Name: techniques for imaging prometaphase and metaphase of meiosis i in fixed drosophila oocytes
Uploaded: 2016-10-31T15:00:00
Description: Watch this Scientific Journal Video about Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes at JoVE.com

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 c...

<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. 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. , 67-85 (2000).</li><li>Colombi&#233;, N., 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=Dual+roles+of+Incenp+crucial+to+the+assembly+of+the+acentrosomal+metaphase+spindle+in+female+meiosis.">Dual roles of Incenp crucial to the assembly of the acentrosomal metaphase spindle in female meiosis.</a> Development. 135 (19), 3239-3246 (2008).</li></ol>

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 ...

The overall goal of this procedure is to prepare Drosophila oocytes for the cytological examination of spindle assembly and chromosome orientation during meiosis I.This method can help answer key questions in the meiosis field, such as how spindle bipolarity is established and how chromosomes become bi-oriented in the absence of centrosomes. The main advantage of this technique is that the oocyte membranes that block antibody penetration are effectively removed, which allows immunofluorescence. To begin pre-coat the inside of a five-millimeter tube and pasture pipette with PTB to prevent oocytes sticking to the surfaces.Add 100 milliliters of ROB's buffer to a blender and then add 100 to 300 anesthetized flies and pulse three times for one second. Filter the resulted slurry into a 250-milliliter beaker through a large mesh with pores of around 1, 500 micrometers. Let the filtrate settle for around two minutes and then aspirate the top layer, removing as many large body parts as possible.Filter the slurry again into a 250-milliliter beaker, through a smaller mesh with the pore size of 300 micrometers. Rinse the first beaker using additional buffer in the coated pipette to collect the remaining oocytes. Let the slurry settle for three minutes and then aspirate all but the bottom 10 milliliters.Pour as much of the 10 milliliters as possible into the coated five-milliliter tube. Let the slurry settle for around two minutes and then carefully remove the liquid. Add the remainder of the 10 milliliters of slurry and let the mix settle again before removing the liquid.Rinse any remaining oocytes out of the beaker using additional buffer in the coated pipette. Let the rinse settle in the five-milliliter tube for three to five minutes. To fix the ovaries, first aspirate off all liquid from the tissues and immediately add one milliliter of Fix Mix.Place the tissues on a nutator for 2 1/2 minutes. Next, add one milliliter of heptane, vortex the tube for one minute, and then allow the oocytes to settle for a further minute. Remove all liquid from the settled tissue, then add one milliliter of 1x PBS.Vortex the tube for 30 seconds and then allow the oocytes to settle for one minute. After that, remove all liquid from the tube and fill it with five milliliters of 1x PBS. Using the coated pipette, add roughly 500 to 1, 000 oocytes to a frosted glass slide.Remove any extraneous body parts or material using forceps. Place a cover slip on top of the tissue and gently roll the oocytes until all membranes are removed. Dragging the edge of the coverslip across the oocytes works well.Check the progress of membrane removal under a microscope periodically. Too much pressure will destroy the oocytes. The most effective oocyte membrane removal is achieved by combining the maximum amount of oocyte movement with the minimum amount of pressure.Use a light touch and quick movements to drag the edge of the coverslip across the oocytes. To begin extraction and blocking, add 15 milliliters of PBS 1%Triton X-100 to a 50-milliliter tube. Rinse the rolled oocytes into this tube and nutate for 1 1/2 to two hours.After allowing the oocytes to settle for two minutes, remove the liquid along with as many membranes as possible and add 50 milliliters of PBS 0.05%Triton X-100. Let the oocytes settle for a further two minutes, and then remove all but around one milliliter of liquid. Transfer the oocytes to a graduated 1.5 milliliter tube and remove the remaining liquid.Add one milliliter of PTB for blocking and nutate for one hour. Remove the liquid from the oocytes and then add 300 microliters of PTB. Next, add the primary antibody of interest at the appropriate dilution, taking care to protect the samples treated with fluorescent antibody from light.Nutute overnight at four degrees Celsius. Wash the oocytes and one milliliter of PTB four times for 15 minutes each. Remove the liquid, and add 200 microliters of supernatant from the secondary antibody-treated embryos.Filter the 300-microliter mark with PTB and then nutate for three to four hours at room temperature. Wash the oocytes once for 15 minutes with one milliliter of PTB. Next, remove the liquid and add 0.5 microliters of Hoechst 33342 and 500 microliters of PTB.Set to nutate for seven minutes. Use one milliliter of PTB to wash the oocytes two times for 15 minutes each. The samples can now be mounted and imaged immediately using standard confocal microscopy techniques, or stored in PTB at four degrees Celsius until needed.Images of late-stage oocytes generated using the demonstrated technique are shown here, capturing three different stages of meiosis. Oocytes in prophase are distinguished by the presence of the nuclear envelope which is visible due to the lack of tubulin signal in the regions surrounding the karyosome. The distinctive elongation of the karyosome in the prometaphase was also observed, and the fourth chromosomes are seen apart from the karyosome mass.Metaphase was also captured and the karyosome is seen retracted into a round shape. Here immunostaining with different antibodies highlights the spindle, centromeres, and central spindle in oocytes fixed with formaldehyde and heptane. Oocytes treated with the drug Colchicine showed non-standard phenotypes with the elimination of most non-kinetic or microtubules, resulting in all microtubules contacting the DNA at centromere labeled Phocyde.Treatment with Paclitaxel resulted in oocytes with excessive microtubules in the cytoplasm. Finally oocytes treated with Binucleine 2 to inhibit Aurora B kinase, showed complete loss of spindle microtubules. While attempting this procedure, it's important to remember not to rush during oocyte membrane removal, because the success of the protocol depends on this step.Generally individuals new to this method will struggle, because the removal of the membranes requires patience and practice. This technique is therefore best learned by watching someone perform it successfully. Once mastered, rolling the oocytes can be done in five to 10 minutes.But it's possible to take up to 30 minutes without affecting the quality of the results. After watching this video, you should have a good understanding of how to prepare late-stage Drosophila oocytes for imaging biotic spindle assembly, spindle organization, chromosome movements, and chromosome bi-orientation. Thanks for watching, and good luck with your experiments.

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SCIENTISTS IN ACTION

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 ...

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  ...

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized microtubules. Organelles move along these processes by microtubule motors. Easy maintenance, high sensitivity to RNAi-mediated protein knock-down and efficient procedure for creating stable cell lines make Drosophila S2 cells an ideal model system to study cargo transport by live imaging. The results obtained with S2 cells can be further applied to a more physiologically relevant system: axonal transport in primary neurons cultured from dissociated Drosophila embryos. Cultured neurons grow long neurites filled with bundled microtubules, very similar to S2 processes. Like in S2 cells, organelles in cultured neurons can be visualized by either organelle-specific fluorescent dyes or by using fluorescent organelle markers encoded by DNA injected into early embryos or expressed in transgenic flies. Therefore, organelle transport can be easily recorded in neurons cultured on glass coverslips using living imaging. Here we describe procedures for culturing and visualizing cargo transport in Drosophila S2 cells and primary neurons. We believe that these protocols make both systems accessible for labs studying cargo transport.

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila S2 cells and cultured neurons are great systems for imaging of motor-driven organelle transport in vivo. Here we describe detailed protocols for culturing both cell types, their imaging and analysis of transport.

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized ...