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Method Article

Techniques for the Analysis of Extracellular Vesicles Using Flow Cytometry

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DOI:

10.3791/52484

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March 17th, 2015

 ,  , 
1Blood Systems Research Institute, 2Department of Medicine, University of California, San Francisco, 3Department of Laboratory Medicine, University of California, San Francisco

In This Article

Summary

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.

Abstract

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–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.

Introduction

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–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 communication2–4. 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 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 “EVs” 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 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 and western blotting22,23. While FCM is the method of choice for many researchers9,24–26 due to its high throughput capabilities, analysis of EVs using FCM has been notoriously difficult due to their size and lack of discrete positive populations27–32. 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 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–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.

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Protocol

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

  1. 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.
  2. Centrifuge the samples at 1,500 x g for 10 min at RT 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.
  3. 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 µl to avoid disturbing the pellet and add the PPP to a new tube.
  4. 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 °C for up to two years for later analysis (refer to Figure 1A for overview).
  5. 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 µm-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.

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.

  1. Remove 12 samples from freezer (if stored at -80 °C) and thaw at 37 °C.
  2. Pipette contents up and down several times to mix. Remove 320 µl 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.

1.3) Staining EV Samples

  1. Prior to staining, filter all antibodies (Abs) to remove aggregates, which can cause positive signals.
    1. Combine antibodies to be used in each of the 3 panels into separate 0.22 µm 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.
  2. Add the appropriate amount of filtered Ab mixture to each well in row 2 (e.g., samples in Panel I are stained with 2 µl of each Ab, so a total of 12 µl 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 µl/well of the Panel II cocktail to row 3 and 11 µl/well of the Panel III cocktail to row 4; refer to Materials List for specific panel information).
  3. Using the multichannel pipet, mix the PPP samples in row 1 up and down and transfer 100 µl from the wells in row 1 to the wells in row 2. Mix up and down. Change tips and repeat, transferring 100 µl from row 1 to rows 3 and 4. Incubate at 4 °C for 30 min.

1.4) Washing MV Samples

  1. Remove the 96-well plate from 4 °C and transfer to biological safety cabinet. Using a multichannel pipette, add 220 µl of PBS/well to rows 6-8 (to be used for rinsing/washing the wells containing stained PPP).
  2. 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 µl of PBS from the wash rows and add to the corresponding wells from which PPP was just removed.
  3. 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.
  4. Repeat this process with the remaining stained samples until all stained PPP samples have been transferred along with their rinse solutions to centrifugal filters.
  5. 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 “dry” 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.
  6. Remove the centrifugal filter tubes and return to the biological safety cabinet. Using the multichannel pipet, resuspend the tops of the filters in 300 µl 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 µl) an exact number of times (e.g., 8 times) for each sample.

1.5) Cytometer Setup

  1. Open the FCM software. Prior to experiment setup, perform daily instrument calibration and setup using instrument setup beads (following manufacturer’s instructions).
  2. 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:
    1. 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.
    2. Incubate at 4 °C for 30 min, wash with PBS and resuspend in 400 µl of PBS.
    3. 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.
    4. Select the tab “Experiment,” then select “Compensation,” then select “Calculate Compensation” to apply compensation values to all samples.
  3. 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.
  4. While running a tube of 0.22 µm-filtered PBS, adjust the FSC & 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).
  5. Next, run a tube containing 0.2 µm – 1.0 µm 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 µm and 1.0 µm. Alternatively, in an SSC-H histogram, draw a gate to include all events smaller than the 1.0 µm beads.
  6. Set the cytometer’s flow rate to “Lo” (approximately 8-12 µl/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.
  7. 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.

1.6) Sample Reading

  1. Set the cytometer’s flow rate to “Lo” (approximately 8-12 µl/min) and run each sample for exactly 1 or 2 min.
  2. After the first reading, add 20 µl 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.
  3. Once all samples have been read, export all .fcs files into a separate file to be used for further analysis using FCM analysis software.

1.7) Data Analysis

  1. Open the FCM analysis software. Import all of the .fcs files into a new experiment file.
  2. Open the beads-only tube. In the FSC-A vs. SSC-A plot, draw a gate around all beads sized between 0.2 µm and 1.0 µm. This is the EV gate. Drag to add to all samples.
  3. 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.
  4. 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.

2. METHOD B: Beads Method

2.1) Processing of Blood Sample/Isolation of EVs

  1. Refer to the blood processing method described in Method A (Section 1.1).

2.2) Preparing Samples for Analysis

  1. If desired, fractionate PPP or ultracentrifuged EVs into exosomes and microvesicles. Add 250 µl of PPP or ultracentrifuged EVs to 0.22 μm 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 “dry” 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.
  2. Wash uncoated 6 µm 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 μl of RPMI media alone to the beads. To all other tubes, add 200 μl of PPP or ultracentrifuged EVs (or their fractions) and 200 μl of RPMI media.
  3. Adjust the final volume of all tubes to 400 μl with media and incubate overnight at 4 °C on a shaker.
  4. The next morning, wash beads with 2 ml of media. Aspirate off the supernatant.
  5. Block with 5% bovine serum albumin (BSA) in media (400 μl) for 3 hr at 4 °C on a shaker.
  6. Wash beads with 2 ml of media. Aspirate and resuspend pellet in 100 μl of medium.

2.3) Staining EV Samples

  1. 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.
  2. Combine all antibodies to be used in a single panel into a 0.22 µm 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.
  3. Add appropriate volume of filtered antibody cocktail to all tubes and incubate for 30 min at 4 °C.
  4. Wash beads with 2 ml of media, resuspend in 400 μl of media and run immediately (or within the same day) on flow cytometer.

2.4) Cytometer Setup and Sample Reading

  1. Open the FCM software. Prior to experiment setup, perform daily instrument calibration and setup using instrument setup beads (following manufacturer’s instructions).
  2. 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:
    1. 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 °C for 30 min, wash with PBS and resuspend in 400 µl of PBS.
    2. 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.
    3. Select the tab “Experiment,” then “Compensation,” then “Calculate Compensation” to apply compensation values to all samples.
  3. 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.
  4. Run samples, gate on the singlet beads population and acquire 2,000 events in this gate. Export .fcs files.
  5. 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.

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Results

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...

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Discussion

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...

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Disclosures

The authors have no conflict of interest to disclose.

Acknowledgements

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.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
LSR II benchtop flow cytometerBD Biosciences3-laser (20 mW Coherent Sapphire 488 nm blue, 25 mW Coherent Vioflame 405 nm violet, and 17 mW JDS Uniphase HeNe 633 nm red)
FACS Diva software BD BiosciencesPC version 6.0
FlowJo software Treestar USMac version 9.6.1 or PC version 7.6.5
Sphero Rainbow fluorescent particlesBD Biosciences556298used to adjust all channel voltages to maintain fluorescence intensity consistency 
Ultra Rainbow fluorescent particles SpherotechURFP-10-5used in addition to Megamix-Plus SSC beads to ensure EV gating consistency from batch to batch
Megmix-Plus SSC beadsBiocytex7803used to adjust FSC and SSC  voltages to maintain  consistency  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
AbC Anti-Mouse Bead KitLife TechnologiesA-10344used for compensation controls & negative AbC beads used for beads-based method
Nonidet P-40 Alternative (NP-40) (CAS 9016-45-9)Santa Cruz sc-281108used 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.
BD TruCOUNT TubesBD Biosciences340334used whenver absolute EV concentrations are needed
Ultrafree-MC, GV 0.22 µm Centrifugal Filter UnitsMillipore UFC30GVNBused to post-stain wash Evs and/or fractionate EVs based on size
Vacutainer glass whole blood tubes ACD-ABD Biosciences364606
Facs tubes 12x75 polystreneBD Biosciences352058
50 ml Reservoirs individually wrapped PhenixRR-50-1s
Green-Pak pipet tips - 10 µlRaininGP-L10S
Green-Pak pipet tips -200 µl RaininGP-L250S
Green -Pak pipet tips - 1,000 µl RaininGP-L1000S
Stable Stack L300 tips presterilizedRaininSS-L300S
Pipet-Lite XLS 8 Channel LTS Adjustable Spacer RaininLA8-300XLS
96 well tissue culture platesE&K ScientificEK-20180
RPMI 1640 Media (without Hepes)UCSF Cell Culture FacilityCCFAE001media used for bead-based detection method
Dulbeccos PBS D-PBS, CaMg-free, 0.2 µm filteredUCSF Cell Culture FacilityCCFAL003
Ultracentrifuge Tube, Thinwall, Ultra-ClearBECKMAN COULTER INC344058
PANEL I
CD3 PerCP-Cy5.5Biolegend3448082 µl
CD14 APC-Cy7Biolegend3018202 µl
CD16 V450BD Biosciences5604742 µl
CD28 FITCbiolegend3029062 µl
CD152 APCBD Biosciences5558552 µl
CD19 A700Biolegend3022262 µl
PANEL II
CD41a PerCP-Cy5.5BD Biosciences3409302 µl
CD62L APCBiolegend3048102 µl
CD108 PE BD Biosciences5528302 µl
CD235a FITCbiolegend3491042 µl
PANEL III
CD11b PE-Cy7Biolegend3013222 µl
CD62p APCBiolegend3049102 µl
CD66b PE Biolegend3051062 µl
CD15 FITCexalphaX1496M5 µl
CD9 PEBiolegend555372
CD63 APCBiolegend353008
APC-Cy7 Ms IgG2a, κBiolegend400230

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Bead Based DetectionIndividual DetectionPlatelet Poor PlasmaAntibody StainingCytometer SetupFluorescent ParticlesPolystyrene BeadsForward Scatter
Techniques for the Analysis of Extracellular Vesicles Using Flow Cytometry
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