We thank Acaimo Gonz&#225;lez Reyes and Mar&#237;a Olmedo L&#243;pez for helpful comments on the manuscript. This study was supported by PID2021-125480NB-I00 (H. S-G), &#34;Ayudas a la contrataci&#243;n de personal Investigador Doctor&#34; from Junta de Andaluc&#237;a (J. G-M), &#34;Ayuda a proyectos de investigaci&#243;n precompetitivos&#34; from VI and VII PPIT-Seville University (P. R-R) and &#34;Contrato de Acceso de I+D+i&#34; from VI PPIT-Seville University (P. R-R). We extend our acknowledgments to the Company of Biologists Ltd for kindly providing rights to share images adapted from the original publication (doi:10.1242/DEV.199716).

NOTE: Step 1.4 must be done at least 2 days in advance. Steps 2.1 and 2.2 can be done 1 day in advance.
1. Pre-experimental setup I
<ol>
	<li>Prepare 100 &#181;L aliquots of streptomycin/penicillin antibiotic mix (10,000 U/mL penicilin and 10 mg/mL streptomycin) and store at -20 &#176;C until use.</li>
	<li>Prepare 15 mL of the following solutions using autoclaved pure water, store them at 4 &#176;C, and use them within a month: Sodium bicarbonate (NaHCO3) 0.1 M pH 8.0 and 0.1% Tween 20.</li>
	<li>Prepare the following culture medium and solution in a laminar flow hood to avoid contamination. Use sterilized filter tips and tubes.
	<ol>
		<li>Prepare 2 mL aliquots of undiluted fetal bovine serum (FBS) and store at -20 &#176;C until use in aseptic conditions.</li>
		<li>Prepare Schneider&#39;s Drosophila medium supplemented with 15% FBS and 0.6% streptomycin/penicillin antibiotic mix. Prepare 7 mL aliquots and store at 4 &#176;C. One aliquot is enough for two MatTek plates (see section 2).
		<ol>
			<li>Prepare Ringer&#39;s solution (128 mM NaCl, 2 mM KCl, 1.8 mM CaCl2, 4 mM MgCl2, 35.5 mM Sucrose, 5 mM Hepes in autoclaved pure water; pH 6.9). Prepare 1 mL aliquots and store at -20 &#176;C until use.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Collect 1-2 days old flies expressing constitutively and ubiquitously the Par-1 protein fused to GFP (poly Ubiquitin-mGFP6::par-1 line) into a new tube with fly food supplemented with yeast powder. Culture them for 2 days at 25 &#176;C in an incubator before dissection. 
	NOTE: This step can be done with any fluorescently tagged reporter of interest.</li>
</ol>
2. Glass bottom plate coating and preparation
<ol>
	<li>Place a 3 &#181;L-drop of a specific Cell-Tak adhesive (henceforth referred to as cell and tissue adhesive) in the center of the coverslip of a 35-mm poly-D-lysine coated plate (glass bottom dish, see Table of Materials) without manual spreading. Use sterile tips. 
	NOTE: The cell and tissue adhesive should be stored at 4 &#176;C. In the case of experiments with two genetic conditions (typically, control and mutant conditions), place two individual, well-separated drops of the adhesive in the same glass bottom plate. This will ensure that both genetic backgrounds are imaged in the same conditions.</li>
	<li>Immediately after, add an equal volume (3 &#181;L in this protocol setup) of 0.1 M NaHCO3 pH 8.0 carefully into the 3 &#181;L adhesive drop as recommended by the manufacturer and mix by pipetting.</li>
	<li>For complete evaporation, incubate the plate at room temperature (RT) for 20 min and then keep at 4 &#176;C overnight to be used the next day. Alternatively, incubate the prepared plate at 37 &#176;C for 25 min until complete evaporation and proceed directly to the next step. In both cases, maintain the plate with its cover. Use sterile tips.</li>
	<li>Allow the prepared glass bottom dish to reach RT by placing it on the bench for 15-20 min before use.</li>
	<li>Wash 3 times the prepared plate carefully. Avoid touching and disturbing the adhesive layer(s). In each wash, add 6 &#181;L of autoclaved pure water and then remove the water using a clean tip for each of the three washes to eliminate residues of the NaHCO3 solution.</li>
	<li>Allow the complete evaporation of the water by placing the clean and prepared plate at RT&#160;while dissecting for 5-10 min with its lid on to avoid dust particles.</li>
</ol>
3. Drosophila ovary dissection and ovariole isolation and mounting
NOTE: The ovary dissection is performed in Ringer&#39;s solution (instead of Schneider&#39;s medium) without FBS to prevent the presence of proteins that could interfere with the sticking of the sample tissue to the adhesive.
<ol>
	<li>Clean a three-well dissection dish, forceps, and dissection needles with 70% ethanol before starting.</li>
	<li>Add a sufficient volume (~200 &#181;L) of Ringer&#39;s solution in each of the three wells.</li>
	<li>Dissect the ovaries of 5 yeast-fed females in 200 &#181;L of Ringer&#39;s solution in the first well with the help of two pairs of forceps following standard procedures10,11. Grab the Drosophila female at the intersection of the thorax and abdomen with one of the forceps. Then, gently tear the cuticle at the posterior abdomen and pull the abdomen cuticle until the two ovaries are exposed.</li>
	<li>Transfer the dissected ovaries to the second well to reduce contamination with remnants of other tissues or debris resulting from dissection.</li>
	<li>Remove the muscle sheath of 15-20 ovarioles with the help of a stereomicroscope (zoom 8X-13.5X) with light-emitting diode (LED) illumination and fine-point forceps. Use one pair of forceps to anchor the entire body of the ovary, and use the second one to smoothly grab a late-stage egg chamber and gently stretch until the muscle breaks and lags behind. 
	NOTE: It is essential to avoid damage to the germaria just by reducing the contact with this part of the ovaries. Remove egg chambers older than stages 8-9.</li>
	<li>Transfer one by one the 15-20 muscle sheath-free ovarioles using the forceps to the third well containing 200 &#181;L of Ringer&#39;s medium.</li>
	<li>Pre-wet a 20-200 &#181;L tip with 0.1% Tween 20 by pipetting. This averts the ovarioles&#39; adhesion to the tip&#39;s plastic walls.</li>
	<li>Transfer 6 &#181;L of of Ringer&#39;s solution containing the muscle sheath-free ovarioles using a pre-wetted tip to the prepared adhesive plate (kept at RT). 
	NOTE: Avoid all contact between the tip and the adhesive layer to prevent germaria detachment after ovariole deposition (next step). Approximately 30% of germaria do not adhere as expected.</li>
	<li>To ensure that the transferred ovarioles adhere to the adhesive surface, carefully press them down to the bottom of the Ringer&#39;s drop until they contact the adhesive surface using a dissection needle or forceps. 
	NOTE: Do not try to rearrange the ovarioles once they settle.</li>
	<li>Fill the MatTek plate carefully with 3 mL of Schneider&#39;s medium supplemented with the 15% FBS and 0.6% streptomycin/penicillin antibiotic mix (step 1.3). To prevent the detachment of the ovarioles, add slowly 1 mL at a time around the outer edge of the plate.</li>
	<li>Transport the plate containing the muscle sheath-free ovarioles on a flat surface to the microscope station.</li>
</ol>
4. Live imaging using a confocal microscope
<ol>
	<li>Place the plate on the 63x objective of a spinning-disk microscope or a confocal microscope (similar equipment can be used) and focus the sample. Select the middle z plane for each germarium.</li>
	<li>Configure the following experimental parameters: capture type: 3D time-lapse, adjust the laser power of 70/100, excitation wavelength: 488 nm, range of the z-stack: 30 &#181;m and distance between Z-stacks: 1.2 &#181;m, duration: 17 h, interval between time-points: every 10 min.</li>
	<li>If available, using a multi-position option, redefine the XY coordinates of the samples and select a Z-plane in the middle of each germarium, as the z-stacks will be centered in this position. Start the acquisition. 
	NOTE: As the normal width of a germarium is 30 &#181;m, each Z-stack will be ~25-27 sections. Depending on the confocal, at least 25 germaria can be imaged within the 10 min intervals. Normally, 40x/1.30 NA or 63x/1.40 NA oil-immersion, planapo-corrected objectives are used.</li>
	<li>Analyze the images in time and Z-axis to visualize specific changes in the spectrosome morphology using Imaris or ImageJ (Fiji) software12,13. Specific times on their observation are displayed (Figure 1B&#39;). To mount the movies, manually select the best Z plane at each time point.</li>
</ol>

Real-Time Imaging of Germline Stem Cells in Drosophila Ex Vivo Germaria Culture

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The authors declare that they have no competing financial interests.

Here we present a protocol to monitor the Drosophila melanogaster GSC niche for a prolonged period of time, up to 16 h. Most of the protocols to film Drosophila biological processes ex vivo focus on shorter time windows and multiple examples can be found for imaginal wing discs15,16,17, embryo gonads18,19 or ovaries20<su...

Live imaging of biological processes is instrumental in obtaining direct experimental evidence. The combination of advanced confocal microscopy and the optimization of methodologies permits the exploration of multiple biological events with high precision. Optimizing steps in protocols such as tissue manipulation and dissection, sample preparation and preservation, and microscopy acquisition settings is critical to maximize the reliability and robustness of the results obtained. Here, we present a protocol to monitor samples for extended imaging, which is specially focused on Drosophila melanogaster ovaries. The Drosophila melanogaster ovary is an excellent model system for the analyses of a wide range of developmental processes. Among many others, this reproductive organ of the fruit fly Drosophila melanogaster contains a very well-defined adult stem cell niche. This GSC niche sustains the development of the female gametes during adulthood. The Drosophila ovaries are composed of approximately 18 ovarioles, where egg chambers are developed in the germarium. In the tip of each germarium, 2-4 GSCs are maintained in a somatic cellular niche mainly formed by a terminal filament of 8-10 cells, a rosette of 5-8 cap cells, and 2-3 anterior escort cells (Figure 1A). This somatic niche provides the GSCs with essential signals and physical support to maintain their stemness, control proliferation, and prevent differentiation1,2,3,4,5.
The GSCs normally divide asymmetrically to generate a new stem cell that keeps contact with the somatic niche and a daughter cell, the cystoblast (CB), which loses direct contact with the somatic cap cells and differentiates. GSCs and CBs contain a highly dynamic and cytoplasmic organelle, the spectrosome, whose main function is the correct orientation of the mitotic spindle during mitosis6. The CB divides 4 times with incomplete cytokinesis to develop 16-cell cysts, where one of the germline cells specifies into the oocyte and the other 15 cells become nurse cells. The CB spectrosome grows into a branched structure called the fusome which connects the 16-cell interconnected germline cells. The spectrosome is enriched in small vesicles and skeletal proteins such as the serine-threonine kinase Par-1 and the membrane component Hu-li tai shao (Hts)7. During the GSC cell cycle, the spectrosome grows by the addition of new material and changes its shape, allowing the identification of the G1, S, G2, and M phases (Figure 1B)8,9.
We have recently implemented an ex vivo culture method of the Drosophila germarium that allows imaging of live GSCs for up to 16 h. Since these cells divide once every 15.5 h on average8, this method permits the filming of large portions of the GSC cell cycle. Thus, in combination with other tools, our culturing method has allowed the description of spectrosome morphology during the GSC cell cycle and the analysis of the duration of the different cell cycle phases in vivo8 (Figure 1C). Here, we provide a detailed protocol of this extended live imaging method supported by a step-by-step guided video that describes the methodology (Figure 2).

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>9-well glass plate</td>
			<td>Corning&#160;</td>
			<td>7220-85</td>
			<td>n/a</td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate</td>
			<td>Sigma Aldrich</td>
			<td>C3881</td>
			<td>CaCl2&#183;2H2O; To prepare Ringer&#39;s solution</td>
		</tr>
		<tr>
			<td>Cell-Tak</td>
			<td>Corning&#160;</td>
			<td>354240</td>
			<td>Cell and Tissue Adhesive used to attach cells or tissue sections to many types of surfaces, including plastic, glass, metal, FEP Polymer, and biological materials.</td>
		</tr>
		<tr>
			<td>Dumont #5 Forceps&#160;</td>
			<td>Finescience</td>
			<td>11252-20</td>
			<td>n/a</td>
		</tr>
		<tr>
			<td>Dumont #55 Forceps&#160;</td>
			<td>Finescience</td>
			<td>11255-20</td>
			<td>Dimensions 0.05 mm x 0.02 mm and length 11 cm</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Gibco</td>
			<td>10500-064</td>
			<td>Qualified, heat inactivated, E.U.-approved, South America Origin&#160;</td>
		</tr>
		<tr>
			<td>Glass bottom dishes</td>
			<td>MatTek</td>
			<td>P35GC-1.5-10-C</td>
			<td>35 mm Dish, No. 1.5 Coverslip, 10 mm Glass Diameter,&#160;Poly-D-Lysine Coated</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma Aldrich</td>
			<td>H4034</td>
			<td>To prepare Ringer&#39;s solution</td>
		</tr>
		<tr>
			<td>Magnesium chloride&#160;</td>
			<td>Sigma Aldrich</td>
			<td>M1028</td>
			<td>MgCl2; To prepare Ringer&#39;s solution</td>
		</tr>
		<tr>
			<td>Needle Holder</td>
			<td>Roboz&#160;</td>
			<td>RS-6061</td>
			<td>Light weight, hollow stainless steel handle; 4 3/4&#34; Long.</td>
		</tr>
		<tr>
			<td>Nikon SMZ18 binocular&#160;</td>
			<td>Nikon</td>
			<td></td>
			<td>https://www.microscope. 
			healthcare.nikon.com/products/stereomicroscopes-macroscopes/smz25-smz18</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Thermo Fisher Scientific</td>
			<td>15140122</td>
			<td>10,000 U/mL</td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Sigma Aldrich</td>
			<td>P3911</td>
			<td>KCl; To prepare Ringer&#39;s solution</td>
		</tr>
		<tr>
			<td>Schneider&#39;s Drosophila Medium</td>
			<td>Biowest&#160;</td>
			<td>L0207</td>
			<td>Cell culture medium for insect cells</td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>Sigma Aldrich</td>
			<td>S8875&#160;</td>
			<td>NaHCO3; &#8805;99.5%, powder</td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Sigma Aldrich</td>
			<td>S9888</td>
			<td>NaCl; To prepare Ringer&#39;s solution</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma Aldrich</td>
			<td>S9378</td>
			<td>&#8805;99.5% (GC); To prepare Ringer&#39;s solution</td>
		</tr>
		<tr>
			<td>Tungsten Dissection Needle</td>
			<td>Roboz&#160;</td>
			<td>RS-6064</td>
			<td>0.25 mm, Ultra Fine, 1 Micron Tip (Pk 10)</td>
		</tr>
		<tr>
			<td>Tween 20</td>
			<td>Sigma Aldrich</td>
			<td>P9416</td>
			<td>for molecular biology, viscous liquid</td>
		</tr>
		<tr>
			<td>Yeast powder</td>
			<td>Sigma Aldrich</td>
			<td>51475</td>
			<td>n/a</td>
		</tr>
	</tbody>
</table>

extended live imaging of female drosophila melanogaster germline stem cell niches

Live imaging methods allow the analysis of dynamic cellular processes in detail and in real-time. The Drosophila ovary represents an excellent model to explore the dynamics of a myriad of developmental processes, such as cell division, stemness, differentiation, migration, apoptosis, autophagy, cellular adhesion, etc., over time. Recently, we have implemented an extended ex vivo culture and live imaging of the female Drosophila GSC niche. Using a Drosophila line harboring a GFP::Par-1 transgene as an example, this method allows the visualization of the GSCs&#39; asymmetric division within their niche and the description of the changes in the spectrosome morphology along the cell cycle. Here, we present a detailed protocol for the ex vivo culture of Drosophila germaria, enabling prolonged visualization of the female GSC niche. Importantly, this protocol is broadly applicable to live imaging GSCs with multiple fluorescently tagged proteins of interest that are available in stock centers and/or in the Drosophila research community.

With this extended live imaging protocol, we can record the asymmetric mitosis of female GSCs inside their niche without apparent biological disturbances. To do so, we use germaria expressing ubiquitously the GFP::Par-1 protein, which discriminates between five distinct spectrosome morphologies through the GSC cell cycle: Round, Plug, Bar, Fusing, and Exclamation point (Figure 1B,C). In a more detailed description, we observed that the strong GFP:Par1 signal of Round-G2 spec...

Extended Live Imaging of Female Drosophila melanogaster Germline Stem Cell Niches

Here, we provide a detailed protocol to perform live imaging of the asymmetric division of germline stem cells (GSCs) in the Drosophila ovarian niche. We use a transgenic line that ubiquitously expresses a green fluorescent protein (GFP) fusion of the spectrosome protein Par-1.

Live imaging methods allow the analysis of dynamic cellular processes in detail and in real-time. The Drosophila ovary represents an excellent model to ...

extended-live-imaging-female-drosophila-melanogaster-germline-stem

Research

JoVE Journal

Developmental Biology

513 Views.  CSIC/Universidad Pablo de Olavide/JA. Here, we provide a detailed protocol to perform live imaging of the asymmetric division of germline stem cells (GSCs) in the Drosophila ovarian niche. We use a transgenic line that ubiquitously expresses a green fluorescent protein (GFP) fusion of the spectrosome protein Par-1.

Video: Extended Live Imaging of Female Drosophila melanogaster Germline Stem Cell Niches

Live-imaging of the Drosophila Pupal Eye

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track during live cell imaging. These processes include: retinal rotation, cell growth and organismal movement. Additionally, irregularities in the topology of the epithelium, including subtle bumps and folds often introduced as the pupa is prepared for imaging, make it challenging to acquire in-focus images of more than a few ommatidia in a single focal plane. The workflow outlined here remedies these issues, allowing easy analysis of cellular processes during Drosophila pupal eye development. Appropriately-staged pupae are arranged in an imaging rig that can be easily assembled in most laboratories. Ubiquitin-DE-Cadherin:GFP and GMR-GAL4&#8211;driven UAS-&#945;-catenin:GFP are used to visualize cell boundaries in the eye epithelium 1-3. After deconvolution is applied to fluorescent images captured at multiple focal planes, maximum projection images are generated for each time point and enhanced using image editing software. Alignment algorithms are used to quickly stabilize superfluous motion, making individual cell behavior easier to track.

Live-imaging of the Drosophila Pupal Eye

This protocol presents an efficient method for imaging the live Drosophila pupal eye neuroepithelium. This method compensates for tissue movement and uneven topology, enhances visualization of cell boundaries through the use of multiple GFP-tagged junction proteins, and uses an easily-assembled imaging rig.

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track ...

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

In vertebrates, hematopoiesis is regulated by inductive microenvironments (niches). Likewise, in the invertebrate model organism&#160;Drosophila melanogaster, inductive microenvironments known as larval Hematopoietic Pockets (HPs) have been identified as anatomical sites for the development and regulation of blood cells (hemocytes), in particular of the self-renewing macrophage lineage.&#160;HPs are segmentally repeated pockets between the epidermis and muscle layers of the larva, which also comprise sensory neurons of the peripheral nervous system. In the larva, resident (sessile) hemocytes are exposed to anti-apoptotic, adhesive and proliferative cues from these sensory neurons and potentially other components of the HPs, such as the lining muscle and epithelial layers. During normal development, gradual release of resident hemocytes from the HPs fuels the population of circulating hemocytes, which culminates in the release of most of the resident hemocytes at the beginning of metamorphosis. Immune assaults, physical injury or mechanical disturbance trigger the premature release of resident hemocytes into circulation. The switch of larval hemocytes between resident locations and circulation raises the need for a common standard/procedure to selectively isolate and quantify these two populations of blood cells from single Drosophila larvae. Accordingly, this protocol describes an automated method to release and quantify the resident and circulating hemocytes from single larvae. The method facilitates ex vivo approaches, and may be adapted to serve a variety of developmental stages of Drosophila and other invertebrate organisms.

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

Drosophila blood cells, or hemocytes, cycle between resident sites and circulation. In the larva, resident (sessile) hemocytes localize to inductive microenvironments, the Hematopoietic Pockets, while circulating hemocytes move freely in the hemolymph. The goal of this protocol is the standardized isolation and quantification of these two, behaviorally distinct but interchanging, hemocyte populations.

In vertebrates, hematopoiesis is regulated by inductive microenvironments (niches). Likewise, in the invertebrate model organism&#160;Drosophila ...

Long-term Live Imaging of Drosophila Eye Disc

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have been used to study many biological processes, such as cell proliferation, differentiation, growth, migration, apoptosis, competition, cell-cell signaling, and compartmental boundary formation. However, methods for the long-term ex vivo culture and live imaging of the imaginal discs have not been satisfactory, despite many efforts. Recently, we developed a method for the long-term ex vivo culture and live imaging of imaginal discs for up to 18 h. In addition to using a high insulin concentration in the culture medium, a low-melting agarose was also used to embed the disc to prevent it from drifting during the imaging period. This report uses the eye-antennal discs as an example. Photoreceptor R3/4-specific m&#948;0.5-Ga4 expression was followed to demonstrate that photoreceptor differentiation and ommatidial rotation can be observed during a 10 h live imaging period. This is a detailed protocol describing this simple method.

Long-term Live Imaging of Drosophila Eye Disc

This protocol describes a detailed method for the long-term ex vivo culture and live imaging of a Drosophila imaginal disc. It demonstrates photoreceptor differentiation and ommatidial rotation within the 10 h period of live imaging of the eye disc. The protocol is simple and does not require expensive setup.

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have ...

Bioengineering of Humanized Bone Marrow Microenvironments in Mouse and Their Visualization by Live Imaging

Human hematopoietic stem cells (HSCs) reside in the bone marrow (BM) niche, an intricate, multifactorial network of components producing cytokines, growth factors, and extracellular matrix. The ability of HSCs to remain quiescent, self-renew or differentiate, and acquire mutations and become malignant depends upon the complex interactions they establish with different stromal components. To observe the crosstalk between human HSCs and the human BM niche in physiological and pathological conditions, we designed a protocol to ectopically model and image a humanized BM niche in immunodeficient mice. We show that the use of different cellular components allows for the formation of humanized structures and the opportunity to sustain long-term human hematopoietic engraftment. Using two-photon microscopy, we can live-image these structures in situ at the single-cell resolution, providing a powerful new tool for the functional characterization of the human BM microenvironment and its role in regulating normal and malignant hematopoiesis.

A method to create and live-image different humanized bone-marrow niches in mice is presented. Based on the supportive niche created by human mesenchymal cells, the addition of human endothelial cells induces the formation of human vessels, while the addition of rhBMP-2 induces the formation of human-mouse chimeric mature bone tissue.

Human hematopoietic stem cells (HSCs) reside in the bone marrow (BM) niche, an intricate, multifactorial network of components producing cytokines, growth ...

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; for example, how the zygote polarizes to establish the body axis and how the embryo specifies various cell fates during organ formation. This manuscript describes an in vitro ovule culture method to perform live-cell imaging of developing zygotes and embryos of Arabidopsis thaliana. The optimized cultivation medium allows zygotes or early embryos to grow into fertile plants. By combining it with a poly(dimethylsiloxane) (PDMS) micropillar array device, the ovule is held in the liquid medium in the same position. This fixation is crucial to observe the same ovule under a microscope for several days from the zygotic division to the late embryo stage. The resulting live-cell imaging can be used to monitor the real-time dynamics of zygote polarization, such as nuclear migration and cytoskeleton rearrangement, and also the cell division timing and cell fate specification during embryo patterning. Furthermore, this ovule cultivation system can be combined with inhibitor treatments to analyze the effects of various factors on embryo development, and with optical manipulations such as laser disruption to examine the role of cell-cell communication.

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

This manuscript describes an in vitro ovule cultivation method that enables live-cell imaging of Arabidopsis zygotes and embryos. This method is utilized to visualize the intracellular dynamics during zygote polarization and the cell fate specification in developing embryos.

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; ...

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and differentiation. Typically, pHi dynamics are determined in cultured cells, which are amenable to measuring and experimentally manipulating pHi. However, the recent development of new tools and methodologies has made it possible to study pHi dynamics within intact, live tissue. For Drosophila research, one important development was the generation of a transgenic line carrying a pHi biosensor, mCherry::pHluorin. Here, we describe a protocol that we routinely use for imaging live Drosophila ovarioles to measure pHi in the epithelial follicle stem cell (FSC) lineage in mCherry::pHluorin transgenic wild type lines; however, the methods described here can be easily adapted for other tissues, including the wing discs and eye epithelium. We describe techniques for expressing mCherry::pHluorin in the FSC lineage, maintaining ovarian tissue during live imaging, and acquiring and analyzing images to obtain pHi values.

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

We provide a protocol for imaging intracellular pH of an epithelial stem cell lineage in live Drosophila ovarian tissue. We describe methods to generate transgenic flies expressing a pH biosensor, mCherry::pHluorin, image the biosensor using quantitative fluorescence imaging, generate standard curves, and convert fluorescence intensity values to pH values.

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and ...

Live Cell Imaging of Chromosome Segregation During Mitosis

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled by multiple mechanisms that include the Spindle Assembly Checkpoint (SAC). The SAC is part of a complex feedback system that is responsible for prevention of a cell progress through mitosis unless all chromosomal kinetochores have attached to spindle microtubules. Chromosomal lagging and abnormal chromosome segregation is an indicator of dysfunctional cell cycle control checkpoints and can be used to measure the genomic stability of dividing cells. Deregulation of the SAC can result in the transformation of a normal cell into a malignant cell through the accumulation of errors during chromosomal segregation. Implementation of the SAC and the formation of the kinetochore complex are tightly regulated by interactions between kinases and phosphatase such as Protein Phosphatase 2A (PP2A). This protocol describes live cell imaging of lagging chromosomes in mouse embryonic fibroblasts isolated from mice that had a knockout of the PP2A-B56&#947; regulatory subunit. This method overcomes the shortcomings of other cell cycle control imaging techniques such as flow cytometry or immunocytochemistry that only provide a snapshot of a cell cytokinesis status, instead of a dynamic spatiotemporal visualization of chromosomes during mitosis.

This protocol describes an easy and convenient method to label and visualize live chromosomes in mitotic cells using Histone2B-GFP BacMam 2.0 label and a spinning disc confocal microscopy system.

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled ...

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits are required to control this behavior. Failure to produce the rhythmic pattern of contractions can be indicative of neurological abnormalities. We previously found that defects in protein O-mannosylation, a posttranslational protein modification, affect the axon morphology of sensory neurons and result in abnormal coordinated muscle contractions in embryos. Here, we present a relatively simple method for recording and analyzing the pattern of peristaltic muscle contractions by live imaging of late stage embryos up to the point of hatching, which we used to characterize the muscle contraction phenotype of protein O-mannosyltransferase mutants. Data obtained from these recordings can be used to analyze muscle contraction waves, including frequency, direction of propagation and relative amplitude of muscle contractions at different body segments. We have also examined body posture and taken advantage of a fluorescent marker expressed specifically in muscles to accurately determine the position of the embryo midline. A similar approach can also be utilized to study various other behaviors during development, such as embryo rolling and hatching.

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Here, we present a method to record embryonic muscle contractions in Drosophila embryos in a non-invasive and detail-oriented manner.

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits ...

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

There are currently over 160 distinct Drosophila cell lines distributed by the Drosophila Genomics Resource Center (DGRC). With genome engineering, the number of novel cell lines is expected to increase. The DGRC aims to familiarize researchers with using Drosophila cell lines as an experimental tool to complement and drive their research agenda. Procedures for working with a variety of Drosophila cell lines with distinct characteristics are provided, including protocols for thawing, culturing, and cryopreserving cell lines. Importantly, this publication demonstrates the best practices required to work with Drosophila cell lines to minimize the risk of contaminations from adventitious microorganisms or from other cell lines. Researchers who become familiar with these procedures will be able to delve into the many applications that use Drosophila cultured cells including biochemistry, cell biology and functional genomics.

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

Drosophila cell lines are important reagents for both fundamental and biomedical research. This article provides protocols for thawing, subculturing, and the cryopreservation of commonly used Drosophila cell lines to assist researchers in incorporating the use of these reagents in their research.

There are currently over 160 distinct Drosophila cell lines distributed by the Drosophila Genomics Resource Center (DGRC). With genome engineering, the ...

Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos

The purpose of this protocol is to visualize intranuclear actin rods that assemble in live Drosophila melanogaster embryos following heat stress. Actin rods are a hallmark of a conserved, inducible Actin Stress Response (ASR) that accompanies human pathologies, including neurodegenerative disease. Previously, we showed that the ASR contributes to morphogenesis failures and reduced viability of developing embryos. This protocol allows the continued study of mechanisms underlying actin rod assembly and the ASR in a model system that is highly amenable to imaging, genetics and biochemistry. Embryos are collected and mounted on a coverslip to prepare them for injection. Rhodamine-conjugated globular actin (G-actinRed) is diluted and loaded into a microneedle. A single injection is made into the center of each embryo. After injection, embryos are incubated at elevated temperature and intranuclear actin rods are then visualized by confocal microscopy. Fluorescence recovery after photobleaching (FRAP) experiments may be performed on the actin rods; and other actin-rich structures in the cytoplasm can also be imaged. We find that G-actinRed polymerizes like endogenous G-actin and does not, on its own, interfere with normal embryo development. One limitation of this protocol is that care must be taken during injection to avoid serious injury to the embryo. However, with practice, injecting G-actinRed into Drosophila embryos is a fast and reliable way to visualize actin rods and can easily be used with flies of any genotype or with the introduction of other cellular stresses, including hypoxia and oxidative stress.

Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos

The goal of this protocol is to inject Rhodamine-conjugated globular actin into Drosophila embryos and image intranuclear actin rod assembly following heat stress.

The purpose of this protocol is to visualize intranuclear actin rods that assemble in live Drosophila melanogaster embryos following heat stress. Actin ...