Paroxysmal nocturnal hemoglobinuria (PNH) is an acquired clonal hematopoietic disorder characterized by the existence of deletions, insertions, or point mutations in the PIG-A (phosphatidylinositolglycan complementation class A) gene located in the human chromosome X at Xp22.1 (1). This genetic abnormality involves pluripotent hematopoietic stem cells, the PNH clone usually coexisting with normal hematopoiesis (2, 3). From the pathogenetic point of view, this genetic alteration translates into a total or partial deficiency in the PNH clone of surface proteins attached to the cell by a glycophosphatidylinositol (GPI) anchor (4). At present, a high number of membrane proteins have been identified which use GPI to be expressed on the cell surface. Among these proteins, both CD55 (decay accelerating factor [DAF]) and CD59 (membrane inhibitor of reactive lysis [MIRL]) have been identified as playing an important role in the regulation of complement activation by preventing it from being attached to self-membranes (4–13). The deficiency of these proteins leads to an increased susceptibility of the PNH cells to complement-mediated cell lysis (5, 6). For many years, the susceptibility of red cells to complement-mediated lysis was used as an indirect indicator for PNH disease. More recently, the development and availability of monoclonal antibodies (mAb) specific for GPI-anchored proteins has allowed a more specific identification and characterization of PNH+ cells present in the peripheral blood (PB) from patients with PNH, using flow cytometry (14). From the deficient GPI-anchored proteins, CD55 and CD59 have been the most widely tested ones (1–10, 12–28). This is due to the fact that both proteins are normally expressed on leucocytes, erythrocytes, and platelets at relatively high levels, at the same time they constantly decrease their expression in PNH+ hematopoietic cells (1, 4). Accordingly, at present, a routine screening of CD55 and CD59 expression on PB hematopoietic cells is used for the diagnosis of PNH (2–4, 6–10, 13–20, 22–24, 26, 28). However, the simultaneous measurement of CD55 and CD59 expression on the different hematopoietic cell types present in whole blood samples shows several limitations due to the great differences that exist on the relative frequencies of red cells, platelets, and leucocytes. In this sense, whereas a few microliters of PB are commonly used to stain red cells and platelets for the analysis of CD55 and CD59 expression, typically between 50 and 200 μL of PB is used to study CD55 and CD59 expression on leucocytes (4, 6–8, 15–19, 23, 24). However, the use of such relatively high sample volumes may be associated with variable levels of CD55 and CD59 expression on leucocytes due to technical artifacts. As suggested by Richards et al. (13), only once prelysing methods are performed to isolate leucocytes, anti-CD55 and anti-CD59 antibodies can be titrated optimally to label a standard number of cells.
In the present study, we analyze the effects of stain-lyse-and-then-wash techniques as compared with lyse-wash-and-then-stain procedures on CD55 and CD59 expression on the major subsets of PB leucocytes, as measured by flow cytometry. Our major goal was to calculate the minimum amount of anti-CD55 and anti-CD59 mAbs required to be added to a minimum volume (3 μL) of PB, which would allow an optimal staining for both antigens on red cells, platelets, and leucocytes, in the same tube.
MATERIALS AND METHODS
Ten normal PB samples from healthy volunteers and a patient diagnosed as suffering from PNH with type II red cells were analyzed in the present study. All samples were obtained by venipuncture in Vacutainer tubes containing tripotasium (K3) ethylene-diaminetetraacetic acid (EDTA) as the anticoagulant (Becton/Dickinson Labware, Franklin Lakes, NJ). The overall number of erythrocytes and leucocytes present in the PB samples analyzed was 4.19 × 106 ± 0.96 × 106/μL (range: 2.67 × 106/μL–5.51 × 106/μL) and 8.40 × 103 ± 1.51 × 103/μL (range: 6.60 × 103/μL–11.6 × 103/μL), respectively.
In 5 of the 10 normal PB samples, sample preparation was performed in parallel using two procedures: a lyse-wash-and-then-stain technique and a stain-lyse-and-then-wash method. In both techniques, the expression of the CD55 and CD59 mAbs was evaluated specifically for each of the four major PB leucocyte subsets (lymphocytes, monocytes, neutrophils, eosinophils) using three volumes of PB per tube (50, 100, and 200 μL) for staining purposes. All these samples were processed within a maximum period of 4 h after they were obtained. Accordingly, each of these five PB samples was aliquoted in 12 tubes to which 200 (four tubes), 100 (four tubes), or 50 μL (four tubes) was added. Two of the four tubes containing identical sample volumes were treated according to the lyse-wash-and-then-stain protocol and the other two tubes with the stain-lyse-and-then-wash technique described below.
The remaining five normal PB samples (either 3, 5, or 10 μL of PB sample per tube) were just stained (with either 3 or 10 μL of the CD55 and with either 3 or 10 μL of the CD59 mAb), incubated (either 15 or 30 min) in the darkness (at room temperature [RT]), and measured in the flow cytometer after adding 0.5 mL of phosphate-buffered saline (PBS). For these latter stainings, the following combinations of mAbs (fluorescein isothiocyanate [FITC]/ phycoerythrin [PE]/peridinin chlorophyll protein [PerCP]) were used: CD64/CD55/CD45+CD61 and CD64/CD59/CD45+CD61. In all cases, the concentration of the CD55 and CD59 mAbs used was 0.05 and 0.2 μg/μL, respectively. The non-lyse-non-wash protocol was used for the PNH+ sample.
Briefly, to each of the six tubes/samples treated according to the lyse-wash-and-then-stain protocol, 2 mL of FACS lysing solution (Becton/Dickinson Biosciences [BDB], San Jose, CA) was added. Following a 10-min incubation period at RT, the tubes were centrifuged for 5 min at 540g, the supernatant was removed, and 2 mL of PBS was added to the cell pellet. After gentle vortexing, a second centrifugation step was performed. Then, 10 μL of an anti-CD45 mAb (clone: HI30) conjugated with FITC (Caltag Laboratories, San Francisco, CA), 10 μL of either an anti-CD55 reagent (clone: IA10) conjugated with PE (BDB), or 10 μL of an anti-CD59 mAb (clone: p282 H19) conjugated with PE (BDB) were added to each tube. After 15 min of incubation at RT in the darkness, the cells were resuspended in 0.5 mL PBS for later analysis in the flow cytometer.
Briefly, to each of the six tubes/samples treated with this protocol, 10 μL of the CD45 plus 10 μL of either the CD59-PE or the CD55-PE mAb reagents were added. After gentle vortexing, the samples were incubated for 15 min at RT in the darkness. Immediately after this incubation period, 2 mL of FACS lysing solution was added to each tube and another incubation for 10 min at RT was performed. Then, the tubes were centrifuged for 5 min at 540g, the supernatant was removed, 2 mL of PBS was added per tube, and a second centrifugation step (5 min, 540g) was performed. The cell pellet was finally resuspended in 0.5 mL of PBS for later analysis in the flow cytometer.
Flow Cytometry Data Acquisition and Analysis
Data acquisition was performed immediately after completion of sample preparation using a FACSort flow cytometer and the CellQuest software program (BDB). Instrument set-up for three-color immunofluorescence adjusting the forward scatter (FSC) threshold to exclude debris (for the simultaneous analysis of red cells, platelets, and leucocytes) was performed prior to data collection according to well-established protocols. For tubes stained with CD64-FITC/CD55-PE or CD59-PE/CD45-PerCP plus CD61-PerCP, a second acquisition step was performed through a side scatter (SSC)/CD45 live gate to enrich in SSChi/CD45+ leucocytes. For each sample/tube, information on at least 5 × 104 events corresponding to either the leucocytes or the whole PB cellularity was collected sequentially and stored. The Paint-A-Gate PRO software program (BDB) was used for data analysis. Color gates based on FSC/SSC and SSC/CD45-CD61 or SSC/CD45 and CD45/CD64 bivariate dot plots were drawn to identify specifically the red cells and platelets or the lymphocytes, monocytes, and neutrophils present in the sample, respectively. Results on the mean fluorescence intensity (MFI expressed in arbitrary relative linear units scaled from 0 to 104) and the coefficient of variation (CV) obtained for both CD55-PE and CD59-PE were calculated for each of the above listed PB cell subsets.
For all parameters under study, their M ± SD, median, and range were calculated using the SPSS 9.0 software program (SPSS, Chicago, IL). In order to explore the potential existence of statistically significant differences between groups, the Student t-test for paired variables was used. The Pearson correlation test (SPSS software program) was used to establish the potential relationship between the number of erythrocytes present in PB samples and the patterns of staining for CD55 and CD59 with the sample preparation techniques compared here (data not shown). P < 0.01 was considered to be associated with statistical significance.
Figures 1 and 2 show the results on CD55 and CD59 expression obtained with the two conventional techniques compared here for each of the four major PB subsets of leucocytes analyzed. For all the PB leucocyte subsets analyzed, the staining for both CD55 and CD59 significantly increased (P < 0.01) when the lyse-wash-and-then-stain technique was used compared with the stain-lyse-and-then-wash method, except for the CD55 MFI on eosinophils which only showed a slight increase (Figs. 1-3). Interestingly, this overall increase in the MFI of CD59 and CD55 was associated with a significantly more homogeneous expression of both antigens on lymphocytes and monocytes as reflected by a significantly lower CV (P < 0.009). No significant differences between both techniques were detected regarding the CD55 and CD59 CV either on neutrophils or on eosinophils. It should be noted that such differences were observed independently of the sample volumes used for staining purposes (50, 100, or 200 μL).
Interestingly, once a lyse-wash-and-then-stain protocol was used, no major differences were detected for either the CD55 and CD59 MFI or their CV according to the volume of sample stained, with the exception of the CD55 MFI (Fig. 1) found on neutrophils that increased (P < 0.002) from 1,859 ± 432 to 2,392 ± 334, once 50 μL of PB was stained instead of 200 μL. In contrast, once the stain-lyse-and-then-wash protocol was employed, significant differences were observed frequently on CD59 and CD55 expression on the different PB leucocyte subsets depending on the amount of sample stained per tube (Figs. 1, 2). Accordingly, a decrease on CD59 MFI for the lymphocytes was observed once 200 μL instead of either 50 or 100 μL of PB was used (17.02 ± 4.72 versus 53.58 ± 13.89 and 28.36 ± 4.92, P = 0.008). In a similar way, the MFI obtained for CD59 increased significantly (P < 0.01) on neutrophils once 50 μL instead of either 200 or 100 μL of PB was used (87.96 ± 26.30 versus 26.04 ± 7.53 and 45.82 ± 8.92, respectively). An increase on CD59 MFI for the eosinophils (P < 0.01) was also observed once 50 μL instead of either 100 or 200 μL of PB was used (293.51 ± 45.91 versus 182.31 ± 28.70 and 93.62 ± 41.75, respectively). Finally, an increase on the CD55 CV for the eosinophils was observed once 200 μL instead of 50 μL of PB was used for staining purposes (26.90 ± 1.94 versus 30.70 ± 2.66, P = 0.009).
Figure 4 shows how red cells, platelets, and leucocytes (including lymphocytes, monocytes, and neutrophils) were identified simultaneously in stained-non-lysed-and-non-washed PB samples after acquiring the same sample in two consecutive steps. As shown in dot plots A and B (Fig. 4) corresponding to the first acquisition step, red cells were identified by their light scatter properties (FSCint/hi/SSChi) in the absence of CD45 and CD61 expression, platelets constantly displayed a low SSC and FSC together with reactivity for CD61, and leucocytes (Fig. 4D,E) had higher light scatter properties (SSChi/FSChi) than platelets at the same time they were CD45+; among them, a CD64/SSC dot plot was used to discriminate among monocytes, neutrophils, and lymphocytes (Fig. 4E). As shown in Figures 4F and 5A-D, the patterns of expression of both CD55 and CD59 within each PB cell subset analyzed are homogeneous whereas the presence of a population of type II (CD59dim) red cells and CD55-deficient leucocytes can be detected with the same technique in the PNH+ sample analyzed (Fig. 5E-H).
Once 3 μL of PB was stained with the CD64/CD55 or CD59/CD45 and CD61 combinations of mAb for 30 min in the darkness (RT; Tables1 and 2), no significantly (P > 0.01) different patterns of CD55 and CD59 expression were observed for the different leucocyte subsets analyzed as compared with the use of prelysing methods (Figs. 1, 2), with the exception of a lower CD55 MFI on neutrophils (P < 0.002).
Tables 1 and 2 show the results obtained with staining protocols for the CD55 and CD59 antigens based on the use of relatively low sample volumes (≤10 μL of PB) in the absence of lysing and washing procedures. As shown in both tables, the strongest and more homogeneous staining patterns obtained for both antigens in the different PB leucocyte subsets analyzed were observed when 10 μL of mAb was used to stain 3 μL of sample for an incubation period of 30 min. Once 3 μL instead of 10 μL of mAb was used, a significant decrease in the MFI obtained for both markers with a parallel increase in the CV was observed for almost every cell population (Tables 1 and 2).
In a similar way, once cells were stained with 10 μL of mAb, a decrease in the MFI with higher CVs was observed for incubation periods of 15 versus 30 min, as well as for sample volumes of 10 and 5 μL versus 3 μL (Tables 1 and 2); however, differences did not reach statistical significance once different sample volumes and incubation periods were compared for the same amount of mAb reagent.
For the last 15 years, flow cytometry-based immunophenotypic analysis of CD55 and CD59 expression on PB red cells, platelets, and leucocytes has been extensively used for the diagnosis of PNH (2–4, 6–10, 13–20, 22–24, 26, 28). Prior to the introduction of flow cytometry, most of the patients screened for PNH displayed a predominantly hemolytic clinical picture. This was mainly due to the fact that diagnostic tests available then were almost exclusively based on the increased susceptibility of PNH red cells to hemolysis (1–16, 20, 21, 25, 27, 28). Even for this purpose, such diagnostic tests were rather limited due to both their low sensitivity for identifying small populations of PNH+ red cells, especially in patients who had been transfused, and their qualitative nature, which did not distinguish between partially and completely deficient cells (3, 4, 6, 7, 9, 13, 24). In contrast, flow cytometry allows the analysis of all PB populations including red cells, platelets, and the leucocyte subsets. At the same time, it also permits a quantitative evaluation of the defective antigen expression and the identification of small cell populations belonging to the PNH clone (3–9, 13, 17, 18, 24). Thus, because of the availability of flow cytometry for the diagnosis of PNH, the cohort of patients screened has increased significantly to include individuals with aplastic anemia, unexplained peripheral cytopenias, infectious diseases, and/or thromboembolic episodes (9, 14–16, 19–22, 25, 26).
At present, many GPI-linked surface proteins have been described as being deficient in PNH+ cells. Of these proteins, the CD55 and CD59 complement-activation regulatory proteins are the most widely used on the flow cytometry-based diagnostic screening for PNH (1–10, 12–28). This is related to the fact that: 1) both proteins are expressed universally in the different cell populations present in normal PB and, 2) abnormal expression of CD55 and CD59 correlates with the clinical behavior of the disease (1, 2, 4–15, 19, 21–28).
Disturbing levels of variability in the levels of CD55 and CD59 expression are observed once different sample preparation protocols are used in combination with flow cytometry (Table 3). MFI levels are higher when prelysing methods are employed for the flow cytometry immunophenotypic analysis of CD55 and CD59 expression on PB leucocytes (13). This is mainly due to the difficulties of obtaining optimal titration curves for the anti-CD55 and anti-CD59 mAbs once a high number of red cells are present in the tube. However, a careful analysis of the literature shows that many sample staining protocols are used, some of which are not optimal (4–10, 14–25, 29).
Table 3. Summary of the Sample Preparation Protocols Used in Papers Reported in the Literature, in which CD55 and/or CD59 Expression Is Analyzed in Different Leucocyte subsets*
Molecules analyzed on leucocytes
Sample/sample preparation protocol used (sample volume)
PMN, polymorphonuclear leucocytes; MNC, mononuclear cells; BM, bone marrow; NS, not specified.
Our results demonstrate that upon comparing a stain-lyse-and-then-wash technique with a lyse-wash-and-then-stain protocol, the presence of hundreds of millions of red cells at the time PB leucocytes are stained with anti-CD55 and anti-CD59 mAb, the MFI obtained for both antigens is decreased significantly; in addition, a more heterogeneous pattern of antigen expression on virtually all major subpopulations of PB leucocytes is also observed. These observations are further supported by the fact that once the stain-lyse-and-then-wash protocol was used, but not the lyse-wash-and-then-stain technique, a significant correlation was observed between the number of red cells present in each stained tube and the CD55 and CD59 MFI and CV obtained (data not shown). In line with these findings, when the lyse-wash-and-then-stain protocol was used, an optimal staining of PB leucocytes was obtained, independently of the volume of sample used. In contrast, when the stain-lyse-and-then-wash protocol was employed, statistically significant differences were observed on CD55 and CD59 MFI obtained depending on the amount of sample stained. These results indicate that optimal staining for red cells, platelets, and leucocytes could not be achieved simultaneously in a single tube unless changes were introduced in the conventional staining procedures. Such modifications could be related to either the use of higher amounts of anti-CD55 and CD59 mAb, a longer incubation period, and/or the use of smaller sample volumes. At the same time, additional mAb reagents should be used in combination with the anti-CD55 and/or anti-CD59 mAbs for the unequivocal identification of the major populations of cells of interest present in the blood. For that purpose, we used a combination of three mAbs conjugated with two fluorochromes (CD64 FITC/CD45-CD61 PerCP) that allowed the simultaneous identification of red cells, platelets, and the three major leucocyte subsets (monocytes, lymphocytes, and neutrophils) present in whole blood samples. A limitation of the specific mAb combination was the presence of giant platelets in the PB; the use of CD61 and CD45 reagents combined with different fluorochromes (i.e., PerCP and allophycocyanin [APC]) would allow, in these cases, a clear distinction between platelets (CD69+/CD45-) and leucocytes (CD45) in these samples (data not shown).
In order to get simultaneous information on red cells, platelets, and leucocytes, a stain-non-lyse-non-wash method should be used. Through this approach, we investigated the optimal conditions regarding sample volume, the amount of mAb reagents, and the incubation period to stain for both CD55 and CD59 in all major blood cell populations. Our results show that the amount of mAb used to stain both the CD55 and CD59 antigens was critical to obtain an optimal staining of all major PB leucocyte subsets and that this approach allows the detection of PNH+ cells.
No significant differences were detected, when the staining patterns obtained after different incubation periods (15 versus 30 min at RT) or distinct sample volumes (3–10 μL) were compared. Nevertheless, a tendency toward higher MFI and lower CV values was observed for both the CD55 and CD59 antigens when 3 μL of sample was stained for 30 min prior to data acquisition in the flow cytometer. Accordingly, our recommendation would be to keep to a maximum the amount of antibody added to assure that saturation is achieved independently of the number of cells present in 3 μL of sample. Additionally, incubation periods should be kept to at least 30 min (RT).
In summary, we show that the sample preparation protocol has a significant impact on the quality of the staining for the CD55 and CD59 antigens obtained for the major PB leucocyte subsets. Additionally, we propose a simple and reliable stain-non-lyse-non-wash method for the simultaneous analysis of the expression of both CD55 and CD59 on PB red cells, platelets, neutrophils, monocytes, and lymphocytes. Although we show the reliability of this technique to detect PNH+ cells in PB, studies on large numbers of clinical samples from known PNH+ patients are ongoing for the evaluation of the assay.