This work is supported by a startup fund from the Department of Biological Sciences at Purdue University.

Part 1: Preparations before microdissection 
<ol>
	<li>Solutions
	<ol class="lalpha">
		<li>E3 medium4 (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2 and 0.33 mM MgSO4).</li> 
		<li>Ringer's solution1, 5 (116 mM NaCl, 2.9 mM KCl, 1.8 mM CaCl2, 5 mM HEPES, pH7.2), filter sterilized.</li> 
		<li>5N NaOH.</li>
	</ol>
	</li> 
	<li>Chemical etching of tungsten needles 
	<ol class="lalpha"> 
		<li>Secure a small beaker containing 5N NaOH on a Petri dish by clay.</li> 
		<li>Attach a paper clip on the side of the beaker, so that it comes in contact with the NaOH solution.</li> 
		<li>Connect the negative electrode from a DC power supply to the paper clip and the positive electrode to one end of a short piece (~2cm) of tungsten wire.</li> 
		<li>Wrap the other end of tungsten wire with a small clay ball and dip this end into the NaOH solution.</li> 
		<li>Increase the voltage to approximately 3V and dip the wire up and down in the NaOH solution until a good etching rate is attained. A needle in good quality can usually be etched in 10 minutes.</li> 
		<li>The tungsten wire exposed to the solution will become gradually thinner. When the clay ball drops off, the other side of the wire that is still attached to the positive electrode will have a sharp needle shape.</li> 
		<li>Rinse the sharp needle with Ringer's solution or E3 to remove salt deposits.</li> <li>Attach the needle to a wooden applicator by tape or a needle holder for easy handling.</li> 
	</ol>
	</li> 
	<li>Culture plates 
	<ol class="lalpha"> 
		<li>The Falcon 60 x 15 mm polystyrene culture plates are ready to use for retinal microdissection.</li>
		<li>For dissecting RPE-attached retina, crush two embryos completely in a culture plate containing Ringer's solution to get rid of the adhesiveness of the surface 30 minutes before dissection. Shortly before dissection, wash the plate extensively with fresh Ringer's solution.</li>
		</ol>
	</li>
	<li>Embryos collection and staging 
	<ol class="lalpha">
		<li>Adult zebrafish are maintained as described4, 5.</li>
		<li>Separate the parent fish in static breeding tanks pairwise by a divider the night before breeding.</li>
		<li>Remove the divider after the room light is turned on in the next morning. Allow the parents to cross at 10-minute intervals every half an hour. Separate the fish after each crossing.</li>
		<li>Maintain the embryos collected from each crossing in E3 medium separately at 28&deg;C.</li>
		<li>Stage embryos at 10 to 12 hours post fertilization (hpf) and restage immediately before dissection at the specific time points, according to established criteria1, 6.</li>
	</ol>
	</li> 
</ol>
Part 2: Dissection - brain removal and eye exposure1
<ul>
	<li>The procedures described in this section are common for dissecting retina (Part 3A) and RPE-attached retina (Part 3B).</li>
	<li>All fine dissections are done by a tip of the Dumont forceps and assisted by another pair of Dumont forceps for positioning the tissue. A chemically-etched tungsten needle can be used for finer manipulations if necessary.</li>
	<li>All fine dissections are performed at ~ 5-8x magnification under an Olympus SZX16 microscope equipped with an 1X objective or equivalent.</li>
	<li>Maintain the embryos in E3 medium in a bench top incubator at 28&deg;C next to the microscope for easy embryo access during dissection.</li>
	<li>The microscope stage is warmed to 28&deg;C by a thermo plate to minimize the influence of temperature fluctuation on gene expression.</li>
</ul>
<ol>
	<li>Transfer an embryo at a specific developmental stage to Ringer's solution in a Falcon 60 X 15 mm culture plate prepared as described in Part 1#3.</li>
	<li>Cut off the head with part of the anterior trunk quickly from the body.</li> 
	<li>Pin the posterior end of this tissue on culture plate with the assisting forceps.</li> 
	<li>Open the head on the dorsal side starting from the anterior forehead.</li>
	<li>Remove the brain so that the medial side of the eyes is exposed and is facing upward for further manipulations.</li>
</ol>
 Part 3A: Retinal dissection1
 <ol> 
	<li>Prepare the embryo as described in Part 2.</li>
	<li>Carefully brush the exposed RPE on the medial side of the eye ball that is facing upward by the tip of the forceps until a small opening to the retina is seen.</li>
	<li>Continue the brushing and peeling action until the medial side of the retina is almost all exposed. Avoid scratching and punching the exposed retina.</li>
	<li>To remove the lateral RPE on the retina which is now facing downward, approach that and brush at an angle of approximately 45&deg; from the culture plate surface.</li>
	<li>Remove the RPE that is attached relatively firmly to the ora serrata by pinning the detached portion of the RPE to the culture plate for dissection, and then lift the retina gently.</li>
	<li>Roll the retina on the bottom of the culture dish to clean up the residual RPE. 
	<ol class="lalpha"> 
		<li>We have noticed that this particular Falcon polystyrene culture plate has a preferential adherence of RPE for approximately 20 minutes once an embryo is crushed. This property is utilized for the removal of RPE remnants. However, the retinal cells do stick to the surface to a lesser extent, which can be inspected closely under high magnification. One has to strike a balance between the completeness of RPE removal and retinal integrity.</li>
		<li>The lens often adheres to the culture plate and detaches from the retina during the rolling process. Occasionally, it is necessary to detach the lens by an etched tungsten needle with a swirling action on the lens surface.</li>
	</ol>
	</li>
	<li>These procedures can successfully remove RPE from the retina without compromising the integrity of the tissue. This is indicated by a good overall morphology (Figure 1B and B') and histology (Figure 1B"). In particular, the preservation of the extracellular matrix in between the photoreceptor layer and the RPE is exceptionally good (Figure 1B", arrow; compare with the histology from the whole embryo (Figure 1A")).</li>
</ol>
Part 3B: RPE-attached retinal dissection3 
<ol>
	<li>Prepare the embryo as described in Part 2.</li>
	<li>Pin the head to the culture plate by the assistant forceps. Lift the eye gently from the posterior lateral side and roll to the anterior side.
	<ol class="lalpha">
		<li>The presumptive choroid and scleral tissue on the outside of the RPE layer are attached relatively tightly to the skin and can be mostly peeled off by rolling the eye carefully to the anterior side.</li>
	</ol>
	</li>
	<li>Remove the lens with a tungsten needle after the lateral side of the eye is exposed and when the eye is still attached to the skin.</li>
	<li>These procedures can successfully preserve the whole RPE layer with the retina. Presumptive choroids and scleral tissue can be largely removed (Figure 1C, C' and C").</li>
</ol>
Part 4: Tissue sample collection for RNA work
<ol>
	<li>The dissected samples can be collected in TRIzol in an RNase-free microfuge tube for downstream RNA characterization as described before1.</li>
</ol>
Representative Results
<img src="/files/ftp_upload/2028/2028fig1.jpg" alt="Figure 1" /> Figure 1. (A) Lateral and (A') dorsal view of a zebrafish larval head at 52 hpf before dissection. (A") The corresponding histological section of the larval head in (A) &amp; (A'). (B) Lateral and (B') dorsal view of a dissected retina at 54 hpf. The surface of retina was intact from both lateral and dorsal views. (B") The corresponding histological section of the dissected retina in (B) &amp; (B'). The structure of the retina and retinal lamination was intact. In particular, the extracellular matrix between the photoreceptor layer and RPE (A", arrows) was well preserved in the dissected retina (B", arrows). (C) Lateral and (C') medial view of a dissected RPE-attached retina at 52 hpf. RPE layer was intact and continuous, which was also indicated by the histological section of the dissected tissue (C"). The white area in C' is the optic nerve (arrow). For histology, tissue samples were collected and fixed in 4% paraformaldehyde. Plastic embedding and sectioning of these samples were performed as described3. Scale bar = 50 &mu;m.

<ol>
	<li>Leung, Y. F., Dowling, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene+expression+profiling+of+zebrafish+embryonic+retina.">Gene expression profiling of zebrafish embryonic retina.</a> Zebrafish. 2, 269-283 (2005).</li><li>Leung, Y. F., Ma, P., Link, B. A., Dowling, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Factorial+microarray+analysis+of+zebrafish+retinal+development.">Factorial microarray analysis of zebrafish retinal development.</a> Proc Natl Acad Sci U S A. 105, 12909-12914 (2008).</li><li>Leung, Y. F., Ma, P., Dowling, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene+expression+profiling+of+zebrafish+embryonic+retinal+pigment+epithelium+in+vivo.">Gene expression profiling of zebrafish embryonic retinal pigment epithelium in vivo.</a> Invest Ophthalmol Vis Sci. 48, 881-890 (2007).</li><li>Nusslein-Volhard, C., Dahm, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Zebrafish : a practical approach. , (2002).</li><li>Westerfield, 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+zebrafish+book+:+a+guide+for+the+laboratory+use+of+zebrafish+(Danio+rerio).">The zebrafish book : a guide for the laboratory use of zebrafish (Danio rerio).</a> , (2000).</li><li>Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B., Schilling, T. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stages+of+embryonic+development+of+the+zebrafish.">Stages of embryonic development of the zebrafish.</a> Dev Dyn. 203, 253-310 (1995).</li><li>Fadool, J. M., Dowling, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish:+a+model+system+for+the+study+of+eye+genetics.">Zebrafish: a model system for the study of eye genetics.</a> Prog Retin Eye Res. 27, 89-110 (2008).</li></ol>

Experiments on animals were performed in accordance with the guidelines and regulations set forth by Purdue Animal Care and Use Committee.

Microdissection of zebrafish eye tissues can effectively obtain intact retinas and RPE-attached retinas. This substantially assists expression studies pertaining to a specific eye tissue (i.e. retina or RPE). In fact, we have successfully utilized these procedures to obtain RNA expression profiles of the whole retina1 and RPE3. The utility of these profiles is strongly supported by our recent identification of pathways and gene families that are perturbed in a retinal differentiation mutant2</s...

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<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<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>Cordless pestle motor</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>47747-370</td>
<td>&nbsp;</td>
<tr>
<td>DC power supply</td>
<td>&nbsp;</td>
<td>Lascar</td>
<td>PSU130</td>
<td>Any DC supply would work. The specific voltage of a different machine will need further optimization.</td>
</tr>
<tr>
<td>Disposable pestle &amp; microtube, 1.5 mL (DNase, RNase and pyrogen-free)</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>47747-366</td>
<td>These are used for tissue collection in TRIzol for expression analysis.</td>
</tr>
<tr>
<td>Dumont #5 forceps, Tips: 0.05 x 0.01mm, Inox</td>
<td>&nbsp;</td>
<td>World Precision Instruments</td>
<td>500341</td>
<td>Fine tip dimension is desirable but is not inflexible, as one may need to sharpen the tip from time to time.</td>
</tr>
<tr>
<td>Dumont #5SF forceps, Tips: 0.025 x 0.005mm, Inox</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>11252-00</td>
<td>Fine tip dimension is desirable but is not inflexible, as one may need to sharpen the tip from time to time.</td>
</tr>
<tr>
<td>Falcon polystyrene culture plates, 60 X 15 mm</td>
<td>&nbsp;</td>
<td>BD Biosciences</td>
<td>351007</td>
<td>These plates are used as dissection plates.</td>
</tr>
<tr>
<td>Olympus SZX16 Stereomicroscope</td>
<td>&nbsp;</td>
<td>Olympus</td>
<td>SZX16</td>
<td>Any stereomicroscope would work. We used Leica stereomicroscope in previous studies1-3 without any issues. We also use the 1X objective exclusively for the dissection even though we have a 2X objective installed. </td>
</tr>
<tr>
<td>Sharpening stone</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>29008-01</td>
<td>Use this to sharpen the tip of the forceps if necessary</td>
</tr>
<tr>
<td>Thermo plate</td>
<td>&nbsp;</td>
<td>Tokai Hit</td>
<td>MATS-U55SZX2B</td>
<td>This is used to maintain the temperature of the tissue throughout dissection and minimize the influence of temperature fluctuation on gene expression. We also put the whole microscope in an environmentally controlled room at 28&deg;C during dissection in previous studies1-3 with good success.</td>
</tr>
<tr>
<td>Trizol, 100 mL</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>15596-026</td>
<td>&nbsp;</td>
<tr>
<td>tungsten wire, 0.015 inch diameter</td>
<td>&nbsp;</td>
<td>World Precision Instruments</td>
<td>TGW1510</td>
<td>&nbsp;</td>
<tr>
<td>Wooden Applicator</td>
<td>&nbsp;</td>
<td>Puritan</td>
<td>807</td>
<td>This is used for holding the chemically-etched tungsten needle.</td>
</tr>		</tbody>
		</table>
		</div>

microdissection of zebrafish embryonic eye tissues

Zebrafish is a popular animal model for research on eye development because of its rapid ex utero development and good fecundity. By 3 days post fertilization (dpf), the larvae will show the first visual response. Many genes have been identified to control a proper eye development, but we are far from a complete understanding of the underlying genetic architecture. Whole genome gene expression profiling is a useful tool to elucidate genetic regulatory network for eye development. However, the small size of the embryonic eye in zebrafish makes it challenging to obtain intact and pure eye tissues for expression analysis. For example, the anterior-posterior length of the eye between day 2 and 3 is only approximately 200-300 &mu;m, while the diameter of the lens is less 100 &mu;m. Also, the retinal pigment epithelium (RPE) underlying the retina is just a single-layer epithelium. While gene expression profiles can be obtained from the whole embryo, they do not accurately represent the expression of these tissues. Therefore pure tissue must be obtained for a successful gene expression profiling of eye development. To address this issue, we have developed an approach to microdissect intact retina and retina with RPE attached from 1-3 dpf, which cover major stages of eye morphogenesis. All procedures can be done with fine forceps and general laboratory supplies under standard stereomicroscopes. For retinal dissection, the single-layer RPE is removed and peeled off by brushing action and the preferential adherence of the RPE remnants to the surface of the culture plate for dissection. For RPE-attached retinal dissection, the adherence of RPE to the dissection plate is removed before the dissection so that the RPE can be completely preserved with the retina. A careful lifting action of this tissue can efficiently separate the presumptive choroid and sclera. The lens can be removed in both cases by a chemically etched tungsten needle. In short, our approach can obtain intact eye tissues and has been successfully utilized to study tissue-specific expression profiles of zebrafish retina1, 2 and retinal pigment epithelium3.

Watch this Scientific Journal Video about Microdissection of Zebrafish Embryonic Eye Tissues at JoVE.com

Microdissection of Zebrafish Embryonic Eye Tissues

This article describes an approach to microdissect zebrafish retinas with and without retinal pigment epithelium attached, from one to three days postfertilization embryos.

Zebrafish is a popular animal model for research on eye development because of its rapid ex utero development and good fecundity. By 3 days post ...

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16.0K Views.  Purdue University. This article describes an approach to microdissect zebrafish retinas with and without retinal pigment epithelium attached, from one to three days postfertilization embryos. 

Video: Microdissection of Zebrafish Embryonic Eye Tissues

Laser Capture Microdissection of Mammalian Tissue

Laser capture microscopy, also known as laser microdissection (LMD), enables the user to isolate small numbers of cells or tissues from frozen or formalin-fixed, paraffin-embedded tissue sections. LMD techniques rely on a thermo labile membrane placed either on top of, or underneath, the tissue section. In one method, focused laser energy is used to melt the membrane onto the underlying cells, which can then be lifted out of the tissue section. In the other, the laser energy vaporizes the foil along a path "drawn" on the tissue, allowing the selected cells to fall into a collection device. Each technique allows the selection of cells with a minimum resolution of several microns. DNA, RNA, protein, and lipid samples may be isolated and analyzed from micro-dissected samples. In this video, we demonstrate the use of the Leica AS-LMD laser microdissection instrument in seven segments, including an introduction to the principles of LMD, initializing the instrument for use, general considerations for sample preparation, mounting the specimen and setting up capture tubes, aligning the microscope, adjusting the capture controls, and capturing tissue specimens. Laser-capture micro-dissection enables the investigator to isolate samples of pure cell populations as small as a few cell-equivalents. This allows the analysis of cells of interest that are free of neighboring contaminants, which may confound experimental results.

Laser capture microscopy, also known as laser microdissection (LMD), enables the user to isolate small numbers of cells or tissues from frozen or ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Lens Transplantation in Zebrafish and its Application in the Analysis of Eye Mutants

The lens plays an important role in the development of the optic cup[1,2]. Using the zebrafish as a model organism, questions regarding lens development can be addressed. The zebrafish is useful for genetic studies due to several advantageous characteristics, including small size, high fecundity, short lifecycle, and ease of care. Lens development occurs rapidly in zebrafish. By 72 hpf, the zebrafish lens is functionally mature [3]. Abundant genetic and molecular resources are available to support research in zebrafish. In addition, the similarity of the zebrafish eye to those of other vertebrates provides basis for its use as an excellent animal model of human defects[4-7]. Several zebrafish mutants exhibit lens abnormalities, including high levels of cell death, which in some cases leads to a complete degeneration of lens tissues [8]. To determine whether lens abnormalities are due to intrinsic causes or to defective interactions with the surrounding tissues, transplantation of a mutant lens into a wild-type eye is performed. Using fire-polished metal needles, mutant or wild-type lenses are carefully dissected from the donor animal, and transferred into the host. To distinguish wild-type and mutant tissues, a transgenic line is used as the donor. This line expresses membrane-bound GFP in all tissues, including the lens. This transplantation technique is an essential tool in the studies of zebrafish lens mutants.

Lens development involves interactions with other tissues. Several zebrafish eye mutants are characterized by an abnormally small lens size. Here we demonstrate a lens transplantation experiment to determine whether this phenotype is due to intrinsic causes or defective interactions with tissues that surround the lens.

The lens plays an important role in the development of the optic cup[1,2]. Using the zebrafish as a model organism, questions regarding lens development ...

Tissue Targeted Embryonic Chimeras: Zebrafish Gastrula Cell Transplantation

Certain fundamental questions in the field of developmental biology can only be answered when cells are placed in novel environments or when small groups of cells in a larger context are altered.  Watching how one cell interacts with and behaves in a unique environment is essential to characterizing cell functions.  Determining how the localized misexpression of a specific protein influences surrounding cells provides insightful information on the roles that protein plays in a variety of developmental processes.  Our lab uses the zebrafish model system to uniquely combine genetic approaches with classical transplantation techniques to generate genotypic or phenotypic chimeras.  We study neuron-glial cell interactions during the formation of forebrain commissures in zebrafish. This video describes a method that allows our lab to investigate the role of astroglial populations in the diencephalon and the roles of specific guidance cues that influence projecting axons as they cross the midline. Due to their transparency zebrafish embryos are ideal models for this type of ectopic cell placement or localized gene misexpression. Tracking transplanted cells can be accomplished using a vital dye or a transgenic fish line expressing a fluorescent protein. We demonstrate here how to prepare donor embryos with a vital dye tracer for transplantation, as well as how to extract and transplant cells from one gastrula staged embryo to another.  We present data showing ectopic GFP+ transgenic cells within the forebrain of zebrafish embryos and characterize the location of these cells with respect to forebrain commissures.  In addition, we show laser scanning confocal timelapse microscopy of Alexa 594 labeled cells transplanted into a GFP+ transgenic host embryo.  These data provide evidence that gastrula staged transplantation enables the targeted positioning of ectopic cells to address a variety of questions in Developmental Biology.

Zebrafish cell transplantation enables the combination of genetics and embryology to generate tissue specific chimeras. This video demonstrates gastrula staged cell transplantations that have allowed our lab to investigate the roles of astroglial populations and specific guidance cues during commissure formation in the forebrain.

Certain fundamental questions in the field of developmental biology can only be answered when cells are placed in novel environments or when small groups ...

RNA Isolation from Embryonic Zebrafish and cDNA Synthesis for Gene Expression Analysis

Many important and complex laboratory procedures require an input of high quality, intact RNA.  A degraded sample or the presence of impurities can lead to disastrous results in downstream experimental applications.  It is therefore, of utmost importance to use solid techniques with numerous safeguards and quality control checks to ensure a superior sample.  Herein, we detail a protocol to isolate total RNA from whole zebrafish embryos using a commercially available chemical denaturant and subsequent cleanup to remove traces of DNA and impurities using a commercial RNA isolation kit. As RNA is relatively unstable and easily prone to cleavage by RNAses, most protocols assay gene expression using a cDNA product that is directly synthesized from an RNA template. We detail a procedure to convert RNA into the more stable cDNA product using a commercially available kit. Throughout these procedures there are numerous quality control checks to ensure that the sample is not degraded or contaminated.  The end product of these protocols is cDNA that is suitable for microarray analysis, RT-PCR or long-term storage.    

The isolation of high quality, intact RNA is an essential step in many laboratory protocols. Here, we demonstrate RNA extraction from whole zebrafish embryos and cDNA synthesis for subsequent application in various experimental procedures including gene expression microarray analysis.

Many important and complex laboratory procedures require an input of high quality, intact RNA.  A degraded sample or the presence of impurities can lead ...

Microdissection of Black Widow Spider Silk-producing Glands

Modern spiders spin high-performance silk fibers with a broad range of biological functions, including locomotion, prey capture and protection of developing offspring 1,2. Spiders accomplish these tasks by spinning several distinct fiber types that have diverse mechanical properties. Such specialization of fiber types has occurred through the evolution of different silk-producing glands, which function as small biofactories. These biofactories manufacture and store large quantities of silk proteins for fiber production. Through a complex series of biochemical events, these silk proteins are converted from a liquid into a solid material upon extrusion.
Mechanical studies have demonstrated that spider silks are stronger than high-tensile steel 3. Analyses to understand the relationship between the structure and function of spider silk threads have revealed that spider silk consists largely of proteins, or fibroins, that have block repeats within their protein sequences 4. Common molecular signatures that contribute to the incredible tensile strength and extensibility of spider silks are being unraveled through the analyses of translated silk cDNAs. Given the extraordinary material properties of spider silks, research labs across the globe are racing to understand and mimic the spinning process to produce synthetic silk fibers for commercial, military and industrial applications. One of the main challenges to spinning artificial spider silk in the research lab involves a complete understanding of the biochemical processes that occur during extrusion of the fibers from the silk-producing glands.
Here we present a method for the isolation of the seven different silk-producing glands from the cobweaving black widow spider, which includes the major and minor ampullate glands [manufactures dragline and scaffolding silk] 5,6, tubuliform [synthesizes egg case silk] 7,8, flagelliform [unknown function in cob-weavers], aggregate [makes glue silk], aciniform [synthesizes prey wrapping and egg case threads] 9 and pyriform [produces attachment disc silk] 10. This approach is based upon anesthetizing the spider with carbon dioxide gas, subsequent separation of the cephalothorax from the abdomen, and microdissection of the abdomen to obtain the silk-producing glands. Following the separation of the different silk-producing glands, these tissues can be used to retrieve different macromolecules for distinct biochemical analyses, including quantitative real-time PCR, northern- and western blotting, mass spectrometry (MS or MS/MS) analyses to identify new silk protein sequences, search for proteins that participate in the silk assembly pathway, or use the intact tissue for cell culture or histological experiments.

Here we describe an efficient strategy to remove the silk-producing glands from the abdomen of female black widow spiders. This procedure allows the rapid isolation of the seven distinct silk-producing glands in a highly purified fashion, an important process for investigators studying spider silk production and fiber assembly.

Modern spiders spin high-performance silk fibers with a broad range of biological functions, including locomotion, prey capture and protection of ...

Immunostaining of Dissected Zebrafish Embryonic Heart

Zebrafish embryo becomes a popular in vivo vertebrate model for studying cardiac development and human heart diseases due to its advantageous embryology and genetics 1,2. About 100-200 embryos are readily available every week from a single pair of adult fish. The transparent embryos that develop ex utero make them ideal for assessing cardiac defects 3. The expression of any gene can be manipulated via morpholino technology or RNA injection 4. Moreover, forward genetic screens have already generated a list of mutants that affect different perspectives of cardiogenesis 5. 
Whole mount immunostaining is an important technique in this animal model to reveal the expression pattern of the targeted protein to a particular tissue 6. However, high resolution images that can reveal cellular or subcellular structures have been difficult, mainly due to the physical location of the heart and the poor penetration of the antibodies. 
 Here, we present a method to address these bottlenecks by dissecting heart first and then conducting the staining process on the surface of a microscope slide. To prevent the loss of small heart samples and to facilitate solution handling, we restricted the heart samples within a circle on the surface of the microscope slides drawn by an immEdge pen. After the staining, the fluorescence signals can be directly observed by a compound microscope. 
Our new method significantly improves the penetration for antibodies, since a heart from an embryonic fish only consists of few cell layers. High quality images from intact hearts can be obtained within a much reduced procession time for zebrafish embryos aged from day 2 to day 6. Our method can be potentially extended to stain other organs dissected from either zebrafish or other small animals.

A rapid way to conduct immunostaining of zebrafish embryonic heart is described. Compared to the whole mount immunostaining approach, this method dramatically increases the penetration of the antibodies, which allows obtaining high resolution images that reveal cellular/subcellular structures in the heart within a much reduced processing time.

Zebrafish embryo becomes a popular in vivo vertebrate model for studying cardiac development and human heart diseases due to its advantageous embryology ...

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal neurons. In contrast, the adult zebrafish retina possesses the robust ability to spontaneously regenerate any neuronal class that is lost in a variety of different retinal damage models, including retinal puncture, chemical ablation, concentrated high temperature, and intense light treatment 1-8. Our lab extensively characterized regeneration of photoreceptors following constant intense light treatment and inner retinal neurons after intravitreal ouabain injection 2, 5, 9. In all cases, resident M&uuml;ller glia re-enter the cell cycle to produce neuronal progenitors, which continue to proliferate and migrate to the proper retinal layer, where they differentiate into the deficient neurons. 
	We characterized five different stages during regeneration of the light-damaged retina that were highlighted by specific cellular responses. We identified several differentially expressed genes at each stage of retinal regeneration by mRNA microarray analysis 10. Many of these genes are also critical for ocular development. To test the role of each candidate gene/protein during retinal regeneration, we needed to develop a method to conditionally limit the expression of a candidate protein only at times during regeneration of the adult retina. 
	Morpholino oligos are widely used to study loss of function of specific proteins during the development of zebrafish, Xenopus, chick, mouse, and tumors in human xenografts 11-14. These modified oligos basepair with complementary RNA sequence to either block the splicing or translation of the target RNA. Morpholinos are stable in the cell and can eliminate or "knockdown" protein expression for three to five days 12. 
	Here, we describe a method to efficiently knockdown target protein expression in the adult zebrafish retina. This method employs lissamine-tagged antisense morpholinos that are injected into the vitreous of the adult zebrafish eye. Using electrode forceps, the morpholino is then electroporated into all the cell types of the dorsal and central retina. Lissamine provides the charge on the morpholino for electroporation and can be visualized to assess the presence of the morpholino in the retinal cells. 
	Conditional knockdown in the retina can be used to examine the role of specific proteins at different times during regeneration. Additionally, this approach can be used to study the role of specific proteins in the undamaged retina, in such processes as visual transduction and visual processing in second order neurons.

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

A method to conditionally knockdown a target protein&#x2019;s expression in the adult zebrafish retina is described, which involves intravitreally injecting antisense morpholinos and electroporating them into the retina. The resulting protein is knocked down for several days, which allows testing the protein&#x2019;s role in the regenerating or intact retina.

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal ...

A System for ex vivo Culturing of Embryonic Pancreas

The pancreas controls vital functions of our body, including the production of digestive enzymes and regulation of blood sugar levels1. Although in the past decade many studies have contributed to a solid foundation for understanding pancreatic organogenesis, important gaps persist in our knowledge of early pancreas formation2. A complete understanding of these early events will provide insight into the development of this organ, but also into incurable diseases that target the pancreas, such as diabetes or pancreatic cancer. Finally, this information will generate a blueprint for developing cell-replacement therapies in the context of diabetes.
 During embryogenesis, the pancreas originates from distinct embryonic outgrowths of the dorsal and ventral foregut endoderm at embryonic day (E) 9.5 in the mouse embryo3,4. Both outgrowths evaginate into the surrounding mesenchyme as solid epithelial buds, which undergo proliferation, branching and differentiation to generate a fully mature organ2,5,6. Recent evidences have suggested that growth and differentiation of pancreatic cell lineages, including the insulin-producing &beta;-cells, depends on proper tissue-architecture, epithelial remodeling and cell positioning within the branching pancreatic epithelium7,8. However, how branching morphogenesis occurs and is coordinated with proliferation and differentiation in the pancreas is largely unknown. This is in part due to the fact that current knowledge about these developmental processes has relied almost exclusively on analysis of fixed specimens, while morphogenetic events are highly dynamic.
 Here, we report a method for dissecting and culturing mouse embryonic pancreatic buds ex vivo on glass bottom dishes, which allow direct visualization of the developing pancreas (Figure 1). This culture system is ideally devised for confocal laser scanning microscopy and, in particular, live-cell imaging. Pancreatic explants can be prepared not only from wild-type mouse embryos, but also from genetically engineered mouse strains (e.g. transgenic or knockout), allowing real-time studies of mutant phenotypes. Moreover, this ex vivo culture system is valuable to study the effects of chemical compounds on pancreatic development, enabling to obtain quantitative data about proliferation and growth, elongation, branching, tubulogenesis and differentiation. In conclusion, the development of an ex vivo pancreatic explant culture method combined with high-resolution imaging provides a strong platform for observing morphogenetic and differentiation events as they occur within the developing mouse embryo.

A System for ex vivo Culturing of Embryonic Pancreas

Here, we describe a method for isolation, culture and manipulation of mouse embryonic pancreas. This represents an excellent ex vivo system for studying various aspects of pancreatic development, including morphogenesis, differentiation and growth. Pancreatic bud explants can be cultured for several days and used in a range of different applications, including whole-mount immunofluorescence and live imaging.

The pancreas controls vital functions of our body,  including the production of digestive enzymes and regulation of blood sugar  levels1. Although in the ...

Laser-inflicted Injury of Zebrafish Embryonic Skeletal Muscle

Various experimental approaches have been used in mouse to induce muscle injury with the aim to study muscle regeneration, including myotoxin injections (bupivacaine, cardiotoxin or notexin), muscle transplantations (denervation-devascularization induced regeneration), intensive exercise, but also murine muscular dystrophy models such as the mdx mouse (for a review of these approaches see 1). In zebrafish, genetic approaches include mutants that exhibit muscular dystrophy phenotypes (such as runzel2 or sapje3) and antisense oligonucleotide morpholinos that block the expression of dystrophy-associated genes4. Besides, chemical approaches are also possible, e.g. with Galanthamine, a chemical compound inhibiting acetylcholinesterase, thereby resulting in hypercontraction, which eventually leads to muscular dystrophy5. However, genetic and pharmacological approaches generally affect all muscles within an individual, whereas the extent of physically inflicted injuries are more easily controlled spatially and temporally1. Localized physical injury allows the assessment of contralateral muscle as an internal control. Indeed, we recently used laser-mediated cell ablation to study skeletal muscle regeneration in the zebrafish embryo6, while another group recently reported the use of a two-photon laser (822 nm) to damage very locally the plasma membrane of individual embryonic zebrafish muscle cells7.
Here, we report a method for using the micropoint laser (Andor Technology) for skeletal muscle cell injury in the zebrafish embryo. The micropoint laser is a high energy laser which is suitable for targeted cell ablation at a wavelength of 435 nm. The laser is connected to a microscope (in our setup, an optical microscope from Zeiss) in such a way that the microscope can be used at the same time for focusing the laser light onto the sample and for visualizing the effects of the wounding (brightfield or fluorescence). The parameters for controlling laser pulses include wavelength, intensity, and number of pulses.
Due to its transparency and external embryonic development, the zebrafish embryo is highly amenable for both laser-induced injury and for studying the subsequent recovery. Between 1 and 2 days post-fertilization, somitic skeletal muscle cells progressively undergo maturation from anterior to posterior due to the progression of somitogenesis from the trunk to the tail8, 9. At these stages, embryos spontaneously twitch and initiate swimming. The zebrafish has recently been recognized as an important vertebrate model organism for the study of tissue regeneration, as many types of tissues (cardiac, neuronal, vascular etc.) can be regenerated after injury in the adult zebrafish10, 11.

The method presented here comprises the precise injury of live zebrafish embryos with high-energy laser pulses and the subsequent analysis of these injuries and their recovery with time. We also show how genetically labeled single or groups of skeletal muscle cells can be tracked during and after laser light induced damage.

Various experimental approaches have been used in mouse to induce muscle injury with the aim to study muscle regeneration, including myotoxin injections ...