We thank Acaimo Gonz&#225;lez Reyes and Mar&#237;a Olmedo L&#243;pez for helpful comments on the manuscript. This study was supported by PID2021-125480NB-I00 (H. S-G), &#34;Ayudas a la contrataci&#243;n de personal Investigador Doctor&#34; from Junta de Andaluc&#237;a (J. G-M), &#34;Ayuda a proyectos de investigaci&#243;n precompetitivos&#34; from VI and VII PPIT-Seville University (P. R-R) and &#34;Contrato de Acceso de I+D+i&#34; from VI PPIT-Seville University (P. R-R). We extend our acknowledgments to the Company of Biologists Ltd for kindly providing rights to share images adapted from the original publication (doi:10.1242/DEV.199716).

NOTE: Step 1.4 must be done at least 2 days in advance. Steps 2.1 and 2.2 can be done 1 day in advance.
1. Pre-experimental setup I
<ol>
	<li>Prepare 100 &#181;L aliquots of streptomycin/penicillin antibiotic mix (10,000 U/mL penicilin and 10 mg/mL streptomycin) and store at -20 &#176;C until use.</li>
	<li>Prepare 15 mL of the following solutions using autoclaved pure water, store them at 4 &#176;C, and use them within a month: Sodium bicarbonate (NaHCO3) 0.1 M pH 8.0 and 0.1% Tween 20.</li>
	<li>Prepare the following culture medium and solution in a laminar flow hood to avoid contamination. Use sterilized filter tips and tubes.
	<ol>
		<li>Prepare 2 mL aliquots of undiluted fetal bovine serum (FBS) and store at -20 &#176;C until use in aseptic conditions.</li>
		<li>Prepare Schneider&#39;s Drosophila medium supplemented with 15% FBS and 0.6% streptomycin/penicillin antibiotic mix. Prepare 7 mL aliquots and store at 4 &#176;C. One aliquot is enough for two MatTek plates (see section 2).
		<ol>
			<li>Prepare Ringer&#39;s solution (128 mM NaCl, 2 mM KCl, 1.8 mM CaCl2, 4 mM MgCl2, 35.5 mM Sucrose, 5 mM Hepes in autoclaved pure water; pH 6.9). Prepare 1 mL aliquots and store at -20 &#176;C until use.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Collect 1-2 days old flies expressing constitutively and ubiquitously the Par-1 protein fused to GFP (poly Ubiquitin-mGFP6::par-1 line) into a new tube with fly food supplemented with yeast powder. Culture them for 2 days at 25 &#176;C in an incubator before dissection. 
	NOTE: This step can be done with any fluorescently tagged reporter of interest.</li>
</ol>
2. Glass bottom plate coating and preparation
<ol>
	<li>Place a 3 &#181;L-drop of a specific Cell-Tak adhesive (henceforth referred to as cell and tissue adhesive) in the center of the coverslip of a 35-mm poly-D-lysine coated plate (glass bottom dish, see Table of Materials) without manual spreading. Use sterile tips. 
	NOTE: The cell and tissue adhesive should be stored at 4 &#176;C. In the case of experiments with two genetic conditions (typically, control and mutant conditions), place two individual, well-separated drops of the adhesive in the same glass bottom plate. This will ensure that both genetic backgrounds are imaged in the same conditions.</li>
	<li>Immediately after, add an equal volume (3 &#181;L in this protocol setup) of 0.1 M NaHCO3 pH 8.0 carefully into the 3 &#181;L adhesive drop as recommended by the manufacturer and mix by pipetting.</li>
	<li>For complete evaporation, incubate the plate at room temperature (RT) for 20 min and then keep at 4 &#176;C overnight to be used the next day. Alternatively, incubate the prepared plate at 37 &#176;C for 25 min until complete evaporation and proceed directly to the next step. In both cases, maintain the plate with its cover. Use sterile tips.</li>
	<li>Allow the prepared glass bottom dish to reach RT by placing it on the bench for 15-20 min before use.</li>
	<li>Wash 3 times the prepared plate carefully. Avoid touching and disturbing the adhesive layer(s). In each wash, add 6 &#181;L of autoclaved pure water and then remove the water using a clean tip for each of the three washes to eliminate residues of the NaHCO3 solution.</li>
	<li>Allow the complete evaporation of the water by placing the clean and prepared plate at RT&#160;while dissecting for 5-10 min with its lid on to avoid dust particles.</li>
</ol>
3. Drosophila ovary dissection and ovariole isolation and mounting
NOTE: The ovary dissection is performed in Ringer&#39;s solution (instead of Schneider&#39;s medium) without FBS to prevent the presence of proteins that could interfere with the sticking of the sample tissue to the adhesive.
<ol>
	<li>Clean a three-well dissection dish, forceps, and dissection needles with 70% ethanol before starting.</li>
	<li>Add a sufficient volume (~200 &#181;L) of Ringer&#39;s solution in each of the three wells.</li>
	<li>Dissect the ovaries of 5 yeast-fed females in 200 &#181;L of Ringer&#39;s solution in the first well with the help of two pairs of forceps following standard procedures10,11. Grab the Drosophila female at the intersection of the thorax and abdomen with one of the forceps. Then, gently tear the cuticle at the posterior abdomen and pull the abdomen cuticle until the two ovaries are exposed.</li>
	<li>Transfer the dissected ovaries to the second well to reduce contamination with remnants of other tissues or debris resulting from dissection.</li>
	<li>Remove the muscle sheath of 15-20 ovarioles with the help of a stereomicroscope (zoom 8X-13.5X) with light-emitting diode (LED) illumination and fine-point forceps. Use one pair of forceps to anchor the entire body of the ovary, and use the second one to smoothly grab a late-stage egg chamber and gently stretch until the muscle breaks and lags behind. 
	NOTE: It is essential to avoid damage to the germaria just by reducing the contact with this part of the ovaries. Remove egg chambers older than stages 8-9.</li>
	<li>Transfer one by one the 15-20 muscle sheath-free ovarioles using the forceps to the third well containing 200 &#181;L of Ringer&#39;s medium.</li>
	<li>Pre-wet a 20-200 &#181;L tip with 0.1% Tween 20 by pipetting. This averts the ovarioles&#39; adhesion to the tip&#39;s plastic walls.</li>
	<li>Transfer 6 &#181;L of of Ringer&#39;s solution containing the muscle sheath-free ovarioles using a pre-wetted tip to the prepared adhesive plate (kept at RT). 
	NOTE: Avoid all contact between the tip and the adhesive layer to prevent germaria detachment after ovariole deposition (next step). Approximately 30% of germaria do not adhere as expected.</li>
	<li>To ensure that the transferred ovarioles adhere to the adhesive surface, carefully press them down to the bottom of the Ringer&#39;s drop until they contact the adhesive surface using a dissection needle or forceps. 
	NOTE: Do not try to rearrange the ovarioles once they settle.</li>
	<li>Fill the MatTek plate carefully with 3 mL of Schneider&#39;s medium supplemented with the 15% FBS and 0.6% streptomycin/penicillin antibiotic mix (step 1.3). To prevent the detachment of the ovarioles, add slowly 1 mL at a time around the outer edge of the plate.</li>
	<li>Transport the plate containing the muscle sheath-free ovarioles on a flat surface to the microscope station.</li>
</ol>
4. Live imaging using a confocal microscope
<ol>
	<li>Place the plate on the 63x objective of a spinning-disk microscope or a confocal microscope (similar equipment can be used) and focus the sample. Select the middle z plane for each germarium.</li>
	<li>Configure the following experimental parameters: capture type: 3D time-lapse, adjust the laser power of 70/100, excitation wavelength: 488 nm, range of the z-stack: 30 &#181;m and distance between Z-stacks: 1.2 &#181;m, duration: 17 h, interval between time-points: every 10 min.</li>
	<li>If available, using a multi-position option, redefine the XY coordinates of the samples and select a Z-plane in the middle of each germarium, as the z-stacks will be centered in this position. Start the acquisition. 
	NOTE: As the normal width of a germarium is 30 &#181;m, each Z-stack will be ~25-27 sections. Depending on the confocal, at least 25 germaria can be imaged within the 10 min intervals. Normally, 40x/1.30 NA or 63x/1.40 NA oil-immersion, planapo-corrected objectives are used.</li>
	<li>Analyze the images in time and Z-axis to visualize specific changes in the spectrosome morphology using Imaris or ImageJ (Fiji) software12,13. Specific times on their observation are displayed (Figure 1B&#39;). To mount the movies, manually select the best Z plane at each time point.</li>
</ol>

Real-Time Imaging of Germline Stem Cells in Drosophila Ex Vivo Germaria Culture

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The authors declare that they have no competing financial interests.

Here we present a protocol to monitor the Drosophila melanogaster GSC niche for a prolonged period of time, up to 16 h. Most of the protocols to film Drosophila biological processes ex vivo focus on shorter time windows and multiple examples can be found for imaginal wing discs15,16,17, embryo gonads18,19 or ovaries20<su...

Live imaging of biological processes is instrumental in obtaining direct experimental evidence. The combination of advanced confocal microscopy and the optimization of methodologies permits the exploration of multiple biological events with high precision. Optimizing steps in protocols such as tissue manipulation and dissection, sample preparation and preservation, and microscopy acquisition settings is critical to maximize the reliability and robustness of the results obtained. Here, we present a protocol to monitor samples for extended imaging, which is specially focused on Drosophila melanogaster ovaries. The Drosophila melanogaster ovary is an excellent model system for the analyses of a wide range of developmental processes. Among many others, this reproductive organ of the fruit fly Drosophila melanogaster contains a very well-defined adult stem cell niche. This GSC niche sustains the development of the female gametes during adulthood. The Drosophila ovaries are composed of approximately 18 ovarioles, where egg chambers are developed in the germarium. In the tip of each germarium, 2-4 GSCs are maintained in a somatic cellular niche mainly formed by a terminal filament of 8-10 cells, a rosette of 5-8 cap cells, and 2-3 anterior escort cells (Figure 1A). This somatic niche provides the GSCs with essential signals and physical support to maintain their stemness, control proliferation, and prevent differentiation1,2,3,4,5.
The GSCs normally divide asymmetrically to generate a new stem cell that keeps contact with the somatic niche and a daughter cell, the cystoblast (CB), which loses direct contact with the somatic cap cells and differentiates. GSCs and CBs contain a highly dynamic and cytoplasmic organelle, the spectrosome, whose main function is the correct orientation of the mitotic spindle during mitosis6. The CB divides 4 times with incomplete cytokinesis to develop 16-cell cysts, where one of the germline cells specifies into the oocyte and the other 15 cells become nurse cells. The CB spectrosome grows into a branched structure called the fusome which connects the 16-cell interconnected germline cells. The spectrosome is enriched in small vesicles and skeletal proteins such as the serine-threonine kinase Par-1 and the membrane component Hu-li tai shao (Hts)7. During the GSC cell cycle, the spectrosome grows by the addition of new material and changes its shape, allowing the identification of the G1, S, G2, and M phases (Figure 1B)8,9.
We have recently implemented an ex vivo culture method of the Drosophila germarium that allows imaging of live GSCs for up to 16 h. Since these cells divide once every 15.5 h on average8, this method permits the filming of large portions of the GSC cell cycle. Thus, in combination with other tools, our culturing method has allowed the description of spectrosome morphology during the GSC cell cycle and the analysis of the duration of the different cell cycle phases in vivo8 (Figure 1C). Here, we provide a detailed protocol of this extended live imaging method supported by a step-by-step guided video that describes the methodology (Figure 2).

<table><tbody><tr><td>9-well glass plate</td><td>Corning&amp;nbsp;</td><td>7220-85</td><td>n/a</td></tr><tr><td>Calcium chloride dihydrate</td><td>Sigma Aldrich</td><td>C3881</td><td>CaCl&lt;sub&gt;2&lt;/sub&gt;&lt;sub&gt;&amp;middot;&lt;/sub&gt;2H&lt;sub&gt;2&lt;/sub&gt;O; To prepare Ringer&#039;s solution</td></tr><tr><td>Cell-Tak</td><td>Corning&amp;nbsp;</td><td>354240</td><td>Cell and Tissue Adhesive used to attach cells or tissue sections to many types of surfaces, including plastic, glass, metal, FEP Polymer, and biological materials.</td></tr><tr><td>Dumont #5 Forceps&amp;nbsp;</td><td>Finescience</td><td>11252-20</td><td>n/a</td></tr><tr><td>Dumont #55 Forceps&amp;nbsp;</td><td>Finescience</td><td>11255-20</td><td>Dimensions 0.05 mm x 0.02 mm and length 11 cm</td></tr><tr><td>Fetal Bovine Serum</td><td>Gibco</td><td>10500-064</td><td>Qualified, heat inactivated, E.U.-approved, South America Origin&amp;nbsp;</td></tr><tr><td>Glass bottom dishes</td><td>MatTek</td><td>P35GC-1.5-10-C</td><td>35 mm Dish, No. 1.5 Coverslip, 10 mm Glass Diameter,&amp;nbsp;Poly-D-Lysine Coated</td></tr><tr><td>HEPES</td><td>Sigma Aldrich</td><td>H4034</td><td>To prepare Ringer&#039;s solution</td></tr><tr><td>Magnesium chloride&amp;nbsp;</td><td>Sigma Aldrich</td><td>M1028</td><td>MgCl&lt;sub&gt;2&lt;/sub&gt;; To prepare Ringer&#039;s solution</td></tr><tr><td>Needle Holder</td><td>Roboz&amp;nbsp;</td><td>RS-6061</td><td>Light weight, hollow stainless steel handle; 4 3/4&quot; Long.</td></tr><tr><td>Nikon SMZ18 binocular&amp;nbsp;</td><td>Nikon</td><td></td><td>https://www.microscope.&lt;br/&gt; healthcare.nikon.com/products/stereomicroscopes-macroscopes/smz25-smz18</td></tr><tr><td>Penicillin-Streptomycin</td><td>Thermo Fisher Scientific</td><td>15140122</td><td>10,000 U/mL</td></tr><tr><td>Potassium chloride</td><td>Sigma Aldrich</td><td>P3911</td><td>KCl; To prepare Ringer&#039;s solution</td></tr><tr><td>Schneider&#039;s Drosophila Medium</td><td>Biowest&amp;nbsp;</td><td>L0207</td><td>Cell culture medium for insect cells</td></tr><tr><td>Sodium bicarbonate</td><td>Sigma Aldrich</td><td>S8875&amp;nbsp;</td><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;; &amp;ge;99.5%, powder</td></tr><tr><td>Sodium chloride</td><td>Sigma Aldrich</td><td>S9888</td><td>NaCl; To prepare Ringer&#039;s solution</td></tr><tr><td>Sucrose</td><td>Sigma Aldrich</td><td>S9378</td><td>&amp;ge;99.5% (GC); To prepare Ringer&#039;s solution</td></tr><tr><td>Tungsten Dissection Needle</td><td>Roboz&amp;nbsp;</td><td>RS-6064</td><td>0.25 mm, Ultra Fine, 1 Micron Tip (Pk 10)</td></tr><tr><td>Tween 20</td><td>Sigma Aldrich</td><td>P9416</td><td>for molecular biology, viscous liquid</td></tr><tr><td>Yeast powder</td><td>Sigma Aldrich</td><td>51475</td><td>n/a</td></tr></tbody></table>

extended live imaging of female drosophila melanogaster germline stem cell niches

Live imaging methods allow the analysis of dynamic cellular processes in detail and in real-time. The Drosophila ovary represents an excellent model to explore the dynamics of a myriad of developmental processes, such as cell division, stemness, differentiation, migration, apoptosis, autophagy, cellular adhesion, etc., over time. Recently, we have implemented an extended ex vivo culture and live imaging of the female Drosophila GSC niche. Using a Drosophila line harboring a GFP::Par-1 transgene as an example, this method allows the visualization of the GSCs&#39; asymmetric division within their niche and the description of the changes in the spectrosome morphology along the cell cycle. Here, we present a detailed protocol for the ex vivo culture of Drosophila germaria, enabling prolonged visualization of the female GSC niche. Importantly, this protocol is broadly applicable to live imaging GSCs with multiple fluorescently tagged proteins of interest that are available in stock centers and/or in the Drosophila research community.

With this extended live imaging protocol, we can record the asymmetric mitosis of female GSCs inside their niche without apparent biological disturbances. To do so, we use germaria expressing ubiquitously the GFP::Par-1 protein, which discriminates between five distinct spectrosome morphologies through the GSC cell cycle: Round, Plug, Bar, Fusing, and Exclamation point (Figure 1B,C). In a more detailed description, we observed that the strong GFP:Par1 signal of Round-G2 spec...

Extended Live Imaging of Female Drosophila melanogaster Germline Stem Cell Niches

Here, we provide a detailed protocol to perform live imaging of the asymmetric division of germline stem cells (GSCs) in the Drosophila ovarian niche. We use a transgenic line that ubiquitously expresses a green fluorescent protein (GFP) fusion of the spectrosome protein Par-1.

Live imaging methods allow the analysis of dynamic cellular processes in detail and in real-time. The Drosophila ovary represents an excellent model to ...

extended-live-imaging-female-drosophila-melanogaster-germline-stem

Research

JoVE Journal

Developmental Biology

578 Views.  CSIC/Universidad Pablo de Olavide/JA. We zijn geïnteresseerd in het ontdekken van nieuwe cellulaire en moleculaire mechanismen die de stam van cellen beheersen. Ons belangrijkste doel is om te ontcijferen hoe stamcellen differentiatie in de niches vermijden. Aangezien we onder experimentele omstandigheden werken, is het essentieel om de weefselintegriteit te behouden om ervoor te zorgen dat GSC's fysiologisch gedrag ontvangen.Ons protocol handhaaft een gezonde en functionele GSC-niche die...

Video: Extended Live Imaging of Female Drosophila melanogaster Germline Stem Cell Niches

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 ...

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a rapid and high-throughput manner. S2 cells have proven amenable to both fixed- and live-cell imaging applications. Notably, live-cell imaging can yield valuable information about how loss or knockdown of a gene can affect the kinetics and dynamics of key events during cell division, including mitotic spindle assembly, chromosome congression, and segregation, as well as overall cell cycle timing. Here we utilize S2 cells stably transfected with fluorescently tagged mCherry:&#945;-tubulin to mark the mitotic spindle and GFP:CENP-A (referred to as &#8216;CID&#8217; gene in Drosophila) to mark the centromere to analyze the effects of key mitotic genes on the timing of cell divisions, from prophase (specifically at Nuclear Envelope Breakdown; NEBD) to the onset of anaphase. This imaging protocol also allows for the visualization of the spindle microtubule and chromosome dynamics throughout mitosis. Herein, we aim to provide a simple yet comprehensive protocol that will allow readers to easily adapt S2 cells for live imaging experiments. Results obtained from such experiments should expand our understanding of genes involved in the cell division by defining their role in several simultaneous and dynamic events. Observations made in this cell culture system can be validated and further investigated in vivo using the impressive toolkit of genetic approaches in flies.

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Cell divisions can be visualized in real time using fluorescently tagged proteins and time-lapse microscopy. Using the protocol presented here, users can analyze cell division timing dynamics, mitotic spindle assembly, and chromosome congression and segregation. Defects in these events following RNA interference (RNAi)-mediated gene knockdown can be assessed and quantified.

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a ...

Genetics

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and differentiation. Typically, pHi dynamics are determined in cultured cells, which are amenable to measuring and experimentally manipulating pHi. However, the recent development of new tools and methodologies has made it possible to study pHi dynamics within intact, live tissue. For Drosophila research, one important development was the generation of a transgenic line carrying a pHi biosensor, mCherry::pHluorin. Here, we describe a protocol that we routinely use for imaging live Drosophila ovarioles to measure pHi in the epithelial follicle stem cell (FSC) lineage in mCherry::pHluorin transgenic wild type lines; however, the methods described here can be easily adapted for other tissues, including the wing discs and eye epithelium. We describe techniques for expressing mCherry::pHluorin in the FSC lineage, maintaining ovarian tissue during live imaging, and acquiring and analyzing images to obtain pHi values.

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

We provide a protocol for imaging intracellular pH of an epithelial stem cell lineage in live Drosophila ovarian tissue. We describe methods to generate transgenic flies expressing a pH biosensor, mCherry::pHluorin, image the biosensor using quantitative fluorescence imaging, generate standard curves, and convert fluorescence intensity values to pH values.

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Biology

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Neuroscience

Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

The Drosophila melanogaster male embryonic gonad is an advantageous model to study various aspects of developmental biology including, but not limited to, germ cell development, piRNA biology, and niche formation. Here, we present a dissection technique to live-image the gonad ex vivo during a period when in vivo live-imaging is highly ineffective. This protocol outlines how to transfer embryos to an imaging dish, choose appropriately-staged male embryos, and dissect the gonad from its surrounding tissue while still maintaining its structural integrity. Following dissection, gonads can be imaged using a confocal microscope to visualize dynamic cellular processes. The dissection procedure requires precise timing and dexterity, but we provide insight on how to prevent common mistakes and how to overcome these challenges. To our knowledge this is the first dissection protocol for the Drosophila embryonic gonad, and will permit live-imaging during an otherwise inaccessible window of time. This technique can be combined with pharmacological or cell-type specific transgenic manipulations to study any dynamic processes occurring within or between the&#160;cells in their natural gonadal environment.

Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

Here, we provide a dissection protocol required to live-image the late embryonic Drosophila male gonad. This protocol will permit observation of dynamic cellular processes under normal conditions or after transgenic or pharmacological manipulation.

The Drosophila melanogaster male embryonic gonad is an advantageous model to study various aspects of developmental biology including, but not limited to, ...

Imaging Live Brain Explants from Drosophila melanogaster Third Instar Larvae

Live-Cell Imaging of Drosophila melanogaster Third Instar Larval Brains

Drosophila neural stem cells (neuroblasts, NBs hereafter) undergo asymmetric divisions, regenerating the self-renewing neuroblast, while also forming a differentiating ganglion mother cell (GMC), which will undergo one additional division to give rise to two neurons or glia. Studies in NBs have uncovered the molecular mechanisms underlying cell polarity, spindle orientation, neural stem cell self-renewal, and differentiation. These asymmetric cell divisions are readily observable via live-cell imaging, making larval NBs ideally suited for investigating the spatiotemporal dynamics of asymmetric cell division in living tissue. When properly dissected and imaged in nutrient-supplemented medium, NBs in explant brains robustly divide for 12-20 h. Previously described methods are technically difficult and may be challenging to those new to the field. Here, a protocol is described for the preparation, dissection, mounting, and imaging of live third-instar larval brain explants using fat body supplements. Potential problems are also discussed, and examples are provided for how this technique can be used.

Author Spotlight: Investigating Asymmetric Cell Division Dynamics: A Protocol for Live-Imaging of Drosophila Larval Brain Explants 

Here, we discuss a workflow to prepare, dissect, mount, and image live explant brains from Drosophila melanogaster third instar larvae to observe the cellular and subcellular dynamics under physiological conditions.

Drosophila neural stem cells (neuroblasts, NBs hereafter) undergo asymmetric divisions, regenerating the self-renewing neuroblast, while also forming a ...

Extended Live Imaging of Drosophila Pupal Leg Morphogenesis Without Puparium

Long-Term Live Imaging of Drosophila Pupal Leg Development After Puparium Removal

Over the past decades, significant progress has been made in understanding the mechanisms of cell fate determination. However, the process by which fate-determined cells form three-dimensional organismal shapes remains unclear. Recent advances in confocal microscopy have facilitated efforts to observe cell dynamics during development through live imaging. The Drosophila melanogaster pupa is ideal for live imaging due to its immobility, transparent pupal cuticle, and the availability of fluorescent reporter lines. A primary challenge for imaging is the puparium, the cuticle surrounding the pupa, which obstructs optical imaging. While previous methods involved either partial or complete removal of the puparium, maintaining pupal viability for extended periods after this procedure has remained challenging. Here, a simple method is presented for days-long live imaging of the Drosophila leg during the pupal stage, involving complete puparium removal. The method includes removing the puparium from a pupa adhered to double-sided tape, followed by assembling a small chamber on a glass-bottom dish to enclose the pupa and a drop of water. This setup is straightforward, reliable, and supports extended pupal survival by preventing desiccation. Long-term live imaging of the Drosophila pupa has significantly contributed to capturing how the adult leg undergoes dramatic three-dimensional structural changes over 2-3 days. These changes include the transient formation of an intriguing structure (the Parthenon-like structure) by epithelial cells, rapid tissue narrowing, joint formation, and bristle elongation. This method is applicable to the observation of various tissues and can potentially be combined with other techniques, such as optical gene induction, to advance the understanding of cell dynamics during the final shape formation of tissues in the pupal stage.

Long-Term Live Imaging of Drosophila Pupal Leg Development After Puparium Removal

Here, a protocol is presented for long-term live imaging of the Drosophila leg during the pupal stage, involving complete removal of the puparium. This protocol can also be applied to other tissues.

Over the past decades, significant progress has been made in understanding the mechanisms of cell fate determination. However, the process by which ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Extended Live Imaging of Female Drosophila melanogaster Germline Stem Cell Niches

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