Inhibition of the mammalian transcription factor LSF induces S-phase-dependent apoptosis by downregulating thymidylate synthase expression
2000; Springer Nature; Volume: 19; Issue: 17 Linguagem: Inglês
10.1093/emboj/19.17.4665
ISSN1460-2075
Autores Tópico(s)RNA modifications and cancer
ResumoArticle1 September 2000free access Inhibition of the mammalian transcription factor LSF induces S-phase-dependent apoptosis by downregulating thymidylate synthase expression Christina M.H. Powell Christina M.H. Powell Committee on Virology and Department of Microbiology and Molecular Genetics, Harvard Medical School and Division of Molecular Genetics, Dana-Farber Cancer Institute, 44 Binney Street, Boston, MA, 02115 USA Present address: Department of Biology, Boston University, 5 Cummington Street, Boston, MA, 02215 USA Search for more papers by this author Thomas L. Rudge Thomas L. Rudge Department of Molecular Genetics, The Ohio State University, Columbus, OH, 43210 USA Search for more papers by this author Quan Zhu Quan Zhu Committee on Virology and Department of Microbiology and Molecular Genetics, Harvard Medical School and Division of Molecular Genetics, Dana-Farber Cancer Institute, 44 Binney Street, Boston, MA, 02115 USA Present address: IntraImmune Therapies, Inc., PO Box 15599, Boston, MA, 02215-0011 USA Search for more papers by this author Lee F. Johnson Lee F. Johnson Department of Molecular Genetics, The Ohio State University, Columbus, OH, 43210 USA Search for more papers by this author Ulla Hansen Corresponding Author Ulla Hansen Committee on Virology and Department of Microbiology and Molecular Genetics, Harvard Medical School and Division of Molecular Genetics, Dana-Farber Cancer Institute, 44 Binney Street, Boston, MA, 02115 USA Present address: Department of Biology, Boston University, 5 Cummington Street, Boston, MA, 02215 USA Search for more papers by this author Christina M.H. Powell Christina M.H. Powell Committee on Virology and Department of Microbiology and Molecular Genetics, Harvard Medical School and Division of Molecular Genetics, Dana-Farber Cancer Institute, 44 Binney Street, Boston, MA, 02115 USA Present address: Department of Biology, Boston University, 5 Cummington Street, Boston, MA, 02215 USA Search for more papers by this author Thomas L. Rudge Thomas L. Rudge Department of Molecular Genetics, The Ohio State University, Columbus, OH, 43210 USA Search for more papers by this author Quan Zhu Quan Zhu Committee on Virology and Department of Microbiology and Molecular Genetics, Harvard Medical School and Division of Molecular Genetics, Dana-Farber Cancer Institute, 44 Binney Street, Boston, MA, 02115 USA Present address: IntraImmune Therapies, Inc., PO Box 15599, Boston, MA, 02215-0011 USA Search for more papers by this author Lee F. Johnson Lee F. Johnson Department of Molecular Genetics, The Ohio State University, Columbus, OH, 43210 USA Search for more papers by this author Ulla Hansen Corresponding Author Ulla Hansen Committee on Virology and Department of Microbiology and Molecular Genetics, Harvard Medical School and Division of Molecular Genetics, Dana-Farber Cancer Institute, 44 Binney Street, Boston, MA, 02115 USA Present address: Department of Biology, Boston University, 5 Cummington Street, Boston, MA, 02215 USA Search for more papers by this author Author Information Christina M.H. Powell1,2, Thomas L. Rudge3, Quan Zhu1,4, Lee F. Johnson3 and Ulla Hansen 1,2 1Committee on Virology and Department of Microbiology and Molecular Genetics, Harvard Medical School and Division of Molecular Genetics, Dana-Farber Cancer Institute, 44 Binney Street, Boston, MA, 02115 USA 2Present address: Department of Biology, Boston University, 5 Cummington Street, Boston, MA, 02215 USA 3Department of Molecular Genetics, The Ohio State University, Columbus, OH, 43210 USA 4Present address: IntraImmune Therapies, Inc., PO Box 15599, Boston, MA, 02215-0011 USA *Corresponding author. E-mail: [email protected] The EMBO Journal (2000)19:4665-4675https://doi.org/10.1093/emboj/19.17.4665 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info The thymidylate synthase (TS) gene, which is induced at the G1–S transition in growth-stimulated cells, encodes an enzyme that is essential for DNA replication and cell survival. Here we demonstrate that LSF (LBP-1c, CP2) binds to sites within the TS promoter and intronic regions that are required for this induction. Mutation of the LSF binding sites inhibits G1–S induction of mRNA derived from a TS minigene. Furthermore, expression of dominant-negative LSF (LSFdn) prevents the increase in TS enzyme levels during G1–S, and induces apoptosis in growth- stimulated mouse and human cell lines. Such apoptosis can be prevented either by circumventing the TS requirement through addition of low concentrations of thymidine, or by coexpression of the TS gene driven by a heterologous promoter. Induction of apoptosis by LSFdn parallels the process known as thymineless death, which is induced by the TS inhibitor and chemotherapeutic drug 5-fluorodeoxyuridine. Thus, LSF is a novel regulatory factor that supports progression through S-phase by targeting a single gene that is critical for cell survival. Introduction Transcription factors integrate environmental signals and genetic information to control cell cycle progression. As cells reach the restriction point in late G1 phase, the decision is made either to progress to DNA replication or revert to quiescence. If cells continue through the cell cycle, preparation for DNA replication requires changes in the pattern of gene expression, including activation of genes that encode enzymes necessary for deoxynucleotide biosynthesis and DNA replication. While key cell cycle regulators that function at the restriction point, such as the G1 cyclins, have been identified, the identity of transcriptional regulators that link cell growth control to transcriptional regulation at the G1–S transition have not yet been fully elucidated. There are at least two parallel regulatory pathways that drive progression into S-phase, one involving cyclin D–kinase complexes (Matsushime et al., 1992; Xiong et al., 1992; Baldin et al., 1993) and the other involving a cyclin E–kinase complex (Dulic et al., 1992; Koff et al., 1992; Resnitzky et al., 1994; Ohtsubo et al., 1995; Resnitzky and Reed, 1995; Lukas et al., 1997). Until now, the major transcriptional regulator known to function in late G1 is E2F, which is directly targeted by the cyclin D pathway (Johnson, 1995; Nevins et al., 1997; Dyson, 1998). It is likely that there are additional regulatory factors that are critical during the G1–S transition. The ability of the mammalian transcription factor, LSF, to transactivate a DNA tumor virus promoter (Kim et al., 1987), as does E2F, suggested a potential role for LSF in cell growth control. LSF (Shirra et al., 1994), a 63 kDa protein also known as LBP-1c (Yoon et al., 1994) and CP2 (Lim et al., 1992), stimulates gene expression from the Simian Virus 40 (SV40) major late promoter (Kim et al., 1987; Huang et al., 1990), which is coordinated with the onset of host cell DNA replication (Acheson, 1976). The consensus DNA-binding site for LSF consists of two direct repeats, with a center-to-center spacing of 10 bp corresponding to adjacent turns of the DNA helix (Lim et al., 1993; Yoon et al., 1994; Shirra, 1995). While LSF is primarily dimeric in solution, it binds its DNA site as an obligate tetramer (Murata et al., 1998; Shirra and Hansen, 1998). Furthermore, LSF is ubiquitously expressed in mouse embryos and adult tissues (Swendeman et al., 1994), suggesting a global role for this transcription factor. We now demonstrate that LSF impacts gene expression at the G1–S transition, through regulation of the gene that encodes thymidylate synthase (TS). TS is an essential enzyme that is responsible for the de novo synthesis of thymidylic acid (Danenberg, 1977), a nucleotide precursor for DNA replication. Inhibition of TS enzyme activity by substrate analogs causes either growth arrest or programmed cell death (Houghton et al., 1994). Regulated expression of TS mRNA requires both promoter and intron sequences within the TS gene (Ash et al., 1993, 1995; Ke et al., 1996). We show that mutation of LSF binding sites in the TS gene inhibits the induction of mRNA from a minimal TS minigene at the G1–S transition. Furthermore, by using dominant-negative LSF (LSFdn) to block endogenous LSF activity, we show that LSF is critical for the S-phase induction of TS enzyme levels. LSFdn expression induces apoptosis in both mouse fibroblast cells and a human prostate cancer cell line, DU145. Using several experimental approaches, we show that this apoptotic phenotype is directly linked to the inhibition of TS expression. Results Identification of LSF DNA-binding sites within TS cell cycle regulatory regions Given the ability of LSF to transactivate the SV40 major late promoter in vitro, we performed a database search for S-phase-regulated genes that contained LSF binding sites. Sequence analysis of the mouse TS gene (Figure 1A), whose mRNA levels increase 10-fold during G1 to S-phase progression in growth-stimulated cells (Jenh et al., 1985), revealed potential LSF binding sites (Figure 1B) in both promoter and intronic cell cycle regulatory regions (inverted triangles, Figure 1A). One site (−93/−75) was located in a region of the promoter that is essential for regulated expression of the gene (Ash et al., 1995) and is highly conserved among mammalian TS genes (Takeishi et al., 1989; Dong et al., 2000). Figure 1.LSF binds within regulatory regions of the mouse TS gene. (A) LSF sites located in promoter and intron regulatory regions of the mouse TS gene. Inverted triangles represent LSF binding sites; black boxes, coding regions; thick lines, promoter region; thin lines, introns; arrows, the positions of the multiple major transcriptional start sites (Deng et al., 1986); the gray bracket from −92 to −14, transcriptional initiation window; and the black bracket from −104 to −75, the essential promoter region (Geng and Johnson, 1993). Position +1 represents the A residue of the codon for translational initiation. The pTTT DNA probe used for S1 analysis (Ash et al., 1993) is indicated below. This probe extends from the BglII site to the XbaI site in the TS gene and lacks introns. The two BamHI sites mark the endpoints of the small 57 bp deletion in the minigenes that enables differentiation of the endogenous and minigene TS mRNAs in the S1 analysis. (B) Sequences of LSF wild-type binding sites in the mouse TS promoter and intron regulatory regions, and of the mutations generated in these sites. Underlined nucleotides in each site represent bases that match the LSF consensus binding site (Shirra, 1995). The locations of Ets-1 and Sp1 binding sites in the promoter −93/−75 region are indicated. The −160/−142 site is written in an inverted orientation relative to its position in the promoter in order to align the site with the consensus sequence. The base pair substitutions generated to mutate each site are indicated below the respective wild-type sequences. (C) EMSA of recombinant LSF with wild-type and mutant mouse TS sites, and the consensus LSF binding site, as indicated. Binding reactions contained either 400 ng of His-LSF (lanes 1 and 2) or 100 ng of His-LSF (lanes 3–8). (D) Mutant TS essential promoter region retains Ets-1 binding activity. Purified recombinant Ets-1 protein and the radiolabeled wild-type −93/−75 DNA were incubated with increasing amounts of wild-type (lanes 2–6) or mutant (lanes 7–11) competitor oligonucleotides, added at the same time as the radiolabeled DNA. The molar excess of competitor DNA was as follows: lanes 2 and 7, 2-fold; lanes 3 and 8, 5-fold; lanes 4 and 9, 10-fold; lanes 5 and 10, 20-fold; lanes 6 and 11, 50-fold. (E) Mutant TS essential promoter region retains Sp1 binding activity. Recombinant Sp1 and the radiolabeled wild-type −93/−75 DNA were incubated with increasing amounts of either wild-type (lanes 2–5) or mutant (lanes 6–9) competitors. The molar excess of competitor DNA was: lanes 2 and 6, 2-fold; lanes 3 and 7, 5-fold; lanes 4 and 8, 20-fold; lanes 5 and 9, 50-fold. Download figure Download PowerPoint To test the relevance of LSF for TS expression, we first demonstrated by electrophoretic mobility shift assay (EMSA) that purified recombinant histidine-tagged LSF specifically bound all of the anticipated sites (Figure 1C, lanes 1, 4, 6 and 8). The ability of cellular LSF to bind these sites was confirmed by EMSA with nuclear extracts from mouse 3T6 fibroblasts, and by DNA footprinting analysis with LSF purified from HeLa cells (data not shown). To determine whether LSF could activate transcription from the TS promoter, an LSF expression construct was transiently cotransfected with a CAT reporter gene driven by the TS promoter region, and the level of gene expression assayed (Figure 2). Expression from the wild-type TS promoter increased 7-fold in the presence of LSF (see Figure 2, compare WT gray bar and black bar). The stimulation was similar to that observed with a synthetic promoter that contained four copies of an LSF binding site (synthetic). Expression from a TS promoter with mutations that improved the LSF site in the essential promoter region (+LSF) increased 12-fold in the presence of LSF. However, expression from a TS promoter with mutations that weakened the LSF site (−LSF) was unaltered by the presence of LSF. Thus, we conclude that LSF can stimulate expression from the TS promoter and that this transactivation is substantially mediated by the LSF binding site within the essential region of the promoter. Figure 2.Transactivation of the TS promoter by wild-type LSF is inhibited by LSFdn. CAT reporter genes were driven by either the wild-type mouse TS promoter region (from −985 to −11 nt relative to the AUG start codon) (WT), the TS promoter region with an improved LSF binding site (+LSF), the TS promoter region with a weakened LSF binding site (−LSF), or a synthetic promoter consisting of four LSF binding sites upstream of a TATA box (synthetic). The sequences of the TS essential promoter region and of the mutations that were introduced (underlined) are shown at the top of the figure. Note that the mutations in the LSF binding site also affect the downstream Ets and Sp1 binding sites, which overlap the LSF binding site. The CAT reporter genes (0.2 μg) were transiently transfected into hamster V79 cells in the absence of an LSF or LSFdn expression construct (gray bars), in the presence of 0.2 μg of pEF-1α LSF alone (black bars), or in the presence of both 0.2 μg of pEF-1α LSF and 0.3 μg of pEF-1α LSFdn (white bars). The amount of transfected pEF-1α promoter was kept constant by adding appropriate amounts of pEF-1α empty vector to the transfection mixture. Cells were harvested 40–48 h after transfection and CAT activity was determined. CAT activity was normalized to the value observed in cells transfected in the absence of LSF or LSFdn, which was set at 100. The standard deviations obtained from duplicate samples are shown. Download figure Download PowerPoint LSF DNA-binding sites are required for cell cycle regulation of TS mRNA levels Due to the ability of LSF to bind cell cycle regulatory regions in the mouse TS gene, and to activate transcription through its binding site in the essential region of the promoter, we tested whether LSF contributed to G1–S stimulation of TS mRNA levels. In both murine and human cells, the coexistence of both the promoter region and an intron is required for normal regulation (Li et al., 1991; Kaneda et al., 1992; Takayanagi et al., 1992; Ash et al., 1993; Ke et al., 1996). In order to focus on contributions from the promoter and to minimize contributing regulatory effects from intron sequences (Ash et al., 1993, 1995; Korb et al., 1993; Ke et al., 1996), we used a minimal TS minigene containing a single internally deleted intron as the parental construct for studies of LSF site mutations. The TS minigene pTI1dT retains S-phase regulation although the fold induction of mRNA at the G1–S transition is reduced from that of the endogenous gene (Ash et al., 1993) (Figure 3A and B, left panels). Point mutations were generated in the −160/ −142 (triple base pair) and the −93/−75 (double base pair) LSF sites, which abolished binding of LSF to these promoter sites (Figure 1C, compare lanes 4 with 5 and 1 with 2). A similar triple base pair mutation of the remaining intron 1 binding site also substantially inhibited binding of LSF to this site in vitro (Figure 1C, compare lanes 6 and 7). The mutation of the −93/−75 site was designed to avoid inactivation of the binding sites for transcription factors Ets and Sp1 (Deng et al., 1989; Jolliff et al., 1991; Geng and Johnson, 1993), which overlapped the LSF site (Figure 1B). To test the relative affinities of these proteins for the wild-type versus mutated −93/−75 sequences, the binding of each of these proteins to the wild-type promoter site was competed in parallel with either the wild-type or the mutated sequence (Figure 1D and E). This type of analysis is the most sensitive in detecting differences in affinities for two different sequences. In contrast to the binding by LSF, the competition curves verified that the introduced point mutations did not prevent the interaction between the −93/−75 TS promoter region and Ets-1 or Sp1. Thus, the only identified factors in nuclear extracts, other than LSF, that bind to the −93/−75 TS promoter region (Ash et al., 1995) were still able to bind the mutated promoter sequences, whereas LSF was not. The situation was more simple at the other sites, as LSF was the only protein detected in nuclear extracts that bound to the −160/−142 site and the intron 1 site (data not shown). Finally, EMSA with nuclear extracts detected no new factors that could bind the mutated LSF sites (data not shown). Figure 3.Mutation of LSF binding sites inhibits G1–S stimulation of TS minigene mRNA levels. (A) Representative S1 analysis of mRNA from endogenous TS and TS minigenes. NIH 3T6 cells were stably transfected with the indicated minigene, and equal amounts of mRNA were analyzed by S1 nuclease protection assays. The number above each lane indicates the time after serum stimulation (in hours) at which mRNA was collected. The R band represents ribosomal protein L32 mRNA and serves as the internal control. The bracket marks the endogenous (E) TS products, which appear as a broad smear due to the protection of the S1 probe into the region of multiple start sites in the TS promoter (Geng and Johnson, 1993). The asterisks mark the minigene (M) products. For visualization purposes, the sections of the autoradiographs containing the TS minigene products were enhanced 8-fold (8× M) and displayed beneath the full-length autoradiographs. (B) Quantitation of the levels of minigene and endogenous TS mRNAs shown in (A). To correct for differences in RNA recovery, TS mRNA levels (E and M) were normalized to those of the control (R). The indicated fold increases were calculated by normalizing to the lowest mRNA levels. Circles represent endogenous TS mRNA levels, triangles represent TS minigene mRNA levels, and squares represent control ribosomal protein L32 mRNA levels. Download figure Download PowerPoint TS minigenes that contained either the wild-type sequence or mutations in various LSF binding sites were transfected into mouse fibroblasts, and pools of stably transfected colonies were generated. RNA was prepared for S1 analysis from quiescent cells and from cells at various times following growth stimulation (Figure 3). A 57 bp deletion in exon 3 of each minigene enabled differentiation between endogenous (E and circles) and minigene (M and triangles) TS mRNA signals (Ash et al., 1993). In addition, a ribosomal protein mRNA was simultaneously analyzed (R and squares) to control for RNA recovery. The striking result was that simultaneous mutation of all LSF sites in the TS minigene regulatory regions nearly abolished cell cycle regulation (Figure 3A and B, right panels). Furthermore, mutation of only the −93/−75 LSF site in the promoter also dramatically inhibited G1–S activation of TS mRNA levels (Figure 3A and B, middle panels). Initial results (data not shown) also show that the mutation of the LSF binding site in intron 1 alone partially blocks the increase in TS mRNA levels as cells enter S-phase. These findings indicate that, whereas other promoter elements direct basal TS mRNA expression, the LSF binding sites are critical for G1–S-regulated TS mRNA expression in this minimal setting. Thymidine overcomes apoptosis induced in S-phase by dominant-negative LSF Because RNA analyses demonstrated that LSF binding sites were essential for the S-phase-specific induction of TS minigene expression, and because TS activity is essential for S-phase progression, the role of LSF in cell survival was investigated more directly. Dominant-negative LSF (LSFdn), a double amino acid substitution mutant of LSF initially named 234QL/236KE, is unable to bind DNA (Shirra et al., 1994). LSFdn also inhibits the DNA-binding activity of comparable levels of wild-type LSF in vitro. Consistent with this observation, LSFdn blocked the ability of LSF to stimulate expression of an LSF-dependent reporter gene in transient transfection experiments, using either human osteosarcoma U2OS cells (data not shown) or hamster V79 cells (Figure 2, synthetic). Furthermore, LSFdn inhibited the expression of a reporter gene driven by the TS promoter (Figure 2, WT), as well as the TS promoter containing an improved LSF binding site (Figure 2, +LSF). This ability of LSFdn to inhibit reporter gene expression was specific for LSF-responsive promoters. When the LSF binding site in the TS essential promoter region was weakened, inhibition by LSFdn was no longer observed (Figure 2, −LSF). Furthermore, expression of other control genes, such as the metallothionein promoter-driven human growth hormone gene, was unaffected by expression of LSFdn (data not shown). Therefore, LSFdn effectively and specifically blocks wild-type LSF function in vivo. By transient coexpression of LSFdn with a cell-surface marker (CD7) (Frangioni et al., 1994), we tested the effects of LSFdn on cell survival in NIH 3T3 mouse fibroblasts. Growth of the transiently transfected cells was arrested by serum withdrawal. At various time points after serum stimulation, transfected cells (expressing the CD7 surface marker) were analyzed by flow cytometry for their cellular DNA content, which is reflective of progressing stages in the cell cycle. Through the beginning of S-phase (16 h), cells transfected with the LSFdn expression construct progressed through the cell cycle in a similar way to cells transfected with vector alone (Figure 4A, compare columns 3 and 1; Table I, compare lines 3 and 4). This was true even in the presence of 20 μM thymidine (see below). Cell cycle progression was represented by the shift in the G1 peak of cellular DNA (peak observed at 0 h) into the broad region of higher DNA content characteristic of S-phase. Control cells continued to progress normally into G2-phase at 22 h, as shown by the accumulation of the peak at double the G1 level of DNA. In stark contrast, a significant proportion of cells transfected with the LSFdn construct accumulated a sub-G1 DNA content at 22 h. The accumulation of sub-G1 DNA content (30% in this experiment) was accompanied by a corresponding decrease in the fraction of cells in the S- and G2/M-phases of the cell cycle (Table I, compare lines 7 and 8), indicating that LSFdn induces DNA fragmentation at some point during, or immediately following, S-phase of the cell cycle. Figure 4.LSFdn induces apoptosis, which is reversible by thymidine. (A) Flow cytometric analysis of the DNA content in transiently transfected NIH 3T3 cells either expressing LSFdn or not. Cells were transfected and growth arrested, as described in Materials and methods, with 2.5 μg of pMARK7 DNA and 12.5 μg of either pEF-1α or pEF-1α-LSFdn DNA. Cells were subsequently propagated in the absence or presence of 20 μM thymidine, which was added at the time of serum stimulation. The histograms represent relative measurements of DNA content in cells expressing exogenous CD7. The bracket indicates cells that contain less than the G1 DNA content, characteristic of apoptosis. Numbers on the left indicate the time (h) after serum stimulation. The number of cells represented in each histogram, left to right, is as follows. Top row: 623, 1023; middle row: 3618, 2419, 2447, 2201; bottom row: 7417, 13434, 6725, 6701. (B) DNA fragmentation in NIH 3T3 cells transfected by LSFdn is inhibited by a peptide caspase inhibitor and by coexpression of LSF. pEF-1α-LSF (or pEF-1α, in control samples) was cotransfected at a 3:1 ratio with pEF-1α-LSFdn (9.4 μg of pEF-1α-LSF DNA, 3.1 μg of either pEF-1α or pEF-1α-LSFdn DNA, and 2.5 μg pMARK7 DNA), as indicated. CD7-positive cells were analyzed 22 h after serum stimulation. The number of cells represented in each histogram, left to right, is as follows: 13347, 5112, 14500, 19124, 10232, 7572. The numbers above the brackets indicate the percentage of cells with a less than G1 cellular DNA content. (C) Induction of TS protein is reduced in cells transfected with LSFdn, even in the presence of the caspase (CPP32) peptide inhibitor, DEVD-CHO. Cytoplasmic extracts were prepared from CD7-positive cells, sorted by flow cytometry and analyzed by western blotting for TS. The blot was subsequently stripped and re-probed with an antibody against β-actin, as indicated. Lane C contains wild-type mouse recombinant TS (Zhang et al., 1989) used as a size marker. Lane 1, 5 μg of cytoplasmic extract from quiescent NIH 3T3 cells (starved 36 h); lane 2, 5 μg of extract from NIH 3T3 cells undergoing logarithmic growth; lane 3, 5 μg of extract from CD7-positive NIH 3T3 cells 22 h post-stimulation with pEF-1α-LSFdn; lane 4, 5 μg of extract from CD7-positive cells 22 h post-stimulation with pEF-1α; lane 5, identical to lane 3 with the exception that the cells were incubated with 2.0 μM DEVD-CHO from the time of serum stimulation. Download figure Download PowerPoint Table 1. LSFdn-induced apoptosis reduces the percentage of 3T3 cells in S + G2 phases of the cell cycle Samplea Time (h) DNA content (range of channel numbers) Sub-G1 (0–124) G1 (125–242) S ± G2 (243–558) Control 0 3.2 86 10 LSFdn 0 3.5 85 11 Control 16 0.6 55 44 LSFdn 16 0.9 59 40 Control + 20 μM thymidine 16 0.7 56 43 LSFdn + 20 μM thymidine 16 1.5 66 33 Control 22 0.8 33 66 LSFdn 22 30 33 37 Control + 20 μM thymidine 22 1.0 28 71 LSFdn + 20 μM thymidine 22 1.3 29 70 a Corresponding to the histograms in Figure 4A. The cause of the sub-G1 cellular DNA content was tested by treating the cells transfected with the LSFdn construct with the caspase (CPP32) peptide inhibitor, DEVD-CHO (Lazebnik et al., 1994; Nicholson et al., 1995) (Figure 4B). This treatment reduced the percentage of cells with a sub-G1 DNA content at 22 h, demonstrating that expression of LSFdn induced an apoptotic program resulting in fragmentation of cellular DNA. Finally, when LSF was transiently coexpressed with LSFdn, the apoptotic phenotype was also circumvented (Figure 4B), suggesting that the induction of apoptosis was due to specific inhibition of wild-type LSF functions. To determine whether the apoptotic phenotype induced by LSFdn was related directly to regulation of TS by LSF, we investigated whether apoptosis could be prevented by exogenous thymidine. Through the salvage pathway, thymidine can replenish the cellular thymidine triphosphate pools needed for DNA replication. Strikingly, 20 μM thymidine, added extracellularly, prevented cells transfected with LSFdn from undergoing DNA fragmentation (Figure 4A, column 4; Table I). This low concentration of thymidine, which is sufficient to overcome the cellular requirement for TS activity, did not significantly affect cell cycle progression of control cells (Figure 4A, compare columns 1 and 2; Table I). Induction of apoptosis by LSFdn therefore parallels a process known as thymineless death, which results from inhibition of TS activity and subsequent depletion of thymidine triphosphate pools (Houghton et al., 1994, 1997; Harwood et al., 1996). To verify that LSFdn is directly regulating TS, the effect of LSFdn on the levels of cellular TS enzyme was measured. Growth-arrested NIH 3T3 cells, starved for 36 h, have extremely low levels of TS protein (Figure 4C, lane 1), as expected based on the 7 h half-life of TS protein (Kitchens et al., 1999). This permitted us to examine G1–S induction of TS gene expression by analysis of TS protein levels. Therefore, NIH 3T3 cells were transfected with either the LSFdn expression construct or the control vector, growth arrested, and stimulated with serum to re-enter the cell cycle. Twenty-two hours after stimulation, transfected cells were sorted on the basis of expression of the CD7 cell-surface marker, to select for cells expressing high levels of LSFdn. Western blotting analysis revealed that TS protein levels in cells expressing high levels of LSFdn remained at the levels observed in quiescent cells (Figure 4C, compare lanes 3 and 1). In contrast, TS protein levels in cells transfected with the control vector (lane 4) were similar to those undergoing logarithmic growth (lane 2). The levels of β-actin remained constant in all cases (Figure 4C). Thus, expression of LSFdn prevented the increase in the level of cellular TS protein during the G1–S transition. To ascertain whether the ability of LSFdn to block induction of TS protein was related to the induction of apoptosi
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