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

Summary

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.

Abstract

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.

Introduction

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 organization^1,2,3,4,5,6. Surrounding the lens tissue is the capsule, a basement membrane essential for maintaining lens structure and biomechanical properties^7,8,9. 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 cells^11,12. 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 modiolus^4,13,14,15. 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 morphology^11,12,16. These fiber cells form the bulk of the lens mass^{11,12,16,17,18}.

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 features^{2,4,19,20,21,22}. 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 NMIIA^{E1841K/E1841K} mice²⁴ obtained originally from Dr. Robert Adelstein (National Heart, Lung, and Blood Institute, National Institutes of Health, Bethesda, MD). As described previously²⁰, NMIIA^{E1841K/E1841K} 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).

Protocol

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

1. Fixation of lenses for whole mount imaging
   1. 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.
   2. 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 °C. Fixed lenses can be stored for up to 5 days.
   3. 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 °C. Fixed lenses can be stored for up to 5 days.
2. Whole mount of anterior lens region (Fixed or live)
   1. For whole-mount imaging of fixed lenses, proceed to step 1.2.3.
   2. 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 °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.
   3. To stain lens capsule, F-actin, and nuclei of fixed lenses, place lenses in a 500 µL 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 °C overnight.
   4. After overnight incubation, wash the lenses 3x (5 min per wash) with 1 mL of 1x PBS. Proceed to imaging lenses.
   5. To stabilize the fixed lens for confocal imaging, create lens immobilization divots in agarose on an imaging dish as previously described²¹ (Figure 1).
      1. Heat and gently mix 2% agarose in PBS using a microwave till the solution is liquified. Pipette 250 µL 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).
      2. Remove excess agarose using a delicate task wipe (Figure 1E). Keep agarose mold hydrated with 1x PBS and maintain at 4 °C until use. Agarose molds have been successfully stored for up to 1 week.
   6. 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.
   7. To invert the lens, use curved forceps and gently rotate the lens ~180° so that the anterior region faces the objective. Acquire images using a confocal microscope.
   8. For visualization of lens capsules, acquire z-stack images using a 40x objective with a step size of 0.3 µm. 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 µm 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.
3. Lens equatorial epithelial and fiber cell staining
   1. 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 °C overnight.
   2. After overnight incubation, wash lenses 3x in 1mL 1x PBS (5 min per wash).
   3. Create lens immobilization agarose wedges as previously described^4,10.
      1. 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).
      2. 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 °C. Wedges can be kept for up to 1 week.
   4. 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.
   5. 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.
   6. Image the lens equatorial epithelial and fiber cells using a laser-scanning confocal microscope (20x objective, NA = 0.8, step size 0.5 µm and/or 40x oil objective, NA = 1.3, step size 0.4 µm). 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.

2. Image analysis methodology

1. Lens capsule thickness measurement
   1. Using z-stack images obtained with a 40x objective (0.3 µm 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.
   2. Open XZ slice (.tif) images in FIJI ImageJ software.
   3. 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 > Options > 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's top surface to beneath the epithelial cells' basal region.
   4. Save the line as a region of interest by pressing Ctrl + T.
   5. Separate channels into tabs for image, color, and split channels.
   6. Measure the intensity along the line for the capsule channel by pressing Ctrl + K.
   7. 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.
   8. Next, calculate capsule thickness by determining the distance between capsule and epithelial fluorescent intensity peaks (Figure 3C, D).
2. Cell area analysis
   1. Using z-stack images obtained using a 63x objective lens (NA = 1.4; 0.3 µm 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.
   2. Export the image as a .tif and open it in FIJI ImageJ.
   3. 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.
   4. 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.
   5. 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.
   6. 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.
3. Cellular nuclear area and shape analysis
   1. 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).
   2. Select a subpopulation of nuclei that are in focus and save a ROI.
   3. 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.
   4. Once all the nuclei are outlined, measure individual cells' nuclear area and shape (i.e., circularity) by pressing Ctrl+M.
   5. Copy and paste data into a spreadsheet and calculate the average nuclear area and circularity (Figure 5C).
4. Meridional row epithelial packing
   1. To measure meridional row disorder, acquire images using a 20x objective (0.5 µm step size).
   2. 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).
   3. Once the fulcrum is identified in 2-month-old mice, select the single optical section ~4.5-5 µm 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 µm 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.
   4. Before performing image analysis, set the scale to µm in ImageJ using step 2.2.2.
   5. 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.
   6. 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.
   7. Measure the outlined disordered area/patched by pressing Ctrl + M in FIJI ImageJ. Sum the total disordered area in a spreadsheet.
   8. 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.
5. Meridional row number of neighboring cells
   1. To measure number of neighboring cells, use F-actin-stained images acquired with 40x oil objective (0.4 µm step size).
   2. 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.
   3. 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.
6. Image analysis of equatorial fiber cells
   1. Take z-stack confocal images of rhodamine phalloidin stained lenses with a 20x objective (0.5 µm step size).
   2. 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 µm inward from the fulcrum in the XY plane (Figure 9A).
   3. Export raw images to FIJI ImageJ. In FIJI ImageJ, draw a line (usually ~300-400 µm long) across several adjacent fiber cells to measure the distance between F-actin-stained peaks (line scan analysis; Figure 9A; pink line).
   4. 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.

Results

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...

Discussion

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 ...

Materials

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