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Endocrinology Vol. 143, No. 8 3044-3059
Copyright © 2002 by The Endocrine Society

ARTICLE

Gestational and Lactational Exposure of Male Mice to Diethylstilbestrol Causes Long-Term Effects on the Testis, Sperm Fertilizing Ability in Vitro, and Testicular Gene Expression

Mark R. Fielden, Robert G. Halgren, Cora J. Fong, Christophe Staub, Larry Johnson, Karen Chou and Tim R. Zacharewski

Department of Biochemistry and Molecular Biology, National Food Safety and Toxicology Center (M.R.F., R.G.H., C.J.F., T.R.Z.), and Department of Animal Science (K.C.), Institute for Environmental Toxicology, Michigan State University, East Lansing, Michigan 48824; and Department of Veterinary Anatomy and Public Health, Texas A&M University College of Veterinary Medicine (C.S., L.J.), College Station, Texas 77843

Address all correspondence and requests for reprints to: Tim Zacharewski, Ph.D., Department of Biochemistry and Molecular Biology, 223 Biochemistry Building, Wilson Road, Michigan State University, East Lansing, Michigan 48824. E-mail: . tzachare{at}msu.edu


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The objective of the study was to determine the long-term effects of gestational and lactational exposure to diethylstilbestrol (DES; 0, 0.1, 1, and 10 µg/kg maternal body weight) on mouse testicular growth, epididymal sperm count, in vitro fertilizing ability, and testicular gene expression using cDNA microarrays and real-time PCR in mice on postnatal day (PND) 21, 105, and 315. In the high dose group there was a persistent decrease in the number of Sertoli cells, and sperm count was decreased on PND315 (P < 0.05). Sperm motion was unaffected; however, the in vitro fertilizing ability of epididymal sperm was decreased in the high dose group on both PND105 (P < 0.001) and PND315 (P < 0.05). Early and latent alterations in the expression of genes involved in estrogen signaling (estrogen receptor {alpha}), steroidogenesis (steroidogenic factor 1, 17{alpha}-hydroxylase/C17,20-lyase, P450 side chain cleavage, steroidogenic acute regulatory protein, and scavenger receptor class B1), lysosomal function (LGP85 and prosaposin), and regulation of testicular development (testicular receptor 2, inhibin/activin ß C, and Hoxa10) were confirmed by real-time PCR. The results demonstrate that early exposure to DES causes long-term adverse effects on testicular development and sperm function, and these effects are associated with changes in testicular gene expression, even long after the cessation of DES exposure.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
DIETHYLSTILBESTROL (DES) is a nonsteroidal synthetic estrogen that was administered to pregnant women to prevent miscarriages in the late 1940s and 1950s. Its use was terminated in 1971 when it was concluded that prenatal exposure to DES causes cervical clear cell adenocarcinoma in female offspring after otherwise normal pubertal development (1). Putative effects in human males include testicular cancer, hypospadias, cryptorchidism, reduced sperm count, and impaired fertility (2). A causal link between estrogen exposure in utero and adverse effects on the male reproductive tract and sperm quality is also supported by studies of wildlife populations exposed to estrogenic chemicals in the environment (3). Furthermore, epidemiological studies support a decrease in human sperm count (4, 5) and an increase in testicular cancer and reproductive tract malformations (6, 7). This has raised a concern over exposure to both synthetic and natural estrogenic endocrine disruptors (EEDs) in the environment and their potential effects on human reproductive health (8). Studies in laboratory animals have confirmed the adverse effect of estrogens on male reproductive health. For example, in utero exposure of male mice to high doses of DES causes sterility resulting from a number of reproductive tract abnormalities, including enlarged and cystic Mullerian remnants, inhibition of gubernaculum development and cryptorchidism, sperm granulomas, hypotrophic testes and epididymides, epididymal cysts of embryonic female origin, and tumors of the rete testis and interstitial cells (9, 10, 11, 12, 13, 14, 15, 16, 17, 18). Nonteratogenic effects induced by high doses of DES include decreases in testicular growth and sperm production in adulthood (19, 20). Changes in organ weight or sperm production, however, do not adequately predict sperm function and quality (21), so it is unknown whether exposure to lower nonteratogenic doses of EEDs can affect sperm function.

The molecular mechanisms responsible for the adverse effects of DES are unclear, although there is evidence that the initiating events involve changes in gene expression. For example, estrogen-induced cryptorchidism is believed to occur as a result of down-regulation of insulin-like factor 3 in Leydig cells of the fetal mouse testis (17, 18). Incomplete regression of the Mullerian ducts in DES-exposed male fetuses may involve increased expression of steroidogenic factor 1 (SF-1) and anti-Mullerian hormone and its receptor (15). By contrast, expression of SF-1 and its target gene 17{alpha}- hydroxylase/C17,20-lyase (Cyp17) were reported to be decreased in the Leydig cells of fetal rats exposed to DES (22, 23). In utero exposure to DES has also been reported to affect the expression of estrogen receptor {alpha} (ER{alpha}), Wnt-7a, epidermal growth factor, and various homeobox genes in the Mullerian duct of female mice (24, 25, 26, 27, 28, 29). Estrogens may also exert long-term reprogramming effects by altering gene expression, cellular differentiation, tissue restructuring, and concomitant responsiveness of various tissues of the reproductive tract (30, 31), thus possibly leading to reproductive tract abnormalities, malfunction of the Wolffian ducts, and adverse effects on spermatogenesis.

The changes in gene expression induced by prenatal exposure to DES are likely to be mediated at least in part by the ER. The distinct expression pattern and ontogeny of ER{alpha} and ERß in the rodent testis (32, 33, 34), and the detrimental effect of genetic disruption of ER{alpha} signaling (35) and estrogen synthesis (36) highlight the importance of estrogen and its receptor for normal spermatogenesis and sperm fertilizing ability. In addition to its ER agonist activity, DES also inhibits the transcriptional activity of the ER-related orphan nuclear receptors in vitro (37, 38) and estradiol (E2)-induced trans-activation of the androgen receptor (AR) in vitro (39), leading to speculation that DES may disrupt male reproductive development through mechanisms independent of the ER. In addition, the mutagenicity of DES (40) and the carcinogenicity in reproductive tissues after developmental exposure to DES (12, 14, 41) suggest that genotoxic mechanisms may contribute to the effects on fertility. Clearly, a more thorough understanding of the molecular changes induced by DES and other EEDs is needed to understand the pathophysiology of DES and other EEDs on the male reproductive system

The purpose of this study was to determine whether gestational and lactational exposure to DES causes long-term effects on testicular development, sperm count, and motility, and in vitro fertilizing ability. Sperm fertilizing ability was assessed using an in vitro fertilization (IVF) assay to avoid confounding treatment effects on sexual behavior, sperm count, or abnormal development of external genitalia and accessory glands. Male offspring were examined at early and mid stages of life to determine the persistence of the treatment-related effects. To test the hypothesis that the long-term effects of DES on sperm quality are the result of persistent changes in testicular gene expression and to potentially identify early biomarkers of adverse effects at later stages of life, a cDNA microarray was used to identify gene expression changes in the testis of affected mice. Doses of DES were chosen to minimize or avoid reproductive tract abnormalities that may confound the interpretation of effects on sperm function or testicular gene expression.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals
Mice were obtained from Charles River Laboratories, Inc. (Portage, MI). They were housed in polycarbonate cages with cellulose fiber chips (Aspen Chip Laboratory Bedding, Northeastern Products, Warrensberg, NY) as bedding and maintained in a humidity- (30–40%) and temperature (23 C)-controlled room on a 12-h light, 12-h dark cycle. All animals were given free access to deionized water in glass bottles with rubber stoppers and AIN-76A rodent feed ad libitum (Research Diets, New Brunswick, NJ). This diet is a casein-based, open formula, purified diet with undetectable levels of the estrogenic isoflavones genistein, diadzein, and glycitein (Fielden, M. R., and T. R. Zacharewski, unpublished data) (42, 43).

F0 treatment
Eleven-week-old virgin C57BL/6 female mice (F0) were housed two per cage upon arrival and acclimatized for at least 3 d. Each pair of females was housed with a 7- to 8-wk-old DBA/2 proven breeder male within a week of arrival. Offspring from this cross, designated B6D2F1, were used because a large number of eggs can be obtained from superovulated B6D2F1 mice, and a high level of fertilization can be consistently and reproducibly obtained in control animals (44, 45). After evidence of pregnancy was obtained, dams were separated from the males, housed individually, and sequentially assigned to one of four treatment groups. Because of the low fecundity of dams in the high dose group, additional pregnant mice were allocated to generate a sufficient number of viable litters per group (see Results). Due to the limited number of male offspring that can be examined for sperm analysis and IVF on any 1 d, breeding was staggered into four cohorts (i.e. replicates) to accommodate sperm analysis for all male offspring. Each cohort included time-matched control animals.

F0 mice were treated by daily gavage with 0.1 ml corn oil (Sigma, St. Louis, MO) for nominal doses of 0, 0.1, 1, and 10 µg DES/kg maternal body weight (0900–1200 h) from gestational d 12 to postnatal d 20 (PND20). There was a 1-d interruption of treatment on the day of parturition (PND0). The dose of test chemical was adjusted daily to body weight for each dam before dosing. Offspring were weaned on PND21, when F0 mice were killed.

F1 development
Litter size and litter weight were recorded on PND0 and PND1, respectively. F1 body weight was measured on PND7 and PND21. Ano-genital distance (AGD; the length of the perineum from the base of the genital tubercle to the center of the anus when skin was naturally extended without stretching) was measured on PND7 and -21. Measurements were recorded to the nearest 0.1 mm and were obtained with a dissecting scope equipped with an ocular micrometer (Nikon, Melville, NY). AGD was measured by the same individual to increase precision and to control for operator variation. F1 mice were weaned on PND21, housed with same sex littermates, and fed AIN-76A ad libitum until necropsy. F1 females were also assessed for oocyte-fertilizing ability in vitro and mammary gland development (to be described elsewhere).

F1 necropsy
On PND21, one F1 male per litter was killed for necropsy. When the number of F1 males in a litter was low, the male(s) was instead withheld for necropsy on PND105 and -315 when male offspring were also killed for sperm assessment. The remaining F1 males from each litter were randomly selected for sperm assessment on either PND105 or -315. At necropsy, both testes were excised, trimmed of adhering connective tissue, and weighed wet. Both seminal vesicles with coagulating glands were excised and weighed wet on PND105 and -315. At all three time points one testis was fixed for histology, and the other testis was stored immediately for subsequent RNA extraction and gene expression analysis.

Testicular histopathology
One testis per animal was fixed in 15–20 volumes of Bouin’s fixative (Sigma) for at least 24 h and cleared in three successive 1-h washes in 70% ethanol. Wet tissue was stored in 70% ethanol at room temperature before being shipped to Experimental Pathology Laboratories (Durham, NC) for analysis. Each testis was embedded in paraffin, and three 5-µm cross-sections from the cranial, median, and caudal parts of the testes were stained with hematoxylin and eosin. All three sections were mounted on the same slide and evaluated microscopically by pathologists blind to the treatment group.

Testicular RNA isolation
One testis per animal was immersed in 5 vol of RNALater (Ambion, Inc., Austin, TX) in a 2-ml tube and immediately placed on dry ice before long-term storage at -20 C. For total RNA isolation, tissue frozen in RNALater was thawed on ice and transferred to 0.75 ml TRIzol reagent (Life Technologies, Inc., Rockville, MD). Tissue was then immediately homogenized, and total RNA was isolated as described in the manufacturer’s instructions. Linear acrylamide (20 µg) was added as a carrier to aid precipitation and increase yield. RNA pellets were resuspended in 25–50 µl 1 mM sodium citrate, pH 6.4, and stored at -20 C until use. Total RNA was quantitated (A260) and assessed for purity (A260/A280) by spectrophotometry.

Stereological analysis of Sertoli cells
Paraffin blocks were deparaffinized, further fixed in 1% osmium in sodium cacodylate buffer, and embedded in epon. Tissues were sectioned at 0.5 µm, stained with toluidine blue, and used for stereological determination of the volume density of Sertoli cell nuclei and the average volume of the Sertoli cell nucleus (46). The number of Sertoli cells per testis was calculated when the product of the volume density of Sertoli cell nuclei, testicular parenchymal volume, and the approximated histological correction factor for section thickness and nuclear diameter (47) was divided by the corrected volume of a single Sertoli cell nucleus.

The volume density of Sertoli cell nuclei was based on random movements of the microscope slide at x1000 magnification with brightfield microscopy. Point counting (6250 points/mouse) by 2 observers was made on 2 blocks of tissue and 2 slides/block in 125 fields randomly chosen throughout the tissue section. Volume density equals the number of points falling on Sertoli cell nuclei divided by the total number of points (6250) counted. Testicular parenchymal volume equals the weight of the testis minus the capsule weight divided by the specific gravity for the testis (1.05 g/ml). The testicular capsule weight was estimated based on its percentage of the total testis weight, as determined previously in the mouse. The average volume of a single Sertoli cell nucleus was calculated from the volume of a sphere based on height and width measurements of 50 Sertoli cell nuclei/mouse that were totally embedded in 20-µm Epon sections observed by Nomarski optics. A nonspherical correction factor for Sertoli cell nuclei has been determined for rats (48), and this factor was shown not to differ during the (prepubertal and pubertal) development of the rat testis. This correction factor was calculated empirically by comparing the estimate of nuclear volume and a reference volume of individual Sertoli cell nuclei determined by reconstruction of serial sections of Sertoli cell nuclei. Area was determined on each of the serial sections, and the volume of an individual nucleus was calculated as the sum of the areas times the section thickness. The ratio of volume based on serial section reconstruction divided by the volume based on height/width measurements and volume formula for sphere was 0.663 ± 0.025.

Epididymal sperm count and motion analysis
Cauda epididymal sperm were collected from F1 males on PND105 and -315 by excising both epididymides and piercing with a 25-gauge needle in an organ culture dish (BD Biosources, Franklin Lakes, NJ) containing 1 ml BMOC-3 medium (Life Technologies, Inc.), a capacitation supporting medium (49). Sperm suspensions were incubated at 37 C in a humidified 5% CO2 air environment for 30 min before sperm concentration and motion analyses and 60 min before insemination (see below). Sperm suspensions (20 µl) were placed on a 20-µm counting chamber and analyzed using a CellSoft computerassisted digital image analysis system (CASA) series 4000 (CRYO Resources Ltd., Montgomery, NY). The number of frames to analyze (n = 15), the number of frames per second (n = 30), the threshold velocity (50 µm/sec), the threshold gray value (85), the maximum velocity (300 µm/sec), the minimum velocity (20 µm/sec), the minimum sampling for velocity (n = 3), and motility (n = 2), and the minimum linearity (3.5) were consistent with optimal settings determined in the rat (50, 51). The cell size range (5–27) and the pixel scale were determined empirically for the mouse. A minimum of 100 cells were analyzed, based on randomly chosen fields, to determine average sperm count, motility, number of motile sperm, velocity, linearity, amplitude of lateral head (ALH) displacement, and beat/cross-frequency for each animal. Counting 100 cells ensures that an equivalent number of sperm per animal was analyzed, and that the sperm spent a constant amount of time in the counting chamber. Sperm count represents the number of sperm in 1 ml medium. Motility is expressed as the percentage of sperm that move faster than 20 µm/sec. The number of motile sperm was the product of sperm count and motility. Velocity is defined as the average distance (microns) traveled by motile sperm in 1 sec. Linearity is the ratio of the straight to actual distance traveled (x10) averaged over all sperm. ALH displacement is a measure of the lateral movement of the sperm head from the curval mean of its track. The beat/cross frequency (hertz) is the numbers of beats (or crosses) of the sperm across its curval mean per second. All measurements for each sperm collection were performed in duplicate and averaged.

IVF assay
The in vitro fertilizing ability of sperm was assessed on PND105 and -315 by inseminating oocytes collected from nontreated, 3- to 4-wk-old, B6D2F1 female mice. Female mice were superovulated with 10 IU pregnant mare’s serum gonadotropin (Sigma), followed 48 h later by 10 IU human chorionic gonadotropin (Sigma). Fourteen to 17 h later the oocytes from each female were collected from the proximal oviducts. Oocytes from each mouse were incubated in a 1-ml organ culture dish containing 0.9 ml Brinster’s BMOC-3 medium. Epididymal sperm from each F1 male was used to inseminate oocytes from two females, each in a separate dish. Sperm were diluted in BMOC-3 medium, and 0.1 ml diluted sperm was added to 0.9 ml medium containing the oocytes to achieve a final sperm concentration of 3 x 104 sperm/ml·dish. This concentration of sperm achieves slightly less than maximum fertilization in naive B6D2F1 mice, thus increasing the assay sensitivity for detecting positive and negative changes in sperm fertilizing ability. It was not possible to pool, randomize, and equally distribute oocytes from all females, because oocytes were contained within a cumulus mass, and oocyte collection was coordinated with sperm collection and CASA analysis for consistency among inseminations. After insemination, oocytes were incubated at 37 C under a humidified 5% CO2 air environment. After a 24-h incubation, 50 µl 35 µM bisbenzimide stain (Sigma) were added to the dish. The oocytes were incubated with stain for at least 30 min before being examined using a Nikon Optophot fluorescent microscope (Melville, NY) equipped with a 100-watt mercury bulb, a 365/10-nm excitation filter, a 400-nm dichromic mirror, and a 400-nm barrier filter. Oocytes were counted and scored as fertilized if the eggs were at the two-cell stage or at the one-cell stage containing two pronuclei and a second polar body. Oocytes were also evaluated for fragmentation and other signs of degeneration. As it was not always possible to distinguish a one-cell from a two-cell fragmented egg, they were not included in the total egg count for calculating percent fertilization. Fertility data for the replicate dishes were averaged.

Statistical analysis
All data analysis was performed using SAS version 8.0 (SAS Institute, Inc., Cary, NC). To estimate experimental error, the litter was considered the experimental unit. There was no significant (P > 0.1) replicate effect; therefore, data from all cohorts were pooled for analyses. All data were tested for normality by the Shapiro-Wilk test. Evidence for nonnormality was declared at the 5% level of significance. When significant, the data were log transformed and retested. Data that were not normally distributed were analyzed by nonparametric one-way ANOVA using the NPAR1WAY procedure of SAS. Comparisons between control and treated groups were made with the Kruskal-Wallis test. Data passing the normality test were analyzed with a repeated measures ANOVA using the MIXED procedure of SAS. Using an analysis of covariance, litter size was found to account for a significant source of the variation (P < 0.05). Therefore, for the analysis of body weight, the model included dose, time, and dose x time interaction as fixed effects and litter size as a covariate. Likewise, for the analysis of AGD and organ weights, body weight was included in the model as a covariate because body weight was found to account for a significant source of the variation (P < 0.05) (52). For comparison, AGD was also analyzed as a ratio of AGD to the cube root of body weight (53), whereas organ weights were analyzed as a ratio of organ weight to body weight. Sperm count, sperm motion parameters, and Sertoli cell morphometry were analyzed using dose, time, and dose x time interactions as fixed effects. Due to the unbalanced nature of the ANOVA, and to adjust group means for the covariate, comparisons between control and treated groups were performed on least square means and adjusted for multiple comparisons using Dunnett’s method. Arithmetic means, SE, and n values for litters are reported in Results. The effect of treatment on discrete data (i.e. proportion of eggs fertilized) was analyzed by logistical regression using a binomial distribution as implemented in the GENMOD procedure of SAS. The level of significance was set at P < 0.05.

Microarray analysis of differential gene expression
cDNA microarrays were used to profile gene expression differences in the testis of male B6D2F1 offspring in the 10 µg/kg DES group relative to offspring in the vehicle control group on PND21, -105, and -315. The 10 µg/kg DES group was chosen for comparison against age-matched vehicle control animals because changes in testicular development and sperm quality were demonstrated at this dose. Each microarray experiment (i.e. hybridization) compared the relative level of gene expression for a single animal in the DES group (Cy5-labeled cDNA) to that of a single animal in the control group (Cy3-labeled cDNA) within each age. One animal per litter was used for the control and DES groups, such that replicate gene expression measurements could be made on independent experimental units (i.e. the litter) on PND21 (n = 5), PND105 (n = 9), and PND315 (n = 4). Differentially expressed genes were then identified at each of the three time points by paired t tests and P values that were adjusted by the step-down Bonferroni method (described below). Both raw and adjusted P values were used to prioritize genes for subsequent experimental verification by real-time PCR, in addition to the persistence of the putative change in gene expression over time, and their possible role in spermatogenesis based on functional information from the literature. Additional genes not on the array were also quantitated by real-time PCR as described below (also see Table 1Go).


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  		Table 1. Genes, primer sequences, amplicon size, and optimal amplification conditions for real-time PCR

 
To maximize the amount of useful gene expression information and increase the probability of detecting relevant changes in testicular gene expression, a list of genes expressed in the testis was generated and used to select genes to be printed on the microarray (http://www.bch. msu.edu/~zacharet/dbZach/dbTestExp.html). A total of 1304 cDNA clones were obtained from Research Genetics, Inc. (Huntsville, AL), and sequence was verified at the Michigan State University Genomic Technology Support Facility (www.genomics. msu.edu). Another 984 sequence-verified cDNA clones were also obtained from Research Genetics, Inc., for a total of 2288 unique cDNA clones (provided by Dr. David Dix, U.S. Environmental Protection Agency, NC; through the E.P.A. Microarray Consortium). The accession number of each cDNA clone was queried against the mouse UniGene database to assign a putative gene identity. Based on build 99 of UniGene, 1873 cDNA clones had an assigned gene name, 334 are expressed sequence tags, and 97 have no match to any gene in the mouse UniGene Database. In total, 2304 cDNA clones (2288 unique) representing 1948 unique UniGene clusters were used in the construction of the microarray. A complete list of genes on the array can be found at http://www.bch.msu.edu/~zacharet/microarray/ supplemental/index.html.

Methods used for construction of the microarrays, labeling of the cDNA probe, and hybridization and washing can be found at http://www.bch.msu.edu/~zacharet/microarray/microarray.html. Briefly, PCR-amplified DNA was robotically arrayed in duplicate onto Superaldehyde slides (Telechem) using an Omnigrid arrayer (GeneMachines, San Carlos, CA) equipped with 16 (4 x 4) Chipmaker 2 pins (Telechem). Total RNA (25 µg) was used as template for a reverse transcriptase reaction to incorporate Cy5-dUTP (DES treated) or Cy3-dUTP (vehicle control) into first strand cDNA. The Fluor-labeled cDNA was purified with the QIAGEN PCR Purification Kit (QIAGEN, Valencia, CA). The Cy3-labeled cDNA was mixed with the Cy5-labeled cDNA, vacuum dried, and resuspended in 32 µl hybridization buffer (40% formamide, 4x standard saline citrate, and 1% sodium dodecyl sulfate) containing 20 µg polyadenylate and 20 µg mouse COT-1 DNA (Life Technologies, Inc.) as competitor. The probe mixture was heated at 95 C for 2 min before being hybridized to the microarray under a 22 x 40-mm coverslip (Corning, Inc., Corning, NY) in a light-protected humidified hybridization chamber (Corning, Inc.). The microarray was hybridized for 16–18 h at 42 C. Slides were washed, dried by centrifugation, and scanned promptly.

Image acquisition and data processing
Slides were scanned at 635 nm (Cy5) and 532 nm (Cy3) on an Affymetrix 428 Array Scanner (Santa Clara, CA). Images were analyzed using GenePix 3.0 (Axon, Foster City, CA). Blocks were manually adjusted, and features were automatically detected and quantitated as suggested in the manufacturer’s instructions. Images were surveyed visually to flag anomalous spots. GenePix results files (i.e. gpr files), which contain raw signal intensity and background signal intensity values for each spot and channel, were further processed in batch using an automated and customizable script called GP3. GP3 filters spots below the limits of detection or at saturating levels, corrects for background signal, and applies a global linear normalization factor in log space to generate valid estimates of gene expression in each channel (54). Details of the algorithm, and downloads can be found at http://www.bch.msu.edu/~zacharet/microarray/gp3.html.

Statistical analysis of microarray data
To assess the degree of experimental, biological, and treatment- induced variation in gene expression measurements, Pearson correlation coefficients (r) were calculated to compare the similarity between sample measurements within and across hybridizations (PRISM 3.0, GraphPad Software, Inc., San Diego, CA). Correlation coefficients were normalized with a Fisher’s z transformation to calculate 95% confidence intervals surrounding r. Differentially expressed genes were identified by computing paired t statistics and P values using SAS version 8.0 (Cary, NC). The paired t test compares the mean of the differences in the observations according to the null hypothesis that the mean difference equals zero. The t statistic is given by the equation: ti = (di - 0)/({varsigma}i/{surd}n), where d is the mean difference between the normalized signal intensity for gene i in channels 1 and 2 (i.e. Cy5 and Cy3), \|[sfgr ]\| is the SD of the paired differences, and n is the number of paired samples. To control the family-wise type 1 error rate, the P values were adjusted using the stepdown Bonferroni method (55).

Real-time PCR
To verify changes in gene expression observed by microarray analyses, real-time PCR was performed on PE Applied Biosystems PRISM 7700 Sequence Detection System (Foster City, CA). Additional genes of interest not on the microarray were also analyzed by real-time PCR (Table 1Go). The expression level for each gene was measured in all dose groups on PND21, -105, and -315 using six animals per dose group, for a total of 72 animals. To synthesize cDNA, total RNA (1 µg/animal) was used as template for a reverse transcriptase reaction in 20 µl 1x First Strand Synthesis buffer (Life Technologies, Inc.) containing 1 µg oligo(T18A/C/GN), 0.2 mM dNTPs, 10 mM dithiothreitol, and 200 U SuperScript II reverse transcriptase (Life Technologies, Inc.). The reaction mixture was incubated at 42 C for 60 min and was stopped by incubation at 75 C for 15 min. Amplification of cDNA (1/20th) was performed using the SYBR Green PCR Core Reagents (PE Applied Biosystems) as suggested by the manufacturer’s instructions. All 72 samples were amplified in a single MicroAmp Optical 96-well Reaction Plate (Applied Biosystems). Primer pairs for each gene were designed using Primer3 (56). Gene names, accession numbers, the forward and reverse primer sequences, amplicon size, and optimal primer and Mg2+ concentrations are listed in Table 1Go. Amplicon size and reaction specificity were confirmed by agarose gel electrophoresis. In addition, a number of stock cultures of the cDNA clones representing the genes on the microarray were used as a template in a PCR to verify gene-specific amplification, clone identity, and the integrity of the clone collection. The PCR cycling conditions were as follows: initial denaturation and enzyme activation for 10 min at 95 C, followed by 40 cycles of 95 C for 15 sec and 60 C for 1 min. Each plate contained duplicate standards of purified PCR products with known template concentration covering at least 6 orders of magnitude to interpolate relative template concentration of the experimental samples from standard curves of log copy number vs. threshold cycle (Ct). No template controls (NTC) were also included on each plate. Experimental samples with a Ct value within 2 SD of the mean Ct value for the NTC were considered below the limits of detection and assigned a value equal to the mean NTC. The relative level of mRNA was standardized to the housekeeping genes ß-actin and ribosomal protein S14 to control for differences in RNA loading, quality, and cDNA synthesis. There were no differences in the results when the data were standardized to either ß-actin or ribosomal S14 alone. As the gene expression data were found to violate the assumptions of normality and homogeneity of variance (data not shown), a nonparametric ANOVA was used to analyze differences in gene expression between control and treated groups. Differences between treatment groups and their age-matched control were determined by the Kruskal- Wallis test. Analyses were performed using SAS version 8.0. The level of significance was set at {alpha} = 0.05. For graphing, the relative expression level was scaled such that the expression level of the time-matched control group was equal to 100.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Reproductive performance of F0 generation
There was no difference in F0 weight gain between premating and PND20 (data not shown). The number of pregnant dams giving birth to live young (gestation index) was significantly decreased in the 10 µg/kg group compared with controls (11 of 19 vs. 14 of 16; P < 0.05). Five of the pregnant females in the 10 µg/kg dose group gave birth to moribund pups, whereas 3 did not give birth at all and were killed. Two (of 16) pregnant females in the control group did not give birth and were also killed. Pregnant females that did not give birth were verified by necropsy to be carrying fetuses. The live pups showed no differences in survival up to PND4 or PND21. There was a significant decrease in litter size in the 10 µg/kg group (9.0 vs. 6.8 pups/litter; P < 0.01). The average pup weight, however, was not significantly affected by DES treatment. The percentage of males in the 1 µg/kg group was slightly greater than that in the control group (17%; P < 0.05), whereas there was a large and significant decrease (32%; P < 0.05) in the percentage of males in the 10 µg/kg group (data not shown).

F1 development
Doses of DES were selected to minimize or avoid reproductive tract abnormalities. Indeed, there was only one incident of unilateral cryptorchidism observed (of 116 DES-exposed offspring) in the 10 µg/kg group on PND315. Both testes and epididymides from this animal were hypotrophic, and no sperm could be obtained. The retained testis was firmly attached to the seminal vesicle. Otherwise, there were no gross reproductive tract abnormalities observed in littermates or any other male offspring in the control or DES-exposed groups. Although epididymis weight was not recorded, abnormally small epididymides were not observed in any animals. Treatment did not have a significant effect on body weight, AGD, or seminal vesicle weight throughout the period of study (data not shown).

Testicular development and CASA analysis
Treatment did not have a significant effect on testis weight throughout the period of study, although there was a 30% decrease on PND21 and an 11% decrease on PND105 and -315 (data not shown). Testes from all dose groups on PND21 were examined microscopically, whereas only samples from the control and 10 µg/kg group on PND105 and -315 were examined microscopically. No treatment-related histological changes could be identified in any sections when assessed blind to the treatment. Stereological analysis of the Sertoli cells was conducted only on the testes from the control group and the 10 µg/kg group due to the observed differences in sperm count at this dose (Fig. 1Go; see below). The number of Sertoli cells per testis was significantly decreased 25%, 17%, and 32% relative to controls on PND21, -105, and -315, respectively (Fig. 1AGo).



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  		Figure 1. Quantitation of Sertoli cells and epididymal sperm from F1 males exposed to DES through gestation and lactation. A, Number of Sertoli cells per testis on PND21, -105, and -315 in male offspring. B, Epididymal sperm count on PND105 and -315 in male offspring. Asterisks indicate significantly different from respective time-matched control group: *, P < 0.05; **, P < 0.01. The results represent the mean ± SE. The number of litters analyzed for each treatment group is shown in Table 2Go.

 
Cauda epididymal sperm count was decreased approximately 19% and 26% on PND105 and -315, respectively, in the 10 µg/kg group, although the difference was only significant on PND315 (Fig. 1BGo). There was a small, but dose-dependent, increase in beat-cross frequency that was significant (P < 0.05) in the 1 and 10 µg/kg groups on PND315 (data not shown). The number of motile sperm, sperm motility, velocity, linearity, and ALH displacement were not significantly affected by DES; however, these parameters were all lower, on the average, in the 10 µg/kg group relative to controls on PND105 and -315 (data not shown).

In vitro fertilizing ability
Fertilizing ability in vitro was measured in duplicate dishes for each animal. The average difference between duplicate measurements in control animals was less than 12%, indicating that in vitro fertility measurements were reproducible. On PND105, there was a modest, but significant, decrease in sperm fertilizing ability in the 1 µg/kg group (P < 0.05) and the 10 µg/kg group (P < 0.001; Table 2Go). There was no significant change in the number of fragmented eggs, but there was a large and significant increase (P < 0.001) in the number of fertilized eggs in the 10 µg/kg group that did not proceed to the two-cell stage. There was also a slight, but significant, decrease (P < 0.05) in the number of one-cell fertilized eggs in the 1 µg/kg group. The significant decrease in sperm fertilizing ability in the 10 µg/kg group persisted to PND315 (P < 0.05). However, there was an unexpected large increase in fertilizing ability in the 0.1 µg/kg group that was significant (P < 0.001). There was a significant decrease in the percentage of one-cell fertilized eggs in the 0.1 and 10 µg/kg groups; however, this was due to the absence of any one-cell fertilized eggs observed in these groups. There was also a slight, but significant, decrease (P < 0.01) in the number of fragmented eggs in the 0.1 and 10 µg/kg groups.


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  		Table 2. In vitro fertilizing (IVF) ability of epididymal sperm from F1 male mice exposed to diethylstilbestrol through gestation and lactation

 
Microarray analysis of testicular gene expression
Before assessing treatment-induced changes in testicular gene expression, it was necessary to first assess the experimental variation of the microarray assay. To assess the reproducibility of the cDNA labeling reaction and the effectiveness of normalization to remove the systematic dye bias, a homotypic hybridization was performed using the same adult testis RNA sample labeled with either Cy3 or Cy5. A scatter plot of the normalized Cy3 signal vs. the normalized Cy5 signal illustrates the reproducibility of the cDNA labeling reaction (Fig. 2AGo). Comparison with the scatter plot of the raw Cy3 signal vs. the raw Cy5 signal (Fig. 2BGo) illustrates the effectiveness of the normalization in removing the systematic dye bias. To assess the reproducibility of replicate spots within each array, a heterotypic hybridization was performed twice on different days using control testis RNA (Cy5) and adult kidney RNA (Cy3). Figure 2CGo illustrates the degree of reproducibility between the ratio measurements within the array, which was similar to the level of reproducibility between ratios measured on different arrays (Fig. 2DGo).



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  		Figure 2. Experimental variation within and between microarray assays. A, Scatterplot of normalized Cy3 and Cy5 signals. The same testis RNA sample was labeled with both Cy3 and Cy5 and hybridized to a single array. Raw signal intensity values were background-corrected, filtered to remove invalid spots, and normalized using a global mean approach, as described in Materials and Methods. The graph depicts data from a representative experiment that was repeated five times. B, Scatterplot of the raw unnormalized Cy3 and Cy5 signal intensity values from the experiment plotted in A. C, Scatterplot of log2 ratio values from duplicate spots within an array. The array was hybridized with cDNA synthesized from kidney (Cy3) and testis (Cy5) RNA. The graph depicts data from a representative experiment that was repeated twice. D, Scatterplot of average log2 ratio values from two different arrays that were both hybridized with cDNA synthesized from kidney (Cy3) and testis (Cy5) RNA.

 
To assess the degree of biological variation in gene expression, a heterotypic hybridization was performed using testis RNA from two 15-wk-old control animals, each from different litters. The heterotypic hybridization was repeated five times using 10 different control animals. A representative scatter plot of the normalized Cy3 signal vs. the normalized Cy5 signal illustrates the high degree of relatedness between the two samples (Fig. 3AGo). The average correlation coefficient (0.94 ± 0.05; n = 5) was lower than the average correlation coefficient of the homotypic hybridizations (0.97 ± 0.01; Fig. 2AGo), suggesting that variability in gene expression between animals was greater than experimental variation alone. To quantitatively compare the correlation coefficients, a Fisher’s z transformation was used to normalize the correlation coefficients and create 95% confidence intervals for r. The 95% confidence interval for the homotypic hybridizations (0.966 <= r <= 0.971; Fig. 2AGo) was greater than the 95% confidence interval for the control vs. control hybridizations (0.935 <= r <= 0.943; Fig. 3AGo), thus confirming that the biological variation is larger than the experimental variation.



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  		Figure 3. Biological and treatment-induced variation in testicular gene expression. Scatterplots are representative graphs from replicate heterotypic hybridizations comparing gene expression in control vs. control animals on PND105 (A; n = 5), control vs. DES animals on PND21 (B; n = 5), control vs. DES animals on PND105 (C; n = 9), and control vs. DES animals on PND315 (D; n = 4). Correlation coefficients (r) from each hybridization were averaged and used to calculate 95% confidence intervals for r.

 
This exploratory approach was extended to determine whether DES treatment had an effect on testicular gene expression. It was reasoned that an average correlation coefficient lower than that observed in control vs. control hybridizations was evidence for a treatment-related effect on testicular gene expression. Comparison of the scatter plots and the 95% confidence intervals for r for control (normalized Cy3) vs. DES (normalized Cy5) hybridizations (Fig. 3Go, B–D) with those of control vs. control hybridizations (Fig. 3AGo) illustrates the larger variation in gene expression due to treatment on PND21 and -105, but not PND315. This exploratory analysis suggests that DES significantly affected testicular gene expression on PND21 and -105, but not on PND315, at least for the genes represented on the array. To further test this hypothesis and to determine the gene- specific sources of variation, paired t tests were performed.

Test statistics were calculated for each gene on PND21, -105, and -315, and P values were used to as a means of prioritizing genes for verification by real-time PCR. Due to the small changes in gene expression and the biological variation, it was expected that the results of the t tests would produce a number of false positives despite the use of adjusted P values. On PND21 there were 644 genes with raw P values less than 5%; however, when the P values were adjusted, the expression of only one gene was significantly altered by DES (P < 0.05). This gene was proteasome subunit {alpha}4, which was repressed an average of 1.7-fold (log2ratio = 0.59 ± 0.16; n = 5). On PND105, 1257 genes with raw P values less than 5% were identified. Adjusted P values revealed 46 genes that were significantly altered by DES (P < 0.05; Table 3Go). Only 2 of the 46 genes were induced by DES (transcobalamin 2 and Duffy blood group). By contrast, the results of paired t tests on the control vs. control heterotypic hybridizations from PND105 revealed only 3 genes (expressed sequence tag, calmodulin, and IL-2 receptor {gamma}, all induced) that were significantly altered in expression (data not shown). On PND315, no genes were significantly altered in expression, which is consistent with the correlation analysis that suggested no treatment-related effects on gene expression (Fig. 3DGo), at least for the genes represented on the array. However, other critical, but unknown, genes may be altered on PND315, as significant changes in Sertoli cell number, epididymal sperm count, and sperm fertilizing ability in vitro were observed at this age. As an exploratory approach to determine whether there were any genes that showed evidence of long-term changes, all genes with raw P values less than 5% on PND21, -105, and -315 were cross-referenced. This resulted in 32 genes that were putatively altered on PND21, -105, and -315 (Table 4Go). Raw data and a complete list of all genes on the array and their normalized expression values, ratios, and P values can be found in the supplementary data file at http://www.bch.msu.edu/~zacharet/ microarray/supplemental/index.html.


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  		Table 3. Genes significantly altered in expression in the testis on PND105 after gestational and lactational exposure to diethylstilbestrol

 

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  		Table 4. Genes significantly (raw P < 0.05) altered in expression on PND21, 105, and 315 in male offspring exposed to diethylstilbestrol through gestation and lactation

 
Real-time PCR verifies changes in testicular gene expression
Although reasonable efforts were made to statistically identify genes that showed evidence of differential expression, the small changes and variability in gene expression measurements on the microarray still warrant verification of the results by an alternative and more quantitative method. Therefore, the expression of selected genes in Tables 3Go and 4Go were verified by real-time PCR. Additional genes were also chosen for analysis by real-time PCR (Table 1Go). All treatment groups (0, 0.1, 1, and 10 µg/kg) were analyzed by real-time PCR to examine dose-dependent effects. The intra- and interassay variations for the real-time PCR assay were less than 1% and 4%, respectively (data not shown). Despite the reproducibility of the assay, there was considerably more variance within treatment groups, as illustrated by the large error bars (Figs. 4Go and 5Go). This indicated large interanimal variation in gene expression, which is consistent with the variation in response to treatment among litters (data not shown). No attempts were made to remove outliers or litters that did not appear to respond to treatment before data analysis.



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  		Figure 4. Real-time PCR for validation of microarray results. The results represent the mean ± SE of six animals per treatment group; each from a different litter. NA, Not analyzed. *, P < 0.05; **, P < 0.01.

 


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  		Figure 5. Real-time PCR of selected genes. The results represent the mean ± SE of six animals per treatment group, each from a different litter. ND, Not detected. Note in E that the expression of ER{alpha} in all animals of the 10 µg/kg group on PND21 was below the limits of detection and therefore repressed below the control value. *, P < 0.05; **, P < 0.01.

 
Genes selected for verification based on observed changes on the microarray are shown in Fig. 4Go. Testicular receptor 2 (Tr2-11) is an orphan nuclear receptor expressed preferentially in advanced germ cell populations and is implicated in germ cell apoptosis in response to retinoic acid (57, 58). The expression of Tr2-11 was significantly increased 1.4-fold on PND105 (P < 0.01; Fig. 4AGo). Inhibin/activin ß C (Inhbc) is a member of the TGFß family of peptides, which are thought to have important paracrine roles in the testis in addition to their role in stimulating FSH secretion in the pituitary (59). Real-time PCR revealed that the expression of Inhbc on PND315 was significantly decreased 9.3- and 3.7-fold (P < 0.05) in the 0.1 and 10 µg/kg DES groups, respectively (Fig. 4BGo). Scavenger receptor class B1 (SR-B1) is a transmembrane protein that mediates selective cholesterol uptake by steroidogenic tissues, including Leydig cells (60). Real-time PCR indicated a significant 3.1-fold increase in SR-B1 expression on PND21 (P < 0.01; Fig. 4CGo). The major lysosomal membrane glycoprotein, LGP85, is thought to play a constitutive role in the lysosomal endocytotic pathway (61). Its expression was significantly decreased 3.0-fold in the 10 µg/kg group on PND21 (P < 0.01). Prosaposin (Psap; also known as Sgp-1) is a sphingolipid activator protein synthesized by Sertoli cells and thought to be involved in phagocytosis of residual bodies (62, 63). The expression of Psap was significantly increased 1.5-fold on PND105 (P < 0.05) in the 10 µg/kg group (Fig. 4EGo). The abdominal B-related Hoxa10 gene is involved in normal development of the male reproductive tract, including testicular descent and development of accessory sex organs (64, 65). There was a significant 5-fold decrease in expression on PND105 (P < 0.05), but not on PND315 (Fig. 4FGo). Due to limiting amounts of RNA, the expression of Hoxa10 was not analyzed on PND21.

The expression of SF-1 was examined by real-time PCR due to the role of SF-1 in regulating the expression of SR-B1 (66), the presence of SF-1 response elements in the Tr2-11 (accession no. U96095) and Hoxa10 (accession no. AF246720) promoters (unpublished observations), as well as the reported down-regulation of SF-1 after in utero exposure to DES (23). The expression of three SF-1 target genes in the steroidogenic pathway was also examined; 17{alpha}-hydroxylase/C17,20-lyase (Cyp17), cytochrome P450 side chain cleavage (Cyp11a), and steroidogenic acute regulatory protein (Star; Fig. 5Go, A–D). The expression of SF-1 was significantly decreased 2.1-fold on PND21 in the 10 µg/kg group (P < 0.01; Fig. 5AGo). Consistent with the decrease in SF-1, the expression of Cyp17, Cyp11a, and Star was significantly decreased 2.3-, 2.4-, and 1.7-fold, respectively, on PND21 (P < 0.05; Fig. 5Go, B–D). However, the expression of each of these genes was unexpectedly increased 1.6- to 1.7-fold on PND105 in the 10 µg/kg group (P < 0.05).

The expression of the nuclear receptors ER{alpha}, ERß, and AR was measured by real-time PCR due to their crucial role in spermatogenesis. ER{alpha} expression was significantly decreased on PND21 in the 1 µg/kg group (P < 0.05; Fig. 5EGo). The expression of ER{alpha} was below detectable levels in the 10 µg/kg group, indicating that expression was also significantly decreased in this treatment group. The expression of ER{alpha} was also below detectable levels in all treatment groups on PND105 and -315 (Fig. 5EGo). Attempts were made to repeat the RT-PCR with more RNA and/or more input cDNA, but expression could not be reproducibly detected to perform statistical analysis. The expression of ERß and AR was not altered by DES treatment (Fig. 5Go, F and G), as were zona pellucida 3 receptor; Xlr-related, meiosis-regulated receptor; CD36 antigen; secreted acidic cysteine-rich glycoprotein; and Bcl-associated death promoter.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
This study has demonstrated a long-term decrease in the number of Sertoli cells, epididymal sperm count, and in vitro fertilizing ability in mice after gestational and lactational exposure to 10 µg/kg·d DES. These results underscore the importance of assessing sperm function, as changes in testes weight and histology or sperm motion and morphology do not adequately predict fertility in vivo or the fertilizing ability of sperm in vitro (21, 45). In the present study there was only one incidence of unilateral cryptorchidism and no evidence of hypoplastic testes or epididymides, or Mullerian remnants. Spermatogenesis was histologically normal, and reproductive development of the offspring was also grossly normal. Therefore, the long-term effects of DES on sperm fertilizing ability are probably due to effects on the development and maturation of the sperm via direct action on the germ cells, the supporting somatic cells of the testes, and/or the epididymides.

The molecular mechanisms responsible for the effects of DES on the testis and spermatogenesis remain elusive, and few estrogen-responsive genes have been described in the testis. Studies by Majdic et al. (22, 23) have demonstrated that in utero exposure to DES alters the expression of SF-1 and Cyp17 in the Leydig cells of fetal rats, suggesting that the effects of DES on spermatogenesis are at least in part a result of inhibition of steroidogenesis. Basal and cAMP-induced expression of Cyp17, Cyp11a, and Star are regulated by SF-1 (67, 68, 69). Therefore, the decreased expression of these genes in the current study is consistent with the decrease in SF-1 mRNA and further supports the evidence that fetal exposure to DES disrupts steroidogenesis in the testis. Neonatal exposure to high doses of DES has also been shown to affect the expression of testicular inhibin {alpha}, Psap, and AR in prepubertal rats (70, 71). The results of the present study add support to the hypothesis that the adverse effects of DES are a result of changes in testicular gene expression. For example, early and latent alterations in the expression of genes involved in estrogen signaling (ER{alpha}), steroidogenesis (SF-1, Star, Cyp17, Cyp11a, and SR-B1), lysosomal function (LGP85 and Psap), and regulation of testicular development (Tr2-11, Inhbc, and Hoxa10) were observed in prepubertal and adult mice, even long after the cessation of DES exposure. Decreased expression of these genes may contribute, together or in part, to the decrease in epididymal sperm count and sperm fertilizing ability as a result of decreased testicular synthesis of C19 steroids or suppression of ER{alpha} signaling in the Leydig cells, both of which are essential for fertility (72). A third mechanism may involve altered function or proliferation of the supporting Sertoli cells of the testis, leading to malfunction of spermatogenesis and/or alterations in the responsiveness of germ cells to maturation cues, which are necessary for the acquisition of sperm fertilizing ability. Possible effects on gene expression in the efferent ducts and/or epididymis also cannot be ruled out, as they play crucial roles in the maturation of spermatozoa. In addition to direct effects on the fetal testis, the impaired reproductive performance of the dams and the apparent intrauterine death of the offspring may indirectly impact the reproductive development of the offspring due to maternal stress (73).

The size of the testis, the number of Sertoli cells, and germ cell production in adulthood are highly correlated with the number of Sertoli cells produced during the perinatal period, which is positively controlled by FSH (74, 75, 76). As expected, the relative decrease in Sertoli cell count in the present study (17% and 32% on PND105 and -315, respectively) was reflected by a comparable decrease in epididymal sperm count (19% and 26%). Although FSH levels were not measured in this study, the decrease in the number of Sertoli cells in the 10 µg/kg group is consistent with a decrease in perinatal levels of FSH and supports the hypothesis of Sharpe and Skakkebaek (77) that estrogens can inhibit sperm production in adulthood by decreasing Sertoli cell proliferation during the perinatal growth phase. However, the suppression of gonadotropin secretion alone is not sufficient to explain the effects on sperm fertilizing ability. Even after normalizing the number of sperm per dish in the IVF assay, decreases in sperm fertilizing ability were still observed. Furthermore, it has been shown that the effects of neonatal DES treatment on Sertoli cell maturation and efficiency of spermatogenesis in adulthood are distinct from those induced by neonatal treatment to a GnRH antagonist and may involve changes in Sertoli cell and/or Leydig cell gene expression (70, 78). The microarray and real-time PCR results further support this hypothesis.

The effects on sperm fertilizing ability are difficult to explain because the critical effects induced by DES may reside in one of many possible cell types within and outside of the testis, all of which contribute to sperm fertilizing ability. Furthermore, the effects may not be evident histologically or may have occurred during periods of development not examined. The long-term effects on sperm fertilizing ability may indicate that DES acts as a mutagen and irreversibly changes the genetic program of the germ cells (reviewed in Ref. 40). Alternatively, there is evidence supporting long-term epigenetic alterations after developmental exposure to DES, which may involve hypomethylation of estrogen- responsive genes (79, 80). For example, administration of the methylation blocker 5- azacytidine to adult rats also caused a decrease in fertility and an increase in the number of abnormal embryos on gestational d 2 (81). Abnormal imprinting of DNA methylation patterns has also been proposed as a possible mechanism for the transgenerational effects of DES (41). Therefore, abnormal imprinting of germ cell DNA during gametogenesis may explain the decrease in sperm fertilizing ability and the higher proportion of eggs that did not proceed to the two-cell stage in the 10 µg/kg group on PND105 (Table 2Go).

Studies in knockout mice have demonstrated a role for ER{alpha} and E2 in controlling sperm fertilizing ability in vitro and fertility in vivo (35, 36). Prenatal DES exposure also decreases the responsiveness of the uterus to prepubertal estrogen stimulation (31). Therefore, prenatal DES exposure may also reduce the responsiveness of the male reproductive tract to estrogens, thereby causing phenotypic changes comparable to those observed in ER{alpha} or aromatase knockout mice. This may occur through down-regulation of ER{alpha} mRNA or through suppression of E2 production by virtue of the decrease in mRNA expression of steroidogenic enzymes. The reduced sperm count and in vitro sperm fertilizing ability in both the DES-exposed mice and ER{alpha} knockout mice (35) suggest a causal link between down-regulation of ER{alpha} and the adverse effects on the testis.

Although a number of changes in gene expression were observed in this study, only a limited number of observations can be adequately addressed due to space considerations. DES caused a transient down-regulation of testicular LGP85 mRNA on PND21, but not on PND105 or -315. LGP85 is a major lysosomal membrane glycoprotein that appears to be expressed constitutively in all mouse tissues (82), although the function of this protein is unknown. As a lysosomal marker, the decrease in LGP85 expression may reflect a general decrease in lysosomal biogenesis, which could have functional consequences in cellular autophagy by Sertoli cells, an important process for the phagocytosis of residual bodies. It is interesting to note that lysosomes and other organelles of the endocytotic pathway are less developed in the nonciliated cells of the efferent duct in ER{alpha} knockout mice (83). If ER{alpha} also plays a role in regulating components of the endocytotic pathway in the testis, the decrease in ER{alpha} expression in the testis of DES-exposed mice on PND21 could explain the decrease in LGP85 mRNA at this time point.

Developmental exposure to DES causes a latent decrease in the expression of Hoxa10 in the adult testis. Homozygous knockouts of Hoxa10 manifest bilateral cryptorchidism due to developmental abnormalities of the gubernaculum, which ultimately results in defects in spermatogenesis and sterility (64, 84). Altered fetal expression of Hoxa10 in the testis may play a role in DES-induced cryptorchidism, as previously observed (9, 10, 17). It is interesting to note that Hoxa10 expression is disrupted in the uterus of adult Wnt-7a knockout mice (85), and that uterine expression of Wnt-7a is down-regulated in fetal mice exposed in utero to DES (27). The link between DES-induced down-regulation of Wnt-7a and its role in maintaining adult expression of Hoxa10 leads to the hypothesis that developmental exposure to DES may cause reproductive abnormalities in gubernaculum and uterine development as a result of disruption of the Wnt-7a/Hoxa10 pathway. Estrogen response element half-sites have also been reported in the mouse Hoxa10 promoter, thus raising speculation that disruption of estrogen signaling may persist to adulthood, causing a decrease in Hoxa10 expression (86).

In summary, the data described in this report demonstrate the potential for developmental exposure to DES, and possibly other estrogenic chemicals, to irreversibly alter testicular growth, sperm function, and testicular gene expression, including genes involved in steroidogenesis, lysosomal function, and testicular development. Establishing effects at the physiological, tissue, and cellular levels and associating them with changes at the molecular level allow us to explore and generate working hypotheses explaining the molecular mechanisms of developmental reproductive toxicants. Similar studies assessing the effects of other potent (17{alpha}-ethynyl estradiol) and weak (genistein) estrogen agonists on testicular development, sperm function, and testicular gene expression are ongoing. Comparisons to DES will allow us to 1) establish biomarkers of effect for estrogens on testicular development and sperm function, 2) determine agonist- specific effects on the testis, and 3) strengthen or dispute the hypotheses regarding the mechanism(s) of action of EEDs on male reproductive health.


   Acknowledgments
 
We thank Experimental Pathology Laboratories for histopathological analyses, P. S. Medadee, M. L. Hodges, M. Lloyd, R. S. Heck, S. L. Van De Wile, and V. B. Hardy for technical assistance with the Sertoli cell stereology; Dr. David Dix (U.S. Environmental Protection Agency, Research Triangle Park, NC) for providing sequence-verified cDNA clones; Drs Pam Green and Jeff Landgraf (Michigan State University, East Lansing, MI) for providing microarray printing facilities and assistance; and K. C. Fertuck, S. M. Samy, D. Boverhof, and L. D. Burgoon and other members of the laboratory for their valuable comments on the manuscript.


   Footnotes
 
This work was supported by a grant from the U.S. Environmental Protection Agency (R827-402-01-0, to K.C. and T.R.Z.) and Science Training Grant T32-ES-07255 (to R.G.H.) from NIEHS, NIH. T.R.Z. is supported in part by the Michigan Agricultural Experiment Station.

Abbreviations: AGD, Anogenital distance; ALH, amplitude of lateral head; AR, androgen receptor; CASA, computer-assisted digital image analysis system; Ct, threshold cycle; Cyp17, 17{alpha}-hydroxylase/C17,20-lyase; Cyp11a, P450 side chain cleavage; DES, diethylstilbestrol; E2, estradiol; EED, estrogenic endocrine disruptor; ER, estrogen receptor; Inhbc, inhibin/activin ß C; IVF, in vitro fertilization; NTC, no template control; PND, postnatal day; Psap, prosaposin; SF-1, steroidogenic factor 1; SR-B1, scavenger receptor class B1; Star, steroidogenic acute regulatory protein; Tr2-11, testicular receptor 2.

Received January 28, 2002.

Accepted for publication April 24, 2002.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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