We would like to thank Alejandro S&#225;nchez Alvarado, who supported the initial optimization and development of this technique. Work in Carolyn Adler's laboratory is supported by Cornell University start-up funds.

1. Preparation
<ol>
	<li>Preparation of planaria water17
	<ol>
		<li>Maintain planarians in a 1X Montju&#239;c salt solution. To prepare planarian water, make individual stock solutions of 1 M CaCl2, 1 M MgSO4, 1 M MgCl2, 1 M KCl and 5 M NaCl in ultrapure water. Filter-sterilize with a 0.2 &#181;m bottle-top filterfor long-term storage. 
		Note: Use only ultrapure deionized water (with a resistivity of 18.2 M&#8486; at 25 &#176;C) to prepare Montju&#239;c salts.</li>
		<li>To prepare a 1 L stock of 5X salt solution, combine 5 mL of 1 M CaCl2, 5 mL of 1 M MgSO4, 0.5 mL of MgCl2, 0.5 mL of KCl and 1.6 mL of 5 M NaCl solutions in 900 mL ultrapure water. To this solution, add 0.504 g of NaHCO3 and stir to mix. Adjust the pH to 7.0 with hydrochloric acid.</li>
		<li>Dilute this 5X stock solution to a 1X working concentration in ultrapure water in a sterile container such as a large capacity carboy (see the Table of Materials). Use this 1X solution for maintaining asexual planarians. 
		Note: As an alternative to Montju&#239;c salt solution, use locally-purchased spring water or ultrapure water containing a commercially available aquarium salt mix (see Table of Materials) at a concentration of0.5 g/L18.</li>
		<li>To prevent bacterial infection in static culture19, maintain planarians in water containing an antibiotic. Prepare a 50 mg/mL stock solution of gentamicin sulfate in ultrapure water and filter-sterilize. To containers where animals are maintained, add gentamicin sulfate to a final concentration of 50 &#181;g/mL (1:1000 dilution).</li>
	</ol>
	</li>
	<li>Preparation of liver paste
	<ol>
		<li>As planarians thrive on a diet of organic, grass-fed beef liver, purchase fresh liver and process within 24 h. Remove the membranous capsule encapsulating the liver by peeling it off gently. Cut the liver into ~1-cm cubes, and use a blade to scrape off and discard all hepatic veins and arteries.</li>
		<li>Macerate liver pieces using a food mill or a food processor and then pass it through a wire mesh food strainer. Combine into pastry bags, and dispense into syringes or 35-mm Petri dishes. Prior to feeding, centrifuge gently to remove air bubbles.</li>
		<li>Store aliquots of liver at -80 &#176;C for up to a year and thaw prior to using. After thawing, re-freeze any leftover liver once, or store at 4 &#176;C for up to 24 h.</li>
	</ol>
	</li>
	<li>Maintenance of animals
	<ol>
		<li>Planarians will grow and shrink to various sizes depending on the frequency of feeding. Feed planarians every other week (once every 14 days). For long-term bulk cultures, use plastic containers of various sizes (see Table of Materials).</li>
		<li>To feed animals, use a metal spatula or plastic transfer pipet to place a pea-sized drop of liver in a box. Allow animals to eat for 1-2 h. Remove remaining food before cleaning the box.</li>
		<li>Clean worms twice a week if fed (once directly after feeding and once two days later), or once a week if not fed. To clean, drain water into a plastic beaker by carefully pouring out the water while retaining planarians in the box. Wipe box surface with a paper towel, then repeat on all sides until the box is clean.</li>
		<li>Replace water (to approximately &#190; of the box volume) and add gentamicin to a final concentration of 50 &#181;g/mL. Store animals in a dark cabinet or in a 20 &#176;C incubator.</li>
	</ol>
	</li>
	<li>Selection of worms
	<ol>
		<li>Select worms that were fed 5-7 days prior. 
		Note: It is significantly more difficult to amputate pharynges from worms that have been recently fed (1-2 days prior to amputation). Fed animals are also more sensitive to sodium azide.</li>
		<li>Select worms that are roughly 6 mm in length. To estimate the size of the worm, transfer it to a Petri dish using a plastic transfer pipette. Place either a flat 6-in ruler or 5 mm x 5 mm graph paper underneath the Petri dish and measure the worm length while it crawls. 
		Note: Worms smaller than 6 mm in length are more difficult to amputate.</li>
	</ol>
	</li>
	<li>Preparation of sodium azide
	<ol>
		<li>Prepare a 100 mM solution by dissolving sodium azide powder in planaria water. 
		CAUTION: Sodium azide is toxic and should be handled carefully. Do not discard sodium azide down the drain. Following use, collect it separately for disposal as hazardous waste.</li>
	</ol>
	</li>
</ol>
2. Pharynx Amputation
<ol>
	<li>Place worms in a 35-mm diameter Petri dish. Carefully remove all planarian water from the dish by using a plastic transfer pipet. 
	Note: A maximum of 20 6-mm worms can be comfortably accommodated in a dish of this size.</li>
	<li>Place the dish under the microscope and adjust magnification so that multiple worms can be viewed simultaneously (e.g.&#160;10X magnification). Replace planarian water with 100 mM sodium azide solution. 
	Note: For a 35-mm Petri dish, approximately 5 mL of sodium azide is sufficient. The solution should completely submerge planarians. Adjust solution volume for larger dishes as necessary.</li>
	<li>Monitor the animals under the microscope&#160;and refrain from moving the dish for the first 3-4 min.</li>
	<li>Observe the extrusion of the pharynx through the microscope; the pharynx will protrude from the pharyngeal pouch and extend fully (approximately 1 mm in length) as shown in Figure 1B. 
	Note: It is important to wait until the pharynx has finished extending completely before proceeding further. Once the pharynx emerges from the animal, full extension takes roughly 1 min.</li>
	<li>Use a plastic transfer pipet to squirt animals around the dish. Suck the animals into the pipet and forcibly release them into the dish a couple of times. If the pharynx is fully extended, this vigorous pipetting will cause it to detach.</li>
	<li>Alternatively, grasp the pharynx either vertically along its length, or horizontally along its circumference using a pair of fine forceps (see Table of Materials) as shown in Figure 1C. With the pharynx pinched in the forceps, lift the animal upwards towards the meniscus. As the animal is raised, surface tension will cause the pharynx to detach from the animal. 
	Note: This method can be used to remove the pharynges of smaller animals (1-3 mm in length) where forceful swirling is not adequate. Avoid poking or otherwise injuring the body of the animal.</li>
	<li>Using a transfer pipet, move amputated animals to a new dish containing fresh planaria water.</li>
	<li>Once transferred, rinse amputated animals thoroughly in planaria water, completely exchanging the water three times. Transfer to a new dish containing planaria water with 50&#181;g/mL of gentamicin. 
	Note: Complete buffer exchange during these three washes is essential to ensure removal of sodium azide.</li>
	<li>The next day, replace water in the dish with fresh planaria water containing 50 &#181;g/mL of gentamicin. Repeat every other day thereafter.</li>
</ol>
3. Assessment of Pharynx Removal After Amputation
<ol>
	<li>Examine animals under a microscope (with 10-20X magnification) to determine whether a dark spot has appeared after successful removal of the pharynx, as shown in Figure 2A.</li>
	<li>Alternatively, fix and bleach animals according to the protocol described by Pearson et al.20. Soak animals overnight in 4&#39;,6-diamidino-2-phenylindole (DAPI) or fluorescently-conjugated streptavidin (1 mg/mL diluted 1:500 in 1X Phosphate Buffered Saline containing 0.3% Triton-X). Mount animals on slides in 80% glycerol/20% 1X Phosphate Buffered Saline and image under a fluorescent stereomicroscope (Figure 2B). 
	Note: All images in this manuscript were captured on a fluorescent stereomicroscope with a 1.0X objective.</li>
</ol>
4. Assessment of Pharynx Regeneration by Measuring Feeding Behavior
<ol>
	<li>Preparation of liver
	<ol>
		<li>Transfer required quantity of liver paste into a microcentrifuge or conical tube. Spin briefly to remove air bubbles. Estimate the volume of liver, and then add planaria water to 1/5 of its volume, and 2% of the total volume in red food coloring. For example, to 1000 &#181;L of liver paste, add 200 &#181;L of planaria water and 24 &#181;L of food coloring.</li>
		<li>Using a plastic pestle, pipette tip or metal spatula, mix thoroughly until food coloring is evenly distributed in the liver paste. Spin again briefly. Liver paste can be stored at 4 &#176;C for 24 h or frozen in aliquots at -80 &#176;C for long-term storage.</li>
		<li>If testing up to 25 animals, prepare 25 &#181;L of liver paste per dish (approximately 1 &#181;L of liver paste per animal).</li>
	</ol>
	</li>
	<li>Feeding animals
	<ol>
		<li>Seven days after chemical amputation, transfer animals to a new Petri dish. If testing 10-15 animals, use a 35 mm dish; more than 15 animals should be tested in a 60-mm Petri dish. Keep animals in the dark, undisturbed, for approximately 1 h prior to feeding.</li>
		<li>Using scissors, increase the width of a P200 pipette tip by trimming roughly &#189; cm off of the narrow end. Pipet 25 &#181;L of red liver paste into the dish. 
		Note: Trimming the end makes pipetting the viscous liver paste easier. To prevent the liver from floating on the surface, touch the tip to the bottom of the dish while dispensing.</li>
		<li>Allow animals to feed for 30 min.</li>
		<li>Score the number of animals that have eaten by placing the animals on a white background or examining them under a microscope with 10-20X magnification.</li>
		<li>After feeding, remove liver from the dish and clean.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

This protocol describes a method of selective ablation of the pharynx using sodium azide. Other targeted ablation studies in planarians have used modified surgery to remove photoreceptors21 or pharmacological treatment to ablate dopaminergic neurons22. One significant advantage of chemical amputation over existing methods is that it does not require surgery. The rigid structure of the pharynx compared to the rest of the planarian body facilitates its complete removal from t...

Regeneration is a phenomenon that occurs throughout the animal kingdom, with regenerative capacities ranging from full body regeneration in certain invertebrates to more restricted abilities in vertebrates1. Replacement of functional tissues is a complex process and often entails the simultaneous restoration of multiple cell types. For example, to regenerate the salamander limb, osteoblasts, chondrocytes, neurons, muscles, and epithelial cells need to be replaced2. These newly generated cell types also need to be organized properly to facilitate new limb function. Understanding these complex processes requires techniques that focus on regeneration of specific cell types and their integration into organs.
One of the strategies employed to simplify the study of regenerative responses is the targeted ablation of either certain cell types or larger collections of tissues. For example, in zebrafish, expression of nitroreductase in specific cell types leads to their destruction after application of metronidazole3,4. In Drosophila larvae, expressing pro-apoptotic genes under tissue-specific promoters can selectively ablate specific regions of the imaginal disc5,6. Both of these strategies cause rapid but controlled damage, and have been used to dissect the molecular and cellular mechanisms responsible for regeneration.
In this manuscript, we describe a method to selectively ablate an entire organ called the pharynx in the planarian Schmidtea mediterranea. Planarians are a classical model of regeneration, known for their prolific regenerative ability, where even minute fragments can regrow whole animals 7,8. They have a large, heterogeneous population of stem cells consisting of both pluripotent cells and lineage-restricted progenitors9,10,11. These cells proliferate and differentiate to replace all missing tissues, including the pharynx, nervous, digestive and excretory systems, and muscle and epithelial cells9,10,12. While we know that these stem cells initiate regeneration, we do not fully comprehend the molecular mechanisms that drive them to replace all these different cell types. Defined wounding methods that elicit precise stem cell responses can help delineate this complex process.
The pharynx is a large, cylindrical tube required for feeding, and contains neurons, muscle, epithelial and secretory cells13,29. Normally hidden in a pouch on the ventral side of the animal, it extends through the animal&#39;s single body opening upon sensing the presence of food. To selectively amputate the pharynx, we soak planarians in a chemical called sodium azide, a commonly used anesthetic in C. elegans14,15,16. Its use in planarians was first reported by Adler et al., in 201412. Within minutes of exposure to sodium azide, planarians extrude their pharynges, and with gentle agitation, the pharynx detaches from the animal. We refer to this complete and selective loss of the pharynx as &#34;chemical amputation&#34;. One week after amputation, a fully functional pharynx is restored12. Because the pharynx is required for feeding, functional regeneration can be measured by monitoring feeding behavior. Below, we describe the protocol for chemical amputation, and for assessing the regeneration of the pharynx and restoration of feeding behavior.

<table border="0" cellpadding="0" cellspacing="0" width="1042">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:447px;">Calcium Chloride Dihydrate (CaCl2)</td>
			<td style="width:204px;">Fisher Chemical</td>
			<td style="width:155px;">C79-3</td>
			<td style="width:236px;">Montju&#239;c salt solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Magnesium Sulfate Anhydrous (MgSO4)</td>
			<td>Fisher Chemical</td>
			<td>M65-3</td>
			<td>Montju&#239;c salt solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Magnesium Chloride Hexahydrate (MgCl2)</td>
			<td>Acros/VWR</td>
			<td>41341-5000</td>
			<td>Montju&#239;c salt solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Potassium Chloride (KCl)</td>
			<td>Acros Organics/VWR</td>
			<td>196770010</td>
			<td>Montju&#239;c salt solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sodium Chloride (NaCl)</td>
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			<td>Acros Organics/VWR</td>
			<td>123360010</td>
			<td>Montju&#239;c salt solution</td>
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			<td height="21" style="height:21px;">Nalgene autoclavable polypropylene copolymer lowboy with spigot</td>
			<td>ThermoFisher Scientific/VWR</td>
			<td>2324-0015</td>
			<td>Storing planaria water</td>
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			<td height="21" style="height:21px;">Instant Ocean Sea Salt</td>
			<td>Spectrum Brands</td>
			<td>Amazon</td>
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			<td>Amazon</td>
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			<td>309647</td>
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			<td height="21" style="height:21px;">Petri dishes-35mm/60mm</td>
			<td>Greiner Bio-One/VWR</td>
			<td>82050-536/82050-544</td>
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			<td height="21" style="height:21px;">Plastic containers (various sizes)</td>
			<td>Ziploc</td>
			<td>Amazon</td>
			<td>Housing planarians in static culture</td>
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			<td>Sigma</td>
			<td>S2002</td>
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			<td height="21" style="height:21px;">Transfer pipette</td>
			<td>Globe Scientific</td>
			<td>138030</td>
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		<tr height="21">
			<td height="21" style="height:21px;">Forceps - Dumont Tweezer, Style 5</td>
			<td>Electron Microscopy Sciences&#160;</td>
			<td>72700-D (0203-5-PO)</td>
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		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Triton X-100</td>
			<td>Sigma</td>
			<td>T8787</td>
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			<td height="21" style="height:21px;">DAPI&#160;</td>
			<td>ThermoFisher</td>
			<td>62247</td>
			<td></td>
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			<td height="21" style="height:21px;">Streptavidin, Alexa Fluor 488 conjugate</td>
			<td>ThermoFisher</td>
			<td>S11223</td>
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			<td height="21" style="height:21px;">Glycerol</td>
			<td>Fisher BioReagents</td>
			<td>BP229-1</td>
			<td></td>
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chemical amputation and regeneration of the pharynx in the planarian schmidtea mediterranea

Planarians are flatworms that are extremely efficient at regeneration. They owe this ability to a large number of stem cells that can rapidly respond to any type of injury. Common injury models in these animals remove large amounts of tissue, which damages multiple organs. To overcome this broad tissue damage, we describe here a method to selectively remove a single organ, the pharynx, in the planarian Schmidtea mediterranea. We achieve this by soaking animals in a solution containing the cytochrome oxidase inhibitor sodium azide. Brief exposure to sodium azide causes extrusion of the pharynx from the animal, which we call &#34;chemical amputation.&#34;&#160;Chemical amputation removes the entire pharynx, and generates a small wound where the pharynx attaches to the intestine. After extensive rinsing, all amputated animals regenerate a fully functional pharynx in approximately one week. Stem cells in the rest of the body drive regeneration of the new pharynx. Here, we provide a detailed protocol for chemical amputation, and describe both histological and behavioral methods to assess successful amputation and regeneration.

Exposure to sodium azide disrupts the normal motility of planarians, causing animals to stretch and writhe. These movements force the pharynx to emerge from the ventral side of the animal, and after approximately 6 min in sodium azide solution, the white tip of the pharynx can be seen (Figure 1B-left panel). A few minutes later, animals actively contract and fully extend the pharynx by forcefully pushing it out of the body. (<strong c...

Watch this Scientific Journal Video about Chemical Amputation and Regeneration of the Pharynx in the Planarian Schmidtea mediterranea at JoVE.com

Chemical Amputation and Regeneration of the Pharynx in the Planarian Schmidtea mediterranea

The planarian Schmidtea mediterranea is an excellent model for studying stem cells and tissue regeneration. This publication describes a method to selectively remove one organ, the pharynx, by exposing animals to the chemical sodium azide. This protocol also outlines methods for monitoring pharynx regeneration.

Chemical Amputation and Regeneration of the Pharynx in the Planarian Schmidtea mediterranea

Planarians are flatworms that are extremely efficient at regeneration. They owe this ability to a large number of stem cells that can rapidly respond to ...

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10.6K Views.  Cornell University. The planarian Schmidtea mediterranea is an excellent model for studying stem cells and tissue regeneration. This publication describes a method to selectively remove one organ, the pharynx, by exposing animals to the chemical sodium azide. This protocol also outlines methods for monitoring pharynx regeneration. 

Video: Chemical Amputation and Regeneration of the Pharynx in the Planarian Schmidtea mediterranea

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

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

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

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

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

Use of the TetON System to Study Molecular Mechanisms of Zebrafish Regeneration

The zebrafish has become a very important model organism for studying vertebrate development, physiology, disease, and tissue regeneration. A thorough understanding of the molecular and cellular mechanisms involved requires experimental tools that allow for inducible, tissue-specific manipulation of gene expression or signaling pathways. Therefore, we and others have recently adapted the TetON system for use in zebrafish. The TetON system facilitates temporally and spatially-controlled gene expression and we have recently used this tool to probe for tissue-specific functions of Wnt/beta&#8211;catenin signaling during zebrafish tail fin regeneration. Here we describe the workflow for using the TetON system to achieve inducible, tissue-specific gene expression in the adult regenerating zebrafish tail fin. This includes the generation of stable transgenic TetActivator and TetResponder lines, transgene induction and techniques for verification of tissue-specific gene expression in the fin regenerate. Thus, this protocol serves as blueprint for setting up a functional TetON system in zebrafish and its subsequent use, in particular for studying fin regeneration.

Here we outline the workflow for using the TetON system to achieve tissue-specific gene expression in the adult regenerating zebrafish tail fin.

The zebrafish has become a very important model organism for studying vertebrate development, physiology, disease, and tissue regeneration. A thorough ...

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Inducing Complete Polyp Regeneration from the Aboral Physa of the Starlet Sea Anemone Nematostella vectensis

Cnidarians, and specifically Hydra, were the first animals shown to regenerate damaged or severed structures, and indeed such studies arguably launched modern biological inquiry through the work of Trembley more than 250 years ago. Presently the study of regeneration has seen a resurgence using both &#34;classic&#34; regenerative organisms, such as the Hydra, planaria and Urodeles, as well as a widening spectrum of species spanning the range of metazoa, from sponges through mammals. Besides its intrinsic interest as a biological phenomenon, understanding how regeneration works in a variety of species will inform us about whether regenerative processes share common features and/or species or context-specific cellular and molecular mechanisms. The starlet sea anemone, Nematostella vectensis, is an emerging model organism for regeneration. Like Hydra, Nematostella is a member of the ancient phylum, cnidaria, but within the class anthozoa, a sister clade to the hydrozoa that is evolutionarily more basal. Thus aspects of regeneration in Nematostella will be interesting to compare and contrast with those of Hydra and other cnidarians. In this article, we present a method to bisect, observe and classify regeneration of the aboral end of the Nematostella adult, which is called the physa. The physa naturally undergoes fission as a means of asexual reproduction, and either natural fission or manual amputation of the physa triggers re-growth and reformation of complex morphologies. Here we have codified these simple morphological changes in a Nematostella Regeneration Staging System (the NRSS). We use the NRSS to test the effects of chloroquine, an inhibitor of lysosomal function that blocks autophagy. The results show that the regeneration of polyp structures, particularly the mesenteries, is abnormal when autophagy is inhibited.

Inducing Complete Polyp Regeneration from the Aboral Physa of the Starlet Sea Anemone Nematostella vectensis

Here we demonstrate how to induce and monitor regeneration in the Starlet Sea Anemone Nematostella vectensis, a model cnidarian anthozoan. We demonstrate how to amputate and categorize regeneration using a morphological staging system, and we use this system to reveal a requirement for autophagy in regenerating polyp structures.

Cnidarians, and specifically Hydra, were the first animals shown to regenerate damaged or severed structures, and indeed such studies arguably launched ...

Imaging of Cell Shape Alteration and Cell Movement in Drosophila Gastrulation Using DE-cadherin Reporter Transgenic Flies

Gastrulation is the first set of morphologically dynamic events that occur during the embryonic development of multicellular animals such as Drosophila. This morphological alteration is also recognized as epithelial to mesenchymal transition (EMT). Dysregulation of EMT is associated with fibrosis and cancer metastasis. There is emerging evidence that EMT is controlled by a number of molecular mechanisms. As such, many key genes that control apical constriction are also known to be important factors in the EMT observed in cancer metastasis. Like EMT during Drosophila gastrulation, epithelial cells can be induced to change their shape and be reprogrammed to redirect cell fate towards various other cell types. Here we provide a robust imaging method of Drosophila gastrulation to assay the initiation of morphogenetic cellular movements and cell fate identification during this stage of embryonic development. Using this method, we identify cell rearrangement at the time of gastrulation and demonstrate the importance of apical constriction during gastrulation using GFP labeled DE-cadherin.

Imaging of Cell Shape Alteration and Cell Movement in Drosophila Gastrulation Using DE-cadherin Reporter Transgenic Flies

Herein we describe a procedure to capture live images of Drosophila gastrulation. This has enabled us to better understand the apical constriction involved in early development and further analyze mechanisms governing cellular movements during tissue structure modification.

Gastrulation is the first set of morphologically dynamic events that occur during the embryonic development of multicellular animals such as Drosophila. ...

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the proteins involved in these pathways are evolutionarily conserved between mammals and other eukaryotes, such as the fruit fly Drosophila melanogaster, suggesting that similar organizing principles exist during the development of these organisms. Importantly, Drosophila has been used extensively to identify cellular and molecular mechanisms regulating processes that are required in mammals including neurogenesis, differentiation, axonal guidance, and synaptogenesis. Flies have also been used successfully to model a variety of human neurodevelopmental diseases. Here we describe a protocol for the step-by-step microdissection, fixation, and immunofluorescent localization of proteins within the adult Drosophila brain. This protocol focuses on two example neuronal populations, mushroom body neurons and retinal photoreceptors, and includes optional steps to trace individual mushroom body neurons using Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. Example data from both wild-type and mutant brains are shown along with a brief description of a scoring criteria for axonal guidance defects. While this protocol highlights two well-established antibodies for investigating the morphology of mushroom body and photoreceptor neurons, other Drosophila brain regions and the localization of proteins within other brain regions can also be investigated using this protocol.

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

This protocol describes the dissection and immunostaining of adult Drosophila melanogaster brain tissues. Specifically, this protocol highlights the use of Drosophila mushroom body and photoreceptor neurons as example neuronal subsets that can be accurately used to uncover general principles underlying many aspects of neuronal development.

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the ...

Application of RNAi and Heat-shock-induced Transcription Factor Expression to Reprogram Germ Cells to Neurons in C. elegans

Studying the cell biological processes during converting the identities of specific cell types provides important insights into mechanism that maintain and protect cellular identities. The conversion of germ cells into specific neurons in the nematode Caenorhabditis elegans (C. elegans) is a powerful tool for performing genetic screens in order to dissect regulatory pathways that safeguard established cell identities. Reprogramming of germ cells to a specific type of neurons termed ASE requires transgenic animals that allow broad over-expression of the Zn-finger transcription factor (TF) CHE-1. Endogenous CHE-1 is expressed exclusively in two head neurons and is required to specify the glutamatergic ASE neurons fate, which can easily be visualized by the gcy-5prom::gfp reporter. A trans gene containing the heat-shock promoter-driven che-1 gene expression construct allows broad mis-expression of CHE-1 in the entire animal upon heat-shock treatment. The combination of RNAi against the chromatin-regulating factor LIN-53 and heat-shock-induced che-1 over-expression leads to reprogramming of germ cell into ASE neuron-like cells. We describe here the specific RNAi procedure and appropriate conditions for heat-shock treatment of transgenic animals in order to successfully induce germ cell to neuron conversion.

Application of RNAi and Heat-shock-induced Transcription Factor Expression to Reprogram Germ Cells to Neurons in C. elegans

This protocol describes how to study cellular processes during cell fate conversion in Caenorhabditis elegans in vivo. Using transgenic animals, allowing heat-shock promoter-driven overexpression of the neuron fate-inducing transcription factor CHE-1 and RNAi-mediated depletion of the chromatin-regulating factor LIN-53 germ cell to neuron reprogramming can be observed in vivo.

Studying the cell biological processes during converting the identities of specific cell types provides important insights into mechanism that maintain ...

Methods for the Study of Regeneration in Stentor

Cells need to be able to regenerate their parts to recover from external perturbations. The unicellular ciliate Stentor coeruleus is an excellent model organism to study wound healing and subsequent cell regeneration. The Stentor genome became available recently, along with modern molecular biology methods, such as RNAi. These tools make it possible to study single-cell regeneration at the molecular level. The first section of the protocol covers establishing Stentor cell cultures from single cells or cell fragments, along with general guidelines for maintaining Stentor cultures. Culturing Stentor in large quantities allows for the use of valuable tools like biochemistry, sequencing, and mass spectrometry. Subsequent sections of the protocol cover different approaches to inducing regeneration in Stentor. Manually cutting cells with a glass needle allows studying the regeneration of large cell parts, while treating cells with either sucrose or urea allows studying the regeneration of specific structures located at the anterior end of the cell. A method for imaging individual regenerating cells is provided, along with a rubric for staging and analyzing the dynamics of regeneration. The entire process of regeneration is divided in three stages. By visualizing the dynamics of the progression of a population of cells through the stages, the heterogeneity in regeneration timing is demonstrated.

Methods for the Study of Regeneration in Stentor

The giant ciliate, Stentor coeruleus, is an excellent system to study regeneration and wound healing. We present procedures for establishing Stentor cell cultures from single cells or cell fragments, inducing regeneration by cutting cells, chemically inducing the regeneration of membranellar band and oral apparatus, imaging, and analysis of cell regeneration.

Cells need to be able to regenerate their parts to recover from external perturbations. The unicellular ciliate Stentor coeruleus is an excellent model ...

Adult Mouse Digit Amputation and Regeneration: A Simple Model to Investigate Mammalian Blastema Formation and Intramembranous Ossification

Here, we present a protocol of adult mouse distal terminal phalanx (P3) amputation, a procedurally simple and reproducible mammalian model of epimorphic regeneration, which involves blastema formation and intramembranous ossification analyzed by fluorescence immunohistochemistry and sequential in-vivo microcomputed tomography (&#956;CT). Mammalian regeneration is restricted to amputations transecting the distal region of the terminal phalanx (P3); digits amputated at more proximal levels fail to regenerate and undergo fibrotic healing and scar formation. The regeneration response is mediated by the formation of a proliferative blastema, followed by bone regeneration via intramembranous ossification to restore the amputated skeletal length. P3 amputation is a preclinical model to investigate epimorphic regeneration in mammals, and is a powerful tool for the design of therapeutic strategies to replace fibrotic healing with a successful regenerative response. Our protocol uses fluorescence immunohistochemistry to 1) identify early-and-late blastema cell populations, 2) study revascularization in the context of regeneration, and 3) investigate intramembranous ossification without the need for complex bone stabilization devices. We also demonstrate the use of sequential in vivo &#956;CT to create high resolution images to examine morphological changes after amputation, as well as quantify volume and length changes in the same digit over the course of regeneration. We believe this protocol offers tremendous utility to investigate both epimorphic and tissue regenerative responses in mammals.

Here, we present a protocol of adult mouse terminal phalanx amputation to investigate mammalian blastema formation and intramembranous ossification, analyzed by fluorescent immunohistochemistry and sequential in-vivo microcomputed tomography.

Here, we present a protocol of adult mouse distal terminal phalanx (P3) amputation, a procedurally simple and reproducible mammalian model of epimorphic ...

Isolation of Rat Adipose Tissue Mesenchymal Stem Cells for Differentiation into Insulin-producing Cells

Mesenchymal stem cells (MSCs)-especially those isolated from adipose tissue (Ad-MSCs)-have gained special attention as a renewable, abundant source of stem cells that does not pose any ethical concerns. However, current methods to isolate Ad-MSCs are not standardized and employ complicated protocols that require special equipment. We isolated Ad-MSCs from the epididymal fat of Sprague-Dawley rats using a simple, reproducible method. The isolated Ad-MSCs usually appear within 3 days post isolation, as adherent cells display fibroblastic morphology. Those cells reach 80% confluency within 1 week of isolation. Afterward, at passage 3-5 (P3-5), a full characterization was carried out for the isolated Ad-MSCs by immunophenotyping for characteristic MSC cluster of differentiation (CD) surface markers such as CD90, CD73, and CD105, as well as inducing differentiation of these cells down the osteogenic, adipogenic, and chondrogenic lineages. This, in turn, implies the multipotency of the isolated cells. Furthermore, we induced the differentiation of the isolated Ad-MSCs toward the insulin-producing cells (IPCs) lineage via a simple, relatively short protocol by incorporating high glucose Dulbecco&#39;s modified Eagle medium (HG-DMEM), &#946;-mercaptoethanol, nicotinamide, and exendin-4. IPCs differentiation was genetically assessed, firstly, via measuring the expression levels of specific &#946;-cell markers such as MafA, NKX6.1, Pdx-1, and Ins1, as well as dithizone staining for the generated IPCs. Secondly, the assessment was also carried out functionally by a glucose-stimulated insulin secretion (GSIS) assay. In conclusion, Ad-MSCs can be easily isolated, exhibiting all MSC characterization criteria, and they can indeed provide an abundant, renewable source of IPCs in the lab for diabetes research.

Adipose tissue-derived mesenchymal stem cells (Ad-MSCs) can be a potential source of MSCs that differentiate into insulin-producing cells (IPCs). In this protocol, we provide detailed steps for the isolation and characterization of rat epididymal Ad-MSCs, followed by a simple, short protocol for the generation of IPCs from the same rat Ad-MSCs.

Mesenchymal stem cells (MSCs)-especially those isolated from adipose tissue (Ad-MSCs)-have gained special attention as a renewable, abundant source of ...