All authors gratefully acknowledge funding support from the National Research Foundation of Singapore through the Singapore-MIT Alliance for Research and Technology (SMART) BioSystems and Micromechanics (BioSyM) interdisciplinary research group. Authors Dr. Jagielska and Dr. Van Vliet also gratefully acknowledge funding and support from the Saks-Kavanaugh Foundation. The authors thank William Ong and Dr. Sing Yian Chew from Nanyang Technological University, Singapore, for providing rat oligodendrocyte progenitor cells for some experiments described in this work, and the authors thank Dr. G. V. Shivashankar from Mechanobiology Institute, National University of Singapore, Singapore, for discussions about nuclear area fluctuations.

1. Design of the single-well polydimethylsiloxane mold for high-resolution imaging
NOTE: The mold for manufacturing polydimethylsiloxane plates is designed with the following features to enable imaging with a 100x oil immersion lens and a correct fit in the custom-build strain device (Figure 1A,B).
<ol>
	<li>Keep the overall plate dimensions such that it fits in the clamps of the strain device. 
	NOTE: Here, they are 60 mm x 73 mm.</li>
	<li>Make a cell compartment or well that is significantly smaller than the whole plate&#8217;s lateral dimensions, to avoid arcing (out-of-plane bending) effects that happen at the unclamped edges of the polydimethylsiloxane plate during applied strain. 
	NOTE: Here, the compartment was designed as a 23 mm x 23 mm square.</li>
	<li>Keep the depth of the cell compartment as minimal as possible (not more than 80 &#181;m), to enable focusing with the 100x oil immersion lens through the cover glass placed on top of the compartment, but make it sufficient enough to contain media and to avoid contacting or squeezing the cells.</li>
	<li>Raise the cell compartment on a square with a height of 3 mm, to bring the cells closer to the lens in the inverted microscope setup to enable easy focusing. 
	NOTE: Molds with the above features can be manufactured by many techniques including, for example, milled aluminum. Alternatively, master molds can also be assembled by using acrylic sheets cut with a laser cutter, cover glass #0 for the cell compartment, and super glue. This master mold is used to make a polydimethylsiloxane imprint, which is then used to make working molds using a casting resin.</li>
</ol>
2. Fabrication of single-well polydimethylsiloxane plates and square compartments
<ol>
	<li>Mix polydimethylsiloxane base and curing agent in a ratio of 20:1 (by weight) in a disposable cup. Weigh a total of 20 g of polydimethylsiloxane per mold (for making one polydimethylsiloxane plate) and 150 g of polydimethylsiloxane per 150 mm-in-diameter plastic dish (for making one batch of square compartments).</li>
	<li>Leave the polydimethylsiloxane mix in a vacuum degasser at -0.8 bar for 30 min (or until all bubbles are removed).</li>
	<li>Pour the polydimethylsiloxane mix into the molds (for plates) or into 150 mm plastic dishes (for square compartments).</li>
	<li>Remove any additional bubbles at this stage, either by blowing air (by mouth) or degassing the polydimethylsiloxane-filled molds/dishes again at -0.8 bar vacuum for 30 min.</li>
	<li>Leave the molds/dishes in an 80 &#176;C oven on a leveling table for&#160;2 h.</li>
	<li>Carefully remove the molds/dishes from the oven and let them cool down to room temperature. Gently peel out the cured polydimethylsiloxane after carefully cutting the edges with a blade (Figure 1C).</li>
	<li>On the 150 mm-diameter polydimethylsiloxane, draw a 2 cm x 2 cm grid with a marker. Within each square, draw a 1 cm x 1 cm square, leaving a margin of 0.5 cm on all sides. Using a blade, carefully cut along the lines to obtain square compartments (Figure 2A).</li>
	<li>Clean the polydimethylsiloxane plates/square compartments by incubating them in 100% acetone for 4&#160;h, followed by 100% ethanol for 4&#160;h, followed by autoclaved water for 4&#160;h.</li>
	<li>Place the plates/square compartments on paraffin film backing paper (not the paraffin film but the paper) inside a 150 mm diameter plastic dish (Figure 2C).</li>
	<li>Leave the polydimethylsiloxane plate in the 80 &#176;C oven to dry for 4&#160;h.</li>
	<li>Seal the 150 mm-diameter dishes containing polydimethylsiloxane plates and container squares with paraffin film, and store them in a cold room until further use.</li>
</ol>
3. Functionalization of polydimethylsiloxane plates
<ol>
	<li>Remove the paraffin film, remove the cover of the plastic dish, and place the polydimethylsiloxane plates and square compartments under ultraviolet light for 30 min.</li>
	<li>Plasma-treat (at 150 W for 5 min) the polydimethylsiloxane plates (with the cell compartment facing up) without the square compartments, to make the culture surface hydrophilic.</li>
	<li>Immediately place the square compartments onto the plasma-treated surface (Figure 2B) and press manually to temporarily stick the two together. 
	NOTE: Do not plasma-treat the square compartments, or else they will stick strongly to the polydimethylsiloxane plate and will not come off easily when they must be peeled off during imaging.</li>
	<li>Add 200 &#181;L of 100 mM (3-aminopropyl)triethoxysilane to the well for 2 h to introduce &#8211;NH2 groups to the polydimethylsiloxane surface. To make a 100 mM solution, dissolve 234 &#181;L of pure (3-aminopropyl)triethoxysilane in 10 mL of water. After a 2 h incubation, wash the wells 3x with deionized water.</li>
	<li>Weigh 2 mg of the molecular crosslinker bissulfosuccinimidyl suberate and dissolve it in 700 &#181;L of 1 M (4-(2-hydroxyethyl)-1-piperazineethane sulfonic acid buffer (pH 8.0) and 2.8 mL of water. 
	NOTE: This will give a 1 mM bissulfosuccinimidyl suberate solution in 200 mM (4-(2-hydroxyethyl)-1-piperazineethane sulfonic acid buffer. Note that a 50 mM concentration buffer also works well.
	<ol>
		<li>Add 135 &#181;L of this solution and 15 &#181;L of 1 mg/mL fibronectin to the well in each polydimethylsiloxane plate for 4 h at room temperature.</li>
	</ol>
	</li>
	<li>Wash 3x with phosphate-buffered saline solution. Add 500 &#181;L of phosphate-buffered saline solution to the well. Cover the 150 mm-diameter dish containing the polydimethylsiloxane with paraffin film and store it in a cold room, until further use.</li>
</ol>
4. Functionalization of plastic dishes and flasks
<ol>
	<li>Incubate plastic dishes and flasks with a 5 &#181;g/mL solution of poly-D-lysine (PDL, in sterile water) for 1 h. Add 1.5 mL of this solution (ligand) per 35 mm-diameter dish, 3 mL of it per 60 mm-diameter dish, and 10 mL per 75 cm2 culture flask.</li>
	<li>Wash the dishes and flasks 2x with sterile water.</li>
	<li>Leave them to dry in the biosafety cabinet for 1 h (or until dry) and store them in the cold room at 4 &#176;C until further use.</li>
</ol>
5. Proliferation and differentiation medium for murine oligodendrocyte progenitor cells
<ol>
	<li>Prepare 50 mL of 100x solutions of the proliferation and differentiation medium, make aliquots of 2.5 mL, and store them at -80 &#176;C. 
	NOTE: The composition of proliferation medium is 0.1 mg/mL bovine serum albumin, 62 ng/mL progesterone, 16 &#181;g/mL putrescine, 5 &#181;g/mL insulin, 50 &#181;g/mL holo-transferrin, 5 ng/mL sodium selenite, 1x penicillin-streptomycin, 10 ng/mL platelet-derived growth factor, and 10 ng/mL fibroblast growth factor in Dulbecco&#8217;s modified Eagle&#8217;s medium (DMEM) with high glucose and pyruvate. While bovine serum albumin, progesterone, putrescine, and sodium selenite can be constituted and stored at 100x, insulin, holo-transferrin, and penicillin-streptomycin must be added fresh while preparing the 1x solution. The growth factors must be constituted at 10 &#181;g/mL and must be added fresh to the cells every day (1 &#181;L/mL of medium). The composition of the differentiation medium is 0.1 mg/mL bovine serum albumin, 62 ng/mL progesterone, 16 &#181;g/mL putrescine, 5 &#181;g/mL insulin, 50 &#181;g/mL holo-transferrin, 5 ng/mL sodium selenite, 1x penicillin-streptomycin, 400 ng/mL triiodothyronine, 400 ng/mL L-thyroxine, and 0.5% fetal bovine serum in DMEM with high glucose and pyruvate. Triiodothyronine and L-thyroxine can be constituted with other reagents and stored at 100x. Insulin, holo-transferrin, and penicillin-streptomycin must be added fresh while preparing the 1x solution.</li>
	<li>For preparing 50 mL of 100x proliferation medium, dissolve 510 mg of bovine serum albumin in 17 mL of Dulbecco&#8217;s medium, 310 &#181;g of progesterone in 31 &#181;L of ethanol, 80 mg of putrescine in 500 &#181;L of Dulbecco&#8217;s medium, 25 &#181;g of sodium selenite in 10 &#181;L of Dulbecco&#8217;s medium, and full DMEM up to 50 mL</li>
	<li>For preparing 50 mL of 100x solution of differentiation medium, additionally dissolve 2 mg of triiodothyronine in 40 &#181;L of sodium hydroxide and 2 mg of L-thyroxine in 40 &#181;L of sodium hydroxide, then fill up the solution to 50 mL with Dulbecco&#8217;s medium. Filter the solution through a sterile disposable filter unit, aliquot, and store the aliquots at -80 &#176;C.</li>
	<li>For preparing 250 mL of 1x proliferation/differentiation medium, mix 2.5 mL of 100x solution of proliferation/differentiation medium with 12.5 mg of holo-transferrin, 125 &#181;L of 10 mg/mL insulin, and 2.5 mL of 100x penicillin-streptomycin. Fill up the solution to 250 mL with Dulbecco&#8217;s medium and filter it through a sterile disposable filter unit. 
	NOTE: The growth factors must be added fresh and directly to the cell culture, not to the whole batch of reconstituted medium.</li>
</ol>
6. Cell culture
<ol>
	<li>Start with oligodendrocyte progenitor cells suspended in proliferation medium. Seed these cells at a density of 35,000 cells/cm2 onto PDL-coated plastic surfaces (dishes or flasks, see section 4). Adjust the volume of the proliferation medium to 3 mL per 10 cm2 of surface area of plastic substratum; a lower volume of medium may lead to cell clumping arising from surface tension forces, while a higher volume of medium in the dishes may cause spillage.</li>
	<li>Add growth factors (platelet-derived growth factor and fibronectin growth factor) every day, maintaining their concentration at 10 ng/mL and change half of the proliferation medium on alternate days.</li>
	<li>On the third day, detach the cells from the plastic surface using a gentle cell detachment solution (e.g., accutase): remove the medium, wash 1x with phosphate-buffered saline solution, add 1 mL of detachment solution per 20 cm2 area, leave the dish/flask in an incubator at 37 &#176;C for 10 min, and gently tap it until all cells have detached (look under a microscope).</li>
	<li>Pipette using a 200 &#181;L tip to break clumps into single cells, transfer them to a 50 mL tube, dilute them in proliferation medium, spin at 0.2 x g for 10 min, discard the supernatant, and resuspend the pellet in proliferation medium to make a total volume of 1 mL. Count the cells using a cytometer.</li>
	<li>Wash the fibronectin-functionalized polydimethylsiloxane plates 2x, 15 min per wash, with proliferation medium.</li>
	<li>Seed 35,000 cells per polydimethylsiloxane plate in 700 &#181;L of proliferation medium.</li>
	<li>Add a plasmid construct of H2B-GFP for labeling histone H2B with green fluorescent protein. 
	NOTE: Here, 1.4 &#181;L of CellLight H2B-GFP BacMam2 transfection mix was added to each plate (2 &#181;L per 50,000 cells).</li>
	<li>After 24 h, mount the polydimethylsiloxane samples with cells to be stretched on the uniaxial strain device (as shown in Figure 2D). Change the medium in all samples to differentiation medium.</li>
	<li>To strain the samples, measure the unstrained cell compartment length (Figure 3A) and turn the stage micrometer screw to increase the cell compartment length by the desired amount (e.g., 10%, see Figure 3B). Leave the stretched and unstretched polydimethylsiloxane samples in the incubator at 37 &#176;C until imaging.</li>
</ol>
7. Imaging
<ol>
	<li>Turn on the microscope. Set the microscope incubator temperature to 37 &#176;C.</li>
	<li>Bring the objective to be used (100x oil immersion) to the central position as the objective turret will not be accessible later. Unscrew the objective and screw it back together with an objective ring (Figure 4A) so that the objective can be brought closer to the cells. If an oil objective is being used here, add a drop of oil at this step.</li>
	<li>Synthesize beforehand (via 3D printing or machining) a plastic or metal holder that has two important features&#8212;an angled window and a step (Figure 4B). 
	NOTE: The window supports a thickness #0 glass coverslip that will hold the medium while the strain device is in an inverted state. The angled cut at the window edges (Figure 4D) allows the objective to come closer to the glass coverslip. The step in the holder allows us to bring the stretched polydimethylsiloxane further down, closer to the objective.</li>
	<li>Place a glass coverslip (thickness #0, with dimensions of 25 mm x 25 mm or 35 mm in diameter) onto the top surface of the holder by spreading vacuum grease at the periphery of the window (Figure 4C), and tape the plastic window onto the microscope stage (Figure 4D and Figure 5A).</li>
	<li>Screw a z-translation stage onto the microscope stage and move it to the topmost z-position (Figure 5A).</li>
	<li>Remove 500 &#181;L of the medium from the stretched polydimethylsiloxane plate to be imaged and add this medium onto the glass coverslip in the white plastic window.</li>
	<li>Carefully detach the square compartment from the polydimethylsiloxane plate with sterile tweezers.</li>
	<li>Hold the strain device in an upright position (cells facing up) above the white plastic window and carefully invert the strain device so as to let any extra medium drop directly in the middle of the glass coverslip (Figure 5B) (cells facing down).</li>
	<li>Place the bottom part of the strain device onto the z-translation stage with the double-sided tape (Figure 5C,D).</li>
	<li>While looking through the eyepiece under brightfield, slowly bring the strain device down (Figure 5E) (by lowering the z-translation stage) and move the objective up to focus on the cells.
	<ol>
		<li>Perform this step extremely slowly, step by step. If the strain device presses too much onto the coverslip, the cells can get compressed (causing them to die) and if the objective presses too much onto the coverslip, the coverslip can break (causing medium to leak and spill onto the objective).</li>
	</ol>
	</li>
	<li>Scan the polydimethylsiloxane plate in x- and y-directions to find a cell that has a fluorescent nucleus (under epifluorescence with 488 nm of wavelength excitation) and appropriate cell morphology (under brightfield).</li>
	<li>If the lateral displacement of the microscope stage does not affect the vertical focusing too much, select multiple regions of interests using a multipoint feature on the software. Sometimes, the focusing is very sensitive to any lateral displacement of the stage. In such cases, image one cell at a time. Mark the x-, y-, and z-positions for each cell of interest.</li>
	<li>Record wide-field (or open-pinhole) images of the nucleus with 488 nm of wavelength excitation and of the cell with brightfield excitation at intervals of 30 s per frame for a total duration of at least 30 min.</li>
</ol>
8. Data analysis
<ol>
	<li>Nuclear fluctuations
	<ol>
		<li>Open the sequence of nucleus images (Figure 6A) in an image analysis software and threshold the nucleus time-lapse images (for example, use the Threshold command in ImageJ or the Graythresh command in the software MATLAB).</li>
		<li>Get the nucleus area (in pixels) as a function of time, plot it in a data plotting software (Figure 6B), and fit a third order polynomial to the data (Figure 6C) (for example, use the Analysis | Fitting | Polynomial Fit command in Origin or the Polyfit command in the software MATLAB).</li>
		<li>Subtract the value of the fitted polynomial from the actual area (in pixels) for each time point. This corrected area is known as the residual area. 
		NOTE: This process removes any increasing or decreasing trend in the data coming from instrumental errors and is called detrending.
		<ol>
			<li>Calculate the percentage of residual area by dividing the residual area at each time point with the value of the fitted polynomial at that time point (Figure 6D).</li>
		</ol>
		</li>
		<li>Calculate the standard deviation of the residual area time series. This standard deviation denotes the amplitude of nuclear fluctuations.</li>
	</ol>
	</li>
	<li>Data plotting and statistical analysis
	<ol>
		<li>Calculate the amplitude of nuclear fluctuations as a mean of at least 20 nuclei per condition.</li>
		<li>Conduct a one-way analysis-of-variance statistical test with Bonferroni correction to determine whether there is a significant difference in the amplitude of fluctuations as a function of conditions of interest (for example, with and without applied strain).</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Bray, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanical+tension+produced+by+nerve+cells+in+tissue+culture.">Mechanical tension produced by nerve cells in tissue culture.</a> Journal of Cell Science. 410, 391-410 (1979).</li><li>Bray, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Axonal+growth+in+response+to+experimentally+applied+mechanical+tension.">Axonal growth in response to experimentally applied mechanical tension.</a> Developmental Biology. 389, (1983).</li><li>Van Essen, D. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+tension-based+theory+of+morphogenesis+and+compact+wiring+in+the+central+nervous+system.">A tension-based theory of morphogenesis and compact wiring in the central nervous system.</a> Nature. , (1997).</li><li>Smith, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stretch+growth+of+integrated+axon+tracts:+Extremes+and+exploitations.">Stretch growth of integrated axon tracts: Extremes and exploitations.</a> Progress in Neurobiology. 89, 231-239 (2011).</li><li>Cullen, D. K., Simon, C. M., LaPlaca, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Strain+rate-dependent+induction+of+reactive+astrogliosis+and+cell+death+in+three-dimensional+neuronal-astrocytic+co-cultures.">Strain rate-dependent induction of reactive astrogliosis and cell death in three-dimensional neuronal-astrocytic co-cultures.</a> Brain Research. , 103-115 (2011).</li><li>Fisher, E., 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=Imaging+correlates+of+axonal+swelling+in+chronic+multiple+sclerosis+brains.">Imaging correlates of axonal swelling in chronic multiple sclerosis brains.</a> Annals of Neurology. 2, 219-228 (2007).</li><li>Niki&#263;, I., 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=A+reversible+form+of+axon+damage+in+experimental+autoimmune+encephalomyelitis+and+multiple+sclerosis.">A reversible form of axon damage in experimental autoimmune encephalomyelitis and multiple sclerosis.</a> Nature Medicine. 17, 495-500 (2011).</li><li>Payne, S. C., Bartlett, C. A., Harvey, A. R., Dunlop, S. A., Fitzgerald, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+loss+during+chronic+secondary+degeneration+in+rat+optic+nerve.">Functional loss during chronic secondary degeneration in rat optic nerve.</a> Investigative Ophthalmology &amp; Visual Science. 53, (2018).</li><li>Zeiger, A. 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=Static+mechanical+strain+induces+capillary+endothelial+cell+cycle+re-entry+and+sprouting.">Static mechanical strain induces capillary endothelial cell cycle re-entry and sprouting.</a> Physical Biology. 13, 1-16 (2017).</li><li>Jagielska, A., 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=Mechanical+strain+promotes+oligodendrocyte+differentiation+by+global+changes+of+gene+expression.">Mechanical strain promotes oligodendrocyte differentiation by global changes of gene expression.</a> Frontiers in Cellular Neuroscience. 11, 93 (2017).</li><li>Pajerowski, J. D., Dahl, K. N., Zhong, F. L., Sammak, P. J., Discher, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Physical+plasticity+of+the+nucleus+in+stem+cell+differentiation.">Physical plasticity of the nucleus in stem cell differentiation.</a> Proceedings of the National Academy of Sciences of the United States of America. 1, 0 (2007).</li><li>Talwar, S., Kumar, A., Rao, M., Menon, G. I., Shivashankar, G. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlated+spatio-temporal+fluctuations+in+chromatin+compaction+states+characterize+stem+cells.">Correlated spatio-temporal fluctuations in chromatin compaction states characterize stem cells.</a> Biophysical Journal. 104, 553-564 (2013).</li><li>Makhija, E., 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=Mechanical+strain+alters+cellular+and+nuclear+dynamics+at+early+stages+of+oligodendrocyte+differentiation.">Mechanical strain alters cellular and nuclear dynamics at early stages of oligodendrocyte differentiation.</a> Frontiers in Cellular Neuroscience. 12, 59 (2018).</li></ol>

The authors have nothing to disclose.

A device has previously1 been designed for the application of extracellular tensile strain on adherent cells. The design of the polydimethylsiloxane substratum in that work was sufficient for biochemical assays, as well as the low-resolution imaging of stretched cells. In this work, the substratum was redesigned, and a novel imaging configuration that facilitates high-resolution subcellular live cell imaging was introduced. The advantages of this system are numerous: it can be built in-lab using s...

Cells and tissues in the body are subjected to various mechanical cues, including tensile strains. However, the effects of these cues on the biology of neural cells have not yet been studied extensively and understood fully. In the central nervous system, sources of mechanical strain include developmental growth1,2,3,4, physiological processes such as spinal cord bending, blood and cerebrospinal fluid pulsation, and pathological conditions such as trauma, axon swelling, glial scarring, or tumor growth5,6,7,8. It is worth investigating how tensile strain affects the differentiation of oligodendrocytes and the subsequent myelination of axons, which is a critical process in the vertebrate central nervous system. Using a custom-designed strain device and elastomeric multiwell plates, previous works9,10 have shown that static uniaxial strain can increase oligodendrocyte differentiation via global changes in gene expression10. To gain further understanding of the mechanisms of strain mechanotransduction in these cells, the previous experimental apparatus must be redesigned as described here, to enable high-resolution fluorescence imaging of nuclear dynamics in living cells under strain. Specifically, a single-well polydimethylsiloxane plate is developed, and the imaging configuration is redesigned to allow for the time-lapse imaging of live cells under strain using a 100x oil immersion lens. To eliminate the negative optical effects of polydimethylsiloxane in the light pathway, cells are imaged not through the polydimethylsiloxane plate, but in the inverted position, through the cover glass covering the cell compartment. Using this new imaging design, hundreds of high-resolution time-lapse movies are recorded, of individual cell nuclei within intact adherent cells, where chromatin is labeled by tagging histone H2B to green fluorescent protein. These movies demonstrate that tensile strain induces changes in chromatin structure and dynamics that are consistent with the progression of oligodendrocyte differentiation.
Live cell imaging under applied strain is technically challenging and requires a device design that is compatible with the microscope system. The custom design described here presents an inexpensive alternative to commercial solutions. Its dimensions enable its installation on microscope stages and live cell imaging at high spatial resolution during applied strain. The imaging setup is designed to facilitate live cell imaging using a 100x oil immersion lens with the highest clarity, through the cover glass, not through the layer of polydimethylsiloxane plate which otherwise decreases the image quality and is common in most imaging setups under strain. The device, with a mounted plate containing cells, can also be stored easily in the incubator. This device is designed to apply uniaxial strain to substrata that facilitate adherent cell culture and maintain a stable and uniform strain over multiple days. The setup described here can be used for the high-resolution imaging of various adherent cell types under strain, making it applicable to mechanotransduction studies in many fields of cell mechanobiology.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Primary cells</td>
			<td></td>
			<td></td>
			<td>Primary Oligodendrocyte Progenitor Cells isolated from Neonatal Rats 
			were provided by Dr. S. Y. Chew&#39;s lab at Nanyang Technological University</td>
		</tr>
		<tr>
			<td>Media</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM high&#160; glucose</td>
			<td>Gibco</td>
			<td>11996-065-500ml</td>
			<td></td>
		</tr>
		<tr>
			<td>Pen/Strep</td>
			<td>Gibco</td>
			<td>15070-063-100ml</td>
			<td></td>
		</tr>
		<tr>
			<td>FGF</td>
			<td>Prospec</td>
			<td>CYT-608-50ug</td>
			<td></td>
		</tr>
		<tr>
			<td>PDGF</td>
			<td>Prospec</td>
			<td>CYT-776-10ug</td>
			<td></td>
		</tr>
		<tr>
			<td>Media Stock components</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma</td>
			<td>A9418-10G</td>
			<td></td>
		</tr>
		<tr>
			<td>Progesterone</td>
			<td>Sigma</td>
			<td>P8783-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>Putrescine</td>
			<td>Sigma</td>
			<td>P5780-5G</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Selenite</td>
			<td>Sigma</td>
			<td>S5261-10G</td>
			<td></td>
		</tr>
		<tr>
			<td>Tri-iodothyronine</td>
			<td>Sigma</td>
			<td>T6397-100MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Thyroxine</td>
			<td>Sigma</td>
			<td>T1775-100MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Holo-Transferrin</td>
			<td>Sigma</td>
			<td>T0665-500MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Insulin</td>
			<td>Sigma</td>
			<td>I9278</td>
			<td></td>
		</tr>
		<tr>
			<td>Mold and PDMS plate</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PDMS - Dow Corning Sylgard 184</td>
			<td>Ellsworth</td>
			<td>184 SIL ELAST KIT 0.5KG</td>
			<td></td>
		</tr>
		<tr>
			<td>Mold - Liquid Plastic</td>
			<td>Smooth-On</td>
			<td>Smooth Cast 310 - Trail Size</td>
			<td></td>
		</tr>
		<tr>
			<td>Substrate Coatings</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>poly-D-lysine</td>
			<td>Sigma</td>
			<td>P6407-5MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Fibronectin</td>
			<td>Sigma</td>
			<td>F1141-2MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Histone staining</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CellLigt H2B-GFP BacMam</td>
			<td>Molecular Probes</td>
			<td>C10594</td>
			<td></td>
		</tr>
		<tr>
			<td>Surface functionalization</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>APTES</td>
			<td>Sigma</td>
			<td>A3648-100ml</td>
			<td></td>
		</tr>
		<tr>
			<td>BS3</td>
			<td>Covachem</td>
			<td>13306-100mg</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES buffer pH 8.0</td>
			<td>Alfa Aesar</td>
			<td>J63578-AK-250ml</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell detachment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Accutase</td>
			<td>Invitrogen</td>
			<td>A1110501-100ml</td>
			<td></td>
		</tr>
	</tbody>
</table>

high-resolution imaging of nuclear dynamics in live cells under uniaxial tensile strain

Extracellular mechanical strain is known to elicit cell phenotypic responses and has physiological relevance in several tissue systems. To capture the effect of applied extracellular tensile strain on cell populations in vitro via biochemical assays, a device has previously been designed which can be fabricated simply and is small enough to fit inside tissue culture incubators, as well as on top of microscope stages. However, the previous design of the polydimethylsiloxane substratum did not allow high-resolution subcellular imaging via oil-immersion objectives. This work describes a redesigned geometry of the polydimethylsiloxane substratum and a customized imaging setup that together can facilitate high-resolution subcellular imaging of live cells while under applied strain. This substratum can be used with the same, earlier designed device and, hence, has the same advantages as listed above, in addition to allowing high-resolution optical imaging. The design of the polydimethylsiloxane substratum can be improved by incorporating a grid which will facilitate tracking the same cell before and after the application of strain. Representative results demonstrate high-resolution time-lapse imaging of fluorescently labeled nuclei within strained cells captured using the method described here. These nuclear dynamics data give insights into the mechanism by which applied tensile strain promotes differentiation of oligodendrocyte progenitor cells.&#160;

Recent work aimed at investigating the effect of tensile strain on oligodendrocytes10 showed that a 10% uniaxial tensile strain promotes the differentiation of oligodendrocyte progenitor cells by global changes in gene expression. The mechanism behind these changes in gene expression can be probed via the imaging of subcellular parameters, such as the cytoskeleton structure, transcription factor localization, nuclear dynamics, and chromatin organization. However, t...

Watch this Scientific Journal Video about High-resolution Imaging of Nuclear Dynamics in Live Cells under Uniaxial Tensile Strain at JoVE.com

High-resolution Imaging of Nuclear Dynamics in Live Cells under Uniaxial Tensile Strain

Using a previously designed device to apply mechanical strain to adherent cells, this paper describes a redesigned substratum geometry and a customized apparatus for high-resolution single-cell imaging of strained cells with a 100x oil immersion objective.

Extracellular mechanical strain is known to elicit cell phenotypic responses and has physiological relevance in several tissue systems. To capture the ...

high-resolution-imaging-nuclear-dynamics-live-cells-under-uniaxial

Research

JoVE Journal

Bioengineering

7.8K Views.  Singapore-MIT Alliance for Research and Technology, CREATE. Using a previously designed device to apply mechanical strain to adherent cells, this paper describes a redesigned substratum geometry and a customized apparatus for high-resolution single-cell imaging of strained cells with a 100x oil immersion objective.

Video: High-resolution Imaging of Nuclear Dynamics in Live Cells under Uniaxial Tensile Strain

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

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

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

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

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

High Resolution 3D Imaging of Ex-Vivo Biological Samples by Micro CT

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized tomography (&mu;CT).
High resolution &mu;CT is a very versatile yet accurate (1-2 microns of resolution) technique for 3D examination of ex-vivo biological samples1, 2. As opposed to electron tomography, the &mu;CT allows the examination of up to 4 cm thick samples. This technique requires only few hours of measurement as compared to weeks in histology. In addition, &mu;CT does not rely on 2D stereologic models, thus it may complement and in some cases can even replace histological methods3, 4, which are both time consuming and destructive. Sample conditioning and positioning in &mu;CT is straightforward and does not require high vacuum or low temperatures, which may adversely affect the structure. The sample is positioned and rotated 180&deg; or 360&deg;between a microfocused x-ray source and a detector, which includes a scintillator and an accurate CCD camera, For each angle a 2D image is taken, and then the entire volume is reconstructed using one of the different available algorithms5-7. The 3D resolution increases with the decrease of the rotation step. The present video protocol shows the main steps in preparation, immobilization and positioning of the sample followed by imaging at high resolution.

High Resolution 3D Imaging of Ex-Vivo Biological Samples by Micro CT

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized tomography ( CT).

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized ...

Live-cell Imaging of Migrating Cells Expressing Fluorescently-tagged Proteins in a Three-dimensional Matrix

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound healing, tissue regeneration, and cancer metastasis, cells must navigate through complex, three-dimensional extracellular tissue. To better understand the mechanisms behind these biological processes, it is important to examine the roles of the proteins responsible for driving cell migration. Here, we outline a protocol to study the mechanisms of cell migration using the epithelial cell line (MDCK), and a three-dimensional, fibrous, self-polymerizing matrix as a model system. This optically clear extracellular matrix is easily amenable to live-cell imaging studies and better mimics the physiological, soft tissue environment. This report demonstrates a technique for directly visualizing protein localization and dynamics, and deformation of the surrounding three-dimensional matrix. 
Examination of protein localization and dynamics during cellular processes provides key insight into protein functions. Genetically encoded fluorescent tags provide a unique method for observing protein localization and dynamics. Using this technique, we can analyze the subcellular accumulation of key, force-generating cytoskeletal components in real-time as the cell maneuvers through the matrix. In addition, using multiple fluorescent tags with different wavelengths, we can examine the localization of multiple proteins simultaneously, thus allowing us to test, for example, whether different proteins have similar or divergent roles. Furthermore, the dynamics of fluorescently tagged proteins can be quantified using Fluorescent Recovery After Photobleaching (FRAP) analysis. This measurement assays the protein mobility and how stably bound the proteins are to the cytoskeletal network.
By combining live-cell imaging with the treatment of protein function inhibitors, we can examine in real-time the changes in the distribution of proteins and morphology of migrating cells. Furthermore, we also combine live-cell imaging with the use of fluorescent tracer particles embedded within the matrix to visualize the matrix deformation during cell migration. Thus, we can visualize how a migrating cell distributes force-generating proteins, and where the traction forces are exerted to the surrounding matrix. Through these techniques, we can gain valuable insight into the roles of specific proteins and their contributions to the mechanisms of cell migration.

Cellular processes such as cell migration have traditionally been studied on two-dimensional, stiff plastic surfaces. This report describes a technique for directly visualizing protein localization and analyzing protein dynamics in cells migrating in a more physiologically relevant, three-dimensional matrix.

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound ...

Continuous High-resolution Microscopic Observation of Replicative Aging in Budding Yeast

We demonstrate the use of a simple microfluidic setup, in which single budding yeast cells can be tracked throughout their entire lifespan. The microfluidic chip exploits the size difference between mother and daughter cells using an array of micropads. Upon loading, cells are trapped underneath these micropads, because the distance between the micropad and cover glass is similar to the diameter of a yeast cell (3-4 &mu;m). After the loading procedure, culture medium is continuously flushed through the chip, which not only creates a constant and defined environment throughout the entire experiment, but also flushes out the emerging daughter cells, which are not retained underneath the pads due to their smaller size. The setup retains mother cells so efficiently that in a single experiment up to 50 individual cells can be monitored in a fully automated manner for 5 days or, if necessary, longer. In addition, the excellent optical properties of the chip allow high-resolution imaging of cells during the entire aging process.

We describe here the operation of a microfluidic device that allows continuous and high-resolution microscopic imaging of single budding yeast cells during their complete replicative and/or chronological lifespan.

We demonstrate the use of a simple microfluidic setup, in which single budding yeast cells can be tracked throughout their entire lifespan. The ...

Magnetically-Assisted Remote Controlled Microcatheter Tip Deflection under Magnetic Resonance Imaging

X-ray fluoroscopy-guided endovascular procedures have several significant limitations, including difficult catheter navigation and use of ionizing radiation, which can potentially be overcome using a magnetically steerable catheter under MR guidance.
	The main goal of this work is to develop a microcatheter whose tip can be remotely controlled using the magnetic field of the MR scanner. This protocol aims to describe the procedures for applying current to the microcoil-tipped microcatheter to produce consistent and controllable deflections.
	A microcoil was fabricated using laser lathe lithography onto a polyimide-tipped endovascular catheter. In vitro testing was performed in a waterbath and vessel phantom under the guidance of a 1.5-T MR system using steady-state free precession (SSFP) sequencing. Various amounts of current were applied to the coils of the microcatheter to produce measureable tip deflections and navigate in vascular phantoms.
	The development of this device provides a platform for future testing and opportunity to revolutionize the endovascular interventional MRI environment.

Current applied to an endovascular microcatheter with microcoil tip made by laser lathe lithography can achieve controllable deflections under magnetic resonance (MR) guidance, which may improve speed and efficacy of navigation of vasculature during various endovascular procedures.

X-ray fluoroscopy-guided  endovascular procedures have several significant limitations, including  difficult catheter navigation and use of ionizing ...

High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e., &#946;2-adrenergic and &#947;-aminobutyric acid type B (GABAB) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.

This protocol describes how to use total internal reflection fluorescence microscopy to visualize and track single receptors on the surface of living cells and thereby analyze receptor lateral mobility, size of receptor complexes as well as to visualize transient receptor-receptor interactions. This protocol can be extended to other membrane proteins.

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., ...

From Fast Fluorescence Imaging to Molecular Diffusion Law on Live Cell Membranes in a Commercial Microscope

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the regulation of many cellular functions. However, due to the fast dynamics and the tiny structures involved, a very high spatio-temporal resolution is required to catch the real behavior of molecules. Here we present the experimental protocol for studying the dynamics of fluorescently-labeled plasma-membrane proteins and lipids in live cells with high spatiotemporal resolution. Notably, this approach doesn&#8217;t need to track each molecule, but it calculates population behavior using all molecules in a given region of the membrane. The starting point is a fast imaging of a given region on the membrane. Afterwards, a complete spatio-temporal autocorrelation function is calculated correlating acquired images at increasing time delays, for example each 2, 3, n repetitions. It is possible to demonstrate that the width of the peak of the spatial autocorrelation function increases at increasing time delay as a function of particle movement due to diffusion. Therefore, fitting of the series of autocorrelation functions enables to extract the actual protein mean square displacement from imaging (iMSD), here presented in the form of apparent diffusivity vs average displacement. This yields a quantitative view of the average dynamics of single molecules with nanometer accuracy. By using a GFP-tagged variant of the Transferrin Receptor (TfR) and an ATTO488 labeled 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphoethanolamine (PPE) it is possible to observe the spatiotemporal regulation of protein and lipid diffusion on &#181;m-sized membrane regions in the micro-to-milli-second time range.

Spatial distribution and temporal dynamics of plasma membrane proteins and lipids is a hot topic in biology. Here this issue is addressed by a spatio-temporal image fluctuation analysis that provides conceptually the same physical quantities of single particle tracking, but it uses small molecular labels and standard microscopy setups.

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the ...

Preparation of Mica Supported Lipid Bilayers for High Resolution Optical Microscopy Imaging

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction with other compounds, such as drugs or peptides. However SLB characteristics differ depending on the support used. 
Commonly used techniques for SLB imaging and measurements are single molecule fluorescence microscopy, FCS and atomic force microscopy (AFM). Because most optical imaging studies are carried out on a glass support, while AFM requires an extremely flat surface (generally mica), results from these techniques cannot be compared directly, since the charge and smoothness properties of these materials strongly influence diffusion. Unfortunately, the high level of manual dexterity required for the cutting and gluing thin slices of mica to the glass slide presents a hurdle to routine use of mica for SLB preparation. Although this would be the method of choice, such prepared mica surfaces often end up being uneven (wavy) and difficult to image, especially with small working distance, high numerical aperture lenses. Here we present a simple and reproducible method for preparing thin, flat mica surfaces for lipid vesicle deposition and SLB preparation. Additionally, our custom made chamber requires only very small volumes of vesicles for SLB formation. The overall procedure results in the efficient, simple and inexpensive production of high quality lipid bilayer surfaces that are directly comparable to those used in AFM studies.

We present a method of preparing mica supported lipid bilayers for high resolution microscopy. Mica is transparent and flat on an atomic scale, but rarely used in imaging because of handling difficulties; our preparation results in even deposition of the mica sheet, and reduces the material used in bilayer preparation.

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction ...

Live Cell Imaging during Mechanical Stretch

There is currently a significant interest in understanding how cells and tissues respond to mechanical stimuli, but current approaches are limited in their capability for measuring responses in real time in live cells or viable tissue. A protocol was developed with the use of a cell actuator to distend live cells grown on or tissues attached to an elastic substrate while imaging with confocal and atomic force microscopy (AFM). Preliminary studies show that tonic stretching of human bronchial epithelial cells caused a significant increase in the production of mitochondrial superoxide. Moreover, using this protocol, alveolar epithelial cells were stretched and imaged, which showed direct damage to the epithelial cells by overdistention simulating one form of lung injury in vitro. A protocol to conduct AFM nano-indentation on stretched cells is also provided.

A novel imaging protocol was developed using a custom motor-driven mechanical actuator to allow the measurement of real time responses to mechanical strain in live cells. Relevant to mechanobiology, the system can apply strains up to 20% while allowing near real-time imaging with confocal or atomic force microscopy.

There is currently a significant interest in understanding how cells and tissues respond to mechanical stimuli, but current approaches are limited in ...

Initial Evaluation of Antibody-conjugates Modified with Viral-derived Peptides for Increasing Cellular Accumulation and Improving Tumor Targeting

Antibody-conjugates (ACs) modified with virus-derived peptides are a potentially powerful class of tumor cell delivery agents for molecular payloads used in cancer treatment and imaging due to increased cellular accumulation over current ACs. During early AC in vitro development, fluorescence techniques and radioimmunoassays are sufficient for determining intracellular localization, accumulation efficiency, and target cell specificity. Currently, there is no consensus on standardized methods for preparing cells for evaluating AC intracellular accumulation and localization. The initial testing of ACs modified with virus-derived peptides is critical especially if several candidates have been constructed. Determining intracellular accumulation by fluorescence can be affected by background signal from ACs at the cell surface and complicate the interpretation of accumulation. For radioimmunoassays, typically treated cells are fractionated and the radioactivity in different cell compartments measured. However, cell lysis varies from cell to cell and often nuclear and cytoplasmic compartments are not adequately isolated. This can produce misleading data on payload delivery properties. The intravenous injection of radiolabeled virus-derived peptide-modified ACs in tumor bearing mice followed by radionuclide imaging is a powerful method for determining tumor targeting and payload delivery properties at the in vivo phase of development. However, this is a relatively recent advancement and few groups have evaluated virus-derived peptide-modified ACs in this manner. We describe the processing of treated cells to more accurately evaluate virus-derived peptide-modified AC accumulation when using confocal microscopy and radioimmunoassays. Specifically, a method for trypsinizing cells to remove cell surface bound ACs. We also provide a method for improving cellular fractionation. Lastly, this protocol provides an in vivo method using positron emission tomography (PET) for evaluating initial tumor targeting properties in tumor-bearing mice. We use the radioisotope 64Cu (t1/2 = 12.7 h) as an example payload in this protocol.

Viral-derived peptides coupled to antibody-conjugates (ACs) is an approach gaining momentum due to the potential of delivering molecular payloads with increased tumor cell accumulation. Utilizing common methods to evaluate peptide conjugation, AC and payload intracellular accumulation, and tumor targeting, this protocol helps researchers during the key initial development phases.

Antibody-conjugates (ACs) modified with virus-derived peptides are a potentially powerful class of tumor cell delivery agents for molecular payloads used ...