Molecular cloning, sequence analysis, and function of the intestinal epithelial stem cell marker Bmi1 in pig intestinal epithelial cells1 C.-M. Li,*2 H.-C. Yan,*2 H.-L. Fu,* G.-F. Xu,* and X.-Q. Wang*3 *Guangdong Provincial Key Laboratory of Agro-Animal Genomics and Molecular Breeding, College of Animal Science, South China Agricultural University, Guangzhou 510642, China

ABSTRACT: In the present work, we cloned the fulllength cDNA of the pig Bmi1 gene (BMI1 polycomb ring finger oncogene), which has been indicated as an intestinal epithelial stem cell (IESC) marker in other mammals. This paper provides the first report of the function of Bmi1 in pig intestinal epithelial cells and a brief description of its underlying mechanism. Rapid amplification of cDNA ends technology was used to clone the complete pig Bmi1 sequence, and a Bmi1-pcDNA3.1 vector was constructed for transfection into an intestinal porcine epithelial cell line (IPEC-1). The proliferation ability of the cells was estimated using the MTT assay and the EdU incorporation method at different time points after seeding. Cell cycle information was detected by flow cytometry. The mRNA abundances of cell cycle-related genes were also measured. The results indicated that the pig Bmi1 cDNA

is 3,193 bp in length and consists of a 981 bp open reading frame, a 256 bp 5´ untranslated region (UTR), and a 1,956 bp 3´ UTR. The transcript contains no signal peptides, and there are no transmembrane regions in the pig Bmi1 coded protein, which has a total of 326 AA. The overexpression of the pig Bmi1 in the IPEC-1 cells led to increased cell proliferation and a lower percentage of cells in the G1 and S phases (P < 0.05), along with a higher percentage of cells in the G2 phase (P < 0.05). Furthermore, the gene expression levels of PCNA, Cyclin D1, Cyclin D2, Cyclin B, CDK1, and CDK2 were all elevated (P < 0.05) by Bmi1 overexpression, while the gene expression levels of Cyclin A2 and p21 showed little difference (P > 0.05). Our data suggested that pig Bmi1 can increase the proliferation of IPEC-1 cells by promoting the G1/S transition and the overall cell cycle process.

Key words: Bmi1, cell cycle, intestinal epithelial stem cell marker, pig, proliferation, RACE © 2014 American Society of Animal Science. All rights reserved. INTRODUCTION The intestinal epithelium is one of the most proliferative tissues in mammals, and it constantly undergoes physiological regeneration. This tissue has an absorptive function and acts as a protective epithelial barrier. The rapid renewal of this tissue is driven by a pool of multipotent intestinal epithelial stem cells (IESC). In recent years, significant progress has been made toward understanding the IESC based on the mouse 1Supported by funding from the National Natural Science Foundation of China (NSFC; grant 31330075), the National Basic Research Program of China (2013CB127302), the National 948 Program of China (2011G35), the Educational Commission of Guangdong Province, China (2012CXZD0015), and the Science and Technology Planning Project of Guangdong Province, China (2012B020305006). 2These authors contributed equally to this work. 3Corresponding author: [email protected] Received August 19, 2013. Accepted November 10, 2013.

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model and studies of human health. Two stem cell populations have been identified in the epithelium of the small intestine: fast-cycling stem cells termed crypt base columnar cells (CBC), which express Lgr5 and are ubiquitous (Barker et al., 2007; Sato et al., 2009), and slower-cycling stem cells, which express the Polycomb protein Bmi1 and can replenish the former cell type when injured (Sangiorgi and Capecchi, 2008; Li and Clevers, 2010; Tian et al., 2012). However, little is known about the identity of pig IESC. While Lgr5-expression (Lgr5+) populations can already be easily isolated using CD24 (von Furstenberg et al., 2011; King et al., 2012) or CD133/CD44 (Hou et al., 2011) with a Fluorescence Activated Cell Sorter (FACS), the isolation of Bmi1 (BMI1 polycomb ring finger oncogene)expressing (Bmi1+) populations have been hampered by the necessity of developing a transgenic animal. Thus, further study of pig IESC should focus on the isolation and identification of Bmi1+ IESC.

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Bmi1, a Polycomb-group transcriptional repressor, has emerged as a key regulator in several cellular processes, including the self-renewal of neural, hematopoietic, intestinal, and mammary stem cells (Park et al., 2003; Reinisch et al., 2006; Pietersen et al., 2008) and cancer cell proliferation. The protein functions largely, but not exclusively, by repressing the expression of p16Ink and p19Arf (He et al., 2009). Bmi1 can also regulate other cell cycle proteins (Zencak et al., 2013). Several studies regarding Bmi1 in the intestine have been published. Reinisch et al. (2006) first recognized Bmi1 as a protein expressed in the stem cells, specialized cells, and tumors of the gastrointestinal tract (Reinisch et al., 2006). Sangiorgi and Capecchi (2008) then identified the protein as an IESC marker in vivo. Tian et al. (2012) supported these previous results and further demonstrated that Bmi1+ stem cells may represent both a reserve stem cell pool in the case of injury to the epithelium of the small intestine and a source for the replenishment of Lgr5+ cells under nonpathological conditions. However, a method for isolating the Bmi1+ IESC has yet to be discovered, and the function of Bmi1 on pig intestine development, which appears to be important for the in vitro culture of Bmi1+ IESC cells, remains unknown. Considering the pivotal importance of Bmi1 to the isolation, identification, and long-term in vitro culturing of pig IESC, this study aimed to clone the pig Bmi1 and to explore the effects of Bmi1 on pig intestinal epithelial cell proliferation. MATERIALS AND METHODS Intestinal Tissue Sample Collection Three male and 3 female 24-d-old LanTang piglets (a kind of Chinese native pig) were euthanized with sodium pentobarbital before sampling. The entire small intestine was then rapidly removed, washed, and frozen in liquid nitrogen. All procedures were approved by the Animal Care Committee of the South China Agricultural University. RNA Isolation and Reverse Transcription Total RNA was isolated from the mixture of duodenum and jejunum epithelium or cells using Trizol reagent (Invitrogen, Carlsbad, CA) following the manufacturer’s instructions. RNA quality was assessed by agarose gel electrophoresis (1%). The RNA had an OD260:OD280 ratio between 1.8 and 2.0. Reverse transcription was performed using 1 mg of total RNA and Moloney murine leukemia virus reverse transcriptase (Invitrogen). The reverse transcription conditions for the cDNA amplification were 65°C for 5 min, 37°C for 30 min, and 70°C for 15 min. Synthesis of the cDNA first

strand was performed using random primers (N10) and Superscript II reverse transcriptase (Invitrogen). Clone of Porcine Bmi1 Full-Length cDNA Sequence The primers for the porcine Bmi1 partial cDNA sequences were designed using Primer Premier 5.0 (Premier Biosoft, Palo Alto, CA) based on a conserved region of the human (NM_005180.8) and Mus musculus (NM_007552.4) sequences. The two primer pairs were 5´-GCCGCTTGGCTCACATTC-3´ (forward), 5´-TTCCTC AACAGTCTCAGGTATCA-3´ (reverse) and 5´-GGTAACCACCAATCTTCCTT-3´ (forward), 5´-AAACACCCTCACAAATAACTG-3´ (reverse). The expected fragment sizes were 1,087 bp and 1,495 bp, respectively. After sequencing and comparison, the 2 fragments were contiged. The 5´ and 3´ rapid amplification of cDNA ends (RACE) primers (Table 1) were then designed based on the contiged sequence. The 5´ and 3´ ends were obtained using the SMART RACE Technique, as previously described (Rajalingam et al., 2001). The resulting 5´ and 3´ end sequences were jointed with the previous partial sequences to construct the full-length Bmi1 cDNA sequence. The open reading frame (ORF) was later cloned to verify these results. The structure and function of the Bmi1 coded protein sequence was then predicted and analyzed. Construction and Identification of Recombinant Plasmid Bmi1-pcDNA3.1 A primer pair (Table 1) for the pig Bmi1 coding sequence (CDS) containing the Kpn1 and Xho1 enzyme digest sites was designed and used to amplify the CDS. The purified PCR product was then linked with pcDNA3.1, which was cut with the same enzymes. The ligation product was transferred into DH5α, which was spread on an antiampicillin LB plate and cultured at 37°C for 12–16 h. Several clones were then chosen from the plate, and their plasmids were extracted for PCR and enzyme digestion and sequencing identification. The purification was performed using the EndoFree Plasmid Kit (TIANGEN, Beijing, China) according to the manufacturer’s instructions. Cell Culture and Transfection of Bmi1-pcDNA3.1 The intestinal porcine epithelial cell line (IPEC-1) donated by Prof. Guoyao Wu (Texas A&M University) was grown in serial passage in uncoated plastic culture flasks (100 mm2) in growth media (GM; DMEM-F12, containing 5% FBS, insulin [5 µg/mL], transferrin ([5 µg/mL], selenium [5 ng/mL], epidermal growth factor [5 µg/L], penicillin [50 µg/mL], and streptomycin [4 µg/mL]) at 37°C in a 5% CO2 incubator. The trans-

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Table 1. Primers used for cloning and quantitative real-time PCR1 Gene/Accession GSP1 NGSP1 GSP2 NGSP2 Bmi1 CDS Cyclin D1 (AK400348.1) Cyclin D2 (NM_214088.1) Cyclin B (GQ184631.1) CyclinA2 (NM_001177926.1) PCNA (DQ473295.1) CDK1 (NM_001159304.2) CDK2 (XM_003481615.1) CDK4 (NM_001123097.1) p21 (XM_001929558.1) 18S (NR_046261.1)

Primer sequence 5´-TACACACATTAGGTGGGGATTTAGC-3´ 5´-GCTCAGTGATCTTGATTCTTGTTGTTCG –3´ 5´-TATGCCCTCTCTGTAATGTCTGGAA –3´ 5´-GTGTGTTCATCACCCATCAGTTATTTG- 3´ F: 5´-CGGGGTACCATGCATCGAACAACAAGAAT- 3´ R:5´-CCGCTCGAGTCAGGTATCAACCAGAAGAAG-3´ F: 5´-GCGAGGAACAGAAGTGCG-3´ R: 5´-TGGAGTTGTCGGTGTAGATGC-3´ F: 5´-TTACCTGGACCGCTTCTTG-3´ R: 5´-GAGGCTTGATGGAGTTGTCG-3´ F: 5´-TGGCTAGTGCAGGTTCAG-3´ R: 5´-CAGTCACAAAGGCAAAGT-3´ F: 5´-TTAGGGAAATGGAGGTTA-3´ R: 5´-TAGTTCACAGCCAAATGC-3´ F: 5´-TACGCTAAGGGCAGAAGATAATG-3´ R: 5´-CTGAGATCTCGGCATATACGTG-3´ F: 5´-CCCTCCTGGTCAGTTCAT-3´ R: 5´-TAGGCTTCCTGGTTTCC-3´ F: 5´-AAACAAGTTGACGGGAGA-3´ R: 5´-GTGAGAATGGCAGAAAGC-3´ F: 5´-GCATCCCAATGTTGTCCG-3´ R: 5´-GGGGTGCCTTGTCCAGATA-3´ F: 5´-ACCCCTTCCCCATACCC-3´ R: 5´-TTCCTAACACCCATGAAACTG-3´ F: 5´-AATTCCGATAACGAACGAGACT-3´ R: 5´-GGACATCTAAGGGCATCACAG-3´

Product size (bp)

981 192 155 199 157 192 466 297 126 247 145

1GSP1 = 5´ RACE out primer; NGSP1 = 5´ RACE nested primer; GSP2 = 3´ RACE out primer; NGSP2 = 3´ RACE nested primer; CDK1, 2, 4 = Cyclindependent kinase 1, 2, 4; PCNA = Proliferation Cell Nuclear Antigen.

fection of Bmi1-pcDNA3.1 was performed according to the instructions for Lipofectamin 2000 (Invitrogen). At 6 h after transfection, cells were digested using trypsin and reseeded at a ratio of 1:10. G418 (400 µg/mL, tested previously before experiment) was then added to the medium. After 7 d, the G418 levels were decreased to 200 µg/mL, when the cells of the nontransfected group had all died. Fourteen days later, the surviving clones were picked and analyzed, and the positive clones were collected. In the following experiments, cells strain transfected with Bmi1 were named the recombinant group (RG) and the nontransfected IPEC-1 cell strain was named the control group (CG), respectively. Proliferation Activity Analysis The MTT Assay. Cells from RG and CG were cultured in a 96-well plate at a density of 4 × 103 cells/mL in GM, as previously described (Wagner et al., 1999; Yang et al., 2008), with minor changes. Twenty microliters of MTT (5 mg/mL; Sigma, St Louis, MO) solution was added to each well and incubated for 4 h. The microtiter plates were centrifuged at 1,400 × g and 4°C for 5 min, and the untransformed MTT solution was carefully re-

moved using an Eppendorf pipette. To each well, 200 mL of DMSO working solution (180 mL DMSO with 20 mL 1 mol/L HCl) was added, and the optical density (OD) of the yellow reaction product was evaluated in an ELISA reader at a wavelength of 490 nm. At least 3 independent experiments were performed to verify the results. EdU Incorporation Assay. Cells from RG and CG were cultured at a density of 4 × 103 cells/mL as previously described (Chehrehasa et al., 2009). To each well, 100 μL of EdU medium (50 μmol/L) was added at 48 h and incubated for 2 h. The cells were then fixed with 4% paraform for 30 min and treated using the Cell-Light EdU Apollo 567 DNA In Vitro Kit (RiboBio, Guangzhou, China) according to the manufacturer’s instructions, as previously described (Salic and Mitchison, 2008). Finally, the newly synthesized DNA incorporated in the EdU and the nuclei of all the cells was labeled, and the cells were photographed. The proliferation ability was assessed by the ratio of EdU+ cells (newly synthesized) to Hoechst 33,342+ cells (all cells). At least 3 independent experiments were performed to verify the results. Flow Cytometry. Cells were cultured in a 6-well plate at a density of 1 × 105 cells/mL in GM and collected at 48, 72, and 96 h after seeding, as previously de-

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scribed (Kang and Alvarado, 2009), with minor changes. Briefly, the cells were fixed with 4 mL cold ethanol and stored at –20°C in fixation buffer until the samples were ready for analysis. The fixed cells were then centrifuged at 200 × g and 4°C for 10 min, resuspended in 1 mL PBS, treated with 100 mL of 200 mg/mL DNase-free RNaseA, and incubated at 37°C for 30 min. Finally, the cells were treated with 100 mL of 1 mg/mL propidium iodide (light sensitive), incubated at room temperature for 5–10 min, and subjected to flow cytometry using BD accuri C6 FACScan (Accuri Cytometers, Ann Arbor, MI) Quantitative Real-Time Polymerase Chain Reaction (qRT-PCR) The mRNA abundance (n = 6) was determined by real-time RT-PCR using a Stratagene MxPro 3005P apparatus (Agilent Technologies, Santa Clara, CA) and the SYBR Green Real-Time PCR Master Mix (TOYOBO, Tokyo, Japan). Primer pairs were designed specifically for each gene using Primer 5.0 software. Melting curve analysis was conducted to confirm the specificity of each product, and the sizes of the products were verified on ethidium bromide-stained 1.0% agarose gels in Tris acetate-EDTA buffer. The relative mRNA expression levels were calculated using 2-ΔCt [ΔCt = Ct of the target gene − Ct of housekeeping gene(18S rRNA)]. The realtime PCR analysis of each sample was repeated 3 times. The primer sequences are provided in Table 1. Western Blot Assay The cells were homogenized in lysis buffer (50  mmol/mL Tris-HCl, pH 7.5, 150 mmol/mL NaCl, 1 mmol/mL EDTA, 0.5% Triton X-100, 1 mmol/mL phenylmethanesulfonyl fluoride (PMSF), 1 mmol/ mL Na3VO4) containing complete protease inhibitor (Invitrogen). The homogenates were centrifuged at 14,000 × g and 4°C for 15 min, and the protein concentrations of the supernatants were determined using a BCA Protein Assay Reagent Kit (Pierce, Rockford, IL). The protein samples were boiled for 5 min and were then subjected to 12% SDS gel electrophoresis. The proteins were separated by electrophoresis at 80 V for 20 min and 120 V for 100 min using Tris-glycine running buffer (0.025 mol/L Tris base, 0.192 mol/L glycine, and 0.1% SDS, pH 8.3). Prestained molecular weight markers (Invitrogen) were used to determine the molecular weight of the proteins. The proteins were subsequently electrotransferred onto polyvinylidene difluoride (PVDF) membranes using a transfer buffer that contained 25 mmol/L Tris base, 192 mmol/L glycine, and 10% methanol, pH 8.1 to 8.3. The membranes were blocked for 1 h in 5% BSA and TBS buf-

fer (20 mmol/L Tris and 500 mmol/L NaCl, pH 7.6) at room temperature. The membranes were incubated overnight at 4°C with 1:300 goat polyclonal antibody against human Bmi1 (Santa Cruz Biotechnology, Santa Cruz, CA) and 1:1,000 mouse monoclonal antibody against β-actin (Santa Cruz Biotechnology). All of the dilutions were made in TBST buffer (TBS buffer with 0.1% Tween-20). The membranes were washed 6 times for 5 min each with TBST buffer, followed by incubation for 1 h with horseradish peroxidase-labeled antigoat IgG (Boisynthesis Biotechnology, Beijing, China) or antimouse IgG (Bioworld Technology, Inc., MN, USA). The proteins were visualized using the BeyoECL Plus chemiluminescence detection kit (Beyotime Institute of Biotechnology, Beyotime, Shanghai, China). Enhanced chemiluminescence (ECL) signals were scanned using a FluorChem M apparatus (ProteinSimple, Inc., Santa Clara, CA). The density of the bands was analyzed using Image Analysis Software (Tanon, Shanghai, China). Statistical Analysis Data are expressed as the mean ± SEM. Student’s t test was conducted to determine differences between 2 groups using SAS (Version 9.2; SAS Inst. Inc., Cary, NC). P values < 0.05 indicate significance. RESULTS Amplification of the Full-Length Pig Bmi1 cDNA Based on the conserved human and mouse sequences, 2 overlapping fragments of 1,087 bp and 1,495 bp were obtained (Fig. 1A). A 321 bp-length 5´ and a 613 bp-length 3´ end sequence were also obtained using RACE (Fig. 1B and 1C). After the alignment and ligation of these sequences, the complete pig Bmi1 cDNA was achieved. The sequence is 3,193 bp in length and consists of a 981 bp ORF, a 256 bp 5´ untranslated region (UTR), and a 1,956 bp 3´ UTR. The homology of the pig Bmi1 CDS with those of the human sequence is 96%, while the homology of the protein with the human Bmi1 is 99% (Fig. 2). The analysis of the pig Bmi1 coded protein indicated that it contains no signal peptides or transmembrane regions; however, the protein contains several zinc finger motifs in the N-terminal region, and the bioinformatics data suggest that the Bmi1 protein is most likely located in the nuclei (69.6%) or the cytoplasm (13.0%). Identification of the Eukaryotic Expression Vector Several clones of DH5α bacteria were selected and amplified for the extraction of plasmids. The M13

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Figure 1. (A, B, C) Cloning of the pig Bmi1 cDNA and (D, E) identification of the recombinant plasmid Bmi1-pcDNA3.1. M: DNA Marker 2000; M1: DNA Marker 10000; Lane 1, 2: partial sequence 1; Lane 3: partial sequence 2 from conserved region; Lane 5: 5´ RACE; Lane 6: 3´ RACE; Lane 4, 7: NUP control; Lane 8, 9: PCR identification of the recombinant plasmid Bmi1-pcDNA3.1; Lane 10, 11: enzyme digesting identification by Kpn1 and Xho1 of the recombinant plasmid Bmi1-pcDNA3.1.

primer was used for PCR identification (Fig. 1D); the Kpn1 and Xho1 enzymes were used for enzyme digesting identifying (Fig. 1E), and sequencing identification was also conducted. All of the data confirmed that the pig Bmi1 had successfully inserted into the pcDNA3.1 vector, creating Bmi1-pcDNA3.1. Expression of Bmi1 mRNA and Protein Cells of both the RG and CG were cultured. Compared with the CG, levels of the Bmi1 mRNA and protein both increased significantly in the RG (P < 0.05; Fig. 3), which means cell strains stably transfected with Bmi1-pcDNA3.1 were achieved successfully.

The Difference in Proliferation Potential after Transfection To investigate the influence of Bmi1 on IPEC-1 cell proliferation, the MTT assay was conducted. Our results indicated that when the different time points were analyzed separately, no significant differences in the number of viable cells were found at 24 h (P > 0.05). From 48 to 120 h after seeding, however, a significantly greater number of viable cells was found in the RG than in the CG (P < 0.05; Fig. 4A). To verify these results, an EdU incorporation experiment was conducted at 48 h, which provided further evidence that pig Bmi1 greatly improved the proliferation ability of IPEC-1 cells (Fig. 4B and 4C).

Figure 2. Comparison of the human and pig Bmi1 protein sequences. The pig Bmi1 protein was predicted from the cloned nucleotide sequence using DNASTAR (www.dnastar.com), and the comparison was conducted using the same software. * means the same amino acid residue.

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Figure 3. Bmi1 overexpression in IPEC-1. Identification of Bmi1 (A) mRNA abundance (n = 6) and (B, C) protein level (n = 3), Lane 1, 2, 3: protein from RG; Lane 4, 5, 6: protein from CG. CG: control group; RG: recombinant group. The results are representative of 3 separate experiments. The bars are means ± SEM. * indicates a significant difference (P < 0.05).

Analysis of Cell Cycle Distribution To further elucidate these proliferation differences, the cells were collected at 48, 72, and 96 h after seeding and tested using flow cytometry. The DNA content distributions of the cells obtained from both the RG and CG consisted of two predominant peaks, corresponding to cells in the G1 and G2 cell cycle phases. The intermediate region between these peaks indicates cells are in the S phase (Fig. 5A and 5B). From these results (Fig. 5C, 5D, and 5E), we observed a decrease in the percentage of cells in the G1 and S phases and an increase in the percentage of cells in the G2 phase for the RG. Although no significant differences were detected for any cell cycle phase at 48 h, at 72 and 96 h after seeding, the percentage of cells in the G1 phase

Figure 4. Assessment of cell proliferation ability after transfection. The cells of the CG and RG were cultured in GM. Cell proliferation ability was (A) assessed using the MTT (n = 20) assay at 24 h, 48 h, 72 h, 96 h, and 120 h and (B, C) confirmed using the EdU incorporation method at 48 h (n = 20). The EdU results were reported as the ratio of newly synthesized cells (EdU+) to total cells (Hoechst33342+). CG: control group; RG: recombinant group. The results are representative of 3 separate experiments. The bars are means ± SEM. * indicates a significant difference (P < 0.05). See online version for figure in color.

in the RG was significantly lower (P < 0.05), while the percentage of cells in the G2 phase was significantly higher (P < 0.05) than in CG. There was no difference in the percentage of cells in the S phase at 48 and 96 h, but the percentage was significantly lower (P < 0.05) in the RG at 72 h compared with the CG. Expression Levels of Cell Cycle-Related Genes The mRNA levels of some cyclins, cyclin-dependent kinases (CDK) and cyclin-dependent kinase inhibitor (CDKI) genes were tested at 48 and 72 h after seeding in the RG and CG. As shown in Fig. 6, the mRNA levels of Cyclin B, Cyclin D1, Cyclin D2,

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Figure 5. Analysis of cell cycle distribution. (A, B) The cell cycle was determined by counting all cells in the sample and plotting their respective DNA contents using flow cytometry. The cells of the CG and RG were cultured in GM. The cell cycle was examined at (C) 48 h, (D) 72 h, and (E) 96 h (n = 6). The statistical analysis of the percentages of cells in the G1, S, and G2 phases at the 3 time points is reported. CG: control group; RG: recombinant group. The results are representative of 3 separate experiments. Bars are means ± SEM. * indicates a significant difference (P < 0.05). See online version for figure in color.

PCNA, CDK1, CDK2, and CDK4 in the RG were significantly higher (P < 0.05) than those in the CG. However, the mRNA levels of Cyclin A2 and p21 exhibited little change at 72 h; what’s more, the change trends at 48 h (data not shown) were the same as at 72 h.

DISCUSSION Currently, cell-sorting technology is the only technique able to isolate IESC. In this study, the pig Bmi1 full-length cDNA was cloned, which will facilitate the future isolation of IESC populations. The bioinformatics analysis of the present work indicated that Bmi1 contains

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Figure 6. Relative mRNA abundance of cell cycle-related genes (n = 6) in cells of the RG and CG at 72 h after seeding. The relative mRNA abundances of cell cycle-related genes were tested. CG: control group; RG: recombinant group. The results are representative of 3 separate experiments. Bars are means ± SEM. * indicates a significant difference (P < 0.05).

no transmembrane regions, suggesting that the sorting of Bmi1+ IESC populations with FACS or magnetic-activated cell sorting (MACS) based on its expression levels is not suitable. Because the production of transgenic pigs is resource-intensive, more traditional strategies for Bmi1+ IESC isolation should be developed. For example (Cammareri et al., 2008), a fluorescence vector with a Bmi1 promoter can be constructed and transfected into crypt cells to aid the recognition of Bmi1+ IESC. To optimize the in vitro culture conditions of pig IESCs, we also investigated the function of Bmi1 in the proliferation of the intestinal epithelial cells. An undifferentiated IPEC-1 cell line was chosen as a model, which was derived from the small intestine of an unsuckled newborn piglet less than 12 h old and has undergone more than 100 passages without any significant changes in cell growth (Gonzalez-Vallina et al., 1996; Lu et al., 2002; Tan et al., 2010). Previous work in our lab has demonstrated that the pcDNA3.1 vector has no effect on IPEC-1 proliferation (data not shown); in the present work, we therefore developed a stably transfected IPEC-1 cell strain with the overexpression of Bmi1-pcDNA3.1 as the RG and designated untransfected IPEC-1 cells as the CG. EdU (5-ethynyl-2´-deoxyuridine) is an analogue of thymidylic acid (T) that can be incorporated into the replicating DNA when cells proliferated, thus the newly synthesized cells can be monitored by the reaction of EdU with fluorescence labeled dyes. What’s more, EdU is too small when compared with BrdU that it can hardly exert any influence to the structure and function of the DNA and the cells. Despite this, the high concentration of EdU and longtime incubation with EdU are not suggested due to the possibility of cytotoxicity. In this experiment, the EdU incorporation method by commercial kit was introduced to evaluate the cell proliferation ability and the method was proved to be effective.

In this study, the proliferative ability of the cells was measured using an MTT assay over a short period of 5 d. Although the difference was not significant at 24 h after seeding, a markedly higher OD value was found in the RG than in the CG for all other time points. The EdU incorporation experiment at 48 h produced a similar result. Furthermore, the PCNA mRNA abundance was also significantly increased after the overexpression of Bmi1, indicating that Bmi1 can improve the proliferation rate of IPEC-1. Similar results have also been observed in neural stem/progenitor cells (He et al., 2009), hematopoietic stem cells (Iwama et al., 2004), and mammary stem cells (Pietersen et al., 2008), although not previously in intestinal cells. Therefore, we conclude that several growth-improvement factors used in current IESC culture conditions may be nonessential in the in vitro culture of pig Bmi1+ IESC cells (Gracz et al., 2012); however, further studies remain necessary. A sustained proliferative response requires the coordination of both cell cycle progression and cell growth (Neufeld and Edgar, 1998; Polymenis and Schmidt, 1999). To remain constant under proliferative conditions, both DNA content and cell mass must double during the course of each cell division cycle. In this study, the IPEC1 cells from the RG and CG were tested for DNA content after 48, 72, and 96 h of proliferation to determine the distribution of cells in the G1, S, and G2 phases of the cell cycle. Our results revealed that after overexpression, the percentage of cells in the G1 phase for the RG was significantly lower than that observed in the CG at 72 and 96 h, while the percentage of cells in the G2 phase was significantly higher. Interestingly, the percentage of cells in the S phase for the RG was also significantly lower than for the CG, implying that pig Bmi1 cannot only improve the rate of cells passing from the G1 phase to the S phase for IPEC-1 but that it can also accelerate the DNA

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duplication processes of the S phase. Previous studies have indicated that Cyclin D can facilitate the cell cycle transition from the G1 to the S phase (Coqueret, 2002; Fu et al., 2004); In all eukaryotic cells, Cyclin A can either arrest the cell cycle transition from the G1 phase to the S phase or promote the transition from the G2 phase to the M phase (Pagano et al., 1992; Garrett, 2001). In the present study, the mRNA levels of Cyclin D1 and Cyclin D2 were both significantly higher in the RG than in the CG, but the Cyclin A2 levels showed little change. The upregulation of Cyclin D1 and Cyclin D2 mRNA abundance, in combination with the reduction of cells in the G1 phase in the RG, indicates that pig Bmi1 can indeed promote the progression of IPEC-1 from the G1 to the S phase and arrest the G2/M phase. Moreover, several positive regulator genes of the cell cycle were upregulated, with the differences in Cyclin B reaching statistical significance, which is also very important to the G2/M transition, as well as some cell cycle cyclin-dependent kinases, including CDK1, CDK2, and CDK4; these results further indicate that the pig Bmi1 increased DNA replication. The expression of the negative regulators of the cell cycle, particularly CDKN1A (p21), was not changed, and CDKN2A (p16/p19) was barely detectable in either of the IPEC-1 strains (data not shown); these results are supported by Hourinaz Behesti’s study on Math1-Bmi1 mice cerebella (Behesti et al., 2013). Although little research exists concerning the effects of Bmi1 on intestinal cells, previous data on other stem cells or progenitor cells had suggested that Bmi1 regulates cell proliferation and senescence through the ink4a locus (p16 and p19; Jacobs et al., 1999; Fasano et al., 2009; Yadirgi et al., 2011). However, Dimri et al. (2002) observed that p14ARF is not downregulated in mammary epithelial cells overexpressing human Bmi1. It is therefore controversial whether Bmi1 operates solely through the ink4a locus, and further studies are required. Several previous studies have also suggested that Bmi1 plays a critical role through p21-Rb pathway (Fasano et al., 2007, 2009; Subkhankulova et al., 2010), and it has been reported that Bmi1 only represses the Ink4a/Arf locus, rather than the p21 gene, in the regulation of the self-renewal, proliferation, and senescence of human fetal neural stem cells in vitro (Wang et al., 2010). Our data suggests that pig Bmi1 does not promote IPEC-1 proliferation through the p21 pathway, and its effects on the ink4a locus require further research. Sangiorgi (2008) observed that the ablation of Bmi1+ cells using a Rosa26 conditional allele, expressing the diphtheria toxin, led to crypt loss, indicating a close relationship between Bmi1 and the intestine; the results of Reinisch (2006) suggest a role for Bmi1 in the tumorigenesis of gastrointestinal cancer. Therefore, the pig Bmi1 may also be involved in pig intestine pathology in addition to its role in the isolation of pig IESC.

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We surmised that Bmi1 may likewise be involved in the atrophy or hypertrophy of villus-crypt, however, more research is required. Li (2010) also observed that the expression level of Bmi1 oncoprotein is associated with progression and prognosis in colon cancer, which reminds us of the theory that intestinal cancers may derive from stem cells. Our results that Bmi1 functions through cell cycle regulation may provide new thoughts about how intestinal cancers come to be, how to search the therapy targets or prevention methods, and even how to reconstruct the villus-crypt axis. In conclusion, we are the first to clone the pig Bmi1 full-length cDNA (GI: JQ012762). From our results, we hypothesize that the Bmi1 promoter vector, along with EGFP, can be used to assist the isolation of Bmi1+ IESC via FACS. Furthermore, pig Bmi1 can promote the G1/S transition and DNA replication in the S phase without repressing p21, thereby accelerating the cell cycle process and IPEC-1 proliferation. LITERATURE CITED Barker, N., J. H. van Es, J. Kuipers, P. Kujala, M. van den Born, M. Cozijnsen, A. Haegebarth, J. Korving, H. Begthel, and P. J. Peters. 2007. Identification of stem cells in small intestine and colon by marker gene Lgr5. Nature. 449:1003–1007. Behesti, H., H. Bhagat, A. M. Dubuc, M. D. Taylor, and S. Marino. 2013. Bmi1 overexpression in the cerebellar granule cell lineage of mice affects cell proliferation and survival without initiating medulloblastoma formation. Dis. Model. Mech.. 6:49–63. Cammareri, P., Y. Lombardo, M. G. Francipane, S. Bonventre, M. Todaro, and G. Stassi. 2008. Isolation and culture of colon cancer stem cells. Methods Cell Biol. 86:311–324. Chehrehasa, F., A. C. Meedeniya, P. Dwyer, G. Abrahamsen, and A. Mackay-Sim. 2009. EdU, a new thymidine analogue for labelling proliferating cells in the nervous system. J. Neurosci. Methods. 177:122–130. Coqueret, O. 2002. Linking cyclins to transcriptional control. Gene. 299:35–55. Dimri, G. P., J. Martinez, J. J. Jacobs, P. Keblusek, K. Itahana, M. van Lohuizen, J. Campisi, D. E. Wazer, and V. Band. 2002. The Bmi-1 oncogene induces telomerase activity and immortalizes human mammary epithelial cells. Cancer Res. 62:4736–4745. Fasano, C. A., J. T. Dimos, N. B. Ivanova, N. Lowry, I. R. Lemischka, and S. Temple. 2007. shRNA knockdown of Bmi-1 reveals a critical role for p21-Rb pathway in NSC self-renewal during development. Cell Stem Cell. 1:87–99. Fasano, C. A., T. N. Phoenix, E. Kokovay, N. Lowry, Y. Elkabetz, J. T. Dimos, I. R. Lemischka, L. Studer, and S. Temple. 2009. Bmi-1 cooperates with Foxg1 to maintain neural stem cell selfrenewal in the forebrain. Genes Dev. 23:561–574. Fu, M., C. Wang, Z. Li, T. Sakamaki, and R. G. Pestell. 2004. Minireview: Cyclin D1: Normal and abnormal functions. Endocrinology. 145:5439–5447. Garrett, M. D. 2001. Cell cycle control and cancer. Curr. Sci. 81:515–522. Gonzalez-Vallina, R., H. Wang, R. Zhan, H. M. Berschneider, R. M. Lee, N. O. Davidson, and D. D. Black. 1996. Lipoprotein and apolipoprotein secretion by a newborn piglet intestinal cell line (IPEC-1). Am. J. Physiol. Gastrointest. Liver Physiol. 271:249–259.

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