Drosophila Oocytes: Preparations for Live Imaging

doi:10.21203/rs.3.pex-2183/v1

Method Article

Drosophila Oocytes: Preparations for Live Imaging

https://doi.org/10.21203/rs.3.pex-2183/v1

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This protocol describes methods for preparing Drosophila oocytes for live imaging by fluorescence microscopy. Live imaging is preferable to fixation for localization of specific proteins to oocytes, given that it can be performed without time-consuming and exacting procedures required to remove the oocyte outer chorion and vitelline membrane. Live imaging is also well suited for analyzing dynamic changes in oocytes that occur rapidly on time scales of seconds. Preparation of oocytes requires only ~12-15 minutes, not including imaging time, which can require 30-60 minutes depending on the experimental objectives and design. Methods are also presented for activating oocytes to induce re-entry into meiosis and progression through the meiotic divisions.

Biological techniques

Oocytes

Drosophila

Live Imaging

Fluorescent Proteins

MI Spindle

Drosophila oocytes develop in clusters of egg chambers that make up the ovary with each oocyte surrounded by a chorion and vitelline membrane, which are secreted by follicle cells that form the outer epithelial layer of the oocyte. The oocytes are easy to collect by dissection from well fed females, but have been studied less extensively than embryos because of the steps involved in preparing fixed oocytes for imaging, which requires removing the chorion and vitelline membrane, either or both of which may be fused to each other or the oocyte cytoplasm after fixation. Moreover, studies of the oocyte meiotic spindle are impeded by the fact that oocytes contain a single spindle in a large volume of cytoplasm, which makes the spindle difficult to locate after fixation and removal of the chorion.

By contrast, the spindle of live oocytes can be located by its position on the anterior dorsal side of the oocyte, near the base of the dorsal appendages. The spindle is present just below the cortical surface of the oocyte, where it can be imaged through the outer layers using a brightly labelled spindle marker by mounting the oocytes so the dorsal side is up for imaging. Mature oocytes are arrested in metaphase of the meiosis I division, but can be activated to re-enter meiosis and progress through the meiotic divisions. This protocol presents procedures for preparing oocytes to acquire live images of the spindle and for activating oocytes to follow spindle dynamics during division. The methods involve dissecting oocytes in buffers adjusted for ionic strength to control for osmotic activation and mounting oocytes for imaging on slides with an O₂-permeable membrane [1] against hypoxia.

Drosophila Stocks

Drosophila stocks that express proteins of interest fused to GFP or other bright fluorophores can be used in this protocol. Lines that express spindle markers for localization of oocyte proteins or for following dynamic changes in oocytes include kinesin-14 Ncd-GFP [2, 3], γTub-GFP [4] and EB1-GFP [5]. These and other lines are available from the Bloomington Drosophila Stock Center. Flies that express new proteins under study are easily produced using standard recombinant DNA methods to construct plasmids and widely available transformation or CRISPR gene editing services.

Dissecting Buffer and Supplies

Robb’s Drosophila Ringer’s solutions A and B [6] are made up separately as 2x or 20x solutions and sterilized, then mixed together to make AB. AB50 contains an additional 50 mM NaCl, whereas AB^– is made without sucrose or glucose.

NB: Schneider’s Drosophila Medium (Gibco #21720024 or Millipore Sigma #S0146) supplemented with 10-15% heat-inactivated FCS can also be used as a nonactivating oocyte dissection and mounting medium, although the background in fluorescence images is slightly higher with Schneider’s medium than Robb’s Ringer’s solution.

Fine dissecting forceps (e.g., Dumont Dumostar No. 55), two pairs

Two-well shallow dissecting slide

Pipettor P20 with tips (e.g., Rainin XLS+ pipettor; Sorenson OneTouch 1-20 μL tips)

Glass microliter pipette (Clay Adams Micropet, 50 μL) or similar glass micropipette

Pipette bulb made from microliter pipette adaptor (e.g., Clay Adams Micropet adaptor) attached to a short length of rubber tubing tied off at the end

Live Imaging Mounting Materials

Metal or 3D printed slide [1] for live imaging (7.6 W x 3.9 D x 0.6 H cm)

Oxygen-permeable Teflon membranes (YSI, Inc.,Yellow Springs, OH, Standard Membrane Kit #5793)

O-rings (e.g., Danco #12 or #67; Standard size 016 or 017)

Silicone vacuum grease (Beckman Instruments, Inc., cat #335148) backloaded into hypodermic syringe (1 mL) fit with cut-off 18 G needle (2 in)

Toothpick, wooden, for spreading the vacuum grease

Stainless steel forceps for picking up clean coverslips, 1 pair

Stainless steel forceps for rolling oocytes, 1 pair

Round coverslips, EtOH-cleaned (e.g., Marienfeld GG-18-1.5H Precision Coverslips)

Stereomicroscope with incident illumination (light from above specimen)

Fluorescence microscope imaging system for acquiring high-resolution images, e.g., Zeiss LSM 780 or 880 laser-scanning confocal microscope

Nonactivated oocytes

1. Collect newly eclosed females and culture for 2-5 da on standard Drosophila cornmeal-glucose-yeast-agar food supplemented with a few granules of live yeast.

2. Assemble a slide for live imaging by cutting a Teflon membrane circle to fit over the slide opening with sufficient margin to allow the membrane to be secured by an O-ring. The membrane can be stretched slightly and smoothed by running a gloved finger around the edge of the opening. Make three short (~0.8 cm) lines of vacuum grease around the opening using a hypodermic syringe with attached cut-off 18 G needle. Slightly spread the vacuum grease lines by cutting through each line with a wooden toothpick.

3. Stun females by placing them in an empty glass vial on ice for 1-2 min. Transfer a female to 10-20 μL of AB50 in a well of a two-well shallow dissecting slide. Detach the abdomen and release the ovaries into the AB50 using fine dissecting forceps. Transfer the ovaries into 10-20 μL of AB50 in the second well and separate the oocytes by pulling gently on the dorsal appendages or by separating the fibrous connections between the follicles, taking care not to apply pressure on the oocytes, which may cause them to leak cytoplasm or burst.

4. Remove excess or unwanted oocytes from the second well, then push 5-10 oocytes for imaging into a cluster using the sides of the forceps tips. Pipette up the oocytes in ~12 μL buffer using a 50-μL glass microliter pipette with pipette bulb made from a pipette adapter and rubber tubing and transfer to the Teflon membrane on the slide for live imaging. A glass microliter pipette is used to transfer the oocytes because they stick to Pipetman yellow tips and can be lost.

5. Using the sides of the tips of a pair of forceps, arrange the oocytes in the drop of solution on the slide so they are spread out and do not touch each other, avoiding making indentations in the Teflon membrane. Five to ten oocytes can be arranged into two rows side-by-side for ease of identification during imaging. Carefully place a clean coverslip over the oocytes so that it barely makes contact with the oocytes.

6. Just before imaging, press down gently on the edges of the coverslip so the oocytes are barely immobilized. Sketch a map of the oocytes, noting their position on the slide with respect to one another, whether the oocyte is stage 13 or 14, and other identifying features (e.g., broken or curved dorsal appendages, attached immature follicles, attached fibers). Immature stage 13 oocytes are readily identified by their fused dorsal appendages associated with nurse cell remnants, whereas mature stage 14 oocytes display closely positioned dorsal appendages without nurse cells (Fig. 1a). The oocytes are usually positioned on their side or ventral side up, neither of which lends itself to imaging the spindle, which is typically located at or near the base of the dorsal appendages. The spindles of stage 13 and 14 oocytes differ in shape and length with stage 13 spindles being longer and with tapered poles, compared to the shorter stage 14 spindles (Fig. 1b).

7. Find oocytes that are dorsal side up and image the spindle in these oocytes first, numbering the oocytes on your map in the order of imaging. Then use the tip of a pair of forceps to roll the remaining oocytes into a dorsal-side-up position by pushing gently to exert force laterally on the edge of the coverslip in the direction in which the oocyte needs to roll. For example, push towards the left on the edge of the coverslip nearest the oocyte if the oocyte is positioned with its dorsal side facing to the right (Fig. 2).

8. Live imaging can be easily performed after orienting the oocytes by placing the slide on an upright compound fluorescence microscope. Slides can also be imaged on inverted Zeiss and Leitz fluorescence microscopes, and possibly other inverted microscopes, by angling the inverted slide in the slide holder and ensuring that it is securely positioned.

Activated Oocytes

Oocytes can be activated to re-enter meiosis and undergo the meiotic divisions by immersion in hypotonic solution, which causes them to swell – this is thought to mimick oocyte activation in vivo during passage through the oviduct [7, 8, 3]. Robb’s AB solution is hypotonic relative to oocytes and causes oocytes to swell and undergo activation. Oocytes can also be activated after imaging in AB50 by pipetting an equivalent volume of AB^– (~12 μL) under the edge of the coverslip. The use of a lightweight pipettor with a tapered 20 μL pipette tip makes the addition of the AB^– solution straightforward to do. The oocyte and spindle swell rapidly after activation and the spindle can be easily located and imaged. Oocytes positioned dorsal-side-up at the time of AB^–addition can be imaged right away. Other oocytes can be identified from prior pre-activation imaging by their stage, position, or other identifying features, and rolled gently into position after activation for imaging of the spindle.

1. The coverslip on the oocytes for live imaging is attached to the slide only by the lines of vacuum grease. If the lines are too short and the slide is imaged with an oil objective, the coverslip may stick to the objective and detach from the slide when the objective is changed. If this occurs, it may be possible to recover the coverslip with the oocytes intact by lifting it off the objective with a pair of forceps, then remount the coverslip on the slide after adding a few μL of additional buffer. However, this situation is best prevented by ensuring that the vacuum grease lines are long enough to hold the coverslip to the slide and the amount of oil used is not excessive, causing the coverslip to adhere to the objective.

2. Rolling oocytes into a dorsal-side-up position requires practice and can be problematic. If an oocyte does not roll but instead slides against the Teflon membrane covering the slide opening, gently touch the coverslip over the posterior end of the oocyte with the forceps tips, then try rolling the oocyte again. This requires only a touch of the forceps tips to the coverslip, which slightly deforms the Teflon membrane near the posterior end of the oocyte, anchoring the end in place so that the oocyte rolls instead of sliding.

If attempts to roll an oocyte cause the coverslip to be pushed so far into the slide opening that the vacuum grease lines supporting the coverslip are carried into the imaging area of the slide, push the coverslip in the opposite direction to move the oocyte back into the imaging area, then touch the coverslip near the posterior end of the oocyte and try rolling the oocyte again. In some cases, the oocyte will roll completely over into a dorsal-side-up position.

If the coverslip is pushed down onto the oocytes too far, the oocytes will be immobilized and will not roll. Lifting the coverslip edge slightly using the tip of a pair of forceps may allow the oocytes to be rolled into position. If not, it will be necessary to make another preparation.

3. Oocytes that are activated too quickly using strongly hypotonic solution will swell rapidly and burst. This occurs if oocytes are dissected and mounted in AB^– solution, which does not contain sucrose or glucose. The use of a less hypotonic solution will result in less rapid swelling and allow imaging of activated oocytes.

Oocytes of lines under study should be tested for activation with AB and nonactivation with AB50, prior to performing extensive experiments. Robb’s AB^– will induce oocyte activation and AB solutions with higher concentration of NaCl (e.g., AB75 containing an additional 75 mM NaCl) can be used to suppress activation.

Females are typically cultured for 3-5 days after eclosion prior to dissection of ovaries. Optimization of the culture time should be performed to ensure that sufficient numbers of oocytes in the stages of interest can be obtained.

Dissection of oocytes from a single female for slide preparation requires ~12-15 min. If the buffer used to prepare the oocytes does not activate the oocytes or does not activate them rapidly, slides can be held for 20-30 min prior to imaging. For buffers that activate oocytes (e.g., AB or AB^–), imaging should be performed as soon as possible after adding a coverslip to the oocytes on the slide. Oocytes can be prepared in nonactivating solution (e.g., AB50) and imaged, then activated by adding an approximately equal volume of AB^–under the coverslip, as described above.

Performed with care, this protocol should result in high-resolution images of oocyte spindles in different stages of assembly and division. Non-spindle proteins can be easily localized and imaged at high resolution.

1. Ly, P. T., Thompson, J. D., & Endow, S. A. Fixation and staining of Drosophila L1 larval brains for immunofluorescence microscopy and preparation for live imaging. Protocol Exchange (2020). https://protocolexchange.researchsquare.com/article/pex-825/v1.

2. Endow, S. A. & Komma, D. J. Centrosome and spindle function of the Drosophila Ncd microtubule motor visualized in live embryos using Ncd-GFP fusion proteins. J Cell Sci. 109, 2429-2442 (1996).

3. Endow, S. A. & Komma, D. J. Spindle dynamics during meiosis in Drosophila oocytes. J Cell Biol. 137, 1321-1336 (1997).

4. Hallen, M. A., Ho, J., Yankel, C. D., & Endow, S. A. Fluorescence recovery kinetic analysis of γ-tubulin binding to the mitotic spindle. Biophys J. 95, 3048-3058 (2008).

5. Liang, Z.-Y., Hallen, M. A., & Endow, S. A. Mature Drosophila meiosis I spindles comprise microtubules of mixed polarity. Curr Biol. 19, 163-168 (2009).

6. Robb, J. A. Maintenance of imaginal discs of Drosophila melanogaster in chemically defined media. J Cell Biol. 41, 876-885 (1969).

7. Mahowald, A. P., Goralski, T. J., & Caulton, J. H. In vitro activation of Drosophila eggs. Dev Biol. 98, 437-445 (1983).

8. Page, A. W. & Orr-Weaver, T. L. Activation of the meiotic divisions in Drosophila oocytes. Dev Biol. 183, 195-207 (1997).

9. Sköld, H. N., Komma, D. J., & Endow, S. A. Assembly pathway of the anastral Drosophila oocyte meiosis I spindle. J Cell Sci. 118, 1745-1755 (2005).

10. Zou, J., Hallen, M. A., Yankel, C. D., & Endow, S. A. A microtubule-destabilizing kinesin motor regulates spindle length and anchoring in oocytes. J Cell Biol. 180, 459-466 (2008).

These methods were developed for studies that led to the publication of reports on the assembly and dynamics of the Drosophila oocyte spindle [9, 3, 5] and the localization of proteins to the spindle [4, 10]. The methods have been used more recently in as yet unpublished studies to develop a sensor to measure forces across a spindle motor protein. H. Sköld, J. Zou, Z-Y Liang and J. Ho contributed to the development of these methods. Microscopes in the Duke University Light Microscopy Core Facility have been used in these studies with training and assistance in imaging provided by B. Carlson. Funding is from a current NSF Award # CMMI-1660924 to SAE; previous studies were supported by NIH R01 GM046225 and March of Dimes Foundation #1-FY07-443 Awards to SAE. Any opinions, findings, conclusions, or recommendations in this paper are those of the author and do not necessarily reflect the views of the National Science Foundation, the National Institutes of Health, or the March of Dimes Foundation.

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Drosophila Oocytes: Preparations for Live Imaging

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