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).

Anmerkung: Verfahren bei Raumtemperatur durchgeführt, sofern nicht anders angegeben. Die temperaturgeregelte Inkubatoren verwendet werden Temperaturen für das Fliegenzucht zu halten und Kreuze, sofern nicht anders vermerkt. 1. Vorbereitungen <ol><li> Bereiten Sie Flies. <ol><li> Oozyten Sammlungen Prometaphase angereicherte. <ol><li> Klar erwachsenen Fliegen von gesunden, jungen Aktien oder Quer Kulturen. Alter Flaschen für zwei Tage bei 25 ° C. HINWEIS: In der Regel zwei gesunde Flaschen genügt, wenn auch für einige Quer Kulturen benötigt werden. </li><li> Nach zwei Tagen sammeln ~ 100 bis 300 Frauen (die zu diesem Zeitpunkt 0 bis 2 Tage alt sind) aus den Flaschen. Die Weibchen müssen nicht Jungfrauen zu sein. Fügen Sie einen Klecks Hefe-Paste auf die Seite eines Fläschchens und legen 30 Frauen und 10 bis 15 Männer jeweils in die yeasted Fläschchen. Alter Phiolen für zwei Tage bei 25 ° C. </li></ol></li><li> Metaphase angereicherte Oozyten Kollektionen. <ol><li> Sammeln Sie ~ 100 bis 300 Frauen ab Lager oder Quer cultures. Fügen Sie einen Klecks Hefe-Paste auf die Seite eines Fläschchens und legen 30 Frauen jeweils (ohne Männchen hinzugefügt) in die yeasted Fläschchen. Alter Phiolen für drei bis fünf Tage bei 25 ° C. </li></ol></li></ol></li><li> Bereiten Sie Lösungen. Hinweis: Die Lösungen werden bei Raumtemperatur unbegrenzt gelagert, wenn nicht anders angegeben. <ol><li> Prepare Modifizierte Robb-Puffer (5x): 500 mM HEPES, 500 mM Sucrose, 275 mM Natriumacetat, 200 mM Kaliumacetat, 50 mM Glucose, 6 mM Magnesiumchlorid und 5 mM Calciumchlorid. Verwenden 10 N 11: 8 Natriumhydroxid: Kaliumhydroxid pH auf 7,4 zu bringen. Sterilisieren durch Filtration; nicht im Autoklaven. Lagerung bei -20 ° C. nach Bedarf auftauen ~ 200 ml 1x Robb prep pro Eizelle vorzubereiten. </li><li> Fixation Solutions. <ol><li> Option # 1: Vorbereiten Formaldehyd / Heptan-Fixierung. Vorbereitung Fixation Buffer: 1x phosphatgepufferter Kochsalzlösung (PBS) plus 150 mM Saccharose. Verwenden zu können, machen frisch mit 687,5 ul Fixation Buffer und 312,5 ul 16% Formaldehydpro Eizelle prep. ACHTUNG: Handschuhe tragen bei der Verwendung von Formaldehydlösungen in einem Abzug. Entsorgen von Abfällen gemäß gültiger Richtlinien. </li><li> Option # 2: Vorbereiten Formaldehyd / Cacodylat Fixierung. Vorbereitung Fix Mix: 250 mM Sucrose, 100 mM Kaliumacetat (pH 7,5), 25 mM Natriumacetat (pH 7,0) und 25 mM EGTA (pH 8,0). Zur Verwendung machen frisch mit 400 ul Fix-Mix, 100 ul Kaliumcacodylat (1 M, pH 7,2) und 500 &amp; mgr; l 16% Formaldehyd pro Eizelle prep. ACHTUNG: Kaliumcacodylat enthält Arsen. </li></ol></li><li> Bereiten PBS / Triton X-100: 1x PBS plus 1% oder 0,05% Triton X-100. Lagerung bei 4 ° C. </li><li> Bereiten PBS-Tween 20-Rinderserumalbumin (BSA) (PTB): 1x PBS, 0,5% BSA (w / v) und 0,1% Tween-20. Kann für eine Woche bei 4 ° C gelagert werden. </li><li> Fluoreszenz in situ Hybridisierung (FISH) Lösungen. <ol><li> Bereiten 20X Natriumchlorid-Natriumcitrat (SSC): 3 M Natriumchlorid und 0,3 M Natriumcitrat. </li><li> Bereiten 2x SSC-Tween-20 (SSCT): 2x SSC plus 0,1% Tween-20. Machen Sie frische, ~ 20 ml pro Eizelle prep. </li><li> Bereiten Formamid-Lösungen: 2 x SSC, 0,1% Tween-20, sowie Formamid. Machen Sie frische, 1 ml 20% Formamid, 0,5 ml 40% Formamid und 2 ml 50% Formamid pro Eizelle prep. ACHTUNG: Handschuhe tragen, während Formamid Lösungen in einem Abzug mit. Entsorgen von Abfällen gemäß gültiger Richtlinien. </li><li> Bereiten Hybridisierungslösung: 2x SSC, 50% Formamid und 10% Dextransulfat (w / v). Lagerung bei 4 ° C. </li></ol></li><li> FISH-Sonden. <ol><li> Bestellen Oligonukleotide (Tabelle 1 für Sequenzen sehen) mit HPLC - Reinigung und Wunsch 5 &#39;fluoreszierende Modifikation (zB Cy3 oder Cy5). Resuspendieren in Tris-EDTA (TE) bei 50 ng / &amp; mgr; l. Hinweis: Schützen Sie Oligos von Lichtexposition zu allen Zeiten. </li></ol></li></ol></li></ol> 2. Sammlung der Late-Stage Drosophila Oozyten <ol><li> Wie <em&gt; Drosophila Oozyten mit Membranen intakt haften auf Kunststoff und Glas, Pre-Beschichtung der Innenseite einer 5 - ml - Röhrchen und einer Pasteur - Pipette pro Eizelle prep mit PTB. </li><li> Anesthetize alle ~ 100 bis 300 yeasted Fliegen mit Kohlendioxid und fügen Sie in einen Mischer enthält ~ 100 ml 1x Robb Buffer. Pulse dreimal (~ 1 Sekunde pro Stück). Halten Sie Eizellen in Robb für &lt;20 min Aktivierung zu vermeiden. HINWEIS: Alternativ Eizellen von Frauen von Hand präpariert werden können. Der Vorteil dieses Verfahrens ist, dass es weniger Weibchen erfordert. Allerdings ist darauf zu Exposition gegenüber Kohlendioxid nur wenige Minuten in Anspruch zu nehmen Artefakte mit Hypoxie assoziiert 7 zu vermeiden. </li><li> Filter durch große Netz (~ 1500 &amp; mgr; m) in 250-ml-Becher große Körperteile zu entfernen. Wenn viele intakte abdomens bleiben auf Mesh, Nachschliff Material zusätzliche Robb Puffer und Filter wieder. Lassen Sie siedeln ~ 2 min, dann obere Schicht absaugen, da viele der großen Körperteile wie möglich zu entfernen. </li><li> Filter durch kleine Maschen (~ 300 &amp; mgr; m) in einen 250-ml-Becher. Spülen restlichen Eizellen aus den ersten Becher mit zusätzlichen Robb und beschichteten Pasteur-Pipette. Lassen Sie settle ~ 3 min; Eizellen werden absetzen. Aspirieren aber ~ 10 ml ab. </li><li> Gießen Sie so viel von den 10 ml wie in beschichteten 5-ml-Röhrchen passen. Lassen Sie, Entfernen Flüssigkeit, und wiederholen Sie mit Rest. Spülen restlichen Eizellen aus Becherglas mit zusätzlichen Robb und beschichteten Pasteur-Pipette. Lassen Sie siedeln in 5-ml-Tube für ~ 3-5 min. </li></ol> 3. Behandlungen mit Arzneimitteln (Optional) <ol><li> Mantel ein zweites 5-ml-Röhrchen mit PTB für jede Eizelle prep. Hinzufügen geeigneten Lösungsmittel (Kontrolle) oder Arzneimittel zu 1 ml Robbs jeweils für jede Oocyte prep (Tabelle 2). </li><li> Split Eizellen in die zweite beschichtete 5-ml-Tube. Lassen Sie, Entfernen Flüssigkeit und 1 ml Robb plus Lösungsmittel in ein Röhrchen und 1 ml Robb plus Medikament in die zweite Röhre. Nutation für entsprechende Menge an Zeit für die Arzneimittelbehandlung (Tabelle 2). Let begleichen. </li></ol> 4. Fixation <ol><li> Absaugen alle Flüssigkeit und fügen sofort 1 ml Fix. </li><li> Option 1: Formaldehyd / Heptan Fixation (Fixation Buffer plus 5% Formaldehyd). <ol><li> Fix für 2,5 min auf einer Nutator. 1 ml Heptan und Wirbel 1 min. Lassen Sie settle ~ 1 min. </li><li> Entfernen Sie alle Flüssigkeit, und fügen Sie dann 1 ml 1x PBS. Vortex 30 Sekunden. Lassen Sie settle ~ 1 min. </li><li> Entfernen Sie alle Flüssigkeit, und dann füllen Rohr mit 1x PBS. HINWEIS: Die Oocyten können sofort oder gehalten auf der Nutator mehrere Stunden bei Raumtemperatur verwendet werden. </li></ol></li><li> Option 2: Formaldehyd / Cacodylat Fixierung (Fix Mix plus 8% Formaldehyd und 100 mM Cacodylat). <ol><li> Fix für 6 min auf einem Nutator. Lassen Sie 2 Minuten absetzen. Entfernen Sie Flüssigkeit, und dann füllen Rohr mit 1x PBS. HINWEIS: Die Oocyten können sofort oder gehalten auf der Nutator mehrere Stunden bei Raumtemperatur verwendet werden. </li></ol></li></ol> 5. Die Membranen Entfernen ( &quot;Rolling&quot;) <ol><li> Mit beschichteten Pasteur-Pipette, fügen ~ 500 bis 1.000 Eizellen zur mattiert (sandgestrahlte) Teil eines Glasträger. Entfernen Sie alle Körperteile und Fremdmaterial mit einer Pinzette. Lassen Sie sich nicht Eizellen austrocknen; 1x PBS bei Bedarf hinzufügen. </li><li> Legen Sie ein Deckglas auf der Eizellen und sanft &quot;rollen&quot; Eizellen, bis alle Membranen werden entfernt (Ziehen der Kante des Deckglases über die Eizellen am besten funktioniert.) Überprüfen Sie regelmäßig die Fortschritte unter dem Mikroskop, das Hinzufügen von mehr 1x PBS wie nötig. Achten Sie darauf, wie zu viel Druck wird die Eizellen zerstören. HINWEIS: Nur für Immunofluoreszenz mit 6. Schritt weiter für Fische (mit oder ohne Immun), fahren Sie mit Schritt 7. </li></ol> 6. Antikörper - Färbung von Drosophila Oozyten <ol><li> Extraktion und Blockierung <ol><li> Spülen gewalzten Oozyten in ein 15 ml konisches Röhrchen mit ~ 15 ml PBS / 1% Triton X-100. Nutation Oozyten in dieser Lösung nicht weniger als 1,5 Stunden und nicht mehr als 2 Stunden.Dieser Schritt ermöglicht Antikörper Penetration. </li><li> Lassen Sie Eizellen absetzen ~ 2 min. Entfernen Flüssigkeit zusammen mit so vielen Membranen wie möglich, dann fügen PBS / 0,05% Triton X-100. HINWEIS: Die Membranen werden siedeln langsamer als gerollt Eizellen. </li><li> Lassen Eizellen absetzen ~ 2 min, dann entfernen Sie alle, aber ~ 1 ml Flüssigkeit. Transfer-Oozyten abgestuft bis 1,5-ml-Röhrchen und restliche Flüssigkeit zu entfernen. 1 ml der PTB für die Sperrung und Nutation für 1 Stunde. </li></ol></li><li> Antikörper-Färbung <ol><li> Pre-Absorption von sekundären Antikörpern gegen Embryonen. HINWEIS: Dieser Schritt eliminiert Hintergrundfärbung von nicht-spezifischen Antikörper - Wechselwirkungen mit Drosophila - Proteine. <ol><li> Sammeln und beheben Drosophila - Embryos (~ 25 &amp; mgr; l) pro typisches Verfahren 8. Speichern in Methanol bei -20 ° C. </li><li> Entfernen von Methanol aus Embryonen, fügen Sie 800 ul Methanol plus 200 ul 1x PBS und Nutation für 15 min. Entfernen Sie 500 ul Überstand und ersetzen mit 500 ul 1x PBS. Dann nutiert für 15 min. Wiederholen Sie zweimal (für insgesamt ca. 1 Stunde von Waschungen) und dann in der PTB zu beenden. HINWEIS: Die Embryonen verwendet werden können, sofort oder auf der Nutator für mehrere Stunden bei Raumtemperatur gehalten. </li><li> Entfernen Sie Flüssigkeit füllen mit PTB bis 200 &amp; mgr; l pro Eizelle prep, und fügen Sie fluoreszenzmarkierten sekundären Antikörper in geeigneten Verdünnungen (wenn man bedenkt, dass das endgültige Volumen von 300 &amp; mgr; l wird, vgl. Schritt 7.4) Nutation bei 4 ° C über Nacht (bevorzugt) oder bei Raumtemperatur für 3 bis 4 Stunden. HINWEIS: Die vorge absorbierte Antikörper werden in Schritt 6.2.4 verwendet werden. HINWEIS: Bewahren Sie Proben im Dunkeln, so viel wie möglich, sobald fluoreszierende Antikörper hinzugefügt wurden. </li></ol></li><li> Entfernen von Flüssigkeit aus Oozyten, füllen mit PTB bis 300 &amp; mgr; l, und fügen Sie primären Antikörper bei geeigneten Verdünnungen. Nutiert bei 4 ° C über Nacht (bevorzugt) oder bei Raumtemperatur für 3 bis 4 Stunden. </li><li> Wasch Oozyten viermal für jeweils 15 min mit 1 ml PTB. </li><li> EntfernenFlüssigkeit aus Oozyten wurden 200 ul Überstand von Embryonen (vorabsorbiert Sekundärantikörper) hinzuzufügen. Dann füllen sich mit PTB bis 300 &amp; mgr; l. über Nacht nutiert für 3 bis 4 h (bevorzugt) oder bei 4 ° C bei Raumtemperatur. </li><li> Waschen Sie Eizellen einmal für 15 Minuten mit 1 ml PTB, entfernen Flüssigkeit. Dann fügen 0,5 ul Hoechst 33342 und 500 ul PTB und Nutation für 7 min. </li><li> Waschen Oozyten zweimal für jeweils 15 min mit 1 ml PTB. HINWEIS: Die Oocyten können unmittelbar an einem Schlitten montiert werden, oder kann für die Bildgebung in PTB bei 4 ° C bis bereit gespeichert werden. </li></ol></li></ol> 7. FISH (weiter von Schritt 5 oben) <ol><li> Spülen Sie rollte Eizellen in eine 15 ml konischen Röhrchen mit ~ 15 ml 2x SSCT-. Lassen Sie Eizellen absetzen ~ 2 min. Entfernen Sie alle ~ 0,5 ml Flüssigkeit zusammen mit so vielen Membranen wie möglich. HINWEIS: Die Membranen werden siedeln langsamer als gerollt Eizellen. </li><li> Transfer-Oozyten bis 0,5-ml-Röhrchen und entfernen restliche Flüssigkeit. Nacheinander hinzufügen und 500 ul entfernen20%, 40% und 50% Formamid-Lösungen, Taumel für 10 min in jeder Lösung. HINWEIS: Die Oozyten werden mit einem höheren Prozentsatz an Formamid langsamer absetzen. </li><li> Entfernen Sie Flüssigkeit, fügen Sie dann 500 ul 50% Formamid-Lösung und Nutation bei 37 ° C für 1 bis 5 Stunden. HINWEIS: Längere Inkubationen führen zu einer besseren Sonde Penetration. </li><li> Entfernen Flüssigkeit und hinterlässt nicht mehr als ~ 100 &amp; mgr; l - Oozyten und fügen 36 ul Hybridisierungslösung plus 2 &amp; mgr; l Sonde (50 ng / ul) und 2 ul Wasser oder 2 &amp; mgr; l einer 2 nd Sonde (Tabelle 1). </li><li> bei 91 ° C inkubieren 3 min (FISH only) oder 80 ° C für 20 min (FISH und Immunofluoreszenz), gefolgt von einer Inkubation über Nacht in einem Wasserbad 37 ° C. Hinweis: diese Schritte in einem Thermocycler durchgeführt werden. </li><li> Entfernen Sie nicht flüssig. Hinzufügen 500 ul 50% Formamid-Lösung und nutiert bei 37 ° C für 1 Stunde. </li><li> Lassen Sie, Entfernen Flüssigkeit, 500 &amp; mgr; l 50% Formamid-Lösung eind nutiert bei 37 ° C für 1 Stunde. </li><li> Lassen Sie, Entfernen Flüssigkeit, 500 &amp; mgr; l 20% Formamid-Lösung und Nutation bei Raumtemperatur für 10 min. </li><li> Führen Sie drei schnelle Waschungen in 500 ul 2x SSCT- (lassen, Entfernen dann Flüssigkeit hinzufügen, kehren mehrmals wiederholen.) Lassen Sie, Entfernen Flüssigkeit, dann fügen Sie 500 ul PTB und Nutation für 4 Stunden. </li></ol> 8. Antikörper-Färbung nach FISH <ol><li> Entfernen von Flüssigkeit aus Oozyten, füllen mit PTB bis 300 &amp; mgr; l, und dann werden 10 &amp; mgr; l anti-α-Tubulin-Antikörper an FITC konjugiert. Alternativ können auch andere Antikörper verwendet werden, nach oben Protokoll. über Nacht bei Raumtemperatur Nutation. </li><li> Waschen Sie Eizellen einmal für 15 Minuten mit 500 ul PTB, entfernen Flüssigkeit, dann fügen 0,5 ul Hoechst 33342 und 500 ul PTB und Nutation für 7 min. </li><li> Waschen Oozyten zweimal für 15 min mit jeweils 500 ul PTB. HINWEIS: Die Oocyten können unmittelbar an einem Schlitten montiert werden, oder sie können für in PTB bei 4 ° C bis bereit gespeichert werdenBildgebung. Halten Sie Eizellen im Dunkeln, so viel wie möglich, sobald fluoreszierende Antikörper hinzugefügt wurden. </li></ol><table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always"><tbody><tr><td fo:width="1in"> Wiederholen Sie Namen </td><td fo:width="1in"> Chromosom </td><td> Oligo Sequenz * </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 - Sequenz von Eric Joyce, persönliche Kommunikation, andere Sequenzen von Sullivan et al. 8 </td&gt; </tr></tbody></table> Tabelle 1: FISH - Sonden für Drosophila zentromerischen wiederholt. <table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always"><tbody><tr><td> Droge </td><td> Lösungsmittel </td><td> Auf Konzentration </td><td> Die Endkonzentration </td><td> Behandlungszeit </td><td> Bewirken </td></tr><tr><td> Colchicin </td><td> Ethanol </td><td> 125 mM </td><td> 150 &amp; mgr; M </td><td> 10 min oder 30 min </td><td> destabilisieren nicht kinetochore (10 min) 9 oder alle (30 min) Mikrotubuli </td></tr><tr><td> Paclitaxel </td><td> DMSO </td><td> 10 mM </td><td> 10 &amp; mgr; M </td><td> 10 Minuten </td><td> stabilisieren microtubules </td></tr><tr><td> Binucleine 2 </td><td> DMSO </td><td> 25 mM </td><td> 25 &amp; mgr; M </td><td> 20 Minuten </td><td> hemmen Aurora B - Kinase 10 </td></tr></tbody></table> Tabelle 2: Die medikamentöse Behandlung.

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The authors have nothing to disclose.

Staging Drosophila Oozyten Obwohl ein längliches karyosome oft in der Prometaphase Oocyten gesehen wird, karyosome Form unter Verwendung von Metaphase Prometaphase Oozyten unterscheiden kann problematisch sein. Während Prometaphase beginnt die karyosome als eine runde Form, längt, und zieht dann in eine runde Form wie die Eizelle die Verhaftung Metaphase nähert. Dies bedeutet, dass viele der Prometaphase Oozyten keine längliche karyosome haben. Darüber hinaus,...

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.

Die Methoden , die wir hier beschrieben haben , in der Sammlung von Spätstadium der Drosophila - Oozyten , die drei Stadien der Meiose (Abbildung 1) wird zur Folge haben . Oozyten in Prophase werden durch das Vorhandensein der Kernhülle aus, die durch das Fehlen von Tubulin - Signal in der Region um die karyosome (1A) sichtbar ist. Prometaphase ist die Zeit, nach der Kernhülle Zusammenbruch während der die Spindel montiert. Während Prometap...

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 und Metaphase der Meiose I in fester<em&gt; Drosophila</em&gt; Oozyten

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

Unit 2

Meiosis

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9.0K Aufrufe.  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.

Serie: 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.

Mit Maus Eizellen menschliche Gen-Funktion während der Meiose zu beurteilen ich

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

Forschung

Genetik

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.

Fluoreszenz In Situ Hybridisierung (FISH) verwenden, um den Zustand der Arm Zusammenhalt im Prometaphase und Metaphase überwachen ich Drosophila Eizellen

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

Entwicklungsbiologie

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.

Mikroinjektion Techniken zur Untersuchung Mitose in der 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 ...

Biologie

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.

Kernmigration in der Drosophila-Eizelle

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.

Konfokale und hochauflösende Bildgebung des polarisierten intrazellulären Transports und der Sekretion von Basalmembranproteinen während der Drosophila-Oogenese

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.

Maus Oocyte Mikroinjektion, Reifung und Ploidie Beurteilung

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.

Studieren mitotischen Checkpoint durch Veranschaulichung Dynamische Kinetochor Protein Verhalten und Chromosome Motion in Lebens-<em&gt; Drosophila</em&gt; Syncytial Embryonen

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.

Live-Imaging von GFP-markierten Proteine ​​in<em&gt; Drosophila</em&gt; Oozyten

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;

Die zytologische Analyse der Spermatogenese: Live-und Fest Zubereitungen aus<em&gt; Drosophila</em&gt; Hoden

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.

Präparation von fixierten Drosophila-Eizellen für die Immunfärbung: eine Hochdurchsatzmethode zur Fixierung und Entfernung der äußeren Membran

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 und Metaphase der Meiose I in fester

Zusammenfassung

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Zusammenfassung

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