We thank J. Kamenz, Y. Chen, and W. Y. C. Huang for comments on the manuscript. This work was supported by grants from the National Institutes of Health (R01 GM110564, P50 GM107615, and R35 GM131792) awarded to James E. Ferrell, Jr.

All methods described here have been approved by the Institutional Animal Care and Use Committee (IACUC) of Stanford University.
1. Preparation of slides and coverslips
<ol>
	<li>Apply a layer of fluorinated ethylene propylene (FEP) adhesive tape to a glass slide with a roller applicator. Cut off excessive tape over the edges with a clean razor blade. Prepare FEP tape-coated coverslips in the same way (Figure 1A).</li>
	<li>Apply a double-sided sticky imaging spacer to the FEP tape-coated side of the slide. Leave the protective liner on the top unpeeled (Figure 1A). 
	&#8203;NOTE: The slides and coverslips should be prepared before the experiment. They can be used immediately, or stored in boxes to prevent dust accumulation on surfaces. The well in the imaging spacer is 120 &#181;m deep and has a diameter of 9 mm.</li>
</ol>
2. Preparation and live imaging of interphase-arrested egg extracts
NOTE: The following protocol is adapted from Deming and Kornbluth19, Murray20, and Smythe and Newport25 with modifications. All steps should be performed at room temperature unless otherwise noted.
<ol>
	<li>Three to ten days before egg collection, inject mature female Xenopus laevis frogs subcutaneously into the dorsal lymph sac with 100 IU of pregnant mare serum gonadotropin (PMSG).</li>
	<li>Sixteen to eighteen hours prior to planned egg collection, inject the frogs from step 2.1 with 500 IU of human chorionic gonadotropin (hCG). Leave frogs at 18 &#176;C in egg laying buffer (100 mM NaCl, 2 mM KCl, 1 mM MgSO4&#183;7H2O, 2.5 mM CaCl2&#183;2H2O, 0.5 mM HEPES, 0.1 mM EDTA, prepare as a 20x stock solution at pH 7.4 and dilute with clean frog tank water to 1x before use) until egg collection.</li>
	<li>On the day of the experiment, collect eggs in a large glass Petri dish and assess egg quality. Discard any eggs that look like white puffy balls or appear in a string (Figure 1B). Examine the eggs under a stereoscope, keep the eggs with normal appearance (Figure 1C), and discard those with irregular or mottled pigment (Figure 1D). 
	NOTE: This protocol works with eggs collected from a single frog, which typically lays 25 mL of eggs by 16 hours after hCG injection. Usually, a total of 3 to 6 frogs are induced by hCG, and the frog with the highest egg quality is chosen for the extract preparation experiment.</li>
	<li>Transfer eggs to a 400 mL glass beaker, and remove as much egg laying buffer as possible by decanting.</li>
	<li>Incubate the eggs in 100 mL of freshly prepared dejellying solution (2% w/v L-cysteine in water, adjust to pH 8.0 with NaOH) and gently swirl them periodically. After about 3 minutes, pour off the solution, and add 100 mL of fresh dejellying solution. Continue the incubation until the eggs are tightly packed (no space between eggs), but avoid leaving eggs in the dejellying solution for more than a total of 5 minutes.</li>
	<li>Remove as much of the dejellying solution as possible by decanting, and wash the eggs in 0.25x MMR buffer (25 mM NaCl, 0.5 mM KCl, 0.25 mM MgCl2, 0.5 mM CaCl2, 0.025 mM EDTA, 1.25 mM HEPES, prepared as a 10x stock solution, adjusted to pH 7.8 with NaOH, and diluted in Milli-Q water before use) by adding the buffer, swirling the eggs, and then pouring off the buffer. Repeat a few times until a total of 1 L of the buffer is used for the washes.</li>
	<li>Wash the eggs a few times with a total of 400 mL of egg lysis buffer (250 mM sucrose, 10 mM HEPES, 50 mM KCl, 2.5 mM MgCl2, 1 mM DTT, made fresh and adjusted to pH 7.7 with KOH). Remove eggs with abnormal appearance using a Pasteur pipette between the washes. 
	NOTE: Eggs with abnormal appearance refer to those that look like white puffy balls (Figure 1B), have mottled pigmentation (Figure 1D), are deteriorating with a growing white region (Figure 1E), or show darkened and contracted pigmented area in the animal hemisphere (Figure 1F).</li>
	<li>Using a transfer pipette with its tip cut wide open, transfer the eggs to a 17 mL round-bottom centrifuge tube containing 1 mL of egg lysis buffer. Spin the tube in a clinical centrifuge at 400 x g for 15 seconds to pack the eggs.</li>
	<li>Remove as much of the egg lysis buffer as possible from the top of packed eggs using a Pasteur pipette. 
	NOTE: It is important to remove as much buffer from the packed eggs as possible, in order to minimize the dilution of the egg extract. Sometimes it is necessary to remove some loose eggs along with the residual buffer to accomplish this.</li>
	<li>Determine the approximate volume of the packed eggs, and then add 5 &#181;g/mL aprotinin, 5 &#181;g/mL leupeptin, 5 &#181;g/mL cytochalasin B, and 50 &#181;g/mL cycloheximide directly on top of the packed eggs. 
	NOTE: Aprotinin and leupeptin are protease inhibitors. Cytochalasin B inhibits actin polymerization, preventing the extract from contracting and gelling26. Cycloheximide inhibits protein synthesis, thereby keeping the extract in the interphase of the cell cycle.</li>
	<li>Crush the eggs by centrifuging the tube at 12,000 x g, 4 &#176;C, for 15 minutes, in a swinging bucket rotor. 
	NOTE: At the end of the centrifugation, the eggs should have ruptured and the lysate separated into three main layers: a yellow lipid layer on top, the cytoplasmic extract (also called crude extract) in the middle, and a dark dense layer containing the pigment granules at the bottom (Figure 1G).</li>
	<li>Attach an 18-gauge needle to a syringe. With the needle tip bevel facing up, puncture the tube from the side at the bottom of the cytoplasmic layer, and recover the extract by drawing slowly. 
	NOTE: Draw the cytoplasmic extract slowly to avoid the inclusion of contaminating content from the yellow lipid layer.</li>
	<li>Transfer the recovered cytoplasmic extract to a new microcentrifuge tube and hold it on ice. Use the extract within 1 hour.</li>
	<li>When ready to image, supplement the extract with desired reagents and fluorescence imaging probes. 
	NOTE: Fluorescence imaging probes label specific cytoplasmic structures so that they can be visualized by a fluorescence microscope.</li>
	<li>Remove the top protective liner from the imaging spacer on the slide prepared in step 1.2, and deposit approximately 7 &#181;L of extract at the center of the well. Immediately apply the FEP tape-coated coverslip with the FEP side facing the extract to seal the well. Quickly proceed to imaging (Figure 1H,I).</li>
	<li>Set the slide on an inverted or upright microscope with a motorized stage and a digital camera. Image the extracts at desired spatial positions and time intervals in both bright-field and fluorescence channels. 
	NOTE: Typically, a 5x objective is used for imaging. The motorized stage enables automated image acquisition at multiple defined spatial positions. Bright-field microscopy visualizes cytoplasmic structures with different degrees of transparency. Fluorescence microscopy visualizes the cytoplasmic structures specifically labeled by the fluorescence imaging probes added in step 2.14. The camera records time-lapse images of these structures, thereby capturing the dynamics of cytoplasmic organization.</li>
</ol>

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The authors have nothing to disclose.

Xenopus laevis egg extracts have emerged as a powerful model system for imaging-based&#160;studies of various subcellular structures10,14,15,16,17,18,21,31,32,33,34,35,36 and cytoplasmic organization on a whole cell scale9. Here we have described a live imaging method suitable for visualizing dynamic cytoplasmic organization in 2D. The effectiveness of the method is demonstrated by representative results.
Several steps are critical to the success of the method. Egg quality is important for extracts, so steps 2.3 and 2.7 are critical. In our experience, the animal hemisphere of high-quality eggs has uniform pigmentation, a distinct white dot at the center (indicative of germinal vesicle breakdown, GVBD), and a clear border with the vegetal hemisphere (Figure 1C). The extracts made from high-quality eggs have better performance under our imaging conditions. If more than 5% of the eggs have mottled pigment (Figure 1D), have darkened and contracted pigmented area (Figure 1F), look like white puffy balls (Figure 1B), appear in a string (Figure 1B), or show signs of deterioration after the washes in steps 2.6 and 2.7 (Figure 1E), then the whole batch of eggs should be discarded. The extracts retain remarkable biological activity because they are essentially undiluted cytoplasm. Therefore, the step that aims to minimize dilution of the extract (step 2.9) is important for the success of the experiment. For imaging, extracts are handled in very small volumes, and will evaporate quickly if in prolonged contact with air. This will negatively affect their activity. Therefore, in step 2.15, after the extract is deposited, it must be sealed with the FEP tape-coated side of the coverslip as soon as possible. Sealing can be visually confirmed by the texture change at the contact site between the adhesive on the spacer and the coverslip. The spacer should be able to create a complete seal if applied properly. However, if an additional seal is desired, mineral oil can be dispensed between the overhang of the coverslip and the glass slide. The oil can form an additional seal around the spacer by capillary action (Figure 3). Passivation of glass surfaces can reduce non-specific adsorption of molecules, and it is important for interphase microtubule asters in extracts21,37. The application of FEP tape over a glass surface described here appears to offer similar benefits, as suggested by the assembly of normal interphase microtubule asters (Figure 2, 6 min). Therefore, step 1.1 is also critical.
We demonstrated the application of an imaging method using interphase-arrested egg extracts following the protocol of Deming and Kornbluth19. By default, the protocol supplements the extracts with the actin polymerization inhibitor cytochalasin B to prevent gelation-contraction in the extracts after extended incubation at room temperature26. To allow actin polymerization and observe actin dynamics, one can leave out the cytochalasin B in step 2.10 of the protocol9. A modification we have made to the Deming and Kornbluth protocol is that we do not supplement extracts with an energy mix to regenerate ATP19. This is because in our hands, in extracts supplemented with this ATP-regenerating mix19 and sperm nuclei, microtubules occasionally form a crosslinked lattice that interferes with cytoplasmic pattern formation. Therefore, the protocol does not include the step that adds the energy mix to the extracts19.
The interphase extract protocol relies on crushing eggs in EGTA-free buffer to release them from meiotic arrest (CSF-arrest)37. The extracts subsequently progress into interphase, and are kept in interphase by cycloheximide. There are other established methods for preparing interphase extracts. Some methods first prepare extracts that maintain meiotic arrest by crushing eggs in lysis buffer with EGTA38, and then release the arrest by adding calcium, driving the extract into interphase21,37,39. The extracts can be subsequently held in interphase by addition of protein synthesis inhibitors such as cycloheximide37,39. Other methods parthenogenetically activate eggs with calcium ionophore (A23187) or electric shock to release them from meiotic arrest, before crushing the eggs in the absence of EGTA20,28 (reviewed in Field et al.37). These extracts can enter interphase but typically will not stay there as they are able to undergo multiple cell cycles. Likewise, well-established methods optimized for preparing extracts with intact actin cytoskeleton have been developed10,37,39. The imaging method may be suitable for these types of extracts, but we have not tested it with them.
For the purpose of imaging the internal organization of the cytoplasm, the imaging system presented here are relatively easy to set up, requiring only applying an FEP tape to glass surfaces. It allows assembly of cytoskeletal structures in extracts9 reported in more sophisticated imaging systems where glass surfaces are passivated with poly-L-lysine-g-polyethylene glycol (PLL-g-PEG) treatment or coated with supported lipid bilayers10,21. The method allows the extract layer to form with a defined thickness (determined by the depth of the spacer, which is 120 &#181;m for the system shown in Figure 1A, 1H, 1I, and Figure 2). We can adjust the thickness by stacking additional spacers. We have stacked up to 6 such spacers (720 &#181;m thick) and the compartments formed normally. This flexibility enables future applications such as imaging of the extracts in 3D using confocal or light-sheet microscopy.

The cytoplasm constitutes the main volume of a cell and has a distinct organization. The ingredients of the eukaryotic cytoplasm can self-assemble into a wide range of spatial structures, such as microtubule asters and the Golgi apparatus, which in turn are dynamically arranged and turned over depending on the cell&#39;s identity and physiological state. Understanding the spatial organization of the cytoplasm and its link to cellular functions is thus important for understanding how the cell works. Xenopus laevis egg extracts have traditionally been used for bulk biochemical assays1,2,3,4,5,6,7,8, but recent work establishes them as a powerful live imaging system for mechanistic studies of cytoplasmic structures and their cellular functions9,10,11,12,13,14,15,16,17,18. These undiluted extracts preserve many structures and functions of the cytoplasm, while allowing direct manipulations of cytoplasmic contents not achievable in conventional cell-based models19,20. This makes them ideal for characterizing cytoplasmic phenomena and dissecting their mechanistic underpinnings.
Existing methods for imaging extracts require chemical surface modifications, or fabrication of microfluidic devices. One coverslip-based method requires polyethylene glycol (PEG) passivation of glass coverslips21. A microemulsion-based method requires vapor deposition of trichloro(1H,1H,2H,2H-perfluorooctyl)silane on glass surfaces22,23. Microfluidic-based systems allow precise control of the volume, geometry and composition of extract droplets, but require specialized microfabrication facilities11,12,24.
Here we introduce an alternative method for imaging egg extracts that is easy to implement and utilizes readily available, low-cost materials. This includes preparation of an imaging chamber with a slide and a coverslip coated with fluorinated ethylene propylene (FEP) tape. The chamber can be used for imaging extracts with a variety of microscopy systems, including stereoscopes and upright and inverted microscopes. This method requires no chemical treatment of surfaces while achieving similar optical clarity obtained with existing glass-based methods discussed above. It is designed to image a layer of extracts with a uniform thickness across a 2D field, and can be easily extended to image a 3D volume of extracts. It is well suited for time-lapse imaging of collective cytoplasmic behavior over a large field of view.
We have used interphase-arrested egg extracts to demonstrate our imaging method. The extract preparation follows the protocol of Deming and Kornbluth19. Briefly, eggs naturally arrested in metaphase of meiosis II are crushed by a low speed spin. This spin releases the cytoplasm from meiotic arrest and allows the extract to proceed into interphase. Normally, cytochalasin B is added prior to the crushing spin to inhibit F-actin formation. However, it can be omitted if F-actin is desired. Cycloheximide is also added prior to the crushing spin to prevent the interphase extract from entering the next mitosis. The extracts are subsequently placed in the aforementioned imaging chambers and placed on a microscope. Finally, images are recorded over time at defined intervals by a camera connected to the microscope, producing time-lapse image series that capture the dynamical behavior of the extract in a 2D field.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>17 ml centrifuge tube</td>
			<td>Beckman Coulter</td>
			<td>337986</td>
			<td></td>
		</tr>
		<tr>
			<td>22x22 mm square #1 cover glass</td>
			<td>Corning</td>
			<td>284522</td>
			<td></td>
		</tr>
		<tr>
			<td>Aprotinin</td>
			<td>MilliporeSigma</td>
			<td>10236624001</td>
			<td>Protease inhibitor</td>
		</tr>
		<tr>
			<td>Cycloheximide</td>
			<td>MilliporeSigma</td>
			<td>01810</td>
			<td>Protein synthesis inhibitor</td>
		</tr>
		<tr>
			<td>Cytochalasin B</td>
			<td>MilliporeSigma</td>
			<td>C6762</td>
			<td>Actin polymerization inhibitor</td>
		</tr>
		<tr>
			<td>Female Xenopus laevis frogs</td>
			<td>Nasco</td>
			<td>LM00535MX</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescent HiLyte 488 labeled tubulin protein</td>
			<td>Cytoskeleton, Inc.</td>
			<td>TL488M-A</td>
			<td>For visualizing the microtubule cytoskeleton</td>
		</tr>
		<tr>
			<td>Fluorescent HiLyte 647 labeled tubulin protein</td>
			<td>Cytoskeleton, Inc.</td>
			<td>TL670M-A</td>
			<td>For visualizing the microtubule cytoskeleton</td>
		</tr>
		<tr>
			<td>Fluorinated ethylene propylene (FEP) optically clear tape</td>
			<td>CS Hyde company</td>
			<td>23-FEP-2-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Pasteur pipette</td>
			<td>Fisher Scientific</td>
			<td>13-678-20C</td>
			<td></td>
		</tr>
		<tr>
			<td>Human chorionic gonadotropin (hCG)</td>
			<td>MilliporeSigma</td>
			<td>CG10</td>
			<td></td>
		</tr>
		<tr>
			<td>Imaging spacer</td>
			<td>Electron Microscopy Sciences</td>
			<td>70327-8S</td>
			<td></td>
		</tr>
		<tr>
			<td>Leupeptin</td>
			<td>MilliporeSigma</td>
			<td>11017101001</td>
			<td>Protease inhibitor</td>
		</tr>
		<tr>
			<td>Microscope slides</td>
			<td>Fisher Scientific</td>
			<td>12-518-100B</td>
			<td></td>
		</tr>
		<tr>
			<td>Mineral oil</td>
			<td>MilliporeSigma</td>
			<td>330760</td>
			<td></td>
		</tr>
		<tr>
			<td>MitoTracker Red CMXRos</td>
			<td>Thermo Fisher Scientific</td>
			<td>M7512</td>
			<td>For visualizing mitochondria</td>
		</tr>
		<tr>
			<td>Pregnant mare serum gonadotropin (PMSG)</td>
			<td>BioVendor</td>
			<td>RP1782725000</td>
			<td></td>
		</tr>
		<tr>
			<td>Roller applicator</td>
			<td>Amazon</td>
			<td>B07HMBJSP8</td>
			<td>For applying the FEP tape to the glass slides and coverslips</td>
		</tr>
		<tr>
			<td>Single-edged razor blades</td>
			<td>Fisher Scientific</td>
			<td>12-640</td>
			<td>For removing excessive FEP tape</td>
		</tr>
		<tr>
			<td>Transfer pipette</td>
			<td>Fisher Scientific</td>
			<td>13-711-7M</td>
			<td></td>
		</tr>
	</tbody>
</table>

xenopus laevis egg extract preparation and live imaging methods for visualizing dynamic cytoplasmic organization

Traditionally used for bulk biochemical assays, Xenopus laevis egg extracts have emerged as a powerful imaging-based tool for studying cytoplasmic phenomena, such as cytokinesis, mitotic spindle formation and assembly of the nucleus. Building upon early methods that imaged fixed extracts sampled at sparse time points, recent approaches image live extracts using time-lapse microscopy, revealing more dynamical features with enhanced temporal resolution. These methods usually require sophisticated surface treatments of the imaging vessel. Here we introduce an alternative method for live imaging of egg extracts that require no chemical surface treatment. It is simple to implement and utilizes mass-produced laboratory consumables for imaging. We describe a system that can be used for both wide-field and confocal microscopy. It is designed for imaging extracts in a 2-dimensional (2D) field, but can be easily extended to imaging in 3D. It is well-suited for studying spatial pattern formation within the cytoplasm. With representative data, we demonstrate the typical dynamic organization of microtubules, nuclei and mitochondria in interphase extracts prepared using this method. These image data can provide quantitative information on cytoplasmic dynamics and spatial organization.

Xenopus laevis egg extracts can be used to study the self-organization of the cytoplasm during interphase. Figure 2A shows results from a successful experiment. We supplemented interphase-arrested extracts with demembranated Xenopus laevis sperm nuclei19 at a concentration of 27 nuclei/&#181;L and 0.38 &#181;M purified GST-GFP-NLS27,28,29,30 (fusion protein consisting of glutathione-S-transferase, green fluorescent protein, and a nuclear localization sequence) to allow reconstitution and visualization of interphase nuclei. We also added 1 &#181;M fluorescently labeled tubulin to visualize microtubules, and 500 nM MitoTracker Red CMXRos to visualize mitochondria. Moments after the extract was placed in the imaging chamber, the cytoplasm appeared disorganized. In the course of the next 30 minutes at room temperature, the cytoplasm started to self-organize into cell-like compartments. Microtubule asters grew from centrosomes introduced with the sperm nuclei, and formed microtubule-depleted border zones when met with microtubules from neighboring asters. GST-GFP-NLS protein translocated into the round interphase nuclei self-assembled from the added demembranated sperm nuclei. Areas depleted of light scattering cytoplasmic components were visible in both bright-field and mitochondria channels (Figure 2A, 20 min and 35 min). Mitochondria also became depleted from the borders established by microtubules, and became enriched in isolated compartments that aligned with microtubule compartments. By 60 min at room temperature, a spatial pattern consisting of cell-like compartments should be well-established, with microtubules forming a hollow wreath-like structure, and mitochondria clearly partitioned into each compartment (Figure 2A, 53 min).
Figure 2B compares extract performance in imaging chambers with and without FEP tapes on glass. We supplemented interphase-arrested extracts with demembranated Xenopus laevis sperm nuclei19 at a concentration of 27 nuclei/&#181;L and 0.35 &#181;M GST-mCherry-NLS27,28,29,30 (fusion protein consisting of glutathione-S-transferase, mCherry fluorescent protein, and a nuclear localization sequence) to allow reconstitution and visualization of interphase nuclei. We also added 1 &#181;M fluorescently labeled tubulin to visualize microtubules. Differences in dynamics became obvious by approximately 20 min at room temperature. In the chamber made with FEP-taped glass, the extract self-organized into normal cell-like patterns (Figure 2B, images in rows 1 and 3). However, in the chamber where glass surfaces were not covered by the FEP tape (unpassivated), the extract showed abnormal bright-field and microtubule patterns that became increasingly disrupted over time (Figure 2B, images in rows 2 and 4). No significant differences were observed in nuclear import of the GST-mCherry-NLS protein (Figure 2B, images in rows 5 and 6).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61923/61923fig01.jpg" /> 
Figure 1: Schematics and photos related to the experimental procedure. (A) Schematic diagram for preparing FEP tape-coated glass coverslips and slides. (B) Xenopus laevis eggs deposited in egg laying buffer, with examples of poor-quality eggs indicated by arrows. Yellow arrows, examples of eggs that look like white puffy balls. Red arrows, examples of eggs that appear in a string. (C) Examples of Xenopus laevis eggs with normal appearance. (D) Examples of poor-quality eggs with irregular or mottled pigment. (E) A deteriorating egg with a growing white region. (F) An egg that shows darkened and contracted pigmented area, possibly due to parthenogenetic activation. (G) The layers formed by ruptured Xenopus laevis eggs after the 12,000 x g centrifugation in step 2.11. (H) Schematics for preparing extract imaging chamber in step 2.15. (I) A photo of a prepared imaging chamber with an egg extract inside. (C) and (D) share the same scale bar at the bottom of (D). (E) and (F) share the same scale bar at the bottom of (F). The scale bars in (B) (C) (D) (E) (F) and (I) are approximate. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61923/61923fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61923/61923fig02.jpg" /> 
Figure 2: Interphase-arrested egg extracts self-organize into cell-like compartments. (A)Time-lapse montage of self-organized pattern formation in a thin layer (120 &#181;m) of interphase-arrested Xenopus laevis egg extract. The extract was supplemented with 27 nuclei/&#181;L of demembranated Xenopus laevis sperm nuclei to allow reconstitution of the interphase nuclei. Microtubules were visualized by HiLyte 647-labeled tubulin (shown in magenta), mitochondria by MitoTracker Red CMXRos (shown in red), and nuclei by GST-GFP-NLS (shown in green). (B) Self-organized pattern formation in interphase-arrested Xenopus laevis egg extracts placed in chambers with and without FEP-tape covered glass surfaces. The extracts were supplemented with 27 nuclei/&#181;L of demembranated Xenopus laevis sperm nuclei to allow reconstitution of the interphase nuclei. Microtubules were visualized by HiLyte 488-labeled tubulin (shown in magenta) and nuclei by GST-mCherry-NLS (shown in green). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61923/61923fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61923/61923fig03.jpg" /> 
Figure 3: Optional secondary seal for the imaging chamber. Schematics for preparing an optional secondary seal with mineral oil to prevent the extract from prolonged contact with air.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61923/61923fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Xenopus laevis Egg Extract Preparation and Live Imaging Methods for Visualizing Dynamic Cytoplasmic Organization at JoVE.com

Xenopus laevis Egg Extract Preparation and Live Imaging Methods for Visualizing Dynamic Cytoplasmic Organization

We describe a method for the preparation and live imaging of undiluted cytoplasmic extracts from Xenopus laevis eggs.

Xenopus laevis Egg Extract Preparation and Live Imaging Methods for Visualizing Dynamic Cytoplasmic Organization

Traditionally used for bulk biochemical assays, Xenopus laevis egg extracts have emerged as a powerful imaging-based tool for studying cytoplasmic ...

xenopus-laevis-egg-extract-preparation-live-imaging-methods-for

Nanyang Technological University

Research

JoVE Journal

Biology

3.0K Views.  Stanford University School of Medicine. We describe a method for the preparation and live imaging of undiluted cytoplasmic extracts from Xenopus laevis eggs.

Video: Xenopus laevis Egg Extract Preparation and Live Imaging Methods for Visualizing Dynamic Cytoplasmic Organization

Plastic Embedding and Sectioning of Xenopus laevis Embryos

Plastic sections maintain true tissue morphology in thin sections of tissue that can be immunostained with fluorescent secondary antibodies, making this method more useful than paraffin-embedded or frozen sections for many types of tissue. The method for staining, plastic embedding, and sectioning is demonstrated in this video.

Obtaining Eggs from Xenopus laevis Females

The eggs of Xenopus laevis intact, lysed, and/or fractionated are useful for a wide variety of experiments. This protocol shows how to induce egg laying, collect and dejelly the eggs, and sort the eggs to remove any damaged eggs.

The eggs of Xenopus laevis intact, lysed, and/or fractionated are useful for a wide variety of experiments. This protocol shows how to induce egg laying, collect and dejelly the eggs, and sort the eggs to remove any damaged eggs.

The eggs of Xenopus laevis intact, lysed, and/or fractionated are useful for a wide variety of experiments. This protocol shows how to induce egg laying, ...

Preparation and Fractionation of Xenopus laevis Egg Extracts

Crude and fractionated Xenopus egg extracts can be used to provide ingredients for reconstituting cellular processes for morphological and biochemical analysis. Egg lysis and differential centrifugation are used to prepare the crude extract which in turn in used to prepare fractionated extracts and light membrane preparations.

Crude and fractionated Xenopus egg extracts can be used to provide ingredients for reconstituting cellular processes for morphological and biochemical analysis. Egg lysis and differential centrifugation are used to prepare the crude extract which in turn in used to prepare fractionated extracts and light membrane preparations.

Crude and fractionated Xenopus egg extracts can be used to provide ingredients for reconstituting cellular processes for morphological and biochemical ...

Microinjection of Xenopus Laevis Oocytes

Microinjection of Xenopus laevis oocytes followed by thin-sectioning electron microscopy (EM) is an excellent system for studying nucleocytoplasmic transport. Because of its large nucleus and high density of nuclear pore complexes (NPCs), nuclear transport can be easily visualized in the Xenopus oocyte. Much insight into the mechanisms of nuclear import and export has been gained through use of this system (reviewed by Pant&eacute;, 2006). In addition, we have used microinjection of Xenopus oocytes to dissect the nuclear import pathways of several viruses that replicate in the host nucleus. 
Here we demonstrate the cytoplasmic microinjection of Xenopus oocytes with a nuclear import substrate. We also show preparation of the injected oocytes for visualization by thin-sectioning EM, including dissection, dehydration, and embedding of the oocytes into an epoxy embedding resin. Finally, we provide representative results for oocytes that have been microinjected with the capsid of the baculovirus Autographa californica nucleopolyhedrovirus (AcMNPV) or the parvovirus Minute Virus of Mice (MVM), and discuss potential applications of the technique.

Here we demonstrate cytoplasmic microinjection of Xenopus laevis oocytes with a nuclear import substrate, as well as preparation of the injected oocytes for visualization by thin-sectioning electron microscopy.

Microinjection of Xenopus laevis oocytes followed by thin-sectioning electron microscopy (EM) is an excellent system for studying nucleocytoplasmic ...

Visualizing RNA Localization in Xenopus Oocytes

RNA localization is a conserved mechanism of establishing cell polarity. Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to spatially restrict gene expression of Vg1 protein. Tight control of Vg1 distribution in this manner is required for proper germ layer specification in the developing embryo. RNA sequence elements in the 3' UTR of the mRNA, the Vg1 localization element (VLE) are required and sufficient to direct transport. To study the recognition and transport of Vg1 mRNA in vivo, we have developed an imaging technique that allows extensive analysis of trans-factor directed transport mechanisms via a simple visual readout. 
To visualize RNA localization, we synthesize fluorescently labeled VLE RNA and microinject this transcript into individual oocytes. After oocyte culture to allow transport of the injected RNA, oocytes are fixed and dehydrated prior to imaging by confocal microscopy. Visualization of mRNA localization patterns provides a readout for monitoring the complete pathway of RNA transport and for identifying roles in directing RNA transport for cis-acting elements within the transcript and trans-acting factors that bind to the VLE (Lewis et al., 2008, Messitt et al., 2008). We have extended this technique through co-localization with additional RNAs and proteins (Gagnon and Mowry, 2009, Messitt et al., 2008), and in combination with disruption of motor proteins and the cytoskeleton (Messitt et al., 2008) to probe mechanisms underlying mRNA localization.

Visualizing RNA Localization in Xenopus Oocytes

Visualization of in vivo RNA transport is accomplished by microinjection of fluorescently labeled RNA transcripts into Xenopus oocytes, followed by confocal microscopy.

RNA localization is a conserved mechanism of establishing cell polarity.  Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to ...

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 ...

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes in cell surface area and cell height, and where cells undergo mitosis and migrate. Furthermore, epithelial cells can also regulate morphogenetic movements in adjacent tissues1. A traditional method to study epithelial cells and tissues involve chemical fixation and histological methods to determine cell morphology or localization of particular proteins of interest. These approaches continue to be useful and provide "snapshots" of cell shapes and tissue architecture, however, much remains to be understood about how cells acquire specific shapes, how various proteins move or localize to specific positions, and what paths cells follow toward their final differentiated fate. High resolution live imaging complements traditional methods and also allows more direct investigation into the dynamic cellular processes involved in the formation, maintenance, and morphogenesis of multicellular epithelial sheets. Here we demonstrate experimental methods from the isolation of animal cap tissues from Xenopus laevis embryos to confocal imaging of epithelial cells and simple measurement approaches that together can augment molecular and cellular studies of epithelial morphogenesis.

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Xenopus embryonic epithelia are an ideal model system to study cell behaviors such as polarity development and shape change during epithelial morphogenesis. Traditional histology of fixed samples is increasingly being complemented by live-cell confocal imaging. Here we demonstrate methods to isolate frog tissues and visualize live epithelial cells and their cytoskeleton using live-cell confocal microscopy.

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, ...

Facial Transplants in Xenopus laevis Embryos

Craniofacial birth defects occur in 1 out of every 700 live births, but etiology is rarely known due to limited understanding of craniofacial development. To identify where signaling pathways and tissues act during patterning of the developing face, a 'face transplant' technique has been developed in embryos of the frog Xenopus laevis. A region of presumptive facial tissue (the "Extreme Anterior Domain" (EAD)) is removed from a donor embryo at tailbud stage, and transplanted to a host embryo of the same stage, from which the equivalent region has been removed. This can be used to generate a chimeric face where the host or donor tissue has a loss or gain of function in a gene, and/or includes a lineage label. After healing, the outcome of development is monitored, and indicates roles of the signaling pathway within the donor or surrounding host tissues. Xenopus is a valuable model for face development, as the facial region is large and readily accessible for micromanipulation. Many embryos can be assayed, over a short time period since development occurs rapidly. Findings in the frog are relevant to human development, since craniofacial processes appear conserved between Xenopus and mammals.

Facial Transplants in Xenopus laevis Embryos

A technique for transplanting "Extreme Anterior Domain" facial tissue between Xenopus laevis embryos has been developed. Tissue can be moved from one gene expression background into another, allowing the study of local requirements for craniofacial development and for signaling interactions between facial regions.

Craniofacial birth defects occur in 1 out of every 700 live births, but etiology is rarely known due to limited understanding of craniofacial development. ...

Reconstitution Of &#946;-catenin Degradation In Xenopus Egg Extract

Xenopus laevis egg extract is a well-characterized, robust system for studying the biochemistry of diverse cellular processes. Xenopus egg extract has been used to study protein turnover in many cellular contexts, including the cell cycle and signal transduction pathways1-3. Herein, a method is described for isolating Xenopus egg extract that has been optimized to promote the degradation of the critical Wnt pathway component, &beta;-catenin. Two different methods are described to assess &beta;-catenin protein degradation in Xenopus egg extract. One method is visually informative ([35S]-radiolabeled proteins), while the other is more readily scaled for high-throughput assays (firefly luciferase-tagged fusion proteins). The techniques described can be used to, but are not limited to, assess &beta;-catenin protein turnover and identify molecular components contributing to its turnover. Additionally, the ability to purify large volumes of homogenous Xenopus egg extract combined with the quantitative and facile readout of luciferase-tagged proteins allows this system to be easily adapted for high-throughput screening for modulators of &beta;-catenin degradation.

Reconstitution Of &#946;-catenin Degradation In Xenopus Egg Extract

A method is described for analyzing protein degradation using radiolabeled and luciferase-fusion proteins in Xenopus egg extract and its adaptation for high-throughput screening for small molecule modulators of protein degradation.

Xenopus laevis egg extract is a well-characterized, robust system for studying the biochemistry of diverse cellular processes. Xenopus egg extract has ...