Expression of mRNAs encoding mammalian chromosomal proteins HMG-I and HMG-Y during cellular proliferation

EXPERIMENTAL

CELL RESEARCH

187,69-76 (1990)

Expression of mRNAs Encoding Mammalian Chromosomal Proteins HMG-I and HMG-Y during Cellular Proliferation KENNETH *Program

R. JOHNSON,* JANE E. DISNEY,* CAROL R. WYATT,? AND RAYMOND REEVES*+? in Genetics and Cell Biology, TDepartment of Microbiology, and *Program in Biochemistry/Biophysics, Washington State University, Pullman, Washington 99164

of nuclear proteins that are thought to be important structural components affecting the conformation of chromatin [4]. The proteins HMG-I and HMG-Y [5,6] are particularly interesting members of this group because they are expressed at elevated levels in proliferating, undifferentiated cells [4-81. Furthermore, our laboratory has recently demonstrated by in situ immunolocalization techniques that these nonhistone proteins are specifically localized to the A/T-rich G/Q- and C-bands of human and mouse metaphase chromosomes [9]. Thus, the HMG-I and HMG-Y proteins most likely play an important in viuo role in cell division [5,9] and/or in maintaining the undifferentiated state of cellular chromatin [ 7-91. Isolation and sequencing of cDNAs encoding the murine [lo] and human [ 111 HMG-I and HMG-Y proteins, along with amino acid analysis of these proteins [ll, 121, have shown that HMG-Y is identical to HMG-I except for an internal deletion of 11 amino acids. The HMG-I and HMG-Y isoforms are most likely the result of alternative processing of pre-mRNAs from a single functional gene [ll]. In this paper, we use the designation HMG-I/Y when referring collectively to both isoforms. We have previously shown by Northern blot analysis that HMG-I/Y mRNAs, like the proteins they encode, are preferentially expressed in proliferating, undifferentiated cells [lo, 111. However, the effects of cellular proliferation are difficult to separate from those of cellular differentiation. In this paper we compare the expression of HMG-I/Y mRNAs in quiescent versus proliferating attached murine NIH/3T3 fibroblasts and suspension cultures of human K-562 erythroleukemic cells. These two cell types were selected for study because under normal growth conditions both lines exhibit a relatively undifferentiated cellular phenotype. We also analyze HMG-I/Y mRNA levels at various stages of the cell cycle in the NIH/3T3 cells partially synchronized by seeding from confluent cultures. We show that HMG-I/ Y mRNA levels gradually increase during the first 16 h after seeding of NIH/3T3 cells but thereafter remain relatively constant. This pattern of mRNA expression is markedly different from that exhibited by the mRNAs of the major histone proteins, whose expression is tightly

The high mobility group chromosomal proteins HMG-I and HMG-Y are closely related isoforms that are expressed at high levels in rapidly dividing, undifferentiated mammalian cells. We analyzed HMG-I/Y mRNA levels at various cell cycle stages in murine NIH/ 3T3 Abroblasts partially synchronized by seeding from quiescent, contact-inhibited cultures. Flow microfluorometic analysis of DNA content demonstrated a comparable degree of synchronization in such seeded NIH/ 3T3 cell populations as is obtained by serum deprivation or other means and has the added advantage of avoiding the use of possibly detrimental inhibitors or metabolic starvation to induce such synchrony. We show that HMG-I/Y mRNA levels gradually increase in NIH/3T3 cells during the first 16 h after seeding (Go/ G1 to late S phase), but thereafter remain constant, in contrast to the cell cycle-regulated expression of the histone H3 gene. Although there is a 6-fold increase in HMG-I/Y expression during the transition from quiescent to proliferating NIH/3T3 cells, there is a much greater difference in expression (15- to 50-fold) among different cell types, possibly related to their state of differentiation. The HMG-I/Y mRNAs appear to be very stable; there was no decrease in their levels 6 h after actinomycin D transcription termination. The proportion of HMG-I to HMG-Y mRNAs was greater in the human than in the murine cells examined, appeared to be greater in proliferating than in quiescent cells, and did not always correspond with the HMG-I to HMG-Y protein ratio. o mso Academic PEWS, IZIG

INTRODUCTION Changes in chromatin structure are thought to affect the regulation of gene expression (reviewed in Refs. [l, 21). Major patterns of transcriptionally active and inactive chromatin appear to be established during the steps of cellular differentiation since they do not seem to change appreciably during the cell cycle [3]. The high mobility group proteins (HMGs) are an abundant class ’ To whom reprint requests should be addressed. 69

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Copyright 0 1990 by Academic Press, Inc. All rights of reproduction in any form reserved.

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coupled to DNA synthesis [ 131. Although we detected a 2- to 6-fold increase in expression of HMG-I/Y mRNA in the transition from cellular quiescence to proliferation in both NIH/3T3 and K562 cells, we detected a much greater difference (15- to 50-fold) in the amount of HMG-I/Y mRNA between different cell types (for example, between K562 and human HUT-78 cells or human placental tissue), possibly related to differences in their state of cellular differentiation. In contrast to the extreme instability of the mRNAs encoding the c-myc proto-oncogene protein (another nuclear phosphoprotein that is expressed at high levels in undifferentiated but not differentiated cells [14]), HMG-I/Y mRNAs appear to be very stable; their levels remained high 6 h after actinomycin D-induced transcription termination. In this study, we also compare relative levels of the individual HMG-I and HMG-Y mRNAs in several different cell types and at different stages of cellular proliferation. We find that the proportion of HMG-I to HMG-Y mRNAs is greater in the human than in the murine cells examined, appears to be greater in proliferating cells than in quiescent cells, and does not always correspond precisely with the HMG-I to HMG-Y protein ratio found within cells. MATERIALS

AND

METHODS

Cell culture and synchronization. NIH/3T3 murine cells obtained from the American Type Culture Collection were maintained as attached monolayers in Dulbecco’s modified Eagle’s (DME) media supplemented with 10% calf serum. Suspension cultures of human K-562 erythroleukemia and human HUT-78 cells were maintained in Roswell Park Memorial Institute (RPMI) media supplemented with 5% fetal bovine serum. All cultures were maintained at 37°C and 5% CO2 and routinely passaged at subconfluence. For experiments with suspension cultures of K562 and HUT-78 cells, exponentially growing log-phase cells were harvested l-2 days after passaging of the cells by dilution into fresh medium. Stationary or quiescent stage K562 or HUT-78 cells were harvested 3-4 days after passaging by dilution. Partial cell synchrony of attached NIH/3T3 cells was achieved by seeding cells from subconfluent cultures into T-75 flasks (15 ml/flask) at a concentration of 1 X 10’ cells per milliliter, growing the cells to confluence, and maintaining the cells at confluence for an additional 36 h. Flow microfluorometric analysis of the cells in such contact-inhibited confluent cultures indicated that they were arrested in a stable but reversible G1/Go quiescent state similar to that induced by low serum concentrations in Swiss 3T3 cells [15]. The confluent cell cultures were trypsinized, washed with fresh media, and seeded at 1 X lo5 cells per milliliter into T-150 flasks. Between 6 X 10s and 1 X lo7 cells were collected by trypsinization at various time points postseeding. For each time point approximately 5 X lo6 cells were washed, fixed with cold 70% ethanol, and stained with chromomycin A3 for flow microfluorometric analysis of DNA content as described by Gray and Coffin0 [16]. The remainder of the cells were centrifuged and the pellet immediately frozen at -90°C for subsequent RNA extraction. Cell cycle progression was monitored by flow microfluorometry using a Becton-Dickinson FACS analyzer and Consort 30 computer (BectonDickinson Immunocytometry Systems, Mountain View, CA). RNA preparation and Northern blot analysis. Cytoplasmic RNA was isolated and purified from cells in culture by a modification of

ET AL. the method of Gough [ 171. RNA from human placenta was prepared according to Cathala et al. [18]. In some cases, poly(A)+ RNA was selected from total RNA on oligo(dT) cellulose columns [ll]. Purified RNA samples (15 pg/lane) were electrophoresed in formaldehyde-denaturing 1.2% agarose gels. Denatured RNAs from the gels were transferred by overnight capillary blotting to ZetaProbe membranes (Bio-Rad) and UV-crosslinked. The membranes were prehybridized for 3 h at 42°C in 40% formamide, 0.5 M sodium phosphate, pH 7.2, 1 mM EDTA, 5% SDS, and 1% bovine serum albumin [19]. Denatured, labeled probe DNA was then added (2 X lo6 cpm/ml), and the hybridization was continued for at least 20 h. Separate blots containing identical quantities of RNA per lane were used for each DNA probe. Final washes were in 1X SSC and 0.1% SDS at 65°C. DNA probes for Northern blot hybridizations were labeled with [32P]dATP by the random primer method [20] to a specific activity of 0.5-l X 10’ cpm/pg. The 455-bp Sac1 to PstI fragment of the murine HMG-Y cDNA [lo], containing the evolutionarily conserved protein coding region, was used for hybridizations with murine and human RNAs. Likewise, the evolutionarily conserved protein coding region of human HMG-17 cDNA (290-bpEcoR1 toPst1 fragment) [21] was used for both murine and human RNA hybridizations. The 1250-bp PstI fragment from chicken fl-tubulin cDNA [22], the entire pSVc-myc-1 plasmid containing the second and third exons of the cellular mouse myc gene [23], and the 450-bp Sac1 to AvaI fragment containing a conserved domain of the Drosophila histone H3 gene [24] were also used as probes in Northern blot hybridizations. Exposure times for autoradiograms were approximately 5 h when the HMG-Y, HMG-17, and P-tubulin cDNA probes were used, and 20 h or longer when the histone H3 and c-myc gene probes were used. of the relative abundance of HMG-I and HMG-Y Determination mRNAs. Northern blot analysis employing agarose gels cannot easily distinguish between the closely related mRNAs that encode the isoform HMG-I and HMG-Y proteins, since these mRNAs differ only by the presence or absence of 33 bases (nucleotides 103-135) [ll]. Therefore, we synthesized distinctive HMG-I and HMG-Y cDNAs by using two synthetic oligonucleotide primers of opposite orientation flanking this 33-nucleotide region-a 5’ primer corresponding to nucleotides 64-102 of HMG-I/Y cDNA [ll] and a 3’primer that is antisense to nucleotides 241-280. The 3’primer was annealed to the total cellular RNA and extended with reverse transcriptase. The 32P-endlabeled 5’ primer was then annealed to the newly synthesized cDNA strand and extended with Klenow polymerase. The reaction products (second strand cDNAs) were separated on a polyacrylamide gel and visualized by autoradiography. The expected sizes of the cDNAs produced by this procedure are 207 bases when the HMG-I mRNA template was copied and 174 bases when the HMG-Y mRNA was copied. Using this procedure we could easily detect and distinguish between the individual HMG-I and HMG-Y mRNA species present within cells by employing only a single round of cDNA synthesis, thereby avoiding potential artifacts that are inherent in quantitative comparisons of the products of multiple rounds of polymerase chain reaction (PCR) amplification. Our procedure in detail was as follows. Twenty micrograms of total RNA from each cell sample was ethanol-precipitated and resuspended with 100 pmol of the 3’ primer in 50 ~1 of hybridization solution A (60% formamide, 0.1% SDS, 20 mMTris-HCl, pH 7.4,400 mMNaC1, 1 mM EDTA) for 15 min at 65°C and then allowed to hybridize for 20 h at 22°C. The RNA and annealed primer were precipitated with ethanol, dried, and suspended in 50 ~1 of an aqueous solution containing 0.5 mM dNTPs, 50 pg/ml of actinomycin D, 1 mM vanadyl ribonucleotide complex, 50 ag/ml of bovine serum albumin, 625 U of Moloney murine leukemia virus reverse transcriptase (Bethesda Research Laboratories, Inc.), and 10 ~1 of 5 X reverse-transcriptase buffer (Bethesda Research Laboratories, Inc.). After 1 h at 37”C, the solution was extracted with phenol-chloroform; nucleic acids in the aqueous phase were then precipitated with ethanol and dried.

HMG-I

100

B g 1 5

0 HR

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HMG-Y

A 20 HR

I

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14 HR

24 HR

F $ a

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kL O

l/!!l

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RELATIVE DNA CONTENT FIG. 1. Cell cycle progression of NIH/3T3 cells. Murine NIH/ 3T3 cells initiated by seeding from a confluent population were analyzed by flow microfluorometry. Time (h) after seeding is given in the upper right-hand portion of each panel. The peak channel numbers for cells with Gi and G2 + M DNA content were 50 and 100, respectively, and are represented on the f: axis as 1 and 2. The peak channels at each time point were: 0 h, 50; 14 h, 65; 20 h, 50 and 90; and 24 h, 55 and 100. An average of 2840 cells from each culture (time point) was analyzed. At time 0, approximately 77% of cells were in G,JG,; at 14 h, 60% of cells were in S and 24% in Gz + M; at 20 h, 33% of cells were in S and 48% in G2 + M. A new cell cycle had begun by 24 h. The RNA-cDNA pellet was suspended in 12 ~1 of water and denatured at 95°C for 5 min. About 1 X lOa cpm of the 5’primer end-labeled with [32P]ATP was added and the solution was adjusted to the concentrations of hybridization solution A. The solution was incubated for 15 min at 65°C and then for 20 h at 22°C. The first strand cDNA and annealed 5’ primer were then ethanol-precipitated, dried, and resuspended in 20 ~1 of 50 mikf Hepes, pH 6.9, 10 mh4 MgClz, 6 m&f /3mercaptoethanol, 1 m&f dNTPs, and 4 U of Klenow enzyme (Boehringer-Mannheim). This reaction mix was incubated for 1 h at 37”C, ethanol-precipitated, and dried. The pellet was suspended in 10 ~1 of loading buffer, denatured, and run on a 6% sequencing gel. The same 5’ primer with cloned HMG-I or HMG-Y cDNAs as templates was used to generate sequencing ladders with Sequenase (U.S. Biochemical Corp.) and 35S-ATP.

RESULTS

The cell cycle progression of attached murine NIH/ 3T3 cells following seeding from confluent cultures, as revealed by flow microfluorometric analysis of cellular DNA content, is shown in Fig. 1. From this data it is evident that following subculturing the NIH/3T3 cells undergo a single, partially synchronous cycle of DNA replication that is comparable in nature to that observed

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in these cells following serum deprivation [15] or other artificial means of synchrony induction. However, this seeding procedure has the significant advantage that it avoids the use of potentially deleterious inhibitors or metabolic starvation methods to induce partial synchronization of the cells. As shown in Fig. 1, immediately following seeding (0 h) approximately 77% of the cells were in the G,/G, phase of the cell cycle whereas by 14 h postseeding approximately 60% of the cells had progressed to the S phase and about 24% to the G2 f M phase of the cell cycle. At 20 h postseeding about 48% of the cells were in the G, + M phase and about 33% in S phase. By 24 h a new cell cycle had begun and the initial partially synchronous wave of DNA synthesis following seeding was beginning to degenerate even further. In partially synchronized NIH/3T3 cells the HMG-I/ Y mRNA levels (relative to total RNA) gradually increased during the first 16 h following seeding (G,/G, to late S phase) but thereafter remained constant (Fig. 2). /3-tubulin and histone H3 mRNA levels were monitored as controls. In contrast to the HMG-I/Y mRNA, levels of the control histone H3 message, known to be correlated with DNA synthesis in mouse 3T6 fibroblasts [ 131, peaked during the S phase of the partially synchronized murine NIH/3T3 cells (12 to 14 h and 20 to 24 h postseeding, Fig. 2). On the other hand, levels of mRNAs encoding @-tubulin, like the HMG-I/Y mRNA, gradually increased until about 20 h postseeding, but then decreased slightly at 24 h (Fig. 2). It is known that variations in tubulin protein levels are determined by the autoregulated instability of their mRNAs [25] and, therefore, should follow a cyclical pattern with peaks occurring during mitosis as has been observed in HeLa cells [26]. The pattern of P-tubulin mRNA expression that we observed in murine NIH/3T3 cells is consistent with this pattern and, along with the observed pattern of histone H3 mRNA, further strengthens our interpretations of the NIH/3T3 cell cycle stages based on flow microfluorometry. We thus conclude that, in NIH/3T3 cells, HMG-I/Y message increases about sixfold during the transition from quiescence to proliferation (Go/G, to late S), but thereafter does not appear to be under cell cycle regulation. To determine whether changes in HMG-I/Y mRNA expression were related to mRNA stability, we measured steady-state mRNA levels in murine NIH/3T3 (Fig. 3) and human K-562 (not shown) cells following treatment with actinomycin D. HMG-I/Y mRNAs appear to be very stable, showing no decrease in expression after prolonged (6 h) actinomycin D treatment. Likewise, HMG17 mRNAs appear quite stable (Fig. 3). Levels of the ptubulin message showed about a 40% decrease at 6 h post actinomycin D treatment (Fig. 3), consistent with the estimated 6-h half-life of tubulin mRNA in mouse 3T6 fibroblasts reported by Caron et al. [27]. The c-myc mRNA was very unstable, disappearing by 1 h after acti-

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ET AL.

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12 14 16 18 20 22 24 36 . d

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Post Serum Stimulation

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FIG. 2. Cell cycle expression of HMG-I/Y mRNAs from partially synchronized murine NIH/3T3 cells. (A) Northern blot autoradiogram of June 8, 1989, experiment. (B) Northern blot autoradiogram of June 19, 1989, experiment. Identical quantities of total RNA (15 .g/lane) from cells at various times (h) after seeding (indicated below each lane) were loaded in triplicate sets of lanes; each set was hybridized with a different probe as indicated. (C) Changes in HMG-I/Y, P-tubulin, and histone H3 mRNA levels. Densitometry scans of autoradiograms shown in A and B and expressed relative to the maximal absorbance values obtained are plotted against time (h) after seeding from confluent cultures.

nomycin D treatment (Fig. 3). Such instability is consistent with previous c-myc mRNA half-life estimates of 10 to 30 min [ 141. HMG-I/Y mRNA levels are about 6-fold higher in proliferating than in quiescent murine NIH/3T3 cells (Fig. 3). Proliferating, log-phase human K-562 cells also exhibit higher levels (2-fold) of HMG-I/Y mRNA than

do quiescent, stationary phase cells (Fig. 4). This 2- to 6fold difference in HMG-I/Y mRNA expression between quiescent and proliferating cells was less than the difference in expression observed between log-phase K562 cells and log-phase HUT-78 cells (X-fold) and much less than that between K-562 cells and placenta tissue (50-fold). These large differences in HMG-I/Y mRNA

HMG-I

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P-tubulin HMG-I 1.2 aI

2 1.0 x ‘0 co0.8 2 a .> Z

I

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-Y aI 0.4 a

P-tubulin \.-.

1

ov21

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Actinomycin

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FIG. 3. Changes in steady-state mRNA levels in NIH/3T3 cells following treatment with 10 pg/ml actinomycin D. (A) Northern blot autoradiograms illustrating the relative amounts of c-myc, @-tubulin, HMG-17, and HMG-I/Y mRNAs at various times (h) following actinomytin D treatment. (B) Graph illustrating the changes in mRNA levels. Densitometry scans of the autoradiograms shown in A and expressed relative to the absorbance value obtained at time point 0 are plotted against time (h) following actinomycin D treatment.

1.0 s 2

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FIG. 4. Changes in HMG-I/Y mRNA levels associated with cellular proliferation (lanes 1 and 2) and with different cell types (lanes 3-8). Twenty micrograms of total RNA from stationary (lane 1) and log-phase (lane 2) human K-562 erythroleukemic cells were hybridized with HMG-I/Y cDNA (lanes 1 and 2). Five micrograms of poly(A)+ mRNA from log-phase human K-562 cells (lane 3), from log-phase human HuT78 T cells (lane 4), and from human placenta tissue (lane 5) were hybridized with HMG-I/Y cDNA (lanes 3-5) and with HMG-17 cDNA (lanes 6-8). (A) Autoradiograms of Northern blots. (B) Bar graph illustrating levels of HMG-I/Y and HMG-17 mRNAs. Densitometry scans of the three autoradiograms shown in A are expressed relative to the maximal absorbance value obtained in each autoradiogram.

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HMG-1 cDNA GATC

a

b

c

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ET AL.

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FIG. 5. Relative proportions of HMG-I and HMG-Y mRNAs in various cell types and in cells dividing at different rates. Double primer cDNA synthesis was used to distinguish HMG-I and HMG-Y mRNA phenotypes. The expected gel migration position of cDNAs produced when HMG-I mRNA templates were copied is indicated at the left of the autoradiogram by an arrow pointing to the corresponding position in the control cDNA sequencing lanes. The expected gel migration position of cDNAs produced when HMG-Y mRNA templates were copied is indicated to the right of the figure by an arrow pointing to the corresponding position in the control HMG-Y cDNA sequencing lanes. The autoradiogram shows the reaction products from 20 pg of total RNA extracted from proliferating murine ascites cells (lane a), proliferating murine Friend cells (lane b-this RNA was partially degraded), quiescent murine NIH/3T3 cells (lane c), murine NIH/3T3 cells 12 h postseeding from confluent cultures (lane d), murine NIH/3T3 cells 24 h postseeding (lane e), proliferating human HUT-78 cells (lane f), quiescent human K-562 cells (lane g), and log-phase human K-562 cells (lane h). The HMG-I:HMG-Y mRNA ratios as measured by densitometry scans of the autoradiogram are given below each lane.

levels cannot be due solely to differences in cellular proliferation. In contrast, the difference in expression of HMG-17 mRNA between log-phase K-562 and HUT-78 cells is less than 2-fold, and the difference between K562 cells and placenta is only about 4-fold (Fig. 4). These rather small differences in HMG-17 mRNA levels probably correspond to different rates of cellular proliferation; a correlation between the rate of cell division and HMG-17 mRNA levels has previously been demonstrated [28]. We were able to detect and quantify differences in HMG-I and HMG-Y mRNA levels by comparing lengths of second strand cDNAs synthesized between primers flanking the 33 nucleotides known to be deleted in HMG-Y [ 111. Thus, cDNAs synthesized from HMGY mRNA templates are 174 nucleotides, whereas cDNAs synthesized from HMG-I mRNA templates are 207 nucleotides (Fig. 5). The HMG-I:HMG-Y mRNA ratio for each of the cellular RNA extracts examined is given below each lane in Fig. 5. An earlier, independent analysis gave comparable ratios (1.17 instead of 1.04 for HUT-78 cells, 1.04 instead of 1.44 for quiescent K-562 cells, and the identical 1.63 ratio for log-phase K-562 cells), demonstrating the reproducibility of this procedure. Overall, RNAs extracted from human cells (HUT-78 or K-562) had greater HMG-I:HMG-Y mRNA ratios (1.04 to 1.63) than did RNAs from murine cells (0.59 to 0.96; ascites, Friend, and NIH/3T3 cells). Log-phase K-562 cells had a higher HMG-I:HMG-Y mRNA ratio (1.63) than did stationary-phase K-562 cells (1.04 and 1.44). Likewise, proliferating NIH/3T3 cells (lanes d and e, Fig. 5) had higher HMG-I:HMG-Y mRNA ratios (0.72 to 0.75) than did quiescent NIH/3T3 cells (0.59; lane c, Fig. 5). The

HMG-I:HMG-Y protein ratios in murine ascites and murine Friend cells as estimated from HPLC elution peak sizes are greater than 1.5 ([6] and unpublished observations), whereas the estimated HMG-I:HMG-Y mRNA ratios in these same cells are only 0.72 to 0.96. Together, these observations suggest two important conclusions: (1) that the ratio of HMG-I:HMG-Y mRNAs is markedly higher in actively proliferating than in nondividing quiescent cells; and, (2) that the cellular concentrations of these two mRNA species may not always directly correlate with the relative amounts of their respective cognate proteins within cells. DISCUSSION

Steady-state levels of mRNA are determined by the equilibrium between rates of transcription and degradation. We have shown that HMG-I/Y mRNAs are very stable; therefore, changes in their steady-state levels are probably due mostly to changes in transcription rates. The unusually long 3’-untranslated region and, in particular, the lack of any (UAUU), mRNA-destabilizing sequences within this region of the HMG-I/Y mRNAs [lo, 111 may contribute to their observed stability as has been shown for other mRNAs [ 29,351. HMG-I/Y mRNA levels in partially synchronized NIH/3T3 cells increase during the transition from quiescence to proliferation, but thereafter remain constant throughout the cell cycle. Cell replication thus influences HMG-I/Y expression but, once activated, this expression does not seem to be under direct cell cycle regulation. It is possible that cellular factors expressed during the transition from quiescence to proliferation (such

HMG-I

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as the c-myc or c-fos proto-oncogene products) may activate or enhance HMG-I/Y transcription. As a result, HMG-I/Y mRNA levels would then increase until they reached an equilibrium level determined by their new rate of transcription and their constant rate of degradation; this steady-state level would remain constant until the transcription rate again changed. Our results are consistent with such a simple equilibrium model and experiments are currently in progress to test these ideas. The differences we observed in HMG-I/Y mRNA levels among different cell types can be only partially explained by differences in rates of cell division. We observed much higher levels of HMG-I/Y mRNAs in NIH/ 3T3 cells and K-562 cells than in HUT-78 cells, even though all of these cells were rapidly dividing with approximately equal generation times. NIH/3T3 cells are undifferentiated mouse embryo fibroblastic cells and K562 cells are undifferentiated, human multipotential hematopoietic cells; in contrast, human HUT-78 cells are a lymphoid cell line exhibiting many phenotypic properties characteristic of overtly differentiated mature T lymphocytes [30]. These results strongly suggest that the level of HMG-I/Y mRNAs may be correlated with the state of differentiation exhibited by cells. In this regard, other workers have also suggested that increased levels of HMG-I and HMG-Y proteins may be associated with undifferentiated or neoplastically transformed cellular phenotypes [ 7,8]. We observed different ratios of HMG-I:HMG-Y mRNAs among different cell types and between quiescent and proliferating cells. HMG-I and HMG-Y mRNAs are the products of alternative splicing [ll]. Differential expression of alternatively spliced mRNAs in different cell types has been documented for other genes [31,32] and has been suggested to play an important gene regulatory role in uiuo. The increased HMG-I: HMG-Y mRNA ratio that we observed in proliferating cells compared to quiescent cells may reflect a preferential HMG-I pattern of RNA splicing associated with an increased rate of transcription. The HMG-I:HMG-Y protein ratios in murine ascites and murine Friend cells (about 1.5; Ref. [6] and unpublished observations) are greater than the HMG-I:HMG-Y mRNA ratios that we have observed (0.72 to 0.96; Fig. 5). This discrepancy may be due to differential rates of protein degradation; the HMG-I isoform may be more stable than the HMGY isoform. HMG-I has an internal hydrophobic region absent in HMG-Y; interior apolar regions have been shown to increase protein stability [33]. In conclusion, we have shown that HMG-I/Y mRNA levels increase during the transition from cellular quiescence to proliferation, but that this increase in expression associated with cell division is not as great as the difference in expression that we observed between differentiated and undifferentiated cells. Although the in vivo functions of the HMG-I and HMG-Y proteins

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are unknown, from the available evidence it seems quite likely that these proteins are both intimately involved with chromosome structure during the division cycle and associated with an undifferentiated cell phenotype. Since these HMG proteins are known to specifically bind to stretches of A/T-rich sequence of DNA both in vitro [34-361 and in viuo [9], it is reasonable to suspect that they can directly affect chromatin conformation and, consequently, may indirectly affect the differential activation/inactivation of genes associated with cell division and/or cellular differentiation. We thank Drs. Michael Bustin and David Landsman (of the U.S. National Institutes of Health) for generously supplying us with the HMG-17 cDNA probe [21]. Thanks are also owed to Drs. Don Lehn and Nancy Magnuson and to Mark Nissen (all of Washington State University) for helpful discussions during the course of this work. This work was supported in part by U.S. National Science Foundation Grant DCB-8602622 (to R.R.) and U.S. National Institutes of Health Training Grant AI-07025 (to K.R.J.).

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