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RESEARCH |
Department of Psychology, Neuroscience and Behaviour, McMaster University, Hamilton, Ontario, L8S 4K1 Canada
Correspondence should be addressed to D deCatanzaro; Email: decatanz{at}mcmaster.ca
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Abstract |
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Introduction |
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Exposure to adult males can induce early onset of sexual maturation in juvenile females of several mammals including mice (Vandenbergh 1967), rats (Vandenbergh 1976), lemmings (Hasler & Banks 1975), deer mice (Teague & Bradley 1978), voles (Spears & Clarke 1986), opossums (Harder & Jackson 2003) and cattle (Roberson et al. 1991), a phenomenon often called the Vandenbergh effect. Within 24 h of male presence, juvenile females begin to show changes in uterine growth followed by surges in serum luteinizing hormone and follicle-stimulating hormone by the third day (Bronson & Stetson 1973). Plasma oestradiol spikes after 6 h of male exposure, while progesterone levels show an influence after about 60 h (Bronson & Desjardins 1974). Following a few weeks of male exposure, cyclical changes in vaginal cytology are evident in samples from repeated vaginal flushing of male-exposed females (Vandenbergh 1967, 1976, Bingel 1972), and the uterine mass is substantially higher than in isolated females (Beaton et al. 2006). These traditional techniques of assessing precocious puberty are invasive, as blood sampling and human handling will influence endogenous steroid levels (deCatanzaro & MacNiven 1992) and repeated vaginal stimulation can introduce artefacts including pseudopregnancy (Diamond 1970) and several hormonal changes (Komisaruk & Steinman 1986). In the present study, we undertook to determine steroid profiles over time in urine collected non-invasively from developing female mice, following recently developed methods for this species (Muir et al. 2001, deCatanzaro et al. 2003, 2004, 2006, Beaton et al. 2006).
Phyto-oestrogens, commonly found in commercial rodent diets, can also significantly influence the onset of puberty (Thigpen et al. 1987, Takashima-Sasaki et al. 2006). These physiologically active compounds are similar in structure to 17β-oestradiol (Kurzer & Xu 1997) and are capable of binding to oestrogen receptors (Kuiper et al. 1998). Given the rich distribution of oestrogen receptors in female reproductive tissue (Cooke et al. 1997, Couse et al. 1997, Fitzpatrick et al. 1999, Jefferson et al. 2002), exposure to phyto-oestrogens can influence oestrogen-related action at the pre-pubertal uterus (Wang et al. 2005, Takashima-Sasaki et al. 2006) and ovaries (Jefferson et al. 2002, Takashima-Sasaki et al. 2006). These compounds are also able to cross the placental barrier (Brown & Setchell 2001, Ikegami et al. 2006), potentially having a significant effect on prenatal hormonal exposure.
The onset of sexual maturation in females can also be influenced by the hormonal milieu in utero (Clark & Galef 1988, Zehr et al. 2001). Testosterone produced by male foetuses in the third week of gestation (Pointis et al. 1980) can be passively transferred to adjacent foetuses (Even et al. 1992, vom Saal & Dhar 1992). A female foetus located between two male foetuses will be exposed to higher levels of androgens than a female surrounded by two females. Differences in intrauterine position have significant implications for developing females, as those located in between two males reach sexual maturity later (Clark & Galef 1988, Zehr et al. 2001) and have later occurrence of vaginal opening (Zielinski & Vandenbergh 1991) and lengthier cycles once oestrus ensues (vom Saal et al. 1981). Vandenbergh & Huggett (1995) have suggested the importance of segregating female subjects based on their prior in utero steroid hormone exposure to reduce variation found in many experimental designs. The ano-genital distance index (AGDI) is a non-invasive bioassay that can be used to assign females accordingly, as it correlates closely with prenatal exposure to androgens (Vandenbergh & Huggett 1995).
The present study was designed to investigate how proximity to adult male conspecifics, dietary phyto-oestrogens and AGDI interacts in sexual maturation of female mice. Repeated non-invasive urinary collections, as previously developed in this laboratory (Muir et al. 2001, deCatanzaro et al. 2003), were conducted throughout a 2-week exposure period. An enzyme-linked immunosorbent assay (ELISA) was used to measure oestradiol and progesterone levels in female urine samples. Following this exposure, females were killed and wet and dry uterine and ovarian mass was recorded.
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Discussion |
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Urinary steroid measures have the advantages of being non-invasive, as they are not altered by human handling and blood sampling. This permits repeated measures and profiling of individuals over time and across conditions (Muir et al. 2001, Vella & deCatanzaro 2001, deCatanzaro et al. 2003, 2004, 2006, Beaton et al. 2006). Unconjugated ovarian and testicular steroids are reliably measurable in the urine of this species, and in general, patterns of urinary excretion have reflected known systemic steroid trends. Levels of testicular and ovarian steroids are substantially reduced by castration (Vella & deCatanzaro 2001, deCatanzaro et al. 2003). Oestradiol and progesterone levels in urine are dynamic over days of the oestrous cycle and rise substantially during mid-pregnancy then decline in later pregnancy (deCatanzaro et al. 2003, 2004). Nevertheless, it remains possible that some variation in excreted levels is distinct from that of systemic levels, given dynamics of receptor binding and excretory mechanisms.
It is a usual practice to adjust urinary steroid output for creatinine in order to compensate for variation in hydration (Munro et al. 1991, Muir et al. 2001, deCatanzaro et al. 2003). We found that urinary creatinine was itself dynamic among conditions, tending to be lower and to decline progressively in male exposed as opposed to isolated females, and to be somewhat higher in females raised on the phyto-oestrogen-free diet. An influence of proximity to males was also reported elsewhere (Beaton et al. 2006). One possibility is that movements of male-exposed females were impeded due to presence of males in adjacent compartments. Isolated and group-housed mice have been found to have a variety of differences in hormone levels in previous experimentation (Brain 1975). In addition, fluid intake can also be influenced by social housing conditions, and urinary output in mice is also influenced by reproductive status (Drickamer 1995). Our major findings with urinary oestradiol and progesterone were observed both with and without creatinine adjustment. We suggest that it may not always be appropriate to adjust urinary steroid levels for creatinine, at least for this species, especially given that laboratory animals have constant access to water.
Urinary oestradiol was substantially reduced in females raised on the phyto-oestrogen-free diet. The amount of phyto-oestrogen content in diet has been shown to directly correlate with phyto-oestrogen concentrations in plasma and urine. Rodents fed diets high in phyto-oestrogen content also display high levels of isoflavone concentrations in their urine (Brown & Setchell 2001). The source of the difference in urinary oestradiol observed here is unknown. The antibodies used in our enzyme assay were polyclonal, and could possibly show some binding with oestrogenic components excreted in urine of females that were fed the regular diet. Nevertheless, the increased intra-individual variability in oestradiol with male exposure was observed despite any such possible direct influence of phyto-oestrogens on urinary measures.
The increase observed here in combined mass of uterus and ovaries among male-exposed females at age 43 is consistent with previous findings (e.g. Beaton et al. 2006). The current data also indicate that AGDI has a significant impact upon dry uterine and ovarian mass, with long-AGDI females having lesser reproductive tract mass. There was also some interaction of diet with this influence of AGDI upon dry uterine and ovarian mass, with the effect being most apparent in females raised on the phyto-oestrogen-free diet. Uterine growth over development and within oestrous cycles is highly influenced by oestrogens, with specific roles depending on timing and species (Gray et al. 2001). In mice, uterine cells proliferate in response to exogenous oestradiol (Ogasawara et al. 1983). Oestrogens regulate growth hormone and insulin-like growth factor-I (IGF-1) action (Kahlert et al. 2000, Leung et al. 2004), and local IGF-1 activity mediates uterine growth in response to oestradiol (Sato et al. 2002). Male urine also contains significant quantities of unconjugated oestradiol (Vella & deCatanzaro 2001, Beaton et al. 2006, deCatanzaro et al. 2006), and male mice actively direct urine droplets at proximate females (Reynolds 1971, deCatanzaro et al. 2006). Since oestrogens are established to play a critical role in male-induced female precocious puberty (Bronson 1975), oestrogens in male excretions could contribute to pubertal development in proximate females.
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Materials and Methods |
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Breeding and nutrition
Female subjects were selected from offspring of 44 CF-1 females fed regular laboratory chow and 40 others fed a nutritionally similar diet designed for very low levels of phyto-oestrogens (Advanced Protocol Verified Casein Diet 1 IF from Purina Mills Inc., Richmond, IN, USA. LabDiet obtained from Ren's Feed & Supply Ltd, Oakville, Ontario, Canada), commencing one week before pairing with a CF-1 breeder male. The ingredients of this ‘phyto-oestrogen-free’ diet are listed by the manufacturer as ground wheat, ground corn, wheat middlings, ground oats, fish meal, casein, corn gluten meal, corn oil, dicalcium phosphate, brewers dried yeast, plus vitamins and minerals, and is specified as consistently containing <1.0 ppm total isoflavones (aglycone equivalents of genistein, daidzein and glycitein).
Litters from regular-diet dams had an average of 11.8 pups, and were 48.8% female with mean body weight of 19.7 g and ano-genital distance of 9.66 mm at 28 days among female pups. Litters from phyto-oestrogen-free diet dams had an average of 11.4 pups, and were 44.3% female with mean body weight of 20.1 g and ano-genital distance of 9.49 mm at 28 days among female pups. Subjects were selected from among female pups with care to distribute representatives from specific litters in a counterbalanced fashion among all conditions, such that no condition had more than two females from a specific litter. Remaining pups were used in other experiments in this laboratory.
Ano-genital distance
Twenty-eight days following birth of litters, an AGDI was generated for each female. Individual ano-genital distance (mm) was determined using a Mastercraft digital caliper to measure the distance between base of the genital papilla and proximal end of the anal opening. An AGDI was calculated by dividing the ano-genital distance (mm) by body mass (g) and multiplying the resultant value by 100 (Vandenbergh & Huggett 1995). Care was taken to ensure that the ano-genital region was neither stretched nor compressed during measurements. All indices were calculated by a single experimenter well trained in animal handling. Since there have been no previous reports on AGDI in females of this age, we first calculated an index that ranged between 37.15 and 66.07. Females with AGDIs of 46 or lower were considered as having short AGDI, while those with an AGDI of 53 or higher were considered as having long AGDI. Remaining pups were excluded from subsequent experimentation.
Experimental treatment
Immediately following calculation of the AGDI, weanling females were each assigned to one of eight conditions in a 2x2x2 factorial design involving diet (regular versus phyto-oestrogen free), social situation (isolated versus male-exposed) and AGDI (short versus long). Among those reared on the regular diet, there were 12 isolated short-AGDI, 14 isolated long-AGDI, 12 male-exposed short-AGDI and 12 male-exposed long-AGDI females. Among those reared on the phyto-oestrogen-free diet, there were 12 isolated short-AGDI, 12 isolated long-AGDI, 11 male-exposed short-AGDI and 11 male-exposed long-AGDI females. Females raised on each specific diet continued on it for the remainder of the experiment.
Apparatus
Each female was placed in the lower compartment of a double-decker cage system described in deCatanzaro et al. (1996). Briefly, this apparatus was constructed from clear Plexiglas, measuring 30x21x27 cm, divided into upper and lower compartments (each measuring 30x21x13) by a stainless steel wire-mesh grid (squares of 0.5 cm2), which allowed for excretions from the males to pass into the females compartment. An opaque Plexiglas partition separated the males to prevent aggression. The female had full olfactory contact with each male but limited behavioural interaction. Each compartment provided continuous independent access to food and water, with food in the female's compartment protected from excretions of the males above. After a 48-h acclimation period to control for urinary hormone dynamics attributable to placement in the novel environment (deCatanzaro et al. 2004), daily urine collection began at day 30 and continued until day 43 of age. On each day of collection, the upper compartment containing the HS males was separated from the lower compartment containing the female for a period of 4±2 h. Each day, a clean tray was placed underneath each subject at the start of the dark phase of the light cycle. Pooled urine was then aspirated via 1-cc syringes with 26-gauge needles. A fresh syringe and needle were used for each animal on each day. Collection was extended up to 6 h wherever necessary to obtain a sufficient urine sample (
0.5 ml). Care was taken to ensure that urine samples were not contaminated with food residue or faeces, by only taking urinary samples clearly free of such deposits. All samples were stored in labelled 1.5 ml Nalgene cryo-tubes and frozen immediately after collection at –20 °C until they were assayed simultaneously for the whole experiment.
Uterine and ovarian mass measurement
After completion of urine collections on day 43 of age, females were killed using a lethal dose of CO2 gas. Ovaries and uterus were extracted through a dorsolateral incision made near the abdominal cavity. Excess fat and mesentery were removed as per standard criteria (Wang et al. 2005). All surgeries were performed by a single experimenter blind to each female's experimental condition. Dry weights were calculated after tissue samples were stored in calcium sulphate crystals for a period of 30 days at 3 °C.
Urinary steroid and creatinine determination
Validations of enzyme immunoassay procedures for this laboratory for adult male and female mice have been reported elsewhere for oestradiol (Muir et al. 2001) and for progesterone (deCatanzaro et al. 2003). Creatinine, 17β-oestradiol, and progesterone were obtained from Sigma Chemical Co. Antibodies to 17β-oestradiol and progesterone and corresponding horseradish peroxidase conjugates were obtained from the Department of Population Health and Reproduction at the University of California (Davis). Cross-reactivities for anti-oestradiol and anti-progesterone have been previously reported (deCatanzaro et al. 2004). The assay was carried out on Nunc Maxisorb plates which were first coated with 50 µl of antibody stock diluted at 1:10 000 in a coating buffer (50 mmol bicarbonate buffer/l; pH 9.6) and stored for 12–14 h at 4 °C. Wash solution (0.15 mol NaCl/l containing 0.5 ml of Tween 20/l) was added five times to each well using an automated strip ( model ELx50 Bio-Tek Instruments Inc., Winooski, VT, USA) to rinse away any unbound antibody and then 50 µl phosphate buffer per well were added. The plates were incubated at room temperature (21 °C) for 2 h for oestradiol determination and 1 h for progesterone determination before adding standards, samples or controls.
For oestradiol and progesterone, urine samples were diluted 1:9 in phosphate buffer before they were added to the plate. Standard curves were derived by serial dilution from a known stock solution. Oestradiol stock was 2000 pg/ml yielding values of 500, 250, 125, 62.5, 31.3, 15.6, 7.8, 3.9, 1.95, 0.98, 0.46 and 0.23 pg/ml. 5000 pg/ml progesterone standard stock was serially diluted to 2500, 1250, 650, 325, 162.5, 81.25, 40.63, 20.31, 10.16, 5.08, 2.54, 1.27 and 0.63. For all assays, 50 µl oestradiol or progesterone–horseradish peroxidase were added to each well, with 20 µl of standard, sample or control for oestradiol, or 50 µl of standard, sample, or control for progesterone. The plates were incubated for 2 h at room temperature. Subsequently, the plates were washed and 100 µl substrate solution of citrate buffer, H2O2 and 2,2'-azino-bis (3-ethylbenzothiazoline-6-sulphonic acid) were added to each well and the plates were covered and incubated while shaking at room temperature for 30–60 min. The plates were then read with a single filter at 405 nm on the microplate reader (Bio-Tek Instruments Inc., model ELX 808).
It is common practice with urinary steroid analyses to compensate for variations in fluid intake and output by adjusting sample values for creatinine (Munro et al. 1991, Muir et al. 2001, deCatanzaro et al. 2003). Standard creatinine values of 100.0, 50.0, 25.0, 12.5, 6.25, 3.12, 1.56, and 0.78 µg/ml were used. All urine samples were diluted 1:41 urine: phosphate buffer (0.1 mol/l sodium phosphate buffer (pH 7.0) containing 8.7 g of NaCl and 1 g of BSA per litre). Dynatech Immulon flat bottom plates were used and 50 µl per well of standard was added together with a solution of 50 µl distilled water, 50 µl 0.75 mol/l NaOH and 50 µl 0.4 mol/l picric acid. The plates were then shaken and incubated at room temperature for 30 min. The plate was measured for absorbance on a plate reader with a single filter at 490 nm. Standard curves were generated, regression lines were fit and the regression equation was applied to the optical density for each sample. Steroid measurements were adjusted for creatinine by dividing the obtained value by the measure of creatinine per ml or urine. Both creatinine-adjusted and unadjusted steroid measures were analysed statistically.
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Acknowledgements |
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TopAbstractIntroductionResultsDiscussionMaterials and MethodsAcknowledgementsReferences |
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Received 6 July 2007
First decision 24 August 2007
Accepted 24 September 2007
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References |
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TopAbstractIntroductionResultsDiscussionMaterials and MethodsAcknowledgementsReferences |
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