A cross-species analysis of the rodent uterotrophic program: elucidation of conserved responses and targets of estrogen signaling

Joshua C. Kwekel1,4, Lyle D. Burgoon2,4, Jeremy W. Burt1,4, Jack R. Harkema3,4 and Timothy R. Zacharewski1,4

1 Departments of Biochemistry and Molecular Biology, Michigan State University, East Lansing, Michigan
2 Departments of Pharmacology and Toxicology, Michigan State University, East Lansing, Michigan
3 Departments of Pathobiology and Diagnostic Investigation, Michigan State University, East Lansing, Michigan
4 Center for Integrative Toxicology and National Food Safety and Toxicology Center, Michigan State University, East Lansing, Michigan


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Physiological, morphological, and transcriptional alterations elicited by ethynyl estradiol in the uteri of Sprague-Dawley rats and C57BL/6 mice were assessed using comparable study designs, microarray platforms, and analysis methods to identify conserved estrogen signaling networks. Comparative analysis identified 153 orthologous gene pairs that were positively correlated, suggesting conserved transcriptional targets important in uterine proliferation. Functional annotation for these responses were associated with angiogenesis, water and solute transport, cell cycle control, redox control, DNA replication, protein synthesis and transport, xenobiotic metabolism, cell-cell communication, energetics, and cholesterol and fatty acid regulation. The identification of conserved temporal expression patterns of these orthologs provides experimental support for the transfer of functional annotation from mouse orthologs to 44 previously unannotated rat expressed sequence tags based on their homology and co-expression patterns. The identification of comparable temporal phenotypic responses linked to related gene expression profiles demonstrates the ability of systematic comparative genomic assessments to elucidate important conserved mechanisms in rodent estrogen signaling during uterine proliferation.

uterotrophic assay; microarray; species comparison; ethynyl estradiol


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
RENEWED INVESTIGATIONS into the modes of estrogen action have emerged to improve the efficacy of selective estrogen receptor modulators (SERMs), a structurally diverse group of drugs, natural products, industrial chemicals, and environmental contaminants that elicit tissue-specific agonist and antagonist responses. Concerns regarding the potential adverse health effects have also resulted in a comprehensive screening program to assess commerce chemicals and environmental contaminants for their potential to inappropriately activate or antagonize normal estrogen receptor (ER) function (31, 49). SERMs typically induce a subset of responsive genes but produce little to no uterotrophic response, while others elicit a strong uterotrophic response but fail to induce the full gene expression spectrum in target tissues (13, 60, 70). This variability warrants further investigation into the tissue-specific transcriptional program of SERMs in relation to their cellular and physiological endpoints to assess the potential adverse effects of exogenous estrogenic compounds and to further elucidate the mechanisms involved in SERM activities. In this study, the rodent uterine response to ethynyl estradiol (EE), a prototypic oral estrogen, is assessed to identify conserved responses and targets of estrogen signaling for further studies of estrogenic compounds.

The rodent uterotrophic assay is the "gold standard" in vivo assay for assessing estrogenicity (14, 29, 48). While it provides a robust physiological endpoint, it lacks utility in further elucidating the mode of action of diverse estrogenic compounds. Estrogens primarily exert their effects via binding and activation of ERs, which function as ligand-dependant transcription factors. Activated ER complexes recruit co-activators or co-repressors to chromatin, leading to the transcriptional modulation of responsive genes (30). Studies have demonstrated that transcriptional responses can vary depending on the ligand structure, which confers differential receptor complex conformations and transactivation activities (23, 45).

The enhanced uterotrophic assay (14, 15) incorporates histological and transcriptional evaluations to complement the uterotrophic endpoint. The current study extends this approach by comparatively examining global gene expression and histological and morphological responses in Sprague-Dawley rats and C57BL/6 mice in comprehensive time-course studies to identify the conserved transcriptional targets critical to the observed molecular and physiological responses to EE. Temporally conserved and divergent transcriptional responses that are phenotypically anchored to histological and morphometric endpoints were identified, providing new insights into the conserved modes of action involved in rodent uterine hypertrophy and hyperplasia.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Husbandry.
Female Sprague-Dawley rats and C57BL/6 mice, ovariectomized on postnatal day 20 and all within 10% of the average body weight, were obtained from Charles River Laboratories (Raleigh, NC) on postnatal day 25. Animals were housed in polycarbonate cages containing cellulose fiber chip bedding (Aspen Chip Laboratory Bedding; Northeastern Products, Warrensberg, NY) and maintained at 40–60% humidity and 23°C on a 12:12-h dark-light cycle (7 AM to 7 PM). Animals were fed deionized water and Harlan Teklad 22/5 Rodent Diet 8640 ad libitum (Madison, WI) and acclimatized for 4 days before treatment. Immature, ovariectomized rodent uteri were examined, since they are more sensitive to estrogen exposure compared with the adult ovariectomized animals (51).

Treatments.
Animals were treated once or once daily for 3 days via oral gavage with 100 µg/kg body wt 17{alpha}-EE in 0.1 ml of sesame oil vehicle (Sigma Chemical, St. Louis, MO) (Fig. 1). This dose was empirically derived, as it elicits a maximal uterotrophic response through the oral route while showing no acute toxic effects. Animals (n = 5) receiving a single dose of vehicle or EE were euthanized 2, 4, 8, 12, 18, or 24 h after treatment. Animals receiving one daily dose on 3 consecutive days were euthanized 72 h after initial dosing, as per the uterotrophic assay. Doses of EE were calculated based on average weights of the animals before dosing.



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Fig. 1. Time-course experimental design. A comprehensive in vivo time-course study was performed in which immature ovariectomized Sprague-Dawley rats and C57BL/6 mice were administered oral doses of 100 µg/kg body wt (bw) ethynyl estradiol (EE) or sesame oil vehicle followed by euthanasia and tissue harvest at 2, 4, 8, 12, 18, and 24 h after dosing. Another group of animals received a single dose, once per day on 3 consecutive days followed by euthanasia and tissue harvest 72 h after the initial dose per the uterotrophic assay.

 
Independent histopathology studies using the same study design (n = 4) were identically performed for each species, with the exception that animals received an intraperitoneal (ip) injection of 50 mg/kg body wt 5-bromo-2-deoxyuridine (BrdU) (Sigma) 2 h before euthanasia. All procedures were performed with the approval of the Michigan State University All-University Committee on Animal Use and Care.

Necropsy.
Animals were euthanized by cervical dislocation, and animal body weights were recorded. The uterine body was dissected at the border of the cervix, and whole uteri were harvested and stripped of extraneous connective tissue and fat. Whole uterine weights were recorded before (wet) and after (blotted) being blotted under pressure with absorbent tissue and were subsequently snap frozen in liquid nitrogen and stored at –80°C. Weight due to water was calculated as the difference between the wet and blotted weights. Necropsy for the histopathology study was performed identically to the gene expression time-course study with the exception that tissues were not blotted and were placed in tubes containing 1 ml of 10% neutral buffered formalin (NBF; VWR, West Chester, PA) and stored at room temperature for at least 24 h before further processing. Statistical analyses of wet weight and water content were conducted using a two-way ANOVA with a Tukey's honestly significant difference (HSD) post hoc test (n = 5, P < 0.05; SAS 9.1, Cary, NC).

RNA isolation.
Total RNA was isolated from whole uteri using Trizol reagent (Invitrogen, Carlsbad, CA) as per the manufacturer's protocol. Uteri were removed from –80°C storage and immediately homogenized in 1 ml of Trizol reagent using a Mixer Mill 300 tissue homogenizer (Retsch, Germany). Total RNA was resuspended in RNA storage solution (Ambion, Austin, TX). RNA concentrations were calculated by spectrophotometric methods (A260, where A is absorption at 260 nm), and purity was assessed by the A260-to-A280 ratio and by visual inspection of 1 µg on a denaturing gel.

Histological processing.
Uteri fixed for 24 h in NBF were dissected and, 6- to 8-mm midhorn sections were embedded in paraffin according to standard histological techniques. Five-micrometer cross sections containing both uterine horns were cut and mounted on glass slides and stained with hematoxylin and eosin. Additionally, a serial section was cut, mounted, and stained with anti-BrdU antibody (BD Biosciences, Palo Alto, CA; Vector Substrate Kit 1, Vector Red, Vector Laboratories, Burlingame, CA) and counterstained with hematoxylin. All embedding, mounting, and staining of tissues were performed at the Histology/Immunohistochemistry Laboratory, Michigan State University (http://humanpathology.msu.edu/histology/index.html).

Histopathological and morphometric assessment.
Histological slides were evaluated according to standardized National Toxicology Program (NTP) pathology codes. Morphometric analyses were performed on midhorn cross sections of both uterine horns for each animal using image analysis software (Scion Image; Scioncorp, Frederick, MD) and standard morphometric techniques. Briefly, the length of basal lamina underlying the luminal epithelium (LE) and corresponding areas of LE, stroma, and myometrium were quantified for multiple representative sectors of each section. Total luminal and glandular circumferences were also quantified. Anti-BrdU labeling indexes were quantified for LE cell height and stromal compartments on a per-cell, per-area (mm2) basis, respectively. Morphometric analyses for the mouse were limited to LE cell height, stromal thickness, and LE BrdU labeling indexes, as these parameters capture the most sensitive histological endpoints in the uterotrophic assay. Statistical analyses on all morphometry data were performed using a two-way ANOVA with a Tukey's HSD post hoc test (n = 4, P < 0.05; SAS 9.1).

Quantitative real-time PCR analysis.
For each sample, 1.0 µg of total RNA was reverse transcribed by SuperScript II using an anchored oligo(dT) primer as described by the manufacturer (Invitrogen). The resultant cDNA (1.0 µl) was used as the template in a 30-µl PCR reaction containing 0.1 µM each of forward and reverse gene-specific primers designed using Primer3 (54), 3 mM MgCl2, 1.0 mM dNTPs, 0.025 IU AmpliTaq Gold and 1 x SYBR Green PCR buffer (Applied Biosystems, Foster City, CA). Gene names, accession numbers, forward and reverse primer sequences, and amplicon sizes are listed in Supplemental Table S1 (available at the Physiological Genomics web site).1 PCR amplification was conducted in MicroAmp Optical 96-well reaction plates (Applied Biosystems) on an Applied Biosystems PRISM 7000 Sequence Detection System using the following conditions: initial denaturation and enzyme activation for 10 min at 95°C followed by 40 cycles of 95°C for 15 s and 60°C for 1 min. A dissociation protocol was performed to assess the specificity of the primers and the uniformity of the PCR-generated products. Each plate contained duplicate standards of purified PCR products of known template concentration covering six orders of magnitude to interpolate relative template concentrations of the samples from the standard curves of log copy number vs. threshold cycle (Ct). No template controls (NTC) were also included on each plate. Samples with a Ct value within 2 SD of the mean Ct values for the NTCs were considered below the limits of detection. The copy number of each unknown sample for each gene was standardized to the geometric mean of two housekeeping genes (IA and Rpl7) to control for differences in RNA loading, quality, and cDNA synthesis. Statistical significance of differentially expressed genes was determined using two-way ANOVA followed by t-test for vehicle treatment comparisons (SAS 9.1). For graphing purposes, the relative expression levels were scaled such that the expression level of the time-matched control group was equal to 1. Correlations of microarray to real-time data were performed using a Pearson's correlation of fold changes relative to vehicle controls (VCs; R Statistical Package 1.9.1) on a per-time-point basis.

Array platform.
Spotted rat cDNA microarrays were produced in-house from the Lion Bioscience's Rat cDNA library (Lion Bioscience, Heidelberg, Germany) consisting of 8,567 clones representing 3,022 unique genes (Unigene build no. 48) that were selected based on their level of annotation as well as sequence similarity to well annotated human and mouse genes. Mouse cDNA arrays of the same platform and construction, consisted of 13,361 features, representing 7,948 unique genes containing clones from multiple sources including the National Institute on Aging (NIA), Environmental Protection Agency (EPA), IMAGE Consortium, and Lion Biosciences. Detailed protocols for microarray construction, labeling of the cDNA probe, sample hybridization, and slide washing can be found at http://dbzach.fst.msu.edu/interfaces/microarray.html. Briefly, PCR-amplified DNA was robotically arrayed onto epoxy-coated glass slides [Nexterion (previously Quantifoil), Jena, Germany], using an Omnigrid arrayer (GeneMachines, San Carlos, CA) equipped with 32 (8 x 4) Chipmaker 2 pins (Telechem) at the Genomics Technology Support Facility at Michigan State University (http://www.genomics.msu.edu).

Array experimental design and protocols.
Temporal changes in gene expression of EE-treated rat and mouse uteri were assessed using an independent reference design in which samples from estrogen-treated animals are co-hybridized with VCs. Comparisons were performed on three biological replicates x2 independent labelings of each sample (incorporating a dye swap) for each time point. For the rat study, total RNA (15 µg) was reverse transcribed in the presence of cysteine-3 (Cy3)- or Cy5-labeled dUTP (Amersham, Piscataway, NJ) to create fluor-labeled cDNA, which was purified using QIAquick PCR purification kit (Qiagen, Valencia, CA). Cy3- and Cy5-labeled samples were mixed, vacuum dried, and resuspended in 32 µl of hybridization buffer (40% formamide, 4x SSC, 1% SDS) with 15 µg of polydA and 15 µg of mouse COT-1 DNA (Invitrogen) as a competitor. This probe mixture was heated at 95°C for 2 min and was then hybridized to the array under a 22 x 40-mm LifterSlip coverslip (Erie Scientific, Portsmouth, NH) in a light-protected and humidified hybridization chamber (Corning, Corning, NY). Samples were hybridized for 18–24 h at 42°C in a water bath. Slides were then washed, dried by centrifugation, and scanned at 635 (Cy5) and 532 nm (Cy3) on an Affymetrix 428 Array Scanner (Santa Clara, CA). Images were analyzed for feature and background intensities using GenePix Pro 3.0 (Axon Instruments, Union City, CA). Because of the limited amount of RNA isolated from mouse uteri (~8 µg/mouse), a 3DNA Array 900 Expression Array Detection Kit (Genisphere, Hatsfield, PA), using 1 µg of total RNA and according to manufacturer's specifications, was used for probe labeling in the mouse microarray experiments. It is well documented that replicate arrays of identical samples within the same array-labeling protocol or platform (i.e., direct label) render correlation coefficients of ~0.85–0.95; this lends confidence in the comparison of array data utilizing direct labeling to those following the Genisphere protocol due to correlations between the two platforms which fall in this same range (http://www.genisphere.com/array_detection_900.html).

Array data normalization and statistical analysis.
Data were normalized using a semiparametric approach (17). Model-based t values were calculated from normalized data, comparing treated from vehicle responses per time point. Empirical Bayes analysis was used to calculate posterior probabilities of activity [P1(t) value] on a per-gene and -time-point basis using the model-based t value (16). Gene lists were filtered for activity based on the P1(t) value, which indicates increasing activity as the value approaches 1.0. A conservative P1(t) cutoff of 0.999999 was used to filter the expression data and to define active gene lists for both the rat and mouse data sets. All arrays in both studies were subjected to quality control assessment to ensure assay performance and data consistency through the study, and stored within dbZach (http://dbzach.fst.msu.edu), a minimum information about a microarray experiment (MIAME)-supportive relational database that ensures proper data management and facilitates data analysis. Complete data sets with annotation and P1(t) values are available in Supplemental Tables S2 and S3.

Gene annotation, clustering, and interpretation.
Features refer to cDNA clones spotted on the array and are assigned a GenBank accession number, gene name, symbol, and LocusLink ID where annotation is available. Gene names of well annotated human or mouse orthologs were adopted for rat clones that have limited or no annotation and where sequence similarity (Bl2seq) according to a given E-value cutoff (10–30) suggests orthology. For brevity and consistency, genes are referenced by their official gene symbol as defined by the National Center for Biotechnology Information (NCBI) Entrez Gene. A full listing of all abbreviated genes with their full names and LocusLink identifiers can be found in Supplemental Tables S2 and S3. Expression data meeting the initial P1(t) cutoff were grouped using a k-means cluster algorithm in GeneSpring 7.1 (Silicon Genetics, Redwood City, CA).


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Because previous gene expression studies (19, 39, 65, 66) have focused on murine responses to estrogen, the focus for this study was placed on the rat.

Uterine wet weights and water content.
A sixfold induction in rat uterine wet weight relative to body weight was observed at 72 h (Fig. 2A), consistent with previous reports (35). A 10-fold increase in water content accounted for ~31% of total uterine wet weight at 72 h. This response was preceded by early increases in wet weight (3.3-fold) due to water imbibition (3, 26) between 4 and 12 h which subsided by 18 h (Fig. 2B). Eotaxin is the estrogen-sensitive chemotactic factor responsible for recruiting eosinophils to the uterine epithelia and stroma where they mediate the edematous response at 6 h (22). The water imbibition response provides an endpoint for estrogen treatment that is not correlated with downstream uterotrophic effects (52) and can therefore serve as a sensitive marker of immediate early gene regulation not causative of proliferation.



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Fig. 2. Increases in uterine wet weight and water content due to EE. Changes elicited in uterine wet weight (A) and water content (B) after oral exposure to 100 µg/kg body wt EE. Changes in uterine wet and blotted weights were recorded at each time point for EE and vehicle control (VC; n = 5) animals. Water content calculated as the difference between the wet and blotted weights was calculated at each time point for EE and vehicle-treated animals. Tissue weights were normalized to body weight for each animal, and average weights per group are indicated at each time point. Error bars represent the SE for the average fold change. *P < 0.05.

 
A comparable uterotrophic response was observed in the mouse with a sevenfold induction in uterine wet weight. However, the water imbibition response was masked or temporally shifted, as increases in weight due to water were not significantly different in EE-treated tissues until 12 h (Fig. 2). Increased water evaporation from mouse uteri during tissue harvest under the dissection microscope may have contributed to this result due to their small size relative to the rat uterus.

Histopathology.
Several histological parameters were modulated by EE at multiple time points (Fig. 3) in both species. Indications of stromal edema (4–18 h) and eosinophil infiltration (8, 12, 24, and 72 h) were evident in both species, consistent with previous reports linking eosinophilia with edema 6 h after treatment in the rat uterus (24, 37, 58). Evidence of both LE cell hypertrophy (increases in cell height) at 24 and 72 h and stromal and LE cell hyperplasia (numbers of mitotic bodies) at 18 and 24 h in EE-treated slides indicates diverse and large-scale cell type-specific proliferation that culminates in the uterotrophic response at 72 h. The marked presence of apoptotic bodies were also observed in stromal, glandular epithelial (GE) cell, and LE cell compartments (72 h), consistent with the marked decrease in uterine weights in rats after 72 h (data not shown). The mouse and rat exhibited similar uterine histology and temporal severity with the exception that endometrial invaginations or ruffling were more pronounced in the mouse at the 72-h time point compared with the rat.



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Fig. 3. Alterations in rat uterine histopathology due to EE. Rat uterine midhorn cross sections (5 µm) were mounted and stained with hematoxylin and eosin. Stromal edema was evident between 4 and 12 h in EE-treated samples. Normal and edematous stromal compartments of vehicle- and EE-treated samples, respectively, are depicted with arrows (A) from the 8-h time point. A second section of each block was also cut and immunohistochemically stained with anti-5-bromo-2-deoxyuridine (anti-BrdU) (red nuclear stain) and hematoxylin. Samples from the 18- and 24-h (B) EE-treated group show marked BrdU staining in the luminal epithelium (LE), stroma, and glandular epithelium (GE) compared with VC samples. Luminal epithelial (LE) cell height, a sensitive morphological marker of estrogen exposure, exhibited a 2.7-fold induction in EE-treated samples (C) compared with VCs at 72 h. Bar = 100 µm.

 
Morphometry.
LE cell height (LECH) is a classical marker of estrogen exposure (46, 47) and has been used to demonstrate the estrogenicity of a number of structurally diverse ligands (42, 44, 46, 47). LECH was significantly induced at 24 and 72 h in both rat (1.6- and 2.7-fold) and mouse (1.4- and 2.1-fold) EE-treated samples (Fig. 4A). Stromal and myometrial thickening are necessary structural modifications and may prove to be equally informative and indicative of estrogen exposure (46). The response of the stromal compartment (Fig. 4B) showed moderate increases in overall thickness at early and late time points compared with VCs. Changes in total luminal circumference (data not shown) loosely paralleled wet weight and LECH changes. In contrast, changes in total glandular circumference (data not shown) were not observed until 72 h. Glandular epithelia are responsible for secretions produced and deposited into the lumen during normal uterine cycling, thus preparing the uterus for pregnancy after implantation (6, 56). Increases in uterine gland circumference at 72 h therefore indicate a developmental and homeostatic role of estrogen in facilitating this response.



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Fig. 4. Morphometric analysis of EE-induced changes in uterine cellular and compartmental structures. Morphometric methods were used to quantify morphological and immunohistochemical indexes of EE response in rat uterine midhorn histological cross sections. Average LE cell height (A) and stromal thickness (B) were calculated for each animal. Averages for each treatment group and time point are indicated for EE and vehicle samples. Immunohistochemical staining for BrdU incorporation into LE (C) was performed on EE- and vehicle-treated samples at each time point, and average percentages of labeled nuclei were calculated. Error bars represent the SE for the average fold change. *P < 0.05 (n = 4).

 
The LE hypertrophic response observed at 24 and 72 h (Fig. 4A) is consistent with the hyperplastic growth at 18 and 24 h as demonstrated by increased anti-BrdU labeling (Fig. 4C). BrdU labeling in VC luminal epithelium exhibited basal level staining of ~4% labeled nuclei per total nuclei in rats and was increased in treated samples to 15% and 34% at 18 and 24 h, respectively. Minimal, nonsignificant staining was observed at all other time points. BrdU labeling in the stromal compartment (data not shown) was strongly correlated with LE cell (LEC) staining and exhibited a 6- and 15-fold induction in BrdU-labeled cells per area at 18 and 24 h, respectively.

Mouse morphometric parameters of LECH, stromal thickness, and LE BrdU incorporation showed similar responses compared with the rat (Fig. 4). LECH showed significant induction of ~1.35-fold at 18 and 24 h and 1.85-fold at 72 h, respectively, whereas stromal thickness showed less reproducible (5, 55, 74) inductions at multiple time points. LECs exhibited 26% and 24% labeling with BrdU at 18 and 24 h, respectively, while minimal staining was observed at other time points relative to baseline vehicle levels.

Microarray data filtering and clustering.
Empirical Bayes analysis identified 2,652 features representing 1,116 unique genes that were active at one or more time points in the rat study. Approximately 51% of the active features (~570 genes) were active between 8 and 24 h (Fig. 5), indicating that the greatest transcriptional activity begins 12–16 h before peak mitotic activity, as indicated in the histopathology and BrdU staining (Fig. 4C). After the 8- to 24-h time points, 4 h was the next most active time point with ~409 significant changes in expression. In the mouse study, 3,102 features representing 2,313 unique genes were found to comprise the active set. The 1,116 active rat genes obtained from Empirical Bayes analysis were used for clustering and for the elucidation of functional pathways affected by EE. k-means clustering revealed six distinct clusters that distinguished up from down, early from late, and transient from sustained changes in expression (Fig. 6). The mouse active gene set was clustered in the same manner, resulting in six comparable clusters (data not shown).



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Fig. 5. Number of active genes per time point. The distribution of active genes [P1(t) > 0.999999] per time point for rat and mouse indicates the most transcriptional activity between 8 and 24 h. P1(t), posterior probability of activity.

 


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Fig. 6. k-means clustering of the rat active gene list. Gene expression data from the 2,844 features comprising the rat active gene list [P1(t) > 0.999999] were best represented by 6 k-means clusters consisting of up immediate early (A), up middle (B), down early (C), up sustained (D), up late (E) and down sustained (F) patterns of expression. Time and fold change are indicated on the x- and y-axes, respectively. The number of features in each cluster is indicated. A black pseudoline representing the general pattern of expression has been superimposed on each cluster.

 
Quantitative RT-PCR verification of microarray results.
Twenty active rat genes identified in the microarray analysis were selected for verification by quantitative RT-PCR (qRT-PCR) analysis (Fig. 7). Genes were selected to represent a majority of both the temporal patterns identified by k-means clustering and their respective functional categories in addition to their robust response. Patterns of temporal expression between the array and qRT-PCR data were correlated using a Pearson's correlation on the fold changes relative to VCs. An average correlation coefficient of 0.83 (range, 0.57–0.96; Stat3 correlation of 0.059 excluded) indicated good agreement between the two methods. Stat3 had a weaker correlation, with maximal upregulation of 3.6-fold at 2 h in the microarray data, while the qRT-PCR data indicated maximal induction of 7-fold at 4 h. Although the reasons for this disparity between the methods are unclear, both are consistent with the role Stat3 plays in immediate early expression in response to growth stimulus in proliferating tissues (36). Compression of microarray expression data, compared with other more sensitive measures of mRNA expression such as qRT-PCR, has been previously documented (73) and was observed with C3, Calb3, Aqp8, and Tk1. Two classically estrogen-induced and -repressed genes, Egr1 and Clu, respectively (8, 11, 15, 69), that were not represented on the array were also analyzed by qRT-PCR. Egr1 was significantly upregulated 7- and 21-fold at 2 and 4 h, respectively (data not shown). Clu was repressed 14-fold at 72 h.



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Fig. 7. Quantitative real-time PCR (qRT-PCR) verification of microarray results. qRT-PCR was used to verify the temporal gene expression profiles of individual genes selected from microarray data representative of each k-means cluster as well as specific functional categories. Bars (left axis) and lines (right axis where present) represent data obtained by qRT-PCR and microarray analysis, respectively, while the x-axis represents time (h). The average fold change relative to time-matched VCs for 3 animals/group is shown. Genes are indicated by their official gene symbols. Error bars represent the SE for the average fold change. *P < 0.05 for qRT-PCR.

 
Functional annotation of rat transcriptional responses.
Further analysis and interpretation of the gene expression data were facilitated by the identification and population of functional gene categories with active genes from the microarray data set. Phenotypically anchored changes in expression were tentatively assigned functions based on associations with the histopathological assessment, manual annotation obtained from the reported literature, and computationally extracted, overrepresented Gene Ontology (http://www.geneontology.org)-associated functional annotations. Categories of response evident from histopathology include edema, hyperplasia, hypertrophy, immune cell response, and apoptosis. Those identified through Gene Ontology and manual annotation include proliferation (e.g., signaling proteins, protein biosynthesis and turnover, cell cycle control, replication), angiogenesis, fatty acid metabolism, cholesterol biosynthesis, redox control, and xenobiotic metabolism (Table 1).


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Table 1. Functional categories of response

 
Comparison of EE-elicited rat and mouse uterine responses.
The development, growth, and regulation of the uterus are critical for reproduction, and thus the regulatory events affecting these processes are expected to be highly conserved among species. Comparative analysis of gene expression serves to verify genome-wide approaches to assay-conserved transcriptional responses to a common stimulus. Rat and mouse orthologous gene pairs were derived from Ensembl and HomoloGene. However, these databases are often incomplete or overly conservative in identifying orthologous gene pairs. Therefore, additional orthologous relationships (17% of those reported) were manually derived based on mRNA sequence similarity as determined by Bl2Seq analysis using an E-value cutoff of 10–30 when the rat and mouse gene also shared a common official gene symbol. Determinations of estrogen responsiveness, orthology, and unique gene annotation were collated, resulting in a list of 211 reciprocally active rat and mouse orthologs (Fig. 8).



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Fig. 8. Species comparison of array composition, gene activity, and orthology. Array features and represented unique genes as determined by LocusLink ID on each array are shown. The number of orthologous relationships between the rat and mouse array as defined by Ensembl and Bl2Seq comparison of RefSeq mRNAs with similar gene symbol and E-value cutoff of 10–30. Active genes determined by a P1(t) >0.999999 at any time point for both species.

 
Orthologous genes that are reciprocally active do not necessarily display similar directional or temporal patterns of expression. Therefore, to determine whether the orthologous gene pairs had comparable temporal expression patterns, a Pearson correlation analysis was performed according to temporal fold change on a gene-by-gene basis (Fig. 9). This analysis identified 153 positively (1.0 to 0.1; Table 2), 12 non- (0.1 to –0.1), and 46 negatively (–0.1 to –1.0; Table 3) correlated orthologous gene pairs. Agglomerative hierarchical clustering (GeneSpring v7.1) of the 153 positively correlated genes indicates that gene expression patterns between 8 and 24 h display similar patterns of expression within species, whereas gene expression at early (2–4 h) and late (72 h) time points clustered by time point rather than species (Fig. 10). \. Of the 153 positively correlated, 95 showed a common pattern of upregulation between 8 and 24 h, indicating a high proportion of conserved gene expression changes and suggesting that the mechanisms of regulation are conserved. A number of these genes have been previously reported in microarray studies (5, 19, 21, 26, 27, 39, 41, 52, 53, 66, 68) to be responsive to estrogen treatment in the rodent uterus (Table 2).



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Fig. 9. Temporal correlation of reciprocally active orthologous genes. Reciprocally active orthologous genes were temporally correlated according to fold change of EE-treated samples relative to time-matched VCs using a Pearson's correlation. Orthologs with a correlation coefficient of –1 to –0.1, –0.1 to 0.1, and 0.1 to 1 were designated as negatively, non-, and positively correlated, respectively. The cDNA array probe sequences for orthologs not positively correlated were subsequently compared using Bl2Seq analysis; sequence alignments having an E-value of 10–30 or less were considered to be overlapping probe pairs.

 

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Table 2. Conserved and coordinated estrogen-responsive genes in the rodent uterus

 

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Table 3. Divergent estrogen-responsive genes in the rodent uterus

 


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Fig. 10. Hierarchical clustering of positively correlated rat-mouse orthologous gene expression profiles. The 153 orthologous pairs were hierarchically clustered by gene and time. Rat and mouse data are indicated in red and black text, respectively, and time points (h) are labeled. Orthologs for 2, 4, and 72 h cluster by time between species, while data points from 8 to 24 h cluster by species, indicating a high level of temporal conservation between species in early and late responses, while data points between 8 and 24 h are not distinguishable between species.

 
cDNA probe sequences for the 54 orthologs not positively correlated were subsequently compared to determine whether they queried comparable regions of the gene using Bl2Seq analysis. Sequence alignments with an E-value of 10–30 were considered to be overlapping probe sets. Thirty of the fifty-eight not positively correlated ortholog pairs were found to have overlapping sequences, indicating that the same region of the gene was queried by microarray analysis and therefore differentially regulated.


    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The enhanced rodent uterotrophic assay was performed using EE as a prototypic estrogen to identify and phenotypically anchor conserved gene expression profiles important for uterotrophy. The analysis of these conserved targets provides baseline transcriptional program information involved in the uterotrophic response through the elucidation of the significance of these gene expression changes and their putative functions as reported within Gene Ontology annotations (http://www.geneontology.org) and the published literature.

Increases in uterine wet weight were preceded by conserved and coordinated gene alterations in multiple functional categories including cell cycle control, redox control, DNA replication, protein synthesis and transport, pro- and anti-apoptotic genes, xenobiotic metabolism, cell-cell communication, and angiogenesis. Several gene expression profiles populate each of these categories and collectively serve as the basis for the development of a uterine molecular finger print of estrogen exposure. Furthermore, these conserved targets of estrogen signaling in the rodent uterus may also support extrapolations to human uterine responses.

Previous rat uterine gene expression studies have reported similar gene expression profiles in response to estrogen (13, 41). However, the study designs used limit the interpretation of the functional significance of the coordinated responses. For example, Naciff et al. (41) utilized pooled uterine and ovarian tissues from rats treated subcutaneously with EE for dose-dependent gene expression studies which confound the elucidation of tissue-specific responses due to the differential distributions of ER{alpha} and ERß (63, 64) in the uterus and ovaries. Moreover, an important consideration in modeling estrogenicity is the fact that the primary route of exposure to diverse estrogenic compounds in human populations is oral. Diel et al. (14) supplemented the uterotrophic assay by evaluating the expression of a select number of known estrogen-responsive genes at 72 h in conjunction with standard gravimetric and histological evaluations (14). However, use of a small number of transcriptional markers to determine estrogenicity limits the ability to phenotypically anchor gene expression changes (50). Global assessments facilitate a more comprehensive determination of transcriptional targets in the physiological context as well as identifying mechanistically based biomarkers. Nevertheless, there is good agreement between these studies that currently can only be appreciated on a qualitative basis due to the limited reporting of the data required to fully correlate their expression profiles.

EE induction of uterine wet weight is a well conserved phenotypic response. The strong conservation of this response is due to the exemplary manner in which the immature ovariectomized rodent uterotrophic assay exhibits synchronized cell cycle progression and subsequent proliferation in response to novel estrogen exposure (9). The wave of uterine cell growth in response to estrogen administration has been previously characterized at the gene expression level in mice (19, 39), whereas responses in the rat and comparative rodent studies have not been reported. In this study, rat and mouse data sets were normalized and statistically analyzed, utilizing identical array platforms to facilitate comparisons. Orthologous gene pairs exhibiting a positive correlation were assigned to functional gene categories (Table 2) to annotate their mechanistic roles and facilitate phenotypic anchoring.

Immediate early genes and signaling molecules are the first line of EE responsive, conserved responders that initiate downstream cascades of transcription and translation. Many genes are regulated through AP-1 (12), a major mitotic effecter and transcription factor composed of c-Jun and Fos, which are both immediate early genes induced by estrogen in the rodent uterus. Igf1, another major estrogen mediator that binds its membrane-bound receptor and activates Mapk signaling cascades (33), was also upregulated between 4 and 24 h. Two members of this Mapk cascade were regulated by EE, with Map2k2 upregulated and Mapk3 downregulated. Mdk, a growth factor with undetermined function, was downregulated at 72 h. Its absence could possibly play a role in halting growth at peak uterine size. Gja1, although not a signaling molecule itself, encodes a gap junction protein crucial for cell-to-cell communication during concerted tissue responses to mitotic stimuli (38) and exhibited conserved upregulation in both species between 8 and 18 h.

Conserved gene expression changes involved in transcription included Atf4, Bcap37, Gata2, Id1, Rere, and Irf7. Rere acts as a transcriptional co-repressor through interactions with Hdac1 and also plays a role in pro-apoptotic signaling (62, 75), consistent with its sustained downregulation at mid time points in rats and mice. Conversely, Bcap37, referred to as REA for repressor of ER activity, interacts with Hdac1 to repress nuclear receptor-mediated transcription (34) and was upregulated for several intermediate time points in both species. There was also conserved induction of Bop1, Hnrpab, and Ppm1g, which are involved in mRNA processing. Hnrpab, a heterogeneous nuclear ribonucleoprotein involved in mRNA splicing and processing, has been shown to be a methylation target of Hrmt1l2 (1), a protein-arginine methyltransferase that was also upregulated by EE in the rat. The Ppmg1 gene encodes a phosphatase required for spliceosome assembly (40).

Following transcriptional activation and mRNA processing is protein synthesis, processing, and turnover. Conserved responses included the following: amino acid biosynthesis genes, Asns, Got1, Fah, and Phgdh; translation-related components, Eif2s2, Eif4e, Eif5, Gspt1, and Dnajc3; protein folding and transport, Cct3, Cct4, Tcp1, and Ran; heat shock proteins, Hspa4, Hspa5, Hspa8, Hspca, and Hspe1; and protein degradation-related genes, 26S proteasome complex members (Psma4, Psma5, Psma6, and Psmb5), and Nedd8. Gspt1 functions as the eukaryotic releasing factor in mediating translational termination via interaction with poly-A binding proteins (61) and was upregulated ~2.5-fold between 4 and 24 h. Tcp1, a component of Cct3 and Cct4 (chaperonin containing Tcp1 subunits 3 and 4) were all coordinately upregulated. These genes encode chaperone proteins involved in stabilization of un- or misfolded proteins and are important in cyclin E stabilization (67), which is important in cell cycle progression. In both rats and mice, Phgdh, an important enzyme in L-serine biosynthesis, was induced at mid time points, while Fah, the rate-limiting and final enzyme step in tyrosine synthesis, was downregulated, suggesting that it may have other yet unknown conserved functions. Specific signals controlling and inducing cell cycle progression (Ccnd1, Cdc2a, Phb, Ndrg2, Sfrp1) and DNA replication (Fen1, Hmgb2, Nme1) were also conserved. Estrogen-responsive Ccnd1 and Cdc2a, key regulators of G1/S-phase and G2/M-phase transition, respectively (10, 74), were both upregulated, suggesting concerted uterine cell cycle transitions induced by EE in the ovariectomized model.

Subsequent to increases in translational activity and protein content are conserved induction of protein degradation and turnover genes (43, 71) such as those involved in the 26S proteasome complex (e.g., Psma4, Psma5, Psma6, and Psmb5). Nedd8 is involved in proteasome-mediated degradation of ER{alpha} (18) and was upregulated at mid time points. App-binding protein-1 (Appbp1)/Uba3, the catalytic subunit of the Nedd8-activating enzyme, was upregulated at similar time points in the mouse. Furthermore, Appbp1 is a binding partner of amyloid precursor protein (App), which has been implicated in Alzheimer's disease (7). App expression was downregulated in both species. Itm2b, which has similar behavior and function as App in diverse brain lesions associated with dementia (2), was also downregulated in parallel with App. This is possibly due to the downregulation of Tgfbi seen in the rat study. Tgfbi has been shown to positively regulate App (7). It is unclear what role App plays in regulating the Nedd8 degradation pathway, but the network of interactions is suggestive of a regulatory mechanism involved in ER degradation via the 26S proteasome.

Increased energy demands were reflected in both species by the induction of genes involved in mitochondrial (Cycs, Atp5g1, Ckmt1, Slc25a4, Slc25a5) as well as cytosolic (Ckb, G6pd, Pgk1) energy production and homeostasis. Slc25a5, an ADP/ATP translocase, catalyzes the exchange of cytosolic ADP for mitochondrial ATP and has been implicated as a marker of cell growth and proliferation due to its key role in regulating cytosolic energy needs (4). G6pd on the other hand functions mainly in generating the other energy currency of the cell, NADPH, which is used in many cell processes including fatty acid and cholesterol biosynthesis via the pentose phosphate pathway (59). Several genes involved in the biosynthesis or transport of membrane precursors were also regulated by EE, including Acadm, Apoc1, Lpl, Sqle, and Sc4mol. Acadm and Lpl, enzymes in the fatty acid ß-oxidation pathway, were downregulated. In contrast, Sc4mol, which catalyzes the final C-4 methyloxidation step in cholesterol formation (20), was upregulated, consistent with the need for building blocks for new membranes demanded by rapid cell division (55).

Despite the overlaps in gene expression profiles, several divergent patterns were identified. Apoe, a known regulator of lipid transport and uptake in the liver, was induced in the rat while repressed in the mouse at intermediate time points. Carbonic anhydrase 2, which catalyzes the reversible hydration of carbon dioxide to carbonate, has recently been shown to be crucial for adenogenesis (gland development) in the uterus (28). The fact that Ca2 expression was repressed in rat while induced in the mouse could explain increased glandular invaginations observed in mouse but not rat at 72 h. Cdc37, a regulator of Hsp90 phosphorylation, as well as Cstb, Cugbp2, Dlat, Smarcd2, and Tcn2 were also differentially regulated at overlapping time points. Bl2Seq sequence comparisons of these differentially regulated orthologous pairs indicated that several of these cDNA probes overlapped comparable transcript regions (Table 3). Therefore, their differential expression cannot solely be attributed to probes querying different transcript regions. Consequently, EE regulation of these gene expression responses are not conserved, suggesting that their role is not critical for the uterotrophic response.

This comparative study has comprehensively assessed the physiological, morphological, and transcriptional programs elicited by EE in the rat and mouse uterus using comparable study designs, assay platforms, and analysis methods. One hundred fifty-three reciprocally active and positively correlated gene expression profiles were identified, suggesting that these are important responses that share a conserved mode of action. Moreover, comparable temporal expression patterns provide further evidence that orthologous genes are functionally related, supporting the transfer of mouse functional annotation to unknown rat expressed sequence tags (ESTs) with tentative annotation. For example, of the 153 reciprocally active and positively correlated gene expression profiles, 36 rat sequences had no official name but a gene symbol, 1 had a name but no symbol, and 7 had neither a name nor symbol. Many of these tentative assignments were based only on sequence homology, but the co-expression data described here provide empirical support that the genes are orthologous and functionally related. Comparable approaches have recently been used to identify functionally related genes across more divergent species [e.g., extrapolations between human, worm, fly, and yeast (57, 72); extrapolating bacteria, yeast, and fly annotation to the worm (57, 72)]. Additional analyses including dose-response studies, promoter response element comparisons, and cell type-specific assessments will not only further elucidate the importance of these conserved and divergent EE-induced uterine proliferation responses but will also provide additional evidence regarding the conservation of estrogen response between rodent and human orthologous gene pairs.


    GRANTS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
J. C. Kwekel is supported by a fellowship from the Center for Integrated Toxicology through National Institute of Environmental Health Sciences (NIEHS) Training Grant T32 ES-07255-16. T. R. Zacharewski is partially supported by the Michigan Agricultural Experimental Station. This work was supported by funds from NIEHS Grant ES-011271 and Environmental Protection Agency Grant RD-83184701.


    ACKNOWLEDGMENTS
 
We thank colleagues that contributed to this work, specifically, Darrell Boverhof, Heather Dalgleish, Edward Dere, and Cora Fong.


    FOOTNOTES
 
Article published online before print. See web site for date of publication (http://physiolgenomics.physiology.org).

Address for reprint requests and other correspondence: T. R. Zacharewski, Michigan State Univ., Dept. of Biochemistry & Molecular Biology, 224 Biochemistry Bldg., Wilson Rd., East Lansing, MI 48824 (e-mail: tzachare{at}msu.edu)

1 The Supplemental Material for this article (Supplemental Tables S1–S3) is available online at http://physiolgenomics.physiology.org/cgi/content/full/00175.2005/DC1. Back


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 DISCUSSION
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