Silicon Master Mold Fabrication Lithography Materials:
<ul>
<li>75 mm Silicon wafers</li>
<li>SU-8 2 Negative Photoresist (Microchem, Boston, MA)</li>
<li>N2</li>
</ul>
Method:
<ol>
<li>Dehydrate Si wafer at 120 &deg;C for 2 hours.</li>
<li>UV ozone treat surface for 7 minutes to remove organics.</li>
<li>N2 blow surface to remove particulates.</li>
<li>Apply SU-8 2 to cover 70% of surface (~10 ml for 75 mm wafer).</li>
<li>Spin at 2000 rpm for 20 seconds with 300 rpm/s acceleration.</li>
<li>Place the wafer on a hotplate and ramp from room temperature to 95 &deg;C.</li>
<li>Softbake the photoresist for 15 minutes at 95 &deg;C.</li>
<li>Flood expose wafer for 20 - 25 mJ/cm2 dose at 365 nm (I-line) to form the bottom layer.</li>
<li>Apply SU-8 2 for second photoresist layer.</li>
<li>Spin at 550 rpm for 20 seconds with 300 rpm/s acceleration for an approximately 11 &mu;m thickness film.</li>
<li>Place the wafer on a hotplate and ramp from room temperature to 65 &deg;C.</li>
<li>Softbake the wafer at 65 &deg;C for 5 minutes.</li>
<li>Ramp the hotplate from 65 &deg;C to 95 &deg;C.</li>
<li>Softbake the wafer for 55 minutes at 95 &deg;C.</li>
<li>Cool wafer to room temperature for 20 minutes.</li>
<li>Align mPAD mask on wafer and expose for 95 - 100 mJ/cm2 dose at 365 nm (I-line).</li>
<li>Place the wafer on a hotplate and ramp from room temperature to 95 &deg;C.</li>
<li>Post-exposure bake the photoresist for 25 minutes at 95 &deg;C.</li>
</ol>
Develop Materials:
<ul>
<li>100mm glass dishes (5)</li>
<li>Propylene glycol methyl ether acetate (PGMEA)</li>
<li>Isopropanol (IPA)</li>
<li>N2</li>
</ul>
Method:
<ol>
<li>In a chemical hood, fill five dishes with PGMEA (2), IPA, and Hexane (2).</li>
<li>Hold wafer in 1st PGMEA for 10 seconds</li>
<li>Agitate wafer for 50 seconds</li>
<li>Transfer wafer to 2nd PGMEA with meniscus of PGMEA.</li>
<li>Agitate for 5 seconds.</li>
<li>Transfer wafer to IPA with meniscus of PGMEA.</li>
<li>Agitate for 20 seconds.</li>
<li>Transfer wafer to 1st IPA.</li>
<li>Agitate for 5 seconds.</li>
<li>Transfer wafer to 2nd IPA.</li>
<li>Agitate for 5 seconds.</li>
<li>Quickly remove wafer from 2nd IPA with meniscus of IPA and dry with N2</li>
<li>Inspect pattern.</li>
<li>Hardbake wafer at 120 &deg;C for 14-20 hours to cross-link SU-8.</li>
<li>Dice the wafer with a diamond cutter</li>
<li>Mount the wafer on a glass slide using epoxy</li>
<li>Fluorosilanize the wafer (see instructions in replication molding)</li>
</ol>
Replication molding of mPADS Negative Mold Materials:
<ul>
<li>Disposable aluminum weighing boat</li>
<li>Polydimethylsiloxane (PDMS) base &amp; curing agent (Dow, Midland, MI)</li>
<li>Razorblade</li>
<li>Silane: (tridecafluoro-1,1,2,2-tetrahyrooctyl)-1-trichlorosilane</li>
<li>Glass slide</li>
<li>Glass Pasteur Pipette</li>
</ul>
Method:
<ol>
<li>Mix PDMS 10:1 by weight and degas in under vacuum for 1 hour to remove bubbles.</li>
<li> Place silicon master mold in aluminum boat.</li>
<li>Pour 50 &ndash; 60 g of PDMS over master mold in aluminum boat.</li>
<li>Degas under vacuum for 30 minutes to remove bubbles.</li>
<li>Blow off surface-trapped air bubbles in PDMS with quick blast of N2.</li>
<li>Place boat in convection oven at 110 &deg;C for 15 minutes until PDMS is firm.</li>
<li>Cut away aluminum boat.</li>
<li> With razorblade, trim away PDMS underneath silicon master.</li>
<li>Peel PDMS away from silicon master slowly and with constant motion in one direction to avoid destroying the master.</li>
<li> Inspect silicon master for damage.</li>
<li>Cut negative mold into 4 pieces.</li>
<li>Repeat steps 1-10 for a large batch of negative molds.</li>
<li>Oxygen plasma treat negative mold for 1.5 minutes to activate surface, or</li>
<li>Ozone treat negative mold for 10 minutes to activate the surface.</li>
<li>Place negative molds in dessicator with mPAD features exposed.</li>
<li>Spread 250-500 &mu;L of silane on glass slide in center of dessicator</li>
<li>Place glass pasture pipette with 150-200 &mu;L of silane in the tip inside dessicator.</li>
<li>Vacuum chamber to evaporate silane and coat negative molds for &gt;14 hours.</li>
</ol>
mPADS Materials:
<ul>
<li>Polydimethylsiloxane (PDMS) base &amp; curing agent (Dow, Midland, MI)</li>
<li>Wide-mouth disposable transfer pipettes</li>
<li>2&rdquo; x 3&rdquo; Glass slides</li>
<li>Glass Scribe</li>
<li>N2</li>
</ul>
Method:
<ol>
<li>Mix PDMS 10:1 by weight and degas in under vacuum for 1 hour to remove bubbles.</li>
<li>lace silanized negative molds into &lsquo;cross&rsquo;</li>
<li>Pipette PDMS into negative molds so that the features are fully covered to a 1mm thickness.</li>
<li>Wait 30 minutes for any air bubbles in the PDMS to rise to the surface.</li>
<li>In 30&rsquo; downtime, dust 22x22mm (No. 2) glass coverslips with a stream of N2.</li>
<li>Oxygen plasma treat glass slides halves for 1.5 minutes to clean and activate glass</li>
<li>Blow off surface-trapped air bubbles in PDMS with quick blast of N2.</li>
<li>Gently lay edge of glass coverslip on PDMS with plasma cleaned side facing the PDMS. Slowly lower the glass slide so that the PDMS fully contacts the entire glass surface and avoid air bubbles getting trapped underneath the glass slide.</li>
<li>Place assembled molds in convection oven at 110 &deg;C for 20 hours to fully cure.</li>
<li>Remove molds from oven and let them cool to RT.</li>
<li>Hold glass slide with one hand on top and slowly peel negative away the negative mold from the bottom to release the mPAD. </li>
<li>Inspect the mPAD under a microscope for collapsed posts.</li>
<li>Re-silanize mold via plasma and silane every four castings. The negative mold lifetime for good replication is approximately 6-20 casts.</li>
</ol>
Seeding Cells onto mPADS Stamping Materials:
<ul>
<li>PDMS stamps for uCP (see Tien, J &amp; Chen, C. S., in Methods of Tissue Engineering, 2001, pp. 113-120.)</li>
<li>Ethanol (100% and 70%)</li>
<li>Sterilized Deionized Water</li>
<li>50 ug/ml Human Fibronectin in sterile deionized water (BD Biosciences)</li>
<li>5 ug/ml DiI Solution filtered with 0.2 &mu;m membrane (D-3886, Molecular Probes, OR)</li>
<li>2% Pluronics F127 (BASF, Mount Olive, NJ)</li>
<li>1X Phosphate Buffer Solution.</li>
<li>100 mm x 25 mm petri dish (4) </li>
<li>150 mm x 25 mm petri dish (2)</li>
<li>Aluminum foil</li>
<li>N2</li>
<li>Tweezers</li>
<li>Sterile Hood</li>
</ul>
Method:
<ol>
<li>Trim excess PDMS from mPADs </li>
<li>If the PDMS stamp has been used previously, sonicate stamps in 100% ethanol for 5 minutes to scrub away contaminating proteins.</li>
<li>In sterile hood, hold stamps on their sides with tweezers and dry with N2 after dipping stamps in fresh 100% ethanol and sterile water</li>
<li>Place in 150 mm petri dish with working side up.</li>
<li>Aliquot drops of fibronectin solution on each stamp. Start with the corners on each of the stamps, then the edges, and lastly the interior regions. Initially the PDMS surface of the stamp is hydrophobic, but becomes hydrophilic as the protein is adsorbed. Working &ldquo;outside to inside&rdquo; changes the contact angle of the PDMS so that a lower volume of fibronectin solution is needed to fully coat the stamp. </li>
<li>Let sit for 1 hour in the hood for protein adsorption.</li>
<li>Wash stamps with DI water by pouring in dish and letting the water level rise over the stamps.</li>
<li>Wash stamps in 100 mm petri dish filled with DI water.</li>
<li>Remove and dry with N2.</li>
<li>Ozone treat mPADS for 7 minutes to activate the surface for stamping.</li>
<li>Under hood, stamp mPADS by placing stamp edge on mPAD and gently lowering to make full contact. Lightly tap the top of the stamp with tweezers to ensure all areas have good contact but be careful not collapse the posts.</li>
<li>Carefully remove stamps with tweezers. </li>
<li>Submerge mPADS in 100 mm petri dish with 100% Ethanol for ~15 seconds. Keep microneedles from dewetting during transfer by transferring quickly between dishes. </li>
<li>Submerge mPADS in 100 mm petri dish with 70% Ethanol for ~15 seconds.</li>
<li>Submerge mPADS in 100 mm petri dish with DI water for ~15 seconds.</li>
<li>Submerge mPADS in second 100 mm petri dish with DI water for ~15 seconds.</li>
<li> Submerge mPADS in 150 mm petri dish with DI water.</li>
<li>Submerge mPADS in 150 mm petri dish with 100 mL of DiI solution.</li>
<li>Cover with aluminum foil and soak for 1 hour.</li>
<li>Submerge mPADS in 100 mm petri dish with DI water for ~15 seconds.</li>
<li>Submerge mPADS in 150 mm petri dish with 10 mL of 2% Pluronics and 90 mL of PBS for total concentration of 0.2% Pluronics.</li>
<li>Cover and soak for 1 hour.</li>
<li>Submerge mPADS in 100 mm petri dish with DI water for ~15 seconds.</li>
<li>Submerge mPADS in second 100 mm petri dish with DI water for ~15 seconds.</li>
<li>Submerge mPADS in third 100 mm petri dish with PBS for ~15 seconds.</li>
</ol>
Seeding Materials:
<ul>
<li>Chosen cell line</li>
<li>Trypsin-EDTA (0.25% Trypsin with EDTA&bull;4Na, Gibco Cat. No. 25200-056)</li>
<li>Growth Medium</li>
<li>1X Phosphate Buffer Solution.</li>
</ul>
Method:
<ol>
<li> In sterile hood, place mPADS in 100 mm petri dish and fill with 17 mL medium. </li>
<li>Place petri dish in incubator to warm to 37&deg;C </li>
<li>Suspend cells via trypsin</li>
<li>Aspirate repeatedly to break up cell clusters. </li>
<li>Pipet cell suspension into the petri dish with the mPADS. Add cells such they will remain single cells (without cell-cell contacts) at time of analysis. The exact amount of cells to add is cell-type specific and depends on cell spread area and growth rate.</li>
<li>Gently disperse cells in petri dish with slow pippetting.</li>
<li>Place in incubator for 2 hours to allow cells to settle and adhere onto microneedles. </li>
<li>Inspect mPADS to see if cells have spread. The cells will look flattened and span six or more posts.</li>
<li>If cells are spread, transfer mPADS to new 100 mm petri dish with 21 mL of fresh medium. </li>
<li>The mPADS are ready for your experiment.</li>
</ol>
Staining and Mounting mPADS Fixing Materials:
<ul>
<li>Deionized water</li>
<li>Paraformeldahyde (16%) vial</li>
<li>10X Phosphate Buffer Solution.</li>
<li>1X Phosphate Buffer Solution.</li>
</ul>
Method:
<ol>
<li> In a 50mL centrifuge tube, add 30 mL of DI, 10 mL of paraformeldahyde (entire vial), and 4.5 mL of 10X PBS for the fixing solution. </li>
<li>Add ~25 mL of fixing solution to a 100 mm petri dish.</li>
<li>Gently submerge mPAD with cells into fixing dish and let incubate for 20 minutes.</li>
<li>Submerge mPADS in 100 mm dish with 1X PBS three times.</li>
<li>Ready for immunostaining and mounting.</li>
</ol>
Analyzing Cellular Forces on mPADS Confocal Imaging Materials:
<ul>
<li>Confocal Microscope</li>
<li>63X Objective</li>
<li>Filter cubes for secondary antibodies</li>
<li>Green filter cube (Rhodamine) for DiI imaging of microneedles.</li>
</ul>
Method:
<ol>
<li> Place mPADs on stage and find single cells with 300 ms exposure, no binning.</li>
<li>Image tops of microneedles for TOP IMAGE.</li>
<li>Using z-control readout, capture image of microneedles for BOTTOM IMAGE.</li>
<li>Image any other channels you have stains in (Hoechst, phalloidin etc.)</li>
</ol>
Image Analysis Materials:
<ul>
<li>MATLAB with Image Processing Toolbox</li>
</ul>
Method:
<ol>
<li>Rotate and register top and bottom images</li>
<li>Measure centroids of bottom (undeflected) and top (deflected) positions</li>
<li>Calculate the displacement</li>
<li>Using the spring constant for the posts (known, depends on post geometry), calculate force vector at each post</li>
<li>Error analysis is based on the ability to resolve forces, and depends on the image resolution and centroid detection resolution</li>
<li>Arrange the data as per your customized needs &ndash; e.g., force magnitudes and/or direction, as a time course, many measurements across different conditions etc.</li>
</ol>

<ol>
	<li>Tan, J. L., Tien, J., Pirone, D., Gray, D. S., Chen, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cells+lying+on+a+bed+of+microneedles:+An+approach+to+isolate+mechanical+force.">Cells lying on a bed of microneedles: An approach to isolate mechanical force.</a> Proc Nat Acad Sci USA. 100, 1484-1489 (2003).</li><li>Sniadecki, N. J., Chen, C. S., Wang, Y. L., 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=Microfabricated+Silicone+Elastomeric+Post+Arrays+for+Measuring+Traction+Forces+of+Adherent+Cells.">Microfabricated Silicone Elastomeric Post Arrays for Measuring Traction Forces of Adherent Cells.</a> Methods in Cell Biology - Cell Mechanics. 83, 313-328 (2007).</li><li>Lemmon, C. A., Sniadecki, N. J., Ruiz, S. A., Tan, J. T., Romer, L. H., Chen, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Shear+Force+at+the+Cell-Matrix+Interface:+Enhanced+Analysis+For+Microfabricated+Post+Array+Detectors.">Shear Force at the Cell-Matrix Interface: Enhanced Analysis For Microfabricated Post Array Detectors.</a> Mechanics &amp; Chemistry of Biosystems. 2, 1-16 (2005).</li></ol>

microfabricated post-array-detectors (mpads): an approach to isolate mechanical forces

In this video, we will present our approach to measure cellular traction forces using a microfabricated array of posts.  Traction forces are generated through myosin-actin interactions and play an important role in our physiology.  During development, they enable cells to move from one location to the next in order to form the early structures of tissue.  Traction forces help in the healing processes.  They are necessary for the proper closure of wounds or the migration and crawling of leukocytes through our body.  These same forces can be detrimental to our health in the case of cancer metastasis or vascular growth towards a tumor.  The most common method by which to study cells in vitro has been to use a glass or polystyrene dish.  However, the rigidity of the substrates makes it impossible to physically measure cell traction forces, and there are relatively few methods to study traction forces.  Our lab has developed a technique to overcome these limitations.  The method is based on a vertical array of flexible cantilevers, the stiffness and size scale of which are such that individual cells spread across many cantilevers and deflect them in the process.  The pillars we use are 3 &mu;m in diameter, 10 &mu;m tall, and are configured in a regular array with 9 &mu;m center-to-center spacing.  But these physical dimensions can be readily varied to accommodate a variety of studies.  We start with a silicon master, but the final posts are made out of silicone rubber called poly (dimethyl siloxane), or PDMS.  We can measure the deflections under a microscope and calculate the magnitude and direction of traction forces required to produce the observed deflections.  We call these substrates microfabricated post-array-detectors, or mPADs.  Here, we will show you how we fabricate and use the mPADs to assess modulations of cellular contractility.

Watch this Scientific Journal Video about Microfabricated Post-Array-Detectors mPADs: an Approach to Isolate Mechanical Forces at JoVE.com

Microfabricated Post-Array-Detectors mPADs: an Approach to Isolate Mechanical Forces

In this video, we demonstrate how to fabricate and utilize microfabricated post array detectors (mPADs) to assess modulations of cellular contractility.

Microfabricated Post-Array-Detectors (mPADs): an Approach to Isolate Mechanical Forces

In this video, we will present our approach to measure cellular traction forces using a microfabricated array of posts.  Traction forces are generated ...

microfabricated-post-array-detectors-mpads-an-approach-to-isolate

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JoVE Journal

Biology

12.4K Views.  University of Pennsylvania. In this video, we demonstrate how to fabricate and utilize microfabricated post array detectors (mPADs) to assess modulations of cellular contractility.

Video: Microfabricated Post-Array-Detectors mPADs: an Approach to Isolate Mechanical Forces

Using Micro-Electro-Mechanical Systems (MEMS) to Develop Diagnostic Tools

Using Micro-Electro-Mechanical Systems MEMS to Develop Diagnostic Tools

An ex-ovo Chicken Embryo Culture System Suitable for Imaging and Microsurgery Applications

Understanding the relationships between genetic and microenvironmental factors that drive normal and malformed embryonic development is fundamental for discovering new therapeutic strategies. Advancements in imaging technology have enabled quantitative investigation of the organization and maturing of the body plan, but later stage embryonic morphogenesis is less clear. Chicken embryos are an attractive vertebrate animal model system for this application because of its ease of culture and surgical manipulation. Early embryos can be cultured for a short time on filter paper rings, which enables complete optical access for cell patterning and fate studies1,2. Studying advanced developmental processes such as cardiac morphogenesis are traditionally performed through a window of the eggshell3-5, but this technique limits optical access due to window size. We previously developed a simple method to culture whole embryos ex-ovo on hexagonal weigh boats for up to 10 days, which enabled high resolution imaging via ultrasonography6,7. These cultures were difficult to transport, limiting the types of imaging tools available for live experiments. We here present an improved shell-less culture system with a cost-effective, portable environmental chamber. Eggs were cracked onto a hammock created by a polyurethane membrane (cling wrap) affixed circumferentially to a plastic cup partially filled with sterile water. The dimensions of the circumference and depth of the hammock were both critical to maintain surface tension, while the mechanics of the hammock and water beneath helped dampen vibrations induced by transportation. A small footprint circulating water bath was also developed to enable continuous temperature control during experimentation. We demonstrate the ability to culture embryos in this way for at least 14 days without morphogenic defect or delay and employ this system in several microsurgical and imaging applications.

An ex-ovo Chicken Embryo Culture System Suitable for Imaging and Microsurgery Applications

In this article, we present a simple methodology to enable long-term ex-ovo avian embryo culture. This technique is ideal for longitudinal experimentation requiring complete optical accessibility and/or sterile transportation in avian embryos.

Understanding the relationships between genetic and microenvironmental factors that drive normal and malformed embryonic development is fundamental for ...

Using SecM Arrest Sequence as a Tool to Isolate Ribosome Bound Polypeptides

Extensive research has provided ample evidences suggesting that protein folding in the cell is a co-translational process1-5. However, the exact pathway that polypeptide chain follows during co-translational folding to achieve its functional form is still an enigma. In order to understand this process and to determine the exact conformation of the co-translational folding intermediates, it is essential to develop techniques that allow the isolation of RNCs carrying nascent chains of predetermined sizes to allow their further structural analysis.
SecM (secretion monitor) is a 170 amino acid E. coli protein that regulates expression of the downstream SecA (secretion driving) ATPase in the secM-secA operon6. Nakatogawa and Ito originally found that a 17 amino acid long sequence (150-FSTPVWISQAQGIRAGP-166) in the C-terminal region of the SecM protein is sufficient and necessary to cause stalling of SecM elongation at Gly165, thereby producing peptidyl-glycyl-tRNA stably bound to the ribosomal P-site7-9. More importantly, it was found that this 17 amino acid long sequence can be fused to the C-terminus of virtually any full-length and/or truncated protein thus allowing the production of RNCs carrying nascent chains of predetermined sizes7. Thus, when fused or inserted into the target protein, SecM stalling sequence produces arrest of the polypeptide chain elongation and generates stable RNCs both in vivo in E. coli cells and in vitro in a cell-free system. Sucrose gradient centrifugation is further utilized to isolate RNCs.
The isolated RNCs can be used to analyze structural and functional features of the co-translational folding intermediates. Recently, this technique has been successfully used to gain insights into the structure of several ribosome bound nascent chains10,11. Here we describe the isolation of bovine Gamma-B Crystallin RNCs fused to SecM and generated in an in vitro translation system.

We describe here a technique that is now routinely used to isolate stably bound ribosome nascent chain complexes (RNCs). This technique takes advantage of the discovery that a 17 amino acid long SecM "arrest sequence" can halt translation elongation in a prokaryotic (E. coli) system, when inserted into (or fused to the C-terminus) of virtually any protein.

Extensive research has provided ample evidences suggesting that protein folding in the cell is a co-translational process1-5. However, the exact pathway ...

An in vivo Crosslinking Approach to Isolate Protein Complexes From Drosophila Embryos

Many cellular processes are controlled by multisubunit protein complexes. Frequently these complexes form transiently and require native environment to assemble. Therefore, to identify these functional protein complexes, it is important to stabilize them in vivo before cell lysis and subsequent purification. Here we describe a method used to isolate large bona fide protein complexes from Drosophila embryos. This method is based on embryo permeabilization and stabilization of the complexes inside the embryos by in vivo crosslinking using a low concentration of formaldehyde, which can easily cross the cell membrane. Subsequently, the protein complex of interest is immunopurified followed by gel purification and analyzed by mass spectrometry. We illustrate this method using purification of a Tudor protein complex, which is essential for germline development. Tudor is a large protein, which contains multiple Tudor domains - small modules that interact with methylated arginines or lysines of target proteins. This method can be adapted for isolation of native protein complexes from different organisms and tissues.

An in vivo Crosslinking Approach to Isolate Protein Complexes From Drosophila Embryos

Multi-component protein complexes play crucial roles during cellular function and development. Here we describe a method used to isolate native protein complexes from Drosophila embryos after in vivo crosslinking followed by purification of the crosslinked complexes for subsequent structure-function analysis.

Many cellular processes are controlled by multisubunit protein complexes. Frequently these complexes form transiently and require native environment to ...

An Alternative Approach for Sample Preparation with Low Cell Number for TEM Analysis

Transmission electron microscopy (TEM) provides details of the cellular organization and ultrastructure. However, TEM analysis of rare cell populations, especially cells in suspension such as hematopoietic stem cells (HSCs), remains limited due to the requirement of a high cell number during sample preparation. There are a few cytospin or monolayer approaches for TEM analysis from scarce samples, but these approaches fail to get significant quantitative data from the limited number of cells. Here, an alternative and novel approach for sample preparation in TEM studies is described for rare cell populations that enables quantitative analysis.
A relatively low cell number, i.e., 10,000 HSCs, was successfully used for TEM analysis compared to the millions of cells typically used for TEM studies. In particular, Evans blue staining was performed after paraformaldehyde-glutaraldehyde (PFA-GA) fixation to visualize the tiny cell pellet, which facilitated embedding in agarose. Clusters of numerous cells were observed in ultra-thin sections. The cells had a well preserved morphology, and the ultra-structural details of the Golgi complex and several mitochondria were visible. This efficient, easy and reproducible protocol allows sample preparation from a low cell number and can be used for qualitative and quantitative TEM analysis on rare cell populations from limited biological samples.

Here, we present a protocol to prepare samples with low cell numbers for transmission electron microscopy (TEM) analysis.

Transmission electron microscopy (TEM) provides details of the cellular organization and ultrastructure. However, TEM analysis of rare cell populations, ...

Analyzing Cell Surface Adhesion Remodeling in Response to Mechanical Tension Using Magnetic Beads

Mechanosensitive cell surface adhesion complexes allow cells to sense the mechanical properties of their surroundings. Recent studies have identified both force-sensing molecules at adhesion sites, and force-dependent transcription factors that regulate lineage-specific gene expression and drive phenotypic outputs. However, the signaling networks converting mechanical tension into biochemical pathways have remained elusive. To explore the signaling pathways engaged upon mechanical tension applied to cell surface receptor, superparamagnetic microbeads can be used. Here we present a protocol for using magnetic beads to apply forces to cell surface adhesion proteins. Using this approach, it is possible to investigate not only force-dependent cytoplasmic signaling pathways by various biochemical approaches, but also adhesion remodeling by magnetic isolation of adhesion complexes attached to the ligand-coated beads. This protocol includes the preparation of ligand-coated superparamagnetic beads, and the application of define tensile forces followed by biochemical analyses. Additionally, we provide a representative sample of data demonstrating that tension applied to integrin-based adhesion triggers adhesion remodeling and alters protein tyrosine phosphorylation.

Cell surface adhesions are central in mechanotransduction, as they transmit mechanical tension and initiate the signaling pathways involved in tissue homeostasis and development. Here, we present a protocol for dissecting the biochemical pathways that are activated in response to tension, using ligand-coated magnetic microbeads and force application to adhesion receptors.

Mechanosensitive cell surface adhesion complexes allow cells to sense the mechanical properties of their surroundings. Recent studies have identified both ...

Assessment of Dictyostelium discoideum Response to Acute Mechanical Stimulation

Chemotaxis, or migration up a gradient of a chemoattractant, is the best understood mode of directed migration. Studies using social amoeba Dictyostelium discoideum revealed that a complex signal transduction network of parallel pathways amplifies the response to chemoattractants, and leads to biased actin polymerization and protrusion of a pseudopod in the direction of a gradient. In contrast, molecular mechanisms driving other types of directed migration, for example, due to exposure to shear flow or electric fields, are not known. Many regulators of chemotaxis exhibit localization at the leading or lagging edge of a migrating cell, as well as show transient changes in localization or activation following global stimulation with a chemoattractant. To understand the molecular mechanisms of other types of directed migration we developed a method that allows examination of cellular response to acute mechanical stimulation based on brief (2 - 5 s) exposure to shear flow. This stimulation can be delivered in a channel while imaging cells expressing fluorescently-labeled biosensors to examine individual cell behavior. Additionally, cell population can be stimulated in a plate, lysed, and immunoblotted using antibodies that recognize active versions of proteins of interest. By combining both assays, one can examine a wide array of molecules activated by changes in subcellular localization and/or phosphorylation. Using this method we determined that acute mechanical stimulation triggers activation of the chemotactic signal transduction and actin cytoskeleton networks. The ability to examine cellular responses to acute mechanical stimulation is important for understanding the initiating events necessary for shear flow-induced motility. This approach also provides a tool for studying the chemotactic signal transduction network without the confounding influence of the chemoattractant receptor.

Assessment of Dictyostelium discoideum Response to Acute Mechanical Stimulation

Here we describe methods for assessing cellular response to acute mechanical stimulation. In the microscopy-based assay, we examine localization of fluorescently-labeled biosensors following brief stimulation with shear flow. We also test activation of various proteins of interest in response to acute mechanical stimulation biochemically.

Chemotaxis, or migration up a gradient of a chemoattractant, is the best understood mode of directed migration. Studies using social amoeba Dictyostelium ...

Magnetic-Activated Cell Sorting Strategies to Isolate and Purify Synovial Fluid-Derived Mesenchymal Stem Cells from a Rabbit Model

Mesenchymal stem cells (MSCs) are the main cell source for cell-based therapy. MSCs from articular cavity synovial fluid could potentially be used for cartilage tissue engineering. MSCs from synovial fluid (SF-MSCs) have been considered promising candidates for articular regeneration, and their potential therapeutic benefit has made them an important research topic of late. SF-MSCs from the knee cavity of the New Zealand white rabbit can be employed as an optimized translational model to assess human regenerative medicine. By means of CD90-based magnetic activated cell sorting (MACS) technologies, this protocol successfully obtains rabbit SF-MSCs (rbSF-MSCs) from this rabbit model and further fully demonstrates the MSC phenotype of these cells by inducing them to differentiate to osteoblasts, adipocytes, and chondrocytes. Therefore, this approach can be applied in cell biology research and tissue engineering using simple equipment and procedures.

This article presents a simple and economic protocol for the straightforward isolation and purification of mesenchymal stem cells from New Zealand white rabbit synovial fluid.

Mesenchymal stem cells (MSCs) are the main cell source for cell-based therapy. MSCs from articular cavity synovial fluid could potentially be used for ...

Protrusion Force Microscopy: A Method to Quantify Forces Developed by Cell Protrusions

In numerous biological contexts, animal cells need to interact physically with their environment by developing mechanical forces. Among these, traction forces have been well-characterized, but there is a lack of techniques allowing the measurement of the protrusion forces exerted by cells orthogonally to their substrate. We designed an experimental setup to measure the protrusion forces exerted by adherent cells on their substrate. Cells plated on a compliant Formvar sheet deform this substrate and the resulting topography is mapped by atomic force microscopy (AFM) at the nanometer scale. Force values are then extracted from an analysis of the deformation profile based on the geometry of the protrusive cellular structures. Hence, the forces exerted by the individual protruding units of a living cell can be measured over time. This technique will enable the study of force generation and its regulation in the many cellular processes involving protrusion. Here, we describe its application to measure the protrusive forces generated by podosomes formed by human macrophages.

Here, we detail the experimental techniques used to evaluate the protrusion forces that podosomes apply on a compliant film, from the preparation of the film to the automated analysis of topographical images.

In numerous biological contexts, animal cells need to interact physically with their environment by developing mechanical forces. Among these, traction ...

A High Output Method to Isolate Cerebral Pericytes from Mouse

In recent years, cerebral pericytes have become the focus of extensive research in vascular biology and pathology. The importance of pericytes in blood brain barrier formation and physiology is now demonstrated but its molecular basis remains largely unknown. As the pathophysiological role of cerebral pericytes in neurological disorders is intriguing and of great importance, the in vitro models are not only sufficiently appropriate but also able to incorporate different techniques for these studies. Several methods have been proposed as in vitro models for the extraction of cerebral pericytes, although an antibiotic-free protocol with high output is desirable. Most importantly, a method that has increased output per extraction reduces the usage of more animals.
Here, we propose a simple and efficient method for extracting cerebral pericytes with sufficiently high output. The mouse brain tissue homogenate is mixed with a BSA-dextran solution for the separation of the tissue debris and microvascular pellet. We propose a three-step separation followed by filtration to obtain a microvessel rich filtrate. With this method, the quantity of microvascular fragments obtained from 10 mice is sufficient to seed 9 wells (9.6 cm2 each) of a 6-well plate. Most interestingly with this protocol, the user can obtain 27 pericyte rich wells (9.6 cm2 each) in passage 2. The purity of the pericyte cultures are confirmed with the expression of classical pericyte markers: NG2, PDGFR-&#946; and CD146. This method demonstrates an efficient and feasible in vitro tool for physiological and pathophysiological studies on pericytes.

We present a protocol for the extraction of murine cerebral pericytes. Based on an antibiotic-free enrichment oriented pericyte extraction, this protocol is a valuable tool for in vitro studies providing high purity and high yield, thus decreasing the number of experimental animals used.

In recent years, cerebral pericytes have become the focus of extensive research in vascular biology and pathology. The importance of pericytes in blood ...