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Molecular and Cellular Biology, December 2006, p. 9256-9267, Vol. 26, No. 24
0270-7306/06/$08.00+0     doi:10.1128/MCB.01125-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

ATM and ATR Pathways Signal Alternative Splicing of Drosophila TAF1 Pre-mRNA in Response to DNA Damage{triangledown}

Rebeccah J. Katzenberger,{dagger} Matthew S. Marengo,{dagger} and David A. Wassarman*

University of Wisconsin School of Medicine and Public Health, Department of Pharmacology, Molecular and Cellular Pharmacology Program, 1300 University Avenue, Madison, Wisconsin 53706

Received 22 June 2006/ Returned for modification 1 August 2006/ Accepted 21 September 2006


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results and Discussion
 References
 
Alternative pre-mRNA splicing is a major mechanism utilized by eukaryotic organisms to expand their protein-coding capacity. To examine the role of cell signaling in regulating alternative splicing, we analyzed the splicing of the Drosophila melanogaster TAF1 pre-mRNA. TAF1 encodes a subunit of TFIID, which is broadly required for RNA polymerase II transcription. We demonstrate that TAF1 alternative splicing generates four mRNAs, TAF1-1, TAF1-2, TAF1-3, and TAF1-4, of which TAF1-2 and TAF1-4 encode proteins that directly bind DNA through AT hooks. TAF1 alternative splicing was regulated in a tissue-specific manner and in response to DNA damage induced by ionizing radiation or camptothecin. Pharmacological inhibitors and RNA interference were used to demonstrate that ionizing-radiation-induced upregulation of TAF1-3 and TAF1-4 splicing in S2 cells was mediated by the ATM (ataxia-telangiectasia mutated) DNA damage response kinase and checkpoint kinase 2 (CHK2), a known ATM substrate. Similarly, camptothecin-induced upregulation of TAF1-3 and TAF1-4 splicing was mediated by ATR (ATM-RAD3 related) and CHK1. These findings suggest that inducible TAF1 alternative splicing is a mechanism to regulate transcription in response to developmental or DNA damage signals and provide the first evidence that the ATM/CHK2 and ATR/CHK1 signaling pathways control gene expression by regulating alternative splicing.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results and Discussion
 References
 
Alternative splicing is a major mechanism utilized by higher eukaryotic organisms to regulate gene expression during development and in response to stress (8, 44, 48, 50). In fact, 35 to 74% of human genes may encode pre-mRNAs that are alternatively spliced (10, 22, 23, 29, 34). Alternative splicing can regulate whether or not a protein is produced, or it can generate pre-mRNAs that encode proteins with distinct functions (7, 17). By analogy to other gene expression-regulatory mechanisms, such as transcription, it is probable that signal transduction pathways play a widespread role in controlling alternative splicing. However, documented examples of this phenomenon are limited, and a complete pathway has not been described.

One of the most thoroughly understood examples of signal-dependent alternative splicing is Ras signal-induced splicing of the CD44 pre-mRNA in humans (28, 32, 57). The Ras GTPase and the downstream mitogen-activated protein kinase (MAPK) signaling cascade specify inclusion of exon 5 (v5) in the mature CD44 mRNA. Stimuli that activate Ras lead to activation of MAPK, which in turn phosphorylates SAM68, an RNA-binding protein that interacts with an exonic splicing silencer element within v5. Phosphorylated SAM68 is then thought to interfere with the repressive activity of hnRNP A1 and allow factors bound to a v5 exonic splicing enhancer element to enhance v5 inclusion. Signal-dependent alternative splicing has also been implicated in the regulation of cellular processes, including apoptosis and the cell cycle (44, 47, 49). For instance, many genes encoding apoptotic regulators are alternatively spliced; however, little is known about how apoptotic signaling pathways interact with the splicing machinery.

In humans, genes encoding TAF (TATA-binding protein [TBP]-associated factor) components of the general transcription factor TFIID are alternatively spliced (4, 5, 12, 52, 59). TFIID is broadly required for RNA polymerase II transcription in eukaryotes and plays a crucial role in recognizing core promoter elements and assembling the preinitiation complex (15, 21, 25, 31, 35). In response to apoptotic signals, human cells produce a TAF6 isoform, TAF6{delta}, by alternative splicing of the TAF6 pre-mRNA and then caspase-dependent cleavage of the encoded protein (5). Increased transcription of proapoptotic genes in TAF6{delta}-expressing cells may result from altered TFIID core promoter recognition. Thus, signal-dependent alternative splicing of TAF pre-mRNAs may be an important determinant of gene-specific transcription, but the secondary messengers that signal these alternative splicing events have not been identified.

In this study, we have investigated alternative splicing of the Drosophila melanogaster TAF1 pre-mRNA. TAF1 (formerly known as TAFII250) is the largest subunit of TFIID (56). In flies, TAF1 null mutations are recessively lethal, and clonal analysis indicates that TAF1 is essential for cell proliferation or viability (55). Knockdown of TAF1 in D. melanogaster embryonic Schneider cell line 2 (S2) tissue culture cells by RNA interference (RNAi) results in cell cycle arrest in the G2/M phase (30). Thus, normal cell physiology is critically dependent on TAF1.

Our data provide evidence that the ATM (ataxia-telangiectasia mutated) and ATR (ATM-RAD3 related) signal transduction pathways regulate TAF1 pre-mRNA alternative splicing in response to DNA damage. ATM and ATR proteins are members of a family of serine/threonine kinases, structurally related to phosphatidylinositol 3-kinases (PIKKs) (2). ATM and ATR function in cell cycle checkpoint pathways activated by DNA damage. ATM activates G1/S, S, and G2/M phase checkpoints in response to DNA double-strand breaks (1, 24, 45, 46). In contrast, ATR activates the S and G2/M phase checkpoints in response to defects in DNA replication, such as stalled replication forks. ATM and ATR induce cell cycle arrest in part by phosphorylating and activating the checkpoint kinases CHK2 and CHK1. These effector kinases phosphorylate proteins such as the transcription factor p53 and CDC25 phosphatase family members to arrest cell cycle progression. Our study reveals that, in response to different DNA damage stimuli, the ATM/CHK2 and ATR/CHK1 signaling pathways are required for alternative splicing of the TAF1 pre-mRNA.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results and Discussion
 References
 
Reverse transcriptase PCR (RT-PCR). Total RNA was isolated from S2 cells using an RNeasy Mini kit (QIAGEN). cDNA was synthesized with 1 µg RNA in a 20-µl RT reaction mixture using an iScript cDNA synthesis kit (Bio-Rad). PCR mixtures contained 0.5 or 1.0 µl cDNA for actin 5C or TAF1 amplification, respectively. Reaction mixtures contained 20 mM Tris-HCl (pH 8.4), 50 mM KCl, 1.5 mM MgCl2, 200 µM deoxynucleoside triphosphates, 1 µM each primer, and 5 U Taq polymerase (Invitrogen). PCR cycling conditions were 94°C for 4 min followed by cycles of 1 min at 94°C, 1 min at 58°C, and 1 min at 72°C. actin 5C and total TAF1 were amplified for 25 cycles, and all TAF1 isoforms and TAF1-4 were amplified for 28 cycles. After PCR, 10-µl samples were resolved on 1.2% agarose gels. Primer sets (oriented 5' to 3') were as follows: actin 5C, CCTTTCAAACCGTGCGGTCG and GGACGTCCCACAATCGATGG; a common region of TAF1 (exon 10), TGGATAAGTTTTCTCGTGGC and GCCATCGTTACCTCTAAAGG; the alternatively spliced region of TAF1, TGAAGCGCGGAAGGGGTAGG and ATCTGAGGGATCGTAGTTGG; and TAF1-4, CAGAATCCGGTTAAGCGTGGand CAACTGCACCATTGCTTCGG. Primers were synthesized by Integrated DNA Technologies.

qPCR. Total RNA was isolated from fly tissues or S2 cells as described above. cDNA was synthesized using 1 µg RNA as described above. Each 25-µl real-time RT-PCR (qPCR) mixture contained 0.5 µl cDNA, 12.5 µl iQ SYBR Green Supermix (Bio-Rad), and 250 nM primers. Reactions were carried out using an iCycler thermal cycler (Bio-Rad). PCR cycling conditions were 95°C for 3 min followed by cycles of 30 s at 95°C, 30 s at 61°C, and 30 s at 72°C. Melt curve analysis and agarose gel electrophoresis were carried out to evaluate the homogeneity of the reaction products. All primer sets produced a single product of the expected size. Amplicons were 75 to 400 bp. Primer sets (oriented 5' to 3') for qPCR were as follows: actin 5C, CGAAGAAGTTGCTGCTCTGGTTGTCG and GGACGTCCCACAATCGATGGGAAG; total TAF1, GGCCAAGTCAAATGATGCATCTAGTCCCand CAGCTTCCGATCCGCATCCTTTG;TAF1-1, CGTGGAGGAGGATCTCCAATGCTC and CATCCATGGATTCATCTGCCATCTGG;TAF1-2, CCAGAATCCGGTTAAGCGTGGTCG and CATCCATGGATTCATCTGCCATCTGG;TAF1-3, CTCAACTGCACCATTGCTTCGGCC and CGTGGAGGAGGATCTCCAATGCTC;TAF1-4, CTGGATGAAGATCTCCAATGCTCCAC and CGATCGGCTCCTCTGCCATCTG; ATM, AGGATATCATCGAGCAGAACCGCC and GCTGCTGCTCATCCAAACTAGCG; ATR, ATCCCTCCGAGCGCTTACGAA and CCCTTGCAAAGCGGATTCACGATG; CHK1, GAACAACTGCAATCCCGGTACACC and TTTACCTCGCCGTAGGCACCTTCG; CHK2, CGAAGATCGGACTCCTCGTTTCCA and GTGTGTCGCGTGCCATAGTGATTC; and dU2AF38, CCACAACAAACCCACTTTCTCGCA and ACTTGTCCTCGCACTCTACGAACA. Experiments were conducted in duplicate or triplicate sets; n (as indicated in the figure legends) represents the number of samples.

Cell culture and drug treatments. S2 cells were maintained at room temperature in Schneider's Drosophila medium containing 10% heat-inactivated fetal bovine serum, 100 U/ml penicillin, and 100 µg/ml of streptomycin (Gibco). For drug treatments, cells were cultured to a density of 2 x 106 cells/ml. Camptothecin (CPT) (Sigma) in dimethyl sulfoxide (DMSO) was added to cells to the concentrations indicated in the legends for Fig. 3 to 8 and incubated at room temperature. DMSO-treated cells served as the mock control. Cells were irradiated using a Mark 1 irradiator and allowed to recover at room temperature. Doses and recovery times are indicated in Fig. 3 to 8. As indicated in the legend for Fig. 6, cells were cotreated with 20 mM caffeine (Sigma) in water or 1 µM wortmannin (Sigma) in DMSO. Where indicated (see the legend for Fig. 5), cells were pretreated with 25 ng/ml actinomycin D (ActD; Sigma) in ethanol 2 h prior to CPT treatment or cotreated with 100 µg/ml cycloheximide (Calbiochem) in methanol and CPT.


Figure 3
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  		FIG. 3. TAF1 pre-mRNA alternative splicing was altered by DNA damage. S2 cells were treated with DMSO or 20 µM CPT (lanes 1 to 8) for the times indicated or unirradiated (untreated) or treated with 40 Gy IR (lanes 9 to 11) and then allowed to recover for the times indicated. RT-PCR from total RNA was used to amplify TAF1-4 (first [top] panel), the unique 3' end of TAF1 isoforms (second panel), the common 5' end of TAF1 isoforms (third panel), or actin 5C (bottom panel). PCR products were resolved on 1.2% agarose gels. TAF1-2 and TAF1-3 products were not resolved by this assay. The positions of PCR products are indicated on the right.

 

Figure 8
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  		FIG. 8. CHK1 is necessary for CPT-induced upregulation of TAF1-3 and TAF1-4. qPCR from cells mock treated or pretreated with CHK1 or CHK2 dsRNA 3 days before treatment with DMSO or 20 µM CPT for 6 h (n = 9). CHK1 and CHK2 levels indicated by a pound sign were significantly lower than mock-treated cells (P < 0.05). TAF1 isoform levels indicated by an asterisk were significantly higher than mock-treated cells, and TAF1 isoform levels indicated by a pound sign were significantly lower than CPT-treated cells (P < 0.05). CHK2 RNAi reduced the IR-induced upregulation of TAF1-3 and TAF1-4 but did not meet the statistical criteria for significance. Error bars represent the standard errors of the means.

 

Figure 6
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  		FIG. 6. Caffeine abrogated both CPT- and IR-induced upregulation of TAF1-3 and TAF1-4, but wortmannin abrogated only the IR effect. (A) qPCR of RNA from cells treated with 20 mM caffeine (caff), 1 µM wortmannin (wort), 20 µM CPT, or combinations as indicated for 6 h (n = 3). (B) qPCR of RNA cells treated with 20 mM caffeine, 1 µM wortmannin, 40 Gy IR, or combinations as indicated, then allowed to recover for 3 h (n = 6). TAF1 isoform levels indicated by an asterisk were significantly higher than mock-treated cells, and levels indicated by a pound sign were significantly lower than CPT- or IR-treated cells (P < 0.05). Error bars represent the standard errors of the means.

 

Figure 5
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  		FIG. 5. CPT-induced upregulation of TAF1-3 and TAF1-4 was abrogated by actinomycin D but not cycloheximide treatment. (A) qPCR of RNA from cells pretreated with 25 ng/ml ActD 2 h prior to treatment with 20 µM CPT for 2, 4, 8, or 16 h (n = 3). (B) qPCR of RNA from cells cotreated with 100 µg/ml cycloheximide (CHX) and 20 µM CPT for 6 h (n = 3). TAF1 isoform levels indicated by an asterisk were significantly higher than mock-treated cells (P < 0.05). Error bars represent the standard errors of the means.

 
RNAi. PCR, using Drosophila melanogaster genomic DNA as a template, was performed to generate RNAi plasmids for ATM, ATR, CHK1, and CHK2. Primer sets (oriented 5' to 3') were as follows: ATM, GTGCTTTGGCACTTGGAGACG and GGAGTGGCGTAGTCCTTGTC; ATR, CCTACTTCGAGAGCTGCCTAAGTGAACCG and GTAACTTGCATCGATGGCTCCCAGTTCTCC; CHK1, GCCAAAGGCTAAGAGGCAGC and GAAGGTGATGCATGAGTTGC; and CHK2, CCTTTGTATTCAAGGATCTCAGCC and CCCGATCGTGTAGGTACTTG. PCR products for ATM (nucleotides [nt] 1063 to 1910 of CG6535-RA), ATR (nt 2903 to 3651 of CG4252-RA), CHK1 (nt 1437 to 2144 of CG17161-RC), and CHK2 (nt 804 to 1241 of CG10895-RC) were cloned in both orientations into the pCRII-TA vector (Invitrogen). DNA linearized at the BamHI site was used as a template to generate single-stranded RNA (ssRNA) with a Ribomax RNA Production T7 kit (Promega). ssRNA was resuspended in annealing buffer containing 5 mM KCl and 10 mM NaH2PO4, and complementary ssRNAs were annealed by heating to 95°C for 5 min and slow cooling 16 to 17 h to generate double-stranded RNA (dsRNA). RNAi was performed by plating 2 x 106 cells for 1 h, removing the serum-containing media, washing the cells with serum-free media, and adding 2 ml serum-free media. Twenty micrograms dsRNA was added per dish and mixed by swirling. After 30 min, 4 ml of serum-containing media was added. The RNAi screen of RNA-binding proteins was carried out using the same protocol except that ssRNAs were generated from plasmid-templated PCR products with Ribomax RNA Production T7 and SP6 kits (Promega), as described by Park et al. (37). Cells were treated with CPT or ionizing radiation (IR) 3 days after addition of dsRNA.

Quantitation and statistics. The cDNA of interest was measured relative to actin 5C by the formula EactinCt(actin 5C)/EtargetCt(target). E is an empirically derived PCR efficiency factor, and Ct is the threshold value for amplification (38). E values were as follows: actin 5C, 2.07; total TAF1, 1.98; TAF1-1, 2.16; TAF1-2, 2.18; TAF1-3, 2.00; TAF1-4, 2.18; ATM, 1.95; ATR, 2.15; CHK1, 2.04; CHK2, 2.00; and dU2AF38, 2.03. One-way analysis of variance was performed in conjunction with Fisher's protected least significant difference with a type I error, {alpha}, of 0.05. A difference greater than Fisher's protected least significant difference was labeled significant. Separate cell cultures were used as independent measurements.

TAF1 immunoprecipitation. Influenza A virus hemagglutinin (HA) epitope-tagged TAF1-4 protein was expressed via the copper-inducible expression vector pRmHa-4 (13). The TAF1-4HA expression plasmid encodes amino acids 1 to 658 of GenBank accession number A47371 linked in frame to amino acids 656 to 2098 of CG17603-CG17603-PC and a carboxy-terminal HA tag (YPYDVPDYA). For stable expression, cells were cotransfected with 0.9 µg of TAF1-4HA expression plasmid and 0.1 µg of a puromycin resistance plasmid (6). After 2 days, cells were selected with 10 µg/ml puromycin (Sigma). More than 99% of mock-transfected cells were dead after 3 days of selection. Induction of TAF1-4HA expression by 500 µM CuSO4 was still observed after 1 month of selection and from freezer stocks of stable S2 cells. Immunoprecipitation of TAF1 was performed from nuclear extracts of uninduced or copper-induced stable S2 cells (16). Immunoprecipitations were carried out as described previously (30), except that the wash buffer contained 500 mM NaCl and 0.1% NP-40 and the following antibodies: DEAE-purified rabbit control immunoglobulin G (IgG; 1:30), TAF1-M (1:30) (30), or affinity-purified rabbit polyclonal anti-HA (1:1,000; Sigma). Western blot analysis was carried out as described previously (55) using antibodies at the following dilutions: TAF1-M, 1:5,000; TAF1(30H9), 1:50 (58); TBP, 1:200 (Santa Cruz Biotechnology); and HA(12CA5), 1:4,000 (Roche).

Flow cytometry. Cells were prepared for flow cytometry analysis by washing 1 x 106 cells twice with phosphate-buffered saline (PBS) and resuspending them in 2 ml PBS, 0.25% Triton X-100, 33 µg/ml propidium iodide, and 50 µg/ml RNase A. Stained cells were sorted with a Becton Dickinson 488-nm laser excitation FACScan machine using CELLQuest software. Data were analyzed by ModFit software. Cells were prepared for fluorescence-activated cell sorting (FACS) by adding 20 µg/ml Hoechst dye to 5.5 x 105 cells/ml and incubating at 37°C for 30 min. Cells were concentrated to 1.0 x 107 cells/ml and run through a 40-µm-pore nylon mesh, and propidium iodide was added to 33 µg/ml. Cells were sorted into G0/G1, S, and G2/M phases with a triple-laser FACSVantage SE using the FACSDiVa digital electronics package. Sorted cells were examined on aFACScan instrument to confirm phase sorting. All FACS analysis was performed at the University of Wisconsin Comprehensive Cancer Center Flow Cytometry Facility.


   RESULTS AND DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results and Discussion
References
 
The TAF1 pre-mRNA is alternatively spliced in D. melanogaster. Comparison of published D. melanogaster TAF1 cDNA sequences to each other and to the TAF1 gene revealed differences that were explainable by alternative splicing of two exons, 12a and 13a (Fig. 1A) (27, 58). Putative exon 12a and 13a splice sites conform to consensus sequences for D. melanogaster and are conserved in eight Drosophila species, which share a common ancestor ~50 million years ago (data not shown) (26). To follow up on these observations, we examined expression of TAF1 mRNAs in adult flies by RT-PCR with oligonucleotide primers complementary to TAF1 exons 12 and 14. Sequencing of clones derived from the RT-PCR products indicated that the TAF1 pre-mRNA is alternatively spliced to produce four mRNAs, TAF1-1 (including neither alternative exon), TAF1-2 (including alternative exon 12a), TAF1-3 (including alternative exon 13a), and TAF1-4 (including alternative exons 12a and 13a).


Figure 1
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  		FIG. 1. TAF1 pre-mRNA is alternatively spliced. (A) Schematic diagram of the TAF1 gene in D. melanogaster. Boxes, exons; lines, introns. Alternative exons 12a and 13a are gray and black, respectively. Below are schematic diagrams of the 3' ends of TAF1-1, TAF1-2, TAF1-3, and TAF1-4. At the bottom is a comparison of exon 12a and exon 13a polypeptide sequences from D. melanogaster and Drosophila virilis. The AT hook motif is underlined, and the dash indicates a gap. (B) qPCR analysis of TAF1 mRNA isoform levels from fly tissues and S2 cells. Graphed are TAF1 isoform/actin 5C levels normalized to the sum of all four TAF1 isoform/actin 5C measurements. Error bars represent the standard errors of the means of three independent experiments.

 
TAF1 isoforms are differentially expressed in fly tissues. To understand the molecular role of TAF1 protein isoforms, we examined the sequences of the alternative-exon-encoded polypeptides. Exon 13a encodes a 31-amino-acid polypeptide with no obvious functional motifs, but exon 12a encodes a 33-amino-acid polypeptide that includes a 9-amino-acid DNA-binding motif called an AT hook (Fig. 1A) (3). AT hooks are present in a variety of nonhistone chromatin-associated proteins but are best characterized in a subgroup of high mobility group (HMG) proteins called HMGA [formerly HMG-I(Y)] (40). HMGA proteins have three AT hook motifs, which preferentially bind the minor groove of adenine-thymine-rich DNA. Since an AT hook is encoded by constitutive exon 12, proteins encoded by TAF1-1 and TAF1-3 are predicted to contain one AT hook whereas proteins encoded by TAF1-2 and TAF1-4 are predicted to contain two AT hooks separated by a 14-amino-acid spacer. Our studies of recombinant TAF1 proteins indicate that two AT hooks are required for DNA binding and that TAF1-2 binds with variable affinity the core promoters of several Drosophila genes (33). These data suggest that alternative splicing is a mechanism to regulate gene-specific transcription through differential DNA-binding properties of TAF1. A prediction that stems from this proposal is that expression of TAF1 isoforms should differ between tissues with different gene expression profiles.

To test the prediction, we used qPCR to quantitate TAF1 mRNA isoform levels in fly tissues. Oligonucleotide primer sets spanning exon boundaries were designed to uniquely detect individual TAF1 mRNA isoforms. This analysis revealed that TAF1 mRNA isoform levels varied greatly between tissues (Fig. 1B). TAF1-3 was the most abundant isoform in all tissues examined, with the exception of testis, where TAF1-2 was most abundant. TAF1-4 levels were higher in adult heads than other tissues. Finally, TAF1-1 was the most abundant isoform in embryonically derived S2 cultured cells. These findings suggest that tissue-specific signals regulate TAF1 pre-mRNA alternative splicing.

Differences in TAF1 isoform expression in tissues with unique cell cycle programs, such as testes, ovaries, and salivary glands, and the requirement of TAF1 for G2/M phase cell cycle progression in S2 cells suggested that TAF1 alternative splicing might be regulated during the cell cycle (30, 51). To examine the extent to which cell cycle-dependent signals regulate TAF1 alternative splicing, we examined S2 cells in more detail. Based on DNA content, asynchronously growing S2 cells were separated into G0/G1, S, and G2/M phases by fluorescence-activated cell sorting. The quality of the cell cycle phase separation was confirmed by subsequent flow cytometry analysis of the samples (data not shown). qPCR analysis revealed that TAF1 isoform levels did not change during the cell cycle (data not shown). Thus, differences in TAF1 isoform levels between S2 cells and fly tissues appear to be independent of normal cell cycle-regulatory signals.

The identification of multiple TAF1 isoforms in Drosophila adds to a growing list of TAF proteins (4, 53). In Drosophila, some TAFs are encoded by two genes; one is ubiquitously expressed, and the other is specifically expressed in the testis. Testis-specific TAFs are important for regulating the unique transcriptional program that occurs during spermatogenesis (19, 20). Interestingly, we found that, relative to other tissues, testes express a unique profile of TAF1 isoforms. Most notably, TAF1-2 is the most abundant isoform in testes, but not in any other tissue that was examined. This finding suggests that TAF1-2, which binds DNA through two AT hooks, contributes to the mechanism of gene-specific transcription during spermatogenesis.

TAF1 protein isoforms may be differentially expressed in Drosophila tissues and are incorporated into TFIID. The identification of four TAF1 mRNA isoforms raised the question of whether the encoded TAF1 protein isoforms are expressed in flies. To address this issue, we performed Western blot analysis of fly tissue extracts with a TAF1 antibody (anti-TAF1-M) that recognized a region common to all TAF1 protein isoforms (30). This analysis revealed a comigrating protein in S2 cells, ovaries, testes, and heads (Fig. 2A). In addition, a second faster- or slower-migrating protein was observed in testes and heads, respectively. In the absence of isoform-specific antibodies, we are unable to definitively identify any of the proteins. But, based on mRNA abundance, the faster-migrating protein in testes is likely TAF1-2 and the slower-migrating protein in heads is likely TAF1-4.


Figure 2
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  		FIG. 2. TAF1 protein isoforms are expressed and associate with TBP. (A) Western blot (WB) analysis of S2 cell, ovary, and head nuclear extracts and testis whole-cell extract probed with anti-TAF1-M antibody, which recognized a region of TAF1 common to all isoforms. TAF1 bands were aligned based on other experiments in which S2 cell extracts were analyzed alongside ovary, testis, or head extracts. The asterisk indicates a potential cross-reacting band that was not observed with anti-TAF1-C antibody, which was raised against a polypeptide encoded by exons 12 to 14 of TAF1-4 (data not shown) (30). (B) Western blot analysis of S2 cell nuclear extracts for coimmunoprecipitation of TBP with TAF1-4. Nuclear extracts were prepared from uninduced (–CuSO4; lanes 1 to 3) or induced (+CuSO4; lanes 4 to 6) S2 cells stably transfected with a TAF1-4HA expression vector. Extracts were then used for immunoprecipitation using a control mouse IgG antibody (IgG) or a rabbit polyclonal antibody to HA (HA). Lanes 1 and 4 contained ~1% of the amount of extract used for the immunoprecipitations. Immunoprecipitates were probed with a mouse monoclonal antibody to TAF1 [{alpha}-TAF1(30H9); top left] or an antibody to TBP ({alpha}-TBP; bottom left). Western blot analysis of total extract (lanes 7 and 8) with an antibody to TAF1 showed that TAF1-4HA was expressed at a low level ({alpha}-TAF1-M; top right) and with an antibody to HA showed that TAF1-4HA was expressed only in the presence of copper ({alpha}-HA; bottom right). Collectively, these experiments showed that TBP coimmunoprecipitated with TAF1-4HA only when TAF1-4HA was expressed and only with an antibody that recognized TAF1-4HA. (C) Western blot analysis of S2 cell nuclear extracts for coimmunoprecipitation of endogenous TBP with TAF1. Nuclear extracts were prepared from S2 cells and used for immunoprecipitation using a control mouse IgG antibody (IgG) or anti-TAF1-M antibody. Lanes 1 and 4 contained ~1% of the amount of extract used for the immunoprecipitations. Immunoprecipitates were probed with a mouse monoclonal antibody to TAF1 [{alpha}-TAF1(30H9); left] or an antibody to TBP ({alpha}-TBP; right). This experiment showed that TBP coimmunoprecipitated with endogenous TAF1. Positions of TAF1 and TBP are indicated on the right, and positions of protein molecular mass markers are indicated on the left.

 
To determine the extent to which TAF1 isoforms associate with TFIID, we examined the association of TAF1-4 with TBP. A stable S2 cell line was generated that expressed HA epitope-tagged TAF1-4 (TAF1-4HA) under the control of a copper-inducible promoter. Western blot analysis of HA immunoprecipitates from induced-cell extracts showed that TAF1-4HA associated with TBP (Fig. 2B). Similarly, Western blot analysis of TAF1 immunoprecipitates from untransfected S2 cells showed that endogenous TAF1 associated with TBP (Fig. 2C). Presumably, endogenous TAF1 is predominantly TAF1-1, as this was the most abundant TAF1 mRNA isoform (Fig. 1B). Thus, TAF1 isoforms with or without alternative-exon-encoded polypeptides can be incorporated into TFIID complexes.

DNA damage signals regulate TAF1 pre-mRNA alternative splicing. With the goal of understanding signaling events that regulate TAF1 alternative splicing, we chose to examine TAF1 splicing in S2 cells exposed to DNA-damaging agents. S2 cells were treated with CPT, which inhibits DNA topoisomerase I to disrupt DNA replication forks, leading to a variety of DNA lesions (14, 18, 39). Cells were treated with either 20 µM CPT or vehicle (DMSO) and incubated for 2 to 16 h. Total RNA was isolated from the cells, and RT-PCR, using oligonucleotide primers to exons 12 and 14, was used to detect all four TAF1 mRNA isoforms in a single reaction. actin 5C mRNA level was used as a normalization control for this and other experiments. Fractionation of PCR products by agarose gel electrophoresis revealed that CPT induced TAF1-4 alternative splicing after 6 h (Fig. 3, lanes 1 to 8). Induction of TAF1-4 was confirmed with isoform-specific primers. This finding is consistent with that of Bell et al. (5), who showed that human TAF6{delta} alternative splicing is induced by treatment of HL-60 cells with 15 µM CPT for 6 h (5). CPT also induced alternative splicing of TAF1-2 and/or TAF1-3 with the same kinetics as TAF1-4. The ambiguity in interpreting this result is due to the fact that the TAF1-2 and TAF1-3 products comigrated in the agarose gel system. Finally, TAF1-1 levels were reduced by CPT treatment. Since CPT did not alter total TAF1 levels, these findings suggest that DNA damage induces a signal that regulates splicing inclusion of exons 12a and 13a.

To determine if alternative splicing of TAF1 is a general response to DNA damage, we exposed S2 cells to IR, which causes double-strand breaks and other replication-independent DNA damage. Cells were exposed to 40 Gy of IR and then allowed to recover for 1 or 3 h. Similar to CPT, IR induced alternative splicing of TAF1-4 and TAF1-2-TAF1-3 and reduced splicing of TAF1-1 (Fig. 3, lanes 9 to 11). Thus, different types of DNA damage had the same effect on TAF1 alternative splicing, suggesting that DNA damage affects the function of a common component of the TAF1 pre-mRNA splicing machinery in S2 cells.

Alternative splicing of TAF1-3 and TAF1-4 is induced by DNA damage. To quantitate the effect of DNA damage on TAF1 alternative splicing and to detect individual isoforms, we turned to the qPCR assay. To examine the time dependence of TAF1 alternative splicing, S2 cells were treated with 20 µM CPT and processed at various times over a 24-h period or treated with 40 Gy of IR and processed over a 3-h recovery period. This analysis revealed that TAF1-3 and TAF1-4 levels significantly increased with similar patterns of time dependence in response to both CPT and IR (Fig. 4A and B). Changes in TAF1 alternative splicing were not due to a general perturbation of splicing, as CPT or IR treatment did not affect TAF4 alternative splicing (data not shown). Thus, induction of TAF1-3 and TAF1-4 alternative splicing appears to be a coordinately regulated response to DNA damage.


Figure 4
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  		FIG. 4. CPT and IR upregulated TAF1-3 and TAF1-4 in a time- and dose-dependent manner. (A) qPCR of RNA from cells treated with DMSO or 20 µM CPT for the times indicated (n = 6). (B) qPCR of RNA from cells treated with 40 Gy IR and then allowed to recover for the times indicated (n = 9). (C) qPCR of RNA from cells treated with the indicated concentrations of CPT for 6 h (n = 2). (D) qPCR of RNA from cells treated with the indicated levels of IR and then allowed to recover for 3 h (n = 6). TAF1 isoform levels, indicated by an asterisk, were significantly greater than mock-treated cells (P < 0.05). Error bars represent the standard errors of the means.

 
To examine the dose dependence of TAF1 alternative splicing, S2 cells were treated with various doses of CPT ranging from 0 to 40 µM and processed after 6 h or with various doses of IR ranging from 0 to 80 Gy and processed after a 3-h recovery period. Dose-dependent increases in TAF1-3 and TAF1-4 splicing were observed in response to both CPT and IR (Fig. 4C and D). Significant increases in TAF1-3 and TAF1-4 occurred at and above 20 and 10 µM CPT, respectively, or with 40 Gy of IR. The increases in TAF1-3 and TAF1-4 levels were compensated by a decrease in TAF1-1 level, and TAF1-2 levels did not significantly change over the dose response of IR treatment (data not shown). Thus, induction of TAF1-3 and TAF1-4 alternative splicing is sensitive to the level of DNA damage.

Transcription but not protein synthesis is required for TAF1 alternative splicing. Since total TAF1 levels were not altered by DNA damage, the change in TAF1 isoform levels was likely due to alternative splicing of newly transcribed TAF1 pre-mRNAs rather than altered stability of existing, mature TAF1 mRNAs. To examine this proposal, cells were treated with the transcription inhibitor ActD for 2 h prior to addition of CPT. As shown in Fig. 5A and B, ActD abrogated CPT-induced upregulation of TAF1-3 and TAF1-4 but ActD treatment alone did not affect TAF1-3 or TAF1-4 levels. This suggested that the change in steady-state levels of TAF1 isoforms was due to transcription and splicing rather than regulated decay of mature mRNAs. Furthermore, splicing of TAF1-3 and TAF1-4 occurred independently of new protein synthesis, as CPT-induced splicing was not affected by cotreatment of S2 cells with the protein synthesis inhibitor cycloheximide, which in parallel experiments was shown to block translation but not transcription of transiently transfected genes (Fig. 5C and data not shown). Thus, we hypothesize that, in response to DNA damage, posttranslational modification of signaling molecules and splicing factors regulates the splicing of TAF1-3 and TAF1-4.

PIKK family members signal TAF1 alternative splicing. The change in TAF1 alternative splicing in response to DNA damage suggested the involvement of ATM and ATR protein kinases (1, 24, 46). To examine this proposal, we employed small-molecule kinase inhibitors. Caffeine inhibits the kinase activity of mammalian PIKK family members, such as ATM and ATR, at low millimolar concentrations, whereas wortmannin inhibits the kinase activity of mammalian ATM but not ATR at low micromolar concentrations (2, 42, 43). S2 cells were treated with 20 µM CPT, 20 mM caffeine, or both CPT and caffeine, and TAF1 isoform levels were determined by qPCR. This analysis revealed that caffeine inhibited CPT-induced upregulation of TAF1-3 and TAF1-4, indicating that PIKK family activity is required for CPT induction of TAF1 alternative splicing (Fig. 6A). Similarly, S2 cells were treated with 20 µM CPT, 1 µM wortmannin, or both CPT and wortmannin. Unlike caffeine, wortmannin did not inhibit the effect of CPT on TAF1-3 and TAF1-4 splicing. These findings suggest that ATR-like kinase activity is required for transducing the CPT signal to the TAF1 splicing machinery.

Similar to the effect on CPT-induced TAF1 splicing, caffeine inhibited IR-induced TAF1-3 and TAF1-4 splicing (Fig. 6B). But whereas wortmannin did not inhibit CPT-induced TAF1-3 and TAF1-4 splicing, it did inhibit IR-induced TAF1-3 and TAF1-4 splicing. These findings suggested that ATM-like kinase activity is required for transducing the IR signal to the TAF1 splicing machinery.

ATR and ATM are required for TAF1 alternative splicing in response to DNA damage. To directly test the requirement for ATR in CPT-induced TAF1-4 alternative splicing, RNAi was used to reduce ATR expression in S2 cells. S2 cells were incubated with ATR or, as a control, ATM dsRNA for 3 days. Cells were then treated with 20 µM CPT for 6 h, and TAF1 isoform levels were determined by qPCR. RNAi reduced ATR and ATM mRNA levels to ~30% of normal levels, but only the reduction in ATR had a significant inhibitory affect on CPT-induced TAF1-3 and TAF1-4 upregulation (Fig. 7A). In contrast, ATM knockdown abrogated IR-induced TAF1-3 and TAF1-4 upregulation, but ATR knockdown had no effect (Fig. 7B). Thus, ATR is necessary for CPT-induced TAF1-3 and TAF1-4 alternative splicing and ATM is necessary for IR-induced TAF1-3 and TAF1-4 alternative splicing.


Figure 7
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  		FIG. 7. ATR is necessary for CPT-induced upregulation of TAF1-3 and TAF1-4, and ATM is necessary for IR-induced upregulation of TAF1-3 and TAF1-4. (A) qPCR of RNA from cells mock treated or pretreated with ATR or ATM dsRNA 3 days before treatment with DMSO or 20 µM CPT for 6 h (n = 9 for the ATR experiments and n = 3 for the ATM experiments). (B) qPCR of RNA from cells mock treated or pretreated with ATR or ATM dsRNA 3 days before treatment with 40 Gy IR and a 3-h recovery (n = 3). ATR and ATM levels indicated by a pound sign were significantly lower than mock-treated cells, TAF1 isoform levels indicated by an asterisk were significantly higher than mock-treated cells, and TAF1 isoform levels indicated by a pound sign were significantly lower than CPT- or IR-treated cells (P < 0.05). Error bars represent the standard errors of the means.

 
Checkpoint kinases are required for TAF1 alternative splicing in response to DNA damage. To examine the extent to which the canonical ATR signaling pathway regulates TAF1 alternative splicing in response to DNA damage, we examined the requirement for downstream checkpoint kinases CHK1 (also known as Grapes) and CHK2 (also known as Loki or MNK) (11). S2 cells were incubated with CHK1 or CHK2 dsRNA for 3 days. Cells were then treated with 20 µM CPT for 6 h, and TAF1 isoform levels were determined by qPCR (Fig. 8). RNAi reduced CHK1 and CHK2 mRNAs to ~25% of normal levels. As with ATM and ATR RNAi, CHK1 and CHK2 RNAi had no effect on TAF1 alternative splicing in the absence of DNA damage. However, reduced levels of CHK1, but not CHK2, significantly inhibited CPT-induced TAF1-3 and TAF1-4 splicing. The analogous experiment with IR did not show a statistically significant effect, at P = 0.05, of CHK1 or CHK2 RNAi on TAF1-3 or TAF1-4 splicing. However, as shown in Fig. 8, we consistently observed that reduced levels of CHK2, but not CHK1, inhibited IR-induced TAF1 alternative splicing. Thus, the ATR/CHK1 and ATM/CHK2 pathways appear to be required for transducing the DNA damage signal to the TAF1 pre-mRNA splicing machinery.

DNA damage signals may inhibit dU2AF38 function to regulate TAF1 alternative splicing. As an unbiased approach to identify RNA-binding proteins involved in regulating TAF1 pre-mRNA alternative splicing, we utilized an RNAi library, developed by Park et al. (37). RNAi knockdown of 15 of the 243 genes examined resulted in an increase or a decrease in TAF1-3 and/or TAF1-4 splicing. One of the identified RNA-binding proteins, Drosophila U2 auxiliary factor 38 (dU2AF38), was examined in more detail. Drosophila U2AF is a heterodimer composed of small (dU2AF38) and large (dU2AF50) subunits, each of which contains an arginine-serine-rich (RS) domain that promotes high-affinity binding to the intron pyrimidine tract between the branch point and the 3' splice site and targets the U2 small nuclear ribonucleoprotein particle to the branch site at an early step in spliceosome assembly (41). The small U2AF subunit has been shown to modulate alternative splicing in Drosophila and human cells (36, 37).

As shown in Fig. 9, reducing the level of dU2AF38 resulted in a significant increase in TAF1-3 splicing, suggesting that dU2AF38 functions as a silencer of exon 13a inclusion. Furthermore, treatment of dU2AF38 RNAi cells with IR or CPT did not significantly increase TAF1-3 levels relative to IR or CPT treatment alone. These data place dU2AF38 in the IR and CPT DNA damage pathways and suggest that these pathways inhibit dU2AF38 function.


Figure 9
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  		FIG. 9. Knockdown of dU2AF38 by RNAi resulted in increased TAF1-3 splicing. The left panel shows that dU2AF38 levels in RNAi-treated cells indicated by a pound sign were significantly lower than mock-treated cells (P < 0.05) (n = 6). The right panel shows that TAF1-3 levels indicated by an asterisk or alpha were upregulated by all of the treatments examined relative to mock-treated cells (P < 0.05 or P < 0.062, respectively). Error bars represent the standard errors of the means.

 
Collectively, these findings suggest a new role for the ATR/CHK1 and ATM/CHK2 signaling pathways in regulating alternative splicing. Our data support a model in which signals induced by DNA damage alter the regulation of TAF1 pre-mRNA alternative splicing. In response to DNA damage, ATR or ATM is activated and phosphorylates CHK1 or CHK2, respectively, which in turn may phosphorylate alternative splicing factors to modulate their activity and promote TAF1-3 and TAF1-4 splicing (Fig. 10). In support of this model, we have found that RNAi knockdown of RNA-binding proteins, including dU2AF38, is sufficient to upregulate TAF1-3 and/or TAF1-4 expression. The link between CHK1/CHK2 and alternative splicing factors with RNA-binding activity may not be direct. For example AKT, a protein kinase that plays an important role in cell survival, may be involved in the TAF1 alternative splicing. ATM mediates full activation of AKT in response to IR, and AKT regulates the function of SR alternative splicing factors by phosphorylation of the RS domain (9, 54). The challenge now is to understand how the ATR/CHK1 and ATM/CHK2 DNA damage signaling pathways modulate the function of the splicing machinery to regulate TAF1 pre-mRNA alternative splicing.


Figure 10
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  		FIG. 10. Model for the transduction of DNA damage signals to the splicing machinery. Our data suggest that distinct DNA damage signaling pathways induced by CPT or IR converge on a common set of splicing factors that function to upregulate TAF1-3 and TAF1-4 alternative splicing. Experiments with pharmacological inhibitors indicate that ATR and ATM kinase activities are necessary to signal alternative splicing of the TAF1 pre-mRNA in response to CPT or IR, respectively. RNAi experiments indicate the requirement for ATR/CHK1 and ATM/CHK2 to signal alternative splicing of the TAF1 pre-mRNA in response to CPT and IR, respectively. CHK2 is indicated in parentheses and in gray lettering because the data are suggestive of a role in the pathway. The CPT and IR pathways may converge on common splicing factors, including dU2AF38, to elicit the same effect on TAF1 pre-mRNA splicing.

 


   ACKNOWLEDGMENTS
 
We thank J. Park and B. Graveley for providing RNAi plasmids for Drosophila RNA-binding proteins, T. Kokubo and R. Tjian for providing TAF1 plasmids and antibodies, and the University of Wisconsin Comprehensive Cancer Center Flow Cytometry Facility for assistance with flow cytometry. We are grateful to D. Brow, C. Metcalf, S. Rimkus, and R. Tibbetts for advice on experiments.

This work was supported by National Institutes of Health grant GM066204 (to D.A.W.) and by National Institutes of Health training grant T32 GM08688 and a University of Wisconsin Prize Fellowship (to M.S.M.).


   FOOTNOTES
 
* Corresponding author. Mailing address: University of Wisconsin School of Medicine and Public Health, Department of Pharmacology, 1300 University Avenue, Madison, WI 53706. Phone: (608) 262-6648. Fax: (608) 262-1257. E-mail: dawassarman{at}wisc.edu. Back

{triangledown} Published ahead of print on 9 October 2006. Back

{dagger} These authors contributed equally to the paper. Back


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 Materials and Methods
 Results and Discussion
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Molecular and Cellular Biology, December 2006, p. 9256-9267, Vol. 26, No. 24
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J. Bacteriol. J. Virol. Eukaryot. Cell