Flow Cytometry–Based Cytotoxicity and Antibody Binding Assay
UNIT 6.34
Mats Alheim1 1
Department of Laboratory Medicine, Division of Clinical Immunology and Transfusion Medicine, Karolinska University Hospital, Stockholm, Sweden
ABSTRACT Human leukocyte antigen (HLA) antibodies with the ability to activate complement are associated with an increased risk of early antibody-mediated graft rejection in kidney transplantation (KTx). Detection of these potentially harmful complement-fixing HLA antibodies is commonly performed via the complement-dependent cytotoxicity (CDC) assay according to protocols that were developed as early as 40 years ago. The readout for this assay is based on manual scoring by visual inspection of cells under a fluorescence microscope. CDC is often used in combination with the flow cytometry– based lymphocyte crossmatch assay (FCXM), which, with high sensitivity, detects HLA antibody binding. Here we describe a new approach wherein both cytotoxicity and antibody binding can be simultaneously assessed with flow cytometry. Two strategies are described, using either magnetic bead–enriched T and B lymphocytes or bulk peripheral blood mononuclear cells (PBMC) as donor target cells. Curr. Protoc. Cytom. 66:6.34.1C 2013 by John Wiley & Sons, Inc. 6.34.11. Keywords: kidney transplantation r HLA antibodies r crossmatching r flow cytometry r complement dependent cytotoxicity
INTRODUCTION The benchmark assay for determination of cytotoxic antibodies, termed complementdependent cytotoxicity (CDC), was developed in the mid-1960s and is still the most important pre-transplantation (pre-Tx) assay (Terasaki and McClelland, 1964). Apart from a few modifications introduced to overcome some of its weaknesses, such as insensitivity and nonspecific responses, it has basically remained unaltered since its introduction (Amos et al., 1969; Johnson et al., 1972). Many transplantation laboratories perform CDC in parallel with a flow cytometry crossmatch (FCXM) assay for assessment of antibody binding to HLA class I and/or class II expressing lymphocytes. The high sensitivity of FCXM enables detection of low-avidity HLA antibodies, previously unrecognized by CDC (Garovoy et al., 1983). The methods presented herein describe novel flow cytometry–based methodologies for simultaneous determination of complement-fixing and non-complement-fixing antibodies. Since the introduction of flow cytometry into transplantation laboratories, there has been a growing interest in developing an assay for detection of cytotoxic antibodies using flow cytometry (Talbot et al., 1987; Lillevang et al., 1992; Wetzsteon et al., 1992). In recent years, the flow cytometry–based approach has been further refined, which has enabled simultaneous detection of both antibody binding and cytotoxicity (Schonemann et al., 2004; Won et al., 2006; Saw et al., 2008). Taken together, all of these studies show that flow cytometry–based assessment of antibody and cytotoxicity in one single assay is sensitive and objective.
Phenotypic Analysis Current Protocols in Cytometry 6.34.1-6.34.11, October 2013 Published online October 2013 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/0471142956.cy0634s66 C 2013 John Wiley & Sons, Inc. Copyright
6.34.1 Supplement 66
BASIC PROTOCOL 1
DETERMINATION OF CYTOTOXICITY AND ANTIBODY BINDING USING MAGNETIC BEAD–PURIFIED T AND B CELLS AS DONOR CELLS In this protocol, simultaneous detection of both cytotoxicity and binding are performed using immunomagnetic bead–enriched T and B cells as target cells. Cells are incubated with serum and thereafter with rabbit complement. FITC-conjugated anti–human IgG secondary antibody detects antibody binding and the 7-amino actinomycin D (7-AAD) probe stain nonviable cells.
Materials Whole blood (drawn in ACD tubes) diluted 1:1 in PBS (e.g., 8 ml blood and 8 ml PBS) Lymphocyte separation medium (LSM; e.g., Lymphoprep Axis-Shield PoC AS) Phosphate-buffered saline (PBS; APPENDIX 2A) PBS (APPENDIX 2A) containing 0.1% bovine serum albumin (BSA, e.g., Sigma-Aldrich) EasySep Human T Cell Enrichment Kit (StemCell Technologies, cat. no. 19051) EasySep Human B Cell Enrichment Kit (StemCell Technologies, cat. no. 19054) HLA antibody–positive control serum (e.g., in-house serum or from commercial source) HLA antibody–negative control serum (e.g., obtained from healthy ABO negative males or commercial source) Patient (test) serum (centrifuge 10 min at 13,000 × g, 4◦ C, prior to use) Standard rabbit complement (e.g., Cedarlane). Fluorescein isothiocyanate (FITC)–conjugated goat anti-human IgG F(ab 2) antibody (e.g., Jackson ImmunoResearch, cat. no. 096-098) 7-aminoactinomycin D (7-AAD) solution (e.g., Beckman Coulter, cat. no. A07704) 15- and 50-ml conical polypropylene centrifuge tubes (e.g., BD Falcon) Refrigerated centrifuge equipped with plate rotor 12 × 75−mm flow cytometry sample tubes Magnet (e.g., EasySep magnet from StemCell Technologies) U-bottom 96-well plate Flow-cytometer tubes Flow cytometer equipped with at least four fluorescence detectors (e.g., Beckman Coulter, cat. no. FC500) Additional reagents and equipment for counting cells (APPENDIX 3A) Isolation of PBMC 1. Prepare PBMC by underlaying diluted blood (16 ml) with 8 ml of LSM in a 50-ml conical centrifuge tube. Alternatively, the diluted blood is gently overlaid onto LSM. For additional information see product instructions provided by LSM supplier.
2. Centrifuge for 20 min at 800 × g, 20◦ C, without brake. 3. Isolate the PBMC from the interphase with a Pasteur pipet. 4. Transfer cells to a 50-ml conical centrifuge tube and fill up with PBS. Centrifuge for 5 min at 420 × g, 4◦ C. 5. Discard supernatant and add 1 ml PBS/0.1% BSA solution. Flow Cytometry Based Cytotoxicity and Antibody Binding Assay
Isolation of T and B cells 6. Transfer 250 μl of cell suspension into one 12 × 75–mm flow cytometry tube labeled “T cells” and transfer 750 μl into another 12 × 75–mm tube labeled “B cells.”
6.34.2 Supplement 66
Current Protocols in Cytometry
7. Centrifuge cells 5 min at 420 × g, 4◦ C. Pour off the supernatant and resuspend cells in 100 μl PBS/0.1% BSA. 8. Add 10 μl of T cell Ab mix (from EasySep Human T Cell Enrichment Kit) to the tube labeled “T cells” and 10 μl of B cell Ab mix (from EasySep Human B Cell Enrichment Kit) to the tube labeled “B cells.” 9. Incubate at room temperature (20◦ to 23◦ C) for 10 min. 10. Add 10 μl of magnetic beads (from either of the EasySep kits) to both tubes. Mix and incubate at room temperature for 10 min. 11. Add 2.5 ml of PBS/0.1% BSA to each tube. Put tubes in magnet for 5 min. 12. Pour off unbound cells (enriched T and B cells) into a 15-ml conical tube. Centrifuge cells for 5 min at 420 × g, 4◦ C. 13. Count cells (APPENDIX 3A) and adjust cell concentration to 3–5 × 106 cells/ml by diluting with PBS/0.1% BSA.
Flow cytometric cytotoxicity and antibody-binding assay 14. Add 20 μl negative and positive control serum and test serum (e.g., patient serum) to designated wells of a 96-well plate. Use of 96-well plate enables processing of large number of samples. The assay can also be performed in flow cytometry sample tubes if preferred.
15. Add 10 μl of sorted T and B cells to separate wells with each set of sera (negative and positive controls and test serum). 16. Incubate 30 min at room temperature. 17. Add 150 μl PBS/0.1% BSA. Centrifuge 5 min 420 × g, 4◦ C, using a plate carrier. Repeat once. 18. Add 20 μl rabbit complement/well. Incubate 30 min at room temperature. The complement activity may differ between lots and suppliers, and it is therefore important to use an optimal predetermined dilution.
19. Repeat step 17 and then continue to step 20. 20. Add 20 μl goat anti-human IgG-FITC (diluted 1:100)/well. Incubate 20 min at 4◦ C in the dark. This step can be omitted if determination of antibody binding is not required.
21. Repeat step 17. 22. Add 100 μl 7-AAD (diluted 1:100)/well. Transfer cell suspension to flow-cytometer tubes. Additional information about assessment of cell viability using 7-AAD and other reagents such as propidium iodide (PI) can be found in UNIT 9.2. If a flow cytometer equipped with a plate loader is used, transfer of cells to tubes is obviously not necessary.
23. Add 250 μl PBS/0.1% BSA to each tube. 24. Acquire samples on flow cytometer.
Sample acquisition and data analysis 25. Perform appropriate quality control of flow cytometer performance. Use optimized instrument settings as determined in pilot-experiments. 26. Create a forward scatter (FSC) versus side scatter (SSC) dot plot (Fig. 6.34.1). Current Protocols in Cytometry
Phenotypic Analysis
6.34.3 Supplement 66
A 1023
1023
1023 MFI:131
SSC
1.4%
R1 0
0 0
1023
0 0
FSC
1023
0
7-AAD
1023 lgG FITC
B 1023
1023
1023 MFI:597
SSC
61%
R1 0
0 0
1023
0 0
FSC
1023
0
7-AAD
1023 lgG FITC
C 1023
1023
1023 MFI:393
SSC
1.8%
R1 0
0 0
1023
0 0
FSC
1023
0
7-AAD
1023 lgG FITC
D 1023
1023
1023 MFI:595
SSC
72%
R1 0
0 0
1023 FSC
0 0
1023 7-AAD
0
1023 lgG FITC
Figure 6.34.1 Sorted T cells (A, B) and B cells (C, D) were crossmatched against donor specific antibody negative (A and C) and positive serum (B and D). Live and dead lymphocytes were gated (R1) as shown in the left-hand panels. The percentage of dead cells (7-AAD+ ) and the mean fluorescence intensity (MFI) of IgG antibody binding for T cells and B cells are shown in the mid and right hand histograms, respectively.
6.34.4 Supplement 66
Current Protocols in Cytometry
27. Set threshold on the FSC/SSC to eliminate cell debris. 28. Set a gate on live and dead lymphocytes (R1). 29. Display R1-gated events in two one-dimensional histograms for 7-AAD and IgG FITC fluorescence, respectively. 30. Acquire and collect data from at least 1000 R1-gated cells. 31. Use the positive control sample to gate on 7-AAD positive cells. Determine the percentage of dead T and B cells for negative and positive control serum and for patient sample. 32. Calculate the percentage of dead cells for patient serum above negative control serum. 33. Determine the level of antibody binding on T and B cells by the mean fluorescence intensity (MFI) of the IgG FITC histogram peak. 34. Calculate the level of IgG binding for patient serum above the negative control serum.
DETERMINATION OF CYTOTOXICITY AND ANTIBODY BINDING USING PBMC AS DONOR CELLS
BASIC PROTOCOL 2
This protocol uses bulk PBMC as donor targets cells. Fluorochrome-conjugated anti–T lymphocyte (CD3) and anti–B lymphocyte (CD19) antibodies are added for the discrimination of HLA class I (T and B cells) and HLA class II (B cells)–expressing cells.
Materials Phycoerythrin (PE)-conjugated anti-human CD3 antibody (clone SK7, BD Bioscience) Phycoerythrin-Cyanin 7 (PC7) conjugated anti-human CD19 antibody (clone J3-119, Beckman Coulter) Fluorescein isothiocyanate (FITC)–conjugated goat anti-human IgG F(ab 2) antibody (e.g., Jackson ImmunoResearch, cat. no. 096-098) Additional reagents and equipment for determination of cytotoxicity and antibody binding using magnetic bead–purified T and B cells as donor cells (Basic Protocol 1) Isolation of PBMC 1. Prepare PBMC as described in Basic Protocol 1, steps 1 to 5. 2. Count cells (APPENDIX 3A) and adjust cell concentration to 15 × 106 cells/ml by dilution in PBS/0.1% BSA.
Flow cytometric cytotoxicity and antibody binding assay 3. Add 50 μl negative and positive control sera and test serum to designated wells of a 96-well plate. Add 10 μl cell suspension to each serum-containing well (150,000 cells/well). 4. Incubate 30 min at room temperature. 5. Add 150 μl PBS/0.1% BSA. Centrifuge 5 min 420 × g, 4◦ C. Repeat once. 6. Add 30 μl of a mixture of phycoerythrin (PE)-conjugated anti-human CD3 antibody and phycoerythrin-cyanin 7 (PC7) conjugated anti–human CD19 antibody to cells. Resuspend. Determine the optimal dilutions of fluorochrome-conjugated antibodies empirically. Recognize that the recommended dilution provided by manufacturer can differ considerably from optimal dilution for a specific assay. Current Protocols in Cytometry
Phenotypic Analysis
6.34.5 Supplement 66
7. Incubate 20 min at 4◦ C in the dark. 8. Repeat step 5 and then continue to step 9. 9. Add 20 μl rabbit complement/well. Incubate 30 min, room temperature. 10. Repeat step 5. 11. Add 20 μl FITC-conjugated goat anti–human IgG (diluted 1:100)/well. Incubate 20 min at 4◦ C in the dark. 12. Repeat step 5. 13. Add 100 μl 7-AAD (diluted 1:100)/well. Transfer to flow-cytometer tubes. 14. Add 250 μl PBS/0.1% BSA to each tube. 15. Acquire samples on flow cytometer.
Sample acquisition and data analysis 16. Perform appropriate quality control of flow cytometer performance. Use optimized instrument settings as determined in pilot experiments. Optimal PMT voltages and compensation settings are determined using single-colorstained samples and fluorescence minus one (FMO) controls as described previously (Maecker and Trotter, 2006).
17. Apply an FSC threshold to minimize the amount of cell debris. 18. Create two-dimensional dot-plots of FSC versus SSC and CD3 versus CD19 (Fig. 6.34.2). Create a one-dimensional histogram for 7-AAD and IgG FITC fluorescence. There are alternative ways to display and gate cells; e.g., CD3 and CD19 cells can be gated using SSC versus CD3 and CD19 dot-plots. It is informative, though, to include an FSC/SSC plot, because dead cells can be distinguished from live cells by their lower FSC signal in addition to bright 7-AAD fluorescence.
19. Gate on live and dead lymphocytes (R1) and apply gate in CD3 versus CD19 dot-plot. Gate on CD3-positive (R2) and CD19-positive cells (R3). 20. Display the 7-AAD and FITC (IgG) fluorescence for CD3 and CD19 gated cells in one-dimensional histograms. 21. Acquire and collect data from at least 1000 gated CD19+ cells. 22. After acquisition, use the positive control sample to gate on 7-AAD positive cells. Determine the percentage of dead T and B cells for negative and positive control serum and for patient sera. 23. Calculate the frequencies of dead T and B cells for patient serum above negative control serum. 24. Determine the level of antibody binding on T and B cells by the mean fluorescence intensity (MFI) of the IgG FITC histogram peak. 25. Calculate the level of IgG binding for patient serum above negative control serum.
COMMENTARY Background Information Flow Cytometry Based Cytotoxicity and Antibody Binding Assay
In the many transplantation laboratories worldwide, the complement-dependent cytotoxicity (CDC) assay is the “gold-standard”
method for determination of antibody-induced cytotoxicity. However, several disadvantages connected with CDC have been recognized such as high assay variability, subjectivity,
6.34.6 Supplement 66
Current Protocols in Cytometry
A 1023
1023
1023
SSC
MFI:234
R2
R2
2.1% R1 0
0 0
1023
0 0
1023
FSC
0
1023
7-AAD
1023
lgG FITC
1023
1023 MFI:246
R3
R3 CD19 PC7
R3 1.6% R2
0
0 0
1023
0 0
1023
0
1023
7-AAD
CD3 PE
lgG FITC
B 1023
1023
1023 MFI:376
R2
SSC
R2
63% R1
0
0 0
1023
0 0
1023
FSC
0
1023
7-AAD
1023
lgG FITC
1023
1023 MFI:464
R3
R3 CD19 PC7
R3 84% R2
0
0 0
1023 CD3 PE
0 0
1023 7-AAD
0
1023 lgG FITC
Figure 6.34.2 PBMC were crossmatched against donor-specific antibody-negative (A) and -positive serum (B). Live and dead lymphocytes were gated (R1) and displayed in a CD3 versus CD19 plot. The percentage of dead cells (7-AAD+ ) and the mean fluorescence intensity (MFI) of IgG antibody binding for T cells (R2) and B cells (R3) are shown in the plots. Note the distorted CD19 expression with positive serum (B, R3). Phenotypic Analysis
6.34.7 Current Protocols in Cytometry
Supplement 66
and high number of false positives. The first two issues are partly due to the assay design; percentage of dead cells is estimated manually by visual inspection under a fluorescence microscope. Because of these (and other) limitations, alternative techniques based on flow cytometry have been developed (Won et al., 2006; Saw et al., 2008). However, to our knowledge, this new procedure has not yet been implemented in any clinical transplantation laboratory. It is noteworthy that all of these studies used multi-color staining of PBMC targets. We present here an alternative version (Basic Protocol 1) where presorted T and B cells are used as targets. This flow-cytometry approach offers several advantages such as low assay variation and straightforward sample acquisition/analysis. Importantly, the novel assay design using 96-well format enables high-throughput analysis of large numbers of samples. In addition, the multi-well format provides the possibility of using pipetting robots, which eliminate sources of variability. The flow cytometry–based cytotoxicity assay makes it possible to automatically transfer data into the laboratory information system (LIS). This strategy minimizes operator errors, and time-consuming manual steps are avoided. It is important to acknowledge that not only assay performance such as sensitivity and specificity play a role in the choice of method, but also ease-of-use, automation, and the possibility for standardization.
Critical Parameters
Flow Cytometry Based Cytotoxicity and Antibody Binding Assay
Choice of protocol–Sorted versus unsorted cells Both protocols presented here exhibit advantages and disadvantages. One major advantage of using pre-sorted targets is that the number of fluorescence antibodies and markers is minimized. In addition, the gating procedures are greatly simplified; it is sufficient to use a FSC/SSC gate and histogram markers to perform data analysis. Furthermore, no compensation is required because there is negligible spectral overlap between the FITC and 7-AAD fluorescence. Note that if phycoerythrin (PE)– conjugated antibody is used together with 7-AAD, adequate compensation needs to be applied. One disadvantage of Basic Protocol 1 is that the pre-enrichment step prolongs the time-to-result by 25 to 30 min and doubles the total number of samples because T and B cells are acquired in separate tubes/wells. On
the other hand, if bulk PBMC are to be used, greater effort needs to be put into adequate gating and compensation, because two additional fluorescence markers (e.g., CD3 PE and CD19 PC7) are added. As discussed further below, there are issues with altered expression of CD19 after complement incubation. This should be taken into account in the choice of protocol. Choice of cell-separation technique If pre-sorted cells are to be used as targets, care needs to be taken to select an appropriate cell-separation strategy. There are multiple, to some extent similar options that can be considered. The initial question that needs to be addressed is whether whole blood or PBMC are to be used as starting material in the cell-isolation procedure. Take into consideration aspects such as easy-of-use, expenditure of time, recovery, and purity. Select a strategy that best fits your purposes and requirements. Here, T and B cells isolated by negative selection (no beads attached to targets) enable care-free sample acquisition without any distortion of FSC/SSC profile. It is advisable, as part of the assay quality control, to check the purity of sorted cells in each experiment. Choice of negative and positive controls The choice of negative and positive controls in flow cytometry is of great importance. The most appropriate choice here is serum to be used as controls. Negative control serum, without any detectable HLA antibodies (IgG or IgM), can be obtained from healthy blood donors (ABO negative males). The positive control serum can consist of pooled sera from highly immunized patients with HLA class I and/or class II antibodies. There are also commercial sources for both negative and positive control sera. An alternative, but not optimal, choice of positive control for cell lysis is to heat treat cells at 45◦ C for 10 min. The pan-HLA class I specific monoclonal antibody clone W6/32 will induce high cell lysis. Note that the W6/32 antibody is made in mouse, and for simultaneous assessment of antibody binding to target cells, a secondary antibody directed against mouse Ig needs to be used. Optimization of serum volume and cell number In the initial phase of assay setup, the choice of serum:cell volume ratio and number of cells per sample needs to be taken into consideration. In Basic Protocol 1, 20 μl of serum and
6.34.8 Supplement 66
Current Protocols in Cytometry
10 μl of cell volume (ratio 2:1) are used. The number of cells per well can affect the level of binding and lysis, and it is therefore advisable to find the optimal serum:cell ratio and cell number to obtain sufficient sensitivity. As an indication, 20,000 to 60,000 cells/well at a 2:1 serum:cell ratio give comparable levels of lysis and antibody binding. In addition, availability of serum controls and, in particular, test serum (e.g., patient serum) will also play a role in deciding on a serum:cell volume ratio. Also, the number of target cells available will obviously also affect the choice of number of cells/well. Optimization of incubation times and assay sensitivity The incubation times used in various steps of the protocol(s) can have a substantial effect on both cytotoxicity and binding. Therefore, it is of utmost importance to determine the optimal incubation time in order to obtain high sensitivity and low background and minimize time-to-results. In both Basic Protocols 1 and 2, cells are incubated with serum and complement for 30 min. Prolonged incubation times (e.g., 45 or 60 min) with serum or complement will give a higher degree of lysis and binding. This increased sensitivity is evident upon serum dilution. Choice of fluorescent antibodies and viability marker There are several options when it comes to selection of fluorescent antibodies and viability markers. Here, FITC-conjugated secondary IgG antibody combined with 7-AAD viability dye is used, and both are excited by a 488-nm laser. The choice of these two fluorochromes enables acquisition of samples without any need for compensation. Note that cells are brightly stained even with highly diluted 7-AAD dye and a short incubation time (5 to 10 min) prior to acquisition. The well known viability marker propidium iodine (PI) can also be used in combination with both FITC and PE, but proper compensation is necessary for both fluorochromes. If a flow cytometer equipped with a red laser (HeNe ∼630 to 640 nm) is preferred, there are alternative farred excited dyes (e.g., TO-PRO-3) that can be used. In Basic Protocol 2, two additional fluorochromes, i.e., CD3 PE and CD19 PC7, are included. In this case, compensation is clearly needed, and adequate consideration must be put into choosing the right combinations of fluorochromes for a given flow cytometry instrument.
Side-by-side comparison with alternative methods Evaluation of Basic Protocol 1 in this unit shows that it is highly reproducible and robust using either fresh or frozen cells. As part of any assay development, it is common practice to perform side-by-side comparisons with widely accepted standard technologies. Our side-byside comparison with conventional methods such as CDC and flow cytometry crossmatch show a high degree of concordance in results between the techniques. Data from other, more recently developed techniques such as single antigen bead (SAB) based anti–HLA, antiIgG/IgM or anti-C1q antibody detection assays (Chen et al., 2011) can be of guidance in the interpretation of crossmatch data, as well as in the determination of appropriate cut-off values for cytotoxicity and antibody binding.
Troubleshooting Low recovery and purity after T and B cell separation There are, as mentioned above, a plethora of techniques for cell separation; whole blood versus PBMC, positive versus negative selection, and column- versus non-column-based systems. Each technique has its advantages and disadvantages, and it is recommended to scrutinize each strategy in relation to your own requirements. The strategy described in Basic Protocol 1 is robust and very reproducible—a necessity for a clinical application. The starting material for the cell-separation strategy described here is PBMC and not whole blood. The benefit of using PBMC instead of whole blood is that in most cases higher purity can be obtained. This advantage outweighs the potential disadvantages, e.g., prolonged assay time. Initial validation may give some indication about the level of purity that is required in order to obtain reliable data. Loss of CD19 expression after complement incubation In the early stage of method development we attempted to confirm previously published reports using PBMC and staining with CD19 and CD3 after complement incubation. However, we repeatedly failed to detect sufficient numbers of B cells in serum samples with positive control serum. Normal numbers of B cells were observed with HLA antibody–negative serum. The CD3 expression level and number of T cells was normal at all times. Thus, for unknown reason(s), CD19 expression appeared to be lost (or masked) after complement incubation irrespective of the CD19 antibody
Phenotypic Analysis
6.34.9 Current Protocols in Cytometry
Supplement 66
clone. Addition of CD19 antibody or alternative B cell marker (e.g., CD20) before complement minimizes (but does not completely eliminate) this problem (see Fig. 6.34.2). Low cytotoxicity and antibody binding Needless to say, the selection of appropriate negative and positive controls is mandatory for correct interpretation of data and quality control of assay performance. Pooled sera from highly immunized patients used as a positive control may show negative cytotoxicity and/or antibody binding. The most plausible explanation (that initially should be checked) is that no donor-specific antibodies are present in the serum. Selection of sera that cover the great majority of all possible HLA alleles present in the donor pool will most likely prevent this from happening. If unexpectedly negative or weak cytotoxicity and/or binding are observed, there are several ways to investigate this further. Firstly, the titer of donor specific HLA antibody may merely be too low to be detected in the flow cytometry crossmatch assay. Prolonged serum and/or complement incubation time can be applied to increase sensitivity. Sera with donor-specific HLA antibodies directed against a single epitope are unable to provide crosslinking with complement factor C1q—a prerequisite for efficient induction of complement-induced cell lysis. This cytotoxicity-negative, adsorptionpositive (CYNAP) phenomenon can be prevented by adding anti–human Ig prior to addition of complement (Fuller et al., 1997). False positive and false negative crossmatch data False positive or negative results are difficult (or even impossible) to avoid entirely. For instance, complement-inhibitory proteins expressed on target cells or present in serum may block induction of lysis (Morgan, 2000). Unspecific antibody binding to Fc receptors on B cells can be avoided by a 30-min enzymatic treatment of PBMC (5 × 106 cells/ml) with 1 mg/ml pronase (Vaidya et al., 2001). Heat treatment of serum is another strategy that may eliminate false positive reactions (Ta and Scornik, 2002).
Anticipated Results
Flow Cytometry Based Cytotoxicity and Antibody Binding Assay
Evaluation of the assays described herein demonstrates a high degree of concordance with CDC and FCXM (Alheim et al., unpub. observ.). In both Basic Protocol 1 and Basic Protocol 2, isolation of PBMC from
whole blood is performed. The recovery of cells varies from donor to donor, but normally 1–2 × 106 PBMC are obtained per ml blood: ∼90% lymphocytes and ∼10% monocytes. The frequencies of CD3+ cells and CD19+ cells within the lymphocyte population are normally 30% to 40% and 5% to 10%, respectively. In Basic Protocol 1, T and B cells are separated by magnetic beads. The recovery is generally >80% and the purity >95% for both T and B cells. If assay is performed as described here, one ACD tube of whole blood (∼8 ml) will yield a sufficient number T and B cells (20,000 to 30,000 cells/well) for crossmatch of at least 10 to 15 different sera per cell type. Using well defined negative control serum lacking detectable IgG and IgM antibodies gives cytotoxicity background of ∼0% to 5% for T cells and 5% to 10% for B cells, using fresh cells. If frozen cells are used, up to 10% higher background lysis can occasionally be observed.
Time Considerations The 96-well format enables convenient processing of large number of samples. To perform Basic Protocol 1 as described here takes approximately 3.5 hr, cell separations and sample acquisition/analysis included. Shorter time (∼3 hr) is required to complete Basic Protocol 2. It should be noted though that the time needed for acquisition of a given number of B cells from PBMC (∼5% B cells) is longer than for pre-sorted B cells with close to 100% purity. Assay time can be reduced by using shorter wash steps (e.g., 1 min, twice) and reduced incubation time for the secondary IgG antibody step (e.g., 10 min).
Literature Cited Amos, D.B., Bashir, H., Boyle, W., MacQueen, M., and Tiilikainen, A. 1969. A simple micro cytotoxicity test. Transplantation 7:220-223. Chen, G., Sequeira, F., and Tyan, D.B. 2011. Novel C1q assay reveals a clinically relevant subset of human leukocyte antigen antibodies independent of immunoglobulin G strength on single antigen beads. Hum. Immunol. 72:849-858. Fuller, T.C., Fuller, A.A., Golden, M., and Rodey, G.E. 1997. HLA alloantibodies and the mechanism of the antiglobulin-augmented lymphocytotoxicity procedure. Hum. Immunol. 56:94105. Garovoy, R.M., Bigos, M., Perkins, H., and Colombe B. 1983. Flow cytometry analysis: A high technology crossmatch technique facilitating transplantation. Transplant Proc. XV:19391944.
6.34.10 Supplement 66
Current Protocols in Cytometry
Johnson, A.H., Rossen, R.D., and Butler, W.T. 1972. Detection of alloantibodies using a sensitive antiglobulin microcytotoxicity test: Identification of low levels of pre-formed antibodies in accelerated allograft rejection. Tissue Antigens 2:215-226. Lillevang, S.T., Steinbruchel, D.A., Kristensen, T., and Kemp, E. 1992. A new flowcytometric CDC assay for detection of cytotoxic antibodies applied to hamster-to-rat cardiac transplantation. Transplant Proc. 24:537-538. Maecker, H.T. and Trotter, J. 2006. Flow cytometry controls, instrument setup, and the determination of positivity. Cytometry A 69:10371042. Morgan, B.P. 2000. The complement system: An overview. Methods Mol. Biol. 150:1-13. Saw, C.L., Bray, R.A., and Gebel, H.M. 2008. Cytotoxicity and antibody binding by flow cytometry: A single assay to simultaneously assess two parameters. Cytometry B 74:287294. Schonemann, C., Lachmann, N., Kiesewetter, H., and Salama, A. 2004. Flow cytometric detection of complement-activating HLA antibodies. Cytometry B 62:39-45. Ta, M. and Scornik, J.C. 2002. Improved flow cytometric detection of donor-specific HLA class II
antibodies by heat inactivation. Transplantation 73:1611-1614. Talbot, D., Shenton, B.K., Givan, A.L., Proud, G., and Taylor, R.M. 1987. A rapid, objective method for the detection of lymphocytotoxic antibodies using flow cytometry. J. Immunol. Methods 99:137-140. Terasaki, P.I. and McClelland, J.D. 1964. Microdroplet assay of human serum cytotoxins. Nature 204:998-1000. Vaidya, S., Cooper, T.Y., Avandsalehi, J., Barnes, T., Brooks, K., Hymel, P., Noor, M., Sellers, R., Thomas, A., Stewart, D., Daller, J., Fish, J.C., Gugliuzza, K.K., and Bray, R.A. 2001. Improved flow cytometric detection of HLA alloantibodies using pronase: Potential implications in renal transplantation. Transplantation 71:422-428. Wetzsteon, P.J., Head, M.A., Fletcher, L.M., Lye, W.C., and Norman, D.J. 1992. Cytotoxic flowcytometric crossmatches (flow-tox): A comparison with conventional cytotoxicity crossmatch techniques. Hum. Immunol. 35:93-99. Won, D.I., Jeong, H.D., Kim, Y.L., and Suh, J.S. 2006. Simultaneous detection of antibody binding and cytotoxicity in flow cytometry crossmatch for renal transplantation. Cytometry B 70:82-90.
Phenotypic Analysis
6.34.11 Current Protocols in Cytometry
Supplement 66
UNIT 6.34
Mats Alheim1 1
Department of Laboratory Medicine, Division of Clinical Immunology and Transfusion Medicine, Karolinska University Hospital, Stockholm, Sweden
ABSTRACT Human leukocyte antigen (HLA) antibodies with the ability to activate complement are associated with an increased risk of early antibody-mediated graft rejection in kidney transplantation (KTx). Detection of these potentially harmful complement-fixing HLA antibodies is commonly performed via the complement-dependent cytotoxicity (CDC) assay according to protocols that were developed as early as 40 years ago. The readout for this assay is based on manual scoring by visual inspection of cells under a fluorescence microscope. CDC is often used in combination with the flow cytometry– based lymphocyte crossmatch assay (FCXM), which, with high sensitivity, detects HLA antibody binding. Here we describe a new approach wherein both cytotoxicity and antibody binding can be simultaneously assessed with flow cytometry. Two strategies are described, using either magnetic bead–enriched T and B lymphocytes or bulk peripheral blood mononuclear cells (PBMC) as donor target cells. Curr. Protoc. Cytom. 66:6.34.1C 2013 by John Wiley & Sons, Inc. 6.34.11. Keywords: kidney transplantation r HLA antibodies r crossmatching r flow cytometry r complement dependent cytotoxicity
INTRODUCTION The benchmark assay for determination of cytotoxic antibodies, termed complementdependent cytotoxicity (CDC), was developed in the mid-1960s and is still the most important pre-transplantation (pre-Tx) assay (Terasaki and McClelland, 1964). Apart from a few modifications introduced to overcome some of its weaknesses, such as insensitivity and nonspecific responses, it has basically remained unaltered since its introduction (Amos et al., 1969; Johnson et al., 1972). Many transplantation laboratories perform CDC in parallel with a flow cytometry crossmatch (FCXM) assay for assessment of antibody binding to HLA class I and/or class II expressing lymphocytes. The high sensitivity of FCXM enables detection of low-avidity HLA antibodies, previously unrecognized by CDC (Garovoy et al., 1983). The methods presented herein describe novel flow cytometry–based methodologies for simultaneous determination of complement-fixing and non-complement-fixing antibodies. Since the introduction of flow cytometry into transplantation laboratories, there has been a growing interest in developing an assay for detection of cytotoxic antibodies using flow cytometry (Talbot et al., 1987; Lillevang et al., 1992; Wetzsteon et al., 1992). In recent years, the flow cytometry–based approach has been further refined, which has enabled simultaneous detection of both antibody binding and cytotoxicity (Schonemann et al., 2004; Won et al., 2006; Saw et al., 2008). Taken together, all of these studies show that flow cytometry–based assessment of antibody and cytotoxicity in one single assay is sensitive and objective.
Phenotypic Analysis Current Protocols in Cytometry 6.34.1-6.34.11, October 2013 Published online October 2013 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/0471142956.cy0634s66 C 2013 John Wiley & Sons, Inc. Copyright
6.34.1 Supplement 66
BASIC PROTOCOL 1
DETERMINATION OF CYTOTOXICITY AND ANTIBODY BINDING USING MAGNETIC BEAD–PURIFIED T AND B CELLS AS DONOR CELLS In this protocol, simultaneous detection of both cytotoxicity and binding are performed using immunomagnetic bead–enriched T and B cells as target cells. Cells are incubated with serum and thereafter with rabbit complement. FITC-conjugated anti–human IgG secondary antibody detects antibody binding and the 7-amino actinomycin D (7-AAD) probe stain nonviable cells.
Materials Whole blood (drawn in ACD tubes) diluted 1:1 in PBS (e.g., 8 ml blood and 8 ml PBS) Lymphocyte separation medium (LSM; e.g., Lymphoprep Axis-Shield PoC AS) Phosphate-buffered saline (PBS; APPENDIX 2A) PBS (APPENDIX 2A) containing 0.1% bovine serum albumin (BSA, e.g., Sigma-Aldrich) EasySep Human T Cell Enrichment Kit (StemCell Technologies, cat. no. 19051) EasySep Human B Cell Enrichment Kit (StemCell Technologies, cat. no. 19054) HLA antibody–positive control serum (e.g., in-house serum or from commercial source) HLA antibody–negative control serum (e.g., obtained from healthy ABO negative males or commercial source) Patient (test) serum (centrifuge 10 min at 13,000 × g, 4◦ C, prior to use) Standard rabbit complement (e.g., Cedarlane). Fluorescein isothiocyanate (FITC)–conjugated goat anti-human IgG F(ab 2) antibody (e.g., Jackson ImmunoResearch, cat. no. 096-098) 7-aminoactinomycin D (7-AAD) solution (e.g., Beckman Coulter, cat. no. A07704) 15- and 50-ml conical polypropylene centrifuge tubes (e.g., BD Falcon) Refrigerated centrifuge equipped with plate rotor 12 × 75−mm flow cytometry sample tubes Magnet (e.g., EasySep magnet from StemCell Technologies) U-bottom 96-well plate Flow-cytometer tubes Flow cytometer equipped with at least four fluorescence detectors (e.g., Beckman Coulter, cat. no. FC500) Additional reagents and equipment for counting cells (APPENDIX 3A) Isolation of PBMC 1. Prepare PBMC by underlaying diluted blood (16 ml) with 8 ml of LSM in a 50-ml conical centrifuge tube. Alternatively, the diluted blood is gently overlaid onto LSM. For additional information see product instructions provided by LSM supplier.
2. Centrifuge for 20 min at 800 × g, 20◦ C, without brake. 3. Isolate the PBMC from the interphase with a Pasteur pipet. 4. Transfer cells to a 50-ml conical centrifuge tube and fill up with PBS. Centrifuge for 5 min at 420 × g, 4◦ C. 5. Discard supernatant and add 1 ml PBS/0.1% BSA solution. Flow Cytometry Based Cytotoxicity and Antibody Binding Assay
Isolation of T and B cells 6. Transfer 250 μl of cell suspension into one 12 × 75–mm flow cytometry tube labeled “T cells” and transfer 750 μl into another 12 × 75–mm tube labeled “B cells.”
6.34.2 Supplement 66
Current Protocols in Cytometry
7. Centrifuge cells 5 min at 420 × g, 4◦ C. Pour off the supernatant and resuspend cells in 100 μl PBS/0.1% BSA. 8. Add 10 μl of T cell Ab mix (from EasySep Human T Cell Enrichment Kit) to the tube labeled “T cells” and 10 μl of B cell Ab mix (from EasySep Human B Cell Enrichment Kit) to the tube labeled “B cells.” 9. Incubate at room temperature (20◦ to 23◦ C) for 10 min. 10. Add 10 μl of magnetic beads (from either of the EasySep kits) to both tubes. Mix and incubate at room temperature for 10 min. 11. Add 2.5 ml of PBS/0.1% BSA to each tube. Put tubes in magnet for 5 min. 12. Pour off unbound cells (enriched T and B cells) into a 15-ml conical tube. Centrifuge cells for 5 min at 420 × g, 4◦ C. 13. Count cells (APPENDIX 3A) and adjust cell concentration to 3–5 × 106 cells/ml by diluting with PBS/0.1% BSA.
Flow cytometric cytotoxicity and antibody-binding assay 14. Add 20 μl negative and positive control serum and test serum (e.g., patient serum) to designated wells of a 96-well plate. Use of 96-well plate enables processing of large number of samples. The assay can also be performed in flow cytometry sample tubes if preferred.
15. Add 10 μl of sorted T and B cells to separate wells with each set of sera (negative and positive controls and test serum). 16. Incubate 30 min at room temperature. 17. Add 150 μl PBS/0.1% BSA. Centrifuge 5 min 420 × g, 4◦ C, using a plate carrier. Repeat once. 18. Add 20 μl rabbit complement/well. Incubate 30 min at room temperature. The complement activity may differ between lots and suppliers, and it is therefore important to use an optimal predetermined dilution.
19. Repeat step 17 and then continue to step 20. 20. Add 20 μl goat anti-human IgG-FITC (diluted 1:100)/well. Incubate 20 min at 4◦ C in the dark. This step can be omitted if determination of antibody binding is not required.
21. Repeat step 17. 22. Add 100 μl 7-AAD (diluted 1:100)/well. Transfer cell suspension to flow-cytometer tubes. Additional information about assessment of cell viability using 7-AAD and other reagents such as propidium iodide (PI) can be found in UNIT 9.2. If a flow cytometer equipped with a plate loader is used, transfer of cells to tubes is obviously not necessary.
23. Add 250 μl PBS/0.1% BSA to each tube. 24. Acquire samples on flow cytometer.
Sample acquisition and data analysis 25. Perform appropriate quality control of flow cytometer performance. Use optimized instrument settings as determined in pilot-experiments. 26. Create a forward scatter (FSC) versus side scatter (SSC) dot plot (Fig. 6.34.1). Current Protocols in Cytometry
Phenotypic Analysis
6.34.3 Supplement 66
A 1023
1023
1023 MFI:131
SSC
1.4%
R1 0
0 0
1023
0 0
FSC
1023
0
7-AAD
1023 lgG FITC
B 1023
1023
1023 MFI:597
SSC
61%
R1 0
0 0
1023
0 0
FSC
1023
0
7-AAD
1023 lgG FITC
C 1023
1023
1023 MFI:393
SSC
1.8%
R1 0
0 0
1023
0 0
FSC
1023
0
7-AAD
1023 lgG FITC
D 1023
1023
1023 MFI:595
SSC
72%
R1 0
0 0
1023 FSC
0 0
1023 7-AAD
0
1023 lgG FITC
Figure 6.34.1 Sorted T cells (A, B) and B cells (C, D) were crossmatched against donor specific antibody negative (A and C) and positive serum (B and D). Live and dead lymphocytes were gated (R1) as shown in the left-hand panels. The percentage of dead cells (7-AAD+ ) and the mean fluorescence intensity (MFI) of IgG antibody binding for T cells and B cells are shown in the mid and right hand histograms, respectively.
6.34.4 Supplement 66
Current Protocols in Cytometry
27. Set threshold on the FSC/SSC to eliminate cell debris. 28. Set a gate on live and dead lymphocytes (R1). 29. Display R1-gated events in two one-dimensional histograms for 7-AAD and IgG FITC fluorescence, respectively. 30. Acquire and collect data from at least 1000 R1-gated cells. 31. Use the positive control sample to gate on 7-AAD positive cells. Determine the percentage of dead T and B cells for negative and positive control serum and for patient sample. 32. Calculate the percentage of dead cells for patient serum above negative control serum. 33. Determine the level of antibody binding on T and B cells by the mean fluorescence intensity (MFI) of the IgG FITC histogram peak. 34. Calculate the level of IgG binding for patient serum above the negative control serum.
DETERMINATION OF CYTOTOXICITY AND ANTIBODY BINDING USING PBMC AS DONOR CELLS
BASIC PROTOCOL 2
This protocol uses bulk PBMC as donor targets cells. Fluorochrome-conjugated anti–T lymphocyte (CD3) and anti–B lymphocyte (CD19) antibodies are added for the discrimination of HLA class I (T and B cells) and HLA class II (B cells)–expressing cells.
Materials Phycoerythrin (PE)-conjugated anti-human CD3 antibody (clone SK7, BD Bioscience) Phycoerythrin-Cyanin 7 (PC7) conjugated anti-human CD19 antibody (clone J3-119, Beckman Coulter) Fluorescein isothiocyanate (FITC)–conjugated goat anti-human IgG F(ab 2) antibody (e.g., Jackson ImmunoResearch, cat. no. 096-098) Additional reagents and equipment for determination of cytotoxicity and antibody binding using magnetic bead–purified T and B cells as donor cells (Basic Protocol 1) Isolation of PBMC 1. Prepare PBMC as described in Basic Protocol 1, steps 1 to 5. 2. Count cells (APPENDIX 3A) and adjust cell concentration to 15 × 106 cells/ml by dilution in PBS/0.1% BSA.
Flow cytometric cytotoxicity and antibody binding assay 3. Add 50 μl negative and positive control sera and test serum to designated wells of a 96-well plate. Add 10 μl cell suspension to each serum-containing well (150,000 cells/well). 4. Incubate 30 min at room temperature. 5. Add 150 μl PBS/0.1% BSA. Centrifuge 5 min 420 × g, 4◦ C. Repeat once. 6. Add 30 μl of a mixture of phycoerythrin (PE)-conjugated anti-human CD3 antibody and phycoerythrin-cyanin 7 (PC7) conjugated anti–human CD19 antibody to cells. Resuspend. Determine the optimal dilutions of fluorochrome-conjugated antibodies empirically. Recognize that the recommended dilution provided by manufacturer can differ considerably from optimal dilution for a specific assay. Current Protocols in Cytometry
Phenotypic Analysis
6.34.5 Supplement 66
7. Incubate 20 min at 4◦ C in the dark. 8. Repeat step 5 and then continue to step 9. 9. Add 20 μl rabbit complement/well. Incubate 30 min, room temperature. 10. Repeat step 5. 11. Add 20 μl FITC-conjugated goat anti–human IgG (diluted 1:100)/well. Incubate 20 min at 4◦ C in the dark. 12. Repeat step 5. 13. Add 100 μl 7-AAD (diluted 1:100)/well. Transfer to flow-cytometer tubes. 14. Add 250 μl PBS/0.1% BSA to each tube. 15. Acquire samples on flow cytometer.
Sample acquisition and data analysis 16. Perform appropriate quality control of flow cytometer performance. Use optimized instrument settings as determined in pilot experiments. Optimal PMT voltages and compensation settings are determined using single-colorstained samples and fluorescence minus one (FMO) controls as described previously (Maecker and Trotter, 2006).
17. Apply an FSC threshold to minimize the amount of cell debris. 18. Create two-dimensional dot-plots of FSC versus SSC and CD3 versus CD19 (Fig. 6.34.2). Create a one-dimensional histogram for 7-AAD and IgG FITC fluorescence. There are alternative ways to display and gate cells; e.g., CD3 and CD19 cells can be gated using SSC versus CD3 and CD19 dot-plots. It is informative, though, to include an FSC/SSC plot, because dead cells can be distinguished from live cells by their lower FSC signal in addition to bright 7-AAD fluorescence.
19. Gate on live and dead lymphocytes (R1) and apply gate in CD3 versus CD19 dot-plot. Gate on CD3-positive (R2) and CD19-positive cells (R3). 20. Display the 7-AAD and FITC (IgG) fluorescence for CD3 and CD19 gated cells in one-dimensional histograms. 21. Acquire and collect data from at least 1000 gated CD19+ cells. 22. After acquisition, use the positive control sample to gate on 7-AAD positive cells. Determine the percentage of dead T and B cells for negative and positive control serum and for patient sera. 23. Calculate the frequencies of dead T and B cells for patient serum above negative control serum. 24. Determine the level of antibody binding on T and B cells by the mean fluorescence intensity (MFI) of the IgG FITC histogram peak. 25. Calculate the level of IgG binding for patient serum above negative control serum.
COMMENTARY Background Information Flow Cytometry Based Cytotoxicity and Antibody Binding Assay
In the many transplantation laboratories worldwide, the complement-dependent cytotoxicity (CDC) assay is the “gold-standard”
method for determination of antibody-induced cytotoxicity. However, several disadvantages connected with CDC have been recognized such as high assay variability, subjectivity,
6.34.6 Supplement 66
Current Protocols in Cytometry
A 1023
1023
1023
SSC
MFI:234
R2
R2
2.1% R1 0
0 0
1023
0 0
1023
FSC
0
1023
7-AAD
1023
lgG FITC
1023
1023 MFI:246
R3
R3 CD19 PC7
R3 1.6% R2
0
0 0
1023
0 0
1023
0
1023
7-AAD
CD3 PE
lgG FITC
B 1023
1023
1023 MFI:376
R2
SSC
R2
63% R1
0
0 0
1023
0 0
1023
FSC
0
1023
7-AAD
1023
lgG FITC
1023
1023 MFI:464
R3
R3 CD19 PC7
R3 84% R2
0
0 0
1023 CD3 PE
0 0
1023 7-AAD
0
1023 lgG FITC
Figure 6.34.2 PBMC were crossmatched against donor-specific antibody-negative (A) and -positive serum (B). Live and dead lymphocytes were gated (R1) and displayed in a CD3 versus CD19 plot. The percentage of dead cells (7-AAD+ ) and the mean fluorescence intensity (MFI) of IgG antibody binding for T cells (R2) and B cells (R3) are shown in the plots. Note the distorted CD19 expression with positive serum (B, R3). Phenotypic Analysis
6.34.7 Current Protocols in Cytometry
Supplement 66
and high number of false positives. The first two issues are partly due to the assay design; percentage of dead cells is estimated manually by visual inspection under a fluorescence microscope. Because of these (and other) limitations, alternative techniques based on flow cytometry have been developed (Won et al., 2006; Saw et al., 2008). However, to our knowledge, this new procedure has not yet been implemented in any clinical transplantation laboratory. It is noteworthy that all of these studies used multi-color staining of PBMC targets. We present here an alternative version (Basic Protocol 1) where presorted T and B cells are used as targets. This flow-cytometry approach offers several advantages such as low assay variation and straightforward sample acquisition/analysis. Importantly, the novel assay design using 96-well format enables high-throughput analysis of large numbers of samples. In addition, the multi-well format provides the possibility of using pipetting robots, which eliminate sources of variability. The flow cytometry–based cytotoxicity assay makes it possible to automatically transfer data into the laboratory information system (LIS). This strategy minimizes operator errors, and time-consuming manual steps are avoided. It is important to acknowledge that not only assay performance such as sensitivity and specificity play a role in the choice of method, but also ease-of-use, automation, and the possibility for standardization.
Critical Parameters
Flow Cytometry Based Cytotoxicity and Antibody Binding Assay
Choice of protocol–Sorted versus unsorted cells Both protocols presented here exhibit advantages and disadvantages. One major advantage of using pre-sorted targets is that the number of fluorescence antibodies and markers is minimized. In addition, the gating procedures are greatly simplified; it is sufficient to use a FSC/SSC gate and histogram markers to perform data analysis. Furthermore, no compensation is required because there is negligible spectral overlap between the FITC and 7-AAD fluorescence. Note that if phycoerythrin (PE)– conjugated antibody is used together with 7-AAD, adequate compensation needs to be applied. One disadvantage of Basic Protocol 1 is that the pre-enrichment step prolongs the time-to-result by 25 to 30 min and doubles the total number of samples because T and B cells are acquired in separate tubes/wells. On
the other hand, if bulk PBMC are to be used, greater effort needs to be put into adequate gating and compensation, because two additional fluorescence markers (e.g., CD3 PE and CD19 PC7) are added. As discussed further below, there are issues with altered expression of CD19 after complement incubation. This should be taken into account in the choice of protocol. Choice of cell-separation technique If pre-sorted cells are to be used as targets, care needs to be taken to select an appropriate cell-separation strategy. There are multiple, to some extent similar options that can be considered. The initial question that needs to be addressed is whether whole blood or PBMC are to be used as starting material in the cell-isolation procedure. Take into consideration aspects such as easy-of-use, expenditure of time, recovery, and purity. Select a strategy that best fits your purposes and requirements. Here, T and B cells isolated by negative selection (no beads attached to targets) enable care-free sample acquisition without any distortion of FSC/SSC profile. It is advisable, as part of the assay quality control, to check the purity of sorted cells in each experiment. Choice of negative and positive controls The choice of negative and positive controls in flow cytometry is of great importance. The most appropriate choice here is serum to be used as controls. Negative control serum, without any detectable HLA antibodies (IgG or IgM), can be obtained from healthy blood donors (ABO negative males). The positive control serum can consist of pooled sera from highly immunized patients with HLA class I and/or class II antibodies. There are also commercial sources for both negative and positive control sera. An alternative, but not optimal, choice of positive control for cell lysis is to heat treat cells at 45◦ C for 10 min. The pan-HLA class I specific monoclonal antibody clone W6/32 will induce high cell lysis. Note that the W6/32 antibody is made in mouse, and for simultaneous assessment of antibody binding to target cells, a secondary antibody directed against mouse Ig needs to be used. Optimization of serum volume and cell number In the initial phase of assay setup, the choice of serum:cell volume ratio and number of cells per sample needs to be taken into consideration. In Basic Protocol 1, 20 μl of serum and
6.34.8 Supplement 66
Current Protocols in Cytometry
10 μl of cell volume (ratio 2:1) are used. The number of cells per well can affect the level of binding and lysis, and it is therefore advisable to find the optimal serum:cell ratio and cell number to obtain sufficient sensitivity. As an indication, 20,000 to 60,000 cells/well at a 2:1 serum:cell ratio give comparable levels of lysis and antibody binding. In addition, availability of serum controls and, in particular, test serum (e.g., patient serum) will also play a role in deciding on a serum:cell volume ratio. Also, the number of target cells available will obviously also affect the choice of number of cells/well. Optimization of incubation times and assay sensitivity The incubation times used in various steps of the protocol(s) can have a substantial effect on both cytotoxicity and binding. Therefore, it is of utmost importance to determine the optimal incubation time in order to obtain high sensitivity and low background and minimize time-to-results. In both Basic Protocols 1 and 2, cells are incubated with serum and complement for 30 min. Prolonged incubation times (e.g., 45 or 60 min) with serum or complement will give a higher degree of lysis and binding. This increased sensitivity is evident upon serum dilution. Choice of fluorescent antibodies and viability marker There are several options when it comes to selection of fluorescent antibodies and viability markers. Here, FITC-conjugated secondary IgG antibody combined with 7-AAD viability dye is used, and both are excited by a 488-nm laser. The choice of these two fluorochromes enables acquisition of samples without any need for compensation. Note that cells are brightly stained even with highly diluted 7-AAD dye and a short incubation time (5 to 10 min) prior to acquisition. The well known viability marker propidium iodine (PI) can also be used in combination with both FITC and PE, but proper compensation is necessary for both fluorochromes. If a flow cytometer equipped with a red laser (HeNe ∼630 to 640 nm) is preferred, there are alternative farred excited dyes (e.g., TO-PRO-3) that can be used. In Basic Protocol 2, two additional fluorochromes, i.e., CD3 PE and CD19 PC7, are included. In this case, compensation is clearly needed, and adequate consideration must be put into choosing the right combinations of fluorochromes for a given flow cytometry instrument.
Side-by-side comparison with alternative methods Evaluation of Basic Protocol 1 in this unit shows that it is highly reproducible and robust using either fresh or frozen cells. As part of any assay development, it is common practice to perform side-by-side comparisons with widely accepted standard technologies. Our side-byside comparison with conventional methods such as CDC and flow cytometry crossmatch show a high degree of concordance in results between the techniques. Data from other, more recently developed techniques such as single antigen bead (SAB) based anti–HLA, antiIgG/IgM or anti-C1q antibody detection assays (Chen et al., 2011) can be of guidance in the interpretation of crossmatch data, as well as in the determination of appropriate cut-off values for cytotoxicity and antibody binding.
Troubleshooting Low recovery and purity after T and B cell separation There are, as mentioned above, a plethora of techniques for cell separation; whole blood versus PBMC, positive versus negative selection, and column- versus non-column-based systems. Each technique has its advantages and disadvantages, and it is recommended to scrutinize each strategy in relation to your own requirements. The strategy described in Basic Protocol 1 is robust and very reproducible—a necessity for a clinical application. The starting material for the cell-separation strategy described here is PBMC and not whole blood. The benefit of using PBMC instead of whole blood is that in most cases higher purity can be obtained. This advantage outweighs the potential disadvantages, e.g., prolonged assay time. Initial validation may give some indication about the level of purity that is required in order to obtain reliable data. Loss of CD19 expression after complement incubation In the early stage of method development we attempted to confirm previously published reports using PBMC and staining with CD19 and CD3 after complement incubation. However, we repeatedly failed to detect sufficient numbers of B cells in serum samples with positive control serum. Normal numbers of B cells were observed with HLA antibody–negative serum. The CD3 expression level and number of T cells was normal at all times. Thus, for unknown reason(s), CD19 expression appeared to be lost (or masked) after complement incubation irrespective of the CD19 antibody
Phenotypic Analysis
6.34.9 Current Protocols in Cytometry
Supplement 66
clone. Addition of CD19 antibody or alternative B cell marker (e.g., CD20) before complement minimizes (but does not completely eliminate) this problem (see Fig. 6.34.2). Low cytotoxicity and antibody binding Needless to say, the selection of appropriate negative and positive controls is mandatory for correct interpretation of data and quality control of assay performance. Pooled sera from highly immunized patients used as a positive control may show negative cytotoxicity and/or antibody binding. The most plausible explanation (that initially should be checked) is that no donor-specific antibodies are present in the serum. Selection of sera that cover the great majority of all possible HLA alleles present in the donor pool will most likely prevent this from happening. If unexpectedly negative or weak cytotoxicity and/or binding are observed, there are several ways to investigate this further. Firstly, the titer of donor specific HLA antibody may merely be too low to be detected in the flow cytometry crossmatch assay. Prolonged serum and/or complement incubation time can be applied to increase sensitivity. Sera with donor-specific HLA antibodies directed against a single epitope are unable to provide crosslinking with complement factor C1q—a prerequisite for efficient induction of complement-induced cell lysis. This cytotoxicity-negative, adsorptionpositive (CYNAP) phenomenon can be prevented by adding anti–human Ig prior to addition of complement (Fuller et al., 1997). False positive and false negative crossmatch data False positive or negative results are difficult (or even impossible) to avoid entirely. For instance, complement-inhibitory proteins expressed on target cells or present in serum may block induction of lysis (Morgan, 2000). Unspecific antibody binding to Fc receptors on B cells can be avoided by a 30-min enzymatic treatment of PBMC (5 × 106 cells/ml) with 1 mg/ml pronase (Vaidya et al., 2001). Heat treatment of serum is another strategy that may eliminate false positive reactions (Ta and Scornik, 2002).
Anticipated Results
Flow Cytometry Based Cytotoxicity and Antibody Binding Assay
Evaluation of the assays described herein demonstrates a high degree of concordance with CDC and FCXM (Alheim et al., unpub. observ.). In both Basic Protocol 1 and Basic Protocol 2, isolation of PBMC from
whole blood is performed. The recovery of cells varies from donor to donor, but normally 1–2 × 106 PBMC are obtained per ml blood: ∼90% lymphocytes and ∼10% monocytes. The frequencies of CD3+ cells and CD19+ cells within the lymphocyte population are normally 30% to 40% and 5% to 10%, respectively. In Basic Protocol 1, T and B cells are separated by magnetic beads. The recovery is generally >80% and the purity >95% for both T and B cells. If assay is performed as described here, one ACD tube of whole blood (∼8 ml) will yield a sufficient number T and B cells (20,000 to 30,000 cells/well) for crossmatch of at least 10 to 15 different sera per cell type. Using well defined negative control serum lacking detectable IgG and IgM antibodies gives cytotoxicity background of ∼0% to 5% for T cells and 5% to 10% for B cells, using fresh cells. If frozen cells are used, up to 10% higher background lysis can occasionally be observed.
Time Considerations The 96-well format enables convenient processing of large number of samples. To perform Basic Protocol 1 as described here takes approximately 3.5 hr, cell separations and sample acquisition/analysis included. Shorter time (∼3 hr) is required to complete Basic Protocol 2. It should be noted though that the time needed for acquisition of a given number of B cells from PBMC (∼5% B cells) is longer than for pre-sorted B cells with close to 100% purity. Assay time can be reduced by using shorter wash steps (e.g., 1 min, twice) and reduced incubation time for the secondary IgG antibody step (e.g., 10 min).
Literature Cited Amos, D.B., Bashir, H., Boyle, W., MacQueen, M., and Tiilikainen, A. 1969. A simple micro cytotoxicity test. Transplantation 7:220-223. Chen, G., Sequeira, F., and Tyan, D.B. 2011. Novel C1q assay reveals a clinically relevant subset of human leukocyte antigen antibodies independent of immunoglobulin G strength on single antigen beads. Hum. Immunol. 72:849-858. Fuller, T.C., Fuller, A.A., Golden, M., and Rodey, G.E. 1997. HLA alloantibodies and the mechanism of the antiglobulin-augmented lymphocytotoxicity procedure. Hum. Immunol. 56:94105. Garovoy, R.M., Bigos, M., Perkins, H., and Colombe B. 1983. Flow cytometry analysis: A high technology crossmatch technique facilitating transplantation. Transplant Proc. XV:19391944.
6.34.10 Supplement 66
Current Protocols in Cytometry
Johnson, A.H., Rossen, R.D., and Butler, W.T. 1972. Detection of alloantibodies using a sensitive antiglobulin microcytotoxicity test: Identification of low levels of pre-formed antibodies in accelerated allograft rejection. Tissue Antigens 2:215-226. Lillevang, S.T., Steinbruchel, D.A., Kristensen, T., and Kemp, E. 1992. A new flowcytometric CDC assay for detection of cytotoxic antibodies applied to hamster-to-rat cardiac transplantation. Transplant Proc. 24:537-538. Maecker, H.T. and Trotter, J. 2006. Flow cytometry controls, instrument setup, and the determination of positivity. Cytometry A 69:10371042. Morgan, B.P. 2000. The complement system: An overview. Methods Mol. Biol. 150:1-13. Saw, C.L., Bray, R.A., and Gebel, H.M. 2008. Cytotoxicity and antibody binding by flow cytometry: A single assay to simultaneously assess two parameters. Cytometry B 74:287294. Schonemann, C., Lachmann, N., Kiesewetter, H., and Salama, A. 2004. Flow cytometric detection of complement-activating HLA antibodies. Cytometry B 62:39-45. Ta, M. and Scornik, J.C. 2002. Improved flow cytometric detection of donor-specific HLA class II
antibodies by heat inactivation. Transplantation 73:1611-1614. Talbot, D., Shenton, B.K., Givan, A.L., Proud, G., and Taylor, R.M. 1987. A rapid, objective method for the detection of lymphocytotoxic antibodies using flow cytometry. J. Immunol. Methods 99:137-140. Terasaki, P.I. and McClelland, J.D. 1964. Microdroplet assay of human serum cytotoxins. Nature 204:998-1000. Vaidya, S., Cooper, T.Y., Avandsalehi, J., Barnes, T., Brooks, K., Hymel, P., Noor, M., Sellers, R., Thomas, A., Stewart, D., Daller, J., Fish, J.C., Gugliuzza, K.K., and Bray, R.A. 2001. Improved flow cytometric detection of HLA alloantibodies using pronase: Potential implications in renal transplantation. Transplantation 71:422-428. Wetzsteon, P.J., Head, M.A., Fletcher, L.M., Lye, W.C., and Norman, D.J. 1992. Cytotoxic flowcytometric crossmatches (flow-tox): A comparison with conventional cytotoxicity crossmatch techniques. Hum. Immunol. 35:93-99. Won, D.I., Jeong, H.D., Kim, Y.L., and Suh, J.S. 2006. Simultaneous detection of antibody binding and cytotoxicity in flow cytometry crossmatch for renal transplantation. Cytometry B 70:82-90.
Phenotypic Analysis
6.34.11 Current Protocols in Cytometry
Supplement 66
