This work was supported by the Eagles Innovation Award (Fraternal Order of Eagles) and by the Mayo Foundation.

**This video protocol is based on an associated publication 1: High-sensitivity detection and quantitative analysis of native protein-protein interactions and multiprotein complexes by flow cytometry. Adam G. Schrum, Diana Gil, Elaine P. Dopfer, David L. Wiest, Laurence A. Turka, Wolfgang W. A. Schamel, Ed Palmer. Science's STKE 2007 (389): pl2, June 5, 2007, [DOI: 10.1126/stke.3892007pl2]. <a href="http://stke.sciencemag.org/cgi/content/abstract/2007/389/pl2">Please click here to see this publication</a>.
Prior to Starting, Prepare the Following Aqueous Stock Solutions:
<table>
 <tbody>
 <tr>
 <td>MES Coupling Buffer:</td>
 <td>Store at 25&deg;C</td>
 </tr>
 <tr>
 <td>MES, pH 6.0</td>
 <td>50 mM</td>
 </tr>
 <tr>
 <td>EDTA</td>
 <td>1 mM</td>
 </tr>
 <tr>
 <td>Quenching/Blocking/Storage (QBS) Buffer:</td>
 <td>Store at 4&deg;C</td>
 </tr>
 <tr>
 <td>BSA</td>
 <td>1%</td>
 </tr>
 <tr>
 <td>Sodium azide</td>
 <td>0.02%</td>
 </tr>
 <tr>
 <td>PBS, pH 7.4</td>
 <td>1x</td>
 </tr>
 <tr>
 <td>FCM Buffer:</td>
 <td>Store at 4&deg;C</td>
 </tr>
 <tr>
 <td>Tris, pH 7.4</td>
 <td>50 mM</td>
 </tr>
 <tr>
 <td>NaCl</td>
 <td>100 mM</td>
 </tr>
 <tr>
 <td>Sodium azide</td>
 <td>0.02%</td>
 </tr>
 <tr>
 <td>FBS</td>
 <td>5%</td>
 </tr>
 <tr>
 <td>PBS, pH 7.4</td>
 <td>1x</td>
 </tr>
 </tbody>
</table>
Prepare Immediately Prior to Use:
<table>
<tbody>
 <tr>
 <td>EDAC-MES:</td>
 <td>Dissolve 50 mg/mL EDAC powder in MES Coupling Buffer</td>
 </tr>
 <tr>
 <td>Digitonin solution (2% w/v):</td>
 <td>Dissolve digitonin powder in dH2O by heating to 95&deg;C for 5min then cooling on ice</td>
 </tr>
 <tr>
 <td>Lysis Buffer:</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Tris, pH 7.4</td>
 <td>50 mM</td>
 </tr>
 <tr>
 <td>NaCl</td>
 <td>150 Mm</td>
 </tr>
 <tr>
 <td>Digitonin solution </td>
 <td>1%</td>
 </tr>
 <tr>
 <td>Protease Inhibitors</td>
 <td>1x</td>
 </tr>
 <tr>
 <td>Keep on ice</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>
 
1. Coupling of mAb to Beads
<ol>
<li>Determine the concentration of beads from the purchased stock by diluting in PBS and counting with a hemacytometer. Start with a 1:10000 dilution of beads for counting.</li>
<li>Pipette 18 x 106 beads into a 1.5-mL microcentrifuge tube.</li>
<li>Wash the beads 2 to 3 times in 0.5 to 1.0 mL MES Coupling Buffer, centrifuging at 20,000g for 3 minutes at 25&deg;C after each wash.</li>
<li>Resuspend the beads in 50 &mu;L MES Coupling Buffer.</li>
<li>Activate the carboxyl groups on the beads by adding 20 &mu;L of freshly prepared EDAC-MES.</li>
<li>Mix gently for 15 min at 25&deg;C by manually pipetting up and down.</li>
<li>Wash the activated beads 2 to 3 times in 0.5 to 1.0 mL PBS, centrifuging at 20,000g for 3 minutes at 25&deg;C after each wash.</li>
<li>Resuspend the activated beads in 50 &mu;L PBS.</li>
<li>Add 50 &mu;L of the IP mAb (at 0.2-1.0 mg/ mL stock concentration) to the activated bead solution.</li>
<li>Mix for 3 to 4 hours at 25&deg;C by placing the tube on a vibrating shaker. Shake sufficiently to prevent settling of the beads on the bottom of the tube.</li>
<li>Wash mAb-coupled beads 2 to 3 times in 0.5 to 1.0 mL PBS, centrifuging at 20,000g for 3 minutes at 25&deg;C after each wash.</li>
<li>Resuspend the beads in 100 &mu;L QBS Buffer. These can be stored at 4&deg;C.</li>
<li>Suspending the beads well, dilute 1:200-1:10,000 in PBS and count with a hemacytometer. The concentration must be measured for each preparative batch, so that the number of beads used in each IP or pull-down experiment can be precisely controlled.</li>
</ol>
2. Post-nuclear Lysate Preparation and Ip
The lysis method and optimal lysis conditions will depend on the cell type and protein-protein interactions being examined. A number of protocols exist and can be modified for your application.
<ol>
<li>Lyse 20 x 106 'small' cells, such as lymphocytes, or 5 x 106 'large' cells, such as macrophages or tumor cells, in 100 &mu;L fresh Lysis buffer in a 1.5-mL microcentrifuge tube for 20min on ice. Scale the lysis volume as needed.</li>
<li>To remove nuclei and insoluble cellular debris, centrifuge the lysate at 20,000g for 10 min at 4&deg;C. Keep the supernatant and discard the pellet.</li>
<li>Add 0.5 x 105 to 2.5 x 105 mAb-coupled beads to the clarified lysate, pipetting gently to mix. The minimum volume we use to perform a single IP is 5 &mu;L.</li>
<li>Place on a vertical rotating wheel 4 hours to overnight in a cold room. Set to sufficient velocity to prevent the beads from settling. It is acceptable for the liquid of low-volume IPs to remain in the bottom of the tube during rotation.</li>
</ol>
3. Probing of Bead-captured Protein with Fluorochrome-conjugated Antibodies
<ol>
<li>Wash the IP beads two times in 0.2 to 1.0 mL ice-cold FCM Buffer, centrifuging at 20,000g at 4&deg;C for 3 min after each wash.</li>
<li>Resuspend the beads in 100 &mu;L FCM Buffer and aliquot 20 &mu;L into each of five or more 1.5-mL microcentrifuge tubes or wells of a chimney-bottom plate.</li>
<li>Add fluorochrome-conjugated mAbs to the samples. Perform the FCM stain according to the vendor's instructions or according to empirically determined concentrations. As a starting point, add 0.2 to 1 &mu;L of stock antibody solution (at 0.2-1.0 mg/ mL) per tube or well, and incubate for 40 min on ice.</li>
<li>Wash probed beads in 1.5-ml tubes two times in 1.0 mL ice-cold FCM Buffer, centrifuging at 20,000g at 4&deg;C for 3 min after each wash. Gently remove the supernatant after each wash, taking care not to disturb the beads. For a chimney-bottom plate, wash two times with 0.2 mL ice-cold FCM Buffer, centrifuging at 1000g for 5 min at 4&deg;C after each wash. A multichannel pipettor is useful for this step. To remove supernatant from beads in a chimney-bottom plate, 'flick' the plate once, then, while still upside-down, blot any drips from the edges of the wells. The residual volume in each well, containing the beads, will be about 20 &mu;L.</li>
<li>Resuspend the beads in 200 &mu;L FCM Buffer per sample. Transfer to labeled FACS tubes. Samples are now ready for FCM.</li>
</ol>
4. FCM Acquisition
The CML beads described in this protocol are 3 to 5 &mu;m in diameter, approximately half the diameter of a quiescent mouse lymphocyte. Therefore, it may be necessary to manually increase the Forward Scatter (FSC) amp gain and the Side Scatter (SSC) voltage in order for the population of bead events to register on scale. Individual IP beads should form a single tightly clustered population. The settings and gate should be adjusted to exclude bead doublets and debris.
<ol>
<li>Prior to running beads or bead standards, remove the droplet containment sleeve that surrounds the sample inlet on some cytometers. Allow the sheath fluid to drip between samples, as this will clear the inlet and prevent beads from being carried over from one sample to another.</li>
<li>Use unlabeled beads, a negative control of fluorescence, and Rainbow Calibration Particles (RCPs), a positive control of fluorescence, to adjust settings so that both the negative and positive fluorescence extremes appear simultaneously on the log-scale x-axis. This ensures that the fluorescence from the experimental samples will also be on-scale. Note that RCPs are smaller than the CML beads, so FSC and SSC parameters, as well as gate position, may need to be temporarily altered for that sample.</li>
<li>Return to unlabeled-bead settings once calibration with RCPs is complete. If only one fluorescent probe per sample is used, no fluorescence compensation is necessary. In general, we probe multiprotein complexes with a single fluorochrome-conjugated mAb per FCM sample, and we examine multiple interaction partners by staining parallel bead samples with mAbs specific for the various subunits.</li>
<li>Acquire and save fluorescence data files. Analysis can then be performed using flow cytometry software such as CellQuest, FlowJo, or CFlow.</li>
</ol>
5. Representative Results
<img src="/files/ftp_upload/2066/2066fig1.jpg" alt="Figure 1" /> Figure 1. IP-FCM for TCR/CD3 multiprotein complex. T cells from the mouse strain BALB/c were lysed in 1% digitonin, and the lysate was subjected to IP-FCM using anti-CD3&#949; beads. The captured complexes contained significant quantities of TCR-&beta; (purple region) and co-associated CD3-&#949; (green), but close to background levels (defined by the irrelevant immunoglobulin probe, pink trace) of other proteins such as Thy1.2, CD45, or H-2K(d) (brown, orange, and blue traces, respectively). See similar previously published results1-6.

<ol>
	<li>Schrum, A. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-sensitivity+detection+and+quantitative+analysis+of+native+protein-protein+interactions+and+multiprotein+complexes+by+flow+cytometry.">High-sensitivity detection and quantitative analysis of native protein-protein interactions and multiprotein complexes by flow cytometry.</a> Sci STKE. , pl2-pl2 (2007).</li><li>Schrum, A. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualization+of+multiprotein+complexes+by+flow+cytometry.">Visualization of multiprotein complexes by flow cytometry.</a> Curr Protoc Immunol. Chapter 5, 9-9 (2009).</li><li>Teixeiro, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=T+cell+division+and+death+are+segregated+by+mutation+of+TCRbeta+chain+constant+domains.">T cell division and death are segregated by mutation of TCRbeta chain constant domains.</a> Immunity. 21, 515-526 (2004).</li><li>Teixeiro, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Different+T+Cell+receptor+signals+determine+CD8++memory+versus+effector+development.">Different T Cell receptor signals determine CD8+ memory versus effector development.</a> Science. 323, 502-505 (2009).</li><li>Treves, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Junctate+is+a+key+element+in+calcium+entry+induced+by+activation+of+InsP3+receptors+and/or+calcium+store+depletion.">Junctate is a key element in calcium entry induced by activation of InsP3 receptors and/or calcium store depletion.</a> J Cell Biol. 166, 537-548 (2004).</li><li>Gil, D., Schrum, A. G., Alarcon, B., Palmer, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=T+cell+receptor+engagement+by+peptide-MHC+ligands+induces+a+conformational+change+in+the+CD3+complex+of+thymocytes.">T cell receptor engagement by peptide-MHC ligands induces a conformational change in the CD3 complex of thymocytes.</a> J Exp Med. 201, 517-522 (2005).</li></ol>

Information about protein-protein interactions is highly relevant to the analysis of many cellular processes such as signal transduction, lineage maturation, cell-cycle progression, and apoptosis cascades. IP-FCM provides a quick, quantitative, and sensitive way to examine the interaction of proteins and define members of multiprotein complexes in their native conformation. Beads can be coupled and incubated with cell lysates in one day and can be probed and analyzed the next day. A 96-well plate format allows for large numbers of samples to be analyzed at one time to provide efficient data collection for statistical or screening purposes. Using fluorescent bead standards, the number of proteins captured by each bead can be estimated. Very little source material is needed for capture and detection, so limited samples and scarce analytes can still be analyzed for multiple interactions. Although IP-FCM does not require genetic engineering, epitope-tagging, denaturation, or in-vitro mixing of proteins in a non-physiological environment, it can be coupled with these and other techniques, making it a valuable and accessible tool with applicability to many biological systems.
Troubleshooting:
Many IP-FCM experiments generate useful protein interaction data the first time they are attempted. However, optimization of IP-FCM can improve antibody conjugation to beads, protein complex capture, and fluorescent probe binding.
The efficiency of IP antibody conjugation can be determined by probing coupled beads directly with an anti-immunoglobulin antibody. If this efficiency is low, increasing the concentration of antibody during the coupling reaction can allow more IP antibodies to attach to each bead. This can increase the binding capacity of the IP bead batch, resulting in enhanced capture and detection of analytes. Other primary-amine containing molecules (e.g. Tris, bovine serum albumin) should not be present during the coupling reaction, as these can compete with the mAb for bead attachment and result in low mAb coupling. If conjugation of mAb to beads proves problematic for other reasons, beads can instead be coupled to avidin/streptavidin, and biotinylated antibodies can subsequently be non-covalently bound and used for immunprecipitation.
Despite good antibody conjugation, initial detection of bead-associated fluorescence may be low. First, complex capture itself may be low. Because IP-FCM depends on the concentration of analytes, increasing the number of cells lysed per unit lysis volume can increase analyte capture and detection. Additionally, capture may be enhanced by decreasing the number of IP beads incubated with the lysate, which distributes captured complexes across fewer beads, and results in increased average fluorescence per bead when probed. Second, it is possible that access of the probe antibody to the captured complexes is sterically hindered by the immunoprecipitating antibody. In this case, the problem can be addressed by using different antibodies to capture and/or probe.

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<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>carboxylate-modified polystyrene latex (CML) beads (5mm surfactant-free)</td>
<td>Reagent</td>
<td>Interfacial Dynamics Corporation/Molecular Probes</td>
<td>2-5000</td>
<td>Store at 4&deg;C.</td>
</tr>
<tr>
<td>Rainbow Calibration Particles</td>
<td>Reagent</td>
<td>Spherotech, Inc.</td>
<td>RCP-30-5A</td>
<td>Store at 4&deg;C.</td>
</tr>
<tr>
<td>2-[N-Morpholino] ethanesulfonic acid (MES)</td>
<td>Reagent</td>
<td>Sigma</td>
<td>M-5287</td>
<td>&nbsp;</td>
<tr>
<td>1-Ethyl-3-(3-dimethylaminopropl) carbodiimide HCl (EDAC)</td>
<td>Reagent</td>
<td>Pierce Sigma</td>
<td>22980 E-6383</td>
<td>Store at -20&deg;C under desiccating conditions.</td>
</tr>
<tr>
<td>Protease Inhibitor General Use Cocktail, containing AEBSF, Bestatin, Aprotinin, EDTA, E-64, and Leupeptin</td>
<td>Reagent</td>
<td>Sigma</td>
<td>P-2714</td>
<td> Prepare 100x stock solution in H2O. Aliquot and store at -20&deg;C. Handle with gloves.</td>
</tr>
<tr>
<td>PCR plates, 96-well 0.2mL, natural, tall chimney</td>
<td>Reagent</td>
<td>Fisher</td>
<td>08-408-230</td>
<td>For use in place of microcentrifuge tubes during FCM staining</td>
</tr>		</tbody>
		</table>
		</div>

ip-fcm:  immunoprecipitation detected by flow cytometry

Immunoprecipitation detected by flow cytometry (IP-FCM) is an efficient method for detecting and quantifying protein-protein interactions. The basic principle extends that of sandwich ELISA, wherein the captured primary analyte can be detected together with other molecules physically associated within multiprotein complexes. The procedure involves covalent coupling of polystyrene latex microbeads with immunoprecipitating monoclonal antibodies (mAb) specific for a protein of interest, incubating these beads with cell lysates, probing captured protein complexes with fluorochrome-conjugated probes, and analyzing bead-associated fluorescence by flow cytometry. IP-FCM is extremely sensitive, allows analysis of proteins in their native (non-denatured) state, and is amenable to either semi-quantitative or quantitative analysis. As additional advantages, IP-FCM requires no genetic engineering or specialized equipment, other than a flow cytometer, and it can be readily adapted for high-throughput applications.

Watch this Scientific Journal Video about IP-FCM:  Immunoprecipitation Detected by Flow Cytometry at JoVE.com

IP-FCM:  Immunoprecipitation Detected by Flow Cytometry

The IP-FCM method is presented, which allows a sensitive, robust, biochemical assessment of native protein-protein interactions, without requiring genetic engineering or large sample sizes.

Immunoprecipitation detected by flow cytometry (IP-FCM) is an efficient method for detecting and quantifying protein-protein interactions. The basic ...

ip-fcm-immunoprecipitation-detected-by-flow-cytometry

Research

JoVE Journal

Biology

17.4K Views.  Mayo Clinic. The IP-FCM method is presented, which allows a sensitive, robust, biochemical assessment of native protein-protein interactions, without requiring genetic engineering or large sample sizes.

Video: IP-FCM:  Immunoprecipitation Detected by Flow Cytometry

Quantitative Measurement of GLUT4 Translocation to the Plasma Membrane by Flow Cytometry

Glucose is the main source of energy for the body, requiring constant regulation of its blood concentration. Insulin release by the pancreas induces glucose uptake by insulin-sensitive tissues, most notably the brain, skeletal muscle, and adipocytes. Patients suffering from type-2 diabetes and/or obesity often develop insulin resistance and are unable to control their glucose homeostasis. New insights into the mechanisms of insulin resistance may provide new treatment strategies for type-2 diabetes.
The GLUT family of glucose transporters consists of thirteen members distributed on different tissues throughout the body1. Glucose transporter type 4 (GLUT4) is the major transporter that mediates glucose uptake by insulin sensitive tissues, such as the skeletal muscle. Upon binding of insulin to its receptor, vesicles containing GLUT4 translocate from the cytoplasm to the plasma membrane, inducing glucose uptake. Reduced GLUT4 translocation is one of the causes of insulin resistance in type-2 diabetes2,3.
The translocation of GLUT4 from the cytoplasm to the plasma membrane can be visualized by immunocytochemistry, using fluorophore-conjugated GLUT4-specific antibodies.
Here, we describe a technique to quantify total amounts of GLUT4 translocation to the plasma membrane of cells during a chosen duration, using flow cytometry. This protocol is rapid (less than 4 hours, including incubation with insulin) and allows the analysis of as few as 3,000 cells or as many as 1 million cells per condition in a single experiment. It relies on anti-GLUT4 antibodies directed to an external epitope of the transporter that bind to it as soon as it is exposed to the extracellular medium after translocation to the plasma membrane.

This protocol describes a rapid technique to quantify the translocation of GLUT4 from the cytoplasm to the plasma membrane of cells by flow cytometry.

Glucose is the main source of energy for the body, requiring constant regulation of its blood concentration. Insulin release by the pancreas induces ...

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 ...

Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry

A variety of cellular processes, both physiological and pathophysiological, require or are governed by calcium, including exocytosis, mitochondrial function, cell death, cell metabolism and cell migration to name but a few. Cytosolic calcium is normally maintained at low nanomolar concentrations; rather it is found in high micromolar to millimolar concentrations in the endoplasmic reticulum, mitochondrial matrix and the extracellular compartment. Upon stimulation, a transient increase in cytosolic calcium serves to signal downstream events. Detecting changes in cytosolic calcium is normally performed using a live cell imaging set up with calcium binding dyes that exhibit either an increase in fluorescence intensity or a shift in the emission wavelength upon calcium binding. However, a live cell imaging set up is not freely accessible to all researchers. Alternative detection methods have been optimized for immunological cells with flow cytometry and for non-immunological adherent cells with a fluorescence microplate reader. Here, we describe an optimized, simple method for detecting changes in epithelial cells with flow cytometry using a single wavelength calcium binding dye. Adherent renal proximal tubule epithelial cells, which are normally difficult to load with dyes, were loaded with a fluorescent cell permeable calcium binding dye in the presence of probenecid, brought into suspension and calcium signals were monitored before and after addition of thapsigargin, tunicamycin and ionomycin.

Calcium is involved in numerous physiological and pathophysiological signaling pathways. Live cell imaging requires specialized equipment and can be time consuming. A quick, simple method using a flow cytometer to determine relative changes in cytosolic calcium in adherent epithelial cells brought into suspension was optimized.

A variety of cellular processes, both physiological and pathophysiological, require or are governed by calcium, including exocytosis, mitochondrial ...

Immunodetection of Outer Membrane Proteins by Flow Cytometry of Isolated Mitochondria

Methods to detect and monitor mitochondrial outer membrane protein components in animal tissues are vital to study mitochondrial physiology and pathophysiology. This protocol describes a technique where mitochondria isolated from rodent tissue are immunolabeled and analyzed by flow cytometry. Mitochondria are isolated from rodent spinal cords and subjected to a rapid enrichment step so as to remove myelin, a major contaminant of mitochondrial fractions prepared from nervous tissue. Isolated mitochondria are then labeled with an antibody of choice and a fluorescently conjugated secondary antibody. Analysis by flow cytometry verifies the relative purity of mitochondrial preparations by staining with a mitochondrial specific dye, followed by detection and quantification of immunolabeled protein. This technique is rapid, quantifiable and high-throughput, allowing for the analysis of hundreds of thousands of mitochondria per sample. It is applicable to assess novel proteins at the mitochondrial surface under normal physiological conditions as well as the proteins that may become mislocalized to this organelle during pathology. Importantly, this method can be coupled to fluorescent indicator dyes to report on certain activities of mitochondrial subpopulations and is feasible for mitochondria from the central nervous system (brain and spinal cord) as well as liver.

Described here is a method to detect and quantify mitochondrial outer membrane proteins by immunolabeling of mitochondria isolated from rodent tissue and analysis by flow cytometry. This method can be extended to assess functional aspects of mitochondrial subpopulations.

Methods to detect and monitor mitochondrial outer membrane protein components in animal tissues are vital to study mitochondrial physiology and ...

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 ...

Measuring Endoreduplication by Flow Cytometry of Isolated Tuber Protoplasts

Endoreduplication, the replication of a cell's nuclear genome without subsequent cytokinesis, yields cells with increased DNA content and is associated with specialization, development and increase in cellular size. In plants, endoreduplication seems to facilitate the growth and expansion of certain tissues and organs. Among them is the tuber of potato (Solanum tuberosum), which undergoes considerable cellular expansion in fulfilling its function of carbohydrate storage. Thus, endoreduplication may play an important role in how tubers are able to accommodate this abundance of carbon. However, the cellular debris resulting from crude nuclear isolation methods of tubers, methods that can be used effectively with leaves, precludes the estimation of the tuber endoreduplication index (EI). This article presents a technique for assessing tuber endoreduplication through the isolation of protoplasts while demonstrating representative results obtained from different genotypes and compartmentalized tuber tissues. The major limitations of the protocol are the time and reagent costs required for sample preparation as well as relatively short lifespan of samples after lysis of protoplasts. While the protocol is sensitive to technical variation, it represents an improvement over traditional methods of nuclear isolation from these large specialized cells. Possibilities for improvements to the protocol such as recycling enzyme, the use of fixatives, and other alterations are proposed.

The protocol described herein is a method for measuring endoreduplication within tubers of potato (Solanum tuberosum). It includes plasmolysis and protoplast extraction steps to decrease the noise and debris in downstream flow cytometric analysis.

Endoreduplication, the replication of a cell's nuclear genome without subsequent cytokinesis, yields cells with increased DNA content and is associated ...

High-throughput Measurement of Dictyostelium discoideum Macropinocytosis by Flow Cytometry

Large-scale non-specific fluid uptake by macropinocytosis is important for the proliferation of certain cancer cells, antigen sampling, host cell invasion and the spread of neurodegenerative diseases. The commonly used laboratory strains of the amoeba Dictyostelium discoideum have extremely high fluid uptake rates when grown in nutrient medium, over 90% of which is due to macropinocytosis. In addition, many of the known core components of mammalian macropinocytosis are also present, making it an excellent model system for studying macropinocytosis. Here, the standard technique to measure internalized fluid using fluorescent dextran as a label is adapted to a 96-well plate format, with the samples analyzed by flow cytometry using a high-throughput sampling (HTS) attachment.
Cells are fed non-quenchable fluorescent dextran for a pre-determined length of time, washed by immersion in ice-cold buffer and detached using 5 mM sodium azide, which also stops exocytosis. Cells in each well are then analyzed by flow cytometry. The method can also be adapted to measure membrane uptake and phagocytosis of fluorescent beads or bacteria.
This method was designed to allow measurement of fluid uptake by Dictyostelium in a high-throughput, labor and resource efficient manner. It allows simultaneous comparison of multiple strains (e.g. knockout mutants of a gene) and conditions (e.g. cells in different media or treated with different concentrations of inhibitor) in parallel and simplifies time-courses.

High-throughput Measurement of Dictyostelium discoideum Macropinocytosis by Flow Cytometry

Macropinocytosis, large-scale non-specific fluid uptake, is important in many areas of clinical biology including immunology, infection, cancer, and neurodegenerative diseases. Here, existing techniques have been adapted to allow high-throughput, single-cell resolution measurement of macropinocytosis in the macropinocytosis model organism Dictyostelium discoideum using flow cytometry.

Large-scale non-specific fluid uptake by macropinocytosis is important for the proliferation of certain cancer cells, antigen sampling, host cell invasion ...

Detecting and Characterizing Protein Self-Assembly In Vivo by Flow Cytometry

Protein self-assembly governs protein function and compartmentalizes cellular processes in space and time. Current methods to study it suffer from low-sensitivity, indirect read-outs, limited throughput, and/or population-level rather than single-cell resolution. We designed a flow cytometry-based single methodology that addresses all of these limitations: Distributed Amphifluoric FRET or DAmFRET. DAmFRET detects and quantifies protein self-assemblies by sensitized emission FRET in vivo, enables deployment across model systems-from yeast to human cells-and achieves sensitive, single-cell, high-throughput read-outs irrespective of protein localization or solubility.

This article describes a FRET-based flow cytometry protocol to quantify protein self-assembly in both S. cerevisiae and HEK293T cells.