We thank Christian Lehner for providing the CENP-C antibody and Eric Joyce for recommendations on FISH. Work in the McKim lab was funded by a grant from NIH (GM101955).

Note: Procedures are performed at room temperature unless otherwise noted. Temperature-controlled incubators are used to maintain temperatures for fly rearing and crosses unless otherwise noted.
1. Preparations
<ol>
	<li>Prepare Flies.
	<ol>
		<li>Prometaphase-enriched oocyte collections.
		<ol>
			<li>Clear adult flies from healthy, young stock or cross cultures. Age bottles for two days at 25 &#176;C. 
			&#8203;NOTE: Generally two healthy bottles will suffice, although more may be needed for some cross cultures.</li>
			<li>After two days, collect ~100 to 300 females (who are 0 to 2 days old at this point) from the bottles. The females do not need to be virgins. Add a dab of yeast paste to the side of a vial and place 30 females and 10 to 15 males each into the yeasted vials. Age vials for two days at 25 &#176;C.</li>
		</ol>
		</li>
		<li>Metaphase-enriched Oocyte Collections.
		<ol>
			<li>Collect ~100 to 300 females from stock or cross cultures. Add a dab of yeast paste to the side of a vial and place 30 females each (with no males added) into the yeasted vials. Age vials for three to five days at 25 &#176;C.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Prepare Solutions. 
	&#8203;Note: Solutions may be stored indefinitely at room temperature, unless otherwise noted.
	<ol>
		<li>Prepare Modified Robb&#39;s Buffer (5x): 500 mM HEPES, 500 mM sucrose, 275 mM sodium acetate, 200 mM potassium acetate, 50 mM glucose, 6 mM magnesium chloride, and 5 mM calcium chloride. Use 10 N 11:8 sodium hydroxide:potassium hydroxide to bring pH to 7.4. Sterilize by filtration; do not autoclave. Store at -20 &#176;C. Thaw as needed to prepare ~200 ml of 1x Robb&#39;s per oocyte prep.</li>
		<li>Fixation Solutions.
		<ol>
			<li>Option #1: Prepare formaldehyde/heptane fixation. Prepare Fixation Buffer: 1x phosphate-buffered saline (PBS) plus 150 mM sucrose. To use, make fresh with 687.5 &#181;l Fixation Buffer and 312.5 &#181;l 16% formaldehyde per oocyte prep. 
			CAUTION: Wear gloves while using formaldehyde solutions in a fume hood. Dispose of waste according to institutional guidelines.</li>
			<li>Option #2: Prepare formaldehyde/cacodylate fixation. Prepare Fix Mix: 250 mM sucrose, 100 mM potassium acetate (pH 7.5), 25 mM sodium acetate (pH 7.0), and 25 mM EGTA (pH 8.0). To use, make fresh with 400 &#181;l Fix Mix, 100 &#181;l potassium cacodylate (1 M, pH 7.2), and 500 &#181;l 16% formaldehyde per oocyte prep. 
			&#8203;CAUTION: Potassium cacodylate contains Arsenic.</li>
		</ol>
		</li>
		<li>Prepare PBS/Triton X-100: 1x PBS plus 1% or 0.05% Triton X-100. Store at 4 &#176;C.</li>
		<li>Prepare PBS-Tween 20-Bovine Serum Albumin (BSA) (PTB): 1x PBS, 0.5% BSA (w/v), and 0.1% Tween-20. May be stored at 4 &#176;C for one week.</li>
		<li>Fluorescence In Situ Hybridization (FISH) Solutions.
		<ol>
			<li>Prepare 20X Sodium chloride-Sodium Citrate (SSC): 3 M sodium chloride and 0.3 M sodium citrate.</li>
			<li>Prepare 2x SSC-Tween-20 (SSCT): 2x SSC plus 0.1% Tween-20. Make fresh, ~20 ml per oocyte prep.</li>
			<li>Prepare formamide solutions: 2x SSC, 0.1% Tween-20, plus formamide. Make fresh, 1 ml 20% formamide, 0.5 ml 40% formamide, and 2 ml 50% formamide per oocyte prep. 
			CAUTION: Wear gloves while using formamide solutions in a fume hood. Dispose of waste according to institutional guidelines.</li>
			<li>Prepare hybridization solution: 2x SSC, 50% formamide, and 10% dextran sulfate (w/v). Store at 4 &#176;C.</li>
		</ol>
		</li>
		<li>FISH Probes.
		<ol>
			<li>Order oligonucleotides (see Table 1 for sequences) with HPLC purification and desired 5&#39; fluorescent modification (e.g., Cy3 or Cy5). Resuspend in Tris-EDTA (TE) at 50 ng/&#181;l. 
			&#8203;NOTE: Protect oligos from light exposure at all times.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Collection of Late-stage Drosophila Oocytes
<ol>
	<li>As Drosophila oocytes with membranes intact will stick to plastic and glass, pre-coat the inside of one 5 ml tube and one Pasteur pipet per oocyte prep with PTB.</li>
	<li>Anesthetize all ~100 to 300 yeasted flies with carbon dioxide and add to a blender containing ~100 ml 1x Robb&#39;s Buffer. Pulse three times (~1 sec each). Keep oocytes in Robb&#39;s for &#60;20 min to avoid activation. 
	NOTE: Alternatively, oocytes may be hand-dissected from females. The advantage of this method is that it requires less females. However, care must be taken to limit exposure to carbon dioxide to only a few minutes to avoid artifacts associated with hypoxia7.</li>
	<li>Filter through large mesh (~1,500 &#181;m) into 250 ml beaker to remove large body parts. If many intact abdomens remain on mesh, re-grind material using additional Robb&#39;s Buffer, and filter again. Let settle ~2 min, then aspirate off top layer, removing as many of the large body parts as possible.</li>
	<li>Filter through small mesh (~300 &#181;m) into a 250 ml beaker. Rinse remaining oocytes out of first beaker using additional Robb&#39;s and coated Pasteur pipet. Let settle ~3 min; oocytes will settle out. Aspirate off all but ~10 ml.</li>
	<li>Pour as much of the 10 ml as will fit into coated 5 ml tube. Let settle, remove liquid, and repeat with remainder. Rinse remaining oocytes out of beaker using additional Robb&#39;s and coated Pasteur pipet. Let settle in 5 ml tube for ~3-5 min.</li>
</ol>
3. Drug Treatments (Optional)
<ol>
	<li>Coat a second 5 ml tube with PTB for each oocyte prep. Add appropriate solvent (control) or drug to 1 ml Robb&#39;s each for each oocyte prep (Table 2).</li>
	<li>Split oocytes into second coated 5 ml tube. Let settle, remove liquid, and add 1 ml Robb&#39;s plus solvent into one tube and 1 ml Robb&#39;s plus drug into second tube. Nutate for appropriate amount of time for drug treatment (Table 2). Let settle.</li>
</ol>
4. Fixation
<ol>
	<li>Aspirate off all liquid and immediately add 1 ml Fix.</li>
	<li>Option #1: Formaldehyde/heptane fixation (Fixation Buffer plus 5% formaldehyde).
	<ol>
		<li>Fix for 2.5 min on a nutator. Add 1 ml heptane and vortex 1 min. Let settle ~1 min.</li>
		<li>Remove all liquid, and then add 1 ml 1x PBS. Vortex 30 sec. Let settle ~1 min.</li>
		<li>Remove all liquid, and then fill tube with 1x PBS. 
		NOTE: Oocytes may be used immediately or kept on the nutator for several hours at room temperature.</li>
	</ol>
	</li>
	<li>Option #2: Formaldehyde/cacodylate fixation (Fix Mix plus 8% formaldehyde and 100 mM cacodylate).
	<ol>
		<li>Fix for 6 min on a nutator. Let settle 2 min. Remove liquid, and then fill tube with 1x PBS. 
		&#8203;NOTE: Oocytes may be used immediately or kept on the nutator for several hours at room temperature.</li>
	</ol>
	</li>
</ol>
5. Removing Membranes (&#34;Rolling&#34;)
<ol>
	<li>Using coated Pasteur pipet, add ~500 to 1,000 oocytes to the frosted (sand-blasted) part of a glass slide. Remove all body parts and extraneous material using forceps. Do not let oocytes dry out; add 1x PBS as necessary.</li>
	<li>Place a coverslip on top of the oocytes and gently &#34;roll&#34; oocytes until all membranes are removed (dragging the edge of the coverslip across the oocytes works best.) Periodically check progress under the microscope, adding more 1x PBS as necessary. Take care as too much pressure will destroy the oocytes. 
	&#8203;NOTE: For immunofluorescence only, continue with Step 6. For FISH (with or without immunofluorescence), continue with Step 7.</li>
</ol>
6. Antibody Staining of Drosophila Oocytes
<ol>
	<li>Extraction and Blocking
	<ol>
		<li>Rinse rolled oocytes into a 15 ml conical tube containing ~15 ml PBS/1% Triton X-100. Nutate oocytes in this solution for no less than 1.5 hr and no more than 2 hr. This step allows antibody penetration.</li>
		<li>Let oocytes settle ~2 min. Remove liquid along with as many membranes as possible, then add PBS/0.05% Triton X-100. 
		&#8203;NOTE: Membranes will settle slower than rolled oocytes.</li>
		<li>Let oocytes settle ~2 min, then remove all but ~1 ml liquid. Transfer oocytes to graduated 1.5 ml tube and remove remaining liquid. Add 1 ml PTB for blocking and nutate for 1 hr.</li>
	</ol>
	</li>
	<li>Antibody Staining
	<ol>
		<li>Pre-absorption of secondary antibodies against embryos. 
		&#8203;NOTE: This step eliminates background staining from non-specific antibody interactions with Drosophila proteins.
		<ol>
			<li>Collect and fix Drosophila embryos (~25 &#181;l) per typical procedure8. Store in methanol at -20 &#176;C.</li>
			<li>Remove methanol from embryos, add 800 &#181;l methanol plus 200 &#181;l 1x PBS, and nutate for 15 min. Remove 500 &#181;l of supernatant and replace with 500 &#181;l 1x PBS. Then nutate for 15 min. Repeat twice (for a total of ~1 hr of washes), and then finish in PTB. 
			&#8203;NOTE: Embryos may be used immediately or kept on the nutator for several hr at room temperature.</li>
			<li>Remove liquid, fill with PTB to 200 &#181;l per oocyte prep, and add fluorescently-labeled secondary antibodies at appropriate dilutions (keeping in mind that the final volume will be 300 &#181;l; see step 7.4.) Nutate at 4 &#176;C overnight (preferable) or at room temperature for 3 to 4 hr. 
			NOTE: The pre-absorbed antibodies will be used in Step 6.2.4. 
			&#8203;NOTE: Keep samples in the dark as much as possible once fluorescent antibodies have been added.</li>
		</ol>
		</li>
		<li>Remove liquid from oocytes, fill with PTB to 300 &#181;l, and add primary antibodies at appropriate dilutions. Nutate at 4 &#176;C overnight (preferable) or at room temperature for 3 to 4 hr.</li>
		<li>Wash oocytes four times for 15 min each with 1 ml PTB.</li>
		<li>Remove liquid from oocytes, add 200 &#181;l of supernatant from embryos (pre-absorbed secondary antibodies). Then fill with PTB to 300 &#181;l. Nutate at room temperature for 3 to 4 hr (preferable) or at 4 &#176;C overnight.</li>
		<li>Wash oocytes once for 15 min with 1 ml PTB, remove liquid. Then add 0.5 &#181;l Hoechst 33342 and 500 &#181;l PTB and nutate for 7 min.</li>
		<li>Wash oocytes twice for 15 min each with 1 ml PTB. 
		&#8203;NOTE: Oocytes may be mounted on a slide immediately or may be stored in PTB at 4 &#176;C until ready for imaging.</li>
	</ol>
	</li>
</ol>
7. FISH (Continue from Step 5 Above)
<ol>
	<li>Rinse rolled oocytes into a 15 ml conical tube containing ~15 ml 2x SSCT. Let oocytes settle ~2 min. Remove all but ~0.5 ml liquid along with as many membranes as possible. 
	NOTE: Membranes will settle slower than rolled oocytes.</li>
	<li>Transfer oocytes to 0.5 ml tube and remove remaining liquid. Successively add and remove 500 &#181;l 20%, 40%, and 50% formamide solutions, nutating for 10 min in each solution. 
	NOTE: Oocytes will settle slower with higher percentages of formamide.</li>
	<li>Remove liquid, then add 500 &#181;l 50% formamide solution, and nutate at 37 &#176;C for 1 to 5 hr. 
	NOTE: Longer incubations result in better probe penetration.</li>
	<li>Remove liquid, leaving behind no more than ~100 &#181;l oocytes, and add 36 &#181;l hybridization solution plus 2 &#181;l of probe (50 ng/&#181;l) and 2 &#181;l of water or 2 &#181;l of a 2nd probe (Table 1).</li>
	<li>Incubate at 91 &#176;C for 3 min (FISH only) or 80 &#176;C for 20 min (FISH plus immunofluorescence), followed by incubation in a 37 &#176;C water bath overnight. 
	NOTE: These steps may be performed in a thermocycler.</li>
	<li>Do not remove liquid. Add 500 &#181;l 50% formamide solution and nutate at 37 &#176;C for 1 hr.</li>
	<li>Let settle, remove liquid, add 500 &#181;l 50% formamide solution, and nutate at 37 &#176;C for 1 hr.</li>
	<li>Let settle, remove liquid, add 500 &#181;l 20% formamide solution, and nutate at room temperature for 10 min.</li>
	<li>Perform three quick washes in 500 &#181;l 2x SSCT (let settle, remove then add liquid, invert several times, repeat.) Let settle, remove liquid, then add 500 &#181;l PTB and nutate for 4 hr.</li>
</ol>
8. Antibody Staining after FISH
<ol>
	<li>Remove liquid from oocytes, fill with PTB to 300 &#181;l, and add 10 &#181;l anti-&#945;-tubulin antibody conjugated to FITC. Alternatively, other antibodies can be used, following above protocol. Nutate at room temperature overnight.</li>
	<li>Wash oocytes once for 15 min with 500 &#181;l PTB, remove liquid, then add 0.5 &#181;l Hoechst 33342 and 500 &#181;l PTB and nutate for 7 min.</li>
	<li>Wash oocytes twice for 15 min each with 500 &#181;l PTB. 
	NOTE: Oocytes may be mounted on a slide immediately or may be stored in PTB at 4 &#176;C until ready for imaging. Keep oocytes in the dark as much as possible once fluorescent antibodies have been added.</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td fo:width="1in">Repeat Name</td>
			<td fo:width="1in">Chromosome</td>
			<td>Oligo Sequence*</td>
		</tr>
		<tr>
			<td>359</td>
			<td>X</td>
			<td>GGGATCGTTAGCACTGGTAATTAGCTGC</td>
		</tr>
		<tr>
			<td>AACAC</td>
			<td>2</td>
			<td>AACACAACACAACACAACACAACACAACACAACACAACAC</td>
		</tr>
		<tr>
			<td>dodeca</td>
			<td>3</td>
			<td>CCCGTACTGGTCCCGTACTCGGTCCCGTACTCGGT</td>
		</tr>
		<tr>
			<td>1.686</td>
			<td>2+3</td>
			<td>AATAACATAGAATAACATAGAATAACATAG</td>
		</tr>
		<tr>
			<td>AATAT</td>
			<td>4(+Y)</td>
			<td>AATATAATATAATATAATATAATATAATAT</td>
		</tr>
		<tr>
			<td colspan="3">*359 sequence from Eric Joyce, personal communication, other sequences from Sullivan et al.8</td>
		</tr>
	</tbody>
</table>
Table 1: FISH probes for Drosophila centromeric repeats.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Drug</td>
			<td>Solvent</td>
			<td>Stock concentration</td>
			<td>Final concentration</td>
			<td>Time of treatment</td>
			<td>Effect</td>
		</tr>
		<tr>
			<td>colchicine</td>
			<td>ethanol</td>
			<td>125 mM</td>
			<td>150 &#181;M</td>
			<td>10 min or 30 min</td>
			<td>destabilize non-kinetochore (10 min)9 or all (30 min) microtubules</td>
		</tr>
		<tr>
			<td>paclitaxel</td>
			<td>DMSO</td>
			<td>10 mM</td>
			<td>10 &#181;M</td>
			<td>10 min</td>
			<td>stabilize microtubules</td>
		</tr>
		<tr>
			<td>Binucleine 2</td>
			<td>DMSO</td>
			<td>25 mM</td>
			<td>25 &#181;M</td>
			<td>20 min</td>
			<td>inhibit Aurora B kinase10</td>
		</tr>
	</tbody>
</table>
Table 2: Drug treatment.

<ol>
	<li>Bridges, C. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-disjunction+as+proof+of+the+chromosome+theory+of+heredity.">Non-disjunction as proof of the chromosome theory of heredity.</a> Genetics. 1 (1), 1-52 (1916).</li><li>Hassold, T., Hunt, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=To+err+(meiotically)+is+human:+the+genesis+of+human+aneuploidy.">To err (meiotically) is human: the genesis of human aneuploidy.</a> Nat Rev Genet. 2 (4), 280-291 (2001).</li><li>Theurkauf, W. E., Hawley, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Meiotic+spindle+assembly+in+Drosophila+females:+behavior+of+nonexchange+chromosomes+and+the+effects+of+mutations+in+the+nod+kinesin-like+protein.">Meiotic spindle assembly in Drosophila females: behavior of nonexchange chromosomes and the effects of mutations in the nod kinesin-like protein.</a> J Cell Biol. 116 (5), 1167-1180 (1992).</li><li>Zou, J., Hallen, M. A., Yankel, C. D., Endow, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+microtubule-destabilizing+kinesin+motor+regulates+spindle+length+and+anchoring+in+oocytes.">A microtubule-destabilizing kinesin motor regulates spindle length and anchoring in oocytes.</a> J Cell Biol. 180 (3), 459-466 (2008).</li><li>Dernburg, A. F., Sedat, J. W., Hawley, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+evidence+of+a+role+for+heterochromatin+in+meiotic+chromosome+segregation.">Direct evidence of a role for heterochromatin in meiotic chromosome segregation.</a> Cell. 86 (1), 135-146 (1996).</li><li>Gilliland, W. D., Hughes, S. F., Vietti, D. R., Hawley, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Congression+of+achiasmate+chromosomes+to+the+metaphase+plate+in+Drosophila+melanogaster+oocytes.">Congression of achiasmate chromosomes to the metaphase plate in Drosophila melanogaster oocytes.</a> Dev Biol. 325 (1), 122-128 (2009).</li><li>Gilliland, W. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hypoxia+transiently+sequesters+mps1+and+polo+to+collagenase-sensitive+filaments+in+Drosophila+prometaphase+oocytes.">Hypoxia transiently sequesters mps1 and polo to collagenase-sensitive filaments in Drosophila prometaphase oocytes.</a> PLoS One. 4 (10), e7544 (2009).</li><li>Sullivan, W., Ashburner, M., Hawley, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Drosophila Protocols. , (2000).</li><li>Radford, S. J., Hoang, T. L., G&#322;uszek, A. A., Ohkura, H., McKim, K. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lateral+and+End-On+Kinetochore+Attachments+Are+Coordinated+to+Achieve+Bi-orientation+in+Drosophila+Oocytes.">Lateral and End-On Kinetochore Attachments Are Coordinated to Achieve Bi-orientation in Drosophila Oocytes.</a> PLoS Genet. 11 (10), e1005605 (2015).</li><li>Smurnyy, Y., Toms, A. V., Hickson, G. R., Eck, M. J., Eggert, U. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Binucleine+2,+an+isoform-specific+inhibitor+of+Drosophila+Aurora+B+kinase,+provides+insights+into+the+mechanism+of+cytokinesis.">Binucleine 2, an isoform-specific inhibitor of Drosophila Aurora B kinase, provides insights into the mechanism of cytokinesis.</a> ACS Chem Biol. 5 (11), 1015-1020 (2010).</li><li>Mahowald, A. P., Goralski, T. J., Caulton, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+activation+of+Drosophila+eggs.">In vitro activation of Drosophila eggs.</a> Dev Biol. 98 (2), 437-445 (1983).</li><li>Page, A. W., Orr-Weaver, T. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activation+of+the+meiotic+divisions+in+Drosophila+oocytes.">Activation of the meiotic divisions in Drosophila oocytes.</a> Dev Biol. 183 (2), 195-207 (1997).</li><li>Tavosanis, G., Llamazares, S., Goulielmos, G., Gonzalez, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Essential+role+for+gamma-tubulin+in+the+acentriolar+female+meiotic+spindle+of+Drosophila.">Essential role for gamma-tubulin in the acentriolar female meiotic spindle of Drosophila.</a> EMBO J. 16 (8), 1809-1819 (1997).</li><li>Endow, S. A., Komma, 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=Spindle+dynamics+during+meiosis+in+Drosophila+oocytes.">Spindle dynamics during meiosis in Drosophila oocytes.</a> J Cell Biol. 137 (6), 1321-1336 (1997).</li><li>Matthies, H. J., Clarkson, M., Saint, R. B., Namba, R., Hawley, R. S., Sullivan, W., Ashburner, M., Hawley, R. 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The authors have nothing to disclose.

Staging Drosophila Oocytes
Although an elongated karyosome is often seen in prometaphase oocytes, using karyosome shape to distinguish prometaphase from metaphase oocytes can be problematic. During prometaphase, the karyosome begins as a round shape, elongates, and then retracts to a round shape as the oocyte approaches the metaphase arrest. This means that many prometaphase oocytes do not have an elongated karyosome. In addition, if mutant or drug-treated oocytes are...

The study of meiosis is sometimes described as the "genetics of genetics". This is because the fundamental properties of chromosome inheritance and independent assortment are carried out through the segregation of chromosomes during gamete production. An important demonstration of the chromosome theory of inheritance came in 1916 from the work of Calvin Bridges in Drosophila melanogaster1. This and other classical genetics studies in Drosophila contributed greatly to our understanding of genetics. Cytological examination of meiotic chromosomes in Drosophila oocytes, however, has been challenging. This is primarily because immunofluorescence of late-stage Drosophila oocytes, when the spindle assembles and chromosomes are oriented for segregation, is hampered by the presence of membranes that render the oocyte impenetrable to antibodies.
Despite this challenge, Drosophila oocytes remain an attractive model for the study of chromosome behavior and spindle assembly. This is because of the powerful genetic tools available in Drosophila, but also because the oocytes arrest at metaphase I, when the chromosomes are oriented and the spindle is fully formed. This facilitates the collection and examination of large numbers of oocytes at this important stage of cell division. In addition, a simple model organism that is amenable to genetic manipulation for the study of oocyte chromosome segregation can provide an important contribution to our understanding of human reproductive health. Errors in chromosome number are the leading genetic cause of miscarriage and birth defects in humans2. A majority of these errors can be traced to the oocyte and are correlated with increasing maternal age. The average age of mothers in the U.S. has been increasing, making this a major public health concern.
We describe here methods for the cytological examination of Drosophila oocytes, including a demonstration of how to remove the oocyte membranes. These methods are modifications of protocols first described by Theurkauf and Hawley3, Zou et al.4, and Dernburg et al.5. We also include methods for the enrichment of different stages of oocytes, based on a protocol first described by Gilliland et al.6. Finally, we add instructions for the drug treatment of Drosophila oocytes. Together, these methods allow the cytological investigation of oocyte chromosome segregation and spindle assembly in Drosophila.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>15 mL conical tubes</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>16% formaldehyde</td>
			<td>Ted Pella, Inc.</td>
			<td>18505</td>
			<td>HAZARDOUS; once opened, discard after one month</td>
		</tr>
		<tr>
			<td>250 mL beakers</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>5 mL tubes</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>active dry yeast</td>
			<td>Various</td>
			<td></td>
			<td>mix with water to make a paste the consistency of peanut butter</td>
		</tr>
		<tr>
			<td>anti-&#945;-tubulin antibody conjugated to FITC</td>
			<td>Sigma</td>
			<td>F2168</td>
			<td>clone DM1A</td>
		</tr>
		<tr>
			<td>Binucleine 2</td>
			<td>Sigma</td>
			<td>B1186</td>
			<td>HAZARDOUS</td>
		</tr>
		<tr>
			<td>blender</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>bovine serum albumin</td>
			<td>Sigma</td>
			<td>A4161</td>
			<td></td>
		</tr>
		<tr>
			<td>calcium chloride</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>colchicine</td>
			<td>Sigma</td>
			<td>C-9754</td>
			<td>HAZARDOUS</td>
		</tr>
		<tr>
			<td>coverslips</td>
			<td>VWR</td>
			<td>48366-227</td>
			<td>No. 1 1/2</td>
		</tr>
		<tr>
			<td>dextran sulfate</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ethanol</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>forceps</td>
			<td>Ted Pella, Inc.</td>
			<td>5622</td>
			<td>Dumont tweezers high precision grade style 5</td>
		</tr>
		<tr>
			<td>formamide</td>
			<td>Sigma</td>
			<td>47670-250ML-F</td>
			<td></td>
		</tr>
		<tr>
			<td>glass slides</td>
			<td>VWR</td>
			<td>48312-003</td>
			<td></td>
		</tr>
		<tr>
			<td>glucose</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>graduated 1.5 mL tubes</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>VWR</td>
			<td>EM-5330</td>
			<td>available from several venders</td>
		</tr>
		<tr>
			<td>Hoechst 33342</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>magnesium chloride</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>methanol</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>large mesh (~1500 &#181;m)</td>
			<td>VWR</td>
			<td>AA43657-NK</td>
			<td>variety of formats and other suppliers, 12 or 14 mesh</td>
		</tr>
		<tr>
			<td>small mesh (~300 &#181;m)</td>
			<td>Spectrum labs</td>
			<td>146 424</td>
			<td>variety of formats, eg 146 422 or 146 486</td>
		</tr>
		<tr>
			<td>nutator</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Pasteur pipets</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>potassium acetate</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cacodylic acid</td>
			<td>Sigma</td>
			<td>C0125</td>
			<td>HAZARDOUS; alternatively, sodium cacodylate may be substituted</td>
		</tr>
		<tr>
			<td>potassium hydroxide</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>sodium acetate</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>sodium chloride</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>sodium citrate</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>sodium hydroxide</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>sucrose</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>taxol (paclitaxel)</td>
			<td>Sigma</td>
			<td>T1912</td>
			<td>HAZARDOUS</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Fisher</td>
			<td>PI-28314</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween 20</td>
			<td>Fisher</td>
			<td>PI-28320</td>
			<td></td>
		</tr>
		<tr>
			<td>vortex</td>
			<td>Various</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

techniques for imaging prometaphase and metaphase of meiosis i in fixed drosophila oocytes

Chromosome segregation in human oocytes is error prone, resulting in aneuploidy, which is the leading genetic cause of miscarriage and birth defects. The study of chromosome behavior in oocytes from model organisms holds much promise to uncover the molecular basis of the susceptibility of human oocytes to aneuploidy. Drosophila melanogaster is amenable to genetic manipulation, with over 100 years of research, community, and technique development. Visualizing chromosome behavior and spindle assembly in Drosophila oocytes has particular challenges, however, due primarily to the presence of membranes surrounding the oocyte that are impenetrable to antibodies. We describe here protocols for the collection, preparation, and imaging of meiosis I spindle assembly and chromosome behavior in Drosophila oocytes, which allow the molecular dissection of chromosome segregation in this important model organism.

The methods we have described here will result in the collection of late-stage Drosophila oocytes representing three stages of meiosis (Figure 1). Oocytes in prophase are distinguished by the presence of the nuclear envelope, which is visible by the lack of tubulin signal in the region surrounding the karyosome (Figure 1A). Prometaphase is the period after nuclear envelope breakdown during which the spindle assembles. During prometaphase, the kar...

Watch this Scientific Journal Video about Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes at JoVE.com

Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes

We present protocols for the collection, preparation, and imaging of mature Drosophila oocytes. These methods allow the visualization of chromosome behavior and spindle assembly and function during meiosis.

Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes

Chromosome segregation in human oocytes is error prone, resulting in aneuploidy, which is the leading genetic cause of miscarriage and birth defects. The ...

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Unit 2

Meiosis

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9.0K Views.  Rutgers University. We present protocols for the collection, preparation, and imaging of mature Drosophila oocytes. These methods allow the visualization of chromosome behavior and spindle assembly and function during meiosis.

Video: Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of the Drosophila melanogaster syncytial embryo for studying mitosis1. Drosophila has useful genetics with a sequenced genome, and it can be easily maintained and manipulated. Many mitotic mutants exist, and transgenic flies expressing functional fluorescently (e.g. GFP) - tagged mitotic proteins have been and are being generated. Targeted gene expression is possible using the GAL4/UAS system2. 

The Drosophila early embryo carries out multiple mitoses very rapidly (cell cycle duration, &asymp;10 min). It is well suited for imaging mitosis, because during cycles 10-13, nuclei divide rapidly and synchronously without intervening cytokinesis at the surface of the embryo in a single monolayer just underneath the cortex. These rapidly dividing nuclei probably use the same mitotic machinery as other cells, but they are optimized for speed; the checkpoint is generally believed to not be stringent, allowing the study of mitotic proteins whose absence would cause cell cycle arrest in cells with a strong checkpoint. Embryos expressing GFP labeled proteins or microinjected with fluorescently labeled proteins can be easily imaged to follow live dynamics (Fig. 1). In addition, embryos can be microinjected with function-blocking antibodies or inhibitors of specific proteins to study the effect of the loss or perturbation of their function3. These reagents can diffuse throughout the embryo, reaching many spindles to produce a gradient of concentration of inhibitor, which in turn results in a gradient of defects comparable to an allelic series of mutants. Ideally, if the target protein is fluorescently labeled, the gradient of inhibition can be directly visualized4. It is assumed that the strongest phenotype is comparable to the null phenotype, although it is hard to formally exclude the possibility that the antibodies may have dominant effects in rare instances, so rigorous controls and cautious interpretation must be applied. Further away from the injection site, protein function is only partially lost allowing other functions of the target protein to become evident.

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of microinjection and high resolution imaging in the Drosophila melanogaster syncytial embryo to study mitosis.

This protocol describes the use of the Drosophila melanogaster syncytial embryo  for studying mitosis1.  Drosophila has useful genetics with a sequenced ...

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

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

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

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

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

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 ...

Techniques for Imaging Ca2+ Signaling in Human Sperm

Fluorescence microscopy of cells loaded with fluorescent, Ca2+-sensitive dyes is used for measurement of spatial and temporal aspects of Ca2+ signaling in live cells. Here we describe the method used in our laboratories for loading suspensions of human sperm with Ca2+-reporting dyes and measuring the fluorescence signal during physiological stimulation. Motile cells are isolated by direct swim-up and incubated under capacitating conditions for 0-24 h, depending upon the experiment. The cell-permeant AM (acetoxy methyl ester) ester form of the Ca2+-reporting dye is then added to a cell aliquot and a period of 1 h is allowed for loading of the dye into the cytoplasm. We use visible wavelength dyes to minimize photo-damage to the cells, but this means that ratiometric recording is not possible. Advantages and disadvantages of this approach are discussed. During the loading period cells are introduced into an imaging chamber and allowed to adhere to a poly-D-lysine coated coverslip. At the end of the loading period excess dye and loose cells are removed by connection of the chamber to the perfusion apparatus. The chamber is perfused continuously, stimuli and modified salines are then added to the perfusion header. Experiments are recorded by time-lapse acquisition of fluorescence images and analyzed in detail offline, by manually drawing regions of interest. Data are normalized to pre-stimulus levels such that, for each cell (or part of a cell), a graph showing the Ca2+ response as % change in fluorescence is obtained.

Techniques for Imaging Ca2+ Signaling in Human Sperm

Stimulus-evoked [Ca2+]i signals of individual human sperm are assessed. Motile cells are loaded with Ca2+-sensitive fluorescent dye (AM-ester method) and immobilised in a perfusable chamber. Cells are imaged by time-lapse fluorescence microscopy and stimulated via the perfusing medium. Responses of single cells (or regions) are analysed offline using Excel.

Fluorescence microscopy of cells loaded with fluorescent, Ca2+-sensitive dyes is used for measurement of spatial and temporal aspects of Ca2+ signaling in ...

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 ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

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.

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...