The authors would like to acknowledge the Department of Biology at Syracuse University and Dr. Marilyn Kerr for their roles in the administration of the Comparative Vertebrate Biology course. The Albertson lab is supported by grant R21DE019223 from the National Institutes of Health/National Institute of Dental and Craniofacial Research, as well as grant R01AG031922 from the National Institutes of Health/National Institute on Aging.

1. Plasmid Transformation of Target cDNA
Part I: Transformation of Plasmid
(Time Required: 3 hours plus overnight incubation)
<ol>
<li>Warm SOC nutrient broth in a 42&deg;C water bath (500 &mu;L per reaction)</li>
<li>Thaw competent cells on ice</li>
<li>Add 1 &mu;L plasmid to 25 &mu;L competent cells</li>
<li>Place on ice for 20 minutes</li>
<li>Heat shock cells in 42&deg;C water bath for 45 seconds</li>
<li>Immediately place cells on ice for 2 minutes</li>
<li>Add 500 &mu;L of 42&deg;C nutrient broth to each vial of cells</li>
<li>Shake at 37&deg;C for 2 hours at 255 rotations per minute (rpm)</li>
<li>Plate 75 &mu;L transformation mix on Luria Bertani (LB) agar plates</li>
<li>Incubate plates inverted at 37&deg;C overnight</li>
</ol>
Part II: E. coli Culture
(Time Required: 15 minutes plus overnight incubation)
<ol>
<li>For each reaction, aliquot 3 mL LB broth and 3 &mu;L of 50 mg/ mL ampicillin into a culture tube</li>
<li>Scrape 1 bacteria colony from the agar plate and add to each culture tube</li>
<li>Shake culture tubes at 37&deg;C at 255 rpm overnight</li>
</ol>
Part III: Plasmid Prep
(Time Required: 1.5 hours)
<ol>
<li>Isolate plasmid from overnight cultures using the 5 Prime FastPlasmid Mini Kit (Catalog #2300000)</li>
<li>To check for the presence of prepared plasmid, run the DNA eluted from the kit on a 1% Tris-Acetate-EDTA (TAE) gel stained with ethidium bromide</li>
</ol>
2. In Situ DIG-labeled RNA Probe Synthesis
Part I: Linearization of Plasmid
(Time Required: 2.5 hours)
<ol>
<li>In a 1.5 mL microfuge tube, combine: 
<table border="1">
<tbody>
 <tr>
 <td>Prepared Plasmid</td>
 <td>20 mL </td>
 </tr>
 <tr>
 <td>Restriction Enzyme*</td>
 <td>2 mL </td>
 </tr>
 <tr>
 <td>Buffer**</td>
 <td>10 mL </td>
 </tr>
 <tr>
 <td>10X Bovine Serum Albumin (BSA)</td>
 <td>10 mL </td>
 </tr>
 <tr>
 <td>Diethyl Pyrocarbonate (DEPC) Water</td>
 <td>58 mL </td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>100 mL </td>
 </tr>
 </tbody>
 </table>
</li>
<li>Incubate at 37&deg;C for 2 hours 
*Varies with plasmid 
**Varies with restriction enzyme</li>
</ol>
Part II: Transcription
(Time Required: 3 hours)
<ol>
<li>In a 1.5 mL microfuge tube, combine: 
<table border="1">
 <tbody>
 <tr>
 <td>Linear Plasmid</td>
 <td>4 mL </td>
 </tr>
 <tr>
 <td>10X Transcription Buffer**</td>
 <td>4 mL </td>
 </tr>
 <tr>
 <td>Digoxigenin Label Mix</td>
 <td>4 mL </td>
 </tr>
 <tr>
 <td>Polymerase*</td>
 <td>2 mL </td>
 </tr>
 <tr>
 <td>RNase Inhibitor</td>
 <td>2 mL</td>
</tr>
<tr>
 <td>DEPC Water</td> 
 <td>24 mL</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>40 mL </td>
 </tr>
 </tbody>
 </table>
</li>
<li>Incubate at 37&deg;C for 1 hour</li>
<li>Add 2 &mu;L of polymerase*</li>
<li>Incubate at 37&deg;C for 1 hour</li>
<li>Add 2 &mu;L of DNase</li>
<li>Incubate at 37&deg;C for 20 minutes 
*Varies with plasmid 
**Varies with polymerase</li>
</ol>
Part III: Precipitation
(Time Required: 5 minutes plus 2 hour to overnight incubation, 1 hour to centrifuge and re-suspend)
<ol>
<li>Add 4 &mu;L of 0.2M EDTA</li>
<li>Add 5 &mu;L of 4M lithium chloride</li>
<li>Add 150 &mu;L of ice cold 100% ethanol</li>
<li>Incubate at -80&deg;C for 2 hours to overnight</li>
<li>Centrifuge at 14,000 rpm for 30 minutes at 4&deg;C, decant away supernatant</li>
<li>Dry pellet for 7 minutes</li>
<li>Re-suspend in 20 &mu;L DEPC water</li>
<li>Incubate at 37&deg;C for 5 minutes</li>
</ol>
Part IV: Fractionation - Perform only if probe size is greater than 0.6 kb
(Time Required: Varies based on probe size, usually no more than 20 minutes)
<ol>
<li>In a 1.5 mL microfuge tube, combine: 
<table border="1">
 <tbody>
 <tr>
 <td>RNA Probe</td>
 <td>20 mL </td>
 </tr>
 <tr>
 <td>DEPC Water</td>
 <td>12 mL </td>
 </tr>
 <tr>
 <td>Sodium Bicarbonate</td>
 <td>4 mL </td>
 </tr>
 <tr>
 <td>Sodium Carbonate</td>
 <td>4 mL </td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>40 mL </td>
 </tr>
</tbody>
</table>
</li>
<li>Incubate in a 60&deg;C water bath. Base the incubation time on the equation: 
Time (min.) = (starting kb - desired kb) / (.11 x starting kb x desired kb) 
Average desired size = 0.6 kb</li>
</ol>
Part V: Final Precipitation
(Time Required: 5 minutes plus 2 hour to overnight incubation, 1 hour to centrifuge and re-suspend, 1 hour for gel electrophoresis)
<ol>
<li>Add 40 &mu;L DEPC water</li>
<li>Add 8 &mu;L Sodium Acetate</li>
<li>Add 1.04 &mu;L Glacial Acetic Acid</li>
<li>Add 240 &mu;L ice cold 100% ethanol</li>
<li>Incubate at -80&deg;C for 2 hours to overnight</li>
<li>Centrifuge at 14,000 rpm for 30 minutes at 4&deg;C, decant away supernatant</li>
<li>Dry pellet for 7 minutes</li>
<li>Re-suspend in 20 &mu;L DEPC water</li>
<li>Vortex to mix</li>
<li>To check for the presence of riboprobe, run the re-suspended solution on a 1% TAE gel stained with ethidium bromide</li>
</ol>
3. Whole Mount In Situ Hybridization
Part I: Fixation of Embryos and Proteinase K Digestion
(Time Required: Fix overnight plus 4 hours the next day for storage, 2.5 hours from storage to Part II)
<ol>
<li>Collect staged zebrafish embryos and fix in 4% paraformaldehyde (PFA) overnight at 4&deg;C</li>
<li>Wash fixed embryos in phosphate buffered saline containing 0.1% Tween-20 (PBSt) 3 times for 10 minutes each</li>
<li>To ensure exposure of the embryos to the experimental reagents, manually dechorionate embryos (if necessary) using fine-tipped forceps </li>
<li>Dehydrate embryos by washing in a graded series of 25% and 50% methanol in PBSt for one hour each wash, then store in 100% methanol at -20&deg;C</li>
<li>When ready to use, rehydrate embryos in PBSt by washing in 50% (2 times) and 25% (1 time) methanol in PBSt for 10 minutes each. Finally wash 2 times for 10 minutes each in 100% PBSt. Exact timing is not important for PBSt/methanol washes</li>
<li>Bleach embryos in a 10% hydrogen peroxide solution in PBSt, if necessary, to remove dark pigments. Embryos should incubate in hydrogen peroxide solution for 10-20 minutes, depending on the age of the embryo, and the cap of the microfuge tube should remain open to prevent build-up of air pressure</li>
<li>Digest embryos with 50 mg/ mL Proteinase K diluted 1:5000 in PBSt for 3-15 minutes, depending on the age of the embryo</li>
<li>Re-fix embryos in 4% PFA for 30 minutes, and then wash 3 times in PBSt for 5 minutes each</li>
</ol>
Part II: Hybridization of Riboprobes
(Time Required: 3 hours plus overnight for hybridization, 1.5 hours the next day to Part III)
<ol start="9">
<li>Prehybridize embryos with prehybridization solution (PHS) in a preheated 70&deg;C water bath for 2-3 hours</li>
<li>Remove PHS, add 0.5 mL hybridization solution and 1.5 &mu;L previously synthesized Digoxigenin labeled riboprobes, incubate at 70&deg;C overnight. Temperature can fluctuate within a few degrees depending on the riboprobe target</li>
<li>The next day, wash embryos at 70&deg;C in graded solutions of 75%, 50%, and 25% PHS in 2X saline-sodium citrate (SSC) for 10 minutes each, then wash in 0.2X SSC for 30 minutes at 68&deg;C</li>
<li>Wash in Maleic Acid Buffer (MAB) 2 times for 10 minutes each at room temperature</li>
</ol>
Part III: Anti-Digoxigenin (&alpha;-DIG) Antibody Incubation
(Time Required: 3 hours plus overnight to block, 2.5 hours the next day to Part IV)
<ol start="13">
<li>Transfer embryos to a 12 well plate</li>
<li>Pre-block embryos in 1-2 mL blocking solution for at least 3 hours at room temperature</li>
<li>Simultaneously, pre-block the antibody by preparing a second volume of blocking solution and diluting the anti-digoxigenin antibody 1:2000 in this solution</li>
<li>Remove pre-block and add 1-2 mL pre-blocked &alpha;-DIG solution, incubate overnight at 4&deg;C</li>
<li>The next day, wash embryos in MAB. Allow the embryos to incubate in MAB for 5 minutes first, then perform buffer changes and incubate for two 10 minute, one 30 minute and one 60 minute interval. Exact timing is not necessary</li>
<li>Wash embryos 3 times for 5 minutes each in Alkaline Phosphatase (AP) buffer</li>
</ol>
Part IV: Staining and Final Processing
(Time Required: 1 hour to overnight for staining, depending on the riboprobe used, 4 hours to beginning of glycerol washes, 6-10 hours per glycerol wash, may be stored in glycerol)
<ol start="19">
<li>Add 1-2 mL staining solution to the embryos, wrap the plate in foil, and check staining at regular intervals (about every 20 minutes) until staining is sufficient</li>
<li>Wash embryos with PBSt 2 times for 5 minutes each to stop the staining reaction</li>
<li>Dehydrate embryos using 10 minute washes in 25% (1 time) and 50% (2 times) methanol in PBSt, then in 100% methanol, to remove background staining</li>
<li>Allow embryos to incubate in 100% methanol for at least 2 hours at room temperature</li>
<li>Rehydrate in PBSt using 10 minute washes in 50% (2 times) and 25% (1 time) methanol in PBSt, then wash in 100% PBSt 2 times</li>
<li>Transfer stained embryos to an 80% glycerol solution in PBSt, using a graded series of washes, and store at 4&deg;C.</li>
</ol>
Recipes:
<ul>
<li>LB agar plates- 10 g LB agar + 250 mL distilled water, autoclave, when cool to the touch add 
250 &mu;L ampicillin, pour 15-20 mL warm solution into each petri dish, allow agar to solidify</li>
<li>LB broth- 12.5 g LB Broth + 200 mL distilled water, autoclave, allow to cool before use</li>
<li>1% TAE gel- 0.4 g agarose, 40 mL 1X TAE, 2 &mu;L ethidium bromide; load 7 &mu;L plasmid (Plasmid Transformation of Target cDNA) or 3 &mu;L riboprobe and 4 &mu;L DEPC water (In situ DIG-labeled RNA Probe Synthesis) + 1 &mu;L loading dye, 5 &mu;L DNA ladder</li>
<li>Prehybridization solution- 50% formamide, 5X SSC, 9.2mM citric acid, 1% Tween-20</li>
<li>Hybridization solution- prehybridization solution plus 500 &mu;g/ mL tRNA and 50 &mu;g/ mL Heparin</li>
<li>MAB- 100mM Maleic Acid, 150mM NaCl, 0.2M NaOH, 0.1% Tween-20, pH to 7.5</li>
<li>Blocking Solution- 3 parts MAB, 1 part 10% Boehringer blocking reagent in MAB, 1 part heat deactivated lamb serum</li>
<li>AP buffer- 60mM Tris-HCl pH to 9.5, 60mM NaCl, 30mM MgCl2, 0.1% Tween-20</li>
<li>Staining Solution- 5-Bromo-4-chloro-3-indolyl phosphate (BCIP) and Nitro blue tetrazolium (NBT) in AP buffer</li>
</ul>
4. REPRESENTATIVE RESULTS
When performed correctly, the reaction between the NBT, BCIP, and alkaline phosphatase will form a purple precipitate that should appear on the zebrafish embryo as a purple stain. Riboprobes should be previously synthesized from cDNA corresponding to the gene of interest. Therefore, it can be concluded any stain visualized represents areas of the zebrafish in which the gene of interest has been transcribed at that particular developmental stage. For the purposes of this course, riboprobes were synthesized from aldh1a2 (previously raldh2; Begemann et al., 2001; Figure 1); fgf8a (Reifers et al., 1998; Figure 2); deltaC (Oates et al., 2005); myod1 (Weinberg et al., 1996); shha (Krauss et al., 1993), pax2a (Brand et al., 1996; Figure 3) and myl7 (previously cmlc2; Yelon et al., 1999) cDNA. Staining was expected in the midline and in anatomical structures including the somites, tailbud, myotome, brain, and heart. Ace/fgf8a mutants were expected to have defects in many of these structures. Staining was easily visualized using a standard dissecting microscope. Additional sources contain more information and troubleshooting on WISH techniques similar to those described here (Clark, 2003; D'Costa et al., 2009; Schmoldt et al., 2009; Schoenwolf, 2009).
<img src="/files/ftp_upload/2533/2533fig1.jpg" alt="Figure 1" /> 
Figure 1. A zebrafish embryo 24 hours post fertilization, which has been hybridized with riboprobes specific for aldh1a2. Specific staining can be seen in the eyes, hindbrain, pectoral fin bud primordia, and somites. Anterior is to the top, posterior is to the bottom.
<img src="/files/ftp_upload/2533/2533fig2.jpg" alt="Figure 2" /> 
Figure 2. A zebrafish embryo at the 13 somite stage of development that has been hybridized with a probe specific for fgf8a. Specific staining is seen in the telencephalon, dorsal diencephalon, midbrain-hindbrain boundary, somites, and tailbud. Ventral is to the left, dorsal is to the right.
<img src="/files/ftp_upload/2533/2533fig3.jpg" alt="Figure 3" /> 
Figure 3. A 22 hours post fertilization zebrafish embryo that has been hybridized with a riboprobe specific for pax2a, a robust marker useful for visualizing the nervous system. Specific staining can be seen in the choroid fissure, midbrain-hindbrain boundary, otic vesicle, and spinal cord neurons. Dorsal view with anterior to the left.

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No conflicts of interest declared.

WISH was used in a Comparative Vertebrate Biology laboratory course to help students understand the roles of genetics in anatomical development through the visualization of known gene expression patterns. For the first part of the course, students performed dissections on organisms representing several different chordate taxa, allowing them ample time to study, understand, compare, and contrast vertebrate anatomy.
As an introduction to the second part of the course, students were given a formal lecture describing zebrafish development and anatomy. The methods and expected results of the WISH experiment were also discussed. Students were then given live zebrafish at somitogenesis and prim-6 stages of development, and at 2 and 5 days post fertilization (dpf) to examine under the dissecting microscope. This was to give students a better understanding of what zebrafish embryos look like and the types of morphological changes that occur over ontogeny.
 In the next laboratory session, students were given zebrafish embryos in which WISH had been previously performed. They were asked to study and describe the gene expression patterns for each gene of interest (riboprobe used). Embryos used for WISH were derived from matings between members of a heterozygous ace/fgf8a line. Based on Mendelian inheritance, 25% of the embryos from the ace/fgf8a matings were expected to be homozygous mutants and to exhibit defects in many of the anatomical structures focused on in this course. Based on published reports and unpublished observations in the Albertson lab, defects in the brain and improper heart looping were expected, as well as defects in the somites (Brand et al., 1996; Albertson and Yelick, 2005; personal observations).
Students were asked to examine all specimens, wild type (heterozygous animals are indistinguishable from wild type siblings at early stages of development) and homozygous mutants, for each gene expression pattern presented. They were then asked to write lab reports describing their results, and based on their knowledge of anatomy and genetics, how defective gene expression may have precipitated anatomical malformations.
Students seemed to receive this laboratory exercise with excitement and curiosity. Most had never used WISH before and were extremely interested in this part of the course. Students found the different patterns of gene expression in the zebrafish embryos intriguing, some even described the staining patterns visualized by associating them with well-known designs and symbols, such as a smiley face. The resulting lab reports showed the students had a general understanding of the WISH protocol and the expression of genes in specific anatomical structures. Students were also required to understand the specific functions of the genes studied using WISH during the lab (Stickney et al., 2000; Huelsken et al., 2002; Geetha-Loganathan et al., 2008a; Geetha-Loganathan et al., 2008b). It was evident however, that certain students had limited background knowledge of the signaling pathways and genes of interest. More information about these concepts in the WISH introductory lecture may be a welcome addition to the use of WISH in future Comparative Vertebrate Biology courses.
Since the protocol generally takes four consecutive days, depending on the schedule of the course, students may only be able to complete a part of the experiment in class and the instructor must be responsible for the remainder. In our Comparative Vertebrate Biology class, students completed the staining reactions in the laboratory, while the Teaching Assistant performed all preceding steps. If it is preferred to have the students perform WISH in class, the protocol may be divided into subunits that can be performed over several laboratory sessions, depending on the time frame of the class. If it is not feasible for students to perform the entire protocol, as it was here due to the lab meeting only once a week, students can add the staining solution at the beginning of class and, depending on the riboprobe used, have the staining complete within an hour. The time it takes for the stain to develop varies greatly with each riboprobe and a variety of experimental conditions, and should be predetermined before class. Notably, if students will only be developing the stain in lab, instructors will be responsible for all preceding steps, which will require significant time and effort outside of the classroom. If desired, a shorter alternative to WISH could be immunohistochemistry, using antibody labels to visualize protein localization, however at this time, zebrafish-specific antibodies for developmental genes are not readily available. Another option would be to perform WISH on different vertebrate species and have the students compare the expression patterns of the same genes in different organisms (Pizard et al., 2004; Aramaki et al., 2007; Emerging Model Organisms, 2008; Emerging Model Organisms, 2010).
The overarching goal of using WISH in a Comparative Vertebrate Biology course was to demonstrate to students how molecular biological techniques are used to study anatomical development. It also provided an opportunity for students to speculate as to how altered gene expression may lead not only to developmental malformations but also to evolutionary change. Formalized as evolutionary developmental biology (often referred to as "evo-devo"), this rapidly growing field of study aims to link genotype and phenotype through development, and to elucidate potential mechanistic bases of evolutionary change. With the rise of this field, more ecologists, organismal biologists, and physiologists are employing molecular techniques in their research. We contend that the use of WISH in a Comparative Vertebrate Biology course will help to keep the curriculum up to date with current technological and conceptual advances in research, and to facilitate a better horizontal alignment of upper level biology courses by combining biological subfields. Moreover, this integrative approach will provide students the opportunity to learn an assortment of biological research techniques in one course, leading to a more diversified education and the promotion of future interdisciplinary scientific research.

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using whole mount in situ hybridization to link molecular and organismal biology

Whole mount in situ hybridization (WISH) is a common technique in molecular biology laboratories used to study gene expression through the localization of specific mRNA transcripts within whole mount specimen. This technique (adapted from Albertson and Yelick, 2005) was used in an upper level undergraduate Comparative Vertebrate Biology laboratory classroom at Syracuse University. The first two thirds of the Comparative Vertebrate Biology lab course gave students the opportunity to study the embryology and gross anatomy of several organisms representing various chordate taxa primarily via traditional dissections and the use of models. The final portion of the course involved an innovative approach to teaching anatomy through observation of vertebrate development employing molecular techniques in which WISH was performed on zebrafish embryos. A heterozygous fibroblast growth factor 8 a (fgf8a) mutant line, ace, was used. Due to Mendelian inheritance, ace intercrosses produced wild type, heterozygous, and homozygous ace/fgf8a mutants in a 1:2:1 ratio. RNA probes with known expression patterns in the midline and in developing anatomical structures such as the heart, somites, tailbud, myotome, and brain were used. WISH was performed using zebrafish at the 13 somite and prim-6 stages, with students performing the staining reaction in class. The study of zebrafish embryos at different stages of development gave students the ability to observe how these anatomical structures changed over ontogeny. In addition, some ace/fgf8a mutants displayed improper heart looping, and defects in somite and brain development. The students in this lab observed the normal development of various organ systems using both external anatomy as well as gene expression patterns. They also identified and described embryos displaying improper anatomical development and gene expression (i.e., putative mutants).
For instructors at institutions that do not already own the necessary equipment or where funds for lab and curricular innovation are limited, the financial cost of the reagents and apparatus may be a factor to consider, as will the time and effort required on the part of the instructor regardless of the setting. Nevertheless, we contend that the use of WISH in this type of classroom laboratory setting can provide an important link between developmental genetics and anatomy. As technology advances and the ability to study organismal development at the molecular level becomes easier, cheaper, and increasingly popular, many evolutionary biologists, ecologists, and physiologists are turning to research strategies in the field of molecular biology. Using WISH in a Comparative Vertebrate Biology laboratory classroom is one example of how molecules and anatomy can converge within a single course. This gives upper level college students the opportunity to practice modern biological research techniques, leading to a more diversified education and the promotion of future interdisciplinary scientific research.

Watch this Scientific Journal Video about Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology at JoVE.com

Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology

Whole mount in situ hybridization (WISH) was used in an upper level undergraduate Comparative Vertebrate Biology course in addition to vertebrate dissections. This gave students the opportunity to study gene expression patterns as well as gross anatomy, linking the study of molecular and organismal biology within one course.

Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology

Whole mount in situ hybridization (WISH) is a common technique in molecular biology laboratories used to study gene expression through the localization of ...

using-whole-mount-situ-hybridization-to-link-molecular-organismal

Research

JoVE Journal

Biology

21.5K Views.  Syracuse University. Whole mount in situ hybridization (WISH) was used in an upper level undergraduate Comparative Vertebrate Biology course in addition to vertebrate dissections. This gave students the opportunity to study gene expression patterns as well as gross anatomy, linking the study of molecular and organismal biology within one course.

Video: Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology

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.

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

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.

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology. Here, we present a high-resolution double fluorescent in situ hybridization protocol for analyzing the precise expression pattern of a single gene and for determining the overlap of the expression domains of two genes. The protocol is a modified version of the standard in situ hybridization using alkaline phosphatase and substrates such as NBT/BCIP and Fast Red 1,2. This protocol utilizes standard digoxygenin and fluorescein labeled probes along with tyramide signal amplification (TSA) 3. The commercially available TSA kits allow flexible experimental design as fluorescence emission from green to far-red can be used in combination with various nuclear stains, such as propidium iodide, or fluorescence immunohistochemistry for proteins. TSA produces a reactive fluorescent substrate that quickly covalently binds to moieties, typically tyrosine residues, in the immediate vicinity of the labeled antisense riboprobe. The resulting staining patterns are high resolution in that subcellular localization of the mRNA can be observed using laser scanning confocal microscopy 3,4. One can observe nascent transcripts at the chromosomal loci, distinguish nuclear and cytoplasmic staining and visualize other patterns such as cortical localization of mRNA. Studies in Drosophila indicate that roughly 70% of mRNAs exhibit specific patterns of subcellular localization that frequently correlate with the function of the encoded protein 5. When combined with computer-aided reconstruction of 3D confocal datasets, our protocol allows the detailed analysis of mRNA distribution with sub-cellular resolution in whole vertebrate embryos.

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology. Here, we present a high-resolution double fluorescent in situ hybridization protocol for analyzing the precise expression pattern of a single gene and for determining the overlap of the expression domains of two genes. We include a propidium iodide nuclear counter-stain to highlight tissue organization.

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology.  Here, we present a high-resolution double ...

Whole Mount in Situ Hybridization of E8.5 to E11.5 Mouse Embryos

Whole mount in situ hybridization is a very informative approach for defining gene expression patterns in embryos. The in situ hybridization procedures are lengthy and technically demanding with multiple important steps that collectively contribute to the quality of the final result. This protocol describes in detail several key quality control steps for optimizing probe labeling and performance. Overall, our protocol provides a detailed description of the critical steps necessary to reproducibly obtain high quality results. First, we describe the generation of digoxygenin (DIG) labeled RNA probes via in vitro transcription of DNA templates generated by PCR. We describe three critical quality control assays to determine the amount, integrity and specific activity of the DIG-labeled probes. These steps are important for generating a probe of sufficient sensitivity to detect endogenous mRNAs in a whole mouse embryo. In addition, we describe methods for the fixation and storage of E8.5-E11.5 day old mouse embryos for in situ hybridization. Then, we describe detailed methods for limited proteinase K digestion of the rehydrated embryos followed by the details of the hybridization conditions, post-hybridization washes and RNase treatment to remove non-specific probe hybridization. An AP-conjugated antibody is used to visualize the labeled probe and reveal the expression pattern of the endogenous transcript. Representative results are shown from successful experiments and typical suboptimal experiments.

Whole Mount in Situ Hybridization of E8.5 to E11.5 Mouse Embryos

This whole mount in situ hybridization protocol discusses critical steps that ensure reproducible high quality results for gene expression studies in E8.5-E11.5 day old mouse embryos.

Whole mount in situ hybridization is a very informative approach for defining gene expression patterns in embryos. The in situ hybridization procedures ...

High-throughput Physical Mapping of Chromosomes using Automated in situ Hybridization

Projects to obtain whole-genome sequences for 10,000 vertebrate species1 and for 5,000 insect and related arthropod species2 are expected to take place over the next 5 years. For example, the sequencing of the genomes for 15 malaria mosquitospecies is currently being done using an Illumina platform3,4. This Anopheles species cluster includes both vectors and non-vectors of malaria. When the genome assemblies become available, researchers will have the unique opportunity to perform comparative analysis for inferring evolutionary changes relevant to vector ability. However, it has proven difficult to use next-generation sequencing reads to generate high-quality de novo genome assemblies5. Moreover, the existing genome assemblies for Anopheles gambiae, although obtained using the Sanger method, are gapped or fragmented4,6.
Success of comparative genomic analyses will be limited if researchers deal with numerous sequencing contigs, rather than with chromosome-based genome assemblies. Fragmented, unmapped sequences create problems for genomic analyses because: (i) unidentified gaps cause incorrect or incomplete annotation of genomic sequences; (ii) unmapped sequences lead to confusion between paralogous genes and genes from different haplotypes; and (iii) the lack of chromosome assignment and orientation of the sequencing contigs does not allow for reconstructing rearrangement phylogeny and studying chromosome evolution. Developing high-resolution physical maps for species with newly sequenced genomes is a timely and cost-effective investment that will facilitate genome annotation, evolutionary analysis, and re-sequencing of individual genomes from natural populations7,8.
Here, we present innovative approaches to chromosome preparation, fluorescent in situ hybridization (FISH), and imaging that facilitate rapid development of physical maps. Using An. gambiae as an example, we demonstrate that the development of physical chromosome maps can potentially improve genome assemblies and, thus, the quality of genomic analyses. First, we use a high-pressure method to prepare polytene chromosome spreads. This method, originally developed for Drosophila9, allows the user to visualize more details on chromosomes than the regular squashing technique10. Second, a fully automated, front-end system for FISH is used for high-throughput physical genome mapping. The automated slide staining system runs multiple assays simultaneously and dramatically reduces hands-on time11. Third, an automatic fluorescent imaging system, which includes a motorized slide stage, automatically scans and photographs labeled chromosomes after FISH12. This system is especially useful for identifying and visualizing multiple chromosomal plates on the same slide. In addition, the scanning process captures a more uniform FISH result. Overall, the automated high-throughput physical mapping protocol is more efficient than a standard manual protocol.

High-throughput Physical Mapping of Chromosomes using Automated in situ Hybridization

Genome assemblies based on massively parallel DNA sequencing technologies are usually highly fragmented. The development of physical chromosome maps can potentially improve genome assemblies. Here, we demonstrate innovative approaches to chromosome preparation, fluorescent in situ hybridization, and imaging that significantly increase throughput of the physical map development.

Projects to obtain whole-genome sequences for 10,000 vertebrate species1 and for 5,000 insect and related arthropod species2 are expected to take place ...

Chromosome Replicating Timing Combined with Fluorescent In situ Hybridization

Mammalian DNA replication initiates at multiple sites along chromosomes at different times during S phase, following a temporal replication program. The specification of replication timing is thought to be a dynamic process regulated by tissue-specific and developmental cues that are responsive to epigenetic modifications. However, the mechanisms regulating where and when DNA replication initiates along chromosomes remains poorly understood. Homologous chromosomes usually replicate synchronously, however there are notable exceptions to this rule. For example, in female mammalian cells one of the two X chromosomes becomes late replicating through a process known as X inactivation1. Along with this delay in replication timing, estimated to be 2-3 hr, the majority of genes become transcriptionally silenced on one X chromosome. In addition, a discrete cis-acting locus, known as the X inactivation center, regulates this X inactivation process, including the induction of delayed replication timing on the entire inactive X chromosome. In addition, certain chromosome rearrangements found in cancer cells and in cells exposed to ionizing radiation display a significant delay in replication timing of &gt;3 hours that affects the entire chromosome2,3. Recent work from our lab indicates that disruption of discrete cis-acting autosomal loci result in an extremely late replicating phenotype that affects the entire chromosome4. Additional 'chromosome engineering' studies indicate that certain chromosome rearrangements affecting many different chromosomes result in this abnormal replication-timing phenotype, suggesting that all mammalian chromosomes contain discrete cis-acting loci that control proper replication timing of individual chromosomes5.
Here, we present a method for the quantitative analysis of chromosome replication timing combined with fluorescent in situ hybridization. This method allows for a direct comparison of replication timing between homologous chromosomes within the same cell, and was adapted from6. In addition, this method allows for the unambiguous identification of chromosomal rearrangements that correlate with changes in replication timing that affect the entire chromosome. This method has advantages over recently developed high throughput micro-array or sequencing protocols that cannot distinguish between homologous alleles present on rearranged and un-rearranged chromosomes. In addition, because the method described here evaluates single cells, it can detect changes in chromosome replication timing on chromosomal rearrangements that are present in only a fraction of the cells in a population.

Chromosome Replicating Timing Combined with Fluorescent In situ Hybridization

A quantitative method for the analysis of chromosome replication timing is described. The method utilizes BrdU incorporation in combination with fluorescent in situ hybridization (FISH) to assess replication timing of mammalian chromosomes. This technique allows for the direct comparison of rearranged and un-rearranged chromosomes within the same cell.

Mammalian DNA replication initiates at multiple sites along chromosomes at different times during S phase, following a temporal replication program. The ...

Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos

Assessing the expression pattern of a gene, as well as the subcellular localization properties of its transcribed RNA, are key features for understanding its biological function during development. RNA in situ hybridization (RNA-ISH) is a powerful method used for visualizing RNA distribution properties, be it at the organismal, cellular or subcellular levels 1. RNA-ISH is based on the hybridization of a labeled nucleic acid probe (e.g. antisense RNA, oligonucleotides) complementary to the sequence of an mRNA or a non-coding RNA target of interest 2. As the procedure requires primary sequence information alone to generate sequence-specific probes, it can be universally applied to a broad range of organisms and tissue specimens 3. Indeed, a number of large-scale ISH studies have been implemented to document gene expression and RNA localization dynamics in various model organisms, which has led to the establishment of important community resources 4-11. While a variety of probe labeling and detection strategies have been developed over the years, the combined usage of fluorescently-labeled detection reagents and enzymatic signal amplification steps offer significant enhancements in the sensitivity and resolution of the procedure 12. Here, we describe an optimized fluorescent in situ hybridization method (FISH) employing tyramide signal amplification (TSA) to visualize RNA expression and localization dynamics in staged Drosophila embryos. The procedure is carried out in 96-well PCR plate format, which greatly facilitates the simultaneous processing of large numbers of samples.

Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos

Here we describe a whole-mount fluorescent in situ hybridization (FISH) protocol for determining the expression and localization properties of RNAs expressed during embryogenesis in the fruit fly, Drosophila melanogaster.

Assessing  the expression pattern of a gene, as well as the subcellular localization  properties of its transcribed RNA, are key features for ...

RNA In situ Hybridization in Whole Mount Embryos and Cell Histology Adapted for Marine Elasmobranchs

Marine elasmobranchs are valued animal models for biomedical and genomic studies as they are the most primitive vertebrates to have adaptive immunity and have unique mechanisms for osmoregulation 1-3. As the most primitive living jawed-vertebrates with paired appendages, elasmobranchs are an evolutionarily important model, especially for studies in evolution and development. Marine elasmobranchs have also been used to study aquatic toxicology and stress physiology in relationship to climate change 4. Thus, development and adaptation of methodologies is needed to facilitate and expand the use of these primitive vertebrates to multiple biological disciplines. Here I present the successful adaptation of RNA whole mount in situ hybridization and histological techniques to study gene expression and cell histology in elasmobranchs.
Monitoring gene expression is a hallmark tool of developmental biologists, and is widely used to investigate developmental processes 5. RNA whole mount in situ hybridization allows for the visualization and localization of specific gene transcripts in tissues of the developing embryo. The expression pattern of a gene's message can provide insight into what developmental processes and cell fate decisions a gene may control. By comparing the expression pattern of a gene at different developmental stages, insight can be gained into how the role of a gene changes during development.
While whole mount in situ's provides a means to localize gene expression to tissue, histological techniques allow for the identification of differentiated cell types and tissues. Histological stains have varied functions. General stains are used to highlight cell morphology, for example hematoxylin and eosin for general staining of nuclei and cytoplasm, respectively. Other stains can highlight specific cell types. For example, the alcian blue stain reported in this paper is a widely used cationic stain to identify mucosaccharides. Staining of the digestive tract with alcian blue can identify the distribution of goblet cells that produce mucosaccharides. Variations in mucosaccharide constituents on short peptides distinguish goblet cells by function within the digestive tract 6. By using RNA whole mount in situ's and histochemical methods concurrently, cell fate decisions can be linked to gene-specific expression.
Although RNA in situ's and histochemistry are widely used by researchers, their adaptation and use in marine elasmobranchs have met limited and varied success. Here I present protocols developed for elasmobranchs and used on a regular basis in my laboratory. Although further modification of the RNA in situ's hybridization method may be needed to adapt to different species, the protocols described here provide a strong starting point for researchers wanting to adapt the use of marine elasmobranchs to their scientific inquiries.

RNA In situ Hybridization in Whole Mount Embryos and Cell Histology Adapted for Marine Elasmobranchs

By combining methods for RNA whole mount in situ hybridization and histology, gene expression can be linked with cell fate decisions in the developing embryo. These methods have been adapted to marine elasmobranchs and facilitate the use of these animals as model organisms for biomedical, toxicology and comparative studies.

Marine  elasmobranchs are valued animal models for biomedical and genomic studies as  they are the most primitive vertebrates to have adaptive immunity ...

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for using FISH to quantify the number of mRNAs in single yeast cells. Cells can be grown in any condition of interest and then fixed and made permeable. Subsequently, multiple single-stranded deoxyoligonucleotides conjugated to fluorescent dyes are used to label and visualize mRNAs. Diffraction-limited fluorescence from single mRNA molecules is quantified using a spot-detection algorithm to identify and count the number of mRNAs per cell. While the more standard quantification methods of northern blots, RT-PCR and gene expression microarrays provide information on average mRNAs in the bulk population, FISH facilitates both the counting and localization of these mRNAs in single cells at single-molecule resolution.

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

This protocol describes an experimental procedure for performing Fluorescence in situ Hybridization (FISH) for counting mRNAs in single cells at single-molecule resolution.

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for ...

High Resolution Whole Mount In Situ Hybridization within Zebrafish Embryos to Study Gene Expression and Function

This article focuses on whole-mount in situ hybridization (WISH) of zebrafish embryos. The WISH technology facilitates the assessment of gene expression both in terms of tissue distribution and developmental stage. Protocols are described for the use of WISH of zebrafish embryos using antisense RNA probes labeled with digoxigenin. Probes are generated by incorporating digoxigenin-linked nucleotides through in vitro transcription of gene templates that have been cloned and linearized. The chorions of embryos harvested at defined developmental stages are removed before incubation with specific probes. Following a washing procedure to remove excess probe, embryos are incubated with anti-digoxigenin antibody conjugated with alkaline phosphatase. By employing a chromogenic substrate for alkaline phosphatase, specific gene expression can be assessed. Depending on the level of gene expression the entire procedure can be completed within 2-3 days.

High Resolution Whole Mount In Situ Hybridization within Zebrafish Embryos to Study Gene Expression and Function

The zebrafish, a small tropical fish, has become a popular model for studying gene function during vertebrate development and disease. The temporal and spatial expression of target genes can be determined by in situ hybridization. Our improved protocol allows for the detection of low abundant transcripts with low non-specific background signal.

This article focuses on whole-mount in situ hybridization (WISH) of zebrafish embryos. The WISH technology facilitates the assessment of gene expression ...