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. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sequential+application+of+glass+coverslips+to+assess+the+compressive+stiffness+of+the+mouse+lens:+Strain+and+morphometric+analyses.">Sequential application of glass coverslips to assess the compressive stiffness of the mouse lens: Strain and morphometric analyses.</a> J Vis Exp. (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).

<table><tbody><tr><td>3 mm Biopsy Punch</td><td>Acuderm Inc</td><td>NC9084780</td><td></td></tr><tr><td>Agarose</td><td>Apex BioResearch Products</td><td>20-102GP</td><td></td></tr><tr><td>Antimycotic/Antibiotic</td><td>Cytiva</td><td>SV30079.01</td><td></td></tr><tr><td>Bovine Serum Albumin (Fraction V)</td><td>Prometheus</td><td>25-529</td><td></td></tr><tr><td>Delicate task wipes</td><td>Kimwipe</td><td></td><td></td></tr><tr><td>Glass bottomed dish (Fluorodish)</td><td>World Precision International</td><td>FD35-100</td><td></td></tr><tr><td>Hoescht 33342</td><td>Biotium</td><td>40046</td><td></td></tr><tr><td>Laser scanning confocal Microscope 880</td><td>Zeiss</td><td></td><td></td></tr><tr><td>MatTek Imaging Dish</td><td>MatTek Life Sciences</td><td>P35G-1.5-14</td><td></td></tr><tr><td>Paraformaldehyde&amp;nbsp;</td><td>Electron Microscopy Sciences</td><td>100503-917</td><td></td></tr><tr><td>PBS</td><td>GenClone</td><td>25-507B</td><td></td></tr><tr><td>Phenol red-free medium 199</td><td>Gibco</td><td>11043023</td><td></td></tr><tr><td>Rhodamine-Phalloidin</td><td>Thermo Fisher</td><td>00027</td><td></td></tr><tr><td>Triton X100</td><td>Sigma-Aldrich</td><td>11332481001</td><td></td></tr><tr><td>WGA-640</td><td>Biotium</td><td>CF 640R</td><td></td></tr></tbody></table>

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

Research

JoVE Journal

Biology

910 Views.  University of Delaware. We streven ernaar om de moleculaire mechanismen te bepalen die de ingewikkelde lensarchitectuur tot stand brengen en hoe deze gevestigde architectuur de lensfunctie reguleert in transparantie en lensvormverandering. Om het onderzoek in ons vakgebied vooruit te helpen, gebruiken we nieuwe beeldvormingsmethoden waarmee we de lenskenmerken met een hoge ruimtelijke resolutie kunnen visualiseren, en dit stelt ons in staat om kwantitatieve beeldanalyse van lensstructuren en cellulaire kenmerken uit...

Video: Author Spotlight: diepgaand morfometrisch onderzoek en kwantificering van de native lensstructuur met behulp van beeldvorming van de hele vatting 

High-resolution Fiber-optic Microendoscopy for in situ Cellular Imaging

Many biological and clinical studies require the longitudinal study and analysis of morphology and function with cellular level resolution. Traditionally, multiple experiments are run in parallel, with individual samples removed from the study at sequential time points for evaluation by light microscopy. Several intravital techniques have been developed, with confocal, multiphoton, and second harmonic microscopy all demonstrating their ability to be used for imaging in situ 1. With these systems, however, the required infrastructure is complex and expensive, involving scanning laser systems and complex light sources. Here we present a protocol for the design and assembly of a high-resolution microendoscope which can be built in a day using off-the-shelf components for under US$5,000. The platform offers flexibility in terms of image resolution, field-of-view, and operating wavelength, and we describe how these parameters can be easily modified to meet the specific needs of the end user.
We and others have explored the use of the high-resolution microendoscope (HRME) in in vitro cell culture 2-5, in excised 6 and living animal tissues 2,5, and in human tissues in vivo 2,7. Users have reported the use of several different fluorescent contrast agents, including proflavine 2-4, benzoporphyrin-derivative monoacid ring A (BPD-MA) 5, and fluoroscein 6,7, all of which have received full, or investigational approval from the FDA for use in human subjects. High-resolution microendoscopy, in the form described here, may appeal to a wide range of researchers working in the basic and clinical sciences. The technique offers an effective and economical approach which complements traditional benchtop microscopy, by enabling the user to perform high-resolution, longitudinal imaging in situ.

High-resolution Fiber-optic Microendoscopy for in situ Cellular Imaging

In many biological and clinical situations it is advantageous to study cellular processes as they evolve in their native microenvironment. Here we describe the assembly and use of a low-cost fiber-optic microscope which can provide real time imaging in cell culture, animal studies, and clinical patient studies.

Many biological and clinical studies require the longitudinal study and analysis of morphology and function with cellular level resolution.  ...

Bioengineering

Single Plane Illumination Module and Micro-capillary Approach for a Wide-field Microscope

A module for light sheet or single plane illumination microscopy (SPIM) is described which is easily adapted to an inverted wide-field microscope and optimized for 3-dimensional cell cultures, e.g., multi-cellular tumor spheroids (MCTS). The SPIM excitation module shapes and deflects the light such that the sample is illuminated by a light sheet perpendicular to the detection path of the microscope. The system is characterized by use of a rectangular capillary for holding (and in an advanced version also by a micro-capillary approach for rotating) the samples, by synchronous adjustment of the illuminating light sheet and the objective lens used for fluorescence detection as well as by adaptation of a microfluidic system for application of fluorescent dyes, pharmaceutical agents or drugs in small quantities. A protocol for working with this system is given, and some technical details are reported. Representative results include (1) measurements of the uptake of a cytostatic drug (doxorubicin) and its partial conversion to a degradation product, (2) redox measurements by use of a genetically encoded glutathione sensor upon addition of an oxidizing agent, and (3) initiation and labeling of cell necrosis upon inhibition of the mitochondrial respiratory chain. Differences and advantages of the present SPIM module in comparison with existing systems are discussed.

A module for single plane illumination microscopy (SPIM) is described which is easily adapted to an inverted wide-field microscope and optimized for 3-dimensional cell cultures. The sample is located within a rectangular capillary, and via a microfluidic system fluorescent dyes, pharmaceutical agents or drugs can be applied in small quantities.

A module for light sheet or single plane illumination microscopy (SPIM) is described which is easily adapted to an inverted wide-field microscope and ...

Engineering

Mitochondria and Endoplasmic Reticulum Imaging by Correlative Light and Volume Electron Microscopy

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved structures are folded into various shapes to form a dynamic network depending on the cellular conditions. Visualization of this network between mitochondria and ER has been attempted using super-resolution fluorescence imaging and light microscopy; however, the limited resolution is insufficient to observe the membranes between the mitochondria and ER in detail. Transmission electron microscopy provides good membrane contrast and nanometer-scale resolution for the observation of cellular organelles; however, it is exceptionally time-consuming when assessing the three-dimensional (3D) structure of highly curved organelles. Therefore, we observed the morphology of mitochondria and ER via correlative light-electron microscopy (CLEM) and volume electron microscopy techniques using enhanced ascorbate peroxidase 2 and horseradish peroxidase staining. An en bloc staining method, ultrathin serial sectioning (array tomography), and volume electron microscopy were applied to observe the 3D structure. In this protocol, we suggest a combination of CLEM and 3D electron microscopy to perform detailed structural studies of mitochondria and ER.

We present a protocol to study the distribution of mitochondria and endoplasmic reticulum in whole cells after genetic modification using correlative light and volume electron microscopy including ascorbate peroxidase 2 and horseradish peroxidase staining, serial sectioning of cells with and without the target gene in the same section, and serial imaging via electron microscopy.

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved ...

Biochemistry

Mapping the Emergent Spatial Organization of Mammalian Cells using Micropatterns and Quantitative Imaging

A fundamental goal in biology is to understand how patterns emerge during development. Several groups have shown that patterning can be achieved in vitro when stem cells are spatially confined onto micropatterns, thus setting up experimental models which offer unique opportunities to identify, in vitro, the fundamental principles of biological organisation.
Here we describe our own implementation of the methodology. We adapted a photo-patterning technique to reduce the need for specialized equipment to make it easier to establish the method in a standard cell biology laboratory. We also developed a free, open-source and easy to install image analysis framework in order to precisely measure the preferential positioning of sub-populations of cells within colonies of standard shapes and sizes. This method makes it possible to reveal the existence of patterning events even in seemingly disorganized populations of cells. The technique provides quantitative insights and can be used to decouple influences of the environment (e.g., physical cues or endogenous signaling), on a given patterning process.

The method presented here uses micropatterning together with quantitative imaging to reveal spatial organization within mammalian cultures. The technique is easy to establish in a standard cell biology laboratory and offers a tractable system to study patterning in vitro.

A fundamental goal in biology is to understand how patterns emerge during development. Several groups have shown that patterning can be achieved in vitro ...

Developmental Biology

Lensless On-chip Imaging of Cells Provides a New Tool for High-throughput Cell-Biology and Medical Diagnostics

Conventional optical microscopes image cells by use of objective lenses that work together with other lenses and optical components. While quite effective, this classical approach has certain limitations for miniaturization of the imaging platform to make it compatible with the advanced state of the art in microfluidics. In this report, we introduce experimental details of a lensless on-chip imaging concept termed LUCAS (Lensless Ultra-wide field-of-view Cell monitoring Array platform based on Shadow imaging) that does not require any microscope objectives or other bulky optical components to image a heterogeneous cell solution over an ultra-wide field of view that can span as large as ~18 cm2. Moreover, unlike conventional microscopes, LUCAS can image a heterogeneous cell solution of interest over a depth-of-field of ~5 mm without the need for refocusing which corresponds to up to ~9 mL sample volume. This imaging platform records the shadows (i.e., lensless digital holograms) of each cell of interest within its field of view, and automated digital processing of these cell shadows can determine the type, the count and the relative positions of cells within the solution. Because it does not require any bulky optical components or mechanical scanning stages it offers a significantly miniaturized platform that at the same time reduces the cost, which is quite important for especially point of care diagnostic tools. Furthermore, the imaging throughput of this platform is orders of magnitude better than conventional optical microscopes, which could be exceedingly valuable for high-throughput cell-biology experiments.

Lensfree on-chip imaging and characterization of cells is illustrated. This on-chip cell imaging approach provides a compact and cost-effective tool for medical diagnostics and high-throughput cell biology applications, making it especially suitable for resource poor settings.

Conventional optical microscopes image cells by use of objective lenses that work together with other lenses and optical components. While quite ...

Whole Ovary Immunofluorescence, Clearing, and Multiphoton Microscopy for Quantitative 3D Analysis of the Developing Ovarian Reserve in Mouse

Female fertility and reproductive lifespan depend on the quality and quantity of the ovarian oocyte reserve. An estimated 80% of female germ cells entering meiotic prophase I are eliminated during Fetal Oocyte Attrition (FOA) and the first week of postnatal life. Three major mechanisms regulate the number of oocytes that survive during development and establish the ovarian reserve in females entering puberty. In the first wave of oocyte loss, 30-50% of the oocytes are eliminated during early FOA, a phenomenon that is attributed to high Long interspersed nuclear element-1 (LINE-1) expression. The second wave of oocyte loss is the elimination of oocytes with meiotic defects by a meiotic quality checkpoint. The third wave of oocyte loss occurs perinatally during primordial follicle formation when some oocytes fail to form follicles. It remains unclear what regulates each of these three waves of oocyte loss and how they shape the ovarian reserve in either mice or humans.
Immunofluorescence and 3D visualization have opened a new avenue to image and analyze oocyte development in the context of the whole ovary rather than in less informative 2D sections. This article provides a comprehensive protocol for whole ovary immunostaining and optical clearing, yielding preparations for imaging using multiphoton microscopy and 3D modeling using commercially available software. It shows how this method can be used to show the dynamics of oocyte attrition during ovarian development in C57BL/6J mice and quantify oocyte loss during the three waves of oocyte elimination. This protocol can be applied to prenatal and early postnatal ovaries for oocyte visualization and quantification, as well as other quantitative approaches. Importantly, the protocol was strategically developed to accommodate high-throughput, reliable, and repeatable processing that can meet the needs in toxicology, clinical diagnostics, and genomic assays of ovarian function.

Here, we present an optimized protocol for imaging entire ovaries for quantitative and qualitative analyses using whole-mount immunostaining, multiphoton microscopy, and 3D visualization and analysis. This protocol accommodates high-throughput, reliable, and repeatable processing that is applicable for toxicology, clinical diagnostics, and genomic assays of ovarian function.

Female fertility and reproductive lifespan depend on the quality and quantity of the ovarian oocyte reserve. An estimated 80% of female germ cells ...

Immunofluorescence Staining of Membrane Interdigitations in Lens Fiber Cells

Preparation and Immunofluorescence Staining of Bundles and Single Fiber Cells from the Cortex and Nucleus of the Eye Lens

The lens is a transparent and ellipsoid organ in the anterior chamber of the eye that changes shape to finely focus light onto the retina to form a clear image. The bulk of this tissue comprises specialized, differentiated fiber cells that have a hexagonal cross section and extend from the anterior to the posterior poles of the lens. These long and skinny cells are tightly opposed to neighboring cells and have complex interdigitations along the length of the cell. The specialized interlocking structures are required for normal biomechanical properties of the lens and have been extensively described using electron microscopy techniques. This protocol demonstrates the first method to preserve and immunostain singular as well as bundles of mouse lens fiber cells to allow the detailed localization of proteins within these complexly shaped cells. The representative data show staining of the peripheral, differentiating, mature, and nuclear fiber cells across all regions of the lens. This method can potentially be used on fiber cells isolated from lenses of other species.

Author Spotlight: Advancing Lens Biomechanics Research Through a Novel Protocol for Imaging Complex Interdigitations and Protein Staining 

This protocol describes methods to prepare peripheral, mature, and nuclear eye lens fiber cells for immunofluorescence staining to study complex cell-to-cell interdigitations and the membrane architecture.

The lens is a transparent and ellipsoid organ in the anterior chamber of the eye that changes shape to finely focus light onto the retina to form a clear ...

Microsurgical Dissection and Tissue Clearing for High Resolution Intact Whole Retina and Vitreous Imaging

Neuronal and vascular structures of the retina in physiologic and pathologic conditions can be better visualized and characterized by using intact whole retina imaging techniques compared to conventional retinal flat mount preparations and sections. However, immunofluorescent imaging of intact whole retina is hindered by the opaque coatings of the eyeball, i.e., sclera, choroid, and retinal pigment epithelium (RPE) and the light scattering properties of retinal layers that prevent full thickness high resolution optical imaging. Chemical bleaching of the pigmented layers and tissue clearing protocols have been described to address these obstacles; however, currently described methods are not suitable for imaging endogenous fluorescent molecules such as green fluorescent protein (GFP) in intact whole retina. Other approaches bypassed this limitation by surgical removal of pigmented layers and the anterior segment of the eyeball allowing intact eye imaging, though the peripheral retina and hyaloid structures were disrupted. Presented here is an intact whole retina and vitreous immunofluorescent imaging protocol that combines surgical dissection of the sclera/choroid/retina pigment epithelium (RPE) layers with a modified tissue clearing method and light sheet fluorescent microscopy (LSFM). The new approach offers an unprecedented view of&#160;unperturbed vascular and neuronal elements of the retina as well as the vitreous and hyaloid vascular system in pathologic conditions.

Presented here is a protocol for intact whole retina imaging in which the outer opaque/pigmented layers of the eyeball are surgically removed, and optical clearing is applied to render retina transparent enabling the visualization of the peripheral retina and hyaloid vasculature in intact retina using light sheet fluorescent microscopy.

Neuronal and vascular structures of the retina in physiologic and pathologic conditions can be better visualized and characterized by using intact whole ...

Neuroscience

Lens-free Video Microscopy for the Dynamic and Quantitative Analysis of Adherent Cell Culture

Here, we demonstrate that lens-free video microscopy enables us to simultaneously capture the kinetics of thousands of cells directly inside the incubator and that it is possible to monitor and quantify single cells along several cell cycles. We describe the full protocol used to monitor and quantify a HeLa cell culture for 2.7 days. First, cell culture acquisition is performed with a lens-free video microscope, and then the data is analyzed following a four-step process: multi-wavelength holographic reconstruction, cell-tracking, cell segmentation and cell division detection algorithms. As a result, we show that it is possible to gather a dataset featuring more than 10,000 cell cycle tracks and more than 2 x 106 cell morphological measurements.

Lens-free video microscopy enables us to monitor cell cultures directly inside the incubator. Here we describe the full protocol used to acquire and analyze a 2.7 day long acquisition of cultured HeLa cells, leading to a dataset of 2.2 x 106 measurements of individual cell morphology and 10584 cell cycle tracks.

Here, we demonstrate that lens-free video microscopy enables us to simultaneously capture the kinetics of thousands of cells directly inside the incubator ...

Novel Protocol for Simultaneous Fluorescence and Ultrastructure Preservation

Epon Post Embedding Correlative Light and Electron Microscopy

Correlative light and electron microscopy (CLEM) is a comprehensive microscopy that combines the localization information provided by fluorescence microscopy (FM) and the context of cellular ultrastructure acquired by electron microscopy (EM). CLEM is a trade-off between fluorescence and ultrastructure, and usually, ultrastructure compromises fluorescence. Compared with other hydrophilic embedding resins, such as glycidyl methacrylate, HM20, or K4M, Epon is superior in ultrastructure preservation and sectioning properties. Previously, we had demonstrated that mEosEM can survive osmium tetroxide fixation and Epon embedding. Using mEosEM, we achieved, for the first time, Epon post embedding CLEM, which maintains the fluorescence and the ultrastructure simultaneously. Here, we provide step-by-step details about the EM sample preparation, the FM imaging, the EM imaging, and the image alignment. We also improve the procedures for identifying the same cell imaged by FM imaging during the EM imaging and detail the registration between the FM and EM images. We believe one can easily achieve Epon post embedding correlative light and electron microscopy following this new protocol in traditional EM facilities.

Author Spotlight: Advancements in Correlative Light and Electron Microscopy with Fluorescent Protein Preservation

We present a detailed protocol for Epon post-embedding correlative light and electron microscopy using a fluorescent protein called mScarlet. This method can maintain the fluorescence and the ultrastructure simultaneously. This technique is amenable to a wide variety of biological applications.

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

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