Name: In Situ Hybridization in Zebrafish Larvae and Juveniles during Mesonephros Development (Video) | JoVE Journal
Uploaded: 2021-08-13T14:45:00.000+00:00
Duration: 8 min 16 s
Description: Learn in situ hybridization for gene expression analysis in zebrafish larvae and juveniles during mesonephros development. This optimized protocol improves probe penetration and visualization of kidney structures.

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5 minutes

Funding was provided by the Pennsylvania Academy of Science, and the Commonwealth of Pennsylvania Biologists, and the Indiana University of Pennsylvania (School of Graduate Studies and Research, Department of Biology, and the Cynthia Sushak Undergraduate Biology Fund for Excellence). The Tg(lhx1a-EGFP) transgenic line was provided by Dr. Neil Hukriede (University of Pittsburgh).

The use of zebrafish larvae and juveniles is approved by the IUP IACUC (protocol #02-1920, #08-1920). Details of the solution content are listed in the Table of Materials.
1. Raising larvae
NOTE: It will take up to 21 days or more to raise larvae and juveniles to the stage of interest.
<ol>
	<li>Set up adult zebrafish to mate by adding 1 male and 1 female fish in a mating tank in the late afternoon after their last meal. 
	&#8203;NOTE: Not all fish pairs will mate. Start by setting up 20 pairs to determine the mating rate and adjust accordingly in future experiments.</li>
	<li>The next day after the fish have finished mating (around 1 pm), collect the embryos with E3 medium in Petri plates.
	<ol>
		<li>Ensure that there are no more than 30 embryos per plate.</li>
		<li>Remove debris (such as feces) from the Petri plates and add 20 mL of E3 medium.</li>
		<li>Incubate at 28.5 &#176;C for 1 day.</li>
	</ol>
	</li>
	<li>Put the adult zebrafish back in the aquatic system.</li>
	<li>At 1 dpf, replace the E3 medium with fresh E3.
	<ol>
		<li>Incubate at 28.5 &#176;C for 4 more days until 5 dpf.</li>
	</ol>
	</li>
	<li>Put a 400 &#181;m screen in a 2.8 L tank and fill it with 2 cm of system water.
	<ol>
		<li>Add the 5 dpf larvae from 1 plate (up to 30 total larvae).</li>
		<li>Put the tank in the aquatic system, but do not start the water flow.</li>
	</ol>
	</li>
	<li>Feed the larvae 4 mL of powder food twice a day until 14 dpf.</li>
	<li>At 6 dpf, start the water flow at 1 drip per second. 
	NOTE: Check the waterflow daily and adjust accordingly.</li>
	<li>At 8 dpf, increase the water flow to a slow, steady stream.</li>
	<li>At 14 dpf, feed live brine shrimp in addition to the powder food.</li>
</ol>
2. Day 1 - 2: Fixing larvae
<ol>
	<li>Remove a tank of larvae at the desired timepoint.</li>
	<li>Fill a Petri plate with 20 mL of system water.</li>
	<li>Use a fine mesh net to gently scoop the larvae and bring them to the water surface.
	<ol>
		<li>Cut off the tip of a transfer pipette to give a wider opening and transfer the larvae to the Petri plate. The larger pipette mouth allows for the transfer of several small larvae or a few larger ones at once.</li>
	</ol>
	</li>
	<li>Add 2 mL of tricaine (2%, pH 7 stock) to immobilize the larvae.
	<ol>
		<li>After the larvae stop moving, remove most of the water and add 10 mL of tricaine. Wait for 15 min for the larvae to be euthanized.</li>
		<li>Optional: For juveniles longer than 8 mm, cut off the heads and tails with a razor blade (after euthanasia) under a dissecting microscope to improve penetration of the fixing solution. Make sure to record the total length of each animal before cutting off the head and tail and isolate each one in its own Petri plate. Refer to step 3 for how to measure the animals.</li>
	</ol>
	</li>
	<li>Replace the tricaine with 20 mL of fixing solution (4% paraformaldehyde, 1% DMSO). Put the lid back on the Petri plate and rock slowly in a fume hood. 
	NOTE: It is important to use a rocking platform. A shaking platform with a circular motion will cause the animal axis to be curved. Use only fresh (not premade frozen) fixing solution. 
	CAUTION: The fixing solution contains paraformaldehyde, which is a probable carcinogen. Use fume hood (or mask) and gloves to measure the powder.</li>
	<li>After 30 min, replace the fixing solution with a fresh fixing solution.
	<ol>
		<li>Collect the used fixing solution in a separate waste container for proper disposal.</li>
		<li>Transfer the larvae into a 50 mL tube containing 25 mL of fresh fixing solution.</li>
		<li>Make sure the cap is tight and rock slowly at 4 &#176;C for 2 days. It is possible to incubate longer for convenience.</li>
	</ol>
	</li>
</ol>
3. Day 3: Measuring larvae
<ol>
	<li>In a fume hood, replace the fixing solution with 20 mL of PBST.</li>
	<li>Transfer the larvae into a Petri plate.</li>
	<li>Put a flat ruler on a dissecting microscope and put the Petri plate on top of the ruler.</li>
	<li>Use an eyelash manipulator to move each larva onto the ruler to measure its total length (from the snout to the tip of the caudal fin) (Figure 1).</li>
	<li>Combine several larvae of similar lengths (e.g., 5.0 mm and 5.1 mm) into one 5.5 mL glass vial, up to 10 larvae per vial.</li>
</ol>
4. Day 3 - 4: Dehydration
<ol>
	<li>Replace the PBST with 4 mL of 100% methanol. Rock the vial for 5 min at room temperature.</li>
	<li>Replace the methanol with fresh methanol and rock for 5 min. Repeat one more time.</li>
	<li>Store the vial at -20 &#176;C for 2 days. It is possible to incubate longer for convenience.</li>
</ol>
5. Day 5: Rehydration
<ol>
	<li>Warm the vial to room temperature.</li>
	<li>Replace the methanol with 4 mL of 75% methanol/25% PBST. Rock for 5 min at room temperature.</li>
	<li>Replace the solution in step 5.2 with 4 mL of 50% methanol/50% PBST and rock for 5 min.</li>
	<li>Replace the solution in step 5.3 with 4 mL of 25% methanol/75% PBST and rock for 5 min.</li>
	<li>Replace the solution in step 5.4 with 4 mL PBST and rock for 10 min.</li>
	<li>Replace the PBST with 4 mL of fresh PBST and rock for 10 min. Repeat one more time.</li>
</ol>
6. Day 5: Proteinase K digest
<ol>
	<li>Replace the PBST with 2 mL of the proteinase K solution (20 &#181;g/mL, 1% DMSO final concentration in PBST) and rock at room temperature for 30 min. 
	NOTE: Larvae longer than 6 mm will need a longer incubation time and/or a higher proteinase K concentration. The exact time and concentration will need to be determined empirically. Start with a 10 min increase in incubation time for every 0.5 mm longer than 6 mm.</li>
	<li>Replace the proteinase K solution with 4 mL of PBST and rock for 10 min.</li>
	<li>Replace the PBST with 4 mL of fresh PBST and rock for 10 min. Repeat one more time.</li>
	<li>Replace the PBST with 4 mL of fresh fixing solution and rock for 1 h. It is possible to incubate longer at 4 &#176;C for convenience.</li>
</ol>
7. Day 5: Bleaching
<ol>
	<li>Replace the fixing solution with 4 mL of PBST and rock at room temperature for 10 min.</li>
	<li>Replace the PBST with 4 mL of fresh PBST and rock for 10 min. Repeat one more time.</li>
	<li>Transfer the larvae to a 6-well plate (up to 10 larvae per well).</li>
	<li>Replace the PBST with 3 mL of fresh bleaching solution.</li>
	<li>Rock at room temperature and monitor the pigmentation under a dissecting microscope every 5 min. Look for the disappearance of pigmentation along the mesonephros (dorsal to the swim bladder and gut). 
	NOTE: For larvae up to 6 mm long, it will take up to 20 min to bleach the pigmentation surrounding the mesonephros. Do not bleach longer than necessary to preserve the integrity of the larvae.</li>
	<li>Transfer the larvae back into a glass vial.</li>
	<li>Replace the bleaching solution with 4 mL of PBST. Rock for 10 min.</li>
	<li>Replace the PBST with 4 mL of fresh PBST and rock for 10 min. Repeat one more time.</li>
	<li>Replace the PBST with 4 mL of fresh fixing solution and rock for 1 h. It is possible to incubate longer at 4 &#176;C for convenience.</li>
</ol>
8. Day 5: Prehybridization
NOTE: For steps done at 70 &#176;C, it is important to work quickly to minimize the vial cooling down.
<ol>
	<li>Replace the fixing solution with 4 mL of PBST and rock at room temperature for 10 min.</li>
	<li>Replace the PBST with 4 mL of fresh PBST and rock for 10 min. Repeat one more time.</li>
	<li>Replace the PBST with 4 mL of the Hyb- solution, and rock at room temperature for 10 min. 
	CAUTION: The Hyb-, Hyb+, and probe solutions contain formamide, which can irritate the skin. Use gloves to handle these solutions.</li>
	<li>Replace the Hyb- with 4 mL of fresh Hyb- and rock for 10 min. Repeat one more time.
	<ol>
		<li>Collect the used Hyb-, Hyb+, and probe solutions in a separate waste container for proper disposal.</li>
	</ol>
	</li>
	<li>Replace the Hyb- with 4 mL of the Hyb+ solution.</li>
	<li>Incubate the vial at 70 &#176;C overnight.</li>
	<li>Dilute the EGFP-fluorescein probe 1:100 in 500 &#181;L of Hyb+ and incubate it O/N at 70 &#176;C. 
	&#8203;NOTE: Follow the manufacturer&#39;s instructions for probe synthesis.</li>
</ol>
9. Day 6: Probe hybridization
<ol>
	<li>Replace the Hyb+ solution in the vial with the preheated probe and incubate at 70 &#176;C overnight. 
	NOTE: Larvae longer than 6 mm need 2 days of incubation.</li>
	<li>Preheat the wash buffer, 2x SSCT, and 0.2x SSCT solutions at 70 &#176;C for overnight.</li>
</ol>
10. Day 7: Probe washing
<ol>
	<li>Replace the probe with 4 mL of the preheated wash buffer and incubate at 70 &#176;C for 30 min.
	<ol>
		<li>Collect the used probe in a separate waste container for proper disposal.</li>
	</ol>
	</li>
	<li>Replace the wash buffer with 4 mL of fresh wash buffer and incubate at 70 &#176;C for 30 min.</li>
	<li>Replace the wash buffer with 4 mL of the preheated 2x SSCT solution and incubate it at 70 &#176;C for 15 min.</li>
	<li>Add 50 mL of preheated 0.2x SSCT into a 50 mL tube.
	<ol>
		<li>Insert a cell strainer (100 &#181;m) into the top of the tube. Ensure that the cell strainer is pushed all the way down until it stops.</li>
		<li>Transfer the larvae from the glass vial into the cell strainer. Ensure that the larvae are submerged in the buffer and incubate at 70 &#176;C for 2 h.</li>
	</ol>
	</li>
	<li>Prepare a new tube containing 50 mL of preheated 0.2x SSCT.
	<ol>
		<li>Transfer the cell strainer from step 10.4.2 into the new tube of 0.2x SSCT and incubate at 70 &#176;C for 2 h.</li>
		<li>Repeat step 10.5 one more time.</li>
	</ol>
	</li>
</ol>
11. Day 7: Blocking
<ol>
	<li>Transfer the larvae to a new glass vial and cool to room temperature.</li>
	<li>Replace the 0.2x SSCT with 4 mL of 67% 0.2x SSCT/33% MABT and rock at room temperature for 10 min.</li>
	<li>Replace the solution in step 11.2 with 4 mL of 33% 0.2x SSCT/67% MABT and rock for 10 min.</li>
	<li>Replace the solution in step 11.3 with 4 mL of MABT and rock for 10 min.</li>
	<li>Replace the MABT with 4 mL of fresh MABT and rock for 10 min. Repeat one more time.</li>
	<li>Replace the MABT with 4 mL of the blocking solution and incubate at 4 &#176;C overnight. It is possible to incubate longer for convenience.</li>
</ol>
12. Day 8 - 9: Antibody incubation
<ol>
	<li>Dilute the anti-fluorescein antibody 1:5,000 in 500 &#181;L blocking solution.</li>
	<li>Replace the blocking solution with the antibody solution and incubate at 4 &#176;C for 2 days. It is possible to incubate longer for convenience.
	<ol>
		<li>Swirl the vial twice a day to agitate the larvae. 
		&#8203;NOTE: Larvae longer than 6 mm need 1-2 more days of incubation.</li>
	</ol>
	</li>
</ol>
13. Day 10 - 11: Antibody washing
<ol>
	<li>Replace the antibody solution with 4 mL of PBST and rock at room temperature for 10 min.</li>
	<li>Replace the PBST with 4 mL of fresh PBST and rock for 10 min. Repeat one more time.</li>
	<li>Transfer the larvae into a 50 mL tube.</li>
	<li>Add 40 mL of PBST2. Lay the tube on its side and rock at 4 &#176;C overnight.</li>
	<li>Replace the PBST2 with 40 mL of fresh PBST2 and rock at 4 &#176;C overnight. It is possible to incubate longer for convenience. 
	&#8203;NOTE: Larvae longer than 6 mm need 1-2 more days of washing.</li>
</ol>
14. Day 12: Staining
<ol>
	<li>Transfer the larvae to a 6-well plate.</li>
	<li>Replace the PBST2 with 3 mL of staining buffer and rock at room temperature for 5 min.</li>
	<li>Replace the staining buffer with 3 mL of fresh staining buffer and rock for 5 min. Repeat one more time.</li>
	<li>Replace the staining buffer with 3 mL of the staining solution. Cover with aluminum foil and monitor the staining over the next few hours.
	<ol>
		<li>For probes with a weak signal, incubate at 4 &#176;C overnight and replace the staining solution with fresh staining solution once in between.</li>
	</ol>
	</li>
	<li>When the desired staining intensity is reached, replace the staining solution with 3 mL of the stopping solution and rock for 30 min.</li>
	<li>Replace the stopping solution with 3 mL of fresh stopping solution and rock for 30 min. Repeat one more time.</li>
	<li>Transfer the larvae to a new glass vial and replace the stopping solution with 4 mL of fresh fixing solution and incubate for 1 h at room temperature. It is possible to incubate longer at 4 &#176;C for convenience.</li>
	<li>Store the larvae in the fixing solution in the dark at 4 &#176;C for up to a year.</li>
</ol>
15. Day 12: Imaging
<ol>
	<li>Replace the fixing solution with 4 mL of PBST and rock at room temperature for 10 min.</li>
	<li>Transfer the larvae to a 6-well plate.</li>
	<li>Replace the PBST with 4 mL of fresh PBST and rock for 10 min. Repeat one more time.</li>
	<li>Replace the PBST with 3 mL of 50% glycerol (in PBST) and rock for 10 min.</li>
	<li>Replace the solution in step 15.4 with 3 mL of 100% glycerol with no rocking for 10 min.</li>
	<li>Use a dissecting microscope to image the larvae directly in the 6-well plate or transfer the larvae into a depression slide to image under a compound microscope. Use an eyelash manipulator to orient the larvae.</li>
</ol>

<ol><li>Kinth, P., Mahesh, G., Panwar, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mapping+of+zebrafish+research%3A+a+global+outlook%5BTitle%5D)+AND+%22Zebrafish%22%5BJournal%5D)">Mapping of zebrafish research: a global outlook.</a> Zebrafish. 10 (4), 510-517 (2013).</li>
<li>Grunwald, D. J., Eisen, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Headwaters+of+the+zebrafish+-+emergence+of+a+new+model+vertebrate%5BTitle%5D)+AND+%22Nature+Reviews.+Genetics%22%5BJournal%5D)">Headwaters of the zebrafish - emergence of a new model vertebrate.</a> Nature Reviews. Genetics. 3 (9), 717-724 (2002).</li>
<li>Wingert, R. A., Davidson, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+zebrafish+pronephros%3A+a+model+to+study+nephron+segmentation%5BTitle%5D)+AND+%22Kidney+International%22%5BJournal%5D)">The zebrafish pronephros: a model to study nephron segmentation.</a> Kidney International. 73 (10), 1120-1127 (2008).</li>
<li>Drummond, I. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Kidney+development+and+disease+in+the+zebrafish%5BTitle%5D)+AND+%22Journal+of+the+American+Society+of+Nephrology%3A+JASN%22%5BJournal%5D)">Kidney development and disease in the zebrafish.</a> Journal of the American Society of Nephrology: JASN. 16 (2), 299-304 (2005).</li>
<li>Swanhart, L. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Zebrafish+kidney+development%3A+basic+science+to+translational+research%5BTitle%5D)+AND+%22Birth+defects+research.+Part+C%2C+Embryo+Today%3A+Reviews%22%5BJournal%5D)">Zebrafish kidney development: basic science to translational research.</a> Birth defects research. Part C, Embryo Today: Reviews. 93 (2), 141-156 (2011).</li>
<li>Dressler, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Advances+in+early+kidney+specification%2C+development+and+patterning%5BTitle%5D)+AND+%22Development%22%5BJournal%5D)">Advances in early kidney specification, development and patterning.</a> Development. 136 (23), 3863-3874 (2009).</li>
<li>Johnson, C. S., Holzemer, N. F., Wingert, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Laser+ablation+of+the+zebrafish+pronephros+to+study+renal+epithelial+regeneration%5BTitle%5D)+AND+%22Journal+of+Visualized+Experiments%3A+JoVE%22%5BJournal%5D)">Laser ablation of the zebrafish pronephros to study renal epithelial regeneration.</a> Journal of Visualized Experiments: JoVE. (54), e2845(2011).</li>
<li>Marra, A. N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Visualizing+gene+expression+during+zebrafish+pronephros+development+and+regeneration%5BTitle%5D)+AND+%22Methods+in+Cell+Biology%22%5BJournal%5D)">Visualizing gene expression during zebrafish pronephros development and regeneration.</a> Methods in Cell Biology. 154, 183-215 (2019).</li>
<li>McMenamin, S. K., Chandless, M. N., Parichy, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Working+with+zebrafish+at+postembryonic+stages%5BTitle%5D)+AND+%22Methods+in+Cell+Biology%22%5BJournal%5D)">Working with zebrafish at postembryonic stages.</a> Methods in Cell Biology. 134, 587-607 (2016).</li>
<li>Diep, C. Q., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Development+of+the+zebrafish+mesonephros%5BTitle%5D)+AND+%22Genesis%22%5BJournal%5D)">Development of the zebrafish mesonephros.</a> Genesis. 53 (3-4), 257-269 (2015).</li>
<li>Zhou, W., Boucher, R. C., Bollig, F., Englert, C., Hildebrandt, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Characterization+of+mesonephric+development+and+regeneration+using+transgenic+zebrafish%5BTitle%5D)+AND+%22American+Journal+of+Physiology.+Renal+Physiology%22%5BJournal%5D)">Characterization of mesonephric development and regeneration using transgenic zebrafish.</a> American Journal of Physiology. Renal Physiology. 299 (5), 1040-1047 (2010).</li>
<li>Diep, C. Q., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Identification+of+adult+nephron+progenitors+capable+of+kidney+regeneration+in+zebrafish%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">Identification of adult nephron progenitors capable of kidney regeneration in zebrafish.</a> Nature. 470 (7332), 95-100 (2011).</li>
<li>Diep, C. Q., Mikeasky, N., Davidson, A. J., et al. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aDiep%2C+CQ+%22The+Zebrafish+in+Biomedical+Research%3A+Biology%2C+Husbandry%2C+Diseases%2C+and+Research+Applications%22">The Zebrafish in Biomedical Research: Biology, Husbandry, Diseases, and Research Applications</a>. Cartner, S. C., et al. , Elsevier. 145-150 (2020).</li>
<li>Parichy, D., Elizondo, M., Mills, M., Gordon, T., Engeszer, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Normal+table+of+postembryonic+zebrafish+development%3A+staging+by+externally+visible+anatomy+of+the+living+fish%5BTitle%5D)+AND+%22Developmental+dynamics%3A+An+Official+Publication+of+the+American+Association+of+Anatomists%22%5BJournal%5D)">Normal table of postembryonic zebrafish development: staging by externally visible anatomy of the living fish.</a> Developmental dynamics: An Official Publication of the American Association of Anatomists. 238 (12), 2975-3015 (2009).</li>
<li>Guo, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((LIM+Homeobox+4+%28lhx4%29+regulates+retinal+neural+differentiation+and+visual+function+in+zebrafish%5BTitle%5D)+AND+%22Science+Reports%22%5BJournal%5D)">LIM Homeobox 4 (lhx4) regulates retinal neural differentiation and visual function in zebrafish.</a> Science Reports. 11 (1), 1977(2021).</li>
<li>Vauti, F., Stegemann, L. A., Vogele, V., Koster, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((All-age+whole+mount+in+situ+hybridization+to+reveal+larval+and+juvenile+expression+patterns+in+zebrafish%5BTitle%5D)+AND+%22PloS+One%22%5BJournal%5D)">All-age whole mount in situ hybridization to reveal larval and juvenile expression patterns in zebrafish.</a> PloS One. 15 (8), 0237167(2020).</li>
<li>Parichy, D. M., Turner, J. M., Parker, N. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Essential+role+for+puma+in+development+of+postembryonic+neural+crest-derived+cell+lineages+in+zebrafish%5BTitle%5D)+AND+%22Developmental+Biology%22%5BJournal%5D)">Essential role for puma in development of postembryonic neural crest-derived cell lineages in zebrafish.</a> Developmental Biology. 256 (2), 221-241 (2003).</li>
<li>Yang, H., Wanner, I. B., Roper, S. D., Chaudhari, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((An+optimized+method+for+in+situ+hybridization+with+signal+amplification+that+allows+the+detection+of+rare+mRNAs%5BTitle%5D)+AND+%22The+Journal+of+Histochemistry+%26+Cytochemistry%22%5BJournal%5D)">An optimized method for in situ hybridization with signal amplification that allows the detection of rare mRNAs.</a> The Journal of Histochemistry & Cytochemistry. 47 (4), 431-445 (1999).</li>
<li>McCampbell, K. K., Springer, K. N., Wingert, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Analysis+of+nephron+composition+and+function+in+the+adult+zebrafish+kidney%5BTitle%5D)+AND+%22Journal+of+Visualized+Experiments%3A+JoVE%22%5BJournal%5D)">Analysis of nephron composition and function in the adult zebrafish kidney.</a> Journal of Visualized Experiments: JoVE. (90), e51644(2014).</li>
</ol>

The authors have no conflicts of interest.

The in situ hybridization method described here is aimed toward studying mesonephros development. However, it can be applied to study the development of other tissues and organs during metamorphosis, such as the gut, nervous system, scales, and pigmentation14. Probes can be generated for endogenous genes or fluorescent markers in transgenic lines.
It is critical for the larvae to remain intact in order to observe the organs and tissues in their native context. ...

The zebrafish embryo is an important model for studying tissue development due to its small size, transparency, available tools, and survival without feeding for up to five days1,2. It has greatly contributed to the understanding of kidney development and the conservation between zebrafish and mammals3,4,5. The kidney plays an essential role in maintaining fluid homeostasis, filtering the blood, and excreting waste6. The nephron, the functional unit of the kidney, comprises a blood filter connected to a long tubule. In zebrafish, two kidney structures form throughout its life. The pronephros (the temporary embryonic kidney) forms first during early development. It consists of two nephrons running along the anterior-posterior axis and becomes functional at around 2 days post fertilization (dpf). The utility of the pronephros lies in its simplicity, having just two nephrons that are mostly linear and easy to visualize (although the proximal convoluted tubule begins to coil at three dpf)3. This has facilitated not only studies of its early development from the intermediate mesoderm, but also the segmentation pattern and tubule repair7,8.
The usefulness of zebrafish becomes limited after five dpf, when the yolk is diminished and the larvae rely on feeding in the aquatic system9. At around 2 weeks old, the larvae undergo metamorphosis into juveniles, where new tissues form and old tissues are lost and/or reorganized9. This is also when the mesonephros (the permanent adult kidney) forms10,11,12. The first adult nephron forms near the sixth somite, and fuses with the distal early tubule of the pronephros. Additional nephrons are added posterior to this position initially, but also toward the anterior later on. The primary nephrons in this first wave fuse to the tubules of the pronephros and share a common lumen to deposit their waste. Secondary nephrons form in the next wave and fuse to primary nephrons. This reiterative process creates a mesonephros that is branched, somewhat akin to the mammalian kidney. Presumably, the pronephros eventually loses its tubule identity and becomes two major collecting ducts where all nephrons drain their waste13.
Prior to the formation of the first adult nephron, single progenitor cells begin to appear in ~4 mm (total length) larvae (~10 dpf). These cells, which are marked in the Tg(lhx1a-EGFP) transgenic line, adhere to the pronephros and seem to migrate to future sites of nephrogenesis. The single cells coalesce into clusters, which differentiate into new nephrons12. It is unclear where these cells reside or what genes they express before the onset of mesonephrogenesis.
Understanding mesonephros development provides insights into the mammalian kidney in ways that the pronephros cannot. These include the formation of nephrogenic aggregates from single progenitor cells, functional integration of new nephrons with existing ones, and branching morphogenesis. However, there are limitations to studying postembryonic development. The larvae are less transparent due to their larger size and having pigmentation. The developmental timeline is not synchronous among individual animals and is highly dependent on environmental factors, such as feeding and crowding9,14. Although knockdown reagents exist, they are less effective in larvae compared to embryos15. In this protocol, an in situ hybridization method to determine gene expression during zebrafish mesonephros development is described. Several steps are optimized to increase visualization of the mesonephros and penetration and washing of the probe and antibody. The Tg(lhx1a-EGFP) transgenic line is used along with a probe for EGFP to label single progenitor cells, nephrogenic aggregates, and the distal tubules of nascent nephrons. This method will provide a deeper understanding of mesonephros development and insight into regenerative therapies.

<table><tbody><tr><td>Anti-fluorescein antibody</td><td>Roche/Sigma-Aldrich</td><td>11426338910</td><td></td></tr><tr><td>Bleaching solution</td><td></td><td></td><td>0.8% KOH, 0.9% H2O2 in PBST</td></tr><tr><td>Blocking reagent</td><td>Roche/Sigma-Aldrich</td><td>11096176001</td><td>Use for blocking solution, prepare according to manufacture&#039;s instruction</td></tr><tr><td>Cell strainer</td><td>Fisher Scientific</td><td>&amp;nbsp;22-363-549</td><td>100 &amp;mu;m</td></tr><tr><td>E3 medium</td><td></td><td></td><td>5 mM NaCl, 0.33 mM CaCl2, 0.33 mM MgSO4, 0.17 mM KCl, 0.0001% methylene blue</td></tr><tr><td>Eyelash manipulator</td><td>Fisher Scientific</td><td>NC1083208</td><td>Use to manipulate larvae</td></tr><tr><td>Fixing solution</td><td></td><td></td><td>4% paraformaldehyde, 1% DMSO in PBS; heat at 65&amp;deg;C while shaking until the powder dissolves, then add DMSO after it cools down</td></tr><tr><td>Fluorescein probe synthesis</td><td>Roche/Sigma-Aldrich</td><td>11685619910</td><td></td></tr><tr><td>Glass vial</td><td>Fisher Scientific</td><td>03-338B</td><td></td></tr><tr><td>Hatchfry encapsulation</td><td>Argent</td><td></td><td></td></tr><tr><td>Hyb- solution</td><td></td><td></td><td>50% formamide, 5X SSC, 0.1% Tween-20</td></tr><tr><td>Hyb+ solution</td><td></td><td></td><td>HYB-, 5 mg/mL torula RNA, 50 ug/mL heparin</td></tr><tr><td>MAB (10X)</td><td></td><td></td><td>1 M maleic acid, 1.5 M NaCl, pH 7.5</td></tr><tr><td>MABT</td><td></td><td></td><td>1X MAB, 0.1% Tween-20</td></tr><tr><td>Maleic acid</td><td>Sigma-Aldrich</td><td>M0375</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Sigma Aldrich</td><td>158127</td><td></td></tr><tr><td>PBS (10X)</td><td></td><td></td><td>8% NaCl, 0.2% KCl, 1.44% Na2HPO4, 0.24% KH2PO4</td></tr><tr><td>PBST</td><td></td><td></td><td>1X PBS, 0.1% Tween-20</td></tr><tr><td>PBST2</td><td></td><td></td><td>1X PBS, 0.2% Tween-20</td></tr><tr><td>Powder food</td><td></td><td></td><td>Mix 2 g of each of spirulina and hatchfry encapsulon in 50 mL of fish system water and shake well</td></tr><tr><td>Proteinase K</td><td>Sigma-Aldrich</td><td>P5568</td><td>Use to permeabilize larvae</td></tr><tr><td>Proteinase K solution</td><td></td><td></td><td>20&amp;nbsp; &amp;mu;g/mL, 1% DMSO final concentration in PBST</td></tr><tr><td>Spirulina microfine powder</td><td>Argent</td><td></td><td></td></tr><tr><td>SSC (20X)</td><td></td><td></td><td>3 M NaCl, 0.3 M sodium acetate anhydrous, pH 7, autoclave</td></tr><tr><td>SSCT (0.2X)</td><td></td><td></td><td>Dilute from 20X SSC, 0.1% Tween-20</td></tr><tr><td>SSCT (2X)</td><td></td><td></td><td>Dilute from 20X SSC, 0.1% Tween-20</td></tr><tr><td>Staining buffer</td><td></td><td></td><td>100 mM Tris pH 9.5, 50 mM MgCl2, 100 mM NaCl, 0.1% Tween-20</td></tr><tr><td>Staining solution</td><td></td><td></td><td>200 &amp;mu;g/mL iodonitrotetrazolium chloride, 200 &amp;mu;g/mL 5-Bromo-4-chloro-3-indolyl phosphate disodium salt, in staining buffer</td></tr><tr><td>Stopping solution</td><td></td><td></td><td>1 mM EDTA, pH 5.5, in PBST</td></tr><tr><td>Torula (yeast) RNA</td><td>Sigma-Aldrich</td><td>R6625</td><td></td></tr><tr><td>Tricaine</td><td>Sigma Aldrich</td><td>E10521</td><td>2%, pH 7</td></tr><tr><td>Wash buffer</td><td></td><td></td><td>50% formamide, 2X SSC, 0.1% Tween-20</td></tr></tbody></table>

in situ hybridization in zebrafish larvae and juveniles during mesonephros development

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.

Using the Tg(lhx1a-EGFP) transgenic line, it was demonstrated that this in situ hybridization protocol is effective in labeling kidney progenitor cells and various nephron structures during mesonephros development. As expected, the central nervous system is also labeled in this transgenic line (not shown). The initial mesonephric nephron forms at approximately 5.2 mm, dorsal to the pronephros (Figure 2A), and the distal tubule of this nephron is labeled by EGFP<sup class="x...

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 ...

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Research

JoVE Journal

Developmental Biology

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.

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

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

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.

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

Developmental processes such as proliferation, patterning, differentiation, and axon guidance can be readily modeled in the zebrafish spinal cord.  In this article, we describe a mounting procedure for zebrafish embryos, which optimizes visualization of these events.

Dissection and Lateral Mounting of Zebrafish Embryos: Analysis of Spinal Cord Development

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.

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

The zebrafish is a powerful model system for developmental biology and human disease research due to their genetic similarity with higher vertebrates. This protocol describes a methodology to create haploid zebrafish embryos that can be utilized for forward screen strategies to identify recessive mutations in genes essential for early embryogenesis.

Production of Haploid Zebrafish Embryos by In Vitro Fertilization

Techniques are described to immunostain phospho-epitopes in whole zebrafish embryos and then conduct two-color fluorescent confocal localization in cellular structures as small as primary cilia. The techniques for fixing and imaging can define the location and kinetics of the appearance or activation of specific proteins.

Immunostaining Phospho-epitopes in Ciliated Organs of Whole Mount Zebrafish Embryos

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

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

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

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

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

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.

In Situ Hybridization Combined with Immunohistochemistry in Cryosectioned Zebrafish Embryos

Learn in situ hybridization for gene expression analysis in zebrafish larvae and juveniles during mesonephros development. This optimized protocol improves probe penetration and visualization of kidney structures.

In Situ Hybridization in Zebrafish Larvae and Juveniles during Mesonephros Development

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