We acknowledge support and advice from Dr. Jill Hutchcroft, Director of the Flow Cytometry Core Facility at Purdue University. This work was supported by R01CA226259 and R01CA205420 to A.L.K., an American Lung Association Innovation Award (ANALA2023) IA-1059916 to A.L.K., a Purdue Shared resource facility grant P30CA023168, and a flagship Fulbright Doctoral Scholarship awarded by the Department of the State, USA to H.H.

NOTE: As a prerequisite to using this technique, identification and validation of selectively exported miRNAs is required. Since different cell lines vary based on the miRNA cargo sorted into their EVs, it is recommended that the cell line of interest and associated EVs be evaluated for miRNAs prior to use. Additionally, lipofectamine-based transfection is one of the critical steps in the protocol; predetermining transfection efficiency prior to setting up the experiment is recommended.
1. Growing cells in culture and transfecting cells with fluorophore-conjugated EV-miRNA
<ol>
	<li>Plate 200,000 to 400,000 cells in triplicate wells in a 6-well plate in an appropriate culture medium. Gently swirl the plate to ensure uniform cell distribution. Incubate the cells until they reach 70%-80% confluency, approximately 24-48 h.</li>
	<li>For each well to be transfected, dilute the transfection reagent in 100&#181;L of serum-free medium in a microcentrifuge tube according to the manufacturer&#39;s instructions. 
	NOTE: The optimal concentration of lipid should be determined for the cell line of interest prior to this setup.</li>
	<li>For each well to be transfected, dilute the fluorophore-conjugated EV-miRNA (2nM) and cell-miRNA (2nM) separately in a microcentrifuge tube in 100 &#181;L of serum-free media. 
	NOTE: The total volume should be calculated for the total number of wells transfected. In this protocol, the fluorescence was evaluated at 7 time points; therefore, the total volume was 700 &#181;L for step 1.2 and 700 &#181;L for step 1.3. Additionally, to confidently capture the positive fluorescent population of cells, the user should transfect cells with a non-fluorescent scrambled miRNA and set up a gate excluding these cells.</li>
	<li>Add the diluted RNA mix (from step 1.3) to the diluted transfection reagent (from step 1.2) and mix gently by inverting the tube 3-4 times. Let the combined lipid-RNA mix incubate for 30 min at room temperature (RT).</li>
	<li>Remove the regular culture media from the cells and add 800 &#181;L of serum-free media to each well. Gently add 200 &#181;L of the lipid-RNA mix (from step 1.4) dropwise to the cells. 
	NOTE: Opti-MEM media was used for transfecting the cell line of interest (Calu6 cells). Try to add the lipid-RNA mix directly to the culture media, avoiding adding the mixture to the sides of the plate.</li>
</ol>
2. Harvesting transfected cells at various time points for analysis of fluorescence
<ol>
	<li>Incubate the transfected cells at 37 &#176;C for 7 h. 
	NOTE: To validate consistent transfection of both miRNAs (EV-miRNA and cell-miRNA), cells are collected from one well 3 h post-transfection to analyze the fluorescence using the flow cytometer.</li>
	<li>After the 7 h incubation period, remove the transfection complex from the wells and replace it with complete media.</li>
	<li>To evaluate the 7-h time-point, harvest the cells and wash the cells three times with 1x PBS.
	<ol>
		<li>If working with adherent cells, add 200 &#181;L of trypsin to one well and wait for the cells to detach from the plate. Transfer the detached cells to a microcentrifuge tube and pellet by centrifugation at 200xg for 5 min.</li>
		<li>Discard the supernatant carefully to avoid disrupting the cell pellet. For the first PBS wash, resuspend the pellet in ~500 &#181;L of 1x PBS and repeat the pelleting using the centrifugation steps above. Repeat the PBS wash an additional two times.</li>
		<li>Resuspend the cell pellet in 200 &#181;L of 2% paraformaldehyde (PFA) solution and evaluate fluorescence. 
		NOTE: After adding 2% PFA, the cell pellet can be stored at 4 &#176;C for 3-4 days. </li>
	</ol>
	</li>
	<li>Harvest the remaining time points following the steps in section 2.3. Collect all of the stored cell pellets and proceed with the steps in section 3.</li>
</ol>
3. Flow cytometry analysis of cell-miRNA and EV-miRNA signal in cells
<ol>
	<li>Prepare cells for flow cytometry analysis. Filter the cell suspension from section 2 through 35 &#181;m strainer caps that are part of the round bottom polystyrene FACS tubes to enable the removal of debris and cell aggregates that might interfere with fluorophore analysis.</li>
	<li>Set up the instrument following calibration instructions. Turn on the flow cytometer and allow it to warm up for at least 30 min before use. Prime the fluidics system with sheath fluid, and check and adjust the laser alignment.</li>
	<li>Run quality control to confirm instrument fluidics. Load ultrapure&#160;filtered water into the flow cytometer and run at an appropriate flow rate for adequate acquisition time. 
	NOTE: The appropriate flow rate and acquisition time will depend on the cell type of interest, sample complexity, and desired data quality.</li>
	<li>Open the flow cytometry data analysis software and confirm the performance of the detector and lasers. Set up threshold and voltage parameters for detection based on the cell line of interest and fluorophores of choice. 
	NOTE: For all downstream experiments, Calu6 cells were used.
	<ol>
		<li>Set the threshold for capturing the signal for Calu6 cells at FSC 5000. Perform the analysis for 1 x 104 cells for each experimental repeat.</li>
		<li>To account for the spectral overlap between different fluorophores, use un-transfected cells and cells independently transfected with only one of the fluorophores to set up compensation controls.</li>
		<li>For detection of miRNA export, use candidate EV-miRNAs (miR-451a and miR-150) conjugated to Alexa-fluor 488, and a control cell-miRNA (cel-miR-67) conjugated to Cy3.</li>
		<li>For each EV-miRNA, conjugate the fluorophore to the 5&#8242; end of the 5p strand. Before transfection, anneal the fluorophore-conjugated strand to its 100% complementary 3p strand.</li>
		<li>For the expression analysis in transfected cells, analyze the samples using the FSC, SSC, FITC, and PE channels set to 258, 192, 269, and 423 voltage units, respectively (Figure 1A, B).</li>
	</ol>
	</li>
	<li>Introduce the negative control sample into the flow cytometer. Capture cell signals after defining forward scatter (FSC) and side scatter (SSC) intensities in a custom worksheet.</li>
	<li>In the plot, choose FSC for the X-axis, SSC for the Y-axis, and gate for the population of interest by drawing a polygon around it (see Figure 1B, bottom panel).</li>
	<li>Create a new plot using the gated population.
	<ol>
		<li>Set the X-axis to fluorescence signal 1 (FS1) for cell-miRNA detection and the Y-axis to fluorescence signal 2 (FS2) for EV-miRNA detection. Set up compensation parameters for the individual fluorophores using cells independently transfected with each of the fluorescently tagged miRNAs. In the new plot, gate on the population of interest.</li>
		<li>For detection of EV-miRNA, use the FITC channel. For the evaluation of cell-miRNA signal, use the PE channel. In the software, create a gate around the cells that were positive for both FITC and PE by drawing a polygon around the double fluorescence population when FITC was selected for the X-axis and PE was selected for the Y-axis.</li>
	</ol>
	</li>
	<li>Record the statistics for the gated population.
	<ol>
		<li>Record the percentage of cells in the gated population that are positive for both FITC and PE, as well as for those that are positive for either FITC or PE.</li>
	</ol>
	</li>
	<li>Repeat the above steps for any additional samples or controls in the experiment.</li>
</ol>
4. Isolating EVs from transfected cells
<ol>
	<li>For cells cultured in serum-containing media, deplete EVs from the serum to prevent cross-contamination.
	<ol>
		<li>To generate EV-depleted serum, dilute the serum to 50% in cell culture media to reduce viscosity and centrifuge the resulting 50% serum at 110,000 x g for 18 h.</li>
		<li>Collect the supernatant (EV-free serum) carefully and filter it using a 0.2 &#181;m filter assembly. Use the resulting 50% EV-free serum to generate complete media, in this case, RPMI plus 10% serum.</li>
		<li>To collect EVs, add the EV-depleted complete media to the cells and collect the EVs 48 h after incubation. 
		NOTE: The protocol outlined below has been modified from Kowal et al.14.</li>
	</ol>
	</li>
	<li>Grow cells in 15 cm tissue culture dishes to 80% confluency. Transfect the cells separately with cell-miRNA and EV-miRNA in 10 mL of Opti-MEM following the transfection protocol in section 1 of the protocol. 
	NOTE: Not all transfection protocols scaleup linearly. It may be essential to first adapt and evaluate the transfection conditions in 15 cm plates.</li>
	<li>After 7 h, aspirate the transfection media and replace it with the EV-depleted media (15 mL). Grow cells in the EV-depleted media for 48 h.</li>
	<li>Remove the conditioned media from the cells and dispense it into a 50 mL centrifuge tube inside the biosafety cabinet. Centrifuge the conditioned media at 2000 x g for 10 min at 4 &#176;C to remove dead cells and cell debris. Transfer the resulting supernatant to a fresh tube.</li>
	<li>Centrifuge the supernatant at 10,000 x g for 40 min at 4 &#176;C. Remove the supernatant and filter it using a 0.2 &#181;m filter assembly.</li>
	<li>Transfer the filtered cell culture media to a sterile ultracentrifuge tube inside the biosafety cabinet. Weigh and balance the tubes against each other. Spin the samples at 110,000 x g for 90 min at 4 &#176;C. Discard the supernatant.</li>
	<li>Resuspend the pellet in an appropriate volume of 1x PBS to wash and centrifuge at 110,000 x g for 90 min 4 &#176;C. 
	NOTE: The EV pellet was resuspended in 60 mL of 1x PBS based on the maximum volume allowed for the fixed-angle rotor tubes used in this study.</li>
	<li>Aspirate PBS and suspend the EV pellet in an appropriate volume (50-100 &#181;L) of fresh 1x PBS and vortex. Store EVs at -80 &#176;C for later use.</li>
</ol>
5. Flow cytometry analysis of cell-miRNA and EV-miRNA signal in EVs
<ol>
	<li>To prepare EV suspension for flow cytometry analysis, dilute EVs in an appropriate volume of ultrafiltered 1x PBS. 
	NOTE: The loading volume for the flow cytometry analysis of EVs should be determined based on the number of EVs in samples. Roughly, if there are ~30,000 particles in the EV suspension, the final volume should be ~500-600 &#181;L in 1x PBS. A clean sample has around 35-50 particles/&#181;L.</li>
	<li>Use the nano-flow cytometer or an alternative equipment powerful enough to analyze nano-sized particles. 
	NOTE: Here, Attune NxT nano-flow cytometer was used.</li>
	<li>Open the software and set the instrument up following the recommended calibration instructions. Perform the Performance Test using performance nano-beads. If the nano-beads are detected in the standard size range, the performance is optimal. Determine the appropriate flow rate for the detection of EV particles.</li>
	<li>Run quality control using ultrapure filtered water to confirm instrument fluidics and performance of detector and lasers.</li>
	<li>Using calibration beads suitable for the experiment, set the flow cytometer to accurately detect different-sized beads and measure fluorescence signals. 
	NOTE: For the setup of EV analysis by flow cytometry, ApogeeMix beads were used. These beads contain a mixture of non-fluorescent silica beads and fluorescent polystyrene beads ranging in sizes from 80 nm to 1300 nm.
	<ol>
		<li>Set up the parameters to assess the sensitivity and resolution of different populations in the bead mix according to the manufacturer&#39;s protocol. Use the appropriate gating to resolve the bead population from the instrument noise.</li>
	</ol>
	</li>
	<li>Vortex the samples thoroughly to evenly distribute EVs before loading for analysis.</li>
	<li>Open the flow cytometry data analysis software and introduce ultra-filtered PBS. Set the gating up for EV particles using ultra-filtered PBS while ensuring that the gating corresponds to the appropriate size of particles based on the beads in step 5.5.</li>
	<li>Load the negative control sample to analyze EVs as they pass through the flow cytometer. Capture EV signals while plotting FSC on the X-axis and SSC on the Y-axis. 
	NOTE: The SSC threshold for capturing EVs was set at 300. The acquisition volume was set at 95 &#181;L from a total of 145 &#181;L drawn volume. The flow rate was set at 12.5 &#181;L/min. The analysis was performed with 1 x 106 EV particles. The EV population is heterogeneous and will result in some debris isolated along with EVs. Overall particle detection in the FSC vs. SSC plot highlights particles in a diverse range, which is expected when using crude EV preparations.</li>
	<li>Create a new plot using the gated population.
	<ol>
		<li>This time, choose fluorescence signal 1 (FS1) for cell-miRNA detection on the X-axis and fluorescence signal 2 (FS2) for EV-miRNA detection on the Y-axis.</li>
		<li>Set up compensation parameters for the fluorophores using the EV samples isolated from cells transfected with a single miRNA. In the new plot, gate on the EV population of interest.</li>
		<li>For the expression analysis of fluorophore-conjugated miRNAs in EVs isolated from the transfected cells, analyze the EV samples using the FSC, SSC, BL1, and YL1 channels set to 370, 370, 400, and 400 voltage units, respectively (Figure 2A, B).</li>
		<li>Record the statistics for the gated population and visualize them using histograms and dot plots.</li>
	</ol>
	</li>
	<li>Repeat the above steps for any additional samples or controls in the experiment.</li>
</ol>
6. Validating the release of miRNAs in EVs through qRT-PCR
<ol>
	<li>Extract RNA from EVs and parent cells using miRVana RNA isolation kit following the manufacturer&#39;s instructions. 
	NOTE: Quantification of RNA isolated from EVs is not possible because the total amount of RNA isolated from EVs does not typically cross the minimal quantity threshold of a standard Nano-drop. This is due to the depletion of ribosomal RNA from EV samples, which can be validated using the Agilent Bioanalyzer. Thus, the integrity score for EV-RNA quantification is usually not reliable and is not the true representation of EV-RNA quantity. Therefore, for qRT-PCR quantification of the EV-RNA and cell-RNA, equal volumes of RNA isolates following isolation from an equal number of cells were used. For normalization of qRT-PCR, miRNAs ubiquitously present across multiple samples as determined from previously acquired RNA sequencing data were used.</li>
	<li>Convert the extracted RNA to cDNA using a reverse transcriptase kit specific for small RNAs.</li>
	<li>Set up the qRT-PCR reaction using a cDNA template and miRNA-specific primers following the manufacturer&#39;s instructions. 
	NOTE: For validation experiments, miRCURY LNA qRT PCR kit and corresponding primers from the manufacturer were used to evaluate the expression of cell-miRNA and EV-miRNA.</li>
</ol>

A Flow Cytometry Protocol for Understanding Export Mechanisms

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The authors have nothing to disclose.

The newly established protocol enables capturing kinetics of miRNA release into EVs post-transfection of EV-miRNAs. The approach allows simultaneous analysis of multiple EV-miRNAs and cell-miRNAs, subject to the capabilities of the cytometer. Moreover, while flow cytometry analysis can provide valuable insights into EV miRNA biology, it is not without limitations. Albeit some of the limitations can be overcome when used in conjunction with other techniques for a more comprehensive understanding. 
<p class="jove_conte...

MicroRNAs (miRNAs) are one of the best-characterized subsets of small noncoding RNAs, which are known for their critical role in post-transcriptional gene regulation. The expression and biogenesis of most miRNAs follow a coordinated series of events that begins with transcription of the primary miRNAs (pri-miRNAs) in the nucleus. Following nuclear processing by the microprocessor complex into precursor-miRNAs (pre-miRNAs), the pre-miRNAs are exported to the cytoplasm, where they undergo further processing by the RNase III endonuclease, dicer into 21-23 nucleotide mature miRNA duplexes1. One of the strands of the processed mature miRNA binds to target messenger RNAs (mRNAs), leading to degradation or translational repression of the targets2. Based on the pleiotropic role of miRNAs in simultaneously regulating multiple diverse target mRNAs, it is not surprising that miRNA expression is tightly regulated3. Indeed, inappropriate expression contributes to various disease states, especially in cancer. The aberrant expression of miRNAs not only represents a disease-specific signature but has also emerged as a target for prognostic and therapeutic potential4,5,6. In addition to their intracellular roles, miRNAs also have non-autonomous roles. For example, miRNAs can be selectively packaged into EVs by donor cells and exported to recipient sites where they elicit diverse phenotypic responses in normal and disease physiology7,8,9.
Despite clear evidence that EV-associated miRNAs are functional biomolecules, it is not completely understood how specific subsets of miRNAs are dysregulated in disease states such as cancer and how cellular machinery selects and sorts miRNAs into Evs10,11. Given the potential role of EV-miRNAs in modulating the microenvironment, it is critical to identify the mechanisms involved in the export of select miRNAs to fully elucidate the role of EVs in intercellular communication and disease pathogenesis. Understanding the process of miRNA release into EVs will not only highlight important mediators of miRNA export but can also provide insights into potential therapeutics. To achieve this level of knowledge, tools need to be adapted to faithfully address these new experimental questions. Indeed, studies that evaluate EVs are growing exponentially due to the introduction of new techniques and controls12,13. In relation to evaluating the abundance of exported miRNAs into EVs, next-generation sequencing and quantitative reverse transcription polymerase chain reaction (qRT-PCR) have been the current standard tools. While these tools are useful for evaluating the abundance of miRNAs in EVs, there are limitations to their sensitivity and specificity, and using these static approaches for evaluating dynamics is insufficient. Delineating the dynamics of miRNA export into EVs requires a combination of specialized tools. Here, we present a comprehensive protocol using fluorescently conjugated miRNAs, to analyze the dynamics of miRNA release from donor cells and incorporation into EVs. We also discuss the advantages and limitations of the method and provide recommendations for optimal use. The versatile method presented in this paper will be useful to researchers interested in studying miRNA release into EVs and their potential roles in cellular communication and disease.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1.5 mL microcentrifuge tubes</td>
			<td>Fisher</td>
			<td>05-408-129</td>
			<td></td>
		</tr>
		<tr>
			<td>15 mL Falcon tubes</td>
			<td>Corning</td>
			<td>352097</td>
			<td></td>
		</tr>
		<tr>
			<td>6-well clear flat bottom surface treated tissue culture plates</td>
			<td>Fisher Scientific</td>
			<td>FB012927</td>
			<td></td>
		</tr>
		<tr>
			<td>ApogeeMix 25 mL, (PS80/110/500 &#38; Si180/240/300/590/880/1300 nm)&#160;</td>
			<td>Apogee Flow Systems</td>
			<td>1527</td>
			<td>To set up gating and parameters for particle detection&#160;</td>
		</tr>
		<tr>
			<td>Attune Nxt Flow Cytometer</td>
			<td>ThermoFisher Scientific</td>
			<td>N/A</td>
			<td>For fluorescence analysis in EVs</td>
		</tr>
		<tr>
			<td>BD Fortessa Cell Analyzer</td>
			<td>BD Biosciences</td>
			<td>N/A</td>
			<td>For fluorescence analysis in cells&#160;</td>
		</tr>
		<tr>
			<td>Cell culture incubator</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Maintaining temperature of 37&#160;&#176;C and 5% CO2</td>
		</tr>
		<tr>
			<td>Cell Culture medium</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>specific for the cell-line of interest</td>
		</tr>
		<tr>
			<td>Cell line of interest&#160;</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Any cell line tested and evaluated for EV and cellular abundance of miRNAs on interest.</td>
		</tr>
		<tr>
			<td>Cell-miRNA (miRIDIAN microRNA Mimic Red Transfection Control)</td>
			<td>Horizon discovery</td>
			<td>CP-004500-01-05</td>
			<td>miRNAs predetermined to be retained by the cell-line of interest and not selectively exported out into EVs</td>
		</tr>
		<tr>
			<td>EV-miRNA (Fluorophore conjugated miRNAs)</td>
			<td>Integrated DNA Technologies (IDT)</td>
			<td></td>
			<td>miRNAs predetermined to be selectively sorted into EVs&#160;</td>
		</tr>
		<tr>
			<td>Fluorescence microscope</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Gibco Opti-MEM Reduced Serum Medium</td>
			<td>Fisher Scientific</td>
			<td>31-985-070</td>
			<td></td>
		</tr>
		<tr>
			<td>Hausser Scientific Hemocytometer</td>
			<td>Fisher</td>
			<td>02-671-54</td>
			<td></td>
		</tr>
		<tr>
			<td>Hyclone 1x PBS (for cell culture)</td>
			<td>Fisher</td>
			<td>SH30256FS</td>
			<td></td>
		</tr>
		<tr>
			<td>Lipofectamine RNAimax</td>
			<td>Fisher Scientific</td>
			<td>13-778-150</td>
			<td></td>
		</tr>
		<tr>
			<td>miRCURY LNA Reverse Transcriptase</td>
			<td>Qiagen</td>
			<td>339340</td>
			<td></td>
		</tr>
		<tr>
			<td>miRCURY LNA SYBR Green PCR</td>
			<td>Qiagen&#160;</td>
			<td>339347</td>
			<td></td>
		</tr>
		<tr>
			<td>mirVana RNA Isolation</td>
			<td>Qiagen</td>
			<td>AM1561</td>
			<td></td>
		</tr>
		<tr>
			<td>Nuclease free water</td>
			<td>Fisher Scientific</td>
			<td>4387936</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde, 4% in PBS</td>
			<td>Fisher</td>
			<td>AAJ61899AK</td>
			<td></td>
		</tr>
		<tr>
			<td>Reagent reservoir nonsterile</td>
			<td>VWR</td>
			<td>89094-684</td>
			<td></td>
		</tr>
		<tr>
			<td>Thermo ABI QuantiStudio DX Real-Time PCR</td>
			<td>ThermoFisher Scientific</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin 0.25%</td>
			<td>Fisher&#160;</td>
			<td>SH3004201</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultracentrifuge</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
	</tbody>
</table>

tracking mirna release into extracellular vesicles using flow cytometry

Extracellular vesicles (EVs) are important mediators of cellular communication that are secreted by a variety of different cells. These EVs shuttle bioactive molecules, including proteins, lipids, and nucleic acids (DNA, mRNAs, microRNAs, and other noncoding RNAs), from one cell to another, leading to phenotypic consequences in the recipient cells. Of all the various EV cargo, microRNAs (miRNAs) have garnered a great deal of attention for their role in shaping the microenvironment and in educating recipient cells because of their clear dysregulation and abundance in EVs. Additional data indicates that many miRNAs are actively loaded into EVs. Despite this clear evidence, research on the dynamics of export and mechanisms of miRNA sorting is limited. Here, we provide a protocol using flow cytometry analysis of EV-miRNA that can be used to understand the dynamics of EV-miRNA loading and identify the machinery involved in miRNA export. In this protocol, miRNAs predetermined to be enriched in EVs and depleted from donor cells are conjugated to a fluorophore and transfected into the donor cells. The fluorescently tagged miRNAs are then verified for loading into EVs and depletion from cells using qRT-PCR. As both a transfection control and a tool for gating the transfected population of cells, a fluorescently labeled cellular RNA (cell-retained and EV-depleted) is included. Cells transfected with both the EV-miRNA and cell-retained-miRNA are evaluated for fluorescent signals over the course of 72 h. The fluorescence signal intensity specific for the EV-miRNAs diminishes rapidly compared to the cell-retained miRNA. Using this straightforward protocol, one could now assess the dynamics of miRNA loading and identify various factors responsible for loading miRNAs into EVs.

Here, we utilize flow cytometry as a powerful tool to investigate the release of miRNA from the cells into EVs. Using this protocol, flow cytometry analysis of cells transfected with cell-miRNA and EV-miRNA revealed a sequential decrease of fluorescence signal corresponding to EV-miRNA, while the signal corresponding to the cell-miRNA was retained in the cells. To ensure that fluorophore conjugation does not interfere with miRNA release into EVs, miR-451a was conjugated to two separate fluorophores (i.e., Alexa fluor 488...

Watch this Scientific Journal Video about Exploring the Mechanisms of MicroRNA Loading into Extracellular Vesicles in Cancer Progression at JoVE.com

Author Spotlight: Exploring the Mechanisms of MicroRNA Loading into Extracellular Vesicles in Cancer Progression 

Here, we describe a straightforward protocol that enables in vitro assessment of the abundance of fluorescently labeled microRNAs to study the dynamics of microRNA packaging and export into extracellular vesicles (EVs).

Tracking miRNA Release into Extracellular Vesicles using Flow Cytometry

Extracellular vesicles (EVs) are important mediators of cellular communication that are secreted by a variety of different cells. These EVs shuttle ...

tracking-mirna-release-into-extracellular-vesicles-using-flow

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JoVE Journal

Biology

1.3K Views.  Purdue University. Here, we describe a straightforward protocol that enables in vitro assessment of the abundance of fluorescently labeled microRNAs to study the dynamics of microRNA packaging and export into extracellular vesicles (EVs).

Video: Author Spotlight: Exploring the Mechanisms of MicroRNA Loading into Extracellular Vesicles in Cancer Progression 

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 ...

Measuring Peptide Translocation into Large Unilamellar Vesicles

There is an active interest in peptides that readily cross cell membranes without the assistance of cell membrane receptors1. Many of these are referred to as cell-penetrating peptides, which are frequently noted for their potential as drug delivery vectors1-3. Moreover, there is increasing interest in antimicrobial peptides that operate via non-membrane lytic mechanisms4,5, particularly those that cross bacterial membranes without causing cell lysis and kill cells by interfering with intracellular processes6,7. In fact, authors have increasingly pointed out the relationship between cell-penetrating and antimicrobial peptides1,8. A firm understanding of the process of membrane translocation and the relationship between peptide structure and its ability to translocate requires effective, reproducible assays for translocation. Several groups have proposed methods to measure translocation into large unilamellar lipid vesicles (LUVs)9-13. LUVs serve as useful models for bacterial and eukaryotic cell membranes and are frequently used in peptide fluorescent studies14,15. Here, we describe our application of the method first developed by Matsuzaki and co-workers to consider antimicrobial peptides, such as magainin and buforin II16,17. In addition to providing our protocol for this method, we also present a straightforward approach to data analysis that quantifies translocation ability using this assay. The advantages of this translocation assay compared to others are that it has the potential to provide information about the rate of membrane translocation and does not require the addition of a fluorescent label, which can alter peptide properties18, to tryptophan-containing peptides. Briefly, translocation ability into lipid vesicles is measured as a function of the Foster Resonance Energy Transfer (FRET) between native tryptophan residues and dansyl phosphatidylethanolamine when proteins are associated with the external LUV membrane (Figure 1). Cell-penetrating peptides are cleaved as they encounter uninhibited trypsin encapsulated with the LUVs, leading to disassociation from the LUV membrane and a drop in FRET signal. The drop in FRET signal observed for a translocating peptide is significantly greater than that observed for the same peptide when the LUVs contain both trypsin and trypsin inhibitor, or when a peptide that does not spontaneously cross lipid membranes is exposed to trypsin-containing LUVs. This change in fluorescence provides a direct quantification of peptide translocation over time.

This protocol details a method for the quantitative measure of peptide translocation into large unilamellar lipid vesicles. This method also provides information about the rate of membrane translocation and can be used to identify peptides that efficiently and spontaneously cross lipid bilayers.

There is an active interest in peptides that readily cross cell membranes without the assistance of cell membrane receptors1.  Many of these are referred ...

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 ...

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.

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 ...

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 ...

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.

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 ...

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 ...

Isolation and Characterization of Cyanobacterial Extracellular Vesicles

Cyanobacteria are a diverse group of photosynthetic, Gram-negative bacteria that play critical roles in global ecosystems and serve as essential biotechnology models. Recent work has demonstrated that both marine and freshwater cyanobacteria produce extracellular vesicles -&#160;small membrane-bound structures released from the outer surface of the microbes. While vesicles likely contribute to diverse biological processes, their specific functional roles in cyanobacterial biology remain largely unknown. To encourage and advance research in this area, a detailed protocol is presented for isolating, concentrating, and purifying cyanobacterial extracellular vesicles. The current work discusses methodologies that have successfully isolated vesicles from large cultures of Prochlorococcus, Synechococcus, and Synechocystis. Methods for quantifying and characterizing vesicle samples from these strains are presented. Approaches for isolating vesicles from aquatic field samples are also described. Finally, typical challenges encountered with cyanobacterial vesicle purification, methodological considerations for different downstream applications, and the trade-offs between approaches are also discussed.

The present protocol providesdetailed descriptions for isolation, concentration,and characterization of extracellular vesicles from cyanobacterial cultures. Approaches for purifying vesicles from cultures at different scales, trade-offs among methodologies, and considerations for working with field samples are also discussed.