1. Pulling Glass Micropipettes
Equipment: Micropipette Puller, Microforge.
Glass: Boroscillicate glass capillaries (~1.5 mm external diameter, ~1.4 mm internal diameter).
<ol>
	<li>Micropipettes are pulled using the same basic approach that is used to prepare glass microelectrodes for electrophysiology recordings. Briefly, a glass capillary is heated in the middle and when the glass starts to melt the two halves of the capillary are pulled apart generating two micropipettes. Multiple commercial pullers are available to perform this process ranging from relatively simple vertical pullers that use gravity to pull the two pipettes apart to highly-sophisticated horizontal pullers that offer multiple programmable options to vary the velocity and other parameters of the pull. Both types of pullers were used in our experiments.</li>
	<li>Requirements for the geometry of the pipette tip: The tips of the pipettes used in these experiments typically range between 2 to 6 &mu;m external diameter depending on the size of the cell. Another important parameter is the shape of the tip, which should approximate a cylindrical tube (see Figure 1). This can be achieved by optimizing the parameters of the pull and verifying the shape of the tip under the microscope until the desired shape is obtained. The optimal length of the pipette shank depends on the amount of expected deformation: if the deformations are small &lt;10 &mu;m, it is enough that the cylinder-like part of the pipette is also relative short (the same order of magnitude), for larger deformations adjust accordingly. In general, increasing the heat and/or increase in "pull" decreases the diameter of the tip. The increase in "pull" also generates a tip with a longer taper. In our experiments, using a Sutter P-97 horizontal pipette puller, the program was optimized to the following parameters: Heat of 473; Pull 22; Velocity 22, Time 200; Pressure 500. It is also possible to create cylindrical tips by pulling a very long shank and then breaking and polishing it. Detailed instructions of how to prepare different types of the pipettes are given in the Sutter manual.</li>
	<li>Microforge: It is also recommended to fire-polish the tip of the pipette to generate a smooth glass surface that makes a good seal to the plasma membrane. This is done by bringing the tip of the pipette to the vicinity of a heated glass ball for a very a fraction of a second using a microforge. Similar technique is routinely used for preparing microelectrodes for electrophysiological recordings.</li>
	<li>Filling the Micropipette: Micropipettes should be filled with a physiological saline solution, such as PBS or non-fluorescent growth media. Importantly the solution should be supplemented with 30% serum that will allow the cell membrane to move smoothly into the pipette. Two approaches can be used to get rid of air bubbles in the tip of the pipette: (i) a tip of the pipette can be immersed in the solution first to allow the liquid to fill the tip by the capillary forces followed by backfilling the pipette from the other end or (ii) the whole pipette can be filled from the back end by gently tapping on the shank of the pipette to remove the bubbles from the tip.</li>
	<li>Note: Pipettes have to be prepared on the day of the experiment.</li>
</ol>
2. Preparation of Cells
<ol>
	<li>Seeding the cells: Microaspiration is performed on single cells that are either maintained in suspension or are attached to the substrate. To aspirate cells in suspension, cells are lifted from their substrates and pipetted into a shallow longitudinal chamber that is mounted on the inverted microscope right before the experiment. To aspirate substrate-attached cells, cells are seeded on small cover-slips (~10 mm diameter) that can also be placed into the microaspiration chamber before the experiment. The rationale to use a shallow longitudinal chamber is to allow a micropipette to approach the cells at a very shallow angle, as close to horizontal as possible. This is done to allow the membrane pulled into the micropipette to be visualized on a single plane of focus (see Figure 2).</li>
	<li>Visualizing cell membrane: To observe the membrane projection into the pipette, cellular membranes are stained with a lipophilic fluorescent dye, such as diI using a standard staining protocol.
	<ol type="i">
		<li>Warm PBS solution.</li>
		<li>Dilute the stock diI to a working concentration (5 &mu;M) with the warmed PBS solution.</li>
		<li>Sonicate for 5 min to break dye aggregates.</li>
		<li>Spin down for 5 min and take supernatant.</li>
		<li>Wash cells in PBS 3 times, 5 min each.</li>
		<li>Incubate cells with the dye solution for 30 min in a 37 &deg;C incubator.</li>
		<li>Wash the cells with PBS 3 times for 5 min each.</li>
	</ol>
	</li>
	<li>Note: It is possible to substitute diI staining of the membrane with visualizing the sub-membrane cytoskeleton that is also being pulled together with the membrane into the pipette. It needs to be taken into the account, however, that perturbing the cytoskeleton may alter the biomechanical properties of the cell. Moreover, when performing microaspiration experiments with substrate attached cells, it is recommended to use 3D imaging to estimate the length of membrane projection into the pipette that is positioned at an angle to the focal plane of the cells.</li>
</ol>
3. Microaspiration and Image Acquisition
Equipment: Inverted Fluorescent Microscope, preferably with 3D deconvolution capabilities (Zeiss Axiovert 200M with computer controlled Z-axis movement of the objectives or an equivalent); videocamera connected to a computer (AxioCam MRm or an equivalent), Pressure Transducer (BioTek or an equivalent), Vibration-free station (TMD or an equivalent), Micromanipulator (Narishige, Sutter, Burleigh or equivalent; manipulators can be mechanical, hydraulic or piezoelectric). It is also important to emphasize that microaspiration can be performed using a microscope without 3D capabilities to estimate the stiffness of cells in suspension, such as red blood cells 1,2 or neutrophils 3, isolated organelles, such as nuclei 4 or artificial liposomes 5.
Image Acquisition software: Zeiss AxioVision or an equivalent.
<ol>
	<li>Mount the cells into a microaspiration chamber, as described above, on an inverted fluorescent microscope. Position the cells choosing a cell for an experiment and place it in the center of the visual field. It is important to perform these experiments in a vibration-free environment, particularly for the experiments with substrate-attached cells because minute vibrations that typically occur on benches and regular tables are likely to completely compromise the creation of the seal, break the tip of the micropipette or result in significant shifts in the position of the tip that will skew the analysis of the results.</li>
	<li>Place a micropipette filled with PBS/media w/ serum solution into a pipette holder connected to a power transducer by flexible tubing with the diameter adjusted to the connecter of the pipette holder for a tight fit. In the beginning of each experiment the pressure in the pipette is equilibrated to the atmospheric pressure. The pipette is mounted onto a micromanipulator that allows fine control of the pipette movements in a micron range. Position a pipette at a shallow angle to the bottom of the chamber and bring the tip of the pipette to the center of the visual field. The shank of the pipette, a cylindrical part of the pipette tip into which the membrane is aspirated, is aligned horizontally to the focal plane by (1) positioning the pipette at the shallowest angle possible (10-15&deg;) and (2) by flexing the shank of the pipette against the bottom of the chamber. Because the shank is very thin it is flexible enough to slide on the bottom of the chamber while approaching a cell, as shown schematically in Figure 1. Slowly bring down the micropipette to the side of a single cell using the course manipulator until near the plane of focus for the cell. Then, using a fine manipulator move the micropipette to the edge of the cell until the tip of the pipette gently touches the membrane. Take one image to observe the position of the pipette. Good seals are created when the whole tip of the pipette is in full contact with the cell surface and the contact is stable. There is no strong objective criterion, however, about how good the seal is except for the visual examination.</li>
	<li>Apply a step of negative pressure using the transducer and maintain it until membrane projection is stabilized. The amount of pressure needed to aspirate the membrane into the pipette varies depending on cell type and specific experimental conditions. In our experiments, initial deformation is typically observed when applying pressure in the range between -2 to -15mm Hg. When the pressure is applied, the membrane is gradually deformed into the pipette until it is stabilized at certain length, a process that typically takes 2-3 min. During this time, images of membrane deformation are acquired every 30 sec to track the progression of the membrane that is pulled into the pipette.</li>
	<li>Increase the pressure to the next level in 2-5 mm Hg steps and repeat the whole procedure until membrane projection detaches from the cell and moves into the pipette, at which point the experiment is stopped.</li>
</ol>
4. Analysis
To quantify the degree of membrane deformation, the aspirated length (L) is measured from the tip of the pipette to the vertex of the circumference of the membrane projection. It is important to note, however, that a larger pipette will apply more force on the cell membrane at the same level of pressure. To account for the variability between the diameters of the pipettes, therefore, the aspirated length is normalized for the pipette diameter (D) measured for each experiment.
The data can be further analyzed using a standard linear viscoelastic half-space model of the endothelial cell, as described in the earlier studies 6,7. Specifically, the elastic modulus of the cells was estimated using the equation:
<img alt="Equation 1" src="/files/ftp_upload/3886/3886eq1.jpg" />
where E is Young's modulus, a is the inner radius of the pipette, &Delta;p is the pressure difference, L is the corresponding aspirated length, and &phi;(&eta;) is a wall function calculated using the force model, as described by Theret et al 7. It is important to note that multiple models have been used to analyze the microaspiration data including a finite element model that assumes that a cell is a deformable sphere with isotropic and homogenous material properties and liquid drop models, which assume that cells form a spherical shape, can deform continuously, and recover upon release, as described in several excellent reviews: 8-10. Microaspiration can also be used to investigate other biomechanical parameters of cells and tissues, such as cellular viscoelastic properties, cortical tension and contribution of different structural elements to cell and tissue biomechanics (see the reviews listed above for more information).
5. Representative Results
In earlier studies, micropipette aspiration was performed either on liposomes 5 or on cells that were not attached to the substrate 2,11-13. In our studies, however, the cells are typically maintained attached to the substrate to avoid changes in the cytoskeletal structure that are likely to occur when cells detach 14-16. To validate the use of microaspiration technique for substrate-attached cells, we tested whether disruption of F-actin results in the decrease in cell stiffness of bovine aortic endothelial cells (BAECs), as estimated by this approach. Figure 3 shows that, as expected, this is indeed the case. Specifically, Figure 3A shows a typical series of fluorescent images of an endothelial membrane undergoing progressive deformation in response to negative pressure applied through a micropipette. As expected, the membrane is gradually aspirated into the pipette and the aspirated length increases as a function of applied pressure. The time-courses of the deformation show that disruption of F-actin significantly increases the aspirated lengths of the projections under all pressure conditions (Figure 3B) 14.
Using this approach, we discovered that cell stiffness increases when cellular membranes are depleted of cholesterol whereas cholesterol enrichment had no effect 14. Figure 4 shows a cholesterol-enriched cell, a control cell, and a cholesterol-depleted cell after reaching maximal aspiration lengths at -15 mm Hg (4A). The projections typically started to develop at -10mmHg and the time-courses of membrane deformation could be measured for the negative pressures of -10, -15 and -20 mm Hg (4B). Application of pressures above -25mmHg resulted in detachment of the aspirated projection forming a separate vesicle. The pressure level that resulted in membrane detachment was similar under different cholesterol conditions. This observation was highly unexpected because previous studies showed that in membrane lipid bilayers an increase in membrane cholesterol increases the stiffness of the membrane 5,17. Our further studies confirmed these observations using several independent approaches, including Atomic Force Microscopy 18,19 and Force Traction Microscopy 20.
<img alt="Figure 1" src="/files/ftp_upload/3886/3886fig1.jpg" /> 
Figure 1. Schematic side view of a recording pipette. The pipette is pulled to generate a cylindrical shank at the tip (side view). Micropipette parameters: D=2a=internal diameter and ED=2b=external diameter.
<img alt="Figure 2" src="/files/ftp_upload/3886/3886fig2.jpg" /> 
Figure 2. Micropipette approaching a substrate-attached cell. (A) Schematic side view; (B) Bright contrast image of a micropipette shank touching a typically shaped cell used in aspiration experiments; (C) Fluorescent image of the same cell labeled with DiIC18. The micropipette is still present but is invisible (From 14).
<img alt="Figure 3" src="/files/ftp_upload/3886/3886fig3.jpg" /> 
Figure 3. Validation of measuring cell stiffness in substrate-attached cells using microaspiration. A: Images of progressive membrane deformation of BAECs under control conditions and after exposure to latrunculin. The pipette is invisible on the images because it does not fluoresce. The cells were exposed to 2 &mu;M latrunculin A for 10 min, which dramatically reduced the amount of F-actin, as measured by rhodamine-phalloidin fluorescence (not shown) but had no significant effect on the cell shape. In a latrunculin-treated cell, there is a thinning of the membrane in the middle of the aspirated projection but the projection is still attached to the cells. B. Effect of latrunculin A on the time courses of membrane deformation where L is aspirated length of the membrane projection and D is the diameter of the pipette for control cells (n=14) and cells exposed to 2 &mu;M latrunculin A for 10 min (n=5). The cells were aspirated with -10 mm Hg (diamonds), -15 mm Hg (squares) and -20 mm Hg (triangles). (From 14.)
<img alt="Figure 4" src="/files/ftp_upload/3886/3886fig4.jpg" /> 
Figure 4. Effect of cellular cholesterol levels on membrane deformation of BAECs. A. Typical images of membrane deformation of cholesterol-enriched, cholesterol-depleted and control cells (control cells were exposed to M&beta;CD:M&beta;CD-cholesterol mixture at 1:1 ratio that had no effect on the level of free cholesterol in the cells (see inset). The images shown depict the maximal deformation at -15 mm Hg. The arrow indicates the position of the aspirated projection. The bar is 30 &mu;m. B. Average time-courses of aspirated lengths for the three experimental cell populations. C. Maximal aspirated lengths plotted as a function of the applied pressure. The maximal normalized length in depleted cells was significantly lower than that of control cells for pressures -15 mm Hg and -20 mm Hg (P &lt; 0.05). (From 14.)

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mapping+the+cytoskeletal+prestress.">Mapping the cytoskeletal prestress.</a> American Journal of Physiology - Cell Physiology. 298, C1245-C1252 (2010).</li><li>Munevar, S., Wang, Y., Dembo, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Traction+force+microscopy+of+migrating+normal+and+H-ras+transformed+3T3+fibroblasts.">Traction force microscopy of migrating normal and H-ras transformed 3T3 fibroblasts.</a> Biophys. J. 80, 1744-1757 (2001).</li><li>Wirtz, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Particle-tracking+microrheology+of+living+cells:+principles+and+applications.">Particle-tracking microrheology of living cells: principles and applications.</a> Annu. Rev. Biophys. 38, 301-326 (2009).</li><li>Shin, D., Athanasiou, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytoindentation+for+obtaining+cell+biomechanical+properties.">Cytoindentation for obtaining cell biomechanical properties.</a> J. Orthop. Res. 17, 880-890 (1999).</li><li>Ou-Yang, H. D., Wei, M. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Complex+fluids:+probing+mechanical+properties+of+biological+systems+with+optical+tweezers.">Complex fluids: probing mechanical properties of biological systems with optical tweezers.</a> Annu. Rev. Phys. Chem. 61, 421-440 (2010).</li><li>Hosu, B. G., Sun, M., Marga, F., Grandbois, M., Forgacs, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Eukaryotic+membrane+tethers+revisited+using+magnetic+tweezers.">Eukaryotic membrane tethers revisited using magnetic tweezers.</a> Phys. Biol. 4, 67-78 (2007).</li></ol>

No conflicts of interest declared.

Microaspiration provides a simple and highly reproducible method to estimate cell stiffness/deformability by applying negative pressure to a cell membrane and measuring membrane deformability in response to well-defined pressure. It was first developed by Mitchison and Swann (1954) to characterize the elastic properties of sea-urchin eggs to provide insights into the mechanisms of cell division 21 and then to look at the mechanical properties in red blood cells 1. This method has been used in multip...

<table border="1">
 <tbody>
 <tr>
 <td>Name of reagent</td>
 <td>Company</td>
 <td>Catalog/Model Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Sutter pipette puller</td>
 <td>Sutter Instruments</td>
 <td>P-97</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Microforge</td>
 <td>Narishige</td>
 <td>MF-830</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Inverted Fluorescent Microscope</td>
 <td>Zeiss</td>
 <td>Axiovert 200M</td>
 <td>The microscope should be preferably equipped with 3D/deconvolution capabilities.</td>
 </tr>
 <tr>
 <td>Videocamera</td>
 <td>Zeiss</td>
 <td>AxioCam MRm</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Image Acquisition sotware</td>
 <td>Zeiss</td>
 <td>AxioVision</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Pneumatic Pressure Transducer</td>
 <td>BioTek</td>
 <td>DPM-1B</td>
 <td>DPM1B Pneumatic Transducer Tester can now be found by FLUKE.</td>
 </tr>
 <tr>
 <td>Pipette glass</td>
 <td>Richland</td>
 <td>Customized glass</td>
 <td>Pipettes were customized with a 1.2 inner diameter and 1.6 outer diameter.</td>
 </tr>
 <tr>
 <td>DiI Dye</td>
 <td>Invitrogen</td>
 <td>D282</td>
 <td>Dissolves well in DMSO</td>
 </tr>
 </tbody>
</table>

micropipette aspiration of substrate-attached cells to estimate cell stiffness

Growing number of studies show that biomechanical properties of individual cells play major roles in multiple cellular functions, including cell proliferation, differentiation, migration and cell-cell interactions. The two key parameters of cellular biomechanics are cellular deformability or stiffness and the ability of the cells to contract and generate force. Here we describe a quick and simple method to estimate cell stiffness by measuring the degree of membrane deformation in response to negative pressure applied by a glass micropipette to the cell surface, a technique that is called Micropipette Aspiration or Microaspiration.
Microaspiration is performed by pulling a glass capillary to create a micropipette with a very small tip (2-50 &mu;m diameter depending on the size of a cell or a tissue sample), which is then connected to a pneumatic pressure transducer and brought to a close vicinity of a cell under a microscope. When the tip of the pipette touches a cell, a step of negative pressure is applied to the pipette by the pneumatic pressure transducer generating well-defined pressure on the cell membrane. In response to pressure, the membrane is aspirated into the pipette and progressive membrane deformation or "membrane projection" into the pipette is measured as a function of time. The basic principle of this experimental approach is that the degree of membrane deformation in response to a defined mechanical force is a function of membrane stiffness. The stiffer the membrane is, the slower the rate of membrane deformation and the shorter the steady-state aspiration length.The technique can be performed on isolated cells, both in suspension and substrate-attached, large organelles, and liposomes.
Analysis is performed by comparing maximal membrane deformations achieved under a given pressure for different cell populations or experimental conditions. A "stiffness coefficient" is estimated by plotting the aspirated length of membrane deformation as a function of the applied pressure. Furthermore, the data can be further analyzed to estimate the Young's modulus of the cells (E), the most common parameter to characterize stiffness of materials. It is important to note that plasma membranes of eukaryotic cells can be viewed as a bi-component system where membrane lipid bilayer is underlied by the sub-membrane cytoskeleton and that it is the cytoskeleton that constitutes the mechanical scaffold of the membrane and dominates the deformability of the cellular envelope. This approach, therefore, allows probing the biomechanical properties of the sub-membrane cytoskeleton.

Watch this Scientific Journal Video about Micropipette Aspiration of Substrate-attached Cells to Estimate Cell Stiffness at JoVE.com

Micropipette Aspiration of Substrate-attached Cells to Estimate Cell Stiffness

Here we describe a quick and simple method to measure cell stiffness. The general principle of this approach is to measure membrane deformation in response to well-defined negative pressure applied through a micropipette to the cell surface. This method provides a powerful tool to study biomechanical properties of substrate-attached cells.

Growing number of studies show that biomechanical properties of individual cells play major roles in multiple cellular functions, including cell ...

micropipette-aspiration-substrate-attached-cells-to-estimate-cell

Research

JoVE Journal

Bioengineering

19.7K Views.  University of Illinois. Here we describe a quick and simple method to measure cell stiffness. The general principle of this approach is to measure membrane deformation in response to well-defined negative pressure applied through a micropipette to the cell surface. This method provides a powerful tool to study biomechanical properties of substrate-attached cells.

Video: Micropipette Aspiration of Substrate-attached Cells to Estimate Cell Stiffness

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

Alginate Microcapsule as a 3D Platform for Propagation and Differentiation of Human Embryonic Stem Cells (hESC) to Different Lineages

Human embryonic stem cells (hESC) are emerging as an attractive alternative source for cell replacement therapy since they can be expanded in culture indefinitely and differentiated to any cell types in the body. Various types of biomaterials have also been used in stem cell cultures to provide a microenvironment mimicking the stem cell niche1-3. The latter is important for promoting cell-to-cell interaction, cell proliferation, and differentiation into specific lineages as well as tissue organization by providing a three-dimensional (3D) environment4 such as encapsulation. The principle of cell encapsulation involves entrapment of living cells within the confines of semi-permeable membranes in 3D cultures2. These membranes allow for the exchange of nutrients, oxygen and stimuli across the membranes, whereas antibodies and immune cells from the host that are larger than the capsule pore size are excluded5. Here, we present an approach to culture and differentiate hESC DA neurons in a 3D microenvironment using alginate microcapsules. We have modified the culture conditions2 to enhance the viability of encapsulated hESC. We have previously shown that the addition of p160-Rho-associated coiled-coil kinase (ROCK) inhibitor, Y-27632 and human fetal fibroblast-conditioned serum replacement medium (hFF-CM) to the 3D platform significantly enhanced the viability of encapsulated hESC in which the cells expressed definitive endoderm marker genes1. We have now used this 3D platform for the propagation of hESC and efficient differentiation to DA neurons. Protein and gene expression analyses after the final stage of DA neuronal differentiation showed an increased expression of tyrosine hydroxylase (TH), a marker for DA neurons, &gt;100 folds after 2 weeks. We hypothesized that our 3D platform using alginate microcapsules may be useful to study the proliferation and directed differentiation of hESC to various lineages. This 3D system also allows the separation of feeder cells from hESC during the process of differentiation and also has potential for immune-isolation during transplantation in the future.

Alginate Microcapsule as a 3D Platform for Propagation and Differentiation of Human Embryonic Stem Cells hESC to Different Lineages

We have optimized a microencapsulation technique as an effective 3D platform for propagation and differentiation of embryonic stem cells to endoderm and dopaminergic (DA) neurons. It also provides an opportunity for immune-isolation of cells from the host during transplantation. This platform can be adapted for other cell types.

Human embryonic stem cells (hESC) are emerging as an attractive alternative source for cell replacement therapy since they can be expanded in culture ...

Spatio-Temporal Manipulation of Small GTPase Activity at Subcellular Level and on Timescale of Seconds in Living Cells

Dynamic regulation of the Rho family of small guanosine triphosphatases (GTPases) with great spatiotemporal precision is essential for various cellular functions and events1, 2. Their spatiotemporally dynamic nature has been revealed by visualization of their activity and localization in real time3. In order to gain deeper understanding of their roles in diverse cellular functions at the molecular level, the next step should be perturbation of protein activities at a precise subcellular location and timing. 
To achieve this goal, we have developed a method for light-induced, spatio-temporally controlled activation of small GTPases by combining two techniques: (1) rapamycin-induced FKBP-FRB heterodimerization and (2) a photo-caging method of rapamycin. With the use of rapamycin-mediated FKBP-FRB heterodimerization, we have developed a method for rapidly inducible activation or inactivation of small GTPases including Rac4, Cdc424, RhoA4 and Ras5, in which rapamycin induces translocation of FKBP-fused GTPases, or their activators, to the plasma membrane where FRB is anchored. For coupling with this heterodimerization system, we have also developed a photo-caging system of rapamycin analogs. A photo-caged compound is a small molecule whose activity is suppressed with a photocleavable protecting group known as a caging group. To suppress heterodimerization activity completely, we designed a caged rapamycin that is tethered to a macromolecule such that the resulting large complex cannot cross the plasma membrane, leading to virtually no background activity as a chemical dimerizer inside cells6. Figure 1 illustrates a scheme of our system. With the combination of these two systems, we locally recruited a Rac activator to the plasma membrane on a timescale of seconds and achieved light-induced Rac activation at the subcellular level6.

A method for spatio-temporal control of small GTPase activity by light is described. This method is based on rapamycin-induced FKBP-FRB heterodimerization and photo-caging systems. Optimization of light-irradiation enables the spatio-temporally controlled activation of small GTPases at the subcellular level.

Dynamic regulation of the Rho family of small guanosine triphosphatases (GTPases) with great spatiotemporal precision is essential for various cellular ...

Development, Expansion, and In vivo Monitoring of Human NK Cells from Human Embryonic Stem Cells (hESCs) and Induced Pluripotent Stem Cells (iPSCs)

We present a method for deriving natural killer (NK) cells from undifferentiated hESCs and iPSCs using a feeder-free approach. This method gives rise to high levels of NK cells after 4 weeks culture and can undergo further 2-log expansion with artificial antigen presenting cells. hESC- and iPSC-derived NK cells developed in this system have a mature phenotype and function. The production of large numbers of genetically modifiable NK cells is applicable for both basic mechanistic as well as anti-tumor studies. Expression of firefly luciferase in hESC-derived NK cells allows a non-invasive approach to follow NK cell engraftment, distribution, and function. We also describe a dual-imaging scheme that allows separate monitoring of two different cell populations to more distinctly characterize their interactions in vivo. This method of derivation, expansion, and dual in vivo imaging provides a reliable approach for producing NK cells and their evaluation which is necessary to improve current NK cell adoptive therapies.

Development, Expansion, and In vivo Monitoring of Human NK Cells from Human Embryonic Stem Cells hESCs and Induced Pluripotent Stem Cells iPSCs

This protocol describes the development, expansion, and in vivo imaging of NK cells derived from hESCs and iPSCs.

We present a method for deriving natural killer (NK) cells from undifferentiated hESCs and iPSCs using a feeder-free approach. This method gives rise to ...

Transplantation of Induced Pluripotent Stem Cell-derived Mesoangioblast-like Myogenic Progenitors in Mouse Models of Muscle Regeneration

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells similar to pericyte-derived mesoangioblasts (iPSC-derived mesoangioblast-like stem/progenitor cells: IDEMs) can be established from iPSCs generated from patients affected by different forms of muscular dystrophy. Patient-specific IDEMs can be genetically corrected with different strategies (e.g. lentiviral vectors, human artificial chromosomes) and enhanced in their myogenic differentiation potential upon overexpression of the myogenesis regulator MyoD. This myogenic potential is then assessed in vitro with specific differentiation assays and analyzed by immunofluorescence. The regenerative potential of IDEMs is further evaluated in vivo, upon intramuscular and intra-arterial transplantation in two representative mouse models displaying acute and chronic muscle regeneration. The contribution of IDEMs to the host skeletal muscle is then confirmed by different functional tests in transplanted mice. In particular, the amelioration of the motor capacity of the animals is studied with treadmill tests. Cell engraftment and differentiation are then assessed by a number of histological and immunofluorescence assays on transplanted muscles. Overall, this paper describes the assays and tools currently utilized to evaluate the differentiation capacity of IDEMs, focusing on the transplantation methods and subsequent outcome measures to analyze the efficacy of cell transplantation.

Induced pluripotent stem cell (iPSC)-derived myogenic progenitors are promising candidates for cell therapy strategies to treat muscular dystrophies. This protocol describes transplantation and functional measurements required to evaluate the engraftment and differentiation of iPSC-derived mesoangioblasts (a type of muscle progenitors) in mouse models of acute and chronic muscle regeneration.

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells ...

Ordering Single Cells and Single Embryos in 3D Confinement: A New Device for High Content Screening

Biological cells are usually observed on flat (2D) surfaces. This condition is not physiological, and phenotypes and shapes are highly variable. Screening based on cells in such environments have therefore serious limitations: cell organelles show extreme phenotypes, cell morphologies and sizes are heterogeneous and/or specific cell organelles cannot be properly visualized. In addition, cells in vivo are located in a 3D environment; in this situation, cells show different phenotypes mainly because of their interaction with the surrounding extracellular matrix of the tissue. In order to standardize and generate order of single cells in a physiologically-relevant 3D environment for cell-based assays, we report here the microfabrication and applications of a device for in vitro 3D cell culture. This device consists of a 2D array of microcavities (typically 105 cavities/cm2), each filled with single cells or embryos. Cell position, shape, polarity and internal cell organization become then normalized showing a 3D architecture. We used replica molding to pattern an array of microcavities, &#8216;eggcups&#8217;, onto a thin polydimethylsiloxane (PDMS) layer adhered on a coverslip. Cavities were covered with fibronectin to facilitate adhesion. Cells were inserted by centrifugation. Filling percentage was optimized for each system allowing up to 80%. Cells and embryos viability was confirmed. We applied this methodology for the visualization of cellular organelles, such as nucleus and Golgi apparatus, and to study active processes, such as the closure of the cytokinetic ring during cell mitosis. This device allowed the identification of new features, such as periodic accumulations and inhomogeneities of myosin and actin during the cytokinetic ring closure and compacted phenotypes for Golgi and nucleus alignment. We characterized the method for mammalian cells, fission yeast, budding yeast, C. elegans with specific adaptation in each case. Finally, the characteristics of this device make it particularly interesting for drug screening assays and personalized medicine.

We report a device and a new method to study cells and embryos. Single cells are precisely ordered in microcavity arrays. Their 3D confinement is a step towards 3D environments encountered in physiological conditions and allows organelle orientation. By controlling cell shape, this setup minimizes variability reported in standard assays.

Biological cells are usually observed on flat (2D) surfaces. This condition is not physiological, and phenotypes and shapes are highly variable. Screening ...

Simple Polyacrylamide-based Multiwell Stiffness Assay for the Study of Stiffness-dependent Cell Responses

Currently, most of the in vitro cell research is performed on rigid tissue culture polystyrene (~1 GPa), while most cells in the body are attached to a matrix that is elastic and much softer (0.1 &#8211; 100 kPa). Since such stiffness mismatch greatly affects cell responses, there is a strong interest in developing hydrogel materials that span a wide range of stiffness to serve as cell substrates. Polyacrylamide gels, which are inexpensive and cover the stiffness range of all soft tissues in the body, are the hydrogel of choice for many research groups. However, polyacrylamide gel preparation is lengthy, tedious, and only suitable for small batches. Here, we describe an assay which by utilizing a permanent flexible plastic film as a structural support for the gels, enables the preparation of polyacrylamide gels in a multiwell plate format. The technique is faster, more efficient, and less costly than current methods and permits the preparation of gels of custom sizes not otherwise available. As it doesn&#8217;t require any specialized equipment, the method could be easily adopted by any research laboratory and would be particularly useful in research focused on understanding stiffness-dependent cell responses.

Here, a method that enables quick, efficient, and inexpensive preparation of polyacrylamide gels in a multiwell plate format is described. The method does not require any specialized equipment and could be easily adopted by any research laboratory. It would be particularly useful in research focused on understanding stiffness-dependent cell responses.

Currently, most of the in vitro cell research is performed on rigid tissue culture polystyrene (~1 GPa), while most cells in the body are attached to a ...

A Method to Estimate Cadaveric Femur Cortical Strains During Fracture Testing Using Digital Image Correlation

This protocol describes the method using digital image correlation to estimate cortical strain from high speed video images of the cadaveric femoral surface obtained from mechanical testing. This optical method requires a texture of many contrasting fiduciary marks on a solid white background for accurate tracking of surface deformation as loading is applied to the specimen. Immediately prior to testing, the surface of interest in the camera view is painted with a water-based white primer and allowed to dry for several minutes. Then, a black paint is speckled carefully over the white background with special consideration for the even size and shape of the droplets. Illumination is carefully designed and set such that there is optimal contrast of these marks while minimizing reflections through the use of filters. Images were obtained through high speed video capture at up to 12,000 frames/s. The key images prior to and including the fracture event are extracted and deformations are estimated between successive frames in carefully sized interrogation windows over a specified region of interest. These deformations are then used to compute surface strain temporally during the fracture test. The strain data is very useful for identifying fracture initiation within the femur, and for eventual validation of proximal femur fracture strength models derived from Quantitative Computed Tomography-based Finite Element Analysis (QCT/FEA).

In this protocol, the femur surface strains are estimated during fracture testing using the digital image correlation technique. The novelty of the method involves application of a high-contrast stochastic speckle pattern on the femur surface, carefully specified illumination, high speed video capture, and digital image correlation analysis for strain calculations.

This protocol describes the method using digital image correlation to estimate cortical strain from high speed video images of the cadaveric femoral ...

Human Pluripotent Stem Cell Culture on Polyvinyl Alcohol-Co-Itaconic Acid Hydrogels with Varying Stiffness Under Xeno-Free Conditions

The effect of physical cues, such as the stiffness of biomaterials on the proliferation and differentiation of stem cells, has been investigated by several researchers. However, most of these investigators have used polyacrylamide hydrogels for stem cell culture in their studies. Therefore, their results are controversial because those results might originate from the specific characteristics of the polyacrylamide and not from the physical cue (stiffness) of the biomaterials. Here, we describe a protocol for preparing hydrogels, which are not based on polyacrylamide, where various stem, cells including human embryonic stem (ES) cells and human induced pluripotent stem (iPS) cells, can be cultured. Hydrogels with varying stiffness were prepared from bioinert polyvinyl alcohol-co-itaconic acid (P-IA), with stiffness controlled by crosslinking degree by changing crosslinking time. The P-IA hydrogels grafted with and without oligopeptides derived from extracellular matrix were investigated as a future platform for stem cell culture and differentiation. The culture and passage of amniotic fluid stem cells, adipose-derived stem cells, human ES cells, and human iPS cells is described in detail here. The oligopeptide P-IA hydrogels showed superior performances, which were induced by their stiffness properties. This protocol reports the synthesis of the biomaterial, their surface manipulation, along with controlling the stiffness properties and finally, their impact on stem cell fate using xeno-free culture conditions. Based on recent studies, such modified substrates can act as future platforms to support and direct the fate of various stem cells line to different linkages; and further, regenerate and restore the functions of the lost organ or tissue.

A protocol is presented to prepare polyvinyl alcohol-co-itaconic acid hydrogels with varying stiffness, which were grafted with and without oligopeptides, to investigate the effect of the stiffness of biomaterials on the differentiation and proliferation of stem cells. The stiffness of the hydrogels was controlled by the crosslinking time.

The effect of physical cues, such as the stiffness of biomaterials on the proliferation and differentiation of stem cells, has been investigated by ...

Stiffness Measurement of Soft Silicone Substrates for Mechanobiology Studies Using a Widefield Fluorescence Microscope

Soft tissues in the human body typically have stiffness in the kilopascal (kPa) range. Accordingly, silicone and hydrogel flexible substrates have been proven to be useful substrates for culturing cells in a physical microenvironment that partially mimics in vivo conditions. Here, we present a simple protocol for characterizing the Young's moduli of isotropic linear elastic substrates typically used for mechanobiology studies. The protocol consists of preparing a soft silicone substrate on a Petri dish or stiff silicone, coating the top surface of the silicone substrate with fluorescent beads, using a millimeter-scale sphere to indent the top surface (by gravity), imaging the fluorescent beads on the indented silicone surface using a fluorescence microscope, and analyzing the resultant images to calculate the Young's modulus of the silicone substrate. Coupling the substrate's top surface with a moduli extracellular matrix protein (in addition to the fluorescent beads) allows the silicone substrate to be readily used for cell plating and subsequent studies using traction force microscopy experiments. The use of stiff silicone, instead of a Petri dish, as the base of the soft silicone, enables the use of mechanobiology studies involving external stretch. A specific advantage of this protocol is that a widefield fluorescence microscope, which is commonly available in many labs, is the major equipment necessary for this procedure. We demonstrate this protocol by measuring the Young's modulus of soft silicone substrates of different elastic moduli.

Substrates with stiffness in the kilopascal-range are useful to study the response of cells to physiologically relevant micro-environment stiffness. Using just a widefield fluorescence microscope, the Young's modulus of soft silicone gels can be determined using an indentation with a suitable sphere.

Soft tissues in the human body typically have stiffness in the kilopascal (kPa) range. Accordingly, silicone and hydrogel flexible substrates have been ...