UD, WFM and BR are supported by the Center for Cellular Construction, a National Science Foundation (NSF) Science and Technology Center, under grant agreement DBI-1548297. BR is also supported through an NSF CAREER award, 1553695.

<ol>
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(2001).</li><li>Voeltz, G., Prinz, W., Shibata, Y., Rist, J., Rapoport, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+class+of+membrane+proteins+shaping+the+tubular+endoplasmic+reticulum.">A class of membrane proteins shaping the tubular endoplasmic reticulum.</a> Cell. 124 (3), 573-586 (2006).</li><li>Zurek, N., Sparks, L., Voeltz, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reticulon+short+hairpin+transmembrane+domains+are+used+to+shape+ER+tubules.">Reticulon short hairpin transmembrane domains are used to shape ER tubules.</a> Traffic. 12 (1), 28-41 (2011).</li><li>Kumar, D., Golchoubian, B., Belevich, I., Jokitalo, E., Schlaitz, A. 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The authors have nothing to disclose.

The protocol described here is highly versatile and easily adaptable for studying biological events in a variety of stages in Drosophila embryonic development. This protocol is well suited for studying biological processes that occur over long periods of time. For example, studies of cellularization29,30 or the formation of heterochromatin domains in Drosophila31,32 could benefit from utilizing the methods described in this protocol since these processes occur over an extended period of time. In addition, sample size can be increased by decreasing magnification to study questions that require less optical resolution, such as the timing between syncytial divisions33 or duration of gastrulation34. &#160;Likewise, a 20x objective allows for two embryos to be imaged within one imaging plane. The methods in this protocol provide a dynamic platform to ask biological questions using embryonic development as a model system for quantitative analysis.
There are 3 critical steps in this protocol. Post dechorionation in 50% bleach it is imperative that embryos are rinsed and dried vigorously 5 times (2.3.4). This step removes any leftover small pieces of chorion. Leftover chorion will obstruct the field of view when imaging the embryo. When mounting the embryos onto the glass coverslip (3.5), it is important to press the embryos onto the glue with uniform pressure. This will place the embryos on the same z plane. When cutting open the microinjection needle, it is important to retract the plunger immediately to adjust the flow rate of the needle or you will lose all your injectant.
Our study was limited by two factors. The first factor was not having an automated segmentation process. We designed a manual segmentation script using MATLAB, however; with a genetically encoded spindle marker this process could be automated. In the future we would like to produce CNN-RFP21, Rtln1-GFP and CNN-RFP; ReepB-GFP transgenic lines to enable automated spindle segmentation. For more information regarding our MATLAB script and analysis pipeline, refer to supplemental materials of Diaz et al.15. Secondly, we were also limited by our acquisition interval of 35 s, which only allowed us to image 6 embryos simultaneously. A longer acquisition interval would allow us to image more embryos simultaneously. Although drug delivery throughput can be increase by replacing microinjection with a permeation step, we found this to be unnecessary since our sample size was already limited by acquisition interval.
While this protocol was used to image Drosophila embryonic development, the overall approach is adaptable for studying embryonic development in other multicellular model systems, such as C. elegans, Sea Urchins, and Xenopus. Embryos are extremely useful for quantitative analysis since they are easily synchronized via collection or fertilization. In addition, embryos don&#8217;t move or swim away from a field of view, enabling the use of a simple multipoint acquisition capable software to produce multiple time lapse videos for quantitative analysis. Although we used a high content analyzer in our study, any inverted microscope system capable of multipoint acquisition can be paired with the methods referred to herein.
Similar results can be obtained using a multipoint acquisition capable inverted confocal microscope; however, the advantages of a high content analyzer emerge as one increases sample size to study questions that require less resolution in time. An obvious advantage to using a high content analyzer is the ability to image 4 separate slides simultaneously compared to 1 slide on traditional systems. With a full set of 4 slides and 40 embryos on each slide, 160 embryos can be imaged over several hours, so long as the acquisition interval is at least 15 min between frames. Another important advantage to using a high content analyzer is that the samples are completely sealed off from any external light sources, which is necessary for fluorescence intensity-based measurements. Lastly, the high content analyzer used in our study has a 5.5-megapixel CMOS camera which provides a generous 221.87 &#956;m field of view at 60x magnification. This larger field of view enabled us to image half an embryo per acquisition point.&#160;&#160;

Over the past 30 years, advances in imaging technologies and molecular probes for live cell imaging have advanced our understanding of the complexity and inner workings of the cell. Accompanying this headway in technology is a greater reliance on the use of ex vivo cell culture models for the acquisition of high throughput imaging data. Cell culture models provide several advantages, including control of the microenvironment through microfluidic systems, and the ability to image multiple cells simultaneously, enabling the use of quantitative approaches necessary for high-throughput screening1,2. However, there are also significant disadvantages associated with cell culture models in the context of morphological studies, such as two-dimensional glass or plastic surfaces producing confounding effects on cell morphology and its regulation3,4,5. These effects can provide false or misleading data with regards to the study of biological processes regulating cellular morphology.
Although the alternative has been to study cellular morphology in the context of an intact organism6, few suitable intact organisms facilitate high throughput quantitative live cell imaging due to difficulties in keeping whole organisms alive through out imaging, and challenges associated with imaging inside a large multicellular organism. As a result, cell biological studies in intact organisms have generally focused on observing a single organism over time, which greatly limits sample size. In addition, this one-animal-at-a-time limitation makes live imaging difficult and costly to employ in the form of a screen and restricts the ability to apply quantitative analytical tools since the number of samples is generally too small. Thus, arises the need for a multicellular model organism that can be used for high content based quantitative approaches.
This protocol adapts the Drosophila embryo for plate-based high content imaging to generate experimental sample sizes large enough for quantitative analysis. Drosophila melanogaster is commonly known as the first model organism. Research using Drosophila has led to breakthroughs in cell and molecular biology, including the transmission of genetic information, identification of cell signaling pathways, and genetic requirements of embryonic development7,8,9. In addition, the Drosophila embryo has been used to explore in vivo microtubule dynamics during cellular division10,11,12. The use of microinjection to deliver small molecules and molecular probes provides temporal control that makes Drosophila embryonic development particularly powerful for probing cellular processes in vivo13,14. However, generating a significant number of experimental repeats with a traditional imaging approach is extremely labor intensive and has limited use in high-throughput imaging screens and quantitative analysis. Our approach makes use of a high content analyzer typically reserved for single cell analysis and adapts in vivo imaging analysis in the syncytium of the early Drosophila embryo.
This protocol was used to collect, stage, and microinject Drosophila embryos to explore the mechanisms that drive morphological changes in the Endoplasmic Reticulum (ER) during mitosis15. Although many studies have outlined the dynamic nature of the ER during the cell cycle16,17,18,19, it has been difficult to compare the effects of multiple perturbations across time on ER morphology. To identify the components that drive changes in ER morphology across cell division, the methods listed in this protocol were used to probe the role of microtubules and other cytoskeletal factors on mitotic ER reorganization using the cortical syncytial division in the early Drosophila embryo. Taken together, the availability of genetic perturbation within the Drosophila community, the ability to microinject drugs and molecular probes at precise times during development, and the inexpensive cost of working with Drosophila makes this approach highly amenable to high-throughput image-based screening. The methods described here are applicable for studying a wide variety of cellular and developmental processes in vivo using the Drosophila embryo20. Specifically, this protocol is well adapted to study biological events that occur over an extended period and within 100 microns of the Drosophila embryo cortex.

<table border="0" cellpadding="0" cellspacing="0" style="width:860px;" width="859">
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			<td height="21" style="height:21px;width:194px;">10 x 35mm Micro Dish</td>
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			<td>N/A</td>
			<td>N/A</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:194px;">Halocarbon 27 Oil</td>
			<td style="width:314px;">Genesee Scientific</td>
			<td>Cat #: 59-133</td>
			<td>N/A</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:194px;">Halocarbon 700 Oil</td>
			<td style="width:314px;">Genesee Scientific</td>
			<td>Cat #: 59-131</td>
			<td>N/A</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:194px;">Heptane Glue</td>
			<td style="width:314px;">REFFER TO RECIPE PREP</td>
			<td>N/A</td>
			<td>N/A</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:194px;">Industrial Razor Blades (Surgical Carbon Steel) Single Edged No. 9</td>
			<td style="width:314px;">VWR</td>
			<td>Cat #: 55411-050</td>
			<td>N/A</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:194px;">Parafilm M Laboratory Sealing Film, 4 inch x 250 foot Roll</td>
			<td style="width:314px;">Fisher Scientific</td>
			<td>Cat #: 50-998-944</td>
			<td>N/A</td>
		</tr>
		<tr height="100">
			<td height="100" style="height:100px;width:194px;">Scotch Double Sided Tape with Dispenser, Narrow Width, Engineered for Bonding, 1/2&#34; x 7 yds., 3/Pack Caddy</td>
			<td style="width:314px;">Fisher Scientific</td>
			<td>Cat #: NC0879005</td>
			<td>N/A</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:194px;">Silica gel beads, desiccant ~ 2 - 5 mm</td>
			<td style="width:314px;">Millipore Sigma</td>
			<td>SKU: 1077351000</td>
			<td>N/A</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:194px;">Stainless Steel Spoon/Spatula</td>
			<td style="width:314px;">VWR</td>
			<td>Cat #: 470149-044</td>
			<td>N/A</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:194px;">W.A. Hammond Drierite Indicating Absorbents</td>
			<td style="width:314px;">Fisher Scientific</td>
			<td>Cat #: 23-116582</td>
			<td>N/A</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:194px;">Yeast Paste</td>
			<td style="width:314px;">REFFER TO RECIPE PREP</td>
			<td>N/A</td>
			<td>N/A</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:194px;">EQUIPMENT</td>
			<td style="width:314px;"></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:194px;">Allegra X-12 Benchtop, Centrifuge</td>
			<td style="width:314px;">Beckman Coulter</td>
			<td>N/A</td>
			<td style="width:206px;">You will need access to 15 ml conical centrifuge inserts.</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:194px;">Autoclave</td>
			<td style="width:314px;">N/A</td>
			<td>N/A</td>
			<td>Any autoclave will do fine.</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:194px;">Drummond Nanoject II Microinjector.</td>
			<td style="width:314px;">Drummond Scientific</td>
			<td>N/A</td>
			<td style="width:206px;">This is a mineral oil based injection system.</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:194px;">GE Healthcare IN Cell Analyzer 6000.</td>
			<td style="width:314px;">GE healthcare</td>
			<td>N/A</td>
			<td style="width:206px;">A fast confocal microscope capable of multipoint acquisition may be substituted.</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:194px;">Refrigerator (4C&#176;)</td>
			<td style="width:314px;">N/A</td>
			<td>N/A</td>
			<td style="width:206px;">Fly media and yeast paste must be stored at (4C&#176;).</td>
		</tr>
		<tr height="106">
			<td height="106" style="height:106px;width:194px;">Zies Stemi 508 Stereo Microscope with Transillumination base 300 and overhead gooseneck lighting.</td>
			<td style="width:314px;">Zeiss Microscopy</td>
			<td>N/A</td>
			<td style="width:206px;">The transillumination base and gooseneck lighting are necessary.</td>
		</tr>
	</tbody>
</table>

drosophila embryo preparation and microinjection for live cell microscopy performed using an automated high content analyzer

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and computational modeling to classify and compare biological processes. Although cell culture model systems are great for high content imaging, high throughput studies of cell morphology suggest that ex vivo cultures are limited in recapitulating the morphological complexity found in cells within living organisms. As such, there is a need for a scalable high throughput model system to image living cells within an intact organism. Described here is a protocol for using a high content image analyzer to simultaneously acquire multiple time-lapse videos of embryonic Drosophila melanogaster development during the syncytial blastoderm stage. The syncytial blastoderm has traditionally served as a great in vivo model for imaging biological events; however, obtaining a significant number of experimental replicates for quantitative and high-throughput approaches has been labor intensive and limited by the imaging of a single embryo per experimental repeat. Presented here is a method to adapt imaging and microinjection approaches to suit a high content imaging system, or any inverted microscope capable of automated multipoint acquisition. This approach enables the simultaneous acquisition of 6-12 embryos, depending on desired acquisition factors, within a single imaging session.

The approach presented in this protocol was used to examine the role of microtubules in Endoplasmic Reticulum (ER) reorganization during mitosis in the Drosophila embryo15. During mitosis, the structure of the ER displays a dramatic reorganization, however, the forces that drive these changes are poorly understood. Recently, a family of ER shaping proteins were shown to promote the formation of ER tubule25,26,27,28, which are known for their close proximity with microtubules28. To study the roll of microtubules on ER morphology, the methods provided in the protocol were used to generate data to quantitatively compare the effects of several microinjected drug treatments on ER reorganization during mitosis. Colchicine, which prevents new microtubule polymerization, was found to drastically reduce the reorganization of the ER during mitosis as shown in Figure 5 and Figure 6. Furthermore, the approach described here enabled the production of time laps imaging data for 32 embryos. Mean and max intensity measurements were produced from 12,800 regions of interest using a custom MATLAB script, which also generated descriptive and inferential statistics vital for making comparisons between drug treatments. Due to the availability of molecular probes and the ease of microinjections, the methods described here are easily adaptable to study a variety of biological processes via fluorescence intensity quantification.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61589/61589fig5.jpg" /> 
Figure 5: Representative Images of Rtnl1-GFP and ReepB-GFP enrichment at the spindle poles during mitosis. (A) 60x field of view of ReepB-GFP during mitosis in cell cycle 10. The blue square represents the field of view used for panels B-E. Panel A is processed with a threshold binary filter. This filter was not applied to Panels B-E since it excludes data from pixels below the set threshold. (B,D) ReepB-GFP in control, and in colchicine. (C,E) Rtnl1-GFP in control, and in colchicine. This figure has been modified from Diaz et al.15. <a href="https://www.jove.com/files/ftp_upload/61589/61589fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61589/61589fig6.jpg" /> 
Figure 6: Quantitation of Rtnl1-GFP and ReepB-GFP enrichment at the spindle poles during mitosis. 20 spindles and 20 cytoplasm ROIs across 10 frames were analyzed for 32 embryos in the following conditions. (A) 10% DMSO control injections for Rtnl1-GFP. Control embryos showed a 0.1-fold increase in enrichment (p&#60;0.001 for all samples at T210) compared to non-injected samples (p &#60; 0.001 for all samples at T210). (B) ReepB-GFP embryos injected with 10% DMSO to serve as control. Control embryos showed a 0.1-fold increase in enrichment (p&#60;0.001 for all samples at T210) compared to non-injected samples (Figure 1E; p &#60; 0.001 for all samples at T210). (C) Rtnl1-GFP embryos injected with 5 &#956;M Colchicine. Here we measured a reduction in Rtnl1-GFP enrichment to spindles (p &#60; 0.001 for all samples at T210) compared to controls (A) (p &#60; 0.001 for all samples at T210). (D) ReepB-GFP embryos injected with 5 &#956;M Colchicine. We measured a reduction in ReepB-GFP enrichment to spindles (p&#60;0.001 for all samples at T210) compared to controls (B) (p&#60;0.001 for all samples at T210). This figure has been modified from Diaz et al.15. <a href="https://www.jove.com/files/ftp_upload/61589/61589fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>1. Yeast Paste21</td>
			<td>2. Grape Plates21</td>
			<td>3. Heptane Glue24</td>
		</tr>
		<tr>
			<td>1.1 Add 7 grams of active dry yeast to a 50 ml conical. Add 9 milliliters of DI water and mix to even constancy with a metal spatula.</td>
			<td>2.1 Make solution a: Add 6 grams of agar to 140ml of DI water in a 250ml flask and autoclave it.</td>
			<td>3.1 Fill a glass scintillation vial with 50 inches of double-sided tape.</td>
		</tr>
		<tr>
			<td>1.1 Add 7 grams of active dry yeast to a 50 ml conical. Add 9 milliliters of DI water and mix to even constancy with a metal spatula.</td>
			<td>2.2 Make solution b: Mix 0.1 grams of methyl paraben in 2ml of ethanol, then add this solution to 60ml of grapefruit juice concentrate.</td>
			<td>3.2 Add 20 ml of heptane and seal glass scintillation vial with parafilm. Rotate overnight at room temperature.</td>
		</tr>
		<tr>
			<td></td>
			<td>2.3 Immediately after autoclaving solution a, mix in solution b and pour mixture into 10 x 35mm perti dishes. (makes 35 grape plates)</td>
			<td>3.3 Remove plastic tape debris by transferring glue into 2 15 ml conical tubes and centrifuging at 3000 RPM for 15 minutes.</td>
		</tr>
		<tr>
			<td></td>
			<td>2.4 Allow grape plates to set at room temperature overnight with plate lids off. Cover plates with lids and store in 4C&#176; after they set.</td>
			<td>3.4 Transfer glue supernatant into 1.5 ml microcentrifuge tubes for storage.</td>
		</tr>
	</tbody>
</table>
Table 1: Recipes of media and solutions.

Watch this Scientific Journal Video about Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer at JoVE.com

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

Presented here is a protocol to microinject and simultaneously image multiple Drosophila embryos during embryonic development using a plate-based, high content imager.

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and ...

drosophila-embryo-preparation-microinjection-for-live-cell-microscopy

Research

JoVE Journal

Developmental Biology

8.4K Views.  San Francisco State University. Presented here is a protocol to microinject and simultaneously image multiple Drosophila embryos during embryonic development using a plate-based, high content imager.

Video: Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance to man of conventional invertebrate model systems and the complexity of genomes in vertebrates are critical challenges to modeling human normal and pathological conditions. The chordate Ciona intestinalis is an invertebrate chordate that emerged from a common ancestor with the vertebrates and may represent features at the interface between invertebrates and vertebrates. A common body plan with much simpler cellular and genomic composition should unveil gene regulatory network (GRN) links and functional genomics readouts explaining phenomena in the vertebrate condition. The compact genome of Ciona, a fixed embryonic lineage with few divisions and large cells, combined with versatile community tools foster efficient gene functional analyses in this organism. Here, we present several crucial methods for this promising model organism, which belongs to the closest sister group to vertebrates. We present protocols for transient transgenesis by electroporation, along with microinjection-mediated gene knockdown, which together provide the means to study gene function and genomic regulatory elements. We extend our protocols to provide information on how community databases are utilized for in silico design of gene regulatory or gene functional experiments. An example study demonstrates how novel information can be gained on the interplay, and its quantification, of selected neural factors conserved between Ciona and man. Furthermore, we show examples of differential subcellular localization in embryonic cells, following DNA electroporation in Ciona zygotes. Finally, we discuss the potential of these protocols to be adapted for tissue specific gene interference with emerging gene editing methods. The in vivo approaches in Ciona overcome major shortcomings of classical model organisms in the quest of unraveling conserved mechanisms in the chordate developmental program, relevant to stem cell research, drug discovery, and subsequent clinical application.

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

We present transient transgenesis and gene knockdown in Ciona intestinalis, a chordate sister group to vertebrates, using microinjection and electroporation techniques. Such methods facilitate functional genomics in this simple invertebrate that features rudimentary characteristics of vertebrates, including notochord and head sensory epithelia, and many orthologs of human disease associated genes.

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance ...

Medium-scale Preparation of Drosophila Embryo Extracts for Proteomic Experiments

Analysis of protein-protein interactions (PPIs) has become an indispensable approach to study biological processes and mechanisms, such as cell signaling, organism development, and disease. It is often desirable to obtain PPI information using in vivo material, to gain the most natural and unbiased view of the interaction networks. The fruit fly Drosophila melanogaster is an excellent platform to study PPIs in vivo, and lends itself to straightforward approaches to isolating material for biochemical experiments. In particular, fruit fly embryos represent a convenient type of tissue to study PPIs, due to the ease of collecting animals at this developmental stage and the fact that the majority of proteins are expressed in embryogenesis, thus providing a relevant environment to reveal most PPIs. Here we present a protocol for collection of Drosophila embryos at medium scale (0.5-1 g), which is an ideal amount for a wide range of proteomic applications, including analysis of PPIs by affinity purification-mass spectrometry (AP-MS). We describe our designs for 1 L and 5 L cages for embryo collections that can be easily and inexpensively set up in any laboratory. We also provide a general protocol for embryo collection and protein extraction to generate lysates that can be directly used in downstream applications such as AP-MS. Our goal is to provide an accessible means for all researchers to carry out the analyses of PPIs in vivo.

Medium-scale Preparation of Drosophila Embryo Extracts for Proteomic Experiments

The goal of this protocol is to provide a straightforward and inexpensive approach to collecting Drosophila embryos at medium scale (0.5-1 g) and preparing protein extracts that can be used in downstream proteomic applications, such as affinity purification-mass spectrometry (AP-MS).

Analysis of protein-protein interactions (PPIs) has become an indispensable approach to study biological processes and mechanisms, such as cell signaling, ...

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; ...

High-resolution Episcopic Microscopy (HREM) - Simple and Robust Protocols for Processing and Visualizing Organic Materials

We provide simple protocols for generating digital volume data with the high-resolution episcopic microscopy (HREM) method. HREM is capable of imaging organic materials with volumes up to 5 x 5 x 7 mm3 in typical numeric resolutions between 1 x 1 x 1 and 5 x 5 x 5 &#181;m3. Specimens are embedded in methacrylate resin and sectioned on a microtome. After each section an image of the block surface is captured with a digital video camera that sits on the phototube connected to the compound microscope head. The optical axis passes through a green fluorescent protein (GFP) filter cube and is aligned with a position, at which the bock holder arm comes to rest after each section. In this way, a series of inherently aligned digital images, displaying subsequent block surfaces are produced. Loading such an image series in three-dimensional (3D) visualization software facilitates the immediate conversion to digital volume data, which permit virtual sectioning in various orthogonal and oblique planes and the creation of volume and surface rendered computer models. We present three simple, tissue specific protocols for processing various groups of organic specimens, including mouse, chick, quail, frog and zebra fish embryos, human biopsy material, uncoated paper and skin replacement material.

High-resolution Episcopic Microscopy HREM - Simple and Robust Protocols for Processing and Visualizing Organic Materials

We provide simple and robust protocols for processing biopsy material of various species, embryos of biomedical model organisms and samples of other organic tissues in order to permit digital volume data generation with the high-resolution episcopic microscopy method.

We provide simple protocols for generating digital volume data with the high-resolution episcopic microscopy (HREM) method. HREM is capable of imaging ...

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 ...

Microinjection-based System for In Vivo Implantation of Embryonic Cardiomyocytes in the Avian Embryo

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a general challenge in developmental biology research. In the embryonic heart, this can be particularly difficult as regional differences in the expression of transcriptional regulators, paracrine/juxtacrine signaling cues, and hemodynamic force are all known to influence cardiomyocyte maturation. A simplified method to alter a developing cardiomyocyte's molecular and biomechanical microenvironment would, therefore, serve as a powerful technique to examine how local conditions influence cell fate and function. To address this, we have optimized a method to physically transplant juvenile cardiomyocytes into ectopic locations in the heart or the surrounding embryonic tissue. This allows us to examine how microenvironmental conditions influence cardiomyocyte fate transitions at single cell resolution within the intact embryo. Here, we describe a protocol in which embryonic myocytes can be isolated from a variety of cardiac sub-domains, dissociated, fluorescently labeled, and microinjected into host embryos with high precision. Cells can then be directly analyzed in situ using a variety of imaging and histological techniques. This protocol is a powerful alternative to traditional grafting experiments that can be prohibitively difficult in a moving tissue such as the heart. The general outline of this method can also be adapted to a variety of donor tissues and host environments, and its ease of use, low cost, and speed make it a potentially useful application for a variety of developmental studies.

In this method, embryonic cardiac tissues are surgically microdissected, dissociated, fluorescently labeled, and implanted into host embryonic tissues. This provides a platform for studying the individual or tissue level developmental organization under ectopic hemodynamic conditions, and/or altered paracrine/juxtacrine environments.

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a ...

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 ...

Isotropic Light-Sheet Microscopy and Automated Cell Lineage Analyses to Catalogue Caenorhabditis elegans Embryogenesis with Subcellular Resolution

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous system can be observed, with single cell resolution, in vivo. Here, we present an integrated protocol for the examination of neurodevelopment in C. elegans embryos. Our protocol combines imaging, lineaging and neuroanatomical tracing of single cells in developing embryos. We achieve long-term, four-dimensional (4D) imaging of living C. elegans&#160;embryos with nearly isotropic spatial resolution through the use of Dual-view Inverted Selective Plane Illumination Microscopy (diSPIM). Nuclei and neuronal structures in the nematode embryos are imaged and isotropically fused to yield images with resolution of ~330 nm in all three dimensions. These minute-by-minute high-resolution 4D data sets are then analyzed to correlate definitive cell-lineage identities with gene expression and morphological dynamics at single-cell and subcellular levels of detail. Our protocol is structured to enable modular implementation of each of the described steps and enhance studies on embryogenesis, gene expression, or neurodevelopment.

Here, we present a combinatorial approach using high-resolution microscopy, computational tools, and single-cell labeling in living C. elegans embryos to understand single cell dynamics during neurodevelopment.

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous ...

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with development, tissue homeostasis, and disease processes. Drosophila spermatocytes undergoing meiosis are ideal for this analysis because (1) whole Drosophila testes containing spermatocytes are relatively easy to prepare for microscopy, (2) the spermatocytes&#8217; large size makes them well suited for high resolution imaging, and (3) powerful Drosophila genetic tools can be integrated with in vivo analysis. Here, we present a readily accessible protocol for the preparation of whole testes from Drosophila third instar larvae and early pupae. We describe how to identify meiotic spermatocytes in prepared whole testes and how to image them live by time-lapse microscopy. Protocols for fixation and immunostaining whole testes are also provided. The use of larval testes has several advantages over available protocols that use adult testes for spermatocyte analysis. Most importantly, larval testes are smaller and less crowded with cells than adult testes, and this greatly facilitates high resolution imaging of spermatocytes. To demonstrate these advantages and the applications of the protocols, we present results showing the redistribution of the endoplasmic reticulum with respect to spindle microtubules during cell division in a single spermatocyte imaged by time-lapse confocal microscopy. The protocols can be combined with expression of any number of fluorescently tagged proteins or organelle markers, as well as gene mutations and other genetic tools, making this approach especially powerful for analysis of cell division mechanisms in the physiological context of whole tissues and organs.

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

The goal of this protocol is to analyze cell division in intact tissue by live and fixed cell microscopy using Drosophila meiotic spermatocytes. The protocol demonstrates how to isolate whole, intact testes from Drosophila larvae and early pupae, and how to process and mount them for microscopy.

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with ...