Name: visualizing multiciliated cells in the zebrafish through a combined protocol of whole mount fluorescent in situ hybridization and immunofluorescence
Uploaded: 2017-11-18T13:00:00.000+00:00
Duration: 9 min 33 s
Description: Watch this Scientific Journal Video about Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence at JoVE.c

This work was supported in part by the grant R01DK100237 to R.A.W. and the National Science Foundation Graduate Research Fellowship No. DGE-1313583 to A.N.M. We also thank the College of Science Summer Undergraduate Research Fellowship Program for support to M.U. We would like to thank the Center for Zebrafish Research at the University of Notre Dame for their dedicated care of our zebrafish. We would also like to thank the Department of Biological Sciences as well as the members of our lab for all of their support and valuable insights.

The following protocol uses zebrafish adults maintained and cared for by the Center for Zebrafish Research at the University of Notre Dame. All methods for working with zebrafish adults and embryos were approved by the Institutional Animal Care and Use Committee.
1. Embryo Fixation
<ol>
	<li>Collect zebrafish embryos using previously described methods39,40. At 24 hpf, add 500 &#181;L of non-specific protease mixture solution (50 mg/mL) to the E3 in an embryo dish Incubate at room temperature (RT).</li>
	<li>Once embryos begin breaking out of the chorion, remove all liquid from the embryo dish.</li>
	<li>Wash the embryos 2&#160;- 3 times with fresh E3 by adding approximately 20 mL of E3, gently swirling the E3 in the dish, and then decanting the fluid into a liquid waste receptacle.</li>
	<li>To euthanize the embryos before fixing, add&#160;2 mL of 0.2% tricaine to the E3.</li>
	<li>Transfer the embryos into a 5 mL glass vial using a plastic Pasteur pipette. Remove most of the tricaine/E3 solution using the pipette after the embryos have settled to the bottom of the vial.</li>
	<li>Fix 24 hpf embryos with 5 mL of 4% paraformaldehyde (PFA) for 2 - 4 h at RT, without agitation. 
	CAUTION: PFA is toxic and PFA solutions should be handled under a chemical hood while the researcher is wearing eye protection, gloves, and a lab coat. Preparation of the PFA fixative from the granulated PFA powder should be performed with exceptional care, as the granulated PFA tends to disperse through static charge. 
	NOTE: Embryos can fix in 4% PFA overnight at 4 &#176;C. Prepare 4% PFA by heating 1x phosphate buffered saline (PBS) to a boil while continually stirring the solution to completely dissolve the granulated PFA. Once the solution has cooled back down to RT, the 4% PFA is filter sterilized by passing it through a 0.45 &#181;m disposable system for vacuum filtration, then and aliquoted into 15 mL or 50 mL portions for storage at -20 &#176;C.</li>
	<li>Remove the 4% PFA and wash embryos twice with 1X phosphate buffered saline with 0.1% Tween-20 (PBST) at RT. Wash the embryos once with 5 mL of 100% methanol (MeOH) at RT. Add 5 mL of fresh 100% MeOH to the embryos and incubate at -20 &#176;C for at least 20 min. 
	NOTE: Embryos can be stored in 100% MeOH at -20&#176;C until ready for embryo preparation.</li>
</ol>
2. Embryo preparation and hybridization
<ol>
	<li>Rehydrate the embryos by washing once in 5 mL of 50% MeOH in 1x PBST for 5 min at RT. Continue rehydration by washing the embryos in 5 mL of 30% MeOH in 1x PBST for 5 min at RT. Wash the embryos in 5 mL of 1x PBST twice for 5 min each at RT.</li>
	<li>Permeabilize the embryos by incubating them in 5 mL of a solution of 1:1,000 proteinase K (pK) (10 mg/mL) in 1x PBST for 2 min at RT. 
	NOTE: Older embryos can be incubated longer than 2 min in the pK concentration stated above, however a concentration of 5 mg/mL and incubation time less than 2 min is suggested for embryos younger than 24 hpf.</li>
	<li>Remove pK solution and wash immediately in 5 mL of 1x PBST twice at RT. Fix the embryos in 5 mL of 4% PFA at RT for at least 20 min. 
	NOTE: Embryos can fix in 4% PFA overnight at 4 &#176;C.</li>
	<li>Remove PFA and wash twice in 5 mL of 1x PBST at RT.</li>
	<li>Transfer embryos in 5 mL of 1x PBST into flat bottom microcentrifuge tubes placed upright into a microcentrifuge rack at RT to facilitate interactions between each embryo and the subsequent hybridization solution.</li>
	<li>Wash the embryos twice in 1.5 mL of hybridization solution (Hyb+) at RT. Add 1.5 mL of fresh Hyb+ and incubate embryos at 70 &#176;C in a hybridization oven for 4-6 h.</li>
	<li>Replace the 1.5 mL of Hyb+ with a volume of probe solution (10 &#181;L RNA probe/500 &#181;L Hyb+) sufficient to fully cover the embryos. 
	NOTE: Synthesize antisense RNA probe using previously described methods39.</li>
	<li>Hybridize the probe overnight in the hybridization oven at 70 &#176;C with the individual tubes positioned upright in the microcentrifuge rack, without shaking.</li>
</ol>
3. Hot Washes and Blocking
NOTE: Hot washes are performed by putting the appropriate solution on the embryos and then incubating in the hybridization oven at 70 &#176;C. To keep the washing solutions at 70 &#176;C, place 50 mL tubes of each solution in the hybridization oven when the probe/Hyb+ mixture is added.
<ol>
	<li>Remove the probe. Wash the embryos twice at 70 &#176;C in 1.5 mL of 50% formamide/2x saline-sodium citrate (SSC) buffer for 20-30 min each. 
	NOTE: The probe can be stored in Hyb+ at -20 &#176;C and re-used at a later date.</li>
	<li>Wash the embryos once at 70 &#176;C in 1.5 mL of 2x SSC for 15 min. Wash the embryos twice at 70 &#176;C in 1.5 mL of 0.2X SSC for 20-30 min each. Replace 0.2x SSC with 1.5 mL of blocking reagent and incubate overnight at 4 &#176;C. 
	NOTE: It is possible to incubate blocking reagent at RT for 4 h instead of overnight.</li>
</ol>
4. Antibody Incubation and Maleic Acid Buffer Washes
NOTE: Keep embryos protected from ambient light.
<ol>
	<li>Replace the blocking reagent with 0.2 mL of anti-DIG-POD in blocking reagent at a ratio of 1:1,000 and incubate for 3 h under foil or another light-blocking cover at RT. 
	NOTE: Alternatively, the antibody can incubate overnight at 4 &#176;C.</li>
	<li>After 3 h, wash the embryos in 1.5 mL of maleic acid buffer 2-4 times for 10-15 min each at RT. Incubate the embryos overnight at 4 &#176;C in 1.5 mL of fresh maleic acid buffer. 
	NOTE: It is not necessary to incubate in maleic acid buffer overnight, but doing so decreases probe background.</li>
</ol>
5. Probe Detection and Removal of Peroxidase
NOTE: Keep embryos protected from ambient light.
<ol>
	<li>Remove maleic acid buffer and replace with 1.5 mL of 1x PBS. Wash embryos twice in 1.5 mL of 1x PBS for 5 min each at RT. Incubate embryos in 0.2 mL of Cy3 fluorescent staining solution for 60 min at RT. 
	NOTE: Incubation time may vary for different probes.</li>
	<li>Wash the embryos once with 1.5 mL of the following percent MeOH in 1X PBS for 10 min at RT: 30% MeOH, 50% MeOH, 75% MeOH and 100% MeOH.</li>
	<li>Incubate the embryos with 1.5 mL of 1% H2O2 in MeOH for 30 min at RT. Wash the embryos once with 1.5 mL of the following MeOH solutions in 1x PBS for 10 min at RT: 75% MeOH, 50% MeOH, and 30% MeOH. Wash the embryos twice for 5 min each at RT in 1.5 mL of 1x PBS. 
	NOTE: Embryos can be stored in PBS overnight at 4&#176;C.</li>
</ol>
6. Immunofluorescence
NOTE: Keep embryos protected from ambient light.
<ol>
	<li>Wash the embryos in 1.5 mL of ddH2O for 5 min at RT. Wash the embryos in 1.5 mL of pre-chilled acetone (kept at -20 &#176;C) for 7 min at RT. Wash the embryos once in 1.5 mL of ddH2O for 5 min at RT. Wash the embryos once in 1.5 mL of 1x PBST with 1% DMSO (PBDT) for 5 min at RT.</li>
	<li>Incubate the embryos in 1.5 mL of PBDT + 10% fetal bovine serum (FBS) on a rocker at RT for 2 h.</li>
	<li>Remove PBDT + FBS and replace with 1.5 mL of primary antibodies, anti-acetylated tubulin (produced from mouse) and anti-&#947;-tubulin (produced from rabbit), diluted together at 1:400 in PBDT + 1% FBS. Incubate the embryos overnight at 4 &#176;C. 
	NOTE: Incubation times may vary depending on the primary antibody. Testing of different time intervals may be necessary to optimize labeling.</li>
</ol>
7. Secondary Antibody
NOTE: Keep embryos protected from ambient light.
<ol>
	<li>To remove excess unbound antibody from the embryos, wash in 1.5 mL of PBDT + 1% FBS + 0.1 M NaCl for 1 min at RT on a rocker.</li>
	<li>Wash the embryos 5 times in 1.5 mL of PBDT + 1% FBS + 0.1 M NaCl for 30 min each on a rocker at RT. Wash the embryos in 1.5 mL of PBDT + 1% FBS for 30 min at RT on a rocker.</li>
	<li>Incubate the embryos overnight at 4 &#176;C in 200 &#181;L of the secondary antibodies, Alexa 488 goat anti-mouse IgG and Alexa 647 goat anti-rabbit IgG, diluted together in PBDT at 1:500.</li>
</ol>
8. DAPI Staining
NOTE: Keep embryos protected from ambient light.
<ol>
	<li>Wash the embryos quickly at RT in 1.5 mL of PBDT two times. Replace PBDT with 1.5 mL of DAPI, diluted 1:15000 in 1X PBST, and incubate 15 min at RT on a rocker.</li>
	<li>Wash the embryos in 1.5 mL of PDBT + 1% FBS + 0.1 M NaCl three times at RT for 15-20 min each. Store embryos in 1.5 mL of PBDT at 4 &#176;C until ready to mount and image.</li>
</ol>
9. Mounting and Imaging for Zebrafish Pronephros
<ol>
	<li>Lay embryo lateral on glass slide in a small volume, approximately 10 &#181;L, of PBDT.</li>
	<li>Squeeze the embryo behind the eyes with a pair of fine forceps to remove the head and some of the yolk ball.</li>
	<li>Use the fine forceps to gently scrape away the remaining yolk ball from the embryo body. Remove the dissociated yolk and extra liquid from the slide with a thin tissue.</li>
	<li>Add a drop of mounting media to the remaining tail of the embryo and position so the tail is laid laterally. Place a coverslip on top of the tail to flatten the sample, then place in a covered slide box.</li>
	<li>Image kidney cilia at 60X magnification on a confocal microscope using the following channels: DAPI in blue (laser set at 408.0 nm), FITC in green (laser set at 488.0 nm), Cy3 dye-labeled in red (laser set at 561.0 nm), and Alexa 680 in white (laser set at 637.0 nm).</li>
</ol>

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J., Sumanas, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-color+fluorescent+in+situ+hybridization+using+chromogenic+substrates+in+zebrafish.">Two-color fluorescent in situ hybridization using chromogenic substrates in zebrafish.</a> Bio Techniques. 57, 254-256 (2014).</li><li>Julich, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=beamter/deltaC+and+the+role+of+Notch+ligands+in+the+zebrafish+somite+segmentation,+hindbrain+neurogenesis+and+hypochord+differentiation.">beamter/deltaC and the role of Notch ligands in the zebrafish somite segmentation, hindbrain neurogenesis and hypochord differentiation.</a> Dev Biol. 286, 391-404 (2005).</li><li>Gross-Thebing, T., Paksa, A., Raz, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+high-resolution+detection+of+multiple+transcripts+combined+with+localization+of+proteins+in+whole-mount+embryos.">Simultaneous high-resolution detection of multiple transcripts combined with localization of proteins in whole-mount embryos.</a> BMC Biology. 12, (2014).</li><li>Cheng, C. N., Li, Y., Marra, A. N., Verdun, V., Wingert, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flat+Mount+Preparation+for+Observation+and+Analysis+of+Zebrafish+Embryo+Specimens+Stained+by+Whole+Mount+In+situ+Hybridization.">Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization.</a> J Vis Exp. (89), e51604 (2014).</li><li>Poureetezadi, S. J., Donahue, E. K., Wingert, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Manual+Small+Molecule+Screen+Approaching+High-throughput+Using+Zebrafish+Embryos.">A Manual Small Molecule Screen Approaching High-throughput Using Zebrafish Embryos.</a> J Vis Exp. (93), e52063 (2014).</li></ol>

The authors have nothing to disclose.

The protocol detailed above is optimized for labeling MCCs in the pronephros with odf3b transcripts and &#945;-tubulin in 24 hpf zebrafish embryos. For the best results, it is recommended to use freshly fixed and prepared embryos. Embryos that have been fixed and stored in either MeOH or Hyb+ at -20 &#176;C for longer than one week can be used, but the probability of unwanted background staining increases with the time that the embryos are kept in storage.
Modifications and troubleshooting of this technique are necessary to tailor this method for other suites of particular markers, e.g. other antisense riboprobes and other antibodies. Many methods for adjusting the parameters of steps in the process of whole mount in situ hybridization have been previously documented by our group and others31,34,36,38,39. Likewise, each protein antibody will require some troubleshooting in terms of staining times, and approximate ranges of dilutions and incubation times for each primary antibody are best estimated by evaluation of documented results28,35. For embryos younger than 24 hpf, pK treatment should be shorter than 2 min, where embryos older than 24 hpf should be treated with pK for longer than 2 min. Also, embryos older than 24 hpf should be bleached of pigment before pK treatment.
After staining with the fluorescent staining solution, it is critical to remove any remaining stain, antibody, and peroxidases by performing the series of methanol and hydrogen peroxide washes. The PBS washes immediately following are vital to remove excess methanol from the embryos. Importantly, before proceeding with the IF antibodies, we have found that the acetone and deionized water washes make the embryos less likely to adhere to one another and/or the centrifuge tubes. In our experience, antibodies for specific transcription factors and other genes, though they may work efficiently in Western blots, do not work well for IF in the zebrafish. However, highly conserved, and abundant proteins, such as &#945;-tubulin and &#946;-catenin, do work well in the zebrafish for IF16,37.
With FISH, it is possible to visualize RNA transcripts of genes that do not yet have specific antibodies in the zebrafish. By combining FISH with IF, as demonstrated by this protocol, co-localization of RNA transcripts and protein can be seen in vivo (Figure 2). The flexible nature of the protocol enables quick troubleshooting for the visualization of various RNA transcripts and proteins in numerous tissues and time points. When combined with the already respected traits of zebrafish embryos, this protocol of FISH + IF provides another tool to explore the expression of genes and proteins important to developmental pathway regulation.

Over the past several decades, the zebrafish (Danio rerio) has emerged as a prime model organism to study developmental biology. The embryos develop outside of the mother and are optically transparent. Also, the formation of vital organs such as the eye, kidney, and forebrain occurs rapidly, with structures formed by just 24 h post fertilization (hpf). Importantly, the zebrafish genome is highly conserved with mammals1,2,3. Additionally, zebrafish and mammalian organs have similar anatomy and physiology. The zebrafish embryonic kidney, or pronephros, demonstrates the value of the model system for examining gene function during early nephrogenesis and fate determination of conserved epithelial cell populations of the vertebrate nephron4,5,6,7,8,9,10. Similarly, the zebrafish embryo has become increasingly important in examining the ontogeny of MCCs11,12,13,14,15,16,17.
As their name suggests, MCCs are epithelial cells characterized by a bundle of motile cilia located on the apical surface17. In the zebrafish, MCCs function in fluid flow and are dispersed in a &#34;salt-and-pepper&#34; like fashion throughout the middle of each nephron of the pronephros by 24 hpf11,12,13,14,15,16,17. Although they have only been noted in a handful of human kidney disease cases18,19,20,21, MCCs are prevalent in other mammalian tissues such as the brain and trachea22,23,24, which poses a host of challenges for experimental design. Elegant studies in various vertebrate models including zebrafish have demonstrated a conserved pathway of MCC fate, with the Notch signaling pathway as an inhibitor of MCC development12,17,25,26,27. Therefore, the zebrafish pronephros provides an easily accessible model to study the genetic mechanisms of MCC development in vivo11,12,13,14,15,16,17.
Both transparency and easy genetic manipulation of zebrafish embryos have proven to be invaluable traits when studying the genetic and molecular pathways that regulate cell fate, tissue growth, and development of the early embryo1,2,3. As such, traditional techniques to visualize protein and gene transcripts, such as in situ hybridization and whole mount IF, have been applied to and optimized for the zebrafish16,28,29,30,31,32,33,34,35,36,37,38. By combining popular protocols for FISH and IF, it is possible to label and analyze MCCs in vivo16,28,37,38.

<table><tbody><tr><td>non-specific protease mixture solution</td><td>Roche</td><td>11459643001, pronase from Streptomyces griseus</td><td>Dilute in E3 &lt;strong&gt;without&lt;/strong&gt; methylene blue to make 50mg/mL stock solution; store at -20&amp;deg;C</td></tr><tr><td>E3 solution</td><td></td><td></td><td>Dilute 50X E3 (250 mM NaCl, 8.5 mM KCl, 16.5 mM CaCl2, 16.5 mM MgSO4 in distilled water) in distilled water; add 1 x 10-5 M methylene blue (sigma M9140) to inhibit contamination</td></tr><tr><td>tricaine (ethyl 3-aminobenzoate methanesulfonate)</td><td>Fluka</td><td>A5040-250G</td><td>To make a 0.2% stock solution, dissolve 1 g of tricaine powder in 10 mL Tris, pH 9.5 and distilled water up to 500 mL</td></tr><tr><td>embryo dishes</td><td>VWR</td><td>351029</td><td></td></tr><tr><td>5 mL glass vials</td><td>Wheaton</td><td>225012</td><td></td></tr><tr><td>disposable plastic Pasteur pippettes</td><td>VWR</td><td>414004-004</td><td></td></tr><tr><td>4% paraformaldehyde (PFA) solution</td><td>Electron Microscopy Services</td><td>19210</td><td>Dissolve 4% PFA in 1X PBS and bring to a boil under a fume hood. Cool and aliquot, then store at -20&amp;deg;C. Do not refreeze once thawed for use.</td></tr><tr><td>10X PBS</td><td>American Bioanalytical</td><td>AB11072</td><td>Dilute in distilled water to make a 1X stock</td></tr><tr><td>Tween-20 stock</td><td>American Bioanalytical</td><td>AB02038</td><td>Make a 0.1% Tween-20 stock by diluting in distilled water.</td></tr><tr><td>1X phosphate buffered saline with 0.1% Tween 20 (PBST)</td><td>Sigma</td><td>P9416</td><td>0.1% Tween-20 in 1X PBS</td></tr><tr><td>methanol (MeOH)</td><td>Sigma</td><td>34860-4L</td><td></td></tr><tr><td>proteinase K</td><td>Roche</td><td>3115879001</td><td>Dissolve in distilled water to make a 10 mg/mL stock; aliquot and store at -20&amp;deg;C</td></tr><tr><td>flat bottom microcentrifuge tubes</td><td>VWR</td><td>87003-300; 87003-298</td><td></td></tr><tr><td>formamide</td><td>American Bioanalytical</td><td>AB00600</td><td>store at -20&amp;deg;C</td></tr><tr><td>hybridization solution (HYB+)</td><td></td><td></td><td>50% formamide, 5X SSC, 0.1% Tween-20, 5 mg/mL yeast torula RNA, 50 &amp;mu;g/&amp;mu;L heparin; store at -20&amp;deg;C</td></tr><tr><td>hybridization oven</td><td>Fisher Scientific</td><td>13-247-10Q</td><td></td></tr><tr><td>20X saline-sodium citrate (SSC) buffer solution</td><td>American Bioanalytical</td><td>AB13156</td><td>dilute in distilled water to make 2X and 0.2X stocks</td></tr><tr><td>blocking reagent solution</td><td>Roche</td><td>11096176001</td><td>dilute in maleic acid buffer to make a 10% stock solution; store at 4&amp;deg;C</td></tr><tr><td>maleic acid buffer solution</td><td>Sigma</td><td>M0375 and S7653</td><td>150mM maleic acid, 100mM NaCl (pH 7.5)</td></tr><tr><td>anti-Digoxigenin-POD, Fab fragments</td><td>Roche</td><td>11207739910</td><td>store at 4&amp;deg;C</td></tr><tr><td>Cy3 fluorescent staining solution</td><td>PerkinElmer, Inc.</td><td>NEL744001KT, TSA Plus Cyanin3 system</td><td>store at 4&amp;deg;C; prepare staining solution fresh by making a 1:50 dilution of TSA reagent (dissolved in 60 uL of DMSO) in the kit buffer</td></tr><tr><td>Hydrogen peroxide (H2O2)</td><td>Sigma</td><td>H1009-500mL</td><td>store at 4&amp;deg;C</td></tr><tr><td>molecular grade distilled water (ddH2O)</td><td>Mediatech</td><td>25-055-CM</td><td></td></tr><tr><td>acetone</td><td>American Bioanalytical</td><td>AB00636-01000</td><td>store an aliquot at -20&amp;deg;C</td></tr><tr><td>DMSO</td><td>American Bioanalytical</td><td>AB00435-01000</td><td></td></tr><tr><td>fetal bovine serum (FBS)</td><td>Gibco</td><td>10438-034</td><td>aliquot and store at -20&amp;deg;C</td></tr><tr><td>monoclonal anti-acetylated tubulin clone 6-11B-1</td><td>Sigma-Aldrich</td><td>T6793</td><td>aliquot and store at -20&amp;deg;C</td></tr><tr><td>anti-&amp;gamma;-tubulin anitbody produced in rabbit</td><td>Sigma-Aldrich</td><td>T5192</td><td>aliquot and store at -20&amp;deg;C</td></tr><tr><td>odf3b cDNA clone MGC:63985</td><td>OpenBiosystems</td><td>IMAGE:6792478</td><td>store bacterial glycerol stock at -80&amp;deg;C</td></tr><tr><td>NaCL</td><td>American Bioanalytical</td><td>AB01915-05000</td><td></td></tr><tr><td>Alexa flour 647 goat anti-rabbit IgG</td><td>Life Technologies</td><td>A21245</td><td>store at 4&amp;deg;C; protect from light</td></tr><tr><td>Alexa fluor 488 goat anti-mouse IgG</td><td>Life Technologies</td><td>A11029</td><td>store at 4&amp;deg;C; protect from light</td></tr><tr><td>DAPI</td><td>Life Technologies</td><td>D1306</td><td>aliquot and store at -20&amp;deg;C</td></tr><tr><td>glass slide</td><td>Thermo-Fisher</td><td>4445</td><td></td></tr><tr><td>glass coverslip</td><td>Thermo-Fisher</td><td>12-540A</td><td>18 x 18 mm</td></tr><tr><td>fine forceps</td><td>Roboz</td><td>RS-1050</td><td>Dumont Tweezers Pattern #55</td></tr><tr><td>mounting media</td><td>Polysciences, Inc.</td><td>18606, Aqua-Poly/Mount</td><td>store at 4&amp;deg;C</td></tr><tr><td>confocal microscope and associated software</td><td></td><td></td><td>We use a Nikon C2plus Confocal Microscope with NIS-elements AR software</td></tr><tr><td>rocker</td><td>Bio-Rad</td><td>1660710EDU</td><td></td></tr></tbody></table>

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.

Wild-type zebrafish embryos were fixed at 24 hpf and immediately prepared as described above. Figure 1 depicts an experimental workflow along with selected illustrated stages. The workflow outlined in Steps 1-8 encompasses the processes of sample procurement, fixation, and manipulation of the fixed tissue to label endogenous transcripts with antisense riboprobes followed by immunofluorescence to label protein(s) of interest. Step 9 in the workflow refers to the mounting and imaging techniques specifically designed to optimize visualization of the zebrafish embryonic trunk. In the drawings that accompany Step 9, we illustrate the method of manipulation for positioning the tissue on a glass slide. In this step, the embryo head and yolk ball are removed, leaving the tail to be laterally positioned between a glass slide and coverslip. Removal of the yolk ball and head is suggested for the best imaging of the resident MCC population in the pronephros, as the yolk ball auto-fluoresces and obstructs the mounting process.
Representative images obtained using a confocal microscope are provided in Figure 2, which shows the same wild-type embryo at 60X magnification and a digital zoom at the same magnification. The top panels provide images of each individual channel, while the bottom two panels are the composite overlay of these imaging data. The white box (in the left column image) outlines the area that is the focus of the digital zoom (the right column image). Highlighted in the final panel of the digital zoom, nuclei were labeled in DAPI, and MCCs were detected based on an antisense riboprobe designed to recognize odf3b, where antibodies to &#947;-tubulin denote the basal bodies and &#945;-tubulin denotes the cilia. In the digital zoom of the composite overlay, the yellow dashed circle indicates the perimeter of an MCC, which was identified due to the possession of odf3b transcripts, multiple basal bodies, and cilia. The adjacent mono-ciliated cell, labeled with a yellow dotted circle, was identified based on the phenotype of possessing a single basal body and a single cilium.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56261/56261fig1.jpg" /> 
Figure 1: Schematic of experimental flowchart. This flowchart shows an experimental workflow, accompanied by illustrations of critical stages in which the snowflake indicates cold incubation, the three curved lines represent heat, and the clock represents long incubation times. The workflow can be performed in a minimum of 6 days (see day number in upper right corner of each stage in the process), though some steps can be performed over longer time periods, as noted in the protocol. A detailed depiction of the mounting process is drawn out, demonstrating the final mounted embryo tail between the glass slide and coverslip. A black dashed box outlines the area that is imaged at 60X magnification. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/56261/56261fig1large.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/56261/56261fig2.jpg" /> 
Figure 2: Representative results for visualizing MCCs of the zebrafish pronephros. Maximum image projections of a 24 hpf wild-type zebrafish embryo at 60X magnification as well as a digital zoom on the confocal at 60X magnification of the same embryo. The white boxes indicate the area focused on for the zoom. Individual stains of DAPI (nuclei), odf3b (MCCs), &#947;-tubulin (basal bodies), and &#945;-tubulin (cilia) are labeled and then merged together in the bottom two panels. In the digital zoom, we provide approximations of the cell locations as follows: an MCC is outlined by the dashed yellow circle, and the dotted yellow circle outlines a mono-ciliated cell. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/56261/56261fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence at JoVE.c

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.

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 ...

visualizing-multiciliated-cells-zebrafish-through-combined-protocol

Universidad San Sebastian

Research

JoVE Journal

Developmental Biology

8.1K Views.  University of Notre Dame. Cilia development is vital to proper organogenesis. This protocol describes an optimized method to label and visualize ciliated cells of the zebrafish.

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

Labeling and Imaging Cells in the Zebrafish Hindbrain

Key to understanding the morphogenetic processes that shape the early vertebrate embryo is the ability to image cells at high resolution. In zebrafish embryos, injection of plasmid DNA results in mosaic expression, allowing for the visualization of single cells or small clusters of cells 1 . We describe how injection of plasmid DNA encoding membrane-targeted Green Fluorescent Protein (mGFP) under the control of a ubiquitous promoter can be used for imaging cells undergoing neurulation. Central to this protocol is the methodology for imaging labeled cells at high resolution in sections and also in real time. This protocol entails the injection of mGFP DNA into young zebrafish embryos. Embryos are then processed for vibratome sectioning, antibody labeling and imaging with a confocal microscope. Alternatively, live embryos expressing mGFP can be imaged using time-lapse confocal microscopy. We have previously used this straightforward approach to analyze the cellular behaviors that drive neural tube formation in the hindbrain region of zebrafish embryos 2. The fixed preparations allowed for unprecedented visualization of cell shapes and organization in the neural tube while live imaging complemented this approach enabling a better understanding of the cellular dynamics that take place during neurulation.

Key to understanding the morphogenetic processes that shape the early embryo is the ability to image cells at high resolution. We describe here a technique for labeling single cells or small clusters of cells in whole zebrafish embryos with membrane-targeted Green Fluorescent Protein.

Key to understanding the morphogenetic processes that shape the early vertebrate embryo is the ability to image cells at high resolution. In zebrafish ...

Neuroscience

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

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 the Interrenal Steroidogenic Tissue and Its Vascular Microenvironment in Zebrafish

This protocol introduces how to detect differentiated interrenal steroidogenic cells through a simple whole-mount enzymatic activity assay. Identifying differentiated steroidogenic tissues through chromogenic histochemical staining of 3-&#946;-Hydroxysteroid dehydrogenase /&#916;5-4 isomerase (3&#946;-Hsd) activity-positive cells is critical for monitoring the morphology and differentiation of adrenocortical and interrenal tissues in mammals and teleosts, respectively. In the zebrafish model, the optical transparency and tissue permeability of the developing embryos and larvae allow for whole-mount staining of 3&#946;-Hsd activity. This staining protocol, as performed on transgenic fluorescent reporter lines marking the developing pronephric and endothelial cells, enables the detection of the steroidogenic interrenal tissue in addition to the kidney and neighboring vasculature. In combination with vibratome sectioning, immunohistochemistry, and confocal microscopy, we can visualize and assay the vascular microenvironment of interrenal steroidogenic tissues. The 3&#946;-Hsd activity assay is essential for studying the cell biology of the zebrafish interrenal gland because to date, no suitable antibody is available for labeling zebrafish steroidogenic cells. Furthermore, this assay is rapid and simple, thus providing a powerful tool for mutant screens targeting adrenal (interrenal) genetic disorders as well as for determining disruption effects of chemicals on steroidogenesis in pharmaceutical or toxicological studies.

The interrenal gland in zebrafish is the teleostean counterpart of the mammalian adrenal gland. This protocol introduces how to perform the 3-&#946;-hydroxysteroid dehydrogenase (&#916;5-4 isomerase; 3&#946;-Hsd) enzymatic activity assay, which detects differentiated steroidogenic cells in the developing zebrafish.

This protocol introduces how to detect differentiated interrenal steroidogenic cells through a simple whole-mount enzymatic activity assay. Identifying ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Biology

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 ...

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling processes of excitable neuronal and muscular cells and many other biological processes, such as embryonic developmental patterning. However, there is a need for in vivo electrical activity monitoring in vertebrate embryogenesis. The advances of genetically encoded fluorescent voltage indicators (GEVIs) have made it possible to provide a solution for this challenge. Here, we describe how to create a transgenic voltage indicator zebrafish using the established voltage indicator, ASAP1 (Accelerated Sensor of Action Potentials 1), as an example. The Tol2 kit and a ubiquitous zebrafish promoter, ubi, were chosen in this study. We also explain&#160;the processes of Gateway site-specific cloning, Tol2 transposon-based zebrafish transgenesis, and the imaging process for early-stage fish embryos and fish tumors using regular epifluorescent microscopes. Using this fish line, we found that there are cellular electric voltage changes during zebrafish embryogenesis, and fish larval movement. Furthermore, it was observed that in a few zebrafish malignant peripheral nerve sheath tumors, the tumor cells were generally polarized compared to the surrounding normal tissues.

Here, we show the process of creating a cellular electric voltage reporter zebrafish line to visualize embryonic development, movement, and fish tumor cells in vivo.

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling ...

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. ...

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 ...

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

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Labeling and Imaging Cells in the Zebrafish Hindbrain

Imaging Subcellular Structures in the Living Zebrafish Embryo

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

Visualizing the Interrenal Steroidogenic Tissue and Its Vascular Microenvironment in Zebrafish

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

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

In Situ Hybridization Combined with Immunohistochemistry in Cryosectioned Zebrafish Embryos