This work was supported by the National Eye Institute Grant R01 EY032056 to CC and R01 EY017724 to VMF, as well as the National Institute of General Medical Sciences under grant number P20GM139760. S.T.I was supported by NIH-NIGMS T32-GM133395 as part of the Chemistry-Biology Interface predoctoral training program, and by a University of Delaware Graduate Scholars Award.

Mice are housed in the University of Delaware animal facility, maintained in a pathogen-free environment. All animal procedures, including euthanasia by CO2 inhalation, were conducted in accordance with approved animal protocols by the University of Delaware Institutional Animal Care and Use Committee (IACUC).
1. Whole lens mount preparation and imaging
<ol>
	<li>Fixation of lenses for whole mount imaging
	<ol>
		<li>Following euthanasia, enucleate eyes and dissect lenses as previously described25. Following dissection, transfer lenses immediately into fresh 1x phosphate buffered saline (PBS; 1.1 mM KH2PO4, 155 mM, NaCl, 3.0 mM Na2HPO4-7H2O; pH 7.4) at room temperature. 
		NOTE: Cellular morphology may be altered if lenses are stored in PBS for an extended period of time, therefore, it is recommended to fix immediately within ~10 min of dissection.</li>
		<li>For whole-mount imaging of the lens anterior region, fix whole lenses by immersing in 0.5 mL of freshly made 4% paraformaldehyde (PFA) in 1x PBS in a microcentrifuge tube at room temperature. After 30 min, wash the lenses 3x (5 min per wash) with 1x PBS. Proceed to step 1.2 or store in 1x PBS at 4 &#176;C. Fixed lenses can be stored for up to 5 days.</li>
		<li>For whole-mount imaging of the lens equatorial region, fix whole lenses by immersing in 0.5 mL of freshly made 4% PFA in 1x PBS in a microcentrifuge tube at room temperature. After 1 h, wash lenses 3x (5 min per wash) with 1x PBS. Proceed to step 1.3 or store in 1x PBS at 4&#160;&#176;C. Fixed lenses can be stored for up to 5 days.</li>
	</ol>
	</li>
	<li>Whole mount of anterior lens region (Fixed or live)
	<ol>
		<li>For whole-mount imaging of fixed lenses, proceed to step 1.2.3.</li>
		<li>For whole-mount imaging of live lenses, transfer lenses into a well of a 48-well plate containing 1 mL of Medium 199 (phenol red-free) containing 1% antibiotic/antimycotic. Incubate lenses at 37 &#176;C and 5% CO2 until imaging. Before imaging, incubate lenses in a solution containing fluorescent-labeled Wheat Germ Agglutinin (WGA-640, 1:500) and Hoechst 33342 (1:500) in Medium 199 for at least 10 min. Proceed to step 1.5.</li>
		<li>To stain lens capsule, F-actin, and nuclei of fixed lenses, place lenses in a 500 &#181;L solution containing WGA-640 (1:500), rhodamine-phalloidin (1:50), and Hoechst 33342 (1:500) in permeabilization/blocking buffer (1x PBS containing 0.3% Triton, 0.3% bovine serum albumin (Fraction V) and 3% goat serum) in a microcentrifuge tube. Stain lenses at 4 &#176;C overnight.</li>
		<li>After overnight incubation, wash the lenses 3x (5 min per wash) with 1 mL of 1x PBS. Proceed to imaging lenses.</li>
		<li>To stabilize the fixed lens for confocal imaging, create lens immobilization divots in agarose on an imaging dish as previously described21 (Figure 1).
		<ol>
			<li>Heat and gently mix 2% agarose in PBS using a microwave till the solution is liquified. Pipette 250 &#181;L of liquefied 2% agarose into a glass bottom dish (Figure 1A) and flatten the agarose across the dish using a flexible plastic coverslip (Figure 1B). Once the agarose is cooled and fully solidified, remove the coverslip using fine-tip forceps (Figure 1C) and use a 3mm biopsy punch to create a hole in agarose at the center of the dish (Figure 1D).</li>
			<li>Remove excess agarose using a delicate task wipe (Figure 1E). Keep agarose mold hydrated with 1x PBS and maintain at 4&#160;&#176;C until use. Agarose molds have been successfully stored for up to 1 week.</li>
		</ol>
		</li>
		<li>Using embryo forceps, gently transfer the live or fixed lens into the divot in the agarose (Figure 1F), containing ~2 mL of 1x PBS (fixed lens) or phenol red-free Medium 199 (live lens), and then place the dish on an inverted microscope stage. To confirm that the lens is situated with the anterior region facing the objective, visualize nuclei staining. If no nuclei are observed, the posterior region may be facing the objective.</li>
		<li>To invert the lens, use curved forceps and gently rotate the lens ~180&#176; so that the anterior region faces the objective. Acquire images using a confocal microscope.</li>
		<li>For visualization of lens capsules, acquire z-stack images using a 40x objective with a step size of 0.3 &#181;m. Acquire the first image before the surface of the lens capsule (indicated by WGA staining) and the last image after the apical surface of the epithelial cells. To visualize epithelial cells, acquire z-stack images using a 63x objective with a step size of 0.3 &#181;m for visualization of lens epithelial cells. 
		NOTE: These microscope parameters allow for adequate three dimensional (3D) reconstruction of images which is essential for image quantification of capsule thickness or epithelial cell area. It is also possible to image sutures in live lens tdTomato lenses by imaging past the epithelial cell monolayer as previously described4.</li>
	</ol>
	</li>
	<li>Lens equatorial epithelial and fiber cell staining
	<ol>
		<li>Place fixed lenses in a 0.5 mL solution of rhodamine-phalloidin (1:300), WGA-640 (1:250), and Hoechst 33342 (1:500) in permeabilization/blocking solution (3% BSA, 3% goat serum and 0.3% Triton) within a microcentrifuge tube. Maintain at 4&#160;&#176;C overnight.</li>
		<li>After overnight incubation, wash lenses 3x in 1mL 1x PBS (5 min per wash).</li>
		<li>Create lens immobilization agarose wedges as previously described4,10.
		<ol>
			<li>Briefly, create agarose wedges in glass bottomed dishes (FD35-100, WPI) by pouring ~5-6 mL of molten 2% agarose in 1x PBS into dishes (Figure 2A). Once the agarose solidifies (Figure 2B), create a triangular divot with a sharp blade (Figure 2C). Remove the agarose wedge and place 1 mL of 1x PBS into the agarose dish (Figure 2D).</li>
			<li>Aspirate any residual agarose that may be left behind from cutting the agarose. Multiple wedges can be created per dish to fit multiple different sizes of lenses (not shown). To store the agarose mold, place 1mL of 1x PBS onto the agarose mold and keep at 4&#160;&#176;C. Wedges can be kept for up to 1 week.</li>
		</ol>
		</li>
		<li>Using curved tweezers, place the lens within an agarose wedge containing 1mL of 1x PBS and adjust so that the equatorial region of the lens is facing down onto microscope glass above the confocal objective (Figure 2E-F). To confirm that the equatorial region is in focus, visualize nuclei and ensure that nuclei are aligned in rows at the equator. The equatorial epithelial cells can also be identified as they are irregularly packed and shaped. As well, the meridional row cells are precisely aligned and hexagonal shaped, indicated by F-actin staining at the cell membranes.</li>
		<li>If nuclei in the field of view are randomly packed, the anterior side of the lens is facing the objective. If nuclei cannot be observed, then the posterior side of the lens is most likely facing the objective. If the lens is not sitting on its equator, re-orient the lens, and use the curved tweezer to rotate the lens until the precisely aligned nuclei at the lens equator is observed.</li>
		<li>Image the lens equatorial epithelial and fiber cells using a laser-scanning confocal microscope (20x objective, NA = 0.8, step size 0.5 &#181;m and/or 40x oil objective, NA = 1.3, step size 0.4 &#181;m). Rotate the images before z-stack collection, in which equatorial epithelial cells are on the top, followed by meridional row and fiber cells right below.</li>
	</ol>
	</li>
</ol>
2. Image analysis methodology
<ol>
	<li>Lens capsule thickness measurement
	<ol>
		<li>Using z-stack images obtained with a 40x objective (0.3 &#181;m step size) of either live tdTomato lenses labelled with WGA or fixed lenses labelled with WGA and rho-phalloidin, obtain an optical 2D projection in the XZ view of the 3D reconstruction and save in .tif format. Process images using Zen software.</li>
		<li>Open XZ slice (.tif) images in FIJI ImageJ software.</li>
		<li>To measure lens capsule thickness, use the straight line function (depicted in Figure 3A,B). Set the line width to 50 pixels by selecting Edit &#62; Options &#62; Line width. This line width was empirically determined to provide a high signal-to-noise ratio with the microscope settings. Optimal line width may differ on other microscopes/microscope settings or when using lenses from other species. Next, draw a line from the capsule&#39;s top surface to beneath the epithelial cells&#39; basal region.</li>
		<li>Save the line as a region of interest by pressing Ctrl + T.</li>
		<li>Separate channels into tabs for image, color, and split channels.</li>
		<li>Measure the intensity along the line for the capsule channel by pressing Ctrl + K.</li>
		<li>In a spreadsheet, plot intensity values as a function of distance and determine intensity peaks for the capsule and F-actin, which represent the capsule surface and basal region of epithelial cells, respectively. On the x-axis is the line distance.</li>
		<li>Next, calculate capsule thickness by determining the distance between capsule and epithelial fluorescent intensity peaks (Figure 3C, D).</li>
	</ol>
	</li>
	<li>Cell area analysis
	<ol>
		<li>Using z-stack images obtained using a 63x objective lens (NA = 1.4; 0.3 &#181;m step size) on live tdTomato lenses or fixed lenses labelled with phalloidin, analyze an XY view slice from a z-stack confocal image corresponding to where the middle (lateral) region of epithelial cells is in focus. Note the lateral membranes are visible using the tdTomato membrane dye or by visualizing phalloidin which are present at the lateral cell membranes. 
		NOTE: Due to the curvature of the lens (as seen in an XZ view; Figure 4A-C), not all epithelial cells will be in focus in the image. The middle region of the epithelium corresponds to the region where both the cell lateral membranes (Figure 4D-E) and nuclei (Figure 4G-I) are in focus.</li>
		<li>Export the image as a .tif and open it in FIJI ImageJ.</li>
		<li>To set the scale in FIJI ImageJ for analysis, use the line tool to create a line that is the length of a scale bar from the confocal image.</li>
		<li>Record the pixel length of the line and the length of the scale bar. Go to Analyze on the toolbar menu and select Set Scale. Input the number of pixels that is the length of the line in Distance in Pixels and in the Known Distance input the known length of the scale bar.</li>
		<li>Using the tdTomato or rhodamine-phalloidin staining as an indication of cell boundaries, manually outline a population of cells that are in focus within the image using the polygonal tool. Only trace cells where lateral membranes are visible (Figure 4E) and avoid tracing partially visible cells (i.e., at the edge of the images). Save ROI by pressing Ctrl+T. Go to Edit on the toolbar menu and select Clear Outside. Now measure total cell area (ROI) by pressing Ctrl + M in FIJI ImageJ.</li>
		<li>Calculate the average cell area by dividing the ROI area by the total cell number. A simple way to determine the number of cells is by counting the number of nuclei. Go to the nuclei channel (blue). On the ROI menu, select the saved ROI outline of cells (Figure 4G). Clear outside of the ROI by going to Edit and then clicking Clear outside (Figure 4H). Using the Multi-point tool, count the nuclei by clicking on individual nuclei.</li>
	</ol>
	</li>
	<li>Cellular nuclear area and shape analysis
	<ol>
		<li>For nuclei area and shape analysis, analyze an XY plane view optical section from a z-stack confocal image corresponding to where the middle (lateral) region of tdTomato epithelial cells are in focus. Analyze a z-stack slice from a confocal image where the nuclear area is the largest (Figure 5A); this represents the mid portion of the nuclei. 
		NOTE: Note that due to the curvature of the lens, not all nuclei will be in focus, as can be seen in the XZ view (Figure 4G and Figure 5A).</li>
		<li>Select a subpopulation of nuclei that are in focus and save a ROI.</li>
		<li>Use the Clear Outside function. Within the ROI, using the Freehand selection tool (fourth menu button from the left), carefully trace the borders of the nuclei (Figure 5B). Save each nuclear trace into the ROI window by pressing Ctrl+T. Only outline nuclei where the complete nuclei can be seen, and the nuclei are not touching each other.</li>
		<li>Once all the nuclei are outlined, measure individual cells&#39; nuclear area and shape (i.e., circularity) by pressing Ctrl+M.</li>
		<li>Copy and paste data into a spreadsheet and calculate the average nuclear area and circularity (Figure 5C).</li>
	</ol>
	</li>
	<li>Meridional row epithelial packing
	<ol>
		<li>To measure meridional row disorder, acquire images using a 20x objective (0.5 &#181;m step size).</li>
		<li>Identify the lens fulcrum/modiolus on images. The fulcrum is the region where the apical tips of elongating epithelial cells constrict to form an anchor point during initial fiber cell differentiation and elongation at the equator. Identify the fulcrum based on bright F-actin intensity (indicated by arrowhead in XZ view in Figure 6A; indicated by a red dashed line in Figure 6B) and change in cellular organization in a single optical section at XY view. 
		NOTE: In addition, F-actin staining of both the basal and apical regions of epithelial cells are apparent above the fulcrum, whereas only the basal region of the elongating cells are apparent below the fulcrum (Figure 6A). The epithelial cells above the fulcrum line are irregularly packed and tend to form rosettes, whereas the fiber cells below the fulcrum are arranged in parallel rows, indicated by bright F-actin staining at the membrane (Figure 6B).</li>
		<li>Once the fulcrum is identified in 2-month-old mice, select the single optical section ~4.5-5 &#181;m peripheral to the fulcrum (toward the lens capsule) in the XY plane. This distance is selected based on the observation that all the nuclei of the meridional row cells in 2-month-old mice are in focus when they are ~5 &#181;m peripheral to the fulcrum. Save the single optical section (.tif). Open the image on FIJI. 
		NOTE: This analysis has only been performed on 2-month-old mice and therefore, it cannot be concluded whether this distance changes with age.</li>
		<li>Before performing image analysis, set the scale to &#181;m in ImageJ using step 2.2.2.</li>
		<li>Manually outline the entire meridional row regions using the freehand line tool (region of interest/ ROI) by identifying nuclear alignment, as shown in Figure 7A. Save ROI by pressing Ctrl+T. Go to Edit on the toolbar menu and select Clear Outside. Measure total cell area (ROI) by pressing Ctrl + M in FIJI ImageJ.</li>
		<li>Identify any regions of disorder indicated by F-actin staining by outlining using the freehand line tool in FIJI ImageJ. Criteria for disordered regions include branching of rows, irregular packing, and misalignment of rows (Figure 7B), as shown previously20.</li>
		<li>Measure the outlined disordered area/patched by pressing Ctrl + M in FIJI ImageJ. Sum the total disordered area in a spreadsheet.</li>
		<li>Divide the total disordered area by the ROI area, then multiply by 100 to get a percent disordered area. If no disorder is observed, place a value of 0% for the disordered area.</li>
	</ol>
	</li>
	<li>Meridional row number of neighboring cells
	<ol>
		<li>To measure number of neighboring cells, use F-actin-stained images acquired with 40x oil objective (0.4 &#181;m step size).</li>
		<li>Identify an optical section (XY plane) at the basal region of the meridional row cells inward from the lens capsule where F-actin is enriched around the entire perimeter of the meridional row cells, all meridional row cells are in focus and are on the same plane.</li>
		<li>Count the number of adjacent cells each meridional row cell has (Figure 8). Measure the average percentage of cells with six adjacent cells in a spreadsheet. 
		NOTE: Determine the number of adjacent cells with a 20x objective. However, it is significantly easier to observe the hexagon shape with a 40x objective.</li>
	</ol>
	</li>
	<li>Image analysis of equatorial fiber cells
	<ol>
		<li>Take z-stack confocal images of rhodamine phalloidin stained lenses with a 20x objective (0.5 &#181;m step size).</li>
		<li>Identify the fulcrum as indicated in step 2.4.2. Once the fulcrum is identified, select a single optical section. For standardization purposes and to compare between lenses, quantify fiber cell widths ~10 &#181;m inward from the fulcrum in the XY plane (Figure 9A).</li>
		<li>Export raw images to FIJI ImageJ. In FIJI ImageJ, draw a line (usually ~300-400 &#181;m long) across several adjacent fiber cells to measure the distance between F-actin-stained peaks (line scan analysis; Figure 9A; pink line).</li>
		<li>Obtain the fluorescent intensity over the line scan distance in FIJI by pressing Ctrl+K. Next, export data into the spreadsheet to calculate the interpeak distance (example shown in Figure 9A-B). This corresponds to fiber cell widths.</li>
	</ol>
	</li>
</ol>

Preparation and Imaging of Murine Ocular Lens Whole Mounts

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D., Tasic, B., Miyamichi, K., Li, L., Luo, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+global+double-fluorescent+Cre+reporter+mouse.">A global double-fluorescent Cre reporter mouse.</a> Genesis. 45 (9), 593-605 (2007).</li><li>Zhang, Y., 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=Mouse+models+of+MYH9-related+disease:+mutations+in+nonmuscle+myosin+II-A.">Mouse models of MYH9-related disease: mutations in nonmuscle myosin II-A.</a> Blood. 119 (1), 238-250 (2012).</li><li>Cheng, C., Gokhin, D. S., Nowak, R. B., Fowler, V. 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(111), e53986 (2016).</li><li>Riedl, J., 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=Lifeact:+a+versatile+marker+to+visualize+F-actin.">Lifeact: a versatile marker to visualize F-actin.</a> Nat Methods. 5 (7), 605-607 (2008).</li><li>Lukinavicius, G., 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=Fluorogenic+probes+for+live-cell+imaging+of+the+cytoskeleton.">Fluorogenic probes for live-cell imaging of the cytoskeleton.</a> Nat Methods. 11 (7), 731-733 (2014).</li></ol>

The authors have nothing to disclose.

The protocols described enable high spatial resolution visualization of peripheral lens structures and cells at the anterior and equatorial regions of the lens. In this study, methods for the visualization of lens peripheral structures using intact (live or fixed) lenses where the overall 3D lens architecture is preserved were shown. Additionally, simple methods for morphometric quantitative analysis using publicly available FIJI ImageJ software were provided. The whole mount visualization and quantification methods has ...

The ocular lens is a flexible, transparent tissue situated at the anterior region of the eye that functions to fine-focus light onto the retina. The ability of the lens to function can be attributed, in part, to its intricate architecture and organization1,2,3,4,5,6. Surrounding the lens tissue is the capsule, a basement membrane essential for maintaining lens structure and biomechanical properties7,8,9. The lens itself is entirely cellular, consisting of two cell types: epithelial and fiber cells. The epithelial layer consists of a monolayer of cuboidal cells that cover the anterior hemisphere of the lens10. Throughout life, the epithelial cells proliferate and migrate along the lens capsule toward the lens equator. Anterior epithelial cells are quiescent and cobble-stone in cross-section, and near the lens equator, epithelial cells proliferate and start to undergo the differentiation process into new fiber cells11,12. Equatorial epithelial cells transform from randomly packed cells into organized meridional rows with hexagon-shaped cells. Hexagonal cell shape is maintained on the basal side of these differentiating cells while the apical side constricts and anchors at the lens fulcrum or modiolus4,13,14,15. As the equatorial epithelial cells start to elongate into newly formed fiber cells, the apical tips of the cells migrate along the apical surface of anterior epithelial cells toward the anterior pole while the basal tips move along the lens capsule toward the posterior pole. New generations of fiber cells overlay previous generations of cells, creating spherical shells of fibers. During the cell elongation and maturation process, fiber cells substantially alter their morphology11,12,16. These fiber cells form the bulk of the lens mass11,12,16,17,18.
The molecular mechanisms that contribute to establishing intricate lens microstructures, cell morphology, and unique cellular organization are not entirely known. Moreover, the contribution of the lens capsule and cell structure to overall lens function (transparency, lens shape change) is unclear. However, these relationships are being elucidated using new imaging methodology and quantitative assessments of lens structural and cellular features2,4,19,20,21,22. New protocols to image whole lenses that allow for high spatial resolution visualization of the lens capsule, epithelial cells, and peripheral fiber cells have been developed. This includes methodology to quantify capsule thickness, cell size, cell nucleus size and circularity, meridional row order, fiber cell packing, and fiber cell widths. These visualization and image quantification methods allow in-depth morphometric examination and have advantages over other visualization methods (imaging of flat mounts or tissue sections) by preserving overall 3D tissue structure. These methods have permitted for the testing of novel hypotheses and will enable continued advancement in understanding of lens cell pattern development and function.
For the following experiments, we use wild-type and Rosa26-tdTomato mice tandem dimer-Tomato (B6.129(Cg)-Gt(ROSA) (tdTomato)23 (Jackson Laboratories) in the C57BL/6J background between the ages of 6 and 10 weeks, of both sexes. The tdTomato mice allow for visualization of cellular plasma membranes in live lenses via expression of tdTomato protein fused to the N-terminal 8 amino acids of a mutated MARCKS protein that targets the plasma membrane via N-terminal myristylation and internal cysteine-palmitoylation sites23. We also use NMIIAE1841K/E1841K mice24 obtained originally from Dr. Robert Adelstein (National Heart, Lung, and Blood Institute, National Institutes of Health, Bethesda, MD). As described previously20, NMIIAE1841K/E1841K&#160;mice in FvBN/129SvEv/C57Bl6 background that has loss of CP49 beaded intermediate filament protein (maintains mature fiber cell morphology and whole lens biomechanics), are backcrossed with C57BL6/J wild-type mice. We screened the offspring for the presence of the wild-type CP49 allele. 
Confocal imaging was performed on a laser-scanning confocal fluorescence microscope with a 20x (NA = 0.8, working distance = 0.55mm, 1x zoom), a 40x (NA = 1.3 oil objective, working distance = 0.2mm, 1x zoom), or a 63x (NA = 1.4 oil objective, working distance = 0.19mm, 1x zoom) magnification. All images were acquired using a pinhole size, which is a determinant of optical section thickness, to 1 Airy Unit (the resultant optical thicknesses are stated in figure legends). Images were processed on Zen Software. Images were exported to .tif format and then imported into FIJI ImageJ Software (imageJ.net).

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			<td>Delicate task wipes</td>
			<td>Kimwipe</td>
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			<td>Glass bottomed dish (Fluorodish)</td>
			<td>World Precision International</td>
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			<td>Hoescht 33342</td>
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			<td>Laser scanning confocal Microscope 880</td>
			<td>Zeiss</td>
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			<td>MatTek Imaging Dish</td>
			<td>MatTek Life Sciences</td>
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			<td>Paraformaldehyde&#160;</td>
			<td>Electron Microscopy Sciences</td>
			<td>100503-917</td>
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			<td>PBS</td>
			<td>GenClone</td>
			<td>25-507B</td>
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			<td>Phenol red-free medium 199</td>
			<td>Gibco</td>
			<td>11043023</td>
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			<td>Rhodamine-Phalloidin</td>
			<td>Thermo Fisher</td>
			<td>00027</td>
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			<td>Triton X100</td>
			<td>Sigma-Aldrich</td>
			<td>11332481001</td>
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			<td>WGA-640</td>
			<td>Biotium</td>
			<td>CF 640R</td>
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whole mount imaging to visualize and quantify peripheral lens structure, cell morphology, and organization

The ocular lens is a transparent flexible tissue that alters its shape to focus light from different distances onto the retina. Aside from a basement membrane surrounding the organ, called the capsule, the lens is entirely cellular consisting of a monolayer of epithelial cells on the anterior hemisphere and a bulk mass of lens fiber cells. Throughout life, epithelial cells proliferate in the germinative zone at the lens equator, and equatorial epithelial cells migrate, elongate, and differentiate into newly formed fiber cells. Equatorial epithelial cells substantially alter morphology from randomly packed cobble-stone-shaped cells into aligned hexagon-shaped cells forming meridional rows. Newly formed lens fiber cells retain the hexagonal cell shape and elongate toward the anterior and posterior poles, forming a new shell of cells that are overlaid onto previous generations of fibers. Little is known about the mechanisms that drive the remarkable morphogenesis of lens epithelial cells to fiber cells. To better understand lens structure, development, and function, new imaging protocols have been developed to image peripheral structures using whole mounts of ocular lenses. Here, methods to quantify capsule thickness, epithelial cell area, cell nuclear area and shape, meridional row cell order and packing, and fiber cell widths are shown. These measurements are essential for elucidating the cellular changes that occur during lifelong lens growth and understanding the changes that occur with age or pathology.

Anterior lens capsule, epithelial cell area, and nuclear area 
To analyze lens capsule thickness, we stained lens capsules, in either live or fixed lenses, with WGA. We identified lens epithelial cells by labeling membranes with tdTomato in live lenses (Figure 2A), or via rhodamine-phalloidin staining for F-actin at the cell membranes in fixed lenses (Figure 2B). In an orthogonal (XZ) projection, staining for WGA and tdTomato/rhodamine-phal...

Watch this Scientific Journal Video about In-Depth Morphometric Examination and Quantification of Native Lens Structure Using Whole Mount Imaging at JoVE.com

Author Spotlight: In-Depth Morphometric Examination and Quantification of Native Lens Structure Using Whole Mount Imaging 

The present protocols describe novel whole mount imaging for the visualization of peripheral structures in the ocular lens with methods for image quantification. These protocols can be used in studies to better understand the relationship between lens microscale structures and lens development/function.

Whole Mount Imaging to Visualize and Quantify Peripheral Lens Structure, Cell Morphology, and Organization

The ocular lens is a transparent flexible tissue that alters its shape to focus light from different distances onto the retina. Aside from a basement ...

whole-mount-imaging-to-visualize-quantify-peripheral-lens-structure

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864 Views.  University of Delaware. The present protocols describe novel whole mount imaging for the visualization of peripheral structures in the ocular lens with methods for image quantification. These protocols can be used in studies to better understand the relationship between lens microscale structures and lens development/function.

Video: Author Spotlight: In-Depth Morphometric Examination and Quantification of Native Lens Structure Using Whole Mount Imaging 

Double Whole Mount in situ Hybridization of Early Chick Embryos

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  gene expression in brain, neural tube, somite and heart primordia formation. This video demonstrates the different steps in 2-color whole mount in situ hybridization; First, the embryo is dissected from the egg and fixed in paraformaldehyde. Second, the embryo is processed for prehybridization. The embryo is then hybridized with two different probes, one coupled to DIG, and one coupled to FITC.  Following overnight hybridization, the embryo is incubated with DIG coupled antibody. Color reaction for DIG substrate is performed, and the region of interest appears blue. The embryo is then incubated with FITC coupled antibody. The embryo is processed for color reaction with FITC, and the region of interest appears red. Finally, the embryo is fixed and processed for phtograph and sectioning. A troubleshooting guide is also presented.

This video demonstrates 2-color whole mount in situ hybridization, a method by which the spatial and temporal expression pattern of 2 different genes can be visualized in young chick embryos. This method was originally introduced by David Wilkinson, Domingos Henrique, Phil Ingham and David Ish -Horowicz.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

Method for Whole Mount Antibody Staining in Chick

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying antibody expression in developing brain, neural tube and somite. This video demonstrates the different steps in whole-mount antibody staining using HRP conjugated secondary antibodies; First, the embryo is dissected from the egg and fixed in paraformaldehyde. Second, endogenous peroxidase is inactivated; The embryo is then exposed to primary antibody. After several washes, the embryo is incubated with secondary antibody conjugated to HRP. Peroxidase activity is revealed using reaction with diaminobenzidine substrate. Finally, the embryo is fixed and processed for photography and sectioning. The advantage of this method over the use of fluorescent antibodies is that embryos can be processed for wax sectioning, thus enabling the study of antigen sites in cross section. This method was originally introduced by Jane Dodd and Tom Jessell 1.

This video demonstrates whole mount immunohistochemistry, a method by which the spatial and temporal expression pattern of an antigen can be visualized in young chick embryos. This method was originally introduced by Jane Dodd and Tom Jessell.

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 ...

Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology

Whole mount in situ hybridization (WISH) is a common technique in molecular biology laboratories used to study gene expression through the localization of specific mRNA transcripts within whole mount specimen. This technique (adapted from Albertson and Yelick, 2005) was used in an upper level undergraduate Comparative Vertebrate Biology laboratory classroom at Syracuse University. The first two thirds of the Comparative Vertebrate Biology lab course gave students the opportunity to study the embryology and gross anatomy of several organisms representing various chordate taxa primarily via traditional dissections and the use of models. The final portion of the course involved an innovative approach to teaching anatomy through observation of vertebrate development employing molecular techniques in which WISH was performed on zebrafish embryos. A heterozygous fibroblast growth factor 8 a (fgf8a) mutant line, ace, was used. Due to Mendelian inheritance, ace intercrosses produced wild type, heterozygous, and homozygous ace/fgf8a mutants in a 1:2:1 ratio. RNA probes with known expression patterns in the midline and in developing anatomical structures such as the heart, somites, tailbud, myotome, and brain were used. WISH was performed using zebrafish at the 13 somite and prim-6 stages, with students performing the staining reaction in class. The study of zebrafish embryos at different stages of development gave students the ability to observe how these anatomical structures changed over ontogeny. In addition, some ace/fgf8a mutants displayed improper heart looping, and defects in somite and brain development. The students in this lab observed the normal development of various organ systems using both external anatomy as well as gene expression patterns. They also identified and described embryos displaying improper anatomical development and gene expression (i.e., putative mutants).
For instructors at institutions that do not already own the necessary equipment or where funds for lab and curricular innovation are limited, the financial cost of the reagents and apparatus may be a factor to consider, as will the time and effort required on the part of the instructor regardless of the setting. Nevertheless, we contend that the use of WISH in this type of classroom laboratory setting can provide an important link between developmental genetics and anatomy. As technology advances and the ability to study organismal development at the molecular level becomes easier, cheaper, and increasingly popular, many evolutionary biologists, ecologists, and physiologists are turning to research strategies in the field of molecular biology. Using WISH in a Comparative Vertebrate Biology laboratory classroom is one example of how molecules and anatomy can converge within a single course. This gives upper level college students the opportunity to practice modern biological research techniques, leading to a more diversified education and the promotion of future interdisciplinary scientific research.

Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology

Whole mount in situ hybridization (WISH) was used in an upper level undergraduate Comparative Vertebrate Biology course in addition to vertebrate dissections. This gave students the opportunity to study gene expression patterns as well as gross anatomy, linking the study of molecular and organismal biology within one course.

Whole mount in situ hybridization (WISH) is a common technique in molecular biology laboratories used to study gene expression through the localization of ...

Whole-mount Immunohistochemical Analysis for Embryonic Limb Skin Vasculature: a Model System to Study Vascular Branching Morphogenesis in Embryo

Whole-mount immunohistochemical analysis for imaging the entire vasculature is pivotal for understanding the cellular mechanisms of branching morphogenesis. We have developed the limb skin vasculature model to study vascular development in which a pre-existing primitive capillary plexus is reorganized into a hierarchically branched vascular network. Whole-mount confocal microscopy with multiple labelling allows for robust imaging of intact blood vessels as well as their cellular components including endothelial cells, pericytes and smooth muscle cells, using specific fluorescent markers. Advances in this limb skin vasculature model with genetic studies have improved understanding molecular mechanisms of vascular development and patterning. The limb skin vasculature model has been used to study how peripheral nerves provide a spatial template for the differentiation and patterning of arteries. This video article describes a simple and robust protocol to stain intact blood vessels with vascular specific antibodies and fluorescent secondary antibodies, which is applicable for vascularized embryonic organs where we are able to follow the process of vascular development.

We introduce a whole-mount immunohistochemistry and laser scanning confocal microscopy with multiple labelling for analyzing intricate vascular network formation in mouse embryonic limb skin.

Whole-mount immunohistochemical analysis for imaging the entire vasculature is pivotal for understanding the cellular mechanisms of branching ...

Whole Mount in Situ Hybridization of E8.5 to E11.5 Mouse Embryos

Whole mount in situ hybridization is a very informative approach for defining gene expression patterns in embryos. The in situ hybridization procedures are lengthy and technically demanding with multiple important steps that collectively contribute to the quality of the final result. This protocol describes in detail several key quality control steps for optimizing probe labeling and performance. Overall, our protocol provides a detailed description of the critical steps necessary to reproducibly obtain high quality results. First, we describe the generation of digoxygenin (DIG) labeled RNA probes via in vitro transcription of DNA templates generated by PCR. We describe three critical quality control assays to determine the amount, integrity and specific activity of the DIG-labeled probes. These steps are important for generating a probe of sufficient sensitivity to detect endogenous mRNAs in a whole mouse embryo. In addition, we describe methods for the fixation and storage of E8.5-E11.5 day old mouse embryos for in situ hybridization. Then, we describe detailed methods for limited proteinase K digestion of the rehydrated embryos followed by the details of the hybridization conditions, post-hybridization washes and RNase treatment to remove non-specific probe hybridization. An AP-conjugated antibody is used to visualize the labeled probe and reveal the expression pattern of the endogenous transcript. Representative results are shown from successful experiments and typical suboptimal experiments.

Whole Mount in Situ Hybridization of E8.5 to E11.5 Mouse Embryos

This whole mount in situ hybridization protocol discusses critical steps that ensure reproducible high quality results for gene expression studies in E8.5-E11.5 day old mouse embryos.

Whole mount in situ hybridization is a very informative approach for defining gene expression patterns in embryos. The in situ hybridization procedures ...

RNA In situ Hybridization in Whole Mount Embryos and Cell Histology Adapted for Marine Elasmobranchs

Marine elasmobranchs are valued animal models for biomedical and genomic studies as they are the most primitive vertebrates to have adaptive immunity and have unique mechanisms for osmoregulation 1-3. As the most primitive living jawed-vertebrates with paired appendages, elasmobranchs are an evolutionarily important model, especially for studies in evolution and development. Marine elasmobranchs have also been used to study aquatic toxicology and stress physiology in relationship to climate change 4. Thus, development and adaptation of methodologies is needed to facilitate and expand the use of these primitive vertebrates to multiple biological disciplines. Here I present the successful adaptation of RNA whole mount in situ hybridization and histological techniques to study gene expression and cell histology in elasmobranchs.
Monitoring gene expression is a hallmark tool of developmental biologists, and is widely used to investigate developmental processes 5. RNA whole mount in situ hybridization allows for the visualization and localization of specific gene transcripts in tissues of the developing embryo. The expression pattern of a gene's message can provide insight into what developmental processes and cell fate decisions a gene may control. By comparing the expression pattern of a gene at different developmental stages, insight can be gained into how the role of a gene changes during development.
While whole mount in situ's provides a means to localize gene expression to tissue, histological techniques allow for the identification of differentiated cell types and tissues. Histological stains have varied functions. General stains are used to highlight cell morphology, for example hematoxylin and eosin for general staining of nuclei and cytoplasm, respectively. Other stains can highlight specific cell types. For example, the alcian blue stain reported in this paper is a widely used cationic stain to identify mucosaccharides. Staining of the digestive tract with alcian blue can identify the distribution of goblet cells that produce mucosaccharides. Variations in mucosaccharide constituents on short peptides distinguish goblet cells by function within the digestive tract 6. By using RNA whole mount in situ's and histochemical methods concurrently, cell fate decisions can be linked to gene-specific expression.
Although RNA in situ's and histochemistry are widely used by researchers, their adaptation and use in marine elasmobranchs have met limited and varied success. Here I present protocols developed for elasmobranchs and used on a regular basis in my laboratory. Although further modification of the RNA in situ's hybridization method may be needed to adapt to different species, the protocols described here provide a strong starting point for researchers wanting to adapt the use of marine elasmobranchs to their scientific inquiries.

RNA In situ Hybridization in Whole Mount Embryos and Cell Histology Adapted for Marine Elasmobranchs

By combining methods for RNA whole mount in situ hybridization and histology, gene expression can be linked with cell fate decisions in the developing embryo. These methods have been adapted to marine elasmobranchs and facilitate the use of these animals as model organisms for biomedical, toxicology and comparative studies.

Marine  elasmobranchs are valued animal models for biomedical and genomic studies as  they are the most primitive vertebrates to have adaptive immunity ...

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

This article focuses on whole-mount in situ hybridization (WISH) of zebrafish embryos. The WISH technology facilitates the assessment of gene expression both in terms of tissue distribution and developmental stage. Protocols are described for the use of WISH of zebrafish embryos using antisense RNA probes labeled with digoxigenin. Probes are generated by incorporating digoxigenin-linked nucleotides through in vitro transcription of gene templates that have been cloned and linearized. The chorions of embryos harvested at defined developmental stages are removed before incubation with specific probes. Following a washing procedure to remove excess probe, embryos are incubated with anti-digoxigenin antibody conjugated with alkaline phosphatase. By employing a chromogenic substrate for alkaline phosphatase, specific gene expression can be assessed. Depending on the level of gene expression the entire procedure can be completed within 2-3 days.

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

The zebrafish, a small tropical fish, has become a popular model for studying gene function during vertebrate development and disease. The temporal and spatial expression of target genes can be determined by in situ hybridization. Our improved protocol allows for the detection of low abundant transcripts with low non-specific background signal.

This article focuses on whole-mount in situ hybridization (WISH) of zebrafish embryos. The WISH technology facilitates the assessment of gene expression ...

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

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model congenital abnormalities. During the first day of life, the zebrafish embryo progresses through many developmental stages including fertilization, cleavage, gastrulation, segmentation, and the organogenesis of structures such as the kidney, heart, and central nervous system. The anatomy of a young zebrafish embryo presents several challenges for the visualization and analysis of the tissues involved in many of these events because the embryo develops in association with a round yolk mass. Thus, for accurate analysis and imaging of experimental phenotypes in fixed embryonic specimens between the tailbud and 20 somite stage (10 and 19 hours post fertilization (hpf), respectively), such as those stained using whole mount in situ hybridization (WISH), it is often desirable to remove the embryo from the yolk ball and to position it flat on a glass slide. However, performing a flat mount procedure can be tedious. Therefore, successful and efficient flat mount preparation is greatly facilitated through the visual demonstration of the dissection technique, and also helped by using reagents that assist in optimal tissue handling. Here, we provide our WISH protocol for one or two-color detection of gene expression in the zebrafish embryo, and demonstrate how the flat mounting procedure can be performed on this example of a stained fixed specimen. This flat mounting protocol is broadly applicable to the study of many embryonic structures that emerge during early zebrafish development, and can be implemented in conjunction with other staining methods performed on fixed embryo samples.

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

The zebrafish embryo is an excellent model for developmental biology research. During embryogenesis, zebrafish develop with a yolk mass, which presents three-dimensional challenges for sample observation and analysis. This protocol describes how to create two-dimensional flat mount preparations of whole mount in situ (WISH) stained zebrafish embryo specimens.

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model ...

Sequential Application of Glass Coverslips to Assess the Compressive Stiffness of the Mouse Lens: Strain and Morphometric Analyses

The eye lens is a transparent organ that refracts and focuses light to form a clear image on the retina. In humans, ciliary muscles contract to deform the lens, leading to an increase in the lens&#39; optical power to focus on nearby objects, a process known as accommodation. Age-related changes in lens stiffness have been linked to presbyopia, a reduction in the lens&#39; ability to accommodate, and, by extension, the need for reading glasses. Even though mouse lenses do not accommodate or develop presbyopia, mouse models can provide an invaluable genetic tool for understanding lens pathologies, and the accelerated aging observed in mice enables the study of age-related changes in the lens. This protocol demonstrates a simple, precise, and cost-effective method for determining mouse lens stiffness, using glass coverslips to apply sequentially increasing compressive loads onto the lens. Representative data confirm that mouse lenses become stiffer with age, like human lenses. This method is highly reproducible and can potentially be scaled up to mechanically test lenses from larger animals.

Age-related increases in eye lens stiffness are linked to presbyopia. This protocol describes a simple, cost-effective method for measuring mouse lens stiffness. Mouse lenses, like human lenses, become stiffer with age. This method is precise and can be adapted for lenses from larger animals.

The eye lens is a transparent organ that refracts and focuses light to form a clear image on the retina. In humans, ciliary muscles contract to deform the ...