We would like to thank Lindsey W. Plasschaert and Justin Sui for their substantial contributions to the early development of this protocol. The authors are grateful to the fly community for their generosity with reagents, and particularly to Ruth Lehmann and Benjamin Lin for their gift of the nos5&#8217;-Lifeact-tdtomato p2a tdkatushka2 Caax nos3&#39; line prior to its publication. Stocks obtained from the Bloomington Drosophila Stock Center (NIH P40OD018537) were used in this study. This work was supported by NIH RO1 GM060804, R33AG04791503 and R35GM136270 (S.D) as well as training grants T32GM007229 (B.W.) and F32GM125123 (L.A.).

1. Day-before-dissection preparation
<ol>
	<li>Electrolytically sharpen a tungsten needle24 so that the resulting diameter is approximately 0.03 mm. Adjust the voltage supplied to approximately 14 V, and use 3.3 M NaOH. Sharpening should take no more than 1 or 2 min. 
	CAUTION: NaOH is highly corrosive and will cause burns upon contact with skin. Wear gloves and goggles while handling, and work inside a fume hood. 
	NOTE: After use, store NaOH in a polypropylene Falcon tube.</li>
	<li>Make the prepared imaging media. In a 15 mL conical tube, combine 4.25 mL of Schneider&#8217;s media with 750 &#181;L of Fetal Bovine Serum (FBS, 15% final concentration) and 27.5 &#181;L of Penicillin-Streptomycin (0.05 U/&#181;L of Penicillin, 0.05 &#181;g/&#181;L final concentration). Store this prepared imaging media at 4 &#176;C.</li>
	<li>Make heptane-glue solution. Add about 0.5 mL heptane to a 20 mL sealable vial stuffed with about 20 cm double-sided tape. Rock on a nutator for about an hour, or stir with a pipette tip until a consistency between that of water and glycerol is achieved. This solution of dissolved glue will last for several days before the heptane evaporates. To freshen the solution, add an additional 0.5 mL heptane, and rock. 
	NOTE: An effective heptane-glue solution can be prepared using various ratios of heptane to tape, as well as varying durations of rocking/stirring. The above specifications are merely suggestions for preparing or freshening one such adequate solution.</li>
</ol>
2. Embryo collection&#8212;15&#8211;17 h before dissection
<ol>
	<li>Add fresh yeast paste to an apple juice agar plate25. Set up an embryo collection cage by adding adult flies (less than 10 days old) to an empty food bottle, perforated with small holes26. Cap the collection cage using the yeasted agar plate, taped to the bottle, and place the cage with the plate side down in a 25 &#176;C dark incubator for 1 h. 
	NOTE: The flies must express a transgene that will fluorescently mark the gonad, for example, six4-eGFP::Moesin20, because dissection of embryos will take place under a stereo-fluorescent microscope. See Table of Materials for a list of genotypes used to mark the gonad.</li>
	<li>Remove the cage from the incubator and discard this first collection. Replace it with a freshly yeasted agar plate and place the cage back in the incubator for 2 h. 
	NOTE: The first embryo collection is used to clear females of fertilized, developing embryos, to achieve a tightly timed second collection of embryos.</li>
	<li>Remove the agar plate from the cage and place it (with yeasted side facing upward) on a moist paper towel inside a sealable plastic container. Place this humid chamber in a 25 &#176;C incubator to age the embryos for 14.5 h (final ages, 14.5&#8211;16.5 h after egg lay). 
	NOTE: Immediately before beginning step 2.4, complete steps 3.1 and 3.2.</li>
	<li>Remove the agar plate from the incubator, and using a squirt bottle, add enough water to the agar plate to dissolve the yeast paste by brushing lightly with a paint brush. Rinse the embryos and dissolved yeast into a small mesh screen basket sitting inside a weigh boat. Rinse the basket with the water squirt bottle until most yeast paste has filtered through the mesh.</li>
	<li>Remove the water from the weigh boat and place the basket back inside. Dechorionate the embryos by immersing them in a 50% bleach solution, using a squirt bottle. The bleach solution should have a depth of 3&#8211;5 mm. Keep the embryos submerged in bleach for ~2 min, with occasional swirling. 
	CAUTION: Bleach is corrosive and can irritate or damage eyes and the respiratory tract. Wear gloves and goggles while handling bleach. 
	NOTE: During this time, complete as much as possible of step 3.3. Dechorionation progress can be checked by placing the basket under a stereo microscope and checking for the absence of the dorsal appendages. Submerge the embryos in the bleach solution for additional time, if necessary.</li>
	<li>Discard the bleach, and thoroughly rinse the embryos inside the basket with water from a squirt bottle (for ~3 s, blotting the mesh basket with paper towels; repeat 2&#8211;3 times).</li>
</ol>
3. Day-of-dissection preparation
<ol>
	<li>Add 30 &#181;L insulin (10 mg/mL) to 1,500 &#181;L prepared imaging media (see step 1.2) in a 1.5 mL Eppendorf tube (0.2 mg/mL final concentration). Mix well and leave the tube on the benchtop to equilibrate to room temperature.</li>
	<li>Prepare coverslip strips coated with glue.
	<ol>
		<li>Use a diamond-tipped knife to cut a 22 mm x 22 mm coverslip into four, equally sized strips (Figure 1A).</li>
		<li>Use forceps to pick up one strip and spread a total of approximately 30 &#181;L of heptane-glue solution onto both sides of the strip (Figure 1B). To achieve an even layer of glue residue, tilt the strip at various angles while the heptane evaporates.</li>
		<li>Store the glue-covered strip in an empty slot of a coverslip box; to maintain its stickiness, place the strip in a slanted, but upright position, leaning against the edges of the box to minimize contact with box surfaces (Figure 1C). Close the box so that particulates in the air do not coat the glue and lessen its stickiness.</li>
	</ol>
	</li>
	<li>Place the following items on the benchtop: a 6-inch glass Pasteur pipette, a microscope slide, a P200 and a P1000 pipetteman with the appropriate pipette tips, Ringer&#8217;s solution27 (pH adjusted to 7.3 with NaOH), and an uncovered, Poly-D-Lysine-coated 35 mm imaging dish.</li>
	<li>Transfer embryos from the mesh screen basket to a small watch glass filled with 500&#8211;750 &#181;L heptane.
	<ol>
		<li>Blot the sides and bottom of the basket dry with a tissue wipe. Moisten a paintbrush in the heptane, touch the paintbrush to the embryos (the hydrophobic vitelline membrane should adhere to the bristles), and dip the paintbrush back into the heptane in the watch glass (embryos should sink to bottom). 
		NOTE: The next steps must be performed as quickly as possible to prevent the embryos from drying out. Embryos should not be exposed to air for more than 20 s.</li>
	</ol>
	</li>
	<li>Transfer the embryos onto the microscope slide using a Pasteur pipette (Figure 1D). Draw embryos into the pipette slowly and limit them to the narrow portion of the pipette. Pipette embryos onto the slide slowly, such that they aggregate just inside the tip of the pipette before flowing onto the slide. Twist the corner of a tissue wipe into a fine tip, and wick away heptane from embryos on the slide. They will aggregate and cover a smaller area, making it easier to capture them on a glue-covered strip in the next step.</li>
	<li>With forceps, pick up a glue-covered strip, and gently touch it to the embryos (Figure 1E). Place the strip in the imaging dish, embryo side-up, and just outside of the Poly-D-Lysine-coated center. Press the strip onto the dish using forceps, to ensure it is fixed in place. Immediately flood the dish with 2&#8211;3.5 mL Ringer&#8217;s solution; submerge the embryos first to prevent them from drying out (Figure 1F).</li>
</ol>
4. Dissection
NOTE: These steps must be carried out under a stereo-fluorescent microscope.
<ol>
	<li>Devitellinize 10&#8211;15 embryos. 
	NOTE: This may range from two embryos, for beginners, to fifteen embryos, for experts.
	<ol>
		<li>Select Stage 16 embryos based on gut morphology (Figure 2). At this stage, embryos have three gut constrictions that create four stacked gut segments (Figure 2B,B&#8217;). To begin devitellinization, pierce the selected embryo at one end, preferably the anterior, with the tungsten needle. The embryo may pop out of its vitelline membrane, but if not, peel the membrane off of the embryo. 
		NOTE: Embryos that are too young to dissect have non-regionalized guts (Figure 2C). Dissection of Stage 17 embryos is possible, but more challenging than dissection of Stage 16 embryos because the cuticle is beginning to develop at this stage. Early Stage 17 embryos present with four gut segments that are shifted relative to one another (Figure 2D,D&#8217;).</li>
		<li>Hook the needle through the embryo in a region far from the gonads. Transfer the hooked embryo to the Poly-lysine-coated region of the dish (from here onwards referred to as simply &#8220;cover slip&#8221;) and drag it against the bottom until it adheres. Repeat these steps, and arrange devitellinized embryos in a row along the top of the Poly-lysine-coated cover slip (Figure 3A), leaving plenty of space below for further dissection. 
		NOTE: The gonads appear as small spherical aggregates of cells that should fluoresce brightly, depending on the specific marker used and transgene copy number. At this stage, the gonads are located laterally in segment A5, at approximately 70%&#8211;80% of embryo length from anterior. Throughout the Dissection protocol, the tissue may stick to the needle. To rid the needle of the debris, raise it just above the surface of the Ringer&#8217;s solution. Devitellinization can be untidy&#160;as long as the gonad proper is&#160;disturbed as little as possible.</li>
	</ol>
	</li>
	<li>Finesse the gonads out of the embryo and onto the dish (Figure 3C,D).
	<ol>
		<li>First, filet the embryo to expose its interior (Figure 3C). Slice through the embryo from its center, moving the needle posteriorly, between the gonads. Tease out some internal tissue, as this will stick to the dish much better than the external cuticle. The stickiness of the tissue will enable the next manipulations to reveal the gonad and allow it to adhere to the cover slip. 
		NOTE: If tissue is not sticking to the dish, try coaxing it to an uncontaminated region of the coated cover slip; the outer regions of the cover slip will be particularly sticky. As stated in step 4.1.2, it does not matter if the dissection manipulations result in a mangled embryo carcass, as long as the gonads remain unscathed.</li>
		<li>Use the needle to slice around a gonad until a piece of tissue, including the gonad, is separated from the remaining carcass. With the needle, draw this tissue to a fresh region of coated cover slip, and coax it against the bottom until it adheres to the dish. 
		NOTE: The best imaging will be achieved for those cases where the gonad itself attaches directly to the cover slip, rather than indirect attachment via overlying tissue. Therefore, as tissue is guided away from the carcass, attempt to have the gonad touch the cover slip first, rather than extraneous tissue.</li>
		<li>Remove as much surrounding tissue from the gonad as possible (Figure 3D) by gently dragging adherent tissue away from the gonad with the needle. Avoid touching the gonad directly, which would damage it (Figure 4E). To ensure the gonad is sufficiently adhered to the dish, move the needle in a gentle circular motion around the gonad&#8212;if it moves, touch the needle to the remaining adherent tissue, and direct the gonad toward a fresh region of sticky cover slip. Repeat this process until there is no detectable movement of the gonad.</li>
		<li>Return to the embryo carcass and dissect out the second gonad. Repeat this process on as many embryos as possible but do NOT exceed 25 min. Tissue viability will be compromised in Ringer&#8217;s solution after more than ~40&#8211;45 min.</li>
	</ol>
	</li>
	<li>After dissections are complete, use an indelible marker to add registration marks to the outer rim of the imaging dish to record its orientation during dissection.</li>
	<li>Remove the glue-coated strip from the dish.
	<ol>
		<li>Gently insert the bottom prong of a pair of forceps underneath the strip, clasp and slowly tilt the strip upwards to free it from the dish with as little sloshing of the Ringer&#8217;s solution as possible to minimize disturbance of the adhered gonads. 
		NOTE: The strip should be tilted away from the dissected-out gonads.</li>
	</ol>
	</li>
	<li>Gently carry the dish to the imaging microscope in a manner that avoids sloshing of the Ringer&#8217;s solution.</li>
</ol>
5. Imaging
<ol>
	<li>Place the imaging dish in the stage holder, using the registration marks to place the dish in the approximate orientation as during dissection. Using brightfield microscopy and a low power (~10x) objective, identify and bring into focus any piece of tissue that is adhered to the cover slip. Switch the eyepiece settings to reveal fluorescence, and using the binocular eyepieces, systematically scan the dish, marking the position of each gonad within the imaging software (see Table of Materials). 
	NOTE: Make sure gonads are centered in the field-of-view before marking positions.</li>
	<li>Gently remove the entire stage holder assembly, with the imaging dish in its holder, and place the assembly on the work bench.</li>
	<li>Replace the Ringer&#8217;s solution with the prepared imaging media containing insulin (see steps 1.2 and 3.1).
	<ol>
		<li>Use a P1000 to remove all Ringer&#8217;s solution from the inside upper ledge of the imaging dish (do not pipette Ringer&#8217;s from the central cover slip region). Next, switch to a P200, and place its tip just under the surface of the remaining Ringer&#8217;s in the central region. Carefully remove 50&#8211;100 &#181;L of Ringer&#8217;s; do NOT remove the entire solution.</li>
		<li>Draw up ~200 &#181;L of imaging media, and, again placing the P200 tip just under the surface of the remaining Ringer&#8217;s, slowly add this imaging media. Next, add the remaining imaging media (~1,300 &#181;L) to the dish, starting at the outermost rim of the upper ledge. While pipetting, move toward the central dome of the fluid, and eventually merge the two by brushing the pipette tip across them both. Place the lid on the dish. 
		NOTE: Imaging media should cover the entire inside diameter of the dish, with a depth of about 2 mm, to prevent evaporation (Figure 4D).</li>
	</ol>
	</li>
	<li>Switch the microscope to a higher power objective (63x, 1.2 NA), apply the proper immersion fluid based on the objective used (immersion fluid type and refractive index required by the objective), and then replace the stage holder assembly. Use brightfield microscopy to focus on tissue adhered to the bottom of the imaging dish. 
	NOTE: If the stage was not moved, the last gonad should come into view once the objective is focused.</li>
	<li>Step through each marked gonad position and adjust that position, as necessary. Select which gonads to image based on image clarity, gonad sex, etc. Customize the imaging settings (multi-channel, exposure times, laser intensities, Z-series increments, time-lapse with appropriate intervals, etc.). Begin imaging. 
	NOTE: Male gonads can be identified by the presence of both a niche, and at the opposite end of the gonad, a cluster of small, highly circular somatic cells called male-specific somatic gonadal precursors (msSGPs). If the six4-eGFP::moesin fluorescent marker is used, the niche cells are the second-brightest cells in the gonad, the brightest being the msSGPs. At this stage, female gonads have neither a niche nor msSGPs.</li>
</ol>

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F., Heidmann, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Incorporation+of+Drosophila+CID/CENP-A+and+CENP-C+into+Centromeres+during+Early+Embryonic+Anaphase.">Incorporation of Drosophila CID/CENP-A and CENP-C into Centromeres during Early Embryonic Anaphase.</a> Current Biology. 17, 237-243 (2007).</li><li>Lin, B., Luo, J., Lehmann, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Collectively+stabilizing+and+orienting+posterior+migratory+forces+disperses+cell+clusters+in+vivo.">Collectively stabilizing and orienting posterior migratory forces disperses cell clusters in vivo.</a> Nature Communications. 11, 1-16 (2020).</li><li>Prasad, M., Jang, A. C. C., Starz-Gaiano, M., Melani, M., 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=A+protocol+for+culturing+drosophila+melanogaster+stage+9+egg+chambers+for+live+imaging.">A protocol for culturing drosophila melanogaster stage 9 egg chambers for live imaging.</a> Nature Protocols. 2, 2467-2473 (2007).</li><li>Gabay, L., Lowell, S., Rubin, L. L., Anderson, 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=Deregulation+of+dorsoventral+patterning+by+FGF+confers+trilineage+differentiation+capacity+on+CNS+stem+cells+in+vitro.">Deregulation of dorsoventral patterning by FGF confers trilineage differentiation capacity on CNS stem cells in vitro.</a> Neuron. 40, 485-499 (2003).</li><li>Engler, A. J., Sen, S., Sweeney, H. L., Discher, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Matrix+elasticity+directs+stem+cell+lineage+specification.">Matrix elasticity directs stem cell lineage specification.</a> Cell. 126, 677-689 (2006).</li></ol>

The authors have nothing to disclose.

During gonadogenesis, the embryonic gonad, and particularly the stem cell niche within the male gonad15, undergoes rapid morphological changes. Developmental mechanisms that underlie these dynamic changes are best understood through live-imaging techniques. However, at embryonic Stage 17, in vivo imaging of the gonad is rendered impossible by the onset of large-scale muscle contractions17. With this protocol, we provide a successful alternative: dissection of the g...

The Drosophila melanogaster testis has served as a paradigm for our understanding of many dynamic cellular processes. Studies of this model have shed light on stem cell division regulation1,2,3, germ cell development4,5, piRNA biology6,7,8, and niche-stem cell signaling events9,10,11,12,13. This model is advantageous because it is genetically tractable14,15 and is one of the few where we can live-image stem cells in their natural environment3,16,17,18. However, live-imaging of this model has been limited to adult tissue and early embryonic stages, leaving a gap in our knowledge of gonadal dynamics in the late embryo, the precise stage when the niche is first forming and beginning to function.
The late stage embryonic gonad is a sphere, consisting of somatic niche cells at the anterior, and germ cells encysted by somatic gonadal cells throughout more posterior regions19. This organ can be imaged live in vivo up until early embryonic Stage 1717,20,21. Further imaging is prevented due to initiation of large-scale muscle contractions. These contractions are so severe that they push the gonad out of the imaging frame, and such movement cannot be corrected with imaging software. Our lab is interested in unveiling the mechanisms of niche formation, which occurs during this elusive period for live-imaging. Therefore, we generated an ex vivo approach to live image the gonad starting at embryonic Stage 16, facilitating the study of the cell dynamics during this crucial period of gonad development. Previous work from our lab shows that this ex vivo imaging faithfully recapitulates in vivo gonad development17. This technique is the first and only of its kind for the Drosophila embryonic gonad.
Here, we present the dissection protocol required for ex vivo live-imaging of the gonad during late embryonic stages. This protocol can be combined with pharmacological treatments, or transgenic manipulation of specific cell lineages within the gonad. Using this technique, we have successfully imaged the steps of stem cell niche formation17. This imaging approach is thus instrumental for the field of stem cell biology, as it will enable visualization of the initial stages of niche formation in real time within its natural environment15,17. While this method is beneficial for the field of stem cell biology, it is additionally applicable for visualizing any dynamic processes occurring in the gonad during this developmental timepoint, including cellular rearrangements22, cell adhesion2,12,23, and cell migration23. This dissection protocol will thus enhance our understanding of many fundamental cell biological processes.

<table>
	<tbody>
		<tr>
			<td>Alfa Aesar&#160;Tungsten wire</td>
			<td>Fisher Scientific</td>
			<td>AA10408G6</td>
			<td>0.25mm (0.01 in.) dia., 99.95% (metals basis)</td>
		</tr>
		<tr>
			<td>D. melanogaster: His2Av::mRFP1</td>
			<td>Bloomington Drosophila Stock Center (BDSC)</td>
			<td>FBtp0056035</td>
			<td>Schuh, Lehner, &#38; Heidmann, Current Biology, 2007</td>
		</tr>
		<tr>
			<td>D. melanogaster: nos-lifeact::tdtomato</td>
			<td>Gift from Ruth Lehmann Lab</td>
			<td></td>
			<td>Lin, Luo, &#38; Lehmann, Nature Communications, 2020: nos5&#39;- Lifeact-tdtomato p2a tdkatushka2 Caax nos3&#39;</td>
		</tr>
		<tr>
			<td>D. melanogaster: P-Dsix4-eGFP::Moesin</td>
			<td></td>
			<td>FBtp0083398</td>
			<td>Sano et al., PLoS One, 2012</td>
		</tr>
		<tr>
			<td>Diamond-tipped knife</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Double-sided tape</td>
			<td>Scotch</td>
			<td>665</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>GIBCO</td>
			<td>10082</td>
			<td></td>
		</tr>
		<tr>
			<td>Imaging dish</td>
			<td>MatTek</td>
			<td>P35GC-1.5-14-C</td>
			<td></td>
		</tr>
		<tr>
			<td>Imaging software</td>
			<td>Molecular Devices</td>
			<td></td>
			<td>MetaMorph Microscopy Automation and Image Analysis Software v7.8.4.0</td>
		</tr>
		<tr>
			<td>Insulin, bovine</td>
			<td>Sigma</td>
			<td>l0516</td>
			<td>Store aliquots at 4 &#176;C</td>
		</tr>
		<tr>
			<td>Needle holder</td>
			<td>Fisher Scientific</td>
			<td>08-955</td>
			<td></td>
		</tr>
		<tr>
			<td>Nytex basket</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin/streptomycin</td>
			<td>Corning</td>
			<td>30-002-Cl</td>
			<td></td>
		</tr>
		<tr>
			<td>Ringer&#39;s solution</td>
			<td></td>
			<td></td>
			<td>2 mM MgCl2, 2 mM CaCl2, 130 mM NaCl, 5mM KCl, 36 mM Sucrose, 5mM Hepe&#8217;s Buffer; adjusted with NaOH until pH of 7.3 is achieved</td>
		</tr>
		<tr>
			<td>Schneider&#39;s Insect Media</td>
			<td>GIBCO</td>
			<td>21720-024</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

We illustrate preparation of the imaging dish in Figure 1, as described in &#8220;Day-of-dissection preparation.&#8221; These methods should ultimately result in well-hydrated embryos adhered to a cover slip strip, which is temporarily fixed to the bottom of the dish and submerged in Ringer&#8217;s solution (Figure 1F). A diamond-tipped knife allows one to cleanly slice a 22 x 22 mm cover slip into three to four smaller strips (Figure 1A</st...

Watch this Scientific Journal Video about Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad at JoVE.com

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.

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, ...

dissection-and-live-imaging-of-the-late-embryonic-drosophila-gonad

Research

JoVE Journal

Developmental Biology

3.1K Views.  University of Pennsylvania. 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.

Video: Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

Large-scale Zebrafish Embryonic Heart Dissection for Transcriptional Analysis

The zebrafish embryonic heart is composed of only a few hundred cells, representing only a small fraction of the entire embryo. Therefore, to prevent the cardiac transcriptome from being masked by the global embryonic transcriptome, it is necessary to collect sufficient numbers of hearts for further analyses. Furthermore, as zebrafish cardiac development proceeds rapidly, heart collection and RNA extraction methods need to be quick in order to ensure homogeneity of the samples. Here, we present a rapid manual dissection protocol for collecting functional/beating hearts from zebrafish embryos. This is an essential prerequisite for subsequent cardiac-specific RNA extraction to determine cardiac-specific gene expression levels by transcriptome analyses, such as quantitative real-time polymerase chain reaction (RT-qPCR). The method is based on differential adhesive properties of the zebrafish embryonic heart compared with other tissues; this allows for the rapid physical separation of cardiac from extracardiac tissue by a combination of fluidic shear force disruption, stepwise filtration and manual collection of transgenic fluorescently labeled hearts.

To analyse cardiac gene expression profiles during zebrafish heart development, total RNA has to be extracted from isolated hearts. Here, we present a protocol for collecting functional/beating hearts by rapid manual dissection from zebrafish embryos to obtain cardiac-specific mRNA.

The zebrafish embryonic heart is composed of only a few hundred cells, representing only a small fraction of the entire embryo. Therefore, to prevent the ...

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 ...

Dissection and Culture of Mouse Embryonic Kidney

The goal of this protocol is to describe a method for the dissection, isolation, and culture of mouse metanephric rudiments. 
During mammalian kidney development, the two progenitor tissues, the ureteric bud and the metanephric mesenchyme, communicate and reciprocally induce cellular mechanisms to eventually form the collecting system and the nephrons of the kidney. As mammalian embryos grow intrauterine and therefore are inaccessible to the observer, an organ culture has been developed. With this method, it is possible to study epithelial-mesenchymal interactions and cellular behavior during kidney organogenesis. Furthermore, the origin of congenital kidney and urogenital tract malformations can be investigated. After careful dissection, the metanephric rudiments are transferred onto a filter that floats on culture medium and can be kept in a cell culture incubator for several days. However, one must be aware that the conditions are artificial and could influence the metabolism in the tissue. Also, the penetration of test substances could be limited due to the extracellular matrix and basal membrane present in the explant.
One main advantage of organ culture is that the experimenter can gain direct access to the organ. This technology is cheap, simple, and allows a large number of modifications, such as the addition of biologically active substances, the study of genetic variants, and the application of advanced imaging techniques.

This protocol describes a method for isolating and culturing metanephric rudiments from mouse embryos.

The goal of this protocol is to describe a method for the dissection, isolation, and culture of mouse metanephric rudiments. 
During mammalian kidney ...

Long-term Live Imaging of Drosophila Eye Disc

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have been used to study many biological processes, such as cell proliferation, differentiation, growth, migration, apoptosis, competition, cell-cell signaling, and compartmental boundary formation. However, methods for the long-term ex vivo culture and live imaging of the imaginal discs have not been satisfactory, despite many efforts. Recently, we developed a method for the long-term ex vivo culture and live imaging of imaginal discs for up to 18 h. In addition to using a high insulin concentration in the culture medium, a low-melting agarose was also used to embed the disc to prevent it from drifting during the imaging period. This report uses the eye-antennal discs as an example. Photoreceptor R3/4-specific m&#948;0.5-Ga4 expression was followed to demonstrate that photoreceptor differentiation and ommatidial rotation can be observed during a 10 h live imaging period. This is a detailed protocol describing this simple method.

Long-term Live Imaging of Drosophila Eye Disc

This protocol describes a detailed method for the long-term ex vivo culture and live imaging of a Drosophila imaginal disc. It demonstrates photoreceptor differentiation and ommatidial rotation within the 10 h period of live imaging of the eye disc. The protocol is simple and does not require expensive setup.

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have ...

Bioengineering of Humanized Bone Marrow Microenvironments in Mouse and Their Visualization by Live Imaging

Human hematopoietic stem cells (HSCs) reside in the bone marrow (BM) niche, an intricate, multifactorial network of components producing cytokines, growth factors, and extracellular matrix. The ability of HSCs to remain quiescent, self-renew or differentiate, and acquire mutations and become malignant depends upon the complex interactions they establish with different stromal components. To observe the crosstalk between human HSCs and the human BM niche in physiological and pathological conditions, we designed a protocol to ectopically model and image a humanized BM niche in immunodeficient mice. We show that the use of different cellular components allows for the formation of humanized structures and the opportunity to sustain long-term human hematopoietic engraftment. Using two-photon microscopy, we can live-image these structures in situ at the single-cell resolution, providing a powerful new tool for the functional characterization of the human BM microenvironment and its role in regulating normal and malignant hematopoiesis.

A method to create and live-image different humanized bone-marrow niches in mice is presented. Based on the supportive niche created by human mesenchymal cells, the addition of human endothelial cells induces the formation of human vessels, while the addition of rhBMP-2 induces the formation of human-mouse chimeric mature bone tissue.

Human hematopoietic stem cells (HSCs) reside in the bone marrow (BM) niche, an intricate, multifactorial network of components producing cytokines, growth ...

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the proteins involved in these pathways are evolutionarily conserved between mammals and other eukaryotes, such as the fruit fly Drosophila melanogaster, suggesting that similar organizing principles exist during the development of these organisms. Importantly, Drosophila has been used extensively to identify cellular and molecular mechanisms regulating processes that are required in mammals including neurogenesis, differentiation, axonal guidance, and synaptogenesis. Flies have also been used successfully to model a variety of human neurodevelopmental diseases. Here we describe a protocol for the step-by-step microdissection, fixation, and immunofluorescent localization of proteins within the adult Drosophila brain. This protocol focuses on two example neuronal populations, mushroom body neurons and retinal photoreceptors, and includes optional steps to trace individual mushroom body neurons using Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. Example data from both wild-type and mutant brains are shown along with a brief description of a scoring criteria for axonal guidance defects. While this protocol highlights two well-established antibodies for investigating the morphology of mushroom body and photoreceptor neurons, other Drosophila brain regions and the localization of proteins within other brain regions can also be investigated using this protocol.

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

This protocol describes the dissection and immunostaining of adult Drosophila melanogaster brain tissues. Specifically, this protocol highlights the use of Drosophila mushroom body and photoreceptor neurons as example neuronal subsets that can be accurately used to uncover general principles underlying many aspects of neuronal development.

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the ...

Dissection and Staining of Drosophila Pupal Ovaries

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a pupal case. The challenge of dissecting pupal ovaries also lies in their physical location within the pupa: the ovaries are surrounded by fat body cells inside the pupal abdomen, and these fat cells must be removed to allow for proper antibody staining. To overcome these challenges, this protocol utilizes customized Pasteur pipets to extract fat body cells from the pupal abdomen. Moreover, a chambered coverglass is used in place of a microcentrifuge tube during the staining process to improve visibility of the pupae. However, despite these and other advantages of the tools used in this protocol, successful execution of these techniques may still involve several days of practice due to the small size of pupal ovaries. The techniques outlined in this protocol could be applied to time course experiments in which ovaries are analyzed at various stages of pupal development.

Dissection and Staining of Drosophila Pupal Ovaries

The Drosophila ovary is an excellent model system for studying stem cell niche development. Though methods for dissecting larval and adult ovaries have been published, pupal ovary dissections require different techniques that have not been published in detail. Here we outline a protocol for dissecting, staining, and mounting pupal ovaries.

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a ...

Dissection of the Drosophila Pupal Retina for Immunohistochemistry, Western Analysis, and RNA Isolation

The Drosophila pupal retina provides an excellent model system for the study of morphogenetic processes during development. In this paper, we present a reliable protocol for the dissection of the delicate Drosophila pupal retina. Our surgical approach utilizes readily-available microdissection tools to open pupae and precisely extract eye-brain complexes. These can be fixed, subjected to immunohistochemistry, and retinas then mounted onto microscope slides and imaged if the goal is to detect cellular or subcellular structures. Alternatively, unfixed retinas can be isolated from brain tissue, lysed in appropriate buffers and utilized for protein gel electrophoresis or mRNA extraction (to assess protein or gene expression, respectively). Significant practice and patience may be required to master the microdissection protocol described, but once mastered, the protocol enables relatively quick isolation of mainly undamaged retinas.

Dissection of the Drosophila Pupal Retina for Immunohistochemistry, Western Analysis, and RNA Isolation

This paper presents a surgical method for dissecting Drosophila pupal retinas along with protocols for the processing of tissue for immunohistochemistry, western analysis, and RNA-extraction.&#160;

The Drosophila pupal retina provides an excellent model system for the study of morphogenetic processes during development. In this paper, we present a ...

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits are required to control this behavior. Failure to produce the rhythmic pattern of contractions can be indicative of neurological abnormalities. We previously found that defects in protein O-mannosylation, a posttranslational protein modification, affect the axon morphology of sensory neurons and result in abnormal coordinated muscle contractions in embryos. Here, we present a relatively simple method for recording and analyzing the pattern of peristaltic muscle contractions by live imaging of late stage embryos up to the point of hatching, which we used to characterize the muscle contraction phenotype of protein O-mannosyltransferase mutants. Data obtained from these recordings can be used to analyze muscle contraction waves, including frequency, direction of propagation and relative amplitude of muscle contractions at different body segments. We have also examined body posture and taken advantage of a fluorescent marker expressed specifically in muscles to accurately determine the position of the embryo midline. A similar approach can also be utilized to study various other behaviors during development, such as embryo rolling and hatching.

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Here, we present a method to record embryonic muscle contractions in Drosophila embryos in a non-invasive and detail-oriented manner.

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits ...