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.

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	<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. 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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><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-&amp;alpha;-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 (~1,500 &amp;micro;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 &amp;micro;m)</td><td>Spectrum labs</td><td>146 424</td><td>variety of formats, &lt;em&gt;e.g.&lt;/em&gt;, 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...

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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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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

Using Mouse Oocytes to Assess Human Gene Function During Meiosis I

Embryonic aneuploidy is the major genetic cause of infertility in humans. Most of these events originate during female meiosis, and albeit positively correlated with maternal age, age alone is not always predictive of the risk of generating an aneuploid embryo. Therefore, gene variants might account for incorrect chromosome segregation during oogenesis. Given that access to human oocytes is limited for research purposes, a series of assays were developed to study human gene function during meiosis I using mouse oocytes. First, messenger RNA (mRNA) of the gene and gene variant of interest are microinjected into prophase I-arrested mouse oocytes. After allowing time for expression, oocytes are synchronously released into meiotic maturation to complete meiosis I. By tagging the mRNA with a sequence of a fluorescent reporter, such as green fluorescent protein (Gfp), the localization of the human protein can be assessed in addition to the phenotypic alterations. For example, gain or loss of function can be investigated by establishing experimental conditions that challenge the gene product to fix meiotic errors. Although this system is advantageous in investigating human protein function during oogenesis, adequate interpretation of results should be undertaken given that protein expression is not at endogenous levels and, unless controlled for (i.e. knocked out or down), murine homologs are also present in the system.

As the genetic variants associated with human disease begin to become uncovered, it is becoming increasingly important to develop systems with which to rapidly evaluate the biological significance of those identified variants. This protocol describes methods for evaluating human gene function during female meiosis I using mouse oocytes.

Embryonic aneuploidy is the major genetic cause of infertility in humans. Most of these events originate during female meiosis, and albeit positively ...

Genetics

Using Fluorescence In Situ Hybridization (FISH) to Monitor the State of Arm Cohesion in Prometaphase and Metaphase I Drosophila Oocytes

In humans, chromosome segregation errors in oocytes are responsible for the majority of miscarriages and birth defects. Moreover, as women age, their risk of conceiving an aneuploid fetus increases dramatically and this phenomenon is known as the maternal age effect. One requirement for accurate chromosome segregation during the meiotic divisions is maintenance of sister chromatid cohesion during the extended prophase period that oocytes experience. Cytological evidence in both humans and model organisms suggests that meiotic cohesion deteriorates during the aging process. In addition, segregation errors in human oocytes are most prevalent during meiosis I, consistent with premature loss of arm cohesion. The use of model organisms is critical for unraveling the mechanisms that underlie age-dependent loss of cohesion. Drosophila melanogaster offers several advantages for studying the regulation of meiotic cohesion in oocytes. However, until recently, only genetic tests were available to assay for loss of arm cohesion in oocytes of different genotypes or under different experimental conditions. Here, a detailed protocol is provided for using fluorescence in situ hybridization (FISH) to directly visualize defects in arm cohesion in prometaphase I and metaphase I arrested Drosophila oocytes. By generating a FISH probe that hybridizes to the distal arm of the X chromosome and collecting confocal Z stacks, a researcher can visualize the number of individual FISH signals in three dimensions and determine whether sister chromatid arms are separated. The procedure outlined makes it possible to quantify arm cohesion defects in hundreds of Drosophila oocytes. As such, this method provides an important tool for investigating the mechanisms that contribute to cohesion maintenance as well as the factors that lead to its demise during the aging process.

Using Fluorescence In Situ Hybridization FISH to Monitor the State of Arm Cohesion in Prometaphase and Metaphase I Drosophila Oocytes

This manuscript presents a detailed method for generating X-chromosome arm probes and performing fluorescence in situ hybridization (FISH) to examine the state of sister chromatid cohesion in prometaphase and metaphase I arrested Drosophila oocytes. This protocol is suitable for determining whether meiotic arm cohesion is intact or disrupted in different genotypes.

In humans, chromosome segregation errors in oocytes are responsible for the majority of miscarriages and birth defects. Moreover, as women age, their risk ...

Developmental Biology

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 ...

Nuclear Migration in the Drosophila Oocyte

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

Nuclear Migration in the Drosophila Oocyte

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

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

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment and maintenance of epithelial tissue morphology and organ morphogenesis. Moreover, the BM is essential for tissue modeling, serving as a signaling platform, and providing external forces to shape tissues and organs. Despite the many important roles that the BM plays during normal development and pathological conditions, the biological pathways controlling the intracellular trafficking of BM-containing vesicles and how basal secretion leads to the polarized deposition of BM proteins are poorly understood. The follicular epithelium of the Drosophila ovary is an excellent model system to study the basal deposition of BM membrane proteins, as it produces and secretes all major components of the BM. Confocal and super-resolution imaging combined with image processing in fixed tissues allows for the identification and characterization of cellular factors specifically involved in the intracellular trafficking and deposition of BM proteins. This article presents a detailed protocol for staining and imaging BM-containing vesicles and deposited BM using endogenously tagged proteins in the follicular epithelium of the Drosophila ovary. This protocol can be applied to address both qualitative and quantitative questions and it was developed to accommodate high-throughput screening, allowing for the rapid and efficient identification of factors involved in the polarized intracellular trafficking and secretion of vesicles during epithelial tissue development.

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane is essential for tissue and organ morphogenesis during development. To better understand the mechanisms leading to proper placement of this structure, the protocol presented describes methods to visualize and characterize the intracellular trafficking and secretion of basement membrane proteins in epithelial cells using confocal and super-resolution microscopy.

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment ...

Mouse Oocyte Microinjection, Maturation and Ploidy Assessment

Mistakes in chromosome segregation lead to aneuploid cells. In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to infertility, miscarriages or developmental disorders like Down syndrome. Haploid gametes form through species-specific developmental programs that are coupled to meiosis. The first meiotic division (MI) is unique to meiosis because sister chromatids remain attached while homologous chromosomes are segregated. For reasons not fully understood, this reductional division is prone to errors and is more commonly the source of aneuploidy than errors in meiosis II (MII) or than errors in male meiosis 1,2. 
In mammals, oocytes arrest at prophase of MI with a large, intact germinal vesicle (GV; nucleus) and only resume meiosis when they receive ovulatory cues. Once meiosis resumes, oocytes complete MI and undergo an asymmetric cell division, arresting again at metaphase of MII. Eggs will not complete MII until they are fertilized by sperm. Oocytes also can undergo meiotic maturation using established in vitro culture conditions 3. Because generation of transgenic and gene-targeted mouse mutants is costly and can take long periods of time, manipulation of female gametes in vitro is a more economical and time-saving strategy. 
Here, we describe methods to isolate prophase-arrested oocytes from mice and for microinjection. Any material of choice may be introduced into the oocyte, but because meiotically-competent oocytes are transcriptionally silent 4,5 cRNA, and not DNA, must be injected for ectopic expression studies. To assess ploidy, we describe our conditions for in vitro maturation of oocytes to MII eggs. Historically, chromosome-spreading techniques are used for counting chromosome number 6. This method is technically challenging and is limited to only identifying hyperploidies. Here, we describe a method to determine hypo-and hyperploidies using intact eggs 7-8. This method uses monastrol, a kinesin-5 inhibitor, that collapses the bipolar spindle into a monopolar spindle 9 thus separating chromosomes such that individual kinetochores can readily be detected and counted by using an anti-CREST autoimmune serum. Because this method is performed in intact eggs, chromosomes are not lost due to operator error.

Oocytes are prone to aneuploidy due to errors in chromosome segregation during meiotic maturation. Aneuploid eggs can cause infertility, miscarriages or developmental disorders like Down syndrome. Here, we describe methods to introduce materials of choice into oocytes and methods to study meiotic maturation and assess ploidy.

Mistakes in chromosome segregation lead to aneuploid cells.  In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to ...

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle ...

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 ...

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both the female and male germ lines of Drosophila have contributed greatly to the current understanding of meiosis as well as stem cell biology. Excellent protocols are available in the literature for the isolation and imaging of Drosophila ovaries&#160;and testes3-12. Herein, methods for the dissection and preparation of Drosophila testes for microscopic analysis are described with an accompanying video demonstration. A protocol for isolating testes from the abdomen of adult males and preparing slides of live tissue for analysis by phase-contrast microscopy as well as a protocol for fixing and immunostaining testes for analysis by fluorescence microscopy are presented. These techniques can be applied in the characterization of Drosophila mutants that exhibit defects in spermatogenesis as well as in the visualization of subcellular localizations of proteins.

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Methods for isolating and preparing Drosophila testes samples (live and fixed) for imaging by phase-contrast and fluorescence microscopy are described herein.&#160;

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both ...

Preparation of Fixed Drosophila Oocytes for Immunostaining: A High-Throughput Method to Fix and Remove the Outer Membrane

Source: Radford, S. J., McKim, K. S. <a href="https://www.jove.com/video/54666/" target="_blank">Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes</a>. J. Vis. Exp. (2016).
Immunostaining of Drosophila late-stage oocytes can prove challenging due to the oocytes&#39; surrounding membranes. This video describes a high-throughput method for obtaining oocytes, their fixation, and membranes&#39; removal for immunostaining. The featured protocol demonstrates these techniques with late-stage oocytes used to visualize different stages of meiosis.

Source: Radford, S. J., McKim, K. S. Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes. J. Vis. Exp. (2016).
...

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

Summary

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Chapters in this video

Using Mouse Oocytes to Assess Human Gene Function During Meiosis I

Using Fluorescence In Situ Hybridization (FISH) to Monitor the State of Arm Cohesion in Prometaphase and Metaphase I Drosophila Oocytes

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

Nuclear Migration in the Drosophila Oocyte

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

Mouse Oocyte Microinjection, Maturation and Ploidy Assessment

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Preparation of Fixed Drosophila Oocytes for Immunostaining: A High-Throughput Method to Fix and Remove the Outer Membrane