The authors are very thankful to Miriam Osterfield for sharing her advice on the in vitro life imaging of egg chambers using an approach adopted from the Celeste Berg laboratory4.&#160;We also thank to Sebastian Tosi for the Fiji script that enables cell segmentation.

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L., Bilder, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Global+tissue+revolutions+in+a+morphogenetic+movement+controlling+elongation.">Global tissue revolutions in a morphogenetic movement controlling elongation.</a> Science. 331, 1071-1074 (2011).</li><li>Andersen, D., Horne-Badovinac, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Influence+of+ovarian+muscle+contraction+and+oocyte+growth+on+egg+chamber+elongation+in+Drosophila.">Influence of ovarian muscle contraction and oocyte growth on egg chamber elongation in Drosophila.</a> Development. 143, 1375-1387 (2016).</li><li>Pokrywka, N. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+imaging+of+GFP-labeled+proteins+in+Drosophila+oocytes.">Live imaging of GFP-labeled proteins in Drosophila oocytes.</a> Journal of Visualized Experiments. (73), e50044 (2013).</li><li>Prasad, M., Jang, A. C., Starz-Gaiano, M., Melani, M., Montell, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+protocol+for+culturing+Drosophila+melanogaster+stage+9+egg+chambers+for+live+imaging.">A protocol for culturing Drosophila melanogaster stage 9 egg chambers for live imaging.</a> Nature Protocols. 2, 2467-2473 (2007).</li><li>Dorman, J. B., James, K. E., Fraser, S. E., Kiehart, D. P., Berg, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=bullwinkle+is+required+for+epithelial+morphogenesis+during+Drosophila+oogenesis.">bullwinkle is required for epithelial morphogenesis during Drosophila oogenesis.</a> Developmental Biology. 267, 320-341 (2004).</li><li>Cetera, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epithelial+rotation+promotes+the+global+alignment+of+contractile+actin+bundles+during+Drosophila+egg+chamber+elongation.">Epithelial rotation promotes the global alignment of contractile actin bundles during Drosophila egg chamber elongation.</a> Nature Communications. 5, 5511 (2014).</li><li>Viktorinova, I., Henry, I., Tomancak, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epithelial+rotation+is+preceded+by+planar+symmetry+breaking+of+actomyosin+and+protects+epithelial+tissue+from+cell+deformations.">Epithelial rotation is preceded by planar symmetry breaking of actomyosin and protects epithelial tissue from cell deformations.</a> PLOS Genetics. 13, e1007107 (2017).</li><li>Rauzi, M., Lenne, P. F., Lecuit, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Planar+polarized+actomyosin+contractile+flows+control+epithelial+junction+remodelling.">Planar polarized actomyosin contractile flows control epithelial junction remodelling.</a> Nature. 468, 1110-1114 (2010).</li><li>Spracklen, A. J., Fagan, T. N., Lovander, K. E., Tootle, T. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+pros+and+cons+of+common+actin+labeling+tools+for+visualizing+actin+dynamics+during+Drosophila+oogenesis.">The pros and cons of common actin labeling tools for visualizing actin dynamics during Drosophila oogenesis.</a> Developmental Biology. 393, 209-226 (2014).</li><li>. Surface manager Available from: <a target="_blank" href="https://git.mpi-cbg.de/tomancaklab/surface_manager">https://git.mpi-cbg.de/tomancaklab/surface_manager</a> (2018)</li><li>. Actomyosin dynamics: Required Software and dependencies Available from: <a target="_blank" href="https://git.mpi-cbg.de/scicomp/viktorinova_et_al_actomyosin_dynamics/blob/master/Software/Software_installation.md#surface-manager-and-ellipsoid-surface-projection-plugins">https://git.mpi-cbg.de/scicomp/viktorinova_et_al_actomyosin_dynamics/blob/master/Software/Software_installation.md#surface-manager-and-ellipsoid-surface-projection-plugins</a> (2018)</li><li>. 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N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+roles+for+the+actin+and+microtubule+cytoskeletons+in+the+morphogenesis+of+epidermal+hairs+during+wing+development+in+Drosophila.">Distinct roles for the actin and microtubule cytoskeletons in the morphogenesis of epidermal hairs during wing development in Drosophila.</a> Mechanisms of Development. 70, 181-192 (1998).</li><li>Heemskerk, I., Streichan, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue+cartography:+compressing+bio-image+data+by+dimensional+reduction.">Tissue cartography: compressing bio-image data by dimensional reduction.</a> Nature Methods. 12, 1139-1142 (2015).</li><li>Chen, D. Y., Lipari, K. R., Dehghan, Y., Streichan, S. J., Bilder, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Symmetry+Breaking+in+an+Edgeless+Epithelium+by+Fat2-Regulated+Microtubule+Polarity.">Symmetry Breaking in an Edgeless Epithelium by Fat2-Regulated Microtubule Polarity.</a> Cell Reports. 15, 1125-1133 (2016).</li><li>Reynaud, E. G., Peychl, J., Huisken, J., Tomancak, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Guide+to+light-sheet+microscopy+for+adventurous+biologists.">Guide to light-sheet microscopy for adventurous biologists.</a> Nature Methods. 12, 30-34 (2015).</li></ol>

The authors have nothing to disclose.

Critical steps and troubleshooting for the dissection and culturing of egg chambers
If too many flies are placed into a small vial, the fly food can turn muddy after 2&#8211;3 days due to extensive amounts of feeding larvae and adult flies getting trapped in the fly food. In such a case, flip the rest of these flies into a new vial with fresh food and downsize their number. In particular, exclude females that were stuck in the food.
The Schneider mi...

The continual development of novel imaging and software technologies with applications in the life sciences has provided an enormous impact on understanding the basic principles of life. One of the main challenges is the reliable visualization of developmental processes in combination with their live imaging in various tissues. Tissues are parts of organs and bodies and, as such, the majority are not easily accessible for imaging. Therefore, protocols that allow their dissection and in vitro culturing have been developed in order to visualize biological events that sufficiently reflect the in vivo situation within a living body.
Over the past decades, the culturing and live imaging of Drosophila egg chambers, acinar-like structures resembling primitive organs, has contributed immensely to the understanding of the basic principles of primitive organ development1,2,3. Currently, there are several culturing protocols available, and their usage depends on acquisition time, cell type to be imaged, and their accessibility (e.g., inner germline vs. outer somatic line)4.
A common feature in all these culturing protocols is the need for the immobilization of analyzed egg chambers that display a high contractile activity in liquid media. The contractile activity of egg chambers is caused mainly by the muscle sheet that covers a long string of connected egg chambers5,6,7. Therefore, to achieve proper immobilization of young egg chambers, various approaches have been developed, involving covering egg chambers with coverslips6,8,9 or flexible blankets4,10 or embedding them in low-melting-point agarose3,11. These approaches are popular as they also allow the imaging of a larger visual plane due to the subtle flattening of the circumferential surface of the egg chambers.
However, recently, it has been shown that young egg chambers (stage 1-8) rotate around their anterior-posterior axis6 and that this tissue motion is powered by a fine actomyosin network close to the circumferential surface of these young egg chambers12. Therefore, artificial alteration of the cellular surface caused by a subtle flattening of this tissue may have a negative impact on the behavior of the force-generating actomyosin machinery. The counterpoint is that if the egg chamber tissue is not flattened, microscopic imaging of proteins at the circumferential surface of egg chambers becomes even more limited by the decreased size of the acquisition plane.
Therefore, we have combined protocols from Prasad et al.9 and the lab of Celeste Berg4,10 and further modified them so that no coverslip/flexible blanket/agarose is used in the developed method. Drosophila egg chambers are freely cultured in media and the protocol presented here applies only inverted microscopy. There are two parts to the protocol. The first part is focused on the analysis of actomyosin signals at the local cellular scale (up to 15 cells) within egg chambers. In the second part, we focus on overcoming the limitations associated with a small acquisition plane caused by the free culturing of egg chambers. In this regard, we have developed a novel Fiji-based computational method with a semi-interactive graphical user interface that selectively extracts and unfolds defined layers of a circumferential tissue surface. This is followed by a protocol that describes how to analyze actomyosin at the tissue scale (i.e., &#62;50 cells). As the selective extraction of a defined thin layer of curved epithelial tissues has not been easily possible using a classical z-stack projection (Figure 1), this easy-to-use method serves as an important prerequisite to comprehensively understand&#160;the behavior of a thin (&#60;1 &#181;m) actomyosin network at the tissue scale in Drosophila egg chambers.
In addition, to facilitate the protocol, we provide example time-lapse movies (TLMs) and sample files of fluorescently tagged nonmuscle conventional Myosin II behavior (see Supplementary File 3). Myosin&#160;II is a motor protein and represents the active contractile part of the actomyosin machinery. In order to image Myosin II, we use Drosophila transgenic lines that contain a modified regulatory light chain of Myosin II called MRLC::GFP (see Table of Materials for details)12,13. In order to visualize cell membranes, the protocol is based on commercial dyes (see Table of Materials). This protocol is suitable not only for the analysis of small subcellular MRLC::GFP signals12 but also for any similar-sized subcellular particles around &#177;300 &#181;M, such as those observed with Life-Act::GFP12,14.
Although both these protocols are presented using in vitro cultured Drosophila egg chambers, the acquisition of actomyosin signals can also be performed using other tissues upon the optimization of the culturing media and depending on the availability of either fluorescently tagged proteins with corresponding commercial dyes or, for example, mRNA microinjections to obtain transient gene expression profiles. Similarly, the Fiji-based protocol for the extraction of a thin layer from a circumferential surface can be applied more generally to ellipsoid and organ-like tissues.

<table>
	<tbody>
		<tr>
			<td>Schneider Medium</td>
			<td>Gibco</td>
			<td>21720-024</td>
			<td>[+]-Glutamine</td>
		</tr>
		<tr>
			<td>Penicillin/Streptomycin</td>
			<td>Gibco</td>
			<td>15140-122</td>
			<td>&#160;[+] 10.000 Units/ml Penicillin [+] 10.000 &#181;g/ml Streptomycin</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Sigma</td>
			<td>&#160;F135</td>
			<td>Heat inactivated</td>
		</tr>
		<tr>
			<td>Insulin Solution Human</td>
			<td>Sigma</td>
			<td>&#160;I9278</td>
			<td></td>
		</tr>
		<tr>
			<td>50 ml Falcon tubes</td>
			<td>Eppendorf</td>
			<td></td>
			<td>sterile</td>
		</tr>
		<tr>
			<td>Millex-GV filter</td>
			<td>Millipore</td>
			<td>SLGV033NS</td>
			<td>33 mm</td>
		</tr>
		<tr>
			<td>Glass Bottom Microwell Dishes</td>
			<td>MatTek Corporation</td>
			<td>P35G-1.5-14-C</td>
			<td>35 mm Petri dish, 14 mm Microwell, No. 1.5 cover glass</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>A. Dumont &#38; Fils</td>
			<td></td>
			<td>&#160;#55</td>
		</tr>
		<tr>
			<td>CellMask Deep Red (cell membrane dye)</td>
			<td>Invitrogen</td>
			<td>C10046</td>
			<td></td>
		</tr>
		<tr>
			<td>FM4-64 (cell membrane dye)</td>
			<td>ThermoFisher/Invitrogen</td>
			<td>T13320</td>
			<td></td>
		</tr>
		<tr>
			<td>Depression dissection slide</td>
			<td>Fisher Scientific</td>
			<td>12-560B</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscopic equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Steromicroscope</td>
			<td>e.g. Zeiss</td>
			<td></td>
			<td>Stemi SV 6</td>
		</tr>
		<tr>
			<td>Inverted confocal microscope</td>
			<td>e.g. Zeiss/Olympus</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Spinning disc microscope</td>
			<td>e.g. Zeiss/Andor</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microscopic glass/cover glass</td>
			<td>e.g. Thermo Scientific/Menzel Glas</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cactus tool</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Wironit needle holder</td>
			<td>Hammacher</td>
			<td>9160020</td>
			<td></td>
		</tr>
		<tr>
			<td>Cactus spine</td>
			<td>Echinocactus grusonii/barrel cactus</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Drosophila stocks used in the manuscript</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>AX3/AX3; sqh-MRLC::GFP/sqh-MRLC::GFP</td>
			<td></td>
			<td></td>
			<td>For detail see Rauzi et al.: Planar polarized actomyosin contractile flows control epithelial junction remodeling. Nature 2010, Vol. 468, pg. 1110-1115.</td>
		</tr>
		<tr>
			<td>AX3/AX3; sqh-MRLC::GFP/sqh-MRLC::GFP; fat258D/fat103C</td>
			<td></td>
			<td></td>
			<td>For detail see Viktorinova et al.: Epithelial rotation is preceded by planar symmetry breaking of actomyosin and protects epithelial tissue from cell deformations. PLoS Genetics 2017, Vol. 13, Issue 11, pg. e1007107.</td>
		</tr>
	</tbody>
</table>

analysis of actomyosin dynamics at local cellular and tissue scales using time-lapse movies of cultured drosophila egg chambers

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ellipsoid shape during development. This developmental process is called oogenesis and is crucial to generating functional mature eggs to secure the next fly generation. For these reasons, egg chambers have served as an ideal and relevant model to understand animal organ development.
Several in vitro culturing protocols have been developed, but there are several disadvantages to these protocols. One involves the application of various covers that exert an artificial pressure on the imaged egg chambers in order to immobilize them and to increase the imaged acquisition plane of the circumferential surface of the analyzed egg chambers. Such an approach may negatively influence the behavior of the thin actomyosin machinery that generates the power to rotate egg chambers around their longer axis.
Thus, to overcome this limitation, we culture Drosophila egg chambers freely in the media in order to reliably analyze actomyosin machinery along the circumference of egg chambers. In the first part of the protocol, we provide a manual detailing how to analyze the actomyosin machinery in a limited acquisition plane at the local cellular scale (up to 15 cells). In the second part of the protocol, we provide users with a new Fiji-based plugin that allows the simple extraction of a defined thin layer of the egg chambers&#8217; circumferential surface. The following protocol then describes how to analyze actomyosin signals at the tissue scale (&#62;50 cells). Finally, we pinpoint the limitations of these approaches at both the local cellular and tissue scales and discuss its potential future development and possible applications.

This protocol enables scientists to investigate the behavior of actomyosin networks in epithelial tissues. This is only possible when a detailed analysis of actomyosin behavior at the local cellular scale (a few cells) is combined with a similar analysis at the tissue scale (many cells). However, epithelial tissues are often curved and the extraction of a thin layer of these tissues was previously not easily possible, as shown in Drosophila egg chambers (Figu...

Watch this Scientific Journal Video about Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers at JoVE.com

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

This protocol provides a Fiji-based, user-friendly methodology along with straightforward instructions explaining how to reliably analyze actomyosin behavior in individual cells and curved epithelial tissues. No programming skills are required to follow the tutorial; all steps are performed in a semi-interactive manner using the graphical user interface of Fiji and associated plugins.

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ...

analysis-actomyosin-dynamics-at-local-cellular-tissue-scales-using

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Developmental Biology

7.4K Views.  Max Planck Institute of Molecular Cell Biology and Genetics. This protocol provides a Fiji-based, user-friendly methodology along with straightforward instructions explaining how to reliably analyze actomyosin behavior in individual cells and curved epithelial tissues. No programming skills are required to follow the tutorial; all steps are performed in a semi-interactive manner using the graphical user interface of Fiji and associated plugins.

Video: Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

Upright Imaging of Drosophila Egg Chambers

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in architecture, is well-characterized, and is amenable to genetic analysis. Each egg chamber consists of germ-line cells surrounded by a single epithelial layer of somatic follicle cells. Subsets of follicle cells undergo differentiation during specific stages to become several different cell types. Standard techniques primarily allow for a lateral view of egg chambers, and therefore a limited view of follicle cell organization and identity. The upright imaging protocol describes a mounting technique that enables a novel, vertical view of egg chambers with a standard confocal microscope. Samples are first mounted between two layers of glycerin jelly in a lateral (horizontal) position on a glass microscope slide. The jelly with encased egg chambers is then cut into blocks, transferred to a coverslip, and flipped to position egg chambers upright. Mounted egg chambers can be imaged on either an upright or an inverted confocal microscope. This technique enables the study of follicle cell specification, organization, molecular markers, and egg development with new detail and from a new perspective.

Upright Imaging of Drosophila Egg Chambers

The upright imaging method described in this protocol allows for the detailed visualization of the poles of a developing Drosophila melanogaster egg. This end-on view provides a new perspective into the arrangements and morphologies of multiple cell types in the follicular epithelium.

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in ...

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two layers of skin that enclose undifferentiated mesenchyme, and because the larval tail fin regenerates rapidly within 2-3 days. Using this system, we demonstrate a method for capturing the repair dynamics of the amputated tail fin with time-lapse video brightfield stereomicroscopy. We demonstrate that fin amputation triggers a contraction of the amputation wound and extrusion of cells around the wound margin, leading to their subsequent clearance. Fin regeneration proceeds from proximal to distal direction after a short delay. In addition, developmental growth of the larva can be observed during all stages. The presented method provides an opportunity for observing and analyzing whole tissue-scale behaviors such as fin development and growth in a simple microscope setting, which is easily adaptable to any stereomicroscope with time-lapse capabilities.

We present a protocol for capturing the dynamics of zebrafish larval tail fin regeneration on a whole-tissue scale using brightfield-based stereomicroscopy. This technique enables capturing the regeneration dynamics with single cell resolution. This methodology can be adapted to any stereomicroscope equipped with a CCD camera and time-lapse software.

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two ...

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

Imaging Calcium in Drosophila at Egg Activation

Egg activation is a universal process that includes a series of events to allow the fertilized egg to complete meiosis and initiate embryonic development. One aspect of egg activation, conserved across all organisms examined, is a change in the intracellular concentration of calcium (Ca2+) often termed a 'Ca2+ wave'. While the speed and number of oscillations of the Ca2+ wave varies between species, the change in intracellular Ca2+ is key in bringing about essential events for embryonic development. These changes include resumption of the cell cycle, mRNA regulation, cortical granule exocytosis, and rearrangement of the cytoskeleton.
In the mature Drosophila egg, activation occurs in the female oviduct prior to fertilization, initiating a series of Ca2+-dependent events. Here we present a protocol for imaging the Ca2+ wave in Drosophila. This approach provides a manipulable model system to interrogate the mechanism of the Ca2+ wave and the downstream changes associated with it.

Imaging Calcium in Drosophila at Egg Activation

This article describes an adaptable ex vivo protocol for visualizing Ca2+ during egg activation in Drosophila.

Egg activation is a universal process that includes a series of events to allow the fertilized egg to complete meiosis and initiate embryonic development. ...

An Ex Vivo Method for Time-Lapse Imaging of Cultured Rat Mesenteric Microvascular Networks

Angiogenesis, defined as the growth of new blood vessels from pre-existing vessels, involves endothelial cells, pericytes, smooth muscle cells, immune cells, and the coordination with lymphatic vessels and nerves. The multi-cell, multi-system interactions necessitate the investigation of angiogenesis in a physiologically relevant environment. Thus, while the use of in vitro cell-culture models have provided mechanistic insights, a common critique is that they do not recapitulate the complexity associated with a microvascular network. The objective of this protocol is to demonstrate the ability to make time-lapse comparisons of intact microvascular networks before and after angiogenesis stimulation in cultured rat mesentery tissues. Cultured tissues contain microvascular networks that maintain their hierarchy. Immunohistochemical labeling confirms the presence of endothelial cells, smooth muscle cells, pericytes, blood vessels and lymphatic vessels. In addition, labeling tissues with BSI-lectin enables time-lapse comparison of local network regions before and after serum or growth factor stimulation characterized by increased capillary sprouting and vessel density. In comparison to common cell culture models, this method provides a tool for endothelial cell lineage studies and tissue specific angiogenic drug evaluation in physiologically relevant microvascular networks.

An Ex Vivo Method for Time-Lapse Imaging of Cultured Rat Mesenteric Microvascular Networks

Angiogenesis involves multi-cell, multi-system interactions that need to be investigated in a physiologically relevant environment. The objective of this study is to demonstrate the ability of the rat mesentery culture model to make time-lapse comparisons of intact microvascular networks during angiogenesis.

Angiogenesis, defined as the growth of new blood vessels from pre-existing vessels, involves endothelial cells, pericytes, smooth muscle cells, immune ...

Kinetic Analysis of Vasculogenesis Quantifies Dynamics of Vasculogenesis and Angiogenesis In Vitro

Vasculogenesis is a complex process by which endothelial stem and progenitor cells undergo de novo vessel formation. Quantitative assessment of vasculogenesis has become a central readout of endothelial progenitor cell functionality, and therefore, several attempts have been made to improve both in vitro and in vivo vasculogenesis models. However, standard methods are limited in scope, with static measurements failing to capture many aspects of this highly dynamic process. Therefore, the goal of developing this novel protocol was to assess the kinetics of in vitro vasculogenesis in order to quantitate rates of network formation and stabilization, as well as provide insight into potential mechanisms underlying vascular dysfunction. Application of this protocol is demonstrated using fetal endothelial colony forming cells (ECFCs) exposed to maternal diabetes mellitus. Fetal ECFCs were derived from umbilical cord blood following birth, cultured, and plated in slides containing basement membrane matrix, where they underwent vasculogenesis. Images of the entire slide wells were acquired using time-lapse phase contrast microscopy over 15 hours. Images were analyzed for derivation of quantitative data using an analysis software called Kinetic Analysis of Vasculogenesis (KAV). KAV uses image segmentation followed by skeletonization to analyze network components from stacks of multi-time point phase contrast images to derive ten parameters (9 measured, 1 calculated) of network structure including: closed networks, network areas, nodes, branches, total branch length, average branch length, triple-branched nodes, quad-branched nodes, network structures, and the branch to node ratio. Application of this protocol identified altered rates of vasculogenesis in ECFCs obtained from pregnancies complicated by diabetes mellitus. However, this technique has broad implications beyond the scope reported here. Implementation of this approach will enhance mechanistic assessment and improve functional readouts of vasculogenesis and other biologically important branching processes in numerous cell types or disease states.

Kinetic Analysis of Vasculogenesis Quantifies Dynamics of Vasculogenesis and Angiogenesis In Vitro

Here, we present a protocol for time-lapse imaging and analysis of vasculogenesis in vitro using phase contrast microscopy coupled with the open source software, Kinetic Analysis of Vasculogenesis. This protocol can be applied to quantitatively assess the vasculogenic potential of numerous cell types or experimental conditions that model vascular disease.

Vasculogenesis is a complex process by which endothelial stem and progenitor cells undergo de novo vessel formation. Quantitative assessment of ...

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

Muscles together with tendons and the skeleton enable animals including humans to move their body parts. Muscle morphogenesis is highly conserved from animals to humans. Therefore, the powerful Drosophila model system can be used to study concepts of muscle-tendon development that can also be applied to human muscle biology. Here, we describe in detail how morphogenesis of the adult muscle-tendon system can be easily imaged in living, developing Drosophila pupae. Hence, the method allows investigating proteins, cells and tissues in their physiological environment. In addition to a step-by-step protocol with helpful tips, we provide a comprehensive overview of fluorescently tagged marker proteins that are suitable for studying the muscle-tendon system. To highlight the versatile applications of the protocol, we show example movies ranging from visualization of long-term morphogenetic events &#8211; occurring on the time scale of hours and days &#8211; to visualization of short-term dynamic processes like muscle twitching occurring on time scale of seconds. Taken together, this protocol should enable the reader to design and perform live-imaging experiments for investigating muscle-tendon morphogenesis in the intact organism.

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

Here, we present an easy-to-use and versatile method to perform live imaging of developmental processes in general and muscle-tendon morphogenesis in particular in living Drosophila pupae.

Muscles together with tendons and the skeleton enable animals including humans to move their body parts. Muscle morphogenesis is highly conserved from ...

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A remarkable example is given by sensory epithelia, where cells of the same or distinct identities are brought together via cell-cell adhesion showing highly organized planar patterns. Cells align to one another in the same direction and display equivalent polarity over large distances. This organization of the mature epithelia is established over the course of morphogenesis. To understand how the planar arrangement of the mature epithelia is achieved, it is crucial to track cell orientation and growth dynamics with high spatiotemporal fidelity during development in vivo. Robust analytical tools are also essential to identify and characterize local-to-global transitions. The Drosophila pupa is an ideal system to evaluate oriented cell shape changes underlying epithelial morphogenesis. The pupal developing epithelium constitutes the external surface of the immobile body, allowing long-term imaging of intact animals. The protocol described here is designed to image and analyze cell behaviors at both global and local levels in the pupal abdominal epidermis as it grows. The methodology described can be easily adapted to the imaging of cell behaviors at other developmental stages, tissues, subcellular structures, or model organisms.

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

This protocol is designed for the imaging and analysis of the dynamics of cell orientation and tissue growth in the Drosophila abdominal epithelia as the fruit fly undergoes metamorphosis. The methodology described here can be applied to the study of different developmental stages, tissues, and subcellular structures in Drosophila or other model organisms.

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A ...

Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes

We describe a technique for retrograde labeling of motor neurons in Drosophila. We use an oil-dissolved lipophilic dye and deliver a small droplet to an embryonic fillet preparation by a microinjector. Each motor neuron whose membrane is contacted by the droplet can then be rapidly labeled. Individual motor neurons are continuously labeled, enabling fine structural details to be clearly visualized. Given that lipophilic dyes come in various colors, the technique also provides a means to get adjacent neurons labeled in multicolor. This tracing technique is therefore useful for studying neuronal morphogenesis and synaptic connectivity in the motor neuron system of Drosophila.

Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes

We describe a method for retrograde tracing of the Drosophila embryonic motor neurons using lipophilic fluorescent dyes.

We describe a technique for retrograde labeling of motor neurons in Drosophila. We use an oil-dissolved lipophilic dye and deliver a small droplet to an ...

Visualizing the Effects of Oxidative Damage on Drosophila Egg Chambers using Live Imaging

Live imaging of Drosophila melanogaster ovaries has been instrumental in understanding a variety of basic cellular processes during development, including ribonucleoprotein particle movement, mRNA localization, organelle movement, and cytoskeletal dynamics. There are several methods for live imaging that have been developed. Due to the fact that each method involves dissecting individual ovarioles placed in media or halocarbon oil, cellular damage due to hypoxia and/or physical manipulation will inevitably occur over time. One downstream effect of hypoxia is to increase oxidative damage in the cells. The purpose of this protocol is to use live imaging to visualize the effects of oxidative damage on the localization and dynamics of subcellular structures in Drosophila ovaries after induction of controlled cellular damage. Here, we use hydrogen peroxide to induce cellular oxidative damage and give examples of the effects of such damage on two subcellular structures, mitochondria and Clu bliss particles. However, this method is applicable to any subcellular structure. The limitations are that hydrogen peroxide can only be added to aqueous media and would not work for imaging that uses halocarbon oil. The advantages are that hydrogen peroxide is readily available and inexpensive, acts quickly, its concentrations can be modulated, and oxidative damage is a good approximation of damage caused by hypoxia as well as general tissue damage due to manipulation.

The objective of this protocol is to use live imaging to visualize the effects of oxidative damage on the localization and dynamics of subcellular structures in Drosophila ovaries.

Live imaging of Drosophila melanogaster ovaries has been instrumental in understanding a variety of basic cellular processes during development, including ...