This work was supported by a Wellcome Trust Senior Research Fellowship to I. Davis.

1. Drosophila preparation prior for dissection (According to E. Gavis, Princeton University)
<ol>
	<li>Discard the adults from a sparsely seeded bottle and transfer the bottle to 25 &deg;C in advance of your experiment. Once new adults have begun to emerge, wait two days.</li>
	<li>Take a fly vial containing approximately 10ml of solid food and add 0.5-0.7 grams of dry yeast (see table) on a 45 degree angle. Do not cover the whole surface of the fly food with yeast.</li>
	<li>Add a few drops of water to the yeast, sufficient to hydrate (Figure 1A). Place the vial at 45 degrees and allow about 3 minutes for the yeast to soak up the water (Figure 1B).</li>
</ol>
Note: Alternatively, premix a yeast paste in a separate container and add the paste to the vial with a spatula.
<ol start="4">
	<li>Anesthetize the flies in the vial by injecting CO2 gas. When they stop moving, tip the unconscious flies onto the CO2 fly pad for sorting (Figure 1C).</li>
	<li>Isolate 10-15 females and 5 males of the desired genotype under a dissecting scope with a fine tipped paint brush (Figure 1D). Males are obvious by the presence of claspers, a dark protruding structure, at their posterior.</li>
	<li>Add selected flies to the yeasted vial (Figure 1E). Place at 25 &deg;C for 24-60 hours. Incubate for shorter time periods for a larger percent of young to mid stage egg chambers and longer time periods for late stage egg chambers.</li>
</ol>
2. Drosophila ovary dissection
<ol>
	<li>Place a small drop of halocarbon oil on a No. 1.5, 22 x 50 mm cover slip (Figure 2A). Set the prepared cover slip to the side where it will not be in the way.</li>
	<li>Anesthetize the flies in the yeasted vile by injecting CO2 gas and tip onto a CO2 pad.</li>
	<li>Use sharp fine-nosed forceps to grasp an individual fattened female at the anterior of the abdomen (not between the abdomen and thorax) (Figure 2B-C).</li>
	<li>Hold the female with the forceps and view under a dissecting microscope, roughly 2-3 cm above the prepared coverslip.</li>
	<li>Still holding at the anterior, use a second set of forceps to remove the tissue at the extreme posterior of the female (Figure 2D). Note: Gut tissue will usually pull out with the posterior end. Wipe the removed tissue from the forceps.</li>
	<li>While still holding the female at the anterior, use the wiped forceps to gently squeeze or massage the abdomen, working from the anterior towards the posterior. Massage the abdomen in the same fashion as removing toothpaste from the end of a tube.</li>
	<li>Two large opaque structures will be pushed from the abdomen and stick on the end of the forceps (Figure 2E-F). These are the paired ovaries.</li>
	<li>Touch the ovaries onto the halocarbon oil on the slide below.</li>
	<li>Repeat the dissection 1-2 times to give a total of 4-6 ovaries in the oil on the coverslip (Figure 2G).</li>
</ol>
3. Isolating ovarioles
Note: Each ovary contains about 16 ovarioles made up of 6 to 10 egg chambers of different stages arranged like beads on a string10.
Note: Prior to isolating the ovarioles, adjust the light source on the dissecting microscope so that the illumination strikes at a shallow angle to the sample. This gives contrast to the sample and allows visualization of the young stage egg chambers.
<ol>
	<li>In the oil, separate the two ovaries by removing the connecting tissue at the posterior end (old stages) with closed forceps (Figure 3A).</li>
	<li>Orient an individual ovary with the oldest stages to the left and youngest stage to the right. The young stage egg chambers in the ovary are transparent and form the more pointed end of the ovary.</li>
	<li>With a pair of forceps in the dominant hand, gently grasp the posterior of an ovary, preferably by the remnants of the connective tissue. While holding the ovary in place with the forceps, use a dissecting probe (or tungsten needle) to tease out individual ovarioles (Figure 3B-D).</li>
</ol>
Note: Individual ovarioles will break between the young stages (germarium to stage 10) as the older stages are too large to be separated from the full ovary.
Note: Puncturing a late stage oocyte with the dissecting probe will result in the cytoplasm leaking into the oil and make extraction of individual ovarioles very challenging. If a late stage oocyte is punctured, move to a fresh ovary.
<ol start="4">
	<li>Once the ovariole is free of the ovary, use the dissecting probe to drag it to the center of the oil drop and orient in the desired orientation for the experiment. Note: Ovarioles in oil will settle and adhere to the glass.</li>
	<li>Repeat this process (3.3-3.4) until there are 15-25 individual ovarioles. Note: This may require the dissection of multiple ovaries. The whole process should take less than 15 minutes.</li>
</ol>
Note: It is advisable to dissect one ovary each from several flies rather than dissecting both ovaries from fewer flies to increase the n number of flies for the experiment.
Note: Rough handling of the ovariole will result in unhealthy oocytes and can lead to artifacts in the experiment.
Note: Oocytes will begin to display phenotypic changes due to stress 40 minutes after being dissected from the ovary (compare Figure 7E to F).
<ol start="6">
	<li>Once ovarioles are in an adequate orientation (Figure 3E), use the dissecting probe and closed forceps to move excess ovarian tissue out of the oil. This material can be wiped from the forceps onto a paper towel.</li>
</ol>
4. Isolating individual late stage egg chambers (According to E. Gavis, Princeton University)
<ol>
	<li>With a pair of forceps in the dominant hand, grasp the posterior of an isolated ovary, preferably by the remnants of the connective tissue. While holding the ovary in place with the forceps, introduce the dissecting probe into the posterior end of the ovary near the forceps (Figure 4A). Avoid puncturing any egg chambers by inserting the probe between late stage egg chambers.</li>
	<li>Pull the dissecting needle through the ovary until it exits at the early stage egg chambers (Figure 4B). When done correctly, the probe will remove the connective tissue holding the ovarioles and late stage egg chambers in place.</li>
	<li>Repeat step 4.2 until the ovarioles are spread out on the coverslip. Note: When egg chambers are no longer stacked on top of each other, this step is complete.</li>
	<li>Using the dissecting probe, separate the late oocytes to allow for easy imaging (Figure 4C).</li>
</ol>
Note: Puncturing an oocyte while dissecting late stage egg chambers is not as condemning as with dissecting individual ovarioles for younger stages.
Note: For stage 14 egg chambers, use the forceps to grasp the dorsal appendages for orientation.
5. Injection preparation
Note: For the injection of mid stage oocytes, orient the ovarioles perpendicularly to the long axis of the cover slip.
<ol>
	<li>Turn on the imaging microscope [in our case a DeltaVision Core from Applied Precision] and load the appropriate imaging settings (Figure 5A-B).</li>
	<li>Turn on the injection apparatus (Figure 5C).</li>
	<li>Load the injection needle with a loading tip (Figure 5D-E). Solutions to be injected should be centrifuged briefly at high speed to avoid deposits that might block the injection needle.</li>
	<li>Attach the loaded injection needle to the injector. Take care to avoid breaking the needle (Figure 5F).</li>
	<li>To assist with unclogging needles, place a small piece of glass 2x10 mm from a cut or broken coverslip at one side of the oil drop containing the ovarioles. Make sure this glass fragment is covered in oil. This glass will serve to ram the needle tip against for breaking or unclogging the needle.</li>
</ol>
6. Injection of fluorescent RNA
Before you begin: Set up the injection apparatus and micro-manipulator controls such that the injection needle is positioned over the center of the field of view.
<ol>
	<li>Mount the sample to be injected on the microscope stage.</li>
	<li>Select the 20x objective. Higher or lower magnification objectives can also be used.</li>
</ol>
Note: If using an automated stage with point visiting capability it is advisable to mark all the stage 8-9 oocytes for injection before beginning. This allows for easy visiting and injection of selected oocytes.
<ol start="3">
	<li>Lower the injection needle until it touches the oil. Center the needle tip over the objective lens.</li>
	<li>Turn off the room lights and turn on the bright-field light for the microscope. Select the first point (oocyte) on the marked points list. While looking through the oculars, focus on the oocyte (Figure 6A).</li>
	<li>Lower the needle manually until it is in the same plane of focus as the oocyte and visible next to the oocyte.</li>
	<li>Using the manipulator controls, move the point of the injection needle until it is inside the oocyte. A fast stabbing movement is often more successful than a slow advance. Once inside, eject a small volume from the needle and visually assess the effects, repeat as required. If the injection is successful, the cytoplasm inside the oocyte will be displaced, if this is not apparent, repeat. It is not necessary to image in fluorescence to determine if material has been injected. After injection, remove the needle from inside the oocyte (Figure 6B-G).</li>
	<li>The precise amount of fluid injected is difficult to quantitate. To increase the injected volume, adjust the pressure or time of the injection pulse, break the needle tip to increase the opening slightly or use repeated injection pulses.</li>
</ol>
Note: The whole injection process should take less than 10 seconds when executed properly. Extensive stabbing or lingering with the needle inside to oocyte will result in severe damage to the egg chamber.
<ol start="8">
	<li>Before selecting the next marked oocyte, raise the injection needle high enough in the oil so as not to contact other oocytes in the travel path.</li>
	<li>At the new oocyte, repeat step 6.6. Carry on in this way until all the required oocytes are injected.</li>
</ol>
Note: If a mistake is made while injecting, move to the next marked oocyte. Do not waste time on damaged oocytes.
Note: The needle may become clogged during an injection session. In this case, move the needle adjacent to the broken piece of glass in the oil on the coverslip. With the glass and needle in the same focal plane, gently ram the needle into the glass (Figure 6H). Test that the needle is unclogged by pushing the injection button and seeing if any fluid exits the tip of the needle.
<ol start="10">
	<li>When all the oocytes have been injected, manually move the needle up, out of the oil and orient it out of the beam path for the microscope. Make sure that it is in a safe orientation, to avoid eye damage.</li>
</ol>
Note: If an extended period of time has elapsed between egg chamber isolation and oocyte injection, oocytes will be difficult to inject and exhibit phenotypic changes associated with stress. Similar issues can arise with rough handling of the oocyte or injection of excess volumes (compare Figures 6I, J, K).
7. Live imaging experimental design
<ol>
	<li>Select the first injected oocyte from the list. Test that the injection was inside the oocyte by imaging one frame in fluorescence (as in Figure 6G).</li>
	<li>Configure the experimental parameters, including: excitation wavelength, emission path, excitation time, Z-stack and time between collections. Note: For more information on this, see Parton R., et al. (2010) 2.</li>
	<li>Gather the data set.</li>
</ol>
8. Representative Results
In vitro synthesized Alexa-546 grk RNA injected into an Me31B::GFP egg chamber (Figure 7A). The RNA localizes to the dorsal anterior of the oocyte and forms a cap around the nucleus (Figure 7B). For more examples of RNA localization see MacDougall N., et al. (2003) 11.
Mid and late stage oocytes expressing fluorescent labeled protein (Tau-GFP, Figure 7C) or RNA tagging systems (grk*mCherry, Figure 7D) can be imaged without injection.
<img alt="Figure 1" src="/files/ftp_upload/3679/3679fig1.jpg" /> 
Figure 1.
 
<img alt="Figure 2" src="/files/ftp_upload/3679/3679fig2.jpg" /> 
Figure 2.
 
<img alt="Figure 3" src="/files/ftp_upload/3679/3679fig3.jpg" /> 
Figure 3.
 
<img alt="Figure 4" src="/files/ftp_upload/3679/3679fig4.jpg" /> 
Figure 4.
 
<img alt="Figure 5" src="/files/ftp_upload/3679/3679fig5.jpg" /> 
Figure 5.
 
<img alt="Figure 6" src="/files/ftp_upload/3679/3679fig6.jpg" /> 
Figure 6.
 
<img alt="Figure 7" src="/files/ftp_upload/3679/3679fig7.jpg" /> 
Figure 7.

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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pushing+the+Limits+of+Live+Cell+Imaging+in+Drosophila.">Pushing the Limits of Live Cell Imaging in Drosophila.</a> Live Cell Imaging: A Laboratory. , 387-418 (2010).</li><li>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=Moving+messages:+the+intracellular+localization+of+mRNAs.">Moving messages: the intracellular localization of mRNAs.</a> Nat. Rev. Mol. Cell Biol. 6 (5), 363-375 (2005).</li><li>Jansen, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=mRNA+Localization:+Message+on+the+Move.">mRNA Localization: Message on the Move.</a> Nat. Rev. Mol. Cell Biol. 2, 247-256 (2001).</li><li>Van De Bor, V., Hartswood, E., Jones, C., Finnegan, D., Davis, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=gurken+and+the+I+factor+retrotransposon+RNAs+share+common+localization+signals+and+machinery.">gurken and the I factor retrotransposon RNAs share common localization signals and machinery.</a> Dev. Cell. 9 (1), 51-62 (2005).</li><li>Delanoue, R., Herpers, B., Soetaert, J., Davis, I., Rabouille, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+Squid&#47;hnRNP+helps+Dynein+switch+from+a+gurken+mRNA+transport+motor+to+an+ultrastructural+static+anchor+in+sponge+bodies.">Drosophila Squid&#47;hnRNP helps Dynein switch from a gurken mRNA transport motor to an ultrastructural static anchor in sponge bodies.</a> Dev. Cell. 13 (4), 523-538 (2007).</li><li>Jaramillo, A. M., Weil, T. T., Goodhouse, J., Gavis, E. R., Schupbach, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+dynamics+of+fluorescently+labeled+endogenous+gurken+mRNA+in+Drosophila.">The dynamics of fluorescently labeled endogenous gurken mRNA in Drosophila.</a> J. Cell. Sci. 121 (Pt. 6), 887-894 (2008).</li><li>Forrest, K. M., Gavis, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+imaging+of+endogenous+mRNA+reveals+a+diffusion+and+entrapment+mechanism+for+nanos+mRNA+localization+in+Drosophila.">Live imaging of endogenous mRNA reveals a diffusion and entrapment mechanism for nanos mRNA localization in Drosophila.</a> Curr. Biol. 13, 1159-1168 (2003).</li><li>Bertrand, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Localization+of+ASH1+mRNA+particles+in+living+yeast.">Localization of ASH1 mRNA particles in living yeast.</a> Mol. Cell. 2 (4), 437-445 (1998).</li><li>Spradling, A. 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Cell. 4 (3), 307-319 (2003).</li><li>Tekotte, H., Tollervey, D., Davis, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+the+migrating+border+cell+cluster+in+living+Drosophila+egg+chambers.">Imaging the migrating border cell cluster in living Drosophila egg chambers.</a> Dev. Dyn. 236 (10), 2818-2824 (2007).</li><li>Weil, T. T., Parton, R., Davis, I., Gavis, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Changes+in+bicoid+mRNA+anchoring+highlight+conserved+mechanisms+during+the+oocyte-to-embryo+transition.">Changes in bicoid mRNA anchoring highlight conserved mechanisms during the oocyte-to-embryo transition.</a> Curr. Biol. 18, 1055-1061 (2008).</li><li>Prasad, M., Jang, A. 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> Nat. Protoc. 2 (10), 2467-2473 (2007).</li><li>Morris, L. X., Spradling, A. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+live+imaging+provides+new+insight+into+stem+cell+regulation+and+germline-soma+coordination+in+the+Drosophila+ovary.">Long-term live imaging provides new insight into stem cell regulation and germline-soma coordination in the Drosophila ovary.</a> Development. 138 (11), 2207-2215 (2011).</li></ol>

We have nothing to disclose.

Live cell imaging is a powerful assay for examining cellular processes in real time. In addition to simple bright field observation, the addition of fluorescent labels to proteins and RNAs of interest has lead to many breakthroughs. Here we have outlined a protocol for imaging individual living oocytes that can be utilized in combination with genetic and biochemical assays. 
 In this protocol, we also explain how to experimentally manipulate living oocytes by injection. There are many possibilities for material to inject, including in vitro synthesized fluorescent labeled RNA to assay the ability of a secondary structure to direct RNA localization (Ball and Davis, unpublished) and antibodies that inhibit the function of proteins 6. Future work will likely see the introduction of other labeling components and cellular machinery to living oocytes enabling molecular mechanisms to be tested. 
 Preserving the viability and health of the tissue is essential when working with live cells. In this protocol, we point out a number of steps that can lead to stress of the egg chamber. For example, despite oil being superior for imaging, extended culture in the oil can lead to stress on the egg chamber. This can be easily monitored under bright field exposure by examining the nuclear morphology and position, oocyte membrane that will distort and bleb under stress (compare Figure 7E (unstressed) to Figure7F (stressed)). Stage 9 egg chambers show RNA localization and border cell migration 12 over multiple hours. However, a stage 7/8 egg chamber will begin to exhibit ill effects in halo carbon oil after approximately 40 minutes of the ovary being removed from the female. Late stage egg chambers, stage 11 to 14, can develop normally in either oil or aqueous media because of the secretion of the egg shell from the follicle cells at these stages 13. It has been reported that the addition of insulin to aqueous insect medium can maintain stage 9 oocytes for up to 6 hours 14 and germaria up to 14 hours 15. In all cases, aggressive maneuvering of the egg chamber should be avoided, as it stresses the oocytes and reduces its viability. Imaging fewer egg chambers and taking care to treat each gently is the best way to ensure optimal viability.

Tools used in dissection should be clean but do not need to be autoclaved. Wipe tools with ETOH and allow them to dry before beginning.
 <table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments </td>
 </tr>
 <tr>
 <td>Fine tipped paint brush 
 (Artists Sable, round, size 00 or 0)</td>
 <td>Available from most art supplies outlets</td>
 <td>&nbsp;</td>
 <td>A good quality brush is important</td>
 </tr>
 <tr>
 <td>Halocarbon Products Corporation, Series 95</td>
 <td>Halocarbon Products Corporation, 887 Kinderkamack Road, River Edge NJ 07661</td>
 <td>&nbsp;</td>
 <td>European distributor: 
 Solvadis (GMBH) 
 Be sure to oxygenate by bubbling air through the oil.</td>
 </tr>
 <tr>
 <td>Cover slip -No. 1.5, 22 x 50mm</td>
 <td>International- No1-VWR Cat=631-0137-22-50mm</td>
 <td>Menzel-Glaser-22-40mm-MNJ 400-070T</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dumont No.5 - Dumostar</td>
 <td>Fine Science Tools</td>
 <td>11253-20</td>
 <td>"Biologie" tip is also good but more fragile</td>
 </tr>
 <tr>
 <td>Dissecting probe </td>
 <td>Fine Science Tools</td>
 <td>10140-01</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dissecting microscope</td>
 <td>Olympus</td>
 <td>SZ61</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>CO2 pad</td>
 <td>Available through 
 <a href="http://www.flystuff.com">http://www.flystuff.com</a>/ 
 (A division of&nbsp; Genesee Scientific)</td>
 <td>59-114 
 (789060 Dutscher)</td>
 <td>European distributor: 
 Dutscher <a href="http://www.dutscherscientific.com">www.dutscherscientific.com</a>/ 
 It is also relatively easy to custom build your own fly pads</td>
 </tr>
 <tr>
 <td>KimCare</td>
 <td>Kimbery Clark-KimCare-Medical wipes</td>
 <td>KLEW3020</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Microscope- DeltaVision</td>
 <td>Applied Precision</td>
 <td>DC-CORE</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Micromanipulator</td>
 <td>Burleigh Micromanipulators 
 from Lumen Dynamics Group inc. 2260 Argentia Road, Mississauga, Ontario, L5N 6H7 Canada</td>
 <td>Burleigh PCS-5000 series, TS-5000-300</td>
 <td><a href="http://www.ldgi-burleigh.com">http://www.ldgi-burleigh.com</a>/ (Formerly Exfo)</td>
 </tr>
 <tr>
 <td>Injection apparatus</td>
 <td>Tritech Research, Inc. 
 2961 Veteran Ave 
 Los Angeles, CA 90064</td>
 <td>MINJ-1</td>
 <td>Modified with a holder to take Eppendorf Femptotips</td>
 </tr>
 <tr>
 <td>Injection needle</td>
 <td>Eppendorf Sterile Femtotips I and II</td>
 <td>930000043</td>
 <td>Different tips are better for different applications</td>
 </tr>
 <tr>
 <td>Loading tips 20&mu;l</td>
 <td>Eppendorf</td>
 <td>5242956.003</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dry active yeast</td>
 <td>Fleischmann's Yeast</td>
 <td>#2192</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
 </table>

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.

Watch this Scientific Journal Video about Preparing Individual Drosophila Egg Chambers for Live Imaging at JoVE.com

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.

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, ...

preparing-individual-drosophila-egg-chambers-for-live-imaging

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19.1K Views.  University of Oxford. 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.

Video: Preparing Individual Drosophila Egg Chambers for Live Imaging

Live Imaging of Glial Cell Migration in the Drosophila Eye Imaginal Disc

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons. While recent progress has been made to describe the live migration of glial cells in the developing pupal wing (1), studies of Drosophila glial cell migration have typically involved the examination of fixed tissue. Live microscopic analysis of motile cells offers the ability to examine cellular behavior throughout the migratory process, including determining the rate of and changes in direction of growth. Paired with use of genetic tools, live imaging can be used to determine more precise roles for specific genes in the process of development. Previous work by Silies et al. (2) has described the migration of glia originating from the optic stalk, a structure that connects the developing eye and brain, into the eye imaginal disc in fixed tissue. Here we outline a protocol for examining the live migration of glial cells into the Drosophila eye imaginal disc. We take advantage of a Drosophila line that expresses GFP in developing glia to follow glial cell progression in wild type and in mutant animals.

Here we describe a protocol to examine the migration of glial cells into the developing Drosophila eye using live microscopic analysis paired with GFP tagged glial cells.

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons.  While ...

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

This technique exposes the Drosophila embryonic neuromusculature for immunohistochemistry or electrophysiological recording. It is useful for studying early events in neuromuscular development or performing electrophysiology in mutants that cannot hatch.

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

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 ...

Live Imaging of Cell Motility and Actin Cytoskeleton of Individual Neurons and Neural Crest Cells in Zebrafish Embryos

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily accessible at all stages of development. Moreover, their optical clarity allows high resolution imaging of cell and molecular dynamics in the natural environment of the intact embryo. We are using a live imaging approach to analyze cell behaviors during neural crest cell migration and the outgrowth and guidance of neuronal axons.
Live imaging is particularly useful for understanding mechanisms that regulate cell motility processes. To visualize details of cell motility, such as protrusive activity and molecular dynamics, it is advantageous to label individual cells. In zebrafish, plasmid DNA injection yields a transient mosaic expression pattern and offers distinct benefits over other cell labeling methods. For example, transgenic lines often label entire cell populations and thus may obscure visualization of the fine protrusions (or changes in molecular distribution) in a single cell. In addition, injection of DNA at the one-cell stage is less invasive and more precise than dye injections at later stages.
Here we describe a method for labeling individual developing neurons or neural crest cells and imaging their behavior in vivo. We inject plasmid DNA into 1-cell stage embryos, which results in mosaic transgene expression. The vectors contain cell-specific promoters that drive expression of a gene of interest in a subset of sensory neurons or neural crest cells. We provide examples of cells labeled with membrane targeted GFP or with a biosensor probe that allows visualization of F-actin in living cells1.
Erica Andersen, Namrata Asuri, and Matthew Clay contributed equally to this work.

This protocol describes imaging of individual neurons or neural crest cells in living zebrafish embryos. This method is used to examine cellular behaviors and actin localization using fluorescence confocal time-lapse microscopy.

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily ...

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three preparations of the brain and visual system that cover the range from the developing eye disc-brain complex in the developing pupae to individual eye and brain dissection from adult flies. All protocols are optimized for the live culture of the preparations. However, we also present the conditions for fixed tissue immunohistochemistry where applicable. Finally, we show live imaging conditions for these preparations using conventional and resonant 4D confocal live imaging in a perfusion chamber. Together, these protocols provide a basis for live imaging on different time scales ranging from functional intracellular assays on the scale of minutes to developmental or degenerative processes on the scale of many hours.

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

This protocol describes three Drosophila preparations: 1) adult brain dissection, 2) adult retina dissection and 3) developing eye disc- brain complexes dissection. Emphasis is laid on special preparation techniques and conditions for live imaging, although all preparations can be used for fixed tissue immunohistochemistry.

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three ...

Electron Spin Resonance Micro-imaging of Live Species for Oxygen Mapping

This protocol describes an electron spin resonance (ESR) micro-imaging method for three-dimensional mapping of oxygen levels in the immediate environment of live cells with micron-scale resolution1. Oxygen is one of the most important molecules in the cycle of life. It serves as the terminal electron acceptor of oxidative phosphorylation in the mitochondria and is used in the production of reactive oxygen species. Measurements of oxygen are important for the study of mitochondrial and metabolic functions, signaling pathways, effects of various stimuli, membrane permeability, and disease differentiation. Oxygen consumption is therefore an informative marker of cellular metabolism, which is broadly applicable to various biological systems from mitochondria to cells to whole organisms. Due to its importance, many methods have been developed for the measurements of oxygen in live systems. Current attempts to provide high-resolution oxygen imaging are based mainly on optical fluorescence and phosphorescence methods that fail to provide satisfactory results as they employ probes with high photo-toxicity and low oxygen sensitivity. ESR, which measures the signal from exogenous paramagnetic probes in the sample, is known to provide very accurate measurements of oxygen concentration. In a typical case, ESR measurements map the probe's lineshape broadening and/or relaxation-time shortening that are linked directly to the local oxygen concentration. (Oxygen is paramagnetic; therefore, when colliding with the exogenous paramagnetic probe, it shortness its relaxation times.) Traditionally, these types of experiments are carried out with low resolution, millimeter-scale ESR for small animals imaging. Here we show how ESR imaging can also be carried out in the micron-scale for the examination of small live samples. ESR micro-imaging is a relatively new methodology that enables the acquisition of spatially-resolved ESR signals with a resolution approaching 1 micron at room temperature2. The main aim of this protocol-paper is to show how this new method, along with newly developed oxygen-sensitive probes, can be applied to the mapping of oxygen levels in small live samples. A spatial resolution of ~30 x 30 x 100 &mu;m is demonstrated, with near-micromolar oxygen concentration sensitivity and sub-femtomole absolute oxygen sensitivity per voxel. The use of ESR micro-imaging for oxygen mapping near cells complements the currently available techniques based on micro-electrodes or fluorescence/phosphorescence. Furthermore, with the proper paramagnetic probe, it will also be readily applicable for intracellular oxygen micro-imaging, a capability which other methods find very difficult to achieve.

This protocol describes a method for micron-scale three-dimensional imaging of oxygen concentration in the immediate environment of live cells by electron spin resonance microscopy.

This protocol describes an electron spin resonance (ESR) micro-imaging method for three-dimensional mapping of oxygen levels in the immediate environment ...

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 ...

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

The Drosophila oocyte has been established as a versatile system for investigating fundamental questions such as cytoskeletal function, cell organization, and organelle structure and function. The availability of various GFP-tagged proteins means that many cellular processes can be monitored in living cells over the course of minutes or hours, and using this technique, processes such as RNP transport, epithelial morphogenesis, and tissue remodeling have been described in great detail in Drosophila oocytes1,2.
The ability to perform video imaging combined with a rich repertoire of mutants allows an enormous variety of genes and processes to be examined in incredible detail. One such example is the process of ooplasmic streaming, which initiates at mid-oogenesis3,4. This vigorous movement of cytoplasmic vesicles is microtubule and kinesin-dependent5 and provides a useful system for investigating cytoskeleton function at these stages.
Here I present a protocol for time lapse imaging of living oocytes using virtually any confocal microscopy setup.

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

A protocol for live imaging of GFP-tagged proteins or autofluorescent structures in individual Drosophila oocytes is described.