We want to thank David Rodriguez for his assistance with the figure preparation. Sputum samples were run on the BD LSR II at the UT Health San Antonio Flow Cytometry Shared Resource Facility, supported by UT Health, NIH-NCI P30 CA054174-20 (CTRC at UT Health) and UL1 TR001120 (CTSA grant).

All steps of the sputum processing are performed in a biological safety cabinet with appropriate personal protective equipment.
1. Reagent preparation before starting sputum dissociation
<ol>
	<li>Thaw 1% Paraformaldehyde (PFA), 25 mL per sample on ice, and keep cold until use. 
	CAUTION: PFA is toxic by inhalation and skin contact. Prepare the fixative according to the manufacturer&#39;s instructions and freeze at -20 &#176;C in 25 mL aliquots until use.</li>
	<li>Approximate the weight of the sample and thaw enough 0.1% Dithiothreitol (DTT) for step 2.2 and bring it to 37 &#176;C. (Aliquots of DTT should be stored at -20 &#176;C before use.)</li>
	<li>Bring enough 0.5% N-acetyl- L-cysteine (NAC) up to 37 &#176;C for step 2.2. (NAC should be made fresh weekly and stored at 4 &#176;C before use.)</li>
</ol>
2. Sputum dissociation
<ol>
	<li>Weigh the sputum sample to determine the volumes of dissociation reagents. 
	NOTE: A sample is considered to be small if the initial weight is &#8804;3 g, medium if &#62;3 g but &#8804;8 g, large if &#62;8 but &#8804;16 g, and extra-large if a sample weighs &#62;16 g. The indications small, medium, large, and extra-large will be used throughout the protocol. The quantity of reagents required for dissociation and labeling differ according to the size of the sputum sample.</li>
	<li>Transfer a small sample to a clean 50 mL conical tube, a medium sample to a clean 250 mL plastic disposable bottle, or a large and extra-large sample to a clean 500 mL plastic disposable bottle. Add 1 mL/g sample weight of 0.5% NAC and 4 mL/g sample weight of 0.1% DTT.</li>
	<li>Vortex at maximum speed (for 15 s), and then rock at room temperature (at maximum speed) for 15 min.</li>
	<li>Dilute the sample with four volumes of Hank&#39;s Balanced Salt Solution (HBSS) (based on the total volume of sample + reagents) to neutralize the NAC and DTT; vortex quickly at maximum speed and rock at room temperature for 5 min at maximum speed.</li>
	<li>Filter the cell suspension through 100 &#181;m nylon mesh cell strainer(s) into one or more 50 mL conical centrifuge tube(s) to create a single-cell suspension.</li>
	<li>Centrifuge the cells at 800 x g for 10 min at 4 &#176;C. Aspirate the supernatant, combine all pellets in one 15 mL conical tube, and then wash the pellets with HBSS using the same conditions.</li>
	<li>Resuspend the cell pellet in a volume of buffer determined by the initial weight of the sputum sample. 
	NOTE: A small sample is resuspended in 250 &#181;L of HBSS. A medium sample is resuspended in 760 &#181;L of HBSS. Large and extra-large samples are resuspended in 1460 &#181;L of HBSS.</li>
	<li>Take an aliquot of the cell suspension for a live/dead cell count using Trypan Blue. 
	NOTE: For a small sample, use 5 &#181;L. For a medium, large or extra-large sample, use 10 &#181;L. Dilute 1:10 with HBSS.
	<ol>
		<li>Mix 10 &#181;L of the sputum dilution with 30 &#181;L of 0.4% Trypan Blue to achieve a final sample dilution of 1:40. Load into the counting chambers of a hemocytometer for a cell count. 
		NOTE: It may be necessary to adjust the final dilution if the cell numbers are too low or too high to achieve an accurate count. Consulting Guiot et al.20 for correctly distinguishing sputum cells from SECs and debris is strongly recommended here. This is essential for an accurate cell count.</li>
	</ol>
	</li>
	<li>From the extra-large sample, remove 50 x 106 cells from the total and add to a new tube with enough added HBSS to create a total volume of 1700 &#181;L. 
	&#8203;NOTE: Consider this a large sample for the remainder of the protocol. Leftover samples can be discarded or used for other purposes.</li>
</ol>
3. Antibody and viability dye staining
<ol>
	<li>Choice of antibody and staining dye 
	NOTE: Table 1 shows the antibodies and viability dye that are used in this protocol and the cell populations they identify.
	<ol>
		<li>Label the tubes containing the sputum cells (see Table 2 for labels).</li>
		<li>Use 5 mL flow cytometry tubes (compatible with the flow cytometer used) for the sample tube with the unstained cells and the tube with the isotype control. Use 15 mL conical tubes for the blood and epithelial tube samples. 
		NOTE: These samples will be transferred to flow cytometry tubes following antibody staining and fixation.</li>
	</ol>
	</li>
	<li>Label the compensation tubes (Table 3). 
	NOTE: Use 5 mL flow cytometry tubes compatible with the flow cytometer being used.</li>
	<li>Add the amount of HBSS, antibody and/or dye to each of the sputum cell tubes and compensation tubes as indicated in Table 2 and Table 3, respectively. 
	NOTE: Add buffer (HBBS), antibodies, and dye to all the tubes before adding cells or compensation beads to ensure that all tubes&#39; staining time is consistent.</li>
	<li>Add the amounts of sputum cell volume listed in Table 2 to the assay tubes.</li>
	<li>Add compensation beads to the compensation tubes as listed in Table 3.</li>
	<li>Incubate all the tubes (assay and compensation tubes) on ice, protected from light, for 35 min. Then, fill the tubes with ice-cold HBSS and centrifuge at 4 &#176;C for 10 min at 800 x g.</li>
	<li>For the compensation tubes, aspirate the supernatant as close to the pellets as possible and flick the pellets to loosen.</li>
	<li>Add 0.5 mL of cold HBSS to the compensation tubes, store them on the ice at 4 &#176;C and protect them from light until needed for flow cytometry analysis.</li>
	<li>Aspirate the supernatant from the unstained isotype, blood, and epithelial tubes after centrifugation (step 3.6 from the antibody staining section) and loosen the pellets by flicking the tubes.</li>
</ol>
4. Fixation with 1% Paraformaldehyde (PFA)
<ol>
	<li>Add cold 1% PFA (which should be thawed by now) to the unstained, isotype, blood, and epithelial tubes; 2 mL to the unstained and isotype tubes, and 10 mL to the blood and epithelial tubes.</li>
	<li>Incubate the tubes on ice, protected from light for 1 h. Vortex quickly at maximum speed after 30 min.</li>
	<li>Fill the tubes with ice-cold HBSS. Then, centrifuge tubes at 4 &#176;C for 10 min at 1600 x g.</li>
	<li>Aspirate as much supernatant as possible without disturbing the cell pellet and flick the tube with the fingers to loosen the cells.</li>
	<li>Add 200 &#181;L of cold HBSS to the unstained and isotype tubes.</li>
	<li>Calculate the volume of HBSS for resuspension of the blood and epithelial tube according to the total cell count. 
	NOTE: Resuspension volume = 0.15 x [total cell count (step 8 of sputum dissociation) / 106]. Use 50 x 106 as the cell count for an extra-large sample.</li>
	<li>Store all sample and compensation tubes, protected from light on the ice at 4 &#176;C until flow cytometry analysis is performed. 
	&#8203;NOTE: This protocol has not been tested for storage for more than 24 h.</li>
</ol>
5. Data acquisition on the flow cytometer
<ol>
	<li>Apply appropriate startup procedures for the flow cytometer being used. 
	NOTE: This section of the protocol assumes that the person operating the flow cytometer is trained in the use of the instrument available to them, especially concerning daily procedures including checking the stability of the optics and fluidics systems, techniques for standardizing light scatter and fluorescence intensity, as well as calculating and applying the correct compensation matrix.</li>
	<li>Use the mixture of National Institute of Standards and Technology (NIST) beads to ensure that forward scatter and side scatter voltages are set to place the NIST beads to span the entire plot without placing the beads too close to the axes. 
	NOTE: This step is crucial to ensure that debris smaller than 5 &#181;m can be eliminated by gating post-acquisition analysis. Depending on the flow cytometer used, be mindful that the smallest beads are not being excluded with the threshold (when using the LSR II or the Lyric) or with the high discriminator (using the Navios EX flow cytometer). For the Navios EX, a gain of 2 was used for the forward and side scatter, a voltage of 236 for forward scatter, and 250 for side scatter. For the LSR II, a forward scatter voltage of 165 and a side scatter voltage of 190 was used.</li>
	<li>Set the flow rate to medium (LSR II) or high (Navios EX). 
	NOTE: The medium or high flow rate for most instruments can be used to acquire the sputum tubes. It is important to note that using too slow of a flow rate or too dilute of the sample may result in the cells settling, resulting in increased vortexing which is not desired. Therefore, adhering to the calculated resuspension volume in step 4.6 should result in cellular density that can be acquired quickly but not clog the machine.</li>
	<li>Adjust the voltages for each parameter used for the scatter and fluorescent parameters to place the cell populations on the scale. Use the figures with the gating strategy as guidance on how to adjust the voltages accordingly. 
	NOTE: Ensure that all the parameters needed are selected in the parameter selection window before the acquisition or that data will not be acquired.</li>
	<li>Acquire data for the unstained sputum sample first, followed by the isotype-stained sample, and then the blood tube and the epithelial tube. 
	NOTE: If the cell suspension is too concentrated to allow the flow cytometer to run the samples without clogging, the samples may be further diluted with HBSS.</li>
</ol>

All the authors are past or current employees of bioAffinity Technologies.

The cellular content of sputum includes a large variety of wide-ranging cells, often accompanied by a lot of debris37. In addition, sputum analysis requires a quality control that confirms the sample is collected from the lung instead of the oral cavity38. Therefore, it is not as simple to analyze sputum by flow cytometry as it is for blood, for example, which releases a much cleaner and homogeneous cell suspension. This protocol has addressed all these issues: providing in...

Technical advancements in the hardware and software of flow cytometers have made it possible to identify many distinct cell populations simultaneously1,2,3,4. The utilization of the flow cytometer in hematopoietic cell research, for example, has led to a much better understanding of the immune system2 and the cellular hierarchy of the hematopoietic system5, as well as the diagnostic distinction of a multitude of different blood cancers6,7,8. Although most sputum cells are of hematopoietic origin9,10,11, flow cytometry has not been widely applied to sputum analysis for diagnostic purposes. However, several studies suggest that the evaluation of immune cell populations in sputum (the most significant subset of cells) may be of great help in diagnosing and/or monitoring diseases such as asthma and chronic obstructive pulmonary disease (COPD)12,13,14,15. Moreover, the existence of epithelial-specific markers that can be used in flow cytometry allows the interrogation of the following most significant subset of cells in sputum, lung epithelial cells.
In addition to the ability to analyze many distinct cell populations of different tissue origins, a flow cytometer can evaluate large numbers of cells in a relatively short period. In comparison, slide-based, cytological types of analyses often require highly specialized personnel and/or equipment. These analyses can be labor-intensive, which leads to only a proportion of the sputum sample being analyzed16.
Three critical issues limit the widespread use of sputum in flow cytometry. The first issue relates to the collection of sputum. Sputum is collected through a huff cough that expels mucus from the lungs into the oral cavity, subsequently spitting into a collection cup. Since the mucus travels through the oral cavity, there is a high chance of SEC contamination. This contamination complicates the specimen analysis, but the problem is easily rectified on a flow cytometric platform, as shown in this study.
Not everyone can produce sputum spontaneously; therefore, several devices have been developed to assist with the sputum collection in a non-invasive manner17. The nebulizer is one such device and has been shown to produce reliable sputum samples18,19,20. Although the nebulizer is a very effective way of non-invasively collecting sputum, its use still requires a setting at a medical facility with specialized personnel21. In contrast, handheld devices such as the lung flute22,23,24 and the acapella16,25 can be used at home since they are very user-friendly. These assist devices are both safe and cost-effective.
For us, the acapella gave consistently better results than the lung flute16, and therefore, the acapella device has been chosen for sputum collections. A 3-day collection sample was decided because the primary purpose for using sputum is to develop a lung cancer detection test16. It has been shown that a 3-day sample increases the likelihood of lung cancer detection compared to a 1- or 2-day sample26,27,28. However, other methods of sputum collection may be preferable for different purposes. If a different sputum collection method is used than the one described here, it is recommended to carefully titrate each antibody or dye used for flow cytometric analysis; very little data is available on how different sputum collection methods affect the targeted proteins for flow cytometry.
The second issue dampening the enthusiasm for using sputum for diagnostics, primarily related to flow cytometry, is cell number. The problem is the collection of sufficient viable cells for a reliable analysis. Two studies demonstrated that sputum samples collected by non-invasive methods, with the help of an assist device, contain enough viable cells that can be utilized in clinical diagnosis or research studies16,24. However, neither of these studies addressed the issue of cell numbers concerning flow cytometry.
For the studies that form the basis for this protocol, sputum samples were collected from participants at high risk for developing lung cancer following approved institutional guidelines for each study site. High-risk participants were defined as between 55-75 years, having smoked 30 pack-years and having not quit smoking within the past 15 years. Patients were shown how to use the acapella device according to the manufacturer&#39;s instructions29 and collected sputum for three consecutive days at home. The sample was kept in the refrigerator until the last collection. On the final collection day, the sample was shipped to the laboratory overnight with a frozen cold pack. The samples were processed into a single cell suspension on the day they were received. With this method of sputum collection, more than enough viable cells are obtained for a reliable flow cytometric analysis.
Lastly, and related to the previous cell number issue, is the question of how to release the sputum cells from its mucinous environment. How can the cells be kept viable and create a single cell suspension that does not clog the flow cytometer? Based on initial work by Pizzichini et al.30 and Miller et al.31, this protocol describes an easy and reliable method for sputum processing into a single cell suspension that is suitable for flow cytometric analysis. This method has used well-established guidelines in flow cytometry32,33,34 to develop an efficient antibody labeling strategy to identify hematopoietic and epithelial cells in sputum and provide instrument settings, quality control measures, and analysis guidelines standardizing sputum analysis on a flow cytometric platform.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1% Paraformaldehyde Flow-Fix</td>
			<td>Polysciences</td>
			<td>25037</td>
			<td></td>
		</tr>
		<tr>
			<td>100 &#181;M nylon cell strainers, Falcon #352360</td>
			<td>Fisher Scientific</td>
			<td>08-771-19</td>
			<td></td>
		</tr>
		<tr>
			<td>3 M NaOH</td>
			<td>EMD</td>
			<td>SX0593-1</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL conical falcon tube</td>
			<td>Fisher Scientific</td>
			<td>14-432-22</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa488 anti-human CD19</td>
			<td>BioLegend</td>
			<td>302219</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa488 anti-human CD3</td>
			<td>BioLegend</td>
			<td>300415</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa488 anti-human cytokeratin</td>
			<td>BioLegend</td>
			<td>628608</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa488 PanCK, CD3, and CD19 Isotype</td>
			<td>BioLegend</td>
			<td>400129</td>
			<td></td>
		</tr>
		<tr>
			<td>BV510 anti-human CD45</td>
			<td>BioLegend</td>
			<td>304036</td>
			<td></td>
		</tr>
		<tr>
			<td>CD66b FITC isotype</td>
			<td>BD Biosciences</td>
			<td>555748</td>
			<td></td>
		</tr>
		<tr>
			<td>CompBead Plus Compensation Beads</td>
			<td>BD Biosciences</td>
			<td>560497</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning Polystyrene dispoable sterile bottle 250 mL</td>
			<td>Fisher Scientific</td>
			<td>09-761-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning Polystyrene dispoable sterile bottle 500 mL</td>
			<td>Fisher Scientific</td>
			<td>09-761-10</td>
			<td></td>
		</tr>
		<tr>
			<td>CS&#38;T beads</td>
			<td>BD Biosciences</td>
			<td>655051</td>
			<td></td>
		</tr>
		<tr>
			<td>DTT</td>
			<td>Fisher Scientific</td>
			<td>BP172-5</td>
			<td></td>
		</tr>
		<tr>
			<td>FITC anti-human CD66b</td>
			<td>GeneTex</td>
			<td>GTX75907</td>
			<td></td>
		</tr>
		<tr>
			<td>Fixable Viability Stain</td>
			<td>BD Biosciences</td>
			<td>564406</td>
			<td></td>
		</tr>
		<tr>
			<td>FlowCheck</td>
			<td>Beckman Coulter</td>
			<td>A69183</td>
			<td></td>
		</tr>
		<tr>
			<td>FlowSet</td>
			<td>Beckman Coulter</td>
			<td>A69184</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS</td>
			<td>Fisher Scientific</td>
			<td>14-175-095</td>
			<td></td>
		</tr>
		<tr>
			<td>NAC</td>
			<td>Sigma-Aldrich</td>
			<td>A9165</td>
			<td></td>
		</tr>
		<tr>
			<td>NIST Beads, 05 &#956;M</td>
			<td>Polysciences</td>
			<td>64080</td>
			<td></td>
		</tr>
		<tr>
			<td>NIST Beads, 20 &#956;M</td>
			<td>Polysciences</td>
			<td>64160</td>
			<td></td>
		</tr>
		<tr>
			<td>NIST Beads, 30 &#956;M</td>
			<td>Polysciences</td>
			<td>64170</td>
			<td></td>
		</tr>
		<tr>
			<td>PE anti-human CD45</td>
			<td>BioLegend</td>
			<td>304039</td>
			<td></td>
		</tr>
		<tr>
			<td>PE-CF594 anti-human EpCAM</td>
			<td>BD Biosciences</td>
			<td>565399</td>
			<td></td>
		</tr>
		<tr>
			<td>PE-CF594 CD206/EpCAM Isotype</td>
			<td>BD Biosciences</td>
			<td>562292</td>
			<td></td>
		</tr>
		<tr>
			<td>PE-CR594 anti-human CD206</td>
			<td>BD Biosciences</td>
			<td>564063</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium citrate dihydrate</td>
			<td>EMD</td>
			<td>SX0445-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypan Blue solution, 0.4%</td>
			<td>Fisher Scientific</td>
			<td>15250061</td>
			<td></td>
		</tr>
	</tbody>
</table>

quality-controlled sputum analysis by flow cytometry

Sputum, widely used to study the cellular content and other microenvironmental features to understand the health of the lung, is traditionally analyzed using cytology-based methodologies. Its utility is limited because reading the slides is time-consuming and requires highly specialized personnel. Moreover, extensive debris and the presence of too many squamous epithelial cells (SECs), or cheek cells, often renders a sample inadequate for diagnosis. In contrast, flow cytometry allows for high-throughput phenotyping of cellular populations while simultaneously excluding debris and SECs.
The protocol presented here describes an efficient method to dissociate sputum into a single cell suspension, antibody stain and fix cellular populations, and acquire samples on a flow cytometric platform. A gating strategy that describes the exclusion of debris, dead cells (including SECs) and cell doublets is presented here. Further, this work also explains how to analyze viable, single sputum cells based on a cluster of differentiation (CD)45 positive and negative populations to characterize hematopoietic and epithelial lineage subsets. A quality control measure is also provided by identifying lung-specific macrophages as evidence that a sample is derived from the lung and is not saliva. Finally, it has been demonstrated that this method can be applied to different cytometric platforms by providing sputum profiles from the same patient analyzed on three flow cytometers; Navios EX, LSR II, and Lyric. Furthermore, this protocol can be modified to include additional cellular markers of interest. A method to analyze an entire sputum sample on a flow cytometric platform is presented here that makes sputum amenable for developing high-throughput diagnostics of lung disease.

This protocol was developed with a clinical laboratory setting in mind. The focus during the development of the protocol was on simplicity, efficiency, and reproducibility. It was found that the most time-consuming step in the processing of sputum was counting the cells. Therefore, the protocol is set up in such a way that sputum processing and cell labeling can be performed independently from cell counting without loss of time. An accurate cell count, which is still necessary to dilute the sample appropriately for an un...

Watch this Scientific Journal Video about Quality-Controlled Sputum Analysis by Flow Cytometry at JoVE.com

Quality-Controlled Sputum Analysis by Flow Cytometry

This protocol describes an efficient method for dissociating sputum into a single cell suspension and the subsequent characterization of cellular subsets on standard flow cytometric platforms.

Sputum, widely used to study the cellular content and other microenvironmental features to understand the health of the lung, is traditionally analyzed ...

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4.9K Views.  bioAffinity Technologies. Questo protocollo risolve le sfide comuni riscontrate con l'elaborazione dell'espettorato che hanno reso difficile in passato l'uso con la citometria a flusso per scopi diagnostici clinici ad alto rendimento. Questo protocollo è progettato per risparmiare tempo utilizzando il peso dell'espettorato per determinare i volumi di colorazione di anticorpi e coloranti. Durante l'incubazione della colorazione, il conteggio delle cellule può essere eseguito per le fasi a valle.È importante o...

Video: Quality-Controlled Sputum Analysis by Flow Cytometry

High-throughput Flow Cytometry Cell-based Assay to Detect Antibodies to N-Methyl-D-aspartate Receptor or Dopamine-2 Receptor in Human Serum

Over the recent years, antibodies against surface and conformational proteins involved in neurotransmission have been detected in autoimmune CNS diseases in children and adults. These antibodies have been used to guide diagnosis and treatment. Cell-based assays have improved the detection of antibodies in patient serum. They are based on the surface expression of brain antigens on eukaryotic cells, which are then incubated with diluted patient sera followed by fluorochrome-conjugated secondary antibodies. After washing, secondary antibody binding is then analyzed by flow cytometry. Our group has developed a high-throughput flow cytometry live cell-based assay to reliably detect antibodies against specific neurotransmitter receptors. This flow cytometry method is straight forward, quantitative, efficient, and the use of a high-throughput sampler system allows for large patient cohorts to be easily assayed in a short space of time. Additionally, this cell-based assay can be easily adapted to detect antibodies to many different antigenic targets, both from the central nervous system and periphery. Discovering additional novel antibody biomarkers will enable prompt and accurate diagnosis and improve treatment of immune-mediated disorders.

Over the recent years, live cell-based assays have been used successfully to detect antibodies against surface and conformational antigens. Here, we describe a method using high-throughput flow cytometry enabling the analysis of large cohorts of patients. Detection of novel antibodies will improve diagnosis and treatment of immune-mediated disorders.

Over the recent years, antibodies against surface and conformational proteins involved in neurotransmission have been detected in autoimmune CNS diseases ...

Adaptation of Semiautomated Circulating Tumor Cell (CTC) Assays for Clinical and Preclinical Research Applications

The majority of cancer-related deaths occur subsequent to the development of metastatic disease. This highly lethal disease stage is associated with the presence of circulating tumor cells (CTCs). These rare cells have been demonstrated to be of clinical significance in metastatic breast, prostate, and colorectal cancers. The current gold standard in clinical CTC detection and enumeration is the FDA-cleared CellSearch system (CSS). This manuscript outlines the standard protocol utilized by this platform as well as two&#160;additional adapted protocols that describe the detailed process of user-defined marker optimization for protein characterization of patient CTCs and a comparable protocol for CTC capture in very low volumes of blood, using standard CSS reagents, for studying in vivo preclinical mouse models of metastasis. In addition, differences in CTC quality between healthy donor blood spiked with cells from tissue culture versus patient blood samples are highlighted. Finally, several commonly discrepant items that can lead to CTC misclassification errors are outlined. Taken together, these protocols will provide a useful resource for users of this platform interested in preclinical and clinical research pertaining to metastasis and CTCs.

Adaptation of Semiautomated Circulating Tumor Cell CTC Assays for Clinical and Preclinical Research Applications

Circulating tumor cells (CTCs) are prognostic in several metastatic cancers. This manuscript describes the gold standard CellSearch system (CSS) CTC enumeration platform and highlights common misclassification errors. In addition, two&#160;adapted protocols are described for user-defined marker characterization of CTCs and CTC enumeration in preclinical mouse models of metastasis using this technology.

The majority of cancer-related deaths occur subsequent to the development of metastatic disease. This highly lethal disease stage is associated with the ...

Collection, Isolation, and Flow Cytometric Analysis of Human Endocervical Samples

Despite the public health importance of mucosal pathogens (including HIV), relatively little is known about mucosal immunity, particularly at the female genital tract (FGT). Because heterosexual transmission now represents the dominant mechanism of HIV transmission, and given the continual spread of sexually transmitted infections (STIs), it is critical to understand the interplay between host and pathogen at the genital mucosa. The substantial gaps in knowledge around FGT immunity are partially due to the difficulty in successfully collecting and processing mucosal samples. In order to facilitate studies with sufficient sample size, collection techniques must be minimally invasive and efficient. To this end, a protocol for the collection of cervical cytobrush samples and subsequent isolation of cervical mononuclear cells (CMC) has been optimized. Using ex vivo flow cytometry-based immunophenotyping, it is possible to accurately and reliably quantify CMC lymphocyte/monocyte population frequencies and phenotypes. This technique can be coupled with the collection of cervical-vaginal lavage (CVL), which contains soluble immune mediators including cytokines, chemokines and anti-proteases, all of which can be used to determine the anti- or pro-inflammatory environment in the vagina.

The use of cytobrush sampling to collect lymphocytes and monocytes from the endocervix is a minimally invasive technique that provides samples for analysis of female genital tract immunity. In this protocol, we describe the collection of cytobrush samples and immune cell isolation for flow cytometry assays.

Despite the public health importance of mucosal pathogens (including HIV), relatively little is known about mucosal immunity, particularly at the female ...

Stereological and Flow Cytometry Characterization of Leukocyte Subpopulations in Models of Transient or Permanent Cerebral Ischemia

Microglia activation, as well as extravasation of haematogenous macrophages and neutrophils, is believed to play a pivotal role in brain injury after stroke. These myeloid cell subpopulations can display different phenotypes and functions and need to be distinguished and characterized to study their regulation and contribution to tissue damage. This protocol provides two different methodologies for brain immune cell characterization: a precise stereological approach and a flow cytometric analysis. The stereological approach is based on the optical fractionator method, which calculates the total number of cells in an area of interest (infarcted brain) estimated by a systematic random sampling. The second characterization approach provides a simple way to isolate brain leukocyte suspensions and to characterize them by flow cytometry, allowing for the characterization of microglia, infiltrated monocytes and neutrophils of the ischemic tissue. In addition, it also details a cerebral ischemia model in mice that exclusively affects brain cortex, generating highly reproducible infarcts with a low rate of mortality, and the procedure for histological brain processing to characterize infarct volume by the Cavalieri method.

Brain myeloid cells characterization following stroke can be performed by stereology using the optical fractionator method, or by a flow cytometric analysis of brain leukocytes suspensions. Both are useful techniques to perform an accurate phenotypical distinction of the main myeloid cell subsets found in the ischemic brain.

Microglia activation, as well as extravasation of haematogenous macrophages and neutrophils, is believed to play a pivotal role in brain injury after ...

Molecular Probe Optimization to Determine Cell Mortality in a Photosynthetic Organism (Microcystis aeruginosa) Using Flow Cytometry

Microbial subpopulations in field and laboratory studies have been shown to display high heterogeneity in morphological and physiological parameters. Determining the real time state of a microbial cell goes beyond live or dead categories, as microbes can exist in a dormant state, whereby cell division and metabolic activities are reduced. Given the need for detection and quantification of microbes, flow cytometry (FCM) with molecular probes provides a rapid and accurate method to help determine overall population viability. By using SYTOX Green and SYTOX Orange in the model cyanobacteria Microcystis aeruginosa to detect membrane integrity, we develop a transferable method for rapid indication of single cell mortality. The molecular probes used within this journal will be referred to as green or orange nucleic acid probes respectively (although there are other products with similar excitation and emission wavelengths that have a comparable modes of action, we specifically refer to the fore mentioned probes). Protocols using molecular probes vary between species, differing principally in concentration and incubation times. Following this protocol set out on M.aeruginosa the green nucleic acid probe was optimized at concentrations of 0.5 &#181;M after 30 min of incubation and the orange nucleic acid probe at 1 &#181;M after 10 min. In both probes concentrations less than the stated optimal led to an under reporting of cells with membrane damage. Conversely, 5 &#181;M concentrations and higher in both probes exhibited a type of non-specific staining, whereby &#39;live&#39; cells produced a target fluorescence, leading to an over representation of &#39;non-viable&#39; cell numbers. The positive controls (heat-killed) provided testable dead biomass, although the appropriateness of control generation remains subject to debate. By demonstrating a logical sequence of steps for optimizing the green and orange nucleic acid probes we demonstrate how to create a protocol that can be used to analyse cyanobacterial physiological state effectively.

Molecular Probe Optimization to Determine Cell Mortality in a Photosynthetic Organism Microcystis aeruginosa Using Flow Cytometry

Microbial populations contain substantial cell heterogeneity, which can dictate overall behavior. Molecular probe analysis through flow cytometry can determine physiological states of cells, however its application varies between species. This study provides a protocol to accurately determine cell mortality within a cyanobacterium population, without underestimating or recording false positive results.

Microbial subpopulations in field and laboratory studies have been shown to display high heterogeneity in morphological and physiological parameters. ...

Imaging- and Flow Cytometry-based Analysis of Cell Position and the Cell Cycle in 3D Melanoma Spheroids

Three-dimensional (3D) tumor spheroids are utilized in cancer research as a more accurate model of the in vivo tumor microenvironment, compared to traditional two-dimensional (2D) cell culture. The spheroid model is able to mimic the effects of cell-cell interaction, hypoxia and nutrient deprivation, and drug penetration. One characteristic of this model is the development of a necrotic core, surrounded by a ring of G1 arrested cells, with proliferating cells on the outer layers of the spheroid. Of interest in the cancer field is how different regions of the spheroid respond to drug therapies as well as genetic or environmental manipulation. We describe here the use of the fluorescence ubiquitination cell cycle indicator (FUCCI) system along with cytometry and image analysis using commercial software to characterize the cell cycle status of cells with respect to their position inside melanoma spheroids. These methods may be used to track changes in cell cycle status, gene/protein expression or cell viability in different sub-regions of tumor spheroids over time and under different conditions.

We describe two complementary methods using the fluorescence ubiquitination cell cycle indicator (FUCCI) and image analysis or flow cytometry to identify and isolate cells in the inner G1 arrested and outer proliferating regions of 3D spheroids.

Three-dimensional (3D) tumor spheroids are utilized in cancer research as a more accurate model of the in vivo tumor microenvironment, compared to ...

Methodology for Sputum Induction and Laboratory Processing

The technique of sputum induction and processing is a recognized non-invasive method allowing the collection and analysis of cells from the airways, which is interesting in various respiratory diseases like asthma, chronic obstructive pulmonary disease (COPD), chronic cough, or idiopathic pulmonary fibrosis. This technique is well tolerated, safe and non-invasive, but is currently limited to research services and specialized centers in clinical practice because it is technically demanding, time-consuming, and requires trained staff. The success rate of sputum induction and analysis is about 80%.
Here, we describe the induction and laboratory processing of sputum samples. Sputum is induced by inhalation of hypertonic or isotonic saline with salbutamol. For the processing, we use the whole sputum technique. Dithiothreitol (DTT) is used to allow mucolysis of sputum samples. The primary aim of sputum processing is to obtain a differential cell count to study the cell types present in the airway lumen. Additional analyses may also be performed on sputum supernatant and sputum cells, which may allow further investigation into inflammatory processes and immune mechanisms. Examples include studying mediators in sputum supernatant and performing a large spectrum of analysis on sputum cells such as flow cytometry, genomics, or proteomics.
Finally, representative results of sputum analysis in healthy controls, asthmatics, and COPD patients are presented.

Here we describe the technique of sputum induction. This protocol also explains the sputum processing to perform a differential cell count and to collect sputum supernatant and cells for further analysis.

The technique of sputum induction and processing is a recognized non-invasive method allowing the collection and analysis of cells from the airways, which ...

Characterization of Immune Cells in Human Adipose Tissue by Using Flow Cytometry

Infiltration of immune cells in the subcutaneous and visceral adipose tissue (AT) deposits leads to a low-grade inflammation contributing to the development of obesity-associated complications such as type 2 diabetes. To quantitatively and qualitatively investigate the immune cell subsets in human AT deposits, we have developed a flow cytometry approach. The stromal vascular fraction (SVF), containing the immune cells, is isolated from subcutaneous and visceral AT biopsies by collagenase digestion. Adipocytes are removed after centrifugation. The SVF cells are stained for multiple membrane-bound markers selected to differentiate between immune cell subsets and analyzed using flow cytometry. As a result of this approach, pro- and anti-inflammatory macrophage subsets, dendritic cells (DCs), B-cells, CD4+ and CD8+ T-cells, and NK cells can be detected and quantified. This method gives detailed information about immune cells in AT and the amount of each specific subset. Since there are numerous fluorescent antibodies available, our flow cytometry approach can be adjusted to measure various other cellular and intracellular markers of interest.

This article describes a method to analyze immune cell content of adipose tissue by isolation of immune cells from adipose tissue and subsequent analysis using flow cytometry.

Infiltration of immune cells in the subcutaneous and visceral adipose tissue (AT) deposits leads to a low-grade inflammation contributing to the ...

A Flow Cytometry-based Assay for Measuring Mitochondrial Membrane Potential in Cardiac Myocytes After Hypoxia/Reoxygenation

Timely and efficient reperfusion of the occluded coronary artery is the best strategy for decreasing myocardial infarct size in patients with a ST-segment elevated myocardial infarction. However, reperfusion per se can result in further cardiomyocyte death, a phenomenon known as reperfusion injury. The opening of the mitochondrial permeability transition pore (mPTP), with the decrease of the mitochondrial membrane potential (MMP), or mitochondrial depolarization, is universally recognized as the final step of reperfusion injury and is responsible for mitochondrial and cardiomyocyte death. JC-1 is a lipophilic cationic dye that accumulates in mitochondria depending on the value of MMP. The higher the MMP is, the more JC-1 accumulates in the mitochondria. The increasing amounts of JC-1 in mitochondria can be reflected by a fluorescence emission shift from green (~530 nm) to red (~590 nm). Therefore, the reduction of the red/green fluorescence intensity ratio can indicate the depolarization of mitochondria. Here, we take advantage of JC-1 to measure MMP, or the opening of mPTP in human cardiac myocytes after hypoxia/reoxygenation, detected by flow cytometry.

Here, we present a protocol that uses JC-1 dye to assess the mitochondrial membrane potential of cells after being exposed to hypoxia/reoxygenation with or without a protective agent.

Timely and efficient reperfusion of the occluded coronary artery is the best strategy for decreasing myocardial infarct size in patients with a ST-segment ...

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.