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Originally published online as doi:10.1189/jlb.0304171 on June 3, 2004

Published online before print June 3, 2004
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(Journal of Leukocyte Biology. 2004;76:609-615.)
© 2004 by Society for Leukocyte Biology

Loss of lineage antigens is a common feature of apoptotic lymphocytes

D. Diaz*, A. Prieto*, H. Barcenilla*, J. Monserrat*, P. Prieto*, M. A. Sánchez*, E. Reyes*,{dagger}, M. P. Hernandez-Fuentes{ddagger}, A. de la Hera*, A. Orfao§ and M. Alvarez-Mon*,¶,1

* CNB-CSIC R&D Associated Unit, Department of Medicine, University of Alcalá, Madrid, Spain;
{dagger} Investigation Unit, Industrial Farmacéutica Cantabria, Madrid, Spain;
{ddagger} Department of Immunology, Imperial College, London, United Kingdom;
§ Flow Cytometry Unit, Cancer Research Center and Department of Medicine, University of Salamanca, Spain; and
Immune System Diseases and Oncology Service, University Hospital "Príncipe de Asturias," Alcalá de Henares, Madrid, Spain

1 Correspondence: Departamento de Medicina, Universidad de Alcalá, Carretera Madrid-Barcelona, Km 33.600, 28871 Alcalá de Henares (Madrid), Spain. E-mail: mams{at}tsai.es


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The analysis of apoptosis in cell populations involves the detection of their specific lineage antigen (LAg) expression. This experimental approach relies on their assumed constant expression, but it is unclear whether such expression is actually maintained during cell death. We examined whether the loss of LAgs is a common feature of apoptotic lymphocytes and whether some might completely lose their LAgs. The changes in the expression of CD3, CD5, CD8, CD4, CD28, CD56, and CD19 were monitored in highly purified lymphocyte populations obtained by negative selection in a fluorescence-activated cell sorter. These were cultured for 24 h with or without phytohemagglutinin or staurosporin. For each LAg-positive subset studied, apoptosis was consistently more common among cells showing partial or total loss of LAg expression compared with cells maintaining their initial LAg levels. The kinetics of expression loss was rapid for CD8, CD56, and CD28, and more than 80% of initial expression was lost in the early stages of apoptosis but was slower for CD3, CD5, and CD4. For CD3 and CD5, expression was dependent on the apoptotic stimulus used. It is interesting that loss of antigen expression was independent of cell size. This phenomenon was also found in nonmanipulated, highly pure CD19 B lymphocytes of peripheral blood mononuclear cells from B chronic lymphocytic leukemia patients. Loss of LAg expression appeared to be a common feature of apoptotic lymphocytes under all the conditions assayed. The different kinetic patterns of LAg loss suggest apoptotic cells might actively regulate this process.

Key Words: FACS • CD • annexin V • 7-amino actinomycin D • physiological • loss


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Apoptosis is a form of programmed cell death, which is essential for normal development, tissue homeostasis, and immune system function [1 ]. Alterations in this process are associated with tumor transformation and autoimmune and degenerative diseases [2 3 4 ]. The measurement of apoptosis in ex vivo cultures of lymphocytes from peripheral blood has provided insights into the pathogenesis of autoimmune and inflammatory diseases [5 ]. Since the early 1990s, multiparameter flow cytometry has facilitated the study of apoptosis in specific subsets of cells within heterogeneous cell samples [6 7 8 9 ]. Several methods have been developed to identify and quantify apoptotic cells within phenotypically defined subsets [6 7 8 9 ]. Typically, apoptotic cells are identified by evaluating the extent of characteristic lesions such as DNA fragmentation [10 , 11 ], altered cell permeability, and phosphatidylserine exposure [7 , 9 , 12 , 13 ]. To accurately measure apoptosis within specific subpopulations of cells in heterogeneous cellular samples, many studies have relied on the supposed maintained expression of lineage antigens (LAgs) characteristically expressed by the subsets of interest [14 ].

Along these lines, the reported increased apoptosis of lymphocytes with weak expression of LAgs such as CD56 or CD94 [15 , 16 ] has been interpreted in terms of specific lymphocyte subsets with increased susceptibility to this cell death mechanism. However, an alternative interpretation could be that apoptosis is physiologically associated with down-regulation in the expression of some LAgs. In support of these interpretations, two studies have shown that the expression of LAgs decreases during apoptosis in CD4+ and CD8+ T lymphocytes [17 , 18 ].

The aim of the present study was to analyze the relationship between apoptosis and LAg expression in lymphocytes. We studied the expression of different LAgs in negatively sorted, homogeneous cell population cultures under different experimental conditions. Furthermore, we analyzed this cellular behavior in a model of in vitro nonmanipulated, highly pure lymphocytes expressing a defined LAg (CD19) in peripheral blood mononuclear cells (PBMCs) from B chronic lymphocytic leukemia. The results show that loss of antigen expression occurs in different stages of apoptosis, the magnitude largely depending on the antigen analyzed. To simultaneously quantify the number of cells in each stage of apoptosis and to evaluate potential changes in the expression of LAgs expressed by these cells, we used a flow cytometry-based method to measure apoptosis, monitoring alterations in cell permeability and phosphatidylserine exposure [7 , 9 , 12 , 13 ]. Our group recently improved this technique [14 ].


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Cell samples
PBMCs from six healthy controls and from four B cell chronic lymphocytic leukemia (B-CLL) patients (diagnosed according to current criteria; ref. [19 ]) were obtained by Ficoll-Hypaque (LymphoprepTM, Axis-Shield, Oslo, Norway) density gradient centrifugation as described previously [20 ]. Purified PBMCs were resuspended in RPMI 1640 (BioWhittaker Products, Verviers, Belgium) supplemented with 10% heat-inactivated fetal calf serum, 25 mM HEPES (BioWhittaker Products), and 1% penicillin-streptomycin (BioWhittaker Products). Initial cell enumeration was performed by conventional light microscopy using a Neubauer chamber and following trypan blue dead cell exclusion criteria and by flow cytometry (FACSCalibur, Becton Dickinson Biosciences, San Jose, CA) as described previously [21 ]. The viability of fresh PBMCs was checked by trypan blue (light microscopy) and 7-amino actinomycin (7-AAD; flow cytometry) exclusion [12 ] and was always greater than 95%. The final cell concentration was adjusted to 0.5 x 106 cells/ml. In all cultures, cells were plated at a density of 2.5 x 105 cells/ml.

Fluorescence-activated cell sorting (FACS)
Lymphocyte subsets were purified by negative selection using flow cytometry cell sorting. For this, fresh PBMCs were incubated with different three- and four-color combinations of fluorescein isothiocyanate (FITC; green), phycoerythrin (PE; orange), peridinin chlorophyll protein conjugate (PerCP; red), and allophycocyanin (APC)-labeled monoclonal antibodies (mAb). To purify CD4+CD28+ cells, CD3+CD8+, and CD3+CD5+ T cell receptor (TCR){alpha}ß+ T cells as well as B cells and natural killer (NK) cells, PBMCs were depleted of different cell subsets using the following combinations of mAb (FITC/PE/PerCP/APC): TCR{gamma}{delta}/CD14–CD16–CD56/CD8/CD19; TCR{gamma}{delta}/CD14–CD16–CD56/CD4/CD19; TCR{gamma}{delta}/CD14–CD16–CD56/–/CD19; –/CD14–CD16–CD56/CD3/–; –/CD14/CD3/CD19, respectively. Anti-CD19-APC, anti-CD56-PE, anti-CD8-PerCP, anti-CD4-PerCP, anti-CD3-PerCP, and anti-TCR{gamma}{delta}-FITC were obtained from Becton Dickinson Biosciences, and anti-CD14-PE, anti-CD16-PE, and anti-CD45-FITC were purchased from Caltag Laboratories (San Francisco, CA).

All sorting experiments were performed in a FACSTARplus flow cytometer using CellQUEST software (Becton Dickinson Biosciences). A biparametric gate in a forward- versus side-scatter (FSC and SSC, respectively) dot plot was drawn around the lymphocyte population with low FSC and SSC characteristics, and no expression of any of the antigens specifically stained in each combination. The purity of the negatively sorted lymphocyte populations were as follows: CD4+CD28+, >98% for CD4 (>96% of these cells being CD28+); CD3+CD8+ T cells, >98% for CD3 (>96% of these cells also being CD8+); CD3+CD5+ T cells, >98% for CD3 (>98% of these cells also being CD5+); and NK cells, >97%. Given that negative FACS sorting of CD19+ B cells from healthy controls only provides low-level purity (>80%), we performed experiments with positively sorted, control CD19+ cells (purity >92%). The purity of CD19+ B cells from a B-CLL patient was always higher than 98%.

Induction of apoptosis in cultured, sorted cell populations
Highly purified, sorted lymphocytes were plated in 96-well, flat-bottomed culture plates at a density of 50,000 cells/well and cultured in triplicate in complete medium for 24 h. Spontaneous apoptosis was studied by culturing T cell subsets from healthy controls and B lymphocytes from B-CLL patients in the absence of any apoptosis inducer. In parallel experiments, phytohemagglutinin (PHA; 2 µg/ml, Difco Laboratories, Detroit, MI) [14 ] or staurosporin (ST; 10–3 M, Sigma Chemical Co., St. Louis, MO) was used to bring about activation-induced cell death. Optimal doses of apoptosis inducers were determined in previous dose/response titrations (data not shown). In all cases, cell cultures were incubated at 37°C in a humidified atmosphere containing 5% CO2. Analysis of the distribution of cultured cells in the G0/G1, S, and G2/M cell-cycle phases was also performed as described previously [22 ]. Tritiated thymidine uptake was also performed to demonstrate the absence of cell proliferation.

Analysis of apoptotic cells
Fresh or cultured lymphocytes were incubated and labeled with mAb (PE/APC) for 20 min at 4°C. The cells were then washed in 2 ml fresh, complete medium by centrifuging for 5 min at 300 g (4°C) and were resuspended in a Ca2+ binding buffer (HEPES 10 mM, NaCl 150 nM, MgCl2 1 mM, CaCl2 1.8 mM, and KCl 5 mM, pH adjusted to 7.4, Sigma Chemical Co.). They were then sequentially incubated with a solution containing annexin V-FITC (Bender MedSystem, Vienna, Austria) in Ca2+ binding buffer (10 min at 4°C) followed by a 3-min incubation with a 7-AAD solution (final concentration of 2.5 µg/ml, Sigma Chemical Co.) in the same buffer to identify early and late apoptotic cells, respectively. The following combinations of mAb (PE/APC) were used: CD5/CD19 for lymphocytes from B-CLL patients and CD28/CD4, CD3/CD8, or CD5/CD3 or CD56/ or CD5/CD19 for lymphocytes from healthy subjects. Control studies comprising unstained cells and cells incubated with isotype-matched, irrelevant PE- and APC-labeled mAb were performed in parallel with each experiment. For these procedures, anti-CD28-PE, anti-CD56-PE, anti-CD3-APC, anti-CD19-APC, and anti-CD4-APC mAb were obtained from Becton Dickinson Biosciences and anti-CD5-PE, anti-CD8-APC, and anti-CD3-PE, from Caltag Laboratories. In all cases, data acquisition and analysis were performed in a FACScalibur flow cytometer (Becton Dickinson Biosciences) using CellQUEST software.

The apoptotic index (AI) for each cell subset analyzed was calculated as the ratio between the percentage of antigen-positive apoptotic cells (annexin V+) and the overall percentage of cells expressing that antigen (annexin V+ plus annexin V– cells).

The percentage of apoptotic cells that had completely lost the expression of the LAgs used for the identification of each cell subset was calculated according to the following equation: %ApoLAgloss = 100 x %ApoLAg/(%ApoLAg–+%ApoLAg+), where %ApoLAgloss is the percentage of apoptotic cells that has lost the expression of LAg, %ApoLAg– represents the percentage of apoptotic cells that has lost their LAgs over the total number of remaining cells, and %ApoLAg+ is the percentage of apoptotic cells that has maintained their LAg over the total number of remaining cells.

Statistical methods
To establish the statistical significance of the differences between the AIs for cell populations with different LAg expressions (strong, weak, or absent), the Wilcoxon matched-pairs signed-ranks test was used, using the Statistical Package for Social Sciences software v. 11.0 (Chicago, IL). Significance was set at P < 0.05.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
LAg expression intensity progressively decreases in apoptotic normal and tumoral lymphocytes
First, we investigated the mean fluorescence intensity (MFI) for several LAgs on highly homogenous lymphocyte populations defined by the expression of a specific LAg, obtained from leukemic B-CLL patients and healthy controls along the different stages of apoptosis. In Figure 1A , we show the different stages of spontaneous apoptosis defined by annexin V/7-AAD-combined staining of a cellular preparation from a B-CLL leukemic patient with a 98% of CD19+ B lymphocytes. As shown in Figure 1B , the intensity of expression of CD19 in viable leukemic B cells (MFIV=131) was greater than that observed in apoptotic cells, independent of apoptotic stage. The mean reduction in the CD19 content per cell, as determined by MFI, was 53%, 89%, and 97% for early, intermediate, and late apoptotic cells, respectively.



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  		Figure 1. The intensity of CD19 expression decreases progressively over the apoptotic cycle in lymphocytes from B-CLL patients. (A) A representative bivariate dot plot of lymphocytes from a B-CLL patient after 24 h of serum-free culture for the definition of different stages of apoptosis by annexin V/7-AAD double-staining. V, Viable cells (annexin V–/7-AAD–); EA, early apoptotic cells (annexin V+/7-AAD–); IA, intermediate apoptotic cells (annexin V+/7-AAD+lo), and LA, late apoptotic cells (annexin V+/7-AAD+hi). (B) Four different fluorescence intensities versus contour histograms corresponding to the four different stages of apoptosis inCD19+ leukemic cells. These include, from top to bottom: V, Viable cells; EA, early apoptotic cells; IA, intermediate apoptotic cells; LA, late apoptotic cells. Arrows indicate the geometric mean of CD19 fluorescence intensity for each histogram.

 
Next, we investigated if this distinctive, decreased expression of a LAg along different stages of lymphocyte apoptosis was not only restricted to neoplasic cells but was also observed in normal lymphocytes. Loss of CD3 and CD5 LAgs was also seen in FACS-purified CD3+CD5+ T cells from healthy controls (Fig. 2A and 2B ). It is interesting that the pattern of loss of both antigens by apoptotic T lymphocytes (Fig. 2A and 2B) depended on the mode of apoptosis induction. In nonstimulated cultures, the expression of CD3 and CD5 was maintained from viable T lymphocytes through to intermediate apoptotic cells and only decreased in late apoptotic cells. After PHA and ST stimulation, the decreased expression of both LAgs was already evident in early apoptotic cells. In these three experimental conditions, the pattern of loss of CD4 expression by CD4+-shorted T cells was similar to that of CD3 and CD5 after stimulation with PHA or ST (Fig. 2C) . In contrast, the other LAgs analyzed, CD8, CD28, and CD56, showed a markedly different, specific pattern of loss: a rapid, initial, and dramatic decrease of expression by early apoptotic cells in sorted, purified CD8+, CD28+, and CD56+ cells, respectively (Fig. 2D 2E 2F) . It should be noted that for CD8, CD4, CD28, and CD56, the pattern of loss of antigen expression was similar for the three different experimental conditions assayed.



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  		Figure 2. Kinetics of loss of different LAgs over apoptosis in purified populations of normal lymphocytes. A dot plot analysis was used to define the four different gates that would enable the identification of the progressive, apoptotic stages in a bivariate dot plot of annexin V versus 7-AAD: viable cells (annexin V–/7-AAD–), early apoptotic cells (EA; annexin V+/7-AAD–), intermediate apoptotic cells (IA; annexin V+/7-AAD+lo), and late apoptotic cells (LA; annexin V+/7-AAD+hi). (A–F) MFI of the LAgs in each apoptotic stage from different sorted cell populations, (A) CD3+, (B) CD5+, (C) CD4+, (D) CD8+ (E) CD28+, and (F) CD56+. Sorted cell populations were cultured 24 h with no stimulation (•), PHA ({triangleup}), or ST ({square}). The coefficients of variation of MFI between replicated samples of all fractions of the antigens studied in all the conditions assayed were always lower than 10%.

 
We further analyzed the percentage decrease in antigen expression between viable and early apoptotic cells from sorted lymphocyte populations cultured with or without PHA and ST (Fig. 3A ). The findings demonstrate that the loss of expression of several LAgs occurs early in apoptosis. Two different patterns of loss of antigen expression can be identified: In unstimulated lymphocytes, the CD5 and CD3 antigens maintained their levels during the early stages of apoptosis, and they were lost by early apoptotic cells in the presence of PHA or ST, and the CD8, CD4, CD28, CD56, and CD19 antigens showed a rapid decrease in their expression by early apoptotic cells, independent of the presence of PHA or ST during culture. Such kinetic differences were particularly clear once the changes in the expression of LAgs were evaluated for different pairs of antigens coexpressed by the same cell (Fig. 3B 3C 3D) .



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  		Figure 3. Patterns of loss of paired antigens coexpressed in the same cells during spontaneous in vitro apoptosis. (A) The percentage decrease in MFI of antigens expressed between the viable and early apoptotic cell stages in different sorted cell populations cultured in the medium alone (open bars), with PHA (hatched bars), or with ST (cross-hatched bars). The different apoptosis stages were determined in a bivariate annexin V versus 7-AAD dot plot. Their corresponding MFI values are represented by symbols: viable cells ({circ}), early apoptotic cells ({triangleup}), intermediate apoptotic cells ({square}), and late apoptotic cells ({diamond}). The mean values of expression of pairs of antigens in the same cell were obtained and are shown in (B) CD4 versus CD28, (C) CD3 versus CD8, and (D) CD3 versus CD5. (E and F) The covariation of the MFI of two LAgs coexpressed in the same cells against the FSC signal over the different stages of apoptosis. (E) CD3+CD8+ negatively sorted cell population: CD3 LAg (continuous line) and CD8 LAg (dotted line). (F) CD4+CD28+ negatively sorted cell population: CD4 LAg (continuous line) and CD28 LAg (dotted line). (A) The coefficients of variation of the percentages between replicated samples of all fractions of the antigens studied, in all conditions, were always lower than 5%. (B–F) The coefficients of variation of mean intensity between replicated samples of all fractions of the antigens, studied in all conditions, were always lower than 10%.

 
A potential mechanism involved in this loss of LAgs by apoptotic lymphocytes could be related to the concomitant cell fragmentation. If this is so, decreased antigen expression should be associated with a loss of cell size, as evaluated by the FSC of light. As shown in Figure 3E and 3F , apoptotic cells showed progressively less FSC as they advanced through the different stages of apoptosis. However, the kinetics of loss differed for different LAgs. For example, CD3+CD8+ T lymphocytes did not suffer loss of CD3 LAg MFI until late apoptosis but lost most of their CD8 LAg in early apoptotic cells. The CD4+CD28+ T cells progressively lost their CD4 LAg but lost most of their CD28 antigen in early apoptotic cells.

Different apoptotic indices are observed in cell fractions defined by the different degree of expression of LAgs
Purified lymphocyte subpopulations obtained by FACS were cultured in different experimental conditions to study the association between LAgs and spontaneous and induced apoptosis. After 24 h of culture and based on the degree of intensity of LAg expression, cells were classified into three different fractions: those with bright MFI signals (strong LAg expression), those with dim signals (weak LAg expression), and those with an undetectable signal (no LAg expression; Fig. 4A ). The AI obtained for each cell fraction is shown in Figure 4B 4C 4D . Under all the experimental conditions assayed (spontaneous, PHA-induced, or ST-induced apoptosis) and for all the studied LAgs (CD5, CD3, CD8, CD4, CD28, CD56, CD19), the cell fraction with high LAg expression showed a lower AI than did the fractions with partial loss of their LAgs (dim MFI signal) or total loss (no MFI signal; no major differences were observed between the two latter fractions).



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  		Figure 4. Apoptotic indices of cell fractions defined by the degree of expression of their LAg. Representative histogram analyses of CD4 in the negatively sorted CD3+CD4+ population before starting the culture (A, upper) and after 24 h of culture in the presence of ST (A, lower). The intensity of expression of the CD4 LAg was divided into three fractions defined by degree of expression: M1 (Bright) represents the CD3+CD4+ cells with strong expression of CD4, M2 (Dim) represents the CD3+CD4+ cells with weak expression of CD4, and M3 (negative) represents the CD3+CD4+ cells lacking expression of CD4. (B–D) The AIs of the three fractions of the different populations studied in which apoptosis was induced by the medium alone, by PHA, and by ST, respectively, are shown. The three bars represent the AI of the three different cell fractions defined: strong expression (open bars), with weak expression (shaded bars), and undetectable expression (solid bars). The coefficients of variation of the proportion of apoptotic cells between replicated samples of all fractions of the antigens studied, in all conditions, were always lower than 5%.

 
The difference in the occurrence of spontaneous apoptosis between the cell populations that completely lost their LAg expression and those that maintained strong LAg expression was dramatically large. The increase in the proportion of apoptotic cells between weak LAg-expressing cells and strong LAg-expressing cells in the CD5+ lymphocytes was 1.4-fold; in the CD56+ cells, this was 1.7-fold, and in the CD3+ lymphocytes, this was 1.9-fold. The increase was even greater in the CD4+ (2.2-fold), CD19+ (2.5-fold), CD28+ (4.3-fold), and CD8+ (9.7-fold) lymphocytes.

Evidence of apoptotic cells with undetectable LAg expression
For all the antigens analyzed, except for CD19, and spontaneously or in the presence of PHA, more than 20% of apoptotic cells completely lost the expression of their LAgs to undetectable levels (Table 1 ). Similar findings were observed in the presence of ST for the indicated LAgs except for CD3 and CD4. NK cells were those with the highest proportion of LAg-negative cells in the apoptotic populations.


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  		Table 1. Percentage of Apoptotic Cells That Completely Lost Detectable Expression of Their LAg

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The results show that the loss of expression of LAgs by apoptotic cells is a common feature of lymphocytes undergoing apoptosis. Different kinetic patterns of LAg loss over the stages of apoptosis were observed. Moreover, in all the conditions assayed, an important fraction of apoptotic lymphocytes saw their expression of LAgs reduced to undetectable levels. With current cytometric methods, such loss of expression could seriously affect any measurement of the proportion of LAg-defined cells undergoing apoptosis.

Previous studies suggest that apoptosis preferentially occurs in cells from subsets with weak expression of LAgs such as CD56 or CD94 rather than in cells expressing the same antigens strongly [15 , 16 ]. Our findings ought to re-evaluate this suggested, special apoptosis susceptibility of lymphocytes with weak LAg expression. Our results show a generalized apoptotic lymphocyte behavior characterized by a decrease in the expression of specific LAgs by apoptotic cell, and consequently, the percentage of apoptotic lymphocytes increases as the intensity of expression of each LAg decreases. To elucidate the exact nature of the relationship between apoptosis and weak LAg expression, we specifically cultured negatively sorted homogeneous cell populations with strong expression of different LAgs. We observed the reduction in LAg expression as lymphocyte apoptosis progresses with a final stage of undetectable LAg levels. As this study shows that the apoptotic pathway itself is associated with reductions in the intensity of antigen expression, any analysis of apoptosis that defines cell subsets by the intensity of specific LAg expression should be interpreted with caution. Along these lines, once cells within a population begin to undergo apoptosis, if cells with strong LAg expression are selected, then, viable cells are being preferentially chosen. The opposite is true for weak LAg-expressing cells. Such arbitrary selection may lead to erroneous evaluation of the occurrence of apoptosis in specific subsets of lymphocytes, especially in those that rapidly lose expression of one or more antigens (as found for CD8 and CD28 antigens in CD3+CD8+ and CD4+CD28+ T lymphocytes, respectively).

Earlier studies have suggested in unpurified, heterogeneous cell preparations that the expression of some antigens (e.g., CD4, CD8, CD45RA, CD56) but not others (e.g., CD20) decreases in apoptotic lymphocytes without specific analysis of the different phases of this dynamic cellular process [17 , 18 ]. Our experimental approach has allowed demonstrating reductions of LAg expression to a final stage of undetectable levels and the loss of LAg expression in early stages of apoptosis. These apoptotic lymphocytes, which appear as LAg-negative, are usually ignored during evaluations of the frequency of apoptotic cells in phenotypically defined cell subsets.

An important issue is whether loss of LAg expression by apoptotic cells is a specific, active process associated with the transition of cells in the different stages of apoptosis or whether it simply reflects a general degradation of cell components. The observation that different markers have distinct loss of expression kinetics might suggest the former may be the case. This is further supported by the fact that antigens coexpressed in the same cells showed different degrees of loss in the different stages of apoptosis. Moreover, the kinetics of this loss was different in viable and early apoptotic cells. Overall, the existence of different rates of antigen loss could indicate that apoptotic cells have different ways of regulating the expression of different antigens.

A potential explanation for the decrease in the expression of surface LAgs is the partial loss of the cell membrane by early apoptotic cells. However, it should be noted that despite CD3+ and CD5+ cells showing decreased FSC signals in the early stages of apoptosis, the expression of both antigens was almost stable during this period. In addition, the patterns of loss of LAgs coexpressed in the same cell over the different stages of apoptosis were clearly different. A recent report has described that apoptosis induces early and active mRNA degradation [23 ]. mRNAs with different half-lives show similar apoptosis-induced degradation kinetics, indicating that this degradation is an active process induced by apoptosis. This suggests that a lack of LAg synthesis could be involved in the loss of LAg by apoptotic cells. Further studies are required regarding the physiological significance of this process.


   ACKNOWLEDGEMENTS
 
This work was partially supported by Grants 98-1431, 00-0806, and 03/1582 from the Fondo de Investigación de la Seguridad Social (FIS; Ministerio de Sanidad y Consumo, Madrid, Spain) and by Grants SAF-99-0099 and GEN 2001-9856-CB-13 (Ministerio de Educación y Ciencia, Madrid, Spain). D. D. and A. P. contributed equally to this study and are joint first authors. This work received the Outstanding Poster Award in the XXII International Congress of the International Society for Analytical Cytology.

Received March 18, 2004; accepted March 29, 2004.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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