This work was supported by National Eye Institute Grant R01 EY017724 (VMF) and National Institute of Arthritis and Musculoskeletal and Skin Diseases Grant K99 AR066534 (DSG).

All animal procedures were performed in accordance with recommendations in the Guide for the Care and Use of Laboratory Animals by the National Institutes of Health and under an approved protocol by the Institutional Animal Care and Use Committee at The Scripps Research Institute.
1. Lens Dissection
<ol>
	<li>Euthanize mice according to recommendations in the National Institutes of Health &#34;Guide for the Care and Use of Laboratory Animals&#34; and approved institution animal use protocols.&#160;</li>
	<li>Enucleate the eye from mice using curved forceps. Depress the tissue around the eye with the forceps to bring the eye out of the socket, and then pluck the eye from the socket with the forceps. Transfer eyes to fresh 1x phosphate buffered saline (PBS) in dissection dish.</li>
	<li>Cut off the optic nerve as close to the eyeball as possible. Gently and carefully insert fine straight tweezers into the eyeball through the hole where the optic nerve exits the posterior.</li>
	<li>Carefully make an incision with scissors into the eyeball from the posterior to the edge of the cornea. Rodent lenses occupy ~30% of the eye. Make these incisions carefully, and do not insert tweezers or scissors too deep into the eye to avoid damaging the lens.</li>
	<li>Cut along the junction between the cornea and sclera at least half way around the eyeball.</li>
	<li>Gently push on the cornea to remove the lens from the eye through the opening made in steps 1.4 and 1.5.</li>
	<li>Use fine tip straight forceps to carefully remove any large debris that is still attached to the lens. Visually inspect the lens for any damage before proceeding to the stiffness measurements.</li>
</ol>
2. Stiffness Measurements
<ol>
	<li>Weigh at least 10 coverslips from the same box using an analytical balance. Find the average weight of the coverslips. For consistency, use the same box of coverslips for all experiments. Pre-wet the coverslips and right-angle mirror in 1x PBS at room temperature for at least 2 hr before starting experiments.</li>
	<li>Fill the measurement chamber (see Figure 1) with 65 - 75 ml of 1x PBS. The measurement chamber was made out of Plexiglas by an in-house machine shop, and divots in the chamber were made by a drill press set to the desired depth with an appropriate drill bit. &#160;Lenses remain transparent in 1x PBS at room temperature for the duration of mechanical testing.</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/53986/53986fig1.jpg" /> 
Figure 1: Stiffness Measurement Chamber. A photo showing the dimensions of the custom-made stiffness measurement chamber with a variety of divots of different depths and shapes. The round divots that are 200 &#181;m or 300 &#181;m deep (yellow arrowheads) are used for the measurements on mouse lenses. Divots are 2 mm in diameter and ~13 - 14 mm from the edge of the chamber. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53986/53986fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="3">
	<li>Place right-angle mirror into the chamber at a constant distance from the divot that will be used to hold the lens. Make sure the mirror does not move during the experiment.</li>
	<li>Transfer dissected lenses to measurement chamber carefully with seizing forceps or curved forceps.</li>
	<li>Take a top-view picture of the unloaded lens from directly overhead. Take mouse lens photos at 30X magnification with illumination from dissecting microscope (bottom) and a fiber optic light source on the left and right sides. Set the fiber optic power supply to 80% of the maximum light intensity. Adjust the power supply output based on ambient lighting, user preference and picture quality as needed.</li>
	<li>Take a side-view picture of the unloaded lens, which can be seen through the right-angle mirror. If camera is not calibrated, take a picture of the mirror edge in focus. The mirror edge is 5 mm long, and this measurement can later be used to determine the pixels/mm and serve as a scale bar in the images.</li>
	<li>Place lens into the divot, and confirm that the lens is seated securely and straight in the divot. Take a picture of the lens before loading. The lens should be resting in the divot on its anterior or posterior pole.</li>
	<li>Place 1 coverslip gently onto the lens. Wait 2 min to permit creep, and take another side-view picture of the loaded lens.</li>
	<li>Continue adding coverslips as in step 2.8 and taking side-view pictures after the addition of each coverslip as in step 2.8 until a total of 10 coverslips are applied.</li>
	<li>Remove all coverslips. Wait 2 min, and take a side-view picture of the lens (inside and outside of the divot) after removing all coverslips.</li>
</ol>
3. Lens Nucleus Measurement
<ol>
	<li>To determine the lens nucleus size, move the lens to a clean Petri dish filled with 1x PBS.</li>
	<li>Gently decapsulate the lens using fine straight forceps.</li>
	<li>Slough off the cortical fiber cells by rolling the lens between gloved fingers. The remaining lens nucleus will feel like a hard marble. Use this procedure to isolate the nucleus on adult lenses starting at 1 month of age. Since the isolated nucleus is a rigid body, further mechanical testing of the lens nucleus cannot be performed using this described method.</li>
	<li>Gently rinse the lens nucleus in 1x PBS in the Petri dish.</li>
	<li>Place the lens nucleus back into the measurement chamber (not in the divot), and take an image of the lens nucleus through the right-angle mirror.</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/53986/53986fig2.jpg" /> 
Figure 2: A Mouse Lenses Compressed by Coverslips. (A) Schematic and (B) photograph of the experimental setup showing a 2-month-old mouse lens in a 200-&#181;m-deep divot in the measurement chamber filled with 1x PBS. A right angle mirror and a digital camera mounted on a dissection microscope were used to collect images of the lens during compression by coverslips. (C) Photos of sagittal views of a 2-month-old wild-type lens compressed by successively increasing numbers of coverslips provided the raw data for measuring axial and equatorial diameters and calculating axial and equatorial strains during coverslip-based compression testing. A reflection of the lens can sometimes be seen in the coverslips (most clearly visible in the 1 coverslip image). When making measurements, ignore the reflection and measure to the apex of the lens.&#160;(D) Photos of sagittal views of the 2-month-old wild-type lens post-compression and the isolated lens nucleus. The post-compression lens and isolated nucleus are sitting outside of the divot. Scale bars, 1 mm. This figure is modified from Gokhin, et al. PloS one, 201219. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53986/53986fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Image Analysis
<ol>
	<li>Measure equatorial and axial diameters of lenses before loading and after each loading step using ImageJ or similar software. Measure the diameter of each lens nucleus. The lens nucleus is nearly spherical so a measurement at any orientation will suffice19,21.</li>
	<li>Correct the lens axial diameters by adding the depth of the divot used. In the measurement chamber, the divot obscured 200 &#181;m (2-month-old mouse lenses) or 300 &#181;m (4-month-old and 8-month-old mouse lenses) of the axial thickness of the lens.</li>
	<li>Calculate the axial and equatorial strains from the lens diameter measurements using the equation, &#949; = (d - d0) / d0, where &#949; is strain, d is the axial or equatorial diameter at a given load, and d0 is the corresponding axial or equatorial diameter at zero load.</li>
	<li>Plot the axial and equatorial strains as functions of the imposed load (in mg).</li>
	<li>Plot the axial, equatorial and nuclear diameters. Calculate and plot the lens aspect ratio by dividing the axial diameter by the equatorial diameter.</li>
	<li>Calculate and plot the lens volume using the equation, volume = 4/3 &#215; &#960; &#215; rE2&#160;&#215; rA, where rE is the equatorial radius and rA is the axial radius measured from the picture taken in step 2.6. This equation assumes the lens is an oblate spheroid (ellipsoid)1,22.</li>
	<li>Calculate and plot the nuclear volume using the equation, volume = 4/3 &#215; &#960; &#215; rN3, where rN is the radius of the lens nucleus as measured from the picture taken in step 3.5. This equation assumes the lens nucleus is a sphere19,21.</li>
	<li>Calculate and plot the nuclear fraction as the ratio of the nuclear volume to the lens volume.</li>
</ol>

<ol>
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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Presbyopia+and+the+optical+changes+in+the+human+crystalline+lens+with+age.">Presbyopia and the optical changes in the human crystalline lens with age.</a> Vision Res. 38 (2), 209-229 (1998).</li><li>Pau, H., Kranz, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+increasing+sclerosis+of+the+human+lens+with+age+and+its+relevance+to+accommodation+and+presbyopia.">The increasing sclerosis of the human lens with age and its relevance to accommodation and presbyopia.</a> Graefes Arch Clin Exp Ophthalmol. 229 (3), 294-296 (1991).</li><li>Hollman, K. W., O'Donnell, M., Erpelding, T. 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S., 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=Intermediate+filaments+regulate+tissue+size+and+stiffness+in+the+murine+lens.">Intermediate filaments regulate tissue size and stiffness in the murine lens.</a> Invest Ophthalmol Vis Sci. 52 (6), 3860-3867 (2011).</li><li>Kuszak, J. R., Mazurkiewicz, M., Zoltoski, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Computer+modeling+of+secondary+fiber+development+and+growth:+I.+Nonprimate+lenses.">Computer modeling of secondary fiber development and growth: I. Nonprimate lenses.</a> Mol Vis. 12, 251-270 (2006).</li><li>Scarcelli, G., Kim, P., Yun, S. 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The authors have nothing to disclose.

There are several key considerations when using this method to measure lens stiffness. First, the coverslips are applied to the lens at a slightly oblique angle (8 - 8.5&#176;) with respect to the bottom of the chamber (&#952;). This will apply a very small component of the load equatorially rather than axially. However, this equatorial load is considered negligible because sin &#952; &#8776; 0.119. If this method is adapted for larger lenses, the angle of the coverslips to the bottom of the chamber would need to be measured to determine whether equatorial load should be factored into strain calculations. Second, it is crucial to allow the lens to equilibrate after the addition of each coverslip. The waiting period of 2 minutes allows time-dependent deformation (i.e., creep) to occur, such that pictures are only taken when the lens is at an equilibrium shape19. Third, this protocol is optimized for measuring compressive strain on mouse lenses across a wide dynamic range of loads. In pilot studies, an applied load of 1,293 mg (i.e., ten 18 x 18 mm coverslips) compressed the mouse lens to a maximum strain, beyond which increased loads do not cause appreciable further deformation. This is due to the presence of the rigid lens nucleus that does not deform appreciably under compression19. Fourth, this protocol avoids irreversible tissue damage. In previously published experiments, no changes in the mechanical properties of mouse lenses were observed upon repeat loading, suggesting that this method does not damage the lens19. When mechanically testing lenses of a different species or mutant lenses, pilot tests should be done to determine the maximum load needed for maximum strain by repeating steps 2.8 - 2.10 and comparing the strain-load curves, lens diameters and lens volumes between the first and second loading. Finally, this method provides an empirical measurement of the stiffness of the whole lens and cannot differentiate the contributions of different cell types (epithelial cells, cortical fibers, nuclear fibers) and the lens capsule to whole-lens mechanical properties.
The axial and equatorial strains are plotted here as functions of the imposed load. Previous studies have quantified mouse lens stiffness19,21, resilience21 or the change in diameter18 upon applied load. Strain is a dimensionless quantity that allows direct comparison between lenses of different sizes. Note that equatorial extension (positive strain) occurs concurrently with the applied axial compression (negative strain) due to conservation of lens volume (i.e., Poisson effect). However, the observed equatorial strains were much smaller in absolute magnitude than the axial strains, indicating that this method has less resolution to detect small changes in equatorial strain as compared to axial strain.
In summary, this simple method with an easily assembled device for measuring mouse lens stiffness can be applied generally and widely in lens research to better understand how mutations in proteins, pathologies and/or aging affect lens stiffness. While mouse lenses do not accommodate, this method can still elucidate the proteins and age-related modifications that contribute to increased lens stiffness, and potentially contribute new knowledge to developing novel treatments for presbyopia.

The lens is a transparent and avascular organ in the anterior chamber of the eye that is responsible for fine focusing of light onto the retina. A clear basement membrane, called the lens capsule, surrounds a bulk of elongated fiber cells covered by an anterior monolayer of epithelial cells1,2. Life-long growth of the lens depends on the continuous proliferation and differentiation of epithelial cells at the lens equator into new fiber cells that are added onto previous generations of fiber cells in a concentric manner2. Over time, lens fiber cells undergo compaction, resulting in a rigid center in the middle of the lens called the nucleus1. Accommodation, defined as a dioptric change in the optical power of the eye, occurs in humans when the ciliary muscles contract to deform the lens and allow a true increase in optical power to focus on near objects3-5. In the unaccommodated eye, the lens is held in a relatively flattened state due to tension from zonular fibers. When the ciliary muscles contract, the tension on the lens is released, leading to decreased lens equatorial diameter and increased axial thickness. Age-related changes in the lens cause presbyopia, a progressive loss of lens accommodation, which leads to the need for reading glasses. 
Several studies have linked presbyopia to age-related increase in the intrinsic stiffness of the lens6-11. Stiffness is defined as the resistance of an elastic object to deform under applied load. A variety of methods have been used to examine stiffness of human lenses, including spin compression12-14, actuator compression15, probe indentation16, dynamic mechanical analysis 6,10 and bubble-based acoustic radiation force17. While mouse lenses do not accommodate or develop presbyopia, mouse models for lens pathologies are valuable tools because mice are less expensive than larger animals, well characterized genetically and undergo accelerated age-related changes due to rapid aging. A handful of studies have examined mouse lens stiffness with compression methods and demonstrated changes in lens stiffness due to aging or targeted genetic disruptions18-21. Thus, mouse lenses are good models for studying age-related changes in lens stiffness. 
This protocol describes a simple and inexpensive, yet precise and reproducible, compression method for determining mouse lens stiffness based on application of glass coverslips onto the lens in conjunction with photographing the lens through a dissection microscope and mirror. This method yields robust strain and morphometric data with an easily fabricated and assembled apparatus. The representative results confirm that mouse lenses increase in stiffness with age.

<table border="0" cellpadding="0" cellspacing="0" width="811">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:361px;">Fine tip straight forceps</td>
			<td style="width:139px;">Fine Scientific Tools</td>
			<td style="width:119px;">11252-40</td>
			<td style="width:193px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Microdissection scissors, straight edge</td>
			<td style="width:139px;">Fine Scientific Tools</td>
			<td style="width:119px;">15000-00</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Curved forceps</td>
			<td style="width:139px;">Fine Scientific Tools</td>
			<td style="width:119px;">11272-40</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Seizing forceps</td>
			<td style="width:139px;">Hammacher</td>
			<td style="width:119px;">HSC 702-93</td>
			<td>Optional</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Dissection dish</td>
			<td style="width:139px;">Fisher Scientific</td>
			<td style="width:119px;">12565154</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">60mm petri dish</td>
			<td style="width:139px;">Fisher Scientific</td>
			<td style="width:119px;">0875713A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">1X phosphate buffered saline (PBS)</td>
			<td style="width:139px;">Life Technologies</td>
			<td style="width:119px;">14190</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">18mm x 18mm glass coverslips</td>
			<td style="width:139px;">Fisher Scientific</td>
			<td style="width:119px;">12-542A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Measurement chamber with divots to hold lenses</td>
			<td style="width:139px;"></td>
			<td style="width:119px;"></td>
			<td>Custom-made (see Figure 1)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Right-angle mirror</td>
			<td style="width:139px;">Edmund Optics</td>
			<td style="width:119px;">45-591</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Light source</td>
			<td style="width:139px;">Schott/Fostec</td>
			<td style="width:119px;">8375</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Illuminated dissecting microscope</td>
			<td style="width:139px;">Olympus</td>
			<td style="width:119px;">SZX-ILLD100</td>
			<td>With SZ-PT phototube</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Digital camera</td>
			<td style="width:139px;">Nikon</td>
			<td style="width:119px;">Coolpix 990</td>
			<td></td>
		</tr>
	</tbody>
</table>

sequential application of glass coverslips to assess the compressive stiffness of the mouse lens: strain and morphometric analyses

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

The stiffness and dimensions of 2-, 4- and 8-month-old mouse lenses were measured. Mice were all wild-type animals on a pure C57BL6 strain background obtained from the TSRI Animal Breeding Facility, and each lens was loaded with 1 to 10 coverslips. The axial and equatorial strains were calculated as a function of applied load by measuring the axial and equatorial diameters of the lens after the addition of each coverslip, and then normalizing each change in diameter to the corresponding unloaded diameter. Eight lenses from each age were tested, and the results are expressed as the average &#177; standard error. As shown previously19, axial strain is a logarithmic function of applied load (Figure 3A). There was a statistically significant age-dependent decrease in axial and equatorial strains under the maximum applied load (Figure 3), indicating that the mouse lens stiffens with age. Strain measurements were highly reproducible across lenses of the same age, as demonstrated by the small standard errors.
The image data collected during this experiment were also used to determine several other lens morphological characteristics (Figure 4). As expected, axial and equatorial diameters and lens volume increased with age (Figure 4A, 4B and 4D). The aspect ratio indicates that the lens has a slightly larger equatorial diameter than axial diameter, and this parameter did not change with age (Figure 4C).&#160;The diameter, volume and fraction of the lens nucleus increased with age (Figure 4E, 4F and 4G). These results suggest that the lens nucleus remodels to increase in relative size as the lens ages.
This data show that mouse lenses increase in stiffness with age, similar to changes in aging human lenses9,15. These data also agree with previous observations made using a similar method18 and by Brillouin optical microscopy23 that mouse lenses increase in stiffness with age. Two other studies have used the described method to show that tropomodulin-1, an actin pointed-end capping protein, CP49, a beaded intermediate filament protein, and aquaporin 0 are needed to maintain lens stiffness19,20. With this method, the plethora of mouse models for lens pathologies and the accelerated aging of mice can be used to understand lens stiffness changes due to genetic variation and/or aging. This method can also be adapted for lenses from other species. The dimensions of the chamber used for these experiments are optimized for mouse lenses, but can easily be scaled up for lenses from larger species. In the future, it would be interesting to determine whether lens size scales with lens stiffness across species.
At the lowest load (1 coverslip, 129.3 mg), there was no obvious difference in axial and equatorial strains between 2-, 4- and 8-month-old lenses. The increase in lens stiffness with age became more apparent at higher loads. It is possible that the outermost cortex of the lens (~100 &#181;m) that is compressed by first coverslip does not change in stiffness with age, or that this method is unable to distinguish small changes in strain at low loads. The morphometrics data indicate that increases in both the whole lens and nuclear volume correlate with increased lens stiffness as a function of age.
<img alt="Figure 3" src="/files/ftp_upload/53986/53986fig3.jpg" /> 
Figure 3: Axial and Equatorial Strain-load Curves for 2-, 4- and 8-month-old (2M, 4M and 8M) Wild-type Mouse Lenses. (A) Axial compressive strain plotted as a function of the applied load (mg). (B) Equatorial compressive strain plotted as a function of the applied load (mg). Four- and 8-month-old lenses exhibited less strain than 2-month-old lenses at equivalent maximum loads, indicating an increase in lens stiffness with age. **, p&#160;&#60;0.01. Note that the Y-axis is different between (A) and (B). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53986/53986fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/53986/53986fig4.jpg" /> 
Figure 4: Morphological Characteristics of 2-, 4- and 8-month-old (2M, 4M and 8M) Wild-type Mouse Lenses. (A) Axial diameter and (B) equatorial diameter progressively increase with age. The lens aspect ratio (C) shows that mouse lenses are slightly wider at the lens equator. Lens volume (D), nuclear diameter (E), nuclear volume (F) and nuclear fraction (G) progressively increase with age. **, p&#160;&#60;0.01. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53986/53986fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Sequential Application of Glass Coverslips to Assess the Compressive Stiffness of the Mouse Lens: Strain and Morphometric Analyses at JoVE.com

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

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

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

sequential-application-glass-coverslips-to-assess-compressive

Nanyang Technological University

Research

JoVE Journal

Biology

7.2K Views.  The Scripps Research Institute. Age-related increases in eye lens stiffness are linked to presbyopia. This protocol describes a simple, cost-effective method for measuring mouse lens stiffness. Mouse lenses, like human lenses, become stiffer with age. This method is precise and can be adapted for lenses from larger animals.

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

Lens Transplantation in Zebrafish and its Application in the Analysis of Eye Mutants

The lens plays an important role in the development of the optic cup[1,2]. Using the zebrafish as a model organism, questions regarding lens development can be addressed. The zebrafish is useful for genetic studies due to several advantageous characteristics, including small size, high fecundity, short lifecycle, and ease of care. Lens development occurs rapidly in zebrafish. By 72 hpf, the zebrafish lens is functionally mature [3]. Abundant genetic and molecular resources are available to support research in zebrafish. In addition, the similarity of the zebrafish eye to those of other vertebrates provides basis for its use as an excellent animal model of human defects[4-7]. Several zebrafish mutants exhibit lens abnormalities, including high levels of cell death, which in some cases leads to a complete degeneration of lens tissues [8]. To determine whether lens abnormalities are due to intrinsic causes or to defective interactions with the surrounding tissues, transplantation of a mutant lens into a wild-type eye is performed. Using fire-polished metal needles, mutant or wild-type lenses are carefully dissected from the donor animal, and transferred into the host. To distinguish wild-type and mutant tissues, a transgenic line is used as the donor. This line expresses membrane-bound GFP in all tissues, including the lens. This transplantation technique is an essential tool in the studies of zebrafish lens mutants.

Lens development involves interactions with other tissues. Several zebrafish eye mutants are characterized by an abnormally small lens size. Here we demonstrate a lens transplantation experiment to determine whether this phenotype is due to intrinsic causes or defective interactions with tissues that surround the lens.

The lens plays an important role in the development of the optic cup[1,2]. Using the zebrafish as a model organism, questions regarding lens development ...

Purification of the M. magneticum Strain AMB-1 Magnetosome Associated Protein MamA&Delta;41

Magnetotactic bacteria comprise a diverse group of aquatic microorganisms that are able to orientate themselves along geomagnetic fields. This behavior is believed to aid their search for suitable environments (1). This capability is conferred by the magnetosome, a subcellular organelle that consists of a linear-chain assembly of lipid vesicles each able to biomineralize and enclose a ~50-nm crystal of magnetite or greigite. A principle component of the magnetosome that was shown to be required for the formation of functional vesicles is MamA. MamA is a highly abundant magnetosome-associated protein which is one of the most characterized magnetosome-associated proteins in vivo (2-6). This article focuses on the purification of MamA, which despite being studied in vivo, no clear functional or structural details have been identified for it. Bioinformatics analysis suggested that MamA is a tetra-tricopeptide repeat (TPR) containing protein. TPR is a structural motif found as such or forming part of a bigger fold in a wide range of proteins, it serves as a template for protein-protein interactions and mediates multi-protein complexes (7). TPRs are involved in many crucial tasks in eukaryotic cell organelle processes and many bacterial pathways (8-14). In order to understand MamA, a unique TPR containing protein, highly purified protein is required as a first step. In this article, we present the purification protocol for a stable MamA deletion mutant (MamA&Delta;41) from M. magneticum AMB-1.

MamA is a unique Magnetosome associated protein which was shown to be involved in magnetosome activation. Here we present the purification protocol of MamA deletion mutant (MamA&Delta;41) from M. magneticum AMB-1.

Magnetotactic bacteria comprise a diverse group of aquatic microorganisms that are able to orientate themselves along geomagnetic fields. This behavior is ...

Evisceration of Mouse Vitreous and Retina for Proteomic Analyses

While the mouse retina has emerged as an important genetic model for inherited retinal disease, the mouse vitreous remains to be explored. The vitreous is a highly aqueous extracellular matrix overlying the retina where intraocular as well as extraocular proteins accumulate during disease.1-3 Abnormal interactions between vitreous and retina underlie several diseases such as retinal detachment, proliferative diabetic retinopathy, uveitis, and proliferative vitreoretinopathy.1,4 The relative mouse vitreous volume is significantly smaller than the human vitreous (Figure 1), since the mouse lens occupies nearly 75% of its eye.5 This has made biochemical studies of mouse vitreous challenging. In this video article, we present a technique to dissect and isolate the mouse vitreous from the retina, which will allow use of transgenic mouse models to more clearly define the role of this extracellular matrix in the development of vitreoretinal diseases.

The dissection technique illustrates evisceration of the vitreous, retina, and lens from the mouse eye, separation by centrifugation, and characterization with protein assays.

While the mouse retina has emerged as an important genetic model for inherited retinal disease, the mouse vitreous remains to be explored. The vitreous is ...

Biophysical Assays to Probe the Mechanical Properties of the Interphase Cell Nucleus: Substrate Strain Application and Microneedle Manipulation

In most eukaryotic cells, the nucleus is the largest organelle and is typically 2 to 10 times stiffer than the surrounding cytoskeleton; consequently, the physical properties of the nucleus contribute significantly to the overall biomechanical behavior of cells under physiological and pathological conditions. For example, in migrating neutrophils and invading cancer cells, nuclear stiffness can pose a major obstacle during extravasation or passage through narrow spaces within tissues.1 On the other hand, the nucleus of cells in mechanically active tissue such as muscle requires sufficient structural support to withstand repetitive mechanical stress. Importantly, the nucleus is tightly integrated into the cellular architecture; it is physically connected to the surrounding cytoskeleton, which is a critical requirement for the intracellular movement and positioning of the nucleus, for example, in polarized cells, synaptic nuclei at neuromuscular junctions, or in migrating cells.2 Not surprisingly, mutations in nuclear envelope proteins such as lamins and nesprins, which play a critical role in determining nuclear stiffness and nucleo-cytoskeletal coupling, have been shown recently to result in a number of human diseases, including Emery-Dreifuss muscular dystrophy, limb-girdle muscular dystrophy, and dilated cardiomyopathy.3 To investigate the biophysical function of diverse nuclear envelope proteins and the effect of specific mutations, we have developed experimental methods to study the physical properties of the nucleus in single, living cells subjected to global or localized mechanical perturbation. Measuring induced nuclear deformations in response to precisely applied substrate strain application yields important information on the deformability of the nucleus and allows quantitative comparison between different mutations or cell lines deficient for specific nuclear envelope proteins. Localized cytoskeletal strain application with a microneedle is used to complement this assay and can yield additional information on intracellular force transmission between the nucleus and the cytoskeleton. Studying nuclear mechanics in intact living cells preserves the normal intracellular architecture and avoids potential artifacts that can arise when working with isolated nuclei. Furthermore, substrate strain application presents a good model for the physiological stress experienced by cells in muscle or other tissues (e.g., vascular smooth muscle cells exposed to vessel strain). Lastly, while these tools have been developed primarily to study nuclear mechanics, they can also be applied to investigate the function of cytoskeletal proteins and mechanotransduction signaling.

We present two independent, microscope-based tools to measure the induced nuclear and cytoskeletal deformations in single, living adherent cells in response to global or localized strain application. These techniques are used to determine nuclear stiffness (i.e., deformability) and to probe intracellular force transmission between the nucleus and the cytoskeleton.

In most eukaryotic cells, the nucleus is the largest organelle and is typically 2 to 10 times stiffer than the surrounding cytoskeleton; consequently, the ...

Generation of Mice Derived from Induced Pluripotent Stem Cells

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also produce a source of patient-matched cells for regenerative therapies. iPSCs may be generated using multiple protocols and derived from multiple cell sources. Once generated, iPSCs are tested using a variety of assays including immunostaining for pluripotency markers, generation of three germ layers in embryoid bodies and teratomas, comparisons of gene expression with embryonic stem cells (ESCs) and production of chimeric mice with or without germline contribution2. Importantly, iPSC lines that pass these tests still vary in their capacity to produce different differentiated cell types2. This has made it difficult to establish which iPSC derivation protocols, donor cell sources or selection methods are most useful for different applications.
The most stringent test of whether a stem cell line has sufficient developmental potential to generate all tissues required for survival of an organism (termed full pluripotency) is tetraploid embryo complementation (TEC)3-5. Technically, TEC involves electrofusion of two-cell embryos to generate tetraploid (4n) one-cell embryos that can be cultured in vitro to the blastocyst stage6. Diploid (2n) pluripotent stem cells (e.g. ESCs or iPSCs) are then injected into the blastocoel cavity of the tetraploid blastocyst and transferred to a recipient female for gestation (see Figure 1). The tetraploid component of the complemented embryo contributes almost exclusively to the extraembryonic tissues (placenta, yolk sac), whereas the diploid cells constitute the embryo proper, resulting in a fetus derived entirely from the injected stem cell line.
Recently, we reported the derivation of iPSC lines that reproducibly generate adult mice via TEC1. These iPSC lines give rise to viable pups with efficiencies of 5-13%, which is comparable to ESCs3,4,7 and higher than that reported for most other iPSC lines8-12. These reports show that direct reprogramming can produce fully pluripotent iPSCs that match ESCs in their developmental potential and efficiency of generating pups in TEC tests. At present, it is not clear what distinguishes between fully pluripotent iPSCs and less potent lines13-15. Nor is it clear which reprogramming methods will produce these lines with the highest efficiency. Here we describe one method that produces fully pluripotent iPSCs and "all- iPSC" mice, which may be helpful for investigators wishing to compare the pluripotency of iPSC lines or establish the equivalence of different reprogramming methods.

Generating induced pluripotent stem cell (iPSC) lines produces lines of differing developmental potential even when they pass standard tests for pluripotency. Here we describe a protocol to produce mice derived entirely from iPSCs, which defines the iPSC lines as possessing full pluripotency1.

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also ...

Gap Junctional Intercellular Communication: A Functional Biomarker to Assess Adverse Effects of Toxicants and Toxins, and Health Benefits of Natural Products

This protocol describes a scalpel loading-fluorescent dye transfer (SL-DT) technique that measures intercellular communication through gap junction channels, which is a major intercellular process by which tissue homeostasis is maintained. Interruption of gap junctional intercellular communication (GJIC) by toxicants, toxins, drugs, etc. has been linked to numerous adverse health effects. Many genetic-based human diseases have been linked to mutations in gap junction genes. The SL-DT technique is a simple functional assay for the simultaneous assessment of GJIC in a large population of cells. The assay involves pre-loading cells with a fluorescent dye by briefly perturbing the cell membrane with a scalpel blade through a population of cells. The fluorescent dye is then allowed to traverse through gap junction channels to neighboring cells for a designated time. The assay is then terminated by the addition of formalin to the cells. The spread of the fluorescent dye through a population of cells is assessed with an epifluorescence microscope and the images are analyzed with any number of morphometric software packages that are available, including free software packages found on the public domain. This assay has also been adapted for in vivo studies using tissue slices from various organs from treated animals. Overall, the SL-DT assay can serve a broad range of in vitro pharmacological and toxicological needs, and can be potentially adapted for high throughput set-up systems with automated fluorescence microscopy imaging and analysis to elucidate more samples in a shorter time.

This protocol describes a scalpel loading-fluorescent dye transfer technique that measures intercellular communication through gap junction channels. Gap junctional intercellular communication is a major cellular process by which tissue homeostasis is maintained and disruption of this cell signaling has adverse health effects.

This protocol describes a scalpel loading-fluorescent dye transfer (SL-DT) technique that measures intercellular communication through gap junction ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

A Morphometric and Cellular Analysis Method for the Murine Mandibular Condyle

The temporomandibular joint (TMJ) has the capacity to adapt to external stimuli, and loading changes can affect the position of condyles, as well as the structural and cellular components of the mandibular condylar cartilage (MCC). This manuscript describes methods for analyzing these changes and a method for altering the loading of the TMJ in mice (i.e., compressive static TMJ loading). The structural evaluation illustrated here is a simple morphometric approach that uses the Digimizer software and is performed in radiographs of small bones. In addition, the analysis of cellular changes leading to alterations in collagen expression, bone remodeling, cell division, and proteoglycan distribution in the MCC is described. The quantification of these changes in histological sections - by counting the positive fluorescent pixels using image software and measuring the distance mapping and stained area with Digimizer - is also demonstrated. The methods shown here are not limited to the murine TMJ, but could be used on additional bones of small experimental animals and in other regions of endochondral ossification.

This manuscript presents methods for analyzing morphometric and cellular changes within the mandibular condyle of rodents.

The temporomandibular joint (TMJ) has the capacity to adapt to external stimuli, and loading changes can affect the position of condyles, as well as the ...

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 ...

Application of Electrophysiology Measurement to Study the Activity of Electro-Neutral Transporters

The transport of ions through cell membranes ensures the fine control of ion content within and outside the cell that is indispensable for cell survival. These transport mechanisms are mediated by the activities of specialized transporter proteins. Specifically,pH dynamics are finely controlled by plasma membrane proton (H+) extrusion systems, such as the Na+/H+ exchanger (NHE) protein family. Despite extensive efforts to study the mechanisms underlying NHE regulation, our current understanding of the biophysical and molecular properties of the NHE family is inadequate because of the limited availability of methods to effectively measure NHE activity. In this manuscript, we used H+-selective electrodes during whole-cell patch clamping recording to measure NHE-induced H+ flux. We proposed this approach to overcome some limitations of typically used methods to measure NHE activity, such as radioactive uptake and fluorescent membrane permeants. Measurement of NHE activity using the described method enables high sensitivity and time resolution and more efficient control of intracellular H+ concentrations. H+-selective electrodes are based on the fact that transporter activity creates an ion gradient in close proximity to the cell membrane. An H+-selective electrode moving up to and away from the cell membrane in a repetitive, oscillatory fashion records a voltage difference that is dependent on H+ flux. While H+-selective electrodes are used to detect H+ flux moving out of the cell, the patch clamp method in the whole-cell configuration is used to control the intracellular ion composition. Moreover, application of the giant patch clamp technique allows modification of the intracellular composition of not only ions but also lipids. The transporter activity of NHE isoform 3 (NHE3) was measured using this technical approach to study the molecular basis of NHE3 regulation by phosphoinositides.

This manuscript describes the applications of proton-selective electrodes and patch clamping methods to measure the activity of proton transport systems. These methods overcome some limitations of techniques commonly used to study proton transport activity, such as moderate sensitivity, time resolution and insufficient intracellular milieu control.

The transport of ions through cell membranes ensures the fine control of ion content within and outside the cell that is indispensable for cell survival. ...