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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

Endocrine disruptor chemicals (EDCs) represent a serious problem for organisms and for natural environments. Drosophila melanogaster represents an ideal model to study EDC effects in vivo. Here, we present methods to investigate endocrine disruption in Drosophila, addressing EDC effects on fecundity, fertility, developmental timing, and lifespan of the fly.

Abstract

In recent years there has been growing evidence that all organisms and the environment are exposed to hormone-like chemicals, known as endocrine disruptor chemicals (EDCs). These chemicals may alter the normal balance of endocrine systems and lead to adverse effects, as well as an increasing number of hormonal disorders in the human population or disturbed growth and reduced reproduction in the wildlife species. For some EDCs, there are documented health effects and restrictions on their use. However, for most of them, there is still no scientific evidence in this sense. In order to verify potential endocrine effects of a chemical in the full organism, we need to test it in appropriate model systems, as well as in the fruit fly, Drosophila melanogaster. Here we report detailed in vivo protocols to study endocrine disruption in Drosophila, addressing EDC effects on the fecundity/fertility, developmental timing, and lifespan of the fly. In the last few years, we used these Drosophila life traits to investigate the effects of exposure to 17-α-ethinylestradiol (EE2), bisphenol A (BPA), and bisphenol AF (BPA F). Altogether, these assays covered all Drosophila life stages and made it possible to evaluate endocrine disruption in all hormone-mediated processes. Fecundity/fertility and developmental timing assays were useful to measure the EDC impact on the fly reproductive performance and on developmental stages, respectively. Finally, the lifespan assay involved chronic EDC exposures to adults and measured their survivorship. However, these life traits can also be influenced by several experimental factors that had to be carefully controlled. So, in this work, we suggest a series of procedures we have optimized for the right outcome of these assays. These methods allow scientists to establish endocrine disruption for any EDC or for a mixture of different EDCs in Drosophila, although to identify the endocrine mechanism responsible for the effect, further essays could be needed.

Introduction

Human activities have been releasing into the environment a massive amount of chemicals, which represent a serious problem for organisms and for natural ecosystems1. Of these pollutants, it is estimated that about 1,000 different chemicals may alter the normal balance of endocrine systems; according to this property, they are classified as endocrine disrupting chemicals (EDCs). Specifically, based on a recent definition by the Endocrine Society, the EDCs are “an exogenous chemical, or mixture of chemicals, that can interfere with any aspect of hormone action”2. Over the last three decades, there has been growing scientific evidence that EDCs can affect the reproduction and development of animals and plants3,4,5,6,7,8. Further, EDC exposure has been related to the increasing prevalence of some human diseases, including cancer, obesity, diabetes, thyroid diseases, and behavioral disorders9,10,11.

General mechanisms of EDC

Due to their molecular properties, EDCs behave like hormones or hormone precursors3,4,5,6,7,8,9,10,11,12. In this sense, they can bind to a hormone’s receptor and disrupt endocrine systems either by mimicking hormone activity or by blocking endogenous hormones binding. In the first case, after binding to the receptor, they can activate it as its natural hormone would do. In the other case, binding of the EDC to the receptor prevents the binding of its natural hormone, so the receptor is blocked and can no longer be activated, even in the presence of its natural hormone3. As a consequence, EDCs can affect several processes, such as the synthesis, secretion, transport, metabolism, or peripheral action of endogenous hormones that are responsible for the maintenance of homeostasis, reproduction, development, and/or behavior of the organism. Receptor binding is not the only way of action described so far for the EDCs. It is now clear that they can also act by recruiting coactivators or corepressors in enzymatic pathways or by modifying epigenetic markers deregulating gene expression10,11,12,13,14, with consequences not only for the current generation but also for the health of generations to come8.

Drosophila hormones

The potential effects of selected EDCs have been studied widely, both in wildlife species and in several model systems in which endocrine mechanisms are reasonably well known. For invertebrates, endocrine systems that influence growth, development, and reproduction have been extensively characterized in insects for several reasons, involving their extensive use in the field of biological research, their economic importance, and finally the development of insecticides able to interfere specifically with the hormone system of pest insects.

In particular, among insects, the fruit fly D. melanogaster has proven to be a very powerful model system to evaluate the potential endocrine effects of EDCs. In D. melanogaster, as well as in vertebrates, hormones play an important role throughout the entire life cycle. In this organism, there are three main hormonal systems, which involve the steroid hormone 20-hydroxyecdysone (20E)15,16, the sesquiterpenoid juvenile hormone (JH)17, and the neuropeptides and peptide/protein hormones18. This third group consists of several peptides discovered more recently but clearly involved in a huge variety of physiological and behavioral processes, such as longevity, homeostasis, metabolism, reproduction, memory, and locomotor control. 20E is homologous to cholesterol-derived steroid hormones such as estradiol, while JH shares some similarities with retinoic acid; both of them are the better-known hormones in Drosophila19,20. Their balance is vital in coordinating molting and metamorphosis, as well as in controlling several postdevelopmental processes, such as reproduction, lifespan, and behavior21, thus offering different possibilities for testing endocrine disruption in Drosophila. Further, ecdysteroid hormones and JHs are the main targets of the so-called third-generation insecticides, developed to interfere with developmental and reproductive endocrine-mediated processes in insects. The agonist or antagonist mode of action of these chemicals is well known, and thus they can serve as reference standards for evaluating the effects of potential EDCs on the growth, reproduction, and development of insects22. For example, methoprene, which has been widely used in controlling mosquitoes and other aquatic insects23,24, works as a JH agonist and represses 20E-induced gene transcription and metamorphosis.

In addition to hormones, the nuclear receptor (NR) superfamily in Drosophila is also well known; it consists of 18 evolutionarily conserved transcription factors involved in controlling hormone-dependent developmental pathways, as well as reproduction and physiology25. These hormone NRs belong to all six NR superfamily subtypes, including those involved in neurotransmission26, two for retinoic acid NRs, and those for steroid NRs that, in vertebrates, represent one of the primary targets of EDCs27.

Drosophila as a model system for studying EDCs

Currently, on the basis of molecular properties, several environmental agencies around the world are attributing the potential to interfere with the endocrine systems to different man-made chemicals. Given that the EDCs are a global and ubiquitous problem for the environment and for organisms, the overall goal of the research in this field is to reduce their disease burden, as well as to protect living organisms from their adverse effects. In order to deepen the understanding about the potential endocrine effects of a chemical, it is necessary to test it in vivo. To this end, D. melanogaster represents a valid model system. To date, the fruit fly has been extensively used as in vivo model to evaluate the effects of several environmental EDCs; it has been reported that exposure to several EDCs, such as dibutyl phthalate (DBP)28, bisphenol A (BPA), 4-nonylphenol (4-NP), 4-tert-octylphenol (4-tert-OP)29, methylparaben (MP)30, ethylparaben (EP)31,32, bis-(2-ethylhexyl) phthalate (DEHP)33, and 17-α-ethinylestradiol (EE2)34, influences metabolism and endocrine functions as in vertebrate models. Several reasons have led to its use as a model in this field of research. Beyond an excellent knowledge of its endocrine systems, further advantages include its short lifecycle, low cost, easily manipulable genome, a long history of research, and several technical possibilities (see the FlyBase website, http://flybase.org/). D. melanogaster also provides a powerful model for easily studying transgenerational effects and population responses to environmental factors8 and avoids ethical issues relevant for in vivo studies in higher animals. In addition, the fruit fly shares a high degree of gene conservation with humans which might make it possible for Drosophila EDC assays to help in predicting or suggesting potential effects of these chemicals for human health. Besides expanding the understanding about human health effects, Drosophila can help to assess risks of EDC exposure to the environment, such as biodiversity loss and environmental degradation. Finally, the fruit fly offers the additional advantage of being used in laboratories, where the factors potentially affecting its development, reproduction, and lifespan can be kept under control in order to attribute any variation to the substance to be tested.

With this in mind, we have optimized simple and robust fitness assays for determining EDC effects on some Drosophila hormonal traits, such as fecundity/fertility, developmental timing, and adult lifespan. These assays have been widely used for some EDCs23,24,25,26,27. In particular, we have used the following protocols to evaluate the effects of the exposure to the synthetic estrogen EE234 and to BPA and to bisphenol AF (BPA F) (unpublished data). These protocols may be easily modified to investigate the effects of a given EDC at a time, as well as the combined effects of multiple EDCs in D. melanogaster.

Protocol

1. Food Preparation

  1. For stock maintenance and for larval growth, use a cornmeal medium containing 3 % powdered yeast, 10 % sucrose, 9 % precooked cornmeal, 0.4 % agar, thereafter called Cornmeal medium (CM).
    1. Put 30 g of yeast into 100 mL of tap water, bring it to a boil and let it boil for 15 min.
    2. Separately, mix well 90 g of precooked cornmeal, 100 g of sugar, and 4 g of agar into 900 mL of tap water.
    3. Bring the solution to a boil, lower the heat and cook for 5 minutes stirring continuously.
    4. After 5 minutes, add the hot yeast solution and simmer for another 15 min.
    5. Turn off the heat source and allow the solution to cool to about 60 °C.
    6. Add 5 mL/L of 10% methyl 4-hydroxybenzoate in ethanol, mix thoroughly and let it sit for 10 min.  
      NOTE: Be careful with the amount of methyl 4-hydroxybenzoate, given that a high concentration of fungicide could be lethal for larvae.
    7. Dispense the medium into vials/bottles:  8 mL into each fly vial (25 mm x 95 mm), 3 mL into each fly vial (22 mm x 63 mm) and 60 mL into each fly bottle (250 mL). 
    8. Cover vials with cheesecloth and allow them to dry at room temperature (RT) for 24 h prior to storage.
    9. Calibrate experimentally right consistency and hydration of the CM by modifying either the amount of agar used and/or the cooling/drying times.
      Note: unplugged, boxed and wrapped vials are stable for about 15 days at 4 °C. 
  2. For Drosophila adults, use a medium containing 10% powdered yeast, 10% sucrose, 2% agar, thereafter called adult medium (AM).
    1. Mix 10 g of powdered yeast, 10 g of sucrose, 2 g of agar into 100 mL of distilled water.
    2. Bring this mixture to a boil two times, with a 3 minutes interval, or until agar is dissolved, by using a microwave. 
    3. Once the solution cools to 60 °C, add 5 mL/L of 10% Methyl 4-hydroxybenzoate in ethanol, mix thoroughly and dispense in vials (10 mL per vial).
    4. Cover vials with cheesecloth and let them dry at RT for 24 h before storing. 
      NOTE: unplugged, boxed and wrapped vials are stable for about 15 days at 4 °C.
  3. For fecundity/fertility assay, use Drosophila tomato juice-cornmeal medium.
    1. Pour off 70 mL of warm cornmeal food into a 100 mL beaker and add 30 mL of tomato juice (30% v/v).
    2. Mix thoroughly with a food processor and pipet 3 mL in small vials.
    3. Cover vials with cheesecloth and allow them to dry at RT for 24 h prior to storage. 
      NOTE: unplugged, boxed and wrapped vials are stable for about 15 days at 4 °C.
  4. For embryo collection, use agar plates, containing 3% agar, 30% tomato sauce, and 3% sugar. 
    NOTE: be careful not to make bubbles when pouring the medium in the plates.

2. Drosophila EDC Dosing

  1. Prepare an appropriate stock solution dissolving the selected EDC in the suitable solvent. For the EE2 (molecular weight 296.403), dissolve 1.48 g in 10 mL of 100% ethanol to make a 0.5 M stock solution and store at -80 °C.
    CAUTION: EDCs are considered environmental pollutant and precautions should be taken in handling them. 
  2. Dilute the EE2 stock solution in 10% ethanol in water (v/v) in order to obtain a 100 mM solution. Make next dilutions (0.1 mM, 0.5 mM and 1 mM) in CM food, starting with lowest concentration and using the same final concentration of solvent for each treatment group. For the control vials use same volume of the solvent alone. 
    Note: it is recommended to keep the final concentration of the solvent as low as possible, bearing in mind that final concentration of ethanol should not exceed 2% in fly food.
  3. Add the solution containing the right dilution of the selected EDC to the cornmeal-based food before solidification, mix thoroughly with a food processor, dispense 10 mL into vials, cover with cotton gauze and let dry at RT for 16 h before using. 
    NOTE: use this medium immediately after its preparation.
  4. For the adult rearing, prepare different working EE2 solutions (10 mM, 50 mM and 100 mM, respectively) in 10% ethanol in water (v/v) and layer 100 µL of each onto the surface of the AM, in order to obtain the desired concentration of the EE2 (0.1 mM, 0.5 mM and 1 mM). For the control use same volume of the solvent alone.
    NOTE: alternatively, add the solution containing the right dilution of the selected EDC to a small amount of AM in a 50 mL conical tube, vortex thoroughly and stratify 1 mL of it onto the surface of the AM vials.
    1. Cover vials with cotton gauze, allow drying at RT for 12-16 h under gentle agitation and use them immediately. 
      NOTE: drying process should be adjusted experimentally because is depending on the ambient humidity.
  5. For feeding assay, add both the solution containing the right dilution of the selected EDC (EE2 0.1 mM, 0.5 mM and 1 mM) and a coloring food (e.g., the red dye no. 40 at 1 mg/mL)35 to the CM before solidification, mix strongly with a food processor and then dispense into vials.

3. Rearing Flies

  1. Use a robust isogenic strain, such as Oregon R, maintained by several generation in the laboratory. 
  2. Keep flies in a humidified, temperature-controlled incubator, with a natural 12 h light: 12 h dark photoperiod at 25 °C in vials containing CM food.
  3. In each assay, use vials at RT.

4. Feeding Assay

NOTE: This assay is recommended to test if the presence of the selected EDC in the medium could affect feeding of flies.

  1. Put 15 young flies in vials containing CM supplemented with different concentrations of the selected EDC and a coloring food. Allow flies to feed on the media for 1 day.
    NOTE:for example, use red dye no. 4035 (1 mg/mL). 
  2. Put 15 young flies in vials containing CM supplemented with the solvent alone and a coloring food for control. Allow flies to feed on the media for 1 day. 
  3. Anesthetize individually each group of flies with ether.
    1. Transfer flies of each group to a cylindrical glass container (etherizer) with a funnel inserted into the open end, inverting the vial over the funnel and gently tapping the two containers together to make the flies fall into the etherizer. 
      NOTE:  The funnel will prevent them from getting out of the etherizer.
    2. Knock flies down by gently tapping the etherizer on a soft surface, such as a mouse-pad, and quickly replace the funnel with an ether-soaked cotton and gauze plug.
    3. Wait about 1 min until the flies fall to the bottom and stop moving. 
      NOTE: do not exceed the time or the flies will die.
  4. Put immobilized flies under a stereo-microscope and compare the abdominal coloring of each treatment group with respect to the control group. 

5. Fecundity/Fertility Assay

  1. For each EDC concentration, prepare 3 vials of flies, thereafter called parental vials, with 8 females and 4 males in 10 mL CM/EDC food; for the control prepare 6 vials of flies with 8 females and 4 males in 10 mL CM food supplemented with solvent. Rear flies in an incubator at 25 °C. 
    NOTE: avoid overcrowding larvae during their development and try to use consistent larval densities across treatments.
  2. After 4 days, remove parents and return the vials into the incubator for 5 more days.
  3. In the late afternoon of the day 9, remove all newly flies from the vials and put the vials in an incubator at 18 °C overnight. 
    NOTE: This removal must be done very carefully, checking the surface of the medium well.
    1. On the morning of day 10, for each treatment group, collect virgin females and young males into two groups, under light CO2 anesthetization. Randomly subdivide each group of flies in small subgroups (10 females and 20 males per vial) in independent vials filled with fresh corresponding CM. 
      1. Repeat step 5.3.1 taking care both to carefully remove all flies from the vials 8-10 h before collection and leave vials at 18 °C, until obtain at least 30 virgin females and 30 males for each EDC concentrations and at least 90 virgin females and 90 males for the control. 
    2. House these groups of flies at 25 °C until they are aged 4 days post-eclosion, transferring them into new vials containing fresh corresponding medium every two days.  
      Note: 4 days is sufficient time for the flies to become mature adults, but it is very far from the beginning of the senescence.
    3. After two days ensure there are no larvae in the vials of females. If they do, the flies are not usable because they are not virgins and must be discarded.
  4. Use 20 single flies of each sex for each treatment group to set up 20 single crosses into small vials containing fresh CM-tomato medium without EDC, as described below. 
    1. To each treatment group assign a different series of sequential numbers, which uniquely identifies it and label the respective vials; e.g., group1 (solvent alone) from 1 to 20, group 2 (EDC concentration x) from 20 to 40, and so on.
    2. Make a fertility spreadsheet to record the different series, each corresponding to a treatment group.
    3. For each sex anesthetize all the flies belonging to each treatment group under light CO2 and randomly transfer them as follows.
      1. Transfer one solvent-treated female into a small vial containing fresh CM-tomato medium without EDC and add one solvent-treated male for the control cross. 
        ​Note: tomato juice should be added to medium during its preparation because dark medium increases the contrast with the white embryos.
      2. Transfer one EDC treated female into a small vial containing fresh CM-tomato without EDC and add one solvent-treated male for each treatment.
      3. Transfer one EDC treated male into a small vial containing fresh CM-tomato medium without EDC and add one solvent-treated female for each treatment.
    4. House all these single crosses at 25°C.
  5. Transfer each mating pair into fresh CM-tomato vials without EDC every day for the subsequent ten days. Label the replicated vials of each series sequentially; e.g. 1-a, 1b, 1c……20a, 20b, 20c and report these number on the fertility spreadsheet. 
  6. Visually inspect each vial every day for the eggs and report their number on the fertility spreadsheet.
  7. Save each vial and, when newly flies start to emerge, also record the daily number of adult progenies over the 10 day period. After 10 days from the initial mating, remove parents. 
    NOTE: discard vial in which one or both parents died; in case of escape of one or both the parents, include in the analysis all data until the day them was lost.
  8. Sum the daily number of eggs and the daily number of adult progenies from each treatment group, to obtain the total fecundity/fertility, the mean egg and adult progeny production by a fly for ten days, and the ratio of total progeny to total number of eggs laid. Calculate the differences in percentage of each treatment values with respect to the control. 
  9. Carry out three independent experiments for each group of flies, by using a minimum of 10 flies for each treatment group. 
  10. Perform statistical analysis to compare the different groups. 

6. Developmental Timing

NOTE: In the two following alternative protocols the developmental timing is evaluated by counting both the number of pupae that form per day and the number of adult progeny eclosing per day.

  1. Eclosion assay protocol 1
    1. For each treatment group, set up 10 vials of young (<2 days), healthy flies, each with 6 females and 3 males in 10 mL cornmeal food without EDC.
    2. Let flies on food for 24 h, and allow them to mate.
    3. Prepare 10 parallel vials per treatment group with 10 mL each of fresh cornmeal food supplemented with different EDC concentrations or the solvent alone for the control. Transfer mated flies to these new vials. 
      NOTE: for each treatment group assign a different series of sequential numbers, which uniquely identifies it and label the respective vials.
    4. Make a developmental spreadsheet to record the different series.
    5. Allow flies to lay eggs for 16 h. Then remove parents from vials. 
      NOTE: The parent flies can be used to repeat the step 6.1.5, by transferring them to other corresponding vials.
    6. Incubate vials for 3-4 days at 25 °C, or until no more pupae form. Every day count the number of newly pupae in each vial and report it on the developmental spreadsheet. To avoid counting the same pupa twice, write a number in sequence at each pupa with a permanent marker on the outside of the vial. 
    7. Starting from the day 9, count daily the number of emerging adults until no more adults emerged, and report it on the developmental spreadsheet.
    8. From these raw data, calculate the mean larval period, the mean pupal period, as well as the differences in percentage of each treatment with respect to the control. 
    9. Carry out three independent experiments for each group of flies, by using a minimum of 5 vials for each treatment group.
    10. Perform statistical analysis to compare the different groups. 
  2. Eclosion assay protocol 2
    1. Rear young and healthy female (about 150) and male (about 50) flies on a collection cage (Table of Materials) with agar-tomato medium supplemented with fresh baker’s yeast paste (3 g of baker’s yeast in 5 mL of water), thereafter called laying tray, for 2 days at 25 °C.
    2. During these 2 days, allow flies to acclimate to the cage in a dark, quiet place, before beginning egg collection, and change the laying tray twice a day.
    3. On the third day, change the laying tray early in the morning. After 1 h, replace the laying tray, discarding these laid eggs. 
    4. Allow flies to lay eggs for 2 h and replace with fresh laying tray. 
      NOTE: By day 3, a good laying tray should produce 100-200 eggs in 2 h. 
    5. For each treatment group, prepare a series of three 60 mm dishes containing tomato cornmeal food supplemented with the corresponding EDC concentration or with solvent alone and report each series in the developmental timing spreadsheet. Alternatively, if preferred, use vials instead of dishes.
    6. Gently pick up eggs under a microscope by using a paintbrush or a probe and transfer them to the top of the medium in each dish/vial. In order to facilitate counting, on the laying tray, arrange eggs in 5 groups of 10 each and transfer them one at a time. 
      NOTE: Repeat step 6.2.4 as many times as is necessary to obtain enough embryos.
    7. House all these dishes/vials at 25 °C. Store also each laying tray at 25 °C, and count the total number of laid eggs.
    8. After 24-30 h, check each dish/vial under a stereomicroscope and count both the number of white, unfertilized eggs and the number of dark dead embryos. 
    9. Subtract the number of white, unfertilized eggs from the 50 transferred eggs value in order to obtain the ‘total embryos’ value per dish/vial. The number of dark dead embryos can be used to determine potential EDC toxic effects during embryogenesis.
    10. Repeat steps 6.1.6-6.1.10 of the Eclosion assay Protocol 1.

7. Lifespan Protocol

  1. Set up 20 vials of flies with 8 females and 4 males and house at 25 °C in CM (10 mL each). 
  2. After 4 days discard flies and place vials back in the incubator. 
    NOTE: these flies can be used to start again to obtain other age-synchronized cohorts of the flies. 
  3. In the late afternoon of the day 9, remove all newly flies from the vials and return vials to the incubator. 
    Note: a few adults should begin to eclose as early as the ninth day; discarding these flies allows to collect a maximum number of synchronized flies, avoiding the careless selection of early emergent. 
  4. 16-24 h later, transfer the adult flies (1 day old) of both sexes into four groups of 250 mL bottles containing AM food supplemented with three different EDC concentrations and one with the solvent alone. If needed, collect another batch the next day.
  5. Maintain flies at 25 °C for 2-3 days to allow them to mate. 
    NOTE: the day of transfer to AM food vials corresponds to the first day of adulthood.
  6. After two-three days, sort each cohort of flies by sex into two groups under light CO2 anesthetization. Randomly subdivide each sex into five vials per treatment at a density of 20 individuals per vial, until there are three replicates of 5 parallel vials for each gender per each treatment.
    NOTE: Work with small groups of flies in order to prevent possible long-lasting health issues due to long exposure time to CO2.
  7. Prepare a lifespan spreadsheet in which the number of dead flies is subtracted from the number of surviving flies to the previous transfer, in such a way as to automatically obtain the number of survivors at each transfer.
  8. Transfer flies into new vials containing the corresponding food every 3 days at the same time and check for death.
    NOTE: the transfer must take place without anesthesia that could have a long-term negative effect on fly longevity.
    1. At each transfer, record the age of the flies and the number of dead flies. 
      NOTE: the number of surviving flies is automatically calculated in the spreadsheet but it is recommended to check it visually. Flies that accidentally both escape or die during the transfer should not be considered. Be careful not to count twice dead flies carried to the new vial reporting this note in the spreadsheet.
    2. Repeat steps 7.8 and 7.8.1 until all flies die.
  9. For each treatment group, create a survival curve as shown in Figure 6, in order to display the survival probability of a fly at any particular time. 
  10. Perform three independent experiments for each treatment group of flies, by using 100 newly eclosed flies for each experiment. 
  11. Prepare a table in which to report the mean lifespan (mean survival days of all flies for each group), the half death time (period of time in days required to reach 50% mortality) and the maximum lifespan (maximum amount of days needed to reach 90% mortality). 
  12. Calculate the differences in percentage between each treatment group with respect to the control group. 
  13. Perform statistical analysis to compare the  different treatment groups.

Results

In this section, key steps of the above protocols are reported in the form of simplified schemes. Given that flies tend to avoid unpalatable compounds, the first thing to do is to assay the taste of the selected EDC. This can be done by mixing a food coloring (for example, red food dye no. 40)35 with the food supplemented with the selected EDC at various doses or with the solvent alone. Flies fed on these media are examined under a stereomicroscope and the food intake is estimated by their abdomin...

Discussion

The fruit fly D. melanogaster has been extensively employed as an in vivo model system to investigate the potential effects of environmental EDCs such as DBP28, BPA, 4-NP, 4-tert-OP29, MP30, EP31,32, DEHP33, and EE234. Several reasons have led its use as a model in this field of research. Apart from its undisputed advantages as a model ...

Disclosures

The authors have nothing to disclose.

Acknowledgements

The authors thank Orsolina Petillo for technical support. The authors thank Dr. Mariarosaria Aletta (CNR) for bibliographic support. The authors thank Dr. Gustavo Damiano Mita for introducing them to the EDC world. The authors thank Leica Microsystems and Pasquale Romano for their assistance. This research was supported by Project PON03PE_00110_1. “Sviluppo di nanotecnologie Orientate alla Rigenerazione e Ricostruzione Tissutale, Implantologia e Sensoristica in Odontoiatria/oculistica” acronimo “SORRISO”; Committente: PO FESR 2014-2020 CAMPANIA; Project PO FESR Campania 2007-2013 “NANOTECNOLOGIE PER IL RILASCIO CONTROLLATO DI MOLECOLE BIO-ATTIVE NANOTECNOLOGIE”.

Materials

NameCompanyCatalog NumberComments
17α-EthinylestradiolSigmaE4876-1G
Agar for Drosophila mediumBIOSIGMA789148
Bisphenol ASigma239658-50G
Bisphenol AFSigma90477-100MG
CornmealCA' BIANCA
Diethyl etherSigma
Drosophila VialsBIOSIGMA78900825x95 mm
Drosophila VialsBIOSIGMA78900929x95 mm
Drosophila VialsKaltek18722X63
Embryo collection cageCraftsPlexiglass cylinder (12,5 x7 cm) with an open end and the other end closed by a rectangular base in which a slot allows the insertion of special trays for laying
EthanolFLUKA2860
EtherizerCraftscylindrical glass container with a cotton plug
Glass Bottle250mL Bottles
Glass VialsMicrotechST 10024FLAT BOTTOM TUBE 100X24
Hand blender PimmyArietefood processor
Instant Success yeastESKAPowdered yeast
Laying trayCraftsplexiglass trays (11 x 2,6 cm) in wich to pour medium for laying
Methyl4-hydroxybenzoateSIGMAH5501
Petri DishFalcon35101660x5
Red dye no. 40SIGMA16035
Stereomicroscope with LED lightsLeicaS4E
SucroseHIMEDIAMB025
Tomato sauceCirio

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