Research in the laboratories is funded by the Fritz-Bender Foundation, Munich, Germany (to T.D.) and a University of Witten/Herdecke internal research grant (to W.-K.L). We would like to thank Prof Dr. Dr. Frank Th&#233;venod (Institute for Physiology, Pathophysiology &#38; Toxicology, University of Witten/Herdecke) for helpful suggestions.

1. Preparation of Reagents and Solutions
<ol>
	<li>Prepare a modified Hank&#8217;s balanced salt solution (HBSS) (in mM: 136.9 NaCl, 5.37 KCl, 0.34 Na2HPO4, 0.44 KH2PO4, 4.17 NaHCO3, pH 7.2, no phenol red). Store at 4 &#176;C.</li>
	<li>Make 1.5 M CaCl2 and 2 M 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (pH 7.2) stock solutions using distilled water.</li>
	<li>Prepare a 250 mM stock solution of probenecid. For the free acid form (often referred to as water insoluble), dissolve probenecid powder in 1 N NaOH, fill to three quarters of the final volume with distilled water and slowly titrate to pH 7.4 with 1 N HCl. Store at 4 &#176;C.</li>
	<li>Dissolve fluorescent membrane permeable calcium binding dye in anhydrous dimethyl sulfoxide (DMSO) to a stock concentration of 1 mM. Store at -20 &#176;C in the dark.</li>
	<li>Prior to each experiment, freshly prepare probenecid-containing HBSS flux (PHF) buffer by combining modified HBSS with (in final concentrations) 1.5 mM CaCl2, 10 mM HEPES, pH 7.2, 5% fetal bovine serum, 2 mM probenecid, and 5.55 mM glucose. Warm to 37 &#176;C.</li>
</ol>
2. Cell Culture
<ol>
	<li>Plate 2.5 x 105 kidney proximal tubule cells (WKPT-0293 Cl.2) per well of a 6 well plate or 35 mm dish in standard culture medium and grow for 2 days reaching a confluency of approximately 70%.</li>
</ol>
3. Calcium Dye Loading
<ol>
	<li>For each well or dish, mix 1 ml culture medium containing 5% fetal bovine serum, 4 &#181;M cell permeable calcium binding dye and 2 mM probenecid.</li>
	<li>Replace the culture medium with 1 ml loading dye mixture in each well and incubate at 37 &#176;C for 30 min in a humidified incubator with 5% CO2. Cover the plate with aluminium foil to prevent bleaching of the fluorescent indicator.</li>
</ol>
4. Preparation of Cells for Flow Cytometry
<ol>
	<li>Warm 0.05% trypsin-ethylenediaminetetraacetic acid (EDTA) solution to 37 &#176;C.</li>
	<li>Aspirate calcium loading dye solution from each well and add 1 ml trypsin along the wall of the well.</li>
	<li>Incubate at 37 &#176;C until all cells are detached.</li>
	<li>Transfer cells to a 1.5 ml tube and pellet by centrifuging in a microcentrifuge at 10,000 g for 1 min.</li>
	<li>Carefully aspirate the supernatant without disturbing the pellet.</li>
	<li>Wash cells by adding 1 ml PHF buffer and resuspend by pipetting up and down.</li>
	<li>Pellet cells by centrifuging at 10,000 g for 1 min.</li>
	<li>Repeat steps 4.6. and 4.7. a further two times.</li>
	<li>Resuspend cells in a final volume of 500 &#181;l PHF buffer and use the cells for experimentation within 1 hr.</li>
	<li>Optional: count cells to determine the cell concentration and use 5 x 104 &#8211; 1 x 105 cells per measurement.</li>
</ol>
5. Flow Cytometer Setup
<ol>
	<li>Start the flow cytometer, open the fluidics drawer and flip the vent valve toggle switch to increase the air pressure of the sheath fluid tank.</li>
	<li>&#8220;Prime&#8221; the instrument to remove potential residing air bubbles from within the inner tubes and the flow chamber.</li>
	<li>In the flow cytometry software, open two dot plot windows.
	<ol>
		<li>The first dot plot requires forward scattered light (FSC) for the X parameter and side scattered light (SSC) for the Y parameter to control for the quality of the cells within the sample through their size and their granularity, respectively.</li>
		<li>The second dot plot measures the fluorescence intensity of the calcium-bound dye. Select the appropriate fluorescence detection channel for the Y parameter using a logarithmic scale and time in seconds for the X parameter on a linear scale.</li>
	</ol>
	</li>
	<li>Set the flow rate to &#8216;high&#8217;.</li>
</ol>
6. Flow Cytometry Measurement
<ol>
	<li>Perform the measurements at RT. In a flow cytometry sample tube (11.5 x 75 mm), add 480 &#181;l PHF buffer.</li>
	<li>Add 20 &#181;l cell suspension and briefly vortex to mix.</li>
	<li>Move the tube support arm to the side and position the sample tube over the sample injection tube until a tight seal is formed. Return the tube support arm to its original centered position.</li>
	<li>Optimize measurement by adjusting the fluorescence channel such that the mean fluorescence intensity of the sample is at approximately 102 on the y-axis. Save and use for subsequent experiments.</li>
	<li>To start an experiment, repeat steps 6.1 to 6.3.</li>
	<li>Record baseline fluorescence for 50 sec.</li>
	<li>Pause measurement, remove the sample tube, add compound of interest, vortex briefly and return to the sample injection tube. This step should not take more than 15 sec.</li>
	<li>Resume measurement and continue for a total of 204.8 sec.</li>
	<li>Use a calcium ionophore, such as ionomycin (10 &#181;M), and a calcium chelator, such as ethylene glycol tetraacetic acid (EGTA), MnCl2 or CoCl2 (2 mM), for positive and negative controls, respectively.</li>
</ol>
7. Data Analysis
<ol>
	<li>Analyze data using dedicated flow cytometry software for offline analysis, such as WinMDI (Windows Multiple Document Interface for Flow Cytometry), a free software package developed by Joe Trotter at The Scripps Institute, Flow Cytometry Core Facility.</li>
	<li>Open a dot plot with SSC and FSC parameters.</li>
	<li>Select the region of cells for analysis using the &#8220;Regions&#8221; tool (right-click on the image) to exclude dead cells or cell debris found in the bottom left corner from the data analysis (Figure 1).</li>
	<li>Quadrant analysis (Figure 2)
	<ol>
		<li>Open a density plot with time on the x-axis and fluorescence intensity on the y-axis. Use all events.</li>
		<li>Use the same gating region as in 7.3.</li>
		<li>Quantify the data by using the &#8220;Quadrant&#8221; tool.</li>
		<li>Adjust the quadrant separators so that the vertical line is placed at the time of compound addition and the horizontal line is placed at the center of the baseline fluorescence in the controls. Ensure that the positioning of the quadrant separators is equal in all samples.</li>
		<li>The percentage of cells in each quadrant is displayed in each corner (Figure 2). Compare the upper right quadrant (UR) between samples.</li>
	</ol>
	</li>
	<li>Histogram analysis (Figure 3)
	<ol>
		<li>Open the control data in a histogram window with fluorescence intensity on the x-axis and events on the y-axis. Use all events.</li>
		<li>For a pictorial overview, add the test data to be compared to the control plot as overlay plots (go to File &#8594; Open File &#8594; Select file &#8594; Overlay).</li>
		<li>For quantitative analysis, single histogram windows must be used. Gate the cell population using the region R1 from step 7.3.</li>
		<li>In the non-treated control, use the marker function (&#8220;Markers&#8221;) to mark the area for analysis starting from the midpoint between the minimum and maximum fluorescence of the control peak and ending at the maximum fluorescence intensity. The percentage of cells in this area is calculated.</li>
		<li>Retrieve data from the statistics window. Repeat for remaining data files using the same marked area for analysis.</li>
	</ol>
	</li>
	<li>Fluorescence intensity analysis (Figure 4)
	<ol>
		<li>Open a dot plot with time on the x-axis and fluorescence intensity on the y-axis. Use all events.</li>
		<li>Enable &#8220;Kinetics Mode&#8221; and &#8220;Overlay Kinetics Line&#8221;. A blue line will appear in the dot plot representing the mean fluorescence intensity.</li>
		<li>Quantify the relative calcium influx by creating multiple rectangular regions. Each region should have a width of about &#8220;50&#8221;, which is equivalent to &#8220;10 sec&#8221;. Define up to 15 rectangular regions.</li>
		<li>Retrieve quantification from the statistics window and copy into a spreadsheet.</li>
		<li>Use the y-mean data for analysis. Calculate the mean of R2 to R6, which is equivalent to 20 to 50 sec. This is the baseline value.</li>
		<li>Choose an appropriate time interval for calcium influx analysis (depends on onset, duration and intensity).</li>
		<li>Calculate the calcium signals from each region within this time interval relative to the mean baseline value, that is, the fluorescence signal before compound addition, which is set to 100%. Determine the mean and standard deviation of the regions post compound addition; this is the mean calcium signal evoked by the compound.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Calcium is a key event in multiple cellular processes acting as a second messenger signaling molecule and also as a means to communicate between the ER and mitochondria. The ability to measure changes in free cytosolic calcium concentrations is a useful technique that can apply to all areas of cell biology. There are various methods to detect cytosolic calcium signals. Classically, signals from a ratiometric calcium indicator, such as Fura-2, are monitored in live cells using a fluorescence imaging setup and absolute cal...

Calcium is an important second messenger responsible for the transmission of cellular physiological and pathophysiological signals1. Cytosolic calcium concentrations are kept low through the activity of calcium pumps and calcium exchangers. The sarcoplasmic/endoplasmic reticulum calcium ATPase (SERCA) refills the endoplasmic reticulum (ER) calcium store as part of a &#8220;pump leak&#8221; system whereas the plasma membrane calcium ATPase (PMCA) extrudes calcium to the extracellular compartment, both in ATP-dependent manners2. Physiological messengers, such as hormones or neurotransmitters, transmit their signals through G-protein coupled receptors, activation of phospholipase C, which hydrolyses phosphatidylinositol 4,5-bisphosphate (PIP2) in the plasma membrane to generate diacylglycerol and inositol triphosphate (IP3)3. While diacylglycerol remains in the plasma membrane, IP3 diffuses into the cytosol and binds to IP3 receptors (IP3Rs), which are ligand activated calcium channels, found in the ER membrane and calcium is released from the ER luminal store culminating in an increase in cytosolic calcium concentrations4. An alternative route for calcium to reach the cytosol is through calcium channels and exchangers present in the plasma membrane. Crosstalk between these two compartments have been described: calcium induced calcium release (CICR) where extracellular calcium induces calcium release from ER stores5, and store operated calcium entry (SOCE) where emptying of the ER stores is sensed by STIM and causes opening of Orai calcium channels in the plasma membrane to promote refilling of the ER stores6.
Under pathophysiological conditions, the cellular calcium response is deregulated and cytosolic calcium increases in the absence of physiological stimulators1. The calcium response may be affected in a number of ways: prolonged calcium signal, delayed calcium removal from the cytosol, depletion of calcium stores or localized changes in calcium. Furthermore, the mitochondria take on a central role in buffering and releasing calcium7. Prolonged and/or supramaximal cytosolic calcium increase leads to cell death. In fact, calcium is more often than not involved in the cellular pathophysiological response and is a key event in a wide array of diseases, such as neurodegenerative disease, heart disease, cancer, bone disease and kidney disease, which are mainly associated with cell death and loss or alteration of organ or tissue function8-10. Moreover, perturbation of calcium signals has been linked to cellular adaptation and changes in cellular functions and responses.
Traditionally, calcium signals are measured with a negatively charged fluorescent calcium indicator that is loaded into cells in a cell permeable acetoxymethyl (AM) ester form. Once cleaved by cellular esterases, the fluorescent indicators remain within the intracellular compartment and increase in fluorescence intensity when calcium is bound. The most well known and used calcium indicator is the ratiometric Fura-2 developed by Roger Tsien and colleagues11. Calcium indicators with differing affinities for calcium allow different calcium pools to be monitored. Detection methods include live cell imaging, fluorescence microplate reader and flow cytometry. The relatively fast kinetics of calcium signals (usually within a few seconds to minutes) makes live cell imaging the best method for gaining the most information about characteristics of the calcium signal. Aside from the fast kinetics of calcium signals (within milliseconds), live cell imaging is a better method for studying cellular compartmentalization of calcium signaling within a single cell 12. Absolute calcium concentrations can be calculated by determining the minimum and maximum fluorescence of the calcium indicator by adding a calcium chelator and permeabilizing the cells, respectively, as described by Grynkiewicz et al.11.
The use of flow cytometry to measure calcium signals was first developed in the 1980s in immunological cells where the opportunity to measure multiple parameters, such as membrane integrity and separation of cell population, and the requirement of cells in suspension combined with the development of single wavelength calcium indicators made flow cytometry an ideal and convenientdetection method13-15. In adherent cells, live cell imaging provides the most information about calcium signaling, most importantly the kinetics, but requires a complicated setup comprising a fluorescence microscope, a perfusion system, maintenance of the cellular environment, such as temperature, and specialized microscope software for acquiring and analyzing data. Alternative methods such as a fluorescence microplate reader or pharmacological means through use of calcium chelators are incomparable in terms of gained information. Flow cytometry is becoming more widely used to monitor cytosolic calcium in adherent cells though, in the majority of studies, only end-point measurements are performed. Using the method initially developed by Gergely et al. in immunological B cells16, changes in cytosolic free calcium concentration by flow cytometry with real-time kinetics and monitoring the fluorescence intensity in individual cells from a sample population have been successfully measured in cancer epithelial cells and cancer stem cell hybrids17. This method was further adapted for use in adherent epithelial cells, which are difficult to load with calcium indicators 18.
Here, using an immortalized adherent epithelial cell line derived from the S1 segment of the rat kidney proximal tubule (WKPT-0293 Cl.2)19, we describe an optimized simplified method for measurement of changes in cytosolic calcium concentration by flow cytometry. Since many epithelial cells possess a number of organic transporters for anionic compounds, loading of cells with a calcium indicator can prove to be a challenge. To prevent the efflux of calcium indicators, probenecid, the prototypical inhibitor of organic anion transporters and originally developed to decrease renal excretion of penicillin20, is applied simultaneously in the incubation medium. The cells are maintained in monolayers for calcium indicator loading, brought into suspension and calcium signals can be detected immediately after compound addition.

<table border="0" cellpadding="0" cellspacing="0" width="735">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Name of Reagent/ Equipment</td>
			<td style="width:79px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:321px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Fluo-3, AM</td>
			<td style="width:79px;">Invitrogen</td>
			<td style="width:119px;">F-1241&#160;</td>
			<td>Cell permeable</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Probenecid</td>
			<td style="width:79px;">Sigma-Aldrich</td>
			<td style="width:119px;">P-8761</td>
			<td>Water insoluble, dissolve in 1 N NaOH</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">0.05% Trypsin-EDTA (1X)</td>
			<td style="width:79px;">Invitrogen</td>
			<td style="width:119px;">25300-062</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Ionomycin</td>
			<td style="width:79px;">Sigma-Aldrich</td>
			<td style="width:119px;">I9657</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Thapsigargin</td>
			<td style="width:79px;">Tocris Bioscience</td>
			<td style="width:119px;">1138</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Tunicamycin</td>
			<td style="width:79px;">Sigma-Aldrich</td>
			<td style="width:119px;">T7765</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">FACSCalibur Flow Cytometer + CELLQuest software</td>
			<td style="width:79px;">Becton Dickinson</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="105">
			<td height="105" style="height:105px;width:216px;">Windows Multiple Document Interface for Flow Cytometry (WinMDI)</td>
			<td style="width:79px;">Joe Trotter, The Scripps Institute</td>
			<td style="width:119px;"></td>
			<td style="width:321px;">Please note that this is an older 16-bit application that reads FCS 2.0 compliant files but does not recognize FCS 3.0 digital data.</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:216px;">WKPT-0293 Cl.2 rat kidney proximal tubule cell line</td>
			<td style="width:79px;"></td>
			<td style="width:119px;"></td>
			<td style="width:321px;">Made available by Dr. Ulrich Hopfer (Department of Physiology &#38; Biophysics, Case 
			Western Reserve University, Cleveland, OH)</td>
		</tr>
	</tbody>
</table>

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.

To increase cytosolic calcium, two pharmacological compounds were applied to renal proximal tubule cells (WKPT-0293 Cl.2) loaded with cell permeable calcium binding dye, Fluo-3-AM. Nontreated control samples were loaded with calcium binding dye in the presence of probenecid and were mixed but without the addition of any compounds. Thapsigargin (TG) is a classical inhibitor of the SERCA pump leading to leakage out of the ER and resulting in increased cytosolic calcium. Tunicamycin (TUN) blocks N-glycosylation of proteins ...

Watch this Scientific Journal Video about Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry at JoVE.com

Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry

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 ...

cytosolic-calcium-measurements-renal-epithelial-cells-flow

Research

JoVE Journal

Biology

15.2K 次观看.  University of Witten/Herdecke. 钙是参与许多生理学和病理生理学的信号通路。活细胞成像，需要专门的设备和非常耗时。使用流式细胞仪来确定在纳入悬浮贴壁的上皮细胞内钙的相对变化快速，简单的方法进行了优化。 

视频: Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry

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 ...

Chromatin Isolation by RNA Purification (ChIRP)

Long noncoding RNAs are key regulators of chromatin states for important biological processes such as dosage compensation, imprinting, and developmental gene expression 1,2,3,4,5,6,7. The recent discovery of thousands of lncRNAs in association with specific chromatin modification complexes, such as Polycomb Repressive Complex 2 (PRC2) that mediates histone H3 lysine 27 trimethylation (H3K27me3), suggests broad roles for numerous lncRNAs in managing chromatin states in a gene-specific fashion 8,9. While some lncRNAs are thought to work in cis on neighboring genes, other lncRNAs work in trans to regulate distantly located genes. For instance, Drosophila lncRNAs roX1 and roX2 bind numerous regions on the X chromosome of male cells, and are critical for dosage compensation 10,11. However, the exact locations of their binding sites are not known at high resolution. Similarly, human lncRNA HOTAIR can affect PRC2 occupancy on hundreds of genes genome-wide 3,12,13, but how specificity is achieved is unclear. LncRNAs can also serve as modular scaffolds to recruit the assembly of multiple protein complexes. The classic trans-acting RNA scaffold is the TERC RNA that serves as the template and scaffold for the telomerase complex 14; HOTAIR can also serve as a scaffold for PRC2 and a H3K4 demethylase complex 13.
Prior studies mapping RNA occupancy at chromatin have revealed substantial insights 15,16, but only at a single gene locus at a time. The occupancy sites of most lncRNAs are not known, and the roles of lncRNAs in chromatin regulation have been mostly inferred from the indirect effects of lncRNA perturbation. Just as chromatin immunoprecipitation followed by microarray or deep sequencing (ChIP-chip or ChIP-seq, respectively) has greatly improved our understanding of protein-DNA interactions on a genomic scale, here we illustrate a recently published strategy to map long RNA occupancy genome-wide at high resolution 17. This method, Chromatin Isolation by RNA Purification (ChIRP) (Figure 1), is based on affinity capture of target lncRNA:chromatin complex by tiling antisense-oligos, which then generates a map of genomic binding sites at a resolution of several hundred bases with high sensitivity and low background. ChIRP is applicable to many lncRNAs because the design of affinity-probes is straightforward given the RNA sequence and requires no knowledge of the RNA's structure or functional domains.

Chromatin Isolation by RNA Purification ChIRP

ChIRP is a novel and rapid technique to map genomic binding sites of long noncoding RNAs (lncRNAs). The method takes advantage of the specificity of anti-sense tiling oligonucleotides to allow the enumeration of lncRNA-bound genomic sites.

Long noncoding RNAs are key regulators of chromatin states for important biological processes such as dosage compensation, imprinting, and developmental ...

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 ...

High Efficiency Differentiation of Human Pluripotent Stem Cells to Cardiomyocytes and Characterization by Flow Cytometry

There is an urgent need to develop approaches for repairing the damaged heart, discovering new therapeutic drugs that do not have toxic effects on the heart, and improving strategies to accurately model heart disease. The potential of exploiting human induced pluripotent stem cell (hiPSC) technology to generate cardiac muscle &#8220;in a dish&#8221; for these applications continues to generate high enthusiasm. In recent years, the ability to efficiently generate cardiomyogenic cells from human pluripotent stem cells (hPSCs) has greatly improved, offering us new opportunities to model very early stages of human cardiac development not otherwise accessible. In contrast to many previous methods, the cardiomyocyte differentiation protocol described here does not require cell aggregation or the addition of Activin A or BMP4 and robustly generates cultures of cells that are highly positive for cardiac troponin I and T (TNNI3, TNNT2), iroquois-class homeodomain protein IRX-4 (IRX4), myosin regulatory light chain 2, ventricular/cardiac muscle isoform (MLC2v) and myosin regulatory light chain 2, atrial isoform (MLC2a) by day 10 across all human embryonic stem cell (hESC) and hiPSC lines tested to date. Cells can be passaged and maintained for more than 90 days in culture. The strategy is technically simple to implement and cost-effective. Characterization of cardiomyocytes derived from pluripotent cells often includes the analysis of reference markers, both at the mRNA and protein level. For protein analysis, flow cytometry is a powerful analytical tool for assessing quality of cells in culture and determining subpopulation homogeneity. However, technical variation in sample preparation can significantly affect quality of flow cytometry data. Thus, standardization of staining protocols should facilitate comparisons among various differentiation strategies. Accordingly, optimized staining protocols for the analysis of IRX4, MLC2v, MLC2a, TNNI3, and TNNT2 by flow cytometry are described.

The article describes the detailed methodology to efficiently differentiate human pluripotent stem cells into cardiomyocytes by selectively modulating the Wnt pathway, followed by flow cytometry analysis of reference markers to assess homogeneity and identity of the population.

There is an urgent need to develop approaches for repairing the damaged heart, discovering new therapeutic drugs that do not have toxic effects on 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.

Protein self-assembly governs protein function and compartmentalizes cellular processes in space and time. Current methods to study it suffer from ...

Measurement of Mitochondrial Mass and Membrane Potential in Hematopoietic Stem Cells and T-cells by Flow Cytometry

A fine balance of quiescence, self-renewal, and differentiation is key to preserve the hematopoietic stem cell (HSC) pool and maintain lifelong production of all mature blood cells. In recent years cellular metabolism has emerged as a crucial regulator of HSC function and fate. We have previously demonstrated that modulation of mitochondrial metabolism influences HSC fate. Specifically, by chemically uncoupling the electron transport chain we were able to maintain HSC function in culture conditions that normally induce rapid differentiation. However, limiting HSC numbers often precludes the use of standard assays to measure HSC metabolism and therefore predict their function. Here, we report a simple flow cytometry assay that allows reliable measurement of mitochondrial membrane potential and mitochondrial mass in scarce cells such as HSCs. We discuss the isolation of HSCs from mouse bone marrow and measurement of mitochondrial mass and membrane potential post ex vivo culture. As an example, we show the modulation of these parameters in HSCs via treatment with a metabolic modulator. Moreover, we extend the application of this methodology on human peripheral blood-derived T cells and human tumor infiltrating lymphocytes (TILs), showing dramatic differences in their mitochondrial profiles, possibly reflecting different T cell functionality. We believe this assay can be employed in screenings to identify modulators of mitochondrial metabolism in various cell types in different contexts.

Here we describe a reliable method to measure mitochondrial mass and membrane potential in ex vivo cultured hematopoietic stem cells and T cells.

A fine balance of quiescence, self-renewal, and differentiation is key to preserve the hematopoietic stem cell (HSC) pool and maintain lifelong production ...