Method Article

Protocols for Visualizing Steroidogenic Organs and Their Interactive Organs with Immunostaining in the Fruit Fly Drosophila melanogaster

DOI:

10.3791/55519

⸱

April 14th, 2017

 ,  ,  , 
1Graduate School of Life and Environmental Sciences, University of Tsukuba, 2Life Science Center of Tsukuba Advanced Research Alliance, University of Tsukuba, 3Faculty of Life and Environmental Sciences, University of Tsukuba

* These authors contributed equally

In This Article

Summary

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We describe a protocol for dissection, fixation, and immunostaining of steroidogenic organs in Drosophila larvae and adult females to study steroid hormone biosynthesis and its regulatory mechanism. In addition to steroidogenic organs, we visualize the innervation of steroidogenic organs as well as steroidogenic target cells such as germline stem cells.

Abstract

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In multicellular organisms, a small group of cells is endowed with a specialized function in their biogenic activity, inducing a systemic response to growth and reproduction. In insects, the larval prothoracic gland (PG) and the adult female ovary play essential roles in biosynthesizing the principal steroid hormones called ecdysteroids. These ecdysteroidogenic organs are innervated from the nervous system, through which the timing of biosynthesis is affected by environmental cues. Here we describe a protocol for visualizing ecdysteroidogenic organs and their interactive organs in larvae and adults of the fruit fly Drosophila melanogaster, which provides a suitable model system for studying steroid hormone biosynthesis and its regulatory mechanism. Skillful dissection allows us to maintain the positions of ecdysteroidogenic organs and their interactive organs including the brain, the ventral nerve cord, and other tissues. Immunostaining with antibodies against ecdysteroidogenic enzymes, along with transgenic fluorescence proteins driven by tissue-specific promoters, are available to label ecdysteroidogenic cells. Moreover, the innervations of the ecdysteroidogenic organs can also be labeled by specific antibodies or a collection of GAL4 drivers in various types of neurons. Therefore, the ecdysteroidogenic organs and their neuronal connections can be visualized simultaneously by immunostaining and transgenic techniques. Finally, we describe how to visualize germline stem cells, whose proliferation and maintenance are controlled by ecdysteroids. This method contributes to comprehensive understanding of steroid hormone biosynthesis and its neuronal regulatory mechanism.

Introduction

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In multicellular organisms, a group of cells is endowed with a specialized function in their biogenic activity that is essential for the whole body. To fulfill their missions, each tissue or organ expresses a series of genes related to their functions and communicates with other tissues to orchestrate their activities in the context of development. To characterize such specialized cellular functions and inter-organ interactions, we need to specify a group of cells along with other types of cells being kept intact in the multicellular architecture.

One example of such specialized organs is a steroidogenic organ, where many biosynthetic enzymes mediate the conversion steps from cholesterol to active steroid hormones1. Most of these enzyme genes are specifically expressed in steroidogenic organs, and the biosynthesis pathway is tightly regulated by many external stimuli via humoral inputs and neuronal inputs. Once synthesized, steroid hormones are secreted into the hemolymph and are targeted to many tissues and organs for regulating the expression of a variety of genes2. Therefore, the action of a steroid hormone induces a systemic response to maintain homeostasis, growth, and reproduction.

To investigate the functions of steroid hormone biosynthesis and the pleiotropic actions of steroid hormones, Drosophila melanogaster can be utilized as a suitable model system. During the larval stages, the insect steroid hormone, ecdysteroid, is biosynthesized in a specialized endocrine organ called the prothoracic gland (PG)3. In the PG, several ecdysteroidogenic enzymes specifically catalyze the multiple conversion steps from cholesterol to ecdysone, which controls molting and metamorphosis at the appropriate developmental stages4. Therefore, a dynamic change in ecdysteroid titer is regulated by many signaling pathways in response to environmental cues. On the other hand, in the adult stage, ecdysteroid plays essential roles in physiology, including reproduction, sleep, memory, and lifespan5,6,7,8. It is known that ecdysteroid is actively biosynthesized in the ovary, regulating the progression of oogenesis6,7,8,9,10,11. Recently we have reported that the number of germline stem cells (GSCs) is affected by ecdysteroid and sex peptide signaling in response to mating stimuli12.

Powerful tools of D. melanogaster genetics and cell biology, including well-annotated genome information, binary gene expression systems, and transgenic RNAi techniques, have enabled us to identify genes essential to ecdysteroid biosynthesis in the PG and the ovary13,14,15. Once the ecdysteroidogenic genes are identified, the transcriptional regulation of these genes and the dynamic localizations of gene products can be examined in the biosynthesis pathway16. For this purpose, quantitative-reverse transcription-PCR, RNA in situ hybridization, and immunohistological analysis are conducted. The application of these techniques includes a challenging task; the elaborate dissection of the PG or the ovary. In particular, the PG of the fruit fly is relatively smaller than that of other insects (e.g. the silkworm and the blow fly), so one needs to practice the vital skill of fruit fly dissection for sampling. Furthermore, both ecdysteroidogenic organs receive innervations from the central nervous system (CNS)17,18,19,20. Thus, for accurate anatomical analyses, the ecdysteroidogenic organs should be kept intact along with the CNS and other organs, not to disrupt their neuronal connections.

Here we provide protocols for the dissection and visualization of steroidogenic organs in D. melanogaster. Learning the dissection technique is the key starting point for these experiments. In addition, one can successfully label the steroidogenic organs as well as their interactive organs with several antibodies and GAL4 driver lines. Taking advantage of these techniques, materials, and genetics, one can study the comprehensive mechanisms of steroid hormone biosynthesis.

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Protocol

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NOTE: The overall scheme of protocols is shown in Figure 1.

1. The Dissection of the Larval Ring Gland (RG)

NOTE: In D. melanogaster, which belongs to cyclorrhaphous Diptera, the PG is within a composite endocrine organ called the ring gland (RG, Figure 2D). Since it is unfeasible that the PG is surgically separated from other types of cells (discussed later), a practical target is to isolate an intact, undamaged RG by dissection.

  1. Preparation of larvae at the appropriate developmental stages
    NOTE: To synchronize the developmental stages of D. melanogaster larvae, it is necessary to collect eggs within a narrow time window and to collect larvae at the appropriate times.
    1. Collect eggs for 2 h on a grape-juice agar plate at 25 °C.
    2. Collect newly-hatched 1st instar larvae under a dissecting microscope with forceps and transfer them to a small vial containing mashed standard yeast/cornmeal fly food.
      NOTE: The age of larvae is represented by "hours after egg laying (hAEL)", "hours after hatch (hAH)", or "hours after L3 ecdysis (hAL3E)".
    3. At the appropriate time point, collect the staged larvae with a disposable plastic loop to avoid undesirable injury. To minimize a difference in developmental progression of the 3rd instar larvae, collect the 2nd instar larvae at 70 hAEL (48 hAH) and allow them to molt in 2 h intervals. Then, collect the 3rd instar larvae within 2 h after the L3 ecdysis; this procedure gives 0-2 hAL3E larvae.
    4. Transfer the staged larvae to a dish filled with phosphate buffered saline (PBS).
    5. Rinse the larvae with PBS to remove residual food from the body.
  2. Coarse dissection of larvae under a dissecting microscope
    1. Hold the mouth hook of a larva with forceps. Sever the larval body at the anterior one-third of the body length using micro scissors.
    2. Hold the cut end of the anterior larval body with one pair of forceps. Using the other pair of forceps, gently push the tip of the head toward the inside of the body; this procedure turns the larval body inside out.
      NOTE: This step can be skipped for the dissection of 1st instar larvae.
    3. Repeat steps 1.2.1-1.2.2 with additional larvae for 5-10 min.
      NOTE: The number of larvae should be equal to or less than 20 to save time for dissection.
  3. Fillet dissection of larvae
    NOTE: This method allows keeping the position of tissues to remain intact.
    1. Leave larvae in water only for 1 h. The larvae are immobilized due to asphyxiation.
    2. Put a larva dorsal side up in a drop of PBS on a silicon-coated dish, with its dorsal side up.
      NOTE: The dorsal side has 2 tracheal tubes running longitudinally.
    3. Under a dissecting microscope, insert an insect pin into the anterior tip of the larva using forceps. Stretch the larval body to the posterior side and put a second pin into the posterior tip of the larva.
      NOTE: In addition to insect pins, a needle made from a tungsten wire can be useful21. A needle can be embedded in a pipette tip through heating for use as a dissecting needle.
    4. Under a dissecting microscope, make a small incision near the tail with micro scissors. From the incision, cut the dorsal cuticle longitudinally along the dorsal midline toward the head. Be careful not to damage tissues under the cuticle.
    5. Stretch the bodywall cuticle on each side and place 4 pins at each corner of the dissected bodywall.
    6. Remove a portion of anterior fat body with forceps, to expose the brain-RG complex be exposed on the surface. Remove a pair of salivary glands, if necessary.
  4. Fixation and staining of tissue with immunohistochemistry
    1. Put the anterior one-third of the larval bodies (step 1.2.2) into a 1.5 mL microtube filled with 500 µL the fix solution (4% paraformaldehyde in PBS). Fix less than 20 samples at one time, otherwise PBS attached to the fixed samples may cause an unfavorable dilution of the fixative. For fillet dissection, remove a drop of PBS and then add a drop of fix solution.
      NOTE: For fix solution, either 3.7% formaldehyde or 4% paraformaldehyde may be used.
    2. Incubate the dissected tissues in the fix solution for 30 min at room temperature (RT) or alternatively for 2 h at 4 °C.
    3. Replace the fix solution with 500 µL PBS and quickly rinse the samples 3 times. Wash the samples with 500 µL 0.3% PBT (PBS + 0.3% octylphenol ethoxylate) for 15 min on a rocker at RT. For fillet dissection, remove the insect pins and transfer the samples into a 1.5 mL microtube.
      NOTE: For increasing the permeability of antibodies into tissues, the samples are treated with 500 µL 2.0% PBT for 1-2 h on a rocker at RT.
    4. Replace PBT with 500 µL blocking solution (2% bovine serum albumin in PBT). Incubate the samples for 1.5 h on a rocker at RT.
    5. Replace the blocking solution with 50 µL the primary antibody solution, i.e. antibodies diluted in the blocking solution.
    6. Incubate the samples overnight on a rocker at 4 °C.
    7. Replace the primary antibody solution with 500 µL 0.3% PBT and quickly rinse the samples 5 times. Wash the samples 3 times with 500 µL 0.3% PBT for 15 min each on a rocker at RT.
    8. Replace 0.3% PBT with 50 µL the secondary antibody solution (dye-conjugated antibodies diluted in the blocking solution). For nuclear staining, 0.5 µL 4',6-diamidino-2-phenylindole (DAPI, 0.1 mg/mL stock solution) is added to 50 µL solution. Incubate the samples on a rocker for 2 h at RT or alternatively at 4 °C overnight.
      NOTE: Keep the samples in the dark.
    9. Replace the antibody solution with 500 µL 0.3% PBT and rinse 5 times. Wash the samples 3 times with 500 µL 0.3% PBT for 15 min each at RT.
  5. Fine dissection of the brain-RG complex containing the PG and mounting
    1. Use a disposable pipet to transfer the immunostained samples to a dish filled with 0.3% PBT.
      NOTE: To avoid inconvenient bubbles during dissection and mounting, 0.3% PBT can be replaced with PBS.
    2. Under a dissecting microscope, hold the cuticle or mouth hook with one pair of forceps. Using a 27 G needle attached to a 1 mL syringe as a "knife", cut the anterior tip of the esophagus and eye discs to remove the brain-RG-eye disc complex from body cuticle.
    3. Separate the esophagus and gut from the brain-RG-eye disc complex with forceps. Since the esophagus passes through the brain above the ventral nerve cord (VNC), pull out the gut to the posterior side.
      NOTE: To remove extra imaginal discs from the samples, cut the connections between the brain, eye discs, and leg discs with a needle knife.
    4. Repeat the steps 1.5.1-1.5.4 for other samples.
      NOTE: To prevent tissue debris from sticking to the samples, transfer each completed sample to a different clean dish filled with 0.3% PBT or PBS.
    5. Transfer the samples to the center of a clean glass slide with a micropipette.
      NOTE: For 2nd or 3rd instar larvae, the end of a micropipette tip should be truncated with a razor blade.
    6. Under a dissecting microscope, align individual samples with their dorsal side up by using forceps.
      NOTE: The RG is located on the dorsal side of the brain (Figure 2A). This alignment facilitates the specification of individual samples during imaging.
    7. Tilt a glass slide and wipe off as much excess PBT as possible. Put a drop of mounting reagent on a side of the slide. Place the edge of a cover slip from the other side of the drop and put the cover slip on the samples slowly with forceps.
      NOTE: This procedure prevents the samples from moving outside the cover slip.
    8. Store the samples at 4 °C. Keep the samples in the dark.

2. The Dissection of the Ovary in Adult Females

  1. Preparation of adult females
    1. Feed female flies with standard yeast/cornmeal fly food for 3-4 days to fatten up the ovary. Maintain at 25 °C.
  2. Dissection of the adult female ovary
    1. Anesthetize female flies with CO2 gas and cut off their heads.
    2. Transfer fly bodies to a 3 cm dish filled with PBS.
    3. Under a dissecting microscope, hold the thorax on the dorsal side using forceps. Grasp the abdominal segment A5 or A6 with the other forceps and peel away the abdominal cuticle toward the posterior side. Wipe off the sticky cuticle from the tip of the forceps.
    4. Pinch an oviduct and take out the ovary from the body.
    5. Comb and spread the tips of the ovary with forceps.
      NOTE: This operation improves the efficiency of immunostaining.
  3. Dissection of adult female brain, VNC and reproductive organs.
    NOTE: This method allows the innervation of the ovary to be kept intact.
    1. Anesthetize female flies with CO2 gas and transfer them to a silicon-coated dish filled with PBS.
    2. Under a dissecting microscope, hold the proboscis and peel away the head cuticle to expose the brain with forceps. Remove the trachea attached to the brain.
      NOTE: The brain is a white and rounded structure. The brain connects to the VNC in the thorax.
    3. Hold the thorax on the dorsal side and cut off the legs and wings with one pair of forceps. Using the other pair of forceps, peel away the thorax ventral cuticle from the bases of the legs. Once the VNC is exposed, remove the residual dorsal cuticle and muscles attached to the VNC using forceps. Be careful not to damage the connection between the brain and the VNC.
    4. Use forceps to peel away the abdominal cuticle and expose the ovary and their interactive organs including neurons, the crop, the gut, the oviduct, the uterus, and the accessary gland.
    5. Remove the residual tissues including the trachea and the fat body using forceps.
  4. Fixation and staining of tissue with immunohistochemistry
    1. Transfer approximately 8-10 pairs of the ovaries to a 1.5 mL microtube filled with 250 µL the fix solution (4% paraformaldehyde in PBS) and incubate the samples for 30 min at RT.
    2. Replace the fix solution with 500 µL PBS and quickly rinse the samples 3 times.
    3. Wash the samples 2 times with 500 µL 0.1% PBT for 5 min each at RT.
    4. Replace PBT with 50 µL blocking solution. Incubate the samples for 1 h on a rocker at RT.
    5. Replace the blocking solution with 50 µL the primary antibody solution.
    6. Incubate the samples overnight on a rocker at 4 °C.
    7. Replace the primary antibody solution with 500 µL 0.1% PBT. Wash the samples 3 times with 500 µL 0.1% PBT for 15 min each on a rocker at RT.
    8. Replace 0.1% PBT with 50 µL secondary antibody solution. For nuclear staining, add 0.5 µL DAPI  to 50 µL the solution. Incubate the samples for 2 h on a rocker at RT.
    9. Replace the secondary antibody solution with 500 µL 0.1% PBT. Wash the samples 3 times with 500 µL 0.1% PBT for 15 min each on a rocker at RT.
      NOTE: The washing procedure can be done at 4 °C overnight. Keep the samples in the dark.
  5. Mounting the samples onto glass slides
    1. Transfer the samples to a glass slide with 0.1% PBT using a micropipette.
      NOTE: To avoid inconvenient bubbles during dissection and mounting, 0.1% PBT can be replaced with PBS.
    2. For the ovary, separate the strings of the ovarioles from each other with forceps under a dissecting microscope. Do not injure the tips of the ovarioles containing the germaria. For the brain-VNC-reproductive organ complex, remove the residual cuticle and arrange the position on a glass slide.
    3. Wipe off as much excess PBT as possible and put a drop of mounting reagent in the center of the samples. Place a cover slip slowly using forceps.
    4. Store the samples at 4 °C. Keep the samples in the dark.

3. Imaging with a Confocal Laser Scanning Microscope

  1. Observe the samples under the microscope and bring the steroidogenic organs in the field of view. Once the view is fixed, switch the imaging system to the image acquisition mode.
    NOTE: The 10-20X objective lenses are used to obtain the images of the brain-RG complex or the whole ovary. Alternatively, the 40-63X objective lenses (water or oil immersion) are used for visualizing the subcellular localization of proteins in GSCs or the neuronal innervations of the PG and the ovary.
  2. Set up the parameters of image acquisition on the attached software. Select the combination of fluorescence (e.g. GFP and RFP). By fast scanning procedure, adjust the laser power, the sensitivity of detectors (Gain/Offset), and the size of the pinhole.
    NOTE: To acquire images at high quality, the scanning laser power and the sensitivity of detector (Gain) should be optimized for each sample. To outline the shape of the tissues, a transmitted-light image can be taken in addition to a fluorescent image.
  3. Take Z stacks of the images. During a continuous fast scan, move the focal plane with the focus drive of the microscope and set the starting position and the end position. The number of slices and the interval distance between slices are adjusted accordingly.
  4. Select the scan speed, the scan method, and the bidirectional scan mode.
  5. "Start the real scans" for image acquisition.
  6. Save the image with the format of the attached software.
    NOTE: The original image files contain the information of image acquisition parameters and scales. Exported images can be processed using an image processing software on the computer22.

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Results

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We used the above protocols to visualize steroidogenic organs and their interactive organs in D. melanogaster larvae and adult females. The overall scheme of protocols is shown in Figure 1.

The RG, including the PG (Figure 2D), is smaller and more transparent than the brain and is located at the anterior-dorsal side of the brain (Figure 2A-C and 3A-E). To label the PG cells, several groups have generated various types of antibodies against ecdysteroidogenic e...

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Discussion

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We studied ecdysteroid biosynthesis and its regulatory mechanism in D. melanogaster, and devised a protocol for dissection and immunostaining. The timing of ecdysteroid biosynthesis is affected by environmental cues through neuronal inputs33, so it is essential to maintain the innervation of the ecdysteroidogenic organs along with the brain, VNC, and other tissues during dissection.

As described above, the D. melanogaster PG forms a complex endocr...

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Disclosures

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No conflicts of interest declared.

Acknowledgements

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We thank Reiko Kise and Tomotsune Ameku for their technical support for this work. We are also grateful to Kei Ito, Olga Alekseyenko, Akiko Koto, Masayuki Miura, the Bloomington Drosophila Stock Center, KYOTO Stock Center (DGRC), and the Developmental Studies Hybridoma Bank for stocks and reagents. This work was supported by grants to Y.S.N. from JSPS KAKENHI Grant Number 16K20945, The Naito Foundation, and Inoue Science Research Award; and by a grant to R.N. from MEXT KAKENHI Grant Number 16H04792.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
egg collection
tissue culture dish (55 mm)AS ONE1-8549-02 for grape-juice agar plates
collection cupHIKARI KAGAKU
yeast pasteOriental dry yeast, Tokyo
100% grape juiceWelch Food Inc.
rearing larvae
small vials (12ml, 40×23.5 mm, PS)SARSTEDT58.487
disposable loopAS ONE6-488-01
standard fly food the recepi us on the website of Blooington stock center.
dissection
dissecting microscopeCarl ZeissStemi 2000-C
dissecting microscopeLeicaS8 AP0
tissue culture dish (35 x 10 mm, non-treated)IWAKI1000-035
SylgardTORAYcoarting silicon inside dishes
Terumo needle (27G, 0.40 x 19 mm) TERUMONN-2719SA "knife" to cut the tissue
forceps, Inox, #5Dumont, Switzerland
insect pin (0.18 mm in diameter)Shiga Brandfor fillet dissection
micro scissorsNATSUME SEISAKUSHO CO LTD. MB-50-10
fixation
ultrapure waterMerck Millipore
phosphate buffered saline (PBS)
FormaldehydeNacalai tesque16222-65
ParaformaldehydeNacalai tesque02890-45
Triton-X100Nacalai tesque35501-15
microtubes (1.5 ml)INA OPTIKACF-0150
Incubation
As one swist mixer TM-300 (rocker)As oneTM-300rocker
Bovine Serum AlbuminSIGMA9048-46-8
primary antibody
anti-Sro (guinea pig), 1:1000
anti-GFP (rabbit), 1:1000Molecular ProbesA6455Shimada-Niwa ans Niwa, 2014
anti-GFP (mouse mAb, GF200), 1:100Nakarai tesque04363-66
anti-5HT (rabbit), 1:500SIGMAS5545
anti-Hts 1B1 (mouse)Developmental Studies Hybridoma Bank (DSHB)1B1
anti-DE-cadherin (rat), 1:20DSHBDCAD2
anti-nc82 (mouse), 1:50DSHBnc82
secondary antibody
Goat anti-Rabbit IgG (H+L) Secondary Antibody, Alexa Fluor 488 conjugateLife TechnologiesA-11008
Goat anti-Mouse IgG (H+L) Secondary Antibody, Alexa Fluor 488 conjugateLife TechnologiesA-11001
Goat anti-Rat IgG (H+L) Secondary Antibody, Alexa Fluor 546 conjugateLife TechnologiesA-11081
Goat anti-Guinea Pig IgG (H+L) Secondary Antibody, Alexa Fluor 555 conjugateLife TechnologiesA-21435
Alexa Fluor 546 dye-conjugated phalloidinLife TechnologiesA-22283
Mounting reagents
Micro slide glassMatsunami Glass Ind.,Ltd.SS7213
Square microscope cover glassMatsunami Glass Ind.,Ltd.C218181
FluorSave reagent (Mounting reagent)Calbiochem345789
Transfer pipette 1 ml (Disposable dropper)WATSON5660-222-1S
imaging
LSM700 laser scanning microscope systemCarl Zeissinverted Axio Observer. Z1 SP left
image processing
LSM700 ZENCarl ZeissIt is a special user interface based on the 64 bit Microsoft Windows7 operating system
ImageJ
Fly stocks
w; GMR45C06-GAL4 from Bloomington Drosophila Stock Center. (#46260)
UAS–GFP; UAS–mCD8::GFPgifts from K. Ito, The University of Tokyo.
w[1118]
w; phantom-GAL4#22/UAS-turboRFP
w; UAS-mCD8::GFP; TRH-GAL4see in Ref29, Alekseyenko, O. V, Lee, C. & Kravitz, E. A.(2010)
w; UAS-mCD8::GFP from Bloomington Drosophila Stock Center. (#32188)
yw;; nSyb-GAL4 from Bloomington Drosophila Stock Center. (#51941)

References

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,
  1. Miller, W. L., Auchus, R. J. The Molecular Biology, Biochemistry, and Physiology of Human Steroidogenesis and Its Disorders. Endocr. Rev. 32 (1), 81-151 (2011).
  2. Rousseau, G. G. Fifty years ago: The quest for steroid hormone receptors. Mol. Cell. Endocrinol. 375 (1), 10-13 (2013).
  3. Gilbert, L. I., Rybczynski, R., Warren, J. T. Control and biochemical nature of the ecdysteroidogenic pathway. Annu. Rev. Entomol. 47, 883-916 (2002).
  4. Niwa, R., Niwa, Y. S. Enzymes for ecdysteroid biosynthesis: their biological functions in insects and beyond. Biosci. Biotechnol. Biochem. 78 (8), 1283-1292 (2014).
  5. Kozlova, T., Thummel, C. S. Steroid regulation of postembryonic development and reproduction in drosophila. Trends Endocrinol. Metab. 11 (7), 276-280 (2000).
  6. Ishimoto, H., Kitamoto, T. Beyond molting-roles of the steroid molting hormone ecdysone in regulation of memory and sleep in adult Drosophila. Fly. 5 (3), 215-220 (2011).
  7. Ishimoto, H., Sakai, T., Kitamoto, T. Ecdysone signaling regulates the formation of long-term courtship memory in adult Drosophila melanogaster. Proc. Natl. Acad. Sci. USA. 106 (15), 6381-6386 (2009).
  8. Simon, A. F., Shih, C., Mack, A., Benzer, S. Steroid control of longevity in Drosophila melanogaster. Science. 299 (5611), 1407-1410 (2003).
  9. Buszczak, M., Freeman, M. R., Carlson, J. R., Bender, M., Cooley, L., Segraves, W. a Ecdysone response genes govern egg chamber development during mid-oogenesis in Drosophila. Development. 126 (20), 4581-4589 (1999).
  10. Carney, G. E., Bender, M. The drosophila ecdysone receptor (EcR) gene is required maternally for normal oogenesis. Genetics. 154 (3), 1203-1211 (2000).
  11. Uryu, O., Ameku, T., Niwa, R. Recent progress in understanding the role of ecdysteroids in adult insects: Germline development and circadian clock in the fruit fly Drosophila melanogaster. Zoological Lett. 1, 32(2015).
  12. Ameku, T., Niwa, R. Mating-Induced Increase in Germline Stem Cells via the Neuroendocrine System in Female Drosophila. PLOS Genet. 12 (6), e1006123(2016).
  13. Danielsen, E. T., et al. A Drosophila Genome-Wide Screen Identifies Regulators of Steroid Hormone Production and Developmental Timing. Dev. Cell. 37 (6), 558-570 (2016).
  14. Ou, Q., Zeng, J., Yamanaka, N., Brakken-Thal, C., O'Connor, M. B., King-Jones, K. The Insect Prothoracic Gland as a Model for Steroid Hormone Biosynthesis and Regulation. Cell Rep. , (2016).
  15. Yamanaka, N., Rewitz, K. F., O'Connor, M. B. Ecdysone control of developmental transitions: lessons from Drosophila research. Annu. Rev. Entomol. 58, 497-516 (2013).
  16. Niwa, Y. S., Niwa, R. Transcriptional regulation of insect steroid hormone biosynthesis and its role in controlling timing of molting and metamorphosis. Dev. Growth Differ. 58, 94-105 (2015).
  17. Monastirioti, M. Distinct octopamine cell population residing in the CNS abdominal ganglion controls ovulation in Drosophila melanogaster. Dev. Biol. 264 (1), 38-49 (2003).
  18. Siegmund, T., Korge, G. Innervation of the ring gland of Drosophila melanogaster. J. Comp. Neurol. 431 (4), 481-491 (2001).
  19. McBrayer, Z., et al. Prothoracicotropic Hormone Regulates Developmental Timing and Body Size in Drosophila. Dev. Cell. 13 (6), 857-871 (1979).
  20. Shimada-Niwa, Y., Niwa, R. Serotonergic neurons respond to nutrients and regulate the timing of steroid hormone biosynthesis in Drosophila. Nat. Commun. 5, 5778(2014).
  21. Brady, J. A simple technique for making very fine, durable dissecting needles by sharpening tungsten wire electrolytically. Bull World Health Organ. 32 (1), 143-144 (1965).
  22. Abramoff, M. D., Magalhães, P. J., Ram, S. J. Image processing with ImageJ. Biophotonics Int. 11 (7), 36-42 (2004).
  23. Ohhara, Y., et al. Autocrine regulation of ecdysone synthesis by β3-octopamine receptor in the prothoracic gland is essential for Drosophila metamorphosis. Proc. Natl. Acad. Sci. USA. 112 (5), 1452-1457 (2015).
  24. Gibbens, Y. Y., Warren, J. T., Gilbert, L. I., O'Connor, M. B. Neuroendocrine regulation of Drosophila metamorphosis requires TGFbeta/Activin signaling. Development. 138 (13), 2693-2703 (2011).
  25. Parvy, J. P., et al. A role for βFTZ-F1 in regulating ecdysteroid titers during post-embryonic development in Drosophila melanogaster. Dev. Biol. 282 (1), 84-94 (2005).
  26. Parvy, J. -P., et al. Forward and feedback regulation of cyclic steroid production in Drosophila melanogaster. Development. 141 (20), 3955-3965 (2014).
  27. Brand, A. H., Perrimon, N. Targeted gene expression as a means of altering cell fates and generating dominant phenotypes. Development. 118 (2), 401-415 (1993).
  28. Rewitz, K. F., Yamanaka, N., Gilbert, L. I., O'Connor, M. B. The Insect Neuropeptide PTTH Activates Receptor Tyrosine Kinase Torso to Initiate Metamorphosis. Science. 326 (5958), 1403-1405 (2009).
  29. Li, H. -H., et al. A GAL4 driver resource for developmental and behavioral studies on the larval CNS of Drosophila. Cell Rep. 8 (3), 897-908 (2014).
  30. Alekseyenko, O. V., Lee, C., Kravitz, E. A. Targeted manipulation of serotonergic neurotransmission affects the escalation of aggression in adult male Drosophila melanogaster. PLOS One. 5 (5), e10806(2010).
  31. Domanitskaya, E., Anllo, L., Schüpbach, T. Phantom, a cytochrome P450 enzyme essential for ecdysone biosynthesis, plays a critical role in the control of border cell migration in in Drosophila. Dev. Biol. 386 (2), 408-418 (2014).
  32. Song, X., Zhu, C. -H., Doan, C., Xie, T. Germline stem cells anchored by adherens junctions in the Drosophila ovary niches. Science. 296 (5574), 1855-1857 (2002).
  33. Niwa, Y. S., Niwa, R. Neural control of steroid hormone biosynthesis during development in the fruit fly Drosophila melanogaster. Genes Genet. Syst. 89 (1), 27-34 (2014).
  34. Yoshiyama-Yanagawa, T., et al. The conserved Rieske oxygenase DAF-36/Neverland is a novel cholesterol-metabolizing enzyme. J. Biol. Chem. 286 (29), 25756-25762 (2011).
  35. Niwa, R., et al. Non-molting glossy/shroud encodes a short-chain dehydrogenase/reductase that functions in the "Black Box" of the ecdysteroid biosynthesis pathway. Development. 137 (12), 1991-1999 (2010).
  36. Komura-Kawa, T., et al. The Drosophila Zinc Finger Transcription Factor Ouija Board Controls Ecdysteroid Biosynthesis through Specific Regulation of spookier. PLOS Genet. 11 (12), e1005712(2015).
  37. Yamanaka, N., Marqués, G., O'Connor, M. B. Vesicle-Mediated Steroid Hormone Secretion in Drosophila melanogaster. Cell. 163 (4), 907-919 (2015).
  38. Riemensperger, T., Pech, U., Dipt, S., Fiala, A. Optical calcium imaging in the nervous system of Drosophila melanogaster. BBA-Gen. Subjects. 1820 (8), 1169-1178 (2012).
  39. Owald, D., Lin, S., Waddell, S. Light, heat, action: neural control of fruit fly behavior. Phil. T. Roy. Soc. B. 370 (1677), 20140211(2015).

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Immunostaining ProtocolLarval DissectionOvary VisualizationBrain Ring GlandEcdysteroid BiosynthesisGermline Stem CellsNeuronal InnervationFluorescent Labeling

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