We thank members of the Kamiyama Lab for comments on the manuscript. This work was supported by an NIH R01 NS107558 (to M.I., K.B., and D.K.).

1. Equipment and Supplies
<ol>
	<li>Materials for collecting embryos and training adults to lay eggs
	<ol>
		<li>Prepare the filtration apparatus by severing a 50 mL tube and cutting open a hole in the cap to set a mesh filter with pores of 100 &#181;m (Table of Materials) in between the tube and the cap. 
		NOTE: Alternatively, cell strainers with pores of 100 &#181;m (Table of Materials) can be used for the filtration step of embryo collection.</li>
		<li>Make agar plates with grape agar premix (Table of Materials) according to the listed instructions. Briefly, gently stir 1 packet of the powder mix into 500 mL of room temperature (RT, 23 &#176;C) dH2O and microwave the dissolved mixture to vigorous boil. After cooling down to 70&#8722;75 &#176;C, pour the mixture into Petri dishes (60 mm). After the agar is solidified, store plates at 4 &#176;C.</li>
		<li>Prepare yeast paste by mixing active dry yeast (Table of Materials) and water to a paste consistency, and keep at 4 &#176;C.</li>
		<li>Use egg-collection cages (for 60 mm Petri dish, Table of Materials) that provide sufficient air flow.</li>
	</ol>
	</li>
	<li>Preparation of dissection needles and dye injection micropipettes
	<ol>
		<li>Prepare dye injection micropipette and dissection needle from the same capillary tubing with an inner diameter of 0.6 mm and an outer diameter of 1.2 mm (Table of Materials). Pull the capillary tubing by a micropipette puller at 7% from 170 V maximum output (Table of Materials) to create a sharp needle with a taper of ~0.4 cm in length.</li>
		<li>For dye injection, adjust the micropipette with a micropipette beveler (Table of Materials) by a bubble beveling technique described in instrument&#39;s manual.
		<ol>
			<li>In short, soak the grinder with a wetting agent (Table of Materials) to prevent the water from &#39;dragging&#39; the needle tip. Place the needle on the micropipette clamp at 25&#8722;30&#176; and lower the tip onto two-thirds of the radius out from the center of the beveling surface. Grind the needle while a syringe with tubing pushes air into the needle, to ensure that the micropipette will be clear of glass shavings.</li>
			<li>Mark the micropipette with a fine-tip permanent marker to indicate the position of the opening at the tip after beveling as it is challenging to locate the narrow opening of the micropipette that is formed at an angle.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Preparation for Embryo Collection
<ol>
	<li>Ensure that the adult flies (20&#8722;40 wild-type Canton-S or white flies), males and females, are maintained in young (&#60;7 days) and healthy conditions for the ideal egg collection. 
	NOTE: To stimulate egg-laying, flies are trained in their egg collection cage a couple of days prior to egg collection on agar plates streaked with yeast paste at least once every day.</li>
</ol>
3. Embryo Staging
<ol>
	<li>Allow the flies to lay eggs overnight (or at least 15 h) at RT to collect the embryos at 15 h AEL, i.e., stage 1618, to view dendritogenesis of the aCC and RP3 motor neurons. In the morning, collect the plate with the eggs. 
	NOTE: The embryos at 15 h AEL will have a distinct 4-chamber gut18. For imaging different stages follow their specific morphological criteria and aging conditions.</li>
	<li>To collect the embryos, dechorionate the eggs laid on the plate with 50% bleach for 5 min.</li>
	<li>Once the chorions have cleared, pour the contents of the plate through the filtration apparatus or cell strainer to isolate the embryos. Using a squeeze bottle of water, dilute the bleach left on the plate and gather as many embryos as possible by decanting the mixture into the filter.</li>
	<li>Wash the embryos on the filter 3&#8722;4x with more water or until the bleach odor dissipates. Remove the filter from the apparatus and wash the embryos onto another clean plate with water. Decant the water from the new plate that the embryos are on.</li>
	<li>Prepare a glass slide by covering it with two layers of vinyl tape in the center, forming a rectangle. Cut a rectangular pool out of the tape using a razor blade. Place a thin strip of double-sided tape towards the upper end of the pool, this is where the embryos will be placed as shown in Figure 1.</li>
	<li>Using fine forceps, individually select 5&#8722;10 embryos at 15 h AEL and place them on the double-sided tape with the dorsal side facing up. Add insect Ringer&#39;s saline19 to the dissection pool to protect the embryos from desiccation (Figure 1).</li>
</ol>
4. Dissection and Staining
<ol>
	<li>Using a glass needle under a dissecting microscope (Table of Materials), cut through the midline of a single embryo at its surface from its posterior to its anterior end. Then drag the embryo out from the vitelline membrane from the tape onto the glass (boxed in Figure 1). Take care not to damage the interior tissues of the embryo.</li>
	<li>Flip the epithelial tissues from the center and attach the epidermal edge onto the surface of the glass slide (Figure 1, inset).</li>
	<li>Using a tube-connected needle with a tip opening of ~300 &#181;m (prepared by breaking the thin tip of a dissection needle), aspirate or blow air to detach and remove the dorsal longitudinal tracheal trunks as well as any remaining guts.</li>
	<li>Use 4% paraformaldehyde (PFA) in phosphate-buffered saline (PBS) to fix the embryos for 5 min at RT. Wash the embryos 3x with PBS.</li>
	<li>Stain the embryos with 1 &#181;L of anti-horseradish peroxidase antibody conjugated with cyanine 3 dye (anti-HRP Cy3) (Table of Materials) in 200 &#181;L of PBS for 1 h. Wash the embryos with PBS 3x after staining. 
	NOTE: The dye of anti-HRP can be changed based on the lipophilic dyes of choice for injection.</li>
</ol>
5. Filling of the Injection Micro-pipette
<ol>
	<li>Heat lipophilic dyes (5 mg/mL of DiO or DiD, Table of Materials) to 60 &#176;C in a 1:10 mixture of ethanol:vegetable oil before use.</li>
	<li>Prepare an oil-dissolved dye slide for the injection micropipette. Place the micropipette into the capillary holder (Figure 2, #1). Using the micromanipulator (Table of Materials), adjust the micropipette to be over the dye slide. Then, adjust the stage to place the micropipette onto the dye (Figure 2, #2).</li>
	<li>To fill up the micropipette, use a microinjector (Table of Materials) (Figure 2, #3). Collect the dye in the micropipette by setting the Pi (injection pressure) between 200&#8722;500 hPa (hectopascal), the Ti (injection time) between 0.1&#8722;0.5 s and Pc (compensation pressure) to 0 hPa for 5 min (Figure 2, #4).</li>
	<li>Once the dye has been collected, remove the dye slide and place the sample onto the microscope stage. Next, increase the Pc to a range of 20&#8722;60 hPa before lowering the micropipette into the sample to prevent contamination of PBS by capillary action.</li>
</ol>
6. Dye Injection into Neurons
<ol>
	<li>Locate the embryo in the center using the microscope with 10x objective lens (Table of Materials) and align the micropipette with the embryo. 
	NOTE: The size of the dye droplet can be adjusted by changing the Pi or the size of the opening of the micropipette tip. The droplet should be 10&#8722;20 &#181;m, which is approximately the width of 1 muscle.</li>
	<li>Change the objective lens to a water-immersion 40x lens (Table of Materials) and submerge the lens into PBS to see the embryo.
	<ol>
		<li>Use fluorescence microscopy to check the neuronal morphology marked by anti-HRP Cy3 and determine the injection site.</li>
		<li>During injection, use brightfield microscopy to see the dye droplet. When the embryo is in focus, change the position of the micropipette to make gentle contact with the tip of the axon of interest (e.g., aCC, RP3).</li>
		<li>Drop the dye in a right abdominal (A2&#8722;A6) hemi-segment at the neuromuscular junction of aCC or RP3 (Figure 3) with either DiD or DiO, by using the neurons marked by anti-HRP Cy3. Using the hand control (mouse; Figure 2, 5) release the dye and remove the micropipette after dropping the dye with the micromanipulator and move onto the next injection site. 
		NOTE: Unlike other dyes (e.g., Lucifer yellow, calcein) which spread into neighboring cells through gap junctions, lipophilic dyes associate with cell membranes and do not transfer to neighbors. Due to the relatively large size of the dye droplet, however, this technique also results in labeling of the partnering muscles (Figure 3A).</li>
	</ol>
	</li>
	<li>Incubate the sample at RT for 1 h after dye-drop before imaging. 
	NOTE: The protocol can be paused here before mounting, and the sample can be kept at 4 &#176;C overnight. Lipophilic dyes can also be delivered using iontophoresis, if an intracellular direct-coupled (DC) amplifier is readily available20.</li>
</ol>
7. Imaging with a Confocal Microscope
<ol>
	<li>Remove the double-sided tape and vinyl tape from the glass slide with the help of forceps.</li>
	<li>Prepare a cover slip (22 x 22 mm2 No.1 cover glass) with a small amount of vacuum grease (Table of Materials) at the four corners and carefully place on the sample, avoiding air bubbles. Remove any excess PBS using task wipers.</li>
	<li>Push down the cover slip to adjust the working distance between the objective lens and the sample. Completely seal the edges of the cover slip with nail polish.</li>
	<li>Image at 10x and 100x magnification using a confocal microscope.</li>
	<li>Use ImageJ software for processing raw images from the microscope (Table of Materials). 
	NOTE: Observation must begin within 10 min after mounting for the best images. Otherwise, at RT, the dye will spread to sites adjacent to the injection site creating unwanted background for imaging. To slow down the diffusion of dye, the sample can be stored at 4 &#176;C for a couple of hours.</li>
</ol>

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M., Bate, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+origin,+location,+and+projections+of+the+embryonic+abdominal+motorneurons+of+Drosophila.">The origin, location, and projections of the embryonic abdominal motorneurons of Drosophila.</a> Journal of Neuroscience. 17 (24), 9642-9655 (1997).</li><li>Kamiyama, D., Chiba, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endogenous+activation+patterns+of+Cdc42+GTPase+within+Drosophila+embryos.">Endogenous activation patterns of Cdc42 GTPase within Drosophila embryos.</a> Science. 324 (5932), 1338-1340 (2009).</li><li>Furrer, M. P., Vasenkova, I., Kamiyama, D., Rosado, Y., Chiba, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Slit+and+Robo+control+the+development+of+dendrites+in+Drosophila+CNS.">Slit and Robo control the development of dendrites in Drosophila CNS.</a> Development. 134 (21), 3795-3804 (2007).</li><li>Kamiyama, D., 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=Specification+of+Dendritogenesis+Site+in+Drosophila+aCC+Motoneuron+by+Membrane+Enrichment+of+Pak1+through+Dscam1.">Specification of Dendritogenesis Site in Drosophila aCC Motoneuron by Membrane Enrichment of Pak1 through Dscam1.</a> Developmental Cell. 35 (1), 93-106 (2015).</li><li>Campos-Ortega, J. A., Hartenstein, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The embryonic development of Drosophila melanogaster. , (1985).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+Ringer's+solution.">Drosophila Ringer's solution.</a> Cold Spring Harbor Protocols. 2007 (4), (2007).</li><li>Rickert, C., Kunz, T., Harris, K. -. L., Whitington, P., Technau, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Labeling+of+single+cells+in+the+central+nervous+system+of+Drosophila+melanogaster.">Labeling of single cells in the central nervous system of Drosophila melanogaster.</a> Journal of Visualized Experiments. (73), e50150 (2013).</li><li>Fujioka, 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=Even-skipped,+acting+as+a+repressor,+regulates+axonal+projections+in+Drosophila.">Even-skipped, acting as a repressor, regulates axonal projections in Drosophila.</a> Development. 130 (22), 5385-5400 (2003).</li><li>Sink, H., Rehm, E. J., Richstone, L., Bulls, Y. M., Goodman, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=sidestep+encodes+a+target-derived+attractant+essential+for+motor+axon+guidance+in+Drosophila.">sidestep encodes a target-derived attractant essential for motor axon guidance in Drosophila.</a> Cell. 105 (1), 57-67 (2001).</li><li>Furrer, M. 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The authors have nothing to disclose.

The use of dye labeling for studying neuronal morphology has several advantages over genetic cell-labeling techniques. The dye labeling technique can minimize the amount of time needed for labeling and imaging the morphologies of motor neurons. The dye labeling process is quite fast as it takes less than 2 h and enables us to define the outline of neuronal projections. As an alternative, one can visualize the aCC motor neuron by choosing a GAL4 line that expresses the yeast GAL4 transcription factor in aCC, and crossing ...

The embryonic motor neuron system of Drosophila offers a powerful experimental model to analyze the mechanisms underlying the development of the central nervous system (CNS)1,2,3. The motor neuron system is amenable to biochemical, genetic, imaging, and electrophysiological techniques. Using the techniques, genetic manipulations and functional analyses can be carried out at the level of single motor neurons2,4,5,6.
During early development of the nervous system, neuroblasts divide and generate a large number of glia and neurons. The spatiotemporal relationship between the delamination and the gene expression profile of neuroblasts has been previously investigated in detail7,8,9. In the case of the motor neuron system, the formation of embryonic neuromuscular junction (NMJ) has been extensively studied using the aCC (anterior corner cell), RP2 (raw prawn 2), and RP5 motor neurons2,10. For instance, when the RP5 motor neuron forms a nascent synaptic junction, the pre-synaptic and post-synaptic filopodia are intermingled11,12,13. Such direct cellular communication is vital to initiate the NMJ formation. Contrary to what we know about the peripheral nerve branches, our knowledge of how motor dendrites initiate synaptic connectivity within the CNS is still primitive.
In this report, we present a technique that allows retrograde labeling of motor neurons in embryos by means of micropipette-mediated delivery of lipophilic dyes. This technique enables us to trace the 38 motor neurons innervating each of the 30 body wall muscles in a hemi-segment at 15 h after egg laying (AEL)14. By using this technique, our group has thoroughly investigated numerous gain-of-function/loss-of-function alleles15,16,17. We have recently unraveled the molecular mechanisms that drive initiation of motor dendrite connectivity and demonstrated that a Dscam1-Dock-Pak interaction defines the site of dendrite outgrowth in the aCC motor neuron17. In general, this technique is adaptable for the phenotypic analysis of any embryonic motor neurons in wild type or mutant strains, enhancing our ability to provide new insights into the functional design of the Drosophila nervous system.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10x objective lens</td>
			<td>Nikon</td>
			<td></td>
			<td>Plan</td>
		</tr>
		<tr>
			<td>40x water-immersion lens</td>
			<td>Nikon</td>
			<td></td>
			<td>NIR Apo</td>
		</tr>
		<tr>
			<td>Capillary tubing</td>
			<td>Frederick Haer&#38;Co</td>
			<td>27-31-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal microscope</td>
			<td>Andor</td>
			<td>N/A</td>
			<td>Dragonfly Spinning disk confocal unit</td>
		</tr>
		<tr>
			<td>Cover glass</td>
			<td>Corning</td>
			<td></td>
			<td>22x22 mm Square #1</td>
		</tr>
		<tr>
			<td>DiD</td>
			<td>ThermoFisher</td>
			<td>V22886</td>
			<td></td>
		</tr>
		<tr>
			<td>DiI</td>
			<td>ThermoFisher</td>
			<td>V22888</td>
			<td></td>
		</tr>
		<tr>
			<td>DiO</td>
			<td>ThermoFisher</td>
			<td>V22887</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting microscope</td>
			<td>Nikon</td>
			<td>N/A</td>
			<td>SMZ-U</td>
		</tr>
		<tr>
			<td>Double Sided Tape</td>
			<td>Scotch</td>
			<td>665</td>
			<td></td>
		</tr>
		<tr>
			<td>Dow Corning High-Vacuum Grease</td>
			<td>Fisher Sci.</td>
			<td>14-635-5D</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont #5 Forceps</td>
			<td>Fine Science Tools</td>
			<td>11252-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Egg collection cage</td>
			<td>FlyStuff</td>
			<td>59-100</td>
			<td></td>
		</tr>
		<tr>
			<td>FemtoJet 5247</td>
			<td>Eppendorf</td>
			<td>discontinued</td>
			<td>FemtoJet 4i (Cat No. 5252000021)</td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td>NIH</td>
			<td></td>
			<td>Image processing software</td>
		</tr>
		<tr>
			<td>Micromanipulator</td>
			<td>Sutter</td>
			<td>MP-225</td>
			<td></td>
		</tr>
		<tr>
			<td>Micropipette beveler</td>
			<td>Sutter</td>
			<td>BV-10-B</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle puller</td>
			<td>Narishige</td>
			<td>PC-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Nutri-Fly Grape Agar Powder Premix Packets</td>
			<td>FlyStuff</td>
			<td>47-102</td>
			<td></td>
		</tr>
		<tr>
			<td>Nylon Net Filter</td>
			<td>Millipore</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde 16% Solution, EM grade</td>
			<td>Electron Microscopy Sciences</td>
			<td>15710</td>
			<td>Any EM grades</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Roche</td>
			<td>11666789001</td>
			<td>Sold on sigmaaldrich, boxed 10x solution</td>
		</tr>
		<tr>
			<td>Photo-Flo 200</td>
			<td>Kodak</td>
			<td>146 4510</td>
			<td>Wetting agent</td>
		</tr>
		<tr>
			<td>Upright fluorescence microscope</td>
			<td>Nikon</td>
			<td>N/A</td>
			<td>Eclipse Ci with a LED light source</td>
		</tr>
		<tr>
			<td>Vinyl Electrical Tape</td>
			<td>Scotch</td>
			<td>6143</td>
			<td></td>
		</tr>
		<tr>
			<td>VWR Cell Strainers</td>
			<td>VWR</td>
			<td>10199-659</td>
			<td></td>
		</tr>
		<tr>
			<td>Yeast</td>
			<td>FlyStuff</td>
			<td>62-103</td>
			<td>Active dry yeast (RED STAR)</td>
		</tr>
	</tbody>
</table>

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.

A representative image of the aCC and RP3 motor neurons is shown in Figure 3C to demonstrate the multicolor labeling of motor neurons at 15 h AEL. Their dendritic morphologies are largely invariant between embryos. The staining pattern obtained with anti-HRP antibody is shown in gray. A small droplet of DiO or DiD was deposited on the NMJ of muscle 1 or 6/7, respectively. Figure 4 demonstrates the capability to quantitatively measure the phenoty...

Watch this Scientific Journal Video about Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes at JoVE.com

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.

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 ...

retrograde-tracing-drosophila-embryonic-motor-neurons-using

Research

JoVE Journal

Developmental Biology

6.1K Views.  University of Georgia. We describe a method for retrograde tracing of the Drosophila embryonic motor neurons using lipophilic fluorescent dyes.

Video: Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes

Analysis of Cardiomyocyte Development using Immunofluorescence in Embryonic Mouse Heart

During heart development, the generation of myocardial-specific structural and functional units including sarcomeres, contractile myofibrils, intercalated discs, and costameres requires the coordinated assembly of multiple components in time and space. Disruption in assembly of these components leads to developmental heart defects. Immunofluorescent staining techniques are used commonly in cultured cardiomyocytes to probe myofibril maturation, but this ex vivo approach is limited by the extent to which myocytes will fully differentiate in culture, lack of normal in vivo mechanical inputs, and absence of endocardial cues. Application of immunofluorescence techniques to the study of developing mouse heart is desirable but more technically challenging, and methods often lack sufficient sensitivity and resolution to visualize sarcomeres in the early stages of heart development. Here, we describe a robust and reproducible method to co-immunostain multiple proteins or to co-visualize a fluorescent protein with immunofluorescent staining in the embryonic mouse heart and use this method to analyze developing myofibrils, intercalated discs, and costameres. This method can be further applied to assess cardiomyocyte structural changes caused by mutations that lead to developmental heart defects.

Mutations that lead to congenital heart defects benefit from in vivo investigation of cardiac structure during development, but high-resolution structural studies in the mouse embryonic heart are technically challenging. Here we present a robust immunofluorescence and image analysis method to assess cardiomyocyte-specific structures in the developing mouse heart.

During heart development, the generation of myocardial-specific structural and functional units including sarcomeres, contractile myofibrils, intercalated ...

A Method for Lineage Tracing of Corneal Cells Using Multi-color Fluorescent Reporter Mice

Lineage tracing experiments define the origin, fate and behavior of cells in a specific tissue or organism. This technique has been successfully applied for many decades, revealing seminal findings in developmental biology. More recently, it was adopted by stem cell biologists to identify and track different stem cell populations with minimal experimental intervention. The recent developments in mouse genetics, the availability of a large number of mouse strains, and the advancements in fluorescent microscopy allow the straightforward design of powerful lineage tracing systems for various tissues with basic expertise, using commercially available tools. We have recently taken advantage of this powerful methodology to explore the origin and fate of stem cells at the ocular surface using R26R-Confetti mouse. This model offers a multi-color genetic system, for the expression of 4 fluorescent genes in a random manner. Here we describe the principles of this methodology and provide an adaptable protocol for designing lineage tracing experiments; specifically for the corneal epithelium as well as for other tissues.

In this article we describe the principles for designing and performing a multi-color lineage tracing experiment using R26R-Confetti mice. We provide a specific protocol for tracking corneal epithelial cells which can be modified for other tissues of interest.

Lineage tracing experiments define the origin, fate and behavior of cells in a specific tissue or organism. This technique has been successfully applied ...

Transcriptome Profiling of In-Vivo Produced Bovine Pre-implantation Embryos Using Two-color Microarray Platform

Early embryonic loss is a large contributor to infertility in cattle. Moreover, bovine becomes an interesting model to study human preimplantation embryo development due to their similar developmental process. Although genetic factors are known to affect early embryonic development, the discovery of such factors has been a serious challenge. Microarray technology allows quantitative measurement and gene expression profiling of transcript levels on a genome-wide basis. One of the main decisions that have to be made when planning a microarray experiment is whether to use a one- or two-color approach. Two-color design increases technical replication, minimizes variability, improves sensitivity and accuracy as well as allows having loop designs, defining the common reference samples. Although microarray is a powerful biological tool, there are potential pitfalls that can attenuate its power. Hence, in this technical paper we demonstrate an optimized protocol for RNA extraction, amplification, labeling, hybridization of the labeled amplified RNA to the array, array scanning and data analysis using the two-color analysis strategy.

Transcriptome Profiling of In-Vivo Produced Bovine Pre-implantation Embryos Using Two-color Microarray Platform

Microarray technology allows quantitative measurement and gene expression profiling of transcripts on a genome-wide basis. Therefore, this protocol provides an optimized technical procedure in a two-color custom made bovine array using Day 7 bovine embryos to demonstrate the feasibility of using low amount of total RNA.

Early embryonic loss is a large contributor to infertility in cattle. Moreover, bovine becomes an interesting model to study human preimplantation embryo ...

In Vivo Tracking of Human Adipose-derived Mesenchymal Stem Cells in a Rat Knee Osteoarthritis Model with Fluorescent Lipophilic Membrane Dye

In order to support the clinical application of human adipose-derived mesenchymal stem cell (haMSC) therapy for knee osteoarthritis (KOA), we examined the efficacy of cell persistence and biodistribution of haMSCs in animal models. We demonstrated a method to label the cell membrane of haMSCs with lipophilic fluorescent dye. Subsequently, intra-articular injection of the labeled cells in rats with surgically induced KOA was monitored dynamically by an in vivo imaging system. We employed the lipophilic carbocyanines DiD (DilC18 (5)), a far-red fluorescent Dil (dialkylcarbocyanines) analog, which utilized a red laser to avoid excitation of the natural green autofluorescence from surrounding tissues. Furthermore, the red-shifted emission spectra of DiD allowed deep-tissue imaging in live animals and the labeling procedure caused no cytotoxic effects or functional damage to haMSCs. This approach has been shown to be an efficient tracking method for haMSCs in a rat KOA model. The application of this method could also be used to determine the optimal administration route and dosage of MSCs from other sources in pre-clinical studies.

In Vivo Tracking of Human Adipose-derived Mesenchymal Stem Cells in a Rat Knee Osteoarthritis Model with Fluorescent Lipophilic Membrane Dye

This protocol describes an efficient way to monitor the cell persistence and biodistribution of human adipose-derived mesenchymal stem cells (haMSCs) by far-red fluorescence labeling in a rat knee osteoarthritis (KOA) model via intra-articular (IA) injection.

In order to support the clinical application of human adipose-derived mesenchymal stem cell (haMSC) therapy for knee osteoarthritis (KOA), we examined the ...

Tracing Gene Expression Through Detection of &#946;-galactosidase Activity in Whole Mouse Embryos

The Escherichia coli LacZ gene, encoding &#946;-galactosidase, is largely used as a reporter for gene expression and as a tracer in cell lineage studies. The classical histochemical reaction is based on the hydrolysis of the substrate X-gal in combination with ferric and ferrous ions, which produces an insoluble blue precipitate that is easy to visualize. Therefore, &#946;-galactosidase activity serves as a marker for the expression pattern of the gene of interest as the development proceeds. Here we describe the standard protocol for the detection of &#946;-galactosidase activity in early whole mouse embryos and the subsequent method for paraffin sectioning and counterstaining. Additionally, a procedure for clarifying whole embryos is provided to better visualize X-gal staining in deeper regions of the embryo. Consistent results are obtained by performing this procedure, although optimization of reaction conditions is needed to minimize background activity. Limitations in the assay should be also considered, particularly regarding the size of the embryo in whole mount staining. Our protocol provides a sensitive and a reliable method for &#946;-galactosidase detection during the mouse development that can be further applied to the cryostat sections as well as whole organs. Thus, the dynamic gene expression patterns throughout development can be easily analyzed by using this protocol in whole embryos, but also detailed expression at the cellular level can be assessed after paraffin sectioning.

Here we describe the standard protocol for the detection of &#946;-galactosidase activity in early whole mouse embryos and the method for paraffin sectioning and counterstaining. This is an easy and quick procedure to monitor gene expression during development that can also be applied to tissue sections, organs or cultured cells.

The Escherichia coli LacZ gene, encoding &#946;-galactosidase, is largely used as a reporter for gene expression and as a tracer in cell lineage studies. ...

Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle

The majority of work on the neuronal specification has been carried out in genetically and physiologically tractable models such as C. elegans, Drosophila larvae, and fish, which all engage in undulatory movements (like crawling or swimming) as their primary mode of locomotion. However, a more sophisticated understanding of the individual motor neuron (MN) specification&#8212;at least in terms of informing disease therapies&#8212;demands an equally tractable system that better models the complex appendage-based locomotion schemes of vertebrates. The adult Drosophila locomotor system in charge of walking meets all of these criteria with ease, since in this model it is possible to study the specification of a small number of easily distinguished leg MNs (approximately 50 MNs per leg) both using a vast array of powerful genetic tools, and in the physiological context of an appendage-based locomotion scheme. Here we describe a protocol to visualize the leg muscle innervation in an adult fly.

Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle

Here we describe a protocol to visualize the axonal targeting with a florescent protein in adult legs of Drosophila by fixation, mounting, imaging, and post-imaging steps.

The majority of work on the neuronal specification has been carried out in genetically and physiologically tractable models such as C. elegans, Drosophila ...

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 ...

Evaluation of Fertilization State by Tracing Sperm Nuclear Morphology in Arabidopsis Double Fertilization

Flowering plants have a unique sexual reproduction system called &#8216;double fertilization&#39;,&#160;in which each of the sperm cells precisely fuses with an egg cell or a central cell. Thus, two independent fertilization events take place almost simultaneously. The fertilized egg cell and central cell develop into zygote and endosperm, respectively. Therefore, precise control of double fertilization is essential for the ensuing seed development. Double fertilization occurs in the female gametophyte (embryo sac), which is deeply hidden and covered with thick ovule and ovary tissues. This pistil tissue construction makes observation and analysis of double fertilization quite difficult and has created the present situation in which many questions regarding the mechanism of double fertilization remain unanswered. For the functional evaluation of a potential candidate for fertilization regulator, phenotypic analysis of fertilization is important. To judge the completion of fertilization in Arabidopsis thaliana, the shapes of fluorescence signals labeling sperm nuclei are used as indicators. A sperm cell that fails to fertilize is indicated by a condensed fluorescence signal outside of the female gametes, whereas a sperm cell that successfully fertilizes is indicated by a decondensed signal due to karyogamy with the female gametes&#8217; nucleus. The method described here provides a tool to determine successful or failed fertilization under in vivo conditions.

Evaluation of Fertilization State by Tracing Sperm Nuclear Morphology in Arabidopsis Double Fertilization

We demonstrate a method to determine successful or failed fertilization on the basis of sperm nuclear morphology in Arabidopsis double fertilization using an epifluorescence microscope.

Flowering plants have a unique sexual reproduction system called &#8216;double fertilization&#39;,&#160;in which each of the sperm cells precisely fuses ...

Clonal Genetic Tracing using the Confetti Mouse to Study Mineralized Tissues

Labeling an individual cell in the body to monitor which cell types it can give rise to and track its migration through the organism or determine its longevity can be a powerful way to reveal mechanisms of tissue development and maintenance. One of the most important tools currently available to monitor cells in vivo is the Confetti mouse model. The Confetti model can be used to genetically label individual cells in living mice with various fluorescent proteins in a cell type-specific manner and monitor their fate, as well as the fate of their progeny over time, in a process called clonal genetic tracing or clonal lineage tracing. This model was generated almost a decade ago and has contributed to an improved understanding of many biological processes, particularly related to stem cell biology, development, and renewal of adult tissues. However, preserving the fluorescent signal until image collection and simultaneous capturing of various fluorescent signals is technically challenging, particularly for mineralized tissue. This publication describes a step-by-step protocol for using the Confetti model to analyze growth plate cartilage that can be applied to any mineralized or nonmineralized tissue.

This method describes the use of the R26R-Confetti (Confetti) mouse model to study mineralized tissues, covering all steps from the breeding strategy to the image acquirements. Included is a general protocol that can be applied to all soft tissues and a modified protocol that can be applied to mineralized tissues.

Labeling an individual cell in the body to monitor which cell types it can give rise to and track its migration through the organism or determine its ...

Efficient Neural Differentiation using Single-Cell Culture of Human Embryonic Stem Cells

In vitro differentiation of human embryonic stem cells (hESCs) has transformed the ability to study human development on both biological and molecular levels and provided cells for use in regenerative applications. Standard approaches for hESC culture using colony type culture to maintain undifferentiated hESCs and embryoid body (EB) and rosette formation for differentiation into different germ layers are inefficient and time-consuming. Presented here is a single-cell culture method using hESCs instead of a colony-type culture. This method allows maintenance of the characteristic features of undifferentiated hESCs, including expression of hESC markers at levels comparable to colony type hESCs. In addition, the protocol presents an efficient method for neural progenitor cell (NPC) generation from single-cell type hESCs that produces NPCs within 1 week. These cells highly express several NPC marker genes and can differentiate into various neural cell types, including dopaminergic neurons and astrocytes. This single-cell culture system for hESCs will be useful in investigating the molecular mechanisms of these processes, studies of certain diseases, and drug discovery screens.

Presented here is a protocol for the generation of a single-cell culture of human embryonic stem cells and their subsequent differentiation into neural progenitor cells. The protocol is simple, robust, scalable, and suitable for drug screening and regenerative medicine applications.

In vitro differentiation of human embryonic stem cells (hESCs) has transformed the ability to study human development on both biological and molecular ...

Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes

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Analysis of Cardiomyocyte Development using Immunofluorescence in Embryonic Mouse Heart

A Method for Lineage Tracing of Corneal Cells Using Multi-color Fluorescent Reporter Mice

Transcriptome Profiling of In-Vivo Produced Bovine Pre-implantation Embryos Using Two-color Microarray Platform

In Vivo Tracking of Human Adipose-derived Mesenchymal Stem Cells in a Rat Knee Osteoarthritis Model with Fluorescent Lipophilic Membrane Dye

Tracing Gene Expression Through Detection of β-galactosidase Activity in Whole Mouse Embryos

Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

Evaluation of Fertilization State by Tracing Sperm Nuclear Morphology in Arabidopsis Double Fertilization

Clonal Genetic Tracing using the Confetti Mouse to Study Mineralized Tissues

Efficient Neural Differentiation using Single-Cell Culture of Human Embryonic Stem Cells