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Cancer Research 67, 9315-9321, October 1, 2007. doi: 10.1158/0008-5472.CAN-07-1128
© 2007 American Association for Cancer Research
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Cell, Tumor, and Stem Cell Biology

ß-Catenin Regulates Multiple Steps of RNA Metabolism as Revealed by the RNA Aptamer in Colon Cancer Cells

Hee Kyu Lee1, Ho Yoon Kwak1, Jung Hur1, In Ae Kim1, Ji Sun Yang1, Min Woo Park1, Jaehoon Yu2 and Sunjoo Jeong1

1 Department of Molecular Biology, BK21 Graduate Program for RNA Biology, Institute of Nanosensor and Biotechnology, Dankook University and 2 Department of Chemistry and Education, Seoul National University, Seoul, Republic of Korea

Requests for reprints: Sunjoo Jeong, Department of Molecular Biology, Dankook University, Seoul, Republic of Korea. Phone: 82-2-709-2819; Fax: 82-2-793-0176; E-mail: sjsj{at}dankook.ac.kr.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Nuclear ß-catenin forms a transcription complex with TCF-4, which is implicated in colon cancer development and progression. Recently, we and others have shown that ß-catenin could be a regulator of RNA splicing and it also stabilizes the cyclooxygenase-2 (COX-2) mRNA. Here, we further explored the role of ß-catenin in the RNA metabolism in colon cancer cells. To specifically modulate the subcellular functions of ß-catenin, we expressed the RNA aptamer in the form of RNA intramers with unique cellular localizations. The nucleus-expressed RNA intramer proved to be effective in reducing the protein-protein interaction between ß-catenin and TCF-4, thus shown to be a specific regulator of ß-catenin–activated transcription. It could also regulate the alternative splicing of E1A minigene in diverse colon cancer cell lines. In addition, we tested whether ß-catenin could stabilize any other mRNAs and found that cyclin D1 mRNA was also bound and stabilized by ß-catenin. Significantly, the cytoplasm-expressed RNA intramer reverted the ß-catenin–induced COX-2 and cyclin D1 mRNA stabilization. We show here that ß-catenin regulated multiple steps of RNA metabolism in colon cancer cells and might be the protein factor coordinating RNA metabolism. We suggest that the RNA intramers could provide useful ways for inhibiting ß-catenin–mediated transcription and RNA metabolism, which might further enhance the antitumorigenic effects of these molecules in colon cancer cells. [Cancer Res 2007;67(19):9315–20]


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Cancer cell formation and progression are regulated by multiple steps of gene expression, which might exist as an extensive network between transcription and posttranscriptional processes. There are emerging evidences that the various steps of gene expression, from transcription to translation, are precisely coordinated and protein factors might be involved in the coordination of gene expression (1, 2). Therefore, it must be important to identify the functionally and sometimes physically connected protein factors responsible for the individual steps of gene expression that may serve as a coordinator of the multiple processes (3). Because cancer cells tend to show aberrant and altered pattern of RNA metabolism, such protein factors are also likely to play profound role in tumorigenesis and cancer progression.

ß-Catenin is part of a cell adhesion complex and a central component of the developmentally important Wnt pathway regulating cell growth and differentiation during embryonic development and tumorigenesis (4). In the absence of Wnt, most of the ß-catenin in epithelial cells is attached to the plasma membrane where it associates with E-cadherin in adherens junctions. Cytosolic ß-catenin is located in a multiprotein complex consisting of the adenomatous polyposis coli (APC) protein, axin/conductin, and glycogen synthase kinase-3ß (GSK-3ß). Mutations of APC or ß-catenin are frequently found in various types of cancer and lead to transcription of target genes, such as cyclin D1 and c-myc, independent of external Wnt signals (5–7). Therefore, the activation of ß-catenin is an important event leading to tumor development and progression thru the transcriptional activation of many oncogenes.

It is also possible for ß-catenin to play multiple other functions during the processes. ß-Catenin interacts with many proteins, including the sequence-specific DNA-binding transcription factor TCF-4 and other proteins implicated in transcription and chromatin remodeling (8, 9). For example, it was recently reported that ß-catenin bound to some splicing factors and may be involved in regulating RNA splicing in colon cancer cells (10, 11). Moreover, we have shown that ß-catenin directly interacts with RNA (12). It stabilized cyclooxygenase-2 (COX-2) mRNA by interacting with the AU-rich element in the 3'-untranslated region (3'-UTR; ref. 12). ß-Catenin could be the protein factor involved in RNA metabolism by its direct interaction to RNA and multiple interactions to other proteins. However, how ß-catenin is involved in the regulation of transcription and posttranscriptional events is not currently known.

Because the Wnt/ß-catenin pathway is an acknowledged molecular target of cancer therapy (13), we isolated an RNA aptamer for ß-catenin and showed that it had high affinity (Kd = 5 nmol/L) and specificity for ß-catenin in vitro and in the cells (11). We also showed it to be an effective inhibitor of ß-catenin–mediated tumorigenesis (11). ß-Catenin–specific small hairpin RNA (shRNA) was also an effective inhibitor of tumor growth (14). However, the exact mechanism of these antitumorigenic RNA inhibitors was not well understood.

As a first step to decipher whether ß-catenin could be a factor involved in the multiple steps of gene expression, we used the ß-catenin–specific RNA intramers and shRNA. Because RNA intramers are effective in vivo (15–17) and specific in their action, the use of various types of intramer can also yield insights into the roles of their target proteins. Here, we have expressed the ß-catenin–specific RNA intramers under two different promoters that allow them to be localized in the nucleus and cytoplasm, respectively, and have further analyzed the roles of ß-catenin in RNA metabolism. Most significantly, the RNA intramer affected ß-catenin–mediated alternative splicing of E1A minigene mRNA as well as the stability of COX-2 and cyclin D1 mRNA. These results led us to propose that ß-catenin is involved in multiple steps of gene expression and plays a key role in coordinating RNA metabolism.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell culture, plasmids, and luciferase assay. Human HCT116 colorectal carcinoma, HT-29 colorectal adenocarcinoma, embryonic kidney 293T, and mouse NIH3T3 cells (American Type Culture Collection) were cultured in DMEM with 10% fetal bovine serum (FBS). Human LoVo colorectal adenocarcinoma cells were cultured in RPMI 1640 with 10% FBS. A retroviral vector expressing human S37A/ß-catenin was previously described (13). The pU6-Vector and pDHFR was previously described (11, 17–19). TCF-responsive luciferase reporters, pGL3-OT (wild-type), and pGL3-OF (mutant) were from Dr. Shivdasani (Dana-Farber Cancer Institute, Boston, MA). To make pSUPER-ß-catenin, specific oligonucleotides were synthesized (Bioneer) as previously reported (20). Transfections were done using LipofectAMINE (Invitrogen). For luciferase assay, cells were scraped into 100 µL of passive lysis buffer (Promega). Luciferase activity in the lysate was determined with a Dual-Luciferase Reporter assay system, according to the manufacturer's instruction, and measured with a Turner Luminometer TD-20/20.

Reverse transcription-PCR and real-time PCR. To confirm the expression of RNA intramers, total RNA was extracted from cells after transfection with pU6-Aptamer or pDHFR-Aptamer by using the TRIzol (Invitrogen). After reverse transcription, cDNA was amplified by using primer pairs U6-F1 (5'-TGATGTCGACTAGGGACGCGTGGT-3') and U6-R1 (5'-GACTCTAGAGGATCCCCG-3') for pU6-Aptamer or DHFR-FOR (5'-TCACCGCGGGAGCTCGGTACC-3') and DHFR-REV (5'-TTGGATCCCCGCGGAAGCTT-3') for pDHFR-Aptamer. Real-time PCR was done with a Rotor-Gene RG-3000A system (Corbett Research). Reactions were amplified using the selective primers described above, and an LC FastStart reaction mix SYBR Green I kit (Roche), according to the manufacturer's instructions. A PCR amplification cycle was at denaturation at 95°C for 15 s, annealing at 47°C for 10 s (pU6-Aptamer) or at 55°C for 10 s (pDHFR-Aptamer), and extension at 72°C for 15 s. Quantification was carried out with the Rotor-Gene 6 software (Corbett Research). Relative levels of RNA intramers were expressed as the ratio of comparative threshold cycle (CT) to internal control glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNA.

Immunoprecipitation and Western blotting. Cytoplasmic and nuclear extracts were prepared as described (21) with minor modification. Cytoplasmic (350 µg) or nuclear (350 µg) extracts were incubated (2 h, 4°C) with a 50% (v/v) suspension of Protein G–Sepharose beads (Amersham) that had been precoated with 3 µg of anti–ß-catenin monoclonal antibody. After beads were washed, the immunoprecipitated proteins resolved by 10% SDS-PAGE were transferred to a polyvinylidene difluoride membrane. The blots were probed with specific antibodies and revealed with enhanced chemiluminescence. Anti–TCF-4 polyclonal (H-125) and anti-HuR monoclonal (3A2) antibodies were from Santa Cruz Biotechnology, and anti–ß-catenin monoclonal and anti–E-cadherin monoclonal antibodies were purchased from Transduction Laboratories. RNA immunoprecipitation assay in HT-29 cells was done as described previously (12).

In vivo splicing analysis of E1A pre-mRNA. Total cellular RNA was isolated with TRIzol (Invitrogen), reverse transcribed with Superscript II Reverse transcriptase (Stratagene), and used in the PCR reaction. The following PCR primers were used: E1A, 5'-GCTCCGACACCGGGACTG-3' (forward), 5'-GTCTCAGGATAGCAGGCACC-3' (reverse); ß-catenin, 5'-CGGGATCCACAAGAAACGGCTTTCA-3' (forward) and 5'-GAGAATTCCAGGTCAGTATCAAACCA-3' (reverse); GAPDH, 5'-TGACATCAAGAAGGTGGTGA-3'(forward), 5'-TCCACCACCCTGTTGCTGTA-3' (reverse).

Analysis of mRNA decay. For in vitro synthesis of RNA transcripts, the full-length COX-2 3'-UTR served as template and was amplified by PCR using primers containing the T7 polymerase promoter. The primers for the F1 UTR were forward, 5'-AGTTAATACGACTCACTATAGGGAAGTCTAATGATCATATT; and reverse, 5'-AGTCTAGATCACAAGTATGACTCCTT. The amplified F1 DNA was incubated in a standard transcription reaction containing [{alpha}-32P]UTP. All the labeled RNA transcripts were gel purified and quantified by liquid scintillation counting. The in vitro decay assay was done as described (12). To measure the decay rate of endogenous COX-2, cyclin D1, and c-myc mRNA, HT-29 or 293T cells were transiently transfected with the various expression vectors, followed by the addition of actinomycin D (10 µg/mL). Total RNA was extracted at intervals for the time course experiments. Remaining mRNA levels were measured by reverse transcription-PCR (RT-PCR) and real-time PCR. The following PCR primers were used: COX-2, 5'-TTCAAATGAGATTGTGGAAAAAT-3' (forward) and 5'-AGATCATCTCTGCCTGAGTATCTT-3' (reverse); cyclin D1, 5'-CTGGCCATGAACTACCTGGA-3' (forward) and 5'-GTCACACTTGATCACTCTGG-3' (reverse); c-myc, 5'-CTTCTGCTGGAGGCCACAGCAAACCTCCTC-3' (forward) and 5'-CCAACTCCGGGATCTGGTCACGCAGGG-3' (reverse).


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Localized expression of RNA intramers in colon cancer cells. We have previously isolated an RNA aptamer that binds to ß-catenin and expressed it as an RNA intramer using a U6-promoter–driven RNA expression vector (11). To explore the effect of ß-catenin on diverse steps of RNA metabolism from the nucleus to the cytoplasm, we used two different kinds of RNA intramers that might have differential localization patterns. In one, the U6-based expression system was again used because it generates small RNA transcripts restricted to the nucleus (19). The DNA sequence encoding the RNA aptamer was inserted between strong stem loops in the U6 RNA to yield pU6-Aptamer (Fig. 1A, top ). In the other case, the aptamer sequence was fused to the 5'-end of the aberrantly spliced DHFR cDNA, yielding pDHFR-Aptamer, to generate stable RNA in the cytoplasm (Fig. 1A, bottom). After transfecting two constructs into HCT116 colon cancer cells, we confirmed the expressions of RNA intramer by RT-PCR (data not shown). We also quantified the amount of intramers by real-time PCR and showed the higher level of pU6-based intramer than that of pDHFR-based intramer (Fig. 1B).


Figure 1
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  		Figure 1. Expression and localization of RNA intramers in HCT116 cells. A, structure of the intramer expression vectors, pU6-Aptamer and pDHFR-Aptamer. One RNA intramer was transcribed from the pU6-Aptamer expression vector with a Pol III–driven U6 promoter, and two strong stems of the human U6 transcript (filled box). The other intramer was transcribed from the pDHFR expression vector with a Pol II–driven cytomegalovirus (CMV) promoter, together with the aberrantly spliced DHFR mRNA (filled box). Bold arrows, the positions and directions of the PCR primers for the RNA aptamer. B, real-time PCR analysis of HCT116 cells transfected with the pU6-Aptamer or the pDHFR-Aptamer. GAPDH mRNA was used as an internal control and the relative expression of the RNA intramers is plotted. C, real-time PCR analysis of intramer localization. After transfection with the pU6-Aptamer (left) or the pDHFR-Aptamer (right) into HCT116 cells, total RNA was isolated from nuclear (NE) and cytoplasmic (CE) extracts, and the subcellular distributions of the RNA intramer were determined by real-time PCR. GAPDH served as an internal control, and the relative expression of the RNA intramers is plotted.

 
Because pU6- and pDHFR-based RNA intramers were expected to be differently localized in the cells, we fractionated the transfected cells and investigated their distributions between nucleus and cytoplasm. Real-time PCR analysis also confirmed the almost exclusive nuclear localization of the intramer when it was expressed from the pU6-Aptamer (Fig. 1C; ref. 19). In contrast, the intramer expressed from the pDHFR-Aptamer was mostly found in the cytoplasm, with smaller amounts in the nucleus of HCT116 cells (Fig. 1C).

Specific disruption of ß-catenin/TCF-4 complex by the nuclear RNA intramer. Because the RNA intramers were designed to bind to the protein-protein interaction domain of ß-catenin and were highly expressed, we tested whether they interfered with the interaction between ß-catenin and other binding proteins. After transfection with the two aptamer expression vectors into HCT116, we fractionated the cells to test whether the formation of subcellular protein complexes was inhibited (Fig. 2A ). Immunoprecipitation of nuclear extracts with ß-catenin antibody showed that the interaction between ß-catenin and the TCF-4 protein complex was completely abolished by the RNA intramer transcribed from pU6-Aptamer, whereas the ß-catenin/E-cadherin complex in cytoplasmic extracts was not significantly affected. In contrast, the RNA intramer transcribed from pDHFR-Aptamer reduced the level of the ß-catenin/E-cadherin complex, but had no effect on the ß-catenin/TCF-4 complex (Fig. 2A).


Figure 2
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  		Figure 2. Differential effects of the two RNA intramers on ß-catenin–protein complexes. A, coimmunoprecipitation (IP) assay of pDHFR- and pU6-derived RNA intramers. HCT116 cells were transfected with the indicated plasmids, and cytoplasmic and nuclear extracts were immunoprecipitated with anti–ß-catenin antibody, followed by Western blotting with anti–ß-catenin, anti–TCF-4, and anti–E-cadherin antibodies. B, HCT116 cells were cotransfected with TCF-responsive wild-type (OT) and mutant (OF) luciferase reporters and increasing amount of the pDHFR-Aptamer or pU6-Aptamer (0.2, 0.5, and 0.7 µg). After overnight incubation, luciferase activities were measured. Five independent experiments were done.

 
Because the nucleus-expressed intramer inhibited the TCF-4/ß-catenin interactions in HCT116 cells, it seemed likely to act as more potent transcription inhibitor than the cytoplasmic expressed intramer did. We therefore analyzed the effects of the two RNA intramers on ß-catenin–mediated transcription using TCF-responsive wild-type (OT) and mutant (OF) luciferase activities in HCT116 colon cancer cells (Fig. 2B). As expected, the pU6-derived intramer suppressed transcription more effectively than the pDHFR-derived intramer, which was mostly cytoplasmic with a little nuclear localization.

Regulation of mRNA alternative splicing by ß-catenin and the nuclear intramer. We used RNA intramers and shRNA to decide whether ß-catenin could mediate posttranscriptional regulation of mRNA. We have previously observed a specific effect of ß-catenin on the alternative splicing of estrogen receptor-ß (ER-ß) mRNA in colon cancer cells (11). We now used a minigene derived from adenovirus E1A mRNA (Fig. 3A ) to test the effects of the ß-catenin and the nucleus-expressed RNA intramer in HCT116 cells (Fig. 3B). Overexpression of ß-catenin significantly changed the alternative splicing pattern of the E1A minigene and led to a reduction of the 12S and 13S isoforms. However, coexpression of the pU6-based RNA intramer restored the normal splicing pattern of the E1A minigene.


Figure 3
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  		Figure 3. Role of ß-catenin and RNA intramer in RNA alternative splicing. A, diagram of the different E1A pre-mRNA splicing isoforms. Numbers, individual exons; dashed lines, spliced sequences. B, alternative splicing assay with an E1A minigene in HCT116 cells. Cells were cotransfected with the E1A reporter and the vector combinations are indicated at top. Alternative splicing of exogenous E1A mRNA was analyzed by RT-PCR. GAPDH is shown as a control. The expected bands resulting from alternative E1A splicing (full-length, 13S, 12S, 10S, and 9S) as well as ß-catenin, aptamer, and GAPDH are indicated on the right of the gels. C and D, alternative splicing assay with the E1A minigene in LoVo (C) and 293T cells (D). Alternative splicing of exogenous E1A mRNA was analyzed by RT-PCR as in (B). Bottom, the relative expression levels of indicated isoforms.

 
To exclude the possibility that the oncogenic mutation of ß-catenin of HCT116 cells was responsible for the alternative splicing of RNA, we used LoVo cells that harbor wild-type ß-catenin gene and higher level of endogenous protein (Fig. 3C). Expression of shRNA reduced the ß-catenin protein level in LoVo cells (data not shown), which also altered the splicing pattern of E1A minigene in LoVo cells. Significantly, the RNA intramer expressed in LoVo cells also showed the altered splicing pattern similar to that of shRNA-induced splicing. The most pronounced effect was observed when the RNA intramer and shRNA were coexpressed. These results show that ß-catenin induces alternative splicing of the E1A minigene, and that the RNA intramer and shRNA specifically reverse this alternative splicing in colon cancer cells, as shown for ER-ß alternative splicing. Similar results were also observed in nontumor 293T cell line (Fig. 3D).

Regulation of COX-2 mRNA stabilization by ß-catenin and the cytoplasmic RNA intramer. Because it was shown that ß-catenin regulated the stability of mRNA, we also tested the effect of the RNA intramer on ß-catenin–induced mRNA stabilization (Fig. 4 ). As previously described, the 3'-UTR of COX-2 transcripts is unstable in NIH3T3 cells and is markedly stabilized by overexpression of ß-catenin (12). In vitro decay assay was done to confirm the previous findings. Using the F1 fragment containing the AU-rich elements of the 3'-UTR of COX-2 mRNA (Fig. 4A), we measured the kinetics of decay and found that ß-catenin induced the stabilization of COX-2 mRNA, and the in vitro transcribed RNA aptamer inhibited the process (Fig. 4B). We next used the in vivo stability assay to investigate whether RNA intramer regulates the stability of endogenous COX-2 mRNA. 293T cells were transfected with ß-catenin–expressing vector in the presence or the absence of pDHFR-Aptamer. Transcription was stopped by actinomycin D treatment, and the stability of COX-2 mRNA was analyzed by RT-PCR (Fig. 4C) and real-time PCR analysis (Fig. 4D). ß-Catenin increased the half-life of endogenous COX-2 mRNA from 1 to >6 h as previously observed, but the coexpression of the RNA intramer decreased it back to 2 h.


Figure 4
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  		Figure 4. Role of ß-catenin and RNA intramer in mRNA stabilization. A, schematic representation of the locations of the F1 fragment in the COX-2 3'-UTR. Vertical lines, AU-rich elements. B, in vitro RNA degradation assay. {alpha}-32P–labeled F1 RNA substrates were incubated with cytoplasmic extracts from either vector- or ß-catenin–expressing NIH3T3 cells, and the reactions were stopped by adding stop buffer at the indicated times. In vitro transcribed RNA aptamer (IVT Aptamer, 1 µg) was supplemented to the ß-catenin–expressing cytoplasmic extracts. The processed RNA was resolved on a 7 mol/L urea/5% acrylamide gel and visualized by autoradiography. C and D, in vivo stability of the endogenous COX-2 mRNA. 293T cells were transfected with the indicated plasmids. Actinomycin D (ActD) was added to the culture medium at time 0, total RNA was isolated at the indicated times, and the amount of COX-2 mRNA was analyzed by RT-PCR. GAPDH served as a loading control. Expressions of ß-catenin and RNA intramer were also shown by RT-PCR. D, the stabilities of COX-2 mRNA in cells as (B) were examined by real-time PCR. The data are representative of three independent experiments.

 
Stabilization of cyclin D1 mRNA by ß-catenin. Because we have previously observed that ß-catenin bound to COX-2 mRNA in vitro as well as in vivo, we tested whether ß-catenin could also bind to other mRNA transcripts. RNA immunoprecipitation assay showed that ß-catenin interacted with endogenous cyclin D1 mRNA in HT-29 cells (Fig. 5A ) as well as in HCT116 cells (data not shown). We also showed that ß-catenin bound to cyclin D1 mRNA with similar extent as HuR-stabilizing protein binding to the same mRNA (21). We next tested whether cyclin D1 mRNA was stabilized by ß-catenin. Real-time PCR analyses showed that ß-catenin activation, by either overexpression (Fig. 5B) or LiCl treatment (data not shown), increased the half-life of cyclin D1 mRNA in HT-29 cells. Similar results were obtained in 293T cells, where coexpression of the cytoplasm-expressed RNA intramer restored ß-catenin–induced stabilization of cyclin D1 mRNA (Fig. 5C). However, we did not find any binding (Fig. 5A) or stabilization of another ß-catenin target transcript, c-myc mRNA, by ß-catenin protein (Fig. 5D). These results showed that ß-catenin bound to cyclin D1 transcript and stabilized the mRNA as did for COX-2 mRNA.


Figure 5
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  		Figure 5. Interaction and stabilization of cyclin D1 mRNA by ß-catenin. A, RNA immunoprecipitation of HT-29 cells. After formaldehyde fixation of the cells, immunoprecipitations were done with normal IgG, anti-HuR, or anti–ß-catenin antibodies. Bound RNA was extracted from the immune complexes and analyzed by RT-PCR. B, in vivo stability of endogenous cyclin D1 in HT-29 cells. Cells were transfected with ß-catenin expression vector or empty vector. Actinomycin D (10 µg/mL) was added to the culture medium at time 0, and total RNA was isolated at the indicated times. Amount of mRNA was analyzed by real-time PCR. C, in vivo stability of endogenous cyclin D1 mRNA in 293T cells. Cells were transfected with ß-catenin as well as pDHFR-intramer as indicated. After ActD treatment, total RNA was isolated at the indicated times. Amount of cyclin D1 mRNA was analyzed by real-time PCR. GAPDH mRNA served as an internal standard. Data are representative of three independent experiments. D, in vivo stability of endogenous c-myc mRNA in HT-29 cells. Cells were treated as in (B). Amount of mRNA was analyzed by real-time PCR.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Despite decades of intensive research on ß-catenin signaling, many questions remain to be answered to explain profound oncogenic effects of ß-catenin during tumor formation and progression (22, 23). Many transcriptional target genes have been identified, but their expression patterns cannot be solely explained by the action of ß-catenin as a transcriptional activator. For example, ß-catenin is shown to bind to TCF-4 consensus motifs near the known target genes as well as to other previously unknown target genes. Interestingly, the ß-catenin binding sites are not only in 5' promoter but also in internal regions and 3'-UTRs of protein-coding genes (24). These findings suggest that ß-catenin binds to multiple regions of genome and may act in many different levels of gene expression than previously envisioned. It is not clear at present whether such bindings are directly related to the transcriptional activity of ß-catenin or to other posttranscriptional activities of the protein.

If ß-catenin binds to nascent RNA just after transcription, it can regulate the splicing pattern of mRNA as shown for ER-ß and E1A minigene. Besides its function in the nucleus (9), ß-catenin may act as a scaffold protein for mRNA-mediated protein complexes, which accompany nuclear to cytoplasmic events. In fact, there are a couple of examples for the single protein involved both in RNA splicing and RNA stability (25, 26). If so, ß-catenin–RNA complex may be involved in the regulation of mRNA stability in the cytoplasm. We have shown here that ß-catenin bound to cyclin D1 mRNA as well as COX-2 mRNA and stabilized their half-lives (Fig. 5). It is interesting to note that ß-catenin was reported to regulate the cyclin D1 and Pitx-2 mRNA stabilities without the knowledge of ß-catenin binding to target mRNA (27). However, we did not observe any association or stabilization of c-myc mRNA by ß-catenin, which may be the reflection of different mRNA decay mechanisms for two transcripts. For example, c-myc mRNA is the very unstable transcript bound to AUF1 destabilization factor but cyclin D1 mRNA is the relatively stable transcript bound to HuR stabilization factor.

There are many critical questions that remain to be answered in the mechanism of ß-catenin–mediated RNA metabolism; it is definitely important to study the mechanism to fully understand the profound oncogenic effect of ß-catenin in cancer cells. Because the alternative splicing as well as aberrant stabilization of mRNA was frequently found in cancer cells, we envision that ß-catenin may also play critical roles in such cancer-specific RNA metabolism in addition to its role in transcription. These multiple functions would further strengthen the oncogenic potential of ß-catenin and Wnt signaling during colon cancer development and progression.

In addition to their use as therapeutics (28, 29), intracellularly expressed RNA aptamers have many uses (30). First, their exquisite specificity makes them selective targeting tools for proteins in complex cellular environment. Second, their high level of expression makes possible effective inhibition of the cellular targets. Thus, it has been reported that an H1 RNA polymerase III promoter-driven RNA aptamer caused strong inhibition of nuclear factor-{kappa}B p50 activity (31). Most significantly, as shown in this study, the controlled localization of an RNA intramer permits one to assess the role of a given protein in different cellular compartments, in contrast to the use of low molecular weight chemical antagonists (32, 33). Finally, the use of intramers can reveal previously unidentified functions of proteins, thereby providing novel insights into their roles.


   Acknowledgments
 
Grant support: Basic Science Research Fund of the Korea Research Foundation (2004-015-C00378), the Korea Science and Engineering Foundation (R01-2006-000-10194-0 and F01-2006-000-10199-0), and the Korean Ministry of Sciences and Technology (2005-01131=M10534040005-05N3404-00500).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 3/27/07. Revised 7/ 2/07. Accepted 7/23/07.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Maniatis T, Reed R. An extensive network of coupling among gene expression machines. Nature 2002;416:499–506.[CrossRef][Medline]
  2. Orphanides G, Reinberg D. A unified theory of gene expression. Cell 2002;108:439–51.[CrossRef][Medline]
  3. Bentley D. The mRNA assembly line: transcription and processing machines in the same factory. Curr Opin Cell Biol 2002;14:336–42.[CrossRef][Medline]
  4. Huelsken J, Birchmeier W. New aspects of Wnt signaling pathways in higher vertebrates. Curr Opin Genet Dev 2001;11:547–53.[CrossRef][Medline]
  5. Polakis P. Wnt signaling and cancer. Genes Dev 2000;14:1837–51.[Free Full Text]
  6. Tetsu O, McCormick F. ß-catenin regulates expression of cyclin D1 in colon carcinoma cells. Nature 1999;398:422–6.[CrossRef][Medline]
  7. He TC, Sparks AB, Rago C, et al. Identification of c-MYC as a target of the APC pathway. Science 1998;281:1509–12.[Abstract/Free Full Text]
  8. Behrens J, von Kries JP, Kuhl M, et al. Functional interaction of ß-catenin with the transcription factor LEF-1. Nature 1996;382:638–42.[CrossRef][Medline]
  9. Willert K, Jones KA. Wnt signaling: is the party in the nucleus? Genes Dev 2006;20:1394–404.[Abstract/Free Full Text]
  10. Sato S, Idogawa M, Honda K, et al. ß-Catenin interacts with the FUS proto-oncogene product and regulates pre-mRNA splicing. Gastroenterology 2005;129:1225–36.[CrossRef][Medline]
  11. Lee HK, Choi YS, Park YA, Jeong S. Modulation of oncogenic transcription and alternative splicing by ß-catenin and an RNA aptamer in colon cancer cells. Cancer Res 2006;66:10560–6.[Abstract/Free Full Text]
  12. Lee HK, Jeong S. ß-Catenin stabilizes Cyclooxygenase-2 mRNA by interacting with AU-rich elements of 3'-UTR. Nucleic Acids Res 2006;34:5705–14.[Abstract/Free Full Text]
  13. Dihlmann S, Doeberitz M. Wnt/ß-catenin-pathway as a molecular target for future anti-cancer therapeutics. Int J Cancer 2005;113:515–24.[CrossRef][Medline]
  14. Takahashi Y, Nishikawa M, Takakura Y. Suppression of tumor growth by intratumoral injection of short hairpin RNA-expressing plasmid DNA targeting ß-catenin or hypoxia-inducible factor 1{alpha}. J Control Release 2006;116:90–5.[CrossRef][Medline]
  15. Blind M, Kolanus W, Famulok M. Cytoplasmic RNA modulators of an inside-out signal-transduction cascade. Proc Natl Acad Sci U S A 1999;96:3606–10.[Abstract/Free Full Text]
  16. Mayer G, Blind M, Nagel W, et al. Controlling small guanine-nucleotide-exchange factor function through cytoplasmic RNA intramers. Proc Natl Acad Sci U S A 2001;98:4961–5.[Abstract/Free Full Text]
  17. Choi KH, Park MW, Lee SY, et al. Intracellular expression of the T-cell factor-1 RNA aptamer as an intramer. Mol Cancer Ther 2006;5:2428–34.[Abstract/Free Full Text]
  18. Kim MY, Jeong S. Inhibition of the functions of the nucleocapsid protein of human immunodeficiency virus-1 by an RNA aptamer. Biochem Biophys Res Commun 2004;320:1181–6.[CrossRef][Medline]
  19. Paul CP, Good PD, Winer I, Engelke DR. Effective expression of small interfering RNA in human cells. Nat Biotechnol 2002;20:505–8.[CrossRef][Medline]
  20. van de Wetering M, Oving I, Muncan V, et al. Specific inhibition of gene expression using a stably integrated, inducible small-interfering-RNA vector. EMBO Rep 2003;4:609–15.[CrossRef][Medline]
  21. Lal A, Mazan-Mamczarz K, Kawai T, Yang X, Martindale JL, Gorospe M. Concurrent versus individual binding of HuR and AUF1 to common labile target mRNAs. EMBO J 2004;23:3092–102.[CrossRef][Medline]
  22. Tolwinski NS, Wieschaus E. Rethinking WNT signaling. Trends Genet 2004;20:177–81.[CrossRef][Medline]
  23. Clevers H. Wnt/ß-catenin signaling in development and disease. Cell 2006;127:469–80.[CrossRef][Medline]
  24. Yochum GS, McWeeney S, Rajaraman V, Cleland R, Peters S, Goodman RH. Serial analysis of chromatin occupancy identifies ß-catenin target genes in colorectal carcinoma cells. Proc Natl Acad Sci U S A 2007;104:3324–9.[Abstract/Free Full Text]
  25. Ji X, Kong J, Carstens RP, Liebhaber SA. The 3'UTR complex involved in stabilization of human {{alpha}}-globin mRNA assembles in the nucleus and serves an independent role as splice-enhancer. Mol Cell Biol 2007;27:3290–302.[Abstract/Free Full Text]
  26. Soller M, White K. ELAV multimerizes on conserved AU4-6 motifs important for ewg splicing regulation. Mol Cell Biol 2005;25:7580–91.[Abstract/Free Full Text]
  27. Briata P, Ilengo C, Corte G, et al. The Wnt/ß-catenin->Pitx2 pathway controls the turnover of Pitx2 and other unstable mRNAs. Mol Cell 2003;12:1201–11.[CrossRef][Medline]
  28. Brody EN, Gold L. Aptamers as therapeutic and diagnostic agents. J Biotechnol 2000;74:5–13.[CrossRef][Medline]
  29. Nimjee SM, Rusconi CP, Sullenger BA. Aptamers: an emerging class of therapeutics. Annu Rev Med 2005;56:555–83.[CrossRef][Medline]
  30. Famulok M, Mayer G. Intramers and aptamers; applications in protein function analyses and potential for drug screening. Chem Biol Chem 2005;5:19–26.
  31. Mi J, Zhang X, Rabbani ZN, et al. H1 RNA polymerase III promoter-driven expression of an RNA aptamer leads to high-level inhibition of intracellular protein activity. Nucleic Acids Res 2006;34:3577–84.[Abstract/Free Full Text]
  32. Lepourcelet M, Chen YN, France DS, et al. Small-molecule antagonists of the oncogenic Tcf/ß-catenin protein complex. Cancer Cell 2004;5:91–102.[CrossRef][Medline]
  33. Nath N, Kashfi K, Chen J, Rigas B. Nitric oxide-donating aspirin inhibits ß-catenin/T cell factor (TCF) signaling in SW480 colon cancer cells by disrupting the nuclear ß-catenin-TCF association. Proc Natl Acad Sci U S A 2003;100:12584–9.[Abstract/Free Full Text]



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