect of ZEA on chicken innate immunity.

Key words: zearalenone, heterophil extracellular trap, reactive oxygen species, glycolysis, autophagy

INTRODUCTION

Zearalenone (ZEA), also known as the F-2 mycotoxin, is a nonsteroidal estrogenic mycotoxin produced as a secondary metabolite by a variety of Fusarium. It was initially reported by Rodney W. Caldwell in 1970 (Caldwell et al., 1970). Numerous studies demonstrated ZEA is genotoxic, immunotoxic, and hepatotoxic (Bai et al., 2018). Moreover, impact of ZEA on gonad function in male animals should not be underestimated. For instance, it has been shown that ZEA-triggered oxidative stress in Leydig cells of male mice (Lin et al., 2015), leading to dysregulation of anti-inflammatory activities and antioxidant balance. This impairment of testicular functions in male mice occurred as a consequence (Boeira et al., 2015). As heterophils play an indispensable role in the innate immune defense of the avian host and ZEA is commonly present in poultry feed. Therefore, the heterophil reaction to ZEA is of interest to poultry health (Miraglia et al., 2009). However, there is limited information about the effects of ZEA on chicken heterophils.

Heterophil extracellular traps were described as a novel effector mechanism of heterophil in 2009. These traps contain DNA, histone H3, and elastase from heterophils cytoplasmic granules and are similar in structure to the neutrophil extracellular traps (NETs) found in mammalian and fish (Chuammitri et al., 2009). Prior research suggested that ROS generation is a crucial factor in ZEA-induced HET formation (Brinkmann et al., 2004). However, the precise pathways that connect ROS to HETs in chickens heterophils, specifically regarding the involvement of ROS regulatory factors like MAPK signaling pathways, are still not fully understood (Cui et al., 2023c; Keshari et al., 2013).

ZEA-induced oxidative stress can trigger cellular autophagy (Liu et al., 2023; Zheng et al., 2019), and some studies have shown that autophagy protects chicken granulosa cells from ZEA via PI3K/AKT/mTOR pathway (Zhu et al., 2021). Although scientists have characterized the molecular process of autophagy, the details of how heterophil autophagy is initiated and controlled during ZEA stimulation remain largely unknown.

Additionally, cellular metabolism and neutrophil autophagy are interconnected (Cui, et al., 2023a; Zhao et al., 2023). However, there is limited knowledge about heterophil metabolism during mycotoxin-induced HETosis. Owing to the relatively small quantity of mitochondria in neutrophils, one controversial issue is whether the energy mainly originates from glycolysis or mitochondrial oxidative phosphorylation (Maianski et al., 2004). Until recently, Bao et al. (2014) have found that mitochondria produce the ATP that initiates purinergic signaling, which leads to the activation of neutrophils. Then glycolysis provides the ATP that sustains neutrophils activation, resulting NETosis being divided into 2 metabolic phases, one independent of exogenous glucose and the other relying solely on glycolysis and exogenous glucose (Rodríguez-Espinosa et al., 2015). In other words, while glycolysis primarily provides energy, there may be another energy resource involved in the activation stage, in addition to glucose. Therefore, we used the inhibitors targeting glucose/lactate transport and glycolysis, respectively, and observed the effects on HETs release.

MATERIALS AND METHODS

The Isolation of Chicken Heterophils

Blood donors were healthy adult Welsummer roosters (n = 6, 14–18-wk old, 1–1.5 kg). Peripheral blood was collected from the large vein under the wing (brachial vein) with sterile glass tubes containing EDTAK2. Chicken heterophils were then isolated by the commercially available Chicken Heterophils Isolation Kit (Cat. #LZS1098C, TianJin HaoYang Biological Manufacture Co., Ltd., Tianjin, China). The procedure follows the manufacturer's instructions. Briefly, blood was diluted with RPMI 1640 medium without phenol red at a 1:1 ratio. The diluted blood was then layered on a separating solution in centrifugal tubes and centrifuged at 800 × g for 40 min. The lower cellular layer was collected and washed with erythrocyte lysis solution until the cell pellet became white. After centrifugation at 650 × g for 15 min, the supernatant was removed by pouring off. Finally, the isolated heterophils from the bottom of the centrifugal tubes were resuspended in RPMI 1640 medium and stored at 37 °C with 5% CO2. For heterophils counting, we took trypan blue-treated cell suspension using a pipette and applied it to the glass hemocytometer; the percentage of living cells was over 95%. According to a previous study, the purity of heterophils can reach over 90% (Han et al., 2019). We set up 5 biological replicates and 2 technical replicates for each experiment. Notably, each rooster was collected 7 mL peripheral blood approximately, once every 2 wk. For each experiment, cell samples from multiple chickens were treated independently. The techniques used in the process require gentle handling.

All procedures followed the Guide for the Care and Use of Laboratory Animals published by the National Institutes of Health (National Research Council Committee for the Update of the Guide for the and Use of Laboratory, 2011) and were approved by the Institutional Animal Care and Use Ethics Committee of Foshan University (Permit NO. SCXK 2018-0002).

Heterophil Viability Assays

Analysis of heterophil viability was performed according to Wu et al. (2023). Heterophils were inoculated (2 × 105/mL) in a 96-well plate and stimulated with different concentrations of ZEA (20, 40, 80, 160 μM; Cat. #Z2125, Sigma-Aldrich, Darmstadt, Germany) for 2 h. One hour later, Then Cell Counting Kit reagent containing water-soluble tetrazolium, monosodium salt (WST-8) was added (10 μL per well) according to manufacturer's instructions (Cat. #GK10001, GLPBIO, California) and the plate was incubated at 37 °C for 1 h. Lastly, the absorbance was measured at 450 nm by the iMark microplate absorbance reader (Serial No. 18855, Bio-Rad Laboratories, Inc., California).

Lactate Dehydrogenase Detection

The damage of the heterophil membrane results in a release of lactate dehydrogenase (LDH) into the surrounding cell culture medium. LDH is a stable cell death marker that can be detected. Heterophils were inoculated into 96-well plates (2 × 105/well), and then stimulated with different concentrations of ZEA (20, 40, 80, 160 μM) for 2 h. Afterward, the heterophils were centrifuged at 400 × g for 5 min, and we collected the 120 μL supernatant, and transferred it to a new 96-well plate. Finally, a commercial LDH cytotoxicity assay kit (Cat. #C0017, Beyotime Biotechnology, Shanghai, China) was used to quantify the extracellular LDH. For a positive control, we used an LDHrelease reagent that lyses cells. Absorbance was detected at 490 nm by the iMark microplate absorbance reader (Serial No. 18855, Bio-Rad Laboratories, Inc., California), and its level is directly proportional to the activity of LDH released into the medium.

Indirect Immunofluorescence Staining and Observation

The heterophils (2 × 105/well) were stimulated with ZEA (80 μM) and zymosan (1 mg/mL) for 2 h at 37 °C on coverslips pretreated with poly-L-Lysine hydrobromide (0.1 mg/mL; Cat. #BS199, Beijing Labgic Technology Co., Ltd., Beijing, China). Then, the samples were fixed with 4% (w/v) paraformaldehyde (Beijing Labgic Technology Co., Ltd., China), washed in PBS 3 times, permeabilized with 0.2% Triton X-100 for 20 min, blocked in 3% goat serum, and soaked in antibody solution (H3 or NE, 1:200 diluted with blocking solution, 4 °C) overnight. The primary antibodies were diluted with 3% goat serum to 1:200. After 3 washes with PBS, the samples were incubated with the secondary antibody (goat anti-rabbit IgG, FITC, 1:200, diluted with blocking solution; Cat. #SA00003-2, Proteintech Group, Inc., Illinois). Finally, the cells were stained with SYTOX Orange Nucleic Acid Stain for 20 min (5 μM, dissolved in PBS; Life Technologies Corporation, Massachusetts). Following incubation, the excess antibody is washed off 3 times with PBS. Then the cells are mounted on coverslips with the antifade mounting medium (Cat. #2100, Beijing Solarbio Science & Technology Co., Ltd., Beijing, China). Visualization was achieved using an inverted fluorescence microscope (Model: RVL-100-G, DISCOVER ECHO Inc., California) equipped with Flat Field Semi-Complex Achromatic objective.

The primary antibody, antihistone polyclonal antibody (Cat. #ab5103) and antineutrophil elastase polyclonal antibody (Cat. #ab68672), were purchase from Abcam in the United Kingdom.

Quantitation of HETs

In the initial set of experiments, we examined the impact of various concentration of ZEA on for HET formation. The heterophils (2 × 105/well) were suspended in 96-well microplates in RPMI 1640 and were stimulated with varying concentrations of ZEA (20, 40, 80, 160 μM) for 2 h at 37 °C in a humidified 5% CO2 incubator. The zymosan (1 mg/mL) group was used as a positive control.

In the subsequent set of experiments, we examined the different stimulation times of ZEA for on HET formation. Heterophils (2 × 105/well) were suspended in 96-well microplates in RPMI 1640 and stimulated with 80 μM ZEA for different lengths of time (30, 60, 120, and 180 min) at 37°C in a humidified 5% CO2 incubator. The zymosan (1 mg/mL) group was used as a positive control.

In the third set of experiments, we examined the effects of specific inhibitors (details provided in the subsequent paragraphs) on HET formation induced by ZEA (80 μM) via HETs quantification. Heterophils (2 × 105/well) were pretreated with inhibitors for 30 min in a 96-well microplate and then treated by RPMI 1640 medium (as control), ZEA (80 μM), and zymosan (1 mg/mL), respectively, at 37 °C for 2 h in a humidified 5% CO2 incubator.

We used previously stated heterophils exposed to zymosan (1 mg/mL, Sigma-Aldrich, Darmstadt, Germany) as a positive control group in a similar manner. For both sampling methods, supernatants were collected and the 1:200 dilution of PicoGreen (Life Technologies Corporation, Massachusetts) in 10 mM Tris-HCl buffered with 1 mM EDTA (Life Technologies Corporation, Massachusetts) was added (50 μL per well). We then detected and quantified extracellular DNA by measuring PicoGreen-derived fluorescence intensities using fluorescence reader (Model: Infiniti M200, Tecan, Zurich, Switzerland) at 485 nm excitation/525 nm emission.

The inhibitor of mTOR (Rapamycin, 50 μM), extracellular regulated protein kinases (ERK) signaling (U0126, 50 μM), PAD enzyme (Cl-amidine, 6 μM), P2X1 receptor (NF449, 10 μM), PI3K class I (wortmannin, 50 μM) were obtained from MedChemExpress (New Jersey). The following inhibitors of PI3K class III (3-MA, 10 μM), GLUT1 (STF-31, 1 μM), PFKFB3 enzyme (3PO, 24 mM), MCT1 (AZD3965, 1.6 μM), Rac-GTPase (NSC23766, 1.6 μM), p38 MAPK signaling (SB202190, 10 μM), NADPH oxidase (diphenyleneiodonium, 10 μM) were obtained from Sigma-Aldrich. The concentrations of used inhibitors were based on our prior studies (Wang et al., 2020; Zhou et al., 2020).

Detection of Antioxidant Enzymes Activity in Chicken Heterophils

Heterophils (2 × 107/well) were treated with 80 μM ZEA in 6-well plates with white, clear-bottomed for 2 h at 37 °C with 5% CO2. Cells were then soaked in PBS and lysed using Non-Contact Ultrasonic Processor with a power of 1,000 W and a pulse length of 7 min (Model: FN-C1500T, Shanghai Sonxi Ultrasonic Instrument Co., Ltd., Shanghai, China). Protein concentrations were measured using Pierce BCA Protein Assay Kit (Cat. #23225, Thermo Fisher Scientific, Massachusetts). We detected the activities of antioxidant enzymes, including SOD (Cat. #A001-3-2) and CAT (Cat. #A007-2-1), as well as the content of GSH (Cat. #A006-2-1), in the samples using corresponding assay kits (NanJing Jian-Cheng Bioengineering Institute, Nanjing, China) according to the manufacturer's instructions. The absorbance was measured by the iMark microplate absorbance reader (Serial No. 18855, Bio-Rad Laboratories, Inc., California).

Briefly, the assay method for SOD activity involves the SOD-mediated inhibition of the formation of nitrite from hydroxylammonium in the presence of O2− generators (xanthine/xanthine oxidase). SOD activity was expressed as units/mg of protein and determined by measuring the reduction of in the optical density of the reaction solution at 550 nm. CAT activity was calculated as H2O2 consumed of tissue protein. The remaining H2O2 reacted with ammonium molybdate to form a yellowish complex that was measurable at 405 nm. CAT's activity was expressed as units/mg of protein. GSH content was determined by measuring the oxidation of GSH using the sulfhydryl reagent 5,5'-dithio-bis (2-nitrobenzoic acid), which forms a yellow derivative called 5'-thio-2-nitrobenzoic acid. The measurement was performed at a wavelength of 405 nm. GSH content was expressed in μmol/g of protein.

Assessment of Lipid Peroxidation

Lipid peroxidation was assessed by quantifying the levels of MDA contents through thiobarbituric acid method using a commercial MDA kit and measuring at 532 nm (Cat. #A003-1, NanJing Jian-Cheng Bioengineering Institute, Nanjing, China). The MDA content was expressed as nmol/mg of protein.

Western Blot Analysis

The Western blot analysis was performed following established protocols as described in previous studies (Cui et al., 2023b, Shi et al., 2023; Miao et al., 2022). Heterophils (2 × 105/well) were stimulated with 80 μM ZEA for 2 h at 37 °C in a humidified 5% CO2 incubator. The cells were lysed using the protein extraction reagent (Thermo Fisher Scientific, Massachusetts) at 4 °C, and the supernatant was collected after centrifugation at 12,000 × g at 4 °C. The total protein concentration was measured with the bicinchoninic acid (BCA) protein assay kit (Cat. #23225, Thermo Fisher Scientific, Massachusetts), and samples with equal amounts of protein were separated by 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis. The separated proteins were then transferred to a polyvinylidene difluoride membrane (0.45 μm Cat. #IPVH00010, MerckMillipore, Darmstadt, Germany). The membranes were blocked in 3% BSA at room temperature for 2 h and then probed with primary antibodies (1:1,000 dilution in TBST) at 4°C overnight. The next day, the membranes were washed 3 times with TBST for 10 min and incubated with HRP-conjugated secondary antibody (1:20,000 dilution in TBST) at room temperature for 2 h. Finally, the membranes were washed 3 times for 10 min and visualized with a bioanalytical imaging system (Azure Biosystems, Inc., California). Chemiluminescence detection was performed using Immobilon Western chemiluminescent HRP Substrate (MerckMillipore, Darmstadt, Germany).

The following primary antibodies were used: anti-phospho-p38 mAb (Cat. #4511), anti-p38 mAb (Cat. #8690), anti-phospho-ERK mAb (Cat. #9101), and anti-ERK mAb (Cat. #4695) from Cell Signaling Technology (Massachusetts). The protein level was normalized with GAPDH expression using GAPDH polyclonal antibody (1:3,000, Cat. #132004, Absin Bioscience Co. Ltd., Shanghai, China). Protein band intensities was quantified using Image J software 1.48.

Measurement of ROS in Chicken Heterophils

We measured ROS level with H2DCFDA (Cat. # HY-D0940, MedChemExpress, New Jersey) in heterophils stimulated with ZEA. To do this, we suspended heterophils (2 × 105/well) in 96-well microplates in RPMI 1640 without phenol red and stimulated them with ZEA (80 μM) for 2 h at 37°C with 5% CO2. In parallel groups, the heterophils were pretreated with specific inhibitors (DPI, SB202190, and U0126) for 30 min before stimulation with ZEA for 2 h. In the final 30 min of the process, DCF-DA (10 μM per well) was added. After 2 h, we measured the fluorescence values of the samples with the fluorometer reader (Model: Infiniti M200, Tecan, Zurich, Switzerland) at 485 nm excitation/525 nm emission. The zymosan (1 mg/mL, Sigma-Aldrich, Darmstadt, Germany) group was used as a positive control.

Autophagosome Detection by Immunofluorescence Analysis

To autophagosome detection, fixed cells were washed in PBS 3 times, permeabilized with ice-cold methanol treatment for 5 min at 4 °C, and blocked with blocking buffer (5% BSA, 0.1% Triton X-100 in sterile PBS) for 50 min at room temperature. Next, cells were incubated at 4 °C overnight in an anti-LC3B antibody solution (diluted 1:100 in a blocking buffer, Cat. #2775, Cell Signaling Technology, Massachusetts). After incubation, samples were washed 3 times with PBS and incubated for 30 min in the dark and room temperature in the secondary antibody (goat anti-rabbit IgG, FITC, 1:100, diluted with blocking solution; Cat. #SA00003-2, Proteintech Group, Inc., Illinois). Samples were washed three times with PBS and then incubated with a 1:1000 dilution of Sytox Orange (Invitrogen, Massachusetts) in sterile PBS for 15 min before mounting. Images were taken using the fluorescent microscope (Model: RVL-100-G, DISCOVER ECHO Inc., California) equipped with Flat Field Semi-Complex Achromatic objective. Image processing was carried out with Fiji ImageJ using merged channel plugins and limited to basic adjustment of brightness and contrast.

Statistical Analysis

All data were expressed as mean ± SEM of at least 3 biological replicates and 2 technical replicates. Graphs and statistical analysis were performed by GraphPad Prism 9. Shapiro-Wilk test was used to determine the normality of samples. Difference between 2 groups was compared by Student 2-tailed t test. When more than one group was compared with the control, statistical analysis was conducted using 1-way ANOVA followed by a Dunnett test in the present study. Statistical significance was defined as a P value < 0.05.

RESULTS

ZEA Did Not Affect the Activity of Heterophils or Induce Heterophil Necrosis

As shown in Figure 1A, the cell viability assays showed that ZEA (20–160 μM) did not significantly affect the viability of heterophils compared to the control group (P > 0.05). To further investigate whether ZEA-induced HETs release involved cell necrosis, we measured extracellular LDH (as shown in Figure 1B). The results showed that absorbance in the ZEA groups (20–160 μM) had no significant difference compared to the control group (P > 0.05), while the lysis group was significantly higher (P < 0.0001), these results indicated that ZEA (20–160 μM) did not induce heterophil necrosis.

Figure 1.

Figure 1

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The effect of ZEA (20–160 μM) on heterophil viability. (A) ZEA did not affect the viability of the heterophils. Heterophils were stimulated for 2 h with ZEA (20, 40, 80, and 160 μM, respectively). Samples were added cell counting reagent and incubated for 1 h, then its absorbance was detected at the wavelength of 450 nm. (B) The level of LDH was analyzed with a commercial LDH cytotoxicity assay kit. The lysed cell group was used as the positive control. Finally, its absorbance was detected at the wavelength of 490 nm. One-way ANOVA was used to determine the significance (n = 6, ****P < 0.0001, “ns” means no significant difference).

ZEA-Triggered HET Formation in Chicken Heterophils with Concentration, and Time-dependent Manners

As shown in Figure 2A, we were able to observe chicken heterophils forming structures in response to ZEA, which were colocalized by DNA, citrullinated histone 3, and elastase staining, indicating that ZEA could trigger HET formation.

Figure 2.

Figure 2

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The influence of ZEA concentration and stimulation time on HET formation induced by ZEA. (A) ZEA-induced HETs formation. Fixed chicken heterophils immunolabeled with antibodies directed against citH3 (FITC channel) and NE (FITC channel) for HETs, and incubated with Nucleic Acid Stain (TxRed channel). Samples were observed, and images (200×) were taken under the inverted fluorescence microscope. White arrows indicated HETs structures in merged photos. Scale bar: 30 µm. (B) ZEA-triggered HET formation was concentration-dependent. Heterophils (2 × 105/well) were incubated with 4 different concentrations of ZEA (20, 40, 80, 160 μM, respectively) for 2 h. One-way ANOVA was used to determine the significance. (C) The time-kinetics of ZEA-triggered HETs. Heterophils were treated with 80 μM ZEA for 30 min, 60 min, 120 min, 180 min, respectively. A Student t test was used to determine the significance. Fluorescence intensity was detected by the fluorescence plate reader at 485 nm excitation/525 nm emission. n = 6, **P < 0.01, ****P < 0.0001, “ns” means no significant difference.

Based on our previous study on the neutrophil in bovine (Wang et al., 2019), we established a range of concentrations of ZEA (20, 40, 80, 160 μM) to assess any potential differences in HET formation. Our results showed a significant increase in HET formation in a concentration-dependent manner, as indicated by the rising trend observed with increasing concentrations of ZEA (mean values of extracellular dsDNA 1.27, 2.10, 3.19, and 3.16 times higher than control, respectively) (see Figure 2B). These findings suggested that ZEA-induced HET formation in a concentration-dependent manner, particularly at concentrations ranging from 20 μM to 80 μM.

To examine the time-dependent effect of ZEA on HET formation, we treated heterophils with 80 μM ZEA for different time intervals ranging from 30 to 180 min. As shown in Figure 2C, we observed a significant increase in HET release by ZEA-treated heterophils at 120 min and 180 min as compared to the control group (P < 0.01). The maximum release of HETs induced by ZEA was observed at the 180-minute time point. However, when comparing it to the 120-minute time point, the P value between ZEA (80 μM) and the control group was 0.0004, which is smaller than the significance threshold of 0.0083 (for the 180-minute time point). Therefore, we conclude that the 2-hour time point was selected to stimulate heterophils.

NADPH Oxidase and MAPK Signaling Pathway Mediated ZEA-induced Oxidative Stress But Had No Effect on ZEA-induced HET Formation

In Figure 3A to D, it can be observed that following ZEA treatment, there was an increase in the activity of SOD and CAT, as well as the content of GSH, along with an increase in MDA content. Specifically, CAT activity and GSH content in heterophils were significantly elevated (P < 0.0001), while SOD activity was also increased (P < 0.01), and MDA content was significantly higher (P < 0.0001) compared to the control group. These results suggested ZEA induced oxidative stress in heterophils, leading to an imbalance between the production of free radicals and the antioxidant defenses.

Figure 3.

Figure 3

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The effect of NADPH oxidase and MAPK signaling pathway in the ZEA-induced oxidative stress/HETs formation. (A) SOD activity was measured. (B) CAT activity was measured. (C) GSH and (D) MDA contents were measured. Heterophils (2 × 107/well) were stimulated by ZEA (80 μM) for 2 h, then we detected the activity of antioxidant enzymes and MDA content via corresponding detection kits. (E) The phosphorylation of ERK and p38 proteins quantified by densitometric analyses after normalization with the GADPH signal. Heterophils (2 × 107/well) were stimulated by ZEA (80 μM) for 2 h, then we detected the expression level of ERK and p38 proteins in samples via Western blotting. A Student t test was used to determine the significance (n = 6, **P < 0.01, ***P < 0.001, ****P < 0.0001). (F) The NADPH oxidase and MAPK signaling pathway-mediated ZEA-induced ROS production in chicken heterophils. Primary chicken heterophils (2 × 105/well) were pretreated with the inhibitor of NADPH oxidase, ERK, p38 for 30 min, then exposed to ZEA (80 μM) for 2 h. DCF-DA (10 μM) was added to each well, and the plate was incubated for the last 30 min. (G) The NADPH oxidase, p38, ERK pathway did not influence ZEA-induced HETs release. Primary chicken heterophils (2 × 105/well) were suspended in 96-well microplates in RPMI 1640 medium. We pretreated them with inhibitors of NADPH oxidase (DPI), ERK signaling (U0126), and the p38 MAPK signaling (SB202190) for 30 min, followed by exposure to ZEA (80 μM) for 2 h. Fluorescence intensities were detected by the fluorescence plate reader at 485 nm excitation/525 nm emission. One-way ANOVA was used to determine the significance (n = 6, ****P < 0.0001, “ns” means no significant difference).

After detecting ZEA-induced cellular oxidative stress, further investigation into MAPKs is warranted, as they are well-known inflammatory signaling cascades. ZEA considerably enhanced phosphorylated ERK and p38 proteins in heterophils, as shown in Figure 3E (P < 0.01).

Then we measured the production of intracellular ROS induced by ZEA and used corresponding inhibitors to determine the source of ROS in heterophils. As shown in Figure 3F, pretreatment with DPI (an inhibitor of NADPH oxidase) significantly decreased ZEA-induced ROS (P < 0.001). Furthermore, the inhibitors of MAPK signaling, SB202190 and U0126, had the same effect (Figure 3F, P < 0.001). These results suggested that ZEA-induced ROS production in heterophils originates from the NADPH oxidase and MAPK signaling pathway.

To further investigate the link between HET formation and NADPH oxidase, as well as the potential role of the MAPK family in HET release, we used specific inhibitors to examine the effects of NADPH oxidase, p38, and ERK signaling on ZEA-induced HET formation. As shown in Figure 3G, pretreatment with SB2012190, U0126, and DPI did not decrease HET release induced by ZEA (P > 0.05). This suggested that ZEA-triggered HET formation was independent on p38, ERK signaling pathways, and NADPH oxidase in chicken heterophils.

ZEA-Triggered Autophagosomes Formation, Which Was Associated With the HET Formation in Chicken Heterophils

We also investigated how ZEA exposure affected the formation of heterophil-derived autophagosome. To do so, we used an anti-LC3B antibody to study the effect of ZEA exposure to heterophil-derived autophagy and effect of autophagy on ZEA-induced HET formation. Figure 4A showed that ZEA exposure resulted in significant autophagosome formation in treated heterophils, compared to that of control group. Interestingly, LC3B-positive and NETotic heterophil against ZEA were captured at the same location in the merged picture, demonstrating a positive correlation between HET formation and autophagy in ZEA-exposed chicken heterophils.

Figure 4.

Figure 4

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The effect of autophagy in ZEA-induced HETs formation. (A) ZEA-triggered HET formation company with the formation of LC3B-coated autophagosomes. Fixed chicken heterophils immunolabeled with antibodies directed against LC3B (FITC channel), and incubated with Nucleic Acid Stain (TxRed channel). Samples were observed, and images (200×) were taken under the fluorescent microscope. The distribution of green fluorescence indicated the formation of autophagosome. In merged photos, the shape of the autophagosome and HETs in the same location (white square frame). Scale bars indicate 30 μm. (B) Autophagy-mediated ZEA-triggered HET formation. We pretreated primary heterophils (2 × 105/well) with the inhibitor of mTOR (rapamycin), PI3K class III (3-methyladenine, wortmannin) for 30 min, followed by exposure to 80 μM ZEA for 2 h. Extracellular DNA was quantified by PicoGreen-derived fluorescence intensities using the fluorescence plate reader at 485 nm excitation/525 nm emission. Statistical analysis by 1-way ANOVA (n = 6, **P < 0.01, ****P < 0.0001).

The pharmacological experiment was performed to prove the relationship between autophagy and the HET formation from different angles under the ZEA stimulation. As shown in Figure 4B, we obtained increased HET formation after rapamycin exposure (P < 0.01) and decreased HET formation after 3-MA/wortmannin treatment (P < 0.0001), indicating that autophagy played an important role in ZEA-triggered HET formation in chicken heterophil.

ZEA-triggered HET Formation Depended on Glycolysis and Glucose Transporter in Chicken Heterophils

It can be seen from the above results that the specific inhibitors failed to control the release of HETs. Thus, we tried to use glycolysis inhibitors pretreated heterophil and found that AZD3965 (an inhibitor of MCT1), bromopyruvic acid (an inhibitor of HK-2), and 3PO (an inhibitor of PFKFB3) were able to significantly reduce the formation of HETs triggered by ZEA (Figure 5A, P < 0.0001), indicating that glycolysis was crucial in ZEA-triggered HET formation in chicken heterophils.

Figure 5.

Figure 5

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The effect of glycolysis in ZEA-triggered HET formation. MCT1, HK-2, and PFKFB3 enzyme-mediated ZEA-triggered HET formation (A). Rac GTPase-, GLUT1-, and GLUT4-mediated ZEA-triggered HET formation (B). Primary chicken heterophils were pretreated (2 × 105/well) with the inhibitor of MCT1 (AZD3965), HK-2 (bromopyruvic acid), PFKFB3 (3PO), Rac GTPase 1 (NSC23766), GLUT1 (STF-31), and GLUT4 (Ritonavir) for 30 min, followed by exposure to ZEA (80 μM) for 2 h. Extracellular DNA was quantified by PicoGreen-derived fluorescence intensities using the fluorescence plate reader at 485 nm excitation/525 nm emission. One-way ANOVA was used to determine the significance (n = 6, ****P < 0.0001).

Additionally, we found a sharp decline in HET formation triggered by ZEA with the use of inhibitors targeting GLUT4 (Ritonavir), GLUT1 (STF-31), and Rac GTPase (NSC23766) compared to the control group (see Figure 5B, P < 0.0001), indicating that glucose uptake also took charge in ZEA-triggered HET formation in chicken heterophils.

ZEA-Triggered HET Formation Depended on P2X1 Receptor and PAD Enzyme in Chicken Heterophils

Finally, we also found that Cl-amidine (an inhibitor of PAD enzyme), NF449 (an inhibitor of P2X1 receptor), could reduce the formation of HETs triggered by ZEA (Figure 6, P < 0.0001), indicating that PAD enzyme and P2X1 receptor also took charge in ZEA-triggered HET formation in chicken heterophils.

Figure 6.

Figure 6

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The effect of PAD enzyme and P2X1 receptor in ZEA-triggered HET formation. Heterophils (2 × 105/well) were pretreated with the inhibitor of PAD (Cl-amidine), P2X1 (NF449) for 30 min, followed by exposure to ZEA (80 μM) for 2 h. Extracellular DNA was quantified by PicoGreen-derived fluorescence intensities using the fluorescence plate reader at 485 nm excitation/525 nm emission. One-way ANOVA was used to determine the significance (n = 6, ****P < 0.0001).

DISCUSSION

ZEA is known to have a long biological half-life, and long-term exposure to it can cause damage to organisms (Kuiper-Goodman et al., 1987; Rai et al., 2019). However, only a few papers have investigated the effects of ZEA in male immune system. Our results indicated that ZEA can trigger HET formation, and this process is associated with ROS production originating from NADPH oxidase and regulating by MAPK signaling. Surprisingly, we found that inhibiting the PAD enzyme and P2X1, rather than the MAPK signaling pathway or NADPH oxidase, was critical for preventing ZEA-induced HET formation. Moreover, we discovered that autophagy in heterophil was associated with ZEA-induced HET formation. Finally, our findings revealed that MCT1, GLUT1, and GLUT4 transport energy substances to heterophil, and then heterophil glycolysis fueled ZEA-induced HET release.

To begin with, we evaluated the impact of ZEA on chicken heterophils' viability and LDH release. Our findings revealed that ZEA did not affect heterophil viability at doses ranging from 20 to 160 μM, These results are consistent with a previous study byWang et al. (2019). Subsequently, we observed that ZEA caused heterophils to release HETs, which were extracellular networks consisting of chromatin and histones along with heterophils elastase. Furthermore, we discovered that as the concentration of ZEA (ranging from 20 to 80 μM) increased, the extracellular DNA also increased gradually. This implies that ZEA-induced HET formation is concentration-dependent until the concentration of ZEA exceeds 80 μM. Interestingly, Jundi Liu's review suggested that poultry may be relatively resilient to ZEA, potentially due to their intrinsically high concentration of estrogen, as demonstrated in a study conducted on male turkeys (Dänicke et al., 2007; Wang et al., 2019; Liu and Applegate, 2020). Given that exposure to 80 μM of ZEA for 2 h was trigger condition for HET release, this might be the best condition for extended research.

Currently, there are 2 main pathways through which HETs can be released by activated heterophils: classical suicidal NETosis, which is dependent on ROS and NADPH oxidase, and vital NETosis, which is ROS and NADPH oxidase independent (Jorch and Kubes, 2017). However, it is still unknown which mechanism is responsible for ZEA-induced HET formation. Our results showed that DPI, an inhibitor of NADPH oxidase, significantly inhibited ZEA-induced ROS generation but did not affect HET release, suggesting that ZEA-induced HET release was the NADPH oxidase-independent ROS. Moreover, ZEA increased the content of MDA in heterophils, indicating that heterophils suffer from lipid accumulation (Ayala et al., 2014). Our results also showed that ZEA significantly increased the production or activation of antioxidant enzymes, including SOD, CAT, and GSH. These enzymes play a crucial role in neutralizing excessive free radicals and protecting cells from oxidative damage. Additionally, antioxidants like GSH contribute to detoxification processes within cells by binding and eliminating toxic substance, potentially mitigating the harmful effects of ZEA (Shang et al., 2023). Additionally, ERK and p38 MAPKs are the core units of the MAPK cascade and are involved in the protection from ZEA-induced cellular ROS (Awasthi et al., 2019; Zhu et al., 2021), and some studies have shown that they played a different role in different contexts (Wei et al., 2018; Wang et al., 2019). Our study found that ZEA significantly increased phosphorylation of ERK and p38, but pharmacological inhibition results showed that blocking ERK, p38 MAPK signaling pathways did not play a significant role in releasing HETs even ZEA-induced ROS generation was decreased by corresponding inhibitors. This suggests that ZEA-induced HETs were ERK, p38 MAPK signaling pathways-independent process, which is similar to the studies on ethylmercury, and Hg2+- or sodium arsenic-induced NETs (Haase et al., 2016; Wei et al., 2018). Some studies mentioned that the endogenous ROS may have another alternative source that is far from essential for NETs formation (Douda et al., 2015; Tatsiy and McDonald, 2018).

HETosis is not cure-all, so finding ways to pharmacologic modulate ZEA-induced HET formation is of great interest. This study used immunostaining analysis to show that autophagosomes coated with LC3B were formed during HET release, suggesting that heterophil autophagy is involved. Therefore, identifying pharmacologic molecules controls autophagy will shed light on HET formation. The master regulator of cellular metabolism and autophagy, mechanistic target of rapamycin complex 1 (mTORC1), is an appealing pharmacologic target for manipulating autophagy, with rapamycin being effective in this regard (Kim and Guan, 2015). In this study, rapamycin, an autophagy activator, increased the amount of HET formation by regulating mTORC1 protein. Immune cells express 3 classes of phosphoinositide 3-kinase (PI3K) and PI3K Class III plays a key role in the initiation of autophagy by generating a lipid signal that recruits downstream autophagy-related proteins to the site of autophagosome formation (Okkenhaug, 2013; Kocaturk et al., 2019). In this study, both wortmannin and 3-MA were used to block autophagosomes formation and significantly reduced the ZEA-induced HET formation. These data suggested that mTOR-dependent autophagy is involved in ZEA-induced HET formation.

A recent study showed that lactate, rather than glucose, can be a primary energy source for the tricarboxylic acid cycle, and a widely expressed transporter allows for this (Hui et al., 2017). Monocarboxylate transporters 1 (MCT1) is crucial for neutrophils and enables them to quickly exchange lactate with other cells (Poole and Halestrap, 1993; Alarcón et al., 2017). In this study, we subjected heterophils to glutamine deprivation and subsequently stimulated them with ZEA. To assess the role of MCT1, we employed AZD3965, a specific inhibitor (Bola et al., 2014), and observed a significant impact on ZEA-induced HET release. Moreover, mitochondrial ATP is critical for triggering neutrophil activation in the first stage (Bao et al., 2014). The ZEA stimulation may cause rapid bursts of ATP release, suggesting that this process requires mitochondria to consume a large amount of lactate to produce ATP in the early stage of heterophil activation. However, MCT1 could not uptake lactate at that time, indicating the importance of MCT1-mediated lactate transport for ZEA-induced HET release.

In addition to the glucose uptake and circulating lactate, there are 2 key steps regulating glucose metabolism in the early stage: HK-2 and phosphofructokinase (PFK). The uptake of glucose is enhanced by the increased recruitment of GLUTs to the cell membrane, and there is a coordinated increase in expression and activity of HK-2 and PFK (Rabbani and Thornalley, 2019). In our study, we used an inhibitor of HK-2, called bromopyruvic acid and found that it significantly reduced ZEA-induced HET formation.

Another important step in glycolysis is the phosphorylation of fructose-6-phosphate to fructose-1,6-diphosphate, which is catalyzed by PFK and is controlled by an enzyme called PFKFB3 (Schoors et al., 2014). We reported that 3-(3-pyridinyl)-1-(4-pyridinyl)-2-propen-1-one (3PO) indirectly reduced glycolysis ATP production, which in turn kept HET release at bay. Unlike 2-deoxy-glucose, a glycolysis inhibitor, PFKFB3 blockade inhibits a defined downstream pathway, reducing glycolytic flux without suppressing the pentose-phosphate pathway that is necessary for NADPH oxidase activity. This makes it a promising target for the treatment of HET-related diseases, as reported by Schoors et al. (2014).

In addition to lactate transport, some members of the glucose transporter (GLUT) family, such as GLUT1 and GLUT4, have been shown to transport glucose under experimental conditions (Mueckler and Thorens, 2013). However, it is still unclear whether these transporters are involved in ZEA-induced HET formation. Inhibition of glucose transport may be a potential pharmacological approach for controlling HET formation. Regarding glucose, the inhibitors of glucose transporter (STF-31 (Chan et al., 2011), ritonavir (McBrayer et al., 2012), NSC23766 (Sylow et al., 2015)) downregulated ZEA-induced HET formation significantly, which in line with our previous study (Wu et al., 2023). In conclusion, our findings suggested that both monocarboxylate and glucose transporters play a role in circulating energy substance for HET formation.

The enzyme PAD is found downstream of ROS and is necessary for chromatin decondensation, which eventually becomes framework for HETs. In vital NETosis, chromatin is expelled through vesicles without harming the cell or requires ROS. This feature allows neutrophils to remain viable, which may also occur during autoimmune development (Pilsczek et al., 2010). Some studies indicated that several isoforms, such as PAD2, PAD3, and PAD4, participate in NETosis (Pisanu et al., 2015; Spengler et al., 2015). However, because chickens do not express the PAD4 gene, we used the general PAD inhibitor cl-amidine, which does not discriminate between PAD isoforms, to pretreat the heterophils in this study. We found that cl-amidine significantly reduced the formation of HETs triggered by ZEA, indicating that PAD mediates this process. Unlike inhibiting the NADPH oxidase, one study showed that inhibiting PAD suppressed NETosis without affecting the associated neutrophil oxidative burst, making it a promising target for HETs-related noninfectious diseases (Jorch and Kubes, 2017).

Finally, ATP serve as the energy currency that drives NETosis and plays a critical role as signal transduction molecule by activating purinergic signaling in an autocrine and paracrine manner (Sofoluwe et al., 2019). One of the purinergic receptors that respond to extracellular ATP binding on neutrophils is P2X1 (North, 2002; Alarcon et al., 2020) and its function in HET formation is still unclear. In this study, the suramin analog NF449, a picomolar potency and reversible antagonist of the P2X1 receptor was used (Hülsmann et al., 2003). We found that P2X1 exerted a strong effect in ZEA-triggered HET formation, as treatment with NF449 strongly blocked HET formation, which was consistent with previous findings (Zhou et al., 2020). Considering its low strength at other P2X receptor subtypes-mediated effects in heterophil, NF449 may hence be a pretty helpful inhibitor for pharmacological discrimination compare to NF279 (Klapperstück et al., 2000). Moreover, the glycolytic ATP is necessary for P2X1 to maintain the cell response in the second phase, as described by Bao et al. (2014). Neutrophil ATP is released through the pannexin-1 channel into the extracellular space, where it combines with extracellular ATP to activate P2X1 receptors (Bao et al., 2014). When the P2X1 receptor is closed, most ATP molecules are expelled, and NETosis does not occur (Junger, 2011).

ACKNOWLEDGMENTS

This work was supported by the National Natural Science Foundation of China (no. 31772721).

DISCLOSURES

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in the present study.

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