This work was supported by grant R01 EY032056 (to CC) from the National Eye Institute. The authors thank Dr. Theresa Fassel and Kimberly Vanderpool at the Scripps Research Core Microscopy Facility for their assistance with the electron microscope images.

Mice have been cared for based on an animal protocol approved by the Institutional Animal Care and Use Committee at Indiana University Bloomington. The mice used to generate representative data were control (wild-type) animals in the C57BL6/J background, female, and 8-12 weeks old. Both male and female mice can be used for this experiment, since the sex of the mice is very unlikely to affect the experiment&#39;s outcome.
1. Lens dissection and decapsulation
<ol>
	<li>Euthanize mice following the National Institutes of Health&#39;s &#34;Guide for the Care and Use of Laboratory Animals&#34; as well as animal use protocols approved by the institution. 
	NOTE: For the present study, mice were euthanized by CO2 overdose followed by cervical dislocation in accordance with an approved animal protocol (Indiana University).</li>
	<li>Enucleate the eyes from the mice using curved forceps by depressing the tissue around the eyes with one side of the forceps to displace the eye out of the socket. Next, close the forceps underneath the eye and lift to remove the eye from the socket. Transfer the eyes to fresh 1x phosphate-buffered saline (PBS) in a dissection tray.</li>
	<li>Cut the optic nerve with ultra-fine scissors as close as possible to the eyeball. Carefully insert fine-tip, straight tweezers into the eyeball through the optic nerve&#39;s exit at the posterior of the eye.</li>
	<li>Carefully insert scissors at the same location as the tweezers in step 1.3 and begin cutting an incision from the posterior toward the corneal-scleral junction. 
	NOTE: Rodent lenses occupy ~30% of their eyes. Accidental damage will occur if the tweezers or scissors are inserted too deep into the eye.</li>
	<li>Continue cutting along the corneal-scleral junction until at least half of the junction has been separated.</li>
	<li>Use tweezers to gently push on the cornea so that the lens can exit through the incision made with steps 1.4 and 1.5.</li>
	<li>Carefully remove any large pieces of tissue from the lens using fine-tip, straight tweezers. Inspect the lens to find the equatorial region.</li>
	<li>Shallowly pierce the lens using fine-tip, straight tweezers, and then remove the lens capsule. The mass of lens fiber cells will remain intact, and the lens epithelial monolayer will remain attached to the lens capsule. Discard the lens capsule.</li>
</ol>
2. Lens single fiber cell staining
<ol>
	<li>Transfer the lens fiber cell mass to a well plate with 500 &#181;L of freshly made 1% paraformaldehyde (PFA; in 1x PBS) solution and incubate the samples at 4 &#176;C overnight with gentle nutation (Figure 2A). 
	NOTE: The volumes for the solutions given are optimized for 48-well plates. If another size well-plate or tubes are used, adjust the volumes of the solutions accordingly. Fixation, blocking, and washing (1x PTX, 0.1% Triton X-100 in 1x PBS) solutions were made with 10x PBS and double distilled water (ddH2O) to a final concentration of 1x PBS.</li>
	<li>Transfer the lens fiber cells to a 60 mm dish with 1% PFA. Use a sharp scalpel to split the ball of fiber cells in half along its anterior-posterior axis (Figure 2B). Cut the halves in half again along the same axis to produce quarters. 
	NOTE: The anterior-posterior axis is easily recognized by the direction of the fiber cells in the tissue mass.</li>
	<li>Use straight tweezers to remove the nucleus region from the lens fiber cell quarters (Figure 2C). 
	NOTE: The lens nucleus region is rigid in rodent lenses, and the center of the lens will separate from the softer cortical fibers easily.</li>
	<li>Post-fix the quarters of the lens cortex region in 200 &#181;L of 1% PFA for 15 min at room temperature (RT) with gentle shaking (300 rpm on a plate shaker).</li>
	<li>Wash the tissue quarters twice in 750 &#181;L of 1x PBS for 5 min each with gentle shaking at RT.</li>
	<li>Block the samples using 200 &#181;L of blocking solution (5% serum, 0.3% Triton X-100, 1x PBS) for 1 h at RT with gentle shaking. 
	NOTE: For the representative data in this protocol, the samples were not stained with primary or secondary antibodies. After the blocking step, the samples were incubated with wheat germ agglutinin (WGA; 1:100) and phalloidin (1:100) (see Table of Materials) for 3 h at RT with gentle shaking and protection from light. Primary antibody staining has been demonstrated in previous publications22,23.</li>
	<li>Incubate with 100 &#181;L of primary antibody solution overnight at 4 &#176;C with gentle shaking. 
	NOTE: The antibodies are diluted in blocking solution. Compared to slides with tissue sections, there are more cells in this type of preparation. Primary antibody concentrations should be increased to provide ample antibodies to stain more cells. Doubling the antibody concentration from what is used on tissue sections is recommended. The same applies to secondary antibodies.</li>
	<li>Wash the tissue quarters three times with 1x PTX (0.1% Triton X-100, 1x PBS) for 5 min each at RT with gentle shaking.</li>
	<li>Incubate the fiber cells with 100 &#181;L of secondary antibody/dye solution for 3 h at RT with gentle shaking. Protect the samples from light during this and subsequent steps. 
	NOTE: WGA, phalloidin, and other fluorescent dyes can be added to the secondary antibody solution for simultaneous labeling of the cell membrane, cytoskeleton, or other organelles while also labeling the primary antibody.</li>
	<li>Wash the fiber cells four times with 1x PTX for 5 min each at RT with gentle shaking.</li>
	<li>Add one drop or 50 &#181;L of mounting media onto a plus-charged microscope slide before transferring the tissue quarters to the slide.</li>
	<li>Use tweezers to gently separate the fiber cells from each other and try to limit overlapping of the cell bundles.</li>
	<li>Gently apply a #1.5 coverslip on top of the sample in mounting media. The mounting media should spread to the edge of the coverslip; if this does not occur, add some additional mounting media to the edge of the coverslip. Aspirate away any excess mounting media around the edge of the coverslip and use nail polish to seal the edges of the coverslip on the slide. 
	&#8203;NOTE: Any type of mounting medium that is formulated for confocal microscopy can be used for these experiments.</li>
</ol>
3. Lens nucleus single fiber cell staining
<ol>
	<li>Complete the dissection outlined in section 1.</li>
	<li>Mechanically remove the cortical fibers from the ball of the lens fiber cells, leaving the lens nucleus, by gently transferring the fiber cell mass to wet, gloved fingertips and gently rolling the tissue mass. 
	NOTE: In mouse lenses, rolling the ball of lens fiber cells between gloved fingertips is an effective method for removal of the cortical fiber cells from the hard lens nucleus28,30. For lenses where this mechanical method is not effective, careful dissection or a vortexing method29 may be used to remove the cortical fiber cells.</li>
	<li>Transfer the lens nucleus to freshly made 1% PFA solution in a well-plate and incubate overnight at 4 &#176;C with gentle shaking.</li>
	<li>Transfer the sample to a 60 mm dish with 1% PFA and use a sharp scalpel to split the lens nucleus along the anterior-posterior axis. Halve the tissue samples again to produce nucleus quarters.</li>
	<li>Follow the process outlined in steps 2.4-2.13 for immunostaining and sample mounting.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65638/65638fig02v2.jpg" /> 
Figure 2: Graphical summary detailing the preparation and immunostaining of lens fiber cells. (A) This 48-well plate has been color-coded by column to demonstrate a sample plate setup for the described methods, allowing easy transfer of samples between the various immunostaining steps by gentle handling using forceps. While the representative data for this protocol is not incubated with a primary antibody, the diagram includes a column for primary antibody incubation, and the wells for washing can be reused after removing used wash buffers by aspiration. (B) After fixation of the lens fiber cell mass, the tissue is split along the anterior-posterior axis (red dashed lines) to preserve the original structure of the cells. Once the tissue mass has been halved, the samples are rotated and the halves split into quarters along the anterior-posterior axis (red dashed lines). (C) Removing the lens nucleus region (in pink) is easily done using tweezers to dig out the dense central tissue from the cortical fiber cells (in blue). Cartoon diagrams were partially created using BioRender.com and not drawn to scale. <a href="https://www.jove.com/files/ftp_upload/65638/65638fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Immunofluorescence Staining of Membrane Interdigitations in Lens Fiber Cells

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The authors have nothing to disclose.

This protocol has demonstrated the fixation, preservation, and immunostaining methods that faithfully preserve the 3D membrane morphology of bundles or singular lens fiber cells from various depths in the lens. The stained lens fibers are compared with SEM preparations that have long been used to study lens fiber cell morphology. The results show comparable membrane structures between both preparations. EM remains the gold standard for studying cell morphology, but immunolabeling is more challenging in SEM samples for lo...

The lens is a clear and ovoid tissue in the anterior chamber of the eye that is made up of two cell types, epithelial and fiber cells1 (Figure 1). There is a monolayer of epithelial cells that covers the anterior hemisphere of the lens. Fiber cells are differentiated from epithelial cells and make up the bulk of the lens. The highly specialized fiber cells undergo an elongation, differentiation, and maturation programming, marked by distinct changes in cell membrane morphology from the lens periphery to the lens center2,3,4,5,6,7,8,9,10,11,12, also known as the lens nucleus. The function of the lens to fine-focus light coming from various distances onto the retina depends on its biomechanical properties, including stiffness and elasticity13,14,15,16,17,18,19. The complex interdigitations of lens fibers have been hypothesized20,21 and recently shown to be important for lens stiffness22,23.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65638/65638fig01.jpg" /> 
Figure 1: Lens anatomy diagrams and representative scanning electron microscopy (SEM) images from lens fibers. The cartoon shows a longitudinal (anterior to posterior from top to bottom) view of the anterior monolayer of epithelial cells (shaded in light blue) and a bulk mass of lens fiber cells (white). The center of the lens (shaded in pink) is known as the nucleus and comprises highly compacted fiber cells. On the right, a cross-section cartoon reveals the elongated hexagon cell shape of lens fibers that are packed into a honeycomb pattern. Fiber cells have two broad sides and four short sides. Representative SEM images along the bottom show the complex membrane interdigitations between lens fiber cells at different depths of the lens. From the right, newly formed lens fibers at the lens periphery have small protrusions along the short sides and balls-and-sockets along the broad side (red boxes). During maturation, lens fibers develop large paddle domains that are decorated by small protrusions along the short sides (blue boxes). Mature fiber cells possess large paddle domains illustrated by small protrusions. These interlocking domains are important for lens biomechanical properties. Fiber cells in the lens nucleus have fewer small protrusions along their short sides and have complex tongue-and-groove interdigitations (purple boxes). The broad sides of the cell display a globular membrane morphology. The cartoon was modified from22,32 and not drawn to scale. Scale bar = 3 &#181;m. <a href="https://www.jove.com/files/ftp_upload/65638/65638fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The lens grows by adding shells of new fiber cells overlaid on top of previous generations of fibers24,25. Fiber cells have an elongated, hexagonal cross section shape with two broad sides and four short sides. These cells extend from the anterior to the posterior pole of the lens, and depending on the species, the lens fibers can be several millimeters in length. To support the structure of these elongated and skinny cells, specialized interdigitations along the broad and short sides create interlocking structures to maintain the lens shape and biomechanical properties. Changes in cell membrane shape during fiber cell differentiation and maturation have been extensively documented by electron microscopy (EM) studies2,3,4,5,6,7,8,9,10,20,26,27,28,29. Newly formed fiber cells have balls-and-sockets along their broad sides with very small protrusions along their short sides, while mature fibers have interlocking protrusions and paddles along their short sides. Nuclear fibers display tongue-and-groove interdigitations and globular membrane morphology. Little is known about the proteins that are required for these complex interlocking membranes. Previous studies on protein localization in fiber cells have relied on lens tissue sections, which do not allow clear visualization of the complex cell architecture.
This work has created and perfected a novel method to fix single and bundles of lens fiber cells to preserve the complex morphology and to allow immunostaining for proteins at the cell membrane and within the cytoplasm. This method faithfully preserves cell membrane architecture, comparable to data from EM studies, and allows staining with primary antibodies for specific proteins. We have previously immunostained cortical lens fibers undergoing differentiation and maturation22,23. In this protocol, there is also a new method to stain fiber cells from the lens nucleus. This protocol opens the door to understanding the mechanisms for formation and changes in membrane interdigitations during fiber cell maturation and lens nucleus compaction.

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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 

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

The lens is a transparent ellipsoid organ in the anterior chamber of the eye, and is responsible for fine focusing of light onto the retina to create a clear image. The function of the lens relies on its biomechanical properties, complex interdigitations between the lens fiber cells has recently been shown to be important for lens stiffness. While the specialized morphology of lens fiber cells has been previously described by electron microscopy, little is known about the proteins that are required for these interlocking membranes.Previous studies have relied on lens tissue sections, which do not allow clear visualization of complex cell architecture. We have created and perfected a novel technique to faithfully preserve the complex membrane interdigitations between lens fiber cells for staining and confocal microscopy imaging of specific proteins. In this protocol, there is also a new method for staining the fiber cells from the center of the lens, which is also known as the lens nucleus.This method opens the door to understanding fiber cell differentiation and maturation by studying the mechanisms of formation and changes in membrane interdigitations.

Lens fiber cells are prepared from the lens cortex (differentiating fibers and mature fibers) and the nucleus, and the cells are stained with phalloidin for F-actin and WGA for the cell membrane. A mixture of bundles of cells or single lens fibers (Figure 3) are observed and imaged. From the lens cortex, two types of cells (Figure 3A) are found. Differentiating fiber cells in the lens periphery are straight, with very small protrusions along their short sides. A...

Watch this Scientific Journal Video about Advancing Lens Biomechanics Research Through a Novel Protocol for Imaging Complex Interdigitations and Protein Staining at JoVE.com

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 ...

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JoVE Journal

Developmental Biology

Mouse Lens Dissection and Processing to Study Complex Cell-to-Cell Interdigitations and the Membrane Architecture

Begin by positioning a euthanized mouse for eye enucleation using curved forceps to press the tissue around the eye to displace the eye out of the socket. Then lift and remove the eye, and transfer it to fresh PBS in the dissection tray. Using ultra fine scissors, cut the optic nerve as close as possible to the eyeball, and carefully insert fine tip straight tweezers into the eyeball through the optic nerve's exit at the posterior of the eye.Now, insert scissors at the posterior of the eye and begin making an incision from the posterior toward the corneal scleral junction, and continue the incision until half of the junction has been separated. Gently push on the cornea so that the lens can exit through the incision. Using fine tip straight tweezers, carefully remove any large pieces of tissue from the lens.After finding the equatorial region, shallowly pierce the lens, and then remove the lens capsule. Transfer the lens fiber cells to a 60 millimeter dish with 1%paraformaldehyde. Using a sharp scalpel, split the ball of the fiber cells in half along its anterior posterior axis.And then further cut the halves along the same axis to produce quarters. Use the straight tweezers to remove the nucleus region from the lens fiber cell quarter. Next, add 200 microliters of 1%paraformaldehyde to a 48-well plate.Transfer the lens cortex to the plate and incubate for 15 minutes at room temperature with gentle shaking at 300 RPM. After blocking, add appropriate primary antibodies to the 48-well plate and transfer samples to the primary antibody solution. Incubate the samples overnight at 4 degrees Celsius with gentle shaking or mutation.The next day, wash the samples with PTX, and similarly, incubate them with secondary antibodies. After washing the samples, add one drop or 50 microliters of mounting media onto a plus charged microscope slide. Place the tissue in the mounting media.And use tweezers to gently separate the fiber cells from each other. Then, gently place a cover slip on top of the separated cells and mounting media. After removing excess media, use nail polish to seal the edges of the cover slip on the slide before confocal microscopy.After the lens dissection and lens capsule removal, transfer the fiber cell mass to wet gloved fingertips and gently roll in all directions to separate the lens nucleus. Then, transfer the lens nucleus to freshly made 1%paraformaldehyde solution in a 48-well plate and incubate overnight at 4 degrees Celsius with gentle shaking or mutation. The next day, transfer the sample to a 60 millimeter dish with 1%paraformaldehyde and use a sharp scalpel to split the lens nucleus along the anterior posterior axis in half, then into quarters.Then, postfix, block, stain, and mount the sample as previously demonstrated In these preparations, bundles of lens fiber cells with unique morphologies are found from different lens regions. The staining of the F-actin network shows enrichment at the cell membrane in differentiating and mature fibers while F-actin signals are present in the cytoplasm of nuclear fibers. Scanning electron microscopy and confocal images of differentiating fiber cells show ball and socket interdigitations with small interlocking protrusions, whereas mature fibers have paddles decorated by small protrusions.Scanning electron microscopy and confocal microscope images of nuclear lens fiber cells reveal infrequent and larger interlocking protrusions on the short sides of the cells where the cell membrane is rough and has tongue and groove interdigitations in these cells from the center of the lens.

1.1K Views.  Indiana University.  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.

Immunofluorescence Staining of Membrane Interdigitations in Lens Fiber Cells

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