Flow Cytometry Assays in Primary Immunodeficiency Diseases Protocol

Introduction of FC Assays in Primary Immunodeficiency Diseases

The collective development of accurate molecular technologies, an increased understanding of complex molecular signaling pathways and the availability of reagents to assess the latter have allowed scientists to elucidate the molecular etiology of an ever increasing number of primary immunodeficiency diseases (PIDs). At the first WHO sponsored meeting of experts involved in the treatment and investigation of PIDs in 1970, 16 PIDs were identified and classified. Prior to the latest meeting of PID experts in Dublin, Ireland in the summer of 2009, over 150 PIDs had been classified and characterized. This large group of disorders is most commonly classified into eight individual categories. More recently, a comprehensive systematic mathematical classification project based on clinical, pathological, and laboratory parameters identified 11 groups for over 200 clinically defined PIDs of which 167 had known genetic etiologies.

Of all of these disorders, the majority have some abnormality that could be detected by a flow cytometry-based application (although it must be acknowledged that not all abnormalities would be "specific" for a particular condition). It would be interesting to attempt to describe each of these applications; however, given that space is limited, we provide only a snapshot of the flow cytometry procedures currently utilized.

To combine an understanding of this inordinately complex group of disorders with an appreciation of how flow cytometry can be applied in their detection, the PIDs are described as one 3 general groups based on their underlying genetic abnormality. Very simply all of the disorders can be grouped according to: (a) mutations in genes that affect the relative representation of a specific subset, i.e., a subset abnormality; (b) mutations in genes that affect the expression of a specific "marker," i.e., marker abnormality, and lastly; (c) mutations in genes that affect a particular cell function, i.e., functional abnormality. Leukocyte subset abnormalities defined by the measurement of the relative and absolute number of specific subsets represent the most common application of clinical flow cytometry. Marker abnormalities are also commonly detected flow cytometrically with the appropriate combination of fluorochromes and monoclonal/ polyclonal antibodies. Lastly, there are several physiologic cell functions whose activity can be assessed with the appropriate reagents. In this chapter, we describe one flow cytometry application that has been utilized in a clinical setting to detect PIDs in one of the three categories (i.e., a subset abnormality, a marker abnormality, a functional abnormality). A more comprehensive treatise of the individual PID can be found in.

Primary Immunodeficiency Diseases

Each discovery of a new primary immunodeficiency can be thought of as an "experiment of nature," a term coined originally by Robert A. Good in the mid 1950s in reference to the fact that each case of a new primary immunodeficiency disease has taught us something about the normal functioning of the inordinately complex immune system. From a clinical perspective, it is common to think of patients with a primary immunodeficiency to suffer from an increased susceptibility to both the frequency and severity of infections with both opportunistic and pathogenic organisms. However, we now know that the scope of clinical symptoms encompassed within the category of primary immunodeficiency encompasses more than an increased susceptibility to infections. Autoimmunity, autoinflammatory disorders, atypical hemolytic uremic syndrome, paroxysmal nocturnal hematuria, for example, are now all classified as primary immunodeficiency states. Increased susceptibility to infection is, however, the most common clinical presentation in all of the primary immunodeficiency diseases. A large international effort spearheaded by the Jeffrey Modell Foundation is aimed at increasing the awareness of the public and medical professionals alike and has led to the posting of PID symptoms in most international airports. The ten warning signs (as summarized in these postings) that warrant an evaluation for the possible detection of a primary immunodeficiency disease are: (1) eight or more new ear infec tions within 1 year; (2) two or more serious sinus infections within 1 year; (3) two or more months on antibiotics with little effect; (4) two or more pneumonias within 1 year; (5) failure of an infant to gain weight or grow normally; (6) recurrent, deep skin or organ abscesses; (7) persistent thrush in mouth or elsewhere on skin, after age 1; (8) need for intravenous antibiotics to clear infections; (9) two or more deep-seated infections; and (10) a family history of primary immunodeficiency.

Initial Evaluation

A patient suspected of a primary immunodeficiency should first be evaluated with a thorough clinical and family history as well as a physical exam. The first pass laboratory tests would include a CBC with a differential followed by the testing of more specific immune parameters, including quantitative serum immunoglobulin levels and specific antibody determinations. The next stage would be to assess the complement system as well as the cellular components of the immune system. The most appropriate and encompassing screening assessment of the cellular immune system is accomplished by what we refer to as "routine immunophenotyping." Based on the results of the history, physical, and other laboratory tests, more specific flow cytometry procedures (i.e., specific subset, marker, or functional abnormalities) can be assessed. Lastly, once a presumptive diagnosis is obtained, we recommend that a patient sample be obtained to ascertain the molecular etiology and confirm the diagnosis.

Methods of FC Assays in Primary Immunodeficiency Diseases

Routine Immunophenotyping Panel for the Screening Diagnosis of Primary Immunodeficiency Disease

Sample Preparation

1. Label an appropriate number of TruCOUNT tubes for each patient and control. Note: Before use, verify that the TruCOUNT bead pellet is intact and within the metal retainer at the bottom of the tube. If not, discard the tube and replace it with another.
2. Add 20 mL of the 6 color MultiTest antibody reagent to the first tube and 10 mL each of the CD3-FITC/HLA-DR-PE Simultest and CD45 PerCP (or CD45-PerCP-Cy5.5 can be substituted) antibodies into the second tube.
3. Add 50 mL of well-mixed EDTA whole blood to each patient and each level of Streck whole blood control tube (we recommend running one set of controls per day) using the BD electronic reverse pipette (see Note 1).
4. Incubate the samples at room temperature in the dark for 15 min.
5. Add 450 mL of 1× FACS Lysing solution to each tube and vortex immediately
6. Incubate the samples at room temperature in the dark for 15 min.
7. Samples should be stored at 2–8°C and acquired on the FACSCanto within 1 hour of lysing using the FACSCanto software for Tube 1 and FACSDiva software for Tube 2 as described below (see Note 2).
8. Use FACSCanto software to check the bead lot numbers. Choose "Tools>Lot IDs." Choose "Absolute Count Beads" and enter or verify the lot number and bead information on the bead-tubes foil pouch is correct. Do not mix lot numbers.

FACSCanto Setup

The FACSCanto cytometer must be calibrated before each use and optimized assay settings must be implemented prior to sample collection. It is highly recommended that manufacturer's instructions be followed. In our laboratory, 7-Color Setup Beads are run before acquisition to automatically set the voltages and compensation for the parameters used in acquiring the 6 color monoclonal antibody combination. These settings are maintained and used for the acquisition of the second tube using the FACSDiva software. List mode data files acquired using both FACSCanto (first tube, 6 color combination) and FACSDiva software (for the second tube, 3 color) are then analyzed in each of the respective software programs. For the 6 color (first tube), the lymphocyte gate (CD45 vs. right angle light scatter) is reviewed and optimized (to include the entire cluster of lymphocytes) and the FACSCanto software automatically calculates the proportions and absolute numbers of the major lymphocyte subsets, i.e., T, B, NK, T helper, and T cytotoxic lymphocyte subset percentages and absolute counts. In summary, we recommend that the manufacturer's instructions for the FACSCanto Setup, acquisition, and analysis of the 6 color-panel first tube be followed. Our second tube, CD45, CD3, and HLA-DR, is a custom combination and is not currently amenable to automated acquisition and analysis of the FACSCanto. FACSDiva software is used for both the acquisition and analysis (i.e., measurement) of the CD3+ HLADR+ lymphocytes. In the final assessment of the routine immunophenotyping panel for each patient, the CD3 lymphocyte percentage and absolute count in each tube are compared to each other and valuable quality control parameters.

FACSCanto Acquisition and Analysis

1. Acquire the 6-color TBNK tubes from both patients and cellular controls using automated FACSCanto software. Verify the accuracy of the "expert gating." All leukocyte populations should be clearly defined, i.e., lymphocytes, monocytes, and granulocytes. The lymphocyte population is gated using low right angle light scatter (also known as side scatter) and bright CD45 fluorescence. Lymphocyte subsets are displayed as distinct populations (see Note 3). Adjust gates by holding down the left mouse button and dragging them to the appropriate location.
2. When all gates are adjusted, print the screen.
3. After printing, go to the next specimen by clicking the➤icon.
4. Repeat steps 1–3 for all specimens in the run. 5. Close FACSCanto software and login to FACSDiva software.

Acquire and Analyze the List Mode Data for the CD3/HLA-DR/45 Tube from Patients and the Cellular Controls Using FACSDiva Software

1. Link the instrument settings to the 7-Color Setup Beads run prior to acquisition using the "Lyse No Wash settings." Collect 2,500 lymphocyte events. Create an acquisition template to include three dot plots. On the first dot plot, display all events in a dot plot of right angle light scatter versus CD45-PerCP, with one analysis gate for the total lymphocyte subset and another for the beads (use this dot plot to calculate the total number of bead events acquired). On the second dot plot, display all events in a dot plot of right angle light scatter versus CD3 and draw an analysis gate around the CD3+ T cell cluster (use this dot plot for total number of T cell events). Draw a third dot plot which displays only those events which fall into the lymphocyte gate. The CD3 versus HLA-DR plot is used to calculate the percentage of lymphocytes that are both CD3+ and HLA-DR+. Save this setup as your experiment template for future use.
   

   Figure 1. Flow cytometry evaluation of the coexpression of CD3 and HLA-DR on lymphocytes.

2. During the analysis, adjust the lymphocyte gate on the dot plot of right angle light scatter (Y-axis) versus CD45-PerCP (X-axis). Ensure clean separation between debris (low CD45 and low right angle light scatter) and monocytes (similar CD45 intensity but higher right angle light scatter) by adjusting the lymphocyte gate as required. Set the second gate to encompass the entire bead population.
3. In the third graph, adjust the quadrants so that the activated T-cells (CD3+HLA-DR+) are in the upper right quadrant (see Note 4).
4. Record the following statistics from the analysis: total number of bead events (first dot plot), total number of CD3+ lymphocyte events (second dot plot), as well as percentage of CD3+ T lymphocytes (third dot plot: upper right plus lower right quadrants) and CD3+HLA-DR+ cells (third dot plot: upper right quadrant only) (see Note 5).
5. Calculate absolute CD3+ T cell counts (see Note 6) and record the percentage of CD3+ events and the percentage of CD3+HLA-DR+ events.

Quality Control

Cellular Controls The percentages and absolute counts of each of the subsets measured on both the low and the regular cellular control samples must fall within the limits established in-house (highly recommended) or printed on the package insert. If there are gross differences between the results obtained on the cellular controls and the defined ranges for each lymphocyte subsets, do not report patient results until/unless the cause of the problem is identified. Significant abnormalities observed with the cellular control material usually signify a problem. This must be addressed before the results of patient samples can be properly interpreted.

Internal Quality Control on Patient Samples

1. The CD3 percentages of Tube 1 (6 color TBNK) and Tube 2 (CD3/HLA-DR) must differ by<5%.
2. The absolute count difference between tubes 1 and 2 must be<15%.
3. The lymphosum must be 100±5.
4. The sum of the CD4 and CD8 subsets must be within 10% of the mean of the CD3 population. If the CD3 mean is 10% or greater, the gamma/delta panel must be run (unless the patient has been tested previously). Acquire and analyze in FACSDiva. CD3+gamma/delta+>15% is one explanation of why the T cell subsets do not add up to the CD3 total.
5. If any of the other criteria above are not within the specified limits, the following actions should be taken:
   
   • All values are reviewed for clerical or transcription errors.
   • FCS files are re-analyzed.
   • If these actions do not correct the criteria into acceptable ranges whether or not the cellular control results are in range, the assay is repeated.

Oxidative Burst Assay for the Screening Diagnosis of Chronic Granulomatous Disease

Sample Preparation

1. Prepare working dilution of DHR 123 (45 mg/mL). Add 30 mL of DHR 123 stock solution (5 mg/mL) to 3.33 mL of PBS and vortex. DHR 123 is extremely light sensitive. Keep stock and working dilutions of DHR 123 in the dark at all times.
2. Prepare working dilution of PMA (10 mg/mL). Add 10 mL of PMA stock (1 mg/mL) to 1 mL of Ca2+Mg2+ free PBS and vortex. PMA is extremely light sensitive. Keep stock and working dilutions of PMA in the dark at all times.
3. Prepare 1× NH₄Cl lysing solution by adding 2.0 mL of 10× NH4 Cl lysing solution and 18.0 mL of distilled, deionized water to a clean beaker or flask.
4. Label three tubes for each patient and control to be assayed. Tube #1: No dye/unstimulated; Tube #2: Plus dye/unstimulated; Tube #3: Plus dye/stimulated. Add 900 mL of PBS and 100 mL of well mixed whole blood to each tube.
5. Add 25 mL of the DHR 123 solution to tubes #2 & 3 (final concentration=1.125 mg/mL). Incubate for 15 min at 37°C in the shaking waterbath.
6. Add 10 mL of PMA to Tube #3 (final concentration = 100 ng/mL). Incubate all three tubes for 15 min at 37°C in a shaking waterbath. After this incubation period, centrifuge the tubes at 400 × g and remove supernatant with a transfer pipette.
7. Lyse the pellet by adding 2.5 mL of the NH4 Cl lysing solution and incubate for 15 min in the dark at room temperature. It is imperative to vortex both at the beginning and at the end of the lysing procedure. If after the first lyse there remains many RBC, repeat the lysing procedure.
8. Wash twice with 2 mL of washing solution.
9. Vortex, add 0.5 mL of 1% PFA and vortex again.
10. Specimen is ready to be acquired on the flow cytometer (see Note 7).

Acquisition and Analysis Using CellQuest Software

1. Standard instrument quality control must be followed prior to acquisition.
2. Open the CellQuest software and load appropriately established instrument "settings." Create an acquisition template of forward versus right angle light scatter and gate on the granulocyte cluster. While in the "Setup," adjust forward and right angle light scatter parameters of Tube #1 such that the lymphocyte, monocyte, and granulocyte clusters are clearly discernable.
3. Set an electronic analysis gate around the granulocyte cluster and adjust the fluorescence (FL1 detector fitted with a 530/30 bandpass filter) to be in the first decade on a 4-decade log scale (
4. Deselect "setup" option and acquire 10,000 events for each of the three tubes after entering the appropriate patient and tube identifying information.
5. Create an analysis document with forward versus right angle light scatter and set an analysis gate around the granulocyte cluster. Create a single parameter histogram to display the FL1 fluorescence of the events in the granulocyte analysis gate and measure the median fluorescence in each of the tubes.
6. Calculate the normal oxidative index (NOI) by dividing the mean fluorescence of Tube #3 by the mean fluorescence obtained in Tube #2 (see Note 9).

CD40-Ligand (CD154) Upregulation Induced In Vitro for the Screening Diagnosis of X-Linked Hyper IgM Syndrome (CD40-Ligand Deficiency)

Detection of CD154 Upregulation Using a Monoclonal Antibody Specific for CD154

1. Prepare working dilution of PMA (final concentration in step 5=15 ng/mL):
   
   • Add 3 mL of 1 mg/mL stock to 5 mL of culture medium.
   • Add 1 mL of (a) to 1 mL of culture medium, mix.
   • Add 1 mL of (b) to 1 mL of culture medium for final dilution.
2. Prepare working dilution of Calcium Ionophore (final concentration in step 5=400 ng/mL):
   
   • Add 18 mL of 1 mg/mL stock to 3 mL of culture medium, mix.
   • Add 1 mL of (a) to 0.5 mL of culture medium for final dilution.
3. For each sample tested, label two tubes: one for the "unstimulated" control and one for the "stimulated" test sample.
4. To the "unstimulated" tube add 800 mL of culture medium and 200 mL of well-mixed whole blood.
5. To the "stimulated" tube add 600 mL of culture medium, 200 mL of whole blood, 100 mL of PMA working dilution and 100 mL of Calcium Ionophore working dilution.
6. Vortex tubes gently, cap loosely or put parafilm loosely over the tops, and place in a dark CO₂ (5%) incubator at 37°C for 4 h.
7. After incubation, vortex gently. Add 2 mL of Ca²⁺Mg²⁺ free PBS and spin at an RFC of 700×g for 5 min.
8. Aspirate supernatant completely. Resuspend pellet in PBS such that final volume is 300 mL (three samples of 100 mL each are stained from each tube).
9. Samples are to be stained with the following monoclonal antibody panel (100 mL of sample and 10 mL of each antibody):
   
   • Tube #1: CD8-FITC/MsIgG1-PE/CD3-PerCP
   • Tube #2: CD8-FITC/CD40L-PE/CD3-PerCP
   • Tube #3: CD3-FITC/CD69-PE.
10. Label three tubes containing the mAb combinations above for the unstimulated cells and three for the stimulated cells.
11. Add 100 mL of cell suspension to each tube, vortex, and incubate at room temperature for 20 min.
12. At the end of the incubation period, add 2 mL of 1× FACS Lysing solution to each tube, vortex, and incubate for 10 min at room temperature. Vortex extensively for 5 min.
13. Spin tubes at an RFC of 700×g for 5 min. Decant and wash two times with flow cytometry wash solution (i.e., add 1 mL of wash, vortex, spin, decant, and repeat).
14. Add 0.5 mL of 1% PFA to each tube and vortex.
15. Acquire and analyze 10,000 events on the FACSCalibur flow cytometer using CellQuest software (see Note 10).

Detection of CD154 Upregulation Using a Chimeric CD40-Receptor-Human IgG Recombinant Protein

1. Prepare working dilution of PMA (final concentration in step 5=15 ng/mL):
   
   • Add 3 mL of 1 mg/mL stock to 5 mL of culture medium.
   • Add 1 mL of (a) to 1 mL of culture medium, mix.
   • Add 1 mL of (b) to 1 mL of culture medium for final dilution.
2. Prepare working dilution of Calcium Ionophore (final concentration in step 5=400 ng/mL):
   
   • Add 18 mL of 1 mg/mL stock to 3 mL of culture medium, mix.
   • Add 1 mL of (a) to 0.5 mL of culture medium for final dilution.
3. For each sample tested, label two tubes: one for the "unstimulated" control and one for the "stimulated" test sample.
4. To the "unstimulated" tube add 800 mL of culture medium and 200 mL of well-mixed whole blood.
5. To the "stimulated" tube add 600 mL of culture medium, 200 mL of whole blood, 100 mL of PMA working dilution and 100 mL of Calcium Ionophore working dilution.
6. Vortex tubes gently, cap loosely or put parafilm loosely over the tops, and place in CO₂ incubator at 37°C for 4 h.
7. After incubation, vortex gently. Add 2 mL of Ca²⁺Mg²⁺ free PBS and spin at an RCF of 700×g for 5 min.
8. Aspirate supernatant completely. Resuspend pellet in PBS such that final volume is 300 mL.
9. Samples are to be stained with the following monoclonal antibody panel:
   
   • Tube #1: CD8-FITC/Human IgG-PE/CD3-PerCP
   • Tube #2: CD8-FITC/Human CD40/Fc Chimera – antihuman IgG-PE/CD3-PerCP
   • Tube #3: CD3-FITC/CD69-PE
10. Label three tubes containing the mAb combinations above for the unstimulated cells and three for the stimulated cells. The staining process is a three step process with complete washing steps after the addition and incubation of each antibody or antibody combination.
11. Add 20 mL of the Recombinant Human CD40/Fc Chimera antibody to Tube #2 for both the stimulated and unstimulated cells. (Tubes #1 and #3 are not stained with any antibody in this step, but undergo the same incubation and washing steps.)
12. Add 100 mL of cell suspension to each tube, vortex, and incu bate at room temperature in the dark for 20 min.
13. Add 1 mL of flow cytometry wash solution. Vortex and centrifuge at an RFC of 700×g for 5 min.
14. Pippet and dispose of supernatant using a transfer pippet.
15. Repeat wash procedure (i.e., add 1 mL of wash solution, vortex, centrifuge, and pippet and dispose of supernatant).
16. Add 10 mL of antihuman IgG-PE to tubes #1 and #2. Incubate at room temperature in the dark for 20 min.
17. Add 1 mL of flow wash solution. Vortex and centrifuge at an RCF of 700×g for 5 min.
18. Pippet and dispose of supernatant using a transfer pippet.
19. Repeat wash procedure (i.e., add 1 mL of wash solution, vortex, centrifuge, and pippet and dispose of supernatant).
20. Add 10 mL of CD8-FITC and CD3-PerCP to tubes #1 and #2 for both the stimulated and unstimulated tubes. Add 10 mL of CD3-FITC and CD69-PE to Tube #3 for both the stimulated and unstimulated tubes.
21. At the end of the incubation period, add 2 mL of 1× FACS Lysing solution to each tube, vortex, and incubate for 10 min at room temperature. Vortex extensively for 5 min.
22. Spin tubes at an RCF of 700×g for 5 min. Decant and wash two times with flow cytometry wash solution (i.e., add 1 mL of wash solution, vortex, spin, decant, and repeat).
23. Add 0.5 mL of 1% PFA to each tube and vortex.
24. Acquire and analyze 2,500 CD3+ CD8− events on the FACSCalibur flow cytometer using CellQuest software.

Flow Cytometric Analysis

The lymphocyte activation protocol used in both procedures results in the downregulation of the CD4 molecule on T cells. The antigen of interest, CD40L is preferentially expressed on CD4+T cells. Since it is not possible to gate on the CD4+ T cells, a negative gating strategy is employed. In both procedures, cells in Tube #1 are labeled with CD8-FITC/IgG-PE and CD3-PerCP. A gate is drawn around the cells which express CD3 but do not express CD8, the majority of which are CD4 positive T cells. Therefore, an initial gate is drawn around the lymphocytes, then using Boolean logic draw a second gate around the population of cells which express CD3 but do not express the CD8 molecule. Alternatively, if it is not possible to draw a gate around the lymphocyte cluster based on light scatter parameters only, the first gate can be drawn around the CD3 positive cells on a histogram

The lymphocyte activation protocol used in both procedures results in the downregulation of the CD4 molecule on T cells. The antigen of interest, CD40L is preferentially expressed on CD4+T cells. Since it is not possible to gate on the CD4+ T cells, a negative gating strategy is employed. In both procedures, cells in Tube #1 are labeled with CD8-FITC/IgG-PE and CD3-PerCP. A gate is drawn around the cells which express CD3 but do not express CD8, the majority of which are CD4 positive T cells. Therefore, an initial gate is drawn around the lymphocytes, then using Boolean logic draw a second gate around the population of cells which express CD3 but do not express the CD8 molecule. Alternatively, if it is not possible to draw a gate around the lymphocyte cluster based on light scatter parameters only, the first gate can be drawn around the CD3 positive cells on a histogram of right angle light scatter versus CD3, and then follow the same steps as above.

1. Using Tube #1, set the positive negative/discriminator such that < 2% of the cells are positive, i.e., > 98% of the cells are negative.
2. The level of CD40L is then determined on the cells in Tube #2 as the percentage of CD3+CD8− cells that express either of the following:
   
   • CD40L fluorescence greater than the isotype control, in the procedure using a monoclonal antibody specific for CD154.
   • Human CD40/Fc Chimera – antihuman IgG PE fluorescence greater than the isotype control, in the procedure using a chimeric CD40-receptor-human IgG recombinant protein.
3. Tube #3 contains a panel of mAb which is used as the in vitro stimulation control. This control consists of measuring the level of CD69 expressed on resting cells versus the expression after 4 h using the same stimulation protocol. Lymphocytes are gated based on light scatter properties and then again based on the positive expression of CD3. The level of CD69 on both resting and activated CD3-positive lymphocytes is then determined.

Quality Control and Normal Reference Range

A normal healthy control is always run in parallel with the patient’s specimen. Failure of the control to fall within established ranges for Human CD40/Fc Chimera expression and CD69 expression necessitates a test repeat. The level of expression of CD69 must be >85% and the level of Human CD40/Fc Chimera expression must be >80% (see Note 11).

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