This work was supported by grants from the Mrs. Clifford Elder White Graham Endowed Research Fund &amp; the NIH (AR059838) to CB, from the NIH (AR052791, AR044387-ARRA, &amp; NS063298) to LTT, and by the Baylor College of Medicine Cytometry and Cell Sorting Core with funding from the NIH (NCRR S10RR024574, NIAID AI036211, &amp; NCI P30CA125123). The content is solely the responsibility of the authors and does not necessarily represent the official views of the NIH.

1. 	Cell Staining
<ol>
<li>Prepare the cells. The cells should be used while still actively growing (60- 80% confluence). Serum starve the cells overnight, plate them cells at 0.1 million per well in 0.5 mL serum-free medium in a 24-well plate and place in the incubator (37&deg;C, 5% CO2) for 30 min - 2 hours to recover from trypsinization.</li>
<li>Mix the primary and secondary antibodies. Mix (5 &mu;L of primary anti-GLUT4 antibody with 1 &mu;L secondary chicken anti-goat IgG antibody conjugated to AlexaFluor 488) x number of wells. Incubate for 10 minutes at room temperature, in the dark.</li>
<li>Prepare a 2X working stock of insulin by diluting it in medium.</li>
<li>Check if the cells have adhered or not.</li>
<li>Gently add 6 &mu;L antibody mix and 0.5 mL of your 2X working stock of insulin to the wells containing the cells. Avoid any turbulence of the medium. Incubate (37&deg;C, 5% CO2) for 30 minutes, in the dark.</li>
<li>Fix the cells by adding 0.5 mL of PBS + 1% PFA to each well, without shaking the cells. Incubate for 20 minutes at room temperature, in the dark.</li>
<li>If your cells have adhered to the wells, gently lift them with a cell lifter. Transfer the cells (1 mL total) into tubes that fit in your flow cytometer and centrifuge to pellet the cells.</li>
<li>Wash the pellet twice with 1 mL PBS and resuspend in 0.4 mL PBS + 1% PFA. At this point, cells can be wrapped in foil and stored in the fridge (4-8&deg;C) or taken to the flow cytometer for immediate data acquisition.</li>
</ol>
2. 	Data Acquisition
<ol>
<li>Set a broad gate around the live cells in the side scatter (SSC) versus forward scatter (FCS) panel. Use a negative tube (cells not treated with insulin or cells "stained" with an isotype control) to set your FL1 parameter. Make sure your peak is completely in the panel, not cut on the lower side.</li>
<li>Acquire data from at least 10,000 cells per tube.</li>
</ol>
3. 	Data Analysis
<ol>
<li>Draw a gate around the live cells in the SSC vs FSC panel (Figure 1A). Plot the histograms of AlexaFluor 488 fluorescence intensity versus number of cells within the live-cell gate (Figure 1B). You can overlay different histograms to visualize differences but for quantitation, determine the mean and median fluorescence intensity for each cell population and use these numbers for your comparisons (Figure 1C).</li> 
<li>Normalize the data to the value obtained in the absence of insulin, as shown in Figures 1C and 1D.</li>
</ol>
4. Representative Results
In myoblasts isolated from a healthy individual (no insulin resistance), insulin induces a dose-dependent translocation of GLUT4 from the cytoplasm to the plasma membrane (Figure 1, left). In contrast, insulin does not induce GLUT4 translocation to the plasma membrane of myoblasts isolated from a patient with insulin resistance (Figure 1, right).
<img src="/files/ftp_upload/2429/2429fig1.jpg" alt="Figure 1" /> 
Figure 1. Example of a staining experiment. Myoblasts were isolated from a donor without insulin resistance (left) and from a patient with insulin resistance (right). A, SSC (side scatter) versus FCS (forward scatter) plot with a gate around the live cells. B, Cells gated in A, stained with the antibody against an external epitope of GLUT4, and left unstimulated (red) or stimulated with insulin 1 nM (blue), 10 nM (orange), or 100 nM (green). For figure clarity, we only show data without and with 100 nM insulin for the cells with insulin resistance. C, Mean fluorescence intensities (MFI) of the histograms shown in B were normalized to the MFI of the cells left unstimulated (no insulin). D, Plots of the normalized MFI data from the histograms shown in B.

<ol>
	<li>Zhao, F. Q., Keating, A. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+properties+and+genomics+of+glucose+transporters.">Functional properties and genomics of glucose transporters.</a> Curr. Genomics. 8, 113-128 (2007).</li><li>Karnieli, E., Armoni, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transcriptional+regulation+of+the+insulin-responsive+glucose+transporter+GLUT4+gene:+from+physiology+to+pathology.">Transcriptional regulation of the insulin-responsive glucose transporter GLUT4 gene: from physiology to pathology.</a> Am. J. Endocrinol. Metab. 295, 38-45 (2008).</li><li>Graham, T. E., Kahn, B. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue-specific+alterations+of+glucose+transport+and+molecular+mechanisms+of+intertissue+communication+in+obesity+and+type+2+diabetes.">Tissue-specific alterations of glucose transport and molecular mechanisms of intertissue communication in obesity and type 2 diabetes.</a> Horm. Metab. Res. 39, 717-721 (2007).</li><li>Alter, G., Malenfant, J. M., Altfeld, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CD107a+as+a+functional+marker+for+the+identification+of+natural+killer+cell+activity.">CD107a as a functional marker for the identification of natural killer cell activity.</a> J. Immunol. Methods. 294, 15-22 (2004).</li><li>Vermes, I., Haanen, C., Steffens-Nakken, H., Reutellingsperger, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+assay+for+apoptosis.+Flow+cytometric+detection+of+phosphatidylserine+expression+on+early+apoptotic+cells+using+fluorescein+labeled+Annexin+V.">A novel assay for apoptosis. Flow cytometric detection of phosphatidylserine expression on early apoptotic cells using fluorescein labeled Annexin V.</a> J. Immunol. Methods. 184, 39-51 (1995).</li><li>Lugli, E., Roederer, M., Cossarizza, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Data+analysis+in+flow+cytometry:+the+future+just+started.">Data analysis in flow cytometry: the future just started.</a> Cytometry A. 77, 705-713 (2010).</li></ol>

No conflicts of interest declared.

We have demonstrated a rapid technique for quantifying the translocation of GLUT4 from the cytoplasm to the plasma membrane of cells by flow cytometry.
GLUT4 is only transiently expressed at the plasma membrane of cells and is endocytosed. Since the bound antibodies remain attached to the GLUT4, even during endocytosis, the fluorescence signal gives the total amount of GLUT4 that was exposed at the plasma membrane during the chosen duration of incubation in the presence of insulin.
This technique can also be applied to studying the translocation of other proteins from an intracellular compartment to the plasma membrane, provided a good antibody directed to an external epitope of the protein of interest is available. For example, a similar assay is now commonly used to assess the rate of degranulation of cytotoxic T lymphocytes and natural killer lymphocytes by measuring the translocation of CD107a to their plasma membrane4,5. The translocation of phosphatidylserine from the inner side of the plasma membrane to the outer side during cell apoptosis or necrosis can also be detected by flow cytometry, using fluorophore-labeled Annexin V6.
Besides rapidity of the assay, flow cytometry presents the advantage of allowing the study of protein translocation to the plasma membrane in mixed cell populations. Indeed, one can combine staining for cell subset markers and for the protein of interest, followed by data acquisition on a multi-laser flow cytometer7.
The amounts of antibodies given in paragraph 1.2 of the protocol are a starting point; we recommend you titrate your antibodies with your cells of interest to determine the minimum antibody concentration that gives you a maximum signal. A good starting range for the primary anti-human GLUT4 antibody we have used here would be 1-10 &mu;L per tube (i.e. 0.2-2 &mu;g antibody per tube), each tube containing no more than 1 million cells. For other primary antibodies, refer to the supplier's instructions.
Normalization of data is necessary as autofluorescence of cells will vary from donor to donor and from experiment to experiment.

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		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Anti-human GLUT4 antibodies</td>
<td>&nbsp;</td>
<td>Santa Cruz Biotechnologies</td>
<td>sc-1606</td>
<td>Against an external epitope of GLUT4</td>
</tr>
<tr>
<td>Chicken anti-goat IgG conjugated to AlexaFluor 488</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>A-21467</td>
<td>&nbsp;</td>
<tr>
<td>Recombinant human insulin</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>I2643</td>
<td>&nbsp;</td>
<tr>
<td>Cell lifters</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>29942-118</td>
<td>Cut to fit the wells</td>
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<td>5 ml tubes for FACS</td>
<td>&nbsp;</td>
<td>VWR</td>
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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.

Watch this Scientific Journal Video about Quantitative Measurement of GLUT4 Translocation to the Plasma Membrane by Flow Cytometry at JoVE.com

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

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 ...

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26.4K Views.  Baylor College of Medicine. This protocol describes a rapid technique to quantify the translocation of GLUT4 from the cytoplasm to the plasma membrane of cells by flow cytometry.

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

Measuring Plasma Membrane Protein Endocytic Rates by Reversible Biotinylation

Plasma membrane proteins are a large, diverse group of proteins comprised of receptors, ion channels, transporters and pumps.  Activity of these proteins is responsible for a variety of key cellular events, including nutrient delivery, cellular excitability, and chemical signaling.  Many plasma membrane proteins are dynamically regulated by endocytic trafficking, which modulates protein function by altering protein surface expression.  The mechanisms that facilitate protein endocytosis are complex and are not fully understood for many membrane proteins.  In order to fully understand the mechanisms that control the endocytic trafficking of a given protein, it is critical that the protein s endocytic rate be precisely measured.  For many receptors, direct endocytic rate measurements are frequently achieved utilizing labeled receptor ligands.  However, for many classes of membrane proteins, such as transporters, pumps and ion channels, there is no convenient ligand that can be used to measure the endocytic rate.  In the present report, we describe a reversible biotinylation method that we employ to measure the dopamine transporter (DAT) endocytic rate.  This method provides a straightforward approach to measuring internalization rates, and can be easily employed for trafficking studies of most membrane proteins.

Regulated endocytosis governs the cell surface expression levels of the majority of membrane proteins. Here we utilize reducible, membrane impermeant biotinylation reagents to measure the endocytic rate of the dopamine transporter (DAT), a polytopic membrane protein. The method facilitates a straightforward approach to measuring the endocytic rate of most plasma membrane proteins.

Plasma membrane proteins are a large, diverse group of proteins comprised of receptors, ion channels, transporters and pumps.  Activity of these proteins ...

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.

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 ...

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 ...

Mapping Molecular Diffusion in the Plasma Membrane by Multiple-Target Tracing (MTT)

Our goal is to obtain a comprehensive description of molecular processes occurring at cellular membranes in different biological functions. We aim at characterizing the complex organization and dynamics of the plasma membrane at single-molecule level, by developing analytic tools dedicated to Single-Particle Tracking (SPT) at high density: Multiple-Target Tracing (MTT)1. Single-molecule videomicroscopy, offering millisecond and nanometric resolution1-11, allows a detailed representation of membrane organization12-14 by accurately mapping descriptors such as cell receptors localization, mobility, confinement or interactions.
We revisited SPT, both experimentally and algorithmically. Experimental aspects included optimizing setup and cell labeling, with a particular emphasis on reaching the highest possible labeling density, in order to provide a dynamic snapshot of molecular dynamics as it occurs within the membrane. Algorithmic issues concerned each step used for rebuilding trajectories: peaks detection, estimation and reconnection, addressed by specific tools from image analysis15,16. Implementing deflation after detection allows rescuing peaks initially hidden by neighboring, stronger peaks. Of note, improving detection directly impacts reconnection, by reducing gaps within trajectories. Performances have been evaluated using Monte-Carlo simulations for various labeling density and noise values, which typically represent the two major limitations for parallel measurements at high spatiotemporal resolution.
The nanometric accuracy17 obtained for single molecules, using either successive on/off photoswitching or non-linear optics, can deliver exhaustive observations. This is the basis of nanoscopy methods17 such as STORM18, PALM19,20, RESOLFT21 or STED22,23, which may often require imaging fixed samples. The central task is the detection and estimation of diffraction-limited peaks emanating from single-molecules. Hence, providing adequate assumptions such as handling a constant positional accuracy instead of Brownian motion, MTT is straightforwardly suited for nanoscopic analyses. Furthermore, MTT can fundamentally be used at any scale: not only for molecules, but also for cells or animals, for instance. Hence, MTT is a powerful tracking algorithm that finds applications at molecular and cellular scales.

Mapping Molecular Diffusion in the Plasma Membrane by Multiple-Target Tracing MTT

Multiple-Target Tracing is a homemade algorithm developed for tracking individually labeled molecules within the plasma membrane of living cells. Efficiently detecting, estimating and tracing molecules over time at high-density provide a user-friendly, comprehensive tool to investigate nanoscale membrane dynamics.

Our goal is to obtain a comprehensive description of molecular processes occurring at cellular membranes in different biological functions. We aim at ...

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 ...

Real-time Analyses of Retinol Transport by the Membrane Receptor of Plasma Retinol Binding Protein

Vitamin A is essential for vision and the growth/differentiation of almost all human organs. Plasma retinol binding protein (RBP) is the principle and specific carrier of vitamin A in the blood. Here we describe an optimized technique to produce and purify holo-RBP and two real-time monitoring techniques to study the transport of vitamin A by the high-affinity RBP receptor STRA6. The first technique makes it possible to produce a large quantity of high quality holo-RBP (100%-loaded with retinol) for vitamin A transport assays. High quality RBP is essential for functional assays because misfolded RBP releases vitamin A readily and bacterial contamination in RBP preparation can cause artifacts. Real-time monitoring techniques like electrophysiology have made critical contributions to the studies of membrane transport. The RBP receptor-mediated retinol transport has not been analyzed in real time until recently. The second technique described here is the real-time analysis of STRA6-catalyzed retinol release or loading. The third technique is real-time analysis of STRA6-catalyzed retinol transport from holo-RBP to cellular retinol binding protein I (CRBP-I). These techniques provide high sensitivity and resolution in revealing RBP receptor's vitamin A uptake mechanism.

Here we describe an optimized technique to produce high-quality vitamin A/RBP complex and two real-time monitoring techniques to study vitamin A transport by STRA6, the RBP receptor.

Vitamin A is essential for vision and the growth/differentiation of almost all human organs.  Plasma retinol binding protein (RBP) is the principle and ...

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 ...

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 ...

Sensitive Measurement of Mitophagy by Flow Cytometry Using the pH-dependent Fluorescent Reporter mt-Keima

Mitophagy is a process of selective removal of damaged or unnecessary mitochondria using autophagy machinery. Close links have been found between defective mitophagy and various human diseases, including neurodegenerative diseases, cancer, and metabolic diseases. In addition, recent studies have shown that mitophagy is involved in normal cellular processes, such as differentiation and development. However, the precise role of and molecular mechanisms underlying mitophagy require further study. Therefore, it is critical to develop a robust and convenient method for measuring changes in mitophagy activity. Here, we describe a detailed protocol for quantitatively assessing mitophagy activity through flow cytometry using the mitochondria-targeted fluorescent protein Keima (mt-Keima). This flow cytometry assay can analyze mitophagy activity more rapidly and sensitively than conventional microscopy- or immunoblotting-based methods. This protocol can be applied to analyze mitophagy activity in various cell types.

Mitophagy, the selective degradation of mitochondria, has been implicated in mitochondrial homeostasis and is deregulated in various human diseases. However, convenient experimental methods for measuring mitophagy activity are very limited. Here, we provide a sensitive assay for measuring mitophagy activity using flow cytometry.

Mitophagy is a process of selective removal of damaged or unnecessary mitochondria using autophagy machinery. Close links have been found between ...

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 ...

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

Summary

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Chapters in this video

Measuring Plasma Membrane Protein Endocytic Rates by Reversible Biotinylation

IP-FCM: Immunoprecipitation Detected by Flow Cytometry

Flow Cytometry Purification of Mouse Meiotic Cells

Mapping Molecular Diffusion in the Plasma Membrane by Multiple-Target Tracing (MTT)

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

Real-time Analyses of Retinol Transport by the Membrane Receptor of Plasma Retinol Binding Protein

Immunodetection of Outer Membrane Proteins by Flow Cytometry of Isolated Mitochondria

Measuring Endoreduplication by Flow Cytometry of Isolated Tuber Protoplasts

Sensitive Measurement of Mitophagy by Flow Cytometry Using the pH-dependent Fluorescent Reporter mt-Keima

High-throughput Measurement of Dictyostelium discoideum Macropinocytosis by Flow Cytometry