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.
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.
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 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).
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
2. Image analysis methodology
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-phalloidin allows us to perform peak-to-peak line scan analysis of fluorescence intensity. The major peak in the WGA channel indicates capsular surface whereas the major peak in the tdTomato/rhodamine-phalloidin channel indicates the basal region of epithelial cells. By calculating the distance between these peaks, we can obtain capsular thickness. The line scan analysis shows that capsules from a 9-week-old mice live lens had a thickness of 11.2 µm, and capsules from a 9-week-old mice fixed lens had a thickness of 12.5 µm. These observed capsule thicknesses are representative of previous findings2,4.
Whole-mount imaging of tdTomato-labeled transgenic mouse lenses (or rhodamine-phalloidin stained lenses; not shown) allows for live visualization of epithelial cell morphology. The orthogonal (XZ) projection provides a side view of the lens epithelial cell, while the planar (XY) view at the lateral regions allows for visualization of the epithelial polygonal shape. In healthy lenses, we do not observe any gaps between cells. We can calculate the average cell area by tracing a population of cells in an image and dividing the area by the number of cells within the ROI. The number of cells is determined by counting the number of Hoechst-stained nuclei in a given ROI. The analysis demonstrates an average cell area of 260 µm2 which is in keeping with previous studies2,4.
Hoechst staining of nuclei also allows for examination of lens epithelial cell nuclear morphometry in epithelial cells. The orthogonal (XZ) projections allow for a side view of nuclei. The planar (XY) view demonstrates the circular/ellipsoid shapes of the nuclei. Tracing nuclei allows for calculating individual cells' nuclear area, and other shape parameters such as circularity. The analysis demonstrates an average nuclear area of 64 µm2 with an average circularity of 0.8. Circularity values close to 1 indicate a perfect circle, whereas values approaching 0 indicate a more elongated morphology.
Equatorial epithelial cell packing, fiber cell hexagonal packing, and fiber cell widths
The planar (XY) view of the lens equatorial region demonstrates hexagon shaped and regularly packed lens epithelial cells which converge at the fulcrum. The fulcrum is where the apical tips of elongating epithelial cells constrict to form an anchor point during initial fiber cell differentiation and elongation at the equator4,13,14,15 (Figure 6B, indicated by a red dashed line). The fulcrum can be localized based on increased F-actin staining forming a continuous line separating the equatorial epithelial cells and fiber cells (Figure 6B, red dashed line). The F-actin staining at the cellular membranes also demonstrates changes in cell shape, in which cells below the fulcrum are precisely aligned and arranged in parallel rows (Figure 6). Cell nuclei are also aligned beneath the fulcrum.
To visualize lens meridional row epithelial cells and fiber cells, an image 4.5-5 µm peripheral to the fulcrum (toward the lens capsule) in the XY plane, where the basal region of meridional row cells are in view, is selected as described previously20. To measure meridional row disorder, a region of interest (ROI) is outlined (Figure 7A; yellow box). The area of the ROI in the wildtype lens image is 15,833 µm2. As there is no observable disorder, the area of disorder percentage is 0. The critical role that non-muscle myosin IIA (NMIIA) plays in cell hexagonal packing using mice with an NMIIA-E1841K mutation has been previously described20. Figure 7A shows a representative NMIIAE1841K/E1841K lens equatorial image to demonstrate meridional row cell disorder. The meridional row ROI was 20,757µm2. Next, the total area of disordered patches was traced. The total area of disorder was 3,185 µm2. The calculated percentage of disorder was determined to be 15.3% (disordered area x 100/total ROI; Figure 7B). This percent disordered area is within the range in a previous study20.
Next, an examination of hexagonal packing in wild-type meridional row cells was demonstrated20. Because F-actin is enriched around the entire perimeter of meridional row cells and at all six vertices of the basal regions of cell membranes in wild-type (i.e., NMIIA+/+) lenses, F-actin staining was used to assess cell shapes and packing organization. In representative image 1, cells were labeled and the number of adjacent cells around each cell was counted. In image 1, all ten cells have six adjacent cells, which suggests that these cells are arranged in a honeycomb packing organization (Figure 8). In contrast, eight out of 10 cells (80% of cells) have six adjacent cells in image 2 that indicate that the cells are irregularly packed (Figure 8).
Finally, to measure the fiber cell width, the peripheral fiber cells located ~10 µm inward from the fulcrum in fixed wild-type lenses labeled with rhodamine-phalloidin were examined (Figure 9A)2,4. Of note, it is also possible to measure fiber cell widths using live tdTomato mice, however, the signal from lenses that are heterozygous for tdTomato tends to be weak at the equatorial regions. Therefore, using mice that are homozygous for tdTomato is recommended as they have been found to have stronger fluorescence (not shown). The distance between the F-actin-stained cell boundaries using line-scan analysis was measured as described previously to indicate fiber cell width2,4 (Figure 9B). This analysis revealed that the average interpeak distance in wild-type lenses is 11.45 ± 2.11 µm (N=117 fiber cells from 4 different mouse lens images, Figure 9B).
Figure 1: Steps to create agarose divot to immobilize lens during imaging. (A) Using a glass bottom dish, pipette liquefied 2% agarose. (B) Flatten with a flexible cover slip and (C) when cooled, remove using fine tip forceps. (D) For the divot, create a hole using a 3 mm biopsy punch. (E) Aspirate residual agaros, rinse with PBS, wipe surface clean using a lint free tissue. (F) Carefully mount whole lens within divot. Please click here to view a larger version of this figure.
Figure 2: Steps to create agarose divot for imaging lens equatorial epithelial and fiber cells. (A) Pour 2% molten agarose in the tissue culture dish. (B) Cool the agarose at roomtemperature until it solidifies completely. (C) With a sharp blade, create a triangulardivot. (D) Put 1 mL of 1x PBS in the tissue culture dish. (E) Place the dissected lens inthe wedge with (F) the lens propped up on its equator wedged between the agarose walls (red arrow). Please click here to view a larger version of this figure.
Figure 3: Determination of lens capsule thickness. Sagittal (X, Z plane view) optical sections from reconstructions of confocal z-stacks of (A) live and (B) fixed lens capsules. Fluorescent intensity of line scan of (C) live and (D) fixed lenses demonstrate a single WGA (green) peak that corresponds to the top surface of the capsule and the basal region of epithelial cells (red) adjacent to the capsule. The distance between the two peaks is measured to quantify capsule thickness. Images acquired using a 40x oil objective with a 1 airy unit pinhole resulting in optical sections of 1.0 μm and 1.2 μm in the tdTomato/Rhodamine-Phalloidin and WGA channels, respectively. Please click here to view a larger version of this figure.
Figure 4: Quantification of epithelial cell area. (A) X, Z plane view of the lens epithelial cells marked with (B) tdTomato for cell membranes and (C) Hoechst for nuclei. (D) X, Y plane view of the middle region of lens epithelial cells. (E) tdTomato signal is used as a guide to define a region of interest corresponding to a group of cells that are in focus. (F) The area of the defined region is determined. (G) Hoechst staining for nuclei is used to determine the number of cells within the defined region. (H) The number of nuclei is counted using (I) FIJI ImageJ's multipoint tool. To calculate the average cell area, divide the total area of the defined region of interest by the total number of cells. Images acquired using a 63x oil objective with a 1 airy unit pinhole resulting in optical sections of 0.7 μm and 1.0 μm in the Hoechst and tdTomato channels, respectively. Please click here to view a larger version of this figure.
Figure 5: Determination of nuclear area and shape. (A) XY plane view of the middle region of nuclei. (B) A region of interest where nuclei are in focus was defined. Nuclei within the ROI are outlined. (C) The area and circularity of nuclei of individual cells were tabulated and averages for area and circularity were calculated. Images acquired using a 63x oil objective with a 1 airy unit pinhole resulting in an optical section of 0.7 μm in the Hoechst channel. Please click here to view a larger version of this figure.
Figure 6: Identification of the lens fulcrum. F-actin and nuclei can be used to identify the fulcrum and meridional rows. (A) XZ view shows enriched phalloidin staining for F-actin which corresponds to the fulcrum. (B) Single optical XY plane view section with the lens fulcrum in focus. Hoechst staining for nuclei shows that the nuclei above the fulcrum are irregularly packed whereas the nuclei below the fulcrum are precisely aligned (Top right). Phalloidin staining for F-actin shows that the cells above the fulcrum are irregularly shaped whereas the cells below the fulcrum are packed in parallel rows (Bottom left). The red dashed line shows the location of the fulcrum. Images acquired using a 20x objective with a 1 airy unit pinhole resulting in optical sections of 1.5 μm, 2.0 μm and 2.2 μm in the Hoechst, Rhodamine-Phalloidin, and WGA-640 channels, respectively. Please click here to view a larger version of this figure.
Figure 7: Determination of meridional row disorder. (A) Single XY plane view optical section of the wildtype and NMIIAE1841K/E1841K meridional row cells ~5 µm away from the fulcrum (toward the lens capsule). The entire meridional row cells are outlined in blue based on nuclear alignment. F-actin (red) staining in wildtype lens shows that meridional row cells are precisely aligned with no signs of disorder (0%). A representative ordered region in wildtype is outlined (orange). In lenses with disordered cells, we outline the disordered regions (yellow). (B) High magnification of ordered region from wildtype (orange box in A). (C) High magnification of disordered areas showing different types of disorder including (I) branching of rows, (II) irregular cell shape and loss of honeycomb packing, and (III) misalignment of rows. Images from NMIIAE1841K/E1841K lens show 15.3% meridional row cells disorder. Images acquired using a 20x objective with a 1 airy unit pinhole resulting in optical sections of 1.5 μm, and 2.0 μm in the Hoechst and Rhodamine-Phalloidin channels, respectively. Please click here to view a larger version of this figure.
Figure 8: Analysis of lens meridional row cell honeycomb packing. (A) Single optical XY sections of meridional row cells stained with rhodamine-phalloidin for F-actin. Low magnification images (Top panel), show the cells being evaluated (numbered in pink). Region outlined in yellow is enlarged (Bottom panel). The yellow roman numerals are the counts of adjacent cells. Image 1 shows cells that are all hexagon in shape, each with 6 adjacent cells. Image 2 has irregularities with cell number 1 and 5 having 5 and 7 adjacent cells, respectively. (B) Data is recorded and tabulated with the percentage of hexagonal cells calculated. Images acquired using a 40x objective with a 1 airy unit pinhole resulting in an optical section of 1.0 μm in the Rhodamine-Phalloidin channel. Please click here to view a larger version of this figure.
Figure 9: Analysis of fiber cell widths. (A) Single optical XY view section of the lens fiber cells ~10 µm in from the fulcrum. Cells were stained with rhodamine-phalloidin for F-actin visualization at the cell membranes. A line (pink; dash) is drawn over a number of fiber cells. (B) Representative line scan of F-actin intensities as a function of distance. The interpeak distance represents the fiber cell width. Images acquired using a 40x objective with a 1 airy unit pinhole resulting in an optical section of 1.0 μm in the Rhodamine-Phalloidin channels. Please click here to view a larger version of this figure.
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 been used in previous studies. These methods allowed us for understanding the response of the anterior capsule and cells to lens shape change or aging2,4. These methods have also been used to examine equatorial fiber cell expansion due to lens shape change or aging2,4 and to determine NMIIA’s role in establishing precise hexagonal shapes and peripheral fiber cell organization20.
Whole-mount imaging allows for high spatial resolution en face imaging of lens structure. At the anterior lens region, whole-mount imaging is advantageous to flat-mount imaging by preventing damage that may alter capsule structure integrity and/or epithelial cellular morphology. Additionally, the interface between epithelial and fiber cells is preserved. This method provides advantages over the visualization of lens sections as en face imaging affords greater spatial resolution and permits analysis of epithelial cell area and nuclear area/shape the selected region (i.e., mid-lateral, as shown here) or other regions of cells. Furthermore, the imaging methods allow for quantifying morphological features at specific points along the lens’s equatorial regions, enabling visualization of hexagonal shapes, meridional row cell packing, fulcrum, and fiber cell widths, which would not be readily achievable via imaging-stained lens sections due to lack of spatial resolution and tissue distortion that can occur during sectioning.
The outlined protocols also allow for the imaging of live lenses to track specific lens structures and cells over time. The live lens imaging protocol was instrumental in a previous study, where repeated measures analysis of capsule thickness and epithelial cell area from a subpopulation of cells from the same lens before and following lens compression to induce lens flattening was performed4. It was determined that lens compression induced a decrease in capsule thickness and an increase in epithelial cell area. Due to substantial variation in capsule thickness and epithelial cell area between individual lenses and the magnitude of effects on these parameters caused by lens compression, it would be difficult to detect differences if an independent measurement design was used (i.e., comparing individual non-compressed lenses versus individual compressed lenses). Using the live imaging methods, whole-mount imaging on live mouse lenses that endogenously express LifeACT-GFP26 to visualize F-actin in epithelial cells22, which enables tracking of F-actin reorganization in live epithelial cells, have been conducted.
While the outlined protocols were developed for imaging mouse lenses and may be adapted to visualize lens structures in other species, the staining protocols to visualize lens both anterior and equatorial peripheral structures in other rodent lenses (rat, guinea pig) as well as in other mammalian lenses (cow, macaque, and human; data not shown). Visualization of structures in larger lenses requires longer fixation (4%PFA, on ice, 4 h), blocking (1 h), and staining (overnight, 4 °C). Of note, the imaging in different species was only conducted on fixed lenses. Live imaging of lenses from other species may be challenging as the live imaging protocol uses lenses from transgenic mice that express fluorogenic proteins. Therefore, this imaging may preclude imaging lenses of other species where genetic modifications are not commonplace. However, we have been able to conduct live visualization of lens epithelial cell membranes or F-actin by prestaining mouse lenses with the lipophilic probe, FM4-64 or an F-actin binding probe, SiR-Jasplakinolide (SiR-actin), respectively (not shown). It may be possible to use such probes to image live lenses from other species. However, caution must be taken when using such probes to avoid off-target effects; for example, SiR-Jasplakinolide can lead to altered F-actin dynamics27. Another limitation of the staining methods, whether on live or fixed lenses, is that such probes have limited penetration. Therefore, imaging is limited to peripheral regions. It is possible to image inner regions (i.e., deep fiber cells) using lenses from tdTomato transgenic mice4, which again relies on the transgenic expression of fluorogenic proteins. The expression also must be high enough for bright signals in deep fiber cells.
Nevertheless, the protocols outlined have enabled effective morphological quantitation of peripheral lens features2,4,20. The quantification methodologies are effective and use open-source, publicly available FIJI ImageJ software. While manual tracing tools were used to identify regions of interest, it may be possible to automate the identification of regions of interest. Automation may be as simple as applying fluorescent thresholding to enhance contrast for FIJI ImageJ-based identification of particular structures (i.e., nuclei boundaries), or more complex machine learning algorithms to detect cell shape differences (i.e., irregular epithelial or fiber cell shapes). More complex analysis may require either specialized plugins or imaging software. Specialized plugins or software may also allow for obtaining 3D volumetric morphological measures from acquired z-stacks. Overall, while the quantification methods described are easily accessible, cost-effective, and suitable for many useful outcome measures2,4,20, the imaging protocol platform, in combination with sophisticated software analysis, could provide a wealth of additional morphological measurements for future studies.
The lens is an intricate biological tissue with specialized functions that rely upon location and depth-dependent geometries of the cells and their associated structures. Using the imaging protocols and quantification methods demonstrated here will allow for a greater understanding of how lens structures and the complex organization of the lens are established. Furthermore, the imaging protocols and quantification methods will help delineate the relationships between lens structure and functions.
The authors have nothing to disclose.
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.
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 |