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Elevated expression of cyclooxygenase-2 (COX-2) and one of its downstream enzymatic products, prostaglandin E2 (PGE2) have been directly linked to colorectal carcinogenesis in a number of ways. Among which, PGE2 promotes cell proliferation, cell cycle progression, and thus tumor growth. All of the mechanism(s) by which PGE2 signaling regulates cell growth are not completely understood. Here, we demonstrate that PGE2 treatment induces human enhancer of filamentation 1 (HEF1) expression and its link with cell cycle machinery in colorectal cancer cells. PGE2 rapidly stimulated the expression of HEF1 mRNA and protein in colorectal cancer cells. Both PGE2 treatment and HEF1 overexpression resulted in similar effects on cell proliferation, cell cycle progression, and tumor growth. Moreover, knockdown of HEF1 using shRNA suppressed PGE2-driven cell proliferation and cell cycle progression. Cell cycle alterations involved HEF1 fragmentation as well as co-distribution of HEF1 and Aurora A along spindle asters during cell division. Furthermore, HEF1 co-immunoprecipitated with and activated Aurora A. Intriguingly, HEF1 expression was increased in 50% of human colorectal cancers compared with expression in paired normal tissue. These data suggest that PGE2 induces HEF1 expression, which in turn promotes cell cycle progression through its interaction and activation of Aurora A. Clearly, HEF1 is a downstream mediator of PGE2 action during colorectal carcinogenesis.
Prostaglandin E2 (PGE2) is the most abundant prostaglandin found in colorectal cancer tissue and is a downstream product of an inducible enzyme cyclooxygenase-2 (COX-2). A significant increase in COX-2 gene expression has been linked to colorectal carcinogenesis (1). Inhibition of COX-2 activity by nonsteroidal anti-inflammatory drugs, particularly selective COX-2 inhibitors such as celecoxib, can suppress colorectal adenoma recurrence (2). PGE2 is the primary mediator of COX-2 in promoting cancer progression. Many studies have shown that PGE2 promotes cell proliferation and tumor growth in colorectal cancer (3–6). Several pathways or molecules have been implicated as the downstream mediators of PGE2 on promoting cell proliferation and tumor growth, which include PI3K/Akt pathway, Ras/MAPK pathway, β-catenin/TCF-4, and PPARδ (3–6). Obviously, more work is needed to reveal the details by which PGE2 signaling affects cell proliferation.
HEF1 is a scaffold protein that encodes multiple protein interaction domains. It has been implicated in numerous biological activities including mediating integrin-dependent signals at focal adhesions. HEF1 is preferentially expressed in epithelial cells and lymphocytes and undergoes substantial regulation during progression through the cell cycle (7). The regulation of HEF1 expression is not fully understood. TGF-β and all-trans retinoic acid have been shown to upregulate HEF1 expression at the transcriptional level (8–10). Serum can also increase HEF1 expression (11). HEF1 localizes not only at focal adhesions but also at the centrosomes and mitotic spindles following stimulation by intrinsic and extrinsic cues (7). Studies have demonstrated that HEF1 becomes fragmented and moves from the cytoplasm and focal adhesions during mitosis to centrosomes and mitotic spindles, where it interacts with and activates Aurora A to promote cell cycle progression (12–14).
Since both PGE2 and HEF1 have been implicated in promoting cell proliferation or cell cycle progression in certain contexts, we hypothesized that PGE2 induces HEF1 expression to promote cell cycle progression and growth of colorectal cancers. In the present study, we demonstrate that HEF1 functions as an important downstream target of PGE2, which promotes the proliferation of colorectal cancer cells.
PGE2 was obtained from Cayman Chemical (Ann Arbor, MI). Cell proliferation reagent WST-1 was purchased from Roche Applied Science (Indianapolis, IN). Antibody to HEF1 (2B11) was purchased from Santa Cruz Biotechnology (Santa Cruz, CA). AuroraA and phospho-Aurora A antibodies were purchased from Cell Signaling Technology (Danvers, MA). Ki-67 antibody was purchased from DAKO (Carpinteria, CA). β-Actin antibody was obtained from Sigma-Aldrich (St. Louis, MO). Retroviral HEF1 expression vector (LZRS-Ires-HEF1) and control vector (LZRS-Ires-GFP) were kind gifts from Dr. Lynda Chin (Dana-Farber Cancer Institute, Boston, MA). shRNA vector targeting HEF1 (pGIPZ-shHEF1) and control nonsilencing vector (pGIPZ-shCon) were purchased from Open Biosystems (Huntsville, AL). Lentivirus packaging vectors pMD2.G and psPAX2 (Addgene plasmids 12259 and 12260) were purchased from Addgene (Cambridge, MA). Human colorectal tumor specimens were obtained as described previously (15).
LS-174T, LoVo, Caco-2, and DLD-1 cells were purchased from the American Type Culture Collection (Manassas, VA). HCA-7 cells were a generous gift from Susan Kirkland. These cells were maintained in McCoy’s 5A medium containing 10% fetal bovine serum in a 5% CO2 atmosphere.
LS-174T cells were cultured in serum-free media for 48h and treated with or without 1 µM PGE2 for 0, 2, 4, 8, and 24 h with three replicates per group. Total RNAs were extracted with Trizol (Invitrogen, Carlsbad, Ca) and subjected to microarray analysis using Affymetrix Human Genome U133 Plus 2.0 Array. Resulting data were analyzed by GeneSpring GX and GeneTraffic softwares. Those genes with significant change in expression (at least two-fold change) as compared with control (0 h) were selected for further study.
Quantitative PCR (Q-PCR) was performed as described previously (15). Primers used were: human Hef1 forward 5'-GATGGGTGTCTCCAGCCTAA-3' and reverse 5'-GGATCTGGTGGGAGTCTTCA-3', human COX-2 forward 5’-CCCTTGGGTGTCAAAGGTAA-3’ and reverse 5’-GCCCTCGCTTATGATCTGTC-3’, and human β-actin forward, 5'-AGAAAATCTGGCACCACACC-3' and reverse, 5'-AGAGGCGTACAGGGATAGCA-3'.
Northern blot analysis was performed as previously described (15). Blot was hybridized in Hybrisol I (InterGen, Burlington, MA) with a 32P-labeled human Hef1 cDNA in coding region. The blot was exposed to film. The dot density on developed film was measured using NIH ImageJ software.
Western blot analysis was performed as described previously (16). HEF1 can be cleaved into smaller fragments (1–405 amino acids, 44kD; The full length of HEF1 is 835 amino acids, 105 kD) under certain conditions (12). The HEF1 antibody used (Santa Cruz Biotechnology, 2B11, which is raised against amino acids 82–398 of human HEF1) can recognize both the full length and the small fragment of HEF1.
Ninety-six well plates were seeded with 3000 cells per well in 0.1 mL of growth medium. After cells were allowed to attach overnight at 37°C, they were washed twice with PBS and then incubated in serum-free medium (SFM) for 2 days. In dose-response studies, cells were treated with different concentrations of PGE2 in SFM for 3 days. Time-point experiments involved a single 1-µM concentration of PGE2. Cell growth was determined by adding 10 µL of WST-1 proliferation reagent in the last 4 h or 20 µL of Brdu Label (Calbiochem, San Diego, Ca) in the last 24 h per well following the assay’s protocol. Absorbance was measured at 450 nm using a SpectraMax M5 microplate reader (Molecular Devices, Sunnyvale, CA).
LZRS-Ires-HEF1 and control LZRS-Ires-GFP retroviral vectors were transfected into Phoenix cells, or pGIPZ-shHEF1 and pGIPZ-shCon along with package vectors psPAX2 and pMD2.G were transfected into 293T cells in 60-mm dishes using Lipofectamine reagent (Invitrogen, Carlsbad CA) according to the manufacturer's protocol. After an overnight incubation, the transfection medium was removed and replaced by 3 mL of growth medium. Culture medium containing virus particles were collected 24 h later and passed through a 0.45-µm filter to remove cell debris. Cells were plated in a 60-mm dish 24 h before they were infected. Medium containing virus was added to cells and repeated one more time with freshly collected virus particles 24 h later. During the viral infection process, the final concentration of polybrene (Sigma, St. Louis, MO) in these experiments was adjusted to 4 µg/mL for Phoenix cells and 8 µg/mL for 293T cells. Puromycin (2 µg/mL) was added for 5 days, or after 3 days of infection, cells were sorted by green fluorescent protein (GFP) positivity to eliminate uninfected cells.
Cells were cultured in SFM for 2 days. Fresh SFM containing 1 µM PGE2 was added for 24 h. Cells were collected by trypsinization and fixed with 100% ethanol on ice for 20 min. After centrifugation (500×g for 3 min), cells were stained with propidium iodide (PI; Invitrogen) in 1 mL of staining solution containing 50 µg/mL PI, 100 µg/mL RNase A (DNase-free), and 70% ethanol prepared in PBS. After a 30-min incubation, the stained cells were subjected to fluorescence-activated cell sorting (FACS) for cell cycle analysis.
Immunofluorescence studies involved plating cells on coverslips in 10% serum medium. After overnight growth, cells were washed twice with PBS, fed with SFM, and incubated at 37°C for 2 more days. After treatment with 1 µM PGE2 in fresh SFM for 24 h, cells were fixed with 4% paraformaldehyde for 15 min at room temperature, permeabilized with methanol at −20°C for 10 min, blocked with 3% BSA, and incubated with antibodies using standard protocols. Primary antibodies included mouse monoclonal antibody anti-HEF1 (2B11) (Santa Cruz Biotechnology, 1:100) and rabbit polyclonal antibody anti-Aurora A (Cell Signaling; 1:100). Secondary antibodies anti-mouse Alexa-488 and anti-rabbit Alexa-594 (Invitrogen, 1:100,000) were applied along with DAPI (Invitrogen, 1:100,000) for 1 h at room temperature.
General procedures for immunohistochemical staining were followed. Briefly, after dewaxing and rehydrating, slides were boiled in citrate buffer using an EZRetriever microwave (Biogenex, 98°C for 5 min). Blocking was performed with 5% normal horse serum and 1% normal goat serum in PBS. Primary antibodies were incubated at 4°C overnight. Antibodies used were anti-HEF1 (2B11, 1:200, Santa Cruz Biotechnology) and anti-Ki-67 (1:50, DAKO, Carpinteria, CA). Secondary antibody was incubated at room temperature for 1 h followed by diaminobenzedine chromogen (Vector Laboratories) and haematoxylin counter staining.
Co-immunoprecipitation analysis was performed as described previously (16) using anti-HEF1 (2B11; Santa Cruz Biotechnology) or anti-Aurora A antibodies (Danvers, MA).
All mice were housed and treated in accordance with protocols approved by the Institutional Animal Care and Use Committee at The University of Texas M. D. Anderson Cancer Center. LS-174T cells (5×105) selected for the stable expression of HEF1 (LS-174T/HEF1) or control GFP (LS-174T/GFP) were injected subcutaneously into the flanks of nude mice. Three weeks after injection, mice were euthanized using CO2 asphyxiation, necropsies were performed to remove tumors, and measurements were taken of tumor weight and size.
Each experiment was performed at least 3 times, and data are presented as the mean ± S.E. Statistical significance was determined using a Student's t test, one-factor ANOVA, or two-factor ANOVA wherever applied. P values < 0.05 were considered statistically significant.
To identify potential target genes regulated by PGE2 in colorectal cancer, LS-174T cells were treated with or without PGE2 and subjected to microarray analysis, which demonstrated a 4-fold increase in HEF1 mRNA expression. To confirm the microarray results, Q-PCR and Western blot analysis revealed that PGE2 rapidly induced HEF1 expression at both RNA (Fig. 1A) and protein (Fig. 1B, upper panels and supplemental Fig. 1) levels. The induction of HEF1 mRNA expression was significant and persisted for at least 24 h. At the protein level, HEF1 expression peaked between 6 and 12 h. HEF1 expression was also increased in HCA-7 cells (Fig. 1B, lower panel). Other colorectal cancer cell lines that express low basal levels of HEF1 also responded to PGE2 treatment by up-regulating HEF1 protein expression, but not to the same degree as observed in LS-174T and HCA-7 cells (Fig. 1C). Because HEF1 induction was most dramatic in LS-174T cells, subsequent experiments were carried out using this cell line.
PGE2 treatment led to increased cell proliferation in LS-174T and HCA-7 cells in a dose-dependent manner (Fig. 2A). To examine whether HEF1 plays a role in PGE2-induced cell proliferation, HEF1 expression was knocked down using shRNA in LS-174T cells. HEF1 expression decreased about 50% with HEF1 shRNA compared with control shRNA (Fig. 2B, left panel). Although the knockdown did not completely eliminate the expression of HEF1 protein, cell proliferation was significantly reduced (Fig. 2B, right panel). These data suggest that the suppression of HEF1 expression limits the ability of parental LS-174T cells to respond to PGE2-induced cell proliferation.
To further demonstrate that HEF1 mediates the effect of PGE2 on cell proliferation, HEF1 protein was overexpressed in LS-174T/HEF1 cells and compared to LS-174T/GFP cells (Fig. 2C, left panel). In the absence of PGE2, stable HEF1 overexpression in LS-174T/HEF1 cells led to increased cell proliferation compared with GFP alone in control cells (Fig. 2C, middle panel, 0 µM). PGE2 treatment increased the proliferation of both LS-174T/GFP and LS-174T/HEF1 cells in a dose- and time-dependent manner (Fig. 2C, right two panels). Overexpression of HEF1 in another colorectal cancer cell line LoVo also resulted to increased cell proliferation as in LS-174T cells (Supplemental Fig. 2). These data suggest that increased HEF1 expression increases cell proliferation similar to PGE2 treatment (Fig. 2A).
To study the mechanism by which PGE2 increases cell proliferation, we performed FACS analysis with PGE2 treated LS-174T cells. Treatment with PGE2 shifted the number of parental LS-174T cells from the G1 phase into the S/G2 phase of the cell cycle (Fig. 3A). To demonstrate the role of HEF1 in PGE2 induced cell cycle progression, we first examined whether HEF1 also promotes cell cycle progression by using the HEF1 overexpressed LS-174T/HEF1 cells. As expected, overexpression of HEF1 in LS-174T cells shifted the number of cell from the G1 phase into the S/G2 phase (Fig. 3B). Next, we examined whether decrease of HEF1 attenuates the effect of PGE2 on cell cycle progression. Indeed, knockdown of HEF1 minimally effected the cell cycle progression of parental LS-174T cells in SFM but blocked PGE2-induced cell cycle progression (Fig. 3C). These results suggest that HEF1 accelerates and mediates the effect of PGE2 on the progression of cells from the G1 to the S/G2 phase of the cell cycle.
HEF1 has been shown to regulate cell cycle progression by fragmenting and relocalizing to spindle asters and activating Aurora A (11, 12, 19). PGE2 treatment led to increased levels of fragmented HEF1 found in LS-174T cells (Fig. 4A). Further examination of these cells by immunofluorescence showed that a single 1-µM dose of PGE2 increased the number of cells observed in metaphase (Fig. 4B). Immunofluorescence analyses also revealed that HEF1 localized in the cytoplasm of quiescent LS-174T cells (Fig. 4C, upper panel). In contrast, close examination of mitotic cells indicated increased instances of metaphase or anaphase along with elevated levels of HEF1 (green) that had redistributed to the area surrounding spindle asters (Fig. 4C). Aurora A (red) staining showed a pattern similar to that of HEF1. When these red and green staining patterns were combined into merged images, the presence of yellow areas indicated that these two proteins were colocalized in LS-174T cells (Fig. 4C, lower panel). Co-immunoprecipitation analysis results indicated a direct interaction between HEF1 and Aurora A and that PGE2 treatment increased Aurora A phosphorylation in LS-174T cells (Fig. 4D, left panel). These data suggested that PGE2 stimulated HEF1 fragmentation and relocalization to spindle asters and the activation of Aurora A to enhance cell cycle progression.
LS-174T/HEF1 or LS-174T/GFP cells were injected into the flanks of nude mice to validate whether our in vitro observations would translate into similar effects in vivo. The mice were euthanized 3 weeks after injection, and tumor weight and size were measured. Cells expressing HEF1 resulted in a significant increase in both tumor weight and size compared with those expressing the GFP control (Fig. 5A). IHC staining revealed that HEF1 and Ki-67 stained the same population of cells in those tumors (Fig. 5B). Furthermore, HEF1 interacted with Aurora A in the tumors formed from LS-174T/HEF1 cells (Fig. 5C). These results indicates that HEF1 also interacts with Aurora A to promote cell proliferation and tumor growth in vivo.
Given that COX-2 overexpression is observed in approximately 70%–80% of colorectal cancers (20), we hypothesized that HEF1 expression is up-regulated in colorectal cancer as well. Q-PCR analysis revealed that HEF1 expression increased in at least 7 of 15 tumor specimens compared with adjacent normal mucosa (Fig. 6A and supplemental Fig. 3A). Meanwhile, higher HEF1 expression (tumor:normal ratio > 1) was found in 15 of 30 tumor tissues using Northern blot analysis (Fig. 6B). Overall, HEF1 expression increased in about 50% of colorectal cancers compared with normal tissues and correlated with COX-2 expression (Supplemental Fig. 2B, r = 0.6235, P = 0.013).
Elevated expression of COX-2 and concomitant overproduction of PGE2 have been directly linked to colorectal carcinogenesis. COX-2 inhibitors can suppress colorectal carcinogenesis, but long-term use can cause adverse effects in a subset of patients (1). Our research has focused on identifying and characterizing the effector molecules downstream of PGE2-driven signaling involved in colorectal carcinogenesis with the ultimate goal of developing approaches that have the same benefit, but result in fewer side effects. Here we report that PGE2 rapidly stimulated the expression of HEF1 in colorectal cancer cells. Both PGE2 treatment and HEF1 expression elicited similar effects on cell proliferation and tumor growth. The alteration of HEF1 levels by PGE2 influenced cell proliferation and cell cycle progression, suggesting that HEF1 can act as a downstream effector of PGE2 on these biological functions.
Reports from our laboratory, as well as others, show that PGE2 affects cell proliferation via activating MAPK or PI3K/Akt pathways (3, 6, 21, 22). How these pathways link to cell cycle and cell division to regulate cell proliferation must be further defined. Here we present the first data indicating that HEF1 can also mediate the effects of PGE2. PGE2 treatment induced HEF1 expression and increased the proliferation of serum-starved LS-174T and HCA-7 cells (Fig. 1B and and2A).2A). Increased HEF1 expression had a similar effect on the proliferation of LS-174T cells not treated with PGE2 (Fig. 2C, middle panel, 0 µM). Moreover, knockdown of HEF1 attenuated the effect of PGE2-enhanced cell proliferation (Fig. 2B). This was even more evident after HEF1 knockdown, which blocked the ability of PGE2 to promote cell cycle progression (Fig. 3C). Together, these results indicate that HEF1 helps mediate the biological effects of PGE2 during cell proliferation and cell cycle progression. Furthermore, mechanistically HEF1 fragmented, relocated, and co-distributed with Aurora A along spindle asters during cell division (Fig. 4A and 4C). HEF1 co-immunoprecipitated with and activated Aurora A (Fig. 4D), which is required for the G2-to-M transition during cell division. Thus, HEF1 bridges the effect of PGE2 on cell proliferation and tumor growth with cell division.
HEF1 protein relocalized from focal adhesions to the mitotic spindle asters in a cell-cycle-regulated manner in MCF-7 cells (14). Increased HEF1 in G2/M phases was accompanied with increased level of fragmentation, and the resulting fragments moved from the cytoplasm or focal adhesions to the centrosomes, where these fragments interacted with and activated molecules required for mitotic progression such as Aurora A (12, 13). These studies have clearly demonstrated the involvement of HEF1 in the regulation of cell cycle progression by using chemically synchronized MCF-7 cells and serum stimulation. In the present study, we tried to synchronize LS-174T cells with serum starvation. However, LS-174T cells were able to grow and divide in the absence of serum (data not shown). Therefore, the cell population we used was a mixed group of cells in different stage of the cell cycle. This could be one of the reasons why only a small group of cells responded in cell cycle progression to PGE2 treatment or HEF1 alteration (Fig. 3). Here we demonstrated that a single bioactive lipid PGE2 has the same effect as serum on HEF1 induction, fragmentation, and relocalization and Aurora A activation.
In conclusion, we provide evidence that HEF1 is induced by PGE2 and mediates its effects on promoting cell proliferation by enhancing cell cycle progression. Given that HEF1 can redistribute between the focal adhesions of migrating cells or the mitotic machinery as cells divide, it may act as a “molecular switch” that causes cells to either migrate or enter cell division. We are currently studying the possible involvement of HEF1 in PGE2-stimulated migration, invasion, and metastasis of colorectal cancer cells. Together, these studies will add to our understanding of how PGE2 regulates cell proliferation, cell cycle progression, cell migration, invasion, and metastasis. The eventual goal is to help identify the possible targets downstream of COX-2 for the prevention and treatment of colorectal cancer.
We thank Drs. Lynda Chin and Minjung Kim (Dana-Farber Cancer Institute) for providing HEF1 expression and control vectors and Dr. Didier Trono (Ecole Polytechnique Fédérale de Lausanne, Switzerland) for the psPAX2 and pMD2.G vectors. This research was supported in part by NCI MERIT award R37 DK47297, NCI P01 CA77839, and The National Colorectal Cancer Research Alliance to R.N.D. and a cancer prevention fellowship supported by the NCI grant R25T CA57730, Robert M. Chamberlain, Ph.D., Principal Investigator and Shine Chang, Ph.D., Co-Principal Investigator (www.CancerPreventionTraining.org).