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>

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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. To retain tissue integrity, it is important to minimize the time of proteinase K treatment. It is important to determine the best treatment time for each new batch of enzyme. Alternatively, acetone can be used instead of proteinase K for improved tissue integrity16. Excessive bleaching also reduces tissue integrity. Minimal bleaching and allowing the eyes to retain some pigmentation help in visualizing the larvae during washes. It is common for the eyes to fall off during the hybridization step, which is an indicator of tissue fragility. The use of DMSO with the fixing and proteinase K solution is crucial for tissue penetration17.
A limitation of this method is the poor penetration of reagents in larger animals. To improve penetration, the head and tail can be removed with a razor blade before fixation17. The gut can be removed with fine tweezers and tungsten needles after fixation to allow direct access of the reagents to the mesonephros. Long probes (greater than 1 KB) can have poor penetration of the tissue, but they can be hydrolyzed into short fragments (around 0.3 KB) to improve penetration18. For probes with weak signals, control probes with the sense sequence can be used to differentiate between the background and the probe signal. Larger animals will have a higher background staining of the somites (Figure 2C, arrows). However, this can be minimized with longer washes of the probe and antibodies.
The protocol here can also be applied to dissected and isolated tissues and organs, such as the adult zebrafish kidney12,19. Although there are published descriptions of similar protocols9,16, none of them describe the entire process from rearing larvae to in situ hybridization with this level of detail. Therefore, this method provides an additional tool in deciphering the development of the vertebrate kidney.

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#39;s instruction</td>
		</tr>
		<tr>
			<td>Cell strainer</td>
			<td>Fisher Scientific</td>
			<td>&#160;22-363-549</td>
			<td>100 &#956;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&#176;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&#160; &#956;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 &#956;g/mL iodonitrotetrazolium chloride, 200 &#956;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

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 EGFP10,12. Progenitor clusters are present at this stage and later on (Figure 2A-B, arrowheads), and single progenitor cells are also labeled (Figure 2C). This method provides an additional tool in studying kidney development and helps shed light on understanding the human kidney.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62930/62930fig01.jpg" /> 
Figure 1: Measuring larvae. Fixed larvae in a Petri plate are placed on top of a flat ruler. Under a dissecting microscope, the larvae are measured and separated by groups of similar sizes. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62930/62930fig01large.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/62930/62930fig02.jpg" /> 
Figure 2: Mesonephros development. At around 5.2 mm in the Tg(lhx1a-EGFP) transgenic line, the first mesonephric nephron is formed dorsal to pronephros and swim bladder (SB) (A). Clusters of progenitor cells are present during mesonephros development (A-B, arrowheads) in addition to single progenitor cells (C, bracket). In larger juveniles, background staining can occur in the somites (C, arrows). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62930/62930fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about In Situ Hybridization in Zebrafish Larvae and Juveniles during Mesonephros Development at JoVE.com

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.

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 ...

in-situ-hybridization-zebrafish-larvae-juveniles-during-mesonephros

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

2.6K Views.  Indiana University of Pennsylvania. 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.

Video: In Situ Hybridization in Zebrafish Larvae and Juveniles during Mesonephros Development

Understanding Early Organogenesis Using a Simplified In Situ Hybridization Protocol in Xenopus

Organogenesis is the study of how organs are specified and then acquire their specific shape and functions during development. The Xenopuslaevis embryo is very useful for studying organogenesis because their large size makes them very suitable for identifying organs at the earliest steps in organogenesis. At this time, the primary method used for identifying a specific organ or primordium is whole mount in situ hybridization with labeled antisense RNA probes specific to a gene that is expressed in the organ of interest. In addition, it is relatively easy to manipulate genes or signaling pathways in Xenopus and in situ hybridization allows one to then assay for changes in the presence or morphology of a target organ. Whole mount in situ hybridization is a multi-day protocol with many steps involved. Here we provide a simplified protocol with reduced numbers of steps and reagents used that works well for routine assays. In situ hybridization robots have greatly facilitated the process and we detail how and when we utilize that technology in the process. Once an in situ hybridization is complete, capturing the best image of the result can be frustrating. We provide advice on how to optimize imaging of in situ hybridization results. Although the protocol describes assessing organogenesis in Xenopus laevis, the same basic protocol can almost certainly be adapted to Xenopus tropicalis and other model systems.

Understanding Early Organogenesis Using a Simplified In Situ Hybridization Protocol in Xenopus

The Xenopus laevis embryo continues to be exceptionally useful in the study of early development due to its large size and ease of manipulation. A simplified protocol for whole mount in situ hybridization protocol is provided that can be used in the identification of specific organs in this model system.

Organogenesis is the study of how organs are specified and then acquire their specific shape and functions during development.  The Xenopuslaevis embryo ...

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two layers of skin that enclose undifferentiated mesenchyme, and because the larval tail fin regenerates rapidly within 2-3 days. Using this system, we demonstrate a method for capturing the repair dynamics of the amputated tail fin with time-lapse video brightfield stereomicroscopy. We demonstrate that fin amputation triggers a contraction of the amputation wound and extrusion of cells around the wound margin, leading to their subsequent clearance. Fin regeneration proceeds from proximal to distal direction after a short delay. In addition, developmental growth of the larva can be observed during all stages. The presented method provides an opportunity for observing and analyzing whole tissue-scale behaviors such as fin development and growth in a simple microscope setting, which is easily adaptable to any stereomicroscope with time-lapse capabilities.

We present a protocol for capturing the dynamics of zebrafish larval tail fin regeneration on a whole-tissue scale using brightfield-based stereomicroscopy. This technique enables capturing the regeneration dynamics with single cell resolution. This methodology can be adapted to any stereomicroscope equipped with a CCD camera and time-lapse software.

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two ...

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Induction of Hypoxia in Living Frog and Zebrafish Embryos

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and zebrafish embryos. Our system comprises a chamber featuring a simple setup that is nevertheless robust to induce and maintain a specific oxygen concentration and temperature in any experimental solution of choice. The presented system is very cost-effective but highly functional, it allows induction and sustainment of hypoxia for direct experiments in vivo and for various time periods up to 48 h.
To monitor and study the effects of hypoxia, we have employed two methods - measurement of levels of hypoxia-inducible factor 1alpha (HIF-1&#945;) in whole embryos or specific tissues and determination of retinal stem cell proliferation by 5-ethynyl-2&#8242;-deoxyuridine (EdU) incorporation into the DNA. HIF-1&#945; levels can serve as a general hypoxia marker in the whole embryo or tissue of choice, here embryonic retina. EdU incorporation into the proliferating cells of embryonic retina is a specific output of hypoxia induction. Thus, we have shown that hypoxic embryonic retinal progenitors decrease proliferation within 1 h of incubation under 5% oxygen of both frog and zebrafish embryos.
Once mastered, our setup can be employed for use with small aquatic model organisms, for direct in vivo experiments, any given time period and under normal, hypoxic or hyperoxic oxygen concentration or under any other given gas mixture.

We introduce a novel hypoxic chamber system for use with aquatic organisms such as frog and zebrafish embryos. Our system is simple, robust, cost-effective and allows the induction and sustainment of hypoxia in vivo and for up to 48 h. We present 2 reproducible methods to monitor the effectiveness of hypoxia.

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and ...

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 ...

Wholemount In Situ Hybridization for Astyanax Embryos

In recent years, a draft genome for the blind Mexican cavefish (Astyanax mexicanus) has been released, revealing the sequence identities for thousands of genes. Prior research into this emerging model system capitalized on comprehensive genome-wide investigations that have identified numerous quantitative trait loci (QTL) associated with various cave-associated phenotypes. However, the ability to connect genes of interest to the heritable basis for phenotypic change remains a significant challenge. One technique that can facilitate deeper understanding of the role of development in troglomorphic evolution is whole-mount in situ hybridization. This technique can be implemented to directly compare gene expression between cave- and surface-dwelling forms, nominate candidate genes underlying established QTL, identify genes of interest from next-generation sequencing studies, or develop other discovery-based approaches. In this report, we present a simple protocol, supported by a flexible checklist, that can be widely adapted for use well beyond the presented study system. It is hoped that this protocol can serve as a broad resource for the Astyanax community and beyond.

Wholemount In Situ Hybridization for Astyanax Embryos

This protocol enables visualization of gene expression in embryonic Astyanax cavefish. This approach has been developed with the goal of maximizing gene expression signal, while minimizing non-specific background staining.

In recent years, a draft genome for the blind Mexican cavefish (Astyanax mexicanus) has been released, revealing the sequence identities for thousands of ...

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.

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 ...

Whole-Mount In Situ Hybridization in Zebrafish Embryos and Tube Formation Assay in iPSC-ECs to Study the Role of Endoglin in Vascular Development

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in zebrafish during different developmental stages. To investigate the role of endoglin(eng) in vessel formation during the development of hereditary hemorrhagic telangiectasia&#160;(HHT), morpholino-mediated targeted knockdown of eng in zebrafish are used to study its temporal expression and associated functions. Here, whole-mount in situ RNA hybridization (WISH) is employed for the analysis of eng and its downstream genes in zebrafish embryos. Also, tube formation assays are performed in HHT patient-derived induced pluripotent stem cell-differentiated&#160;endothelial cells (iPSC-ECs; with eng mutations). A specific signal amplifying system using the whole amount In Situ Hybridization &#8211; WISH provides higher resolution and lower background results compared to traditional methods. To obtain a better signal, the post-fixation time is adjusted to 30 min after probe hybridization. Because fluorescence staining is not sensitive in zebrafish embryos, it is replaced with diaminobezidine (DAB) staining here. In this protocol, HHT&#160;patient-derived iPSC lines containing an eng mutation are differentiated into endothelial cells. After coating a plate with basement membrane matrix for 30 min at 37 &#176;C, iPSC-ECs are seeded as a monolayer into wells and kept at 37 &#176;C for 3 h. Then, the tube length and number of branches are calculated using microscopic images. Thus, with this improved WISH protocol, it is shown that reduced eng expression affects endothelial progenitor formation in zebrafish embryos. This is further confirmed by tube formation assays using iPSC-ECs derived from a patient with HHT. These assays confirm the role for eng in early vascular development.

Presented here is a protocol for whole-mount in situ RNA hybridization analysis in zebrafish and tube formation assay in patient-derived induced pluripotent stem cell-derived endothelial cells to study the role of endoglin in vascular formation.

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in ...

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. ...

Fluorescent Calcium Imaging and Subsequent In Situ Hybridization for Neuronal Precursor Characterization in Xenopus laevis

Spontaneous intracellular calcium activity can be observed in a variety of cell types and is proposed to play critical roles in a variety of physiological processes. In particular, appropriate regulation of calcium activity patterns during embryogenesis is necessary for many aspects of vertebrate neural development, including proper neural tube closure, synaptogenesis, and neurotransmitter phenotype specification. While the observation that calcium activity patterns can differ in both frequency and amplitude suggests a compelling mechanism by which these fluxes might transmit encoded signals to downstream effectors and regulate gene expression, existing population-level approaches have lacked the precision necessary to further explore this possibility. Furthermore, these approaches limit studies of the role of cell-cell interactions by precluding the ability to assay the state of neuronal determination in the absence of cell-cell contact. Therefore, we have established an experimental workflow that pairs time-lapse calcium imaging of dissociated neuronal explants with a fluorescence in situ hybridization assay, allowing the unambiguous correlation of calcium activity pattern with molecular phenotype on a single-cell level. We were successfully able to use this approach to distinguish and characterize specific calcium activity patterns associated with differentiating neural cells and neural progenitor cells, respectively; beyond this, however, the experimental framework described in this article could be readily adapted to investigate correlations between any time-series activity profile and expression of a gene or genes of interest.

Fluorescent Calcium Imaging and Subsequent In Situ Hybridization for Neuronal Precursor Characterization in Xenopus laevis

We present a two-part protocol that combines fluorescent calcium imaging with in situ hybridization, allowing the experimenter to correlate patterns of calcium activity with gene expression profiles on a single-cell level.

Spontaneous intracellular calcium activity can be observed in a variety of cell types and is proposed to play critical roles in a variety of physiological ...

In Situ Hybridization in Zebrafish Larvae and Juveniles during Mesonephros Development

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Understanding Early Organogenesis Using a Simplified In Situ Hybridization Protocol in Xenopus

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

Induction of Hypoxia in Living Frog and Zebrafish Embryos

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

Wholemount In Situ Hybridization for Astyanax Embryos

Genotyping and Quantification of In Situ Hybridization Staining in Zebrafish

Whole-Mount In Situ Hybridization in Zebrafish Embryos and Tube Formation Assay in iPSC-ECs to Study the Role of Endoglin in Vascular Development

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

Fluorescent Calcium Imaging and Subsequent In Situ Hybridization for Neuronal Precursor Characterization in Xenopus laevis