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• W1974590591 abstract "Covalent histone modifications and the incorporation of histone variants bring about changes in chromatin structure that in turn alter gene expression. Interest in non-allelic histone variants has been renewed, in part because of recent work on H3 (and other) histone variants. However, only in mammals do three non-centromeric H3 variants (H3.1, H3.2, and H3.3) exist. Here, we show that mammalian cell lines can be separated into two different groups based on their expression of H3.1, H3.2, and H3.3 at both mRNA and protein levels. Additionally, the ratio of these variants changes slightly during neuronal differentiation of murine ES cells. This difference in H3 variant expression between cell lines could not be explained by changes in growth rate, cell cycle stages, or chromosomal ploidy, but rather suggests other possibilities, such as changes in H3 variant incorporation during differentiation and tissue- or species-specific H3 variant expression. Moreover, quantitative mass spectrometry analysis of human H3.1, H3.2, and H3.3 showed modification differences between these three H3 variants, suggesting that they may have different biological functions. Specifically, H3.3 contains marks associated with transcriptionally active chromatin, whereas H3.2, in contrast, contains mostly silencing modifications that have been associated with facultative heterochromatin. Interestingly, H3.1 is enriched in both active and repressive marks, although the latter marks are different from those observed in H3.2. Although the biological significance as to why mammalian cells differentially employ three highly similar H3 variants remains unclear, our results underscore potential functional differences between them and reinforce the general view that H3.1 and H3.2 in mammalian cells should not be treated as equivalent proteins. Covalent histone modifications and the incorporation of histone variants bring about changes in chromatin structure that in turn alter gene expression. Interest in non-allelic histone variants has been renewed, in part because of recent work on H3 (and other) histone variants. However, only in mammals do three non-centromeric H3 variants (H3.1, H3.2, and H3.3) exist. Here, we show that mammalian cell lines can be separated into two different groups based on their expression of H3.1, H3.2, and H3.3 at both mRNA and protein levels. Additionally, the ratio of these variants changes slightly during neuronal differentiation of murine ES cells. This difference in H3 variant expression between cell lines could not be explained by changes in growth rate, cell cycle stages, or chromosomal ploidy, but rather suggests other possibilities, such as changes in H3 variant incorporation during differentiation and tissue- or species-specific H3 variant expression. Moreover, quantitative mass spectrometry analysis of human H3.1, H3.2, and H3.3 showed modification differences between these three H3 variants, suggesting that they may have different biological functions. Specifically, H3.3 contains marks associated with transcriptionally active chromatin, whereas H3.2, in contrast, contains mostly silencing modifications that have been associated with facultative heterochromatin. Interestingly, H3.1 is enriched in both active and repressive marks, although the latter marks are different from those observed in H3.2. Although the biological significance as to why mammalian cells differentially employ three highly similar H3 variants remains unclear, our results underscore potential functional differences between them and reinforce the general view that H3.1 and H3.2 in mammalian cells should not be treated as equivalent proteins. Eukaryotic organisms depend on complex and highly regulated mechanisms to activate or silence genes in response to a variety of stimuli, including environmental changes, cell cycle regulators, and developmental cues. An increasing body of evidence suggests that epigenetic mechanisms involving chromatin remodeling alter the accessibility of proteins, such as transcription factors, to the DNA template. The fundamental repeating unit of chromatin is the nucleosome core particle, which consists of DNA in close association with an octameric unit of core histones (H2A, H2B, H3, and H4). However, in some instances, specialized histone variants are found in place of the canonical histones, enabling the encoding of epigenetic information through defined or “specialized” nucleosome arrays (reviewed in Ref. 1Hake S.B. Xiao A. Allis C.D. Br. J. Cancer. 2004; 90: 761-769Crossref PubMed Scopus (305) Google Scholar). Histones are subject to a diverse array of covalent modifications that occur mostly at the N- and C-terminal tail domains. The histone “code” hypothesis (2Strahl B.D. Allis C.D. Nature. 2000; 403: 41-45Crossref PubMed Scopus (6533) Google Scholar, 3Fischle W. Wang Y. Allis C.D. Curr. Opin. Cell. Biol. 2003; 15: 172-183Crossref PubMed Scopus (979) Google Scholar) has been put forward to explain the seemingly complex nature of the reported patterns of histone modifications. Formally, this hypothesis states that a specific histone modification, or combination of modifications, can affect distinct downstream cellular events by altering the structure of chromatin and/or generating a binding platform for effector proteins, which specifically recognize the modification(s) and initiate events that lead to gene transcription or silencing. Expanding the scope of this code, a large number of variant histones has been identified, including some that are unique to vertebrates and some that are highly conserved among all eukaryotes (reviewed in Ref. 4Malik H.S. Henikoff S. Nat. Struct. Biol. 2003; 10: 882-891Crossref PubMed Scopus (412) Google Scholar). It has been shown that replacement of the replication-dependent (RD) 4The abbreviations used are: RDreplication-dependentRP-HPLCreverse phase-HPLCMSmass spectrometryRIreplication-independentHEK293human embryonic kidney 293FACSfluorescence-activated cell sorterPBSphosphate-buffered salineTAUTriton-acid-ureaRAretinoic acid.4The abbreviations used are: RDreplication-dependentRP-HPLCreverse phase-HPLCMSmass spectrometryRIreplication-independentHEK293human embryonic kidney 293FACSfluorescence-activated cell sorterPBSphosphate-buffered salineTAUTriton-acid-ureaRAretinoic acid. histone H3 (formally H3.2, see supplemental Fig. 1A, top) with its replication-independent (RI) variant H3.3 in Drosophila cells occurs at transcriptionally active loci (5Ahmad K. Henikoff S. Mol. Cell. 2002; 9: 1191-1200Abstract Full Text Full Text PDF PubMed Scopus (869) Google Scholar, 6Smith M.M. Mol. Cell. 2002; 9: 1158-1160Abstract Full Text Full Text PDF PubMed Scopus (13) Google Scholar). Furthermore, characterization of Drosophila and Arabidopsis histones by mass spectrometry (MS) revealed enrichment of modifications associated with transcriptional activity, such as methylation of lysine 4 (Lys4) and Lys79 and acetylation of Lys14, Lys18, and Lys23, in H3.3 compared with H3.2 (7McKittrick E. Gafken P.R. Ahmad K. Henikoff S. Proc. Natl. Acad. Sci. U. S. A. 2004; 101: 1525-1530Crossref PubMed Scopus (406) Google Scholar, 8Johnson L. Mollah S. Garcia B.A. Muratore T.L. Shabanowitz J. Hunt D.F. Jacobsen S.E. Nucleic Acids Res. 2004; 32: 6511-6518Crossref PubMed Scopus (178) Google Scholar). These results suggest that, at least in plant and Drosophila cells, H3.2 and its variant H3.3 play different roles in remodeling chromatin, in part by altering covalent histone modification patterns associated with transcriptional silencing and activation. replication-dependent reverse phase-HPLC mass spectrometry replication-independent human embryonic kidney 293 fluorescence-activated cell sorter phosphate-buffered saline Triton-acid-urea retinoic acid. replication-dependent reverse phase-HPLC mass spectrometry replication-independent human embryonic kidney 293 fluorescence-activated cell sorter phosphate-buffered saline Triton-acid-urea retinoic acid. Unlike Drosophila, which contains only two different histone H3 molecules, mammalian cells contain three non-centromeric H3 variants: H3.1, H3.2, and H3.3, which differ only in a few amino acids (see supplemental Fig. 1, A). The function of these three mammalian H3s, especially H3.1 and H3.2, is poorly understood. In this report, we investigate the expression patterns and post-translational modifications associated with these three mammalian H3 variants. Analyses of multiple mammalian cell lines revealed that they can be divided into two groups based upon the relative amounts of the individual H3 variants in chromatin. Although the functional significance of this grouping remains unclear, cellular differentiation appears to alter these ratios in at least one ES cell line in a modest, but reproducible, fashion. We also show that these cell line-specific differences in H3 variant expression do not originate from changes in growth rate, cell cycle stages, or chromosomal ploidy. Possible mechanisms are discussed. Additionally, the different human H3 variants were subjected to quantitative tandem MS analyses. As expected from studies in Drosophila (7McKittrick E. Gafken P.R. Ahmad K. Henikoff S. Proc. Natl. Acad. Sci. U. S. A. 2004; 101: 1525-1530Crossref PubMed Scopus (406) Google Scholar), transcriptionally active marks are associated with H3.3; those often associated with gene silencing, e.g. Lys27 di- and trimethylation, are found on H3.2. Surprisingly, H3.1 is enriched both with modifications that are largely associated with gene silencing, e.g. Lys9 dimethylation, as well as those linked to gene activation, e.g. Lys14 acetylation. These data reinforce the general view that alterations in the covalent modification patterns associated with histone variants provide additional regulatory options for epigenomic “indexing” of biological processes, many of which remain unclear. As well, our data lend support to a poorly appreciated notion that H3.1 and H3.2 variants, while highly similar at the level of the primary sequence, differing in only one amino acid position (Cys96 in H3.1 versus Ser96 in H3.2; see supplemental Fig. 1, A), are not equivalent at the level of post-translational modifications. Thus, our studies underscore the potential need for caution when interpreting H3-related studies in mammalian models. Cell Lines and Culture—All mammalian cell lines, with the exception of mouse LF2 cells were grown in Iscove's Dulbecco's modified Eagle's medium supplemented with 10% fetal calf serum and penicillin/streptomycin at 37 °C and 5% CO2. Cell lines used in this study are described under supplemental material and methods. Cell Synchronization—HeLa cells were grown to 70% confluency and treated with 3 mm thymidine for 15 h. Thymidine containing medium was removed, cells washed once with fresh thymidine-free medium and grown for 9 h in regular medium. This process was repeated and then, after release from the double thymidine block, cells were harvested from individual plates every 2 h by trypsinization, washed with PBS, and split into three samples to prepare for cell cycle analysis by FACS and isolation of RNA and histones (see below). Preparation of Histones—Nuclei and histones were isolated as described earlier (9Hake S.B. Garcia B.A. Kauer M. Baker S.P. Shabanowitz J. Hunt D.F. Allis C.D. Proc. Natl. Acad. Sci. U. S. A. 2005; 102: 6344-6349Crossref PubMed Scopus (149) Google Scholar). Cell nuclei were isolated by hypotonic lysis in buffer containing 10 mm Tris-HCl, pH 8.0, 1 mm KCl, 1.5 mm MgCl2, 1 mm dithiothreitol, 0.4 mm phenylmethylsulfonyl fluoride, protease and phosphatase inhibitors. Pelleted nuclei were acid-extracted using 0.4 n sulfuric acid, precipitated with trichloroacetic acid and resuspended in water. Reverse Phase HPLC (RP-HPLC)—Separation of mammalian core histones by RP-HPLC was done as described (9Hake S.B. Garcia B.A. Kauer M. Baker S.P. Shabanowitz J. Hunt D.F. Allis C.D. Proc. Natl. Acad. Sci. U. S. A. 2005; 102: 6344-6349Crossref PubMed Scopus (149) Google Scholar). Briefly, acid-extracted histones were separated by RP-HPLC on a C8 column (220 by 4.6 mm Aquapore RP-300, PerkinElmer Life Sciences) using a linear ascending gradient of 35–60% solvent B (solvent A: 5% acetonitrile, 0.1% trifluoroacetic acid, solvent B: 90% acetonitrile) over 75 min at 1.0 ml/min on a Beckman Coulter System Gold 126 Pump Module and 166/168 Detector. Under these conditions histones H3 split into two peaks. The H3-containing fractions were dried under vacuum and stored at -80 °C. RP-HPLC fractions were resuspended in water, analyzed by SDS-PAGE and control-stained with Coomassie Brilliant Blue. The identified fractions were then subjected to MS analysis. Two-dimensional Triton-Acid-Urea (TAU) Gels—Total histones were dried under vacuum and resuspended in loading buffer (6 m urea, 0.02% (w/v) pyronin Y, 5% (v/v) acetic acid, 12.5 mg/ml protamin sulfate). Samples were separated on TAU mini-gels (15% PAGE, 6 m urea, 5% acetic acid, 0.37% Triton X-100; 300 V in 5% acetic acid for 1.5 h). Lanes containing the samples were cut out, adjusted in 0.125 m Tris, pH 6.6, and the TAU gel slice was assembled on top of a 15% SDS-PAGE mini-gel. After the run, the gel was stained with Coomassie Blue and destained with 5% methanol, 7.5% acetic acid overnight. The gels were scanned and quantified using Image Gauge software (Science Lab), with the subtraction of background staining. Growth Rate Analysis—1 × 105 cells (HeLa and HEK293) were grown in 6-well plates, and every 24 h samples were collected and counted, with exclusion of dead cells (staining with Trypan Blue). Half of the cells were discarded to avoid contact inhibition of the cells as they become too confluent, fresh medium was added, and the cells were allowed to grow for another 24 h before the next sample was collected. With this method, we could measure the doubling time of HeLa and HEK293 cells while still maintaining them at normal confluency (i.e. the same confluency at which we grow them for all other experiments). Cell numbers were calculated to present the cell growth over days of both cell lines. HeLa and HEK293 cells were seeded as duplicates, and this experiment was performed twice. Cell Cycle Analysis by FACS—1 × 106 cells were collected, washed with PBS, and fixed overnight at -20 °C in 70% ethanol, diluted in PBS. The next day, cells were washed with PBS and incubated for 30 min at 37 °C in PBS containing RNase A (10 μg/ml), followed by the addition of propidium iodide (10 μg/ml) and another incubation for 30 min at 37 °C. Stained cells were analyzed on a FACSort instrument (BD Immunocytometry Systems) with the exclusion of doublets. Analysis of the results was performed with CellQuest software (BD Bioscience). Immunoblots—Acid-extracted histones were separated on 15% SDS-PAGE mini-gels and either stained with Coomassie Brilliant Blue or transferred onto poly(vinylidene difluoride) membranes (Millipore), and stained with Ponceau S (Sigma) to ensure proper protein transfer. After incubation with primary antibody (anti-H3 S28P: 1:1000 (Upstate Biotechnology) or anti-H3: 1:10000; Abcam)) and addition of a horseradish peroxidase-conjugated secondary antibody (Amersham Biosciences), membranes were incubated with ECL-Plus substrate (Amersham Biosciences), and proteins were detected by exposure to x-ray film (Amersham Biosciences). Sample Preparation of Histone H3 Variants for MS—Purified histone H3 protein from pooled RP-HPLC fractions were derivatized by treatment with propionyl anhydride reagent (8Johnson L. Mollah S. Garcia B.A. Muratore T.L. Shabanowitz J. Hunt D.F. Jacobsen S.E. Nucleic Acids Res. 2004; 32: 6511-6518Crossref PubMed Scopus (178) Google Scholar). The reagent was created using 75 μl of MeOH and 25 μl of propionic anhydride (Aldrich, Milwaukee, WI). Equal volumes of reagent and each H3 variant were mixed and allowed to react at 51 °C for 15 min. Propionylated histone H3s were then digested with trypsin (Promega) at a substrate/enzyme ratio of 20:1 for 5 h at 37°C after dilution of the sample with 100 mm ammonium bicarbonate buffer solution (pH 8). The reaction was quenched by the addition of concentrated acetic acid and freezing. A second round of propionylation was then performed to propionylate the newly created peptide N termini. Mass Spectrometry—Propionylated histone digest mixtures were loaded onto capillary precolumns (360 μm outer diameter × 75 μm inner diameter, Polymicro Technologies, Phoenix, AZ) packed with irregular C18 resin (5–20 μm, YMC Inc., Wilmington, NC) and washed with 0.1% acetic acid for 10 min. Precolumns were connected with Teflon tubing to analytical columns (360 μm outer diameter × 50 μm inner diameter, Polymicro Technologies) packed with regular C18 resin (5 μm, YMC Inc.) structured with an integrated electrospray tip as previously described (10Martin S.E. Shabanowitz J. Hunt D.F. Marto J.A. Anal Chem. 2000; 72: 4266-4274Crossref PubMed Scopus (302) Google Scholar). Samples were analyzed by nanoflow HPLC-μ-electrospray ionization on a linear quadrupole ion trap-Fourier Transform Ion Cyclotron Resonance (LTQ-FT-ICR) mass spectrometer (Thermo Electron, San Jose, CA). The gradient used on an Agilent 1100 series HPLC solvent delivery system (Palo Alto, CA) consisted of 0–40% B in 60 min, 45–100% B in 15 min (A, 0.1% acetic acid, B, 70% acetonitrile in 0.1% acetic acid) or other similar gradients. The LTQ-FT mass spectrometer was operated in the data-dependent mode with the 10 most abundant ions being isolated and fragmented in the linear ion trap. All MS/MS spectra were manually interpreted. Stable Isotope Labeling for Relative Quantitative Analysis—For a differential expression comparison of histone post-translational modifications from the three H3 variants, stable isotope labeling based on conversion of peptide carboxylic groups to their corresponding methyl esters was used (11Syka J.E. Marto J.A. Bai D.L. Horning S. Senko M.W. Schwartz J.C. Ueberheide B. Garcia B. Busby S. Muratore T. Shabanowitz J. Hunt D.F. J. Proteome Res. 2004; 3: 621-626Crossref PubMed Scopus (333) Google Scholar). First, all samples were dried to dryness by lyophilization. Aliquots of solutions from propionylated histone peptides from H3.1, H3.2, or H3.3 were converted to d0-methyl esters by reconstituting the lyophilized sample in 100 μl of 2 m d0-methanol/HCl, or converted to d3-ethyl esters by reconstituting the lyophilized sample in 100 μlof2 m d4-methanol/HCl. Reaction mixtures were allowed to stand for 1 h at room temperature. Methyl ester solvent was removed from each sample by lyophilization, and the procedures were repeated using a second 100-μl aliquot of methyl ester reagents. Solvent was then removed again by lyophilization, and samples were dissolved in 20 μl of 0.1% acetic acid. Aliquots of each solution were then equally mixed for comparative analysis by MS. Quantitative PCR—Total RNA isolation was performed using TRIzol Reagent (Invitrogen). Single-stranded cDNA was generated with the Superscript First-Strand Synthesis kit (Invitrogen). Quantitative PCR was performed with SYBR green dye according to the manufacturer's instructions (Stratagene). HeLa cDNA was used to generate a standard curve from which the amount of cDNA amplified in each sample was determined as indicated. mRNA levels were normalized to H3.2 mRNA expression. All oligos were synthesized by Sigma, and the sequences of the primer pairs for quantitative PCR used in this study are listed under supplemental material and methods. Only mammalian cells contain three different non-centromeric H3 variants: RD H3.1 and H3.2 variants versus a single RI H3.3. Other organisms contain only one type of H3, H3.3 (Saccharomyces cerevisiae), or two, H3.2 and H3.3 (e.g. Arabidopsis, Xenopus, and Drosophila) (supplemental Fig. 1A, top). The three mammalian H3 variants are almost identical in their amino acids sequence (see supplemental Fig. 1, A, bottom). H3.1 and H3.2 have only a single amino acid difference (amino acid 96, cysteine/serine, respectively; gray box), whereas H3.1 and H3.3 differ in five amino acids (amino acid 31, alanine/serine; amino acid 87, serine/alanine; amino acid 89, valine/isoleucine; amino acid 90, methionine/glycine; amino acid 96, cysteine/serine, respectively). Because there are no available antibodies that distinguish the three mammalian H3 variants, the investigation of endogenous expression of these variants is restricted largely to chromatographic and electrophoretic separations (7McKittrick E. Gafken P.R. Ahmad K. Henikoff S. Proc. Natl. Acad. Sci. U. S. A. 2004; 101: 1525-1530Crossref PubMed Scopus (406) Google Scholar, 9Hake S.B. Garcia B.A. Kauer M. Baker S.P. Shabanowitz J. Hunt D.F. Allis C.D. Proc. Natl. Acad. Sci. U. S. A. 2005; 102: 6344-6349Crossref PubMed Scopus (149) Google Scholar, 12Waterborg J.H. J. Biol. Chem. 1990; 265: 17157-17161Abstract Full Text PDF PubMed Google Scholar). Thus, to test whether the different mammalian H3 variants are present in similar abundance in different cell types, we first turned to a chromatographic method. Acid-extracted total histones from different mammalian cell lines were resolved by RP-HPLC; two distinctive H3 peaks were typically observed (supplemental Fig. 1, A). Based on the H3 RP-HPLC profiles, we were able to separate mammalian cell lines into two groups (A and B) based on their peak height (absorbance) and peak area differences. Interestingly, members of group A show an absorbance that is higher for peak 2 than peak 1; a reverse relationship is evident for group B (i.e. an absorbance that is higher for peak 1 than peak 2 (supplemental Fig. 1, C). To gain further insight into the protein compositions of the two RP-HPLC peaks, MS was employed (supplemental Fig. 2). Our MS analyses demonstrate that the first fractions of peak 1 contain H3.2, and the later “shoulder” fractions contain H3.3; peak 2, in contrast, contains almost entirely H3.1 variant (supplemental Fig. 2 and supplemental Tables 1–3). To confirm the observed cell type-specific H3 differences and to additionally identify the expression levels of each H3 variant in the different cell lines examined, we separated acid-extracted histones on two-dimensional TAU gels (Fig. 1A) and visualized the histones by staining with Coomassie Blue. Three of the histone spots also stained with H3 specific antibodies (data not shown). To determine the identity of the variant in each of the three H3 spots, each protein sample was digested in gel and the resulting peptides were then characterized by tandem MS as described above (data not shown and Fig. 1A). Two-dimensional TAU gels were then employed to examine the distribution of H3.1, H3.2, and H3.3 in six different mammalian cell lines, three from each of the group A and B categories (Fig. 1B). We find that cell lines from group A are enriched in H3.1 and those in group B are enriched in H3.3. Next, we quantified the proportion of protein in each of the different H3 spots using Image Gauge software (Fig. 1C). Interestingly, while the proportion of H3.2 (dark gray bars) did not change dramatically between the two different groups, cell lines from group A were enriched in H3.1 (light gray bars). In contrast, cell lines from group B contained less or equal proportions of H3.1 compared with H3.3 protein (black bars). Furthermore, as seen before by RP-HPLC analysis (supplemental Fig. 1), several modest changes in H3 variant composition were observed in murine ES cells (LF2) that were treated with retinoic acid (RA) to induce neuronal differentiation (Fig. 1C, see LF2 columns). First, an immediate increase in H3.3 and a corresponding drop in H3.1 occurred during the first 6 days of treatment, whereas H3.2 levels remained largely the same. However, by day 10 post-RA treatment, when the majority of the ES cells have taken on a neuronal phenotype, the levels of H3.2 increased marginally and the levels of H3.3 dropped slightly, whereas the levels of H3.1 remained about the same. These results confirm and extend the observations made by RP-HPLC (supplemental Fig. 1, C). Because all cell lines from group A are derived from cancer cells, we wondered whether the high level of H3.1 expression arises from differences in chromosomal ploidy. Therefore, we used RP-HPLC to separate histones from mouse embryonic fibroblast cells where the chromosomal status was either diploid (P-CUT MEF) or tetraploid (10T1/2). Results of this experiment are shown in Fig. 1D. Both of these cell lines had equal peak area ratios in RP-HPLC analysis and were assigned to group B because the observed ratio of peak areas (peak 1/peak 2) was 3. We then separated the H3 variants by two-dimensional TAU gels and found that the levels of H3.1, H3.2, and H3.3 were very similar between diploid (P-CUT MEF) and tetraploid (10T1/2) cell lines (Fig. 1E). H3.3 was the most highly expressed variant in both cell lines, followed by H3.1 and then H3.2. These results suggest that ploidy is not responsible for the different expression levels observed for H3 variant proteins. However, because differences in the peak 1/peak 2 ratio were observed in human (group A) versus mouse (group B) species (Fig. 1 and supplemental Fig. 1), we cannot rule out the formal possibility that variable copy numbers of the H3.1 and H3.2 genes might contribute, at least in part, to some of the differences in H3 variant expression profiles. Another explanation for the above observations is that cells from embryonic origin contain high levels of H3.3, and cells derived from adult tissue have more H3.1. Because H3.1 and H3.2 are expressed only in S-phase whereas H3.3 is expressed and incorporated into chromatin independent of the cell cycle (5Ahmad K. Henikoff S. Mol. Cell. 2002; 9: 1191-1200Abstract Full Text Full Text PDF PubMed Scopus (869) Google Scholar), we next wondered whether the observed differences in H3 variant expression arise from differences in growth rates and/or time spent in S-phase. Therefore, we tested a representative cell line from group A and B (HeLa and HEK293, respectively) in growth assays and cell cycle analyses. While somewhat variable, these cells showed a similar growth curve (Fig. 2A), and similar numbers of cells in S-phase by FACS analysis (Fig. 2B). It is also important to note that the growth rates of cell lines within a single group was very different, e.g. within group A, HT-29 grew extremely slowly, whereas Raji cells grew extremely fast (data not shown). Therefore, we conclude that the observed differences in the proportions of the three H3 variant proteins between groups A and B are not likely explained by the RD expression of H3.1 and H3.2 alone. We wondered whether differences in the proportion of H3.1, H3.2, and H3.3 between group A and B cells originated from differences in steady state levels of mRNA expression. To address this possibility, we performed quantitative analyses of mRNA expression levels of five human cell lines used in this study. We could not include other cell lines from group B in this study, because these are of mouse origin and differ in their nucleotide sequence from human H3 variant genes. Fig. 2, C and D show the mRNA expression levels of one H3.2, nine different H3.1, and both H3.3A and H3.3B genes normalized to 18 S rRNA expression. Because we do not know if the 18 S rRNA expression level is the same in all human cell lines examined, we also normalized our data to H3.2 mRNA expression because the proportion of H3.2 protein did not change as drastically between groups A and B compared with H3.1 and H3.3 protein (supplemental Fig. 3). Because it is still possible that different cell lines express different amounts of 18 S rRNA, these results should be viewed with caution. Nonetheless, we observed a similar pattern in H3 variant gene expression when normalized to 18S rRNA expression (Fig. 2, C and D) as when normalized to H3.2 gene expression (supplemental Fig. 3). Interestingly, H3.1 genes of the five different human cell lines were expressed at relative low level and did not exhibit dramatic differences in their expression, with the exception of CEM cells, which seem to express H3.1C. On the other hand, HEK293 cells, which belong to group B, showed a reproducible increase in the expression level of H3.3A (almost 2-fold compared with other human cell lines from group A). The lack of significant differences in growth rates from HEK293 and HeLa cells together with the mRNA expression data suggest that the differences in H3.1, H3.2, and H3.3 protein expression that we observed by both RP-HPLC and two-dimensional TAU gel analyses might originate at the transcriptional level and are independent of growth rates. Because we observed slight differences in the proportions of cells in G1 or G2/M between HeLa and HEK293 cells, we wondered if cell cycle phases could account for the observed differences in H3.1, H3.2, and H3.3 proportions between HeLa and HEK293 cell lines. We therefore performed a detailed analysis of H3 variant expression on both mRNA and protein levels during G1, S, and G2/M phases in HeLa cells. The results from one of two independently conducted, highly reproducible experiments are shown in Fig. 3. HeLa cells were synchronized in G1 by a double thymidine block and released from this block to continue through different cell cycle phases. Every 2 h, cells were harvested and samples prepared for cell cycle analysis by FACS, mRNA isolation, and cDNA generation or acid-extraction of histones. Fig. 3A shows the cell cycle profile of these cells analyzed by FACS, and quantitative analysis of the amount of cells in each cell phase is depicted in Fig. 3B. The majority of asynchronously growing cells was found to be in G1 (∼75%), but also cells in S and G2/M phase were observed. Treatment of HeLa cells with Nocodazole led to an arrest in G2/M (∼50%). ∼75% of cells were found to be i" @default.
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• W1974590591 date "2006-01-01" @default.
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• W1974590591 title "Expression Patterns and Post-translational Modifications Associated with Mammalian Histone H3 Variants" @default.
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• W1974590591 doi "https://doi.org/10.1074/jbc.m509266200" @default.
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