The authors would like to thank Dale Hirschkorn from Blood Systems Research Institute for his help with flow cytometer instrument settings. This work was supported by NIH grants HL095470 and U01 HL072268 and DoD contracts W81XWH-10-1-0023 and W81XWH-2-0028.

NOTE: The following protocols have been performed in compliance with all institutional, national and international guidelines for human welfare. All human subject samples were tested under an institutional review board (IRB)-approved protocol and with informed consent of the subjects.
1. METHOD A: Individual Detection Method
1.1) Processing of Blood Sample/Isolation of EVs
<ol>
	<li>Draw blood from donor/patient into two 10 ml glass tubes containing 1.5 ml of ACD-Solution A or other suitable anticoagulant and process immediately (within 30 min max) using the following 2-step differential centrifugation protocol. 
	NOTE: This protocol will yield approximately 10 ml of platelet poor plasma (PPP) from the combined ~17 ml of blood drawn. If more or less PPP is needed, the number of tubes of blood collected may be adjusted accordingly.</li>
	<li>Centrifuge the samples at 1,500 x g for 10 min at&#160;RT&#160;to separate the plasma from the buffy coat and red cells. Transfer 1.2 ml aliquots of the plasma supernatant to 1.5 ml centrifuge tubes, being careful not to disturb the bottom layers containing the buffy coat and red cells.</li>
	<li>Spin at 13,000 x g for 10 min at RT to remove platelets and large cell fragments. Carefully transfer the PPP, leaving behind 200 &#181;l to avoid disturbing the pellet and add the PPP to a new tube.</li>
	<li>At this point, use PPP immediately for analysis or transfer in 1.0 ml aliquots to new 1.5 ml centrifuge tubes and store at -80 &#176;C for up to two years for later analysis (refer to Figure 1A for overview).</li>
	<li>If purified EVs are needed for functional experiments, transfer 6 ml of the PPP to an ultracentrifuge tube and add 28 ml of 0.2 &#181;m-filtered phosphate buffered saline (PBS). Spin for 60 min at 100,000 x g at RT using an ultracentrifuge equipped with a swinging bucket rotor. Aspirate supernatant and resuspend EV pellet in 1.5 ml media. 
	NOTE: For highest reproducibility, blood samples should be processed as consistently as possible from donor to donor. Any variation in EV isolation method could significantly impact the number and type of EVs detected.</li>
</ol>
1.2) Preparing Samples for Analysis
NOTE: From this point on, the steps explain a high throughput protocol for analyzing 12 samples for 14 markers in 3 panels. However, other combinations of antibodies can be used here; the protocol can be adapted to study other EV populations by substituting the suggested markers for those of interest.
<ol>
	<li>Remove 12 samples from freezer (if stored at -80 &#176;C) and thaw at 37 &#176;C.</li>
	<li>Pipette contents up and down several times to mix. Remove 320 &#181;l from each sample and add to the top row of a 96 well plate. 
	NOTE: A width-adjustable multi-well pipet is extremely helpful for this and many other steps throughout the assay, particularly when analyzing multiple samples at once.</li>
</ol>
1.3) Staining EV Samples
<ol>
	<li>Prior to staining, filter all antibodies (Abs) to remove aggregates, which can cause positive signals.
	<ol>
		<li>Combine antibodies to be used in each of the 3 panels into separate 0.22 &#181;m centrifugal filter tubes and centrifuge using a fixed angle single speed centrifuge (~750 x g) at RT for 2 min, or until all of the Ab mixture has passed through the filter and no antibody liquid remains on the surface of the filter. Store Ab cocktails in the fridge for up to two weeks but re-filter each time before use.</li>
	</ol>
	</li>
	<li>Add the appropriate amount of filtered Ab mixture to each well in row 2 (e.g., samples in Panel I are stained with 2 &#181;l of each Ab, so a total of 12 &#181;l of the filtered Ab cocktail is added per well to row 2). Refer to Figure 1B for an outline of the suggested plate map. Repeat these additions to the rows beneath if more panels are run (here, add 8 &#181;l/well of the Panel II cocktail to row 3 and 11 &#181;l/well of the Panel III cocktail to row 4; refer to Materials List for specific panel information).</li>
	<li>Using the multichannel pipet, mix the PPP samples in row 1 up and down and transfer 100&#160;&#181;l from the wells in row 1 to the wells in row 2. Mix up and down. Change tips and repeat, transferring 100 &#181;l from row 1 to rows 3 and 4. Incubate at 4 &#176;C for 30 min.</li>
</ol>
1.4) Washing MV Samples
<ol>
	<li>Remove the 96-well plate from 4 &#176;C and transfer to biological safety cabinet. Using a multichannel pipette, add 220 &#181;l of PBS/well to rows 6-8 (to be used for rinsing/washing the wells containing stained PPP).</li>
	<li>Transfer the contents of each well to pre-labeled centrifugal filter tubes using the width-adjustable multichannel pipet (For 12 samples, with 3 panels of antibodies, 12 x 3 = 36 filter tubes will be needed). Using the same tips, remove 200 &#181;l of PBS from the wash rows and add to the corresponding wells from which PPP was just removed.</li>
	<li>Mix up and down to rinse the wells and transfer the rinse solution to the same filters to which the PPP was previously added. Close tops of centrifugal filters. Change tips.</li>
	<li>Repeat this process with the remaining stained samples until all stained PPP samples have been transferred along with their rinse solutions to centrifugal filters.</li>
	<li>Transfer the centrifugal filters to a fixed rotor centrifuge and spin at 850 x g for 3 min at RT. 
	NOTE: Ensure that no liquid remains on the filter tops. After centrifugation, the filter should appear to be &#8220;dry&#8221; with no visible fluid layer remaining on top. While unlikely, certain PPP samples may require a longer centrifugation time to effectively move through the filter.</li>
	<li>Remove the centrifugal filter tubes and return to the biological safety cabinet. Using the multichannel pipet, resuspend the tops of the filters in 300 &#181;l of PBS. Transfer the resuspended contents to pre-labeled tubes for immediate FCM analysis. 
	NOTE: It is very important to keep the force and number of pipette plunger depressions consistent among samples to avoid sample-to-sample variation. This should ideally be done using an electronic pipet that has been programmed to pipet up and down a specific volume (e.g., 280 &#181;l) an exact number of times (e.g., 8 times) for each sample.</li>
</ol>
1.5) Cytometer Setup
<ol>
	<li>Open the FCM software. Prior to experiment setup, perform daily instrument calibration and setup using instrument setup beads (following manufacturer&#8217;s instructions).</li>
	<li>If EV samples have been stained with more than one antibody and multiple fluorochromes are to be measured at once, calculate compensation values as follows:
	<ol>
		<li>Add 2 drops of compensation beads to pre-labeled tubes (1 tube for each fluorochrome-conjugated antibody) and add the recommended amount of antibody. Add 2 drops of negative compensation beads to another tube to use as the unstained compensation control.</li>
		<li>Incubate at 4 &#176;C for 30 min, wash with PBS and resuspend in 400 &#181;l of PBS.</li>
		<li>Using the flow cytometer software included with the instrument, run each compensation tube and adjust fluorescent voltages to place each peak at approximately 104 on a 5-decade log scale. Ensure that the fluorescence peak is highest (brightest) in its own fluorescent channel compared to all other channels, and adjust voltages of fluorescent parameters again if necessary. Run each individually-stained comp tube and capture at least 5,000 events per tube.</li>
		<li>Select the tab &#8220;Experiment,&#8221; then select &#8220;Compensation,&#8221; then select &#8220;Calculate Compensation&#8221; to apply compensation values to all samples.</li>
	</ol>
	</li>
	<li>Set the forward scatter (FSC) and side scatter (SSC) voltage parameters to log scale and select the lowest thresholds allowed by the cytometer (FSC=200 and SSC=200) for each.</li>
	<li>While running a tube of 0.22 &#181;m-filtered PBS, adjust the FSC &#38; SSC voltages to the highest values that exclude the majority of background noise (i.e., just below the voltage threshold at which event rate surpasses 5 events/sec).</li>
	<li>Next, run a tube containing 0.2 &#181;m &#8211; 1.0 &#181;m beads, diluted in PBS if necessary. In an FCS vs. SSC plot, draw a gate around the bead population to capture events between 0.2 &#181;m and 1.0 &#181;m. Alternatively, in an SSC-H histogram, draw a gate to include all events smaller than the 1.0 &#181;m beads.</li>
	<li>Set the cytometer&#8217;s flow rate to &#8220;Lo&#8221; (approximately 8-12 &#181;l/min). Using the beads tube (or other tube containing a known concentration of beads) adjust the flow rate dial on the cytometer until the event rate reaches approximately 200 events/sec. Read all sample tubes at the same flow rate and use the same bead concentration in future experiments to ensure that flow rates remain consistent between runs.</li>
	<li>Run a tube of rainbow fluorescent particles diluted in PBS. Acquire 5,000 events. Record the mean intensity values for FSC, SSC, and each color channel. Use these values to adjust voltages in future experiments to ensure that fluorescence intensities remain consistent between experiments.</li>
</ol>
1.6) Sample Reading
<ol>
	<li>Set the cytometer&#8217;s flow rate to &#8220;Lo&#8221; (approximately 8-12 &#181;l/min) and run each sample for exactly 1 or 2 min.</li>
	<li>After the first reading, add 20 &#181;l of 10% NP-40 to each sample, pipette up and down, and re-read for the same amount of time (either 1 or 2 min) to allow for the subtraction of positive events detected in the lysed sample over an equal time frame. 
	NOTE: It is extremely important that samples be mixed using a pipet rather than a vortex. In our experience, vortexing can cause self-aggregation of some antibodies, leading to EV-mimicking positive events.</li>
	<li>Once all samples have been read, export all .fcs files into a separate file to be used for further analysis using FCM analysis software.</li>
</ol>
1.7) Data Analysis
<ol>
	<li>Open the FCM analysis software. Import all of the .fcs files into a new experiment file.</li>
	<li>Open the beads-only tube. In the FSC-A vs. SSC-A plot, draw a gate around all beads sized between 0.2 &#181;m and 1.0 &#181;m. This is the EV gate. Drag to add to all samples.</li>
	<li>Using the lysed samples as negative controls, draw gates at the edge of background fluorescence in each fluorescence channel used. Drag fluorescent gates into the EV gate of each corresponding non-lysed sample. 
	NOTE: At this point it is also useful to examine dual fluorescent bi-parameter plots. Rocket shapes in the double-positive quadrant (see Figure 8), particularly if the markers are known to reside on unrelated cell types, may be indicative of artifact from aggregation or other vesicle-mimicking events.</li>
	<li>For each fluorescent marker, subtract the number of events in the lysed sample from the number of events in the non-lysed sample. Optionally, divide this number by the total number of EVs within the non-lysed EV gate to get % positive values.</li>
</ol>
2. METHOD B: Beads Method
2.1) Processing of Blood Sample/Isolation of EVs
<ol>
	<li>Refer to the blood processing method described in Method A (Section 1.1).</li>
</ol>
2.2) Preparing Samples for Analysis
<ol>
	<li>If desired, fractionate PPP or ultracentrifuged EVs into exosomes and microvesicles. Add 250 &#181;l of PPP or ultracentrifuged EVs to 0.22 &#956;m centrifugal filters and transfer to a fixed rotor centrifuge and spin at 750 x g for 2 min at RT. 
	NOTE: Ensure that no liquid remains on the filter tops. After centrifugation, the filter should appear to be &#8220;dry&#8221; with no visible fluid layer remaining on top. While unlikely, certain samples may require a slightly longer centrifugation time for the fluid to effectively move through the filter.</li>
	<li>Wash uncoated 6 &#181;m polystyrene beads (e.g., negative AbC beads) 2x with RPMI media, and resuspend in 2 ml. Add 6,000 beads to each FACS tube. To the negative control tube, add 400 &#956;l of RPMI media alone to the beads. To all other tubes, add 200 &#956;l of PPP or ultracentrifuged EVs (or their fractions) and 200 &#956;l of RPMI media.</li>
	<li>Adjust the final volume of all tubes to 400 &#956;l with media and incubate overnight at 4 &#176;C on a shaker.</li>
	<li>The next morning, wash beads with 2 ml of media. Aspirate off the supernatant.</li>
	<li>Block with 5% bovine serum albumin (BSA) in media (400 &#956;l) for 3 hr at 4 &#176;C on a shaker.</li>
	<li>Wash beads with 2 ml of media. Aspirate and resuspend pellet in 100 &#956;l of medium.</li>
</ol>
2.3) Staining EV Samples
<ol>
	<li>Filter all antibodies. Use the same antibody panels used in Method A, or if desired, create a different combination of antibodies as long as their fluorochromes are compatible with one another.</li>
	<li>Combine all antibodies to be used in a single panel into a 0.22 &#181;m centrifugal filter tube. Centrifuge using a fixed angle single speed centrifuge for 2 min or until all of the Ab mixture has passed through the filter and no antibody liquid remains on the surface of the filter.</li>
	<li>Add appropriate volume of filtered antibody cocktail to all tubes and incubate for 30 min at 4 &#176;C.</li>
	<li>Wash beads with 2 ml of media, resuspend in 400 &#956;l of media and run immediately (or within the same day) on flow cytometer.</li>
</ol>
2.4) Cytometer Setup and Sample Reading
<ol>
	<li>Open the FCM software. Prior to experiment setup, perform daily instrument calibration and setup using instrument setup beads (following manufacturer&#8217;s instructions).</li>
	<li>If EV samples have been stained with more than one antibody and multiple fluorochromes are to be measured at once, calculate compensation values as follows:
	<ol>
		<li>Add 2 drops of compensation beads to pre-labeled tubes (1 tube for each fluorochrome-conjugated antibody) and add the recommended amount of antibody. Add 2 drops of negative compensation beads to another tube to use as the unstained compensation control. Incubate at 4 &#176;C for 30 min, wash with PBS and resuspend in 400 &#181;l of PBS.</li>
		<li>Using the FCM software, run each compensation tube and adjust fluorescent voltages to place each peak at approximately 104 on a 5-decade log scale. Ensure that the fluorescence peak is highest (brightest) in its own fluorescent channel compared to all other channels, and adjust voltages of fluorescent parameters again if necessary. Run each individually-stained comp tube and capture at least 5,000 events per tube.</li>
		<li>Select the tab &#8220;Experiment,&#8221; then &#8220;Compensation,&#8221; then &#8220;Calculate Compensation&#8221; to apply compensation values to all samples.</li>
	</ol>
	</li>
	<li>Change the FSC and SSC voltage parameters to log scale and select the lowest thresholds allowed by the cytometer (FSC = 200 and SSC = 200) for each.</li>
	<li>Run samples, gate on the singlet beads population and acquire 2,000 events in this gate. Export .fcs files.</li>
	<li>Use FCM analysis software to analyze .fcs files. Gate on singlet beads. Calculate the geometric mean fluorescent intensity (MFI) for each fluorochrome and compare with the MFI of the negative control.</li>
</ol>

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G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improved+Flow+Cytometric+Assessment+Reveals+Distinct+Microvesicle+(Cell-Derived+Microparticle)+Signatures+in+Joint+Diseases.">Improved Flow Cytometric Assessment Reveals Distinct Microvesicle (Cell-Derived Microparticle) Signatures in Joint Diseases.</a> PLoS ONE. 7 (11), e49726 (2012).</li><li>Gy&#246;rgy, B., M&#243;dos, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+and+isolation+of+cell-derived+microparticles+are+compromised+by+protein+complexes+resulting+from+shared+biophysical+parameters.">Detection and isolation of cell-derived microparticles are compromised by protein complexes resulting from shared biophysical parameters.</a> Blood. 117 (4), e39-e48 (2011).</li><li>Gy&#246;rgy, B., Pasztoi, M., Buzas, E. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Response:+systematic+use+of+Triton+lysis+as+a+control+for+microvesicle+labeling.">Response: systematic use of Triton lysis as a control for microvesicle labeling.</a> Blood. 119 (9), 2175-2176 (2012).</li><li>Trummer, A., De Rop, C., Tiede, A., Ganser, A., Eisert, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isotype+controls+in+phenotyping+and+quantification+of+microparticles:+a+major+source+of+error+and+how+to+evade+it.">Isotype controls in phenotyping and quantification of microparticles: a major source of error and how to evade it.</a> Thrombosis Research. 122 (5), 691-700 (2008).</li><li>Shet, A. S., Aras, O., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sickle+blood+contains+tissue+factor-positive+microparticles+derived+from+endothelial+cells+and+monocytes.">Sickle blood contains tissue factor-positive microparticles derived from endothelial cells and monocytes.</a> Blood. 102 (7), 2678-2683 (2003).</li><li>Connor, D. E., Exner, T., Ma, D. D. F., Joseph, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+majority+of+circulating+platelet-derived+microparticles+fail+to+bind+annexin+V,+lack+phospholipid-dependent+procoagulant+activity+and+demonstrate+greater+expression+of+glycoprotein+Ib.">The majority of circulating platelet-derived microparticles fail to bind annexin V, lack phospholipid-dependent procoagulant activity and demonstrate greater expression of glycoprotein Ib.</a> Thrombosis and Haemostasis. 103 (5), 1044-1052 (2010).</li><li>Nolte-&#8217;t Hoen, E. N. M., van der Vlist, E. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+and+qualitative+flow+cytometric+analysis+of+nanosized+cell-derived+membrane+vesicles.">Quantitative and qualitative flow cytometric analysis of nanosized cell-derived membrane vesicles.</a> Nanomedicine: Nanotechnology, Biology, And Medicine. 8 (5), 712-720 (2012).</li><li>Van der Vlist, E. J., Nolte-&#8217;t Hoen, E. N. M., Stoorvogel, W., Arkesteijn, G. J. A., Wauben, M. H. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+labeling+of+nano-sized+vesicles+released+by+cells+and+subsequent+quantitative+and+qualitative+analysis+by+high-resolution+flow+cytometry.">Fluorescent labeling of nano-sized vesicles released by cells and subsequent quantitative and qualitative analysis by high-resolution flow cytometry.</a> Nature Protocols. 7 (7), 1311-1326 (2012).</li><li>Orozco, A. F., Lewis, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flow+cytometric+analysis+of+circulating+microparticles+in+plasma.">Flow cytometric analysis of circulating microparticles in plasma.</a> Cytometry Part A. 77 A. 77A (6), 502-514 (2010).</li></ol>

The authors have no conflict of interest to disclose.

Two different protocols for the isolation, treatment and analysis of EVs were presented, using either an individual detection or bead-based approach. Selecting the most appropriate method to use is not always straightforward and requires an understanding of the sample being tested as well as the individual subpopulations of interest. Furthermore, the sensitivity of the cytometer used for acquisition must be considered when choosing the most appropriate method. Oftentimes there is no single best protocol to use, rather, a...

Extracellular Vesicles (EVs) are small, membrane-derived vesicles found in bodily fluids that are highly involved in cell-cell communication and help regulate a diverse range of biological processes. Analysis of EVs using flow cytometry (FCM) has been notoriously difficult due to their small size and lack of discrete populations positive for markers of interest. Methods for EV analysis, while considerably improved over the last decade, are still a work in progress. Unfortunately, there is no one-size-fits-all protocol, and several aspects must be considered when determining the most appropriate method to use. Presented here are several different techniques for processing EVs and two protocols for analyzing EVs using either individual detection or a bead-based approach. The methods described here will assist with eliminating the antibody aggregates commonly found in commercial preparations, increasing signal&#8211;to-noise ratio, and setting gates in a rational fashion that minimizes detection of background fluorescence. The first protocol uses an individual detection method that is especially well suited for analyzing a high volume of clinical samples, while the second protocol uses a bead-based approach to capture and detect smaller EVs and exosomes. 
EVs, also known as microparticles, are small, membrane-derived vesicles found in bodily fluids that are involved in cell-cell communication and help regulate a diverse range of biological processes1. Through expression of various surface markers and/or direct transfer of biological material, EVs are able to alter the function of recipient cells to play either activating or suppressing roles in intercellular communication2&#8211;4. Clinically, platelet-derived EVs are known to have strong anticoagulant activity5, while others have been shown to contribute to a wide range of conditions, from promoting tumor metastasis6&#160;to protecting against disease7. EVs can be classified into smaller categories of cell-derived vesicles such as exosomes and microvesicles (MVs), depending on their size and mechanism of generation8. The nomenclature of cell-derived vesicle subpopulations continues to be a topic of ongoing debate8,9, however, exosomes are generally described as small, 40 to 100 nm particles derived from endosomal fusion with the plasma membrane, while MVs are larger 100 to 1,000 nm particles formed by shedding of the plasma membrane10. Here, the general term &#8220;EVs&#8221; will be used to refer to all types of extracellular biological vesicles released by cells.
Isolation of EVs from whole blood is a multi-step procedure and many different processing variables have been shown to affect EV content, including storage temperature and duration11,12, anticoagulant/preservative used13&#160;and centrifugation method used14. A need for standardization of these variables has led to recommendations by the International Society on Thrombosis and Haemostasis Scientific and Standardization Committee (ISTH SSC) for proper blood processing and EV isolation procedures15,16, yet there exists no consensus among researchers on the optimal protocol to use 12. Most agree, however, that tightly controlled pre-analytical variables are crucial for accurate and reproducible data.
In order to analyze EVs, researchers have utilized various methods, including transmission electron microscopy17, scanning electron microscopy18,19, atomic force microscopy, dynamic light scattering20,21&#160;and western blotting22,23. While FCM is the method of choice for many researchers9,24&#8211;26&#160;due to its high throughput capabilities, analysis of EVs using FCM has been notoriously difficult due to their size and lack of discrete positive populations27&#8211;32. Compared to analysis of cells, the small size of the EVs results in 1) less fluorescence emitted due to the fewer number of antigens per particle and 2) limited feasibility of post-stain washing, which is necessary to reduce background fluorescence. Common challenges among researchers include signals arising from immunoglobulin aggregates27,28and self-aggregation of antibodies29. Furthermore, the long processing times and lengthy washing/isolation procedures used by many of the current protocols33,34&#160;require multi-day time commitments to analyze a small number of samples, making them less than ideal for high throughput applications. Some researchers forgo a wash step altogether, rendering traditionally used FCM negative controls such as fluorescence minus one (FMO) and antibody isotypes useless for accurately assessing background fluorescence30.
Our protocols address three common problems that can impede proper FCM analysis of EVs: signals arising from antibody aggregates and other non-vesicles, difficulty in removing unbound antibody, and lack of discernible positive populations. The techniques described here will assist with eliminating the antibody aggregates commonly found in commercial preparations, increasing signal&#8211;to-noise ratio, and setting gates in a rational fashion that minimizes detection of background fluorescence. Two different detection methods are presented here: the first protocol uses an individual detection method that is especially well suited for analyzing a high volume of clinical samples, while the second protocol uses a beads-based approach to capture and detect smaller EVs and exosomes.

<table border="0" cellpadding="0" cellspacing="0" >
	<tbody>
		<tr >
			<td >LSR II benchtop flow cytometer</td>
			<td >BD Biosciences</td>
			<td ></td>
			<td >3-laser (20 mW Coherent Sapphire 488 nm blue, 25 mW Coherent Vioflame 405 nm violet, and 17 mW JDS Uniphase HeNe 633nm red)</td>
		</tr>
		<tr >
			<td >FACS Diva software&#160;</td>
			<td >BD Biosciences</td>
			<td ></td>
			<td >PC version 6.0</td>
		</tr>
		<tr >
			<td >FlowJo software&#160;</td>
			<td >Treestar US</td>
			<td ></td>
			<td >Mac version 9.6.1 or PC version 7.6.5</td>
		</tr>
		<tr >
			<td >Sphero Rainbow fluorescent particles</td>
			<td >BD Biosciences</td>
			<td >556298</td>
			<td >used to adjust all channel voltages to maintain fluorescence intensity consistency&#160;</td>
		</tr>
		<tr >
			<td >Ultra Rainbow fluorescent particles&#160;</td>
			<td >Spherotech</td>
			<td >URFP-10-5</td>
			<td >used in addition to Megamix-Plus SSC beads to ensure EV gating consistency from batch to batch</td>
		</tr>
		<tr >
			<td >Megmix-Plus SSC beads</td>
			<td >Biocytex</td>
			<td >7803</td>
			<td >used to adjust FSC and SSC&#160; voltages to maintain&#160; consistency&#160; between runs. Can also used to monitor flow rate and ajust flow rate dial in order to ensure that same flow rate is used in all runs</td>
		</tr>
		<tr >
			<td >AbC Anti-Mouse Bead Kit</td>
			<td >Life Technologies</td>
			<td >A-10344</td>
			<td >used for compensation controls &#38; negative AbC beads used for beads-based method</td>
		</tr>
		<tr >
			<td >Nonidet P-40 Alternative (NP-40) (CAS 9016-45-9)</td>
			<td >Santa Cruz&#160;</td>
			<td >sc-281108</td>
			<td >used in the individual detection method only to lyse samples after initial reading for use as negative controls. Stock may be diluted to 1:10 in PBS and stored in fridge for up to 1 month.</td>
		</tr>
		<tr >
			<td >BD TruCOUNT Tubes</td>
			<td >BD Biosciences</td>
			<td >340334</td>
			<td >used whenver absolute EV concentrations are needed</td>
		</tr>
		<tr >
			<td >Ultrafree-MC, GV 0.22 &#181;m Centrifugal Filter Units</td>
			<td >Millipore&#160;</td>
			<td >UFC30GVNB</td>
			<td >used to post-stain wash Evs and/or fractionate EVs based on size</td>
		</tr>
		<tr >
			<td >Vacutainer glass whole blood tubes ACD-A</td>
			<td >BD Biosciences</td>
			<td >364606</td>
			<td ></td>
		</tr>
		<tr >
			<td >Facs tubes 12x75 polystrene</td>
			<td >BD Biosciences</td>
			<td >352058</td>
			<td ></td>
		</tr>
		<tr >
			<td >50mL Reservoirs individually wrapped&#160;</td>
			<td >Phenix</td>
			<td >RR-50-1s</td>
			<td ></td>
		</tr>
		<tr >
			<td >Green-Pak pipet tips - 10&#181;L</td>
			<td >Rainin</td>
			<td >GP-L10S</td>
			<td ></td>
		</tr>
		<tr >
			<td >Green-Pak pipet tips -200&#181;L&#160;</td>
			<td >Rainin</td>
			<td >GP-L250S</td>
			<td ></td>
		</tr>
		<tr >
			<td >Green -Pak pipet tips - 1000&#181;L&#160;</td>
			<td >Rainin</td>
			<td >GP-L1000S</td>
			<td ></td>
		</tr>
		<tr >
			<td >Stable Stack L300 tips presterilized</td>
			<td >Rainin</td>
			<td >SS-L300S</td>
			<td ></td>
		</tr>
		<tr >
			<td >Pipet-Lite&#160;XLS 8 Channel LTS&#160;Adjustable&#160;Spacer&#160;</td>
			<td >Rainin</td>
			<td >LA8-300XLS</td>
			<td ></td>
		</tr>
		<tr >
			<td >96 well tissue culture plates</td>
			<td >E&#38;K Scientific</td>
			<td >EK-20180</td>
			<td ></td>
		</tr>
		<tr >
			<td >RPMI 1640 Media (without Hepes)</td>
			<td >UCSF Cell Culture Facility</td>
			<td >CCFAE001</td>
			<td >media used for bead-based detection method</td>
		</tr>
		<tr >
			<td >Dulbeccos PBS D-PBS, CaMg-free, 0.2&#181;m filtered</td>
			<td >UCSF Cell Culture Facility</td>
			<td >CCFAL003</td>
			<td ></td>
		</tr>
		<tr >
			<td >Ultracentrifuge Tube, Thinwall, Ultra-Clear</td>
			<td >BECKMAN COULTER INC</td>
			<td >344058</td>
			<td ></td>
		</tr>
		<tr >
			<td >PANEL I</td>
			<td ></td>
			<td ></td>
			<td ></td>
		</tr>
		<tr >
			<td >CD3 PerCP-Cy5.5</td>
			<td >Biolegend</td>
			<td >344808</td>
			<td >2 &#181;l</td>
		</tr>
		<tr >
			<td >CD14 APC-Cy7</td>
			<td >Biolegend</td>
			<td >301820</td>
			<td >2 &#181;l</td>
		</tr>
		<tr >
			<td >CD16 V450</td>
			<td >BD Biosciences</td>
			<td >560474</td>
			<td >2 &#181;l</td>
		</tr>
		<tr >
			<td >CD28 FITC</td>
			<td >biolegend</td>
			<td >302906</td>
			<td >2 &#181;l</td>
		</tr>
		<tr >
			<td >CD152 APC</td>
			<td >BD Biosciences</td>
			<td >555855</td>
			<td >2 &#181;l</td>
		</tr>
		<tr >
			<td >CD19 A700</td>
			<td >Biolegend</td>
			<td >302226</td>
			<td >2 &#181;l</td>
		</tr>
		<tr >
			<td >PANEL II</td>
			<td ></td>
			<td ></td>
			<td ></td>
		</tr>
		<tr >
			<td >CD41a PerCP-Cy5.5</td>
			<td >BD Biosciences</td>
			<td >340930</td>
			<td >2 &#181;l</td>
		</tr>
		<tr >
			<td >CD62L APC</td>
			<td >Biolegend</td>
			<td >304810</td>
			<td >2 &#181;l</td>
		</tr>
		<tr >
			<td >CD108 PE&#160;</td>
			<td >BD Biosciences</td>
			<td >552830</td>
			<td >2 &#181;l</td>
		</tr>
		<tr >
			<td >CD235a FITC</td>
			<td >biolegend</td>
			<td >349104</td>
			<td >2 &#181;l</td>
		</tr>
		<tr >
			<td >PANEL III</td>
			<td ></td>
			<td ></td>
			<td ></td>
		</tr>
		<tr >
			<td >CD11b PE-Cy7</td>
			<td >Biolegend</td>
			<td >301322</td>
			<td >2 &#181;l</td>
		</tr>
		<tr >
			<td >CD62p APC</td>
			<td >Biolegend</td>
			<td >304910</td>
			<td >2 &#181;l</td>
		</tr>
		<tr >
			<td >CD66b PE&#160;</td>
			<td >Biolegend</td>
			<td >305106</td>
			<td >2 &#181;l</td>
		</tr>
		<tr >
			<td >CD15 FITC</td>
			<td >exalpha</td>
			<td >X1496M</td>
			<td >5 &#181;l</td>
		</tr>
		<tr >
			<td >CD9 PE</td>
			<td >Biolegend</td>
			<td >555372</td>
			<td ></td>
		</tr>
		<tr >
			<td >CD63 APC</td>
			<td >Biolegend</td>
			<td >353008</td>
			<td ></td>
		</tr>
		<tr >
			<td >APC-Cy7 Ms IgG2a, &#954;</td>
			<td >Biolegend</td>
			<td >400230</td>
			<td ></td>
		</tr>
	</tbody>
</table>

techniques for the analysis of extracellular vesicles using flow cytometry

Extracellular Vesicles (EVs) are small, membrane-derived vesicles found in bodily fluids that are highly involved in cell-cell communication and help regulate a diverse range of biological processes. Analysis of EVs using flow cytometry (FCM) has been notoriously difficult due to their small size and lack of discrete populations positive for markers of interest. Methods for EV analysis, while considerably improved over the last decade, are still a work in progress. Unfortunately, there is no one-size-fits-all protocol, and several aspects must be considered when determining the most appropriate method to use. Presented here are several different techniques for processing EVs and two protocols for analyzing EVs using either individual detection or a bead-based approach. The methods described here will assist with eliminating the antibody aggregates commonly found in commercial preparations, increasing signal&#8211;to-noise ratio, and setting gates in a rational fashion that minimizes detection of background fluorescence. The first protocol uses an individual detection method that is especially well suited for analyzing a high volume of clinical samples, while the second protocol uses a bead-based approach to capture and detect smaller EVs and exosomes.

Figure 1 outlines the overall processing scheme for the isolation and detection of EVs using either the bead-based method or individual detection method. Individual detection of EVs using FCM works well for analyzing larger EVs but most cytometers are not capable of individually detecting particles as small as exosomes. A bead-based approach allows small EVs to be detected, however, there are drawbacks associated with using this method, as outlined in Table 1. Generally, isolation of EVs...

Watch this Scientific Journal Video about Techniques for the Analysis of Extracellular Vesicles Using Flow Cytometry at JoVE.com

Techniques for the Analysis of Extracellular Vesicles Using Flow Cytometry

Many different methods exist for the measurement of extracellular vesicles (EVs) using flow cytometry (FCM). Several aspects should be considered when determining the most appropriate method to use. Two protocols for measuring EVs are presented, using either individual detection or a bead-based approach.

Extracellular Vesicles (EVs) are small, membrane-derived vesicles found in bodily fluids that are highly involved in cell-cell communication and help ...

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22.9K Views.  Blood Systems Research Institute. Many different methods exist for the measurement of extracellular vesicles (EVs) using flow cytometry (FCM). Several aspects should be considered when determining the most appropriate method to use. Two protocols for measuring EVs are presented, using either individual detection or a bead-based approach. 

Video: Techniques for the Analysis of Extracellular Vesicles Using Flow Cytometry

Flow Cytometry Purification of Mouse Meiotic Cells

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the unique differentiation programs that are manifest during meiosis. Two principal methods have been developed to purify, to varying degrees, various meiotic fractions from both adult and immature animals: elutriation or Staput (sedimentation) using BSA and/or percoll gradients. Both of these methods rely on cell size and density to separate meiotic cells1-5. Overall, except for few cell populations6, these protocols fail to yield sufficient purity of the numerous meiotic cell populations that are necessary for detailed molecular analyses. Moreover, with such methods usually one type of meiotic cells can be purified at a given time, which adds an extra level of complexity regarding the reproducibility and homogeneity when comparing meiotic cell samples.
Here, we describe a refined method that allows one to easily visualize, identify, and purify meiotic cells, from germ cells to round spermatids, using FACS combined with Hoechst 33342 staining7,8. This method provides an overall snapshot of the entire meiotic process and allows one to highly purify viable cells from most stage of meiosis. These purified cells can then be analyzed in detail for molecular changes that accompany progression through meiosis, for example changes in gene expression9,10and the dynamics of nucleosome occupancy at hotspots of meiotic recombination11.

An efficient method to obtain highly purified viable meiotic fractions from mouse testis is described, which combines a refined cell dissociation protocol with fluorescent activated cell sorting (FACS). This method takes advantage of differences in the DNA content and nuclear density of discrete meiotic fractions.

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the ...

Optimized Staining and Proliferation Modeling Methods for Cell Division Monitoring using Cell Tracking Dyes

Fluorescent cell tracking dyes, in combination with flow and image cytometry, are powerful tools with which to study the interactions and fates of different cell types in vitro and in vivo.1-5 Although there are literally thousands of publications using such dyes, some of the most commonly encountered cell tracking applications include monitoring of:
<ol>
	<li>stem and progenitor cell quiescence, proliferation and/or differentiation6-8</li>
	<li>antigen-driven membrane transfer9 and/or precursor cell proliferation3,4,10-18 and</li>
	<li>immune regulatory and effector cell function1,18-21.</li>
</ol>
Commercially available cell tracking dyes vary widely in their chemistries and fluorescence properties but the great majority fall into one of two classes based on their mechanism of cell labeling. "Membrane dyes", typified by PKH26, are highly lipophilic dyes that partition stably but non-covalently into cell membranes1,2,11. "Protein dyes", typified by CFSE, are amino-reactive dyes that form stable covalent bonds with cell proteins4,16,18. Each class has its own advantages and limitations. The key to their successful use, particularly in multicolor studies where multiple dyes are used to track different cell types, is therefore to understand the critical issues enabling optimal use of each class2-4,16,18,24.
The protocols included here highlight three common causes of poor or variable results when using cell-tracking dyes. These are:
<ol>
	<li>Failure to achieve bright, uniform, reproducible labeling. This is a necessary starting point for any cell tracking study but requires attention to different variables when using membrane dyes than when using protein dyes or equilibrium binding reagents such as antibodies.</li>
	<li>Suboptimal fluorochrome combinations and/or failure to include critical compensation controls. Tracking dye fluorescence is typically 102 - 103 times brighter than antibody fluorescence. It is therefore essential to verify that the presence of tracking dye does not compromise the ability to detect other probes being used.</li>
	<li>Failure to obtain a good fit with peak modeling software. Such software allows quantitative comparison of proliferative responses across different populations or stimuli based on precursor frequency or other metrics. Obtaining a good fit, however, requires exclusion of dead/dying cells that can distort dye dilution profiles and matching of the assumptions underlying the model with characteristics of the observed dye dilution profile.</li>
</ol>
Examples given here illustrate how these variables can affect results when using membrane and/or protein dyes to monitor cell proliferation.

Successful use of cell tracking dyes to monitor immune cell function and proliferation involves several critical steps. We describe methods for: 1) obtaining bright, uniform, reproducible label-ing with membrane dyes; 2) selecting fluorochromes and data acquisition conditions; and 3) choosing a model to quantify cell proliferation based on dye dilution.

Fluorescent cell tracking dyes, in combination with flow and image cytometry, are powerful tools with which to study the interactions and fates of ...

Temporal Tracking of Cell Cycle Progression Using Flow Cytometry without the Need for Synchronization

This protocol describes a method to permit the tracking of cells through the cell cycle without requiring the cells to be synchronized. Achieving cell synchronization can be difficult for many cell systems. Standard practice is to block cell cycle progression at a specific stage and then release the accumulated cells producing a wave of cells progressing through the cycle in unison. However, some cell types find this block toxic resulting in abnormal cell cycling, or even mass death. Bromodeoxyuridine (BrdU) uptake can be used to track the cell cycle stage of individual cells. Cells incorporate this synthetic thymidine analog, while synthesizing new DNA during S phase. By providing BrdU for a brief period it is possible to mark a pool of cells that were in S phase while the BrdU was present. These cells can then be tracked through the remainder of the cell cycle and into the next round of replication, permitting the duration of the cell cycle phases to be determined without the need to induce a potentially toxic cell cycle block. It is also possible to determine and correlate the expression of both internal and external proteins during subsequent stages of the cell cycle. These can be used to further refine the assignment of cell cycle stage or assess effects on other cellular functions such as checkpoint activation or cell death.

This protocol describes the use of bromodeoxyuridine (BrdU) uptake to permit the temporal tracking of cells that were in S phase at a specific point in time. Addition of DNA dyes and antibody labeling facilitates detailed analysis of the fate of the S phase cells at later times.

This protocol describes a method to permit the tracking of cells through the cell cycle without requiring the cells to be synchronized. Achieving cell ...

Sample Preparation for Mass Cytometry Analysis

Mass cytometry utilizes antibodies conjugated with heavy metal labels, an approach that has greatly increased the number of parameters and opportunities for deep analysis well beyond what is possible with conventional fluorescence-based flow cytometry. As with any new technology, there are critical steps that help ensure the reliable generation of high-quality data. Presented here is an optimized protocol that incorporates multiple techniques for the processing of cell samples for mass cytometry analysis. The methods described here will help the user avoid common pitfalls and achieve consistent results by minimizing variability, which can lead to inaccurate data. To inform experimental design, the rationale behind optional or alternative steps in the protocol and their efficacy in uncovering new findings in the biology of the system being investigated is covered. Lastly, representative data is presented to illustrate expected results from the techniques presented here.

This article describes the collection and processing of samples for mass cytometry analysis.

Mass cytometry utilizes antibodies conjugated with heavy metal labels, an approach that has greatly increased the number of parameters and opportunities ...

LED Thermo Flow &#8212; Combining Optogenetics with Flow Cytometry

Optogenetic tools allow isolated, functional investigations of almost any signaling molecule within complex signaling pathways. A major obstacle is the controlled delivery of light to the cell sample and hence the most popular tools for optogenetic studies are microscopy-based cell analyses and in vitro experiments. The flow cytometer has major advantages over a microscope, including the ability to rapidly measure thousands of cells at single cell resolution. However, it is not yet widely used in optogenetics. Here, we present a device that combines the power of optogenetics and flow cytometry: the LED Thermo Flow. This device illuminates cells at specific wavelengths, light intensities and temperatures during flow cytometric measurements. It can be built at low cost and be used with most common flow cytometers. To demonstrate its utility, we characterized the photoswitching kinetics of Dronpa proteins in vivo and in real time. This protocol can be adapted to almost all optically controlled substances and substantially expands the set of possible experiments. More importantly, it will greatly simplify the discovery and development of new optogenetic tools.

LED Thermo Flow &amp;#8212; Combining Optogenetics with Flow Cytometry

Optically controlled substances are powerful tools to study signaling pathways. To expand the spectrum of possible experiments, we developed a device for studying optically controlled substances in real time using flow cytometry: the LED Thermo Flow.

Optogenetic tools allow isolated, functional investigations of almost any signaling molecule within complex signaling pathways. A major obstacle is the ...

Analysis of Cell Suspensions Isolated from Solid Tissues by Spectral Flow Cytometry

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to the analysis of a few fluorochromes, currently there are dozens of different fluorescent dyes, and up to 14-18 different dyes can be combined at a time. However, several limitations still impair the analytical capabilities. Because of the multiplicity of fluorescent probes, data analysis has become increasingly complex due to the need of large, multi-parametric compensation matrices. Moreover, mutant mouse models carrying fluorescent proteins to detect and trace specific cell types in different tissues have become available, so the analysis (by flow cytometry) of auto-fluorescent cell suspensions obtained from solid organs is required. Spectral flow cytometry, which distinguishes the shapes of emission spectra along a wide range of continuous wavelengths, addresses some of these problems. The data is analyzed with an algorithm that replaces compensation matrices and treats auto-fluorescence as an independent parameter. Thus, spectral flow cytometry should be capable of discriminating fluorochromes with similar emission peaks and can provide a multi-parametric analysis without compensation requirements.
This protocol describes the spectral flow cytometry analysis, allowing for a 21-parameter (19 fluorescent probes) characterization and the management of an auto-fluorescent signal, providing high resolution in minor population detection. The results presented here show that spectral flow cytometry presents advantages in the analysis of cell populations from tissues difficult to characterize in conventional flow cytometry, such as the heart and the intestine. Spectral flow cytometry thus demonstrates the multi-parametric analytical capacity of high-performing conventional flow cytometry without the requirement for compensation and enables auto-fluorescence management.

This article describes spectral cytometry, a new approach in flow cytometry that uses the shapes of emission spectra to distinguish fluorochromes. An algorithm replaces compensations and can treat auto-fluorescence as an independent parameter. This new approach allows for the proper analysis of cells isolated from solid organs.

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to ...

Flow Cytometric Analysis of Extracellular Vesicles from Cell-conditioned Media

Flow cytometry (FC) is the method of choice for semi-quantitative measurement of cell-surface antigen markers. Recently, this technique has been used for phenotypic analyses of extracellular vesicles (EV) including exosomes (Exo) in the peripheral blood and other body fluids. The small size of EV mandates the use of dedicated instruments having a detection threshold around 50-100 nm. Alternatively, EV can be bound to latex microbeads that can be detected by FC. Microbeads, conjugated with antibodies that recognize EV-associated markers/Cluster of Differentiation CD63, CD9, and CD81 can be used for EV capture. Exo isolated from CM can be analyzed with or without pre-enrichment by ultracentrifugation. This approach is suitable for EV analyses using conventional FC instruments. Our results demonstrate a linear correlation between Mean Fluorescence Intensity (MFI) values and EV concentration. Disrupting EV through sonication dramatically decreased MFI, indicating that the method does not detect membrane debris. We report an accurate and reliable method for the analysis of EV surface antigens, which can be easily implemented in any laboratory.

The protocol describes a reproducible method designed for use with cell culture supernatants to detect surface epitopes on small extracellular vesicles (EV). It utilizes specific EV immunoprecipitation using beads coupled with antibodies that recognize surface antigen CD9, CD63, and CD81. The method is optimized for downstream flow cytometry analysis.

Flow cytometry (FC) is the method of choice for semi-quantitative measurement of cell-surface antigen markers. Recently, this technique has been used for ...

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 ...

Nanoparticle Tracking Analysis for the Quantification and Size Determination of Extracellular Vesicles

The physiological and pathophysiological roles of extracellular vesicles (EVs) have become increasingly recognized, making the EV field a quickly evolving area of research. There are many different methods for EV isolation, each with distinct advantages and disadvantages that affect the downstream yield and purity of EVs. Thus, characterizing the EV prep isolated from a given source by a chosen method is important for interpretation of downstream results and comparison of results across laboratories. Various methods exist for determining the size and quantity of EVs, which can be altered by disease states or in response to external conditions. Nanoparticle tracking analysis (NTA) is one of the prominent technologies used for high-throughput analysis of individual EVs. Here, we present a detailed protocol for quantification and size determination of EVs isolated from mouse perigonadal adipose tissue and human plasma using a breakthrough technology for NTA representing major advances in the field. The results demonstrate that this method can deliver reproducible and valid total particle concentration and size distribution data for EVs isolated from different sources using different methods, as confirmed by transmission electron microscopy. The adaptation of this instrument for NTA will address the need for standardization in NTA methods to increase rigor and reproducibility in EV research.

We demonstrate how to use a novel nanoparticle tracking analysis instrument to estimate the size distribution and total particle concentration of extracellular vesicles isolated from mouse perigonadal adipose tissue and human plasma.

The physiological and pathophysiological roles of extracellular vesicles (EVs) have become increasingly recognized, making the EV field a quickly evolving ...

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 ...