We would like to thank David Rodriguez for assistance with the figure preparation and Precision Pathology Services (San Antonio, TX) for the use of its Navios EX flow cytometer.

All procedures need to be done using appropriate personal protective equipment.
1. Preparation of the TCPP stock solution, 1.0 mg/ mL
NOTE: This can be prepared monthly.
<ol>
	<li>Using an analytical balance, spatula, and weighing paper, weigh 49.0-50.9 mg of TCPP. Round the weight to 1/10 of a milligram. Set the measured amount of TCPP aside protected from light. 
	NOTE: Use a static gun if the weight reading is unstable.</li>
	<li>Determine the required amounts of purified water and isopropanol (IPA) from Table 1 based on the amount of TCPP weighed in step 1.1. Add the purified water and isopropanol to a 100 mL glass beaker, and cover with parafilm to protect from evaporation.</li>
	<li>Determine the required amount of sodium bicarbonate from Table 1 based on the amount of TCPP weighed in step 1.1.</li>
	<li>Using the analytical balance, spatula, and weighing paper, weigh the required amount of sodium bicarbonate determined in step 1.3. Round the weight to 1/10 of a milligram. 
	NOTE: Use a static gun if the weight reading is unstable.</li>
	<li>Add the sodium bicarbonate to the&#160;100 mL beaker containing purified water and isopropanol. Cover the solution with parafilm to protect from evaporation.</li>
	<li>Place the solution from step 1.5 on a stir plate, and stir until dissolved (approx. 10 min)</li>
	<li>Measure the pH to ensure that the sodium bicarbonate-containing solution from step 1.6 has a pH between 9 and 10.</li>
	<li>Slowly add the TCPP weighed in step 1.1 to the solution from step 1.7, and continue stirring until dissolved (~30 min). Protect from light during this step.</li>
	<li>Store in a glass or polypropylene container at room temperature and protected from light.</li>
</ol>
2. Preparation of 2-(N -morpholino)-ethanesulfonic acid (MES) and hemisodium salt buffer solution, 0.1 M, pH 6.0-6.2 (&#34;MES buffer&#34;)
NOTE: This must be prepared on the day of use and kept at room temperature.
<ol>
	<li>Weigh out 2.50 g of MES hemisodium salt, and add this to a 150 mL plastic bottle.</li>
	<li>Add 121 mL of purified water, and dissolve by manual shaking until no solid is visible.</li>
	<li>Measure the pH of the MES buffer to ensure that it is between 6 and 6.2.</li>
	<li>Maintain at room temperature for use on the same day.</li>
</ol>
3. N-(3-Dimethlyaminopropyl)-N&#39;-ethylcarbodiimide (EDC) powder
<ol>
	<li>Take the EDC powder out of the freezer, and let sit at room temperature until use in step 5.</li>
</ol>
4. Combining amine-functionalized polystyrene beads with TCPP solution
<ol>
	<li>Add 4.3 mL of the 0.1 M MES buffer solution prepared in step 2 to a 15 mL polypropylene tube.</li>
	<li>Vortex the amine-functionalized polystyrene bead suspension (10 &#181;m, 2.5% w/v) for 60 s at maximum speed.</li>
	<li>Add 288 &#181;L of this freshly vortexed bead suspension to the MES buffer from step 4.1.</li>
	<li>Vortex the MES/bead solution for 15 s at maximum speed.</li>
	<li>Vortex the 1 mg/mL TCPP solution prepared in step 1 for 60 s at maximum speed.</li>
	<li>Add 1.20 mL of this freshly vortexed TCPP stock solution to the MES/bead suspension from step 4.4.</li>
	<li>Vortex the MES/bead/TCPP suspension for 15 s at maximum speed.</li>
	<li>Cover the tube with foil while the EDC solution is prepared.</li>
</ol>
5. Preparation of N-(3-dimethlyaminopropyl)-N&#39;-ethylcarbodiimide (EDC) hydrocholoride (HCl) stock solution
NOTE: The EDC solution is perishable and should be used immediately following preparation.
<ol>
	<li>Add 20.0 mL of purified water to a new 50 mL conical tube.</li>
	<li>Weigh out 200 mg of EDC HCl from step 3, and add it to the water (step 5.1).</li>
	<li>Vortex the EDC HCL for 15 s at maximum speed to generate a clear solution.</li>
</ol>
6. Preparation of the EDC HCl/MES working solution
NOTE: The EDC HCl/MES solution is perishable and should be used immediately following preparation.
<ol>
	<li>Add 54.0 mL of MES buffer solution (prepared in step 2) to a 150 mL plastic bottle.</li>
	<li>Add 6.0 mL of the EDC HCl stock solution (prepared in step 5) to the MES buffer solution, and mix by shaking for 10 s.</li>
</ol>
7. Labeling the beads with TCPP
<ol>
	<li>Add 4.5 mL of EDC working solution (from step 6) to the 15 mL polypropylene tube containing the beads and TCPP in MES buffer (step 4.7).</li>
	<li>Place the tube in an inverting rotator at 35 rpm for 16 h at room temperature and protected from light.</li>
	<li>Centrifuge the tube at room temperature for 10 min at 1,000 &#215; g.</li>
	<li>Aspirate the supernatant, and resuspend the beads in 0.8 mL of Hanks&#39; balanced salt solution (HBSS).</li>
	<li>Transfer the bead solution to an amber polypropylene vial with a 1 mL pipette, and store at 4 &#176;C until further use. 
	&#8203;NOTE: The beads are stable for at least 3 months at this point.</li>
</ol>
8. Quality check (QC) of the TCPP beads by flow cytometry
NOTE: The QC should be centered on whether the median fluorescence intensity (MFI) of the TCPP beads is sufficiently bright for their intended use and the amount of particulates generated by the procedure. See the representative results section for more details.
<ol>
	<li>Label one 5 mL polystyrene tube as &#34;TCPP negative beads.&#34; 
	NOTE: The negative beads are different from the amine-functionalized beads used for labeling. See the Table of Materials.</li>
	<li>Label another tube as &#34;TCPP positive beads.&#34;</li>
	<li>Label another tube as &#34;Rainbow beads.&#34;</li>
	<li>Aliquot 300 &#181;L of ice-cold HBSS into the tubes labeled with &#34;TCPP negative beads&#34; and &#34;TCPP positive beads.&#34;</li>
	<li>Add 500 &#181;L of ice-cold HBSS into the tube labeled as &#34;Rainbow beads.&#34;</li>
	<li>Vortex the non-functionalized polystyrene (unlabeled) bead suspension briefly at maximum speed (2-3 s), and add 10 &#181;L of it to the tube labeled &#34;TCPP negative beads.&#34;</li>
	<li>Vortex the TCPP-labeled bead suspension (finalized in step 7.5) briefly at maximum speed, and add 3 &#181;L of it to the &#34;TCPP positive beads&#34; tube.</li>
	<li>Vortex the Rainbow bead suspension briefly at maximum speed, and add two drops of it into the &#34;Rainbow beads&#34; tube.</li>
	<li>Keep all tubes on ice, covered, and protected from light.</li>
	<li>Initiate the appropriate daily startup procedures of the flow cytometer, and perform a QC to verify the optimal fluids and laser alignment. 
	NOTE: For this part of the protocol, it is assumed that the operator is trained in the use of the flow cytometer that is available, including the procedures of standardizing the light scatter and fluorescence intensity, as well as the basic principles of calculating the correct compensation matrix.</li>
	<li>Run the Rainbow beads and the TCPP beads without changing the voltage settings in between the different runs.
	<ol>
		<li>Run and collect 10,000 events of the Rainbow beads.</li>
		<li>Perform a rinse with water, and collect 10,000 events of the TCPP-negative beads.</li>
		<li>Perform a rinse with water, and collect 10,000 events of the TCPP-positive beads.</li>
		<li>Perform a 1 min water rinse. 
		NOTE: It is important to perform a rinse with water after running the TCPP beads. If the TCPP is not rinsed from the lines in the cytometer, there is the possibility that residual TCPP can label cells in the next tube to be acquired.</li>
		<li>Perform the appropriate cleaning and shutdown protocols specific to the manufacturer&#39;s instructions for the cytometer. 
		&#8203;NOTE: For representative results, see Figure 1.</li>
	</ol>
	</li>
</ol>
9. Bead filtration
NOTE: If the QC of the beads by flow cytometry (step 8) shows a high proportion of particulates (70% or higher), consider filtering the bead suspension using the protocol below (Figure 2).
<ol>
	<li>Add 3.20 mL of ice-cold HBSS to 0.8 mL of the TCPP bead suspension finalized in step 7.5 (creating a five-fold dilution).</li>
	<li>Vortex the diluted bead suspension at maximum speed for 15 s.</li>
	<li>Remove the plunger from a disposable 5 mL syringe.</li>
	<li>Fit the syringe with a glass fiber tip filter (5 &#181;m, 13 mm diameter).</li>
	<li>Add 4 mL of HBSS to the syringe.</li>
	<li>Add 0.5 mL of the vortexed diluted bead suspension (step 9.2).</li>
	<li>Use the plunger to filter the suspension through the syringe/filter setup at approximately 2 drops/s.</li>
	<li>Wash the beads by drawing 5 mL of fresh HBSS into the syringe through the filter at approximately 2 drops/s.</li>
	<li>Push the HBSS out again into the waste container at approximately 2 drops/s.</li>
	<li>To remove the beads from the filter, draw another 5 mL of fresh HBSS into the syringe through the filter.</li>
	<li>Carefully remove the filter from the syringe.</li>
	<li>Eject the bead suspension from the syringe into a 50 mL conical centrifuge tube.</li>
	<li>Place the filter back on the syringe, and repeat steps 9.10-9.12 four more times. Then&#160;discard the filter and syringe.</li>
	<li>Repeat steps 9.2-9.13 until all the beads from step 9.1 have been filtered. Use a fresh syringe and filter each time.</li>
	<li>Centrifuge the filtered bead suspensions for 10 min at 1,000 &#215; g at room temperature.</li>
	<li>Aspirate the supernatant of each 50 mL tube, and gently resuspend the beads, combining them in 0.5 mL of fresh HBSS.</li>
	<li>Transfer the beads with a p1,000 micropipette to a new amber, glass, or polypropylene vial, and store at 4 &#176;C.</li>
	<li>Repeat step 8 to determine if the proportion of TCPP-related particulates in the filtered bead suspension has decreased. For representative results, see Figure 3.</li>
</ol>

Porphyrin-Modified Beads in Flow Cytometry

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Next-generation+photodynamic+therapy:+New+probes+for+cancer+imaging+and+treatment.">Next-generation photodynamic therapy: New probes for cancer imaging and treatment.</a> Biochemistry. 57 (2), 173-174 (2018).</li><li>El-Far, M., Pimstone, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+comparative+study+of+28+porphyrins+and+their+abilities+to+localize+in+mammary+mouse+carcinoma:+Uroporphyrin+I+superior+to+hematoporphyrin+derivative.">A comparative study of 28 porphyrins and their abilities to localize in mammary mouse carcinoma: Uroporphyrin I superior to hematoporphyrin derivative.</a> Progress in Clinical and Biological Research. 170, 661-672 (1984).</li></ol>

All authors are employees of bioAffinity Technologies.

Despite the many applications of porphyrins in cancer diagnosis and therapeutics2, there is limited literature on their potential use as a flow cytometric reagent for the identification of cancerous versus non-cancerous cell populations in primary human tissues24,25,26. Our research on the flow cytometric analysis of human sputum24,27 requires the...

Porphyrins have been of interest for many years in the biomedical field owing to their fluorescence and tumor-targeting properties1,2,3. Therapeutic applications such as photodynamic therapy (PDT) and sonodynamic therapy (SDT) entail the systemic administration of a porphyrin to a cancer patient, the accumulation of the drug in the tumor, and the localized exposure of the tumor to a laser light of a specific wavelength or ultrasound. The exposure to laser light or ultrasound leads to the generation of reactive oxygen species by the porphyrin and subsequent cell death4,5. In photodynamic diagnosis (PDD), porphyrin fluorescence is used to distinguish cancer cells from normal cells6. In this context, protoporphyrin IX, a natural fluorescent porphyrin that accumulates in tumors upon the systemic or local injection of its precursor, 5-aminolevulinic acid (5-ALA), is used to identify gastrointestinal stromal tumors, bladder cancer, and brain cancer7,8. More recently, 5-ALA treatment was explored as an approach to detect minimal residual disease in multiple myeloma9. Our laboratory has been using the tetraaryl porphyrin TCPP (5,10,15,20-tetrakis-(4-carboxyphenyl)-21,23H-porphine) for its ability to selectively stain lung cancer cells and cancer-associated cells in human sputum samples, which is a property that has been exploited in slide-based and flow cytometric diagnostic assays10.
Some porphyrins are bifunctional in that they can be used as therapeutic and diagnostic agents2,11. In biomedical research, such bifunctional porphyrins are used to evaluate how their ability to selectively target and kill cancer cells is a function of their structure as well as how it is affected by the presence of other compounds12,13,14,15,16. Both the cellular uptake of porphyrins and their cytotoxicity can be measured on a flow cytometric platform in a high-throughput manner. The absorption and emission spectra of fluorescent porphyrins are complex, but most flow cytometric platforms are equipped to correctly identify them. The absorption spectrum of fluorescent porphyrins is characterized by a strong absorption band in the 380-500 nm range, known as the Soret band. Two to four weaker absorption bands are generally observed in the 500-750 nm range (Q bands)17. A blue&#160;488 nm laser, present in most flow cytometers, or a violet laser (405 nm) can generate light of the appropriate wavelength to excite porphyrins. The emission spectra of porphyrins typically display peaks in the 600-800 nm range18, which results in very little spectral overlap with fluorescein isothiocyanate or phycoerythrin (PE) fluorophores but considerable overlap with other often-used fluorophores, such as allophycocyanin (APC), as well as tandem fluorophores, such as PE-Cy5 and others. Therefore, when using porphyrins in multi-color flow cytometry assays, single-fluorophore controls are essential to adequately correct the spillover of fluorescence in channels other than the one designated to measure the porphyrin&#39;s fluorescence.
Ideally, the single-fluorophore controls used to calculate the spillover matrix for a panel of fluorophores (also called &#34;compensation controls&#34;) should consist of the same cell type(s) as the sample. However, using the sample for this purpose is not optimal if there is very little sample to begin with or if the target population within the sample is very small&#160;(for example, if one wants to look at minimal residual disease or cancer cells at the early stages of the disease). A useful alternative to cells is beads coupled with the same fluorophore that is used to analyze the sample. Many such beads are commercially available; these beads are either prelabeled with the desired fluorophore (prelabeled fluorophore-specific beads)19,20, or a fluorescently labeled antibody can be attached to them (antibody capture beads)20,21. While commercial compensation beads are available for many fluorophores, such beads are unavailable for porphyrins, despite their increasing use in basic and clinical research.
In addition to sample preservation and appropriately sized positive versus negative populations, the other advantages of using beads as compensation controls are the ease of preparation, low background fluorescence, and excellent stability over time22. The potential disadvantage of using beads as a compensation control is that the emission spectrum of the fluorescent antibody captured on beads may differ from that of the same antibody used to label the cells. This may be of specific importance when using a spectral flow cytometer20. Therefore, the development of beads as a compensation control needs to be performed on the flow cytometer that will be used for the assay for which the beads are developed. Moreover, the development of the beads needs to include a comparison with cells labeled with the same fluorescent staining reagent.
Here, we describe the preparation of TCPP amine-functionalized polystyrene compensation beads, whose median fluorescence intensity in the detection channel was comparable to that of TCPP-labeled cells in sputum, and their use as compensation controls for flow cytometry. The autofluorescence of equivalent, non-functionalized beads was sufficiently low for their use as negative fluorescence compensation controls. In addition, these beads demonstrated stability in storage for nearly 1 year.

<table><tbody><tr><td>Amber plastic vials, 2 mL, U- bottom, polypropylene</td><td>Research Products International&amp;nbsp;&amp;nbsp;</td><td>ZC1028-500</td><td></td></tr><tr><td>Amine-funtionalized polystyrene divinylbenzene crosslinked (PS/DVB) beads, 10.6 &amp;mu;m diameter, 2.5% w/v aqueous suspension, 3.82 x 10&lt;sup&gt;7&lt;/sup&gt; beads/mL, 7.11 x 10&lt;sup&gt;11 &lt;/sup&gt;amine groups/ bead</td><td>Spherotech</td><td>APX-100-10</td><td>Diameter spec. 8.0-12.9 um, suspension 2.5% w/v 3.82 x 10&lt;sup&gt;7&lt;/sup&gt; beads/mL, 7.11 x 10&lt;sup&gt;11&lt;/sup&gt; amine groups/ bead</td></tr><tr><td>Conical tubes, 50 mL, Falcon</td><td>Fisher Scientific</td><td>14-432-22</td><td></td></tr><tr><td>Centrifuge</td><td></td><td></td><td>with appropriate rotor</td></tr><tr><td>Disposable polystyrene bottle with cap, 150 mL</td><td>Fisher Scientific</td><td>09-761-140</td><td></td></tr><tr><td>EDC (N- (3- dimethylaminopropyl)- N&#039;- ethylcarbodiimide hydrochloride), &amp;ge;98%</td><td>Sigma</td><td>03450-1G</td><td>CAS No:&amp;nbsp; 25952-53-8</td></tr><tr><td>FlowJo Single Cell Analysis Software (v10.6.1)</td><td>BD</td><td></td><td></td></tr><tr><td>Glass coverslips, 22 x 22 mm</td><td>Fisher Scientific</td><td>12-540-BP</td><td></td></tr><tr><td>Glass fiber syringe filters (Finneran, 5 &amp;micro;m, 13 mm diameter)</td><td>Thomas Scientific</td><td>1190M60</td><td></td></tr><tr><td>Glass microscope slides, 275 x 75 x 1 mm</td><td>Fisher Scientific</td><td>12-550-143</td><td></td></tr><tr><td>Hanks Balanced Salt Solution (HBSS)</td><td>Fisher Scientific</td><td>14-175-095</td><td></td></tr><tr><td>Isopropanol, ACS grade</td><td>Fisher Scientific</td><td>AC423830010</td><td></td></tr><tr><td>Mechanical pipette, 1 channel, 100-1000 uL with tips</td><td>Eppendorf</td><td>3123000918</td><td></td></tr><tr><td>MES (22- (N- mopholino)- N&#039;- ethanesulfonic acid, hemisodium salt</td><td>Sigma</td><td>M0164</td><td>CAS No:&amp;nbsp; 117961-21-4</td></tr><tr><td>Navios EX flow cytometer</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>Olympus BX-40 microscope with DP73 camera and 40X objective with cellSens software</td><td>Olympus</td><td></td><td>or similar</td></tr><tr><td>Pasteur pipettes, glass, 5.75&quot;</td><td>Fisher Scientific</td><td>13-678-6B</td><td></td></tr><tr><td>pH meter (UB 10 Ultra Basic)</td><td>Denver Instruments</td><td></td><td></td></tr><tr><td>Pipette controller (Drummond)</td><td>Pipete.com</td><td>DP101</td><td></td></tr><tr><td>Plastic Syringe, 5 mL</td><td>Fisher Scientific</td><td>14955452</td><td></td></tr><tr><td>Polystyrene Particles (non-functionalized), SPHERO,&amp;nbsp; 2.5% w/v, 8.0-12.9 &amp;micro;m</td><td>Spherotech</td><td>PP-100-10&amp;nbsp;</td><td></td></tr><tr><td>Polypropylene tubes, 15mL, conical</td><td>Fisher Scientific</td><td>14-959-53A</td><td></td></tr><tr><td>Polystyrene tubes, round bottom&amp;nbsp;</td><td>Fisher Scientific</td><td>14-959-2A</td><td></td></tr><tr><td>Rainbow Beads (Spherotech URCP-50-2K)</td><td>Fisher Scientific</td><td>NC9207381</td><td></td></tr><tr><td>Serological pipettes, disposable - 10 mL</td><td>Fisher Scientific</td><td>07-200-574</td><td></td></tr><tr><td>Serological pipettes, disposable - 25 mL</td><td>Fisher Scientific</td><td>07-200-576</td><td></td></tr><tr><td>Sodium bicarbonate (NaHCO&lt;sub&gt;3&lt;/sub&gt;)</td><td>Sigma</td><td>S6014</td><td>CAS No:&amp;nbsp; 144-55-8</td></tr><tr><td>TCPP (meso-tetra(4-carboxyphenyl)porphine)&amp;nbsp; Frontier Scientific&amp;nbsp;</td><td>Fisher Scientific</td><td>50-393-68</td><td>CAS No:&amp;nbsp; 14609-54-2</td></tr><tr><td>Tecan Spark Plate Reader (or similar)</td><td>Tecan Life Sciences</td><td></td><td></td></tr><tr><td>Tube revolver/rotator</td><td>Thermo Fisher</td><td>88881001</td><td></td></tr><tr><td>Vortex mixer</td><td>Fisher Scientific</td><td>2215365</td><td></td></tr></tbody></table>

porphyrin-modified beads for use as compensation controls in flow cytometry

Flow cytometry can rapidly characterize and quantify diverse cell populations based on fluorescence measurements. The cells are first stained with one or more fluorescent reagents, each functionalized with a different fluorescent molecule (fluorophore) that binds to cells selectively based on their phenotypic characteristics, such as cell surface antigen expression. The intensity of fluorescence from each reagent bound to cells can be measured on the flow cytometer using channels that detect a specified range of wavelengths. When multiple fluorophores are used, the light from individual fluorophores often spills over into undesired detection channels, which requires a correction to the fluorescence intensity data in a process called compensation. 
Compensation control particles, typically polymer beads bound to a single fluorophore, are needed for each fluorophore used in a cell labeling experiment. Data from compensation particles from the flow cytometer are used to apply a correction to the fluorescence intensity measurements. This protocol describes the preparation and purification of polystyrene compensation beads covalently functionalized with the fluorescent reagent meso-tetra(4-carboxyphenyl) porphine (TCPP) and their application in flow cytometry compensation. In this work, amine-functionalized polystyrene beads were treated with TCPP and the amide coupling reagent EDC (N-(3-dimethylaminopropyl)-N&#8242;-ethylcarbodiimide hydrochloride) at pH 6 and at room temperature for 16 h with agitation. The TCPP beads were isolated by centrifugation and resuspended in a pH 7 buffer for storage. TCPP-related particulates were observed as a byproduct. The number of these particulates could be reduced using an optional filtration protocol. The resultant TCPP beads were successfully used on a flow cytometer for compensation in experiments with human sputum cells labeled with multiple fluorophores. The TCPP beads proved stable following storage in a refrigerator for 300 days.

This protocol for the TCPP labeling of beads is relatively fast and efficient. Figure 1 shows a representative outcome of the TCPP bead-labeling process as determined by flow cytometry. Figure 1A shows the standardized profile of Rainbow beads, as detected in the appropriate channel for detecting TCPP. These beads serve as a QC for the standardization of the laser voltages for the detection of TCPP by the flow cytometer. Figure 1B s...

Watch this Scientific Journal Video about Porphyrin-Modified Beads for Use as Compensation Controls in Flow Cytometry at JoVE.com

Author Spotlight: Porphyrin-Modified Beads for Use as Compensation Controls in Flow Cytometry  

The protocol describes how porphyrin-based compensation beads for flow cytometry are prepared by the reaction of amine-functionalized polystyrene beads with the porphyrin TCPP and the amide coupling reagent EDC. A filtration procedure is used to reduce the particulate byproducts.

Porphyrin-Modified Beads for Use as Compensation Controls in Flow Cytometry

Flow cytometry can rapidly characterize and quantify diverse cell populations based on fluorescence measurements. The cells are first stained with one or ...

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Research

JoVE Journal

Chemistry

2.0K Views.  bioAffinity Technologies. The protocol describes how porphyrin-based compensation beads for flow cytometry are prepared by the reaction of amine-functionalized polystyrene beads with the porphyrin TCPP and the amide coupling reagent EDC. A filtration procedure is used to reduce the particulate byproducts. 

Video: Author Spotlight: Porphyrin-Modified Beads for Use as Compensation Controls in Flow Cytometry  

Comprehensive Protocol to Sample and Process Bone Marrow for Measuring Measurable Residual Disease and Leukemic Stem Cells in Acute Myeloid Leukemia

Response criteria in acute myeloid leukemia (AML) has recently been re-established, with morphologic examination utilized to determine whether patients have achieved complete remission (CR). Approximately half of the adult patients who entered CR will relapse within 12 months due to the outgrowth of residual AML cells in the bone marrow. The quantitation of these remaining leukemia cells, known as minimal or measurable residual disease (MRD), can be a robust biomarker for the prediction of these relapses. Moreover, retrospective analysis of several studies has shown that the presence of MRD in the bone marrow of AML patients correlates with poor survival. Not only is the total leukemic population, reflected by cells harboring a leukemia associated immune-phenotype (LAIP), associated with clinical outcome, but so is the immature low frequency subpopulation of leukemia stem cells (LSC), both of which can be monitored through flow cytometry MRD or MRD-like approaches. The availability of sensitive assays that enable detection of residual leukemia (stem) cells on the basis of disease-specific or disease-associated features (abnormal molecular markers or aberrant immunophenotypes) have drastically improved MRD assessment in AML. However, given the inherent heterogeneity and complexity of AML as a disease, methods for sampling bone marrow and performing MRD and LSC analysis should be harmonized when possible. In this manuscript we describe a detailed methodology for adequate bone marrow aspirate sampling, transport, sample processing for optimal multi-color flow cytometry assessment, and gating strategies to assess MRD and LSC to aid in therapeutic decision making for AML patients.

Detection of minimal or measurable residual disease (MRD) is an important prognostic biomarker for refining risk assessment and predicting relapse in acute myeloid leukemia (AML). These comprehensive guidelines and recommendations with best practices for consistent and accurate identification and detection of MRD, may aid in making effective AML treatment decisions.

Response criteria in acute myeloid leukemia (AML) has recently been re-established, with morphologic examination utilized to determine whether patients ...

Cancer Research

Phospho Flow Cytometry with Fluorescent Cell Barcoding for Single Cell Signaling Analysis and Biomarker Discovery

Aberrant cell signaling plays a central role in cancer development and progression. Most novel targeted therapies are indeed directed at proteins and protein functions, and cell signaling aberrations may therefore serve as biomarkers to indicate personalized treatment options. As opposed to DNA and RNA analyses, changes in protein activity can more efficiently evaluate the mechanisms underlying drug sensitivity and resistance. Phospho flow cytometry is a powerful technique that measures protein phosphorylation events at the cellular level, an important feature that distinguishes this method from other antibody-based approaches. The method allows for simultaneous analysis of multiple signaling proteins. In combination with fluorescent cell barcoding, larger medium- to high-throughput data-sets can be acquired by standard cytometer hardware in short time. Phospho flow cytometry has applications both in studies of basic biology and in clinical research, including signaling analysis, biomarker discovery and assessment of pharmacodynamics. Here, a detailed experimental protocol is provided for phospho flow analysis of purified peripheral blood mononuclear cells, using chronic lymphocytic leukemia cells as an example.

Here, a protocol for medium- to high-throughput analysis of protein phosphorylation events at the cellular level is presented. Phospho flow cytometry is a powerful approach to characterize signaling aberrations, identify and validate biomarkers, and assess pharmacodynamics.

Aberrant cell signaling plays a central role in cancer development and progression. Most novel targeted therapies are indeed directed at proteins and ...

Identification Of Erythromyeloid Progenitors And Their Progeny In The Mouse Embryo By Flow Cytometry

Macrophages are professional phagocytes from the innate arm of the immune system. In steady-state, sessile macrophages are found in adult tissues where they act as front line sentinels of infection and tissue damage. While other immune cells are continuously renewed from hematopoietic stem and progenitor cells (HSPC) located in the bone marrow, a lineage of macrophages, known as resident macrophages, have been shown to be self-maintained in tissues without input from bone marrow HSPCs. This lineage is exemplified by microglia in the brain, Kupffer cells in the liver and Langerhans cells in the epidermis among others. The intestinal and colon lamina propria are the only adult tissues devoid of HSPC-independent resident macrophages. Recent investigations have identified that resident macrophages originate from the extra-embryonic yolk sac hematopoiesis from progenitor(s) distinct from fetal hematopoietic stem cells (HSC). Among yolk sac definitive hematopoiesis, erythromyeloid progenitors (EMP) give rise both to erythroid and myeloid cells, in particular resident macrophages. EMP are only generated within the yolk sac between E8.5 and E10.5 days of development and they migrate to the fetal liver as early as circulation is connected, where they expand and differentiate until at least E16.5. Their progeny includes erythrocytes, macrophages, neutrophils and mast cells but only EMP-derived macrophages persist until adulthood in tissues. The transient nature of EMP emergence and the temporal overlap with HSC generation renders the analysis of these progenitors difficult. We have established a tamoxifen-inducible fate mapping protocol based on expression of the macrophage cytokine receptor Csf1r promoter to characterize EMP and EMP-derived cells in vivo by flow cytometry.

While infiltrating macrophages are continuously recruited to adult tissues from circulating precursors, resident macrophages seed their tissue during development, where they are maintained without further input from progenitors. The progenitors for resident macrophages were recently identified. Here, we present methods for the genetic fate mapping of the resident macrophage progenitors.

Macrophages are professional phagocytes from the innate arm of the immune system. In steady-state, sessile macrophages are found in adult tissues where ...

Developmental Biology

Two Flow Cytometric Approaches of NKG2D Ligand Surface Detection to Distinguish Stem Cells from Bulk Subpopulations in Acute Myeloid Leukemia

Within the same patient, absence of NKG2D ligands (NKG2DL) surface expression was shown to distinguish leukemic subpopulations with stem cell properties (so called leukemic stem cells, LSCs) from more differentiated counterpart leukemic cells that lack disease initiation potential although they carry similar leukemia specific genetic mutations. NKG2DL are biochemically highly diverse MHC class I-like self-molecules. Healthy cells in homeostatic conditions generally do not express NKG2DL on the cell surface. Instead, expression of these ligands is induced upon exposure to cellular stress (e.g., oncogenic transformation or infectious stimuli) to trigger elimination of damaged cells via lysis through NKG2D-receptor-expressing immune cells such as natural killer (NK) cells. Interestingly, NKG2DL surface expression is selectively suppressed in LSC subpopulations, allowing these cells to evade NKG2D-mediated immune surveillance. Here, we present a side-by-side analysis of two different flow cytometry methods that allow the investigation of NKG2DL surface expression on cancer cells i.e., a method involving pan-ligand recognition and a method involving staining with multiple antibodies against single ligands. These methods can be used to separate viable NKG2DL negative cellular subpopulations with putative cancer stem cell properties from NKG2DL positive non-LSC.

We present two different staining protocols for NKG2D ligand (NKG2DL) detection in human primary acute myeloid leukemia (AML) samples. The first approach is based on a fusion protein, able to recognize all known and potentially yet unknown ligands, while the second protocol relies on the addition of multiple anti-NKG2DL antibodies.

Within the same patient, absence of NKG2D ligands (NKG2DL) surface expression was shown to distinguish leukemic subpopulations with stem cell properties ...

High-Dimensionality Flow Cytometry for Immune Function Analysis of Dissected Implant Tissues

The success of implanting laboratory-grown tissue or a medical device in an individual is subject to the immune response of the recipient host. Considering an implant as a foreign body, a hostile and dysregulated immune response may result in the rejection of the implant, while a regulated response and regaining of homeostasis can lead to its acceptance. Analyzing the microenvironments of implants dissected out under in vivo or ex vivo&#160;settings can help in understanding the pattern of immune response, which can ultimately help in developing new generations of biomaterials. Flow cytometry is a well-known technique for characterizing immune cells and their subsets based on their cell surface markers. This review describes a protocol based on manual dicing, enzymatic digestion, and filtration through a cell strainer for the isolation of uniform cell suspensions from dissected implant tissue. Further, a multicolor flow cytometry staining protocol has been explained, along with steps for initial cytometer settings to characterize and quantify these isolated cells by flow cytometry.

Isolation of cells from dissected implants and their characterization by flow cytometry can significantly contribute to understanding the pattern of immune response against implants. This paper describes a precise method for the isolation of cells from dissected implants and their staining for flow cytometric analysis.

The success of implanting laboratory-grown tissue or a medical device in an individual is subject to the immune response of the recipient host. ...

Bioengineering

Flow Cytometric Measurement Of ROS Production In Macrophages In Response To Fc&#947;R Cross-linking

The oxidative or respiratory burst is used to describe the rapid consumption of oxygen and generation of reactive oxygen species (ROS) by phagocytes in response to various immune stimuli. ROS generated during immune activation exerts potent antimicrobial activity primarily through the ability of ROS to damage DNA and proteins, causing death of microorganisms. Being able to measure ROS production reproducibly and with ease is necessary in order to assess the contribution of various pathways and molecules to this mechanism of host defense. In this paper, we demonstrate the use of fluorescent probes and flow cytometry to detect ROS production. Although widely used, fluorescent measurement of ROS is notoriously problematic, especially with regards to measurement of ROS induced by specific and not mitogenic stimuli. We present a detailed methodology to detect ROS generated as a result of specific Fc&#947;R stimulation beginning with macrophage generation, priming, staining, Fc&#947;R cross-linking, and ending with flow cytometric analysis.

This study demonstrates the use of flow cytometry to detect reactive oxygen species (ROS) production resulting from activation of the Fc&#947;R. This method can be used to assess changes in the antimicrobial and redox signaling function of phagocytes in response to immune complexes, opsonized microorganisms, or direct Fc&#947;R cross-linking.

The oxidative or respiratory burst is used to describe the rapid consumption of oxygen and generation of reactive oxygen species (ROS) by phagocytes in ...

Immunology and Infection

Database-guided Flow-cytometry for Evaluation of Bone Marrow Myeloid Cell Maturation

A working group initiated within the French Cytometry Association (AFC) was developed in order to harmonize the application of multiparameter flow cytometry (MFC) for myeloid disease diagnosis in France. The protocol presented here was agreed-upon and applied between September 2013 and November 2015 in six French diagnostic laboratories (University Hospitals of Saint-Etienne, Grenoble, Clermont-Ferrand, Nice, and Lille and Institut Paoli-Calmettes in Marseille) and allowed the standardization of bone marrow sample preparation and data acquisition. Three maturation databases were developed for neutrophil, monocytic, and erythroid lineages with bone marrow from "healthy" donor individuals (individuals without any evidence of a hematopoietic disease). A robust method of analysis for each myeloid lineage should be applicable for routine diagnostic use. New cases can be analyzed in the same manner and compared against the usual databases. Thus, quantitative and qualitative phenotypic abnormalities can be identified and those above 2SD compared with data of normal bone marrow samples should be considered indicative of pathology. The major limitation is the higher variability between the data achieved using the monoclonal antibodies obtained with the methods based on hybridoma technologies and currently used in clinical diagnosis. Setting criteria for technical validation of the data acquired may help improve the utility of MFC for MDS diagnostics. The establishment of these criteria requires analysis against a database. The reduction of investigator subjectivity in data analysis is an important advantage of this method.

The MDS diagnosis is difficult in the absence of morphological criteria or non-informative cytogenetics. MFC could help refine the MDS diagnostic process. To become useful for clinical practice, the MFC analysis must be based on parameters with sufficient specificity and sensitivity, and data should be reproducible between different operators.

A working group initiated within the French Cytometry Association (AFC) was developed in order to harmonize the application of multiparameter flow ...

Medicine

Isolation and Purification of Murine Cardiac Pericytes

Pericytes, perivascular cells of microvessels and capillaries, are known to play a part in angiogenesis, vessel stabilization, and endothelial barrier integrity. However, their tissue-specific functions in the heart are not well understood. Moreover, there is currently no protocol utilizing readily accessible materials to isolate and purify pericytes of cardiac origin. Our protocol focuses on using the widely used mammalian model, the mouse, as our source of cells. Using the enzymatic digestion and mechanical dissociation of heart tissue, we obtained a crude cell mixture that was further purified by fluorescence activating cell sorting (FACS) by a plethora of markers. Because there is no single unequivocal marker for pericytes, we gated for cells that were CD31-CD34-CD45-CD140b+NG2+CD146+. Following purification, these primary cells were cultured and passaged multiple times without any changes in morphology and marker expression. With the ability to regularly obtain primary murine cardiac pericytes using our protocol, we hope to further understand the role of pericytes in cardiovascular physiology and their therapeutic potential.

We have optimized a protocol to isolate and purify murine cardiac pericytes for basic research and investigation of their biology and therapeutic potential.

Pericytes, perivascular cells of microvessels and capillaries, are known to play a part in angiogenesis, vessel stabilization, and endothelial barrier ...

Immunofluorescent Microscopy and Analysis for High-Parameter Tissue Imaging

Generating and Analyzing High-Parameter Histology Images with Histoflow Cytometry

The usage of histology to investigate immune cell diversity in tissue sections such as those derived from the central nervous system (CNS) is critically limited by the number of fluorescent parameters that can be imaged at a single time. Most immune cell subsets have been defined using flow cytometry by using complex combinations of protein markers, often requiring four or more parameters to conclusively identify, which is beyond the capabilities of most conventional microscopes. As flow cytometry dissociates tissues and loses spatial information, there is a need for techniques that can retain spatial information while interrogating the roles of complex cell types. These issues are addressed here by creating a method for expanding the number of fluorescent parameters that can be imaged by collecting the signals of spectrally overlapping fluorophores and using spectral unmixing to separate the signals of each individual fluorophore. These images are then processed using an analysis pipeline to take high-parameter histology images and extract single cells from these images so that the unique fluorescent properties of each cell can be analyzed at a single-cell level. Using flow cytometry-like gating strategies, cells can then be profiled into subsets and mapped back onto the histology sections to not only quantify their abundance, but also establish how they interact with the tissue environment. Overall, the simplicity and potential of using histoflow cytometry to study complex immune populations in histology sections is demonstrated.

Author Spotlight: Enhanced Multiplex Immunofluorescent Microscopy Protocol for Neuroscience Research

Described here is a method that can be used to image five or more fluorescent parameters by immunofluorescent microscopy. An analysis pipeline for extracting single cells from these images and conducting single-cell analysis through flow cytometry-like gating strategies is outlined, which can identify cell subsets in tissue sections.

The usage of histology to investigate immune cell diversity in tissue sections such as those derived from the central nervous system (CNS) is critically ...

Neuroscience

Human Adipose Tissue Micro-fragmentation for Cell Phenotyping and Secretome Characterization

In the past decade, adipose tissue transplants have been widely used in plastic surgery and orthopaedics to enhance tissue repletion and/or regeneration. Accordingly, techniques for harvesting and processing human adipose tissue have evolved in order to quickly and efficiently obtain large amounts of tissue. Among these, the closed system technology represents an innovative and easy-to-use system to harvest, process, and re-inject refined fat tissue in a short time and in the same intervention (intra-operatively). Adipose tissue is collected by liposuction, washed, emulsified, rinsed and minced mechanically into cell clusters of 0.3 to 0.8 mm. Autologous transplantation of mechanically fragmented adipose tissue has shown remarkable efficacy in different therapeutic indications such as aesthetic medicine and surgery, orthopedic and general surgery. Characterization of micro-fragmented adipose tissue revealed the presence of intact small vessels within the adipocyte clusters; hence, the perivascular niche is left unperturbed. These clusters are enriched in perivascular cells (i.e., mesenchymal stem cell (MSC) ancestors) and in vitro analysis showed an increased release of growth factors and cytokines involved in tissue repair and regeneration, compared to enzymatically derived MSCs. This suggests that the superior therapeutic potential of microfragmented adipose tissue is explained by a higher frequency of presumptive MSCs and enhanced secretory activity. Whether these added pericytes directly contribute to higher growth factor and cytokine production is not known. This clinically approved procedure allows the transplantation of presumptive MSCs without the need for expansion and/or enzymatic treatment, thus bypassing the requirements of GMP guidelines, and reducing the costs for cell-based therapies.

Here, we present human adipose tissue enzyme-free micro-fragmentation using a closed system device. This new method allows the obtainment of sub-millimeter clusters of adipose tissue suitable for in vivo transplantation, in vitro culture, and further cell isolation and characterization.

In the past decade, adipose tissue transplants have been widely used in plastic surgery and orthopaedics to enhance tissue repletion and/or regeneration. ...