Regulation of Chromatin Assembly and Cell Transformation by Formaldehyde Exposure in Human Cells

Cell Transfection

Transfections were carried out using Lipofectamine 2000 (Invitrogen) according to the manufacturer’s instructions. Control short interfering RNA (siRNA) or two distinct siRNAs for H3.3 (H3.3 siRNA1 and H3.3 siRNA 2) (Sigma) were transfected into BEAS-2B cells to knock down H3.3 expression. For H3.3 overexpression, the pcDNA3.1-FLAG-H3.3 plasmid was transfected into BEAS-2B cells. pcDNA3.1 empty vector was used as the control.

Antibodies

Anti-histone H3 (ab1791), antiacetyl-histone H4 Lys-12 (ab61238), and anti - β - actin (ab8226) were purchased from Abcam; anti-dinitrophenyl (DNP; D9656), which reacts with DNP–bovine serum albumin (BSA) and DNP–rabbit serum albumin but not with BSA or rabbit serum albumin, was purchased from Sigma-Aldrich; antiacetyl-histone H3 (06-599), anti-trimethyl-histone H3 Lys-4 (07-473), anti-dimethyl-histone H3 Lys-9 (07-441), anti-trimethyl-histone H3 Lys-27 (07-499), antiacetyl-histone H3 Lys-18 (07-354), and anti-histone H4 (07-108) were purchased from Millipore; and anticaspase-3 (CST9662) was purchased from Cell Signaling Technology.

Trypan Blue Staining and Cell Viability Assay

Postexposure, cells were trypsinized, centrifuged, and resuspended in DMEM culture medium. The total number of viable cells was determined by trypan blue staining. Cell viability was assessed by calculating the percentage of live cells within five randomly selected fields. Independent experiments were repeated three times.

Soft-Agar Assays

After transfection and FA treatment, cells were seeded in soft agar growth medium in 6-well plates (5,000 cells/well) and incubated at 37°C for 4–5 wk. Colonies were stained with INT/BCIP (Roche) and photographed. Colonies larger than 50 μm were counted. Independent experiments were performed three times. The results are presented as ±fold change.

Cell Lysates, Acid Extraction, and Western Blot Analysis

After FA exposure, cells were washed with ice-cold phosphate-buffered saline (PBS) and collected by centrifugation. Whole-cell lysates were prepared using Triton X-100 lysis buffer [pH 7.4; 50 mM Tris-HCl, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate (SDS), 500 mM sodium chloride (NaCl), 10 mM magnesium chloride (MgCl₂), 10 mM sodium butyrate, protease inhibitors]. For acid extraction, cells were resuspended in lysis buffer (pH 7.5; 10 mM HEPES, 1.5 mM MgCl₂, 10 mM potassium chloride (KCl), 0.2 M hydrochloric acid (HCl), 0.5 mM dithiothreitol (DTT), 10 mM sodium butyrate, and protease inhibitors], incubated on ice for 60 min, and centrifuged at 11,000×g for 10 min. The supernatant was collected and neutralized using one-fifth volume of 1.5 M Tris-HCl buffer (pH 8.8). Total protein (50 μg) was separated on 14% polyacrylamide gel by electrophoresis and transferred to a polyvinylidene difluoride (PVDF) membrane (Bio-Rad). The membrane was cut according to the predicted size of the target proteins. Nonspecific binding was blocked by incubation in Tris-buffered saline (TBS) containing 0.1% TWEEN®-20 and 5% dry nonfat milk. Immunoblotting was performed with primary antibodies overnight at 4°C, followed by incubation with a peroxidase-conjugated second antibody. The reagent for enhanced chemiluminescence (Bio-Rad) was used for detection and developed on X-ray film. Independent experiments were performed three times.

Cellular Fractionation

Cells (4 × 10⁷) were suspended in 1.5 mL of hypotonic buffer (pH 7.4; 10 mM Tris-HCl, 10 mM KCl, 1.5 mM MgCl₂, 1 mM DTT, and protease inhibitors) for 10 min on ice and centrifuged at 2,500×g for 10 min after disruption through a 25-gauge needle. The supernatant was retained as the cytosolic fraction. The remaining pellet was resuspended in 0.5 pellet volume of low-salt buffer [pH 7.4; 20 mM Tris-HCl, 20 mM KCl, 1.5 mM MgCl₂, 1 mM DTT, 0.2 mM ethylenediaminetetraacetic acid (EDTA), 25% glycerol, and protease inhibitors] and homogenized using a 25-gauge needle. The sample volume was carefully measured, and 0.5 vol of high-salt buffer (pH 7.4; 20 mM Tris-HCl, 1.2 M KCl, 1.5 mM MgCl₂, 1 mM DTT, 0.2 mM EDTA, 25% glycerol, and protease inhibitors) was added to obtain a final KCl concentration of 0.42 M. Samples were incubated on a rotator for 30 min at 4°C; then, the suspension was centrifuged at 14,000×g for 15 min, and the supernatant was retained as nuclear extract. The insoluble pellet was resuspended in Tris buffer [pH 7.4; 10 mM Tris-HCl, 10 mM sodium chloride (NaCl), 3 mM MgCl₂, and protease inhibitors] containing 1.5 mM calcium chloride (CaCl₂). The sample was disrupted by four passes through a 20-gauge needle and another four passes through a 25-gauge needle. The suspension was then adjusted until the amount of nucleic acid in 1 mL of the sample had an optical density of 100 (A₂₆₀ = 100) and digested with 12 × 10⁻² units/μL micrococcal nuclease (MNase) for 12 min at 37°C. The reaction was terminated by the addition of EDTA (10 mM, final concentration). The suspensions were then incubated on ice for 30 min. After incubation, the supernatant was collected as the S1 subfraction. The remaining pellet was resuspended in the Tris buffer plus 0.25 mM EDTA, incubated on ice for 15 min, and passed four times through a 25-gauge needle. After centrifugation at 14,000×g for 10 min, the supernatant was collected as the S2 fraction and combined with S1 as the chromatin fraction.

Histone Carbonyl Assays

Histone carbonyl assays were performed as described by Thompson and Burcham (2008), using 15 μg of acid-extracted total histones as substrates.

Nucleosome Preparation and Chromatin Immunoprecipitation

Mono- and dinucleosomes were isolated by MNase digestion and sucrose gradient purification. Chromatin immunoprecipitation (ChIP) analysis was performed as described previously (Jin and Felsenfeld 2006). See Table 1 for primers.

MNase Digestion Assay

Nuclei were isolated from FA-treated and control BEAS-2B cells as described previously (Jin and Felsenfeld 2006). The A₂₆₀ of the suspension was adjusted to 1.25. For the digestion, MNase was added (3 × 10⁻³, 6 × 10⁻³, 12 × 10⁻³, or 24 × 10⁻³ units/μL, final concentration), and the suspension was incubated for 10 min at 37°C. The reaction was stopped by the addition of EDTA to a final concentration of 10 mM. Proteinase K (1%, v/v) and SDS (1%, w/v) were also added, and the mixture was incubated overnight at 37°C. The DNA was purified by phenol/chloroform extraction and ethanol precipitation. Purified DNA (3 μg) was separated on a 2% agarose gel and visualized by ethidium bromide staining. Independent experiments were performed three times.

MNase–Real-Time Quantitative Polymerase Chain Reaction to Determine Nucleosome Occupancy

Nuclei were prepared from FA-treated and untreated cells and were treated with different concentrations of MNase. Mononucleosomes were prepared and purified on a 5–30% sucrose gradient as described previously (Jin and Felsenfeld 2007). DNA fragments from MNase-digested nucleosomes were purified by phenol/chloroform extraction and ethanol precipitation and were quantified using a PicoGreen fluorescence kit (Molecular Probes). The same amount of DNA in each sample was used in real-time quantitative polymerase chain reaction (qPCR) to determine the differences in the abundance of specific target DNA sequences obtained from untreated cells and from cells treated with FA. If a genomic locus is occupied by nucleosomes in untreated cells, the corresponding DNA fragment will be protected from the MNase treatment and can easily be amplified by qPCR. However, if nucleosome occupancy of the region is reduced following FA exposure, the DNA fragment is largely removed by MNase digestion and cannot be properly amplified. Accordingly, the ratio of the concentration of a target sequence protected from MNase digestion in FA-treated cells versus that in untreated cells will be  < 1.

RNA Sequencing

Total RNA from FA-treated BEAS-2B cells was converted into complementary DNA (cDNA) libraries using a Truseq RNA Sample Preparation V2 Kit (Illumina). Validation of library preparations was performed on an Agilent Bioanalyzer using the DNA1000 kit. Library concentrations were adjusted to 4 nM, and libraries were pooled for multiplex sequencing. Pooled libraries were denatured and diluted to 15 pM and then clonally clustered onto the sequencing flow cell using the Illumina cBOT Cluster Generation Station and a TruSeq Paired-End Cluster Kit v3-cBot-HS. Sequencing was performed on an Illumina HiSeq2500 Sequencing System using a TruSeq SBS Kit v3-HS. Quality control of FASTQ data was performed using FastQC (version 0.10.1; Babraham Bioinformatics group), and RNA reads were mapped to the human genome (UCSC hg19; February 2009 release; Genome Reference Consortium GRCh37) using STAR (version 2.5.2; Alexander Dobin) (Dobin et al. 2013) with the human reference GTF annotation file (GRCh37). Transcript counts were calculated and normalized using Gfold (version 1.0.8; Jianxing Feng) and DESeq (version 1.6.1; Simon Anders) (Anders and Huber 2010; Feng et al. 2012). The DESeq negative binomial distribution was used to call differentially expressed genes. A total of 654 genes were identified as differentially expressed genes (p < 0.05). Differentially expressed genes were further investigated for biological function and pathway enrichment using Ingenuity Pathway Analysis (IPA, Qiagen). RNA-seq data have been deposited in the Gene Expression Omnibus database (accession number GSE87541).

RNA Extraction and Real Time Quantitative RT-PCR

Total RNA was extracted using TRIzol reagent (Invitrogen) according to the manufacturer’s protocol. The quantity and purity of the RNA prepared from each sample were determined by UV absorbance spectroscopy. Reverse transcription (RT) was performed using the SuperScript III First-Strand Synthesis System (Invitrogen) with 1 μg of RNA in a final reaction volume of 20 μL. After incubation at 50°C for 50 min, RT was terminated by heating at 85°C for 5 min. Quantitative real-time PCR analysis was performed using Power SYBR Green PCR Master Mix (Applied Biosystems). PCR was performed in triplicate. Transcripts of target genes were calculated with the ΔΔCt method using the formula R = 2^−ΔΔCt, where R is the relative expression ratio of a target gene in comparison to a reference gene, and ΔΔCt = ΔCt(target) − ΔCt(reference). ΔCt (target) is the Ct deviation of untreated – treated sample of the target gene transcript; ΔCt (reference) is the Ct deviation of untreated – treated sample of the reference gene transcript. In this method, the relative levels of the target gene expression are presented as a fold change relative to the reference gene expression. A relative quantity of 1 indicates no change in expression levels. For the ΔΔCt method to be valid, the PCR efficiencies of the target and the reference must be approximately equal. Thus, we assessed the efficiencies of the target amplifications and the reference (Tubulin) amplification using the equation E = 10^[−1/slope] as described by Pfaffl (2001). The real-time PCR efficiency of Tubulin amplification was 2.18, and the efficiencies of most of the target genes we tested were between 2.14 and 2.21. For these genes, we used the ΔΔCt method to calculate their relative gene expressions given that the efficiencies were approximately equal. However, we analyzed the data for the genes HMGA2, SERPINB5, MYEOV, HES1, and LAMA4 using the Pfaffl method (Pfaffl 2001) because the amplication efficiencies of these genes were between 1.54 and 1.99. See Table 2 for primers.

Effects of FA on Acetylation of the N-Terminal Tails of Cytosolic Histones

The formation of FA-histone adducts on lysine residues prevents the acetylation of the same site by histone acetyltransferase (HAT) (Lu et al. 2008; Rager et al. 2013). Given that FA reacts with histones in BEAS-2B cells (Figure 1), we hypothesized that FA may inhibit the induction of physiologic PTMs on histone proteins. To test this hypothesis, total histones were isolated by acid extraction from BEAS-2B cells treated with or without FA. Histone PTMs were detected by Western blot analysis. Unexpectedly, no significant alterations were observed (Figure 2A). The previous observation that FA cannot react with acetylated sites on histones (Lu et al. 2008) suggests that FA likely reacts with unmodified, newly synthesized histones. Because such histones localize to the cytoplasm and represent only a small percentage of total histones (Loyola et al. 2006), this change may not be detectable when total histones are analyzed. To confirm this possibility, the cytosolic fraction (the water-soluble components of cytoplasm), the nuclear extract, and soluble chromatin fractions were isolated as previously described (Chen et al. 2013) and analyzed by Western blotting. Figure 2B shows that FA causes a drastic decrease in the acetylation (Ac) of H3K9, H3K14, and H4K12 residues in the cytosolic fraction. Interestingly, the levels of these two modifications were increased in nuclear extract fractions but were slightly decreased in chromatin fractions following FA treatment (Figure 2B).

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Figure 2.

Effects of formaldehyde (FA) on acetylation of N-terminal tails of cytosolic histones. (A) BEAS-2B cells were exposed to FA for 6 h. Total histones were prepared by acid extraction and subjected to Western blot analysis. No marked decrease of histone modification was observed (n = 2). The band intensities were quantified using ImageJ (version 1.46r; National Institutes of Health) software. (B) Cytosolic fractions, nuclear extracts, and chromatin fractions were isolated from BEAS-2B cells treated with or without FA for 6 h and subjected to Western blot analysis. FA caused drastic decreases in cytosolic levels of H4K12Ac and H3K9&K14Ac (n = 2). (C) Cytosolic fractions were isolated from cells treated with or without FA (0.5 mM, 6 h) and the proteasome inhibitor MG132 (20 μM, 2 h) and then subjected to Western blot analysis. FA exposure increased cytosolic H3 in the presence of MG132 (n = 2).

Effects of FA on the Deposition of Histone H3 into Chromatin

Acetylation of the N-terminal tails of H3 and H4 is critical for histone nuclear import and assembly into chromatin (Burgess and Zhang 2013). Thus, the reduction of cytosolic H3K9Ac, H3K14Ac and H4K12Ac by FA treatment may compromise the assembly of newly synthesized histones into chromatin. If this occurs, it should be expected that levels of nucleosomal histone H3 and H4 will be reduced after FA exposure. In fact, the amount of histone H3 in the chromatin fraction decreased by approximately 20% (Figure 2B). The level of total H3 protein remained unchanged following FA exposure, making it unlikely that the observed reduction of H3 in the chromatin fraction is due to reduced expression of H3 (Figure 2A). Interestingly, FA increased the levels of total histone H3, H3K9Ac, H3K14Ac, and H4K12Ac in the nuclear extracts (Figure 2B). This finding implies that the delivery of histones to chromatin is blocked by FA exposure, resulting in the accumulation of FA-modified histones in the nuclear extract fraction. The levels of cytosolic H3 were also increased following FA exposure in the presence of the proteasome inhibitor MG132 (lanes 1 and 3; 2 and 4 in Figure 2C), suggesting that a) exposure to FA reduces histone nuclear import and b) aggregated FA-modified histones are degraded by the proteasome pathway. These critical observations reveal the activation of quality control mechanisms in histone homeostasis.

To further investigate the effects of FA exposure on chromatin assembly, we compared the levels of histones at a number of genomic loci before and after FA treatment. It was hypothesized that the inhibition of chromatin assembly would lead to a reduction of histone enrichment at specific genomic loci. ChIP assays were used to measure the amount of histone variant H3.3 at several genomic loci. The noncanonical histone variant H3.3 was used because it primarily localizes to active regions of the genome with high rates of histone turnover; thus, the impact of lack of histone supplies should be greatest in the H3.3 regions (Deal et al. 2010). The results of the ChIP assays clearly showed that the amount of H3.3 was significantly reduced at the majority of loci tested (Figure 3A). Taken together, we conclude that FA compromises chromatin assembly.

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Figure 3.

Effects of formaldehyde (FA) on H3.3-containing nucleosome assembly. (A) BEAS-2B cells with stable expression of FLAG-H3.3 were treated with or without FA for 6 h. Mono- and dinucleosomes were prepared and subjected to chromatin immunoprecipitation (ChIP) assays with FLAG antibodies to isolate FLAG-H3.3 nucleosomes. The data shown are the mean ± standard deviation (SD) from real-time quantitative polymerase chain reactions (qPCRs) performed in triplicate. *p < 0.05; **p < 0.01. The loci tested are mostly promoters except for GAPDH (gene body) and CSRP3 (gene end). Relative fold change was calculated after normalization to input and no antibody control. (B) Exposure to FA at physiologically relevant concentrations inhibits N-terminal tail acetylation of cytosolic histones. Cytosolic cell fractions were isolated from BEAS-2B cells, UTA6 cells, and RPMI 2650 cells treated with or without 100 μM FA and were subjected to Western blot analysis (n = 2). (C) Exposure of cells to physiologically relevant concentrations of FA compromises assembly of H3.3-containing nucleosomes. BEAS-2B cells with stable expression of FLAG-H3.3 were treated with or without FA for 48 h. Mono- and dinucleosomes were prepared and were subjected to ChIP assays to isolate FLAG-H3.3 nucleosomes. Data are the mean ± standard deviation (SD) from qPCRs performed in triplicate. *p < 0.05; **p < 0.01.

Effects of Physiologically Relevant Concentrations of FA on Acetylation of the N-Terminal Tails of Cytosolic Histones

We have used a relatively high dose of FA (0.5 mM) to demonstrate inhibition of histone modifications and chromatin assembly following FA exposure. Because epigenetic effects are considered to be nonlinear, it is important to test whether similar outcomes can be seen at a more physiologically relevant dose. It is reported that endogenous concentrations of FA in the blood are approximately 0.1 mM in rats, monkeys, and humans. Additionally, FA concentrations in the liver and nasal mucosa of the rat are 0.2 and 0.4 mM, respectively (Andersen et al. 2010; Casanova et al. 1988; Casanova-Schmitz et al. 1984; Heck and Casanova 2004; Heck et al. 1982, 1985). Based on these reports, it was determined that 100 μM FA is a physiologically relevant concentration. Relatively high levels of endogenous FA raised the question of whether the formation of FA–histone adducts could be a normal process. If so, environmental exposure to a physiologically relevant dose of FA may only slightly increase the burden of cellular toxicity. Exposure to FA could interfere with normal cellular processes because a major source of endogenous FA is from the demethylation of histones, RNA, and DNA within the nucleus (Walport et al. 2012), indicating that the level of endogenous FA in the cytoplasm might be relatively low. Thus, exposure to FA, which mainly targets cytoplasmic proteins, would have a significant impact on cytoplasmic processes even at physiological concentrations. Moreover, FA scavengers such as glutathione would be rapidly consumed by continuous exposure to the FA that we used in the experiments. To determine the effects of exogenous exposure to a physiologically relevant concentration of FA on cellular processes, we measured the levels of cytosolic H4K12Ac in several cell lines after they were exposed to 100 μM FA. The treatment significantly reduced the levels of cytosolic H4K12Ac in BEAS-2B cells, UTA6 cells (a human osteosarcoma cell line), and RPMI 2650 cells (a human nasal epithelial cell line) (Figure 3B); however, the time needed to induce the change varied from 24 h to 72 h among the different cell lines tested. The data suggest that continuous exposure of cells to physiological concentrations of FA significantly affects posttranslational modification of cytosolic histones at critical sites.

To determine whether nucleosome assembly is compromised after exposure to 100 μM FA, ChIP assays were performed to measure the amount of histone H3.3 in several genomic loci before and after FA exposure. H3.3 was depleted by FA treatment at the promoters of the genes that encode EGR2, S100A10, and H3.3 (Figure 3C). This result indicates that nucleosome assembly in these sites is compromised following FA exposure at a physiologically relevant concentration.

Effects of FA on Chromatin Structure

Repression of histone expression interferes with chromatin assembly and enhances chromatin accessibility (Gossett and Lieb 2012). Chromatin accessibility could be affected by FA exposure through inhibition of chromatin assembly. MNase digestion assays were used to investigate the influence of FA exposure on chromatin accessibility. Nuclei from BEAS-2B cells treated with or without FA were extracted and digested with MNase. Extracted DNA was then monitored for changes of protective monomer ladders by agarose gel electrophoresis. No changes were observed in cells treated with 0.25 mM FA (Figure 4A, lanes 1–6). However, exposure to 0.5 mM FA increased overall sensitivity to MNase. This effect was obvious when the nuclei were digested with 12 × 10⁻³ U/μL and 24 × 10⁻³ U/μL MNase, where the bands indicative of polynucleosomes (larger than 4-mers and 3-mers, respectively) were missing from FA-treated cells (arrowheads when comparing lanes 2 and 8; stars when comparing lanes 3 and 9). The general increase in chromatin accessibility with FA treatment suggests that nucleosome occupancy at genomic loci changed after FA exposure. MNase-qPCR was then utilized to determine the nucleosome occupancy at several genomic loci by measuring the ratio of the abundance of target DNA sequences in MNase-digested mononucleosomes isolated from FA-treated versus control cells (Jin and Felsenfeld 2006). As shown in Figure 4B, nucleosome occupancy decreased in the majority of sites we tested. Next, to investigate how MNase sensitivity is changed by exposure to physiological levels of FA, we performed MNase digestion assays after treating cells with 0.1 mM FA for 48 h. Figure 4C shows that when digested with 6 × 10⁻³ U/μL and 12 × 10⁻³ U/μL MNase, the intensities of the trinucleosome band (arrowheads when comparing lanes 2 and 5) and the dinucleosome band (stars when comparing lanes 3 and 6), respectively, were greatly reduced following low-dose FA exposure (Figure 4C). Taken together, these data demonstrate that FA exposure can increase chromatin accessibility.

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Figure 4.

Effects of formaldehyde (FA) exposure on chromatin structure. (A) FA exposure increases chromatin accessibility. BEAS-2B cells were treated with or without FA for 6 h. Nuclei were isolated and digested with micrococcal nuclease (MNase). Digested DNA was extracted, electrophoretically separated on a 2% agarose gel, and visualized by ethidium bromide staining. Representative results from two independent experiments are shown. (B) Changes in nucleosome occupancy. MNase-digested monomeric DNA bands from untreated and FA-treated BEAS-2B cells were excised, and the DNA was extracted. The abundance of a sequence in protected nucleosomes was determined by real-time quantitative polymerase chain reactions (qPCRs) performed in triplicate. *p < 0.05; **p < 0.01. (C) Chromatin accessibility is increased with the exposure of cells to physiologically relevant doses of FA. BEAS-2B cells were treated with or without 0.1 mM FA for 48 h followed by MNase digest analysis as described in (A). Representative results from two independent experiments are shown.

Characterization of Cancer-Related Genes Mediated by FA Exposure

Aberrant chromatin assembly can result in transcriptional defects. Thus, we hypothesized that the inhibition of chromatin assembly resulting from the formation of FA–histone adducts contributes to FA-mediated transcriptional dysregulation. RNA-seq was first used to characterize FA-inducible genes in BEAS-2B cells exposed to 100 μM FA for 48 h. DESeq (Anders and Huber 2010) was used to call differentially expressed genes in the RNA-seq data generated from two biological replicates (Feng et al. 2012) (p < 0.05). A total of 654 genes were identified as FA-responsive genes (see Table S1). Figure 5A shows an RNA-Seq heatmap for differentially expressed genes. Ingenuity Pathway Analysis (IPA) was then used to annotate the data set and to analyze the data in the context of biological processes, pathways, and networks. Top diseases and biological functions related to this data set were dermatological diseases, cancer, inflammatory response, tumor morphology, and neurological disease (Figure 5B).

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Figure 5.

Ingenuity Pathway Analysis (IPA) of formaldehyde (FA)-induced genes. (A) RNA sequencing (RNA-Seq) heatmap for differentially expressed genes. Color represents the log₂-fold changes between FA-treated and untreated cells. (B) The top five diseases and biological functions related to 654 FA-responsive genes are shown. (C) Signaling pathways, including chronic myeloid leukemia signaling, are associated with FA-responsive cancer-related genes.

It is important to note that of 654 FA-responsive genes, 361 were identified as cancer-related. Signaling pathway analysis showed that these genes were involved in several pathways, including p53 signaling, HER-2 signaling, and Wnt/beta-catenin signaling (Figure 5C). Interestingly, leukemia signaling was present in these cancer pathways, providing a potential mode of action for FA-induced leukemia. To validate the RNA-seq results, eight up-regulated and eight down-regulated genes were selected for qPCR analysis. The selected genes included well-known oncogenes such as FOS and JUN and tumor suppressors such as CDKN1A and SERPINB5 from the top and bottom 60 FA-responsive genes (Tables 3 and ​and4).4). All 16 selected genes are associated with head/neck neoplasia, hematological neoplasia, or both (Tables 3 and ​and4).4). The similar tendency of differential expression with FA treatment of all of the selected genes was observed with RT-qPCR as shown with RNA-seq (Figure 6A, B). The functional and RT-qPCR analyses demonstrate that the RNA-seq data were reliable and confirm that the selected 16 cancer-related genes were dysregulated following FA exposure.

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Figure 6.

Effects of H3.3 knockdown on the expression of formaldehyde (FA)-responsive genes. (A, B) Real-time polymerase chain reaction (PCR) validation of RNA sequencing (RNA-seq) results. Eight up-regulated (A) and eight down-regulated (B) cancer-related genes following exposure to FA (100 μM for 48 h) in BEAS-2B cells were selected from RNA-seq results. mRNA levels of these genes were analyzed by reverse transcriptase quantitative real-time PCR (RT-qPCR). Differential expression of these genes was in accord with the RNA-seq results. Data are the mean ± standard deviation (SD). (n = 3). *p < 0.05; **p < 0.01. (C) RT-qPCR analysis of H3.3 mRNA levels in BEAS-2B cells after 48-h transfection with control (Ctrl) short interfering RNA (siRNA), H3.3 siRNA 1, or H3.3 siRNA 2. (D) Western blot analysis of ectopic H3.3 protein levels in BEAS-2B cells after 48-h transfection with control (Ctrl) siRNA or with two distinct siRNAs for H3.3 (H3.3 siRNA 1 and H3.3 siRNA 2). Antibodies against β - actin (top), FLAG (middle), or H3 (bottom) were used. (E, F) RT-qPCR measurements of transcripts of the indicated FA-responsive cancer-related genes in H3.3 knockdown and control cells. The data shown are the mean ± standard deviation (SD). (n = 3). *p < 0.05; **p < 0.01.

Effects of Chromatin Assembly Inhibition on Expression of FA-Responsive Cancer-Related Genes

To delineate the contribution of defective chromatin assembly in FA-mediated dysregulation of gene expression, we examined how the expression of the selected 16 cancer-related genes was influenced by knockdown of the histone variant H3.3. Because knockdown of H3.3 disrupts chromatin assembly, the overlap between transcriptional patterns generated by H3.3 knockdown and FA exposure may represent the genes regulated by FA-induced aberrant chromatin assembly. Expression of canonical histones H3.1 and H3.2 peaks during S phase, and their assembly is replication-dependent. By contrast, H3 variant H3.3 is expressed throughout the cell cycle, and its assembly is not dependent on DNA replication (Szenker et al. 2011). Moreover, H3.3 is mainly localized at active regions with high rates of histone turnover (Deal et al. 2010). Thus, the assembly of H3.3 should be affected first and to the greatest extent by a lack of histone supplies. Knockdown of histone variant H3.3 was carried out using two distinct siRNAs. The knockdown efficiencies of H3.3 mRNA were approximately 74% and 60% in cells transfected with H3.3 siRNA 1 and H3.3 siRNA 2, respectively, compared with control siRNA (Figure 6c). Western blot analysis showed that the protein levels of FLAG-H3.3 and most likely endogenous H3.3 were reduced by transfection of both siRNAs for H3.3 (Figure 6D). Six of the eight genes that were highly up-regulated by FA exposure (including FOS, JUN, and JUNB) and three of the eight highly down-regulated genes (including CDKN1A and SERPINB5) were also abnormally expressed as a result of H3.3 knockdown (Figure 6E, F; see also Figure S2). These results suggest that aberrant chromatin assembly dysregulates expression of FA-responsive cancer-related genes and may play an important role in FA-induced carcinogenesis.