This research was funded by National Science Foundation Award Number 1237969, &#34;Unraveling the Heterozygosity, Allelic Composition, and Copy Number Variation of Potato&#34; and USDA Special Grant 2014-34141-22266 (University of Maine) to RV.

NOTE: It is important to include appropriate controls, usually in the form of leaf tissue of the same ploidy as experimental samples. This is because the 2C peak of tuber samples may be small and difficult to identify.
1. Preparations
NOTE: The solutions listed here may be prepared ahead of time and stored at 4 &#176;C but those presented in subsequent steps must be made fresh on the day of use.
<ol>
	<li>Make an appropriate volume of Plasmolysis Incubation Solution (PIS) (15 mL/sample): 0.55 M mannitol, 2 mM CaCl2, 1 mM KH2PO4, 1 mM MgCl2, 50 mM Tris buffer pH 7.5 (adjusted with HCl).</li>
	<li>Make an appropriate volume of Plasmolysis Wash Solution (PWS) (15 mL/sample) which is identical to PIS except for addition of 0.71 M mannitol.</li>
	<li>Make 250 mL of modified Galbraith&#39;s Flow Cytometry Buffer11 (FCB): 13.6 mM sodium citrate (trisodium), 8 mM MOPS, 18 mM MgCl2, 0.4% v/v Triton X-100. Stir for at least 30 min to distribute the Triton X-100.</li>
	<li>Filter sterilize PIS and PWS solutions with a 0.22 &#181;m asymmetric polyethersulfone (aPES) filter. Use vacuum driven filtration units.</li>
	<li>Prepare 106 &#181;m mesh filters for lysed protoplasts. Prepare 1.5 mL microcentrifuge tubes which may be directly nested into collection tubes, however any method of passing suspensions through a 106 &#181;m filter may be used. 
	NOTE: These do not need to be sterile.
	<ol>
		<li>If using 1.5 mL microcentrifuge tubes, cut 1 cm off the tip each microcentrifuge tube. Using a hot plate or other heat source heat a 1 cm2 piece 106 &#181;m steel mesh on a piece of aluminum foil. Press the cut end of the tube into the mesh until they have fused.</li>
	</ol>
	</li>
	<li>Wash potatoes to be sampled, removing all soil and debris. 
	NOTE: Tuber size and age should be appropriate for the goals of the experiment but do not seem to affect quality of results. We have successfully applied this protocol to tubers which have been in cold storage (4 &#176;C, 95% RH) up to 8 months. While tubers with defects (e.g. tuber end rot, hollow heart, brown necrotic areas) may be responsive, they should be avoided, if possible.</li>
</ol>
2. Day 1: Plasmolyze Tuber Tissue
NOTE: Control leaf samples can be included here to produce protoplasts or at step 5.2 if crude preparations are preferred. This must be performed with sterile tools in an aseptic environment such as a laminar flow hood.
<ol>
	<li>Surface sterilize potato tubers via immersion in 75% ethanol for at least 5 min.</li>
	<li>Remove tubers from ethanol and air dry. This usually takes ~5 min, longer if the tuber is larger than 100 g.</li>
	<li>Prepare and label sterile 50 mL conical tubes for each sample and aliquot 15 mL of filter sterilized PIS into each.</li>
	<li>Sample approximately 1 cm3 of the desired tissue from sterilized tubers using either sterilized scalpel or cork borer. 
	NOTE: Depending on the tissue being sampled one may use either a sterilized cork borer or knife/scalpel. For instance, if the pith is desired the cork borer may be used to take a core from the apical to distal end of the tuber and the central fraction of the core may be sampled. However, due to the asymmetric internal morphology of potato tubers, it may be more precise to sample parenchyma or cortex by halving the tuber and excising the desired tissue. See Figure 1 for a depiction of internal tuber morphology.
	<ol>
		<li>Coarsely chop tissue using a sterilized scalpel into approx. 3 mm3 pieces and transfer tissue to conical tube containing PIS. Incubate overnight at 4 &#176;C. 
		NOTE: This is to maximize surface area so that the PIS, and later the enzyme solution, may fully permeate the tissue resulting in more complete digestion.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57134/57134fig1.jpg" /> 
Figure 1: Internal morphology of a potato tuber. The separation between pith (P) and perimedullary parenchyma (PM) is illustrated with black lines. The vascular ring, which separates parenchyma from the cortex (C) is denoted with a black arrow. The stolon end (S) and tuber eyes (E) are labeled as well. <a href="//cloudfront-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57134/57134fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Day 2: Generate Potato Protoplasts
NOTE: This must be performed with sterile tools in an aseptic environment such as a laminar flow hood.
<ol>
	<li>Prepare and filter (0.45 &#181;m aPES filter) an appropriate amount of Enzyme Solution (ES) (10 mL/sample): 0.71 M mannitol, 3 mM CaCl2, 1 mM KH2PO4, 1 mM MgCl2, 4% Onozuka R-10 cellulase, 0.8% macerozyme R-10, 1% hemicululase, 10 mM MES buffer, pH 5.8. Ensure that appropriate filters are used in this step; filters with pore size smaller than 0.45 &#181;m are prone to clogging whereas a low protein binding membrane, such as aPES, minimizes loss of enzyme during filtration.</li>
	<li>Aspirate off PIS from samples in conical tubes using a sterile serological pipette.</li>
	<li>Add 10 mL of ES to each sample and invert 2&#8211;3 times.</li>
	<li>Incubate samples overnight at 29 &#176;C with 180 rpm horizontal shaking. Incubate samples for at least 16 h.</li>
</ol>
4. Day 3: Harvest Protoplasts and Wash Protoplasts
NOTE: Aseptic conditions are no long necessary at this point.
<ol>
	<li>Remove samples from shaker and allow them to settle for ~10 min.</li>
	<li>Aspirate off as much of the ES solution as possible.
	<ol>
		<li>Remove as much of the ES solution as feasible with a serological pipette, taking care to avoid removing the digested tissue.</li>
		<li>Using a micropipette remove any remaining ES solution, again avoiding the digested tissue. This is most easily done by holding the tube at an angle so that the ES solution flows toward the lid while the digested tissue remains at the bottom.</li>
	</ol>
	</li>
	<li>Add 15 mL of PWS to each sample and invert gently 2&#8211;3 times. Allow protoplasts to settle for 10 min.</li>
	<li>Remove PWS with a serological pipette. Use a micropipette to remove any remaining liquid. 
	NOTE: This step is absolutely critical to ensure the integrity of the sample nuclei during flow cytometry. In troubleshooting, this step was found to be the most likely source of failure in a sample.</li>
</ol>
5. Day 3: Prepare Samples for Flow Cytometry
NOTE: The samples should be kept on ice from here on unless otherwise noted.
<ol>
	<li>Prepare propidium iodide (0.4 mg propidium iodide/mL FCB) and RNase solutions (0.8 mg RNase A/mL FCB) by dissolving each in FCB. 
	CAUTION: Propidium iodide is highly toxic. Avoid contact with skin or eyes. Wear appropriate PPE.</li>
	<li>Add 1.5 mL of ice cold FCB to each sample. Briefly shake or vortex each sample to break up aggregated tissue. It is important to break up the clumps of tuber tissue so that the FCB may fully permeate the cells and release the nuclei. Different samples may require more or less agitation depending on tissue and genotype. 
	NOTE: The FCB lyses the protoplasts, releasing the nuclei. Hence the necessity of keeping samples on ice to prevent degradation.
	<ol>
		<li>If control flow cytometry samples were not included in the protoplast generation step they may be included here. If a control is to be added, finely chop a small leaf (~3 cm2) in 1.5 mL of ice cold FCB using a razor blade. Control samples should be the same ploidy as the experimental samples, preferably the same genotype. In vitro plantlets tend to give cleaner peaks than greenhouse or field-grown plants.</li>
	</ol>
	</li>
	<li>Pass 1 mL of the FCB/tissue suspension through a 106 &#181;m mesh filter. Use a 1.5 mL microcentrifuge tube with the tip cut off and metal mesh melted to the bottom as described in step 1.5.1. This microcentrifuge tube may be nested directly into a 2 mL microcentrifuge tube and the sample passed directly through. If using another method of filtration, the filtrate may be collected in an ice-cold Petri plate and then transferred to a 2 mL tube. 
	NOTE: sometimes the remaining aggregates and cellular debris may clog the mesh filter. In this case, simply tap the two nested microcentrifuge tubes a few times to dislodge the debris.</li>
	<li>Add 250 &#181;L of the RNase solution to each sample. Invert and incubate for 30 min at room temperature (RT). 
	NOTE: This step is intended to remove RNA from the samples, leading to less noise during flow cytometry. However, as the nuclei are short-lived, researchers may decide to decrease the incubation time or perform it on ice if they are experiencing severe degradation of their samples.</li>
	<li>Add 125 &#181;L of the propidium iodide solution to each sample. Invert and incubate on ice for 30 min. 
	NOTE: Samples should be used for flow cytometry as soon as possible as degradation is apparent within 2 h, even on ice.</li>
</ol>
6. Day 3: Flow Cytometry of Potato Tuber Nuclei
NOTE: Flow cytometer operation and software will vary depending on the instrument and manufacturer. Detailed instructions on the specific operation of the flow cytometer will be provided in the instrument&#39;s User Guide.
<ol>
	<li>Create two dot plots using logarithmic scale of forward scatter vs. side scatter and propidium iodide (PI) vs side scatter. Also create a histogram with PI on the x-axis. Logarithmic scale is required to ensure all events are within scale as nuclei may differ vastly in fluorescence.</li>
	<li>Load a known control sample tube and adjust the voltage so that all events are on scale. We use in vitro leaf tissue from a sample genotype. If samples of different ploidies are to be run, a control sample for each should be included.
	<ol>
		<li>Make note of the channel of the 2C peak in the control sample. The 2C peaks of the experimental samples should fall in the same location.</li>
	</ol>
	</li>
	<li>Load an experimental (tuber) sample and again ensure that all events are on scale. If adjustments are required, repeat step 6.2 to identify the channel of 2C peaks.</li>
	<li>Manually gate the protoplast nuclei using the side scatter vs PI plot.</li>
	<li>Set the PI histogram to only show the gated protoplast nuclei region.</li>
	<li>Collect the desired number of events from each sample. Frequently researchers use 10,000 gated events for flow cytometry; however we use 2,000 events for tuber protoplast samples to accommodate more samples and samples with low concentrations.</li>
	<li>Calculate each sample&#39;s EI from the PI histograms using the following formula: 
	EI = <img alt="Equation 1" src="/files/ftp_upload/57134/57134eq1.jpg" /> 
	where 4C is the percentage of nuclei which are 4C (representing a single round of endoreduplication), 8C is the percentage which are 8C, and so on. See Figure 4 for examples of histograms and C-values of peaks.</li>
</ol>

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The authors declare that they have no competing financial interests.

The protocol presented herein provides researchers with a means to assess endoreduplication within potato tubers, whose modified cellular content and increased cell size seemingly preclude other flow cytometry preparations. The protocol relies upon protoplast generation as a means to reduce noise and debris while maintaining nuclear integrity. Previously, researchers have described similar preparations for particularly recalcitrant flow cytometry samples as well as utilized tuber protoplasts to study a variety of topics such as pathogenesis19,20. However, to our knowledge, none have combined the use of such tuber protoplasts with flow cytometry for the purpose of studying endoreduplication. Furthermore, we found the use of protoplasts to be more reliable than typical crude preparations as well as the technique utilized in the two only other studies to assess tuber endoreduplication6,7. Here we discuss the shortcomings of the protocol, potential pitfalls in its execution and sample preparation, and results of a typical experiment which employs it.
Despite the utility and repeatability of the tuber flow cytometry protocol, it does have a few weaknesses which should be discussed. To begin, the protocol is time intensive, requiring two overnight incubations. Furthermore, the protocol requires some expensive reagents, particularly the cellulase and macerozyme. Additionally, the preparations are highly time-sensitive, degrading within a few hours, which may limit throughput within a single day, especially if many events are desired. Lastly, the protocol, while reliable, is sensitive to errors within sample preparation all of which seem to result in damage to the nuclei and lower quality results. For example, microbial contamination may occur during the plasmolysis and protoplast generation steps (1&#8211;3.4) due to improper aseptic technique. Sample contamination, while not always precluding the success of a sample, seems to dramatically decrease quality of obtained histograms, again likely due to damage to the nuclei.
As previously stated, the major drawbacks of the protocol are costs, time, and sensitivity to errors. Researchers may wish to take steps to mitigate each of these. As for cost, researchers intending to apply the protocol to many samples may consider modifying the enzyme concentrations and/or duration of the digestion step as different tissues and genotypes may vary in responsiveness. This includes the possibility of filtering and recycling the ES which has previously been demonstrated but not employed here21. As for time, in our observation, it is possible to dispense with the first incubation (the plasmolysis step) and still extract protoplasts; however, their integrity and abundance will suffer, which negatively impacts results. To reduce the deterioration of samples researchers may consider that some flow cytometry approaches involve the use of fixatives (e.g., formaldehyde) to preserve sample integrity22. This may be a useful means to both prevent deterioration and allow for sample storage rather than protoplast exaction and flow cytometry occurring on the same day as presented. Another means of preventing the attrition of nuclei may be to include a nuclease inhibitor to the ES and/or FBC; while 2-mercaptoethanol effected no noticeable changes during development, other inhibitors have not been tested herein.
In our observation, the single most critical step is step 4.4, the removal of all plasmolysis wash solution before the addition of the flow cytometry buffer (FCB). If even a small volume (&#60;10 &#181;L) remains, the samples are much more likely to be degraded which can lead to a poor or even failed sample. This is likely due to impurities within the enzyme solution (e.g. nucleases, proteases)23,24 which quickly degrade the nuclei during the incubation periods and time between sample runs, even at low concentrations. Another important consideration is the duration of the PI and RNase incubation steps. As noted in the protocol, the samples do not remain stable for more than a few hours after addition of the FCB so researchers may decide to reduce the duration of these steps to accommodate more samples or lengthy flow cytometry runs resulting from low nuclei concentrations. This also requires the researcher to consider the tradeoff between number of events per sample and total number of samples to be run or consider the use of fixatives as previously mentioned.
Differences between Tissues and Genotypes
To provide results representative of the protocol we designed two simple experiments to confirm previously reported variation by tissue and examine the influence of ploidy and genotype. The results of the tissue experiment demonstrate that the protocol yields reproducible results, as pith tissue was once again found to have the highest EI. Somewhat surprisingly parenchyma tissue, which had not previously been evaluated, had an EI value similar to cortical tissue and significantly lower than pith. This was unanticipated as parenchyma cells are typically larger than either pith or cortical cells, at least at maturity. One possible explanation is that the tubers (90&#8211;130 g) were too immature for the parenchyma cells, which comprise the majority of tuber volume at maturity, to have fully expanded and reached their maximum C-values25. This may also explain why the dihaploid (VT_SUP_19) and the doubled dihaploid (VT_SUP_19 4x) demonstrated significantly greater EI than cv. Superior tubers; while tubers of approximately equal size were sampled from each genotype, the maximum size of tubers from either VT_SUP_19 or the isogenic tetraploid is much smaller than that of the progenitor. Thus, it may be that they displayed greater EI simply because they were larger relative to their maximum attainable size. Alternatively, the difference may be a consequence of the genetic complement VT_SUP_19 received from its tetraploid progenitor. Another possibility is that the genomic reduction that occurred on extraction of the dihaploid from the tetraploid may have unmasked deleterious alleles resulting in plant-wide stress, which has also been shown to contribute to elevated EI26. Nevertheless, this demonstrates the careful consideration researchers must employ when selecting tubers and tissues to be used for comparison of EI values, especially between genotypes.
Future applications
The protocol described in this article provides researchers with an important tool for understanding endoreduplication in potato. It may allow for studies into the genetic and environmental components of endoreduplication, the time-course of development across tuber tissues, and assessment of natural variation. Ultimately, endoreduplication may make a promising target for potato improvement, an undertaking which will require a reliable means of assessment.

Endoreduplication is the process by which a cell forgoes the typical cell cycle and instead undergoes an alternate course of development consisting of repeated rounds of DNA replication without cellular division. The resulting cell will have increased DNA content and nuclear size which is thought to play a role in cellular regulation, expansion, and specialization. Generally, a round of endoreduplication (termed an endocycle) and the corresponding increase in DNA content are associated with larger cell volume, an observation that precipitated the &#34;karyoplasmic theory&#34; that increased DNA content is required to properly regulate a larger, perhaps more complex, cell1. This phenomenon is common in higher plants, having been observed in a range of tissues including those with structural/defensive (trichomes)2,3, nutritive (maize endosperm)4,5, and sink/storage (tomato pericarp; potato tuber)6,7,8 functions. In fruit, it has been suggested that endoreduplication plays a role in facilitating the rapid expansion of the pericarp as evidenced by the negative relationship between endoreduplication and fruit developmental period9. For instance cells with DNA content up to 512C (512 times the haploid genome) have been observed in the tomato pericarp8. Furthermore Chevalier et al. (2014) demonstrated that alterations in expression of cell cycle genes can lead to increases in endoreduplication levels within the pericarp which then results in larger fruit10. Thus, alteration of genes promoting endoreduplication provides a potential target for improvement of biomass or yield through plant breeding or genetic manipulation. However, such improvement is contingent upon greater understanding of the causes and consequences of endoreduplication.
Endoreduplication is most often measured via flow cytometry whereby nuclei, released in crude tissue preparations11, are incubated with a DNA-binding fluorophore, such as propidium iodide12 (PI). The filtered samples are then passed by the laser of a flow cytometer where emission wavelengths specific to the fluorophore can be observed. The intensity of fluorescence in each event (i.e., nucleus) is directly correlated with the DNA-content of the particle. Thus, by comparing to a known standard, the relative and absolute DNA content of cells in a given sample may be calculated. Endoreduplication indices (EI) are determined by establishing the average number of endocycles per cell within a sample by observing cellular DNA content (C-value) where 1C is the DNA content of a haploid cell (formula presented in step 6.7 of the protocol). For instance, in a diploid organism, the base DNA content of somatic cells is 2C. If a sample has few cells with 4C, corresponding to a single round of endoreduplication, or greater it would have an EI near 0; however if nearly all the cells are 4C the EI would be approximately 1. However, as multiple rounds of endoreduplication are common in higher plants observed EI values may be much greater. While calculating endoreduplication indices may be relatively straightforward, it requires that relative abundances of nuclei C-values be reliably ascertained, which is precluded in certain species and tissues (including potato tubers). It is likely that differences in cellular anatomy, chemistry or cell wall composition13 cause these samples to be recalcitrant to the typical preparations using a razor blade to release the nuclei directly into appropriate buffers that are commonly used with tissues such as tomato pericarp, a model for endoreduplication studies8.
In potato, the examination of endoreduplication within the tuber remains limited to just a couple studies, perhaps due in part to a lack of a reliable protocol as the aforementioned crude preparations yield inconsistent results. While such approaches work well for leaves the tuber samples experience severe degradation and an abundance of noise in the form of debris, making differentiation of peaks comprised of nuclei with differing C-values nearly impossible. This is perhaps due to differences in cellular composition and a profusion of cellular debris within tuber cells (e.g. starch-storing amyloplasts). To overcome this hurdle, we recently developed a more reliable protocol for acquiring distinguishable peaks in flow cytometry of potato tubers. The frequency of cells within each peak can then be used for calculating accurate endoreduplication levels for different genotypes or different tissues that comprise a tuber. The protocol, which employs a modified protoplast extraction method14,15 antecedent previously described flow cytometry11, showed considerable variation within potato relatives, cultivars, and tissues16. Here we present the protocol in detail by evaluating EI in the contexts of ploidy, tissue, and tuber size.

<table>
	<tbody>
		<tr>
			<td>Serological pipette</td>
			<td>Fisher</td>
			<td>13-676-10k</td>
			<td>(sterile)</td>
		</tr>
		<tr>
			<td>Pipet Aid XP (autopipettor)</td>
			<td>Drummond Scientific</td>
			<td>4-000-101</td>
			<td></td>
		</tr>
		<tr>
			<td>Conical tubes (50 ml)</td>
			<td>Thermo Fisher Scientific</td>
			<td>339652</td>
			<td>(sterile)</td>
		</tr>
		<tr>
			<td>Microcentrifuge tubes (1.5 ml)</td>
			<td>Globe Scientific</td>
			<td>111558</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcentrifuge tubes (2 ml)</td>
			<td>Globe Scientific</td>
			<td>111552</td>
			<td></td>
		</tr>
		<tr>
			<td>Vacuum filtration unit (0.22&#181;m)(aPES)</td>
			<td>Thermo Fisher Scientific</td>
			<td>564-0020</td>
			<td>(sterile)</td>
		</tr>
		<tr>
			<td>Vacuum filtration unit (0.45&#181;m)(aPES)</td>
			<td>Thermo Fisher Scientific</td>
			<td>168-0045</td>
			<td>(sterile)</td>
		</tr>
		<tr>
			<td>Cellulase &#34;Onozuka R-10&#34;</td>
			<td>Yakult</td>
			<td></td>
			<td>contact company</td>
		</tr>
		<tr>
			<td>Macerozyme &#34;Onozuka R-10&#34;</td>
			<td>Yakult</td>
			<td></td>
			<td>contact company</td>
		</tr>
		<tr>
			<td>Hemicellulase</td>
			<td>Sigma</td>
			<td>H2125</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris base</td>
			<td>Fisher Scientific</td>
			<td>BP152-1</td>
			<td></td>
		</tr>
		<tr>
			<td>MES buffer</td>
			<td>Acros Organics</td>
			<td>172590250</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>MP Biochemicals</td>
			<td>194854</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium Chloride (hexahydrate)</td>
			<td>Fisher Scientific</td>
			<td>M35-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium Chloride (dihydrate)</td>
			<td>Fisher Scientific</td>
			<td>C79-500</td>
			<td></td>
		</tr>
		<tr>
			<td>D-Mannitol</td>
			<td>Acros Organics</td>
			<td>423922500</td>
			<td></td>
		</tr>
		<tr>
			<td>MOPS</td>
			<td>Acros Organics</td>
			<td>17263-0250</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Phosphate (monobasic, anhydrous)</td>
			<td>Sigma-Aldrich</td>
			<td>P-5379</td>
			<td></td>
		</tr>
		<tr>
			<td>Propidium iodide</td>
			<td>Sigma-Aldrich</td>
			<td>P4170</td>
			<td>For flow cytometry preparations</td>
		</tr>
		<tr>
			<td>Ribonuclease A</td>
			<td>Sigma-Aldrich</td>
			<td>R5000</td>
			<td>For flow cytometry preparations</td>
		</tr>
		<tr>
			<td>Cork borers with punch</td>
			<td>American Scientific</td>
			<td>2093-02</td>
			<td>For smapling desired tissue</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Thermo Fisher Scientific</td>
			<td>16-31</td>
			<td>For smapling desired tissue</td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>Thermo Fisher Scientific</td>
			<td>08-920B</td>
			<td>For smapling desired tissue</td>
		</tr>
		<tr>
			<td>106um stainless steel mesh or other seive of same pore size</td>
			<td>New Jersey Wire Mesh Co.</td>
			<td></td>
			<td>contact company</td>
		</tr>
	</tbody>
</table>

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.

Production of protoplasts
The generation of protoplasts is necessary to achieve repeatable flow cytometry results from potato tubers, the general morphology of which is displayed in Figure 1. Researchers may wish to ensure high quality protoplasts have been produced prior to the addition of the FCP, especially in the event that troubleshooting is required. The majority of the protoplasts should be spherical with minimal protrusions (Figure 2) and may differ greatly in size, which is perhaps reflective of differences in EI.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57134/57134fig2.jpg" /> 
Figure 2: Representative leaf and tuber protoplasts acquired at step 4.4 of the protocol. Isolated protoplasts should be spherical and symmetrical with plasma membrane intact. Note the size difference between leaf (A-C) and tuber (D, E) protoplasts as well as the differences in size within a tissue which may indicate different levels of endoreduplication. Leaf protoplasts contain chloroplasts whereas amyloplasts are visible within the tuber protoplasts. Scale bars = 1,000 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57134/57134fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Evaluation of flow cytometry results
The success or failure of an experiment may be gauged from the width of peaks and their separation in the flow cytometry histograms. As measuring endoreduplication requires calculating relative abundance of nuclei in each peak, mere presence or absence of peaks is insufficient if there is too much noise to draw meaningful boundaries between them. Figure 3 displays the variation in results researchers may encounter in both the flow cytometry scatter plots and histograms. Potential causes for failed samples are presented within the discussion.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57134/57134fig3.jpg" /> 
Figure 3: Representative results obtained from flow cytometry of protoplast nuclei of intact and degraded pith samples. Intact nuclei (A) show clean separation between peaks to quantify relative abundance of cells in each using the appropriate software whereas peaks in degraded samples (B) show shifted, wide and overlapping bases. In the scatterplots of intact (C) and degraded (D) samples, black boxes indicate nuclear event gating for histograms and the clustering of events is reflected in the width of the histogram peaks. Events outside the gates indicate debris, severely degraded nuclei, or other aggregates. PI-H corresponds to intensity of fluorescence which is measured in arbitrary units. Troubleshooting methods to prevent sample degradation are discussed within the text. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57134/57134fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Endoreduplication differences between tissues
Previously we reported that tuber pith tissue has significantly greater levels of endoreduplication than cortex tissue16. To confirm and expand on this we looked at endoreduplication levels of three different tissues in cv. Superior: pith, perimedullary parenchyma, and cortex, replicated in triplicate with 2,000 gated events per sample. Our results confirmed our earlier observation that pith tissue has significantly greater EI than cortical tissue with means of 1.79 and 1.12 endocycles per cell, respectively (p = 0.018; Student&#39;s t-test). Somewhat surprisingly, the parenchyma tissue demonstrated a profile similar to cortex and was also significantly different from pith tissue (mean = 1.14; p = 0.013; Student&#39;s t-test). These results are summarized in Figure 4.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57134/57134fig4.jpg" /> 
Figure 4: Endoreduplication in three tissues of cv. Superior. Histograms of leaf (A), tuber cortex (B), parenchyma (C), and pith (D) tissues are shown along with the EI value calculated from those histograms. Differences in relative abundance of nuclei comprising each peak are readily apparent, especially between leaf and tuber tissues. C-values for each peak are also displayed. PI-H corresponds to intensity of fluorescence which is measured in arbitrary units. Mean comparisons of EI of tuber tissues are displayed in E where asterisk indicates significant difference (p &#60;0.05) and error bars show standard deviation. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57134/57134fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Influence of tuber size and ploidy
We previously reported that, for a given genotype, tubers of different size but similar maturity did not display a corresponding difference in EI16. We aimed to confirm this result as well as evaluate the relationship between ploidy and endoreduplication. For this experiment, we used three replicates of parenchyma tissue from three different genotypes: cv. Superior (4x), VT_SUP_19 (2x) which is a dihaploid extracted from cv. Superior by prickle pollination17, and VT_SUP_19 4x which is a doubled dihaploid18 isogenic to VT_SUP_19. We included a set of replicates for large (90&#8211;130 g) and small (&#60;35 g) tubers for cv. Superior while tubers for the other two genotypes were 90&#8211;130 g. The tubers were all harvested from greenhouse grown plants at full maturity, i.e. the tops of the plants had senesced. We observed a significant difference between VT_SUP_19 and its progenitor Superior (p = 0.04); however, there was no significant difference between VT_SUP_19 and VT_SUP_19 4x (p = 0.69). This indicates that while there is a likely genetic component to endoreduplication, as unmasked by the genomic reduction, it is not dictated by ploidy, at least in this background. Lastly, we once again observed no significant differences between large and small cv. Superior tubers as demonstrated in Figure 5.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/57134/57134fig5.jpg" /> 
Figure 5: Endoreduplication in two sizes of tubers of cv. Superior and its diploid (VT_Sup_19) and tetraploid (VT_Sup_19 4x) derivatives. The bar chart shows the mean for small (&#60;35 g) and large (90&#8211;130 g) cv. Superior tubers as well as large (90&#8211;130 g) tubers of the dihaploid VT_SUP_19 and the isogenic doubled dihaploid VT_SUP_19 4x. Significant differences (p &#60;0.05) are indicated by the connecting letter report. Error bars depict standard deviation. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57134/57134fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Measuring Endoreduplication by Flow Cytometry of Isolated Tuber Protoplasts at JoVE.com

Measuring Endoreduplication by Flow Cytometry of Isolated Tuber Protoplasts

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 ...

measuring-endoreduplication-flow-cytometry-isolated-tuber

Nanyang Technological University

Research

JoVE Journal

Biology

10.1K Views.  Virginia Tech. 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.

Video: Measuring Endoreduplication by Flow Cytometry of Isolated Tuber Protoplasts

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 ...

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

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

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

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

Flow Cytometry Purification of Mouse Meiotic Cells

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the unique differentiation programs that are manifest during meiosis. Two principal methods have been developed to purify, to varying degrees, various meiotic fractions from both adult and immature animals: elutriation or Staput (sedimentation) using BSA and/or percoll gradients. Both of these methods rely on cell size and density to separate meiotic cells1-5. Overall, except for few cell populations6, these protocols fail to yield sufficient purity of the numerous meiotic cell populations that are necessary for detailed molecular analyses. Moreover, with such methods usually one type of meiotic cells can be purified at a given time, which adds an extra level of complexity regarding the reproducibility and homogeneity when comparing meiotic cell samples.
Here, we describe a refined method that allows one to easily visualize, identify, and purify meiotic cells, from germ cells to round spermatids, using FACS combined with Hoechst 33342 staining7,8. This method provides an overall snapshot of the entire meiotic process and allows one to highly purify viable cells from most stage of meiosis. These purified cells can then be analyzed in detail for molecular changes that accompany progression through meiosis, for example changes in gene expression9,10and the dynamics of nucleosome occupancy at hotspots of meiotic recombination11.

An efficient method to obtain highly purified viable meiotic fractions from mouse testis is described, which combines a refined cell dissociation protocol with fluorescent activated cell sorting (FACS). This method takes advantage of differences in the DNA content and nuclear density of discrete meiotic fractions.

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the ...

Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating cells. Cell cycle division is an integral part of growth and reproduction and deregulation of key cell cycle components have been implicated in the precipitating events of carcinogenesis 1,2. Molecular agents in anti-cancer therapies frequently target biological pathways responsible for the regulation and coordination of cell cycle division 3. Although cell cycle kinetics tend to vary according to cell type, the distribution of cells amongst the four stages of the cell cycle is rather consistent within a particular cell line due to the consistent pattern of mitogen and growth factor expression. Genotoxic events and other cellular stressors can result in a temporary block of cell cycle progression, resulting in arrest or a temporary pause in a particular cell cycle phase to allow for instigation of the appropriate response mechanism. 
The ability to experimentally observe the behavior of a cell population with reference to their cell cycle progression stage is an important advance in cell biology. Common procedures such as mitotic shake off, differential centrifugation or flow cytometry-based sorting are used to isolate cells at specific stages of the cell cycle 4-6. These fractionated, cell cycle phase-enriched populations are then subjected to experimental treatments. Yield, purity and viability of the separated fractions can often be compromised using these physical separation methods. As well, the time lapse between separation of the cell populations and the start of experimental treatment, whereby the fractionated cells can progress from the selected cell cycle stage, can pose significant challenges in the successful implementation and interpretation of these experiments.
Other approaches to study cell cycle stages include the use of chemicals to synchronize cells. Treatment of cells with chemical inhibitors of key metabolic processes for each cell cycle stage are useful in blocking the progression of the cell cycle to the next stage. For example, the ribonucleotide reductase inhibitor hydroxyurea halts cells at the G1/S juncture by limiting the supply of deoxynucleotides, the building blocks of DNA. Other notable chemicals include treatment with aphidicolin, a polymerase alpha inhibitor for G1 arrest, treatment with colchicine and nocodazole, both of which interfere with mitotic spindle formation to halt cells in M phase and finally, treatment with the DNA chain terminator 5-fluorodeoxyridine to initiate S phase arrest 7-9. Treatment with these chemicals is an effective means of synchronizing an entire population of cells at a particular phase. With removal of the chemical, cells rejoin the cell cycle in unison. Treatment of the test agent following release from the cell cycle blocking chemical ensures that the drug response elicited is from a uniform, cell cycle stage-specific population. However, since many of the chemical synchronizers are known genotoxic compounds, teasing apart the participation of various response pathways (to the synchronizers vs. the test agents) is challenging. 
Here we describe a metabolic labeling method for following a subpopulation of actively cycling cells through their progression from the DNA replication phase, through to the division and separation of their daughter cells. Coupled with flow cytometry quantification, this protocol enables for measurement of kinetic progression of the cell cycle in the absence of either mechanically- or chemically- induced cellular stresses commonly associated with other cell cycle synchronization methodologies 10. In the following sections we will discuss the methodology, as well as some of its applications in biomedical research.

Tracking subtle changes in the progression and kinetics of cell cycle stages can be accomplished by use of a combination of metabolic labeling of nucleic acids with BrdU and total genomic DNA staining via Propidium Iodide. This method avoids the need of chemical synchronization of cycling cells, thereby preventing the introduction of non-specific DNA damage, which in turn affects cell cycle progression.

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating ...

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 ...

Measuring the Osmotic Water Permeability Coefficient (Pf) of Spherical Cells: Isolated Plant Protoplasts as an Example

Studying AQP regulation mechanisms is crucial for the understanding of water relations at both the cellular and the whole plant levels. Presented here is a simple and very efficient method for the determination of the osmotic water permeability coefficient (Pf) in plant protoplasts, applicable in principle also to other spherical cells such as frog oocytes. The first step of the assay is the isolation of protoplasts from the plant tissue of interest by enzymatic digestion into a chamber with an appropriate isotonic solution. The second step consists of an osmotic challenge assay: protoplasts immobilized on the bottom of the chamber are submitted to a constant perfusion starting with an isotonic solution and followed by a hypotonic solution. The cell swelling is video recorded. In the third step, the images are processed offline to yield volume changes, and the time course of the volume changes is correlated with the time course of the change in osmolarity of the chamber perfusion medium, using a curve fitting procedure written in Matlab (the &#8216;PfFit&#8217;), to yield Pf.

Measuring the Osmotic Water Permeability Coefficient Pf of Spherical Cells: Isolated Plant Protoplasts as an Example

Measuring the osmotic water permeability coefficient (Pf) of cells can help understand the regulatory mechanisms of aquaporins (AQPs). Pf determination in spherical plant cell protoplasts presented here involves protoplasts isolation and numerical analysis of their initial rate of volume change as a result of an osmotic challenge during constant bath perfusion.

Studying AQP regulation mechanisms is crucial for the understanding of water relations at both the cellular and the whole plant levels. Presented here is ...

Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry

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

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

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

Immunodetection of Outer Membrane Proteins by Flow Cytometry of Isolated Mitochondria

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

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

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

LED Thermo Flow &#8212; Combining Optogenetics with Flow Cytometry

Optogenetic tools allow isolated, functional investigations of almost any signaling molecule within complex signaling pathways. A major obstacle is the controlled delivery of light to the cell sample and hence the most popular tools for optogenetic studies are microscopy-based cell analyses and in vitro experiments. The flow cytometer has major advantages over a microscope, including the ability to rapidly measure thousands of cells at single cell resolution. However, it is not yet widely used in optogenetics. Here, we present a device that combines the power of optogenetics and flow cytometry: the LED Thermo Flow. This device illuminates cells at specific wavelengths, light intensities and temperatures during flow cytometric measurements. It can be built at low cost and be used with most common flow cytometers. To demonstrate its utility, we characterized the photoswitching kinetics of Dronpa proteins in vivo and in real time. This protocol can be adapted to almost all optically controlled substances and substantially expands the set of possible experiments. More importantly, it will greatly simplify the discovery and development of new optogenetic tools.

Optically controlled substances are powerful tools to study signaling pathways. To expand the spectrum of possible experiments, we developed a device for studying optically controlled substances in real time using flow cytometry: the LED Thermo Flow.

Optogenetic tools allow isolated, functional investigations of almost any signaling molecule within complex signaling pathways. A major obstacle is the ...

Analysis of Cell Suspensions Isolated from Solid Tissues by Spectral Flow Cytometry

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to the analysis of a few fluorochromes, currently there are dozens of different fluorescent dyes, and up to 14-18 different dyes can be combined at a time. However, several limitations still impair the analytical capabilities. Because of the multiplicity of fluorescent probes, data analysis has become increasingly complex due to the need of large, multi-parametric compensation matrices. Moreover, mutant mouse models carrying fluorescent proteins to detect and trace specific cell types in different tissues have become available, so the analysis (by flow cytometry) of auto-fluorescent cell suspensions obtained from solid organs is required. Spectral flow cytometry, which distinguishes the shapes of emission spectra along a wide range of continuous wavelengths, addresses some of these problems. The data is analyzed with an algorithm that replaces compensation matrices and treats auto-fluorescence as an independent parameter. Thus, spectral flow cytometry should be capable of discriminating fluorochromes with similar emission peaks and can provide a multi-parametric analysis without compensation requirements.
This protocol describes the spectral flow cytometry analysis, allowing for a 21-parameter (19 fluorescent probes) characterization and the management of an auto-fluorescent signal, providing high resolution in minor population detection. The results presented here show that spectral flow cytometry presents advantages in the analysis of cell populations from tissues difficult to characterize in conventional flow cytometry, such as the heart and the intestine. Spectral flow cytometry thus demonstrates the multi-parametric analytical capacity of high-performing conventional flow cytometry without the requirement for compensation and enables auto-fluorescence management.

This article describes spectral cytometry, a new approach in flow cytometry that uses the shapes of emission spectra to distinguish fluorochromes. An algorithm replaces compensations and can treat auto-fluorescence as an independent parameter. This new approach allows for the proper analysis of cells isolated from solid organs.

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to ...