We would like to acknowledge our funding source: NIH R01 EY05661 to J.E.H, Ines Gehring for assisting with generating the aqp0a/b double mutants and zebrafish husbandry, Dr Daniel Dranow for discussions leading to generation of the transgenics, Dr Bruce Blumberg and Dr Ken Cho&#8217;s labs for use of their dissecting microscopes, and Dr Megan Smith for help with statistical analysis.

The animal protocols used in this study adhere to the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research and have been approved by the Institutional Animal Care and Use Committee (IACUC) of University of California, Irvine.

1. Zebrafish Husbandry and Euthanasia
<ol>
	<li>Raise and maintain zebrafish (AB strain) under standard laboratory conditions12. Raise embryos in embryo medium (EM)12. Add 0.003% PTU to EM from 20-24 h postfertilization (before the prim-5 stage) to prevent pigment formation in embryos12 used for imaging at embryonic stages9. Raise larvae from 6-30 days postfertilization (dpf) on a diet of live rotifers until 14 dpf, and live artemia after 14 dpf12. 
	CAUTION: PTU is very toxic</li>
	<li>Anesthetize fish using tricaine until non-responsive to touch.
	<ol>
		<li>Prepare tricaine stock by combining 400 mg of 3-amino benzoic acidethylester, with 2.1 mL of 1 M Tris (pH 9), in 100 mL of ddH2O. Adjust to pH 7.0 and store at -20 &#176;C. 
		CAUTION: Tricaine is toxic.</li>
		<li>Dilute 4.2 mL of tricaine stock in 100 mL of tank water to anesthetize larvae/adults or in EM to anesthetize embryos (final concentration of tricaine at 0.0165% w/v).</li>
	</ol>
	</li>
</ol>
2. Fixation of Embryos and Larvae
NOTE: Immunohistochemical protocols were adapted from previously published materials7.
<ol>
	<li>Fix dechorionated embryos or larvae up to 2 weeks postfertilization in 4% (v/v) paraformaldehyde (PFA) in phosphate buffered saline (PBS) overnight at 4 &#176;C on a rocker. 
	CAUTION: PFA is combustible and is carcinogenic.</li>
	<li>Wash embryos three times for 10 min in PBS, and permeabilize in PBS with 10% Triton and 1% DMSO (PBS-T) overnight at 4 &#176;C. 
	CAUTION: DMSO is toxic, is harmful by ingestion or skin absorption, carries hazardous materials through skin, and is combustible. Triton is toxic, corrosive and is hazardous to aquatic environments.</li>
</ol>
3. Dissection of Larval and Adult Zebrafish Lenses
<ol>
	<li>Anesthetize fish from 6 dpf larvae to adulthood with tricaine until non-responsive to touch but still showing a strong heartbeat. Measure fish standard length as per Schilling (2002) for staging13.</li>
	<li>Immediately excise eyes using micro-dissection scissors and place into a dissection dish in PBS (Figure 2 shown for adult eyes).
	<ol>
		<li>Make a custom 35 mm dish filled with silicone for lens dissections. Once silicone has set, excise a divot around 2-3 mm in diameter and 0.5 mm deep for immobilizing adult eyes/lenses during dissection.</li>
	</ol>
	</li>
	<li>Dissection of adult zebrafish lenses
	<ol>
		<li>Place an adult eye into the dish divot posterior side up filled with PBS. Immobilize the eye by inserting forceps at &#60;45&#176; angle through the optic disc. Be careful not to nick or compress the lens. Make two or three radial incisions through retina and sclera from the optic disc to the ciliary zone with dissection scissors.</li>
		<li>Peel back the retina and sclera like flower petals and invert the eye, cornea side up. Immobilize the lens indirectly via manipulation of the sclera and cornea with the flat side of the scissors, while pulling away the retina and attached tissues with forceps. Carefully trim excess tissue from the lens. 
		NOTE: It is vital to dissect carefully to obtain consistently healthy lenses.</li>
	</ol>
	</li>
	<li>Dissection of larval zebrafish lenses
	<ol>
		<li>Place a larval eye posterior side up onto the flat part of the silicone dish filled with PBS and use a sharpened tungsten needle to make radial cuts through the retina and sclera while immobilizing the eye with another tungsten needle or forceps. 
		NOTE: Be careful not to damage the lens.
		<ol>
			<li>Sharpen the tip of a 2 cm length of 0.1 mm tungsten wire electrolytically by suspending the wire tip into 10% (w/v) NaOH and applying a low voltage alternating-current14. Secure the needle into a Pasteur pipette by melting the glass end using a Bunsen burner. 
			NOTE: Alternative fine dissection tools can also be used. 
			CAUTION: NaOH is corrosive.</li>
		</ol>
		</li>
		<li>Gently scoop out the lens from the dissociated eye with a blunt side of the needle, and carefully pull away attached tissue.</li>
	</ol>
	</li>
</ol>
4. Fixation of Dissected Lenses
<ol>
	<li>Immediately fix dissected lenses in 1.5% (v/v) PFA in PBS for 24 h at room temperature (RT). Wash lenses three times in PBS for 10 min each.</li>
	<li>Permeabilize lenses for whole mount analysis or cryoprotect for cryosectioning (see section 8).
	<ol>
		<li>Permeabilize lenses in PBS-T overnight at 4 &#176;C.</li>
	</ol>
	</li>
</ol>
5. Lens Immunohistochemistry
<ol>
	<li>Label membranes of fixed embryo/larval or adult lenses by incubation in Phalloidin- Alexa Fluor 546 (1:200) and cell nuclei with DAPI (1:1,000 of 5 mg/mL stock) in PBS-T overnight at 4 &#176;C. 
	CAUTION: DAPI is a skin and eye irritant.</li>
	<li>Wash three times with PBS-T for 10 min each. Clear tissue by incubation in 30%, 50% and 70% glycerol in PBS-T (v/v) for at least 1 h at RT each, or overnight at 4 &#176;C.</li>
	<li>Mount fish in 70% glycerol in PBS-T (v/v) onto glass bottom 35 mm microwell dishes with eyes as flat and straight against the cover slip as possible. For younger fish, a slight anterior-lateral tilt may help to orient the lens suture to be parallel to the coverslip.</li>
</ol>
6. Analysis of Zebrafish Anterior Lens Sutures Using a Transgenic Line In Vivo
<ol>
	<li>Generate Tg(&#946;B1cry:mAppleCAAX) lines using the Tol2 kit15. The Tol2 transposable element system15 enables stable integration of the construct where mApple is driven by 300 bp of the human &#946;B1-crystallin promoter16 tethered to the membrane by the CAAX sequence resulting in expression specifically in lens fiber cell membranes. 
	NOTE: This construct (ID:122451) is available for purchase.
	<ol>
		<li>On the morning of injection, prepare the injection mixture and keep on ice. To reach a final volume of 10 &#181;L of containing RNase- and DNase-free H2O, add: 1.5 &#181;L of hu&#946;B1cry:mAppleCAAX DNA construct at 50 ng/&#181;L - [0.375-0.75 pg/final injection], 1.5 &#181;L of Tol2 transposase mRNA at 300 ng/&#181;L (Tol2 kit)15 [15-30 pg/final injection], and 1 &#181;L of phenol red indicator at 1% w/v [1 x10-5% (w/v)/final injection].</li>
		<li>Inject 50-100 pL of injection mixture into 1-cell stage embryos12.</li>
	</ol>
	</li>
	<li>Measure transgenesis efficiency in F0 injected embryos by measuring frequency of embryos with mApple positive lenses at 3 dpf. 
	NOTE: Expect to have &#62;80% transgenesis efficiency using this method. F0 mosaics allow detailed analysis of membrane morphologies of individual cells. Varying levels of mosaicism allows one to image the morphologies of single or small groups of cells, while more global lens expression allows analysis of multicellular structures like lens sutures in vivo.</li>
	<li>Outcross F0 mosaic fish to generate stable lines, which label all fiber cell membranes. F0 founders with the transgene integrated into the germline will generate offspring with stable integrations that can be maintained as transgenic lines.</li>
</ol>
7. Analysis of Zebrafish Anterior Lens Sutures Using a Transgenic Line In Vivo
<ol>
	<li>Anesthetize 3 dpf or adult mosaic or stable transgenic fish and mount in 1% low melt agarose (LMA) with tricaine with the eye flat against the cover slip of glass bottom microwell dishes.
	<ol>
		<li>Dissolve 1 g of LMA in 100 mL of EM (without methylene blue) to make 1% LMA. Store long term as 15 mL stocks at 4 &#176;C. Microwave to dissolve stock and store at 42 &#176;C until each tube is used.</li>
	</ol>
	</li>
	<li>Cover fish up to 6 dpf with EM with tricaine, or with tank water with tricaine for adults when LMA is set.
	<ol>
		<li>Mix 5 mL of EM without methylene blue or tank water with 250 &#181;L of tricaine stock and 7 &#181;L of PTU if imaging PTU treated fish.</li>
	</ol>
	</li>
</ol>
8. Lens Cryosectioning and Immunohistochemistry
<ol>
	<li>Cryoprotect fixed embryos or adult lenses by placing into 10% sucrose in PBS (w/v), 20% sucrose for 1 h at RT each (or overnight at 4 &#176;C) and overnight in 30% sucrose at 4 &#176;C.</li>
	<li>Embed tissue in OCT in base molds, and then freeze onto chucks with OCT. Cryosection at 12-14 &#181;m and collect onto a Superfrost/Plus microscope slides. Warm sections onto slide for 10 min on a slide warmer prewarmed to ~35 &#176;C.</li>
	<li>Wash sections three times with PBS for 10 min each. Label sections with Phalloidin-Alexa Flour 546 (1:200) with DAPI (1:1,000) or WGA-Alexa Flour 594 (1:200), followed by three PBS washes. Mount with anti-fade mounting medium, and seal coverslip with nail polish.</li>
</ol>
9. Imaging
<ol>
	<li>Acquire images with a confocal microscope. Acquire z-stacks or optical slices using a 60x N/A 1.2 water-immersion objective, or similar, at specific regions from the anterior to posterior lens pole. Use the following acquisition settings: 1.2 airy disc using the 561 nm laser, Texas red filter for phalloidin-Alexa Flour 546 or mApple, or the 405 laser with the UV filter for DAPI.</li>
	<li>Correct z-intensity laser power to counteract signal loss with depth into the lens. This allows for a strong signal at the posterior pole of fixed and live 3 dpf embryos, and strong signal in equatorial optical slices in adult fish lenses.</li>
	<li>Compile and view images using image processing software.</li>
</ol>
10. Measurement of Lens Nucleus Localization in the Anterior-posterior Axis
<ol>
	<li>Orient freshly excised lenses axially in PBS in a 35 mm dish with a coverglass bottom, with poles and sutures oriented parallel to the plane of focus. Look for a difference in the refractive index, which usually occurs at the interface of the lens cortex and lens nucleus to identify the lens nucleus. 
	NOTE: Healthy wild type adult lenses are extremely transparent making it difficult to see the lens sutures and nucleus, or to determine lens orientation. In this case, a slight nick with forceps to the lens capsule causes minor damage, which makes the sutures and lens nucleus apparent in about 10 min helping to orient the lens for this measurement. However, the lenses become unusable for downstream applications as they are now damaged.</li>
	<li>Take images of lenses with the lens nucleus in focus under bright field illumination with or without DIC optics using a dissection microscope with a camera attached. Image a micrometer under the same magnification for calibration.
	<ol>
		<li>Click on the live view button, adjust the exposure settings to visualize the lens nucleus and lens periphery, and take an image by clicking the snap shot button. Save as a tif file.</li>
		<li>Use image processing software to calibrate the acquired lens images. Select the straight-line tool and draw a line of known length on the micrometer image, click analyze | set scale. Enter known distance, units and select &#39;global&#39; calibration, and click ok.</li>
	</ol>
	</li>
	<li>Measure the distance of the center of the lens nucleus to the anterior pole (a - r).
	<ol>
		<li>Use the straight-line tool in image processing software to draw a line across the imaged center of the lens nucleus in an axial orientation. Take the center of this line as the center of the lens nucleus.</li>
		<li>Draw another line from this point to the anterior pole of the lens and select &#39;measure&#39; in the &#39;analyze&#39; menu to measure the distance (a - r), where a is the radius of the lens and r is the distance from the center of the lens to the center of the nucleus.</li>
		<li>Draw another line from the anterior to the posterior poles and measure this distance as the lens diameter (2a). Copy these lengths from the &#39;results&#39; window and transfer to a spreadsheet or statistics program. 
		NOTE: It is easier to precisely measure from the center of the nucleus to the anterior pole, than measuring from the center of the lens nucleus to the center of the lens. This measurement is converted by the formula in step 10.3.5&#160;to report the distance of the center of the lens nucleus to the center of the lens. Always use the adult nucleus for measurements, even if the embryonic nucleus is evident.</li>
		<li>Divide the diameter by 2, to calculate the lens radius, a.</li>
		<li>Calculate r/a, as <img alt="Equation 1" src="/files/ftp_upload/59528/59528eq1.jpg" style="opacity: 0.9;" />, which is the normalized localization of the lens nucleus with respect to the lens radius. 
		NOTE: For a nucleus placed closer to the anterior pole, r/a&#160;will be &#62;0.0, while a centrally localized nucleus will have an&#160;r/a of 0.0.</li>
	</ol>
	</li>
	<li>Graph the log of the normalized axial nucleus measurement as a function of the standard length to determine changes in lens nucleus localization during zebrafish development.</li>
</ol>

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<li>Chee, K. N., Vorontsova, I., Lim, J. C., Kistler, J., Donaldson, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Expression+of+the+sodium+potassium+chloride+cotransporter+%28NKCC1%29+and+sodium+chloride+cotransporter+%28NCC%29+and+their+effects+on+rat+lens+transparency%5BTitle%5D)+AND+%22Molecular+Vision%22%5BJournal%5D)">Expression of the sodium potassium chloride cotransporter (NKCC1) and sodium chloride cotransporter (NCC) and their effects on rat lens transparency.</a> Molecular Vision. 16, 800-812 (2010).</li>
<li>Hayes, J. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Integrin+%CE%B15%2Ffibronectin1+and+focal+adhesion+kinase+are+required+for+lens+fiber+morphogenesis+in+zebrafish%5BTitle%5D)+AND+%22Molecular+Biology+of+the+Cell%22%5BJournal%5D)">Integrin α5/fibronectin1 and focal adhesion kinase are required for lens fiber morphogenesis in zebrafish.</a> Molecular Biology of the Cell. 23 (24), 4725-4738 (2012).</li>
<li>Imai, F., Yoshizawa, A., Fujimori-Tonou, N., Kawakami, K., Masai, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+ubiquitin+proteasome+system+is+required+for+cell+proliferation+of+the+lens+epithelium+and+for+differentiation+of+lens+fiber+cells+in+zebrafish%5BTitle%5D)+AND+%22Development%22%5BJournal%5D)">The ubiquitin proteasome system is required for cell proliferation of the lens epithelium and for differentiation of lens fiber cells in zebrafish.</a> Development. 137 (19), 3257-3268 (2010).</li>
<li>Biswas, S. K., Lee, J. E., Brako, L., Jiang, J. X., Lo, W. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Gap+junctions+are+selectively+associated+with+interlocking+ball-and-sockets+but+not+protrusions+in+the+lens%5BTitle%5D)+AND+%22Molecular+Vision%22%5BJournal%5D)">Gap junctions are selectively associated with interlocking ball-and-sockets but not protrusions in the lens.</a> Molecular Vision. 16, 2328-2341 (2010).</li>
</ol>

We have no disclosures.

Analysis of zebrafish lens morphology is the initial step in understanding phenotypes in mutants, or the effects of pharmacological interventions aimed at studying biology of the ocular lens. We outline methods to analyze lens sutures, cortical fiber cell morphology and aspects of the lens nucleus. These approaches are a combination of in vitro and in vivo (compared in Table 1). The in vitro methods allow for greater detail of the outer cortical cell morphology, as well as acce...

The zebrafish is an excellent model for studying lens development and physiology due to the anatomical similarities to mammalian lenses, relative ease of genetic and pharmacological manipulation, speed of embryonic eye development, small size and transparency at early stages allowing for in vivo imaging. The methods described here were developed to analyze zebrafish lens morphology at embryonic and adult stages with a focus on sutural integrity, cortical membrane morphology in vitro and in vivo, and location of lens nuclear position along the anterior-posterior axis ex vivo. These methods serve as a starting point for functional studies of lens development and physiology, as well as reverse genetic screens for lens phenotypes in zebrafish.
Imaging zebrafish lens morphology
Aquaporin 0 (AQP0) is the most abundant lens membrane protein and is critical for both, lens development and clarity, due to multiple essential functions in mammals. Zebrafish have two Aqp0s (Aqp0a and Aqp0b) and we have developed methods to analyze loss of their functions in both embryonic and adult lenses. Our studies reveal that aqp0a-/- mutants develop anterior polar opacity due to instability of the anterior suture, and aqp0a/b double mutants develop nuclear opacity1. AQP0 has been shown to play roles in water transport2, adhesion3,4, cytoskeletal anchoring5 and generation of the refractive index gradient6, but these studies have largely been performed in vitro. The zebrafish provides a unique opportunity to study how loss of function, or perturbed function of Aqp0a or Aqp0b would affect morphology and function in a living lens. To assess lens cell morphology and sutural integrity during development, we modified existing in vitro immunohistochemical methods7 for use in embryonic and adult lenses, and generated transgenics to monitor this process in vivo.
Immunohistochemical analysis of plasma membrane structure and sutural integrity was performed in whole fixed embryos and adult lenses. Zebrafish lenses are extremely small (lens diameter is ~100 &#181;m in embryos and up to 1 mm in adults) compared with their mammalian counterparts and have point sutures8, which are infrequently captured in cryosections. Thus, whole lenses are essential for analyzing sutural integrity. For in vivo analysis of anterior suture formation, and imaging of precise lens cell architecture, we generated transgenics expressing mApple specifically labeling lens membranes.
Advantages of live imaging of lens membrane transgenics include: 1) avoiding fixation artifacts, 2) studying dynamic morphological changes in time-lapse movies, and 3) enabling longitudinal studies in which earlier events can be correlated with later phenotypes. Pigmentation of the iris normally prevents clear imaging of the lens periphery. Addition of 1-phenyl 2-thiourea (PTU) before the primordia-5 (prim-5) stage9 prevents melanogenesis and eye pigmentation up to around 4 days postfertilization (dpf). However, after 4 dpf, the lens periphery is obscured in vivo, particularly at older stages. Furthermore, the density of the lens itself obscures imaging of its posterior pole. Therefore, to study morphology of the lens periphery, or the posterior suture, after 4 dpf, lenses need to be excised and fixed.
Transgenic zebrafish lines have been used to analyze embryonic lens membrane structure in vivo10. The Q01 transgenic expresses a cyan fluorescent protein fused to a membrane targeting sequence, Gap43, driven by the EF1&#945; promoter and a hexamer of the DF4 pax6 enhancer element ubiquitously in lens fiber cells11. Q01 does have extra-lenticular expression, including amacrine cells in the retina, which adds background signal if the primary interest is the lens. We developed a novel transgenic line that expresses a membrane-tethered mApple specifically in the lens, with the aim of avoiding any extra-lenticular signal.
Lens nucleus localization
We discovered that the lens nucleus moves from an initial anterior location in larval zebrafish to a central location in the anterior-posterior axis in adult lenses. This shift in the position of the high refractive index lens nucleus is thought to be a requirement for normal development of zebrafish lens optics1. Our methods allow quantification of lens nuclear centralization in relation to the lens radius. Using this method, we showed that Aqp0a is required for lens nuclear centralization1, and this simple method can be applied to other studies of the development and physiology of the lens and its optical properties in the zebrafish model.

<table><tbody><tr><td>1-phenyl-2-thiourea (PTU)</td><td>Sigma</td><td>P7629</td><td>CAUTION &amp;ndash; very toxic</td></tr><tr><td>4% Paraformaldehyde aqueous solution</td><td>Electron Microscopy Sciences</td><td>RT 157-4</td><td>CAUTION &amp;ndash; health hazard, combustible</td></tr><tr><td>Confocal microscope</td><td>Nikon</td><td>Eclipse Ti-E</td><td></td></tr><tr><td>Cryostat</td><td>Leica</td><td>CM3050S</td><td>Objective and chamber temperature set to -21 &amp;deg;C</td></tr><tr><td>DAPI</td><td>Invitrogen</td><td>D1306</td><td>CAUTION &amp;ndash; irritating to eyes and skin</td></tr><tr><td>Dimethyl Sulfoxide (DMSO)</td><td>Fisher Scientific</td><td>D128</td><td>CAUTION &amp;ndash; combustible, penetrates skin&amp;nbsp;</td></tr><tr><td>Disposable base mold</td><td>VWR Scientific</td><td>15154-631</td><td></td></tr><tr><td>Disposable Pasteur glass pipets</td><td>Fisherbrand</td><td>13-678-20A</td><td></td></tr><tr><td>Dumont #5 forceps</td><td>Dumont &amp;amp; Fils</td><td></td><td>Keep forceps sharpened</td></tr><tr><td>Ethyl 3-aminobenzoate methanesulfonate salt (Tricaine)</td><td>Sigma-Aldrich</td><td>A5040</td><td>CAUTION - toxic</td></tr><tr><td>Glass bottom microwell dish (35mm Petri dish, 14 mm microwell, #1.5 coverglass)</td><td>MatTek Corporation</td><td>P35G-1.5-14-C</td><td></td></tr><tr><td>Glycerol</td><td>Sigma</td><td>G2025</td><td></td></tr><tr><td>hu&amp;beta;B1cry:mAppleCAAX DNA construct</td><td>Addgene</td><td>ID:122451</td><td></td></tr><tr><td>ImageJ</td><td>Wayne Rasband, NIH</td><td>v1.51n</td><td></td></tr><tr><td>Low Melt Agarose (LMA)</td><td>Apex</td><td>902-36-6</td><td></td></tr><tr><td>NIS-Elements</td><td>Nikon</td><td>V 4.5</td><td></td></tr><tr><td>NIS-Elements AR software</td><td>Nikon</td><td></td><td></td></tr><tr><td>Olympus&amp;nbsp; with a model 2.1.1.183 controller</td><td>Olympus Corp</td><td>DP70</td><td></td></tr><tr><td>Olympus microscope&amp;nbsp;</td><td>Olympus Corp</td><td>SZX12</td><td></td></tr><tr><td>Phalloidin-Alexa Flour 546</td><td>Thermo Fisher</td><td>A22283</td><td></td></tr><tr><td>Phenol Red indicator (1% w/v)</td><td>Ricca Chemical Company</td><td>5725-16</td><td></td></tr><tr><td>Phosphate buffered saline (PBS)</td><td>Fisher Scientific</td><td>BP399</td><td></td></tr><tr><td>Photoshop</td><td>Adobe</td><td>CS5 v12.0</td><td></td></tr><tr><td>Photoshop software</td><td>Adobe</td><td>CS5 v12.0</td><td></td></tr><tr><td>Plan Apo 60x/1.2 WD objective</td><td>Nikon</td><td></td><td></td></tr><tr><td>Power source</td><td>Wild Heerbrugg</td><td>MTr 22</td><td>Or equivalent power source&amp;nbsp;</td></tr><tr><td>Slide warmer model No. 26020FS</td><td>Fisher Scientific</td><td>12-594</td><td></td></tr><tr><td>Sodium Hydroxide beads</td><td>Fisher Scientific</td><td>S612-3</td><td>CAUTION - corrosive/irritating to eyes and skin, target organ - respiratory system, corrosive to metals</td></tr><tr><td>Superfrost/Plus microscope slide</td><td>Fisher Scientific</td><td>12-550-15</td><td></td></tr><tr><td>Sylgard 184 silicone Dow Corning</td><td>World Prevision Instruments</td><td>SYLG184</td><td></td></tr><tr><td>Tissue-Tek O.C.T. Compound</td><td>Sakura Finetek</td><td>4583</td><td></td></tr><tr><td>Triton X-100 BioXtra</td><td>Sigma</td><td>T9284</td><td>CAUTION &amp;ndash; Toxic, hazardous to aquatic environment, corrosive</td></tr><tr><td>Vannas micro-dissection scissors</td><td>Ted Pella Inc</td><td>1346</td><td>Sharp/sharp straight tips</td></tr><tr><td>Vectashield antifade mouting medium</td><td>Vector laboratories</td><td>H-1000</td><td></td></tr><tr><td>Wheat Germ Agglutinin (WGA)-Alexa Flour-594</td><td>Life Technologies</td><td>W11262</td><td></td></tr></tbody></table>

assessment of zebrafish lens nucleus localization and sutural integrity

The zebrafish is uniquely suited to genetic manipulation and in vivo imaging, making it an increasingly popular model for reverse genetic studies and for generation of transgenics for in vivo imaging. These unique capabilities make the zebrafish an ideal platform to study ocular lens development and physiology. Our recent findings that an Aquaporin-0, Aqp0a, is required for stability of the anterior lens suture, as well as for the shift of the lens nucleus to the lens center with age led us to develop tools especially suited to analyzing the properties of zebrafish lenses. Here we outline detailed methods for lens dissection that can be applied to both larval and adult lenses, to prepare them for histological analysis, immunohistochemistry and imaging. We focus on analysis of lens suture integrity and cortical cell morphology and compare data generated from dissected lenses with data obtained from in vivo imaging of lens morphology made possible by a novel transgenic zebrafish line with a genetically encoded fluorescent marker. Analysis of dissected lenses perpendicular to their optical axis allows quantification of the relative position of the lens nucleus along the anterior-posterior axis. Movement of the lens nucleus from an initial anterior position to the center is required for normal lens optics in adult zebrafish. Thus, a quantitative measure of lens nuclear position directly correlates with its optical properties.

Adult zebrafish eye anatomy closely resembles that of mammals (Figure 1A). Despite some differences between zebrafish and mammalian eyes, such as having a ciliary zone instead of a ciliary body17, differences in optical properties18, and differences in morphogenesis during embryonic development19, the zebrafish eye is an excellent model for studying eye development and understanding ophth...

Watch this Scientific Journal Video about Assessment of Zebrafish Lens Nucleus Localization and Sutural Integrity at JoVE.com

Assessment of Zebrafish Lens Nucleus Localization and Sutural Integrity

These protocols were developed to analyze cortical lens morphology, structural integrity of the zebrafish lens sutures in fixed and live lenses and to measure the position of the zebrafish lens nucleus along the anterior-posterior axis.

The zebrafish is uniquely suited to genetic manipulation and in vivo imaging, making it an increasingly popular model for reverse genetic studies and for ...

assessment-zebrafish-lens-nucleus-localization-sutural

Research

JoVE Journal

Biology

8.5K Views.  University of California, Irvine. Onze protocollen vormen een uitgangspunt voor functionele studies van lensontwikkeling en fysiologie en voor omgekeerde genetische screening van lensfenotypes bij zebravissen. Onze protocollen overbruggen in vitro en in vivo analyses van zebrafish lens membraan morfologie en bieden een nieuwe en eenvoudige methode voor het kwantificeren van de ontwikkeling van zebravis lens optica. Het analyseren van zebravis lens morfologie en optica leidt tot inzichten in mechanismen die de ontwikkeling van...

Video: Assessment of Zebrafish Lens Nucleus Localization and Sutural Integrity

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 ...

Developmental Biology

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM compared to confocal systems are its low phototoxicity, gentle mounting strategies, fast acquisition with high signal to noise ratio and the possibility of imaging samples from various angles (views) for long periods of time. Imaging from multiple views unleashes the full potential of LSFM, but at the same time it can create terabyte-sized datasets. Processing such datasets is the biggest challenge of using LSFM. In this protocol we outline some solutions to this problem. Until recently, LSFM was mostly performed in laboratories that had the expertise to build and operate their own light sheet microscopes. However, in the last three years several commercial implementations of LSFM became available, which are multipurpose and easy to use for any developmental biologist. This article is primarily directed to those researchers, who are not LSFM technology developers, but want to employ LSFM as a tool to answer specific developmental biology questions. 
Here, we use imaging of zebrafish eye development as an example to introduce the reader to LSFM technology and we demonstrate applications of LSFM across multiple spatial and temporal scales. This article describes a complete experimental protocol starting with the mounting of zebrafish embryos for LSFM. We then outline the options for imaging using the commercially available light sheet microscope. Importantly, we also explain a pipeline for subsequent registration and fusion of multiview datasets using an open source solution implemented as a Fiji plugin. While this protocol focuses on imaging the developing zebrafish eye and processing data from a particular imaging setup, most of the insights and troubleshooting suggestions presented here are of general use and the protocol can be adapted to a variety of light sheet microscopy experiments.

Light sheet fluorescence microscopy is an excellent tool for imaging embryonic development. It allows recording of long time-lapse movies of live embryos in near physiological conditions. We demonstrate its application for imaging zebrafish eye development across wide spatio-temporal scales and present a pipeline for fusion and deconvolution of multiview datasets.

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM ...

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 ...

Lens Transplantation in Zebrafish and its Application in the Analysis of Eye Mutants

The lens plays an important role in the development of the optic cup[1,2]. Using the zebrafish as a model organism, questions regarding lens development can be addressed. The zebrafish is useful for genetic studies due to several advantageous characteristics, including small size, high fecundity, short lifecycle, and ease of care. Lens development occurs rapidly in zebrafish. By 72 hpf, the zebrafish lens is functionally mature [3]. Abundant genetic and molecular resources are available to support research in zebrafish. In addition, the similarity of the zebrafish eye to those of other vertebrates provides basis for its use as an excellent animal model of human defects[4-7]. Several zebrafish mutants exhibit lens abnormalities, including high levels of cell death, which in some cases leads to a complete degeneration of lens tissues [8]. To determine whether lens abnormalities are due to intrinsic causes or to defective interactions with the surrounding tissues, transplantation of a mutant lens into a wild-type eye is performed. Using fire-polished metal needles, mutant or wild-type lenses are carefully dissected from the donor animal, and transferred into the host. To distinguish wild-type and mutant tissues, a transgenic line is used as the donor. This line expresses membrane-bound GFP in all tissues, including the lens. This transplantation technique is an essential tool in the studies of zebrafish lens mutants.

Lens development involves interactions with other tissues. Several zebrafish eye mutants are characterized by an abnormally small lens size. Here we demonstrate a lens transplantation experiment to determine whether this phenotype is due to intrinsic causes or defective interactions with tissues that surround the lens.

The lens plays an important role in the development of the optic cup[1,2]. Using the zebrafish as a model organism, questions regarding lens development ...

Visualization of the Superior Ocular Sulcus during Danio rerio Embryogenesis

Congenital ocular coloboma is a genetic disorder that is typically observed as a cleft in the inferior aspect of the eye resulting from incomplete choroid fissure closure. Recently, the identification of individuals with coloboma in the superior aspect of the iris, retina, and lens led to the discovery of a novel structure, referred to as the superior fissure or superior ocular sulcus (SOS), that is transiently present on the dorsal aspect of the optic cup during vertebrate eye development. Although this structure is conserved across mice, chick, fish, and newt, our current understanding of the SOS is limited. In order to elucidate factors that contribute to its formation and closure, it is imperative to be able to observe it and identify abnormalities, such as delay in the closure of the SOS. Here, we set out to create a standardized series of protocols that can be used to efficiently visualize the SOS by combining widely available microscopy techniques with common molecular biology techniques such as immunofluorescent staining and mRNA overexpression. While this set of protocols focuses on the ability to observe SOS closure delay, it is adaptable to the experimenter&#39;s needs and can be easily modified. Overall, we hope to create an approachable method through which our understanding of the SOS can be advanced to expand the current knowledge of vertebrate eye development.

Visualization of the Superior Ocular Sulcus during Danio rerio Embryogenesis

Here, we present a standardized series of protocols to observe the superior ocular sulcus, a recently-identified, evolutionarily-conserved structure in the vertebrate eye. Using zebrafish larvae, we demonstrate techniques necessary to identify factors that contribute to the formation and closure of the superior ocular sulcus.

Congenital ocular coloboma is a genetic disorder that is typically observed as a cleft in the inferior aspect of the eye resulting from incomplete choroid ...

Refined Multiview Light Sheet Microscopy Protocol for Zebrafish Embryos

Light Sheet Microscopy Imaging and Mounting Strategies for Early Zebrafish Embryos

Light sheet microscopy has become the methodology of choice for live imaging of zebrafish embryos over long time scales with minimal phototoxicity. In particular, a multiview system, which allows sample rotation, enables imaging of entire embryos from different angles. However, in most imaging sessions with a multiview system, sample mounting is a troublesome process as samples are usually prepared in a polymer tube. To aid in this process, this protocol describes basic mounting strategies for imaging early zebrafish development between the 70% epiboly and early somite stages. Specifically, the study provides statistics on the various positions the embryos default to at the 70% epiboly and bud stages within the chorion. Furthermore, it discusses the optimum number of angles and the interval between angles required for imaging whole zebrafish embryos at the early stages of development so that cellular-scale information can be extracted by fusing the different views. Finally, since the embryo covers the entire field of view of the camera, which is required to obtain a cellular-scale resolution, this protocol details the process of using bead information from above or below the embryo for the registration of the different views.

Author Spotlight: Strategies for Mounting Zebrafish Embryos for High-Resolution Multiview Light-Sheet Microscopy &#8212; Techniques for Imaging and Image Reconstruction

A sample preparation strategy for imaging early zebrafish embryos within an intact chorion using a light-sheet microscope is described. It analyzes the different orientations that embryos acquire within the chorion at the 70% epiboly and bud stages and details imaging strategies for obtaining cellular-scale resolution throughout the embryo using the light-sheet system.

Light sheet microscopy has become the methodology of choice for live imaging of zebrafish embryos over long time scales with minimal phototoxicity. In ...

Analysis of Skeletal Muscle Defects in Larval Zebrafish by Birefringence and Touch-evoke Escape Response Assays

Zebrafish (Danio rerio) have become a particularly effective tool for modeling human diseases affecting skeletal muscle, including muscular dystrophies1-3, congenital myopathies4,5, and disruptions in sarcomeric assembly6,7, due to high genomic and structural conservation with mammals8. Muscular disorganization and locomotive impairment can be quickly assessed in the zebrafish over the first few days post-fertilization. Two assays to help characterize skeletal muscle defects in zebrafish are birefringence (structural) and touch-evoked escape response (behavioral).
Birefringence is a physical property in which light is rotated as it passes through ordered matter, such as the pseudo-crystalline array of muscle sarcomeres9. It is a simple, noninvasive approach to assess muscle integrity in translucent zebrafish larvae early in development. Wild-type zebrafish with highly organized skeletal muscle appear very bright amidst a dark background when visualized between two polarized light filters, whereas muscle mutants have birefringence patterns specific to the primary muscular disorder they model. Zebrafish modeling muscular dystrophies, diseases characterized by myofiber degeneration followed by repeated rounds of regeneration, exhibit degenerative dark patches in skeletal muscle under polarized light. Nondystrophic myopathies are not associated with necrosis or regenerative changes, but result in disorganized myofibers and skeletal muscle weakness. Myopathic zebrafish typically show an overall reduction in birefringence, reflecting the disorganization of sarcomeres.
The touch-evoked escape assay involves observing an embryo&#39;s swimming behavior in response to tactile stimulation10-12. In comparison to wild-type larvae, mutant larvae frequently display a weak escape contraction, followed by slow swimming or other type of impaired motion that fails to propel the larvae more than a short distance12. The advantage of these assays is that disease progression in the same fish type can be monitored in vivo for several days, and that large numbers of fish can be analyzed in a short time relative to higher vertebrates.

The zebrafish is now an established and powerful tool for modeling muscular dystrophies, congenital myopathies, and related neuromuscular diseases. Birefringence and touch-evoked escape behavior are two common noninvasive assays used to determine the degree of muscular disorganization and locomotive impairment of zebrafish embryos during early development.

Zebrafish (Danio rerio) have become a particularly effective tool for modeling human diseases affecting skeletal muscle, including muscular ...

4-Dimensional Imaging of Zebrafish Optic Cup Morphogenesis

Visual system function requires the establishment of precise tissue and organ structures. In the vertebrate eye, structural defects are a common cause of visual impairment, yet mechanisms of eye morphogenesis are still poorly understood. The basic organization of the embryonic eye is conserved throughout vertebrates, thus live imaging of zebrafish embryos has become a powerful approach to directly observe eye development at real time under normal and pathological conditions. Dynamic cell processes including movements, morphologies, interactions, division, and death can be visualized in the embryo. We have developed methods for uniform labeling of subcellular structures and timelapse confocal microscopy of early eye development in zebrafish. This protocol outlines the method of generating capped mRNA for injection into the 1-cell zebrafish embryo, mounting embryos at optic vesicle stage (~12 hours post fertilization, hpf), and performing multi-dimensional timelapse imaging of optic cup morphogenesis on a laser scanning confocal microscope, such that multiple datasets are acquired sequentially in the same imaging session. Such an approach yields data that can be used for a variety of purposes, including cell tracking, volume measurements, three-dimensional (3D) rendering, and visualization. Our approaches allow us to pinpoint the cellular and molecular mechanisms driving optic cup development, in both wild type and genetic mutant conditions. These methods can be employed directly by other groups or adapted to visualize many additional aspects of zebrafish eye development.

This protocol describes an approach for in toto labeling and multidimensional imaging of zebrafish early eye development. We describe labeling, embedding, and four dimensional (4D) imaging using laser scanning confocal microscopy, and considerations for optimizing acquisition of datasets for dissecting mechanisms of optic cup morphogenesis.

Visual system function requires the establishment of precise tissue and organ structures. In the vertebrate eye, structural defects are a common cause of ...

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 ...

Assessment of Zebrafish Lens Nucleus Localization and Sutural Integrity

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