This research was supported in part by the Intramural Research Program of the NIH, including the National Institute of Biomedical Imaging and Bioengineering. Disclaimer: The NIH, its officers, and employees do no recommend or endorse any company, product, or service.

NOTE: Figure 1 gives an overview of the flow cytometry protocol.
1) Reagent preparation
<ol>
	<li>Prepare media for diluting enzymes and for tissue culture.
	<ol>
		<li>Add 5 mL of 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer solution into 500 mL of RPMI medium and shake well. Store the medium at 4 &#176;C until further use.</li>
	</ol>
	</li>
	<li>Calculate the volume of the enzyme solution. 
	&#8203;NOTE: The volume of the enzyme solution is the volume of the medium containing the enzymes (collagenase and DNase I) that will be needed to digest diced tissue in 6-well plates; it depends on the number of samples.
	<ol>
		<li>Use the following equation to calculate the required volume: 
		(Number of samples to be digested) &#215; 5 mL of serum-free RPMI with 10 mM HEPES buffer 
		&#8203;NOTE: For example, for 0.1 to 0.3 g of dissected spleen, use 5 mL of medium to prepare the enzyme solution. The enzymatic digestion protocol described in this manuscript is primarily for soft tissues (ranging from 0.1 to 1 g) dissected from mice which include the brain, kidney, skin, liver, lungs, spleen, muscles, as well as capsules around subcutaneous implants made of synthetic materials or extracellular matrix proteins. Digesting hard cartilaginous tissues might require different conditions and volumes of digestive enzymes, which can only be ascertained by optimization.</li>
		<li>Calculate the amount of collagenase and DNase I needed for the enzyme solution. 
		NOTE: In this protocol, 0.25 mg/mL of collagenase and 0.2 mg/mL of DNase I were used. The working concentrations required for collagenase and DNase can vary for different types of tissues19.</li>
	</ol>
	</li>
	<li>Prepare a 1:1000 viability dye solution by adding 1 &#181;L of the viability dye (see Table of Materials) to 999 &#181;L of phosphate-buffered saline (PBS) and vortex the solution. Store the solution in the dark at 4 &#176;C until further use.</li>
	<li>Prepare the staining buffer by adding 1 g of bovine serum albumin (BSA) to 100 mL of PBS, and vortex the solution until BSA is completely dissolved. Store the solution at 4 &#176;C until further use.</li>
</ol>
2) Setting up enzymatic digestion plates
<ol>
	<li>Set up a 6-well plate with 70 &#181;m cell strainers in each well. Add 3 mL of the prepared medium from step 1.1.1 into each well, and incubate on ice.</li>
	<li>Store the remaining media (2 mL/well) in an incubator or water bath at 37 &#176;C for suspending the calculated amount of collagenase and DNase I (step 1.2.2).</li>
</ol>
3) Isolation of cells
<ol>
	<li>Place the dissected implants/tissue into plates, and dice finely using scissors. Alternatively, utilize a mechanical disruption method using a tissue dissociator or hand-held homogenizer. Owing to the high number of leukocytes, dissect out the spleen to utilize as a staining control in each run. 
	NOTE: As some materials induce higher levels of fibrosis, this protocol is applicable for both highly fibrotic and minimally fibrotic materials. However, each processing method should be evaluated for both cell yield and viability after digestion prior to staining. Avoid peripheral blood contamination during dissection.</li>
	<li>Add collagenase and DNase I (volume calculated in step 1.2.2) to the remaining RPMI (2 mL) that has been warmed to 37 &#176;C. Add 2 mL of the complete enzyme solution to each well.</li>
	<li>Place the plates in an incubated shaker for 45 min at 37 &#176;C and 100 rpm. 
	NOTE: Be cautious when stacking plates as this can inhibit proper heat transfer.</li>
	<li>Meanwhile, prepare a suspension-dispensing tip by chopping 3-4 mm from the end of a 1000 &#181;L tip with scissors. After incubation, pipette the suspension in the wells up and down to mix it thoroughly using the chopped tip. Place the cell strainer on the top of a 50 mL conical tube and pass the digested solution through the cell strainer. 
	NOTE: The 70 &#181;m cell strainer is sufficient to separate cells from implanted biomaterial, such as alginate. If greater purity is required, use processes such as density gradient separation to obtain an enriched population of leukocytes.</li>
	<li>Rinse the wells with 1x PBS solution, and transfer the rinse volume through the strainer, followed by adding the washes of the strainer with 1x PBS solution until the volume in the tube reaches 50 mL. 
	NOTE: The 1x PBS must be at room temperature to prevent any increase in the viscosity of the digest and to allow cell pelleting during centrifugation.</li>
	<li>Centrifuge the tube at 300 &#215; g for 5 min at room temperature. Following centrifugation, aspirate the supernatant carefully using a serological pipette without disturbing the pellet. Re-suspend the pellet in 1000 &#181;L of 1x PBS, and transfer the suspension to a microcentrifuge tube.</li>
	<li>Mix 10 &#181;L of the cell suspension with 10 &#181;L of trypan blue, and load the suspension on counting chamber slides for counting cells using an automatic cell counter or on a hemocytometer to count via the conventional method under a microscope. Alternatively, use flow cytometry cell counting beads.</li>
</ol>
4) Staining for flow cytometry
<ol>
	<li>See Figure 2 for an example plate layout for a 14-color experiment with 45 samples, fluorescence minus one (FMO) controls, compensation controls, and a fully stained spleen sample control. After estimating the total number of cells, calculate the volume of the cell suspension required for staining 1 &#215; 106 cells for each sample, and 0.5 &#215; 106 cells for each compensation and FMO control. Dispense the required volume of the cell suspension into the wells of a V-bottom 96-well plate, and make up the volume to 100 &#181;L with 1x PBS. 
	NOTE: As an example, if 1000 &#181;L of cell suspension obtained at step 3.6 contain 40 &#215; 106 cells, then for 1 &#215; 106 cells, 1000/40 &#215; 106 = 25 &#181;L of cell suspension will be required.</li>
	<li>Centrifuge the plate at 300 &#215; g for 5 min at 4 &#176;C, aspirate the supernatant, and then resuspend the cells in the sample wells and in a compensation control well to determine viability, with 100 &#181;L of a 1:1000 solution of the viability dye (see Table of Materials).</li>
	<li>For cells in the other compensation wells as well as FMO and unstained wells, resuspend them with 100 &#181;L of 1x PBS.</li>
	<li>Incubate the cells in the dark for 30 min at 4 &#176;C.</li>
	<li>In the meantime, prepare a surface antibody cocktail for staining the sample and FMO controls: 50 &#181;L per sample or FMO. See Table 1 for an example of the antibody cocktail.</li>
	<li>After incubation, add 100 &#181;l of 1x PBS to each well, spin down at 300 &#215; g for 5 min at 4 &#176;C.</li>
	<li>Aspirate the supernatant and resuspend in 200 &#181;L of staining buffer (1x PBS + 1% BSA); centrifuge at 300 &#215; g for 5 min at 4 &#176;C.</li>
	<li>Aspirate the supernatant, and resuspend the cells in the compensation, FMO, and unstained wells with 50 &#181;L of 1x PBS. Add 50 &#181;L of the antibody cocktail to the respective sample wells and FMO wells. Add the respective antibodies to the different compensation wells.</li>
	<li>Incubate the plate for 45 min in the dark at 4 &#176;C. After incubation, add 150 &#181;L of staining buffer in each well, and centrifuge the cell suspension at 300 &#215; g for 5 min at 4 &#176;C.</li>
	<li>Aspirate the supernatant, resuspend the cells with 200 &#181;L of staining buffer, and centrifuge the cell suspension at 300 &#215; g for 5 min at 4 &#176;C.</li>
	<li>If analyzing samples without fixation, resuspend them in 200 &#181;L of staining buffer, and proceed to section 6 on cytometer setup and sample analysis. If fixing the cells, aspirate the supernatant after washing, and add 100 &#181;L of a fixative such as 4% paraformaldehyde. If staining for intracellular markers, proceed to section 5 on intracellular staining, and fix and permeabilize the cells (see Table of Materials). Incubate cells for 20 min in the dark at 4 &#176;C. 
	NOTE: As fixation can affect the fluorescence intensity of some fluorophores, evaluate each panel both with and without fixation.</li>
	<li>After incubation, add 100 &#181;L of 1x PBS, followed by centrifugation at 300 &#215; g for 5 min at 4 &#176;C. Aspirate the supernatant, resuspend in 1x PBS, and centrifuge at 300 &#215; g for 5 min at 4 &#176;C.</li>
	<li>Re-suspend the cells in 200 &#181;L of staining buffer, and store at 4 &#176;C prior to flow cytometric analysis.</li>
</ol>
5) Intracellular staining
<ol>
	<li>Continuing from step 4.11 for fixation and permeabilization for intracellular markers, add 100 &#181;L of the appropriate buffer. Centrifuge the cells at 350 &#215; g for 5 min at 4 &#176;C. Aspirate the supernatant, and re-suspend the pellets in 200 &#181;L of the fixative-permeabilizing agent solution; centrifuge the cells at 350 &#215; g for 5 min at 4 &#176;C. Aspirate the supernatant, and resuspend the pellets with intracellular antibodies diluted in the appropriate buffer. 
	NOTE: Antibody types and dilutions will depend on the target cells. For example, one common intracellular marker is forkhead box P3 for regulatory T cells, which is frequently used at a 1:100 dilution.</li>
	<li>Incubate the cell suspension in the dark for 45 min at 4 &#176;C. After incubation, add 150 &#181;L of the appropriate buffer, and centrifuge the suspension at 350 &#215; g for 5 min at 4 &#176;C.</li>
	<li>Aspirate the supernatant, and re-suspend the cells in 200 &#181;L of staining buffer followed by centrifugation at 300 &#215; g for 5 min at 4 &#176;C. Aspirate and re-suspend the cells in 200 &#181;L of staining buffer for flow cytometric analysis.</li>
</ol>
6) Cytometer and compensation setup
<ol>
	<li>Immediately before running the samples, prepare compensation beads (see Table of Materials) if utilizing bead-based compensation as opposed to cell-based compensation.
	<ol>
		<li>Label separate microcentrifuge tubes for each fluorochrome-conjugated antibody, and add 100 &#181;L of 1x PBS followed by one full drop of anti-mouse negative control compensation beads and one drop of positive control compensation beads to each tube. Add 1 &#181;L of the appropriate antibody to each separate tube. Vortex and incubate for 5 min before acquisition. 
		NOTE: As some dyes, such as BUV737, cannot be bound to beads, use a cell-based control.</li>
	</ol>
	</li>
	<li>Calibrate the flow cytometer before each experiment by running the cytometer setup with the tracking beads, and maintain the flow cytometer settings for each experiment.</li>
	<li>Before analyzing the sample, adjust the flow cytometer by running an unstained sample to adjust the cell population on a side scatter (SSC) versus forward scatter (FSC) plot so that the cell population falls in the center of the plot and is not off-scale.</li>
	<li>Run the stained sample briefly to adjust the voltages for FSC and SSC and for each channel to make sure that no events fall outside the logarithmic scale of each channel. Record 5,000 events for each compensation control, followed by gating of positive and negative populations. Calculate the compensation matrix and then run the samples, gathering at least 1,000 events for the populations of interest. 
	NOTE: Compensation on larger-color panels should be verified, as automated compensation can be prone to over- or under-compensation. Each parameter should be graphed against every other parameter to monitor for signs of over- or under-compensation, and each single-color control should be evaluated. Complex compensation of larger-color experiments should be carried out with the assistance of a collaborator or a core facility with extensive flow cytometry experience. Newer types of flow cytometers such as spectral cytometers utilize the full spectrum of a fluorophore through a process known as spectral unmixing, which can yield a cleaner distinction of fluorescent overlap as compared to standard compensation alone22.</li>
</ol>

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</ol>

The authors have nothing to disclose.

This review describes a detailed methodology for isolating cells from biomaterial implants to obtain a uniform cell suspension. In addition, a detailed protocol has been provided for staining the cell suspension for multicolor flow cytometry, along with the steps for configuring a flow cytometer for optimal results. Cell isolation methods can involve multiple steps, often utilizing manual tissue dissection followed by enzymatic digestion with proteolytic enzymes to dissociate the extracellular matrix in the tissue and di...

Advances in the field of medicine have led to the frequent use of implanted materials for supporting the function or re-growth of damaged tissue1,2. These include devices such as pacemakers, reconstructive cosmetic implants, and orthopedic plates used for bone fracture fixation3,4. However, the materials used to make these implants and the locations in which they are implanted play important roles in determining the success of these implants5,6,7. As foreign bodies, these implants can generate an immune response from the host that can either lead to rejection or tolerance8. This factor has driven biomaterial research to generate materials that can attract the desired immune response after implantation9,10,11,12.
The immune response is an essential requirement in the field of regenerative medicine, where a tissue or an organ is grown around a biomaterial skeleton (scaffold) in a laboratory for the replacement of a damaged tissue or organ13,14,15,16. In regenerative medicine, the goal is to replace missing or damaged tissue through the use of cells, signals, and scaffolds, each of which can be greatly modulated by immune responses17. Furthermore, even when a lack of immune response is desired, it is very rarely an absence of immune activity rather than the presence of a regulatory profile that is desired18. Techniques such as flow cytometry can play a significant role in characterizing the pattern of immune response to various biomaterials used for coating implant devices or for developing scaffolds for tissue engineering19.
This information, in turn, will ultimately help in developing biomaterials for implants that can be well-tolerated by the immune system or in developing scaffolds that can play a constructive role in tissue engineering. Proper preparation of samples for analysis by flow cytometry is an important step for avoiding inaccurate results in immune characterization via fluorescence activated cell sorting20,21. Therefore, this review presents a detailed methodology that can be utilized for the isolation of cells from scaffold tissue, staining the cell suspension, and analysis by flow cytometry.

<table><tbody><tr><td>50 mL conical tubes</td><td>Fisher Scientific</td><td>14-432-22</td><td></td></tr><tr><td>6 Well Plate</td><td>Fisher Scientific</td><td>07-000-646</td><td></td></tr><tr><td>BD Brilliant Stain Buffer Plus</td><td>BD Biosciences</td><td>566385</td><td></td></tr><tr><td>BD Cytofix</td><td>BD Biosciences</td><td>554655</td><td>For only fixing cells</td></tr><tr><td>Bovine serum albumin</td><td>Millipore Sigma</td><td>A7906</td><td>For preparing FACS staining buffer</td></tr><tr><td>CD11b AF700</td><td>Biolegend</td><td>101222</td><td>Clone: M1/70</td></tr><tr><td>CD11c PerCP/Cy5.5</td><td>Biolegend</td><td>117325</td><td>Clone: N418</td></tr><tr><td>CD197 PE/Dazzle594</td><td>Biolegend</td><td>120121</td><td>Clone: 4B12</td></tr><tr><td>CD200R3 APC</td><td>Biolegend</td><td>142207</td><td>Clone: Ba13</td></tr><tr><td>CD206 PE</td><td>Biolegend</td><td>141705</td><td>Clone: C068C2</td></tr><tr><td>CD45 BUV737</td><td>BD Biosciences</td><td>612778</td><td>Clone: 104/A20</td></tr><tr><td>CD86 BUV395</td><td>BD Biosciences</td><td>564199</td><td>Clone: GL1</td></tr><tr><td>CD8a BV421</td><td>Biolegend</td><td>100737</td><td>Clone: 53-6.7</td></tr><tr><td>Comp Bead anti-mouse</td><td>BD Biosciences</td><td>552843</td><td>For compensation control</td></tr><tr><td>DNase I</td><td>Millipore Sigma</td><td>11284932001</td><td>Bovine pancreatic deoxyribonuclease I (DNase I)</td></tr><tr><td>F4/80 PE/Cy7</td><td>Biolegend</td><td>123113</td><td>Clone: BM8</td></tr><tr><td>Fc Block</td><td>Biolegend</td><td>101301</td><td>Clone: 93</td></tr><tr><td>Fixation/Permeabilization Solution Kit</td><td>BD Biosciences</td><td>554714</td><td>For fixing and permeabilization of cells.</td></tr><tr><td>HEPES buffer</td><td>Thermo Fisher</td><td>15630080</td><td>Buffer to supplement cell media</td></tr><tr><td>Liberase</td><td>Millipore Sigma</td><td>5401127001</td><td>Blend of purified Collagenase I and Collagenase II</td></tr><tr><td>LIVE/DEAD Fixable Blue Dead Cell Stain Kit</td><td>Thermo Fisher</td><td>L23105</td><td>Viability dye</td></tr><tr><td>Ly6c AF488</td><td>Biolegend</td><td>128015</td><td>Clone: HK1.4</td></tr><tr><td>Ly6g BV510</td><td>Biolegend</td><td>127633</td><td>Clone: 1A8</td></tr><tr><td>MHCII BV786</td><td>BD Biosciences</td><td>742894</td><td>Clone: M5/114.15.2</td></tr><tr><td>Phosphate buffer saline</td><td>Thermo Fisher</td><td>D8537</td><td></td></tr><tr><td>RPMI</td><td>Thermo Fisher</td><td>11875176</td><td>Cell culture media</td></tr><tr><td>Siglec F BV605</td><td>BD Biosciences</td><td>740388</td><td>Clone: E50-2440</td></tr><tr><td>V-bottom 96-well plate</td><td></td><td></td><td></td></tr></tbody></table>

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.

The process of development of flow cytometry panels for immune analysis often relies on the comparison of results to existing data and the literature in the field. Knowledge of how populations may present in flow cytometry is critical for proper interpretation of data. Regardless, populations and cell types can appear differently in different tissues, so some variability is to be expected. In the context of well-defined control tissues, such staining optimization can be evaluated against ...

Watch this Scientific Journal Video about High-Dimensionality Flow Cytometry for Immune Function Analysis of Dissected Implant Tissues at JoVE.com

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

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. ...

high-dimensionality-flow-cytometry-for-immune-function-analysis

Research

JoVE Journal

Bioengineering

2.1K Views.  National Institutes of Health. 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.

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

Immunophenotyping of Orthotopic Homograft (Syngeneic) of Murine Primary KPC Pancreatic Ductal Adenocarcinoma by Flow Cytometry

Homograft (syngeneic) tumors are the workhorse of today&#39;s immuno-oncology (I/O) preclinical research. The tumor microenvironment (TME), particularly its immune-components, is vital to the prognosis and prediction of treatment outcomes, especially those of immunotherapy. TME immune-components are composed of different subsets of tumor-infiltrating immune cells assessable by multi-color FACS. Pancreatic ductal adenocarcinoma (PDAC) is among the deadliest malignances lacking good treatment options, thus an urgent and unmet medical need. One important reason for its non-responsiveness to various therapies (chemo-, targeted, I/O) has been its abundant TME, consisting of fibroblasts and leukocytes that protect tumor cells from these therapies. Orthotopically implanted PDAC is believed to more accurately recapture the TME of human pancreatic cancers than conventional subcutaneous (SC) models.
Homograft tumors (KPC) are transplants of mouse spontaneous PDAC originating from genetically engineered KPC-mice (KrasG12D/+/P53-/-/Pdx1-Cre) (KPC-GEMM). The primary tumor tissue is cut into small fragments (~2 mm3) and transplanted subcutaneously (SC) to the syngeneic recipients (C57BL/6, 7&#8211;9 weeks old). The homografts were then surgically orthotopically transplanted onto the pancreas of new C57BL/6 mice, along with SC-implantation, which reached tumor volumes of 300&#8211;1,000 mm3 by 17 days. Only tumors of 400&#8211;600 mm3 were harvested per approved autopsy procedure and cleaned to remove the adjacent non-tumor tissues. They were dissociated per protocol using a tissue dissociator into single-cell suspensions, followed by staining with designated panels of fluorescently-labeled antibodies for various markers of different immune cells (lymphoid, myeloid and NK, DCs). The stained samples were analyzed using multi-color FACS to determine numbers of immune cells of different lineages, as well as their relative percentage within tumors. The immune profiles of orthotopic tumors were then compared to those of SC tumors. The preliminary data demonstrated significantly elevated infiltrating TILs/TAMs in tumors over the pancreas, and higher B-cell infiltration into orthotopic rather than SC tumors.

Immunophenotyping of Orthotopic Homograft Syngeneic of Murine Primary KPC Pancreatic Ductal Adenocarcinoma by Flow Cytometry

The experimental procedure on the immunophenotyping of murine orthotopic PDAC homografts aims at profiling the tumor immuno-microenvironment. Tumors are orthotopically implanted via surgery. Tumors of 200&#8211;600 mm3 in size were harvested and dissociated to prepare single-cell suspensions, followed by multi-immune marker FACS analysis using different fluorescently-labeled antibodies.

Homograft (syngeneic) tumors are the workhorse of today&#39;s immuno-oncology (I/O) preclinical research. The tumor microenvironment (TME), particularly ...

Cancer Research

Functional Analysis of Tumor-Infiltrating Myeloid Cells by Flow Cytometry and Adoptive Transfer

The tumor-infiltrating myeloid cell compartment represents a heterogeneous population of broadly immunosuppressive cells that have been exploited by the tumor to support its growth. Their accumulation in tumor and secondary lymphoid tissue leads to the suppression of antitumor immune responses and is thus a target for therapeutic intervention. As it is known that the local cytokine milieu can dictate the functional programming of tumor-infiltrating myeloid cells, strategies have been devised to manipulate the tumor microenvironment (TME) to express a cytokine landscape more conducive to antitumor myeloid cell activity. To evaluate therapy-induced changes in tumor-infiltrating myeloid cells, this paper will outline the procedure to dissociate intradermal/subcutaneous tumor tissue from solid tumor-bearing mice in preparation for leukocyte recovery. Strategies for flow cytometric analysis will be provided to enable the identification of heterogeneous myeloid populations within isolated leukocytes and the characterization of unique myeloid phenotypes. Lastly, this paper will describe a means of purifying viable myeloid cells for functional assays and determining their therapeutic value in the context of adoptive transfer.

This protocol provides reliable methods of solid tumor dissociation and myeloid cell isolation in murine intradermal or subcutaneous tumor models. Flow cytometry allows for phenotypic characterization of heterogeneous myeloid populations within the tumor microenvironment and sorting will demonstrate their functionality in the context of adoptive transfer.

The tumor-infiltrating myeloid cell compartment represents a heterogeneous population of broadly immunosuppressive cells that have been exploited by the ...

Isolation and Flow Cytometric Analysis of Immune Cells from the Ischemic Mouse Brain

Ischemic stroke initiates a robust inflammatory response that starts in the intravascular compartment and involves rapid activation of brain resident cells. A key mechanism of this inflammatory response is the migration of circulating immune cells to the ischemic brain facilitated by chemokine release and increased endothelial adhesion molecule expression. Brain-invading leukocytes are well-known contributing to early-stage secondary ischemic injury, but their significance for the termination of inflammation and later brain repair has only recently been noticed.
Here, a simple protocol for the efficient isolation of immune cells from the ischemic mouse brain is provided. After transcardial perfusion, brain hemispheres are dissected and mechanically dissociated. Enzymatic digestion with Liberase&#160;is followed by density gradient (such as Percoll) centrifugation to remove myelin and cell debris. One major advantage of this protocol is the single-layer density gradient procedure which does not require time-consuming preparation of gradients and can be reliably performed. The approach yields highly reproducible cell counts per brain hemisphere and allows for measuring several flow cytometry panels in one biological replicate. Phenotypic characterization and quantification of brain-invading leukocytes after experimental stroke may contribute to a better understanding of their multifaceted roles in ischemic injury and repair.

Inflammation plays a central role in the pathogenesis of ischemic stroke. Increasing evidence suggests that it acts as a double-edged sword which exacerbates early brain injury, but also contributes to later repair. This protocol describes the isolation of immune cells from the ischemic brain and their subsequent flow cytometric phenotyping.

Ischemic stroke initiates a robust inflammatory response that starts in the intravascular compartment and involves rapid activation of brain resident ...

Immunology and Infection

Qualitative and Quantitative Analysis of the Immune Synapse in the Human System Using Imaging Flow Cytometry

The immune synapse is the area of communication between T cells and antigen-presenting cells (APCs). T cells polarize surface receptors and proteins towards the immune synapse to assure a stable binding and signal exchange. Classical confocal, TIRF, or super-resolution microscopy have been used to study the immune synapse. Since these methods require manual image acquisition and time-consuming quantification, the imaging of rare events is challenging. Here, we describe a workflow that enables the morphological analysis of tens of thousands of cells. Immune synapses are induced between primary human T cells in pan-leukocyte preparations and Staphylococcus aureus enterotoxin B (SEB)-loaded Raji cells as APCs. Image acquisition is performed with imaging flow cytometry, also called In-Flow microscopy, which combines features of a flow cytometer and a fluorescence microscope. A complete gating strategy for identifying T cell/APC couples and analyzing the immune synapses is provided. As this workflow allows the analysis of immune synapses in unpurified pan-leukocyte preparations and hence requires only a small volume of blood (i.e., 1 mL), it can be applied to samples from patients. Importantly, several samples can be prepared, measured, and analyzed in parallel.

Here, we describe a complete workflow for the qualitative and quantitative analysis of immune synapses between primary human T cells and antigen-presenting cells. The method is based on imaging flow cytometry, which allows the acquisition and evaluation of several thousand cell images within a relatively short period of time.

The immune synapse is the area of communication between T cells and antigen-presenting cells (APCs). T cells polarize surface receptors and proteins ...

Determination of Regulatory T Cell Subsets in Murine Thymus, Pancreatic Draining Lymph Node and Spleen Using Flow Cytometry

Our immune system consists of a number and variety of immune cells including regulatory T cells (Treg) cells. Treg cells can be divided into two subsets, thymic derived Treg (tTreg) cells and peripherally induced Treg (pTreg) cells. They are present in different organs of our body and can be distinguished by specific markers, such as Helios and Neuropilin 1. It has been reported that tTreg cells are functionally more suppressive than pTreg cells. Therefore, it is important to determine the proportion of both tTreg and pTreg cells when investigating heterogeneous cell populations. Herein, we collected thymic glands, pancreatic draining lymph nodes and spleens from normoglycemic non-obese diabetic mice to distinguish tTreg cells from pTreg cells using flow cytometry. We manually prepared single cell suspensions from these organs. Fluorochrome conjugated surface CD4, CD8, CD25, and Neuropilin 1 antibodies were used to stain the cells. They were kept in the fridge overnight. On the next day, the cells were stained with fluorochrome conjugated intracellular Foxp3 and Helios antibodies. These markers were used to characterize the two subsets of Treg cells. This protocol demonstrates a simple but practical way to prepare single cells from murine thymus, pancreatic draining lymph node and spleen and use them for subsequent flow cytometric analysis.

Herein, we present a protocol to prepare single cells from murine thymus, pancreatic draining lymph node and spleen to further study these cells using flow cytometry. In addition, this protocol was used for determining the subsets of regulatory T cells using flow cytometry.

Our immune system consists of a number and variety of immune cells including regulatory T cells (Treg) cells. Treg cells can be divided into two subsets, ...

Discrimination of Seven Immune Cell Subsets by Two-fluorochrome Flow Cytometry

Immune cell characterization heavily relies on multicolor flow cytometry to identify subpopulations based on differential expression of surface markers. Setup of a classic multicolor panel requires high-end instruments, custom labeled antibodies, and careful study design to minimize spectral overlap. We developed a multiparametric analysis to identify major human immune populations (CD4+ and CD8+ T cells, &#947;&#948; T cells, B cells, NK cells and monocytes) in peripheral blood by combining seven lineage markers using only two fluorochromes. Our strategy is based on the observation that lineage markers are constantly expressed in a unique combination by each cell population. Combining this information with a careful titration of the antibodies allows investigators to record five additional markers, expanding the optical limit of most flow cytometers. Head-to-head comparison demonstrated that the vast majority of immune cell populations in peripheral blood can be characterized with comparable accuracy between our method and the classic &#34;one fluorochrome-one marker approach&#34;, although the latter is still more precise for identifying populations such as NKT cells and &#947;&#948; T cells. Combining seven markers using two fluorochromes allows for the analysis of complex immune cell populations and clinical samples on affordable 6-10 fluorochrome flow cytometers, and even on 2-3 fluorochrome field instruments in areas with limited resources. High-end instruments can also benefit from this approach by using extra fluorochromes to accomplish deeper flow cytometry analysis. This approach is also very well suited for screening several cell populations in the case of clinical samples with limited number of cells.

Here, we present a flow cytometric protocol to identify CD4+ and CD8+ T cells, &#947;&#948; T cells, B cells, NK cells and monocytes in human peripheral blood by using only two fluorochromes instead of seven. With this approach, five additional markers can be recorded on most flow cytometers.

Immune cell characterization heavily relies on multicolor flow cytometry to identify subpopulations based on differential expression of surface markers. ...

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 ...

Medicine

Visualization, Quantification, and Mapping of Immune Cell Populations in the Tumor Microenvironment

The immune landscape of the tumor microenvironment (TME) is a determining factor in cancer progression and response to therapy. Specifically, the density and the location of immune cells in the TME have important diagnostic and prognostic values. Multiomic profiling of the TME has exponentially increased our understanding of the numerous cellular and molecular networks regulating tumor initiation and progression. However, these techniques do not provide information about the spatial organization of cells or cell-cell interactions. Affordable, accessible, and easy to execute multiplexing techniques that allow spatial resolution of immune cells in tissue sections are needed to complement single cell-based high-throughput technologies. Here, we describe a strategy that integrates serial imaging, sequential labeling, and image alignment to generate virtual multiparameter slides of whole tissue sections. Virtual slides are subsequently analyzed in an automated fashion using user-defined protocols that enable identification, quantification, and mapping of cell populations of interest. The image analysis is done, in this case using the analysis modules Tissuealign, Author, and HISTOmap. We present an example where we applied this strategy successfully to one clinical specimen, maximizing the information that can be obtained from limited tissue samples and providing an unbiased view of the TME in the entire tissue section.

Here, we describe a simple and accessible strategy for visualizing, quantifying, and mapping immune cells in formalin-fixed paraffin-embedded tumor tissue sections. This methodology combines existing imaging and digital analysis techniques with the purpose of expanding the multiplexing capability and multiparameter analysis of imaging assays.

The immune landscape of the tumor microenvironment (TME) is a determining factor in cancer progression and response to therapy. Specifically, the density ...

Flow Cytometric Analysis for Identification of the Innate and Adaptive Immune Cells of Murine Lung

The respiratory tract is in direct contact with the outside environment and requires a precisely regulated immune system to provide protection while suppressing unwanted reactions to environmental antigens. Lungs host several populations of innate and adaptive immune cells that provide immune surveillance but also mediate protective immune responses. These cells, which keep the healthy pulmonary immune system in balance, also participate in several pathological conditions such as asthma, infections, autoimmune diseases, and cancer. Selective expression of surface and intracellular proteins provides unique immunophenotypic properties to the immune cells of the lung. Consequently, flow cytometry has an instrumental role in the identification of such cell populations during steady-state and pathological conditions. This paper presents a protocol that describes a consistent and reproducible method to identify the immune cells that reside in the lungs of healthy mice under steady-state conditions. However, this protocol can also be used to identify changes in these cell populations in various disease models to help identify disease-specific changes in the lung immune landscape.

In this study, we present an effective and reproducible protocol to isolate the immune populations of the murine respiratory system. We also provide a method for the identification of all innate and adaptive immune cells that reside in the lungs of healthy mice, using a 9-color-based flow cytometry panel.

The respiratory tract is in direct contact with the outside environment and requires a precisely regulated immune system to provide protection while ...

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 ...