INTRODUCTION

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The c-Myc proto-oncogene, which is frequently overexpressed in human cancer, is a central regulator of cell proliferation and can also sensitize cells to apoptosis (for a review, see reference 20). The carboxy terminus of the c-Myc protein contains a basic-helix-loop-helix-leucine zipper (b-HLH-LZ) domain characteristic of known transcription factors. This region is required for binding to the b-HLH-LZ protein Max to form a heterodimeric protein complex capable of sequence-specific DNA binding (9, 10, 44). Myc-Max heterodimers recognize the E-box related consensus sequence CACGTG and can induce transactivation when this sequence is placed proximal to a minimal TATA box containing promoter (2, 3, 18, 31). Consistent with the role of the Myc-Max heterodimer as a sequence-specific transactivating complex, c-Myc expression has been shown to directly transactivate a number of genes associated with cellular proliferation and metabolism including ornithine decarboxylase (ODC), cad, lactate dehydrogenase A (LDH-A), and eIF4E (reviewed in reference 13).

The amino terminus of c-Myc (amino acids [aa] 1 to 144) has both transcriptional activation and repression activities. Within this domain are two evolutionarily conserved regions termed Myc Box (MB) I (aa 47 to 62) and MB II (aa 106 to 143). Deletions within MB I have been shown to diminish Myc-mediated transactivation, while MB II deletion mutations result in diminished transrepression (4, 33, 34, 36). Transcriptional repression by c-Myc is hypothesized to act through a pyrimidine-rich cis-initiator element termed the Inr, although the exact molecular mechanism through which c-Myc mediates Inr-dependent repression remains unknown (5, 11, 33). Genes that are repressed by c-Myc include cEBP, gadd45, and gas1 (reviewed in reference 11).

The ability of c-Myc to repress transcription has been recently linked to its ability to mediate cellular transformation (11). Furthermore, several lines of evidence suggest that there is no absolute correlation between transcriptional activation by c-Myc and its function in growth regulation. First, partial deletion of MB I (aa 41 to 53) results in diminished transactivation but does not significantly abrogate either transformation of Rat1a cells, cotransformation of primary rat embryo cells (51), or cell cycle progression in the presence of limiting serum (17). Second, structure-function analyses have suggested that deletion of MB II (aa 106 to 143), the region required for transcriptional repression (33, 34), completely abrogates transformation and cell cycle progression (17, 51). Third, a recent report has demonstrated that a transactivation-defective Myc S (short) protein (lacking the first 100 aa of c-Myc) retains the ability to enhance both proliferation and apoptosis (57). While collectively these experiments suggest that transcriptional activation may be dispensable for some c-Myc functions, the variable growth conditions, cell lines, and experimental systems used in previous experimental systems makes drawing firm conclusions about transcriptional activation and apoptosis difficult. Therefore, in the current study, we characterize the functional requirements for MB I and MB II in a single cell culture system. We show that c-Myc-induced transcriptional activation of ODC clearly does not directly correlate with either c-Myc-accelerated cell cycle progression or apoptosis. However, the presence of the putative transcriptional repression domain, MB II, is absolutely required for c-Myc-accelerated cell cycle progression, apoptosis, and transformation. Moreover, we provide evidence that expression of a lymphoma-derived mutant of c-Myc, in which threonine 58 is mutated, results in wild-type levels of ODC transcription and cell cycle progression while failing to accelerate apoptosis or repress gadd45 as efficiently as wild-type c-Myc. This observation suggests that separable c-Myc-dependent pathways may execute cell cycle progression and apoptosis. Taken together, these data provide further evidence that, in immortalized fibroblasts, (i) transactivation of ODC expression is unlikely to be a primary mechanism of c-Myc-induced cell cycle progression or apoptosis, (ii) transcriptional repression may be required to mediate c-Myc-induced apoptosis, and (iii) the mechanisms of c-Myc-accelerated cycle progression and apoptosis are separable, although both appear to require a function dependent upon MB II. Finally, our results show that acceleration of apoptosis by c-Myc can be compromised not only by lesions in other cellular genes but also by mutations in the c-Myc protein itself.

	MATERIALS AND METHODS

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Plasmids and vectors. The c-MycER deletion mutants were constructed by subcloning the human c-MycER fusion gene (14) into the HindIII site of pBKS(+) vector. Deletion mutants (51) were then constructed in pBKS(+) by replacing the wild-type 5' end of c-Myc with the various 5' ends of mutant c-Myc alleles using the flanking EcoRI site and the ClaI site at nucleotide 784 of the c-Myc cDNA. The mutant c-Myc-ER fusion genes were then subcloned back into the HindIII site of the pMV7 retroviral vector (56). The Thr58Ala c-MycER mutant was also constructed by subcloning the 5' end of aa-58 point mutant c-Myc gene (a gift of J. Woodgett, Ontario Cancer Center) into the EcoRI/ClaI sites of c-MycER in the pBKS(+) vector. VP16-MycER was constructed by inserting a BglII-ClaI polylinker between EcoRV (aa 47) and ClaI (aa 262) sites of c-Myc cDNA and then inserting the transactivation domain of VP16 in frame into this sites. VP16-Myc chimeric cDNA was cloned in frame with estrogen receptor (ER) ligand-binding domain into the pMV7 retroviral vector to generate pMV7-VP16MycER containing the activation domain of VP16 substituting for aa 47 to 262 of c-Myc. The pBabePuro retroviral vector was used to deliver T58AMycER and wild-type MycER (35) into mouse embryo fibroblasts (MEF).

Cell culture. Rat1a fibroblasts were grown in Dulbecco modified Eagle medium (DMEM) with 10% fetal calf serum (FCS). Cells were infected with either pMV7, pMV7-c-MycER, or pMV7-mutant c-MycER retroviruses as described before (56). Cells with integrated viruses were selected in 500 µg of G418 per ml and maintained in phenol red-free DMEM with 10% certified low-estrogen content FCS (Atlanta Biologicals). For apoptosis assays, cells were plated at density of 100,000 cell per 3-cm dish. Quantitation of apoptosis by DAPI (4',6'-diamidino-2-phenylindole) staining was performed as previously described (29). For cell cycle progression cells were plated at high density of 10⁶ cells per 10-cm dish to prevent spontaneous apoptosis during prolonged serum deprivation period. MEF were derived from 14.5-day embryos of C57/B6 mice. Passage three MEF were infected with pBabePuro retroviruses followed by selection with 1 µg of puromycin per ml.

Flow cytometric analysis. For flow cytometric analysis, cells (approximately 10⁴ per time point) were quiesced by serum deprivation (0.5% FCS) for 60 h in phenol red-free DMEM. Subsequently, 4-hydroxytamoxifen (4-OHT) was added to the media for 18 h prior to harvesting cells by trypsinization. Cells were then pelleted in a clinical centrifuge at 1,000 rpm, resuspended in 300 µl of phosphate-buffered saline (PBS), and then fixed by adding 700 µl of cold ethanol while vortexing the mixture. Fixed cells were then repelleted, resuspended in 1 ml of PBS, and pelleted again. Cells were resuspended in 1 ml of PBS containing 10 µg of propidium iodide and 100 µg of RNase-free DNase per ml. Cell cycle profiles were determined using the Lysis II program on the FACScan flow cytometer (Beckton-Dickinson).

BrdU analysis. Cells were plated on sterile glass coverslips in 30-mm-diameter wells and starved for 60 h in phenol red-free DMEM containing 0.5% FBS. Cells were then treated for 18 h with 10⁷ M 4-OHT. During the last hour of treatment 10 µM 5-bromo-2'-deoxyuridine (BrdU) was added to the media. Immunofluorescent detection of BrdU incorporation as performed according to the manufacturer's protocol (Boehringer Mannheim). Nuclei were counterstained with DAPI, and slides were visualized by fluorescent microscopy and photographed with a digital camera. Cells stained with DAPI and with BrdU incorporated were then counted by printing the digitized images of random fields and scoring at least 300 cells per experimental group.

RNA analysis. Rat1a cells were grown to 80% confluence and serum starved in 0.5% FCS with phenol red-free DMEM for 60 h. Cells were then treated with 10⁷ M 4-OHT, and RNA was harvested 8 h later. RNA was isolated by using the Qiagen method according to the manufacturer's description and then fractionated on 1% agarose-6% formaldehyde gels and transferred to Duralon-UV membranes (Stratagene) by capillary blotting. After UV cross-linking, membranes were hybridized sequentially to cDNA probes for ODC, gadd45, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) that had been labeled with [³²P]dCTP by random priming (Stratagene). Northern blot analysis for ODC mRNA was done by using a 2-kb EcoRI/BamHI fragment from pBS-ODC (25) and for GAPDH was done by using a 1.4-kb PstI fragment of pBS-GAPDH. Quantitation of autoradiograph signals was performed by using the NIH Image software.

Western blot analysis. Equal numbers of exponentially growing Rat1a cells expressing the pMV7 control, wild type, or c-Myc mutants were lysed in 2× Laemmli buffer (28). After sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transfer to nitrocellulose, equal loading of protein was confirmed by Ponceau S staining. Nitrocellulose was then blotted with the rat monoclonal anti-ER antibody H222 (a kind gift of Geoffrey Greene), detected by Anti-Rat-HRP (Zymed), and developed using chemiluminescence according to the manufacturer's instructions (Amersham).

Cytochrome c immunostaining. Cells were plated at a density of 125,000 cells per 3-cm dish on glass coverslips. Cells were fixed in PBS containing 4% formaldehyde-0.2% saponin and stained with 1 µg of Hoechst 33258 per ml for 20 min. The fixed cells were incubated in blocking buffer (PBS containing 10% FCS and 0.2% Triton X-100) for 30 min and then for an additional 30 min in PBS containing 0.2% saponin, 2% bovine serum albumin (BSA), and 1 µg of anti-cytochrome c antibody (clone 6H2.B4; PharMingen, La Jolla, Calif.) per ml. Cells were then washed three times with blocking buffer and incubated for 30 min in PBS containing 2% BSA, 0.2% saponin, and 1 µg of tetramethyl rhodamine isocyanate (TRITC)-conjugated anti-mouse antibody (Jackson ImmunoResearch, West Grove, Pa.) per ml. Cells were then rinsed three times with blocking buffer, and coverslips were mounted onto slides.

Soft agarose assays. Cells (10⁵) were plated subconfluently in 60-mm plates in 0.7% agarose on a 1.4% agarose bed in the presence of 4-OHT with various serum concentrations. Plates were fed once per week with 2 ml of 2% or 10% FCS-DMEM plus 4-OHT. Colonies were scored by projecting plates onto a dry erase board using an overhead projector. Only colonies with a diameter of >1.0 mm were scored.

	RESULTS

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The ability of c-Myc to accelerate apoptosis does not correlate with the integrity of Myc's known transcriptional activating domains. Ectopic expression of c-Myc accelerates apoptosis upon growth factor withdrawal (6, 8, 15, 56). The ability of c-Myc to accelerate apoptosis is thought to be associated with its transcriptional activation capability and with its ability to induce cell cycle progression (40). However, treatment of cells with cycloheximide does not diminish c-Myc-accelerated apoptosis (15, 54), suggesting that transcriptional activation and the subsequent induction of de novo protein synthesis by c-Myc are not required to accelerate apoptosis. To directly address this apparent paradox, we initiated studies to define the transcriptional regulatory domains required for c-Myc-accelerated apoptosis.

View larger version (25K): [in this window] [in a new window]   FIG. 1.   Expression levels of wild-type and mutant c-Myc proteins in stably infected Rat1a polyclonal cell lines. (A) Schematic illustration of wild-type and mutant c-MycER fusion proteins. Hatched boxes indicate MB I and MB II; potential phosphorylation sites (Thr58 and Ser62) in MB I are also indicated. In the VP16-Myc construct, the activation domain of VP16 replaces aa 47 to 263 and is fused in frame with the DNA binding domain of c-Myc. (B) Western blot analysis of MycER proteins using the ER-specific H222 antibody.

View larger version (25K): [in this window] [in a new window]   FIG. 2.   Induction of apoptosis by c-Myc mutants. Time course of percentages of apoptotic Rat1a cells scored by DAPI staining. Cells (105) were plated in 30-mm wells, allowed to adhere overnight, and then serum deprived in the presence of 4-OHT in order to activate c-Myc. Cells were then fixed directly in tissue culture plates, stained with DAPI, and scored for nuclear condensation. Averages of at least 300 cells from three independent experiments are shown ± the standard error (SE).

c-Myc-accelerated apoptosis correlates with transrepression and not transactivation of gene expression. The results described above led us to examine directly the correlation between transactivation, transrepression, and apoptosis functions of c-Myc. The most well-characterized target of c-Myc transcriptional activation is the ODC gene. ODC has been shown to be induced by c-Myc in transient and stably expressing cell populations (reviewed in reference 13). Both the rat and human ODC genes contain two consensus CACGTG Myc-binding sites located in the same position in the first intron. We have previously shown 4-OHT-dependent activation of ODC expression in BALB/c 3T3 and Rat1a fibroblasts following expression of the conditionally active c-MycER chimeric protein (55). Therefore, we measured the relative induction of endogenous ODC gene expression and the acceleration of apoptosis in cells expressing individual mutant c-MycER proteins. To determine the extent of ODC transactivation by c-Myc, Rat1a cell pools were cell cycle arrested by serum deprivation, and c-Myc was activated by the addition of 10⁷ M 4-OHT. Analysis of the induction of ODC mRNA transcript normalized for GAPDH expression (Fig. 3) revealed that transactivation of ODC in cells expressing the MB I mutant dl41-53 was inhibited approximately 30% compared with cells expressing wild-type c-Myc. Thus, the MB I deletion mutant dl41-53 was unable to transactivate ODC as robustly as wild-type c-Myc, even though it retained the ability to accelerate apoptosis. In contrast, expression of the phosphorylation site mutant T58A resulted in potent transactivation of ODC and a significant loss of apoptotic function. Similarly, the VP16-Myc chimeric protein also induced significant ODC expression (Fig. 3A) and yet expression of this protein had no effect on apoptosis (Fig. 2). In summary, these results indicate that c-Myc-mediated transcriptional activation has no direct correlation with acceleration of apoptosis and suggest that induction of apoptosis may be mediated by mechanisms independent of transactivation using ODC as a readout for transactivation.

View larger version (44K): [in this window] [in a new window]   FIG. 3.   Induction of ODC mRNA by c-Myc mutants. (A) Northern blot analysis of ODC mRNA was performed using 10 µg of Rat1a RNA from cell pools expressing various mutant c-Myc proteins. Cells were made quiescent with serum deprivation, and c-Myc was activated by the addition of 4-OHT. RNA was harvested 6 h later and assayed for induction of ODC mRNA. (B) ODC mRNA fold induction relative to control (GAPDH) RNA, as quantified by densitometry. The fold induction following c-Myc activation with 4-OHT was averaged from three individual experiments.

View larger version (43K): [in this window] [in a new window]   FIG. 4.   Repression of gadd45 mRNA by c-Myc mutants. RNA (15 µg) from Rat1a cell pools deprived of serum for 60 h and then exposed to 4-OHT was analyzed by RNase protection assay to detect the presence of endogenous gadd45 and GAPDH-specific sequences. Experiment is a representative of three independent experiments.

Induction of cell cycle progression and acceleration of apoptosis are separable functions of c-Myc. The c-Myc mutants enabled us to determine whether there is a strict correlation between the ability of c-Myc to induce cell cycle progression and to accelerate apoptosis. In order to measure cell cycle progression and apoptosis simultaneously, cells expressing different c-Myc alleles were serum deprived for 60 h so that approximately 80% of cells were in G₀/G₁ prior to c-Myc activation (Table 1). At 18 h after c-Myc activation with 4-OHT, we employed two techniques to measure induction of S phase: (i) evaluation of individual cell BrdU incorporation, as measured by indirect immunofluorescent microscopy, and (ii) fluorescence-activated cell-sorting (FACS) examination of cell populations (10⁴ cells) for DNA content (Fig. 5). Table 1 illustrates that the two techniques correlated well; with either method cells evaluated 18 h after activating ectopically expressed mutant c-Myc proteins either showed robust induction of S phase similar to wild-type c-Myc or no S-phase induction (Table 1). Wild-type c-Myc and dl41-53 and T58A mutants were able to induce S phase equally well, while expression of VP16-Myc, dl106-143, and dl41-178 did not stimulate cell cycle progression (Table 1 and Fig. 5) and the cells remained arrested indefinitely (data not shown). Since VP16-Myc led to a robust transactivation of ODC but could not induce cell cycle progression, ODC induction is clearly not sufficient to drive fibroblasts into the cell cycle. Also, despite diminished ODC induction in dl41-53-expressing cells, these cells were able to enter the cell cycle as efficiently as cells expressing wild-type c-Myc. Most interestingly, S-phase entry and apoptosis induced by c-Myc could be genetically dissociated in cells expressing the T58A c-Myc protein. This lymphoma-associated mutation in MB I showed robust induction of S-phase, diminished acceleration of apoptosis, and diminished ability to repress transcription. Taken together, these results suggest that the induction of cell cycle progression and acceleration of apoptosis are separable functions of c-Myc.

View larger version (35K): [in this window] [in a new window]   FIG. 5.   Cell cycle analysis after activation of c-Myc mutants. (A) Cells were deprived of serum for 60 h until ~80% were in G0/G1. The percentage of cells was determined by FACS in three independent experiments, and the SE was calculated. (B) Histogram showing cell cycle distributions of c-Myc pools prior to and following activation of Myc by 4-OHT.

View larger version (39K): [in this window] [in a new window]   FIG. 6.   Acceleration of apoptosis in MEF by wild-type and T58A mutant c-Myc. The percentages of apoptotic MEF were scored by DAPI staining. Cells (3 × 104) were plated in 30-mm wells, allowed to adhere overnight, and then placed in either 0.5 or 2% FCS in the presence of 4-OHT in order to activate c-Myc. Cells were then fixed directly in tissue culture plates for 24 or 48 h after addition of 4-OHT, stained with DAPI, and scored for nuclear condensation. Averages of at least 300 cells from three independent experiments are shown ± the SE.

The diminished ability of Thr58Ala mutant to accelerate apoptosis correlates with its reduced ability to induce the release of cytochrome c from mitochondria. In most cases, apoptosis is initiated by the loss of integrity of mitochondria and the release of cytochrome c (47). The released cytochrome c acts as a cofactor to initiate the apoptotic cascade and the activation of caspases that execute apoptosis. We and others have recently shown that c-Myc is able to induce the release of cytochrome c from mitochondria in a caspase-independent manner (27, 30). It was shown that the mechanism by which c-Myc is sensitizing cells to apoptotic stimuli is dependent on its ability to release cytochrome c from mitochondria (27). We therefore compared the abilities of wild-type c-Myc and T58A mutant to release cytochrome c. Quantitation of cytochrome c release was performed as previously described (30). After activation of c-MycER with OHT the cells were maintained in 0.5% FCS in the presence the caspase inhibitor zVAD for 3 h and then fixed and immunostained with cytochrome c antibodies. As a negative control in this experiment we used the cells expressing the deletion mutant dl106-143 that showed even less apoptosis than did the control Rat1a cells (Fig. 2). Rat1a cells coexpressing Bcl-2 and wild-type c-Myc served as a positive control. As shown in Fig. 7, the release of cytochrome c is directly correlated with the level of apoptosis (Fig. 2). The T58A mutant is about 70% less effective in releasing cytochrome c in comparison with wild-type c-Myc (Fig. 7). These results suggest that the primary defect in the ability of T58A mutant to elicit apoptosis is its inability to effectively mediate the release cytochrome c from mitochondria.

View larger version (19K): [in this window] [in a new window]   FIG. 7.   Induction of cytochrome c release by c-Myc. The percentages of cells demonstrating diffuse cytochrome c immunostaining were quantitated. Rat1a, Rat1a/MycER, Rat1a/T58AMycER, Rat1a/dl106-143MycER, and Rat1a/MycER/Bcl-2 cells were incubated overnight in DMEM with 2.5% FCS with 1 µM 4-OHT. The next day cells were placed in DMEM with 0.5% FCS and 100 µM zVAD for 3 h. Cells were then fixed and immunostained for cytochrome c. Averages (±SE) of at least 100 cells from four independent experiments are shown.

The transformation potential of wild-type and mutant c-Myc proteins reflects a balance between c-Myc-induced cell cycle progression and apoptosis. The balance between cell cycle progression and apoptosis is thought to contribute to the ability of cells to establish tumors and to anchorage-independent growth (38, 46, 50). The transformation potential of cells expressing the various mutant c-Myc proteins was assessed by measuring anchorage-independent growth in low serum, in which apoptosis is accelerated in wild-type c-Myc-expressing cells, versus high serum, in which growth and survival factors protect against c-Myc-accelerated apoptosis. Rat1a cells can be transformed to anchorage-independent growth by wild-type c-Myc alone (49, 51). Previously published reports of transformation assays comparing cells expressing different mutant c-Myc alleles were done in high (10% FCS) under conditions in which the differences in apoptotic potential were suppressed. We therefore assessed the effect of variable serum concentrations (i.e., high [10%] or limited [2%] FCS) and, in turn, variable apoptotic rates on anchorage-independent growth of colonies by performing the agarose assays in high and low serum concentrations. As a control a polyclonal cell line that coexpresses MycER and Bcl-2 was also included in these studies. The level of MycER expression in the MycER/Bcl-2 cell line was comparable to the level in the MycER cell line (data not shown). We found that, in 10% serum, the cell pools expressing wild-type Myc, T58A mutant, and Myc/Bcl-2 formed similar numbers of colonies on soft agarose and the dl41-53 mutant formed colonies 76% of the wild type (Fig. 8). The deletion of aa 41 to 53 does not affect apoptosis and cell cycle progression, but it inhibited transactivation of ODC and cellular transformation, implying that activation of ODC or other growth-related genes is required for full transformation by c-Myc.

View larger version (37K): [in this window] [in a new window]   FIG. 8.   Anchorage-independent growth on soft agarose of Rat1a polyclonal cell lines expressing different c-Myc alleles. Cells were plated in 2 or 10% FCS in the presence of 4-OHT and allowed to grow for 21 days in soft agarose. Colonies were counted by projecting plates on a white board using an overhead projector. Only colonies 1 mm in diameter or larger were scored. (A) Bar graph of the results as a relative percentage of growth of cell lines in soft agarose, where the number of colonies formed by wild-type-Myc-expressing cells is 100%. (B) Light microscopy pictures of soft agarose colonies formed in 2% FCS by either wild-type c-Myc, T58A-expressing cells, or cells infected with empty retrovirus.

	DISCUSSION

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Ectopic expression of c-Myc promotes cellular transformation and cell cycle progression. However, unless compromised by the activation of complementary pathways, c-Myc-overexpressing cells are sensitized to cell death (apoptosis) upon growth factor withdrawal. Thus, the ability of c-Myc to elicit neoplastic transformation is also dependent on the sensitivity of the cells to apoptosis.

In this report we described a comprehensive analysis of apoptosis, cell cycle progression, and transformation in Rat1a fibroblasts to understand what role regulatory domains in the amino-terminus region of c-Myc play in these processes. Early structure-function studies implicated both the DNA-binding carboxyl-terminus region and the amino-terminal region (aa 1 to 147) as being required for cell cycle progression and apoptosis. This bolstered the prediction that transcriptional activation was essential for c-Myc function. However, recently Xiong et al. described a transactivation-defective mutant of c-Myc lacking the first 100 aa that retains the ability to regulate cell cycle progression and apoptosis (57). In addition, mounting evidence suggests that c-Myc is also a potent transcriptional repressor (32, 33, 34, 36; see also the review in reference 11). Therefore, genes identified as being targets of transcriptional repression by c-Myc, such as the gadd45 and gas genes, could potentially be important in the regulation of growth control by c-Myc. In four instances (gas1, AdML promoter, c/EBP-alpha, and gadd45 genes) transcriptional repression by c-Myc has been shown to be dependent on the conserved MB II region, although other regions of c-Myc may affect transrepression (for a review, see reference 11).

In the studies presented here we showed that deletion of part of MB I significantly diminishes transactivation of the ODC gene and yet does not affect Myc's ability to drive the cell cycle from G₀/G₁ into S phase or to accelerate apoptosis in Rat1a cells. Because the diminished ability of MB I deletion mutant to activate ODC correlates with its diminished ability to induce cellular transformation, we concluded that activation of ODC and other growth-related genes is required for full transformation by c-Myc. However, substitution mutation of threonine 58 to alanine, a phosphorylatable amino acid immediately downstream of this region, inhibited apoptosis without affecting transactivation. The augmented ability of this mutant to induce cellular transformation appears to be due to its diminished ability to accelerate apoptosis. Interestingly, this T58A mutation was found to inhibit c-Myc-mediated transrepression. Deletion of the repression domain, MB II, was found to be required for cell cycle progression, apoptosis, and transformation, suggesting that transcriptional repression may play an essential role in most Myc functions. The functionality of MB II might be partially dependent on MB I since mutation of the potential phosphorylation site in this region diminishes the ability of c-Myc to repress transcription. Phosphorylation of residues in MB I might modulate MB II activity through regulation of binding of accessory proteins to MB II. Overall, our results clearly demonstrate a correlation between Myc-mediated transcriptional repression and apoptosis and dissociation of transcriptional activation and apoptosis, whereas cellular transformation is dependent on both transrepression and transactivation.

Our results also showed that the ability of c-Myc to induce cellular transformation, as measured by soft agarose assay, is a result of a balance between acceleration of apoptosis and cell cycle progression. The loss of apoptotic activity and the induction of Myc-mediated cell cycle progression both contribute to cellular transformation and can occur through several possible combinations of genetic events, including (i) activation of genes that inhibit apoptosis (e.g., Bcl-2 or Akt activation [8, 16, 28, 29, 56]) or (ii) loss of function of apoptosis-promoting genes (e.g., p53 mutation [22, 54]); both can result in a diminution of cell death despite c-Myc overexpression. In this report we present a third possibility for enhanced transformation by c-Myc: somatic mutation of c-Myc itself (aa 58) can contribute to c-Myc transformation potential, through reduced acceleration of apoptosis, while maintaining the ability to promote the cell cycle progression. Moreover, this observation suggests an additional level of regulation of c-Myc-mediated transformation: kinases or phosphatases targeting c-Myc might contribute to transformation by varying the phosphorylation state. This hypothesis is strengthened by the high frequency of MB I mutations in the region of phosphorylation in Burkitt's lymphomas and cell lines and the v-myc alleles in the avian lymphoma viruses (7, 52, 53). Thus, while overexpression of Bcl-2 or other antiapoptotic genes may encourage the growth of c-Myc-overexpressing tumors, somatic mutations within the overexpressed or amplified c-Myc gene may ultimately lead to a similar end result.

How does c-Myc induce cell cycle progression? Many studies have shown that ectopic expression of c-Myc can drive cells into all phases of the cell cycle in the absence of growth factors (20). However, the mechanism by which Myc drives cell cycle progression remains elusive. The emphasis in the past was to identify genes that can be transcriptionally activated by c-Myc. Indeed, there are many target genes that can be transcriptionally activated by c-Myc and are required for cell cycle progression (reviewed in reference 13). In agreement with previous reports (12, 57), the data presented here suggest that transcriptional repression by c-Myc is also important for c-Myc-mediated cell cycle progression. Several genes that are negative regulators of cell cycle progression have been identified as c-Myc targets, and these include the cyclin kinase inhibitors p27 (11) and p21 (37; A. Gartel and N. Hay, unpublished results), gadd45 (36), and gas1 (33). Identification of other genes that are transcriptionally repressed by c-Myc may uncover key downstream effectors of c-Myc-induced cell cycle progression and acceleration of apoptosis.

How does c-Myc accelerate apoptosis? Our results suggest that the ability of c-Myc to activate the transcription of cellular genes is unlikely to be the main mechanism by which c-Myc sensitizes immortalized cells to apoptosis. The acceleration of apoptosis induced by c-Myc could be through repression of transcription of certain cellular genes and/or through interaction with cellular proteins that are associated with apoptosis. We have previously shown that the execution of c-Myc-accelerated apoptosis is dependent on wild-type p53 (54). More recently, it was shown that the acceleration of apoptosis by c-Myc is mediated, at least in part, through the stabilization of p53 as a consequence of elevating p19-ARF expression by c-Myc (59). While this mechanism clearly contributes to the acceleration of apoptosis by c-Myc in primary cells, it might not be the major mechanism by which c-Myc accelerates apoptosis in immortalized cells. This is mainly because c-Myc was shown to accelerate apoptosis in a variety of established cell lines that lack functional ARF. Furthermore, we have previously shown that overexpression of p53 in primary cells in which endogenous p53 was spontaneously deleted is not sufficient to elicit apoptosis. Only the overexpression of both p53 and c-Myc could elicit apoptosis in these cells (54). Another mechanism by which c-Myc accelerates apoptosis could be through the activation of FADD, the downstream effector of Fas-mediated apoptosis (24). Although the precise molecular link between c-Myc and FADD has not been established yet, it is possible that c-Myc can activate this pathway by repressing the expression of negative regulators of this pathway such as FLIP (26). Finally, it has been recently shown that the primary function of c-Myc in acceleration of apoptosis is the enhancement of the release of cytochrome c from mitochondria (27, 30). The induction of cytochrome c release is not sufficient by itself to accelerate apoptosis, and p53 and FADD are required after cytochrome c release (27). We showed here that the T58A mutant of c-Myc, which is defective in transrepression, has also significantly reduced ability to induce cytochrome c release, accounting for its diminished ability to accelerate apoptosis in Rat1a fibroblasts. Because the decreased ability of T58A mutant to accelerate apoptosis is more profound in immortalized cells, it is possible that in immortalized cells cytochrome c release is more critical for Myc-mediated apoptosis, whereas in primary MEF activation of p19-ARF/p53 pathway is more critical.

Induction of cell cycle progression and apoptosis are two separable functions of c-Myc. Previous results showed that the ability of c-Myc to accelerate apoptosis is not dependent on the cell cycle phase of the cells (15, 19, 41, 54). However, the ability of c-Myc to induce cell cycle progression has always coincided with its ability to accelerate apoptosis. The analysis presented here allowed the separation of these two functions of c-Myc. The results obtained with the various deletion mutants in the amino-terminus region of c-Myc and with the T58A mutant clearly show that the ability of c-Myc to induce entry into cell cycle and to accelerate apoptosis are separable functions of c-Myc. These properties of c-Myc have a remarkable resemblance to the ability of E2F-1 to induce entry into cell cycle and apoptosis. Analysis of E2F-1 mutants showed that, although DNA binding is required, transcriptional activation is not necessary for the induction of apoptosis by E2F-1 (42). It was concluded that E2F-1 can show independent cell cycle progression and apoptotic functions. Interestingly, E2F-1 also activates the ARF-p53 pathway (25) and as with c-Myc this may account, at least in part, for the ability of E2F-1 to induce apoptosis in primary cells but not in established cell lines.