Relationship between Eimeria tenella development and host cell apoptosis in chickens

Relationship between Eimeria tenella development and host cell apoptosis in chickens Zhang Yan,∗,1 Zheng Ming-xue,∗,2 Xu Zhi-yong,∗,†,1 Xu Huan-cheng,∗ Cui Xiao-zhen,∗ Yang Sha-sha,∗ Zhao Wen-long,∗ Li Shan,∗ Lv Qiang-hua,∗ and Bai Rui∗ ∗

College of Animal Science and Technology, Shanxi Agricultural University, Taigu, 030801, China; and † College of Animal Science, Henan Institute of Science and Technology, Xinxiang, 453003, China

Key words: apoptosis, chick embryo, development, Eimeria tenella, cecal epithelial cell 2015 Poultry Science 00:1–10 http://dx.doi.org/10.3382/ps/pev293

INTRODUCTION

Cell apoptosis is an elementary process that, along with proliferation, growth, and differentiation, maintains tissue and organ homeostasis by deleting unrepairably damaged or mutated cells that have lost their functions. Moreover, apoptosis is an active, genetically controlled process and an important regulator of a host’s response to infection by a variety of intracellular protozoan parasites (L¨ uder et al., 2001; Balamurugan et al., 2002). Chick cecal epithelial cells are the host cells of E. tenella in vivo, so the mechanism by which E. tenella infects chickens and injures cells can be studied using a primary culture of chick embryo cecal cells to develop E. tenella in vitro. Some apoptosis-inducing factors are also essential for parasite development. Previous studies showed that parasitic pathogens spread in the host by inducing or inhibiting host-cell apoptosis (Bienvenu et al., 2010; Engelbrecht et al., 2012; La¨etitia et al., 2013). These systems work to regulate the host’s immune response, aid transmission within the host, and facilitate general intracellular survival (L¨ uder et al., 2001). To protect its host cell from apoptosis, Eimeria bovis sporozoites can increase the expression of the

Chicken coccidiosis, a protozoan disease caused by several varieties of Eimeria parasites in chicken intestinal mucosa epithelial cells, is responsible for enormous economic losses in the global poultry industry (Huang, 2009; Li, 2011; Hady and Zaki, 2012; Martynova-Van et al., 2012; Ritzi et al., 2014; Yin et al., 2015). Eimeria tenella (E. tenella) is one of the most pathogenic coccidia to infect chickens. Infections may be characterized by the presence of blood in droppings and bring high morbidity and mortality rates. Death may occur unexpectedly owing to excessive blood loss. Currently, coccidiosis is controlled mainly with drugs and vaccines; however, these are economically restrictive because of the huge cost to the international poultry industry (Shirley et al., 2005; Wallach et al., 2008).  C 2015 Poultry Science Association Inc. Received April 18, 2015. Accepted August 19, 2015. 1 Indication: Xu Zhi-yong and Zhang Yan are joint first authors. 2 Corresponding author: [email protected]

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we demonstrated that untreated cells had less apoptosis at 4 h and, inversely, more apoptosis at 24 to 120 h compared with control cells. Furthermore, after the application of Z-LEHD-FMK, terminal deoxynucleotidyl transferase–mediated dUTP nick-end labeling assays, and translation of phosphatidyl serines to the host cell plasma membrane surface, the treated group chick embryo cecal epithelial cells exhibited decreased apoptosis and DNA injuries (P < 0.01) at 24 to 120 h. However, light microscopy showed that E. tenella infection rates of treated cells were higher (P < 0.01) than untreated cells during the whole experimental period. Together, these observations suggest that E. tenella can protect host cells from apoptosis at early stages of development but can promote apoptosis during the middle to late stages. In addition, the inhibition of host cell apoptosis can be beneficial to the intracellular growth and development of E. tenella.

ABSTRACT Coccidiosis causes considerable economic losses in the poultry industry. At present, the pathology of coccidiosis is preventable with anticoccidials and vaccination, although at considerable cost to the international poultry industry. The purpose of the present study was to elucidate the relationship between Eimeria tenella development and host cell apoptosis in chickens, which provides a theoretical basis for further study of the injury mechanism of E. tenella and the prevention and treatment of coccidiosis. Cecal epithelial cells from chick embryo were used as host cells in vitro. In addition, flow cytometry, terminal deoxynucleotidyl transferase–mediated deoxyuridine triphosphate (dUTP) nick-end labeling, and histopathological assays were used to detect the dynamic changes in E. tenella infection rates, DNA injury rates, and apoptosis rates in groups treated with and without the caspase-9 inhibitor Z-LEHD-FMK. Following E. tenella infection,

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MATERIALS AND METHODS Experimental Animals A total of twenty 1-d-old chicks and sixty-six 15-dold specific pathogen-free (SPF) chicken embryos were used in this study; they were provided by Beijing Meri Avignon Laboratory Animal Technology Co., Ltd. (Beijing, China). The 1-d-old chicks were raised under strict pathogen-free conditions.

Parasite The E. tenella Shanxi virulent strain (EtSX01) used in the current study was provided by the Laboratory of Veterinary Pathology in the College of Animal Science and Technology (Shanxi Agricultural University, Shanxi, China).

Preparation of E. tenella Sporozoites Twenty 20-d-old SPF chicks were infected orally with 6,000 sporulated E. tenella oocysts. After oocysts were isolated and sporulated, sporozoites were excysted as previously described (Zheng et al., 2011). In brief, from day 7 to day 8 following oral infection, the oocysts were isolated from feces by using standard salt flotation techniques. Isolated oocysts were sporulated in 2.5% dichromicum kalium (Fengchuan Corp, Tianjin, China) to attain 85% or more sporulation (at 28◦ C with aerobic atmosphere, 2 to 3 d). Sporocysts were released from sporulated oocysts by mechanical rubbing with a glass homogenizer. The released sporocysts were digested with excystation fluid (10% chicken bile from 20-d-old SPF chickens, 0.75% trypsin, Solarbio, Beijing, China, v/v) at 41◦ C, shaken for 1 h, centrifuged at 950 g for 10 min, and finally purified

with a 1,400 mesh filter. Free sporozoites were collected and dissolved to a final concentration of 1.5 × 105 sporozoites/mL in a culture medium consisting of 33% low glucose Dulbecco’s modified Eagle medium (DMEM) (HyClone, Logan, America), 65.5% MEM199/EBSS (Hyclone), and 1.5% chicken serum (20-d-old SPF chicken) (v/v) supplemented with 100 U/mL penicillin (Huabei Corp., Shijiazhuang, China), 100 U/mL streptomycin (Weierkong Corp., Sichuan, China), 0.02 μg/mL epidermal growth factor (PEPRO TECH), 0.1 mg/mL Heparin (Solarbio), 1.1 mg/mL sodium pyruvate (Solarbio), 0.05 μg/mL insulin (Solarbio), 0.5 mmol/L L-glutamine (Solarbio), 0.05 μg/mL folic acid (Solarbio), 0.05 μg/mL VB6 (Solarbio), 0.05 μg/mL VB1 (Solarbio), and 0.3 mg/mL glucose (Tianjin Chemical Agent Corp., Tianjin, China).

Primary Culture of Chick Embryo Cecal Cells and Parasite Infection Chick embryo cecal epithelial cells were taken from sixty-six 15-d-old SPF chick embryos (Merial Vital Corp, Beijing, China) and cultured as described previously (Gu et al., 2009b; Gu et al., 2011). Briefly, the ceca were removed and placed in PBS, minced to 1mm3 , digested by 50mg/l thermolysin (Sigma) at 37 ◦ C for 2 h, rinsed with PBS, and centrifuged at 220 g for 5 min to remove single cells. Before being plated, based on the distinction of different cell’s adherence speed, the precipitated dissociated cells were cultivated for 70 min at 41◦ C in a humidified, 8% CO2 -air incubator to remove all other cells except the cecal epithelial cells in a culture medium consisting of low glucose DMEM supplemented with 10% FBS (Sijiqing Corp., Hangzhou, China), 100 U/mL penicillin, 100 U/mL streptomycin, 0.02 μg/mL epidermal growth factor, 0.1 mg/mL Heparin, 1.1 mg/mL sodium pyruvate, 0.05 μg/mL insulin, and 0.5 mmol/l L-glutamine. Maintained at 41◦ C in a humidified, 8% CO2 -air incubator, the nonadherent cecal epithelial cell aggregates were plated at concentrations of either 2 × 105 live cell aggregates per well on a 6-well tissue culture plate or at 3.1 × 104 live cell aggregates per well on a 48-well chamber slide with 2.5% FBS instead of 10% FBS in addition to the culture medium described earlier. Confluent chick embryo cecal epithelial cell monolayers in 6-well tissue culture plates and 48-well chamber slides were infected with 3 × 105 and 1 × 105 freshly excysted E. tenella sporozoites per well, respectively.

Experimental Protocol When the adherence rate exceeded 90%, chick embryo cecal epithelial cells were randomly divided into three experimental groups, as follows: (1) control group (group C); (2) untreated group (cells infected with E. tenella sporozoites, group T0); and (3) treated group (cells infected with E. tenella sporozoites and treated

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anti-apoptotic factors cellular FLICE inhibitory protein and cellular inhibition of apoptosis protein 1 (Lang et al., 2009). In chick cells infected with E. tenella, apoptosis generally occurs during the initial stage of infection, although necrosis occurs during the final stage (Gu et al., 2010). In Madin-Darby bovine kidney cells infected with E. tenella, apoptosis can be inhibited, which suggests that there is a unique mechanism by which E. tenella sporozoites may ensure their own survival and development in the host cell by extending the survival time of host cells (Deng et al., 2011). To date, the relationship between E. tenella development and host-cell apoptosis in chickens has not yet been intensively studied or systematically summarized. In the present study, flow cytometry (FCM), terminal deoxynucleotidyl transferase–mediated dUTP nick-end labeling (TUNEL), and histopathological techniques were performed in vitro. This research provides a theoretical basis for further studying the injury mechanism of E. tenella and the prevention and treatment of coccidiosis.

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with 20 μmol Z-LEHD-FMK [Biovision, California, America], group T1).

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DNA injury rate of uninfected cells (%) = number of apoptotic cells uninfected

Adherence rate (%) = area of hole bottom attached

by E. tenella/200 × 100

by cells/total area of hole bottom × 100

Hematoxylin and Eosin Stain

TUNEL Assays Apoptosis-induced DNA strand breakage was detected by an In Situ Cell Death Detection Kit (Roche, Bael, Switzerland). At 4, 24, 48, 72, 96, and 120 h following infection, 48-well chamber slides fixed by paraformaldehyde (4% w/v, 1 h, room temperature; Solarbio) were rinsed twice with PBS for 5 min, blocked in confining liquid (10 min, room temperature), and rinsed again twice with PBS for 5 min. Chamber slides were permeabilized with 0.1% (v/v) Triton X-100 (Solarbio) (4◦ C, 2 min), rinsed twice with PBS, and exposed to TUNEL reaction mixture in a dark humidified chamber at 37◦ C for 1 h. After rinsing three times with PBS and air dried, slides were added to 50 μL converterPeroxidase. Slides were then incubated in a humidified atmosphere in the dark at 37◦ C for 30 min and rinsed with PBS. Dry slides reacted with 100 μL 3,3 diaminobenzidine (10 min, room temperature), rinsed with PBS, and stained with hematoxylin and eosin. Two hundred cell samples selected randomly were analyzed by light microscopy to observe DNA injury. The broken DNA appeared brown in the apoptotic cells. DNA injury rate of total cells (%) = number of apoptotic cells/200 × 100

DNA injury of E. tenella − infected cells (%) = number of apoptotic cells infected by E. tenella/200 × 100

Phosphatidyl serine (PS) is embedded in the cytoplasmic surface of cell membranes but translocates from the inner to the outer leaflet of the plasma membrane during apoptosis. Annexin V, a protein that has a high affinity for PS, is used to stain apoptotic cells with green fluorescence. Propidium iodide (PI) is impermeable to live cells and early apoptotic cells but stains dead cells with red fluorescence by tightly binding to the nucleic acids in the cell. The employed detection kit (Biovision) applies fluorescein isothiocyanate (FITC)conjugated annexin V. At 4, 24, 48, 72, 96, and 120 h following infection, chick embryo cecal cells in 6-well tissue culture plates were harvested by 0.25% trypsin, rinsed in PBS, centrifuged at 600 g for 5 min, suspended with 200 μL binding buffer, and incubated with 5 μL annexin V-FITC and 1 μL PI (30 min, room temperature, in the dark). Afterward, cells were added to 200 μL binding buffer and subjected to FCM analyses. Annexin V−/PI− quadrant: viable cells; annexin V+/PI− quadrant: early apoptotic cells; annexin V+/PI+ quadrant: late apoptotic and necrotic cells.

Statistical Analysis All quantitative data were analyzed by ANOVA in a SPSS 19.0 (SPSS Inc., Chicago, IL, USA) and expressed as mean ± SE. A P-value of <0.05 was considered significant.

RESULTS AND DISCUSSION This study demonstrated that inhibiting apoptosis in E. tenella-infected chick embryo cecal epithelial cells increases E. tenella infection rates and may be beneficial to the intracellular development of E. tenella. TUNEL assays showed that the total cells of group T0 had fewer (P < 0.01) DNA injuries at 4 h (2.60 ± 0.19,4 h) and, conversely, more DNA injuries (P < 0.01) from 24 h to 120 h (6.20 ± 0.20, 24 h; 7.80 ± 0.12, 48 h; 8.70 ± 0.20, 72 h; 8.80 ± 0.12, 96 h; 9.90 ± 0.19, 120 h) compared with group C (4.50 ± 0.22, 4 h; 3.00 ± 0.16, 24 h; 3.90 ± 0.19, 48 h; 4.30 ± 0.26, 72 h; 4.70 ± 0.26, 96 h; 5.30 ± 0.26, 120 h) (Figure 1A-D). Likewise, the DNA injury rates of E. tenella-infected cells were lower (P < 0.01) at 4 h (1.40 ± 0.19, 4 h) and higher (P < 0.01) at 24 to 120 h (7.90 ± 0.19, 24 h; 10.10 ± 0.19, 48 h; 11.90 ± 0.19, 72 h; 12.00 ± 0.22, 96 h; 13.60 ± 0.19, 120 h) compared to uninfected cells (3.20 ± 0.12, 4 h; 5.60 ± 0.10, 24 h; 6.90 ± 0.19, 48 h; 7.60 ± 0.10, 72 h; 8.10 ± 0.19, 96 h; 9.40 ± 0.19, 120 h) in group

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At 4, 24, 48, 72, 96, and 120 h following infection, 48-well chamber slides were fixed with Bouin’s solution for 8 min, rinsed with 70% alcohol, and maintained for 30 min. Following hydration with distilled water for 15 min, chamber slides were stained with Lillie-Mayer’s hematoxylin (Solarbio) for 30 min, washed in distilled water, differentiated with a mixture of 1% hydrochloric acid and 70% alcohol for 1.5 min, blued in running tap water for 15 min, and dehydrated with 50, 70, 80, 90, and 95% alcohol in sequence. Slides were exposed to 1% eosin (Solarbio) for 25 min, dehydrated with 100% alcohol, cleaned in xylene solution, and mounted with neutral balsam. Sample E. tenella infections were observed in 200 cells selected randomly by light microscopy. Infection rates at each time point (%) = number of infected cells on each time point/200 × 100.

Annexin V and PI Assays

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Figure 1. TUNEL stains in chick embryo cecal epithelial cells. A1 to C1: TUNEL-positive nuclei of total cells in groups C, T0, and T1 at 4 h. A2 to C2: TUNEL-positive nuclei of total cells in groups C, T0, and T1 at 24 h. A3 to C3: TUNEL-positive nuclei of total cells in groups C, T0, and T1 at 48 h. A4 to C4: TUNEL-positive nuclei of total cells in groups C, T0, and T1 at 72 h. A5 to C5: TUNEL-positive nuclei of total cells in groups C, T0, and T1 at 96 h. A6 to C6: TUNEL-positive nuclei of total cells in groups C, T0, and T1 at 120 h. D: quantitative determination of apoptotic nuclei of total cells (n = 5). E: quantitative determination of apoptotic nuclei of infected and noninfected cells (n = 5). F: Quantitative determination of apoptotic nuclei of group C cells and noninfected cells in group T0 (n = 5). ∗∗ P < 0.01 vs. C, ++ P < 0.01 vs.T0, ## P < 0.01 vs. infected cells in the same group,  P < 0.01 vs. group C cells, the same as following figures. Magnification 400×.

T0 (Figure 1E). Compared with group C cells (8.50 ± 0.41, 4 h; 2.71 ± 0.21, 24 h; 3.97 ± 0.31, 48 h; 4.59 ± 0.26, 72 h; 5.84 ± 0.13, 96 h; 5.57 ± 0.27, 120 h), PS translocation also indicated that group T0 cells had less (P < 0.01) early apoptosis at 4 h (4.61 ± 0.15, 4 h)

and, in contrast, more early apoptosis (8.82 ± 0.36, 24 h; 6.88 ± 0.21, 48 h; 7.11 ± 0.18, 72 h; 9.57 ± 0.45, 96 h; 10.60 ± 0.45, 120 h) (P < 0.01) at 24 to 120 h (Figure 2A-D). In addition, group T0 cells (19.07 ± 0.80, 48 h; 19.32 ± 1.15, 72 h; 19.44 ± 0.61, 96 h;

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D 15

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Figure 1. Continued.

23.06 ± 1.46,120 h) had more late apoptosis and necrosis than group C cells (7.25 ± 0.37, 48 h; 10.34 ± 0.54, 72 h; 10.52 ± 0.39, 96 h; 10.90 ± 0.41, 120 h) from 48 h to 120 h (Figure 2A to C, E). Furthermore, the apoptosis rates in both groups directly increased with the development of E. tenella (Figure 2A to E).

The results obtained by the aforementioned two methods were consistent with each other, strongly implying that E. tenella may protect host cells from apoptosis at early development stages but promote apoptosis during middle to late development stages. Via ultrastructural analysis as well as the TUNEL method, it was found

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Figure 2. Annexin V/PI-based apoptosis detection in chick embryo cecal epithelial cells. A1 to 6: annexin V/PI-based apoptosis detection in group C at 4, 24, 48, 72, 96, and 120 h; B1 to 6: annexin V/PI-based apoptosis detection in group T0 at 4, 24, 48, 72, 96, and 120 h; C1 to 6: annexin V/PI-based apoptosis detection in group T1 at 4, 24, 48, 72, 96, and 120 h; D: quantitative determination of early apoptosis (n = 5). E: quantitative determination of late apoptosis and necrosis (n = 5). + P < 0.05 vs. T0.

that cecal mucosa epithelial cell apoptosis increased at 2 d after chicks were infected with E. tenella, and lesions were the most serious from 4 to 6 d postinfection (Gu et al., 2010). Our results were in agreement

with previous research in vivo (Gu et al., 2010). Group T0 had decreased apoptosis at 4 h, which may be due to the fact that during early development, E. tenella requires a stable intracellular environment to obtain

EIMERIA TENELLA AND CELL APOPTOSIS

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essential nutrients and evade host immune attack (Lang et al., 2009). The NF-κB signaling system is a crucial cellular survival mechanism (Salminen et al., 2012). Before the development of second-generation schizonts is completed, E. tenella may directly activate the NF-κB pathway in host cells to further inhibit host-cell apoptosis but after development is completed may induce host-cell apoptosis (Del Cacho et al., 2004). In other words, E. tenella may prevent the expression of the NFκB response gene at its middle to late developmental stage (Del Cacho et al., 2004) then further decrease the expression of the anti-apoptotic proteins Bcl-2 and BclXL, which accelerate host-cell apoptosis and promote the release of merozoite (Rasul et al., 2012). Apoptosis, which is one of the main ways in which E. tenella can damage cells, can be triggered by E. tenella sporozoite invasion, the subsequent schizogony, and the eventual rupture of the host cells (Gu et al., 2010). Host cells

supply necessary nutrients for the growth and development of E. tenella during infection, and both nutrients and energy are also consumed to maintain cell apoptosis. Therefore, apoptotic cells are not conducive to E. tenella development (Gu et al., 2009a). As seen in this study, at 24 to 120 h following E. tenella infection, the parasites have already reached the phase of schizogamy or gametogony (Figure 3A to F). The apoptosis process of host cells may be sped up by the loss of nutrients (Gu et al., 2009a), the destruction of cell structure (Gu et al., 2010), or even the inhibition of NF-kB expression (Del Cacho et al., 2004). Caspase-9, a member of the caspase family of cysteine proteases that have been implicated in apoptosis and cytokine processing, is essential for apoptosis during the normal development of the central nervous system (Kuida, 2000). Because caspase-9 inhibits apoptosis (W¨ urstle et al., 2012; Wang et al.,

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Figure 2. Continued.

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Figure 3. E. tenella infection rates. A: cecal epithelial cells in group C at 48 h; B: cecal epithelial cells in group T0 at 48 h, and “→” represents trophozoites; C: cecal epithelial cells in group T1 at 48 h, and “→” represents immature schizont; D: cecal epithelial cells in group C at 72 h; E: cecal epithelial cells in group T0 at 72 h, and “→” represents mature schizont; F: cecal epithelial cells in group T1 at 72 h, and “→” represents mature schizont. G: quantitative determination of E. tenella infection (n = 5). Magnification 400×.

2013; Xue et al., 2013), it is an important therapeutic target of various diseases related to apoptosis. Furthermore, a variety of apoptotic stimuli can regulate caspase-9 (Kim et al., 2015). For example, cells stimulated by internal or external factors can release various apoptotic factors to partially activate caspase-9 or caspase-3, thereby inducing cell apoptosis by causing DNA fragmentation, the dissolution of protein skeletons, or other biological processes (Zheng et al., 2011;

Jochen et al., 2013). Mitochondria play a vital role in regulating and transmitting apoptotic signals initiated by multiple stressors (Green and Reed, 1998). Z-LEHD-FMK, a specific inhibitor against caspase-9, can specifically bind and inactivate caspase-9, thereby mediating cell apoptosis (Zhu et al., 2007). Metoprolol and Z-LEHD-FMK may protect rat myocardium during coronary microembolization by inhibiting apoptosis and improving cardiac function, most likely by

EIMERIA TENELLA AND CELL APOPTOSIS

We further found by a TUNEL assay that the noninfected cells of group T0 had fewer DNA injuries at 4 h and, conversely, had more DNA injuries from 24 to 120 h compared with those of noninfected control cells (Figure 1F), which showed that the culture medium used may have generated some unknown components during the development of E. tenella that influenced apoptosis in noninfected cells of group T0. In addition, there was no significant difference between the DNA injury rates of E. tenella–infected and uninfected cells of group T1 (Figure 1E); this may be a result of the joint action of both Z-LEHD-FMK and the unknown components in the culture medium. Further studies are needed to investigate the specific components. In conclusion, chick embryo cecal epithelial cells infected with E. tenella exhibited decreased apoptosis and increased E. tenella infection rates at early developmental stages of E. tenella. Furthermore, along with the development of E. tenella, the E. tenella–infected cells had increased apoptosis in addition to decreased infection rates. Inhibition of host-cell apoptosis is beneficial to the intracellular growth and development of E. tenella.

ACKNOWLEDGMENTS This study was funded by a grant supported by the National Natural Science Foundation of China (31272536).

INSTRUCTIONS TO CONTRIBUTORS Y. Zhang and Z. Y. Xu carried out most of the experiments and wrote the manuscript and should be considered primary authors. M. X. Zheng critically revised the manuscript and the experiment design. H. C. Xu, X. Z. Cui, S. S. Yang, W. L. Zhao, S. Li, Q. H. Lv, and R. Bai helped with the experiment. All of the authors read and approved the final version of the manuscript.

REFERENCES Balamurugan, K., R. Rajaram, T. Ramasami, and S. Narayanan. 2002. Chromium(III)-induced apoptosis of lymphocytes: death decision by ROS and Src-family tyrosine kinases. Free Radic. Biol. Med. 33:1622–1640. Bienvenu, A. L., E. Gonzalez-Rey, and S. Picot. 2010. Parasit. Vectors. 3:106. doi:10.1186/1756-3305-3-106. Del Cacho, E., M. Gallego, F. L´ opez-Bernad, J. Qu´ılez, and C. S´ anchez-Acedo. 2004. Expression of anti-apoptotic factors in cells parasitized by second-generation schizonts of Eimeria tenella and Eimeria necatrix. Vet. Parasitol. 125:287– 300. Deng, J. J., L. X. Wang, and J. An. 2011. Eimeria Tenella Sporozoite inhibits apoptosis in infected host cells. J. Beijing Agric. College. 26:24–27. Engelbrecht, D., P. M. Durand, and T. L. Coetzer. 2012. On programmed cell death in Plasmodium falciparum: status quo. J. Trop. Med. doi:10.1155/2012/ 646534.

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inhibiting apoptosis or the mitochondrial apoptotic pathway (Su et al., 2013). Flow cytometric analysis has shown that inhibition of the intrinsic pathway by a caspase-9 inhibitor (Z-LEHD-FMK) significantly suppresses serum deprivation–induced apoptosis; however, a caspase-8 inhibitor (Z-IETD-FMK) did not reduce apoptotic cell death (Li et al., 2014). In our study, a TUNEL assay illustrated that the total cells in group T1 (1.40 ± 0.19, 4 h; 3.90 ± 0.10, 24 h; 5.70 ± 0.12, 48 h; 6.70 ± 0.12, 72 h; 6.80 ± 0.20, 96 h; 7.30 ± 0.12, 120 h) had fewer DNA injuries than group T0 at 4 to 120 h (Figure 1A to D), which suggests that Z-LEHD-FMK significantly inhibited the apoptosis of host cells. This result was consistent with previous research (Zhu et al., 2007; Li et al., 2014). In terms of early apoptosis, FCM analysis also showed no significant difference between groups T0 and T1 (4.29 ± 0.13, 4 h) (P > 0.05) at 4 h following infection, whereas the early apoptosis rates of group T1 (3.79 ± 0.33, 24 h; 3.49 ± 0.21, 48 h; 5.78 ± 0.08, 72 h; 6.22 ± 0.29, 96 h; 6.83 ± 0.11, 120 h) were significantly lower (P < 0.01) than those of group T0 at 24 to 120 h (Figure 2D). As for late apoptosis and necrosis, no statistically significant difference was observed between groups T0 and T1 (4.97 ± 0.14, 4 h) at 4 h following infection (P > 0.05) (Figure 2E). We also found no significant difference among the three groups at 24 h (P > 0.05), although at 48, 72, 96, and 120 h postinfection, group T1 (12.03 ± 0.35, 48 h; 13.74 ± 0.20, 72 h; 13.63 ± 0.40, 96 h; 15.30 ± 0.29, 120 h) had fewer (P < 0.01 or P < 0.05) damaged cells than group T0 (19.07 ± 0.80, 48 h; 19.32 ± 1.15, 72 h; 19.44 ± 0.61, 96 h; 23.06 ± 1.46, 120 h) (Figure 2E); this further implies that Z-LEHD-FMK significantly suppresses E. tenella– induced apoptosis. Although no significant difference in E. tenella infection rates was observed between group T1 (40.80 ± 0.46, 4 h) and group T0 (41.30 ± 0.37, 4 h) at 4 h following infection by 1 × 105 E. tenella sporozoites per chamber slide (P > 0.05), group T1 (34.90 ± 0.19, 24 h; 31.90 ± 0.19, 48 h; 30.60 ± 0.19, 72 h; 25.50 ± 0.22, 96 h; 22.60 ± 0.19, 120 h) exhibited higher E. tenella infection rates (P < 0.01) than group T0 (33.10 ± 0.19, 24 h; 26.10 ± 0.29, 48 h; 25.10 ± 0.29, 72 h; 19.50 ± 0.27, 96 h; 17.00 ± 0.35, 120 h) at 24, 48, 72, 96, and 120 h following infection (Figure 3G); this shows that there were sufficient levels of well-developed E. tenella that could be released to invade other cells, resulting in group T1 having a higher infection rate than group T0. In other words, E. tenella infection rates were higher when host-cell apoptosis rates were low. It is likely that inhibiting caspase-9 activity may have blocked the mitochondrial apoptotic pathway, thereby decreasing the host-cell apoptosis rates. Furthermore, when apoptosis rates were lower, more host cells could be invaded by merozoites, indirectly increasing infection rates. Together with the reasons discussed earlier, inhibition of host-cell apoptosis is likely conducive to prolonging E. tenella development in cells, leading to more infection.

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