Distinct Roles for MAX Protein Isoforms in Proliferation and Apoptosis*

(Received for publication, April 24, 1997)

Hong Zhang Dagger , Saijun Fan § and Edward V. Prochownik Dagger §par

From the Dagger  Department of Molecular Genetics and Biochemistry, The University of Pittsburgh Medical Center, § Section of Hematology/Oncology, Children's Hospital of Pittsburgh, and  The University of Pittsburgh Cancer Institute, Pittsburgh, Pennsylvania 15213

ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES


ABSTRACT

MAX is a basic helix-loop-helix-leucine zipper protein that plays a central role in the transcriptional control of Myc oncoproteins. MYC-MAX heterodimers stimulate transcription, whereas MAX homodimers, or heterodimers between MAX and members of the MAD family of basic helix-loop-helix-leucine zipper proteins, repress transcription. Max exists in two major isomeric forms, MAX(L) and MAX(S), which differ from one another only by a 9-amino acid insertion/deletion. We show here that MAX(L) is much more effective at homodimeric DNA binding than MAX(S). In NIH3T3 cells, MAX(L) was able to repress a c-Myc-responsive reporter gene whereas MAX(S) either stimulated the reporter gene or had little effect on its expression. In comparison to control cell lines or those stably over-expressing MAX(S), MAX(L)-over-expressing cell lines showed reduced expression of transiently expressed or endogenous c-Myc responsive genes, grew more slowly, possessed a higher growth factor requirement, and showed accelerated apoptosis following growth factor deprivation. Differential effects on growth and apoptosis represent two previously unrecognized properties of MAX proteins. These can at least partly be explained by the differences in their DNA binding abilities and their effects on target gene expression.


INTRODUCTION

The c-MYC oncoprotein is an important member of the basic helix-loop-helix-leucine zipper family of transcription factors and is involved in such cellular processes as proliferation, transformation, differentiation, and apoptosis (1-4). Like most proteins of this class, c-MYC possesses sequence-specific DNA binding properties (5-7) and can transactivate synthetic or natural promoters containing c-MYC binding sites (8-12). At extremely high protein concentrations in vitro, c-MYC binds DNA as a homodimer (13). However, at lower concentrations and within cells, c-MYC homodimerization does not occur to any detectable degree (14, 15). Thus, the active DNA binding moiety appears to be a heterodimer comprised of c-MYC and another basic helix-loop-helix-leucine zipper protein, MAX (16, 17). Most of the biological properties of c-MYC have been attributed to its association with MAX (18-21). These heterodimers bind DNA strongly and activate transcription (18). Max also homodimerizes avidly, and in vitro and in vivo, both c-MYC-MAX heterodimers and MAX homodimers can be easily detected (14, 22-25). However, because MAX lacks a transactivation domain (26), MAX over-expression has been reported to repress the transcription of genes bearing c-MYC binding sites (8, 11, 27-29). Several model systems have demonstrated that the expression of such genes is directly dependent upon the levels of c-MYC-MAX heterodimers and inversely dependent upon the levels of MAX (8, 10, 11, 28, 29).

Alternate mRNA splicing produces two major MAX isoforms, the shorter of which (MAX(S)) is a 151-amino acid polypeptide with an apparent molecular mass of 21 kDa as determined by SDS-PAGE1 (16, 17, 23). The longer form MAX(L) is a 160-amino acid, 22-kDa polypeptide, identical to MAX(S) except for a 9-amino acid insert between residues 12 and 13. Both proteins, and their respective transcripts, are expressed at approximately equal levels in most cell types examined and, unlike c-MYC, are highly stable and unresponsive to the proliferative state of the cell (23, 30, 31). Both MAX isoforms are highly conserved between humans and such lower species as chickens, Xenopus, and zebrafish (32, 33).

Little is known regarding how the two MAX proteins differ from one another functionally, as most studies have been performed with only one of the two isoforms and/or have been of restricted biological scope. These studies are also difficult to compare as they have utilized different types of expression vectors, growth conditions, and biochemical and biological read-outs. Recent work has also emphasized the seemingly contradictory results of c-MYC reporter gene assays that can occur due to differences in cell density at the time of transfection (34). We have previously reported that MAX(L) and MAX(S) differ significantly in their DNA binding abilities (35). Using several synthetic oligonucleotides whose c-MYC binding sites differed only in their flanking regions, we showed that, in all cases, MAX(L) bound >20-fold more strongly to these sites than MAX(S). However, we could not absolutely rule out that these differences were, at least in part, the consequence of amino-terminal hexahistidine tags that had been introduced to facilitate purification of the recombinant, bacterially expressed proteins.

In the present study, we have compared the abilities of full-length, unmodified MAX(L) and MAX(S) to bind DNA and to modulate transiently transfected and endogenous c-MYC-responsive genes. We have also examined the effects of the two proteins on proliferation, growth factor requirement, and apoptosis in two different cell lines. We find that MAX(L) and MAX(S) differ markedly from one another in their abilities to affect these cellular properties. Because not all of the observed effects appear to be the result of the antagonism of c-MYC-MAX heterodimers, our results suggest that, depending upon the identity of the expressed isoform, as well as on its biological context, MAX proteins can either enhance or oppose many of the known activities of c-MYC. Because MAX plays a central role in both the positive and negative regulation of c-Myc activity, such opposing effects may provide a more sensitive control over this process.


MATERIALS AND METHODS

Expression and Purification of Recombinant Proteins

MAX cDNAs were amplified by the polymerase chain reaction using the following primers: forward, 5'-CGC TCA TGA GCG ATA ACG ATG ACA TC-3'; reverse, 5'-GCG CAA GCT TGC CTG CCC CGA GTG GCT TAG-3'. In the former case, the italics denote "GC clamps" and NcoI-compatible BspHI sites that were added to facilitate cloning into the NcoI site of the QE60 vector (Qiagen, Chatsworth, CA). Following this, cloned cDNAs were sequenced to confirm their identity. The cloning step permitted the expression of IPTG-inducible, full-length, non-hexahistidine-tagged MAX proteins in the pREP4 Escherichia coli strain (Qiagen). Cultures were induced for 4-6 h with IPTG (final concentration 1 mM) at which time the bacteria were concentrated by centrifugation, washed in PBS, and lysed by three rounds of freeze-thawing and sonication in 2 × binding buffer (35) that, for MAX(L), consisted of 100 mM KCl; 40 mM Hepes, pH 6.5; 2 mM EDTA; 2 mM dithiothreitol; 10% glycerol plus 1 mM phenylmethylsulfonyl fluoride (Sigma); and 1 µg/ml each of aprotinin, pepstatin, and leupeptin (Boehringer Mannheim). For MAX(S), the identical buffer was used except it's pH was adjusted to 7.6. Following clarification of each lysate at 10,000 × g for 10 min, a portion of the supernatant ("crude lysate") was used directly in DNA binding assays after adjusting for differences in MAX protein content using the pH 7.6 binding buffer. The remainder of the supernatant was applied to a DEAE-cellulose column equilibrated with the above pH 6.5 or pH 7.6 2 × binding buffer. MAX(L) and MAX(S) proteins were eluted with a stepwise KCl gradient, dialyzed against pH 7.6 2 × binding buffer, and stored in small aliquots at -80 °C. Hexahistidine-tagged N-Myc protein (35) was purified by nickel-agarose affinity chromatography essentially to homogeneity and dialyzed against 2 × binding buffer. Preliminary control experiments indicated that the different pH values used to bind MAX(L) and MAX(S) to DEAE-cellulose did not influence their DNA binding properties following their adjustment to pH 7.6 (not shown).

Electrophoretic Mobility Shift Assays

Electrophoretic mobility shift assays were performed essentially as described (35) using 50 ng of each purified protein or approximately 200 ng of crude bacterial lysate, equivalent to 20-50 ng of MAX. 5 µl of each protein was incubated at room temperature for 30 min with 5 µl of 2 × binding buffer containing 1 µg of poly(dI-dC) (Pharmacia Biotech Inc.) and 10,000 dpm of the 32P-end-labeled, double-stranded EO(GAC) oligonucleotide containing a single c-MYC consensus binding site (specific activity approximately 108 dpm/µg). Bound and free labeled complexes were resolved at room temperature on native 4% polyacrylamide gels in 1 × TBE buffer as described (35).

Plasmids for Mammalian Cell Expression

cDNA inserts for c-Myc (1.5-kilobase XhoI fragment, Ref. 36), MAX(L), and MAX(S) (16) were excised from their parental vectors and cloned into the XhoI site of the pSVLneo vector. This was derived from the pSVL vector (Pharmacia) by inserting into the unique EcoRI site a 2.5-kilobase EcoRI-linkered DNA fragment containing the neomycin resistance gene under the control of the SV40 early promoter (37). The p3xMyc-E1b-luc wild-type and p0Myc-E1b-luc mutant c-Myc reporter plasmid (11) were kind gifts from Dr. Roger Davis. The pSV2-beta gal plasmid was routinely included in all transfections to control for transfection efficiency. Plasmid DNAs were purified using a standard alkaline lysis method followed by two cycles of purification by CsCl/ethidium bromide equilibrium centrifugation.

Cell Culture and DNA Transfections

NIH3T3 cells were routinely grown in Dulbecco's modified minimal essential medium supplemented with 1 mM glutamine, 100 µg/ml streptomycin, 100 units/ml penicillin G, and 10% supplemented calf serum (Hyclone, Logan, UT). The 32D clone 3 murine myeloid cell line and the WEHI-238 murine B cell line were routinely grown in RPMI 1640 medium supplemented as described above except that fetal calf serum was substituted for supplemented calf serum. In addition, the medium used for 32D cell growth was supplemented with 10% (v/v) conditioned medium obtained from the IL-3-producing WEHI-238 cell line.

For transient expression assays, NIH3T3 cells were transfected using a standard calcium-phosphate-based procedure (38). Cultures were routinely plated at 5 × 105 cells/100-mm dish, transfected the following day, and harvested 2 days later for the assay of beta -galactosidase and luciferase (Analytical Laboratories, Ann Arbor, MI). Stable NIH3T3 transfectants were obtained using the above procedure except that 2 days after transfection, the monolayers were split 1:5 and cultured in the presence of 400 µg/ml G-418 (Life Technologies, Inc.). Individual G-418-resistant clones were isolated using glass cloning cylinders.

To obtain stably transfected 32D clones, 2 × 107 cells were collected by low speed centrifugation, washed twice in tissue culture medium containing IL-3 and fetal calf serum, and resuspended in 0.3 ml of the same medium. 10 µg of NdeI-linearized vector DNA in 25 µl of water was added. Electroporation was carried out in a 0.4-cm disposable cuvette (Bio-Rad) using a Bio-Rad apparatus set at 300 V and a capacitance extender set at 960 microfarads. Following electroporation, cells were incubated on ice for 10 min before being replated in standard IL-3 containing medium. Two days later G-418 was added to a final concentration of 400 µg/ml and maintained continuously. G-418-resistant clones were generally observed after about 2 weeks. Pooled cells representing at least several hundred independent clones were used for all experiments.

Analysis of MAX Protein Expression

NIH3T3 cell monolayers (approximately 80% confluency) were washed in methionine + cysteine-free Dulbecco's modified minimal essential medium (ICN Biomedical, Costa Mesa, CA). The medium was removed and replaced with 1 ml of the same medium containing 100 µCi of 35S-Translabel (ICN, specific activity 1.21 Ci/mmol) for 6 h. The monolayers were then washed exhaustively in ice-cold PBS and collected in 1 ml of standard RIPA buffer containing the same protease inhibitors as used for bacterial cultures (see above). Following centrifugation at 14,000 × g at 4 °C, 2 × 107 dpm of lysate supernatant in a total volume of 1 ml of RIPA buffer was incubated at room temperature for 2 h with a rabbit polyclonal anti-human MAX antibody at a final dilution of 1:300. Antigen-antibody complexes were removed by the addition of 50 µl of protein A-Sepharose (Bio-Rad) for an additional 2 h. Precipitates were washed three times in RIPA buffer, disrupted in loading buffer, and resolved by 12% SDS-polyacrylamide gel electrophoresis (SDS-PAGE). 32D cells were processed in an identical manner except that labeling was performed following the resuspension of 5 × 106 cells in 1 ml of methionine/cysteine-free Dulbecco's modified minimal essential medium containing 100 µCi of Translabel.

Cell Cycle Analysis

Cells were pelleted by centrifugation at 4 °C, washed in PBS, and fixed in 1 ml of ice-cold 70% ethanol for at least 1 h. The cells were pelleted twice in ice-cold PBS and resuspended in PBS containing 100 µg/ml RNase A and 50 µg/ml propidium iodide (Sigma). Cell cycle analyses were performed on a Becton-Dickinson FACSTAR fluorescence-activated cell sorter. 20,000 cells were analyzed for each assay with quantitation of cell cycle parameters being performed using single histogram statistics.

Apoptosis Studies

32D cells were cultured at 3 × 104 cells/ml in 30 ml of IL-3-supplemented medium containing 10% fetal calf serum. 24 h later, the cells were pelleted by low speed centrifugation and resuspended in the above medium minus IL-3. Aliquots of cells were removed at timed intervals, and the fraction of viable cells was determined by trypan blue exclusion. To evaluate the integrity of DNA, cells were washed twice in PBS and resuspended in 1 ml of lysis buffer (10 mM Tris-HCl, pH 8.0; 0.1 M EDTA; 0.5% SDS, 20 µg/ml RNase A, and 100 µg/ml proteinase K). After incubating at 37 °C for 4 h overnight, the lysates were phenol-extracted and ethanol-precipitated. DNA was redissolved in water and 10 µg was electrophoresed on a 2% agarose gel.

Ornithine Decarboxylase (ODC) Assays

NIH3T3 cells were plated at 105 cells/ml and maintained in log-phase growth for 2 days. Cells were then washed twice with PBS, scraped into conical centrifuge tubes, and pelleted at 1,000 × g for 10 min. ODC assays, using 200 µg of cellular extract, were performed as described (39) except that an incubation period of 2 h was used.


RESULTS

DNA Binding by MAX Proteins

We previously demonstrated that bacterially expressed MAX(L) binds strongly to a c-Myc binding site, whereas MAX(S) shows little detectable binding (35). In these experiments, however, each protein was expressed as a hexahistidine fusion to facilitate its purification, and it was therefore formally possible that these extraneous amino acids altered the DNA binding properties. To address this, we expressed both MAX proteins as non-fusion polypeptides in E. coli. Examination of IPTG-induced E. coli lysates by SDS-PAGE showed that MAX accounted for approximately 25% of the total protein. Following purification by DEAE-cellulose chromatography under non-denaturing conditions, MAX proteins were tested for their ability to bind the above c-Myc site. MAX(L) bound the labeled oligonucleotide strongly, whereas binding by the equivalent amount of MAX(S) was barely detectable (Fig. 1A). This effect was not due to an inadvertent loss of DNA binding ability by MAX(S) during its purification as it retained the ability to bind the probe in association with purified N-Myc protein (lane 4). The differences in DNA binding by the two MAX proteins were also apparent prior to purification (Fig. 1B). In some experiments, and at higher protein concentrations, we have observed some DNA binding by MAX(S), but it has always been at least 20-fold less than seen with a comparable amount of MAX(L). These results, with both purified and crude MAX proteins, confirmed our previous observations (35) and indicated that the observed differences in DNA binding were not the artifactual result of changes in polypeptide structure.


Fig. 1. DNA binding by purified and crude MAX proteins. MAX proteins were produced as unmodified, full-length polypeptides in E. coli using the QE-60 expression vector (Qiagen). MAX proteins were purified from IPTG-induced cultures by non-denaturing DEAE-cellulose chromatography to yield individual MAX proteins of >90% purity (A) or were used directly from crude bacterial lysates of approximately 25% purity (B). N-MYC was produced as a hexahistidine fusion protein (50) and purified using nickel-agarose chromatography. Each protein (approximately 20 ng) was tested for DNA binding with the 32P-labeled double-stranded, palindromic EO(GAC) oligonucleotide (50) using non-denaturing polyacrylamide gel electrophoresis. A, MAX(L) showed strong binding to the 32-P-labeled probe (lane 2), whereas MAX(S) bound poorly (lane 3). The ability of MAX(S) to dimerize with purified N-MYC and bind the probe (lane 4) indicated that MAX(S) was in an active state. Similar results were obtained with crude extracts of the proteins (B), indicating that the differences in DNA binding were not related to protein purification. Lane 2 shows the absence of MAX(L) DNA binding activity in crude extracts of uninduced E. coli.
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Reporter Gene Response to MAX(L) and MAX(S)

The observed differences in DNA binding by MAX(L) and MAX(S) in vitro suggested that the two proteins might have dissimilar effects on a c-Myc-responsive reporter gene in vivo. We tested this idea using transient transfections in NIH3T3 cells. In the first series of experiments, a constant amount of a c-Myc responsive reporter construct (11) was co-transfected with increasing amounts of either a MAX(L) or MAX(S) expression vector. We assumed that the basal expression of luciferase activity by the reporter in the absence of any other co-transfected plasmids at least partly reflected the levels of endogenous c-Myc and MAX protein activities in these cells. As seen in Fig. 2A, MAX(L) suppressed luciferase activity in a dose-dependent manner, consistent with the idea that at high DNA concentrations, MAX(L) homodimers predominate over c-Myc-MAX heterodimers, compete for c-Myc binding sites, and repress reporter gene transcription (10, 11, 29). The small amount of residual luciferase activity seen at the highest MAX(L) concentrations probably represents the c-Myc/MAX-independent activity of the core E1b promoter. In contrast, co-transfected MAX(S) resulted in an increase in reporter gene activity which, at the highest concentrations tested, was 3-4-fold over basal levels. At these concentrations the differences between MAX(L) and MAX(S) were over 7-fold. This observation suggested that high concentrations MAX(S) allowed its heterodimerization with endogenous c-Myc, resulting in an increase in the number of transcriptionally active c-Myc-MAX heterodimers. However, once all available c-Myc was dimerized, further increases in MAX(S) were unable to repress transcription. This is consistent with the weak DNA binding activity of MAX(S) protein purified from E. coli (Fig. 1).


Fig. 2. Regulation of a c-Myc-regulated reporter by MAX(L) and MAX(S). A, NIH3T3 cells were transiently transfected with 5 µg of the 3xMyc-E1b-luc reporter plasmid and the indicated amounts of MAX(L) (dark boxes) or MAX(S) (open boxes) expression vectors. Luciferase activity was measured 2 days later. The values shown are the averages of three separate experiments ± 1 S.E. B, 5 µg of 3xMyc-E1b-luc was expressed either in the absence (-) or presence (+) of an equivalent amount of pSVL-Myc expression vector (hatched boxes) along with the indicated amounts of pSVL-MAX(L) or MAX(S) expression vectors. Luciferase activities were determined as described in A.
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We next tested the effect of expression of each MAX isoform on reporter gene expression in the presence of co-expressed c-Myc. As shown in Fig. 2B, c-Myc alone stimulated reporter gene activity an average of 4-fold, consistent with the degree of activation reported by many other groups (8, 10, 11, 29). With increasing amounts of the co-transfected MAX(S) expression plasmid, only minimal (1.5-2 fold) additional stimulation of the reporter was observed. In contrast, the transfection of the MAX(L) expression plasmid resulted in a dose-dependent decrease in reporter gene expression of up to 8-fold. The difference between MAX(L) and MAX(S) at the highest concentrations examined was over 10-fold. These results were consistent with those presented in Fig. 2A and indicated that, under the conditions employed for these experiments, MAX(S) was stimulatory whereas MAX(L) was generally inhibitory toward c-Myc reporter genes. Similar results were observed with a CAT reporter construct driven by an ornithine decarboxylase (ODC) promoter, a known physiologic c-Myc target (9) (not shown). In control experiments, no regulation by c-Myc or MAX was seen in transient transfections employing either E1b-luciferase or ODC-CAT vectors containing absent or mutant c-Myc binding sites (9, 11) (not shown).

Stable Expression of MAX Proteins in NIH3T3 Cells

To study the biological consequences of MAX protein over-expression in more detail, we established lines of NIH3T3 cells stably transfected with either MAX(L) or MAX(S) expression plasmids. We selected representative clones and measured the amount of MAX protein by metabolic labeling. Fig. 3 shows that both the parental NIH3T3 cell line as well as single cell clones transfected with the empty vector alone expressed very low levels of Max. In contrast, MAX-transfected clones expressed the expected individual isoform at high levels.


Fig. 3. Immunoprecipitation of MAX proteins from NIH3T3 cells. NIH3T3 cells were stably transfected with MAX(L) or MAX(S) expression vectors and selected for G-418 resistance. Single cell clones were expanded and metabolically labeled with [35S]methionine/cysteine. MAX proteins were immunoprecipitated from equivalent amounts of total cell lysate and resolved by SDS-PAGE. Note that parental NIH3T3 cells and vector-transfected control clones contain barely detectable amounts of Max.
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A representative clone from each group was transfected with the p3xMyc-E1b-luc plasmid to determine the effect of stable over-expression of each MAX isoform on a c-Myc-responsive gene. As seen in Fig. 4A, the expression of the plasmid was only slightly elevated above the control in MAX(S)-over-expressing cells, whereas in MAX(L)-over-expressing cells, plasmid expression was reduced by about 7-fold. In control experiments, a reporter plasmid with mutant c-Myc binding sites (11) was expressed at equivalent levels in all three cell lines thus indicating that the above results did not simply reflect a generalized lower rate of transcription in the MAX(L)-over-expressing cell line.


Fig. 4. Expression of c-Myc-responsive genes in representative NIH3T3 cell clones over-expressing MAX(L) (clone L2) or MAX(S) (clone S18) plus a vector-transfected clone (clone Neo7). A, cells were transiently transfected with the c-Myc-responsive 3xMyc-E1b-luc reporter plasmid (32) (filled boxes) or the mutant p0Myc-E1b-luc mutant plasmid that lacks functional c-Myc binding sites (32) (hatched boxes). Each plate received 5 µg of each plasmid as well as 5 µg of pSV2beta -gal to control for transfection efficiency. Two days later, cells were harvested and assayed for luciferase activity following correction for differences in beta -galactosidase activity. The results shown represent the average of three independent experiments, each performed in duplicate, ± 1 S.E. B, each of the indicated clones was plated at a density of 3 × 104 cells/ml and allowed to grow for 3 days before harvesting. Endogenous Odc activity was then measured as described previously (47). The results shown represent the average of three experiments ± 1 S.E.
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To determine how each MAX isoform affected the expression of an endogenous, c-Myc-responsive gene, we also measured ODC enzyme levels in each of the above clones. As seen in Fig. 4B, the average ODC level was reduced more than 7-fold in the MAX(L)-over-expressing cell line, whereas no significant change from the control was seen in the MAX(S)-over-expressing cells.

Growth rates for each of the clones shown in Fig. 3 were also determined. Fig. 5 shows that each of the MAX(S) over-expressing cell lines grew at rates that were slightly faster than those of controls. More strikingly, each of the three MAX(L)-over-expressing clones was growth retarded, showing 8-20-fold fewer cells compared with controls after 7 days in culture. These differences were not due to variations in initial plating efficiencies of MAX(L)-over-expressing cells2 and persisted throughout the duration of the experiment.


Fig. 5. Growth curves of MAX-over-expressing NIH3T3 clones. 105 cells from the clones analyzed in Fig. 3 were seeded onto 100-mm tissue culture dishes and maintained in 10% serum. Cells from triplicate plates were counted at the indicated times.
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To determine the basis for the differences in growth rates among the three groups of cell lines, we studied their cell cycle distribution at various times after plating. Surprisingly, only small and transient differences were seen in cell cycle patterns among all of the clones during logarithmic growth (not shown). This suggested that the reduced growth rate of MAX(L)-over-expressing cells was attributable to a reduced rate of progression through each phase of the cell cycle without altering the overall distribution pattern. To test this directly, control, MAX(L), or MAX(S)-over-expressing cells were arrested in G0 by serum deprivation for 24 h. The cells were then stimulated with serum in the presence of nocodazole to prevent cell cycle progression beyond the first mitotic stage. This allowed us to clearly measure the rate of progression through a single cell cycle by flow cytometry. As seen in Fig. 6, MAX(S)-over-expressing cells traversed the cell cycle slightly faster than control cells. This was seen best at 16 and 18 h after serum stimulation. At the latter time, virtually all of the MAX(S) over-expressing cells had reached G2/M, whereas nearly half the control cells remained in G1 or S phase. By 24 h, however, both these lines had completely entered G2/M. This finding was consistent with the slightly faster growth rate of MAX(S)-over-expressing cells seen in Fig. 5. In contrast to these subtle findings, MAX(L)-over-expressing cells showed a significant delay in cell cycle progression at all stages. For example, at 14 h after serum addition, the MAX(L) population had barely begun to exit the G0/G1 phase, whereas a majority of cells in both the control and MAX(S) population had entered S phase and even progressed to G2/M. Similarly, by 20 h, a significant fraction of the MAX(L) population remained in G0/G1 and S phases, whereas virtually the entire control population had traversed the entire cell cycle. Independent confirmation of this lag was obtained by determining the mitotic index of each of the three populations of cells 20 h after serum stimulation. At this time, both control and MAX(S)-over-expressing cultures showed 90-95% of the populations to be in mitosis whereas, in MAX(L)-over-expressing cultures, only 46% of the cells had entered mitosis. Two additional experiments gave virtually identical results as did similar studies performed with each of the other clonal cell populations (not shown). Our findings thus indicate that the slower growth of MAX(L)-over-expressing cells is largely attributable to a prolonged cell cycle that results from a lengthening of all phases without any change in the overall distribution pattern.


Fig. 6. MAX(L)-over-expressing cells show a slower rate of transit through all phases of the cell cycle. Representative NIH3T3 clones were serum-starved for 24 h. At time 0, the cells were stimulated with medium containing 10% serum and 0.4 µg/ml nocodazole to prevent passage past the first G2/M phase. At the indicated times after serum re-stimulation, cultures were harvested to assess nuclear DNA content by fluorescence-activated cell sorting.
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We also examined each of the MAX(L)-over-expressing clones to determine if their slower rate of growth might be explained by a higher rate of apoptotic cell death. No such differences in the rates of cell death were seen under standard conditions of logarithmic growth (not shown), and this was confirmed by the absence of any cells with subdiploid amounts of DNA in our cell cycle analyses (Fig. 6). Nevertheless, in a number of settings following growth factor withdraw, c-Myc over-expression accelerates apoptotic cell death through a process that is dependent upon Max (3, 20, 40-42). Therefore, to determine whether either of the MAX protein isoforms could influence cell death, we deprived representative clones of serum and monitored the fraction of surviving cells. Under these conditions, control and MAX(S)-over-expressing cells showed identical rates of cell death, whereas MAX(L)-over-expressing cells died at a 2-3-fold faster rate (Fig. 7A). This was confirmed by light microscopic studies of cell monolayers that showed a significantly greater number of rounded, refractile cells among the MAX(L) population (Fig. 7B) and by electron microscopic studies demonstrating the morphological features typical of apoptosis, including nuclear condensation and cytoplasmic "blebbing" (not shown). Despite repeated attempts, we were unable to demonstrate the presence of the nucleosome-sized DNA fragments that are a cardinal feature of apoptosis. The inability of some murine fibroblasts, including NIH3T3 cells, to display this property following treatment with known apoptotic stimuli has been previously noted (43-45). Nevertheless, our results are consistent with the idea that MAX(L) over-expression could, under appropriate conditions, modulate cell death in NIH3T3 cells and that this process probably utilized apoptotic pathways.


Fig. 7. Accelerated death of MAX(L)-over-expressing NIH3T3 cells following serum removal. A, vector control clone Neo7 (black-square), MAX(S) clone S3 (bullet ), and MAX(L) clone L2 (square ) were seeded at an initial density of 2 × 105 cells/100-mm plate and allowed to grow for approximately 24 h. The plates were washed and replaced with serum-free medium at time 0. At daily intervals thereafter, cells from both the monolayer and supernatant were harvested and stained with trypan blue to determine the viable fraction. B, phase-contrast photomicrographs of monolayers from each clone 4 days after removal of serum. Note the relative paucity of viable, fusiform-shaped cells in the MAX(L)-over-expressing cell line.
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Expression of MAX Proteins in 32D Cells

To study the effect of MAX protein over-expression on apoptosis in a better system, as well as to generalize our findings in NIH3T3 cells, we utilized the 32D murine myeloid cell line (46). Proliferation and survival of 32D cells are dependent on the hematopoietic cytokine IL-3 whose withdrawal results in rapid and easily quantifiable apoptotic cell death (40). Previous work has shown that c-Myc over-expression accelerates apoptosis in these cells following IL-3 withdrawal, thus suggesting that MAX proteins might be involved in this process (40, 47). Stably transfected populations of 32D cells were therefore generated, and the over-expression of MAX proteins was confirmed in pooled clones by immunoprecipitation of [35S]methionine-labeled cell extracts (Fig. 8). We then compared the growth rates of control and MAX-overexpressing 32D cells at several IL-3 concentrations. At the highest IL-3 concentration tested, control and MAX(S)-overexpressing cells were indistinguishable and showed robust growth, whereas MAX(L)-overexpressing cells grew at a rate one-half to two-thirds that of control cells (Fig. 9A). More marked differences were seen as the IL-3 concentration became limiting. For example, when the amount of cytokine was reduced by one-half, MAX(L)-over-expressing cells showed significant growth impairment, whereas MAX(S)-over-expressing and control cells remained indistinguishable (Fig. 9B). Even more profound differences among the three cell lines became apparent at the lowest IL-3 concentration tested (Fig. 9C). Although control cultures showed a significant impairment in growth, they still proliferated significantly faster than MAX(L)-over-expressing cells. In marked contrast, MAX(S)-over-expressing cells proliferated 4-fold more rapidly than control cultures and 10-fold more rapidly than MAX(L)-over-expressing cells. These results indicate that, compared with control 32D cells, MAX(L)-over-expressing cells have decreased IL-3 sensitivity, whereas MAX(S)-over-expressing cells have an increased sensitivity.


Fig. 8. Immunoprecipitation of MAX proteins from 32D cells. Pooled G-418-resistant clones transfected with the indicated plasmids were metabolically labeled with [35S]methionine/cysteine. MAX proteins were immunoprecipitated and analyzed by SDS-PAGE as described in the legend to Fig. 3.
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Fig. 9. Growth curves for 32D cells in different IL-3 concentrations. 2 × 104 pooled G-418-resistant clones of 32D cells transfected with the empty vector alone (black-square), with pSVL-MAX(S) (bullet ), or with pSVL-MAX(L) (square ) were plated in medium supplemented with 10% (A), 5% (B), or 2% (C) WEHI-238-conditioned, IL-3-containing medium. Viable cell counts were determined in triplicate cultures at the indicated intervals and are represented as the average number of cells/plate ± 1 S.E.
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The increased requirement for IL-3 by MAX(L)-over-expressing 32D cells suggested that they might be more sensitive to apoptotic cell death following complete removal of the cytokine. This was confirmed when all three cell lines were studied in parallel (Fig. 10A). Over the 24-h course of the experiment, MAX(L)-over-expressing cultures showed a 5-8-fold greater accumulation of dead cells than either control or MAX(S)-over-expressing cells. That death was due to apoptosis was confirmed by examination of nuclear DNA which showed the expected increase in nucleosome-sized fragments (Fig. 10B).


Fig. 10. Accelerated apoptosis in MAX(L)-over-expressing 32D cells. A, logarithmically growing control (black-square), MAX(S) (bullet ), or MAX(L) (square ) over-expressing 32D cells (>95% viability) were washed free of IL-3-containing medium and cultured in IL-3-depleted medium for the indicated times. The percentage of viable cells was determined by trypan blue staining. B, nuclear DNA fragmentation was assessed in each cell line either at the time of IL-3 withdrawal (0 h) or 24 h later. Note that, at the latter time, MAX(L)-over-expressing cells demonstrate a greater amount of DNA fragmentation, consistent with the results in A.
[View Larger Version of this Image (19K GIF file)]


DISCUSSION

The ability of MAX to homodimerize or heterodimerize with c-MYC suggests that MAX can participate in both the transcriptional activation and repression of c-MYC-responsive genes (29). Additional modes of MAX regulation, however, have made it apparent that the model is subject to many other positive and negative influences and is thus more complicated than originally proposed. For example, either CKII phosphorylation or dimerization with a MAX isoform lacking a basic domain can negatively regulate DNA binding (30, 48-50). At least four additional members of the basic helix-loop-helix-leucine zipper family, known as MAD proteins, can also heterodimerize with MAX and repress transcription (51-53). Their contribution to MYC-regulated transcription may depend upon their abundance as well as certain cell type-specific factors. Qualitative differences among the various MAD-MAX complexes may also exist but have not been explored.

Although MAX(L) and MAX(S) represent the major isoforms of the protein and are highly evolutionarily conserved (33), how they differ functionally from one another has not been determined. Most published studies on the effects of c-MYC and MAX have examined only one MAX isoform and/or have assessed a limited number of properties (8, 10, 28, 29, 31). To our knowledge, none of the work has compared the properties of cell lines stably expressing either MAX isoform.

We have re-examined several of the known properties of MAX proteins and have shown that MAX(L) and MAX(S) regulate DNA binding, cell cycle progression, and apoptosis in distinct ways. One of the most striking differences between MAX(L) and MAX(S) is in their intrinsic DNA binding properties (Fig. 1). These differences, previously noted with hexahistidine-tagged polypeptides (35), have now been confirmed with full-length, unmodified proteins. Although we have occasionally detected DNA binding by MAX(S), it is observed at a level similar to that seen with "forced" c-MYC homodimers (5) and, in our hands, is always at least 20-fold less than obtained with comparable amounts of MAX(L).

It is possible to understand our in vivo results in light of the above DNA binding studies. Thus, the findings in transient transfection assays that MAX(L) is a more potent repressor of a c-MYC reporter construct than MAX(S) (Fig. 2) can be interpreted as indicating that MAX(L) homodimers compete with c-Myc-MAX heterodimers for the same DNA binding sites. In contrast, DNA binding by c-Myc-MAX(S) remains largely unopposed by MAX(S) homodimers so that only reporter gene stimulation is seen. However, the extent of this effect may depend upon the amount of endogenous c-Myc protein, and at sufficiently high MAX(S):c-Myc ratios, repression may also occur. Similar findings were obtained in cells stably over-expressing individual MAX isoforms. Thus, fibroblasts over-expressing MAX(L) showed significantly less expression of either a transiently expressed or endogenous c-Myc-responsive gene (Fig. 4). In a more biological setting, those lines expressing MAX(L) were growth retarded, whereas those expressing MAX(S) either behaved no differently from control lines or were growth-stimulated (Figs. 5 and 9). However, the nature and extent of these opposing effects was dependent upon the cellular context (for example, see Fig. 9).

The contrasting effects of MAX(L) and MAX(S) on cell cycle progression and apoptosis represent heretofore unrecognized properties of these proteins. In retrospect, these attributes are not unexpected, given the known role for c-MYC in G0/G1 progression and apoptosis (40, 41, 50, 54, 55). Our observations argue that cellular decisions pertaining to these processes may hinge upon the proper levels of both c-MYC and MAX, as well as other factors. In the case where mitogenic stimulation results in high levels of endogenous c-MYC, MAX proteins, particularly MAX(S), may exert either little negative effect on proliferation and apoptosis or may even stimulate proliferation. However, as c-MYC becomes limiting, as in the case of serum-deprived fibroblasts (Fig. 7) or IL-3-deprived 32D cells (Fig. 9), MAX(L) may act to limit the proliferative response. With more profound c-MYC depletion, severe growth impairment and/or apoptosis may ensue as a result of the suppression of c-MYC-responsive genes by a relative excess of MAX(L) (Fig. 10). These results suggest that the control of cellular proliferation and apoptosis by the c-MYC-MAX network can tolerate some variation in the relative levels of these proteins but ultimately results in cell death when these limits are violated in either direction. Apoptosis may thus be viewed as a decision based upon the cell's assessment and integration of absolute and relative c-MYC and MAX levels together with its ability to proliferate or enter a quiescent state in response to these levels. In turn, these latter activities will be determined by the availability of growth factors which by themselves may affect proliferative and apoptotic pathways.

The observation that MAX(L)-over-expressing cells contain lower levels of Odc (Fig. 4B) provides a partial, although necessarily incomplete, biochemical explanation for their biological behavior. Odc, the rate-limiting enzyme in polyamine biosynthesis, is cell cycle-regulated and is required for S phase entry (39, 56). Thus, the lower Odc levels in MAX(L)-over-expressing cells might be sufficient to account for their slower rate of cell cycle progression, particularly through S phase. On the other hand, at least some of the pro-apoptotic activity of c-Myc has been linked to its direct up-regulation of Odc (39) although other c-Myc target genes are likely to be important for this process (42). Although the specific relationship between Odc levels, proliferation, and apoptosis remains to be established with regard to each of the Max isoforms, the cell lines reported here should be useful in establishing these associations.

Because c-Myc over-expression has been associated with accelerated apoptosis in IL-3-deprived 32D cells (40), we examined c-Myc transcript levels in all three cell lines following IL-3 removal. In all three cases, c-Myc was expressed at equivalent levels and became undetectable within 1-3 h following cytokine withdrawal.2 These observations argue that the apoptosis mediated by MAX(L) over-expression not only does not require concomitant c-Myc over-expression but may well be c-Myc-independent. However, we cannot rule out the possibility that small amounts of c-Myc-MAX(L) heterodimer may be sufficient to promote apoptosis. At the very least, our results suggest that the apoptotic pathways utilized by MAX(L) are distinct from those utilized by c-Myc, which generally, but not invariably, require high level and inappropriate expression of the protein (40, 41, 57).

Our results are consistent with a model in which three types of DNA binding complexes exist in subconfluent cultures of mammalian cells: c-Myc-MAX(S) and c-Myc-MAX(L) heterodimers, both of which will be stimulatory, and MAX(L) homodimers which will be repressive. MAX(S) homodimers will also be present but will be relatively ineffective at DNA binding. The overall transcriptional state of a c-Myc-responsive gene will thus reflect the relative abundance of the DNA binding dimers (34). Over-expressed MAX(L) will be repressive due to the greater abundance of MAX(L) homodimers than c-Myc-MAX(L) heterodimers. When co-expressed with c-Myc, low concentrations of MAX(L) might initially be stimulatory due to its predilection to heterodimerize. However, repression will predominate at higher MAX(L) levels as homodimers form and compete with c-Myc-MAX heterodimers. MAX(S) excess will have little inhibitory effect on target gene expression due to the poor intrinsic DNA binding activity of MAX(S) homodimers. Co-expression of c-Myc and MAX(S) will stimulate reporter gene expression due both to increased c-Myc-MAX(S) heterodimer formation and the weak repressive activity of MAX(S) homodimers. High levels of MAX(S) expression, perhaps coupled with limiting c-Myc expression, might eventually produce some repression and could account for the few reports of MAX(S)-mediated suppression that have been described.

It is important to bear in mind that the above model is likely to be oversimplified as it considers DNA binding only by unmodified MAX homodimers. CKII-mediated phosphorylation and associations with MAD proteins may affect each MAX isoform differently (27, 35, 48, 49, 51-53). Differences in the affinities of various MAX-MAD heterodimers for the mSin3 repressor may also exist but have not been explored (32, 58). Such differences, if they exist, may well be highly cell type-specific and could also account for much of the variability in MAX function that has been observed.

Despite these questions, our results indicate distinct roles for MAX(L) and MAX(S) in c-Myc target gene expression, cellular proliferation, and apoptosis.


FOOTNOTES

*   This work was supported by National Institutes of Health Predoctoral Training Grant 538572 (to H. Z.) and by National Institutes of Health Grant HL33741 (to E. V. P.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
par    To whom correspondence should be addressed: Section of Hematology/Oncology, Children's Hospital of Pittsburgh, 3705 Fifth Ave., Pittsburgh, PA 15213. Tel.: 412-692-6797; Fax: 412-692-5723.
1   The abbreviations used are: PAGE, polyacrylamide gel electrophoresis; IPTG, isopropyl-1-thio-beta -D-galactopyranoside; PBS, phosphate-buffered saline; ODC, ornithine decarboxylase; IL, interleukin.
2   H. Zhang and E. V. Prochownik, unpublished observations.

ACKNOWLEDGEMENTS

We thank Seth Corey for providing 32D and WEHI-238 cells and Xiaoying Yin for assistance in Northern blotting.


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