This work was supported by a grant from the MRC.

1. Mating and collecting eggs
<ol>
 <li>Female and male medaka can easily be distinguished by the size and shape of the anal and dorsal fins. The anal fins of the males are larger and parallelogram-shaped compared to that of the females. The female anal fin is smaller and triangle shaped. The male dorsal fin has a clearly visible deep notch between the last two rays.</li>
 <li>Medaka lay up to 40 eggs every day and medaka females carry eggs clustered at their belly by attachment filaments for several hours before they are stripped off at plants in the tank.</li>
 <li>Females can be caught in a net and held inside gently from both sides by one hand. Eggs can then be gently teased from the belly with the index finger of the other hand. The female can then be returned to the tank. Eggs should be transferred to a 6 cm Petri dish filled with embryo media by blunt forceps or a finger.</li>
 <li>Medaka pairs rarely fight when they are kept in a small mating box. Therefore, they can be kept in one small tank with water flow so that they can be fed and eggs can be collected from the same pair every day for a couple of months.</li>
 <li>It is important that female fish should not be kept without males, since ovulation occurs everyday in medaka females. Otherwise, females cannot deliver eggs and become unwell.</li>
</ol>
2. Cleaning embryos
<ol>
 <li>Transfer unclustered eggs with a glass pipette to sandpaper (p2000 grit size, waterproof) placed in the lid of a 9cm Petri dish. Remove excess medium but ensure a sufficient volume remains as to prevent drying of embryos.</li>
 <li>Gently roll embryos on sandpaper using the forefinger, applying minimal pressure and keeping finger parallel to the surface of the sand paper for around 45-60 seconds to remove some of the outer surface hairs of the chorion. Do not roll more than 5-7 embryos at once this precaution will minimize the risk of crushing embryos beneath each other.</li>
 <li>Transfer cleaned embryos to a fresh dish of embryo medium.</li>
</ol>
3. Making agarose plates for holding embryos during microinjection
Embryos are held in troughs during injection made with a plexiglass mould in 1.5% agarose (Figure 1). The concentration of agarose is important to hold embryos properly.
<ol>
 <li>First, make a frame of agarose to hold the plexiglass mould. Pour 1.5% agarose in water to a 0.5cm depth in a 9cm Petri dish and wait until it completely solidifies. Cut out an area of the agarose just larger than the size of the mould. </li>
 <li>Pour 1.5% agarose to fill the cut area and place the mould so that no bubbles are trapped and wait until agarose solidifies. This agarose plate can be stored in the refrigerator to speed up solidifaction.</li>
 <li>Remove the mould to create troughs to hold the eggs. Add enough embryo medium to immerse the mould, cover and store at 4&deg;C until needed. 
 <img src="/files/ftp_upload/1937/1937fig1.jpg" alt="Figure 1" /> 
 
 Figure 1. Schematic of custom-made mould used to prepare troughs in agarose plates for microinjection. </li>
</ol>
4. Microinjection of medaka embryos
In medaka, DNA, RNA, morpholino oligonucleotides and tracer dyes are microinjected through the chorion into the cytoplasm of the 1 to 64-cell stage embryo rather than the yolk as in zebrafish. Since the cytoplasm is smaller and less visible in medaka, practice by injecting dyes such as rhodamine-dextran is recommended to develop this skill. Addition of phenol red as a tracer is also encouraged to allow confirmation of injections.
As the chorion surrounding medaka embryos is tough, a short, strong and wide injection needle is required (Figure 2). Prepare this strong wide injection needle from a filament-containing glass capillary.
<ol>
 <li>Prepare injection solution with phenol red as a tracer and load it into the needle from the back with an Eppendorf microloader. Connect the needle to the air pump injector.</li>
 <li>Eggs must be collected and unclustered as soon as possible after fertilization to inject at the one-cell stage. More embryos can be injected by storing them on ice (not more than 30 minutes) to stop cell division. 
 <img src="/files/ftp_upload/1937/1937fig2.jpg" alt="Figure 2" /> 
 
 Figure 2. Comparison of medaka and zebrafish needles. Note the tip of the MF needle (top) is shorter and wider hence stronger for puncturing the tough chorion surrounding medaka embryos.</li>
 <li>Transfer eggs into the agarose plate and align them in the troughs with forceps. Just before injection, five eggs should be rotated so the cell membrane of the one-cell cytoplasm can be hit by the needle perpendicularly. The cytoplasm of the one-cell can be found by looking for an area devoid of small oil droplets which is opposite to the side of the egg with densest oil droplets. The identified area can be confirmed to be the cytoplasm by viewing from the side.</li>
 <li>Place the needle close to the eggs. Open the tip of the needle by gently touching the tip of the needle to the chorion or using microforceps (Figure 3). A small amount of injection substance should be seen to flow out.</li>
 <li>Puncture through the chorion to place the tip of the needle inside of the chorion. Adjust the holding pressure of the injector to be relatively high, to ensure no reflux occurs due to the high pressure upon puncture through the tough chorion (settings of 80-100hPA for holding pressure and 500-700hPA for injection pressure).</li>
 <li>Insert the tip of the needle into the cytoplasm by sharply poking its membrane. 
 
 Avoid inserting the tip of the needle into the yolk. Unlike zebrafish, material injected into the yolk will not be transferred into the cytoplasm. Injected liquids have a distinct boundary when injected into the yolk whereas the boundary is blurred when in the cytoplasm (Figure 4). 
 
 Occasionally the needle may become blocked by agarose/debris. Forceps can be used to touch/rebreak the end of the needle and remove the blockage. If the needle breaks at any point during injection, bubbling will be seen in the embryo medium and the substance you are injecting will be lost. Injections should be carried out systematically from top to bottom along agarose channels and from left to right across the mould.</li>
 <li>Injected embryos should be transferred to a clean dish containing embryo medium and developed at the appropriate temperature. 
 <img src="/files/ftp_upload/1937/1937fig3.jpg" alt="Figure 3" /> 
 
 Figure 3. Breaking the needle tip prior to injection. 1: Move tip of the needle close to the chorion of embryo; 2: Touch the chorion and move needle backwards and forwards gently to break tip; 3: When a small amount of injection substance flows out of the end of the needle this confirms breakage of the tip.</li>
</ol>
5. Representative results
<img src="/files/ftp_upload/1937/1937fig4.jpg" alt="Figure 4" /> 
 Figure 4. Representative images of injected medaka embryos. Panel A shows a representative image of correct injection of rhodamine-dextran into the cytoplasm of a one-cell stage medaka embryo. Panel B shows a representative image of incorrect injection of rhodamine-dextran into the yolk sac of a one-cell stage medaka embryo. Panels C and D show the embryos from panels A and B respectively at approximately 30 hours post-injection. EM: embryo, anterior is upwards in panels C and D and view is dorsal. Note the body of the embryo in panel D is not red as the dye was incorrectly injected into the yolk sac. Arrows in B and D indicate rhodamine-dextran injected incorrectly into the yolk sac note the distinct circular boundary. Dotted lines in A and B indicate outline of cytoplasm of one-cell, view is lateral.

<ol>
	<li>Furutani-Seiki, M., Sasado, T., Morinaga, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+systematic+genome-wide+screen+for+mutations+affecting+organogenesis+in+Medaka,+Oryzias+latipes.">A systematic genome-wide screen for mutations affecting organogenesis in Medaka, Oryzias latipes.</a> Mech Dev. 121, 647-658 (2004).</li><li>Furutani-Seiki, M., Wittbrodt, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Medaka+and+zebrafish,+an+evolutionary+twin+study.">Medaka and zebrafish, an evolutionary twin study.</a> Mech. Dev. 121, 629-6237 (2004).</li></ol>

In this video, we demonstrate a method for cell tracer, mRNA, DNA or anti-sense oligonucleotide injection into one-cell stage medaka embryos. This technique has allowed us to study various developmental processes in-vivo in real-time. This process and the subsequent detailed analysis it affords has the potential to significantly enhance our understanding of vertebrate organogenesis and the underlying cell biology. Such knowledge may be important for, and yield therapeutic applications in regenerative medicine.

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>4" fine net</td>
<td>Tools</td>
<td>J &amp; K Aquatics Ltd.</td>
<td>MD001</td>
<td>Used to capture fish during egg harvesting.</td>
</tr>
<tr>
<td>Embryo medium</td>
<td>Reagent</td>
<td>Made in-house</td>
<td>&nbsp;</td>
<td>200ml 50X stock solution, 1ml 1%methyleneblue in H2O/10 liter RO water (50X stock solution: NaCl 14.7g, KCl 0.6g, CaCl2 2H2O 2.4g, MgSO4 7H2O 4.0g/1 litre RO water.</td>
</tr>
<tr>
<td>Fine waterproof sandpaper - p2000 grit size </td>
<td>Tools</td>
<td>Hermes Abrasives Ltd.</td>
<td>&nbsp;</td>
<td>Very fine grit size for rolling/cleaning embryos with chorions.</td>
</tr>
<tr>
<td>Borosilicate glass capillaries</td>
<td>Tool</td>
<td>Harvard Apparatus</td>
<td>GC100-10F</td>
<td>For making microinjection needles.</td>
</tr>
<tr>
<td>Micropipette puller</td>
<td>Equipment</td>
<td>Narishige scientific instrument lab</td>
<td>Model PP-830</td>
<td>For making microinjection needles.</td>
</tr>
<tr>
<td>Eppendorf microloader</td>
<td>Tool</td>
<td>Eppendorf</td>
<td>5242956.003</td>
<td>For loading microinjection needles.</td>
</tr>
<tr>
<td>Microinjector apparatus </td>
<td>Equipment</td>
<td>Eppendorf</td>
<td>&nbsp;</td>
<td>Model 5242.</td>
</tr>
<tr>
<td>Micromanipulator</td>
<td>Equipment</td>
<td>Narishige</td>
<td>&nbsp;</td>
<td>Model MN151.</td>
</tr>
<tr>
<td>Agarose injection plate</td>
<td>Tools</td>
<td>Made in-house</td>
<td>&nbsp;</td>
<td>For holding embryos during injection.</td>
</tr>		</tbody>
		</table>
		</div>

microinjection of medaka embryos for use as a model genetic organism

In this video, we demonstrate the technique of microinjection into one-cell stage medaka embryos. Medaka is a small egg-laying freshwater fish that allows both genetic and embryological analyses and is one of the vertebrate model organisms in which genome-wide phenotype-driven mutant screens were carried out 1, as in zebrafish and the mouse. Divergence of functional overlap of related genes between medaka and zebrafish allows identification of novel phenotypes that are unidentifiable in a single species 2, thus medaka and zebrafish are complementary for genetic dissection of vertebrate genome functions.
To take advantage of medaka fish whose embryos are transparent and develop externally, microinjection is an essential technique to inject cell-tracers for labeling cells, mRNAs or anti-sense oligonucleotides for over-expressing and knocking-down genes of interest, and DNAs for making transgenic lines.

Watch this Scientific Journal Video about Microinjection of Medaka Embryos for use as a Model Genetic Organism at JoVE.com

Microinjection of Medaka Embryos for use as a Model Genetic Organism

Medaka and zebrafish are complementary for genetic dissection of vertebrate genome functions. This protocol highlights the key points for successful microinjection into medaka embryos, an important technique for embryological and genetic analysis using medaka and zebrafish in a laboratory.

In this video, we demonstrate the technique of microinjection into one-cell stage medaka embryos. Medaka is a small egg-laying freshwater fish that allows ...

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25.6K Views.  University of Bath. Medaka and zebrafish are complementary for genetic dissection of vertebrate genome functions. This protocol highlights the key points for successful microinjection into medaka embryos, an important technique for embryological and genetic analysis using medaka and zebrafish in a laboratory.

Video: Microinjection of Medaka Embryos for use as a Model Genetic Organism

Microinjection of A. aegypti Embryos to Obtain Transgenic Mosquitoes

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever.  The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever. The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue ...

Method for Culture of Early Chick Embryos ex vivo (New Culture)

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  the formation and patterning of brain, neural tube, somite and heart primordia. Applications of chick embryo culture include electroporation of DNA or RNA constructs in order to analyze gene function, grafts of growth factor coated beads such as FGFs and BMPs , as well as whole mount in situ hybridization and immunohistochemistry. This video demonstrates the different steps in chick embryo culture; First, the embryo is explanted in saline. Then, the embryo is centered on a glass ring. The membranes surrounding the embryo are lifted along the walls of the ring. The ring is then placed in a culture dish containing a pool of albumine. The culture dish is sealed and placed in a humid chamber, where the embryo is cultured for up to 24 hrs. Finally, the embryo is removed from the ring, fixed and processed for further applications. A troubleshooting guide is also presented.

Method for Culture of Early Chick Embryos ex vivo New Culture

This video demonstrates New culture, a method by which chick embryos are cultured outside the egg for up to 24 hr. This method enables one to study early development (primitive streak to 14 som.), a period corresponding to E7-9 in mouse. Applications of this technique include electroporation, in situ hybridization and immunohistochemistry.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

Double Whole Mount in situ Hybridization of Early Chick Embryos

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  gene expression in brain, neural tube, somite and heart primordia formation. This video demonstrates the different steps in 2-color whole mount in situ hybridization; First, the embryo is dissected from the egg and fixed in paraformaldehyde. Second, the embryo is processed for prehybridization. The embryo is then hybridized with two different probes, one coupled to DIG, and one coupled to FITC.  Following overnight hybridization, the embryo is incubated with DIG coupled antibody. Color reaction for DIG substrate is performed, and the region of interest appears blue. The embryo is then incubated with FITC coupled antibody. The embryo is processed for color reaction with FITC, and the region of interest appears red. Finally, the embryo is fixed and processed for phtograph and sectioning. A troubleshooting guide is also presented.

This video demonstrates 2-color whole mount in situ hybridization, a method by which the spatial and temporal expression pattern of 2 different genes can be visualized in young chick embryos. This method was originally introduced by David Wilkinson, Domingos Henrique, Phil Ingham and David Ish -Horowicz.

Method for Whole Mount Antibody Staining in Chick

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying antibody expression in developing brain, neural tube and somite. This video demonstrates the different steps in whole-mount antibody staining using HRP conjugated secondary antibodies; First, the embryo is dissected from the egg and fixed in paraformaldehyde. Second, endogenous peroxidase is inactivated; The embryo is then exposed to primary antibody. After several washes, the embryo is incubated with secondary antibody conjugated to HRP. Peroxidase activity is revealed using reaction with diaminobenzidine substrate. Finally, the embryo is fixed and processed for photography and sectioning. The advantage of this method over the use of fluorescent antibodies is that embryos can be processed for wax sectioning, thus enabling the study of antigen sites in cross section. This method was originally introduced by Jane Dodd and Tom Jessell 1.

This video demonstrates whole mount immunohistochemistry, a method by which the spatial and temporal expression pattern of an antigen can be visualized in young chick embryos. This method was originally introduced by Jane Dodd and Tom Jessell.

Assay for Neural Induction in the Chick Embryo

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying the formation and initial patterning of the nervous system. This video demonstrates how to graft organizer tissue into a host, a method by which Hensen s node (the organizer in the chick embryo) is grafted to a host competent ectoderm. The organizer graft instructs overlying na ve tissue to adopt a neural fate via neural inducing signals. This mechanism is referred to as neural induction, and constitutes the initial step in the formation of brain and spinal cord in amniotes. This method is essentially used for the characterization of putative neural inducing molecules in chick. This video demonstrates the different steps in the assay for neural induction; First, the donnor embryo is explanted and pinned on a dish. Then, the host embryo is prepared for New culture. The graft is excised and transplanted to the host area pellucida margin. The host is cultured for 18-22 hrs. The assembly is fixed and processed for further applications (e.g. in situ hybridization). This method was originally devised by Waddington 1,2 and Gallera 3,4.

Neural induction is the first step in the formation of the brain. It is a mechanism by which Hensen's node (organizer), instructs adjacent tissue to adopt a neural fate, i.e. to give rise to the nervous system. This video demonstrates an assay for neural induction in chick embryo.

Use of Rotorod as a Method for the Qualitative Analysis of Walking in Rat

High speed videoanalysis of the details of movement can provide a source of information about qualitative aspects of walking movements. When walking on a rotorod, animals remain in approximately the same place making repetitive movements of stepping. Thus the task provides a rich source of information on the details of foot stepping movements. Subjects were hemi-Parkinson analogue rats, produced by injection of 6-hydroxydopamine (6-OHDA) into the right nigrostriatal bundle to deplete nigrostriatal dopamine (DA). The present report provides a video analysis illustration of animals previously were filmed from frontal, lateral, and posterior views as they walked (15). Rating scales and frame-by-frame replay of the video records of stepping behavior indicated that the hemi-Parkinson rats were chronically impaired in posture and limb use contralateral to the DA-depletion. The contralateral limbs participated less in initiating and sustaining propulsion than the ipsilateral limbs. These deficits secondary to unilateral DA-depletion show that the rotorod provides a use task for the analysis of stepping movements.

The rotorod test is used to assess motor status in the walking movement of hemi-Parkinson analogue rats.

High speed videoanalysis of the details of movement can provide a source of information about qualitative aspects of walking movements. When walking on a ...

Microinjection of Zebrafish Embryos to Analyze Gene Function

One of the advantages of studying zebrafish is the ease and speed of manipulating protein levels in the embryo.  Morpholinos, which are synthetic oligonucleotides with antisense complementarity to target RNAs, can be added to the embryo to reduce the expression of a particular gene product.  Conversely, processed mRNA can be added to the embryo to increase levels of a gene product.  The vehicle for adding either mRNA or morpholino to an embryo is microinjection.  Microinjection is efficient and rapid, allowing for the injection of hundreds of embryos per hour.  This video shows all the steps involved in microinjection.  Briefly, eggs are collected immediately after being laid and lined up against a microscope slide in a Petri dish.  Next, a fine-tipped needle loaded with injection material is connected to a microinjector and an air source, and the microinjector controls are adjusted to produce a desirable injection volume.  Finally, the needle is plunged into the embryo's yolk and the morpholino or mRNA is expelled.

This video shows how morpholino or mRNA can be injected into zebrafish embryos at the one-cell stage to decrease or increase the level of specific gene products during subsequent development.

One of the advantages of studying zebrafish is the ease and speed of manipulating protein levels in the embryo.  Morpholinos, which are synthetic ...

Genetic Studies of Human DNA Repair Proteins Using Yeast as a Model System 

Understanding the roles of human DNA repair proteins in genetic pathways is a formidable challenge to many researchers. Genetic studies in mammalian systems have been limited due to the lack of readily available tools including defined mutant genetic cell lines, regulatory expression systems, and appropriate selectable markers. To circumvent these difficulties, model genetic systems in lower eukaryotes have become an attractive choice for the study of functionally conserved DNA repair proteins and pathways. We have developed a model yeast system to study the poorly defined genetic functions of the Werner syndrome helicase-nuclease (WRN) in nucleic acid metabolism. Cellular phenotypes associated with defined genetic mutant backgrounds can be investigated to clarify the cellular and molecular functions of WRN through its catalytic activities and protein interactions. The human WRN gene and associated variants, cloned into DNA plasmids for expression in yeast, can be placed under the control of a regulatory plasmid element. The expression construct can then be transformed into the appropriate yeast mutant background, and genetic function assayed by a variety of methodologies. Using this approach, we determined that WRN, like its related RecQ family members BLM and Sgs1, operates in a Top3-dependent pathway that is likely to be important for genomic stability. This is described in our recent publication [1] at <a href="http://www.impactaging.com">www.impactaging.com</a>. Detailed methods of specific assays for genetic complementation studies in yeast are provided in this paper.

Genetic Studies of Human DNA Repair Proteins Using Yeast as a Model System

Genetic studies in yeast can be employed to investigate the molecular and cellular functions of human genes in cellular DNA metabolism. Methods are described for the genetic characterization of the human WRN gene product defective in the premature aging disorder Werner syndrome in functionally conserved pathways using yeast as a tractable model system.

Understanding the roles of human DNA repair proteins in genetic pathways is a formidable challenge to many researchers.  Genetic studies in mammalian ...

Dechorionation of Medaka Embryos and Cell Transplantation for the Generation of Chimeras

Medaka is a small egg-laying freshwater fish that allows both genetic and embryological analyses and is one of the three vertebrate model organisms in which genome-wide phenotype-driven mutant screens were carried out 1. Divergence of functional overlap of related genes between medaka and zebrafish allows identification of novel phenotypes that are unidentifiable in a single species 2, thus medaka and zebrafish are complementary for genetic dissection of the vertebrate genome functions. Manipulation of medaka embryos, such as dechorionation, mounting embryos for imaging and cell transplantation, are key procedures to work on both medaka and zebrafish in a laboratory. Cell transplantation examines cell autonomy of medaka mutations. Chimeras are generated by transplanting labeled cells from donor embryos into unlabeled recipient embryos. Donor cells can be transplanted to specific areas of the recipient embryos based on the fate maps 3 so that clones from transplanted cells can be integrated in the tissue of interest during development. Due to the hard chorion and soft embryos, manipulation of medaka embryos is more involved than in zebrafish. In this video, we show detailed procedures to manipulate medaka embryos.

Due to the hard chorion and soft embryos, manipulation of medaka embryos is more involved than in zebrafish. This video shows step-by-step procedures for how to manipulate medaka embryos, including dechorionation, mounting in agarose for imaging and cell transplantation for the production of chimeras. These procedures are essential to use medaka and zebrafish in a laboratory to take full advantage of their complementary features for the genetic dissection of vertebrate genome functions.

Medaka is a small egg-laying freshwater fish that allows both genetic and embryological analyses and is one of the three vertebrate model organisms in ...

Use of Time Lapse Microscopy to Visualize Anoxia-induced Suspended Animation in C. elegans Embryos

Caenorhabdits elegans has been used extensively in the study of stress resistance, which is facilitated by the transparency of the adult and embryo stages as well as by the availability of genetic mutants and transgenic strains expressing a myriad of fusion proteins1-4. In addition, dynamic processes such as cell division can be viewed using fluorescently labeled reporter proteins. The study of mitosis can be facilitated through the use of time-lapse experiments in various systems including intact organisms; thus the early C. elegans embryo is well suited for this study. Presented here is a technique by which in vivo imaging of sub-cellular structures in response to anoxic (99.999% N2; &lt;2 ppm O2) stress is possible using a simple gas flow through setup on a high-powered microscope. A microincubation chamber is used in conjunction with nitrogen gas flow through and a spinning disc confocal microscope to create a controlled environment in which animals can be imaged in vivo. Using GFP-tagged gamma tubulin and histone, the dynamics and arrest of cell division can be monitored before, during and after exposure to an oxygen-deprived environment. The results of this technique are high resolution, detailed videos and images of cellular structures within blastomeres of embryos exposed to oxygen deprivation.

Use of Time Lapse Microscopy to Visualize Anoxia-induced Suspended Animation in C. elegans Embryos

Described here is an in vivo technique to image sub-cellular structures in animals exposed to anoxia using a gas flow through microincubation chamber in conjunction with a spinning disc confocal microscope. This method is straightforward and flexible enough to suit a variety of experimental parameters and model systems.

Caenorhabdits elegans has been used extensively in the study of stress resistance, which is facilitated by the transparency of the adult and embryo stages ...