Butyrate response factor 1 enhances cisplatin sensitivity in human head and neck squamous cell carcinoma cell lines



Cisplatin is a widely used chemotherapeutic agent in head and neck squamous cell carcinoma (HNSCC). Resistance to cisplatin is a common feature of HNSCC. To identify genes that may regulate cisplatin sensitivity, we carried out a cDNA microarray analysis of gene expression in cisplatin-sensitive and cisplatin-resistant HNSCC-derived cell lines. Among genes differentially expressed by cisplatin treatment, we have confirmed the elevated expression of butyrate responsive factor 1 (BRF1) in cisplatin-sensitive HNSCC cells and have demonstrated that the expression level of BRF1 is associated with cisplatin-sensitivity. Specific inhibition of BRF1 expression using an antisense oligodeoxynucleotide (ODN) decreased the cisplatin-sensitivity and, on the contrary, overexpression of BRF1 increased cisplatin-sensitivity in HNSCC cells. Elevated expression of BRF1 decreased the level of the human inhibitor of apoptosis protein-2 (cIAP2) and increased the caspase-3 activity in HNSCC cells. In addition, elevated expression of BRF1 decreased the expression level of enhanced green fluorescent protein (EGFP) linked to a 3′ terminal AU-rich element (ARE) of cIAP2 mRNA. These findings demonstrate that BRF1 expression enhanced cisplatin sensitivity in HNSCC cells by reducing the levels of cIAP2 mRNA. © 2005 Wiley-Liss, Inc.

The regulation of mRNA stability is an important factor in modulating gene expression, in particular for transiently-expressed genes that require tightly controlled mRNA levels. The best characterized cis-elements mediating control of mRNA stability have been the AU-rich element (ARE) contained in the 3′ untranslated region (3′UTR) of cytokines, growth factors, and proto-oncogenes.1 The list of mRNA containing the ARE has increased considerably with genome sequencing programs and several hundred ARE-containing genes have been compiled in the ARE mRNA database (ARED).2

Transcripts containing an ARE usually exhibit rapid decay in resting cells, but are readily stabilized upon exposure to exogenous signals. Importantly, defective ARE function causing increased transcript stability has been reported to be a pathogenic factor in human malignancies,3, 4, 5, 6, 7, 8 experimental tumors9 and inflammatory disorders.10, 11 The mechanism of mRNA stabilization is likely to involve ARE-binding proteins (AUBP) as adaptors between the signalling cascade and the decay machinery.12, 13 AUBP with assigned functions in vivo include TTP, butyrate response factor-1 (BRF1) and AUF1, all of which promote degradation, as well as HuR that exerts an mRNA stabilizing effect.14, 15, 16, 17, 18, 19 Alteration in the stabilizing and destabilizing activities of AUBP can be linked to the malignant state, survival and clinical recurrence of tumors. AUBP were implicated recently in tumorigenesis, causing tumor development when altered in their stabilizing or destabilizing activities.8, 20, 21, 22, 23

Initially, BRF1 was cloned from an epithelial rat cell line as an EGF-inducible cDNA of unknown function termed cMG1.24 The murine and human homologues were then cloned as TIS11b25 and EGF-response factor-1 (ERF-1),26 respectively. The human gene, renamed BRF1 by the Human Genome Database Nomenclature Committee, was located on chromosome 14q22.27BRF1 has been identified using functional genetic screening19 and was shown to mediate rapid decay of an ARE-containing reporter transcript18, 19, 28 and to enhance apoptosis.29, 30

The inhibitor of the apoptosis (IAP) family consists of 5 members: XIAP,31, 32, 33 cIAP1 and cIAP2,34 NAIP,31, 35 and Survivin.36 IAP are found to protect against a broad spectrum of apoptotic signals and overexpression of IAP in tumor cells has been shown to cause an inhibitory effect on cell death induced by a variety of apoptotic stimuli and induce resistance to chemotherapy.37

Cisplatin (cis-diamminedichloroplatinum) is a DNA-damaging agent that has been used in head and neck cancer, both alone and in combination with other chemotherapeutic agents or radiation therapy.38 Development of cisplatin resistance is a major obstacle in clinical treatment.39 Cisplatin resistance may be mediated by a number of different mechanisms, including reduced intracellular drug accumulation, drug detoxification, increase in DNA damage repair and overexpression of antiapoptotic proteins such as Bcl-2 and IAP.40

There are still wide gaps, however, in a fuller appreciation of the process leading to cisplatin resistance. An understanding of the cellular determinants of cisplatin sensitivity is indeed desirable in refining therapeutic approaches that further enhance the antitumor activity of the cisplatin.

We conducted a cDNA microarray to screen genes responsible for cisplatin sensitivity and identified BRF1 as a key gene that regulates cisplatin sensitivity in HNSCC cell lines. BRF1 inhibition using BRF1 antisense oligodeoxynucleotide (ODN) decreased the cisplatin sensitivity of the HNSCC cell line and, on the contrary, an overexpression of BRF1 induced apoptosis and increased the cisplatin sensitivity of the HNSCC cell line. We also show that BRF1 promotes a decrease in the level of cIAP2 mRNA and an increased caspase-3 activity. We propose that BRF1 enhanced cisplatin sensitivity by reducing the level of cIAP2 mRNA.

Material and methods

Cell culture and cell line selection

Four HNSCC cell lines, HN2, HN3, HN4 and HN7, derived from patients with HNSCC were studied41 and maintained in DMEM supplemented with 10% FBS (Gibco BRL, Carlsbad, CA), 100 U penicillin and 100 μg streptomycin/ml. They were cultured at 37°C in a humidified chamber containing 5% CO2. For the induction of apoptosis, cells were plated in 60-mm dishes 1 day before cisplatin treatment. Relative cisplatin cytotoxicity of the 4 cell lines was evaluated using an MTT colorimetric assay. Briefly, HNSCC cells were plated in triplicate at 1.2 × 104 cells/well in 96-well culture plates in DMEM. On the following day, the cells were treated with cisplatin at concentrations ranging from 0–50 μM. On the consecutive days, the cells were incubated with MTT dye (1 mg/ml) at 37°C for 4 hr and lysed in a buffer containing 20% (w/v) SDS, 50% (v/v) N,N-dimethylformamide (pH = 4.5). Absorbance at 600 nm (OD600) was determined for each well using an ELX 808 automated microplate reader (Bio-Tek Instrument, Inc., Winooski, VT).

TUNEL staining

TUNEL staining was conducted using an in situ cell death detection kit, TMR Red, according to the protocol supplied by the manufacturer (Roche Molecular Biochemicals, Mannheim, Germany). HNSCC cells were plated in 25 cm2 flasks at 2 × 105 cells/ml DMEM. On the following day, the cells were treated with 40 μM cisplatin, harvested and were fixed with 2% paraformaldehyde solution and permeabilized with 0.1% Triton X-100 in 0.1% sodium citrate. After washing twice with PBS, cells were incubated in a TUNEL reaction mixture containing terminal deoxynucleotidyl transferase and tetramethyl-rhodamine-dUTP. Cells were analyzed for fluorescence intensity using a FACS flow cytometer (Becton Dickinson, Inc., Franklin Lakes, NJ) and a FluoView™ 500 confocal microscope (Olympus, Tokyo, Japan).

The effect of caspase-3 inhibitor, Ac-DEVD-CHO (Calbiochem, Darmstadt, Germany), on cisplatin-induced apoptosis was determined by adding 20 μM or 40 μM Ac-DEVD-CHO to HN4 cells 1 hr before treatment with 40 μM cisplatin.

cDNA synthesis and array labeling

Gene expression was analyzed using the Atlas™ human 1.2, 1.2 II, and 1.2 III Arrays (Clontech, Franklin Lakes, NJ) that contain cDNA fragments of 3,528 human genes. To make radiolabeled cDNA pool probes of HNSCC cells, 10 μg of HN4 and HN7 cells total RNA were heated at 70°C for 2 min, cooled to 50°C, and incubated for an additional 2 min. Eight microliters of labeling reaction mixture, containing 50 mM Tris-HCl (pH = 8.3), 75 mM KCl, 3 mM MgCl2, dNTP mix (5 mM each dCTP, dGTP, dTTP; 50 μM dATP), 35 μCi of [α-32P]dATP, 10 mM DTT, and 50 U of MMLV reverse transcriptase, were added to the preheated total RNA and the reaction was incubated at 50°C for 20 min. cDNA was separated from unincorporated nucleotides using Microspin SR-400 columns (Amersham, Buckinghamshire, UK). The labeled probe was denatured by incubation in 0.1 M NaOH-0.1 mM EDTA at 68°C and then neutralized with 1 M NaH2PO4. Human Cot-1 DNA (20 μg/ml; Gibco BRL) was added to the probe to suppress cross-hybridization to repetitive DNA. The arrays were preincubated for 30 min in ExpressHyb hybridization solution (Clontech) with denatured salmon sperm DNA (100μg/ml; Roche) at 42°C. The labeled oligonucleotides were hybridized to the array for 18 hr at 42°C in 5 ml of ExpressHyb with salmon sperm DNA (100 μg/ml). The arrays were washed twice at 50°C in 2× SSC (0.3 M NaCl plus 0.03 M sodium citrate)-0.1% sodium dodecyl sulfate (SDS), with a final wash in 0.1× SSC-0.1% SDS. The hybridization signals in the arrays were visualized by autoradiography (BioMax film, Eastman Kodak, Rochester, NY), using different exposure times (1 hr, 3 hr, 6 hr, 1 day, 2 days, 1 week). Arrays were stored at 4°C on filter paper (Whatman, Brentford, UK) saturated with 0.1× SSC-0.1% SDS and were rinsed in 0.1× SSC-0.1% SDS at 68°C before being used again.

Data analysis of cDNA array

The signal generated by cDNA arrays were analyzed as described previously42 with minor modification. Briefly, autoradiographic signals were scanned with a flat-bed scanner (model ES-1000C; Epson, Nagano, Japan) in a grey mode of Adobe Photoshop 5.5. The 2 scanned autoradiograms (hybridized with HN4 and HN7 probes) were transformed to the RGB color mode with an 8-bit channel. A new canvas with an 8-bit RGB color mode was prepared, and the autoradiogram hybridized with HN4 cDNA probes was copied and pasted onto the prepared canvas with the green channel. The autoradiogram with HN7 cDNA probes was copied and superimposed upon the first using the red channel. Now the corresponding spots in the 2 autoradiograms appear as either green, red or a combination according to their composition of HN4 or HN7 probes. Using the information function of Adobe Photoshop, each corresponding spots was analyzed as to its color intensity, and the ratio of the resulting values were used to compare the 2 probe sets.

Semi-quantitative RT-PCR and real-time PCR

Five micrograms of DNase I-treated total RNA was reverse transcribed using oligo-dT and Superscript II reverse transcriptase (Invitrogen, Carlsbad, CA), according to the manufacturer's instructions. Semi-quantitative RT-PCR was carried out using Taq polymerase (Qiagen, Hilden, Germany) and appropriate primers. Quantitative PCR was carried out by monitoring in real-time the increase in fluorescence of the SYBR Green dye (Molecular Probes, Eugene, OR) on a DNA Engine Opticon Continuous Fluorescence Detection System (MJ Research Inc., Waltham, MA) according to the manufacturer's instructions. Specificities of each primer pair were confirmed by melting curve analysis and agarose-gel electrophoresis.

PCR primers


Plasmid construction and transfection

HN7 cell lines that overexpressed human BRF1 were generated using the pcDNA3.1/Myc-His A vector (Invitrogen). Full-length human cDNA of BRF1 was cloned by RT-PCR from the RNA of HN4 cell lines using a forward primer GGATTCATGACCACCACCCTC and a reverse primer CTCGAGGTC ATCTGAGATGGA and subcloned into the pcDNA3.1/Myc-His A vector. HN7 cells were transfected with pcDNA3.1-BRF1 construct using Lipofectamine (Gibco BRL). HN7/BRF1 cells stably transfected with BRF1 were selected by adding G418 (100μg of G418/ml; Gibco BRL) 3 days after transfection and stably transfected cells were tested for an overexpression of BRF1 by RT-PCR. A control cell line, HN7/pcDNA was generated by transfection with pcDNA3.1/Myc-His vector. A region of BRF1 (5′-TGGTGGTCATCCTGTGCGTT-3′) was chosen as the target for BRF1 antisense ODN. A BRF1 scrambled ODN with the same nucleotide composition as BRF1 antisense ODN but which lacked significant sequence homology to the BRF1 was also designed as a negative control. The phosphorothioate/phosphodiester chimeric ODN were purchased from Integrated DNA Technologies (Coralville, IA). Cells (5 × 105) were plated on culture flask (25 cm2) on Day 0 and transfection was carried out on Day 1 using oligofectamine (GIBCO-BRL) according to the manufacturer's protocol in the presence of 200 nM of ODN.

To generate pEGFP-cIAP2 3′UTR, the 3′UTR of human cIAP2 was amplified by PCR using primer cIAP2-UP (5′-TCTCGAGCTCTTTCATGAAGA-3′) and cIAP2-DOWN (5′-GGATCCCCCAGGATGGATTCA-3′) from cDNA of HN7. The 459-bp fragment was ligated into the SacI/BamHI site of pEGFP-C1 (Clontech). HN4 and HN7 cells were transiently transfected with pEGFP-cIAP2 3′UTR or empty pEGFP-C1 vector using Lipofectamine. Transfected cells were analyzed for fluorescence intensity using a FACS flow cytometer (Becton Dickinson, Inc.)

Caspase-3 activity assay

Cells were lysed in lysis buffer (10 mM HEPES, pH = 7.4, 10% sucrose, 2 mM EDTA, 0.1% 3-[(3-cholamidopropyl)dimethyl-ammonio]-1-propanesulfonate, 10 mM dithiothreitol, 1 mM phenylmethylsulphonyl fluoride and 20 μg/ml each of pepstatin A, leupeptin and aprotinin) for 15 min on ice. After centrifugation at 15,000g for 15 min at 4°C, supernatants containing 50 μg of protein were incubated with 50 μM Ac-DEVD-AFC for 1 hr. Caspase-3 activity was determined by fluorometric detection of the hydrolyzed product using a microplate spectrofluorometer (VICTOR2 Multilabel Reader, Wallac, Boston, MA).


Cisplatin cytotoxicity assay

To select cell lines with different cisplatin sensitivity, we determined cisplatin sensitivity among 4 HNSCC cell lines. There was variable cisplatin sensitivity among those 4 cell lines. The IC50 for the most resistant cell line, HN7, was approximately 10 times higher than the IC50 for the most sensitive cell line, HN4 (Fig. 1a). To examine the ability of cisplatin to induce apoptosis in HN4 and HN7 cells, cultures were treated with 40 μM cisplatin, after which they were stained with TUNEL and examined by FACS analysis. As shown in Figure 1b, cisplatin-induced apoptosis began to be detected in HN4 cells after 5 hr of treatment of cisplatin and increased markedly thereafter resulting in the death of >62% of the cell population by 12 hr of treatment. Cisplatin treatment did not cause a significant increase in cisplatin-induced apoptosis of HN7 cells, however, until 12 hr of cisplatin treatment.

Figure 1.

Selection of cisplatin-sensitive (HN4) and cisplatin-resistant (HN7) cell lines. (a) Cisplatin cytotoxicity assay for the 4 HNSCC cell lines. Cell lines were plated at equal density on Day 0, and treated with increasing concentrations of cisplatin on Day 1. Viability for all cell lines was determined by MTT on Day 2. (b) Cisplatin induced apoptosis of HN4 cells in a time-dependent manner. HN4 and HN7 cells were treated with 40 μM cisplatin for the indicated times. Cells were then fixed with 2% paraformaldehyde, stained with TUNEL, and analyzed by FACS. The results are presented as the means ± SD from 3 separate experiments.

Microarray analysis of genes differentially expressed between HN4 and HN7 cell lines

To determine genes that are differentially expressed between HN4 and HN7 cells, our next approach was to probe commercially available Atlas cDNA expression arrays (Clontech) with the cDNA pools derived from cisplatin-treated HN4 and HN7 cells. The arrays contain fragments of 3,528 known genes that encode a variety of proteins (for a complete listing, see www.clontech.com). A pair-wise comparison was made between a cisplatin-treated HN4 and HN7 cDNA probed arrays. Only transcripts that showed a net difference of more than 2-fold in hybridization intensity were considered for confirmation by real-time PCR. Of the 3,528 genes, 43 genes showed a net difference of more than 2-fold in hybridization intensity between HN4-array and HN7-array (Table I). Nineteen genes of the 43 transcripts showed a difference of more than 2-fold in real-time PCR analysis. Among the 19 genes whose expression was altered both in cDNA microarray and real-time PCR, BRF1 was prominent and was selected for further study.

Table 1. Differentially Expressed Genes Between Cisplatin-Treated HN4 and HN7 Cells
GenBank accession numberGene name or productFold change
cDnA arrayReal-time PCR
Experiment 1Experiment 2
 X79067Butyrate response factor 1 (BRF1)18.797.396.97
 X93921Dual-specificity protein phosphatase 77.732.932.37
 X54297Histidine decarboxylase (HDC)6.132.652.31
 D16826Somatostatin receptor 4 (SSTR4)5.972.542.37
 AF052941DAP-Kinase related protein 14.862.242.07
 M83234Nuclease-sensitive element DNA-binding protein (NSEP)4.792.202.18
 U15979Delta-like protein (DLK)4.513.322.99
 M22490Bone morphogenetic protein 4 (BMP4)4.321.431.31
 M60314Bone morphogenetic protein 5 (BMP5)4.301.281.27
 U40462Ikaros/LyF-1 homolog hlk-14.201.651.51
 X00588Epidermal growth factor receptor (EGFR)
 D15057Defender against cell death 1 protein4.061.321.27
 M12529Apolipoprotein E (APOE)3.971.281.22
 Z35141Paired box protein 7 (PAX7)3.061.491.42
 U20759Extracellular calcium-sensing receptor (CASR)
 Y00109Mitochondrial aldehyde dehydrogenase2.841.271.18
 Z31718Myelin protein zero (MPZ)2.301.131.11
 L35240Enigma protein2.241.151.12
 X08058Glutathione S-transferase pi (GSTPi)15.282.502.43
 U07418mutL protein homolog 1 (MLH1)10.542.152.03
 X5407927-kDa heat shock protein (HSP27)9.863.172.97
 AB021288Homo sapiens mRNA for beta 2-microglobulin9.262.142.01
 D38076RAN binding protein 1 (RANBP1)9.122.332.06
 M17733Thymosin beta 4 X chromosome (TMSB4X)7.692.272.23
 X01402IL2 receptor alpha subunit (IL2RA)7.672.312.20
 X95404Non-muscle cofilin 1 (CFL1)7.391.951.87
 X13238Cytochrome c oxidase polypeptide Vic5.111.981.90
 U43586Kinase suppressor of ras-1 (KSR1)4.921.741.65
 U0995360S ribosomal protein L9 (RPL9)4.591.921.66
 M98343emsl oncogene4.181.951.93
 AF092124ATPase subunit G4.171.791.52
 M26708Prothymosin alpha (PTMA)4.031.531.38
 X07549Cathepsin H (CTSH)3.743.153.06
 K02770Interleukin 1 beta (IL1B)3.612.272.21
 M35368Galactoside-binding protein (GALBP)3.271.431.26
 X16707FOS-related antigen 1 (FRA1)3.231.391.27
 X0727090-kDa heat-shock protein A (HSP90A)2.772.162.04
 J03746Microsomal glutathione S-transferase 1 (MGST1)2.751.321.26
 X51841Integrin beta 4 (ITGB4)
 X13227Integrin alpha L (ITGAL)2.191.321.14
 L19779Histone 2A (H2A)2.151.411.21

BRF1 expression and cisplatin sensitivity in 4 HNSCC cell lines

To determine whether or not BRF1 expression was correlated with cisplatin sensitivity, we analyzed BRF1 expression in 4 HNSCC cell lines with different cisplatin sensitivities. Cell lines showing cisplatin sensitivity (such as HN4 and HN3) showed elevated expression levels of BRF1 compared to cisplatin resistant cell lines (such as HN2 and HN7) (Fig. 2a), suggesting an association of BRF1 expression with cisplatin sensitivity of HNSCC cell lines.

Figure 2..

Effects of BRF1 on the cisplatin-induced apoptosis in HNSCC cell lines. Levels of BRF1 expression in 4 HNSCC cell lines were determined by RT-PCR (a). Cisplatin-sensitive cell, HN4, was transfected with 200 nM of BRF1 antisense or scrambled ODN and then 2 days after the transfection, cells were treated with 40 μM cisplatin for 8 hr. Cisplatin-resistant cell, HN7, was stably transfected with full-length cDNA of BRF1 and, as a control, HN7 cells were stably transfected with pcDNA3.1 vector. The stable transfectants of HN7 were treated with 40 μM cisplatin for 8 hr. The expression of BRF1 was determined by RT-PCR before the cisplatin treatment (b) and the cisplatin-induced apoptosis was determined by TUNEL staining at 8 hr after cisplatin treatment (c). (d) Caspase-3 activity was measured in cell lysates at 8 hr after cisplatin treatment as described under Material and Methods. The caspase-3 activity of parental HN7 cells was defined as one. (e) Effect of the caspase-3 inhibitor Ac-DEVD-CHO on cisplatin-induced apoptosis in HN4 cells. HN4 cells were treated with 40 μM cisplatin in the absence or presence of Ac-DEVD-CHO (20 μM and 40 μM). Ac-DEVD-CHO was added 1 hr before cisplatin treatment. Cisplatin-induced apoptosis was determined by TUNEL staining at 8 hr after cisplatin treatment. Data represent means ± SD of 3 independent experiments.

BRF1 overexpression enhanced the sensitivity of HNSCC cell lines to cisplatin

Although our results demonstrated that elevated levels of BRF1 were associated with cisplatin sensitivity, they do not address whether BRF1 was involved directly in cisplatin sensitivity in HNSCC cells. We sought to determine if an overexpression of BRF1 could induce cisplatin sensitivity in cisplatin resistant HNSCC cells, HN7 cells. To test this possibility, BRF1 expression vector (pcDNA3.1-BRF1) was transfected into HN7 cells to establish a stable transfectant derivative of HN7, HN7/BRF1. As a negative control, an HN7/pcDNA cell line stably transfected with empty vector was included.

Overexpression of BRF1 in HN7/BRF1 cell line was confirmed by RT-PCR (Fig. 2b). BRF1 expression of HN7/BRF1 cells was increased by 5-fold compared to HN7 (Fig. 2b). As shown in Figure 2c, the cisplatin sensitivity of HN7/pcDNA cells was slightly increased compared to that of HN7 cells, which may be explained by the nonspecific effects of vectors. The HN7/BRF1 showed an increased cisplatin sensitivity by 8-fold compared to HN7 cells.

To test whether or not downregulation of BRF1 affected cisplatin-sensitivity in HN4 cells, we used antisense ODN. Treatment of the HN4 cells with antisense ODN against BRF1 significantly reduced BRF1 mRNA level to 15% of control (Fig. 2b). Downregulation of BRF1 by treatment with antisense ODN reduced the cisplatin-induced apoptosis to 17% of control (Fig. 2c). Treatment with scrambled ODN did not affect the endogenous BRF1 mRNA level, however, and also did not induce any change in the cisplatin-induced apoptosis. These results indicated that the expression of BRF1 was associated directly with the cisplatin-induced apoptosis of HNSCC cell lines.

BRF1 expression enhanced the caspase-3 activity

There was a significant difference in caspase-3 activity between cisplatin-sensitive HN4 and cisplatin-resistant HN7 cells. Although caspase-3 activity was hardly detected in cisplatin-treated HN7 cells, cisplatin-treatment induced a 2.1-fold increase in caspase-3 activity in HN4 cells (Fig. 2d). Treatment of caspase-3 inhibitor, Ac-DEVD-CHO, suppressed the cisplatin-induced apoptosis of cisplatin-sensitive HN4 cells (Fig. 2e), which suggested that caspase-3 activity was required for the cisplatin-induced apoptosis of HN4 cells. To determine whether or not BRF1 expression was associated with caspase-3 activity, we analyzed caspase-3 activities among HN7/BRF1 cells and antisense ODN-treated HN4 cells after cisplatin treatment. Although caspase-3 activity of HN7/BRF1 cells was increased by 1.6-fold compared to that of HN7 cells, that of antisense ODN-treated HN4 cells was reduced to 53% of untreated HN4 cells (Fig. 2d). These results showed that BRF1 expression increased caspase-3 activity, which coincided with an increased cisplatin-induced apoptosis.

BRF1 expression enhanced the cisplatin sensitivity of HNSCC cell lines by reducing the level of cIAP2

BRF1 has been found to be a member of AUBP and to accelerate the decay rates of ARE-containing mRNA.18, 19, 28 and to induce apoptosis.29, 30 In HNSCC cells, BRF1 expression increased caspase-3 activity and cisplatin-induced apoptosis. It is possible to speculate that BRF1 facilitated the mRNA degradation of anti-apoptotic genes and increased the cisplatin-induced apoptosis. To determine whether or not BRF1 expression regulated mRNA degradation of anti-apoptotic genes, we analyzed the mRNA levels of anti-apoptotic genes of HNSCC cells by RT-PCR, including Bcl-2, Bcl-xL, XIAP, cIAP1, cIAP2, NAIP and Survivin. In addition, we analyzed several apoptotic genes within the Bcl-2 family, such as Bak, Bax, Bid and Bik. Among the genes we tested, cIAP2 expression was found to be associated with the expression level of BRF1 (Fig. 3). The expression level of cIAP2 of HN4 cells was severely reduced to 20% of that of HN7. To determine whether or not BRF1 expression was associated directly with the expression level of cIAP2, we analyzed the level of cIAP2 mRNA in HN7/BRF1 cells and antisense ODN-treated HN4 cells. There was no change in the level of cIAP2 mRNA between HN7 and HN7/pcDNA cells. The level of cIAP2 mRNA of HN7/BRF1, however, was reduced to 44% of that of HN7. In addition, the suppression of BRF1 expression in HN4 cells using antisense ODN increased the level of cIAP2 mRNA to 90% of that of HN7 (Fig. 3). Our results suggest the possibility that BRF1 expression decreased the expression level of cIAP2 mRNA and, thus, increased the caspase-3 activity and the cisplatin-induced apoptosis.

Figure 3.

Effects of BRF1 on the expressions of apoptotic and anti-apoptotic genes. Total RNA samples were isolated from parental HN4 and HN7 cells, antisense ODN-treated or scrambled ODN-treated HN4 cells and HN7 derivatives stably transfected with pcDNA3.1-BRF1, HN7/BRF1 or empty vector pcDNA3.1, HN7/pcDNA. Expressions of apoptotic and anti-apoptotic genes were determined by semiquantitative RT-PCR analysis and the PCR products were analyzed by 1% agarose gel electrophoresis and stained with ethidium bromide (left panels). The band densities in the agarose gel were quantified by PhosphorImager, normalized to the internal control GAPDH and expressed as percentage (%) of the HN7 value. Data represent means ± SD of 3 independent experiments (right panels).

BRF1 expression decreased the level of cIAP2 3′UTR-containing mRNA of EGFP

Although our studies demonstrated that elevated levels of BRF1 were associated with cellular states displaying a reduced level of cIAP2 mRNA, they did not address whether or not BRF1 was sufficient to decrease cIAP2 expression in HNSCC cells. To evaluate the possibility that BRF1 expression and cIAP2 mRNA reduction were associated events, we made use of an EGFP reporter gene that linked it to the cIAP2 3′UTR in plasmid pEGFP-C1 (Fig. 4). Upon transient transfection into HNSCC cells, the cIAP2 3′UTR inhibited EGFP expression in HN4 cells but did not inhibit EGFP expression in HN7 cells. When HN7 cells were transfected to over express BRF1 in HN7/BRF1, however, the expression of EGFP containing cIAP2 3′UTR was inhibited. The results of a representative experiment are shown in Figure 5a and quantification of at least 3 independent experiments is exhibited in Figure 5b. Collectively, the results shown in Figures 3 and 5 indicated that the elevated expression of BRF1 contributed to an enhancement of the cisplatin sensitivity by lowering the cIAP2 level.

Figure 4.

Schematic representation of the reporter construct EGFP-cIAP2 ARE. To create EGFP-cIAP2 ARE, cIAP2 ARE, a 446-nt segment of cIAP2 3′UTR ranging from nucleotide 2255–2700 of the cIAP2 cDNA sequence (GenBank accession no. U45878) and containing the ARE, was ligated into the SacI/BamHI site of pEGFP-C1 vector. Dark-shaded box, EGFP; light-shaded box, cIAP2 ARE; white boxes, AUUUA motifs.

Figure 5.

Expression of EGFP reporter construct in HNSCC cell lines. EGFP-reporter gene expression constructs containing the cIAP2 3′UTR (EGFP-cIAP2 3′UTR) or no 3′UTR (EGFP) were transfected in the parental HN4 and HN7 cells and HN7 derivatives, HN7/BRF1 and HN7/pcDNA. (a) EGFP expression was analyzed by FACS. (b) Results shown on the graph represent means ± SD of 3 independent experiments.


We have demonstrated a strong positive correlation between the cellular BRF1 levels and cisplatin sensitivity in HNSCC cell lines. This suggests that BRF1 expression is a major determinant of cisplatin sensitivity in HNSCC cells. This conclusion was established by our results showing that the expression level of BRF1 was increased in cisplatin-sensitive HNSCC cells. In addition, an overexpression of BRF1 reverted the cisplatin-resistant cells to cisplatin-sensitive cells and, vice versa, a suppression of BRF1 using antisense ODN reverted the cisplatin-sensitive cells to cisplatin resistant cells. There was an inverse correlation between the expression levels of BRF1 and that of cIAP2 mRNA. This suggests that an expression of BRF1 decreases the accumulation level of cIAP2 mRNA, which then leads to the cisplatin-sensitivity of HNSCC cells.

BRF1 has been described as a protein able to bind to the ARE contained in the 3′UTR of mRNAs and to participate in their destabilization.18, 19, 28 ARE is clustered into 5 groups depending on the number of motifs in the ARE stretch. Groups 1–4 contain 5, 4, 3 and 2 pentameric (AUUUA) repeats, respectively, whereas Group 5 contains only one repeat within the 13-bp pattern.43 Based on this grouping, mRNA of cIAP2 contains Group 3 ARE (http://rc.kfshrc.edu.sa/ared/diversity.htm). Thus, it is possible that BRF1 binds to the ARE in the 3′UTR of cIAP2 mRNA and shortens cIAP2 mRNA half-lives. Several lines of evidence show a direct correlation between BRF1 expression and the modulation of cIAP2 mRNA expression: (i) elevated levels of BRF1 are associated with cellular states displaying reduced levels of cIAP2 mRNA; (ii) overexpression of BRF1 is sufficient to decrease cIAP2 expression in HNSCC cells and, vice versa, suppression of BRF1 expression using antisense ODN increases cIAP2 expression in HNSCC cells; and (iii) the expression of EGFP reporter gene containing cIAP2 3′UTR was inhibited when BRF1 was overexpressed. cIAP2 was shown to bind and inhibit caspase 3 and suppress apoptosis.44 Our study provides evidence that the caspase 3 activity of HNSCC cells exhibited a positive correlation with the expression levels of BRF1 and showed an inverse correlation with that of cIAP2 mRNA. It is possible to speculate that reduced expression levels of cIAP2 by BRF1 expression can cause an increase of caspase 3 activity and lead to the apoptotic cell death of HNSCC cells.

It was surprising to observe that there was no change in the levels of Bcl-2 mRNAs containing ARE. The Bcl-2 mRNA contained the same Group 3 ARE as cIAP2 mRNA does43 and binding of AUBP such as AUF121, 45, 46 or TINO47 to this Bcl-2 ARE was reported to modulate Bcl-2 mRNA stability. In our study, although the expression level of cIAP2 mRNA containing Group 3 ARE was reduced by BRF1 expression, that of Bcl-2 mRNA containing the same Group 3 ARE was not affected by BRF1 expression. Similar phenomena were reported by other researchers. AUF1, for example, has been clearly linked to RNA destabilization,17, 48 but can enhance mRNA stability under stress responses such as heat shock49 or in a transgenic mouse.20 In addition, an overexpression of HuR in TNF-α-stimulated glioma cells significantly augmented the mRNA stabilization of VEGF, TNF-α, and IL-8 but not c-myc,23 yet HuR has been shown to bind avidly to the 3′UTR of c-myc.50, 51 These findings suggest that the interaction between these factors to confer stability or instability to the transcript is likely to be complex and the determinants of mRNA stability or instability cannot be predicted by any one individual RNA-binding factor but requires auxiliary factors with which it interacts to exert its modulating activity.

An analysis of the human ARE-containing mRNA database suggests that the proportion of mRNA with ARE could be as high as 8%.2 Besides the cIAP2, there could be many candidates for post-transcriptional regulation by BRF1. Because a series of regulators of relevant for oncogenic growth including cyclins, growth factors and proto-oncogenes contain ARE,1 it is conceivable that AUBP, in fact, has a broader spectrum of target mRNA in tumor cells and that it might play a role in human carcinogenesis. Recently, AUBP were implicated in tumorigenesis, causing tumor development when altered in their stabilizing or destabilizing activities.8, 20, 21, 22, 23 Alteration in the stabilizing and destabilizing activities of BRF1 can be linked not only to cisplatin sensitivity as described in our study but also to carcinogenesis. The cIAP2, a target gene of BRF1, contains Group 3 ARE and it is possible that BRF1 regulates the expression levels of mRNAs containing Group 3 ARE. It does not, however, seem that BRF1 can regulate all members of Group 3 because, in our study, expression of BRF1 did not cause any changes in the level of Bcl-2 mRNA containing Group 3 ARE. Further study of the BRF1 target genes may provide insight into the relation between BRF1 expression and cisplatin resistance and carcinogenesis.

We have shown for the first time that BRF1 act as a trans-acting factor regulating the expression level of cIAP2 mRNA and its expression is associated with cisplatin-sensitivity in head and neck cancer. Our results may provide important information to understand the molecular mechanisms that lead to cisplatin resistance.


This project was supported by a Korean Research Foundation Grant. J.S. Kim was partly supported by the BK21 program of the Korean Research Foundation.