We thank Dr. Grant Sumida for critical reading of the manuscript. This work was supported by a Beckman Young Investigator Award (SY), a Hellman Family New Faculty Award (SY), a NIH EUREKA, the University of California Cancer Research Coordinating Committee.

1. Generation of stable cell line (e.g. MDCK cells)
<ol>
	<li>Plate cells at 80-90% confluency in a p35 dish. Do not let cells form 100% confluent monolayer, which will reduce transfection efficiency.</li>
	<li>Transfect the cells with the plasmid of interest using Lipofectamine 2000. Optimize transfection conditions using manufacturer's protocol.</li>
	<li>Next day, passage the cells into two p150 petri dishes. The large dish is recommended to allow enough spacing between the stable colonies.</li>
	<li>Next day, change the media and add 500 &mu;g/ml of G418 to each dish. The G418 concentration should be optimized for individual cell lines.</li>
	<li>Change media every other day for approximately 2 weeks. After 2 weeks, G418-resistant colonies should start to form, and will be visible by the naked eye.</li>
	<li>Using an inverted fluorescent microscope, identify GFP positive colonies. Mark these colonies on the plate using a sharpie pen.</li>
	<li>To selectively trypsinize the colonies from the tissue culture plate, aspirate out the media, and wash the cells twice with PBS or trypsin solution. On the second wash, do not aspirate out all the solution. Leave a thin layer of liquid on the bottom of the plate to prevent cells from drying.</li>
	<li>For each marked colony: use a sterilized cotton swab to wipe as close as possible around the edge of the colony. This will create an island of wet area containing the cell colony. Pipette 10 &mu;l of trypsin onto the colony. Repeat for every colony, and proceed quickly to avoid drying. An experienced researcher can usually pick ~12 colonies per p150 plate.</li>
	<li>Incubate plate at 37 &deg;C for 5-10 minutes until cells detach from the plate and appear round.</li>
	<li>For each colony: Pipette 10 &mu;l of trypsin onto the colony, and pipette up and down a couple times to detach cells from the plate. Then pipette all the cells from the colony into a single well in a 24 well dish.</li>
	<li>After stable colonies have grown, protein expression of each colony is analyzed using standard western blot and immunofluorescence. Expand these cell lines for further analysis.</li>
</ol>
2. Surface modification of glass bottom dish for optimal collagen binding (Optional)
<ol>
	<li>To silanize the glass, pipette 300 &mu;l of 2% 3-Aminopropyltrimethoxysilane solution onto the glass portion of each p35 dish with a 10 mm opening. See Figure 1 for the silanization and cross-linking schematic. 3-Aminopropyltrimethoxysilane is diluted in filtered water.</li>
	<li>Incubate for 1 hour at room temperature.</li>
	<li>Aspirate out the 3-Aminopropyltrimethoxysilane solution and wash with filtered water three times for 10 minutes each.</li>
	<li>Aspirate out the water and place dishes on hot plate set to 50 &deg;C for 1.5 hours. Place tops of dishes slightly off the dish so that moisture in the dish can escape.</li>
	<li>Remove dishes from heat and allow to cool.</li>
	<li>Pipette 300 &mu;l of 2% glutaraldehyde solution onto the glass portion of each dish. The glutaraldehyde is diluted in PBS.</li>
	<li>Incubate for 1 hour.</li>
	<li>Aspirate out the glutaraldehyde solution and wash dishes with PBS three times for 10 minutes each.</li>
	<li>Sterilize dishes by exposure to UV light for 1 hour. The silanized dishes may be stored at room temperature.</li>
</ol>
3. Preparation of 3D collagen gel with tracer particles
<ol>
	<li>Wash fluorescent tracer particles by spinning down 100 &mu;l of stock tracer particles (1010 particles/ml) in a centrifuge at 15,000 rpm for 5 minutes, aspirate out liquid, and add 500 &mu;l of media. Repeat 5 times. After last wash, resuspend particles in 30 &mu;l of media.</li>
	<li>Trypsinize GFP expressing cells as usual.</li>
	<li>Resuspend cell pellet to approximately 2 x 106 cells/ml.</li>
	<li>Pipette 240 &mu;l of growth media into an eppendorf tube.</li>
	<li>Add 12.6 &mu;l of 1M Hepes, 20 &mu;l of filtered water, 50 &mu;l of cell solution, then 10 &mu;l of fluorescent particles.</li>
	<li>Lastly, add 167 &mu;l of bovine dermis Collagen I solution (to obtain a working concentration of 1 mg/ml).</li>
	<li>Mix solution thoroughly, then pipette 80 &mu;l of the solution onto the glass portion of the silanized glass bottom dish.</li>
	<li>Place dish into incubator and allow the gel to polymerize at 37 &deg;C for 30 minutes. See Figure 1A for typical collagen gel in a p35 glass bottom dish.</li>
	<li>Add 2 ml of growth media carefully to each dish.</li>
</ol>
4. Procedure for time-lapse image acquisition
<ol>
	<li>Turn on the temperature control of the microscope enclosure and let the scope chamber equilibrate to a steady state temperature to 37 &deg;C. See Figure 2 for an example of live-cell imaging system.</li>
	<li>Change media to the new media supplemented with 25 mM Hepes to maintain a neutral pH. For DIC imaging, exchange the top of the dish with one that has a glass top. Using a thin strip of parafilm, cover the side of the dish to prevent media evaporation.</li>
	<li>For an oil immersion objective, place one drop of immersion fluid on the objective.</li>
	<li>Place the collagen gel containing dish on microscope stage, and make sure the dish makes contact with the immersion liquid.</li>
	<li>Focus the sample and search for cells of interest to image. To minimize stage drift, allow dish to settle for about 45 minutes before starting a long capture.</li>
	<li>Specify the parameters of image acquisition. Minimize laser exposure, which could damage the cells, by modulating the laser power, exposure time, frequency and duration of capture time. The actual parameters will vary with the microscope system and cell type.</li>
	<li>During the time-lapse acquisition, an inhibitor may be added. For the inhibitor addition experiment, do not seal the dish with parafilm.
	<ol>
		<li>Prepare Hepes supplemented media with drug at a desired working concentration.</li>
		<li>When ready to add drug to media, pause the image capture, and carefully remove the dish top without disturbing the dish.</li>
		<li>Aspirate out the media in the dish, and pipette the drug containing media into the dish. Then, carefully replace the dish top.</li>
		<li>The sample may be out-of-focus due to the addition of new media, readjust the focus.</li>
		<li>Restart the capture, then monitor the capture for the next 30 minutes, and readjust the focus as necessary. See Figure 4 for an example of drug addition experiment.</li>
	</ol>
	</li>
</ol>
5. FRAP procedure and analysis
<ol>
	<li>The FRAP setup varies between systems, follow manufacturer's instruction.</li>
	<li>Set up parameters for live-cell imaging.</li>
	<li>Set up parameters for photobleaching. Use sufficient laser power to photobleach the fluorescent signal without damaging the cell. Test these parameters on practice cells.</li>
	<li>Start image capture, and allow at least 5 frames of image capture before photobleaching the region of interest.</li>
	<li>Continue capture, and recovery time should be sufficient to capture the full fluorescence recovery. See Figure 5 for an example of FRAP experiment.</li>
	<li>Analyze the fluorescence recovery by measuring the average fluorescent intensity of the photobleached area (before and after photobleaching) over time. Using a statistical analysis software (e.g. Excel), fit the recovery curve to the exponential equation: I=If(1-e-kt), where I is the intensity, If is the final intensity, t is time, and &tau;&frac12; is the time it takes for intensity to reach half the final value: &tau;&frac12;=ln(2)/k. The half-time is a measure of the rate of mobility of the protein.</li>
	<li>Obtain the parameters, half-time and final intensity from the exponential-fit curve. Calculate the mobile fraction by taking the ratio of the final to initial fluorescence intensities.</li>
</ol>
6. Representative Results
An example of the live-cell imaging of healthy epithelial cells within the matrix is shown in Figure 3. Healthy cells exhibit a smooth, continuous membrane, and distinct nucleus, whereas unhealthy cells often have a disrupted membrane, and an excessive number of vacuoles. In a 3D matrix, single epithelial cells migrate1, and over the course of several days, epithelial cells form three-dimensional, spherical, multi-cellular cysts within the matrix2. The cells are also highly dynamic, and migrate within the cyst (Figure 3). The matrix deformation as a result of the traction forces exerted by migrating cells is analyzed through the displacement of tracer particles embedded in the surrounding matrix (Figure 4). The tracer particle movements are displayed as a maximum projection image of different time points, and each time point is pseudo-colored to indicate the time (Figure 4B). Alternatively, the images of an individual tracer particle are displayed as a montage to show the movement of the tracer particle over time (Figure 4C). All image analysis was done using ImageJ. These qualitative assessments of matrix deformation, and therefore the forces exerted by migrating cells, are useful first approximation of traction force distribution. The quantitative estimation of traction forces exerted by migrating cells in 3D is beyond the scope of this protocol and is described elsewhere3,4.
A typical fluorescence recovery after photobleaching experiment is shown in Figure 5. The region of interest must be photobleached using optimized laser settings so that the fluorescence intensity is visibly diminished compared to background levels (to maximize signal-to-noise ratio), while maintaining healthy and undamaged cells. The optimal setting must be determined empirically as each FRAP setup is different (e.g., FRAP using laser-scanning confocal system).
The exposure and time-interval of post-FRAP image acquisition must be carefully controlled to avoid background photobleaching. The use of lower laser power, short exposure time, and the use of sensitive camera and high efficient optics are essential for high quality FRAP imaging. If the fluorescence bleaching during the image acquisition is significant, this background fading must be quantified and normalized from the observed fluorescence intensity recovery. Due to 3D nature of environment, the z-component of fluorescence intensity recovery may become significant. Therefore, it is important to define a photobleached volume by taking pre and post 3D-stack images of photobleached region. Further analysis and modeling of fluorescence recovery is discussed elsewhere5,6.
The unique combination of live-cell imaging techniques: GFP to follow the dynamics of cytoskeletal proteins, red fluorescence tracer particles to monitor the matrix deformation, and the addition of inhibitors, can be used simultaneously for the analysis of protein dynamics, traction force and molecular pathways.
<img alt="Figure 1" src="/files/ftp_upload/3589/3589fig1.jpg" /> 
Figure 1. Preparation of collagen gel. A) Polymerized collagen matrix on a glass-bottom dish. The pink color of the gel is due to embedded fluorescent particles. B) Procedure to treat the glass bottom dishes to crosslink the collagen gel to the glass surface. First, the glass bottom dish is treated with 3-Aminopropyltrimethoxysilane solution, then glutaraldehyde solution that crosslinks the collagen matrix to the glass.
<img alt="Figure 2" src="/files/ftp_upload/3589/3589fig2.jpg" /> 
Figure 2. Schematics of confocal/FRAP microscope setup. The confocal microscope is based on a Zeiss AxioObserver with a CoolSnap HQ II CCD camera and completely automated by Slidebook software (Intelligent Imaging Innovations). The confocal unit is custom designed and based on Yokogawa spinning disk unit CSU10 and two solid-state lasers (488 nm with 50 mW and 561 nm with 40 mW) with Acousto-Optical Tunable Filter (AOTF) to allow milliseconds switching between two lasers. The emission filters are 525/50 nm and 620/60 nm (#118661 and #118085, Chroma Technology) and dichroic mirror is 488-568 BrightLine Dual band (Semrock). The objectives are a long working distance 40x C-Apochromat objective with NA 1.1 and working distance of 0.62 mm, and a 63x Plan-Apochromat objective with NA 1.4 and a working distance of 0.19 mm. The microscope also includes a FRAP photoablation system that consists of a fiber optically pumped dye laser, a computer controlled beam position and intensity, and a diffraction limited spot size. Furthermore, the microscope is equipped with an x-y motorized stage that includes 0.1 micron linear encoders on each axis. During the time-lapse imaging, the environmental temperature is maintained by a custom designed microscope chamber and a heater with a feedback temperature control. To isolate any noise and vibration, the entire microscope system is on a vibration-free table.
<img alt="Figure 3" src="/files/ftp_upload/3589/3589fig3.jpg" /> 
Figure 3. Live cell imaging of epithelial cells expressing GFP-actin. These cells formed a cyst after 4 days of culture in a 3D collagen matrix. Some cells move along the surface of cyst (yellow arrowhead), while others migrate within the interior of cyst (red arrowhead). Scale bar 10 &mu;m, time in hours.
<img alt="Figure 4" src="/files/ftp_upload/3589/3589fig4.jpg" /> 
Figure 4. The effect of Rho-kinase inhibition on traction force. A) DIC image of a migrating MDCK cell expressing GFP tagged nuclear marker. The image was taken immediately before the treatment of Rho-kinase inhibitor Y-27632. Scale bar 10 &mu;m. B) Particle displacement resulting from the addition of Y-27632. The particle positions at various time points (0 - 52 min) were pseudo-colored according to the intensity scale, then projected onto a single image. The white region is the GFP positive nucleus in the migrating cells. Scale bar 10 &mu;m. Time in minutes. C) The particle movement at the trailing edge of the cell (see arrow in B). Asterisk denotes the last frame captured before Y-27632 addition. The tracer particle moved toward and away from the trailing edge of the migrating cell before and after the addition of Y-27632, respectively. Scale bar 1 &mu;m.
<img alt="Figure 5" src="/files/ftp_upload/3589/3589fig5.jpg" /> 
Figure 5. FRAP analysis of GFP-actin expressing cell migrating in a three-dimensional matrix. A) Timelapse images of the GFP-actin expressing cell of before and after the photobleaching. A small region of the GFP-actin accumulated at the cell rear was photobleached (red arrowhead) at timepoint 0. Time in seconds, scale bar 5 &mu;m. B) The average fluorescence intensity of the photobleached region (solid line), and the exponential fit of the fluorescence recovery (circles) plotted over time. The fluorescence intensity was normalized to the initial value before photobleaching.

<ol>
	<li>Shih, W., Yamada, S., Myosin, . <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+dependent+retrograde+flow+drives+3D+cell+migration.">A dependent retrograde flow drives 3D cell migration.</a> Biophys. J. 98 (8), L29-L31 (2010).</li><li>O'Brien, L. E., Yu, W., Tang, K., Jou, T. S., Zegers, M. M., Mostov, K. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphological+and+biochemical+analysis+of+Rac1+in+three-dimensional+epithelial+cell+cultures.">Morphological and biochemical analysis of Rac1 in three-dimensional epithelial cell cultures.</a> Methods Enzymol. 406, 676-691 (2006).</li><li>Legant, W. R., Miller, J. S., Blakely, B. L., Cohen, D. M., Genin, G. M., 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=Measurement+of+mechanical+tractions+exerted+by+cells+in+three-dimensional+matrices.">Measurement of mechanical tractions exerted by cells in three-dimensional matrices.</a> Nat. Methods. 7 (12), 969-971 (2010).</li><li>Del Alamo, J. C., Meili, R., Alonso-Latorre, B., Rodriguez-Rodriguez, J., Aliseda, A., Firtel, R. A., Lasheras, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatio-temporal+analysis+of+eurkaryotic+cell+motility+by+improved+force+cytometry.">Spatio-temporal analysis of eurkaryotic cell motility by improved force cytometry.</a> Proc. Natl. Acad. Sci. U.S.A. 104 (33), 13343-13348 (2007).</li><li>Lippincott-Schwartz, J., Snapp, E., Kenworthy, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Studying+protein+dynamics+in+living+cells.">Studying protein dynamics in living cells.</a> Nat. Rev. Mol. Cell Biol. 2 (6), 444-456 (2001).</li><li>Phair, R. D., Misteli, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinetic+modeling+approaches+to+in+vivo+imaging.">Kinetic modeling approaches to in vivo imaging.</a> Nat. Rev. Mol. Cell Biol. 2 (12), 898-907 (2001).</li><li>Giepmans, B. N., Adams, S. R., Ellisman, M. H., Tsien, R. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+toolbox+for+assessing+protein+location+and+function.">Fluorescent toolbox for assessing protein location and function.</a> Science. 312 (5771), 217-224 (2006).</li><li>Shaner, N. C., Steinbach, P. A., Tsien, R. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+guide+to+choosing+fluorescent+proteins.">A guide to choosing fluorescent proteins.</a> Nat. Methods. 2, 905-909 (2005).</li><li>Cukierman, E., Pankov, R., Stevens, D. R., Yamada, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Taking+cell-matrix+adhesions+to+the+third+dimension.">Taking cell-matrix adhesions to the third dimension.</a> Science. 294 (5547), 1708-1712 (2001).</li></ol>

No conflicts of interest declared.

Here we describe a method for using live-cell imaging to study the mechanisms of cell migration in a three-dimensional matrix. The success of this technique depends on obtaining "good" clones stably expressing GFP-tagged proteins. The low level of GFP proteins will require an excess excitation exposure that compromises cell health, while too high GFP level will have undesirable side effects on the cell. Thus, the vector choice to transfect genes into cells is important (e.g., the promoter, fluorescence tag, etc). There a...

<table border="1">
<tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments </td>
 </tr>
 <tr>
 <td>Collagen, bovine, Type I</td>
 <td>BD Biosciences</td>
 <td>354231</td>
 <td>Stock is about 3 mg/ml</td>
 </tr>
 <tr>
 <td>3-aminopropyltrimethoxysilane</td>
 <td>Sigma Aldrich</td>
 <td>281778</td>
 <td>Dilute in water</td>
 </tr>
 <tr>
 <td>glutaraldehyde</td>
 <td>Sigma Aldrich</td>
 <td>340855</td>
 <td>Dilute in PBS</td>
 </tr>
 <tr>
 <td>&nbsp;1M Hepes</td>
 <td>Invitrogen</td>
 <td>15630-080</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Fluospheres polystyrene microspheres 1 &micro;m, red fluorescence (580/605)</td>
 <td>Invitrogen</td>
 <td>F13083</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Geneticin (G418)</td>
 <td>Invitrogen</td>
 <td>11811-031</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td colspan="4">Culture media components:</td>
 </tr>
 <tr>
 <td>DMEM</td>
 <td>Invitrogen</td>
 <td>31600-034</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Fetal Bovine Serum</td>
 <td>Atlanta Biologicals</td>
 <td>S115500</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Penicillin/Streptomycin</td>
 <td>Invitrogen</td>
 <td>15140-122</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Kanamycin</td>
 <td>Invitrogen</td>
 <td>15160-054</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Live-cell Imaging of Migrating Cells Expressing Fluorescently-tagged Proteins in a Three-dimensional Matrix at JoVE.com

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

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 ...

live-cell-imaging-migrating-cells-expressing-fluorescently-tagged

Research

JoVE Journal

Bioengineering

13.0K Views.  University of California, Davis. 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.

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

Three-dimensional Optical-resolution Photoacoustic Microscopy

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As the mainstays of in vivo three-dimensional (3-D) optical microscopy, single-/multi-photon fluorescence microscopy and optical coherence tomography (OCT) have demonstrated their extraordinary sensitivities to fluorescence and optical scattering contrasts, respectively. However, the optical absorption contrast of biological tissues, which encodes essential physiological/pathological information, has not yet been assessable. 
The emergence of biomedical photoacoustics has led to a new branch of optical microscopy optical-resolution photoacoustic microscopy (OR-PAM)1, where the optical irradiation is focused to the diffraction limit to achieve cellular1 or even subcellular2 level lateral resolution. As a valuable complement to existing optical microscopy technologies, OR-PAM brings in at least two novelties. First and most importantly, OR-PAM detects optical absorption contrasts with extraordinary sensitivity (i.e., 100%). Combining OR-PAM with fluorescence microscopy3 or with optical-scattering-based OCT4 (or with both) provides comprehensive optical properties of biological tissues. Second, OR-PAM encodes optical absorption into acoustic waves, in contrast to the pure optical processes in fluorescence microscopy and OCT, and provides background-free detection. The acoustic detection in OR-PAM mitigates the impacts of optical scattering on signal degradation and naturally eliminates possible interferences (i.e., crosstalks) between excitation and detection, which is a common problem in fluorescence microscopy due to the overlap between the excitation and fluorescence spectra. 
Unique for optical absorption imaging, OR-PAM has demonstrated broad biomedical applications since its invention, including, but not limited to, neurology5, 6, ophthalmology7, 8, vascular biology9, and dermatology10. In this video, we teach the system configuration and alignment of OR-PAM as well as the experimental procedures for in vivo functional microvascular imaging.

Optical-resolution photoacoustic microscopy (OR-PAM) is an emerging technology capable of imaging optical absorption contrasts in vivo with cellular resolution and sensitivity. Here, we provide a visualized instruction on the experimental protocols of OR-PAM, including system configuration, system alignment, typical in vivo experimental procedures, and functional imaging schemes.

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As ...

Planar and Three-Dimensional Printing of Conductive Inks

Printed electronics rely on low-cost, large-area fabrication routes to create flexible or multidimensional electronic, optoelectronic, and biomedical devices1-3. In this paper, we focus on one- (1D), two- (2D), and three-dimensional (3D) printing of conductive metallic inks in the form of flexible, stretchable, and spanning microelectrodes.
Direct-write assembly4,5 is a 1-to-3D printing technique that enables the fabrication of features ranging from simple lines to complex structures by the deposition of concentrated inks through fine nozzles (~0.1 - 250 &mu;m). This printing method consists of a computer-controlled 3-axis translation stage, an ink reservoir and nozzle, and 10x telescopic lens for visualization. Unlike inkjet printing, a droplet-based process, direct-write assembly involves the extrusion of ink filaments either in- or out-of-plane. The printed filaments typically conform to the nozzle size. Hence, microscale features (&lt; 1 &mu;m) can be patterned and assembled into larger arrays and multidimensional architectures.
In this paper, we first synthesize a highly concentrated silver nanoparticle ink for planar and 3D printing via direct-write assembly. Next, a standard protocol for printing microelectrodes in multidimensional motifs is demonstrated. Finally, applications of printed microelectrodes for electrically small antennas, solar cells, and light-emitting diodes are highlighted.

Planar and three-dimensional printing of conductive metallic inks is described. Our approach provides new avenues for fabricating printed electronic, optoelectronic, and biomedical devices in unusual layouts at the microscale.

Printed electronics rely on low-cost, large-area fabrication routes to create flexible or multidimensional electronic, optoelectronic, and biomedical ...

Universal Hand-held Three-dimensional Optoacoustic Imaging Probe for Deep Tissue Human Angiography and Functional Preclinical Studies in Real Time

The exclusive combination of high optical contrast and excellent spatial resolution makes optoacoustics (photoacoustics) ideal for simultaneously attaining anatomical, functional and molecular contrast in deep optically opaque tissues. While enormous potential has been recently demonstrated in the application of optoacoustics for small animal research, vast efforts have also been undertaken in translating this imaging technology into clinical practice. We present here a newly developed optoacoustic tomography approach capable of delivering high resolution and spectrally enriched volumetric images of tissue morphology and function in real time. A detailed description of the experimental protocol for operating with the imaging system in both hand-held and stationary modes is provided and showcased for different potential scenarios involving functional and molecular studies in murine models and humans. The possibility for real time visualization in three dimensions along with the versatile handheld design of the imaging probe make the newly developed approach unique among the pantheon of imaging modalities used in today&#8217;s preclinical research and clinical practice.

We provide herein a detailed description of the experimental protocol for imaging with a newly developed hand-held optoacoustic (photoacoustic) system for three-dimensional functional and molecular imaging in real time. The demonstrated powerful performance and versatility may define new application areas of the optoacoustic technology in preclinical research and clinical practice.

The exclusive combination of high optical contrast and excellent spatial resolution makes optoacoustics (photoacoustics) ideal for simultaneously ...

Analysis of Cell Migration within a Three-dimensional Collagen Matrix

The ability to migrate is a hallmark of various cell types and plays a crucial role in several physiological processes, including&#160;embryonic development, wound healing, and immune responses. However, cell migration is also a key mechanism in cancer enabling these cancer cells to detach from the primary tumor to start metastatic spreading. Within the past years various cell migration assays have been developed to analyze the migratory behavior of different cell types. Because the locomotory behavior of cells markedly differs between a two-dimensional (2D) and three-dimensional (3D) environment it can be assumed that the analysis of the migration of cells that are embedded within a 3D environment would yield in more significant cell migration data. The advantage of the described 3D collagen matrix migration assay is that cells are embedded within a physiological 3D network of collagen fibers representing the major component of the extracellular matrix. Due to time-lapse video microscopy real cell migration is measured allowing the determination of several migration parameters as well as their alterations in response to pro-migratory factors or inhibitors. Various cell types could be analyzed using this technique, including lymphocytes/leukocytes, stem cells, and tumor cells. Likewise, also cell clusters or spheroids could be embedded within the collagen matrix concomitant with analysis of the emigration of single cells from the cell cluster/ spheroid into the collagen lattice. We conclude that the 3D collagen matrix migration assay is a versatile method to analyze the migration of cells within a physiological-like 3D environment.

Cell migration is a biological phenomenon that is involved in a plethora of physiological, such as wound healing and immune responses, and pathophysiological processes, like cancer. The 3D-collagen matrix migration assay is a versatile tool to analyze the migratory properties of different cell types within in a 3D physiological-like environment.

The ability to migrate is a hallmark of various cell types and plays a crucial role in several physiological processes, including&#160;embryonic ...

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 ...

Generation of a Three-dimensional Full Thickness Skin Equivalent and Automated Wounding

In vitro models are a cost effective and ethical alternative to study cutaneous wound healing processes. Moreover, by using human cells, these models reflect the human wound situation better than animal models. Although two-dimensional models are widely used to investigate processes such as cellular migration and proliferation, models that are more complex are required to gain a deeper knowledge about wound healing. Besides a suitable model system, the generation of precise and reproducible wounds is crucial to ensure comparable results between different test runs. In this study, the generation of a three-dimensional full thickness skin equivalent to study wound healing is shown. The dermal part of the models is comprised of human dermal fibroblast embedded in a rat-tail collagen type I hydrogel. Following the inoculation with human epidermal keratinocytes and consequent culture at the air-liquid interface, a multilayered epidermis is formed on top of the models. To study the wound healing process, we additionally developed an automated wounding device, which generates standardized wounds in a sterile atmosphere.

The goal of this protocol is to build up a three-dimensional full thickness skin equivalent, which resembles natural skin. With a specifically constructed automated wounding device, precise and reproducible wounds can be generated under maintenance of sterility.

In vitro models are a cost effective and ethical alternative to study cutaneous wound healing processes. Moreover, by using human cells, these models ...

Live Cell Imaging during Mechanical Stretch

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

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

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

Engineering Three-dimensional Epithelial Tissues Embedded within Extracellular Matrix

The architecture of branched organs such as the lungs, kidneys, and mammary glands arises through the developmental process of branching morphogenesis, which is regulated by a variety of soluble and physical signals in the microenvironment. Described here is a method created to study the process of branching morphogenesis by forming engineered three-dimensional (3D) epithelial tissues of defined shape and size that are completely embedded within an extracellular matrix (ECM). This method enables the formation of arrays of identical tissues and enables the control of a variety of environmental factors, including tissue geometry, spacing, and ECM composition. This method can also be combined with widely used techniques such as traction force microscopy (TFM) to gain more information about the interactions between cells and their surrounding ECM. The protocol can be used to investigate a variety of cell and tissue processes beyond branching morphogenesis, including cancer invasion.

This manuscript describes a soft lithography-based technique to engineer uniform arrays of three-dimensional (3D) epithelial tissues of defined geometry surrounded by extracellular matrix. This method is amenable to a wide variety of cell types and experimental contexts and allows for high-throughput screening of identical replicates.

The architecture of branched organs such as the lungs, kidneys, and mammary glands arises through the developmental process of branching morphogenesis, ...

Melt Electrospinning Writing of Three-dimensional Poly(&#949;-caprolactone) Scaffolds with Controllable Morphologies for Tissue Engineering Applications

This tutorial reflects on the fundamental principles and guidelines for electrospinning writing with polymer melts, an additive manufacturing technology with great potential for biomedical applications. The technique facilitates the direct deposition of biocompatible polymer fibers to fabricate well-ordered scaffolds in the sub-micron to micro scale range. The establishment of a stable, viscoelastic, polymer jet between a spinneret and a collector is achieved using an applied voltage and can be direct-written. A significant benefit of a typical porous scaffold is a high surface-to-volume ratio which provides increased effective adhesion sites for cell attachment and growth. Controlling the printing process by fine-tuning the system parameters enables high reproducibility in the quality of the printed scaffolds. It also provides a flexible manufacturing platform for users to tailor the morphological structures of the scaffolds to their specific requirements. For this purpose, we present a protocol to obtain different fiber diameters using melt electrospinning writing (MEW) with a guided amendment of the parameters, including flow rate, voltage and collection speed. Furthermore, we demonstrate how to optimize the jet, discuss often experienced technical challenges, explain troubleshooting techniques and showcase a wide range of printable scaffold architectures.

Melt Electrospinning Writing of Three-dimensional Poly(&amp;#949;-caprolactone) Scaffolds with Controllable Morphologies for Tissue Engineering Applications

This protocol serves as a comprehensive guideline to fabricate scaffolds via electrospinning with polymer melts in a direct writing mode. We systematically outline the process and define the appropriate parameter settings for achieving targeted scaffold architectures.

This tutorial reflects on the fundamental principles and guidelines for electrospinning writing with polymer melts, an additive manufacturing technology ...

Culturing Mammalian Cells in Three-dimensional Peptide Scaffolds

A useful technique for culturing cells in a self-assembling nanofiber three-dimensional (3D) scaffold is described. This culture system recreates an environment that closely mimics the structural features of non-polarized tissue. Furthermore, the particular intrinsic nanofiber structure of the scaffold makes it transparent to visual light, which allows for easy visualization of the sample under microscopy. This advantage was largely used to study cell migration, organization, proliferation, and differentiation and thus any development of their particular cellular function by staining with specific dyes or probes. Furthermore, in this work, we describe the good performance of this system to easily study the redifferentiation of expanded human articular chondrocytes into cartilaginous tissue. Cells were encapsulated into self-assembling peptide scaffolds and cultured under specific conditions to promote chondrogenesis. Three-dimensional cultures showed good viability during the 4 weeks of the experiment. As expected, samples cultured with chondrogenic inducers (compared to non-induced controls) stained strongly positive for toluidine blue (which stains glycosaminoglycans (GAGs) that are highly present in cartilage extracellular matrix) and expressed specific molecular markers, including collagen type I, II and X, according to Western Blot analysis. This protocol is easy to perform and can be used at research laboratories, industries and for educational purposes in laboratory courses.

Here, we present a protocol to obtain 3D-culture systems in self-assembling peptide scaffolds to promote the differentiation of dedifferentiated human articular chondrocytes into cartilage-like tissue.

A useful technique for culturing cells in a self-assembling nanofiber three-dimensional (3D) scaffold is described. This culture system recreates an ...