Hum. Reprod. Advance Access published online on August 29, 2007
Human Reproduction, doi:10.1093/humrep/dem241
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Class I histone deacetylase expression in the human cyclic endometrium and endometrial adenocarcinomas
1 Department of Anatomy and Reproductive Biology, RWTH University of Aachen, Wendlingweg 2, 52074 Aachen, Germany 2 Department of Child and Adolescent Psychiatry, RWTH Aachen University, Aachen, Germany 3 Institute of Pathology, Düren, Germany
4 Correspondence address. Tel: +49-241-8088926; Fax: +49-241-8082508; E-mail: ckrusche{at}ukaachen.de
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Abstract |
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BACKGROUND: Class I histone deacetylases (HDACs) and acetylases (HATs) are members of transcriptional pre-initiation complexes assembled by steroid hormone receptors. Recently, HDAC inhibitors were shown to enhance differentiation of endometrial fibroblasts and endometrial adenocarcinomas. However, there is only rare information on HDAC and HAT expression in the human endometrium.
METHODS: HDAC-1, -2, -3 and HAT (PCAF and GCN5) mRNA expression was studied in tissue from premenopausal women undergoing hysterectomy by real-time or semiquantitative RT–PCR. HDAC protein expression was assessed by Western Blot and immunohistochemistry. In endometrial adenocarcinomas (n = 17), HDAC-1 expression was studied by immunohistochemistry.
RESULTS: In the human endometrium, HDAC-1, -2, -3 and PCAF mRNA are expressed without cyclical changes. Western blot analysis demonstrated that HDAC-2 protein expression was slightly, but significantly elevated in the secretory phase (P < 0.01 versus day 5–8), whereas HDAC-1 and -3 protein expression was constitutive throughout the menstrual cycle. By immunohistochemistry, nuclear expression of HDAC proteins was detected in all endometrial cell types. In the case of HDAC-3, immunostaining was significantly reduced in the endometrial surface epithelium on day 6–10 (P < 0.01 versus days 15–18 and 24–28). Compared to normal endometrium, a high proportion of endometrial adenocarcinomas showed impaired HDAC-1 protein expression in the epithelial and stromal compartment.
CONCLUSIONS: Class I HDACs and HATs are expressed in the human endometrium throughout the menstrual cycle, suggesting the cyclic endometrium as a potential target for HDAC inhibitors. We hypothesis that alterations of HDAC and/or HAT expression are potentially involved in impaired endometrial differentiation.
Key words: cancer/endometrium/histone deacetylases/steroid hormones/uterus
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Introduction |
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The overall and local structure of the chromatin plays a pivotal role in the regulation of gene transcription. Chromatin structure is modulated by epigenetic modifications like DNA-methylation, histone-phosphorylation and -acetylation (Momparler, 2003
; Thiagalingam et al., 2003). In particular, the acetylation status of the N-terminal lysine residues of histone proteins seem to correlate with gene transcription in eukaryotes (Allfrey et al. 1964; Santos-Rosa and Caldas, 2005; Joo et al., 2006). The extent of histone acetylation is influenced by the antagonistic activities of histone acetylases (HATs) and histone deacetylases (HDACs). However, these enzymes also acetylate and deacetylate specific non-histone proteins, e.g. the tumor suppressor protein p53 and 
-tubulin (Hubbert et al., 2002; Langley et al., 2002; Glozak et al., 2005).
Three different classes of HDACs, which are categorized according to sequence and functional homology to the yeast counterparts, have been described so far, namely HDAC-1, -2, -3 and -8 which belong to Class 1 HDACs and HDAC-4, -5, -6, and -7 which belong to HDAC class II. The third class comprises the sirtuins, which are NAD+ dependent (Thiagalingam et al., 2003).
The class I HDACs, HDAC-1, -2 and -3, play an important role in steroid-hormone dependent gene expression by directly interacting with proteins which are recruited to steroid-hormone receptor proteins after ligand-binding (Giangrande et al., 2000; Guenther et al. 2001; Shang and Brown, 2002; Liu and Bagchi, 2004). In addition, HDACs specifically interact with a variety of proteins belonging to multicomponent complexes, which control gene transcription like NURD, Sin3, CoREST (Strahl and Allis, 2000; Grozinger and Schreiber, 2002; Sengupta and Seto, 2004). Consequently, HDACs are involved in the modulation of proliferation, differentiation and in cases of aberrant expression in carcinogenesis (Lagger et al. 2002; Bradbury et al., 2005; Krusche et al., 2005; Phillips et al., 2005).
The potential relevance of HDACs in endometrial differentiation was demonstrated by the effects exerted by HDAC inhibitors on uterine cells. Sakai et al. (2003) showed that HDAC inhibitors enhance the progesterone-induced decidualization of human endometrial fibroblasts cultured in vitro. In mice, HDAC inhibitors like Trichostatin A and sodium butyrate enhance the estradiol (E2) induced proliferation as well as the estrogen- (ER) and progesterone receptor (PR) expression of uterine cell populations (Gunin et al., 2005). Furthermore, it has to be questioned whether the anticonvulsive drug valproic acid, which is also a HDAC inhibitor, affects fertility in women suffering from epilepsy (Artama et al., 2006).
HDACs also came into focus as therapeutic targets in cancer. The application of HDAC inhibitors leads to growth arrest and the induction of differentiation in a variety of in vitro cultured tumor cells, including endometrial cancer cell lines, in experimental animal tumor models and in first clinical trials (Marks et al., 2001; Vigushin et al., 2001; Takai et al., 2004; Hess-Stumpp, 2005; Uchida et al., 2005; Xiong et al., 2005; Takai et al., 2006; Minucci and Pelicci, 2006).
Very recently, it was shown that the class I HDAC-8 is expressed in human myometrium and in the vascular smooth muscle cells of the endometrium (Waltregny et al., 2004; de Leval et al., 2006). However, the expression of the other class I HDACs in the human endometrium is largely unknown but of considerable interest with regard to basic research and new therapeutic strategies. Therefore, we assessed HDAC-1, -2 and -3 mRNA and protein expression in the human cyclic endometrium. Since HDAC-1 expression plays a crucial role in cell cycle control, we studied HDAC-1 protein expression in human endometrial carcinomas by immunohistochemistry. Furthermore, mRNA expression of the two main HATs, PCAF and GCN5, was also studied in human cyclic endometrium.
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Materials and Methods |
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Samples from human cyclic endometrium
Endometrial samples were obtained throughout the menstrual cycle from normal fertile, premenopausal women undergoing hysterectomy in collaboration with the Departments of Gynaecology and Obstetrics of Marienhospital (Aachen), Luisenhospital (Aachen) and St Antonius Hospital (Eschweiler). 24 to 31 endometrial samples were used dependinig on the employed method. ‘Patients’ age was between 30 and 46 years. All women had a regular menstrual cycle (25–35 days) and did not take steroid hormones at least 6 month prior to surgery. The reason for the hysterectomy was in nearly all cases leiomyomata with menorrhagia and dysmenorrhea.
Dating of each specimen was done by menstrual history, histological examination (Noyes et al., 1950) and PR as well as Ki67 (marker of cell proliferation) immunohistochemistry. In addition, 17
- estradiol progesterone and LH blood serum levels were assessed on the day of hysterectomy by routine laboratory diagnostics of the Department of Gynaecological Endocrinology and Reproductive Medicine, University Hospital, RWTH Aachen.
The human endometrial samples have been derived as anonymous samples from incidental hysterectomies performed exclusely for medical reasons. The appropriate ethical approval by the Ethics Committee of the Medical Faculty of the RWTH University of Aachen (Nr. EK 347) has been obtained for these studies.
Endometrial cell culture
Endometrial cell cultures originating from proliferative phase endometrium were performed from three different donors. Endometrial epithelial and stromal cell culture was performed as described previously (Classen-Linke et al., 1997). One group of endometrial epithelial cells and fibroblasts was treated only with E2 (Sigma; 10–8 M); the other group with E2 (10–8 M) in combination with medroxyprogesterone acetate (10–6 mol/l MPA; Sigma, Deisenhofen, Germany), a metabolically stable progestin. Both hormones were diluted in ethanol. The final ethanol content of the culture medium was <0.1%. After recovery, cells were cultured at first for six days without steroid hormones. Thereafter, cells were treated for six days either with E2 or E2 and MPA to mimic the hormonal conditions of the proliferative and secretory phase of the menstrual cycle.
Human endometrial adenocarcinomas
The carcinomas were predominantly endometrioid adenocarcinomas, hallmarked by post-menopausal bleeding, bleeding-dysfunctions or discharge with normal cervical finding. We assessed 17 endometrial adenocarcinomas, which were kindly provided by the Department of Pathology, RWTH Aachen University, Aachen, Germany and Institute of Pathology, Düren, Germany. The carcinomas were rated according to the pathological tumor-node-metastasis (pTNM) and International Federation of Gynecology and Obstetrics (FIGO) classification. Furthermore, the PR and ER protein expression and the histological differentiation status were determined.
Semiquantitative RT–PCR
Endometrial tissue samples used for semiquantitative RT–PCR were stored and homogenized in RNAwiz solution (WAK Chemie; Steinbach/Ts., Germany) at –35°C. After addition of 100 µl chloroform to 600 µl of tissue homogenate and a centrifugation step (10 600 xg, 10 min, 4°C), the supernatant was used to isolate total RNA by the RNeasy Mini Kit (Qiagen, Hilden, Germany). The obtained endometrial RNA was treated with DNAse (DNA-freeTM Kit, Ambion, Huntingdon, UK) to eliminate contaminating DNA.
RNA isolation from cultured and freshly separated endometrial epithelial and stromal cells for semiquantitative RT–PCR was performed with the High Pure RNA isolation kit (Roche, Mannheim, Germany) according to the manufacturer's recommendations.
Complementary DNA (cDNA) was prepared from endometrial RNA using Ready-to-goTM You-Prime First-Strand Beat-Kit (Amersham Biosciences, Freiburg, Germany). About 5 µg of total RNA were used and priming was performed with 1.6 µg oligo(dT)15-primer according to Kit instructions. Equal RNA input in the cDNA synthesis was controlled by loading 4 µl of the cDNA synthesis reaction on an agarose gel and measurement of the optical density (OD) of 18S and 28S ribosomal RNA bands by the Kodak Image Station 440 (PerkinElmerLifeSciences, Boston, USA). The remaining 29 µl of synthesis solution were diluted with 121 µl DEPC-water and 2 µl (equivalent to 60 ng of RNA) were used in the following PCR reactions.
Primers used for semiquantitative PCR reaction were made by GeneFisher Software (http://bibiserv.techfak.uni-bielefeld.de/genefisher). Table 1 provides a summary of the primer pairs used. To guarantee reliable quantification of the relative mRNA expression, the logarithmic phase of each PCR reaction was determined in separate experiments. Amplification of the S26 ribosomal protein was employed to normalize the PCR reaction, e.g. to eliminate pipetting failures and/or unequal RNA/cDNA load between different samples in the PCR reaction.
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The 50 µl PCR reaction contained 2 µl of diluted cDNA, 5 µl 10x Taq reaction buffer plus Mg2+ (final concentration 1.5 mM), 2 µl 10 mM dNTP and 2 units of Taq DNA polymerase (Roche) as well as 10 pM of specific forward and reverse primer (Table 1).
PCR running program was as follows: (i) initial denaturation at 94°C (2 min) followed by (ii) 27–35 cycles (Table 1): 94°C (1 min), primer annealing at 62–67°C (1 min), elongation at 72°C (1 min) and (iii) one final elongation step at 72°C for 3 min.
Identity of PCR products was determined by sequencing of representative bands (SeqLab, Göttingen, Germany).
About 15 µl of each PCR product were loaded to a 1.2% agarose gel (1x TAE buffer = 40 mM Tris-acetate, 1 mM EDTA, pH 7.5; with 0.4 µg ethidium bromide/ml gel). Afterwards, the OD of the bands was measured by the Kodak Image Station 440 CF. The obtained data were analyzed with the 1D Image Analysis software, version 3 (PerkinElmerLifeSciences, Boston, USA). The ODs of the PCR products was divided by the OD of the housekeeping genes. This ratio is named ‘arbitrary unit’.
We performed two negative control experiments: instead of cDNA, 2 µl of water or 60 ng RNA was used in the PCR reactions. Both control reactions did not generate products.
RNA isolation, cDNA synthesis and real-time RT–PCR
For the real-time RT–PCR, total RNA was isolated from endometrial samples by using the High Pure RNA Tissue Kit (Roche). About 25 mg endometrial tissue was transferred to 900 µl lysis-/binding-buffer and was homogenized for 90 s with a glass-teflon-homogeniser. Thereafter, 700 µl of water-saturated phenol/chloroform/isoamyl alcohol mix (Ambion) was added, mixed and centrifuged (13 000g, 2 min, room temperature). The aqueous supernatant was mixed with 0.5 volumes of ethanol and loaded onto the column provided with the kit. The subsequent steps of RNA isolation were made according to the kit protocol with a prolongation of DNAse I treatment to 45 min at 37°C.
One microgram RNA was reverse transcribed into cDNA according to the manufacture's specifications [first strand cDNA synthesis (AMV) Kit (Roche)], using oligo-(dT)15 primer and the cDNA was diluted 1:1 with sterile water. Real-time PCR was performed with the LightCycler (Roche).
5-Aminolevulinat-Synthetase (ALAS), which is expressed with 
500 mRNA molecules/cell, was used as housekeeping gene. ALAS expression was measured by the LightCycler-h-ALAS housekeeping gene set and the LightCycler FastStart DNA MasterPlus HybProbe Kit (both Roche) according to the kit protocol.
For the target genes, LightCycler FastStart DNA MasterPlus SYBR Green I Kit (Roche) was used. Primer pairs were created with the Primer3 primer design program (http://frodo.wi.mit.edu/cgi-bin/primer3/primer3_www.cgi; Table 1).
The PCR volume was 20 µl and consisted of 2 µl cDNA and 18 µl PCR reaction mix. The MgCl2 concentration was 3 mM. Water and an equivalent of 60 ng RNA were employed as negative controls in each PCR reaction.
Analysis of PCR products resulting from melting curves and product size was checked on ethidium bromide-containing agarose gels.
The PCR runs were programmed with the LightCycler Software, version 3.5.
Each LightCycler run included the following four steps:
- Denaturation and activation of the Taq polymerase at 95°C, 10 min
- Amplification and quantification: denaturation at 95°C, 10 s; annealing 5 s (annealing temperature see Table 2), extension at 72°C, 20 s
- Melting curve profile: 95°C, 5 s, cooling to 77°C, 30 s and slow heating to 95°C (0.1°C per second) with continuous measurement
- Cooling to 40°C
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Quantification of RT–PCR results
To correct RT–PCR results of HDAC-1, -2 and -3 mRNA expression for pipetting errors and inter-sample variations of the cDNA synthesis reaction, ALAS was used as housekeeping gene.
To determine the threshold cycle (Ct) the LightCycler software used the ‘second derivative maximum’ method.
The efficiency of target and housekeeping gene PCRs was assessed with LightCycler runs using a dilution series of an endometrial cDNA pool (1:1, 1:10, 1:100 and 1:1000) with each dilution step utilized in triplicate. The slope of the PCR reaction is calculated from the average Ct of each dilution step by the LightCycler software. The slope of the HDAC-1, HDAC-2 and HDAC-3 PCR was –4.565, –4.420 and –3.467, respectively. The slope of the ALAS PCR was –3.346. The efficiency is then calculated as E = 10–1/slope.
For further efficiency corrections of target and housekeeping gene RT–PCRs, a so called ‘coefficient file’ is generated by the LightCycler software.
To assess endometrial target and housekeeping gene mRNA expression, each cDNA sample was used in duplicate. For all genes, the average Ct was calculated. With these data and the respective coefficient files, the RelQuant Software, version 1.01 (Roche) calculates the efficiency-corrected ratio of target gene and housekeeping gene. The results of this calculation are given as ‘arbitrary units’.
To determine the relative quantity of class I HDAC-1, -2 and -3 mRNA expression in the endometrium, at first the respective Ct values were normalized by subtracting the Ct-value of the ALAS housekeeping gene (
Ct = Ct target – Ct ALAS).
To compare the HDAC-1, -2 and -3 expression levels, the 2–Ct value of HDAC-1, -2 and -3 was calculated and HDAC-3 mRNA expression was set arbitrarily to ‘1’.
Northern hybridization
Amplification of partial sequences of the HDAC-1, -2 and -3 cDNA was done with the specific cloning primer pairs listed in Table 1 and PCR products were cloned with the TOPO TA Cloning Kit according to the manufacturer's recommendations (Invitrogene, Paisley, UK).
Plasmids from transformants were isolated with the GeneElute Plasmid Miniprep Kit (Sigma-Aldrich, München, Germany). The orientation of the inserts was tested by PCR reaction using the insert specific forward or reverse primers in combination with the M13 forward primer. Additionally, plasmids were sequenced (Seqlab).
The MAXIscriptTM in-vitro-transcription Kit (Ambion) and SP6 RNA Polymerase (Ambion) were used to obtain the antisense transcript, labeled non-radioactively by the addition of 0.4 µl DIG-UTP (Roche). The efficiency of labeling and the size of the in vitro transcripts were checked by gel electrophoresis, Northern blot and immunological blot detection with the Anti-DIG-AP-conjugate (Roche).
Approximately 5 µg total RNA was separated on 1.2% agarose gels containing 0.66 M formaldehyde and 1x MSE buffer (10x MSE buffer: 0.2 M morpholinopropansulfonic acid (MOPS, pH 8), 50 mM sodium acetate and 5 mM EDTA, pH 8). As running buffer 1x MOPS pH 7 was used.
The RNA was transferred by capillary blotting to a neutral nylon membrane (Hybond N, AmershamBiosciences, Freiburg, Germany) using 10x SSC buffer (20x SSC buffer: 3.6 M NaCl, 0.3 M Na3Citrate, pH 7) and fixed under UV light (120 mJ; UV Stratalinker, Stratagene).
Blots were prehybridized in DIG Easy Hyb hybridization solution (Roche) at 60°C for 1 h. The DIG-UTP labeled in-vitro-antisense transcripts were denaturated at 65°C for 10 min in 50% formamide. After short cooling on ice, the transcripts were transferred to DIG Easy Hyb Hybridization solution (final concentration: 100 ng/ml). Hybridization was carried out overnight and stringent washing was performed with 0.2x SSC/0.1% sodium dodecyl sulphate (SDS) buffer (2 x 15 min, 60°C).
HDAC-1, -2 and -3 immunohistochemistry
Immunohistochemistry was performed on 5 µm thick paraffin sections after microwave pretreatment (10 mM citrate buffer, pH 6; 4 x 5 min; 600 W). After blocking unspecific binding sites (Histostain Plus Broad Spectrum Kit; Zymed Laboratories, South San Francisco, USA), sections were incubated for 1 h with the first antibody (either anti-HDAC-1 or HDAC-3 polyclonal rabbit antibody; supplier and dilution see Table 2). Detection of the first antibody was accomplished by using the Histostain Plus Broad Spectrum and AEC Kit (both Zymed Laboratories). Two negative control experiments were performed: (i) the first antibody was omitted and (ii) the first antibody was replaced by a polyclonal rabbit non-immune serum. Both negative control experiments did not generate an immunostaining.
Quantitative assessment was performed as follows. Cells were counted in hematoxylin- counterstained sections using a magnification of x400.
The surface epithelium and the glandular epithelium of the functional layer, as well as the epithelium of the basal glands, were assessed separately. In three areas per section, 100 epithelial cells were counted and the percentage of HDAC positive cells was calculated. Furthermore, the percentage of HDAC positive stromal cells was determined in the functional endometrial layer. Six endometrial samples from the proliferative and six samples from the secretory phase were studied. Within each immunostained section three different, randomly chosen areas were assessed. Endothelial and smooth muscles cells of the blood vessel walls as well as cells located within lymph follicles were not quantified.
Western blots
SDS-disc polyacrylamide gel electrophoresis was performed using 10% gels (20 µg protein per lane) under reducing conditions (5% mercaptoethanol) according to Laemmli (1970). The separated proteins were transferred to a polyvinylidene difluoride membrane (Immobilon P, Millipore, Hamburg, Germany, pore diameter 0.45 µm) by a semidry electroblotting procedure (2 mA/cm2) for 40 min. The membrane was blocked in TBS (5 mM Tris-buffered saline; pH 7.6) supplemented with 5% milk powder and 0.1% Tween (1 h). The detection of HDAC-1, -2 and -3 protein was performed by incubating the blocked membrane overnight at 4°C with the first antibody (dilution of antibodies; see Table 2). After washing with TBS/0.1% Tween, membranes were incubated for 1 h with a horse-radish peroxidase (HRP)-conjugated second antibody. For the detection of the three HDAC antibodies, goat anti-rabbit immunoglobulin (Ig)-HRP (DAKO Cytomatics) diluted 1:5000 in phosphate-buffered saline (PBS/1% milk powder/0.1%Tween) was used. Immunoreactive proteins were detected with the enhanced chemiluminescence kit (Amersham Pharmacia Biotech, Freiburg, Germany) according to the manufacture's instructions.
For relative quantification, -actin expression was analyzed on all blots after the detection of HDAC-1, -2 or -3. The bound -actin antibody was detected by goat anti-mouse IgG (Santa Cruz, CA, USA) diluted 1:5000 in PBS/1% milk powder/0.1%Tween.
The ODs of the HDAC-1, -2, -3 and -actin protein bands were measured using the Kodak Image Station 440 CF in the ‘luminescence’ modus and the 1D Image Analysis software, version 3. The OD of HDAC protein bands were divided by that of the corresponding -actin. The ratio is named ‘relative expression’.
Statistical analyses
Statistical analyses were performed using GraphPad Prism, version 3 for Windows (GraphPad Software, Inc., San Diego, USA).
Because the number of samples used in the five subphases of the menstrual cycle (early proliferative, late proliferative, early secretory, mid secretory and late secretory phase) was lower than 10, Gaussian distribution of values could not be estimated. Therefore, the non-parametric Kruskal–Wallis test was used if more than two groups were compared, as in the case of HDAC-1, -2 and -3 mRNA and protein expression as well as GCN5 and PCAF mRNA expression. The Dunn's multiple comparison test was used as post hoc test.
For comparison of two groups, the non-parametric Mann–Whitney U-test was used, e.g. when HDAC-1, -2 and -3 protein expression studied by immunohistochemistry was compared in stromal cells of the proliferative and secretory phase, and to compare endometrial GCN5 mRNA expression beween the proliferative and secretory phase, after pooling the proliferative phase (day 7–14) data and the secretory phase (day 15–28) data.
Non-parametric test results are given as median and range. To compare relative HDAC-1, -2 and -3 mRNA expression levels, the one-way analysis of variance (ANOVA) test was used. Each group contained 26 samples. The Bonferroni multiple comparison test was employed as post hoc test. The values of this assessment are given as mean ± SD. In all tests, P-values <0.05 were considered as significantly different.
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Results |
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HDAC-1, -2 and -3 mRNA expression in the human cyclic endometrium
The mRNA expression of class I HDAC-1, -2 and -3 was assessed by real-time RT–PCR and Northern Blot (Fig. 1A–C, respectively). All three HDACs were constitutively expressed in the human endometrium throughout the menstrual cycle (Kruskal–Wallis tests: PHDAC-1 = 0.9598; PHDAC-2 = 0.3434; PHDAC-3 = 0.0802).
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Northern Blot analysis of representative endometrial samples showed bands of the expected size of approximately 2–2.2 kb (Fig. 1A–C).
To compare the HDAC-1, -2 and -3 mRNA expression levels, the 2–Ct value of each HDAC was calculated from all endometrial samples (Fig. 2). HDAC-3 mRNA expression was lowest. HDAC-2 mRNA expression was 22.78 ± 7.77-fold, and HDAC-1 mRNA expression was 9.99 ± 4.48-fold, higher than HDAC-3 mRNA expression. The differences in HDAC mRNA expression levels were statistically significant (ANOVA: P < 0.0001; post hoc Bonferroni's multiple comparison tests: HDAC-1 versus HDAC-2, HDAC-2 versus HDAC-3 and HDAC-1 versus HDAC-3: P < 0.001).
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Detection of HDAC-1, -2 and -3 protein expression by Western blot analysis
HDAC-1, -2 and -3 protein expression was studied semiquantitatively by Western blot analysis (Fig. 3A–C, respectively).
-Actin expression served as loading control. Proteins bands were found at the expected size (HDAC-1: 62 kDa, HDAC-2: 60 kDa and HDAC-3: 49 kDa). The level of HDAC-1 and -3 protein expression did not vary significantly and was not obviously following hormonal changes throughout the menstrual cycle (Kruskal–Wallis tests: PHDAC-1 = 0.3226; PHDAC-3 = 0.2517). It should be emphasized that HDAC-1 protein expression was highly variable between individual women especially on day 5–8 and day 19–24 (Fig. 3A).
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HDAC-2 protein showed a trend towards higher expression in the secretory phase (Kruskal–Wallis test: PHDAC-2 = 0.0216). The Dunn's multiple comparison post hoc tests revealed significantly elevated HDAC-2 protein expression on cycle day 15–18 and day 19–24 compared with day 5–8 (P < 0.01), and on cycle day 19–24 compared with day 9–14 (P < 0.01).
HDAC-1 immunohistochemistry in human cyclic endometrium and endometrial adenocarcinomas
HDAC-1 protein expression was analyzed in 28 endometrial samples derived from day 7 to day 28 of the cycle as well as in 17 endometrial adenocarcinomas (Fig. 4). These results were confirmed with a second anti-HDAC-1 antibody (data not shown).
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The nuclei of epithelial, stromal, smooth muscle cells and endothelial cells were HDAC-1 positive (Fig. 4A–D). Furthermore, the nuclei of cells located within lymph-follicles of the basal endometrial layers expressed the HDAC-1 protein (data not shown).
Table 3 shows the quantitative results of HDAC-1 protein expression in different populations of endometrial epithelial and stromal cells. Endometrial epithelial cells covering the surface as well as the epithelial cells of the upper and basal glands did not show significant differences in the number of HDAC-1 positive cells during the menstrual cycle (Kruskal–Wallis-tests: Psurface = 0.1278, Pupper glands = 0.1785 and Pbasal glands = 0.1224). Furthermore, the percentage of HDAC-1 positive endometrial stromal cells did not vary between the proliferative and secretory phase (Table 3; Mann–Whitney U-test: P = 0.0649).
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HDAC-1 protein expression was also assessed in 17 endometrial adenocarcinomas (Table 4 and Fig. 4E and F). As in the normal endometrium, HDAC-1 immunostaining was confined to the nuclei of the various cell types (Fig. 3E and F). The percentage of HDAC-1 positive cells was found to be reduced in epithelial and stromal cells of the carcinomas compared with the normal situation (Table 4; Fig. 4E–H). Six out of the 17 tumors showed less than 10% HDAC-1 positive epithelial tumor cells in contrast to the normal endometrial epithelium with averages of 56.8–88.8% HDAC-1 positive cells depending on epithelial cell compartment and cycle phase. The stroma within 15 of the 17 carcinomas exhibited fewer HDAC-1 stained cells (<50% positive cells; Fig. 4E–H; Table 4) than the stroma of normal endometrium with an average of 81.6% and 86.9% (Fig. 4A–D, Table 3).
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In the carcinomas, there was no trend towards a correlation between the tumor grade and percentage of tumor HDAC-1 positive cells. Grade 3 adenocarcinomas were found in the group depicting >80% HDAC-1 positive tumor cells as well as in the group with <10% of HDAC-1 positive tumor cells.
HDAC-2 and -3 immunohistochemistry
The distribution of the HDAC-2 (Fig. 5A–D; Table 5) and HDAC-3 protein (Fig. 6A–D; Table 6) was studied in 20 and 30 endometrial samples, respectively.
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HDAC-2 and HDAC-3 protein expression was found exclusively in the nuclei of epithelial, stromal, endothelial and smooth muscle cells. Furthermore, prominent HDAC-3 immunostaining was detected in cells within lymph-follicles located in the basal endometrial layer (Fig. 6D).
The percentage of HDAC-2 positive epithelial cells did not change during the menstrual cycle (Table 5; Kruskal–Wallis tests: Psurface = 0.4623, Pupper glands = 0.2371 and Pbasal glands = 0.5478). Furthermore, the percentage of HDAC-2 positive stromal cells did not differ between the proliferative and secretory phase (Mann–Whitney U-test: P = 0.4848).
The percentage of HDAC-3 positive epithelial cells located in the upper and the basal parts of the glands did not change during the menstrual cycle (Kruskal–Wallis tests: Pupper glands = 0.0667; Pbasal glands = 0.0287, but all post hoc Dunn's multiple comparison test P > 0.05). However, in the surface epithelium, the percentage of HDAC-3 positive cells was significantly lower on cycle day 6–10 than on day 15–18 and 24–28 (Kruskal–Wallis tests: Psurface = 0.008; Dunn's multiple comparison tests: days 6–10 versus 15–18 and days 6–10 versus 24–28, both P < 0.01; Table 6).
In contrast, the percentage of HDAC-3 positive stromal cells did not change between the proliferative and secretory phase (Mann–Whitney U-test: P = 0.6991).
HAT mRNA expression
Since HDAC-activity is antagonized by HATs, we analyzed mRNA expression of PCAF and GCN5 in the human cyclic endometrium by semiquantitative RT–PCR (Fig. 7, Table 7). PCAF mRNA was expressed constitutively in the endometrium throughout the menstrual cycle (Kruskal–Wallis test: PPCAF = 0.1071) However, there is a obvious trend towards reduced GCN5 mRNA expression during the secretory phase (Kruskal–Wallis test: PGCN5 = 0.058). This trend reaches statistical significance, when data from the proliferative phase are pooled (3.27; 1.68–6.38; n = 12) and compared with the pooled data of the secretory phase (1.33; 0.52–5.58; n = 13) using the Mann–Whitney U-test (P = 0.0134).
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Localization of mRNA expression was assessed by performing RT–PCRs on cDNA obtained from in vitro cultured endometrial fibroblasts and epithelial cells. GCN5 as well as PCAF was expressed in endometrial epithelial cells and fibroblast supplemented with E2 alone or with E2 and medroxyprogesterone acetate (Fig. 7B).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionAcknowledgementsReferences |
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Recently, it was shown that HDAC inhibitors affect proliferation, differentiation and the progesterone response of endometrial cells and endometrial tumor cell lines (Sakai et al., 2003
; Gunin et al., 2005; Takai et al., 2006; Uchida et al., 2006). However, there is only rare information on the type of HDACs expressed in the human endometrium. Consequently, we studied for the first time systematically the temporal and spatial expression of class I HDAC-1, -2 and -3 mRNAs and proteins and the mRNA expression of two counteracting HATs, GCN5 and PCAF, in the human cyclic endometrium and its specific cell types. Furthermore, we showed for the first time HDAC-1 protein expression in endometrial adenocarcinomas. The results demonstrate that in the human endometrium HDAC-1, -2 and -3 mRNAs and proteins are expressed throughout the menstrual cycle. Although HDAC-1, -2 and -3 mRNA is expressed constitutively, the expression of the corresponding proteins show subtle variations during the cycle. There is a significant trend towards (i) higher HDAC-2 protein expression during the secretory phase, shown by Western Blot analysis and (ii) reduced HDAC-3 immunostaining in the surface epithelium on day 6–10 of the menstrual cycle. Furthermore, especially during the proliferative phase, HDAC-1 and -3 mRNA and protein levels showed considerable variations among women.
All endometrial cell types—epithelial and stromal cells as well as endothelial and smooth muscle cells of the vessel wall—showed nuclear HDAC-1, -2 and -3 protein expression. However, the number of HDAC-2 positive stromal cells was lower than the number of HDAC-1 and -3 positive stromal cells. Further studies are needed to assess the differentiation status of endometrial cells that do not express certain HDAC proteins and the functional relevance. In summary, more or less all endometrial cell types are potential targets of HDAC inhibitors. Recently, it became evident that the anticonvulsant drug valproic acid is an inhibitor of class I and II HDACs (Phiel et al., 2001; Hess-Stumpp, 2005). On the background of the presented data, it has to be considered that valproic acid, which affects fertility at the endocrine level (Artama et al., 2006), may also exert direct effects on endometrial function.
Disturbing the HDAC/HATs balance alters the status of histone acetylation and consequently global and local chromatin structure. These epigenetic changes finally lead to the impairment of specific gene expression and cell function (Thiagalingiam et al., 2003). Since HDACs and HATs are members of steroid hormone receptor complexes, they are involved in the steroid hormone mediated gene expression. Therefore, it might be speculated that decreased or elevated endometrial HDAC-1, -2 and -3 protein expression may alter endometrial hormone responsiveness and secretory transformation. Furthermore, we hypothesize that a disturbed HDAC/HAT ratio is involved in the endometrial maturation defect called ‘progesterone resistance’. Affected women showed adequate phase specific E2 and progesterone serum levels and normal endometrial steroid hormone receptor expression, however, the endometrium remains flat and does not show secretory transformation (Alfer et al., 2000).
The observed trends towards elevated HDAC-2 protein expression in parallel with reduced GCN5 mRNA expression possibly shifts the balance between histone deacetylation and acetylation towards deacetylation which may lead to the repression of gene expression. Further studies are needed to explore the histone acetylation status of endometrial cells during the menstrual cycle as well as the impact of histone acetylation status on endometrial function.
The dominant role of HDAC-1 in cell cycle control (Lagger et al., 2002; Zhu et al., 2004) and the fact that HDAC-1 expression is a marker for higher differentiation in breast cancer (Krusche et al., 2005) prompted us to analyze HDAC-1 expression in endometrioid adenocarcinomas. We found that 35% of endometrial adenocarcinomas showed a massive decrease or a complete loss of epithelial HDAC-1 protein expression. Furthermore, in nearly 90% of the assessed adenocarcinomas, the stroma depicted a reduced number of HDAC-1 stained cells compared with the stroma of the normal cyclic endometrium. This may be indicative of an impaired epigenetic status of epithelial and stromal cells within these tumors. We proposed that the variability of HDAC-1 expression leads to different sensitivity and responsiveness of tumors to HDAC inhibitors, which has to be taken into account during therapeutic HDAC inhibitor administration and this issue needs further assessment.
Taken together, we have shown that class I HDACs-1, -2 and -3 are expressed throughout the menstrual cycle in the human endometrium, suggesting the endometrium as a potential target for HDAC inhibitors. In addition, we have shown altered expression of HDAC-1 in endometrial adenocarcinomas.
Further studies are needed to determine which genes are regulated by HDACs in the different endometrial cells to explore if the application of HDAC inhibitors represents an option for the treatment of endometrial disorders, e.g. the lack of hormone response, endometriosis or cancer.
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We thank Dr. Nicole Heussen (Institute of Medical Statistics, RWTH Aachen University) for supporting the statistical assessment. We thank Bärbel Bonn, Sabine Eisner, Sabina Hennes-Mades and Diana Seelis-Schmidt for excellent technical assistance. This work was supported by the START program ‘Molekulare Endokrinologie’ (TP6, 1/2000–12/2003) of the School of Medicine, RWTH University of Aachen and by a grant of the Schering AG, Berlin, Germany.
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|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionAcknowledgementsReferences |
|---|
Alfer J, Muller-Schottle F, Classen-Linke I, von Rango U, Happel L, Beier-Hellwig K, Rath W, Beier HM. The endometrium as a novel target for leptin: differences in fertility and subfertility. Mol Hum Reprod (2000) 6:595–601.
Allfrey V G, Faulkner R, Mirsky A E. Acetylation and methylation of histones and their possible role in the regulation of RNA synthesis. Proc Natl Acad Sci USA (1964) 51:786–794.
Artama M, Isojärvi JIT, Auvinen A. Antiepileptic drug use and birth rate in patients with epilepsy—a population-based cohort study in Finland. Hum Reprod (2006) 21:2290–2295.
Bradbury CA, Khanim FL, Hayden R, Bunce CM, White DA, Drayson MT, Craddock C, Turner BM. Histone deacetylases in acute myeloid leukaemia show a distinctive pattern of expression that changes selectively in response to deacetylase inhibitors. Leukemia (2005) 19:751–759.
Classen-Linke I, Kusche M, Knauthe R, Beier HM. Establishment of a human endometrial cell culture system and characterization of its polarized hormone responsive epithelial cells. Cell Tissue Res (1997) 287:171–185.[CrossRef][Web of Science][Medline]
Giangrande PH, Kimbrel EA, Edwards DP, McDonnell DP. The opposing transcriptional activities of the two isoforms of the human progesterone receptor are due to differential cofactor binding. Mol Cell Biol (2000) 20:3102–3115.
Glozak MA, Sengupta N, Zhang X, Seto E. Acetylation and deacetylation of non-histone proteins. Gene (2005) 363:15–23.[CrossRef][Web of Science][Medline]
Grozinger CM, Schreiber SL. Deacetylase enzymes: biological functions and the use of small-molecule inhibitors. Chem Biol (2002) 9:3–16.[CrossRef][Web of Science][Medline]
Guenther MG, Yu J, Kao GD, Yen TJ, Lazar MA. Assembly of the SMRT-histone deacetylase 3 repression complex requires the TCP-1 ring complex. Genes Dev (2002) 16:3130–3135.
Gunin AG, Kapitova IN, Suslonova NV. Effects of histone deacetylase inhibitors on estradiol-induced proliferation and hyperplasia formation in the mouse uterus. J Endocrinol (2005) 185:539–549.
Hess-Stumpp H. Histone deacetylase inhibitors and cancer: from cell biology to the clinic. Eur J Cell Biol (2005) 84:109–121.[CrossRef][Web of Science][Medline]
Hubbert C, Guardiola A, Shao R, Kawaguchi Y, Ito A, Nixon A, Yoshida M, Wang XF, Yao TP. HDAC6 is a microtubule-associated deacetylase. Nature (2002) 417:455–458.[CrossRef][Medline]
Joo EJ, Chung JJ, Choe SS, Kim KH, Kim JB. Down-regulation of histone deacetylases stimulates adipocyte differentiation. J Biol Chem (2006) 281:6608–66015.
Krusche CA, Wulfing P, Kersting C, Vloet A, Bocker W, Kiesel L, Beier HM, Alfer J. Histone deacetylase-1 and -3 protein expression in human breast cancer: a tissue microarray analysis. Breast Cancer Res Treat (2005) 90:15–23.[CrossRef][Web of Science][Medline]
Laemmli UK. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature (1970) 227:680–685.[CrossRef][Medline]
Lagger G, O'Carroll D, Rembold M, Khier H, Tischler J, Weitzer G, Schuettengruber B, Hauser C, Brunmeir R, Jenuwein T, et al. Essential function of histone deacetylase 1 in proliferation control and CDK inhibitor repression. EMBO J (2002) 3:2672–2681.
Langley E, Pearson M, Faretta M, Bauer UM, Frye RA, Minucci S, Pelicci PG, Kouzarides T. Human SIR2 deacetylates p53 and antagonizes PML/p53-induced cellular senescence. EMBO J (2002) 21:2383–2396.[CrossRef][Web of Science][Medline]
de Leval L, Waltregny D, Boniver J, Young RH, Castronovo V, Oliva E. Use of histone deacetylase 8 (HDAC8), a new marker of smooth muscle differentiation, in the classification of mesenchymal tumors of the uterus. Am J Surg Pathol (2006) 30:319–327.[Web of Science][Medline]
Liu XF, Bagchi MK. Recruitment of distinct chromatin-modifying complexes by tamoxifen-complexed estrogen receptor at natural target gene promoters in vivo. J Biol Chem (2004) 279:15050–15058.
Marks PA, Richon VM, Breslow R, Rifkind RA. Histone deacetylase inhibitors as new cancer drugs. Curr Opin Oncol (2001) 13:477–483.[CrossRef][Web of Science][Medline]
Minucci S, Pelicci PG. Histone deacetylase inhibitors and the promise of epigenetic (and more) treatments for cancer. Nat Rev Cancer (2006) 6:38–51.[CrossRef][Web of Science][Medline]
Momparler RL. Cancer epigenetics. Oncogene (2003) 22:6479–6483.[CrossRef][Web of Science][Medline]
Noyes RW, Hertig AT, Rock J. Dating the endometrial biopsy. Fertil Steril (1950) 1:3–25.[Medline]
Phiel CJ, Zhang F, Huang EY, Guenther MG, Lazar MA, Klein PS. Histone deacetylase is a direct target of valproic acid, a potent anticonvulsant, mood stabilizer, and teratogen. J Biol Chem (2001) 276:36734–36741.
Phillips RJ, Tyson-Capper Nee Pollard AJ, Bailey J, Robson SC, Europe-Finner GN. Regulation of expression of the chorionic gonadotropin/luteinizing hormone receptor gene in the human myometrium: involvement of specificity protein-1 (Sp1), Sp3, Sp4, Sp-like proteins, and histone deacetylases. J Clin Endocrinol Metab (2005) 90:3479–3490.
Sakai N, Maruyama T, Sakurai R, Masuda H, Yamamoto Y, Shimizu A, Kishi I, Asada H, Yamagoe S, Yoshimura Y. Involvement of histone acetylation in ovarian steroid-induced decidualization of human endometrial stromal cells. J Biol Chem (2003) 278:16675–16682.
Santos-Rosa H, Caldas C. Chromatin modifier enzymes, the histone code and cancer. Eur J Cancer (2005) 41:2381–2402.[CrossRef][Web of Science][Medline]
Sengupta N, Seto E. Regulation of histone deacetylase activities. J Cell Biochem (2004) 93:57–67.[CrossRef][Web of Science][Medline]
Shang Y, Brown M. Molecular determinants for the tissue specificity of SERMs. Science (2002) 295:2465–2468.
Strahl BD, Allis CD. The language of covalent histone modifications. Nature (2000) 403:41–45.[CrossRef][Medline]
Takai N, Desmond JC, Kumagai T, Gui D, Said JW, Whittaker S, Miyakawa I, Koeffler HP. Histone deacetylase inhibitors have a profound antigrowth activity in endometrial cancer cells. Clin Cancer Res (2004) 10:1141–1149.
Takai N, Ueda T, Nishida M, Nasu K, Narahara H. A novel histone deacetylase inhibitor, Scriptaid, induces growth inhibition, cell cycle arrest and apoptosis in human endometrial cancer and ovarian cancer cells. Int J Mol Med (2006) 17:323–329.[Web of Science][Medline]
Thiagalingam S, Cheng KH, Lee HJ, Mineva N, Thiagalingam A, Ponte JF. Histone deacetylases: unique players in shaping the epigenetic histone code. Ann N Y Acad Sci (2003) 983:84–100.[Web of Science][Medline]
Uchida H, Maruyama T, Nagashima T, Asada H, Yoshimura Y. Histone deacetylase inhibitors induce differentiation of human endometrial adenocarcinoma cells through up-regulation of glycodelin. Endocrinology (2005) 146:5365–5373.
Uchida H, Maruyama T, Nagashima T, Ono M, Masuda H, Arase T, Sugiura I, Onouchi M, Kajitani T, Asada H, et al. Human endometrial cytodifferentiation by histone deacetylase inhibitors. Hum Cell (2006) 19:38–42.[CrossRef][Medline]
Vigushin DM, Ali S, Pace PE, Mirsaidi N, Ito K, Adcock I, Coombes RC. Trichostatin A is a histone deacetylase inhibitor with potent antitumor activity against breast cancer in vivo. Clin Cancer Res (2001) 7:971–976.
Waltregny D, De Leval L, Glenisson W, Ly Tran S, North BJ, Bellahcene A, Weidle U, Verdin E, Castronovo V. Expression of histone deacetylase 8, a class I histone deacetylase, is restricted to cells showing smooth muscle differentiation in normal human tissues. Am J Pathol (2004) 165:553–564.
Xiong Y, Dowdy SC, Podratz KC, Jin F, Attewell JR, Eberhardt NL, Jiang SW. Histone deacetylase inhibitors decrease DNA methyltransferase-3B messenger RNA stability and down-regulate de novo DNA methyltransferase activity in human endometrial cells. Cancer Res (2005) 65:2684–2689.
Zhu P, Huber E, Kiefer F, Gottlicher M. Specific and redundant functions of histone deacetylases in regulation of cell cycle and apoptosis. Cell Cycle (2004) 3:1240–1242.[Web of Science][Medline]
Submitted on October 17, 2006; resubmitted on June 22, 2007; accepted on June 28, 2007.
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