The authors would like that thank Fedex Institute of Technology at the University of Memphis for their support. The authors would like to acknowledge students of the senior design project group in the Mechanical Engineering Department at the University of Memphis (David Butler, Jackie Carter, Dominick Cleveland, Jacob Shaffer), Daniel Kohn from the University of Memphis Engineering Technology department for motor control, and Dr. Bin Teng and Ms. Charlean Luellen for their help in cell culture. This work was supported by K01 HL120912 (ER) and R01 HL123540 (CMW).

1. Construction of Membrane with Well Walls for Retention of Cell Culture Media (see Figure 1D for the final product)
<ol>
	<li>Using polydimethylsiloxane (PDMS) sheets coated with Collagen I, cut the outline of the flexible membrane with a scalpel or a die.</li>
	<li>Place each membrane in a 60 mm Petri dish for storage.</li>
	<li>Creation of walls:
	<ol>
		<li>Mix PDMS at a 10:1 weight ratio of elastomer A to elastomer B (curing agent).</li>
		<li>Pour 5 ml of fully mixed PDMS into 50 ml tubes.</li>
		<li>Place 50 ml tubes with uncured PDMS horizontally in a hybridization oven.</li>
		<li>Use the rotor function to coat the inner walls of the tubes at 8 rpm during the curing time. At RT, the PDMS will be fully cured in 2 days.</li>
		<li>Remove the PDMS from the tube in a sterile cell culture hood.</li>
		<li>Using a new razor blade, partition the cylinder of PDMS into sections 4 mm in height.</li>
		<li>Place the sections, which will serve as the membrane walls, into a Petri dish container without allowing the walls to become creased.</li>
	</ol>
	</li>
	<li>Select and center one wall of PDMS on one membrane while maintaining the two in the Petri dish.</li>
	<li>Gently place uncured PDMS (10:1 ratio) on the outside perimeter of the wall to be used as glue. Prevent formation of gaps between the wall and the membrane since it is there to retain liquid.</li>
	<li>Place each Petri dish containing the completed membrane and covered in an oven at 70 &#176;C for 24 hr to cure.</li>
</ol>
2. Correlation of Motor Rotations with Clamp Displacement or Radial Growth for Calibration (Figure 2)
<ol>
	<li>Start the software controlling the motor.</li>
	<li>Displace the clamps of the device using the manual setting in the motor software.</li>
	<li>Measure the distance and record the motor count position between opposing clamps at both minimum and maximum displacement of the clamps.</li>
	<li>Calculate the percent change in the distance between the clamps as a function of the motor count (~0-75k counts). This will indicate the maximum potential strain achieved on the membrane.</li>
</ol>
3. Application of Stretch on Mouse Lung Epithelial Cell Line (MLE12)
<ol>
	<li>Seed MLE12 cells at 2.5 million cells on the flexible membrane (Step 1) per well to be confluent within two days. The seeding density may vary for different cell types.</li>
	<li>Make sure the mechanical actuator is in a fully relaxed position.
	<ol>
		<li>Remove cell culture media.</li>
		<li>Using 1.5 mm biopsy punches, punch two holes in each of the six clamp (see Figure 1D) tabs of the membrane. The radial placement of the holes determines the amount of pre-tension the membrane experiences initially. Punch the holes at a radius of 20.5 mm on the membrane tabs if no pre-tension is desired.</li>
		<li>Position the membrane on the stretcher with the punched holes lining up with the pins within the clamps. Place top clamps in place. Tighten the screws one at a time alternating sides.</li>
		<li>Add 1 ml cell culture media.</li>
	</ol>
	</li>
	<li>Place the device on the microscope stage centering the middle of the membrane with the light path.</li>
	<li>Use tape or magnets (if possible) to fix the device to the stage. Once fixed, control the in-plane (parallel to the membrane) and vertical (z-direction) of the mechanical device with the stage controller of the microscope.</li>
	<li>Apply stretch with the software provided by the manufacturer by manually controlling the position and speed of motor rotation20.</li>
</ol>
4. Measurement of Mitochondrial Reactive Oxygen Species (ROS)
<ol>
	<li>Once cells are confluent, add 1-2 ml of RT DMEM with mitochondrial superoxide indicator (5 &#181;M final concentration) directly on the cells.</li>
	<li>Incubate for 10 min at 37 &#176;C.</li>
	<li>Wash cells gently three times with buffer warmed in a water bath to 37 &#176;C.</li>
	<li>Immediately place the flexible membrane on the stretcher (Step 2.2) already in place under an upright confocal microscope.</li>
	<li>Add 1 ml of DMEM with phenol-red free medium and 25 mM of HEPES.</li>
	<li>Set excitation/emission filters to 510/580 nm.</li>
	<li>Image multiple fields every 15 min to create the desired time course of mitochondrial superoxide production.</li>
	<li>From captured images, record fluorescence intensity histograms at each time interval using software capable of quantifying fluorescence intensity.</li>
</ol>
5. Application of Stretch and Atomic Force Microscopy (AFM)
NOTE: These steps are provided for a specific AFM and optical microscope combination (Figure 4 and Materials List).
<ol>
	<li>Prepare the AFM for the experiment.
	<ol>
		<li>Increase the height of the AFM head to its maximum position in the z-direction.</li>
		<li>Put extenders on the legs of the AFM to lift the plane at which AFM cantilever contacts the sample. The specimen and the AFM head needs to be lifted to accommodate the height of the mechanical device.</li>
	</ol>
	</li>
	<li>Prepare the optical microscope (if available).
	<ol>
		<li>Remove the AFM scanner plate. Remove the desired objective.</li>
		<li>Add a spacer to the objective. The height of the spacer would depend on the objective and the specific AFM set-up, but it is necessary if optical imaging is desired since observation plane will be shift in the z-direction by an amount equal to the stretcher and adapter height (Figure 5A). Note that the AFM usually provides low magnification imaging from an optical path above the device.</li>
		<li>Mount the desired objective back in its location. Place the scanner back on to the AFM.</li>
		<li>Start the AFM software. Start all necessary light sources including the light source for fluorescence measurements.</li>
		<li>Mount a chip with a cantilever beam that is appropriate for the desired measurements. 200 pN/nm or less stiffness is preferred when measuring the elastic modulus of live cells.</li>
		<li>Align the laser and calibrate the cantilever stiffness according to manufacturer&#8217;s suggestions on a glass coverslip mounted on the device.</li>
	</ol>
	</li>
	<li>Prepare a membrane as in Step 1, but with the following modifications.
	<ol>
		<li>Immediately before mounting the membrane on to the stretching device, cut the walls to about 1 mm in height. This prevents the interference experienced between the membrane walls and the load cell.</li>
		<li>Mount the membrane on the mechanical device as described 3.2.1-3.2.3.</li>
		<li>Remove cell culture media to prevent damage to the AFM scanner in case of a spill.</li>
		<li>Couple the device with an adapter (Figure 5A). Place the mechanical device with the adapter on the scanner.</li>
		<li>Stretch the membrane to the desired tensile strain level.</li>
	</ol>
	</li>
	<li>Nano-indentation of stretched cells.
	<ol>
		<li>Add a limited (&#60;0.5 ml) volume of media on the cells to avoid AFM scanner or microscope damage due to a spill.</li>
		<li>Engage the cantilever beam with the membrane.</li>
		<li>Follow the protocol of the particular AFM device to scan areas of interest.&#160;</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Tschumperlin, D. J., Boudreault, F., Liu, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recent+advances+and+new+opportunities+in+lung+mechanobiology.">Recent advances and new opportunities in lung mechanobiology.</a> J Biomech. 43, 99-107 (2010).</li><li>Waters, C. M., Roan, E., Navajas, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Comprehensive Physiology. , (2011).</li><li>Majkut, S., Dingal, P. C. D. P., 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=Stress+Sensitivity+and+Mechanotransduction+during+Heart+Development.">Stress Sensitivity and Mechanotransduction during Heart Development.</a> Current Biology. 24, R495-R501 (2014).</li><li>Hoffman, B. D., Grashoff, C., Schwartz, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+molecular+processes+mediate+cellular+mechanotransduction.">Dynamic molecular processes mediate cellular mechanotransduction.</a> Nature. 475, 316-323 (2011).</li><li>Wang, N., Butler, J. P., Ingber, 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=Mechanotransduction+across+the+cell-surface+and+through+the+cytoskeleton.">Mechanotransduction across the cell-surface and through the cytoskeleton.</a> Science. 260, 1124-1127 (1993).</li><li>Liu, M., Tanswell, A. K., Post, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanical+force-induced+signal+transduction+in+lung+cells.">Mechanical force-induced signal transduction in lung cells.</a> Am J Physiol. 277, L667-L683 (1999).</li><li>Janmey, P. A., McCulloch, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+mechanics:+integrating+cell+responses+to+mechanical+stimuli.">Cell mechanics: integrating cell responses to mechanical stimuli.</a> Annu Rev Biomed Eng. 9, 1-34 (2007).</li><li>Waters, 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=Reactive+oxygen+species+in+mechanotransduction.">Reactive oxygen species in mechanotransduction.</a> Am J Physiol Lung Cell Mol Physiol. 287, L484-L485 (2004).</li><li>Chapman, K. 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=Cyclic+mechanical+strain+increases+reactive+oxygen+species+production+in+pulmonary+epithelial+cells.">Cyclic mechanical strain increases reactive oxygen species production in pulmonary epithelial cells.</a> Am J Physiol Lung Cell Mol Physiol. 289, L834-L841 (2005).</li><li>Chu, E. K., Whitehead, T., Slutsky, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+cyclic+opening+and+closing+at+low-+and+high-volume+ventilation+on+bronchoalveolar+lavage+cytokines.">Effects of cyclic opening and closing at low- and high-volume ventilation on bronchoalveolar lavage cytokines.</a> Crit Car Med. 32, 168-174 (2004).</li><li>Tschumperlin, D., Margulies, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Equibiaxial+deformation-induced+injury+of+alveolar+epithelial+cells+in+vitro.">Equibiaxial deformation-induced injury of alveolar epithelial cells in vitro.</a> Am J Physiol. 275, L1173-L1183 (1998).</li><li>Vlahakis, N. E., Hubmayr, R. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cellular+stress+failure+in+ventilator-injured+lungs.">Cellular stress failure in ventilator-injured lungs.</a> Am J Respir Crit Care Med. 171, 1328-1342 (2005).</li><li>Roan, 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=Hyperoxia+alters+the+mechanical+properties+of+alveolar+epithelial+cells.">Hyperoxia alters the mechanical properties of alveolar epithelial cells.</a> Am J Physiol Lung Cell Mol Physiol. 302, L1235-L1241 (2012).</li><li>Gamerdinger, K., 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+load+and+mechanical+integrity+of+lung+cells+-+Experimental+mechanostimulation+of+epithelial+cell-+and+fibroblast-monolayers.">Mechanical load and mechanical integrity of lung cells - Experimental mechanostimulation of epithelial cell- and fibroblast-monolayers.</a> J Mech Behav Biomed Mater. 4, 201-209 (2014).</li><li>Hayashi, K., Naiki, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adaptation+and+remodeling+of+vascular+wall;+biomechanical+response+to+hypertension.">Adaptation and remodeling of vascular wall; biomechanical response to hypertension.</a> J Mech Behav Biomed Mater. 2, 3-19 (2009).</li><li>Villemure, I., Stokes, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Growth+plate+mechanics+and+mechanobiology.+A+survey+of+present+understanding.">Growth plate mechanics and mechanobiology. A survey of present understanding.</a> J Biomech. 42, 1793-1803 (2009).</li><li>Waters, C. M., 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+system+to+impose+prescribed+homogenous+strains+on+cultured+cells.">A system to impose prescribed homogenous strains on cultured cells.</a> J Appl Physiol (1985). 91, 1600-1610 (2001).</li><li>Gerstmair, A., Fois, G., Innerbichler, S., Dietl, P., Felder, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+device+for+simultaneous+live+cell+imaging+during+uni-axial+mechanical+strain+or+compression.">A device for simultaneous live cell imaging during uni-axial mechanical strain or compression.</a> J Appl Physiol (1985). 107, 613-620 (1985).</li><li>Dassow, C., 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+method+to+measure+mechanical+properties+of+pulmonary+epithelial+cell+layers.">A method to measure mechanical properties of pulmonary epithelial cell layers.</a> J. Biomed. Mater. Res. Part B Appl. Biomater. 101, 1164-1171 (2013).</li><li>Chapman, K., 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=Cyclic+mechanical+strain+increases+reactive+oxygen+species+production+in+pulmonary+epithelial+cells.">Cyclic mechanical strain increases reactive oxygen species production in pulmonary epithelial cells.</a> Am J Physiol Lung Cell Mol Physiol. 289, L834-L841 (2005).</li><li>Birukov, K. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cyclic+stretch,+reactive+oxygen+species,+and+vascular+remodeling.">Cyclic stretch, reactive oxygen species, and vascular remodeling.</a> Antioxid Redox Signal. 11, 1651-1667 (2009).</li><li>Turrens, J. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+formation+of+reactive+oxygen+species.">Mitochondrial formation of reactive oxygen species.</a> J Physiol. 552, 335-344 (2003).</li><li>Wang, W., 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=Superoxide+flashes+in+single+mitochondria.">Superoxide flashes in single mitochondria.</a> Cell. 134, 279-290 (2008).</li><li>Pouvreau, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Superoxide+flashes+in+mouse+skeletal+muscle+are+produced+by+discrete+arrays+of+active+mitochondria+operating+coherently.">Superoxide flashes in mouse skeletal muscle are produced by discrete arrays of active mitochondria operating coherently.</a> PLoS One. 5, (2010).</li><li>Yalcin, H. C., 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=Influence+of+cytoskeletal+structure+and+mechanics+on+epithelial+cell+injury+during+cyclic+airway+reopening.">Influence of cytoskeletal structure and mechanics on epithelial cell injury during cyclic airway reopening.</a> Am J Physiol Lung Cell Mol Physiol. 297, L881-L891 (2009).</li><li>Jacob, A. M., Gaver, D. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atelectrauma+disrupts+pulmonary+epithelial+barrier+integrity+and+alters+the+distribution+of+tight+junction+proteins+ZO-1+and+claudin+4.">Atelectrauma disrupts pulmonary epithelial barrier integrity and alters the distribution of tight junction proteins ZO-1 and claudin 4.</a> J Appl Physiol. 113, 1377-1387 (2012).</li><li>DiPaolo, B. C., Lenormand, G., Fredberg, J. J., Margulies, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stretch+magnitude+and+frequency-dependent+actin+cytoskeleton+remodeling+in+alveolar+epithelia.">Stretch magnitude and frequency-dependent actin cytoskeleton remodeling in alveolar epithelia.</a> Am J Physiol Cell Physiol. 299, C345-C353 (2010).</li></ol>

The authors declare that they have no competing financial interests.

A unique device for live cell imaging during mechanical stretch was developed; and this device was used in a protocol to study lung epithelial cell mechanobiology. In preliminary studies, it was found that a single held stretch stimulated the production of mitochondrial superoxide in bronchial epithelial cells. In addition, it was demonstrated that increased levels of mechanical strain caused direct damage to the integrity of a monolayer of alveolar epithelial cells.
To conduct these preliminary experiments, the device was first calibrated, and then it was shown that the membrane strain was transmitted to cells (Figures 2-3). Overall the device performed well, as indicated by the close correlation between the membrane strain (measured by marks on the membrane) and the cell strain (measured by distances between nuclei) shown in Figure 3E. However, there were some minor differences observed between strain measured on the membranes during the approach and the return curves of the markers, as shown in Figure 2D. This was likely due to backlash in the device and possibly related to gripping issues. In addition, a universal mounting mechanism for the device is needed so that it can efficiently be used on microscopes with different configurations. This would reduce the time spent between positioning of the device and image acquisition while maintaining a robust connection to the microscope. Holes punched in the membranes were used to apply a minor pre-strain to the membrane to prevent formation of an initial compressive state due to clamping of an elastic material. The compressive state also leads to a reduction of maximum applicable strain.
The custom device was designed to fit in an Asylum MFP3D Atomic Force Microscope to allow nano-indentation of live cells during mechanical stretch. We are not aware of any other device that is capable of inducing an isotropic stress state in the imaging plane that can function within an AFM. The device was also designed for continuous use in a cell-culture incubator (kept at 37 &#176;C in a humid, sterile environment with 5% CO2). The ability to function within an incubator allows the study of cells undergoing long-term stretch regimens.
There are several limitations to the presented approach. First, as the membrane stretches, the field of focus moves out of plane, although minimally (~100 &#181;m). This becomes a limitation when tracking a field of cells through a stretch regimen, because the field of view changes over the course of the membrane distention. Even though membrane within the 22 mm diameter experiences 0.23-0.25 strain, the regions outside of this inner diameter experience a more variable strain field. Although imaging studies are easily limited to this central region, it is possible that the cells responding to higher levels of strain at outer regions can impact the response of cells within the central region. A uniform strain distribution over a larger area of interest can be achieved with improvements in the membrane design. The PDMS membrane is auto-fluorescent at wavelengths commonly used for fluorescent indicators, and this limits the capability for imaging fluorescence in cells from beneath the membrane. For this reason an upright microscope was utilized with water immersion objectives in our studies of ROS production described above. This can be a limitation for studies in which AFM and fluorescence imaging are combined.
The most common commercial device (FlexCell FX 5000 Tension System) for applying mechanical strain to cells incorporates a controllable vacuum system applying tensile strain on a membrane while moving in and out of the optical plane. The membrane and clamp design presented herein create a bi-axial strain field where strain is isotropic and remains in plane allowing real-time imaging. With minor modifications to the membrane design, the stretcher can produce uniaxial stretch modes as well. If the cells are first seeded on a distended membrane, in-plane compressive loads can also be applied. The device is able to reach both physiological and pathological levels of strain. In summary, a new device and protocols were developed that can be used to apply mechanical strain to live cells that can be imaged using fluorescence microscopy and AFM nano-indentation.

Cells are subjected to mechanical loads in many tissues, and this mechanical stimulation has been shown to promote changes in patterns of gene expression, release of growth factors, cytokines, or remodeling of the extracellular matrix and cytoskeleton1-4. The intracellular signals transduced from such mechanical stimuli occur through the process of mechanotransduction5-7. In the respiratory system, one outcome of mechanotransduction is the increase in reactive oxygen species (ROS)8,9 and pro-inflammatory cytokines10 in pulmonary epithelial cells in the presence of cyclic tensile strain. Strong evidence also suggests that excessive tensile strain leads to direct injury to the alveolar epithelium, in addition to the biochemical responses of cells11-14. Although the focus here is primarily on the response of lung cells to mechanical deformation, pathways induced by mechanotransduction play a key role in the basic function of many tissues in the human body, including the regulation of vascular tone15 and the development of the growth plate16.

The growing interest in mechanotransduction has resulted in the development of numerous devices for the application of physiologically relevant mechanical loads to cultured cells and tissue. In particular, devices applying a tensile strain, which is a common form of mechanical loading experienced by tissue, are popular 11,17-19. However, many of the available devices are either designed as a bioreactor for tissue engineering applications or are not conducive to real time imaging with stretch. As such, there is a need to develop tools and methods that can visualize cells and tissues in tension to facilitate the investigation of pathways of mechanotransduction.
Herein, an in-plane mechanical stretching device was designed and protocols were developed to apply multiple forms of strain to tissues and cells while allowing imaging of the biochemical and mechanical responses in real time (Figure 1A-D). The device utilizes six evenly spaced clamps arranged circumferentially to grasp a flexible membrane and apply an in-plane, radial distention up to approximately 20% (Figure 1B). The actuating device can be placed in a cell culture incubator for an extended period of time, while the motor (Figure 1C) is positioned outside the incubator and controlled by proprietary software provided by the motor supplier. The motor is connected to a linear driver, which rotates an internal cam, driving the six stretcher clamps uniformly in tension and relaxation.
In addition to the mechanical device, customized flexible membranes were created from commercially available cell culture ready membranes to be used in the mechanical system. Then circular walls (with a diameter of approximately 28 mm) were made and attached on to the flexible membrane so that cells could be cultured only in this region of well-described strain profile. In order to determine whether the placement of these membranes within the actuating device would provide uniform and isotropic strain in the center of the flexible membrane, finite element analysis was conducted using commercially available software (Figure 1E-F). The flexible membrane was modeled with symmetric boundary conditions and utilizing all quadrilateral elements for the mesh. The concentric rings seen in the contour plot of maximum principal strain shown in Figure 1F indicate the isotropic distribution of the strain.
The strain experienced by the membrane was measured by recording images of markings through loading (Figure 2). Figure 2D shows that the average membrane strain measured in radial and axial directions was approximately linear with respect to the applied motor counts up to a maximum linear strain of 20%. There was no significant difference between the strain levels measured during distention compared with those measured during retraction back to the resting position. Next, the displacement of human bronchial epithelial cells (16HBE) and their nuclei cultured on the custom flexible membrane were measured. Fluorescently labeled (DAPI) nuclei of the 16HBE cells were imaged using a 20X objective under a confocal microscope, whereas whole cell displacement was measured with phase contrast images recorded with a digital microscope. As seen in Figure 3, the strain measured by displacement of nuclei was similar to that measured by displacement of markings on the membrane, up to ~20% linear strain. This confirms that the strain applied to the membranes was transmitted to the adherent cells. The protocols describing the use of the custom device on a traditional microscope and an atomic force microscope are provided in the following steps.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>SmartMotor NEMA 34: 3400 Series</td>
			<td>MOOG Animatics</td>
			<td>SM3416D</td>
			<td>Integrated motor, controller, amplifier, encoder and communications bus</td>
		</tr>
		<tr>
			<td>Flexcell Membrane (Collagen I coated)</td>
			<td>Flexcell International Corp</td>
			<td>SM2-1010C</td>
			<td>3.5x5.25x0.020&#34;</td>
		</tr>
		<tr>
			<td>Sylgard 184&#160;</td>
			<td>Dow Corning Corporation</td>
			<td></td>
			<td>10:1</td>
		</tr>
		<tr>
			<td>Hoechst 33342&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>H1399</td>
			<td>DAPI stain</td>
		</tr>
		<tr>
			<td>MitoSOX</td>
			<td>Sigma-Aldrich</td>
			<td>M36008</td>
			<td></td>
		</tr>
		<tr>
			<td>Tiron</td>
			<td>Sigma-Aldrich</td>
			<td>D7389&#160;</td>
			<td>mitochondrial superoxide label</td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td></td>
			<td></td>
			<td>superoxide inhibitor</td>
		</tr>
		<tr>
			<td>FBS</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>50 ml tubes</td>
			<td>Fisher Scientific</td>
			<td>06-443-19</td>
			<td>Any centriguge tube can be used to create an area for imaging.</td>
		</tr>
		<tr>
			<td>Hybridization oven</td>
			<td>Bellco Glass</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MLE12 Cells</td>
			<td>ATCC</td>
			<td>CRL-2110</td>
			<td>Mouse Lung Epithelial Cells&#160;</td>
		</tr>
		<tr>
			<td>16HBE cells</td>
			<td>ATCC</td>
			<td>CRL-2741</td>
			<td>Human Bronchial Epithelial Cells</td>
		</tr>
		<tr>
			<td>AFM Indentation Experiments</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cantilever Beams for Nano-indentation</td>
			<td>Budget Sensors</td>
			<td>Si-Ni30</td>
			<td></td>
		</tr>
		<tr>
			<td>AFM&#160;</td>
			<td>Asylum Research</td>
			<td>MFP3D</td>
			<td></td>
		</tr>
		<tr>
			<td>Olympus microscope</td>
			<td>Olympus</td>
			<td>IX-71</td>
			<td>Inverted microscope with 20X and 40X objectives.</td>
		</tr>
		<tr>
			<td>AFM Leg Extenders</td>
			<td>Asylum Research</td>
			<td>Not available</td>
			<td>AFM microscope</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Finite Element Analyses</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ABAQUS</td>
			<td>Simulia</td>
			<td>6.12</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Software</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td>NIH</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microscopes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Digital microscope</td>
			<td>Life Technologies</td>
			<td>EVOS XL Core</td>
			<td>Initially a self standing company, now owned by Life Technologies.</td>
		</tr>
		<tr>
			<td>Confocal microscope</td>
			<td>Zeiss</td>
			<td>LSM 710</td>
			<td>2-photon upright microscope</td>
		</tr>
	</tbody>
</table>

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.

Reactive Oxygen Species and Deformation
Previous studies have shown increases in reactive oxygen species (ROS) in airway and alveolar epithelial cells in response to cyclic stretch21. Reactive oxygen species include molecules and free radicals derived from molecular oxygen with high reactivity to lipids, proteins, polysaccharides, and nucleic acids22-24. ROS serve as a common intracellular signal to regulate ion channel function, protein kinase/phosphatase activation, and gene expression, but excessive or unregulated production can contribute to apoptotic and necrotic cell death, neurodegeneration, atherosclerosis, diabetes, and cancer24,25. Superoxide, a pre-cursor for other forms of ROS, can be produced as a by-product of mitochondrial respiration when electrons leak from the electron transfer chain. Previous studies suggested that cyclic mechanical stretch stimulated the production of superoxide through a combination of the NADPH oxidase system (nicotinamide adenine dinucleotide phosphate-oxidase) and the mitochondrial electron transport chain (complex I and III). It has been proposed that this stimulation was due to a direct distortion of the mitochondria21 possibly due to connections to the cell cytoskeleton. In order to test this hypothesis, the cell stretching device was developed so that we could image changes in ROS production in live cells in response to mechanical stretch. Here, the mitochondrial ROS produced by bronchial epithelial cells due to stretch was measured using the custom device and protocol. In the absence of mechanical stretch, ROS production did not significantly increase over 60 min (Figure 4). When a single 17% stretch was applied and maintained, there was an increase in mitochondrial ROS that persisted for another 60 min.
Direct Epithelial Monolayer Damage Due to Stretch
Indirect mechanisms by which excessive stretch in the lung epithelium harms the lungs have been discussed in depth elsewhere2. Several studies suggest that excessive mechanical stretch of lung epithelial cells can cause direct injury involving damage to cells11,13,26,27. This type of damage is difficult to capture without real-time imaging during the mechanical distention. As such, utilizing the custom device and phase contrast microscopy consecutive images of cells that were mechanically stretched were recorded before and after treatment with a pro-inflammatory cytokine (50 ng/ml TNF-&#945; for 6 hr) (Figure 5). Images of the same field of cells were captured at increasing strain levels. The images show the formation of a gap when the cells were stretched between 10 and 15% strain as indicated by the red arrows. It is important to note that these images were recorded in near real-time, &#60;10 sec of lag between the end of stretch and imaging, while the cells were stretched. In previous studies of cell imaging of stretched cells, images were obtained before and after stretch comparing only the images of cells not at stretched condition11,13,28.
AFM Nano-indentation on stretched cells
The mechanical device was placed in the AFM with the use of an adapter plate (see Figure 6A). Using previously published methods13, elastic modulus maps (E-maps) of MLE12 cells before and after 10% tensile strain were obtained (Figure 6D-E). Although phase contrast image at 10% strain (Figure 6D) was obtained within 10 sec of the stretch, the E-maps shown in Figure 6D-E consumed approximately 22 min&#160;with 300 individual force-deflection curves recorded over a 40 &#181;m X 40 &#181;m area with a 2.5 &#181;m/s velocity of the tip in the z-direction. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52737/52737fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 1" src="/files/ftp_upload/52737/52737fig1.jpg" /> 
Figure 1. The custom designed device for cell culture and stretching. (A) Computer-generated drawing of the mechanical design of the device, (B) photograph of the completed device, (C) photograph of device connected to the motor and linear driver, which converts the motor rotation into a 1-dimensional motion which controls the clamp biaxial stretch of the (D) silicone rubber membrane. The substrate contains two holes on each clamp tab to be placed on posts on each clamp. (E) Dimensioned FEA model of the flexible membrane that is 1/4 of the whole structure due to use of symmetric boundary conditions. (F) Maximum principal strain is shown in the full membrane depicting the formation of rings indicative of isotropic strain field. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52737/52737fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52737/52737fig2.jpg" /> 
Figure 2. (A) The device supporting a flexible membrane with markings applied to measure applied strain is shown. Close-up images of the markings, before (B) and after (C) stretch, were labeled with an arrow to clearly indicate the displacement of the markings on the membrane. (D) Membrane strain as a function of the motor counts during approach and retraction. Membrane strain was similar in both approach and retraction with retraction producing slightly higher strains. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52737/52737fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52737/52737fig3.jpg" /> 
Figure 3. Membrane strain transmission to the cells. (A-B) Phase contrast images of human bronchial epithelial cells (16HBE) on the flexible membrane using a 10X objective, before (A) and after (B) 20% strain. (C-D) Fluorescently labeled (DAPI) nuclei of the 16HBE cells were imaged using a 20X objective under a confocal microscope, before (C) and after (D) 20% strain. (E) The membrane strain experienced by cells was linear and homogenous. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52737/52737fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/52737/52737fig4.jpg" /> 
Figure 4. The impact of a single stretch on the ROS production of 16HBE cells was observed using a cumulative mitochondrial superoxide sensor using a 20X objective under a confocal microscope. (A-C) Snapshots of the time course (0, 60 min, and ~65 min) of superoxide production during stretch in the same field of cells. (D) Relative fluorescence intensity of 16HBE cells over time indicating a 50% increase in fluorescence intensity from baseline production of superoxide within the first hour (A to B). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52737/52737fig4highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/52737/52737fig5.jpg" /> 
Figure 5. Phase contrast images of an alveolar epithelial (MLE12) monolayer response to increasing stretch.&#160;(A-D) were all recorded using a digital microscope with a 20X objective. The arrows indicate the locations where cell-to-cell separation occurred. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52737/52737fig5highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" src="/files/ftp_upload/52737/52737fig6.jpg" /> 
Figure 6. (A) Mechanical device shown in the AFM for nano-indentation of cells with stretch. The scanner must be removed for the placement of objective extender. Three leg extenders are attached to all three legs of the AFM head to allow for device placement. An adapter plate is utilized to mount the stretcher on to the AFM scanner (black plate). The cantilever beam scans in the direction of the arrow shown in the before stretch phase contrast image an MLE12 cell (B). The red squares in the before (B) and after (C) stretch images depict the area that was captured during nano-indentation (40X objective). The elastic modulus maps, before (D) and after (E) stretch, are obtained with previously published analysis techniques13. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52737/52737fig6highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Live Cell Imaging during Mechanical Stretch at JoVE.com

Live Cell Imaging during Mechanical Stretch

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 ...

live-cell-imaging-during-mechanical-stretch

Nanyang Technological University

Research

JoVE Journal

Bioengineering

10.3K Views.  University of Tennessee Health Science Center. 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. 

Video: Live Cell Imaging during Mechanical Stretch

Axon Stretch Growth: The Mechanotransduction of Neuronal Growth

During pre-synaptic embryonic development, neuronal processes traverse short distances to reach their targets via growth cone. Over time, neuronal somata are separated from their axon terminals due to skeletal growth of the enlarging organism (Weiss 1941; Gray, Hukkanen et al. 1992). This mechanotransduction induces a secondary mode of neuronal growth capable of accommodating continual elongation of the axon (Bray 1984; Heidemann and Buxbaum 1994; Heidemann, Lamoureux et al. 1995; Pfister, Iwata et al. 2004).
Axon Stretch Growth (ASG) is conceivably a central factor in the maturation of short embryonic processes into the long nerves and white matter tracts characteristic of the adult nervous system. To study ASG in vitro, we engineered bioreactors to apply tension to the short axonal processes of neuronal cultures (Loverde, Ozoka et al. 2011). Here, we detail the methods we use to prepare bioreactors and conduct ASG. First, within each stretching lane of the bioreactor, neurons are plated upon a micro-manipulated towing substrate. Next, neurons regenerate their axonal processes, via growth cone extension, onto a stationary substrate. Finally, stretch growth is performed by towing the plated cell bodies away from the axon terminals adhered to the stationary substrate; recapitulating skeletal growth after growth cone extension.
Previous work has shown that ASG of embryonic rat dorsal root ganglia neurons are capable of unprecedented growth rates up to 10mm/day, reaching lengths of up to 10cm; while concurrently resulting in increased axonal diameters (Smith, Wolf et al. 2001; Pfister, Iwata et al. 2004; Pfister, Bonislawski et al. 2006; Pfister, Iwata et al. 2006; Smith 2009). This is in dramatic contrast to regenerative growth cone extension (in absence of mechanical stimuli) where growth rates average 1mm/day with successful regeneration limited to lengths of less than 3cm (Fu and Gordon 1997; Pfister, Gordon et al. 2011). Accordingly, further study of ASG may help to reveal dysregulated growth mechanisms that limit regeneration in the absence of mechanical stimuli.

A unique tissue engineering method was developed to elongate numerous nerve fibers in culture by recapitulating axon stretch growth; a form of nervous system growth whereby nerves elongate in conjunction with growth of the enlarging body.

During pre-synaptic embryonic development, neuronal processes traverse short distances to reach their targets via growth cone. Over time, neuronal somata ...

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 ...

Monitoring the Wall Mechanics During Stent Deployment in a Vessel

Clinical trials have reported different restenosis rates for various stent designs1. It is speculated that stent-induced strain concentrations on the arterial wall lead to tissue injury, which initiates restenosis2-7. This hypothesis needs further investigations including better quantifications of non-uniform strain distribution on the artery following stent implantation. A non-contact surface strain measurement method for the stented artery is presented in this work. ARAMIS stereo optical surface strain measurement system uses two optical high speed cameras to capture the motion of each reference point, and resolve three dimensional strains over the deforming surface8,9. As a mesh stent is deployed into a latex vessel with a random contrasting pattern sprayed or drawn on its outer surface, the surface strain is recorded at every instant of the deformation. The calculated strain distributions can then be used to understand the local lesion response, validate the computational models, and formulate hypotheses for further in vivo study.

Stent-induced arterial strain distributions are characterized using an optical surface strain measurement system. This visualization technique is used to gain insights into the impact of stent implantation on the host vessel.

Clinical trials have reported different restenosis rates for various stent designs1. It is speculated that stent-induced strain concentrations on the ...

High Throughput Single-cell and Multiple-cell Micro-encapsulation

Microfluidic encapsulation methods have been previously utilized to capture cells in picoliter-scale aqueous, monodisperse drops, providing confinement from a bulk fluid environment with applications in high throughput screening, cytometry, and mass spectrometry. We describe a method to not only encapsulate single cells, but to repeatedly capture a set number of cells (here we demonstrate one- and two-cell encapsulation) to study both isolation and the interactions between cells in groups of controlled sizes. By combining drop generation techniques with cell and particle ordering, we demonstrate controlled encapsulation of cell-sized particles for efficient, continuous encapsulation. Using an aqueous particle suspension and immiscible fluorocarbon oil, we generate aqueous drops in oil with a flow focusing nozzle. The aqueous flow rate is sufficiently high to create ordering of particles which reach the nozzle at integer multiple frequencies of the drop generation frequency, encapsulating a controlled number of cells in each drop. For representative results, 9.9 &mu;m polystyrene particles are used as cell surrogates. This study shows a single-particle encapsulation efficiency Pk=1 of 83.7% and a double-particle encapsulation efficiency Pk=2 of 79.5% as compared to their respective Poisson efficiencies of 39.3% and 33.3%, respectively. The effect of consistent cell and particle concentration is demonstrated to be of major importance for efficient encapsulation, and dripping to jetting transitions are also addressed. 
Introduction 
Continuous media aqueous cell suspensions share a common fluid environment which allows cells to interact in parallel and also homogenizes the effects of specific cells in measurements from the media. High-throughput encapsulation of cells into picoliter-scale drops confines the samples to protect drops from cross-contamination, enable a measure of cellular diversity within samples, prevent dilution of reagents and expressed biomarkers, and amplify signals from bioreactor products. Drops also provide the ability to re-merge drops into larger aqueous samples or with other drops for intercellular signaling studies.1,2 The reduction in dilution implies stronger detection signals for higher accuracy measurements as well as the ability to reduce potentially costly sample and reagent volumes.3 Encapsulation of cells in drops has been utilized to improve detection of protein expression,4 antibodies,5,6 enzymes,7 and metabolic activity8 for high throughput screening, and could be used to improve high throughput cytometry.9 Additional studies present applications in bio-electrospraying of cell containing drops for mass spectrometry10 and targeted surface cell coatings.11 Some applications, however, have been limited by the lack of ability to control the number of cells encapsulated in drops. Here we present a method of ordered encapsulation12 which increases the demonstrated encapsulation efficiencies for one and two cells and may be extrapolated for encapsulation of a larger number of cells.
To achieve monodisperse drop generation, microfluidic "flow focusing" enables the creation of controllable-size drops of one fluid (an aqueous cell mixture) within another (a continuous oil phase) by using a nozzle at which the streams converge.13 For a given nozzle geometry, the drop generation frequency f and drop size can be altered by adjusting oil and aqueous flow rates Qoil and Qaq. As the flow rates increase, the flows may transition from drop generation to unstable jetting of aqueous fluid from the nozzle.14 
When the aqueous solution contains suspended particles, particles become encapsulated and isolated from one another at the nozzle. For drop generation using a randomly distributed aqueous cell suspension, the average fraction of drops Dk containing k cells is dictated by Poisson statistics, where Dk = &lambda;k exp(-&lambda;)/(k!) and &lambda; is the average number of cells per drop. The fraction of cells which end up in the "correctly" encapsulated drops is calculated using Pk = (k x Dk)/&Sigma;(k' x Dk'). The subtle difference between the two metrics is that Dk relates to the utilization of aqueous fluid and the amount of drop sorting that must be completed following encapsulation, and Pk relates to the utilization of the cell sample. As an example, one could use a dilute cell suspension (low &lambda;) to encapsulate drops where most drops containing cells would contain just one cell. While the efficiency metric Pk would be high, the majority of drops would be empty (low Dk), thus requiring a sorting mechanism to remove empty drops, also reducing throughput.15
Combining drop generation with inertial ordering provides the ability to encapsulate drops with more predictable numbers of cells per drop and higher throughputs than random encapsulation. Inertial focusing was first discovered by Segre and Silberberg16 and refers to the tendency of finite-sized particles to migrate to lateral equilibrium positions in channel flow. Inertial ordering refers to the tendency of the particles and cells to passively organize into equally spaced, staggered, constant velocity trains. Both focusing and ordering require sufficiently high flow rates (high Reynolds number) and particle sizes (high Particle Reynolds number).17,18 Here, the Reynolds number Re =uDh/&nu; and particle Reynolds number Rep =Re(a/Dh)2, where u is a characteristic flow velocity, Dh [=2wh/(w+h)] is the hydraulic diameter, &nu; is the kinematic viscosity, a is the particle diameter, w is the channel width, and h is the channel height. Empirically, the length required to achieve fully ordered trains decreases as Re and Rep increase. Note that the high Re and Rep requirements (for this study on the order of 5 and 0.5, respectively) may conflict with the need to keep aqueous flow rates low to avoid jetting at the drop generation nozzle. Additionally, high flow rates lead to higher shear stresses on cells, which are not addressed in this protocol. The previous ordered encapsulation study demonstrated that over 90% of singly encapsulated HL60 cells under similar flow conditions to those in this study maintained cell membrane integrity.12 However, the effect of the magnitude and time scales of shear stresses will need to be carefully considered when extrapolating to different cell types and flow parameters. The overlapping of the cell ordering, drop generation, and cell viability aqueous flow rate constraints provides an ideal operational regime for controlled encapsulation of single and multiple cells.
Because very few studies address inter-particle train spacing,19,20 determining the spacing is most easily done empirically and will depend on channel geometry, flow rate, particle size, and particle concentration. Nonetheless, the equal lateral spacing between trains implies that cells arrive at predictable, consistent time intervals. When drop generation occurs at the same rate at which ordered cells arrive at the nozzle, the cells become encapsulated within the drop in a controlled manner. This technique has been utilized to encapsulate single cells with throughputs on the order of 15 kHz,12 a significant improvement over previous studies reporting encapsulation rates on the order of 60-160 Hz.4,15 In the controlled encapsulation work, over 80% of drops contained one and only one cell, a significant efficiency improvement over Poisson (random) statistics, which predicts less than 40% efficiency on average.12
In previous controlled encapsulation work,12 the average number of particles per drop &lambda; was tuned to provide single-cell encapsulation. We hypothesize that through tuning of flow rates, we can efficiently encapsulate any number of cells per drop when &lambda; is equal or close to the number of desired cells per drop. While single-cell encapsulation is valuable in determining individual cell responses from stimuli, multiple-cell encapsulation provides information relating to the interaction of controlled numbers and types of cells. Here we present a protocol, representative results using polystyrene microspheres, and discussion for controlled encapsulation of multiple cells using a passive inertial ordering channel and drop generation nozzle.

Combining monodisperse drop generation with inertial ordering of cells and particles, we describe a method to encapsulate a desired number of cells or particles in a single drop at kHz rates. We demonstrate efficiencies twice exceeding those of unordered encapsulation for single- and double-particle drops.

Microfluidic encapsulation methods have been previously utilized to capture cells in picoliter-scale aqueous, monodisperse drops, providing confinement ...

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of the Drosophila larva makes it an attractive model organism for live imaging studies. However, an important challenge for live imaging techniques is to noninvasively immobilize and position an animal on the microscope. This protocol presents a simple and easy to use method for immobilizing and imaging Drosophila larvae on a polydimethylsiloxane (PDMS) microfluidic device, which we call the 'larva chip'. The larva chip is comprised of a snug-fitting PDMS microchamber that is attached to a thin glass coverslip, which, upon application of a vacuum via a syringe, immobilizes the animal and brings ventral structures such as the nerve cord, segmental nerves, and body wall muscles, within close proximity to the coverslip. This allows for high-resolution imaging, and importantly, avoids the use of anesthetics and chemicals, which facilitates the study of a broad range of physiological processes. Since larvae recover easily from the immobilization, they can be readily subjected to multiple imaging sessions. This allows for longitudinal studies over time courses ranging from hours to days. This protocol describes step-by-step how to prepare the chip and how to utilize the chip for live imaging of neuronal events in 3rd instar larvae. These events include the rapid transport of organelles in axons, calcium responses to injury, and time-lapse studies of the trafficking of photo-convertible proteins over long distances and time scales. Another application of the chip is to study regenerative and degenerative responses to axonal injury, so the second part of this protocol describes a new and simple procedure for injuring axons within peripheral nerves by a segmental nerve crush.

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Drosophila larvae are an attractive model system for live imaging due to their translucent cuticle and powerful genetics. This protocol describes how to utilize a single-layer PDMS device, called the &#39;larva chip&#39; for live imaging of cellular processes within neurons of 3rd instar Drosophila larvae.

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of ...

A Microfluidic Technique to Probe Cell Deformability

Here we detail the design, fabrication, and use of a microfluidic device to evaluate the deformability of a large number of individual cells in an efficient manner. Typically, data for ~102 cells can be acquired within a 1 hr experiment. An automated image analysis program enables efficient post-experiment analysis of image data, enabling processing to be complete within a few hours. Our device geometry is unique in that cells must deform through a series of micron-scale constrictions, thereby enabling the initial deformation and time-dependent relaxation of individual cells to be assayed. The applicability of this method to human promyelocytic leukemia (HL-60) cells is demonstrated. Driving cells to deform through micron-scale constrictions using pressure-driven flow, we observe that human promyelocytic (HL-60) cells momentarily occlude the first constriction for a median time of 9.3 msec before passaging more quickly through the subsequent constrictions with a median transit time of 4.0 msec per constriction. By contrast, all-trans retinoic acid-treated (neutrophil-type) HL-60 cells occlude the first constriction for only 4.3 msec before passaging through the subsequent constrictions with a median transit time of 3.3 msec. This method can provide insight into the viscoelastic nature of cells, and ultimately reveal the molecular origins of this behavior.

We demonstrate a microfluidics-based assay to measure the timescale for cells to transit through a sequence of micron-scale constrictions.

Here we detail the design, fabrication, and use of a microfluidic device to evaluate the deformability of a large number of individual cells in an ...

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 ...

Using Cell-substrate Impedance and Live Cell Imaging to Measure Real-time Changes in Cellular Adhesion and De-adhesion Induced by Matrix Modification

Cell-matrix adhesion plays a key role in controlling cell morphology and signaling. Stimuli that disrupt cell-matrix adhesion (e.g., myeloperoxidase and other matrix-modifying oxidants/enzymes released during inflammation) are implicated in triggering pathological changes in cellular function, phenotype and viability in a number of diseases. Here, we describe how cell-substrate impedance and live cell imaging approaches can be readily employed to accurately quantify real-time changes in cell adhesion and de-adhesion induced by matrix modification (using endothelial cells and myeloperoxidase as a pathophysiological matrix-modifying stimulus) with high temporal resolution and in a non-invasive manner. The xCELLigence cell-substrate impedance system continuously quantifies the area of cell-matrix adhesion by measuring the electrical impedance at the cell-substrate interface in cells grown on gold microelectrode arrays. Image analysis of time-lapse differential interference contrast movies quantifies changes in the projected area of individual cells over time, representing changes in the area of cell-matrix contact. Both techniques accurately quantify rapid changes to cellular adhesion and de-adhesion processes. Cell-substrate impedance on microelectrode biosensor arrays provides a platform for robust, high-throughput measurements. Live cell imaging analyses provide additional detail regarding the nature and dynamics of the morphological changes quantified by cell-substrate impedance measurements. These complementary approaches provide valuable new insights into how myeloperoxidase-catalyzed oxidative modification of subcellular extracellular matrix components triggers rapid changes in cell adhesion, morphology and signaling in endothelial cells. These approaches are also applicable for studying cellular adhesion dynamics in response to other matrix-modifying stimuli and in related adherent cells (e.g., epithelial cells).

Here, we present a protocol to continuously quantify cell adhesion and de-adhesion processes with high temporal resolution in a non-invasive manner by cell-substrate impedance and live cell imaging analyses. These approaches reveal the dynamics of cell adhesion/de-adhesion processes triggered by matrix modification and their temporal relationship to adhesion-dependent signaling events.

Cell-matrix adhesion plays a key role in controlling cell morphology and signaling. Stimuli that disrupt cell-matrix adhesion (e.g., myeloperoxidase and ...

Magnetic Levitation Coupled with Portable Imaging and Analysis for Disease Diagnostics

Currently, many clinical diagnostic procedures are complex, costly, inefficient, and inaccessible to a large population in the world. The requirements for specialized equipment and trained personnel require that many diagnostic tests be performed at remote, centralized clinical laboratories. Magnetic levitation is a simple yet powerful technique and can be applied to levitate cells, which are suspended in a paramagnetic solution and placed in a magnetic field, at a position determined by equilibrium between a magnetic force and a buoyancy force. Here, we present a versatile platform technology designed for point-of-care diagnostics which uses magnetic levitation coupled to microscopic imaging and automated analysis to determine the density distribution of a patient's cells as a useful diagnostic indicator. We present two platforms operating on this principle: (i) a smartphone-compatible version of the technology, where the built-in smartphone camera is used to image cells in the magnetic field and a smartphone application processes the images and to measures the density distribution of the cells and (ii) a self-contained version where a camera board is used to capture images and an embedded processing unit with attached thin-film-transistor (TFT) screen measures and displays the results. Demonstrated applications include: (i) measuring the altered distribution of a cell population with a disease phenotype compared to a healthy phenotype, which is applied to sickle cell disease diagnosis, and (ii) separation of different cell types based on their characteristic densities, which is applied to separate white blood cells from red blood cells for white blood cell cytometry. These applications, as well as future extensions of the essential density-based measurements enabled by this portable, user-friendly platform technology, will significantly enhance disease diagnostic capabilities at the point of care.

We present a magnetic levitation technique coupled with automated imaging and analysis in both a smartphone-compatible device and a device with embedded imaging and processing. This is applied to measure the density distribution of cells with two demonstrated biomedical applications: sickle cell disease diagnosis and separating white and red blood cells.

Currently, many clinical diagnostic procedures are complex, costly, inefficient, and inaccessible to a large population in the world. The requirements for ...

Isokinetic Robotic Device to Improve Test-Retest and Inter-Rater Reliability for Stretch Reflex Measurements in Stroke Patients with Spasticity

Measuring spasticity is important in treatment planning and determining efficacy after treatment. However, the current tool used in clinical settings has been shown to be limited in inter-rater reliability. One factor in this poor inter-rater reliability is the variability of passive motion while measuring the angle of catch (AoC) measurements. Therefore, an isokinetic device has been proposed to standardize the manual joint motion; however, the benefits of isokinetic motion for AoC measurements has not been tested in a standardized manner. This protocol investigates whether isokinetic motion itself can improve inter-rater reliability for AoC measurements. For this purpose, a robotic isokinetic device was developed that is combined with surface electromyography (EMG). Two conditions, manual and isokinetic motions, are compared with the standardized method to measure the angle and subjective feeling of catch. It is shown that in 17 stroke patients with mild elbow flexor spasticity, isokinetic motion improved the intraclass correlation coefficient (ICC) for inter-rater reliability of AoC measurements to 0.890 [95% confidence interval (CI): 0.685&#8211;0.961] by the EMG criteria, and 0.931 (95% CI: 0.791&#8211;0.978) by the torque criteria, from 0.788 (95% CI: 0.493&#8211;0.920) by manual motion. In conclusion, isokinetic motion itself can improve inter-rater reliability of AoC measurements in stroke patients with mild spasticity. Given that this system may provide greater standardized angle measurements and catch of feeling, it may be a good option for the evaluation of spasticity in a clinical setting.

Using a robotic isokinetic device with electromyography (EMG) measurements, this protocol illustrates that isokinetic motion itself can improve inter-rater reliability for the angle of catch measurements in stroke patients with mild elbow flexor spasticity.

Measuring spasticity is important in treatment planning and determining efficacy after treatment. However, the current tool used in clinical settings has ...