This work was supported by NCI grant nos. U54CA143803, CA163124, CA093900, and CA143055 to K. J. P. This research was supported by a grant of the Korea Health Technology R&#38;D Project through the Korea Health Industry Development Institute (KHIDI), funded by the Ministry of Health &#38; Welfare, Republic of Korea (grant number: HI19C1122). Work by J. Kim and Y.-K. Cho was supported by Institute for Basic Science (IBS-R020-D1), funded by the Korean Government. The authors thank the current and past members of the Brady Urological Institute, especially members of the Pienta-Amend laboratory, for the critical reading of the manuscript.

1. EV isolation and on-chip immuno-fluorescent EV labeling
<ol>
	<li>Collection of cell culture media (CCM) and pre-processing of CCM for EV isolation
	<ol>
		<li>Seed PC3 cells at 30% confluency in a 75 cm2 cell culture flask. Allow control cells to grow to 90% confluency (~48 h) in standard media and cell-line-specific supplements. 
		NOTE: To prevent EV-containing components from affecting cellular uptake (i.e., fetal bovine serum), use exosome-depleted media and supplements.</li>
		<li>Harvest the CCM.</li>
		<li>Centrifuge the CCM at 1000 x g for 10 min at room temperature (RT) to pellet any unattached cells and large debris harvested with the media. Transfer the supernatant to a new conical tube.</li>
		<li>In the new tube, centrifuge the supernatant at 10,000 x g for 20 min at 4 &#176;C to pellet smaller debris and apoptotic bodies remaining in the media. Some larger EVs will pellet. Transfer the supernatant to a new tube.</li>
		<li>Filter the supernatant through a 0.45 &#181;m hydrophilic Polyvinylidene fluoride (PVDF) membrane syringe filter. 
		NOTE: If not immediately processing CCM for EV isolation, store the pre-processed CCM at -80 &#176;C until isolation is performed. If frozen, limit freeze-thaw cycles to one.</li>
	</ol>
	</li>
	<li>EV isolation from CCM using a nano-filtration based microfluidic device
	<ol>
		<li>If frozen, completely thaw CCM and vortex for 30 s before step 1.2.2.</li>
		<li>Inject 1 mL of pre-processed CCM (step 1.1) into the sample chamber of the nano-filtration-based microfluidic device (see Table of Materials)17,26. 
		NOTE: Follow the standard operating procedure for the nano-filtration-based microfluidic device17,26.</li>
		<li>Spin at 3000 rpm for 10 min in the bench-top spinning machine (see Table of Materials) to operate the microfluidic device. 
		NOTE: If CCM remains on the sample chamber following the initial run, perform additional spins until all CCM has emptied from the sample chamber.</li>
		<li>Remove the fluid from the waste chamber by pipetting and repeat steps 1.2.1, 1.2.2, and 1.2.3 twice. 
		NOTE: In total, 3 mL of CCM will be processed for EV isolation.</li>
		<li>Inject 1 mL phosphate-buffered saline (PBS) into the sample chamber to wash the isolated EVs. Spin in the bench-top spinning machine for operating the microfluidic device as mentioned in step 1.2.3. Locate the pure EVs on the membrane of the device. 
		NOTE: The quality of EVs isolated from the nano-filtration based microfluidic device, specifically, was confirmed and compared to the conventional UC method by transmission electron microscopy (TEM), scanning electron microscope (SEM), nanoparticle tracking analysis (NTA), structured illumination microscopy, enzyme-linked immunosorbent assay and real-time PCR in the previous research17,26.</li>
	</ol>
	</li>
	<li>Immunofluorescent labeling of EV using nano-filtration based microfluidic device (Figure 1)
	<ol>
		<li>Select an EV-specific antibody according to the purpose of the assay (see Table of Materials). 
		NOTE: Certain antibodies may interfere with ligand binding sites specific to EV-uptake pathways (i.e., endocytosis).</li>
		<li>Inject 1 &#181;g/mL of the EV-specific antibody into the elution hole of the device containing 100 &#181;L of isolated EVs.</li>
		<li>Incubate for 1 h in the dark at RT on a plate shaker to ensure the even distribution of the antibody across the sample.</li>
		<li>Attach an adhesive tape to the elution hole. Inject 1 mL PBS into the sample chamber to wash out any residual antibodies.</li>
		<li>Spin the device at 3000 rpm until the sample chamber is empty. Remove any fluid from the waste chamber by pipetting. Inject 1 mL PBS into the sample chamber. 
		NOTE: Fluorescently labeled EVs will be located in the membrane chamber.</li>
		<li>Pipette the fluorescently labeled EVs (Figure 2) from the membrane chamber to an amber tube. Block from light until use.</li>
	</ol>
	</li>
</ol>
2. Incubation of the cells with fluorescently labeled EVs for the EV-uptake assay
<ol>
	<li>Target cell seeding and culture on the cell-culture compatible dishes.
	<ol>
		<li>Seed 1 x 104 PC3 cells into the microslide 8-well plate (9.4 x 10.7 mm for each well) with 0.2 mL of media or 4 x 104 PC3 cells into a 35 mm dish with 1 mL of media. Plate the cells into a cell-culture compatible dish consisting of a thin coverslip (thickness: 0.18 mm). 
		NOTE: The thin coverslip minimizes the adverse scattering of light.</li>
		<li>Allow cells to adhere overnight in optimal cell-culture conditions (37 &#176;C, 5% CO2 concentration, 90% humidity).</li>
		<li>Wash adhered cells twice with exosome-depleted media (described in step 1.1.1).</li>
	</ol>
	</li>
	<li>Cell incubation with fluorescently labeled EVs.
	<ol>
		<li>Measure the concentration of fluorescently labeled EVs (step 1.3.5) by nanoparticle tracking analysis (NTA, Supplementary Figure 1). Determine the optimal concentration of fluorescently labeled EVs to be added to the cultured cells (step 2.1.2.).</li>
		<li>Dilute the fluorescently labeled EVs with exosome-depleted media to match the desired concentration measured in step 2.2.1. (i.e., 7.80 x 109 EVs (in NTA value) in 200 &#181;L of exosome-depleted media.)</li>
		<li>Add the diluted EVs (Step 2.2.2) to the adhered target cells prepared at 2.1.2. Incubate for experimental time (i.e., 4, 8, or 12 h).</li>
		<li>Wash cells thrice with exosome-free media to remove any non-internalized EVs. 
		NOTE: Optional: Cells can be fixated following wash.</li>
		<li>Label the cytoplasm of the adhered cells with 1 &#181;g/mL of CMTMR ((5-(and-6)-(((4-chloromethyl)benzoyl)amino) tetramethylrhodamine) (see Table of Materials) and incubate in optimal cell-culture conditions (37 &#176;C, 5% CO2 concentration, 90% humidity). 
		NOTE: Cell area dyes should fluoresce separately from labeled EVs to aid in determining the spatial location (internalized or superficial) of the spiked EVs during the EV uptake assay.</li>
		<li>Wash labeled cells twice with exosome-depleted media to remove the residual dye. Add fresh exosome-depleted media to the cells in preparation for live-cell confocal imaging.</li>
	</ol>
	</li>
</ol>
3. Confocal microscopy
<ol>
	<li>To perform live-cell imaging, utilize an on-stage incubator to maintain optimal cell-culture conditions (37 &#176;C, 5% CO2 concentration, 90% humidity).</li>
	<li>Place the prepared cells in the on-stage incubator.</li>
	<li>Set the imaging parameters based on control samples. 
	NOTE: Suggested control samples include: Fluorescently labeled EVs only, fluorescently labeled cells, unlabeled EVs, and unlabeled cells.</li>
	<li>Determine the depth of the target cells and the range of stacking size in the z-direction to acquire 3D confocal images. 
	NOTE: The thickness of a Z-stack is 1 &#181;m. The confocal 3D image acquisition lasted 2 min 34 s (each Z-plane image acquisition took approximately 8 s; a total of twenty Z-stack images).</li>
	<li>Set image acquisition to multiple z-stacked images of both cell-specific dye (i.e., red) and EV-specific dye (i.e., green) simultaneously (Figure 3 and Figure 4A).</li>
</ol>
4. Image processing
<ol>
	<li>Utilize automatic image-processing software to analyze the raw z-stacked confocal images and determine the EV uptake by cells (see Table of Materials).</li>
	<li>Set thresholding parameters to the fluorescent signal of the cells and EV-specific dyes. Build the virtual surfaces of cells (Figure 4A,B).
	<ol>
		<li>To build the virtual surfaces of cells, click the button Add new Surfaces.</li>
		<li>Select Shortest Distance Calculation as &#34;Algorithm Settings&#34; to use the provided algorithm by the software, then click Next: Source Channel.</li>
		<li>Select Channel 2 - CMTMR as &#34;Source Channel&#34; in this experiment.</li>
		<li>Select Smooth and put the appropriate value into &#34;Surfaces Detail&#34; for surface smoothing. 
		NOTE: 0.57 &#181;m in this experiment since 1 pixel represents 0.57 &#181;m in raw imaging data.</li>
		<li>Select Absolute Intensity as &#34;Thresholding.&#34;</li>
		<li>To automatically threshold the fluorescent image by the provided algorithm, click Threshold (Absolute Intensity): The value is automatically set.</li>
		<li>Select Enable as &#34;Split touching Objects (Region Growing)&#34; and put the value of estimated cell size into &#34;Seed Points Diameter,&#34; 10.0 &#181;m in this experiment. Then click Next: Filter Seed Points.</li>
		<li>To configure the virtual cell surfaces, click + Add button, then select Quality as &#34;Filter Type.&#34; Threshold the appropriate value (210 in this experiment) for the low limit by a visual inspection and the maximum value (1485) for the upper limit, then click the Finish button. 
		NOTE: A visual inspection means that a researcher can discriminate the cellular area from a raw fluorescent image.</li>
		<li>Next, to build the virtual dots of EVs, click the button Add new Spots.</li>
		<li>Select Different Spot Sizes (Region Growing) and Shortest Distance Calculation as &#34;Algorithm Settings,&#34; then click Next: Source Channel.</li>
		<li>Select Channel 1 - Alexa Fluor 488 as &#34;Source Channel&#34; in this experiment.</li>
		<li>Put the appropriate value into &#34;Estimated XY Diameter&#34; for the spot detection, 1 &#181;m in this experiment. Then, click Next: Filter Spots.</li>
		<li>To configure the virtual EV dots, click + Add button, select Quality as &#34;Filter Type,&#34; and set &#34;Lower Threshold&#34; by a visual inspection, 100 in this experiment. Then, click the Next: Spot Region Type button.</li>
		<li>Select Absolute Intensity as &#34;Spot Regions Type,&#34; then click Next: Spot Regions.</li>
		<li>To threshold, the region of EV dots, put the appropriate value into &#34;Region Threshold&#34; by a visual inspection, 100 as &#34;Region Threshold&#34; in this experiment. 
		NOTE: A visual inspection means that a researcher can discriminate the EVs area from a raw fluorescent image.</li>
		<li>Select Region Volume as &#34;Diameter from,&#34; then click Finish.</li>
	</ol>
	</li>
	<li>Use the software&#39;s provided algorithms to split the grouped spots inside the built surface at step 4.2 (Figure 4C, i-iv).
	<ol>
		<li>Click the built Spots, then go into Filters.</li>
		<li>Click + Add button, then select Shortest Distance to Surfaces Surfaces = Surface 1 as Filter Type, then click Duplicate Selection to new Spots button. The lowest threshold (-7.0 in this experiment) for the low limit and the appropriate value (-0.5) for the upper limit. 
		NOTE: Set the upper limit with the estimated radius of Spots. In this experiment, the estimated diameter of Spots,i.e., EV dots, was set to 1 &#181;m in step 4.2.11.; thus, the upper limit can be 0.5.</li>
	</ol>
	</li>
	<li>Automatic count of EVs inside the cells 
	NOTE: The software will automatically count the number of EVs inside the cells, indicating the number internalized by the target cells.
	<ol>
		<li>Click the built Spots 1 selection [Shortest Distance to Surfaces Surfaces = Surfaces 1 between -7.00 and -0.500].</li>
		<li>Go to the Statistics, and export the value from &#34;Total Number of Spots&#34; 
		NOTE: The software&#39;s provided algorithms will automatically calculate the number and volume of cells.</li>
	</ol>
	</li>
	<li>Determine the yield of EV uptake per incubation period based on the above-calculated values (Figure 5).
	<ol>
		<li>To obtain the number of cells, click the built Surfaces 1, then go to the Statistics, and export the value of &#34;Total Number of Surfaces&#34; from Overall.</li>
		<li>Go to the Detailed into Statistics to export the Volume from Detailed.</li>
	</ol>
	</li>
</ol>

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Y.-K. Cho is an inventor of the patents on the nano-filtration-based microfluidic device, Exodisc, which are licensed to Labspinner (Ulsan, Korea). All other authors have nothing to disclose.

An EV uptake assay based on 3D fluorescence imaging via confocal microscopy provides an efficient methodology and sensitive analysis. This fluorescent EV labeling facilitates the visualization of EVs and successfully performs a precise EV uptake assay. Previous methods for labeling EVs and removing the residual dye have been reported by removing precipitation using ultracentrifugation (UC); however, UC may co-precipitate EVs, and the immobilized dye may lead to a false-positive signal12<s...

Extracellular vesicles (EVs) are nano-sized, lipid membrane-bound particles that are categorized by their sizes: ectosomes (100-500 nm) and exosomes (50-150 nm)1. EVs contain various biomolecules, such as proteins, nucleic acids, and lipids. These biomolecules originate from the cells before being encapsulated as cargo and released into the extracellular space via EVs1,2,3.
Due to the variety of their cargo, EVs are believed to play an active role in intercellular communication. The release and uptake of EVs by cells allow the transfer of biomolecules between the cells4,5. The introduction of EV cargo to a cell may alter the recipient cell&#39;s functions and homeostatic state4,5,6. EVs are internalized through multiple pathways; however, the exact mechanisms have not been accurately demonstrated.
The majority of the EV uptake assays, such as genetic tagging, fluorescently label individual EVs7. The resulting signal can be measured by microplate photometer, flow cytometry, or microscopy, with each technology having substantial limitations. Microplate photometers, flow cytometry, or standard two-dimensional (2D) microscopy cannot distinguish between internalized and superficially attached EVs8,9. Additionally, the necessary sample preparation for each of these techniques may introduce additional issues to EV uptake evaluation. For example, lifting adhered cells with trypsin before EV uptake analysis may cleave some superficially attached EVs on the cell&#39;s surface10,11. Trypsin may also interact with the cell surface, affecting cell and EV phenotype. Additionally, trypsin may not detach superficial EVs entirely, skewing isolated populations.
To accurately label EVs with fluorescent dyes, additional wash steps are required to remove the residual dye7. Accepted isolation techniques can also contribute to false-positive signals due to coagulation that occurs during EV isolation. For example, serial ultracentrifugation (UC) is widely used to isolate EVs and remove the immobilized dye. However, UC may co-precipitate EVs, and the residual dye may lead to a false-positive signal12,13. Other nano-filtration methods, such as column-based filtration, are also widely used for non-immobilized dye removal. The complex nature of EVs and dye interacting within the column matrix may lead to incomplete removal of residual dye due to the molecular cut-off of the column being altered by the complex input14,15,16.
The current protocol proposes a nano-filtration-based microfluidic device to isolate and wash fluorescently labeled isolated EVs. The nano-filtration-based microfluidic device can provide efficient filtration via fluid-assisted separation technology (FAST)17,18. FAST reduces the pressure drop across the filter, thus reducing potential aggregation between EVs and dyes. By efficiently removing residual dye, it is possible to enhance the quality of fluorescently labeled EVs and the assay&#39;s specificity.
Confocal microscopy can distinguish between internalized and superficially attached EVs on the cell surface and comprehensively investigate the cellular mechanisms of EV uptake in a spatiotemporal resolution19,20,21,22,23,24,25. For example, Sung et al. described the visualization of the exosome lifecycle using their developed live-cell reporter. The location of the internalized EVs was detected and analyzed using a confocal microscope in three-dimension (3D) and post-image processing tools20. Although the size of small EVs (40-200 nm) is below the resolution limit of the optical microscope, the fluorescently labeled EVs can be detected by confocal microscopy since the photodetector can detect the enhanced fluorescence emission. Therefore, the subcellular localization of the fluorescently labeled EVs within a cell can be precisely determined by acquiring multiple z-stacked images of the EVs and the surrounding cellular organelles.
Additionally, 3D reconstruction and post-data processing can provide further insight into the positioning of the internalized, superficial, and free-floating EVs. By utilizing these processes in conjunction with the time-lapse live-cell imaging offered by confocal microscopy, the level of EV uptake can be precisely evaluated, and the real-time tracking of EV uptake is also possible. Further, EV trafficking analysis can be performed using confocal microscopy by assessing the co-localization of EVs with organelles, a first step to determine how internalized EVs are involved in the intracellular function. This protocol describes the methodology for performing an EV uptake assay using the nano-filtration-based microfluidic device17,26, confocal microscopy, and post-image analysis.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Alexa Fluor 488 anti-human CD63 Antibody</td>
			<td>Biolegend</td>
			<td>353038</td>
			<td>Fluorescent dye conjugated EV-specific antibody</td>
		</tr>
		<tr>
			<td>CellTracker Orange CMTMR Dye</td>
			<td>Thermo Fisher Scientific</td>
			<td>C2927</td>
			<td>Live cell (cytoplasm) fluoresent labeling reagent</td>
		</tr>
		<tr>
			<td>CFI Apo Lambda S 40XC WI</td>
			<td>Nikon</td>
			<td>MRD77400</td>
			<td>Objective for confocal imaging, NA=1.25</td>
		</tr>
		<tr>
			<td>CFI Plan Apo VC 20X</td>
			<td>Nikon</td>
			<td>MRD70200</td>
			<td>Objective for confocal imaging, NA=0.75</td>
		</tr>
		<tr>
			<td>Exodisc</td>
			<td>Labspinner Inc.</td>
			<td>EX-D1001</td>
			<td>A nano-filtration based microfluidic device for EV isolation</td>
		</tr>
		<tr>
			<td>ExoDiscovery</td>
			<td>Labspinner Inc.</td>
			<td>EX-R1001</td>
			<td>Operation device for Exodisc</td>
		</tr>
		<tr>
			<td>Exosome-depleted FBS</td>
			<td>Thermo Fisher Scientific</td>
			<td>A2720801</td>
			<td>Nutrient of cell culture media for PC3 cell line derived EV collection</td>
		</tr>
		<tr>
			<td>Fetal bovine serum (FBS)</td>
			<td>VWR</td>
			<td>1500-500</td>
			<td>Nutrient for cell cultivation</td>
		</tr>
		<tr>
			<td>Goat Anti-Mouse IgG H&#38;L preadsorbed</td>
			<td>abcam</td>
			<td>&#160;ab7063</td>
			<td>Mouse IgG antibody for negatvie control of EV labeling</td>
		</tr>
		<tr>
			<td>Ibidi USA&#160;U DISH &#956;-Dish 35 mm</td>
			<td>Ibidi</td>
			<td>81156</td>
			<td>Culture dish for confocal imaging</td>
		</tr>
		<tr>
			<td>Imaris 9.7.1</td>
			<td>Oxford Instruments</td>
			<td>9.7.1</td>
			<td>Post-image processing software</td>
		</tr>
		<tr>
			<td>Incubator System+ CO2/O2/N2 gas mixer</td>
			<td>Live Cell Instrument</td>
			<td>TU-O-20</td>
			<td>Incubator system for live cell imaging</td>
		</tr>
		<tr>
			<td>Nikon A1 HD25 / A1R HD25 camera</td>
			<td>Nikon</td>
			<td>NA</td>
			<td>Camera for confocal imaging</td>
		</tr>
		<tr>
			<td>Nikon Eclipse Ti microscope</td>
			<td>Nikon</td>
			<td>NA</td>
			<td>Inverted microscope for confocal imaging</td>
		</tr>
		<tr>
			<td>NIS-Elements AR 4.50.00</td>
			<td>Nikon</td>
			<td>4.50.00</td>
			<td>Image processing software for Nikon microscope</td>
		</tr>
		<tr>
			<td>NTA, NanoSight NS500</td>
			<td>Malvern Panalytical</td>
			<td>NS500</td>
			<td>Measurement device for EV concentration</td>
		</tr>
		<tr>
			<td>OriginPro 2020</td>
			<td>OriginLab</td>
			<td>9.7.0.185</td>
			<td>Graphing software</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Thermo Fisher Scientific</td>
			<td>15140122</td>
			<td>Antibiotics for cell cultivation</td>
		</tr>
		<tr>
			<td>RPMI 1640</td>
			<td>Thermo Fisher Scientific</td>
			<td>21875034</td>
			<td>Cell culture media for PC3 cell line cultivation</td>
		</tr>
		<tr>
			<td>SYTO RNASelect Green Fluorescent cell Stain - 5 mM Solution in DMSO</td>
			<td>Thermo Fisher Scientific</td>
			<td>S32703</td>
			<td>RNA staining fluorescent dye for the EV labeling</td>
		</tr>
	</tbody>
</table>

extracellular vesicle uptake assay via confocal microscope imaging analysis

There is a need for practical assays to visualize and quantify the cells' extracellular vesicle (EV) uptake. EV uptake plays a role in intercellular communication in various research fields; cancer biology, neuroscience, and drug delivery. Many EV uptake assays have been reported in the literature; however, there is a lack of practical, detailed experimental methodology. EV uptake can be assessed by fluorescently labeling EVs to detect their location within cells. Distinguishing between internalized EVs in cells and the superficial EVs on cells is difficult, yet critical, to accurately determine the EV uptake. Therefore, an assay that efficiently quantifies EV uptake through three-dimensional (3D) fluorescence confocal microscopy is proposed in this work. Fluorescently labeled EVs were prepared using a nano-filtration-based microfluidic device, visualized by 3D confocal microscopy, and then analyzed through advanced image-processing software. The protocol provides a robust methodology for analyzing EVs on a cellular level and a practical approach for efficient analysis.

Using a nano-filtration-based microfluidic device, EVs were isolated from PC3 CCM and labeled with a fluorophore-conjugated EV-specific (CD63) antibody (Figure 1). The labeled EVs were successfully visualized by the 3D confocal microscopy (Figure 2). The labeled EVs were incubated with cells for several hours in exosome-depleted media. Following incubation, cells were washed with exosome depleted media. The remaining EVs were internalized or adhered to cells dur...

Watch this Scientific Journal Video about Extracellular Vesicle Uptake Assay via Confocal Microscope Imaging Analysis at JoVE.com

Extracellular Vesicle Uptake Assay via Confocal Microscope Imaging Analysis

Extracellular vesicles (EVs) contribute to cellular biology and intercellular communications. There is a need for practical assays to visualize and quantify EVs uptake by the cells. The current protocol proposes the EV uptake assay by utilizing three-dimensional fluorescence imaging via confocal microscopy, following EV isolation by a nano-filtration-based microfluidic device.

Extracellular Vesicle Uptake Assay via Confocal Microscope Imaging Analysis

There is a need for practical assays to visualize and quantify the cells' extracellular vesicle (EV) uptake. EV uptake plays a role in intercellular ...

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7.3K Views.  Johns Hopkins University School of Medicine. Extracellular vesicles (EVs) contribute to cellular biology and intercellular communications. There is a need for practical assays to visualize and quantify EVs uptake by the cells. The current protocol proposes the EV uptake assay by utilizing three-dimensional fluorescence imaging via confocal microscopy, following EV isolation by a nano-filtration-based microfluidic device. 

Video: Extracellular Vesicle Uptake Assay via Confocal Microscope Imaging Analysis

Proper Care and Cleaning of the Microscope

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a ...

Major Components of the Light Microscope

The light microscope is a basic tool for the cell biologist, who should have a thorough understanding of how it works, how it should be aligned for different applications, and how it should be maintained as required to obtain maximum image-forming capacity and resolution. The components of the microscope are described in detail here.

The light microscope is a basic tool for the cell biologist, who should have a thorough understanding of how it works, how it should be aligned for different applications, and how it should be maintained as required to obtain maximum image-forming capacity and resolution. The components of the microscope are described in detail here.

The light microscope is a basic tool for the cell biologist, who should have a thorough understanding of how it works, how it should be aligned for ...

A Neuronal and Astrocyte Co-Culture Assay for High Content Analysis of Neurotoxicity

High Content Analysis (HCA) assays combine cells and detection reagents with automated imaging and powerful image analysis algorithms, allowing measurement of multiple cellular phenotypes within a single assay. In this study, we utilized HCA to develop a novel assay for neurotoxicity. Neurotoxicity assessment represents an important part of drug safety evaluation, as well as being a significant focus of environmental protection efforts. Additionally, neurotoxicity is also a well-accepted in vitro marker of the development of neurodegenerative diseases such as Alzheimer's and Parkinson's diseases. Recently, the application of HCA to neuronal screening has been reported. By labeling neuronal cells with &beta;III-tubulin, HCA assays can provide high-throughput, non-subjective, quantitative measurements of parameters such as neuronal number, neurite count and neurite length, all of which can indicate neurotoxic effects. However, the role of astrocytes remains unexplored in these models. Astrocytes have an integral role in the maintenance of central nervous system (CNS) homeostasis, and are associated with both neuroprotection and neurodegradation when they are activated in response to toxic substances or disease states. GFAP is an intermediate filament protein expressed predominantly in the astrocytes of the CNS. Astrocytic activation (gliosis) leads to the upregulation of GFAP, commonly accompanied by astrocyte proliferation and hypertrophy. This process of reactive gliosis has been proposed as an early marker of damage to the nervous system. The traditional method for GFAP quantitation is by immunoassay. This approach is limited by an inability to provide information on cellular localization, morphology and cell number. We determined that HCA could be used to overcome these limitations and to simultaneously measure multiple features associated with gliosis - changes in GFAP expression, astrocyte hypertrophy, and astrocyte proliferation - within a single assay. In co-culture studies, astrocytes have been shown to protect neurons against several types of toxic insult and to critically influence neuronal survival. Recent studies have suggested that the use of astrocytes in an in vitro neurotoxicity test system may prove more relevant to human CNS structure and function than neuronal cells alone. Accordingly, we have developed an HCA assay for co-culture of neurons and astrocytes, comprised of protocols and validated, target-specific detection reagents for profiling &beta;III-tubulin and glial fibrillary acidic protein (GFAP). This assay enables simultaneous analysis of neurotoxicity, neurite outgrowth, gliosis, neuronal and astrocytic morphology and neuronal and astrocytic development in a wide variety of cellular models, representing a novel, non-subjective, high-throughput assay for neurotoxicity assessment. The assay holds great potential for enhanced detection of neurotoxicity and improved productivity in neuroscience research and drug discovery.

This article describes a novel protocol and reagent set designed for sensitive measurement of neurotoxic effects of compounds and treatments on co-cultures of neurons and astrocytes using high content analysis. Results demonstrate that high content analysis represents an exciting novel technology for neurotoxicity assessment.

High Content Analysis (HCA) assays combine cells and detection reagents with automated imaging and powerful image analysis algorithms, allowing ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software

Cellular processes, such as chromosome assembly, segregation and cytokinesis,are inherently dynamic. Time-lapse imaging of living cells, using fluorescent-labeled reporter proteins or differential interference contrast (DIC) microscopy, allows for the examination of the temporal progression of these dynamic events which is otherwise inferred from analysis of fixed samples1,2. Moreover, the study of the developmental regulations of cellular processes necessitates conducting time-lapse experiments on an intact organism during development. The Caenorhabiditis elegans embryo is light-transparent and has a rapid, invariant developmental program with a known cell lineage3, thus providing an ideal experiment model for studying questions in cell biology4,5and development6-9. C. elegans is amendable to genetic manipulation by forward genetics (based on random mutagenesis10,11) and reverse genetics to target specific genes (based on RNAi-mediated interference and targeted mutagenesis12-15). In addition, transgenic animals can be readily created to express fluorescently tagged proteins or reporters16,17. These traits combine to make it easy to identify the genetic pathways regulating fundamental cellular and developmental processes in vivo18-21. In this protocol we present methods for live imaging of C. elegans embryos using DIC optics or GFP fluorescence on a compound epifluorescent microscope. We demonstrate the ease with which readily available microscopes, typically used for fixed sample imaging, can also be applied for time-lapse analysis using open-source software to automate the imaging process.

Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software

The C. elegans embryo is a powerful system for studying cell biology and development. We present a protocol for live imaging of C. elegans embryos utilizing DIC optics or fluorescence using readily available epifluorescent microscopes and open-source software.

Cellular processes, such as chromosome assembly, segregation and cytokinesis,are inherently dynamic. Time-lapse imaging of living cells, using ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

LDL Cholesterol Uptake Assay Using Live Cell Imaging Analysis with Cell Health Monitoring

The regulation of LDL cholesterol uptake through LDLR-mediated endocytosis is an important area of study in various major pathologies including metabolic disorder, cardiovascular disease, and kidney disease. Currently, there is no available method to assess LDL uptake while simultaneously monitoring for health of the cells. The current study presents a protocol, using a live cell imaging analysis system, to acquire serial measurements of LDL influx with concurrent monitoring for cell health. This novel technique is tested in three human cell lines (hepatic, renal tubular epithelial, and coronary artery endothelial cells) over a four-hour time course. Moreover, the sensitivity of this technique is validated with well-known LDL uptake inhibitors, Dynasore and recombinant PCSK9 protein, as well as by an LDL uptake promoter, Simvastatin. Taken together, this method provides a medium-to-high throughput platform for simultaneously screening pharmacological activity as well as monitoring of cell morphology, hence cytotoxicity of compounds regulating LDL influx. The analysis can be used with different imaging systems and analytical software.

This protocol provides an efficient approach to measuring LDL cholesterol uptake with real time influx rates using a live cell imaging system in various cell types. This technique provides a platform to screen the pharmacological activity of compounds affecting LDL influx while monitoring for cell morphology and hence potential cytotoxicity.

The regulation of LDL cholesterol uptake through LDLR-mediated endocytosis is an important area of study in various major pathologies including metabolic ...

Purification and Analysis of Caenorhabditis elegans Extracellular Vesicles

The secretion of small membrane-bound vesicles into the external environment is a fundamental physiological process of all cells. These extracellular vesicles (EVs) function outside the cell to regulate global physiological processes by transferring proteins, nucleic acids, metabolites, and lipids between tissues. EVs reflect the physiological state of their cells of origin. EVs are implicated to have fundamental roles in virtually every aspect of human health. Thus, EV protein and genetic cargos are being increasingly analyzed for biomarkers of health and disease. However, the EV field still lacks a tractable invertebrate model system that permits the study of EV cargo composition. C. elegans is well suited for EV research because it actively secretes EVs outside of its body into its external environment, permitting facile isolation. This article provides all the necessary information for generating, purifying, and quantifying these environmentally secreted C. elegans EVs including how to work quantitatively with very large populations of age-synchronized worms, purifying EVs, and a flow cytometry protocol that directly measures the number of intact EVs in the purified sample. Thus, the large library of genetic reagents available for C. elegans research can be tapped into for investigating the impacts of genetic pathways and physiological processes on EV cargo composition.

Purification and Analysis of Caenorhabditis elegans Extracellular Vesicles

This article presents methods for generating, purifying, and quantifying Caenorhabditis elegans extracellular vesicles.

The secretion of small membrane-bound vesicles into the external environment is a fundamental physiological process of all cells. These extracellular ...

Single Extracellular Vesicle Transmembrane Protein Characterization by Nano-Flow Cytometry

Single particle characterization has become increasingly relevant for research into extracellular vesicles, progressing from bulk analysis techniques and first-generation particle analysis to comprehensive multi-parameter measurements such as nano-flow cytometry (nFCM). nFCM is a form of flow cytometry that utilizes instrumentation specifically designed for nano-particle analysis, allowing for thousands of EVs to be characterized per minute both with and without the use of staining techniques. High resolution side scatter (SS) detection allows for size and concentration to be determined for all biological particles larger than 45 nm, while simultaneous fluorescence (FL) detection identifies the presence of labeled markers and targets of interest. Labeled subpopulations can then be described in quantitative units of particles/mL or as a percentage of the total particles identified by side scatter.
Here, EVs derived from conditioned cell culture media (CCM) are labeled with both a lipid dye, to identify particles with a membrane, and antibodies specific for CD9, CD63, and CD81 as common EV markers. Measurements of comparison material, a concentration standard and a size standard of silica nanospheres, as well as labeled sample material are analyzed in a 1-minute analysis. The software is then used to measure the concentration and size distribution profile of all particles, independent of labeling, before determining the particles that are positive for each of the labels.
Simultaneous SS and FL detection can be utilized flexibly with many different EV sources and labeling targets, both external and internal, describing EV samples in a comprehensive and quantitative manner.

The latest generation of EV characterization tools are capable of single EV analysis across multiple parameters simultaneously. Nano-flow cytometry measures all biological particles larger than 45 nm without labeling and identifies specific characteristics of subpopulations by a variety of fluorescent labeling techniques.

Single particle characterization has become increasingly relevant for research into extracellular vesicles, progressing from bulk analysis techniques and ...

Analysis of Extracellular Vesicle-Mediated Vascular Calcification Using In Vitro and In Vivo Models

Cardiovascular disease is the leading cause of death in the world, and vascular calcification is the most significant predictor of cardiovascular events; however, there are currently no treatment or therapeutic options for vascular calcification. Calcification begins within specialized extracellular vesicles (EVs), which serve as nucleating foci by aggregating calcium and phosphate ions. This protocol describes methods for obtaining and assessing calcification in murine aortas and analyzing the associated extracted EVs. First, gross dissection of the mouse is performed to collect any relevant organs, such as the kidneys, liver, and lungs. Then, the murine aorta is isolated and excised from the aortic root to the femoral artery. Two to three aortas are then pooled and incubated in a digestive solution before undergoing ultracentrifugation to isolate the EVs of interest. Next, the mineralization potential of the EVs is determined through incubation in a high-phosphate solution and measuring the light absorbance at a wavelength of 340 nm. Finally, collagen hydrogels are used to observe the calcified mineral formation and maturation produced by the EVs in vitro.

Analysis of Extracellular Vesicle-Mediated Vascular Calcification Using In Vitro and In Vivo Models

This paper presents the methodology for obtaining and assessing vascular calcification by isolating murine aortas followed by extracting calcified extracellular vesicles to observe the mineralization potential.

Cardiovascular disease is the leading cause of death in the world, and vascular calcification is the most significant predictor of cardiovascular events; ...