NTK would like to acknowledge the University of Miami Research Awards in Natural Sciences and Engineering for funding this research. BR would like to thank Alex and Ann Lauterbach for funding the Comparative and Evolutionary Immunology Laboratory.&#160;The work of BR was supported by Israel Science Foundation (ISF) numbers: 1416/19 and 2841/19, and HFSP Research Grant, RGY0085/2019. We would like to thank Zhanna Kozhekbaeva and Mike Connelly for technical assistance. We would also like to thank the University of Miami, Miller School of Medicine&#8217;s Flow Cytometry Shared Resource at the Sylvester Comprehensive Cancer Center for access to the FACS cytometer and to Shannon Saigh for technical support.

1. Dissociation of tissues from coral skeleton via airbrush and compressor
NOTE: Perform steps on ice and protect hands with gloves.
<ol>
	<li>Assemble the airbrush kit by connecting the air compressor, hose, and airbrush (Figure 2). Set the pressure gauge between 276&#8722;483 kPa. 
	NOTE: The recommended compressor and air hose used for this study was preset to a maximum pressure of 393 kPa. Use outside of this 276-483 kPa range may result in either inadequate cell removal from the skeleton or rupture of the cell membranes. This range may differ for each species and may require a viability test. A viability test may be performed with a subset of the cell slurry with a simple hemocytometer and a 4&#8242;,6-diamidino-2-phenylindole (DAPI) dye exclusion assay, visualized on a compound microscope.</li>
	<li>Prepare 100 mL of cell staining media in an autoclaved, sterile glass container by adding 33 mL of 10x phosphate-buffered saline (PBS; without calcium and magnesium) to a final concentration of 3.3x, 2 mL of fetal calf serum (FCS; heat inactivated at 56 &#176;C for 30 min) to 2% of total volume, and 2 mL of 1 M HEPES buffer to a final concentration of 20 mM. Then top off at 100 mL with sterile water. 
	NOTE: The 3.3x PBS concentration was chosen to mimic the salinity of seawater, as demonstrated in Rosental et al.18. This concentration needs to be adjusted for species found in other environments to mimic their salinity.</li>
	<li>Transfer 10 mL of cell staining media to an airbrush bottle (labeled &#34;F&#34; in Figure 2) and attach the adaptor lid for the airbrush connector. 
	NOTE: Larger fragments and species with thicker tissue layers may require more media for adequate tissue removal.</li>
	<li>For the branching coral species demonstrated here, use a bone cutter to clip or cut a coral fragment approximately 3&#8722;5 cm in length or 4 cm2 in area. 
	NOTE: The fragment must be clear of macroalgae and other multicellular organisms because these may not be removed from the sample downstream and will contaminate the sample during the FACS analysis and sorting processes. A 1 cm fragment of P. damicornis gives approximately 2&#8722;10 x 106 cells after filtering, staining, and washing.</li>
	<li>Place the coral fragment inside a plastic collection bag and use the airbrush to spray the cell staining media onto the coral (Figure 2). Continue this process until most of the skeleton is exposed. Remove the remaining skeleton from the collection bag.</li>
</ol>
2. Dissociation of cells from coral tissue
NOTE: Perform all steps on ice and protect hands with gloves.
<ol>
	<li>Filter the cell slurry through a 40 &#181;m nylon cell strainer into a 50 mL centrifuge tube in order to obtain a single cell suspension. 
	NOTE: Different cell strainer mesh sizes may be more appropriate for different species. However, larger sizes may result in inadequate tissue dissociation due to the passage of debris and cell clumps.
	<ol>
		<li>For specimens with thicker mucus layers, use the sterilized plunger of a 1 mL plastic syringe and gently grind it against the filter to break up clumps and help the cells pass through the strainer. Rinse the strainer and plunger with cell staining media into the centrifuge tube.</li>
	</ol>
	</li>
	<li>Pellet the cells by centrifugation at 4 &#176;C at 450 x g for 5 min. Remove the supernatant and resuspend the cells in 1 mL of cell staining media.</li>
</ol>
3. Cell staining
NOTE: Perform all steps on ice and protect hands with gloves. Stains featured in this protocol are for representation purposes. Alternative stains will require different concentrations and incubation times.
<ol>
	<li>Transfer 500 &#181;L of resuspended cells to a 5 mL round-bottom tube and set aside as a control sample. Do not add stain to this aliquot.</li>
	<li>To the remaining cell suspension, add 0.42 &#181;L of 12 mM DAPI viability dye (Table of Materials), 1 &#181;L of 5 &#181;M reactive oxygen species (ROS) stain (Table of Materials), and 0.1 &#181;L of 0.2 &#181;M lysosome stain (Table of Materials). Pipette well to mix and incubate on ice for 30 min in the dark to prevent photobleaching. 
	NOTE: DAPI is used in this example to work as a DNA stain and viability dye for the differential gating of dead cells on the FACS machine. This assumes that DAPI penetrates dying cells due to membrane integrity. The ROS stain emits light in the 520 nm range (green), while the lysosomal marker emits light at approximately 668 nm (red).</li>
	<li>Pellet 500 &#181;L of the stained cells by centrifugation at 4 &#176;C at 450 x g for 5 min. Remove the supernatant and resuspend the pellet in 500 &#181;L of cell staining media. Transfer to a new 5 mL round-bottom tube and store on ice.</li>
</ol>
4. FACS startup
NOTE: The steps may vary according to the make and model of the cytometer due to differences in the lasers and channels. For this protocol, a cytometer with 405, 488, 535, and 640 nm wavelength lasers was used. Filters featured in this protocol are for representation purposes. Alternative cell stains may require a different set of filters and lasers.
<ol>
	<li>Begin creating a new project template on the sorter software and choose the laser panel. For the represented combination of stains and natural fluorescence, select all four lasers (405, 488, 535, and 640 nm).</li>
	<li>Select the appropriate filters for each expected color represented in the experiment.
	<ol>
		<li>To detect the DAPI emission and aid in the separation of live and dead cells, use a filter with a bandpass (BP) of 450/50 (425&#8722;475 nm range) such as a DAPI, Hoeschst, or Pacific Blue filter.</li>
		<li>For the detection of light emitted by the green ROS signal, use a filter with a BP of 530/30 (wavelength range of 515-545 nm) such as a fluorescein isothiocyanate (FITC) or green fluorescent protein (GFP) filter. This filter will measure the concentration of ROS in each cell.</li>
		<li>Add an additional filter with a BP of 670/30 (655&#8722;685 nm range) such as an allophycocyanin (APC) filter for the detection of the lysosomal stain emission. 
		NOTE: The APC channel will allow for the measurement of phagocytic activity. This channel will also detect the autofluorescence generated by the coral symbiotic algae from the family Symbiodiniaceae. This signal can be separated using an additional channel that has a longer wavelength than the APC filter, however.</li>
		<li>Include a filter that has a BP of 780/60 (750-810 nm range), such as the most commonly used phycoerythrin, cyanine filter (APC-Cy7), which detects the emission of far-red autofluorescence from Symbiodiniaceae and aids in the isolation of aposymbiotic cells.</li>
	</ol>
	</li>
</ol>
5. FACS gating setup
NOTE: Steps may vary according to make and model of the cytometer and the acquisition program coupled with the cytometer.
<ol>
	<li>On the project experimental screen of the cytometry software, create a scatter plot and select forward scatter (FCS) as the metric for the X-axis and side scatter (SSC) for the Y-axis. 
	NOTE: FCS correlates with the size of the cell and SSC correlates with granularity. This will allow for the removal of most debris, as most will be smaller in size than intact cells.</li>
	<li>Set the axes to either logarithmic or biexponential scales, as cell sizes and granularity may vary by several orders of magnitude (Figure 3A), particularly in corals.</li>
</ol>
6. FACS analysis and cell isolation
NOTE: Steps may vary according to the make and model of the cytometer and the coupled acquisition program.
<ol>
	<li>Place the control (i.e., the unstained cell subset) in the sample chamber of the cytometer and start the reading process. When the computer starts to receive data from each cell, or event, dots will begin to appear on the scatter plot. To clear any debris left in the cytometer chambers from previous experiments, allow approximately 30 s to pass before starting the analysis.</li>
	<li>In the scatter plot, adjust the photomultiplier tube (PMT) voltage in order to center the points. 
	NOTE: The PMT voltage is used to amplify the signal strength of photons being measured; if the PMT voltage is too weak, fewer cells will be read, and if the PMT voltage is too strong, cells will be measured near maximum value and different cell types will be indistinguishable from one another. An adequate PMT voltage ensures that independently distinguishable events will populate the scatter plot. Event separation is easier to visualize by projecting logarithmic or biexponential scales on FSC and SSC scatter plots.</li>
	<li>Start recording the events. To conserve the sample, pause data acquisition after approximately 15,000 events have been read. In most cases, this will be adequate to visualize the overall cell population patterns. Record more events if analyzing and isolating small, specialized cell populations.</li>
	<li>On the first graph of FSC and SSC, create a selection, or gate, of the cells around the 102 mark and higher on the FSC X-axis. Anything below this threshold is likely cellular debris. Draw rectangular gates, which is a regular option on flow cytometry software programs and good for clear, distinct cell populations. For cell populations that take a more irregular shape on the scatter plots, use a polygonal gating option. 
	NOTE: A minimum cutoff of 102 on the forward scatter should select for the smallest coral cells but may allow contamination of debris. To amend this, increase the lower limit of the gating to be more conservative.</li>
	<li>Create a new scatter plot expressing the DAPI filter on the X-axis and the APC-Cy7 filter on the Y-axis. To do this, select the gate for intact cells created in step 6.4, right-click, and select the option to create a new scatter plot. Adjust the axes to either logarithmic or biexponential scales. 
	NOTE: The steps needed to create a new plot may vary with different software programs.</li>
	<li>Now remove the control sample from the cytometer chamber and replace with the stained cells. Allow 30 s to pass before starting the analysis and recording the data. Record approximately 15,000 cells before pausing. 
	NOTE: The distinct population(s) on the higher end of the APC-Cy7 scale are those with high red autofluorescence from Symbiodiniaceae. No stain is needed to visualize these population breaks, and they can also be viewed on the control sample recording (Figure 3C). Select for the coral-only populations, those lower on the Y-axis, for further differentiation.</li>
	<li>Create a new scatter plot of the aposymbiotic cells to visualize the ROS concentrations (FITC filter) and the cells positive to lysosomes (APC filter), again utilizing logarithmic or biexponential scales. 
	NOTE: If there are multiple distinct cell populations that are aposymbiotic, each one can be further distinguished in its own scatter plot (Figure 3C,D).</li>
	<li>Compare the stained sample group against the control, unstained subset by either using the first recording of the control sample or reinserting and rerunning the control group. Gate the events that are unique to the stained sample group.</li>
</ol>
7. FACS sorting and collection
NOTE: Steps may vary according to make and model of the cytometer and the acquisition program coupled with the cytometer.
<ol>
	<li>With a selection of cells chosen to collect, place microcentrifuge tubes (containing 250&#8722;500 &#181;L of cell culture media or lysis buffer based on number of cells collected and method of storage) in the cytometer collection chamber. 
	NOTE: The represented flow cytometer is capable of sorting four different populations at a time, and the machine can be configured to hold a variety of collection devices, including various centrifuge tubes and 96 well plates.</li>
	<li>Once the cytometer starts to read the sample and an appropriate amount of time has passed to flush out debris, start sorting the desired populations and collect anywhere from 20,000 cells to several millions.
	<ol>
		<li>For more than 500,000 cells, adjust the volume of cell culture media or lysis buffer, and the volume of the collection device.</li>
		<li>For in vitro studies, store cells on ice until placed in an incubator or sterilized environment.</li>
		<li>For molecular studies, either immediately start the DNA/RNA/protein isolation process or flash freeze and store at -80 &#176;C until extraction.</li>
	</ol>
	</li>
	<li>Each time a new stain, species, or sorter is used, perform a purity check on the population of interest by sorting at least 20,000 cells of interest into 500 &#181;L of staining media, then re-analyzing the sorted cells and confirming that the cells are being read within the gate used to initially sort (Figure 4).</li>
</ol>

<ol>
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P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regional-scale+assembly+rules+and+biodiversity+of+coral+reefs.">Regional-scale assembly rules and biodiversity of coral reefs.</a> Science. 292 (5521), 1532-1535 (2001).</li><li>Ferrario, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effectiveness+of+coral+reefs+for+coastal+hazard+risk+reduction+and+adaptation.">The effectiveness of coral reefs for coastal hazard risk reduction and adaptation.</a> Nature Communications. 5 (3794), 1-9 (2014).</li><li>Wegley, L., Edwards, R., Rodriguez-Brito, B., Liu, H., Rohwer, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Metagenomic+analysis+of+the+microbial+community+associated+with+the+coral+Porites+astreoides.">Metagenomic analysis of the microbial community associated with the coral Porites astreoides.</a> Environmental Microbiology. 9 (11), 2707-2719 (2007).</li><li>Gates, R. 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The authors have nothing to disclose.

This protocol was adapted from Rosental et al.18 and developed for the identification and isolation of P. damicornis cells. The methodology focuses on the process of filtering samples to remove debris, nonviable cells, and Symbiodiniaceae-hosted cells through the examination of cell intrinsic factors, including relative cell size, relative cell granularity, cell autofluorescence, and the presence of intact cellular membranes. These techniques can be applied to other coral species. However...

Coral reefs are one of the most important ecosystems on Earth. They facilitate biodiversity by providing critical habitats for fish and invertebrates and are crucial for sustaining anthropogenic communities by providing food and economic livelihood through tourism1. As the key builder of coral reefs, the coral animal (Phylum: Cnidaria) also aids coastal communities by creating large calcium carbonate frameworks that mitigate wave and storm damage2.
Corals as adults are sessile animals that host a wide array of endosymbiotic partners, including viruses, archaea, bacteria, protists, fungi, and most notably, members of the algal dinoflagellate family Symbiodiniaceae3. Changes in the environment can cause imbalances in this community, often leading to disease outbreaks and coral bleaching in which the symbiotic Symbiodiniaceae are expelled from the coral colony, thus eliminating the major source of nutrition for the coral. Both of these scenarios often cause death of the coral host4,5,6. Effects of anthropogenic-induced stressors, such as rapid climate change, are accelerating mass coral death events, leading to a global decline of coral reefs7.
Recently, many different methods have been developedto help mitigate coral reef loss. These methods includeoutplanting of corals on existing reefs, genetic crossing using thermally tolerant genotypes, and cellular manipulation of the microbial and symbiotic communities hosted within the coral8,9. Despite these efforts, much remains unknown about coral cell diversity and cell function10,11,12,13. A thorough understanding of coral cell type diversity and cell function is necessary to understand how the coral organism behaves under normative and stressful conditions. Efforts to maximize restoration and preservation efficiency will benefit from an enhanced understanding of how cell diversity and gene function are coupled.
Previous work on cell diversity and function has primarily focused on histological studies and whole-tissue RNA sampling14,15,16,17. To obtain greater detail on specific cell type function in corals, there need to be methods for the isolation of specific populations of live coral cells. This has been done successfully in nonclassical model organisms by means of fluorescence-activated cell sorting (FACS) flow cytometry technology18. FACS utilizes a combination of lasers tuned to varying wavelengths to measure different endogenous cellular properties at the single cell level such as relative cell size, cell granularity, and autofluorescence. Additionally, the cells may be marked by fluorescently labeled compounds to measure specific, desired properties18,19.
Thus far, the application of flow cytometry to coral cells has mainly been for the analysis of symbiotic Symbiodiniaceae and other bacterial populations by utilizing their strong, natural autofluorescence20,21,22. FACS has also been used to estimate coral genome size by using fluorescent DNA marker signal compared against reference model organism cells23,24. The efficient application of FACS provides three distinct tools that are useful for cell biology studies: 1) morphological and functional description of single cells; 2) identification, separation, and isolation of specific cell populations for downstream studies; and 3) the analysis of functional assays at the single cell level.
The development and application of various exogenous fluorescent markers for the study of coral cells remains almost unexplored. Such markers may include tagged proteins, tagged substrates for enzymes, or fluorescent responses to other compounds. These markers can be used to identify cell types that have unique properties, such as highlighting cells that produce varying amounts of a specific cellular compartment feature, like lysosomes. An additional example is the use of fluorescently labeled beads to functionally identify cells competent for phagocytosis, or the engulfment of a targeted pathogen25. Populations of cells active in immunity responses can be easily identified by FACS after engulfment of these exogenously applied beads. While traditional histological methods require preserved tissue and many hours to approximate the percentage of cells positive for bead engulfment, a FACS-based functional assay for pathogen engulfment can be performed relatively quickly on isolated live cells. In addition to studying cell-specific responses to stress, this technology has the potential to clarify gene-specific expression and illuminate the evolutionary and developmental history of cell types entirely unique to cnidarians, such as calicoblasts and cnidocytes.
Recently, we performed an intensive screening of over 30 cellular markers that resulted in identifying 24 that are capable of labeling coral cells, 16 of which are useful for distinguishing unique populations18, making them clusters of differentiation (CD). Here we describe the process of coral cell isolation in Pocillopora damicornis from removing cells from the calcium carbonate skeleton to the identification and isolation of specific cell populations with FACS (Figure 1).

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Airbrush Kit &#38; Compressor</td>
			<td>TCP Global</td>
			<td>ABD KIT-H-SET</td>
			<td>Paasche H Series Single-Action Siphon Feed Airbrush Kit with Master TC-20 Compressor &#38; Air Hose</td>
		</tr>
		<tr>
			<td>BD FACSAria II</td>
			<td>BD</td>
			<td>644832</td>
			<td></td>
		</tr>
		<tr>
			<td>Bone Cutters</td>
			<td>Bulk Reef Supply</td>
			<td>205357</td>
			<td>Oceans Wonders Coral Stony Bone Cutter</td>
		</tr>
		<tr>
			<td>Cell Strainer</td>
			<td>Corning</td>
			<td>352340</td>
			<td>40 um; BD Falcon; individually wrapped; sterile; nylon</td>
		</tr>
		<tr>
			<td>CellRox Green</td>
			<td>Life Technologies</td>
			<td>C10444</td>
			<td>2.5 mM in DMSO; Excitation/Emission: 485/520 nm</td>
		</tr>
		<tr>
			<td>Collection bag</td>
			<td>Grainger</td>
			<td>38UV35</td>
			<td>Reloc Zippit 6&#34;L x 4&#34;W Standard Reclosable Poly Bag with Zip Seal Closure, Clear; 2 mil Thickness</td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Invitrogen</td>
			<td>D1306</td>
			<td>10mg in H2O; Excitation/Emission: 358/461 nm</td>
		</tr>
		<tr>
			<td>Fetal Calf Serum</td>
			<td>Sigma-Aldrich</td>
			<td>F2442-100ML</td>
			<td>Heat-inactivated at 57 &#176;C for 30 minutes</td>
		</tr>
		<tr>
			<td>Hemacytometer</td>
			<td>Sigma-Aldrich</td>
			<td>Z359629</td>
			<td>Bright-Line Hemacytometer</td>
		</tr>
		<tr>
			<td>HEPES Buffer</td>
			<td>Sigma-Aldrich</td>
			<td>H0887</td>
			<td></td>
		</tr>
		<tr>
			<td>LysoTracker Deep Red</td>
			<td>Life Technologies</td>
			<td>L12492</td>
			<td>1mM in DMSO; Absorption/Emission: 647/668 nm</td>
		</tr>
		<tr>
			<td>Microcentrifuge tubes</td>
			<td>VWR</td>
			<td>87003-294</td>
			<td>1.7 mL</td>
		</tr>
		<tr>
			<td>Phophate Buffered Saline (PBS)</td>
			<td>Gibco</td>
			<td>70011-044</td>
			<td>pH 7.4; 10X</td>
		</tr>
		<tr>
			<td>Round-bottom tubes</td>
			<td>VWR</td>
			<td>352063</td>
			<td>5 mL Polypropylene Round-Bottom Tube</td>
		</tr>
		<tr>
			<td>Syringe</td>
			<td>BD</td>
			<td>309628</td>
			<td>1 mL BD Luer-Lok Syringe sterile, singe use polycarbonate</td>
		</tr>
	</tbody>
</table>

fluorescence-activated cell sorting for the isolation of scleractinian cell populations

Coral reefs are under threat due to anthropogenic stressors. The biological response of coral to these stressors may occur at a cellular level, but the mechanisms are not well understood. To investigate coral response to stressors, we need tools for analyzing cellular responses. In particular, we need tools that facilitate the application of functional assays to better understand how cell populations are reacting to stress. In the current study, we use fluorescence-activated cell sorting (FACS) to isolate and separate different cell populations in stony corals. This protocol includes: (1) the separation of coral tissues from the skeleton, (2) creation of a single cell suspension, (3) labeling the coral cells using various markers for flow cytometry, and (4) gating and cell sorting strategies. This method will enable researchers to work on corals at the cellular level for analysis, functional assays, and gene expression studies of different cell populations.

Overall, this protocol is useful because it facilitates the identification and collection of live coral cell populations that can be used for functional analyses. The workflow started with the mechanical separation of coral tissues from the underlying calcium carbonate skeleton (Figure 1). This is one of the most important initial steps because improper technique results in high cell mortality and can create large amounts of debris. Enzymatic separation is no...

Watch this Scientific Journal Video about Fluorescence-Activated Cell Sorting for the Isolation of Scleractinian Cell Populations at JoVE.com

Fluorescence-Activated Cell Sorting for the Isolation of Scleractinian Cell Populations

Corals create biodiverse ecosystems important for both humans and marine organisms. However, we still do not understand the full potential and function of many coral cells. Here, we present a protocol developed for the isolation, labeling, and separation of stony coral cell populations.

Coral reefs are under threat due to anthropogenic stressors. The biological response of coral to these stressors may occur at a cellular level, but the ...

fluorescence-activated-cell-sorting-for-isolation-scleractinian-cell

Research

JoVE Journal

Biology

7.9K Views.  University of Miami. Corals create biodiverse ecosystems important for both humans and marine organisms. However, we still do not understand the full potential and function of many coral cells. Here, we present a protocol developed for the isolation, labeling, and separation of stony coral cell populations.

Video: Fluorescence-Activated Cell Sorting for the Isolation of Scleractinian Cell Populations

Fluorescence Activated Cell Sorting of Plant Protoplasts

High-resolution, cell type-specific analysis of gene expression greatly enhances understanding of developmental regulation and responses to environmental stimuli in any multicellular organism. In situ hybridization and reporter gene visualization can to a limited extent be used to this end but for high resolution quantitative RT-PCR or high-throughput transcriptome-wide analysis the isolation of RNA from particular cell types is requisite. Cellular dissociation of tissue expressing a fluorescent protein marker in a specific cell type and subsequent Fluorescence Activated Cell Sorting (FACS) makes it possible to collect sufficient amounts of material for RNA extraction, cDNA synthesis/amplification and microarray analysis.
An extensive set of cell type-specific fluorescent reporter lines is available to the plant research community. In this case, two marker lines of the Arabidopsis thaliana root are used: PSCR::GFP (endodermis and quiescent center) and PWOX5::GFP (quiescent center). Large numbers (thousands) of seedlings are grown hydroponically or on agar plates and harvested to obtain enough root material for further analysis. Cellular dissociation of plant material is achieved by enzymatic digestion of the cell wall. This procedure makes use of high osmolarity-induced plasmolysis and commercially available cellulases, pectinases and hemicellulases to release protoplasts into solution.
FACS of GFP-positive cells makes use of the visualization of the green versus the red emission spectra of protoplasts excited by a 488 nm laser. GFP-positive protoplasts can be distinguished by their increased ratio of green to red emission. Protoplasts are typically sorted directly into RNA extraction buffer and stored for further processing at a later time.
This technique is revealed to be straightforward and practicable. Furthermore, it is shown that it can be used without difficulty to isolate sufficient numbers of cells for transcriptome analysis, even for very scarce cell types (e.g. quiescent center cells). Lastly, a growth setup for Arabidopsis seedlings is demonstrated that enables uncomplicated treatment of the plants prior to cell sorting (e.g. for the cell type-specific analysis of biotic or abiotic stress responses). Potential supplementary uses for FACS of plant protoplasts are discussed.

A method for isolating specific cell types from plant material is demonstrated. This technique employs transgenic marker lines expressing fluorescent proteins in particular cell types, cellular dissociation and Fluorescence Activated Cell Sorting. Additionally, a growth setup is established here that facilitates treatment of Arabidopsis thaliana seedlings prior to cell sorting.

High-resolution, cell type-specific analysis of gene expression greatly enhances understanding of developmental regulation and responses to environmental ...

Bromodeoxyuridine (BrdU) Labeling and Subsequent Fluorescence Activated Cell Sorting for Culture-independent Identification of Dissolved Organic Carbon-degrading Bacterioplankton

Microbes are major agents mediating the degradation of numerous dissolved organic carbon (DOC) substrates in aquatic environments. However, identification of bacterial taxa that transform specific pools of DOC in nature poses a technical challenge. 
Here we describe an approach that couples bromodeoxyuridine (BrdU) incorporation, fluorescence activated cell sorting (FACS), and 16S rRNA gene-based molecular analysis that allows culture-independent identification of bacterioplankton capable of degrading a specific DOC compound in aquatic environments. Triplicate bacterioplankton microcosms are set up to receive both BrdU and a model DOC compound (DOC amendments), or only BrdU (no-addition control). BrdU substitutes the positions of thymidine in newly synthesized bacterial DNA and BrdU-labeled DNA can be readily immunodetected 1,2. Through a 24-hr incubation, bacterioplankton that are able to use the added DOC compound are expected to be selectively activated, and therefore have higher levels of BrdU incorporation (HI cells) than non-responsive cells in the DOC amendments and cells in no-addition controls (low BrdU incorporation cells, LI cells). After fluorescence immunodetection, HI cells are distinguished and physically separated from the LI cells by fluorescence activated cell sorting (FACS) 3. Sorted DOC-responsive cells (HI cells) are extracted for DNA and taxonomically identified through subsequent 16S rRNA gene-based analyses including PCR, clone library construction and sequencing.

Bromodeoxyuridine BrdU Labeling and Subsequent Fluorescence Activated Cell Sorting for Culture-independent Identification of Dissolved Organic Carbon-degrading Bacterioplankton

Environmental bacterioplankton are incubated with a model dissolved organic carbon (DOC) compound and a DNA labeling reagent, bromodeoxyuridine (BrdU). Afterward, DOC-degrading cells are separated from the bulk community based on their elevated BrdU incorporation using fluorescence activated cell sorting (FACS). These cells are then identified by subsequent molecular analyses.

Microbes are major agents mediating the degradation of numerous dissolved organic carbon (DOC) substrates in aquatic environments. However, identification ...

Robust Generation of Hepatocyte-like Cells from Human Embryonic Stem Cell Populations

Despite progress in modelling human drug toxicity, many compounds fail during clinical trials due to unpredicted side effects. The cost of clinical studies are substantial, therefore it is essential that more predictive toxicology screens are developed and deployed early on in drug development (Greenhough et al 2010). Human hepatocytes represent the current gold standard model for evaluating drug toxicity, but are a limited resource that exhibit variable function. Therefore, the use of immortalised cell lines and animal tissue models are routinely employed due to their abundance. While both sources are informative, they are limited by poor function, species variability and/or instability in culture (Dalgetty et al 2009). Pluripotent stem cells (PSCs) are an attractive alternative source of human hepatocyte like cells (HLCs) (Medine et al 2010). PSCs are capable of self renewal and differentiation to all somatic cell types found in the adult and thereby represent a potentially inexhaustible source of differentiated cells. We have developed a procedure that is simple, highly efficient, amenable to automation and yields functional human HLCs (Hay et al 2008 ; Fletcher et al 2008 ; Hannoun et al 2010 ; Payne et al 2011 and Hay et al 2011). We believe our technology will lead to the scalable production of HLCs for drug discovery, disease modeling, the construction of extra-corporeal devices and possibly cell based transplantation therapies.

This article will focus on the generation of human hepatic endoderm from human embryonic stem cell populations.

Despite progress in modelling human drug toxicity, many compounds fail during clinical trials due to unpredicted side effects. The cost of clinical ...

Cell Specific Analysis of Arabidopsis Leaves Using Fluorescence Activated Cell Sorting

After initiation of the leaf primordium, biomass accumulation is controlled mainly by cell proliferation and expansion in the leaves1. However, the Arabidopsis leaf is a complex organ made up of many different cell types and several structures. At the same time, the growing leaf contains cells at different stages of development, with the cells furthest from the petiole being the first to stop expanding and undergo senescence1. Different cells within the leaf are therefore dividing, elongating or differentiating; active, stressed or dead; and/or responding to stimuli in sub-sets of their cellular type at any one time. This makes genomic study of the leaf challenging: for example when analyzing expression data from whole leaves, signals from genetic networks operating in distinct cellular response zones or cell types will be confounded, resulting in an inaccurate profile being generated.
To address this, several methods have been described which enable studies of cell specific gene expression. These include laser-capture microdissection (LCM)2 or GFP expressing plants used for protoplast generation and subsequent fluorescence activated cell sorting (FACS)3,4, the recently described INTACT system for nuclear precipitation5 and immunoprecipitation of polysomes6.
FACS has been successfully used for a number of studies, including showing that the cell identity and distance from the root tip had a significant effect on the expression profiles of a large number of genes3,7. FACS of GFP lines have also been used to demonstrate cell-specific transcriptional regulation during root nitrogen responses and lateral root development8, salt stress9 auxin distribution in the root10 and to create a gene expression map of the Arabidopsis shoot apical meristem11. Although FACS has previously been used to sort Arabidopsis leaf derived protoplasts based on autofluorescence12,13, so far the use of FACS on Arabidopsis lines expressing GFP in the leaves has been very limited4. In the following protocol we describe a method for obtaining Arabidopsis leaf protoplasts that are compatible with FACS while minimizing the impact of the protoplast generation regime. We demonstrate the method using the KC464 Arabidopsis line, which express GFP in the adaxial epidermis14, the KC274 line, which express GFP in the vascular tissue14 and the TP382 Arabidopsis line, which express a double GFP construct linked to a nuclear localization signal in the guard cells (data not shown; Figure 2). We are currently using this method to study both cell-type specific expression during development and stress, as well as heterogeneous cell populations at various stages of senescence.

Cell Specific Analysis of Arabidopsis Leaves Using Fluorescence Activated Cell Sorting

A method for producing Arabidopsis leaf protoplasts that are compatible with fluorescence activated cell sorting (FACS), allowing for studies of specific cell populations. This method is compatible with any Arabidopsis line that expresses GFP in a subset of cells.

After initiation of the leaf primordium, biomass accumulation is controlled mainly by cell proliferation and expansion in the leaves1. However, the ...

Studying Proteolysis of Cyclin B at the Single Cell Level in Whole Cell Populations

Equal distribution of chromosomes between the two daughter cells during cell division is a prerequisite for guaranteeing genetic stability 1. Inaccuracies during chromosome separation are a hallmark of malignancy and associated with progressive disease 2-4. The spindle assembly checkpoint (SAC) is a mitotic surveillance mechanism that holds back cells at metaphase until every single chromosome has established a stable bipolar attachment to the mitotic spindle1. The SAC exerts its function by interference with the activating APC/C subunit Cdc20 to block proteolysis of securin and cyclin B and thus chromosome separation and mitotic exit. Improper attachment of chromosomes prevents silencing of SAC signaling and causes continued inhibition of APC/CCdc20 until the problem is solved to avoid chromosome missegregation, aneuploidy and malignant growths1.
Most studies that addressed the influence of improper chromosomal attachment on APC/C-dependent proteolysis took advantage of spindle disruption using depolymerizing or microtubule-stabilizing drugs to interfere with chromosomal attachment to microtubules. Since interference with microtubule kinetics can affect the transport and localization of critical regulators, these procedures bear a risk of inducing artificial effects 5.
To study how the SAC interferes with APC/C-dependent proteolysis of cyclin B during mitosis in unperturbed cell populations, we established a histone H2-GFP-based system which allowed the simultaneous monitoring of metaphase alignment of mitotic chromosomes and proteolysis of cyclin B 6.
To depict proteolytic profiles, we generated a chimeric cyclin B reporter molecule with a C-terminal SNAP moiety 6 (Figure 1). In a self-labeling reaction, the SNAP-moiety is able to form covalent bonds with alkylguanine-carriers (SNAP substrate) 7,8 (Figure 1). SNAP substrate molecules are readily available and carry a broad spectrum of different fluorochromes. Chimeric cyclin B-SNAP molecules become labeled upon addition of the membrane-permeable SNAP substrate to the growth medium 7 (Figure 1). Following the labeling reaction, the cyclin B-SNAP fluorescence intensity drops in a pulse-chase reaction-like manner and fluorescence intensities reflect levels of cyclin B degradation 6 (Figure 1). Our system facilitates the monitoring of mitotic APC/C-dependent proteolysis in large numbers of cells (or several cell populations) in parallel. Thereby, the system may be a valuable tool to identify agents/small molecules that are able to interfere with proteolytic activity at the metaphase to anaphase transition. Moreover, as synthesis of cyclin B during mitosis has recently been suggested as an important mechanism in fostering a mitotic block in mice and humans by keeping cyclin B expression levels stable 9,10, this system enabled us to analyze cyclin B proteolysis as one element of a balanced equilibrium 6.

Metaphase to anaphase transition is triggered through anaphase-promoting complex (APC/C)-dependent ubiquitination and subsequent destruction of cyclin B. Here, we established a system which, following pulse-chase labeling, allows monitoring cyclin B proteolysis in entire cell populations and facilitates the detection of interference by the mitotic checkpoint.

Equal distribution of chromosomes between the two daughter cells during cell division is a prerequisite for guaranteeing genetic stability 1. Inaccuracies ...

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 ...

Isolation of Normal and Cancer-associated Fibroblasts from Fresh Tissues by Fluorescence Activated Cell Sorting (FACS)

Cancer-associated fibroblasts (CAFs) are the most prominent cell type within the tumor stroma of many cancers, in particular breast carcinoma, and their prominent presence is often associated with poor prognosis1,2. CAFs are an activated subpopulation of stromal fibroblasts, many of which express the myofibroblast marker &alpha;-SMA3. CAFs originate from local tissue fibroblasts as well as from bone marrow-derived cells recruited into the developing tumor and adopt a CAF phenotype under the influence of the tumor microenvironment4. CAFs were shown to facilitate tumor initiation, growth and progression through signaling that promotes tumor cell proliferation, angiogenesis, and invasion5-8. We demonstrated that CAFs enhance tumor growth by mediating tumor-promoting inflammation, starting at the earliest pre-neoplastic stages9. Despite increasing evidence of the key role CAFs play in facilitating tumor growth, studying CAFs has been an on-going challenge due to the lack of CAF-specific markers and the vast heterogeneity of these cells, with many subtypes co-existing in the tumor microenvironment10. Moreover, studying fibroblasts in vitro is hindered by the fact that their gene expression profile is often altered in tissue culture11,12 . To address this problem and to allow unbiased gene expression profiling of fibroblasts from fresh mouse and human tissues, we developed a method based on previous protocols for Fluorescence-Activated Cell Sorting (FACS)13,14. Our approach relies on utilizing PDGFR&alpha; as a surface marker to isolate fibroblasts from fresh mouse and human tissue. PDGFR&alpha; is abundantly expressed by both normal fibroblasts and CAFs9,15 . This method allows isolation of pure populations of normal fibroblasts and CAFs, including, but not restricted to &alpha;-SMA+ activated myofibroblasts. Isolated fibroblasts can then be used for characterization and comparison of the evolution of gene expression that occurs in CAFs during tumorigenesis. Indeed, we and others reported expression profiling of fibroblasts isolated by cell sorting16. This protocol was successfully performed to isolate and profile highly enriched populations of fibroblasts from skin, mammary, pancreas and lung tissues. Moreover, our method also allows culturing of sorted cells, in order to perform functional experiments and to avoid contamination by tumor cells, which is often a big obstacle when trying to culture CAFs.

Isolation of Normal and Cancer-associated Fibroblasts from Fresh Tissues by Fluorescence Activated Cell Sorting FACS

Cancer Associated Fibroblasts (CAFs) facilitate tumor initiation, growth and progression through signaling that promotes proliferation, angiogenesis, and inflammation. Here we describe a method to isolate pure populations of normal fibroblasts and CAFs from fresh mouse and human tissues by cell sorting, using PDGFR&#x03B1; as a surface marker.

Cancer-associated fibroblasts (CAFs) are the most prominent cell type within the tumor stroma of many cancers, in particular breast carcinoma, and their ...

Isolation of Blood-vessel-derived Multipotent Precursors from Human Skeletal Muscle

Since the discovery of mesenchymal stem/stromal cells (MSCs), the native identity and localization&#160;of MSCs have been obscured by their retrospective isolation in culture. Recently, using fluorescence-activated cell sorting (FACS), we and other researchers prospectively identified and purified three subpopulations of multipotent precursor cells associated with the vasculature of human skeletal muscle. These three cell populations: myogenic endothelial cells (MECs), pericytes (PCs), and adventitial cells (ACs), are localized respectively to the three structural layers of blood vessels: intima, media, and adventitia. All of these human blood-vessel-derived stem cell (hBVSC) populations not only express classic MSC markers but also possess mesodermal developmental potentials similar to typical MSCs. Previously, MECs, PCs, and ACs have been isolated through distinct protocols and subsequently characterized in separate studies. The current isolation protocol, through modifications to the isolation process and adjustments in the selective cell surface markers, allows us to simultaneously purify all three hBVSC subpopulations by FACS from a single human muscle biopsy. This new method will not only streamline the isolation of multiple BVSC subpopulations but also facilitate future clinical applications of hBVSCs for distinct therapeutic purposes.

Blood vessels within human skeletal muscle harbor several multi-lineage precursor populations that are ideal for regenerative applications. This isolation method allows simultaneous purification of three multipotent precursor cell populations respectively from three structural layers of blood vessels: myogenic endothelial cells from intima, pericytes from media, and adventitial cells from adventitia.

Since the discovery of mesenchymal stem/stromal cells (MSCs), the native identity and localization&#160;of MSCs have been obscured by their retrospective ...

Single Cell Micro-aspiration as an Alternative Strategy to Fluorescence-activated Cell Sorting for Giant Virus Mixture Separation

During the amoeba co-culture process, more than one virus may be isolated in a single well. We previously solved this issue by end point dilution and/or fluorescence activated cell sorting (FACS) applied to the viral population. However, when the viruses in the mixture have similar morphologic properties and one of the viruses multiplies slowly, the presence of two viruses is discovered at the stage of genome assembly and the viruses cannot be separated for further characterization. To solve this problem, we developed a single cell micro-aspiration procedure that allows for separation and cloning of highly similar viruses. In the present work, we present how this alternative strategy allowed us to separate the small viral subpopulations of Clandestinovirus ST1 and Usurpativirus LCD7, giant viruses that grow slowly and do not lead to amoebal lysis compared to the lytic and fast-growing Faustovirus. Purity control was assessed by specific gene amplification and viruses were produced for further characterization.

Here, we describe a single cell micro-aspiration method for the separation of infected amoebae. In order to separate viral subpopulations in Vermamoeba vermiformis infected by Faustoviruses and unknown giant viruses, we developed the protocol detailed below and demonstrated its ability to separate two low-abundance novel giant viruses.

During the amoeba co-culture process, more than one virus may be isolated in a single well. We previously solved this issue by end point dilution and/or ...

Isolation of Quiescent Stem Cell Populations from Individual Skeletal Muscles

Skeletal muscle harbors distinct populations of adult stem cells that contribute to the homeostasis and repair of the tissue. Skeletal muscle stem cells (MuSCs) have the ability to make new muscle, whereas fibro-adipogenic progenitors (FAPs) contribute to stromal supporting tissues and have the ability to make fibroblasts and adipocytes. Both MuSCs and FAPs reside in a state of prolonged reversible cell cycle exit, called quiescence. The quiescent state is key to their function. Quiescent stem cells are commonly purified from multiple muscle tissues pooled together in a single sample. However, recent studies have revealed distinct differences in the molecular profiles and quiescence depth of MuSCs isolated from different muscles. The present protocol describes the isolation and study of MuSCs and FAPs from individual skeletal muscles and presents strategies to perform molecular analysis of stem cell activation. It details how to isolate and digest muscles of different developmental origin, thicknesses, and functions, such as the diaphragm, triceps, gracilis, tibialis anterior (TA), gastrocnemius (GA), soleus, extensor digitorum longus (EDL), and the masseter muscles. MuSCs and FAPs are purified by fluorescence-activated cell sorting (FACS) and analyzed by immunofluorescence staining and 5-ethynyl-2&#180;-deoxyuridine (EdU) incorporation assay.

This protocol describes the isolation of muscle stem cells and fibro-adipogenic progenitors from individual skeletal muscles in mice. The protocol involves single muscle dissection, stem cell isolation by fluorescence activated cell sorting, purity assessment by immunofluorescence staining, and quantitative measurement of S-phase entry by 5-ethynyl-2&#180;-deoxyuridine incorporation assay.

Skeletal muscle harbors distinct populations of adult stem cells that contribute to the homeostasis and repair of the tissue. Skeletal muscle stem cells ...