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><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

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. Dit protocol is belangrijk omdat het de bepaling van genexpressie in oudere zebravislarven en juvenielen tijdens metamorfose mogelijk maakt. Het voordeel van deze techniek is dat verschillende stappen zijn geoptimaliseerd voor sondepenetratie en visualisatie van de nier. Om te beginnen, stel volwassen zebravissen in om te paren door een mannelijke en een vrouwelijke vis toe te voegen in een paringstank in de late namiddag na hun laatste maaltijd.Verzamel de volgende dag de embryo's in ...

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

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

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

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.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

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

The rapid proliferation of cells, the tissue-specific expression of genes and the emergence of signaling networks characterize early embryonic development of all vertebrates. The kinetics and location of signals - even within single cells - in the developing embryo complements the identification of important developmental genes. Immunostaining techniques are described that have been shown to define the kinetics of intracellular and whole animal signals in structures as small as primary cilia. The techniques for fixing, imaging and processing images using a laser-scanning confocal compound microscope can be completed in as few as 36 hr.
Zebrafish (Danio rerio) is a desirable organism for investigators who seek to conduct studies in a vertebrate species that is affordable and relevant to human disease. Genetic knockouts or knockdowns must be confirmed by the loss of the actual protein product. Such confirmation of protein loss can be achieved using the techniques described here. Clues into signaling pathways can also be deciphered by using antibodies that are reactive with proteins that have been post-translationally modified by phosphorylation. Preserving and optimizing the phosphorylated state of an epitope is therefore critical to this determination and is accomplished by this protocol.
This study describes techniques to fix embryos during the first 72 hr of development and co-localize a variety of relevant epitopes with cilia in the Kupffer&#39;s Vesicle (KV), the kidney and the inner ear. These techniques are straightforward, do not require dissection and can be completed in a relatively short period of time. Projecting confocal image stacks into a single image is a useful means of presenting these data.

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.

The rapid proliferation of cells, the tissue-specific expression of genes and the emergence of signaling networks characterize early embryonic development ...

Chemistry

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

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

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene knockdown approaches can be utilized to examine the molecular mechanisms underlying nervous system development. Second, large clutches of developmentally synchronized embryos provide large experimental sample sizes. Third, the optical clarity of the zebrafish embryo permits researchers to visualize progenitor, glial, and neuronal populations. Although zebrafish embryos are transparent, specimen thickness can impede effective microscopic visualization. One reason for this is the tandem development of the spinal cord and overlying somite tissue. Another reason is the large yolk ball, which is still present during periods of early neurogenesis. In this article, we demonstrate microdissection and removal of the yolk in fixed embryos, which allows microscopic visualization while preserving surrounding somite tissue. We also demonstrate semipermanent mounting of zebrafish embryos. This permits observation of neurodevelopment in the dorso-ventral and anterior-posterior axes, as it preserves the three-dimensionality of the tissue.

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.

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene ...

Neuroscience

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 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 ...

In Situ Hybridization in Zebrafish Larvae and Juveniles during Mesonephros Development

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