Evaluation of Prenatal Exposure to Bisphenol Analogues on Development and Long-Term Health of the Mammary Gland in Female Mice
2018; National Institute of Environmental Health Sciences; Volume: 126; Issue: 8 Linguagem: Inglês
10.1289/ehp3189
ISSN1552-9924
AutoresDeirdre K. Tucker, Schantel Hayes Bouknight, Sukhdev S. Brar, Grace E. Kissling, Suzanne E. Fenton,
Tópico(s)Effects and risks of endocrine disrupting chemicals
ResumoVol. 126, No. 8 ResearchOpen AccessEvaluation of Prenatal Exposure to Bisphenol Analogues on Development and Long-Term Health of the Mammary Gland in Female Mice Deirdre K. Tucker, Schantel Hayes Bouknight, Sukhdev S. Brar, Grace E. Kissling, and Suzanne E. Fenton Deirdre K. Tucker Curriculum in Toxicology, University of North Carolina at Chapel Hill, Chapel Hill, North Carolina, USA Division of the National Toxicology Program (DNTP), NTP Laboratory, National Institute of Environmental Health Sciences (NIEHS), National Institute of Health (NIH), Research Triangle Park, North Carolina, USA , Schantel Hayes Bouknight Charles River Laboratories, Inc., Durham, North Carolina, USA , Sukhdev S. Brar DNTP, Cellular and Molecular Pathology Branch, NIEHS, Research Triangle Park, North Carolina, USA , Grace E. Kissling Division of Intramural Research, Biostatistics and Computational Biology Branch, NIEHS, Research Triangle Park, North Carolina, USA , and Suzanne E. Fenton Address correspondence to S. E. Fenton, PhD, 111 T.W. Alexander Dr., MD E1-08, ResearchTriangle Park, NC 27709, USA. Telephone: (984) 287-4182. Email: E-mail Address: [email protected] Division of the National Toxicology Program (DNTP), NTP Laboratory, National Institute of Environmental Health Sciences (NIEHS), National Institute of Health (NIH), Research Triangle Park, North Carolina, USA Published:10 August 2018CID: 087003https://doi.org/10.1289/EHP3189Cited by:5AboutSectionsPDF Supplemental Materials ToolsDownload CitationsTrack CitationsCopy LTI LinkHTMLAbstractPDF ShareShare onFacebookTwitterLinked InRedditEmail AbstractBackground:Continued efforts to phase out bisphenol A (BPA) from consumer products have been met with the challenges of finding safer alternatives.Objectives:This study aimed to determine whether early-life exposure to BPA and its related analogues, bisphenol AF (BPAF) and bisphenol S (BPS), could affect female pubertal mammary gland development and long-term mammary health in mice.Methods:Timed pregnant CD-1 mice were exposed to vehicle, BPA (0.5, 5, 50mg/kg), BPAF (0.05, 0.5, 5mg/kg), or BPS (0.05, 0.5, 5mg/kg) via oral gavage between gestation days 10–17. Mammary glands were collected from resulting female offspring at postnatal day (PND) 20, 28, 35, and 56, and at 3, 8, and 14 months for whole mount, histopathological evaluation, and quantitative real-time polymerase chain reaction (qPCR); serum steroid concentrations were also measured at these time points.Results:In the bisphenol-exposed mice, accelerated mammary gland development was evident during early puberty and persisted into adulthood. By late adulthood, mammary glands from bisphenol-exposed female offspring exhibited adverse morphology in comparison with controls; most prominent were undifferentiated duct ends, significantly more lobuloalveolar hyperplasia and perivascular inflammation, and various tumors, including adenocarcinomas. Effects were especially prominent in the BPAF 5mg/kg and BPS 0.5mg/kg groups. Serum steroid concentrations and mammary mRNA levels of Esr1, Pgr, Ar, and Gper1 were similar to controls.Conclusions:These data demonstrate that prenatal exposure of mice to BPAF or BPS induced precocious development of the mammary gland, and that siblings were significantly more susceptible to spontaneous preneoplastic epithelial lesions and inflammation, with an incidence greater than that observed in vehicle- and BPA-exposed animals. https://doi.org/10.1289/EHP3189IntroductionThe mammary gland is an essential tissue that is required for lactation and infant nourishment. Like other female reproductive tissues, many hormones and growth factors influence mammary tissue, with the primary drivers depending on the stage of development. Steroid hormones regulate ductal growth, elongation, branching, and differentiation, and exposure to environmental chemicals that mimic or interfere with the action of endogenous hormones can alter normal mammary development (Rudel et al. 2011). The earliest stages of mammary development, during fetal mammary bud formation in late gestation and the primary stages of ductal branching that occur just prior to and after birth, have been reported as susceptible time points for the effects of endocrine-disrupting chemicals (EDCs) (Enoch et al. 2007; Macon et al. 2011; Padilla-Banks et al. 2006; Tucker et al. 2015). Studies of prenatal and perinatal exposure to human relevant doses of bisphenol A (BPA) in rodents have reported increased epithelial tissue growth, decreased apoptosis, decreased latency, and increased incidence of preneoplastic and neoplastic lesions in the mammary gland (Acevedo et al. 2013; Durando et al. 2007; Markey et al. 2001; Muñoz-de-Toro et al. 2005). Several rodent studies have also linked BPA exposure at human-relevant doses to neurobehavioral deficits (Dessi-Fulgheri et al. 2002; Palanza et al. 2008; Ryan and Vandenbergh 2006), reproductive alterations (Markey et al. 2005; Timms et al. 2005), and hepatic tumors (Weinhouse et al. 2014).BPA is a chemical commonly used in the manufacturing of plasticizers, epoxy resins, thermal paper, dental sealants, and linings of canned foods. The fourth National Report on Human Exposure to Environmental Chemicals (CDC 2017) suggested that the highest human urinary geometric mean for total BPA in all age groups occurred in the United States between 2003 and 2004, a time when BPA was detected in urine samples of >90% of the general population (Calafat et al. 2008; Ye et al. 2015). BPA has also been recovered in human maternal serum (Schönfelder et al. 2002), maternal urine (Gerona et al. 2016), amniotic fluid (Pinney et al. 2017), and fetal cord blood (Schönfelder et al. 2002; Todaka and Mori 2002), suggesting that BPA readily crosses the placenta to the unborn child. Distribution into milk in humans (Mendonca et al. 2014; Sun et al. 2004) and rodents (Doerge et al. 2010; Okabayashi and Watanabe 2010) has also been reported. Exposure to BPA during the fetal or perinatal period is of concern, based on many reports demonstrating BPA's estrogenic and endocrine-disrupting properties in both the toxicology and epidemiological literature (reviewed in Gore et al. 2015). There remains some controversy surrounding BPA's impact on human health; as human exposures are low when measured in urine, these chemicals are thought to be quickly cleared, and biological activity of BPA metabolites in humans is poorly understood (Vandenberg et al. 2013). However, numerous epidemiological studies report a range of health effects associated with BPA exposure, such as increased effects related to metabolic disease, neurobehavior, growth and development, and reproductive tissue anomalies or dysfunction (reviewed in Rochester et al. 2013 and Gore et al. 2015).In 2010, the World Health Organization (WHO 2010) estimated that breastfed infants between the ages of 0–6 months were exposed to 0.3μg/kg body weight (BW) BPA daily, and BPA levels in infants fed formula from polycarbonate bottles were estimated at eight times breast-fed concentrations, and those given canned formula in polycarbonate-free bottles were exposed to 0.5μg/kg BW (WHO 2010). This finding suggested that most of the exposure that infants acquired was through plastic bottles. Petitioning from baby bottle manufacturers prompted the FDA to ban the use of BPA in baby bottles and sippy cups in 2012 and phase out use in infant formula packaging in 2013 (U.S. FDA 2014). However, infants and children were also exposed through additional routes, including gestational and lactational transfer, inhalation and ingestion of contaminated dust, and oral exposures through food and beverage containers/films (Liao and Kannan 2013; Liao et al. 2012b; Mendonca et al. 2014). The combination of FDA's BPA use restrictions and voluntary recalls by some manufacturers have led to the integration of other bisphenol analogues that demonstrate similarity in structure and activity to BPA (Pelch et al. 2017) into paper- and food-containing products, dental sealants, or food-processing equipment.The fluorinated bisphenol analogue, Bisphenol AF (BPAF), was nominated for health effects assessment by the National Toxicology Program (NTP) in 2008 because estrogenic activity assays reported that BPAF (53.4 nM) is an agonist and binds to estrogen receptor alpha (ERα) with ∼20x greater affinity than BPA (1030 nM), and BPAF is a full antagonist for ERβ (Kitamura et al. 2005; Matsushima et al. 2010). Production of BPAF is 10,000–500,000 pounds per year and considered to be moderate (NTP 2008); but BPAF has been detected in contaminated air, soil, water, and sediment downstream of factories involved in fluoroelastomer production (Song et al. 2012; Wang et al. 2015). Female workers in the molding and casting machine operations industry are exposed to BPAF (NIOSH 1990), and BPAF was detected in the urine of a Chinese population downstream from a manufacturing plant (Yang et al. 2014). The general population may be exposed to BPAF through products used for dental sealants and composites, as well as food containers and films (Hashimoto et al. 2001; Liao and Kannan 2013).Bisphenol S (BPS), a common sulfated BPA analogue, has been measured in thermal receipt papers, currency bills, and canned foods, and it is also used as a modifier for leather fiber, for polymers, and as an epoxy curing agent (Liao and Kannan 2013; Liao et al. 2012c). Similar to BPA, BPS was detected in 100% of indoor dust samples from the United States, China, Japan, and Korea, indicating that oral ingestion, dermal exposure, and inhalation are all potential routes of exposure (Liao et al. 2012b). Human exposure to BPS has been confirmed by urinary measurements in the United States and seven Asian countries, with the highest concentrations found in Japan, followed by the United States (Liao et al. 2012a, 2012c). In a study of cashiers, an occupation that requires extensively handling receipt paper, urine BPS concentrations were significantly increased in post-shift measurements in comparison with pre-shift measurements (Thayer et al. 2016).Recent data from the Center for Disease Control (with human samples collected in 2013–2014; see CDC 2017, Volume 1, pp. 42–43) has shown an overall decline in the creatinine-adjusted urinary BPA geometric mean concentration in both sexes, and all ethnicities and age groups, between 2003 and 2014 (CDC 2017). Similarly, urinary measurements from convenience sampling of the U.S. general population between 2000 and 2014 showed a decline in the percent detection and geometric mean urinary BPA concentration, but the percentage of samples with detectable BPS, and the average concentrations measured, steadily increased, suggesting a change in exposure trends for BPS and BPA (Ye et al. 2015). Although BPAF was below the limit of detection (<LOD) in most urine samples tested (detected in 14 months when exhibiting abnormal cycles), and within 4 h of lights on, to minimize effects of hormonal fluctuation. Necropsy usually occurred between 0900 and 1100. Data from mice necropsied at 3, 8, and 14 months were reported herein, but accounting of all animals was made in Table S1.Mammary Gland Preparation and AnalysisThe fourth and fifth inguinal mammary glands were removed and processed for whole mounts, and contralateral glands were fixed in 10% neutral buffered formalin (Fisher Scientific) for histopathology. Fixed glands were embedded in paraffin and cut into 5-μm sections for hematoxylin and eosin (H&E) staining or immunohistochemistry. Whole mounts were prepared by flattening glands on a charged slide for 1 h, followed by fixation in Carnoy's solution overnight, rinsing in 70% ethanol and transitioning to water, staining with carmine alum (2g/L carmine and 5g/L aluminum potassium sulfate) overnight, rinsing in water, then increasing gradually to 100% ethanol, and finally defatting in xylene until visibly clear (Davis and Fenton 2013, Appendix). Qualitative and quantitative assessments were performed at PND 20 and 35, hallmark timepoints associated with weaning and puberty. Developmental scores were assigned separately by two individuals, using a scale from 1–4 (1=poor development and 4=best development) (Hilakivi-Clarke et al. 1997). Depending on age and level of development, scores were based on lateral and longitudinal epithelial growth, presence or absence of terminal end buds (TEBs), branching density, budding, and appearance of ductal ends. In whole mounts, TEBs were defined as ends that were ≥2x the diameter of its duct (Macon et al. 2011). Because the samples were compared with vehicle controls, the assessors were un-blinded to the vehicle control group, initially, and then they evaluated all glands within an age group blinded to treatment group.Longitudinal growth, mammary epithelial area (MEA), duct length, and TEB count (Macon et al. 2011) were quantitatively measured using ImageJ ( https://imagej.nih.gov/ij/). Glands were also quantitatively assessed for branching density using ImageJ and the modified Sholl analysis method (Stanko et al. 2015). A detailed visual description of this method is illustrated by Stanko and Fenton (Stanko and Fenton 2017). Briefly, images of glandular epithelium are skeletonized, binarized, dilated, and used to measure the MEA and longitudinal distance, defined as the most anterior position of the collecting duct to the most distal branch on the gland, using ImageJ. The total number of radial intersections (N) in the MEA was determined by Sholl analysis. The branching density of the gland was then calculated using the formula N/MEA. All quantitative measurements were performed on the fourth inguinal gland.Numerous undifferentiated terminal ends, presumed to be TEBs, were noted upon examining whole mounts at 3 months, which is unusual for this age of CD-1 mice (S. Fenton, oral communication, June 2016). Therefore, all whole mounted glands were visualized by light microscopy (Leica Z16 APO, Leica Microsystems), terminal epithelial duct ends were measured and if greater than 2 times the diameter of the duct, they were recorded as a TEB. Whole mounts from 8- and 14-month-old mice were also examined for abnormalities (potential lesions) using light microscopy as described above. Unusual development was recorded. It should be noted that the Sholl method and certain quantitative measurements were unable to be performed on glands of animals >3 months old because of extensive growth and tissue density.Contralateral fixed glands from mice of 3, 8, and 14 months old were embedded in paraffin, cut into 5-μm sections, stained with H&E, visualized on an Olympus BX41 (Olympus Scientific Solutions Americas Corp.) and digitally captured on an Olympus DP70 camera. A board-certified veterinary pathologist (S.H.B.) evaluated all histopathology samples and was blind to treatment.Neoplastic and nonneoplastic mammary lesions were diagnosed using standardized nomenclature proposed by the International Harmonization of Nomenclature and Diagnostic Criteria for Lesions in Rats and Mice (INHAND) (Rudmann et al. 2012). On occasions where a definitive diagnosis could not be made, a pathology peer review group was convened to resolve the diagnosis. Microscopic lesions were graded using a standard four-point scale of minimal, mild, moderate, and marked severity grade criteria (Rudmann et al. 2012), representing the extent of involvement and size within the tissue.Subsequent to evaluation and image capture of whole mounts from 3-, 8-, and 14-month time points, it was noted that the whole mounts contained gross abnormalities that were often not noted in the single section from the contralateral gland. Therefore, all mammary gland whole mounts from these ages were sectioned to observe the cellular environment surrounding the abnormal lesion. A detailed description and a visual tutorial of this method have recently been published (Tucker et al. 2016, 2017). Briefly, mammary whole mounts were immersed in xylene overnight to remove excess Permount (Fisher Scientific) and allow for ease of removing tissue from the slide. Glands were halved at the midline, placed in cassettes, and processed in a succession of xylene and xylene:molten paraffin steps before final embedding. Glands were sectioned at 4μm, stained with H&E and evaluated using the same criteria as the contralateral glands, by the same pathologist. The sectioned whole mount samples from 3-month-old mice were also evaluated for presence of retained TEB or intraductal hyperplasia, as stated above for formalin-fixed, contralateral glands in those same animals.Detection and Monitoring of Puberty and Estrous CycleBeginning on PND 16, all female offspring were checked daily for vaginal opening (VO) by trained observers, using the methods described in Goldman et al. (Goldman et al. 2007). Upon vaginal opening, lavage was conducted, using 1X PBS, pH 7.4 (Gibco), to determine the timing of first estrus and their estrous stage at VO. Morning lavage was continued and read daily until each animal exhibited estrus. Beginning when all animals had achieved VO and for the next 4 wk, soiled bedding from age-matched male control mice was collected into a homogenous mix and dispersed into all female cages weekly to circulate male urinary proteins known to promote cyclity. From PND 63 until PND 84, the same groups of females were reassessed for estrous cyclity by examining cytology using the above referenced methods (Goldman et al. 2007).Hormone AnalysisBriefly, following trunk blood collection in a BD Vacutainer SST Plus tube (BD Bioscience), the blood was inverted and stored at room temperature for 30 min. Samples were centrifuged at 1,100g for 15 min, and serum was collected in a fresh tube and stored at −80°C until processing.Serum preparation and analysis were performed per the manufacturer's protocols. Samples were run on a multiplex Steroid Hormone Panel Kit from Meso Scale Discovery (MSD) to determine estradiol, progesterone, testosterone, and dehydroepiandrosterone (DHEA) concentrations. Samples and standards were added to 96-well plates precoated with antibody and incubated for 2 h at room temperature. A SULFOTAG label tracer was added to each well, which generates a signal to determine the analyte concentration. The plates were washed three times with 1X PBS-T and after final wash, 150μL of a 1X Read-Buffer added to each well, and plates were immediately imaged on a Sector Imager 2,400 System (Meso Scale Discovery). All samples (50μL) were run in duplicate, on the same day. Standards for all hormones were supplied in the MSD reagent kit (lowest standards were 0.005, 0.07, and 2.7 ng/ml for estradiol, progesterone, and DHEA, respectively), except for testosterone (range 0.02 ng/ml–16 ng/ml), which was purchased from Steraloids, Inc. The coefficient of variation varied from 2% (E2) to 11% (T) for the four assay endpoints.RNA PreparationRNA was extracted from frozen mammary tissue by homogenizing the gland with Trizol using Lysing D Matrix tubes (MP Biomedicals). Samples were homogenized in a MP Biomedical Fast Prep-24 SG at 6.0m/ sec for 40-s intervals. Samples were homogenized a total of 2–3 times and placed on ice between each interval. Samples were transferred to a clean tube and centrifuged to remove debris and the lipid layer. RNA was isolated following the manufacturer's protocol. Samples underwent on-column DNAse I digestion using the RNA Mini Kit (Qiaqen). RNA quantity and integrity were measured on a Nano Drop 2000c (ThermoScientific) and Agilent Bioanalyzer. Samples with RNA integrity numbers ≥7.9 were used for PCR. One microgram of RNA was reverse transcribed with the High Capacity cDNA Synthesis Kit (Applied Biosystems) using the MyCycler (BioRad Thermocycler, Hercules). Complimentary DNA was amplified with Taqman Universal PCR Master mix, No AmpErase UNG, with the following Taqman primers (Roche): Esr1 (Mm00433149_m1), Pgr (Mm00435628_m1), Ar (Mm00442688_m1), Gper1 (Mm02620446_s1). Amplification was performed on a QuantStudio 7 Flex PCR (Applied Biosystems) and analyzed using QuantStudio Real-Time PCR Software and Microsoft Excel™ 2010 (Microsoft Office Suite). Mean Ct values ≥35 or with a standard deviation of ≥0.5 between duplicates were not included in the final analyses. Cdkna1 was used as the housekeeping gene for all samples. All analyses were performed using the 2−ΔΔCt method and are illustrated as the fold change relative to vehicle control.Statistical AnalysisUnless noted, all data are represented as mean±SEM. The dam was considered the unit of measurement. In nearly all cases, one pup per dam was sampled at any given time point. When multiple pups per dam were evaluated in the same analysis of quantitative endpoints, such as timing of VO and first estrus, mixed effects analysis of variance (ANOVA) with Dunnett's test was used to account for potential litter effects. TEB (or retained undifferentiated ends) occurrence was statistically evaluated using one-sided Cochran-Armitage trend tests. ANOVA with Dunnett's multiple comparisons test was applied to quantitative endpoints for which one pup per litter was assessed. The Mann-Whitney test was used to compare chemical dose group to control for Sholl analysis and mammary gland quantitative measurements. One-sided Jonckheere's test (SAS/STAT® 9.2 User's Guide) was performed to determine dose-related trends. Hormone measurements were log transformed prior to statistical analyses to improve normality. Mean severity scores were calculated for 3-, 8-, and 14-month lesions whenever applicable, and one-sided Fisher's exact tests were performed to compare lesion incidences in each dose group with the vehicle control group. Cochran-Armitage tests (SAS/STAT® 9.2 User's Guide) were used to analyze for trends across dose groups. All analyses were performed using SAS 9.3 by G.E.K. All Graphs and tables were generated using Microsoft Excel™ and GraphPad Prism, and statistical significances were denoted at p≤0.05.ResultsFemale Offspring BWBody weight was assessed in vehicle, BPA, BPAF, or BPS prenatally exposed female offspring necropsied on PND 20, 28, 35, and 56 and at 3, 8, and 14 months (Figure S1). Few differences among doses within a treatment group were observed; at PND 35, the BPAF 0.5mg/kg group was significantly smaller in comparison with controls and transitioned back to control-comp
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