Flow cytometric measurement of apoptosis and necrosis in cryopreserved PBPC concentrates from patients with malignant diseases

Abstract

The number of viable precursor cells actually reinfused into patients after high-dose chemotherapy is one of the most clinically important variables determining graft success or failure. A modified, previously described flow cytometric method based on annexin V staining was therefore applied to assess the degree of apoptosis and necrosis in cryopreserved PBPC concentrates from patients with malignant diseases. Twenty-two samples of unmanipulated cryopreserved PBPC concentrates were analyzed by flow cytometry. The samples were triple-stained with anti-CD34 PE, annexin V-FITC and actinomycin D, which enabled the separation of viable, early apoptotic and late apoptotic/necrotic CD34+ precursor cells. Apotosis and necrosis were also measured in the total cell population of the concentrates. Eighty-one percent (range 49–97) of the CD34+ cells were viable, while 7% (range 1–15) were early apoptotic and 12% (range 2–36) were late apoptotic/necrotic after freeze/thaw. There was no difference in apoptosis and necrosis in CD34+ cells harvested from mildly pretreated patients with multiple myeloma and heavily pre-treated patients with non-Hodgkin's lymphoma. Apoptosis and necrosis were higher in the total mature cell population of the concentrates. Thirty-two percent (range 7–69) of the cells were apoptotic and 33% (range 12–60) were necrotic. We conclude that flow cytometric analysis of annexinV/actinomycin D binding in PBPC concentrates is a simple technique that can give additional information of the viability status of the cells post thaw. The present study confirms the relative robustness of human CD34+ precursor cells concerning the freeze/thaw procedure, which are carried out in daily clinical practice.

Bone Marrow Transplantation (2002) 29, 165–171. doi:10.1038/sj.bmt.1703346

Main

Over the last few years, the number of clinical applications for precursor cell transplantation has increased considerably.1 Clinical studies have repeatedly demonstrated that peripheral blood precursor cells (PBPC) are capable of long-term engraftment following myeloablative chemotherapy.2,3 PBPC are transplanted both in the autologous and allogenic setting.1

Until recently, the most common quality assessment of PBPC concentrates has been to quantify the total number of CD34+ cells present in the concentrate before freezing. However, the addition of the cryoprotectant DMSO, cryopreservation and thawing may alter the biological characteristics of cells.4,5,6,7 Ultimately, the number of viable CD34+ cells actually reinfused to the patient, is the most clinically important variable determining graft success or failure.

Keeny et al8 have recently described a new, flow cytometric method for quantitation of the actual number of viable, non-necrotic CD34+ cells in cryopreserved PBPC, using the non-vital dye actinomycin D. Applying this simple, fast method gave an excellent measurement of the fraction and actual number of viable, non-necrotic CD34+ cells. However, PBPC may also, as a consequence of the cryopreservation procedure, die by apoptosis.5,6 The entry into apoptosis is accompanied by a loss of membrane phopholipid asymmetry, resulting in the exposure of phophatidylserine (PS) at the surface of the cells.9 PS exposure during apoptosis is a universal phenomenon of apoptosis occurring in most if not all cell types.10,11,12 The protein annexin V preferentially binds to negatively charged phopholipids like PS.13 Based on this finding, Koopman et al14 in 1994 and Vermes et al15 in 1995, developed a flow cytometric method for detection of apoptosis by combinding annexin V staining with a nonvital dye. During the initial stages of apoptosis, the cell membrane remains intact, while in late apoptosis/necrosis, the cell membrane loses its integrity and becomes leaky. The fluorescent DNA binding agent actinomycin D will enter late apoptotic/necrotic cells, but not early apoptotic cells with an intact cell membrane. Thus viable cells will stain annexin Vneg/actinomycin Dneg, early apoptotic cells will stain annexin Vpos/actinomycin Dneg and late apoptotic/necrotic cells will stain annexin Vpos/actinomycin Dpos.16,17 The flow cytometric annexin V-based method has been applied and verified in several cell types by conventional methods for apoptosis detection such as light microscopy, flow cytometric measurements of hypodiploid nuclei and measurements of DNA fragmentation by DNA electrophoresis.10,11,12,13,14,15,17 However, studies on apotosis and necrosis in PBPC are scant.7,18

In the present study we have measured apoptosis and necrosis in frozen–thawed PBPC collected from 22 patients with malignant diseases using the well-documented annexin V-based method of Vermes et al.15 Since our main interest has been to investigate specifically the status of the CD34+ cells, we have modified the method slightly by staining the cells with PE-conjugated anti-CD34 and substituting the original nonvital-dye PI with actinomycin D, which has less spectral overlap with PE. The same modification has been made by Herault et al, who validated this method on HL-60 cells and human bone marrow CD34+ cells.19

To our knowledge, this is the first time apoptosis has been measured in human cryopreserved peripheral blood CD34+ cells, using the annexin V flow cytometric method. Our main interest was to get an impression of the degree of apoptosis and necrosis in cryopreserved PBPC that are being reinfused to patients in daily clinical practice.

Materials and methods

Patients

Twenty-two autologous PBPC concentrates were collected from 22 patients, most of them with a diagnosis of multiple myeloma or non-Hodgkin's lymphoma (Table 1). The nine patients with multiple myeloma (mean age 58 years), had received only three courses of chemotherapy before mobilization and precursor cell harvest. Most of the lymphoma patients (mean age 32 years), were treated for a relapse and had received a mean of 8.8 courses (range 5–11) of chemotherapy before precursor cell harvest.

Table 1 Flow cytometric measurements of apoptotis and necrosis in CD34+ cells of frozen/thawed PBPC concentrates from 22 patients

PBPC were mobilized with G-CSF and the following chemotherapy regimens. The patients with multiple myeloma were mobilized with cyclophosphamide; the lymphoma patients with MIME (methyl-GAG, iphosphamide, metothrexate, etoposide); the testicular cancer patients with BEF-If (bleomycin, etoposide, cisplatin, iphosphamide) and the sarcoma patients with EVAIA (etoposide, vincristin, adriamycin, iphosphamide, actinomycin D). None of the patients were bad mobilizors (mobilizing <10 × 103 CD34+ cells/ml blood). Eleven of the 22 patients had more than one PBPC collection, but only the first collection was included in the present investigation.

PBPC collections and cryopreservation

The PBPC were collected on a Cobe Spectra (Cobe Laboratories, Gloucester, UK) into citrated autologous plasma.

Before freezing, the cells were concentrated or diluted with autologous plasma to a mean concentration of 100–200 × 106/ml and mixed with DMSO to a final concentration of 10% DMSO. Cells from two fresh samples were stained for flow cytometry (as described below) and analyzed before freezing. After DMSO admixture, portions of 0.5 ml of the PBPC concentrate were transferred to small test tubes and frozen to −180°C in a controlled programmed freezer (Planer Cryo 10; Planer Concentrate Ltd, Sunbury-on-Thames, UK). Thereafter the samples were transferred to a liquid N2 storage container.

Flow cytometry

A small frozen test tube with the PBPC was removed from the N2 storage container and immediately placed in a water bath at 37°C. The tube was gently shaken in the water bath and removed when only a few ice crystals were left. The sample was immediately diluted 1:20 in phosphate-buffered saline (PBS). This dilution ratio was chosen to simulate the dilution that occurs in vivo, when a PBPC concentrate of approximately 200 ml is reinfused into a patient with a blood volume of 4–5 l.

Subsequently, 600 μl of the diluted PBPC was mixed with 5 ml of PBS and centrifuged at 400 g for 10 min. The cell pellet was mixed with 20 μl anti-CD34 PE (HPCA-2; Becton Dickinson, San Jose, CA, USA) and incubated on ice in the dark for 30 min. The cells were washed again and the cell pellet resuspended in 800 μl diluted binding buffer (APOTEST-FITC, Nexin Research, Kattendijke, The Netherlands). Five μl FITC-conjugated annexin V solution, 25 μg/ml (APOTEST-FITC) was added to the test tube together with 25 μl of a stock solution (100 μg/ml in methanol) of actinomycin D (Molecular Probes, Eugene, OR, USA). One negative control without annexin V and one control without actinomycin D were also added in each experiment. The cells were incubated on ice for 15 min in the dark before flow cytometric measurements were performed on a FACSCalibur flow cytometer (Becton Dickinson), using the Cellquest software program. Fluorescence compensation on the flow cytometer was adjusted to minimize spectral overlap. Between 100 and 500000 cells were analyzed and mean CD34+ events were 2700 (range 800–10000).

As indicated at the top of Figure 1, quadrants to define the actinomycin D-positive cells were based on the fluorescence of a fresh PBPC concentrate stained only with anti- CD34 PE to detect the upper limit of background staining of the PE signals into the FL3 channel used to measure actinomycin D.

Figure 1
figure1

Flow cytometric analysis of control cells and a cryopreserved PBPC concentrate to demonstrate the basis for the gating of viable, apoptotic and necrotic cells. (1) A fresh PBPC concentrate stained only with anti-CD34 PE to indicate the upper limit of background staining from anti-CD34 PE into the FL3 channel used to measure actinomycin D fluorescence. (2) Fresh PBMNC from two blood donors stained with annexin V-FITC and actinomycin D to determine the upper limit of background staining of annexin V in normal viable cells. (3) A cryopreserved PBPC concentrate triple-stained with anti CD34 PE, annexin V-FITC and actinomycin D. According to the control samples, the following quadrants of the cytograms were defined. (A) The lower left quadrant shows the viable cells, neither binding annexin V nor showing actinomycin D uptake. (B) The lower right quadrant represents the early apototic cells, binding annexin V but still retaining their cytoplasmic integrity and excluding actinomycin D. (C) The upper right quadrant represents nonviable, late apoptotic/necrotic cells, positive for annexin V and actinomycin D staining. The upper left quadrant shows nonviable necrotic cells/nuclear fragments with no annexin V binding but actinomycin D uptake. The gate R1 was set to define the CD34+ cells which were further analyzed for annexin V and actinomycin D fluorescence.

To get an impression of the background staining of annexin V in normal viable cells,18 control samples of fresh peripheral blood were collected from six healthy blood donors. PBMNCs were prepared by ficoll/metrizoate (Nycomed, Oslo, Norway) density gradient centrifugation, and the PBMNCs were stained with annexin V-FITC and actinomycin D as described above. Quadrants to define the annexin V-positive cells were based on the background staining of these six control samples, as shown in Figure 1.

Statistical analysis

Date were analyzed by the paired-samples T-test, comparing means of two independent samples using the SPSS software package (SPSS, Chicago, IL, USA).

Results

Flow cytometric measurements of apoptosis and necrosis

The annexin V-FITC/actinomycin D bivariate flow cytometry of three control samples and one frozen/thawed PBPC concentrate are shown in Figure 1. The quadrants defining the fractions of viable, apoptotic and necrotic cells are described in the figure.

In the six control samples of PBMC from healthy blood donors that were examined to get an impression of the background staining of annexin V in normal viable cells, 1.1% (mean) (range 0–3.0) of the PBMC cells were apoptotic and 3.2% (mean) (range 0.3–7.3) of the PBMC were necrotic after density centrifugation. These figures were not subtracted from the measurements of the frozen/thawed PBPC concentrate shown in Tables 1 and 2.

Apoptosis and necrosis in the frozen/thawed CD34+ precursor cell population

For two-color analysis, CD34+ cells were first gated on side-scatter and CD34 fluorescence characteristics, and further analyzed for annexin V-FITC fluorescence and actinomycin D uptake, as shown in Figures 1 and 2.

Figure 2
figure2

Flow cytometric analysis demonstrating viable, apoptotic and necrotic CD34+ cells of three cryopreserved PBPC concentrates. (From Figure 1: patients Nos 15, 20 and 14 in Table 1) The diagrams show, from left, the gate R1 set to include the CD34+ cells, which were gated on to the other plots. The plot of annexin V-FITC vs actinomycin D was used to define the early apoptotic CD34+ cells in R2 colored green, and the late apoptotic/necrotic CD34 cells in R3 colored blue. The rest of the viable cells are colored red. The two color plots to the right demonstrate that both the apoptotic and necrotic cells have less forward scatter than the viable cells. Furthermore, one can get an approximate estimate of the fraction of viable, apoptotic and necrotic cells by inspecting the plot of forward scatter vs actinomycin D.

The fraction of viable, early apoptotic and late apoptotic/necrotic CD34+ cells are shown in Tables 1 and 2. The fraction of early apoptotic CD34+ cells was small (mean 7%), and in none of the samples was this fraction higher than 15%. There was no more apoptosis nor necrosis in the samples that were stored for a longer time period. The magnitude of late apoptotic/necrotic cells varied more between the different samples (range 3–36%). No particular diagnosis was predominant in the five samples with more than 10% early apoptotic or 20% late apoptotic/necrotic CD34+ cells. Although the patients with multiple myeloma and NHL differed concerning age, previous chemotherapy and mobilization chemotherapy, no significant difference was found concerning apoptosis and necrosis in the cryopreserved CD34+ cells from either patient group, as shown in Table 3.

Table 3 Comparison of apotosis and necrosis in non-manipulated cryopreserved CD34+ cells collected from patients with non-Hodgkin's lymphoma and multiple myeloma

Twenty-one of the patients have received high-dose chemotherapy and reinfusion of the PBPC concentrate. Only 11 of the patients received one PBPC concentrate, thus only in these 11 patients could the number of viable CD34 cells reinfused be related to hematopoietic recovery after high-dose therapy. These 11 patients received between 1.7 and 11 × 106 annexin Vneg/actinomycin Dneg viable CD34+ cells/kg, and all obtained normal hematological recovery (granulocytes >0.5 × 109/l days 7–12 and platelets >20 × 109/l days 9–13. No significant relationship between viable CD34+ cell count and days to hematological recovery could be demonstrated in these 11 patients.

Figure 2 shows bivariate flow cytometry plots of CD34+ cells from three different precursor cell concentrates with varying degree of apoptosis and necrosis. This Figure demonstrates that most of the apoptotic cells had a somewhat higher actinomycin D uptake than the viable cells. Furthermore, the annexin V-positive apoptotic and necrotic CD34+ cells generally had less forward scatter than the viable cells.

Apotosis and necrosis in the frozen/thawed total cell population

As shown in Figure 1, a large proportion of the total cell population, consisting mainly of mature cells, were annexin V-positive. The scatter characteristics of the frozen/thawed cells also differed from the scatter profile of cells from fresh PBPC concentrates, as demonstrated in Figure 1. As demonstrated in Table 2, less than half of the total cell population was annexin V-negative.

Table 2 Flow cytometric measurements of apotosis and necrosis in frozen/thawed PBPC concentrates from 22 patients

Discussion

In the present study we have shown that a rather small fraction of peripheral blood CD34+ cells enter into apoptosis during the process of DMSO addition, freezing and thawing. On the other hand, the fraction of apoptosis and necrosis was high in the total cell population (Figure 1 and Table 2).

To our knowledge this is the first time apoptosis and necrosis have been measured in human cryopreserved peripheral blood CD34+ cells, using the well-documented annexin V flow cytometric method. Anthony et al18 have stained fresh autologous PBPC concentrates with annexin V (without the double staining with nonvital dye), and report that 87.6% (range 30–96.6) of the CD34+ cells were annexin V-negative, which is almost identical to our figure of 81% (range 49–97). However, the figures are not strictly comparable, since the cells in our study were investigated after DMSO addition and freeze/thaw. De Boer et al5 did flow cytometric measurements of frozen/thawed MACS-isolated CD34+ cells and showed that both loss of L-selectin expression and emergence of apoptosis occurred after freeze–thawing. Recently, Schuurhuis et al,7 have extended this work by measuring early apoptosis in both purified and nonpurified CD34+ cells from cryopreserved leukapheresis samples. Using the early apoptotic marker SytoR16 in combination with actinomycin D, they report that in unpurified frozen/thawed CD34+ PBPC from 15 patients, as many as 37% of the CD34+ cells were early apoptotic and 29% were necrotic. Their figure of apoptotic CD34+ cells is clearly higher than the figure in our investigation. Their measurements of apoptosis and necrosis in the CD34+ cells are more in accordance with our measurements of apoptosis and necrosis in the total cell population. According to Schuurhuis et al, SytoR16 detects a somewhat higher fraction of early apoptotic cells than annexin V. However, we do not believe this can be the only reason for the large discrepancy of measurements. The addition of DMSO, freeze/thaw process, dilution and handling of the sample post thaw may influence the post-thaw viability. More flow cytometric studies of post-freeze/thaw PBPC apoptosis are needed to clarify the difference between the two studies.

The exposure of PS on the outer cell membrane acts as a recognition signal for macrophages to phagocytose the apoptotic cells. Thus, the annexin V-positive CD34 cells will probably be eliminated from the hematopoietic system and thus have no effect on the long-term engraftment following myeloablative chemotherapy. Quantitative measurements of annexin V-negative CD34 cells prior to reinfusion of PBPC may be important in the clinical setting to identify patients at risk of graft failure. This may be especially important for precursor cell concentrates with a low CD34+ cell count or for concentrates that have been manipulated by purging/positive selection. In the present study, none of the 11 patients were reinfused with PBPC concentrates containing less than 1.7 × 106 annexin V negative CD34+ cells/kg, and all the patients had a normal hematological recovery.

As shown in Figure 2, the present study indicates that one can get a rough estimate of the combined fraction of apoptotic + necrotic CD34+ cells, just by looking at the scatter characteristics. This is in accordance with other studies.5,12 The observation that the apoptotic cells had a somewhat higher actinomycin D uptake than the viable cells and lower actinomycin D uptake than the necrotic cells, is also in accordance with other flow cytometric investigations.16,20

We were surprised to find that as many as 31% (mean) of the total cell population in the PBPC concentrate (mainly mature blood cells), was in early apoptosis after cryopreservation. The late apoptotic/necrotic total cell fraction in our study was 33%, which is not too far from the findings of others.21 Possible unwanted clinical implications of reinfusing a concentrate with 49–81% apoptotic and necrotic cells are unknown. The common clinical side-effects of having a thawed cryopreserved PBPC concentrate reinfused are flushing, nausea, dyspnea, post-infusion rigors and hypotension. These symptoms are usually attributed to DMSO-induced histamine release. However, a small clinical study has shown that these symptoms were also present in patients who had DMSO removed from the PBPC product before reinfusion, suggesting a possible harmful effect of reinfusing poorly cryopreserved non-viable cells.22 On the other hand, one could speculate whether contaminating tumor cells in the PBPC concentrate may also become apoptotic and necrotic after the freeze/thaw process, suggesting a possible advantageous purging effect of the freezing procedure.

In conclusion, this study confirms the relative robustness of human CD34+ cells during the freeze/thaw procedures which are carried out in daily clinical practice. The flow cytometric three-color measurements of peripheral blood precursor cell concentrates stained with anti-CD34, annexin V and actinomycin D is a simple, fast and reproducible method. This investigation might be useful to assess the potential of progenitor cells for in vivo hematopoietic engraftment and further ex vivo manipulations, such as precursor cell expansion and gene therapy.

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Acknowledgements

This work was supported by grants from the Norwegian Cancer Society.

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Correspondence to JF Abrahamsen.

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Abrahamsen, J., Bakken, A., Bruserud, Ø. et al. Flow cytometric measurement of apoptosis and necrosis in cryopreserved PBPC concentrates from patients with malignant diseases. Bone Marrow Transplant 29, 165–171 (2002). https://doi.org/10.1038/sj.bmt.1703346

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Keywords

  • apoptosis
  • annexin V
  • actinomycin D
  • CD34-positive cells
  • cryopreservation

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