The authors wish to thank Dr. Douglas Goetz (Department of Chemical and Biomolecular Engineering, Ohio University) and Dr. Fabian Benencia (Department of Biomedical Sciences, Ohio University) for insightful discussions and manuscript review. We also thank Dr. Christopher Huppenbauer for helpful technical discussions (W. Nuhsbaum Inc.). This work was supported by grants from the National Science Foundation (CBET-1106118) and the National Institutes of Health (1R15CA161830-01).

1. Software Installation
<ol>
	<li>Purchase StreamPix 5 Multi-Camera software with SimulPix module or other imaging software capable of collecting and combining images captured by monochrome CCD cameras.</li>
	<li>Minimum requirements for StreamPix 5 include: A PC equipped with Core 2 Duo with 2.4 GHz or better with 2 GB RAM or higher. A supported IEEE digital or analog camera and compatible frame grabber. Windows XP/7/Vista 32 or 64 bit. A monitor supporting resolution 1,024 x 768 or higher. A graphic adapter with good 2D performance (OCU express 16x or better is recommended) and 7,200 RPM hard disk drive for recording with RAID-O configuration.</li>
</ol>
2. Installation of Dual Camera System Capable of Two Color Simultaneous Real-time Imaging
<ol>
	<li>Connect the DC2 Photometrics emission splitter, or similar dual camera emission splitting device, to the microscope by sliding the DC2 C-mount adapter into the video port of the microscope such that the housing of the dichroic on the device is accessible.</li>
	<li>Insert two monochrome CCD cameras into female C-mount 1 and female C-mount 2. Identical cameras are preferable. Similar cameras may be used, but compatibility is dependent on internal hardware, most importantly the CCD image sensor.</li>
	<li>Use imaging software to display images from both cameras. The following steps (2.3.1 - 2.3.4) apply to StreamPix 5 with SimulPix module and must be adapted for other multi-camera imaging software.
	<ol>
		<li>Open StreamPix 5 and select &#34;workspace&#34; in the task ribbon.</li>
		<li>Open &#34;workspace manager&#34; located at far right of the screen and select &#34;new workspace&#34;.</li>
		<li>After naming a camera, follow software prompts to select a grabber from a pre-populated list. Select the grabber matching the camera model and repeat for the second camera workspace. For the merged workspace, repeat once more but &#34;all cameras are in use&#34; error message will appear. This message is normal.</li>
		<li>Once the merged workspace is loaded, assign cameras from workspace 1 and workspace 2 to the merged workspace in the docking panel located on the right-hand side of the viewing screen.</li>
	</ol>
	</li>
	<li>Align cameras in the dual camera emission splitting system.
	<ol>
		<li>Align cameras in bright field or phase contrast with the calibration grid provided by the manufacturer using a PlanApo 40X objective (or greater).</li>
		<li>Send light to the microscope eyepieces, place the metallic-coated calibration grid that was provided by the manufacturer on the microscope stage, and square the grid on the microscope stage.</li>
		<li>Bring the grid into focus.</li>
		<li>Displace, but do not remove, the dichroic housing to align camera 1 (bypass camera). Display the grid without binning and without autoscaling. Focus the image using the focus adjustment dial on C-mount port 1.</li>
		<li>Push the dichroic housing into the dual channel acquisition device fully, so that the image is split by the dichroic to both cameras.</li>
		<li>Loosen the three 5/64&#34; set screws and rotate the second camera on the female C-mount 2 until the orientation of the grid is identical in both images. Using the focus adjustment dial on C-mount port 2, bring the image from camera 2 into focus.</li>
		<li>Finalize the alignment using the combined image in the merged workspace of the imaging software. This allows one to see a real-time pseudocolor overlay of the two images of the calibration grid. Adjust the combined image positions using only the R/L knob on the right side of the DC2. Adjust this knob until right/left vertical boundary of the images is precisely aligned.</li>
		<li>Use the U/D knob to adjust the up/down alignment of the second image until the horizontal grid bars are properly aligned.</li>
		<li>If during the up/down alignment a slight camera rotation occurs, correct this issue by loosening the three 5/64&#34; inch screws for camera 2 and rotate until the displayed image is properly aligned.</li>
	</ol>
	</li>
</ol>
3. Preparation of Cancer Cells for Parallel Plate Flow Chamber Adhesion Assay
<ol>
	<li>Culture BT-20 breast cancer cells in minimum essential medium (MEM) supplemented to complete medium with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin at 37 &#176;C and 5.0% CO2 as previously described 2.</li>
	<li>Harvest cancer cells with a dilute trypsin treatment and count cells with a hemocytometer.</li>
	<li>Wash and suspend cells at 107 cells/ml in 0.1% albumin from bovine serum (BSA) in DPBS with calcium and magnesium (DPBS+).</li>
	<li>Dilute primary antibodies, purified mouse anti-human CD24 and purified rat HECA-452 (anti-sialofucosylated antigens), to a predetermined optimal concentration in 0.1% BSA DPBS+. Incubate cells for 30 min 9, 10.</li>
	<li>Centrifuge cells at 1,200 rpm to obtain a cell pellet. Aspirate primary antibody, and wash cells twice with 0.1% BSA DBPS+.</li>
	<li>Dilute secondary antibodies, goat anti-mouse IgG AlexaFluor 568 (emission max, 603 nm), and goat anti-rat IgM AlexaFluor 488 (emission max, 519 nm), to a predetermined optimal concentration in 0.1% BSA DPBS+. Incubate cells for 30 min.</li>
	<li>Wash cells twice and suspend in 0.1% BSA DPBS+ at 106 cells/ml in 0.1% BSA DPBS+.</li>
</ol>
4. Preparation of Reactive Substrate for Parallel Plate Flow Chamber Adhesion Assay
<ol>
	<li>Connect two separate lengths of tubing to the inlet and outlet ports of the parallel plate flow chamber that fits inside a 35 mm tissue culture dish. One tube will connect to the reservoir of labeled cells suspended in 0.1% BSA DPBS+ and the other to the syringe pump, which will withdraw fluid at a specified flow rate.</li>
	<li>Connect a vacuum line to the flow chamber 1.</li>
	<li>Position the flow chamber over the 35 mm tissue culture dish containing a monolayer of Chinese hamster ovary cells expressing E-selectin (CHO-E). CHO-E cells were grown in MEM complete medium at 37 &#176;C and 5.0% CO2 2.</li>
	<li>Adjust the flow chamber and/or flow chamber gasket to ensure an adequate vacuum seal 1.</li>
	<li>Withdraw fluid from reservoir, monitoring the flow chamber assembly for proper seal and no visual evidence of air bubble formation 1.</li>
	<li>Place flow chamber assembly on microscope (Leica DMI 6000B)1.</li>
</ol>
5. Two-color Image Acquisition
<ol>
	<li>Power a Leica EL-6000 compact light source, or similar device, to generate high intensity light and couple it into a light guide. Use an inverted microscope to pass this light through a combination filter cube, i.e. G/R (green/red) filter cube, to excite fluorophores of a desired color/intensity.</li>
	<li>Ensure dichroic housing is in correct operating position (depressed) and is not in bypass mode.</li>
	<li>Operate microscope in fluorescence mode and select proper fluorescence filter cube (green/red).</li>
	<li>Determine the exposure time and gain in camera 1 (red; 620&#177;60 nm) and camera 2 (green; 535&#177;40 nm).
	<ol>
		<li>Depress the dichroic to obtain images in both the red and green emission channels. Use a flow chamber assay with cells labeled with the red fluorophore only, and obtain an image in the red channel by adjusting the exposure time of camera 1 in &#34;live settings&#34; of the imaging software.</li>
		<li>Match the exposure time of camera 2 to the exposure time of camera 1. If an image is observed in the green channel, bleed-through artifacts are present and exposure time in both cameras needs to be reduced.</li>
		<li>Keeping the exposure time equivalent in camera 1 and camera 2, reduce the exposure time until the bleed-through artifact is no longer visible in the green channel. Adjust the gain of camera 1 to improve image visibility in the red channel if necessary.</li>
		<li>Repeat steps 5.4.1 - 5.4.3 with cells labeled with the green fluorophore only, but define exposure time with camera 2 to image cells in the green channel without bleed-through artifacts in the red channel collected by camera 1.</li>
		<li>Retitrate antibodies if bleed-through artifacts cannot be eliminated 11, or if exposure times of camera 1 and camera 2 are substantially different such that merged images may be temporally misaligned.</li>
	</ol>
	</li>
 <li>Use imaging software to view images simultaneously captured from two monochrome CCD cameras in the flow chamber experiment.</li>
 <li>Merge images collected from both cameras using the SimulPix module of Streampix 5 or an alternative software package.</li>
 <li>Use digital correction adjustments in SimulPix or alternative software to spatially align the merged images, if necessary. Images can be corrected with the SimulPix module for horizontal, vertical, and rotational adjustments.</li>
</ol>

<ol>
	<li>Wiese, G., Barthel, S. R., Dimitroff, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+physiologic+E-selectin-mediated+leukocyte+rolling+on+microvascular+endothelium.">Analysis of physiologic E-selectin-mediated leukocyte rolling on microvascular endothelium.</a> J. Vis. Exp. (24), e1009 (2009).</li><li>Shirure, V. S., Reynolds, N. M., Burdick, M. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mac-2+binding+protein+is+a+novel+e-selectin+ligand+expressed+by+breast+cancer+cells.">Mac-2 binding protein is a novel e-selectin ligand expressed by breast cancer cells.</a> PLoS One. 7 (9), e44529 (2012).</li><li>Burdick, M. M., McCaffery, J. M., Kim, Y. S., Bochner, B. S., Colon Konstantopoulos, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=carcinoma+cell+glycolipids,+integrins,+and+other+glycoproteins+mediate+adhesion+to+HUVECs+under+flow.">carcinoma cell glycolipids, integrins, and other glycoproteins mediate adhesion to HUVECs under flow.</a> Am. J. Physiol. Cell Physiol. 284 (4), C977-C987 (2003).</li><li>Resto, V. A., Burdick, M. M., Dagia, N. M., McCammon, S. D., Fennewald, S. M., Sackstein, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=L-selectin-mediated+lymphocyte-cancer+cell+interactions+under+low+fluid+shear+conditions.">L-selectin-mediated lymphocyte-cancer cell interactions under low fluid shear conditions.</a> J. Biol. Chem. 283 (23), 15816-15824 (2008).</li><li>Shirure, V. S., Henson, K. A., Schnaar, R. L., Nimrichter, L., Burdick, M. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gangliosides+expressed+on+breast+cancer+cells+are+E-selectin+ligands.">Gangliosides expressed on breast cancer cells are E-selectin ligands.</a> Biochem Biophys. Res. Commun. 406 (3), 423-429 (2011).</li><li>Tees, D. F., Goetz, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Leukocyte+adhesion:+an+exquisite+balance+of+hydrodynamic+and+molecular+forces.">Leukocyte adhesion: an exquisite balance of hydrodynamic and molecular forces.</a> News Physiol. Sci. 18, 186-190 (2003).</li><li>Chang, K. C., Tees, D. F., Hammer, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+state+diagram+for+cell+adhesion+under+flow:+leukocyte+rolling+and+firm+adhesion.">The state diagram for cell adhesion under flow: leukocyte rolling and firm adhesion.</a> Proc. Natl. Acad. Sci. U.S.A. 97 (21), 11262-11267 (2000).</li><li>Lichtman, J. W., Conchello, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescence+microscopy.">Fluorescence microscopy.</a> Nat. Methods. 2 (12), 910-919 (2005).</li><li>Kummitha, C. M., Shirure, V. S., Delgadillo, L. F., Deosarkar, S. P., Tees, D. F., Burdick, M. M., Goetz, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=HECA-452+is+a+non-function+blocking+antibody+for+isolated+sialyl+Lewis+x+adhesion+to+endothelial+expressed+E-selectin+under+flow+conditions.">HECA-452 is a non-function blocking antibody for isolated sialyl Lewis x adhesion to endothelial expressed E-selectin under flow conditions.</a> J. Immunol. Methods. 384 (1-2), 43-50 (2012).</li><li>Burdick, M. M., Henson, K. A., Delgadillo, L. F., Choi, Y. E., Goetz, D. J., Tees, D. F., Benencia, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expression+of+E-selectin+ligands+on+circulating+tumor+cells:+cross-regulation+with+cancer+stem+cell+regulatory+pathways?.">Expression of E-selectin ligands on circulating tumor cells: cross-regulation with cancer stem cell regulatory pathways?.</a> Front. Oncol. 2, 103 (2012).</li><li>Hoffman, G. E., Le, W. W., Sita, L. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Importance+of+Titrating+Antibodies+for+Immunocytochemical+Methods.">The Importance of Titrating Antibodies for Immunocytochemical Methods.</a> Current Protocols in Neuroscience. , (2001).</li><li>Weinberg, S., Lipke, E. A., Tung, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+electrophysiological+mapping+of+stem+cells.">In vitro electrophysiological mapping of stem cells.</a> Methods Mol. Biol. 660, 215-237 (2010).</li><li>Kong, K. V., Leong, W. K., Lim, L. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Osmium+carbonyl+clusters+containing+labile+ligands+hyperstabilize+microtubules.">Osmium carbonyl clusters containing labile ligands hyperstabilize microtubules.</a> Chem. Res. Toxicol. 22 (6), 1116-1122 (2009).</li><li>Vermot, J., Fraser, S. E., Liebling, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fast+fluorescence+microscopy+for+imaging+the+dynamics+of+embryonic+development.">Fast fluorescence microscopy for imaging the dynamics of embryonic development.</a> HFSP J. 2 (3), 143-155 (2008).</li><li>Bullen, A., Friedman, R. S., Krummel, M. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+imaging+of+the+immune+system:+a+custom+technology+platform+for+high-speed,+multicolor+tissue+imaging+of+immune+responses.">Two-photon imaging of the immune system: a custom technology platform for high-speed, multicolor tissue imaging of immune responses.</a> Curr. Top. Microbiol. Immunol. 334, 1-29 (2009).</li><li>Worth, D. C., Parsons, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advances+in+imaging+cell-matrix+adhesions.">Advances in imaging cell-matrix adhesions.</a> J. Cell Sci. 123, 3629-3638 (2010).</li><li>Xiao, Z., Visentin, G. P., Dayananda, K. M., Neelamegham, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immune+complexes+formed+following+the+binding+of+anti-platelet+factor+4+(CXCL4)+antibodies+to+CXCL4+stimulate+human+neutrophil+activation+and+cell+adhesion.">Immune complexes formed following the binding of anti-platelet factor 4 (CXCL4) antibodies to CXCL4 stimulate human neutrophil activation and cell adhesion.</a> Blood. 112 (4), 1091-1100 (2008).</li></ol>

The authors declare that they have no competing financial interests.

The dual camera emission splitting system has the spatial, temporal, and optical resolution needed to capture high quality images in applications where cell or molecular movement is rapid. In generating the representative results, parameters in the dual camera emission splitting system, including software settings and camera settings, were optimized to obtain a merged image sequence in which rolling and adherent cells were spatially and temporally aligned. This optimization step is critical, as misalignment can result in images that do not properly convey the physical phenomenon under investigation, e.g. a single cell observed to roll in both the green and red channels may appear as two rolling cells in the merged workspace. Spatial alignment should be achieved in the setup of the dual camera emission splitting system when cameras are physically aligned as described in step 2.4 of the protocol. Digital corrections to the spatial alignment of images captured by the cameras can be applied for horizontal, vertical, or rotational offset(s) through digital alignment settings in the imaging software (step 5.7 and Figure 5).
Temporal alignment between images captured by camera 1 and camera 2 can be achieved by adjusting the "live settings" of the cameras: exposure time, offset, and gain. Gain and offset can be adjusted without affecting the temporal alignment of the merged images, yet deviations in the user-defined exposure time between cameras can result in misalignment. Thus gain can be adjusted to compensate for differences between exposure times used to collect images in each camera. Ideally, differences in the exposure time between cameras should only be allowed when gain adjustments appear to significantly compromise image quality. Alternatively, antibody titration experiments can be performed to optimize cell labeling to compensate for differences in exposure time required by each camera (a) due to inherent properties of each fluorophore or (b) due to differences in the molecular expression levels detected by each antibody/fluorophore. However, it should also be noted that antibody titration experiments are best-practice to prevent bleed-through artifacts (step 5.4.5)11.
Although the dual camera emission splitting system was used to simultaneously image two target molecules in separate emission channels, alternative technologies are able to simultaneously capture images in multiple emission channels. These imaging systems range from single color cameras to confocal microscopes, and each has advantages and disadvantages relative to image quality, speed, and cost 12-17. In particular, state-of-the-art laser scanning confocal microscopes that incorporate resonant scanners are impressive multi-spectral imaging systems capable of acquiring images at hundreds of frames per second. These confocal systems are extremely powerful, but their cost may limit the number of researchers that have access to these systems through core facilities. Ultimately, the investigational needs and resources of each lab dictate which multi-channel imaging system is most appropriate.
The methods described herein explain how to obtain real-time image sequences in two emission channels using a dual camera emission splitting system and imaging software. Spatial and temporal alignment of images captured in two emission channels by similar monochrome CCD cameras is essential for production of high quality real-time merged image sequences that simultaneously reveal multiple target molecules on cells. Applications for the dual camera emission splitting system include any two-color live cell analysis that demands exceptional optical and temporal resolution of a full field of view, such as flow assays, translocation of cell surface molecules, cytoskeletal remodeling, vesicular transport, and 3D particle tracking .

Methods for analysis of molecules on the cell surface, such as immunostaining, employ probes that are chemically conjugated to fluorophores, allowing detection of target molecules. Live cell imaging and hydrodynamic flow-based cell adhesion assays are typically recorded with monochrome CCD cameras designed to capture physiological processes at the cellular and/or molecular level 1, 2. These cameras are highly sensitive, deliver fast frame rates (greater than 30 frames per second), and provide exceptional temporal resolution (due to fast frame rates and short exposure times). However, monochrome cameras can only capture a single emission channel (detecting a single fluorophore) to collect images. Single camera emission splitting systems can be incorporated to capture multiple emission channels but often reduce the field of view and require the same exposure time for imaging all channels. To capture the full color spectrum from cells labeled with multiple fluorophores, a color camera can be used as an alternative. However, color cameras are not generally capable of providing the temporal resolution desired for live cell imaging in certain applications. Another imaging device is needed for applications in which it is advantageous to image live cells at multiple wavelengths while retaining a high temporal resolution. A prime experimental application is the parallel plate flow chamber adhesion assay, in which cells are perfused at physiologically relevant conditions over a potentially reactive substrate 1, 3. Cells in flow that express specific cell surface molecules may adhere and roll on the substrate, such as a cell monolayer expressing adhesion molecules or surface-adsorbed extracellular matrix proteins 4, 5. Rolling cells may undergo rotational and translational movement in fractions of a second. Molecular features on rolling and adherent cells, such as clusters of cell surface molecules, also have the potential to undergo active reorganization on the cell surface. Thus, imaging systems must provide an exceptional temporal resolution (30 frames per second or greater and "near zero" exposure times) to generate an image sequence that illustrates the step-by-step progression of cell rolling 6, 7. Dual camera emission splitting systems are capable of meeting these demands for imaging cells labeled with multiple fluorophores.
Dual camera emission splitting systems split and filter fluorescence channels into two similar cameras to simultaneously capture two spatially identical but fluorophore-specific images while retaining the full field of view. This technology enables direct comparison of the image captured in real-time in each channel and allows the user to quickly switch between camera models with different imaging capabilities. This feature is useful for making adjustments to image capture settings in one camera that better allow the system to capture fluorophores with different intensities, lifetimes, and extinction coefficients 8. Coupled with imaging software, dual camera emission splitting systems allow the real-time recording of live cell imaging assays in multiple wavelengths and may enhance in vitro assays that use fluorescence to study cell behavior.

<table border="1">
 <tbody>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Reagent/Material</td>
 </tr>
 <tr>
 <td>BT-20 cells</td>
 <td>ATCC</td>
 <td>HTB-19</td>
 <td></td>
 </tr>
 <tr>
 <td>CHO-E cells</td>
 <td>Gift from Dr. R. Sackstein (Brigham and Women's Hospital, Harvard Medical School)</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>MEM </td>
 <td>Thermo Scientific</td>
 <td>SH30024.01</td>
 <td></td>
 </tr>
 <tr>
 <td>FBS</td>
 <td>Thermo Scientific</td>
 <td>SH30396.03</td>
 <td></td>
 </tr>
 <tr>
 <td>Penicillin-streptomycin</td>
 <td>Thermo Scientific</td>
 <td>SV30010</td>
 <td></td>
 </tr>
 <tr>
 <td>0.25% Trypsin / 0.1% EDTA</td>
 <td>Thermo Scientific</td>
 <td>SV30031.01</td>
 <td></td>
 </tr>
 <tr>
 <td>DPBS</td>
 <td>Thermo Scientific </td>
 <td>SH30028.02</td>
 <td></td>
 </tr>
 <tr>
 <td>DPBS+</td>
 <td>Life Technologies</td>
 <td>14080-055</td>
 <td></td>
 </tr>
 <tr>
 <td>BSA</td>
 <td>Sigma</td>
 <td>A9647</td>
 <td></td>
 </tr>
 <tr>
 <td>HECA-452 monoclonal antibody</td>
 <td>BD Biosciences</td>
 <td>555946</td>
 <td></td>
 </tr>
 <tr>
 <td>Anti-human CD24 monoclonal antibody</td>
 <td>BD Biosciences</td>
 <td>555426</td>
 <td></td>
 </tr>
 <tr>
 <td>Anti-rat IgM AlexaFluor 488</td>
 <td>Invitrogen</td>
 <td>A21212</td>
 <td></td>
 </tr>
 <tr>
 <td>Anti-mouse IgG AlexaFluor 568</td>
 <td>Invitrogen</td>
 <td>A11004</td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>[header]</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Equipment</td>
 </tr>
 <tr>
 <td>EXi Blue Fluorescence Microscopy Digital CCD Camera</td>
 <td>Q Imaging</td>
 <td>EXI-BLU-R-F-M-14-C</td>
 <td></td>
 </tr>
 <tr>
 <td>Retiga EXi FAST 1394 Digital CCD Camera</td>
 <td>Q Imaging</td>
 <td>RET-EXi-F-M-12-C</td>
 <td></td>
 </tr>
 <tr>
 <td>DC2 Emission Splitter</td>
 <td>Photometrics</td>
 <td>DC2 </td>
 <td></td>
 </tr>
 <tr>
 <td>Inverted Fluorescence Microscope</td>
 <td>Leica</td>
 <td> DMI6000 B</td>
 <td></td>
 </tr>
 <tr>
 <td>Streampix 5 software</td>
 <td>Norpix</td>
 <td></td>
 <td></td>
 </tr>
 </tbody>
</table>

simultaneously capturing real-time images in two emission channels using a dual camera emission splitting system: applications to cell adhesion

Multi-color immunofluorescence microscopy to detect specific molecules in the cell membrane can be coupled with parallel plate flow chamber assays to investigate mechanisms governing cell adhesion under dynamic flow conditions. For instance, cancer cells labeled with multiple fluorophores can be perfused over a potentially reactive substrate to model mechanisms of cancer metastasis. However, multi-channel single camera systems and color cameras exhibit shortcomings in image acquisition for real-time live cell analysis. To overcome these limitations, we used a dual camera emission splitting system to simultaneously capture real-time image sequences of fluorescently labeled cells in the flow chamber. Dual camera emission splitting systems filter defined wavelength ranges into two monochrome CCD cameras, thereby simultaneously capturing two spatially identical but fluorophore-specific images. Subsequently, psuedocolored one-channel images are combined into a single real-time merged sequence that can reveal multiple target molecules on cells moving rapidly across a region of interest.

A parallel plate flow chamber adhesion assay was used to demonstrate a dual camera emission splitting system that simultaneously captured real-time image sequences in two emission channels (Figure 1). The dual camera emission splitting system detected BT-20 cells that were fluorescently labeled with anti-human CD24 and HECA-452 (detecting sialofucosylated antigens) monoclonal antibodies and appropriate secondary antibodies (Figure 2). Some rolling cells displayed red signals (CD24) yet undetectable green signals (HECA-452), others displayed green signals and undetectable red signals, and yet other cells displayed both red and green signals (Figure 2). Merged emission channels revealed the distribution and colocalization of CD24 and sialofucosylated molecules on the surface of BT-20 cells rolling on and adhering to CHO-E monolayers (Figures 3 and 4). The dual camera emission splitting system originally had a slight image alignment error due to the alignment of the cameras, but this error was corrected with digital adjustments (Figure 5) using the SimulPix module of StreamPix 5 . Altogether, the dual camera emission splitting system has the spatial, temporal, and optical resolution needed to reveal cellular and molecular features in applications such as flow chamber adhesion assays, in which cells move rapidly through the field of view.
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/50604/50604fig1.jpg" /> 
Figure 1. Illustration of a dual camera emission splitting system for acquiring images in a parallel plate flow chamber adhesion assay. A syringe pump is used to perfuse cells over substrate in the parallel plate flow chamber. The dual camera emission splitting system connected to an inverted Epifluorescence microscope uses a dichroic mirror and filter cubes to split and filter emission channels into two cameras. These cameras simultaneously capture two spatially identical but fluorophore-specific image sequences. A computer equipped with multi-camera imaging software is used to merge and record image sequences from each emission channel.
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/50604/50604fig2.jpg" /> 
Figure 2. A dual camera emission splitting system was used for real-time simultaneous imaging of a parallel plate flow chamber adhesion assay. BT-20 cells labeled with (1) anti-human CD24 and anti-mouse IgG AlexaFluor 568 (pseudocolored red, emission max, 603 nm; camera 1, 620&#177;60 nm) and (2) HECA-452 and anti-rat IgM AlexaFluor 488 (pseudocolored green, emission max 519 nm; camera 2, 535&#177;40 nm) were perfused over a CHO-E monolayer at wall shear stress = 1 dyne/cm2. Dual camera merged images reveal heterogeneity within a rolling cell population and can be used to characterize rolling behaviors based on expression of cell surface molecules. Scale bar = 100 &#956;m. Image acquired with a 10X objective.
<img alt="Figure 3" fo:content-width="6in" src="/files/ftp_upload/50604/50604fig3.jpg" /> 
Figure 3. A time sequence of merged images captured by a dual camera emission splitting system showing BT-20 cells expressing CD24 (pseudocolored red) and sialofucosylated molecules (pseudocolored green) rolling on a CHO-E monolayer at wall shear stress = 1 dyne/cm2. Cameras maintained the optical and temporal resolution to track cell features on a cell tumbling &#34;end over end&#34; as it rolled on the reactive substrate in the direction of flow. White arrows indicate a tracked cluster of cell surface molecules. Emission signals in each channel were pseudocolored and merged in the imaging software. Note that colocalized signals appear yellow/orange. Still images were exported from real-time image sequences at time points shown. Scale bar = 10 &#956;m. Images acquired with a 40X objective.
<img alt="Figure 4" fo:content-width="6in" src="/files/ftp_upload/50604/50604fig4.jpg" /> 
Figure 4. Dual camera emission splitting system reveals colocalization of CD24 (pseudocolored red) and sialofucosylated molecules (pseudocolored green) expressed by a representative BT-20 breast cancer cell rolling on a CHO-E monolayer. Heterogeneous expression of (A) CD24 and (B) sialofucosylated molecules on the cell surface is emphasized in each image. (C) Colocalization of molecules appears as yellow/orange pseudocolored spots when red and green pseudocolored emission channels are merged. Cell diameter is approximately 10 &#956;m. Images acquired with a 40X objective.
<img alt="Figure 5" fo:content-width="6in" src="/files/ftp_upload/50604/50604fig5.jpg" /> 
Figure 5.Adjustments to the horizontal, vertical, and rotational spatial settings of images captured in camera 1 and camera 2 can improve the spatial alignment in the merged image. Proper alignment is critical to accurately portray how cells are adhering and rolling in the adhesion assay. (A) A merged image of a single BT-20 cell is misaligned due to camera positioning. (B) Spatial alignment is improved with software adjustments in the SimulPix module of StreamPix 5. (C) Further software adjustments achieve near-perfect alignment. Scale bar = 2 &#956;m. Images acquired with a 40X objective.

Watch this Scientific Journal Video about Simultaneously Capturing Real-time Images in Two Emission Channels Using a Dual Camera Emission Splitting System: Applications to Cell Adhesion at JoVE.com

Simultaneously Capturing Real-time Images in Two Emission Channels Using a Dual Camera Emission Splitting System: Applications to Cell Adhesion

Dual camera emission splitting systems for two-color fluorescence microscopy generate real-time image sequences with exceptional optical and temporal resolution, a requirement of certain live cell assays including parallel plate flow chamber adhesion assays. When software is employed to merge images from simultaneously acquired emission channels, pseudocolored image sequences are produced.

Multi-color immunofluorescence microscopy to detect specific molecules in the cell membrane can be coupled with parallel plate flow chamber assays to ...

simultaneously-capturing-real-time-images-two-emission-channels-using

Nanyang Technological University

Research

JoVE Journal

Bioengineering

9.5K Views.  Ohio University. Dual camera emission splitting systems for two-color fluorescence microscopy generate real-time image sequences with exceptional optical and temporal resolution, a requirement of certain live cell assays including parallel plate flow chamber adhesion assays. When software is employed to merge images from simultaneously acquired emission channels, pseudocolored image sequences are produced.

Video: Simultaneously Capturing Real-time Images in Two Emission Channels Using a Dual Camera Emission Splitting System: Applications to Cell Adhesion

Creating Two-Dimensional Patterned Substrates for Protein and Cell Confinement

Microcontact printing provides a rapid, highly reproducible method for the creation of well-defined patterned substrates.1 While microcontact printing can be employed to directly print a large number of molecules, including proteins,2 DNA,3 and silanes,4 the formation of self-assembled monolayers (SAMs) from long chain alkane thiols on gold provides a simple way to confine proteins and cells to specific patterns containing adhesive and resistant regions. This confinement can be used to control cell morphology and is useful for examining a variety of questions in protein and cell biology. Here, we describe a general method for the creation of well-defined protein patterns for cellular studies.5 This process involves three steps: the production of a patterned master using photolithography, the creation of a PDMS stamp, and microcontact printing of a gold-coated substrate. Once patterned, these cell culture substrates are capable of confining proteins and/or cells (primary cells or cell lines) to the pattern.
The use of self-assembled monolayer chemistry allows for precise control over the patterned protein/cell adhesive regions and non-adhesive regions; this cannot be achieved using direct protein stamping. Hexadecanethiol, the long chain alkane thiol used in the microcontact printing step, produces a hydrophobic surface that readily adsorbs protein from solution. The glycol-terminated thiol, used for backfilling the non-printed regions of the substrate, creates a monolayer that is resistant to protein adsorption and therefore cell growth.6 These thiol monomers produce highly structured monolayers that precisely define regions of the substrate that can support protein adsorption and cell growth. As a result, these substrates are useful for a wide variety of applications from the study of intercellular behavior7 to the creation of microelectronics.8
While other types of monolayer chemistry have been used for cell culture studies, including work from our group using trichlorosilanes to create patterns directly on glass substrates,9 patterned monolayers formed from alkane thiols on gold are straight-forward to prepare. Moreover, the monomers used for monolayer preparation are commercially available, stable, and do not require storage or handling under inert atmosphere. Patterned substrates prepared from alkane thiols can also be recycled and reused several times, maintaining cell confinement.10

Self-assembled monolayers (SAMs) formed from long chain alkane thiols on gold provide well-defined substrates for the formation of protein patterns and cell confinement. Microcontact printing of hexadecanethiol using a polydimethylsiloxane (PDMS) stamp followed by backfilling with a glycol-terminated alkane thiol monomer produces a pattern where protein and cells adsorb only to the stamped hexadecanethiol region.

Microcontact printing provides a rapid, highly reproducible method for the creation of well-defined patterned substrates.1  While microcontact printing ...

Cell Co-culture Patterning Using Aqueous Two-phase Systems

Cell patterning technologies that are fast, easy to use and affordable will be required for the future development of high throughput cell assays, platforms for studying cell-cell interactions and tissue engineered systems. This detailed protocol describes a method for generating co-cultures of cells using biocompatible solutions of dextran (DEX) and polyethylene glycol (PEG) that phase-separate when combined above threshold concentrations. Cells can be patterned in a variety of configurations using this method. Cell exclusion patterning can be performed by printing droplets of DEX on a substrate and covering them with a solution of PEG containing cells. The interfacial tension formed between the two polymer solutions causes cells to fall around the outside of the DEX droplet and form a circular clearing that can be used for migration assays. Cell islands can be patterned by dispensing a cell-rich DEX phase into a PEG solution or by covering the DEX droplet with a solution of PEG. Co-cultures can be formed directly by combining cell exclusion with DEX island patterning. These methods are compatible with a variety of liquid handling approaches, including manual micropipetting, and can be used with virtually any adherent cell type.

Aqueous two-phase systems were used to simultaneously pattern multiple populations of cells. This fast and easy method for cell patterning takes advantage of the phase separation of aqueous solutions of dextran and polyethylene glycol and the interfacial tension that exists between the two polymer solutions.

Cell patterning  technologies that are fast, easy to use and affordable will be required for the  future development of high throughput cell assays, ...

Study of Cell Migration in Microfabricated Channels

The method described here allows the study of cell migration under confinement in one dimension. It is based on the use of microfabricated channels, which impose a polarized phenotype to cells by physical constraints. Once inside channels, cells have only two possibilities: move forward or backward. This simplified migration in which directionality is restricted facilitates the automatic tracking of cells and the extraction of quantitative parameters to describe cell movement. These parameters include cell velocity, changes in direction, and pauses during motion. Microchannels are also compatible with the use of fluorescent markers and are therefore suitable to study localization of intracellular organelles and structures during cell migration at high resolution. Finally, the surface of the channels can be functionalized with different substrates, allowing the control of the adhesive properties of the channels or the study of haptotaxis. In summary, the system here described is intended to analyze the migration of large cell numbers in conditions in which both the geometry and the biochemical nature of the environment are controlled, facilitating the normalization and reproducibility of independent experiments.

A quantitative method to study spontaneous migration of cells in a one-dimensional confined microenvironment is described. This method takes advantage of microfabricated channels and can be used to study migration of large number of cells under different conditions in single experiments.

The method described here allows the study of cell migration under confinement in one dimension. It is based on the use of microfabricated channels, which ...

A Novel Stretching Platform for Applications in Cell and Tissue Mechanobiology

Tools that allow the application of mechanical forces to cells and tissues or that can quantify the mechanical properties of biological tissues have contributed dramatically to the understanding of basic mechanobiology. These techniques have been extensively used to demonstrate how the onset and progression of various diseases are heavily influenced by mechanical cues. This article presents a multi-functional biaxial stretching (BAXS) platform that can either mechanically stimulate single cells or quantify the mechanical stiffness of tissues. The BAXS platform consists of four voice coil motors that can be controlled independently. Single cells can be cultured on a flexible substrate that can be attached to the motors allowing one to expose the cells to complex, dynamic, and spatially varying strain fields. Conversely, by incorporating a force load cell, one can also quantify the mechanical properties of primary tissues as they are exposed to deformation cycles. In both cases, a proper set of clamps must be designed and mounted to the BAXS platform motors in order to firmly hold the flexible substrate or the tissue of interest. The BAXS platform can be mounted on an inverted microscope to perform simultaneous transmitted light and/or fluorescence imaging to examine the structural or biochemical response of the sample during stretching experiments. This article provides experimental details of the design and usage of the BAXS platform and presents results for single cell and whole tissue studies. The BAXS platform was used to measure the deformation of nuclei in single mouse myoblast cells in response to substrate strain and to measure the stiffness of isolated mouse aortas. The BAXS platform is a versatile tool that can be combined with various optical microscopies in order to provide novel mechanobiological insights at the sub-cellular, cellular and whole tissue levels.

We present in this article a novel stretching platform that can be used to investigate single cell responses to complex anisotropic biaxial mechanical deformation and quantify the mechanical properties of biological tissues.

Tools that allow the application of mechanical forces to cells and tissues or that can quantify the mechanical properties of biological tissues have ...

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 ...

Electrotaxis Studies of Lung Cancer Cells using a Multichannel Dual-electric-field Microfluidic Chip

The behavior of directional cell migration under a direct current electric-field (dcEF) is referred to as electrotaxis. The significant role of physiological dcEF in guiding cell movement during embryo development, cell differentiation, and wound healing has been demonstrated in many studies. By applying microfluidic chips to an electrotaxis assay, the investigation process is shortened and experimental errors are minimized. In recent years, microfluidic devices made of polymeric substances (e.g., polymethylmethacrylate, PMMA, or acrylic) or polydimethylsiloxane (PDMS) have been widely used in studying the responses of cells to electrical stimulation. However, unlike the numerous steps required to fabricate a PDMS device, the simple and rapid construction of the acrylic micro&#64258;uidic chip makes it suitable for both device prototyping and production. Yet none of the reported devices facilitate the efficient study of the simultaneous chemical and dcEF effects on cells. In this report, we describe our design and fabrication of an acrylic-based multichannel dual-electric-field (MDF) chip to investigate the concurrent effect of chemical and electrical stimulation on lung cancer cells. The MDF chip provides eight combinations of electrical/chemical stimulations in a single test. The chip not only greatly shortens the required experimental time but also increases accuracy in electrotaxis studies.

Many microfluidic devices have been developed for use in the study of electrotaxis. Yet, none of these chips allows the efficient study of the simultaneous chemical and electric-field (EF) effects on cells. We developed a polymethylmethacrylate-based device that offers better-controlled coexisting EF and chemical stimulation for use in electrotaxis research.

The behavior of directional cell migration under a direct current electric-field (dcEF) is referred to as electrotaxis. The significant role of ...

Real Time Monitoring of Intracellular Bile Acid Dynamics Using a Genetically Encoded FRET-based Bile Acid Sensor

F&#246;rster Resonance Energy Transfer (FRET) has become a powerful tool for monitoring protein folding, interaction and localization in single cells. Biosensors relying on the principle of FRET have enabled real-time visualization of subcellular signaling events in live cells with high temporal and spatial resolution. Here, we describe the application of a genetically encoded Bile Acid Sensor (BAS) that consists of two fluorophores fused to the farnesoid X receptor ligand binding domain (FXR-LBD), thereby forming a bile acid sensor that can be activated by a large number of bile acids species and other (synthetic) FXR ligands. This sensor can be targeted to different cellular compartments including the nucleus (NucleoBAS) and cytosol (CytoBAS) to measure bile acid concentrations locally. It allows rapid and simple quantitation of cellular bile acid influx, efflux and subcellular distribution of endogenous bile acids without the need for labeling with fluorescent tags or radionuclei. Furthermore, the BAS FRET sensors can be useful for monitoring FXR ligand binding. Finally, we show that this FRET biosensor can be combined with imaging of other spectrally distinct fluorophores. This allows for combined analysis of intracellular bile acid dynamics and i) localization and/or abundance of proteins of interest, or ii) intracellular signaling in a single cell.

We provide a detailed protocol to study bile acid dynamics in living cells using a genetically encoded BAS FRET sensor. This Bile Acid Sensor represents a unique tool to study (regulation of) bile acid transport and FXR activation in a wide range of cell types.

F&#246;rster Resonance Energy Transfer (FRET) has become a powerful tool for monitoring protein folding, interaction and localization in single cells. ...

Hand-held Clinical Photoacoustic Imaging System for Real-time Non-invasive Small Animal Imaging

Translation of photoacoustic imaging into the clinic is a major challenge. Handheld real-time clinical photoacoustic imaging systems are very rare. Here, we report a combined photoacoustic and clinical ultrasound imaging system by integrating an ultrasound probe with light delivery for small animal imaging. We demonstrate this by showing sentinel lymph node imaging in small animals along with minimally invasive real-time needle guidance. A clinical ultrasound platform with access to raw channel data allows the integration of photoacoustic imaging leading to a handheld real-time clinical photoacoustic imaging system. Methylene blue was used for sentinel lymph node imaging at 675 nm wavelength. Additionally, needle guidance with dual modal ultrasound and photoacoustic imaging was shown using the imaging system. Depth imaging of up to 1.5 cm was demonstrated with a 10 Hz laser at a photoacoustic imaging frame rate of 5 frames per second.

A clinical handheld photoacoustic imaging system will be demonstrated for real-time non-invasive small animal imaging.

Translation of photoacoustic imaging into the clinic is a major challenge. Handheld real-time clinical photoacoustic imaging systems are very rare. Here, ...

A Protocol for Real-time 3D Single Particle Tracking

Real-time three-dimensional single particle tracking (RT-3D-SPT) has the potential to shed light on fast, 3D processes in cellular systems. Although various RT-3D-SPT methods have been put forward in recent years, tracking high speed 3D diffusing particles at low photon count rates remains a challenge. Moreover, RT-3D-SPT setups are generally complex and difficult to implement, limiting their widespread application to biological problems. This protocol presents a RT-3D-SPT system named 3D Dynamic Photon Localization Tracking (3D-DyPLoT), which can track particles with high diffusive speed (up to 20 &#181;m2/s) at low photon count rates (down to 10 kHz). 3D-DyPLoT employs a 2D electro-optic deflector (2D-EOD) and a tunable acoustic gradient (TAG) lens to drive a single focused laser spot dynamically in 3D. Combined with an optimized position estimation algorithm, 3D-DyPLoT can lock onto single particles with high tracking speed and high localization precision. Owing to the single excitation and single detection path layout, 3D-DyPLoT is robust and easy to set up. This protocol discusses how to build 3D-DyPLoT step by step. First, the optical layout is described. Next, the system is calibrated and optimized by raster scanning a 190 nm fluorescent bead with the piezoelectric nanopositioner. Finally, to demonstrate real-time 3D tracking ability, 110 nm fluorescent beads are tracked in water.

This protocol details the construction and operation of a real-time 3D single particle tracking microscope capable of tracking nanoscale fluorescent probes at high diffusive speeds and low photon count rates.

Real-time three-dimensional single particle tracking (RT-3D-SPT) has the potential to shed light on fast, 3D processes in cellular systems. Although ...

In Situ Microscopy for Real-time Determination of Single-cell Morphology in Bioprocesses

In situ monitoring in microbial bioprocesses is mostly restricted to chemical and physical properties of the medium (e.g.,pH value and the dissolved oxygen concentration). Nevertheless, the morphology of cells can be a suitable indicator for optimal conditions, since it changes with dependence on the growth state, product accumulation and cell stress. Furthermore, the single-cell size distribution provides not only information about the cultivation conditions, but also about the population heterogeneity. To gain such information, a photo-optical in situ microscopy device1 was developed to enable the monitoring of the single-cell size distribution directly in the cell suspension in bioreactors. An automated image analysis is coupled to the microscopy based on a neural network model, which is trained with user-annotated images. Several parameters, which are gained from the captures of the microscope, are correlated to process relevant features of the cells, like their metabolic activity. Until now, the presented in situ microscopy probe series was applied to measure the pellet size in filamentous fungi suspensions. It was used to distinguish the single-cell size in microalgae cultivation and relate it to lipid accumulation. The shape of cellular particles was related to budding in yeast cultures. The microscopy analysis can be generally split into three steps: (i) image acquisition, (ii) particle identification, and (iii) data analysis, respectively. All steps have to be adapted to the organism, and therefore specific annotated information is required in order to achieve reliable results. The ability to monitor changes in cell morphology directly in line or on line (in a by-pass) enables real-time values for monitoring and control, in process development as well as in production scale. If the off line data correlates with the real-time data, the current tedious off line measurements with unknown influences on the cell size become needless.

In Situ Microscopy for Real-time Determination of Single-cell Morphology in Bioprocesses

A photo-optical in situ microscopy device was developed to monitor the size of single cells directly in the cell suspension. The real-time measurement is conducted by coupling the photo-optical sterilizable probe to an automated image analysis. Morphological changes appear with dependence on the growth state and cultivation conditions.

In situ monitoring in microbial bioprocesses is mostly restricted to chemical and physical properties of the medium (e.g.,pH value and the dissolved ...

Simultaneously Capturing Real-time Images in Two Emission Channels Using a Dual Camera Emission Splitting System: Applications to Cell Adhesion

Summary

Explore More Videos

Chapters in this video

Creating Two-Dimensional Patterned Substrates for Protein and Cell Confinement

Cell Co-culture Patterning Using Aqueous Two-phase Systems

Study of Cell Migration in Microfabricated Channels

A Novel Stretching Platform for Applications in Cell and Tissue Mechanobiology

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

Electrotaxis Studies of Lung Cancer Cells using a Multichannel Dual-electric-field Microfluidic Chip

Real Time Monitoring of Intracellular Bile Acid Dynamics Using a Genetically Encoded FRET-based Bile Acid Sensor

Hand-held Clinical Photoacoustic Imaging System for Real-time Non-invasive Small Animal Imaging

A Protocol for Real-time 3D Single Particle Tracking

In Situ Microscopy for Real-time Determination of Single-cell Morphology in Bioprocesses