We appreciate assistance from members of the fly community, particularly Dr. Denise Montell, Dr. Lynn Cooley, and Dr. Pernille Rorth, for reagents. We thank Flybase, the Bloomington Drosophila Stock Center, and Developmental Studies Hybridoma Bank for information and providing fly stocks and antibodies, respectively. LM is supported by the Department of Education&#160;Grant, Graduate Assistance in the Areas of National Need (GAANN) training fellowship (P200A120017) and by a NIGMS Initiative for Maximizing Student Development Grant (2 R25-GM55036). A portion of the microscopy work was supported by NSF MRI grant DBI-0722569 and the Keith R. Porter Core Imaging Facility. Research was supported in part by a NSF CAREER Award (1054422) and a Basil O&#8217;Connor Starter Scholar Award from the March of Dimes, both awarded to MSG.

1. Dissection of Ovarioles
<ol>
	<li>Transfer about fifteen 2-4-day-old female flies and a few males to a fresh fly food vial with added active dry yeast. Use cotton as a plug for the vials, and add a few drops of water to the cotton to keep the humidity high.</li>
	<li>Place vial in a 25 &#176;C incubator for 14-16 hr to maximize the number of stage 8-10 egg chambers. 
	NOTE: Incubation time varies depending on temperature and desired stage.</li>
	<li>Prepare dissection media, 0.1M KPO4 and NP40 solutions as needed for the next day (see Materials).</li>
	<li>Anesthetize the flies using CO2, and place under a dissection microscope. Pipet several drops of dissection media into each cavity in the glass depression slide.</li>
	<li>Hold forceps in each hand at an approximately 30&#176; angle to the table top and orient one female fly with the wings down. With your non-dominant hand, pick up the female using forceps, grasping her at the anterior of the abdomen, near the thorax, and submerge her into the dissection media in the depression slide (Figure 1A inset).</li>
	<li>With the other pair of forceps, grasp the exoskeleton at the ventral posterior of the abdomen and pierce through. Grab the ovaries at the posterior end, then pull them out of the abdomen and place into the fresh media on the other side of the depression slide. (Figure 1A inset). Avoid pulling apart the gut. Discard the carcass.</li>
	<li>Repeat with the other female flies until all ovaries are dissected. Discard the males.</li>
	<li>With one pair of forceps, hold an ovary at the posterior end (near the largest egg chambers), and with the other hand use a pair of forceps to pinch one of the most anterior egg chambers (germarium). (Figure 1A).</li>
	<li>Slowly and steadily, pull an ovariole chain out of the ovary sheath. Continue to dissect out ovarioles individually until all desired egg chambers are removed.</li>
	<li>Repeat steps 1.7-1.8 for all dissected ovaries.</li>
	<li>Once complete, transfer ovarioles to a 0.6ml tube using a plastic transfer pipette.</li>
	<li>Let ovarioles settle to the bottom of tube, then proceed to fixing and staining steps.</li>
</ol>
2. Immunofluorescent (IF) Staining of Ovarioles
<ol>
	<li>Carefully remove dissection media from ovarioles with a pipette. Be sure not to remove any ovarioles or egg chambers. Leave approximately 50 &#956;l&#160;of ovarioles and dissection media.</li>
	<li>Add 50 &#956;l of 16% paraformaldehyde to 150 &#956;l of 0.1 M KPO4 Buffer in a tube, and add solution to egg chambers to fix them. Incubate while rocking for 10 min. CAUTION: paraformaldehyde is toxic.</li>
	<li>Remove fixative and rinse twice with 0.5 ml NP40 wash buffer.</li>
	<li>Wash two times for 15 min each at RT.</li>
	<li>Refer to standard IF protocol15 for subsequent steps.
	<ol>
		<li>Briefly, after fixation, add primary antibodies at the appropriate dilutions and incubate O/N at 4 &#176;C. Wash in NP40 several times, then add secondary antibodies (1:200 dilutions).</li>
		<li>Wash several times in NP40, then add DAPI (1:1,000) for 10 min, then repeat washes. Once IF protocol is complete, add 200&#160;&#956;l&#160;of 50% glycerol in PBS to egg chambers and let sit at RT for 2 hr.</li>
		<li>Remove solution, replace it with 70% glycerol in PBS, and cover with foil. Place sample at 4 &#7506;C until ready for mounting. Meanwhile, continue with step 3.</li>
	</ol>
	</li>
</ol>
3. Advance Preparation of Glycerin Jelly Slides
<ol>
	<li>Measure at least 4 ml&#160;of glycerin jelly using a spatula and fill a glass cell culture tube to the half way mark. Melt glycerin jelly in a water bath at 55 &#176;C for 30 min.</li>
	<li>Pour a small drop of glycerol, approximately 300 &#956;l, from the stock solution onto two microscope slides. Using a tissue, spread glycerol in a thin layer across each of the slides. This provides a base and allows for easy removal of glycerin jelly at step 5.6.</li>
	<li>Cut the tip off of a filtered 1,000 &#956;l pipette tip. Use cut pipette tip to transfer 1,000-1,500 &#956;l of warm glycerin jelly slowly to each microscope slide. Take care to prevent bubbles.</li>
	<li>Let glycerin jelly slides sit for 5 min at RT&#160;to set.</li>
	<li>Place each glycerin jelly slide in 60&#160;mm &#215; 15&#160;mm petri dish and cover with plastic wrap. Place at 4 &#176;C O/N. Optionally, store glycerin jelly slides for up to 5 days at 4 &#176;C.</li>
</ol>
4. Mounting Egg Chambers
<ol>
	<li>Using samples from step 2.5, pipette several 3 &#956;l drops of egg chambers in 70% glycerol across one glycerin jelly slide. Continue pipetting 3 &#956;l drops until all egg chambers are on this slide. Make sure each drop contains at least ten egg chambers. Meanwhile, heat a culture tube of glycerin jelly in a 55 &#176;C water bath.</li>
	<li>Place the glycerin jelly slide with egg chambers under a dissection microscope. Adjust magnification so that only one drop of egg chambers is in the field of view (Figure 1B).</li>
	<li>Using a 30 gauge needle as a spoon, pick up the desired stage egg chamber. When necessary, separate desired egg chambers from an ovariole chain using the needle as a knife.</li>
	<li>To avoid debris that may interfere with imaging, transfer the egg chamber to the second glycerin jelly slide, with the anterior facing left (Figure 1E-1). Repeat until there are at least seven egg chambers lined up in a column (Figure 1C).</li>
	<li>Form new columns of seven until all desired egg chambers are transferred to the second glycerin jelly slide.</li>
	<li>Tear off a small piece of tissue and twist. Use this to absorb the excess PBS-Glycerol from the egg chambers by lightly touching each one. Take care to not remove egg chambers.</li>
	<li>Cut off the tip of a filtered 200 &#956;l pipette tip. Use this to transfer 150 &#956;l of glycerin jelly to cover all the columns of egg chambers. 
	NOTE: The amount of glycerin jelly needed to cover the columns is based on egg chamber stages and number of columns. Amount may be adjusted.</li>
	<li>Let mounted egg chambers sit for 5 min at RT until the top layer of glycerin jelly is completely solidified (Figure 1D).</li>
	<li>Place slide of mounted egg chambers in a 60&#160;mm &#215; 15&#160;mm petri dish and cover with plastic wrap. Store at 4 &#176;C O/N to allow the glycerin jelly layers to solidify together (place in the dark if there is concern about photobleaching).</li>
</ol>
5. Preparation of Egg Chambers for Imaging
<ol>
	<li>Place slide of mounted egg chambers under a dissection microscope. Adjust magnification so that only one column of egg chambers is in the field of view.</li>
	<li>Using a 45&#176; angle miniature scalpel, cut a straight line along the right side of the first column of egg chambers (close to the posterior side of the egg chambers). (Figure 1D-E)</li>
	<li>Next, cut above the first egg chamber at the top and below the last egg chamber in the column. At this point the column of egg chambers should be cut on three sides.</li>
	<li>Using a microknife, slice along the left side of the column, as close as possible to the anterior side of the egg chambers (Figure 1E-3).</li>
	<li>Pipette 100 &#956;l of cold 1X PBT over the cut column.</li>
	<li>Use a round edged spatula to remove the excess glycerin jelly around three sides of the cut column of egg chambers and discard, leaving the rest on the slide.</li>
	<li>Insert the spatula in the cut made at the anterior side of egg chambers and separate the column from the primary glycerin jelly block.</li>
	<li>Prepare samples for either inverted (5.8.1) or upright (5.8.2) microscopy.
	<ol>
		<li>For Inverted Microscope: Transfer the column of egg chambers to a coverslip with anterior side of egg chambers facing down. Repeat steps 5.2-5.8 until all columns of glycerin jelly have been removed and mounted. Add a couple of drops of 50% PBS-Glycerol onto each block of egg chambers to prevent them from drying out. Image egg chambers directly on coverslip.</li>
		<li>For Upright Microscope: Transfer the column of egg chambers to a microscope slide with the anterior side facing up. Repeat steps 5.2-5.9 until all columns of glycerin jelly have been removed and mounted on microscope slides. Add a couple of drops of 50% PBS-Glycerol onto each block of egg chambers and cover blocks with a #1 &#189; coverslip.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Hudson, A. M., Cooley, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+for+studying+oogenesis.">Methods for studying oogenesis.</a> Methods. 68 (1), 207-217 (2014).</li><li>Bastock, R., St Johnston, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+oogenesis.">Drosophila oogenesis.</a> Curr Biol. 18 (23), R1082-R1087 (2008).</li><li>Wu, X., Tanwar, P. S., Raftery, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+follicle+cells:+morphogenesis+in+an+eggshell.">Drosophila follicle cells: morphogenesis in an eggshell.</a> Semin Cell Dev Biol. 19 (3), 271-282 (2008).</li><li>Bilder, D., Haigo, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expanding+the+morphogenetic+repertoire:+perspectives+from+the+Drosophila+egg.">Expanding the morphogenetic repertoire: perspectives from the Drosophila egg.</a> Dev Cell. 22 (1), 12-23 (2012).</li><li>Montell, D. J., Yoon, W. H., Starz-Gaiano, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Group+choreography:+mechanisms+orchestrating+the+collective+movement+of+border+cells.">Group choreography: mechanisms orchestrating the collective movement of border cells.</a> Nat Rev Mol Cell Biol. 13 (10), 631-645 (2012).</li><li>Spradling, A. C., Bate, M., Marinez-Arias, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developmental+genetics+of+oogenesis.">Developmental genetics of oogenesis.</a> The Development of Drosophila melanogaster. , 1-70 (1993).</li><li>Montell, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Border-cell+migration:+the+race+is+on.">Border-cell migration: the race is on.</a> Nat Rev Mol Cell Biol. 4 (1), 13-24 (2003).</li><li>Silver, D., Geisbrecht, E., Montell, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Requirement+for+JAK/STAT+signaling+throughout+border+cell+migration+in+Drosophila.">Requirement for JAK/STAT signaling throughout border cell migration in Drosophila.</a> Development. 132 (15), 3483-3492 (2005).</li><li>Ghiglione, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Drosophila+cytokine+receptor+Domeless+controls+border+cell+migration+and+epithelial+polarization+during+oogenesis.">The Drosophila cytokine receptor Domeless controls border cell migration and epithelial polarization during oogenesis.</a> Development. 129 (23), 5437-5447 (2002).</li><li>Denef, N., Sch&#252;pbach, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patterning:+JAK-STAT+signalling+in+the+Drosophila+follicular+epithelium.">Patterning: JAK-STAT signalling in the Drosophila follicular epithelium.</a> Curr Biol. 13 (10), R388-R390 (2003).</li><li>Beccari, S., Teixeira, L., R&#248;rth, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+JAK/STAT+pathway+is+required+for+border+cell+migration+during+Drosophila+oogenesis.">The JAK/STAT pathway is required for border cell migration during Drosophila oogenesis.</a> Mech Dev. 111 (1-2), 115-123 (2002).</li><li>Montell, D., Rorth, P., Spradling, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=slow+border+cells,+a+locus+required+for+a+developmentally+regulated+cell+migration+during+oogenesis,+encodes+Drosophila+C/EBP.">slow border cells, a locus required for a developmentally regulated cell migration during oogenesis, encodes Drosophila C/EBP.</a> Cell. 71 (1), 51-62 (1992).</li><li>Xi, R., McGregor, J., Harrison, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+gradient+of+JAK+pathway+activity+patterns+the+anterior-posterior+axis+of+the+follicular+epithelium.">A gradient of JAK pathway activity patterns the anterior-posterior axis of the follicular epithelium.</a> Dev Cell. 4 (2), 167-177 (2003).</li><li>Toomre, D., Bewersdorf, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+wave+of+cellular+imaging.">A new wave of cellular imaging.</a> Annu Rev Cell Dev Biol. 26, 285-314 (2010).</li><li>McDonald, J., Pinheiro, E., Kadlec, L., Schupbach, T., Montell, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiple+EGFR+ligands+participate+in+guiding+migrating+border+cells.">Multiple EGFR ligands participate in guiding migrating border cells.</a> Dev Biol. 296 (1), 94-103 (2006).</li><li>Van de Bor, V., Zimniak, G., C&#233;r&#233;zo, D., Schaub, S., Noselli, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Asymmetric+localisation+of+cytokine+mRNA+is+essential+for+JAK/STAT+activation+during+cell+invasiveness.">Asymmetric localisation of cytokine mRNA is essential for JAK/STAT activation during cell invasiveness.</a> Development. 138 (7), 1383-1393 (2011).</li><li>Hayashi, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glypicans+regulate+JAK/STAT+signaling+and+distribution+of+the+Unpaired+morphogen.">Glypicans regulate JAK/STAT signaling and distribution of the Unpaired morphogen.</a> Development. 139 (22), 4162-4171 (2012).</li><li>Airoldi, S. J., McLean, P. F., Shimada, Y., Cooley, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intercellular+protein+movement+in+syncytial+Drosophila+follicle+cells.">Intercellular protein movement in syncytial Drosophila follicle cells.</a> J Cell Sci. 124 (Pt 23), 4077-4086 (2011).</li><li>R&#248;rth, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Systematic+gain-of-function+genetics+in+Drosophila).">Systematic gain-of-function genetics in Drosophila).</a> Development. 125 (6), 1049-1057 (1998).</li><li>Lee, T., Lee, A., Luo, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+the+Drosophila+mushroom+bodies:+sequential+generation+of+three+distinct+types+of+neurons+from+a+neuroblast.">Development of the Drosophila mushroom bodies: sequential generation of three distinct types of neurons from a neuroblast.</a> Development. 126 (18), 4065-4076 (1999).</li><li>Shimada, Y., Burn, K. M., Niwa, R., Cooley, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reversible+response+of+protein+localization+and+microtubule+organization+to+nutrient+stress+during+Drosophila+early+oogenesis.">Reversible response of protein localization and microtubule organization to nutrient stress during Drosophila early oogenesis.</a> Dev Biol. 355 (2), 250-262 (2011).</li><li>Qui&#241;ones-Coello, A. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Exploring+strategies+for+protein+trapping+in+Drosophila.">Exploring strategies for protein trapping in Drosophila.</a> Genetics. 175 (3), 1089-1104 (2007).</li><li>Buszczak, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+carnegie+protein+trap+library:+a+versatile+tool+for+Drosophila+developmental+studies.">The carnegie protein trap library: a versatile tool for Drosophila developmental studies.</a> Genetics. 175 (3), 1505-1531 (2007).</li><li>Belu, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Upright+imaging+of+Drosophila+embryos.">Upright imaging of Drosophila embryos.</a> J Vis Exp. (43), (2010).</li><li>Witzberger, M. M., Fitzpatrick, J. A., Crowley, J. C., Minden, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=End-on+imaging:+a+new+perspective+on+dorsoventral+development+in+Drosophila+embryos.">End-on imaging: a new perspective on dorsoventral development in Drosophila embryos.</a> Dev Dyn. 237 (11), 3252-3259 (2008).</li></ol>

The authors have no competing financial interests.

Here we describe a method to mount and image small, developing egg chambers from a end-on perspective. Common techniques for imaging egg chambers are optimized for lateral views and primarily allow precise visualization of medio-lateral follicle cells when stained with fluorescent antibodies. The use of Z-stacks or 3-D reconstructions aids in viewing multiple focal planes, but is still inadequate for sub-cellular resolution of the poles of elliptical egg chambers (Figure 2D). While this can be overcome p...

Study of Drosophila melanogaster has provided great insights into the genetic regulation of a wide range of phenomena. In particular, there has been extensive research on egg development, because oogenesis provides a tractable way to investigate many different developmental processes, including tissue patterning, cell polarity changes, cell cycle switching, and translational regulation1,2,3,4. One important morphogenic event during oogenesis is the specification, acquisition of motility, and migration of a set of cells called border cells (reviewed in 5). Since cell migration is a key feature of animal morphogenesis, and because the genetic regulation of this process is well-conserved, mechanisms determined in flies are likely to be important in other contexts. Thus, we are investigating the molecular control of border cell migration. For our studies, we have developed a new method to observe the anterior poles of developing eggs, where border cells arise, to examine how they develop in detail.
Within the ovary, egg chambers at various developmental stages exist along chains called ovarioles, which are encased in a thin sheath (Figure 1A). Each egg chamber will go on to form one egg. During oogenesis, several different cell types must develop in a coordinated manner. The D. melanogaster egg chamber consists of 16 germ-line cells, including one oocyte, surrounded by a single-layer follicular epithelium6. A small number of specialized cells, called polar cells, arise at the anterior and posterior poles of the epithelium. The 6-8 border cells originate in the anterior epithelium of the egg chamber, induced by the polar cells5,7. In mid-oogenesis (stage 9), the border cells detach from their neighbors and migrate between the nurse cells to reach the oocyte at the posterior of the egg chamber5,7. This movement must be accomplished while the border cells remain in a cluster surrounding two non-motile polar cells, making this a type of collective cell migration. Successful migration of the border cell cluster ensures proper development of the micropyle of the egg shell, which is necessary for fertilization.
The anterior polar cells instruct border cell fate by activating a signal transduction cascade. Polar cells secrete a cytokine, Unpaired (UPD), which binds to a transmembrane receptor, Domeless (DOME), on neighboring follicle cells during stage 8 of oocyte development8,9. The binding of UPD causes Janus tyrosine kinase (JAK) to phosphorylate the Signal Transducer and Activator of Transcription (STAT)8,10,11. STAT then moves to the nucleus to activate transcription. Slow Border Cells (SLBO) is a transcription factor that is a direct transcriptional target of STAT and is also required for border cell migration12. Lateral views of egg chambers indicate that STAT activity is regulated in a gradient across the anterior epithelium8,11,13. Follicle cells closest to the polar cells have the highest levels of activated STAT, thus they become border cells and invade the adjacent germ-line tissue.
To understand how the border cells are specified within and detach from the epithelium, we need to observe how the tissue is organized. If we view egg chambers from an anterior-on perspective, we would expect radial symmetry of STAT activity in the follicle cells surrounding the polar cells. An end-on view would also more accurately show differences of membrane proteins and cell-cell interfaces prior to and during detachment than comparing cells in different focal planes. Because egg chambers are oblong and attached to each other by stalk cells, they settle onto slides laterally, making it difficult to observe the anterior architecture. Thus, much information about the cells at the poles of the egg chamber has been inferred from lateral views. Although some information can be obtained through algorithmic 3-D reconstructions of optical sections, light scattering, photobleaching, and poorer limits of resolution in the Z-axis make this information less detailed and reliable in the absence of expensive techniques like super-resolution microscopy14. Other kinds of section-based imaging (for example, electron microscopy or microtome sectioning) require extensive manipulation of tissues, including dehydration, increasing the likelihood for artifacts. Thus, we developed a new method to image D. melanogaster egg chambers while upright. This method has already proven useful in elucidating how motile cells are fated (see Representative results and Manning et al, under review), and is likely to be more broadly valuable in studies of other aspects of oogenesis.

<table><tbody><tr><td>0.1 M Potassium phosphate Buffer (KPO&lt;sub&gt;4&lt;/sub&gt; Buffer)</td><td></td><td></td><td>Add 3.1 g of NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;&amp;bull;H&lt;sub&gt;2&lt;/sub&gt;O and 10.9 g of Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt; (anhydrous) to distilled H&lt;sub&gt;2&lt;/sub&gt;O to make a volume of 1 L. The pH of the final solution will be 7.4. This buffer can be stored for up to 1 month at 4&amp;deg;C.</td></tr><tr><td>16% Paraformaldehyde aqueous solution, EM grade</td><td>Electron Microscopy Sciences</td><td>15710</td><td>methanol-free to preserve GFP fluorescence</td></tr><tr><td>30G x 1/2 inch needle&amp;nbsp;</td><td>VWR</td><td>BD305106</td><td>Regular bevel</td></tr><tr><td>Active Dry Yeast</td><td>Genesee Scientific</td><td>62-103</td><td>To fatten female flies</td></tr><tr><td>Bovine Serum Albumin</td><td>PAA-cell culture company</td><td>A15-701</td><td>Used in NP40 Wash Buffer</td></tr><tr><td>Dumont #5 Forceps</td><td>Fine Science Tools</td><td>11295-10</td><td>Dumostar alloy, biologie tip; sharp tips are essential for ovariole dissection</td></tr><tr><td>DAPI (4&amp;prime;,6-Diamidino-2-phenylindole dihydrochloride)</td><td>Sigma Aldrich</td><td>D9542</td><td>5 mg/ml stock solution&amp;nbsp;&amp;nbsp;</td></tr><tr><td>Fetal Bovine Serum (FBS)</td><td>Life Technologies</td><td>16140-071</td><td>Added to supplement dissection medium (10%)&amp;nbsp;</td></tr><tr><td>Glass culture tube</td><td>VWR</td><td>47729-576</td><td>14 ml</td></tr><tr><td>Glass Depression Slides</td><td>VWR</td><td>470019-020</td><td>1.2 mm Thick, Double Cavity</td></tr><tr><td>Glycerin Jelly&amp;nbsp;</td><td>Electron Microsocpy Sciences</td><td>17998-10</td><td>Mounting media</td></tr><tr><td>Glycerol</td><td>IBI Scientific</td><td>IB15760</td><td>70% in PBS</td></tr><tr><td>IGEPAL CA-630</td><td>Sigma Aldrich</td><td>I3021-500ML</td><td>(interchangable for Nonidet P-40)&amp;nbsp; Used in NP40 Wash Buffer&amp;nbsp;</td></tr><tr><td>Leica Fluorescent Stereoscope&amp;nbsp;</td><td>Leica Microsystems</td><td></td><td></td></tr><tr><td>Leica SP5 Confocal Microscope</td><td>Leica Microsystems</td><td></td><td>40x/0.55NA dry objective&amp;nbsp;</td></tr><tr><td>Micro spatula&amp;nbsp;</td><td>VWR</td><td>82027-518</td><td>Stainless steel</td></tr><tr><td>Microscope Slide</td><td>VWR</td><td>16004-368</td><td>75x25x1 mm</td></tr><tr><td>NP40 Wash Buffer</td><td></td><td></td><td>50 mM TRIS-HCl, 150 mM NaCl, 0.5% Ipegal, 1 mg/ml BSA, and 0.02% sodium azide</td></tr><tr><td>Penicillin-Streptomycin-Glutamine</td><td>Life Technologies</td><td>10378-016</td><td>Added to supplement dissection medium (0.6X)</td></tr><tr><td>Petridish- Polysterine, sterile</td><td>VWR</td><td>82050-548</td><td>60 W x 15 H mm</td></tr><tr><td>Phosphate Buffer Saline</td><td>Sigma Aldrich</td><td>P3813-10PAK</td><td>10 packs of Powder</td></tr><tr><td>Potassium phosphate dibasic</td><td>Sigma Aldrich</td><td>P3786-500G</td><td>Used in 0.1 M KPO&lt;sub&gt;4&lt;/sub&gt; Buffer&amp;nbsp;</td></tr><tr><td>Potassium phosphate monobasic</td><td>Sigma Aldrich</td><td>P9791-500G</td><td>Used in 0.1 M KPO&lt;sub&gt;4&lt;/sub&gt; Buffer&amp;nbsp;</td></tr><tr><td colspan="4">[header]</td></tr><tr><td>Schneider&amp;rsquo;s Insect Medium</td><td>Life Technologies</td><td>&lt;br/&gt; 11720018</td><td>With L-glutamine and sodium bicarbonate</td></tr><tr><td>Sodium azide</td><td>Sigma Aldrich</td><td>S2002-25G</td><td>Used in fluorescent antibody staining</td></tr><tr><td>Sodium chloride</td><td>Sigma Aldrich</td><td>S3014-500G</td><td>Used in NP40 Wash Buffer</td></tr><tr><td>TRIS-HCl</td><td>IBI Scientific</td><td>IB70144</td><td>1 M TRIS-HCl, pH 7.4</td></tr><tr><td>Volocity 3D Image Analysis Software</td><td>PerkinElmer</td><td></td><td>For processing confocal Z-stacks</td></tr></tbody></table>

upright imaging of drosophila egg chambers

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in architecture, is well-characterized, and is amenable to genetic analysis. Each egg chamber consists of germ-line cells surrounded by a single epithelial layer of somatic follicle cells. Subsets of follicle cells undergo differentiation during specific stages to become several different cell types. Standard techniques primarily allow for a lateral view of egg chambers, and therefore a limited view of follicle cell organization and identity. The upright imaging protocol describes a mounting technique that enables a novel, vertical view of egg chambers with a standard confocal microscope. Samples are first mounted between two layers of glycerin jelly in a lateral (horizontal) position on a glass microscope slide. The jelly with encased egg chambers is then cut into blocks, transferred to a coverslip, and flipped to position egg chambers upright. Mounted egg chambers can be imaged on either an upright or an inverted confocal microscope. This technique enables the study of follicle cell specification, organization, molecular markers, and egg development with new detail and from a new perspective.

The upright imaging method allowed us to see directly the organization of cells in the anterior follicular epithelium at stage 8. A general marker for follicle cell fate, the Eyes Absent (EYA) protein, as well as the nuclear DNA marker DAPI, showed even expression across this field of cells, and demonstrated that all cells could be seen with similar staining intensities (Figure 2B&#8221;). Proteins regulated in response to the cytokine UPD, however, showed variable patterns and expression levels. Polar c...

Watch this Scientific Journal Video about Upright Imaging of Drosophila Egg Chambers at JoVE.com

Upright Imaging of Drosophila Egg Chambers

The upright imaging method described in this protocol allows for the detailed visualization of the poles of a developing Drosophila melanogaster egg. This end-on view provides a new perspective into the arrangements and morphologies of multiple cell types in the follicular epithelium.

Upright Imaging of Drosophila Egg Chambers

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in ...

upright-imaging-of-drosophila-egg-chambers

Research

JoVE Journal

Developmental Biology

7.6K Views.  University of Maryland Baltimore County. The upright imaging method described in this protocol allows for the detailed visualization of the poles of a developing Drosophila melanogaster egg. This end-on view provides a new perspective into the arrangements and morphologies of multiple cell types in the follicular epithelium.

Video: Upright Imaging of Drosophila Egg Chambers

Live-imaging of the Drosophila Pupal Eye

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track during live cell imaging. These processes include: retinal rotation, cell growth and organismal movement. Additionally, irregularities in the topology of the epithelium, including subtle bumps and folds often introduced as the pupa is prepared for imaging, make it challenging to acquire in-focus images of more than a few ommatidia in a single focal plane. The workflow outlined here remedies these issues, allowing easy analysis of cellular processes during Drosophila pupal eye development. Appropriately-staged pupae are arranged in an imaging rig that can be easily assembled in most laboratories. Ubiquitin-DE-Cadherin:GFP and GMR-GAL4&#8211;driven UAS-&#945;-catenin:GFP are used to visualize cell boundaries in the eye epithelium 1-3. After deconvolution is applied to fluorescent images captured at multiple focal planes, maximum projection images are generated for each time point and enhanced using image editing software. Alignment algorithms are used to quickly stabilize superfluous motion, making individual cell behavior easier to track.

Live-imaging of the Drosophila Pupal Eye

This protocol presents an efficient method for imaging the live Drosophila pupal eye neuroepithelium. This method compensates for tissue movement and uneven topology, enhances visualization of cell boundaries through the use of multiple GFP-tagged junction proteins, and uses an easily-assembled imaging rig.

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track ...

Imaging Calcium in Drosophila at Egg Activation

Egg activation is a universal process that includes a series of events to allow the fertilized egg to complete meiosis and initiate embryonic development. One aspect of egg activation, conserved across all organisms examined, is a change in the intracellular concentration of calcium (Ca2+) often termed a 'Ca2+ wave'. While the speed and number of oscillations of the Ca2+ wave varies between species, the change in intracellular Ca2+ is key in bringing about essential events for embryonic development. These changes include resumption of the cell cycle, mRNA regulation, cortical granule exocytosis, and rearrangement of the cytoskeleton.
In the mature Drosophila egg, activation occurs in the female oviduct prior to fertilization, initiating a series of Ca2+-dependent events. Here we present a protocol for imaging the Ca2+ wave in Drosophila. This approach provides a manipulable model system to interrogate the mechanism of the Ca2+ wave and the downstream changes associated with it.

Imaging Calcium in Drosophila at Egg Activation

This article describes an adaptable ex vivo protocol for visualizing Ca2+ during egg activation in Drosophila.

Egg activation is a universal process that includes a series of events to allow the fertilized egg to complete meiosis and initiate embryonic development. ...

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

Biology

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A remarkable example is given by sensory epithelia, where cells of the same or distinct identities are brought together via cell-cell adhesion showing highly organized planar patterns. Cells align to one another in the same direction and display equivalent polarity over large distances. This organization of the mature epithelia is established over the course of morphogenesis. To understand how the planar arrangement of the mature epithelia is achieved, it is crucial to track cell orientation and growth dynamics with high spatiotemporal fidelity during development in vivo. Robust analytical tools are also essential to identify and characterize local-to-global transitions. The Drosophila pupa is an ideal system to evaluate oriented cell shape changes underlying epithelial morphogenesis. The pupal developing epithelium constitutes the external surface of the immobile body, allowing long-term imaging of intact animals. The protocol described here is designed to image and analyze cell behaviors at both global and local levels in the pupal abdominal epidermis as it grows. The methodology described can be easily adapted to the imaging of cell behaviors at other developmental stages, tissues, subcellular structures, or model organisms.

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

This protocol is designed for the imaging and analysis of the dynamics of cell orientation and tissue growth in the Drosophila abdominal epithelia as the fruit fly undergoes metamorphosis. The methodology described here can be applied to the study of different developmental stages, tissues, and subcellular structures in Drosophila or other model organisms.

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A ...

Whole Animal Imaging of Drosophila melanogaster using Microcomputed Tomography

Biomedical imaging tools permit investigation of molecular mechanisms across spatial scales, from genes to organisms. Drosophila melanogaster, a well-characterized model organism, has benefited from the use of light and electron microscopy to understand gene function at the level of cells and tissues. The application of imaging platforms that allow for an understanding of gene function at the level of the entire intact organism would further enhance our knowledge of genetic mechanisms. Here a whole animal imaging method is presented that outlines the steps needed to visualize Drosophila at any developmental stage using microcomputed tomography (&#181;-CT). The advantages of &#181;-CT include commercially available instrumentation and minimal hands-on time to produce accurate 3D information at micron-level resolution without the need for tissue dissection or clearing methods. Paired with software that accelerate image analysis and 3D rendering, detailed morphometric analysis of any tissue or organ system can be performed to better understand mechanisms of development, physiology, and anatomy for both descriptive and hypothesis testing studies. By utilizing an imaging workflow that incorporates the use of electron microscopy, light microscopy, and &#181;-CT, a thorough evaluation of gene function can be performed, thus furthering the usefulness of this powerful model organism.

Whole Animal Imaging of Drosophila melanogaster using Microcomputed Tomography

A protocol is presented that allows for the visualization of intact Drosophila melanogaster at any stage of development using microcomputed tomography.

Biomedical imaging tools permit investigation of molecular mechanisms across spatial scales, from genes to organisms. Drosophila melanogaster, a ...

Visualizing the Effects of Oxidative Damage on Drosophila Egg Chambers using Live Imaging

Live imaging of Drosophila melanogaster ovaries has been instrumental in understanding a variety of basic cellular processes during development, including ribonucleoprotein particle movement, mRNA localization, organelle movement, and cytoskeletal dynamics. There are several methods for live imaging that have been developed. Due to the fact that each method involves dissecting individual ovarioles placed in media or halocarbon oil, cellular damage due to hypoxia and/or physical manipulation will inevitably occur over time. One downstream effect of hypoxia is to increase oxidative damage in the cells. The purpose of this protocol is to use live imaging to visualize the effects of oxidative damage on the localization and dynamics of subcellular structures in Drosophila ovaries after induction of controlled cellular damage. Here, we use hydrogen peroxide to induce cellular oxidative damage and give examples of the effects of such damage on two subcellular structures, mitochondria and Clu bliss particles. However, this method is applicable to any subcellular structure. The limitations are that hydrogen peroxide can only be added to aqueous media and would not work for imaging that uses halocarbon oil. The advantages are that hydrogen peroxide is readily available and inexpensive, acts quickly, its concentrations can be modulated, and oxidative damage is a good approximation of damage caused by hypoxia as well as general tissue damage due to manipulation.

The objective of this protocol is to use live imaging to visualize the effects of oxidative damage on the localization and dynamics of subcellular structures in Drosophila ovaries.

Live imaging of Drosophila melanogaster ovaries has been instrumental in understanding a variety of basic cellular processes during development, including ...

Nuclear Migration in the Drosophila Oocyte

Live cell imaging is particularly necessary to understand the cellular and molecular mechanisms that regulate organelle movements, cytoskeleton rearrangements, or polarity patterning within the cells. When studying oocyte nucleus positioning, live-imaging techniques are essential to capture the dynamic events of this process. The Drosophila egg chamber is a multicellular structure and an excellent model system to study this phenomenon because of its large size and availability of numerous genetic tools. During Drosophila mid-oogenesis, the nucleus migrates from a central position within the oocyte to adopt an asymmetric position mediated by microtubule-generated forces. This migration and positioning of the nucleus are necessary to determine the polarity axes of the embryo and the subsequent adult fly. One characteristic of this migration is that it occurs in three dimensions (3D), creating a necessity for live imaging. Thus, to study the mechanisms that regulate nuclear migration, we have developed a protocol to culture the dissected egg chambers and perform live imaging for 12 h by time-lapse acquisitions using spinning-disk confocal microscopy. Overall, our conditions allow us to preserve Drosophila egg chambers alive for a long period of time, thereby enabling the completion of nuclear migration to be visualized in a large number of samples in 3D.

Nuclear Migration in the Drosophila Oocyte

In Drosophila, the oocyte nucleus undergoes microtubule-dependent migration during oogenesis. Here, we describe a protocol that was developed to follow the migration by performing live imaging on egg chambers ex-vivo. Our procedure maintains egg chambers alive for 12 h to acquire multi-position 3D time-lapse movies using spinning-disk confocal microscopy.

Live cell imaging is particularly necessary to understand the cellular and molecular mechanisms that regulate organelle movements, cytoskeleton ...

Upright Imaging of Drosophila Embryos

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current techniques used to view such processes are somewhat limited. The following protocol describes a new technique for mounting fixed and labeled Drosophila embryos for coronal viewing with confocal imaging. This method consists of embedding embryos between two layers of glycerin jelly mounting media, and imaging jelly strips positioned upright. The first step for sandwiching the embryos is to make a thin bedding of glycerin jelly on a slide. Next, embryos are carefully aligned on this surface and covered with a second layer of jelly. After the second layer is solidified, strips of jelly are cut and flipped upright for imaging. Alternatives are described for visualizing the embryos depending upon the type of microscope stand to be used. Since all cells along the dorsal-ventral axis are imaged within a single confocal Z-plane, our method allows precise measurement and comparison of fluorescent signals without photobleaching or light scattering common to 3D reconstructions of longitudinally mounted embryos.

Upright Imaging of Drosophila Embryos

Here we present a mounting protocol for stained Drosophila embryos in an upright position that allows imaging of cross-sections using Confocal microscopy.

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current ...

Detecting Anastasis In Vivo by CaspaseTracker Biosensor

Anastasis (Greek for &#8220;rising to life&#8221;) is a recently discovered cell recovery phenomenon whereby dying cells can reverse late-stage cell death processes that are generally assumed to be intrinsically irreversible. Promoting anastasis could in principle rescue or preserve injured cells that are difficult to replace such as cardiomyocytes or neurons, thereby facilitating tissue recovery. Conversely, suppressing anastasis in cancer cells, undergoing apoptosis after anti-cancer therapies, may ensure cancer cell death and reduce the chances of recurrence. However, these studies have been hampered by the lack of tools for tracking the fate of cells that undergo anastasis in live animals. The challenge is to identify the cells that have reversed the cell death process despite their morphologically normal appearance after recovery. To overcome this difficulty, we have developed Drosophila and mammalian CaspaseTracker biosensor systems that can identify and permanently track the anastatic cells in vitro or in vivo. Here, we present in vivo protocols for the generation and use of the CaspaseTracker dual biosensor system to detect and track anastasis in Drosophila melanogaster after transient exposure to cell death stimuli. While conventional biosensors and protocols can label cells actively undergoing apoptotic cell death, the CaspaseTracker biosensor can permanently label cells that have recovered after caspase activation - a hallmark of late-stage apoptosis, and simultaneously identify active apoptotic processes. This biosensor can also track the recovery of the cells that attempted other forms of cell death that directly or indirectly involved caspase activity. Therefore, this protocol enables us to continuously track the fate of these cells and their progeny, facilitating future studies of the biological functions, molecular mechanisms, physiological and pathological consequences, and therapeutic implications of anastasis. We also discuss the appropriate controls to distinguish cells that undergo anastasis from those that display non-apoptotic caspase activity in vivo.

Detecting Anastasis In Vivo by CaspaseTracker Biosensor

Anastasis is technically challenging to detect in vivo because the cells that have reversed the cell death process can be morphologically indistinguishable from normal healthy cells. Here we describe protocols for detecting and tracking cells that undergo anastasis in live animals by using our newly developed in vivo CaspaseTracker biosensor system.

Anastasis (Greek for &#8220;rising to life&#8221;) is a recently discovered cell recovery phenomenon whereby dying cells can reverse late-stage cell death ...

Medicine

Drosophila Pupal Abdomen Immunohistochemistry

The Drosophila pupal abdomen is an established model system for the study of epithelial morphogenesis and the development of sexually dimorphic morphologies 1-3. During pupation, which spans approximately 96 hours (at 25 &deg;C), proliferating populations of imaginal cells replace the larval epidermis to generate the adult abdominal segments. These imaginal cells, born during embryogenesis, exist as lateral pairs of histoblast nests in each abdominal segment of the larvae. Four pairs of histoblast nests give rise to the adult dorsal cuticle (anterior and posterior dorsal nests), the ventral cuticle (ventral nests) and the spiracles associated with each segment (spiracle nests) 4. Upon puparation, these diploid cells (distinguishable by size from the larger polyploid larval epidermal cells- LECs) begin a stereotypical process of proliferation, migration and replacement of the LECs. Various molecular and genetic tools can be employed to investigate the contributions of genetic pathways involved in morphogenesis of the adult abdomen. Ultimate adult phenotypes are typically analyzed following dissection of adult abdominal cuticles. However, investigation of the underlying molecular processes requires immunohistochemical analyses of the pupal epithelium, which present unique challenges. Temporally dynamic morphogenesis and the interactions of two distinct epithelial populations (larval and imaginal) generate a fragile tissue prone to excessive cell loss during dissection and subsequent processing. We have developed methods of dissection, fixation, mounting and imaging of the Drosophila pupal abdominem epithelium for immunohistochemical studies that generate consistent high quality samples suitable for confocal or standard fluorescent microscopy.

Drosophila Pupal Abdomen Immunohistochemistry

Antibody staining of the Drosophila pupae can enhance genetic analyses of adult abdominal developmental genetics. We present our protocol for dissection, fixation and antibody staining of staged Drosophila pupal abdomen.

Upright Imaging of Drosophila Egg Chambers

Summary

Explore More Videos

Chapters in this video

Live-imaging of the Drosophila Pupal Eye

Imaging Calcium in Drosophila at Egg Activation

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

Whole Animal Imaging of Drosophila melanogaster using Microcomputed Tomography

Visualizing the Effects of Oxidative Damage on Drosophila Egg Chambers using Live Imaging

Nuclear Migration in the Drosophila Oocyte

Upright Imaging of Drosophila Embryos

Detecting Anastasis In Vivo by CaspaseTracker Biosensor

Drosophila Pupal Abdomen Immunohistochemistry