We would like to thank the staff in Biomedical Services in Oxford and Birmingham for excellent zebrafish husbandry. T.D. was funded by a Wellcome Trust Chromosome and Developmental Biology PhD Scholarship (#WT102345/Z/13/Z). R.M. and M.K. were funded by the British Heart Foundation (BHF IBSR Fellowship FS/13/50/30436) and are thankful for their generous support. R.M. acknowledges support from the BHF Centre of Research Excellence (RE/13/1/30181), Oxford.

Procedures involving animal subjects are regulated by the Animals (Scientific Procedures) Act 1986 and have been approved by the Home Office and the local Animal Welfare and Ethical Review Body.
1. Image ISH-stained embryos
<ol>
	<li>Prepare a glycerol solution (50%&#8211;80% in 1x PBS buffer) and mix to homogenize the solution (e.g., leave in a roller for at least 5 min). This solution can be kept for months at room temperature.</li>
	<li>After in situ hybridization14,15,16,17, transfer embryos to the glycerol solution with a 3 mL Pasteur pipette and leave to settle for at least 5 min. If imaging embryos older than 24 hpf (hours post fertilization), they may be bleached as described18.
	<ol>
		<li>Prepare and label enough PCR tubes to transfer ISH-stained embryos after imaging.</li>
	</ol>
	</li>
	<li>Add 100% glycerol to the bottom of the well in a glass depression slide with a 3 mL Pasteur pipette.</li>
	<li>Using a 3 mL Pasteur pipette, transfer a single ISH-stained embryo to the glass slide and orient as required under a stereomicroscope equipped with a digital camera and bottom and top illumination. 
	NOTE: Use gel loading pipette tips to position the embryos for imaging, but other tools (e.g., forceps, dissecting needle) will be equally adequate.</li>
	<li>Using the first embryo, adjust the illumination and exposure time at the desired magnification. Use these conditions for ALL of the embryos in the same experiment (i.e., if imaging 40 embryos from a heterozygous mutant incross, make sure the illumination, exposure time and magnification are the same for all).</li>
	<li>Image as many embryos as required. Label each image with a unique number. After imaging, transfer the embryo to a PCR tube/plate labelled with the same number. 
	NOTE: Images should be saved as TIF files, but other formats are also adequate.
	<ol>
		<li>If required, remove excess glycerol in the PCR tubes/plates. 
		NOTE: At this point, the embryos can be stored in the PCR tubes for several weeks at room temperature.</li>
	</ol>
	</li>
</ol>
2. Extract DNA and genotype the ISH-stained embryos
NOTE: Here, use a reliable and inexpensive method to isolate genomic DNA based on the HoTSHOT method19 with a DNA extraction efficiency of 95%-100%3.
<ol>
	<li>After the imaging is complete, add 40-75 &#181;L of alkaline lysis buffer (e.g., HoTSHOT) to each tube.</li>
	<li>Incubate at 95 &#176;C for approximately 30 min and cool the tubes to 4 &#176;C before adding an equal volume of neutralization buffer. An overnight incubation at 4 &#176;C may improve PCR efficiency. 
	NOTE: At this point, the genomic DNA can be used for genotyping or stored at -20 &#176;C until it is required.</li>
	<li>Genotype samples with an appropriate method (e.g., HRMA, RFLP)3,20,21 as required for the mutation of interest.</li>
	<li>Note the genotype corresponding to each sample (e.g., using a spreadsheet software).</li>
</ol>
3. Quantify the pixel intensity of ISH-stained embryos (image analysis using Fiji software)
<ol>
	<li>To quantify the in situ hybridization (ISH) staining signal intensity, convert all images to 8-bit grayscale as described3. If the images were saved as .TIF files, use a Fiji macro for batch conversion3. Alternatively, convert images in other formats (e.g., .JPG) to .TIF using appropriate software and then convert to 8-bit grayscale using Fiji. For convenience, here is the step-by-step procedure that was published previously3, with some alterations.</li>
	<li>Open images in Fiji and invert the image with Edit &#62; Invert. Then change the image type to 8-bit (Image &#62; Type &#62; 8-bit).</li>
	<li>Using the polygon selection tool, draw the region of interest (ROI) manually on the image around the region containing the signal.</li>
	<li>Press t to open the ROI manager. Use the ROI manager&#39;s Measure command to measure the intensity of the ROI. Copy the mean value from the Results window to a spreadsheet software.</li>
	<li>Move the same ROI, ensuring the same size and shape as the original region, to a region of the zebrafish not containing any staining. Repeat step 3.4 to measure the background.</li>
	<li>To obtain the mean pixel intensity of the ISH signal, subtract the mean intensity value of the background region from that of the stained region for each embryo.</li>
	<li>Assign each intensity value to a genotype (from step 2.3).</li>
</ol>
4. Analyze the results with appropriate statistical tests
<ol>
	<li>Plot all the values on one Q-Q plot to identify any deviations from normal distribution. 
	NOTE: Normal distribution can also be verified with the Kolmogorov-Smirnov test or the Shapiro-Wilk test. However, for large sample sizes there is a high risk of false positives in these tests.</li>
	<li>If there are strong deviations from normal distribution, transform all the values (using ln or sqrt functions) to make sure they are normally distributed before proceeding.</li>
	<li>Analyze the differences between the values (transformed if necessary) assigned to each genotype (wt vs. heterozygote vs. mutant) with 2-tailed ANOVA with 95% confidence levels, accounting for the equality of variances with a Levene&#39;s test and Welch correction. For pairwise comparisons between each pair of genotypes, use Tukey&#39;s (equal variances) or Games-Howell (unequal variances) post-hoc test.
	<ol>
		<li>If the values are not normally distributed despite the transformation, use a non-parametric test (Kruskall-Wallis) to analyze the differences between ranked values and a post-hoc Dunn&#39;s multiple comparisons test with Bonferroni correction for pairwise comparisons.</li>
	</ol>
	</li>
	<li>Plot the untransformed values (from step 3.6) as dot plots for the best representation of the results.</li>
</ol>

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The authors declare they have no competing financial interests.

A few factors should be considered when using this method to quantify gene expression. The imaging conditions must be maintained throughout the experiment (for example illumination, exposure times and embryo positioning) to reduce variability between measurements. A critical point is to avoid overstaining of the samples, as differences in staining between samples may be masked. For example, the decrease in VegfA expression in the absence of Eto2 in Xenopus laevis embryos30 could only be detected by carefully monitoring staining over a 24 hour period. Thus, it is good practice to empirically determine adequate staining levels for each gene that best represent its expression, without reaching saturation. Overstaining will also artificially increase the background pixel intensity in the converted 8-bit grayscale images and skew the quantitation results. In extreme cases, the background level in embryonic tissues might be higher than the ISH signal in the selected ROI and these samples should be excluded from the analysis. A similar phenomenon was observed when we tested the suitability of the unstained yolk for background correction (Figure 4). After inversion and conversion to 8-bit the darker pixels in the yolk region become brighter than the ISH signal in the embryo and render the background corrected values negative. Thus, avoid the use of the yolk for background correction. Measuring the background signal in pigmented areas in the embryo (e.g., the eyes or the dorsal part of the trunk from 26/28 hpf onwards) will equally skew the quantitation results and should also be avoided. There are protocols available for bleaching zebrafish embryos, either before or after ISH18 and bleaching embryos older than 24 hpf before imaging is recommended.
Because this method relies on measuring the pixel intensity in a defined area against a background pixel intensity in an equivalent non-stained area, it is not appropriate for quantitation of ubiquitous or near-ubiquitously expressed genes as is. Instead, it is well suited for measuring expression of genes with a spatially restricted distribution where an area for measuring background pixel intensity can be readily identified. Our additional analysis now suggests that using a smaller area (3-4x smaller) for background correction yields similar outcomes to using an equivalent area to that of the ROI. This extends the applicability of the method to genes expressed in wider spatial domains (and thus requiring larger ROIs for intensity measurements), as long as one can use clearly unstained areas of the embryo for background correction.
Finally, we suggest that the genotyping be performed in parallel or after the image quantitation to minimize experimenter bias. Asking a second experimenter to repeat the quantitation on anonymized samples and compare with the first set of measurements will also help reduce experimenter bias. If the images to be quantified are from a comparison between treatments that does not require genotyping (e.g., wild type vs chemical inhibitor or wild type vs. MO knockdown), the experimenter performing the measurements should be blinded to the identity of the sample.

The introduction of genome editing technologies (ZFN, TALENs and more recently, CRISPR/Cas9) has led to a massive increase in the number of laboratories around the world that make use of these systems to study the function of specific genes in vivo. Zebrafish in particular are amenable to genetic manipulation and many mutants have been generated in the recent past1,2. For developmental biologists, one of the most common methods to assess the phenotypical consequences of gene mutations in embryonic development is in situ hybridization (ISH). In the absence of obvious morphological defects that separate homozygous mutants from their wild type or heterozygous siblings, it is essential to be able to correctly identify different genotypes accurately.
Classical ISH relies on qualitative analyses of signal intensities to derive conclusions about regulatory interactions between the gene of interest and selected marker genes. Although useful, these analyses suffer from technical variation and may be biased by researcher expectations. Thus, a method was developed to quantify gene expression after imaging ISH-stained embryos, without prior knowledge of the corresponding genotype. This was followed by an efficient DNA extraction and genotyping that allowed us to quantitatively correlate genotype with gene expression3. While the genotyping of embryos post ISH has been used before4,5, image-based quantification of ISH patterns has not been widely used apart from a couple of studies6,7. The most popular alternatives rely on visual scoring or counting of ISH-stained cells8,9,10, both prone to poor reproducibility and researcher bias. This method is particularly useful to study changes in genes with expression patterns that are spatially restricted, such as runx1 or gata2b, both expressed in a restricted subset of aortic floor cells called the haemogenic endothelium11,12.
Here, we aim to provide a practical guide to the implementation of the quantification by image analysis using Fiji13, as well as the DNA extraction and genotyping protocol. This is meant to illustrate visually our previously published method3. Our method allows an accurate representation of the variation in gene expression detected by ISH and an unbiased assignment of gene expression levels to specific genotypes.

<table><tbody><tr><td>0.2 mL PCR tubes (8-strips with lids)</td><td>StarLab</td><td>A1402-3700</td><td>96-well plates are equally appropriate for sample handling but beware of cross contamination between samples</td></tr><tr><td>3 mL Pasteur pipettes</td><td>Alpha Laboratories</td><td>LW4114</td><td></td></tr><tr><td>Cavity slides</td><td>Brand</td><td>BR475535-50EA</td><td></td></tr><tr><td>Digital Camera (Qimaging Micropublisher 5.0)</td><td>Qimaging</td><td></td><td></td></tr><tr><td>Eppendorf Microloader tips</td><td>Eppendorf</td><td>10289651</td><td>the tips are used to orient the embryos for imaging in glycerol</td></tr><tr><td>Excel</td><td>Microsoft</td><td></td><td></td></tr><tr><td>F3000 Fiber Optic Cold Light Source</td><td>Photonic</td><td></td><td></td></tr><tr><td>Fiji</td><td></td><td></td><td></td></tr><tr><td>Glycerol</td><td>Sigma</td><td>G5516-1L</td><td></td></tr><tr><td>Graphpad Prism 8.01</td><td>GraphPad Software, Inc.</td><td></td><td>we prefer to use Graphpad, but other statistics software packages are also suitable (e.g. SigmaPlot or SPSS)</td></tr><tr><td>HotSHOT alkaline lysis buffer</td><td></td><td>25 mM NaOH, 0.2 mM disodium EDTA, pH 12</td><td></td></tr><tr><td>HotSHOT neutralization buffer</td><td></td><td>Tris HCl 40 mM, pH 5</td><td></td></tr><tr><td>PBS (10X) pH 7.4</td><td>Thermofisher</td><td>70011044</td><td></td></tr><tr><td>Stereomicroscope with illumination stand (Nikon SMZ800N)</td><td>Nikon</td><td></td><td></td></tr><tr><td>Thermocycler</td><td>Thermofisher</td><td></td><td></td></tr></tbody></table>

genotyping and quantification of in situ hybridization staining in zebrafish

In situ hybridization (ISH) is an important technique that enables researchers to study mRNA distribution in situ and has been a critical technique in developmental biology for decades. Traditionally, most gene expression studies relied on visual evaluation of the ISH signal, a method that is prone to bias, particularly in cases where sample identities are known a priori. We have previously reported on a method to circumvent this bias and provide a more accurate quantification of ISH signals. Here, we present a simple guide to apply this method to quantify the expression levels of genes of interest in ISH-stained embryos and correlate that with their corresponding genotypes. The method is particularly useful to quantify spatially restricted gene expression signals in samples of mixed genotypes and it provides an unbiased and accurate alternative to the traditional visual scoring methods.

Here, we describe the practical application of the pipeline for image quantitation and embryo genotyping as published elsewhere3. The workflow for the method is shown in Figure 1. To illustrate how to use this method, ISH was performed for dnmt3bb.1 in 33 hpf embryos from a runx1W84X/+ 22 incross (Figure 2). 130 embryos were imaged using the same illumination conditions as detailed in the protocol and labeling them with a unique number. After imaging, each embryo was transferred to a PCR tube for genotyping. At this point, the image analysis was performed to attribute a pixel intensity value to each image. The genotype was then assigned to its corresponding image and the pixel intensity values grouped according to their genotype for statistical analysis. A decrease in dnmt3bb.1 expression was detected in runx1W84X/W84X mutants (Figure 2A,B)3, in agreement with previous observations5. Interestingly, runx1W84X/+ heterozygous embryos showed no significant differences in dnmt3bb.1 expression (Figure 2A,B) compared to its wild type siblings, suggesting that one copy of Runx1 is sufficient to maintain dnmt3bb.1 expression at appropriate levels.
Many zebrafish mutants fail to show an embryonic phenotype that can otherwise be detected using other loss of function technologies like morpholino oligonucleotides (MOs). This discrepancy can be attributed to a number of causes including off-target effects, maternal protein compensation, a hypomorphic allele23 or the recently discovered phenomenon of genetic compensation24,25,26,27. In this example, we asked whether runx1 expression was reduced or lost in lmo4auob100 mutants since previously published data using an lmo4a MO suggested that runx1 is decreased in lmo4a morphants28. Here, the analysis revealed no significant differences in runx1 expression between wild type and lmo4auob100 homozygous mutants3 (Figure 3A,B). Further analysis by single embryo qPCR showed that there was a small but significant decrease in runx1 expression in lmo4auob100 mutants (Figure 3C). Thus, it is possible that image quantification might not be able to detect small differences in expression levels. Alternatively, the lack of difference between genotypes that we detected is real and the qPCR experiments are detecting changes in runx1 expression in other tissues like the telenchephalon where both lmo4a and runx1 are expressed. Researchers should always verify their results with an independent method like qPCR, but ideally enriching for the tissue of interest by flow cytometry, for example.
In rare instances where the ISH has high background (Figure 3D), the pixel intensity value of this area is so high that subtraction from the signal value produces a negative number and in such instances those embryos would be excluded from the analysis. In our experience, this occurred in about 0.4% of the runx1-probed embryos3 but may vary between experiments, probes or batches of reagents. Although that might be a limitation of the method, the low frequency of high background is very unlikely to influence the overall results.
To test the effect of selecting different areas for background corrections, we first measured the pixel intensity of the runx1 ISH signal in 28 hpf embryos, using different regions for background corrections (Figure 4). Four different regions were selected: two in the trunk region (R1, and R2), one in the yolk region (unstained, but likely to accumulate background staining) and a smaller area anterior to the ROI (R4, Figure 4B). Measuring pixel intensity in these regions showed a relatively stable difference in intensity between ROI and either background area (Figure 4C). However, R3 always showed very high values (above those in the ROI). After inversion and conversion to 8-bit, the yolk region appears very bright and thus is not suitable for use as a background correction. R2 was closer to the ROI but contained some ISH signal, and using it for correction decreased the mean pixel intensity when compared with either R1 (located further dorsally, away from the ISH signal) or R4. Thus, either R1 or R4 are appropriate areas that can be used for background correction (despite the area of R4 being smaller than that of R1). Next, we wished to compare how using R1 or R4 affected the outcomes when comparing runx1 expression. For this, we incrossed dll4+/- heterozygotes29 and analyzed runx1 expression in randomly selected wild type and dll4-/-embryos (Figure 4E). Although using R1 or R4 for background correction affected individual values, the mean pixel intensities within the same genotype were not significantly different (Figure 4E). Moreover, comparing runx1 expression still yields similar mean intensity values between genotypes using either R1 or R4 areas as background correction (&#181;R1=16.3 and &#181;R4=18.2, respectively). Taken together, we concluded that although the choice of background area is important, the main criteria is that it does not include yolk regions (prone to accumulation of background staining) and that it should not contain any (specific) staining that might skew the pixel intensity values of the background.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59956/59956fig1a.jpg" /> 
Figure 1: Workflow of the parallel image quantitation and genotyping protocol. Embryos collected from an incross of fish heterozygous for a mutant allele are probed for the measured gene with a standard ISH protocol. After imaging, genomic DNA is extracted using the HotSHOT protocol by adding the lysis buffer directly to the embryo in a 0.2 mL PCR tube, followed by a 30 min incubation at 95 &#176;C. This DNA is used for genotyping of the embryos by PCR, PCR and restriction fragment length polymorphism (RFLP), KASP assays or any other appropriate method. In parallel, the images for each embryo are inverted and converted to 8-bit greyscale. ROIs of identical shape and size containing the ISH signal (yellow) and background (blue) are manually selected and measured. The measurements, assigned to corresponding genotypes, are statistically analyzed. Figure adapted from Dobrzycki et al.3 <a href="https://www.jove.com/files/ftp_upload/59956/59956fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59956/59956fig02.jpg" /> 
Figure 2: Image quantitation in runx1 mutants reveals reduced levels of dnmt3bb.1 expression by ISH. (A) Example images of ISH in 33 hpf wild type (blue), runx1+/W84X (green) and runx1W84X/W84X (orange) embryos, showing dnmt3bb.1 expression in the dorsal aorta. (B) Pixel intensity values of dnmt3bb.1 mRNA in runx1W84X/W84Xembryos (n=36) are significantly decreased compared to wild types (n=32) and heterozygotes (n=62) (ANOVA, p &#60; 0.001). The coefficients of variation are 24%, 22% and 21% for wild type, heterozygote and mutant groups, respectively. Blue, green and orange data point correspond to the example images from panel A. The bars represent mean &#177; s.d. ***p&#60;0.001 (Games-Howell post-hoc test). Figure adapted from Dobrzycki et al.3 <a href="https://www.jove.com/files/ftp_upload/59956/59956fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59956/59956fig03.jpg" /> 
Figure 3: Measuring runx1 expression levels by ISH in lmo4auob100mutants. (A) Representative images of ISH for runx1 in 28 hpf wild type (blue), heterozygous (green) and lmo4auob100/uob100 (orange) embryos, showing the expression in the dorsal aorta. (B) Quantification of the runx1 mRNA signal, detected by ISH, from 28 hpf wild type (n=15), heterozygous lmo4a+/- (het) (n=34) and lmo4auob100/uob100 mutant (n=18) embryos from one clutch shows no significant difference in runx1 pixel intensity among the different genotypes (ANOVA,&#62; p 0.6). Blue, green and orange data point correspond to the example images from panel A. The bars represent mean &#177; s.d. (C) Boxplots displaying normalized runx1 mRNA levels (2-&#916;Ct) in single wild type (blue; n=12) and lmo4auob100/uob100 (mut, orange; n=12) embryos, measured by qRT-PCR, showing decreased levels of runx1 in the mutants compared to wild type. *p &#60; 0.05 (t test). (D) Example of an ISH experiment on a 28 hpf embryo (stained for runx1, yellow arrowheads) showing high background. Figure adapted from Dobrzycki et al.3 <a href="https://www.jove.com/files/ftp_upload/59956/59956fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59956/59956fig04.jpg" /> 
Figure 4: Effect of background intensity correction on measurement outcomes. (A) Representative image of runx1 ISH staining in a wild type embryo at 28 hpf. (B) Same image after inversion and conversion to 8-bit. The region of interest (ROI) is highlighted in green and four different areas used for background correction (R1-R4) are highlighted in yellow. (C) Raw pixel intensity measurements in all regions shown in panel B. Note the intensity in R3 (yolk) is consistently higher than the actual ISH signal in the ROI (n=11). (D) Runx1 expression levels in the ROI using R1, R2 and R4 background areas. Areas for ROI, R1, R2 and R3~28500 pixels; R4~8500 pixels. Note that R3 background was not used for this comparison as the background correction (ROI-R3) consistently yielded negative values. (E) Runx1 expression levels in wild type and dll4-/- mutants using either R1 or R4 for background correction (n=10 for each sample). Statistical analysis in panels D and E was performed using a non-parametric Kruskal-Wallis test, assuming that the pixel intensity values are not normally distributed. <a href="https://www.jove.com/files/ftp_upload/59956/59956fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Genotyping and Quantification of In Situ Hybridization Staining in Zebrafish at JoVE.com

Genotyping and Quantification of In Situ Hybridization Staining in Zebrafish

Gene editing technologies have enabled researchers to generate zebrafish mutants to investigate gene function with relative ease. Here, we provide a guide to perform parallel embryo genotyping and quantification of in situ hybridization signals in zebrafish. This unbiased approach provides greater accuracy in phenotypical analyses based on in situ hybridization.

In situ hybridization (ISH) is an important technique that enables researchers to study mRNA distribution in situ and has been a critical technique in ...

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9.4K Views.  University of Oxford. Gene editing technologies have enabled researchers to generate zebrafish mutants to investigate gene function with relative ease. Here, we provide a guide to perform parallel embryo genotyping and quantification of in situ hybridization signals in zebrafish. This unbiased approach provides greater accuracy in phenotypical analyses based on in situ hybridization.

Video: Genotyping and Quantification of In Situ Hybridization Staining in Zebrafish

Multi-target Chromogenic Whole-mount In Situ Hybridization for Comparing Gene Expression Domains in Drosophila Embryos

To analyze gene regulatory networks active during embryonic development and organogenesis it is essential to precisely define how the different genes are expressed in spatial relation to each other in situ. Multi-target chromogenic whole-mount in situ hybridization (MC-WISH) greatly facilitates the instant comparison of gene expression patterns, as it allows distinctive visualization of different mRNA species in contrasting colors in the same sample specimen. This provides the possibility to relate gene expression domains topographically to each other with high accuracy and to define unique and overlapping expression sites. In the presented protocol, we describe a MC-WISH procedure for comparing mRNA expression patterns of different genes in Drosophila embryos. Up to three RNA probes, each specific for another gene and labeled by a different hapten, are simultaneously hybridized to the embryo samples and subsequently detected by alkaline phosphatase-based colorimetric immunohistochemistry. The described procedure is detailed here for Drosophila, but works equally well with zebrafish embryos.

Multi-target Chromogenic Whole-mount In Situ Hybridization for Comparing Gene Expression Domains in Drosophila Embryos

We describe a multi-target chromogenic whole-mount in situ hybridization (MC-WISH) procedure in intact Drosophila embryos allowing simultaneous and specific detection of three different mRNA distribution patterns by contrasting color precipitates.

To analyze gene regulatory networks active during embryonic development and organogenesis it is essential to precisely define how the different genes are ...

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

In recent years, the zebrafish embryo has emerged as a popular model to study developmental biology due to traits such as ex utero embryo development and optical transparency. In particular, the zebrafish embryo has become an important organism to study vertebrate kidney organogenesis as well as multiciliated cell (MCC) development. To visualize MCCs in the embryonic zebrafish kidney, we have developed a combined protocol of whole-mount fluorescent in situ hybridization (FISH) and whole mount immunofluorescence (IF) that enables high resolution imaging. This manuscript describes our technique for co-localizing RNA transcripts and protein as a tool to better understand the regulation of developmental programs through the expression of various lineage factors.

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

Cilia development is vital to proper organogenesis. This protocol describes an optimized method to label and visualize ciliated cells of the zebrafish.

In recent years, the zebrafish embryo has emerged as a popular model to study developmental biology due to traits such as ex utero embryo development and ...

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 ...

Biology

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. Although many immunohistochemistry protocols have been optimized for mammalian tissue and tissue sections, these protocols often require modification and optimization for non-mammalian model organisms. Zebrafish are increasingly used as a model system in basic, biomedical, and translational research to investigate the molecular, genetic, and cell biological mechanisms of developmental processes. Zebrafish offer many advantages as a model system but also require modified techniques for optimal protein detection. Here, we provide our protocol for whole-mount fluorescence immunohistochemistry in zebrafish embryos and larvae. This protocol additionally describes several different mounting strategies that can be employed and an overview of the advantages and disadvantages each strategy provides. We also describe modifications to this protocol to allow detection of chromogenic substrates in whole mount tissue and fluorescence detection in sectioned larval tissue. This protocol is broadly applicable to the study of many developmental stages and embryonic structures.

Here, we present a protocol for fluorescent antibody-mediated detection of proteins in whole preparations of zebrafish embryos and larvae.

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. ...

Using Flow Cytometry to Detect and Quantitate Altered Blood Formation in the Developing Zebrafish

The diversity of cell lineages that comprise mature blood in vertebrate animals arise from the differentiation of hematopoietic stem and progenitor cells (HSPCs). This is a critical process that occurs throughout the lifespan of organisms, and disruption of the molecular pathways involved during embryogenesis can have catastrophic long-term consequences. For a multitude of reasons, zebrafish (Danio rerio) has become a model organism to study hematopoiesis. Zebrafish embryos develop externally, and by 7 days postfertilization (dpf) have produced most of the subtypes of definitive blood cells that will persist for their lifetime. Assays to assess the number of hematopoietic cells have been developed, mainly utilizing specific histological stains, in situ hybridization techniques, and microscopy of transgenic animals that utilize blood cell-specific promoters driving the expression of fluorescent proteins. However, most staining assays and in situ hybridization techniques do not accurately quantitate the number of blood cells present; only large differences in cell numbers are easily visualized. Utilizing transgenic animals and analyzing individuals with fluorescent or confocal microscopy can be performed, but the quantitation of these assays relies on either counting manually or utilizing expensive imaging software, both of which can make errors. Development of additional methods to assess blood cell numbers would be economical, faster, and could even be automated to quickly assess the effect of CRISPR-mediated genetic modification, morpholino-mediated transcript reduction, and the effect of drug compounds that affect hematopoiesis on a large scale. This novel assay to quantitate blood cells is performed by dissociating whole zebrafish embryos and analyzing the amount of fluorescently labelled blood cells present. These assays should allow elucidation of molecular pathways responsible for blood cell generation, expansion, and regulation during embryogenesis, which will allow researchers to further discover novel factors altered during blood diseases, as well as pathways essential during the evolution of vertebrate hematopoiesis.

This assay is a simple method to quantitate hematopoietic cells in developing embryonic zebrafish. Blood cells from dissociated zebrafish are subjected to flow cytometry analysis. This allows the detection of blood defects in mutant animals and after genetic modification.

The diversity of cell lineages that comprise mature blood in vertebrate animals arise from the differentiation of hematopoietic stem and progenitor cells ...

In Situ Hybridization in Zebrafish Larvae and Juveniles during Mesonephros Development

The zebrafish forms two kidney structures in its lifetime. The pronephros (embryonic kidney) forms during embryonic development and begins to function at 2 days post fertilization. Consisting of only two nephrons, the pronephros serves as the sole kidney during larval life until more renal function is required due to the increasing body mass. To cope with this higher demand, the mesonephros (adult kidney) begins to form during metamorphosis. The new primary nephrons fuse to the pronephros and form connected lumens. Then, secondary nephrons fuse to primary ones (and so on) to create a branching network in the mesonephros. The vast majority of research is focused on the pronephros due to the ease of using embryos. Thus, there is a need to develop techniques to study older and larger larvae and juvenile fish to better understand mesonephros development. Here, an in situ hybridization protocol for gene expression analysis is optimized for probe penetration, washing of probes and antibodies, and bleaching of pigments to better visualize the mesonephros. The Tg(lhx1a-EGFP) transgenic line is used to label progenitor cells and the distal tubules of nascent nephrons. This protocol fills a gap in mesonephros research. It is a crucial model for understanding how new kidney tissues form and integrate with existing nephrons and provide insights into regenerative therapies.

In Situ Hybridization in Zebrafish Larvae and Juveniles during Mesonephros Development

The zebrafish is an important model for understanding kidney development. Here, an in situ hybridization protocol is optimized to detect gene expression in zebrafish larvae and juveniles during mesonephros development.

The zebrafish forms two kidney structures in its lifetime. The pronephros (embryonic kidney) forms during embryonic development and begins to function at ...

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.

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.

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

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 ...

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model congenital abnormalities. During the first day of life, the zebrafish embryo progresses through many developmental stages including fertilization, cleavage, gastrulation, segmentation, and the organogenesis of structures such as the kidney, heart, and central nervous system. The anatomy of a young zebrafish embryo presents several challenges for the visualization and analysis of the tissues involved in many of these events because the embryo develops in association with a round yolk mass. Thus, for accurate analysis and imaging of experimental phenotypes in fixed embryonic specimens between the tailbud and 20 somite stage (10 and 19 hours post fertilization (hpf), respectively), such as those stained using whole mount in situ hybridization (WISH), it is often desirable to remove the embryo from the yolk ball and to position it flat on a glass slide. However, performing a flat mount procedure can be tedious. Therefore, successful and efficient flat mount preparation is greatly facilitated through the visual demonstration of the dissection technique, and also helped by using reagents that assist in optimal tissue handling. Here, we provide our WISH protocol for one or two-color detection of gene expression in the zebrafish embryo, and demonstrate how the flat mounting procedure can be performed on this example of a stained fixed specimen. This flat mounting protocol is broadly applicable to the study of many embryonic structures that emerge during early zebrafish development, and can be implemented in conjunction with other staining methods performed on fixed embryo samples.

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

The zebrafish embryo is an excellent model for developmental biology research. During embryogenesis, zebrafish develop with a yolk mass, which presents three-dimensional challenges for sample observation and analysis. This protocol describes how to create two-dimensional flat mount preparations of whole mount in situ (WISH) stained zebrafish embryo specimens.

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model ...

In Situ Hybridization Combined with Immunohistochemistry in Cryosectioned Zebrafish Embryos

As a vertebrate, the zebrafish has been widely used in biological studies. Zebrafish and humans share high genetic homology, which allows its use as a model for human diseases. Gene function study is based on the detection of gene expression patterns. Although immunohistochemistry offers a powerful way to assay protein expression, the limited number of commercially available antibodies in zebrafish restricts the application of costaining. In situ hybridization is widely used in zebrafish embryos to detect mRNA expression. This protocol describes how to obtain images by combining in situ hybridization and immunohistochemistry for zebrafish embryo sections. In situ hybridization was performed prior to cryosectioning, followed by antibody staining. Immunohistochemistry and the imaging of a single cryosection were performed after in situ hybridization. The protocol is helpful to unravel the expression pattern of two genes, first by in situ transcript detection and then by immunohistochemistry against a protein in the same section.

In Situ Hybridization Combined with Immunohistochemistry in Cryosectioned Zebrafish Embryos

This protocol describes how to obtain images by combining in situ hybridization and immunohistochemistry of zebrafish embryonic sections. In situ hybridization was performed prior to cryosectioning, followed by antibody staining. It is useful to detect the expression patterns of two genes in zebrafish if there is a paucity of antibodies.

As a vertebrate, the zebrafish has been widely used in biological studies. Zebrafish and humans share high genetic homology, which allows its use as a ...

Genotyping and Quantification of In Situ Hybridization Staining in Zebrafish

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Chapters in this video

Multi-target Chromogenic Whole-mount In Situ Hybridization for Comparing Gene Expression Domains in Drosophila Embryos

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

Using Flow Cytometry to Detect and Quantitate Altered Blood Formation in the Developing Zebrafish

In Situ Hybridization in Zebrafish Larvae and Juveniles during Mesonephros Development

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

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

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

In Situ Hybridization Combined with Immunohistochemistry in Cryosectioned Zebrafish Embryos