|
|||||||||
|
|
|
|
|
|
|
|
|
|
|
IMMUNOBIOLOGY
From the Laboratory of Molecular Immunology, Institute
for Molecular and Cell Biology, University of Porto; the Department of
Hematology, Santo António General Hospital, Porto, Portugal; and
the Laboratory of Food and Biodynamics, Nagoya University Graduate
School of BioAgricultural Sciences, Japan.
Red blood cells (RBCs) are known to perform one prominent function:
to carry and deliver oxygen to the tissues. Earlier studies, however,
suggested a role for RBCs in potentiating T-cell proliferation in
vitro. Here it is shown that the presence of RBCs in cultures of
stimulated human peripheral blood lymphocytes strengthens T-cell proliferation and survival. Analysis of phosphatidylserine
externalization and DNA fragmentation showed that RBCs inhibit T-cell
apoptosis. This inhibition correlated with a reduction in CD71 but not
CD95 expression. RBCs enhanced T-cell proliferation and survival upon activation with phytohemagglutinin and with OKT3 antibodies. Studies aimed at characterizing the cellular and molecular basis of the protection afforded to T cells by RBCs showed that (1) optimal protection required intact RBCs and red cell/T-cell contact but not
monocytes; (2) RBCs markedly reduced the level of intracellular reactive oxygen species; and (3) RBCs inhibited the formation of protein-bound acrolein, a peroxidation adduct in biologic systems. Overall, these data indicate that human RBCs protect T cells from activation-induced cell death, at least in part by reducing the pro-oxidant state, and suggest a role for RBCs as conceivable modulators of T-cell homeostasis.
(Blood. 2001;97:3152-3160) Programmed cell death, or apoptosis, is a
physiological process that contributes to the homeostasis of
multicellular organisms. Apoptosis resulting from activation of T
cells Despite the important role played by the death receptors, growing
evidence indicates that environmental constituents, such as nonlymphoid
secreted factors and cytokines, can modulate apoptosis of activated T
cells, thus emphasizing the significance of the environment in the
maintenance of T-cell homeostasis.10-12 Any imbalance in
the apoptotic process may result in either of 2 possible scenarios:
lymphocyte accumulation or lymphocyte depletion. In the former,
lymphoproliferative disorders Other factors implicated in different forms of apoptosis are
reactive oxygen species (ROS). Several studies have provided evidence
for the involvement of ROS in apoptosis of T-cell blasts and hybridomas
by using antioxidants such as N-acetyl cysteine and
glutathione.19,20 Apoptosis in these cells is the
consequence of changes in mitochondrial permeability and the subsequent
release of ROS.21 Studies performed with primary T cells,
however, indicate that the formation of intracellular ROS is necessary
for T-cell activation and IL-2 secretion but also regulates
activation-induced T-cell apoptosis, therefore suggesting that
intracellular ROS could play a role in peripheral T-cell
homeostasis.22-24 Nevertheless, studies with primary T
cells are usually performed in cultures lacking other "nonlymphoid"
cells, albeit activation-induced T-cell apoptosis is thought to occur
in organs and tissues where other cell types, such as red blood cells
(RBCs), are present (for review, see 25). In addition to
their main function in oxygen and CO2 transport,26 RBCs also have the capacity to scavenge
exogenous ROS because of the permeability of their membranes to oxygen
radicals and the presence of high levels of intracellular antioxidant
enzymes27-29 (for review, see 30). In addition,
RBCs are also known to increase T lymphocyte proliferation upon
stimulation in vitro.31-33 These features confer RBCs the
potential to function as ROS scavengers produced within their milieu,
thereby protecting neighboring cells from ROS-mediated
damage.34-37 Whether the potentiating effect of RBCs on
T-cell proliferation is related to their capacity to scavenge ROS is
unknown. The main goal of the present study was to perform a
comprehensive study to examine the effect of RBCs on T lymphocytes
after activation. Results showed that RBCs enhance T-cell expansion and
survival by inhibiting activation-induced T-cell death, a finding
associated with a decrease in oxidative stress within the activated T cells.
Reagents and monoclonal antibodies
Cell preparation and culture
PBLs were cultured at a final concentration of 0.25 × 106 cells/mL either in 96- or 6-well plates (TPP, Trasadingen, Switzerland) in a final volume of 0.2 mL or 5 mL CM, respectively, giving a density of 1650 cells/mm2 in either culture plate. PBLs were either left unstimulated or stimulated with 5 µg/mL PHA-P and cultured in the absence or presence of autologous RBCs in an incubator at 37°C, 5% CO2, and 95% humidity for 1 to 5 days. In some experiments PBLs were stimulated with 5 µg/mL PHA-L and with 0.5 µg/mL OKT3. For proliferation studies, 5 × 104 cells were cultured in triplicate in 96-well flat-bottom microplates. For other studies 1.5 × 106 cells were cultured in 6-well macroplates. RBC suspensions were prepared in CM and adjusted to give an RBC:PBL ratio of 100:1 after addition to the PBL cultures. In some experiments, RBCs were lysed by brief sonication before they were added to the PBL cultures. Sonic cell disruption was performed by 10 bursts of 1 second each at 0.1 W power output. In experiments with cell culture inserts (pore size, 0.2 µm; Nunc, Roskilde, Denmark), PBLs were placed in the lower chamber and RBCs in the upper chamber. Red cell lysis procedure For the determination of parameters that required the use of flow cytometry (eg, T-cell activation or death, intracellular ROS
production)only activated T cells that received RBCs were treated
with lysis solution (10 mM Tris, 150 mM NH4Cl, pH 7.4) to
remove RBCs. This treatment did not significantly alter the activation
parameters studied when compared with nontreated activated T cells. In
some experiments, however, flow cytometry determinations (eg, Annexin V
and ROS) were performed without the removal of RBCs.
Determination of T-cell activation and proliferation T-cell activation and proliferation was studied by 3 methods: (1) thymidine uptake; (2) determination of cell size; and (3) examination of activation markers. For thymidine uptake, 0.5 µCi [3H]-TdR (specific activity, 5.0 Ci/mmol; Amersham-Pharmacia Biotech, United Kingdom) was added 4 hours before the end of the culture, and cells were harvested on glass fiber filters (Filter MAT; Skatron Instruments, Suffolk, United Kingdom), using a semi-automatic cell harvester (Skatron, Norway). The incorporated [3H]-TdR was measured in a Beckman liquid scintillation counter, and results were expressed as counts per minute (cpm). For determination of cell size and activation, forward and side scatter characteristics (FSC/SSC) and expression of CD69, CD25, CD71, CD95, and HLA-DR, respectively, were analyzed by flow cytometry. Proliferation index was defined as follows: cpm in the presence of RBC/cpm in the absence of RBC.Detection of lymphocyte apoptosis and death Apoptosis was studied by characterizing phosphatidylserine expression and DNA fragmentation. T cells were harvested 1 to 5 days after activation, washed with PBS, and counted. For phosphatidylserine determination, cells were washed twice with binding buffer (10 mM HEPES, 140 mM NaCl, and 2.5 mM CaCl2, pH 7.4) and incubated with Annexin V-FITC for 15 minutes at room temperature. Cells were immediately harvested and analyzed by flow cytometry. For DNA fragmentation, cells were lysed by overnight incubation at 37°C in lysis buffer (100 mM Tris, pH 8, 5 mM EDTA, 200 mM NaCl, 0.2% SDS, 0.2 mg/mL proteinase K). After centrifugation at 13 000g for 15 minutes, the supernatant containing fragmented DNA was transferred to another tube and precipitated with an equal volume of 100% ethanol. The pellet was washed twice with cold 70% ethanol and then dissolved in 20 µL Tris-EDTA buffer containing 0.2 mg/mL RNAse A. After incubation at 37°C for 2 hours, DNA was resolved on 2% agarose gels and visualized with ethidium bromide staining.T-cell death was determined by PI and trypan blue (TB) staining (PI and TB are 2 dyes that enter cells with damaged plasma membranes) and by a decrease in cell size (shrinking). For PI staining, 2 µL/sample PI (25 µg/mL) was added to each cell sample before flow cytometry analysis. For TP staining, aliquots of activated lymphocytes were resuspended in PBS-containing trypan blue. Dead and alive cells were counted in a Neubauer chamber under a light microscope. For cell shrinking, FSC/SSC characteristics of activated lymphocytes were analyzed by flow cytometry. Measurement of oxidative stress Oxidative stress in T cells induced by PHA was measured by the detection of ROS and the detection of protein-bound acrolein. ROS produced within activated T cells were detected with the membrane-permeant probe 2',7'-dichlorofluorescein-diacetate (DCFH-DA). The probe freely enters the cell and is incorporated into hydrophobic regions, and the acetate moiety is cleaved off by cellular esterases leaving a nonfluorescent and impermeant form of DCFH.40,41 ROS produced by the cell oxidize DCFH to DCF, which, after excitation at 488 nm, emits fluorescence at 530 nm (FL1 channel). Resting PBLs were incubated with 100 µM DCFH-DA in CM for 30 minutes at 37°C and were washed 3 times with the same media. Then PBLs were left unstimulated or were stimulated with 5 µg/mL PHA-P. At time intervals after activation, cells were harvested, washed with PBS, and directly analyzed by flow cytometry. Protein-bound acrolein was also detected by flow cytometry using the mouse mAb 5F6 (see 38 for a detailed description of the mAb), followed by the proper fluorochrome-conjugated rabbit antimouse immunoglobulins.Flow cytometry analysis Cells were stained on day 0 (resting cells) and at different time points after mitogenic stimulation (T-cell blasts), as described previously.42 Briefly, cells were harvested from cultures by gentle pipetting and washed twice with PBS. In cultures with RBCs, erythrocytes were first lysed in lysis solution as indicated above. Staining was performed at 4°C for 30 minutes in staining solution (PBS, 0.2% BSA, 0.1% NaN3) in 96-well round-bottom plates (Greiner, Nürtingen, Germany) with approximately 0.5 × 106 cells/well. Cells were stained in one (direct), 2 (indirect), or 3 steps depending on the combination of mAbs and the antigens studied. Irrelevant mouse mAbs were used as negative controls to define background staining. Rabbit antimouse FITC or R-phycoerythrin (RPE)-conjugated F(ab')2 fragments were used as second-step antibodies. After they were stained, cells were washed 3 times with staining buffer and were immediately acquired without fixation in a FACSort (Becton Dickinson, Mountain View, CA) equipped with an argon laser that emitted at 488 nm. For each sample 10 000 cells were usually acquired using FSC/SSC characteristics and analyzed using the Lysys II software.Western blot analysis Immunodetection was performed as previously described.43 Briefly, resting and activated T cells were solubilized in 1% Triton X-100 lysis buffer, and cell debris was removed by centrifugation at 11 000g. Cell lysates were boiled for 5 minutes in 2× SDS buffer and resolved by 12% SDS-PAGE. Proteins were transferred to nitrocellulose membranes (Hybond C-super, Amersham), and filters were blocked with TBS-T containing 5% (wt/vol) nonfat dry milk for 1 hour. After washing with TBS-T, filters were incubated for 1 hour with a 1:1000 dilution of an mAb against the transferrin receptor (clone H68.4). Filters were washed and incubated with horseradish peroxidase-conjugated goat-antimouse antibodies (Transduction Laboratories, Lexington, United Kingdom) for 1 additional hour. All incubations were performed at room temperature. Finally, membranes were extensively washed, and detection was accomplished using enhanced chemiluminescence (Amersham) and exposure to Biomax MR-1 Kodak films (Sigma-Aldrich).Statistical analysis The Student t test was used to test the significance of the differences between group means. Statistical significance was defined as P < .05. Kolgomorov-Smirnov statistics were used for analysis of the statistical significance of differences seen between histograms.44,45 Two histograms were considered significantly different when D/s(n) was greater than 15, as described.46
Activation-induced T-cell death in normal human PBLs Activation of resting human PBLs with the T-cell mitogen PHA-P increased thymidine uptake (141 ± 105 in resting vs 4606 ± 1914 in activated; mean cpm ± 1 SD; P < .0005), a measure of cell division. Morphologically, however, PHA-P stimulation was characterized by the appearance of 2 populations of activated T cells with an overall increase in cell size when compared to resting lymphocytes (Figure 1, gates R1 and R2). Blasts within gate R1 were usually more than 95% CD3+ and PI . On the contrary, the characteristics of the cells
within gate R2 clearly show that they were undergoing cell death. They
had a reduced size (cell shrinking), showed reduced CD3 expression, and
a high percentage (more than 90%) were PI+, all features
of T cells that have lost plasma membrane integrity. Activation-induced
T-cell death was observed in all PBL samples studied and started as
early as 1 hour after mitogenic stimulation (data not shown). The data
illustrate the T-cell specificity of the mitogen used and that AICD
takes place in primary cultures of human peripheral blood T cells.
Characterization of early (eg, CD69) and late (eg, HLA-DR) T-cell
activation markers after activation by flow cytometry further
demonstrated that, under the culture conditions used, human PBLs
responded optimally to PHA-P stimulation (see below).
RBCs enhance T-cell proliferation by inhibiting activation-induced T-cell death To examine the effect of RBCs on PHA-P-driven T-cell proliferation and survival, PBL cultures were stimulated with or without the presence of RBCs for 5 days, as indicated in "Materials and methods." As illustrated in Figure 2, T cells proliferated approximately 5 times better when cocultured with RBCs. This enhanced T-cell proliferation was observed at low (1% FCS) and high (10% FCS) serum concentrations (data not shown). In the absence of the mitogenic stimulus, RBCs were unable to drive T cells into cell division. The significantly enhanced proliferation observed in cultures with RBCs (P < .05) was also observed with 2 other mitogens, PHA-L (ie, leuco-agglutinin) and OKT3 antibodies (Figure 2). Flow cytometry analysis showed that the percentage of T-cell blasts undergoing activation-induced cell death was significantly decreased by the presence of RBCs (59.7 ± 6.9 vs 30.4 ± 8.7; P < .002; n = 7). An illustration of the inhibition of AICD in the presence of RBCs is shown in Figure 3A and is further corroborated by the impact that an enriched RBC environment had on total cell recovery. Thus, T-cell activation of human PBLs in an enriched RBC environment resulted in a 5-fold increase in viable cell recovery 5 days after activation when compared with cultures of PBL alone (Figure 3B, P < .01).
Enriched RBC environment reduces T-cell apoptosis and CD71 expression The above data showed that the presence of RBCs inhibited the death of activated human T cells. To ascertain whether the decreased T-cell death was associated with changes in the expression of early markers of T-cell apoptosis, phosphatidylserine externalization was studied by flow cytometry. Kinetic experiments showed that the presence of RBCs reduced the percentage of Annexin V+ "apoptotic" T-cell blasts along the culture period after mitogenic activation (Figure 4). Despite interindividual variation in Annexin V labeling, RBCs consistently reduced phosphatidylserine externalization. The protection of T-cell apoptosis by RBCs was substantiated by results of DNA fragmentation (data not shown).
Considering that high expression of CD95 has been associated with
accelerated lymphocyte apoptosis and that triggering of CD95 on
activated T cells and T-cell lines leads to apoptosis, we tested the
possibility that the protection provided to T lymphocytes by RBCs was
associated with a decrease in CD95 expression. However, the expression
of CD95 in activated T cells cultured either in the absence or the
presence of RBCs was not significantly different. In contrast, the
presence of RBCs markedly reduced the expression of CD71, the
transferrin receptor (Figure 5). Thus, a
2-fold reduction in the mean fluorescence intensity of CD71 in
activated T cells was observed in cultures of PBLs stimulated in the
presence of RBCs when compared with cultures of PBLs alone
(40.9 ± 12.3 vs 96.7 ± 20.3, respectively;
P < .001; n = 5). Kinetic experiments showed that the
RBC-mediated decrease in cell surface CD71 expression takes place along
the entire culture period. CD71 down-modulation in the presence of RBCs
was confirmed by Western blot analysis (data not shown). Analysis of
other T-cell activation markers showed that RBCs slightly enhanced the
expression of CD25 and HLA-DR but not of the early activation marker
CD69 (Figure 5).
Inhibition of T-cell apoptosis by RBCs correlates with a reduction in oxidative stress ROS have been implicated in the apoptotic process in a number of systems, including activated T lymphocytes, and previous studies have shown that T cells produce intracellular ROS after mitogenic stimulation.22-24 In our hands, human peripheral blood CD3+ T cells activated in vitro with PHA also produced intracellular ROS, as demonstrated by a marked increase in DCF fluorescence (data not shown). Importantly, the presence of RBCs significantly reduced ROS levels. As illustrated in Figure 6, RBCs decreased by approximately 4-fold the level of intracellular ROS among activated CD3+ T cells (D/s(n) = 39.07; D = 0.83). This decrease was observed regardless of the use of lysis solution to remove RBCs before analysis. However, in cultures not treated with the lysis solution, the high number of RBCs present interfered with the detection of activated CD3+ T cells by the flow cytometer (Figure 6 and data not shown). In 7 separate experiments and 16 different determinations performed within the first 24 hours after stimulation, the presence of RBCs decreased DCF mean fluorescence intensity of PHA-activated T cells on average by 3.5-fold (range, 1.4-9.7; P = .0013). Kinetic experiments demonstrated that intracellular ROS are detectable as soon as 30 minutes after mitogenic activation of PBLs with PHA and that RBCs are capable of counteracting ROS production (data not shown).
Next, we examined whether intracellular ROS production by activated T
cells led to the expression of protein-bound acrolein, a marker of
oxidative stress. Acrolein was indeed detected in activated, but not in
resting, PBLs. We observed maximal expression of acrolein adducts by
activated T cells 3 days after mitogenic activation (Figure
7A). The presence of RBCs decreased both
the percentage of acrolein-positive cells and the amount of reactive acrolein adducts, as indicated by a marked (approximately 3-fold) decrease in the mean fluorescence intensity (Figure 7A). These results
suggested that acrolein expression among activated T cells was a late
event in T-cell apoptosis. To clarify this possibility, double labeling
(ie, Annexin V vs acrolein) kinetic experiments were performed. As
illustrated in Figure 7B, cells expressing cell surface
phosphatidylserine were observed as soon as 1 hour after T-cell
activation, and they constituted more than half the activated T cells
for 24 hours, when this percentage dropped drastically. In marked
contrast, the percentage of Annexin V+ "apoptotic" cells expressing
cell surface acrolein adducts was very low during the first 4 hours
after activation. Cell surface acrolein expression started to increase
24 hours after activation and was maximal by day 3 (see Figure 7A).
RBCs markedly reduced the percentage of both Annexin V and
acrolein-positive cells after T-cell activation at all time points
studied. Characterization of acrolein adducts among activated T cells
undergoing T-cell death (ie, PI+) showed that acrolein
expression during the early phases of T-cell activation is mainly
intracellular (data not shown).
Enhanced proliferation induced by RBCs requires T-cell contact and intact RBCs The latter results suggested that the enhancement of T-cell proliferation and protection from apoptosis by RBCs could be due, at least in part, to the ROS-scavenging properties of RBCs. Thus, we first investigated whether the effect of RBCs on T-cell expansion and survival required cell contact by using cell culture inserts. As illustrated in Figure 8, RBCs had their highest enhancing effect on T-cell proliferation and survival when they were cultured with the activated PBLs. When activated PBLs and RBCs were cultured separately using cell inserts, T-cell proliferation was reduced by 50%. In accordance with the proliferation results, AICD in cultures with cell inserts was also decreased (data not shown). Next, we examined whether intact red blood cells were required. Proliferation in the presence of red cell lysates was reduced by 30% when compared to intact RBCs (Figure 8). Finally, the addition of purified catalase, an abundant erythrocyte antioxidant enzyme, to stimulated cultures of PBLs did not reproduce the marked enhancing effect of RBCs, either in T-cell proliferation or in survival (Figure 8 and data not shown).
Monocytes are not involved in the enhancing effect of RBCs in T-cell proliferation To ascertain whether the RBC-enhancing effect on T-cell proliferation and survival was dependent on the presence of accessory cells such as monocytes, experiments with highly enriched T-cell preparations (more than 95% CD3+, less than 2% CD14+) were performed. As shown in Figure 9, the depletion of monocytes resulted in a very low proliferative response by T cells to mitogen stimulation. However, the proliferative response by purified T cells was restored by the addition of increasing amounts of monocyte-enriched populations (more than 70% CD14+). Adding RBCs not only increased T-cell proliferation in cultures of PBLs, as expected, but also in cultures of purified T cells (Figure 8). Interestingly, adding RBCs to the purified T cells resulted in a significantly higher proliferation index than that seen with PBLs (26.9% ± 5.8% vs 15% ± 3.1%; P < .01; n = 3 ). Adding monocytes to the activated T-cell/RBC cultures had no further effects on T-cell proliferation.
The maintenance of accurate peripheral T-lymphocyte homeostasis is
crucial to a patient's health, and a balance between T-cell death and
survival is central to the homeostatic process.47 Most of
the research performed on T-cell apoptosis has focused on the
characterization of cytoplasmic, secreted, and membrane-anchored molecules. Less attention has been paid to the role that nonlymphoid cells play in this process. This is a relevant point if one takes into
consideration that T-cell death or survival in the periphery takes
place in organs and tissues populated by different cell types,25,48 some of which The main finding of this study was that RBCs inhibit the T-cell
apoptotic process started after mitogenic activation of resting human
peripheral blood T cells. RBCs reduced exposure of phosphatidylserine on the outer leaflet of the plasma membrane of T lymphocytes, a
hallmark of apoptosis. Several studies have shown that intracellular oxidative stress, as a result of activation, also initiates apoptotic processes in primary T cells.22-24,53 More recently, it
has been proposed that selective oxidation of phosphatidylserine during oxidative stress promotes its externalization.54 By using
dichlorofluorescein, an indicator of generalized oxidative stress, we
have shown that human peripheral blood T cells activated in vitro
produced intracellular ROS. It is likely that intracellular ROS
reduction in activated T cells was in part due to the removal of ROS by
RBCs. Numerous studies have shown that RBCs can scavenge radicals such
as H2O2 and ONOO The mechanism by which RBCs counteract intracellular ROS production in T cells, and hence enhance T-cell proliferation, could depend on the high antioxidant activity of the RBCs.30 Yet, exogenous added catalase (10 µg/mL), though increasing T-cell proliferation by approximately 2-fold, did not completely reproduce the proliferation levels observed with RBCs. Previous studies with human PBLs have also shown that catalase had no significant influence, either in the proliferative response of T cells to PHA or in the inhibition of monocyte-dependent T-cell death.55,56 These findings are in accordance with the reported inability of catalase to cross the lymphocyte membrane and to inhibit the intracellular production of ROS.23,57 Optimal T-cell proliferation and survival was only observed with intact RBCs and when RBCs were in close contact or proximity with the activated T cells. The need for proximity and for RBC integrity might imply the involvement of interactions between receptors present on each cell type, as suggested by some authors.33,58 The overall results, however, do not support that scenario. First, though reduced by 50%, the proliferative responses with RBCs in cell inserts were still much higher than with PHA alone. Second, red cell lysates were almost as efficient as intact RBCs in enhancing T-cell proliferation. Third, preliminary results indicate that RBCs are not required during the early phases of T-cell activation to fully exert their protective effect (Fonseca et al, unpublished data). Among the T-cell receptors studied, RBCs significantly modulated only the expression of CD71, the transferrin receptor. Thus, the presence of RBCs in cultures of stimulated PBLs resulted in a decrease in the level of expression of CD71. The transferrin receptor is considered a marker of T-cell activation that is expressed by activated T lymphocytes to import extracellular iron for metabolic needs (reviewed in 59). However, earlier studies demonstrated that the expression of CD71 in mitogen-activated human peripheral blood T cells is mainly regulated by the intracellular iron level rather than the rate of proliferation.60 It is now well established that CD71 expression is regulated at the post-transcriptional level not only by iron availability but also by extracellular and intracellular oxidative stress induced by reactive oxygen species such as H2O2 and NO.61,62 Therefore, CD71 down-regulation in activated T cells cultured in the presence of RBCs may reflect a reduction of the pro-oxidant state generated after T-cell activation, an increase in the intracellular iron pool, or a combination of both.60-62 In this context, the possibility that the enhancing effect on T-cell proliferation and survival is due to the costimulatory effect of RBC components, such as heme and iron, released during the culture period cannot be ruled out. Iron compounds are costimulatory agents of T-cell proliferation and induce a decrease in CD71 expression.50,60,63,64 Although we do not have evidence that hemolysis takes place in the cultures, some authors have pointed out the possibility of a T-cell-dependent RBC elimination.65 Given the existence of a close interplay between the intracellular iron pool and sensitivity to oxidative stress,30,66 the intracellular iron status of the resting human PBLs before stimulation could also have influenced the final outcome of the RBC effect. Finally, we have demonstrated that human T cells activated in vitro generated protein-bound acrolein. Acrolein is a lipid-derived product that functions as a marker of oxidative stress and that has recently been found in several human diseases such as diabetic nephropathy and Alzheimer disease.39,67 A recent report showed that acrolein inhibits glucose and glutamate uptake in primary neuronal cultures.68 Whether acrolein also inhibits glucose uptake by activated T cells is presently unknown. Acrolein adducts were only significantly detected 24 hours after stimulation and were maximal by day 3. This suggests that acrolein formation in activated T cells is a late event during the apoptotic process. The presence of RBCs reduced the expression of acrolein adducts among activated T cells, therefore reinforcing the view that RBCs play an important role in reducing oxidative stress and apoptosis in activated T cells. Overall, these results are in agreement with a recent study showing that RBCs inhibit apoptosis of human neutrophils.69 They add to the growing evidence indicating that hemoglobin-containing RBCs may perform unforeseen functions in addition to O2 and CO2 transport. For example, RBCs can regulate vascular vasodilatation by reacting with endothelial NO, a process regulated by intravascular flow.70 In a series of elegant studies, Jain et al71,72 have demonstrated that RBCs are crucial elements that facilitate the engagement of circulating lymphocytes with the vascular endothelium. It is well known that during inflammation and local immune response, capillary diameter and blood flow increase.73 This may allow the extravasation of RBCs, together with activated T cells, to inflammatory places, which would favor T-cell survival and enhance T-cell effector functions. Moreover, the reported increase in natural killer cytotoxicity against tumors after interaction with intact RBCs74 adds further support to the data presented here. In summary, we have presented evidence that red blood cells can function as modulators of T-cell apoptosis in vitro, a finding associated with a reduction in intracellular oxidative stress. Whether protection from T-cell apoptosis also takes place, or is hampered, in vivo if erythrocyte anomalies exist remains the topic of future studies. In our view, the correlation found between anomalies in erythrocyte antioxidant defense and abnormalities in T-cell phenotype and function in certain chronic disorders, such as thalassemia,75-77 emphasizes the potential that mature red blood cells may have in regulating T-cell death or survival in vivo.
We thank the personnel of the Blood Bank of the Santo António General Hospital for their help in collecting blood samples. We thank Dr Alexandre M. Carmo for helpful comments during the performance of this work, Dr Rui Appelberg for reviewing the manuscript, and Dr Angelo Cardoso for valuable help in providing antibodies used in this study and for critical reading of the manuscript. We also thank Prof Maria de Sousa for continuous support. We thank the University of Porto for supporting the publication costs of this manuscript.
Submitted August 14, 2000; accepted January 17, 2001.
This work is part of the PhD thesis of A.M.F., who is the recipient of a Praxis XXI fellowship from Fundaçâo para a Ciêucia e a Tecnologia (BD 18503/98).
The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked "advertisement" in accordance with 18 U.S.C. section 1734.
Reprints: Fernando A. Arosa, Laboratory of Molecular Immunology, Institute for Molecular and Cell Biology, Rua do Campo Alegre, 823, 4150 Porto, Portugal; e-mail:farosa{at}ibmc.up.pt.
1.
Green DR, Ware CF.
Fas-ligand: privilege and peril.
Proc Natl Acad Sci U S A.
1997;94:5986-5990
2.
Russell JH, White CL, Loh DY, Meeledy-Rey P.
Receptor-stimulated death pathway is opened by antigen in mature T cells.
Proc Natl Acad Sci U S A.
1991;88:2151-2155 3. Wesselborg S, Janssen O, Kabelitz D. Induction of activation-driven death (apoptosis) in activated but not resting peripheral blood T cells. J Immunol. 1993;150:4338-4345[Abstract]. 4. Salmon M, Pilling D, Borthwick NJ, et al. The progressive differentiation of primed T cells is associated with an increasing susceptibility to apoptosis. Eur J Immunol. 1994;24:892-899[Medline] [Order article via Infotrieve]. 5. Dhein J, Walczak H, Baumler C, Debatin KM, Krammer PH. Autocrine T-cell suicide mediated by APO1(Fas/CD95). Nature. 1995;373:348-441. 6. Zheng L, Fisher G, Miller RE, Peschon J, Lynch RE, Lenardo M.J.. Induction of apoptosis in mature T cells by tumor necrosis factor. Nature. 1995;377:348-351[CrossRef][Medline] [Order article via Infotrieve].
7.
Ashkenazi A, Dixit VM.
Death receptors: signaling and modulation.
Science.
1998;281:1305-1308 8. Suda T, Okazaki T, Naito Y, et al. Expression of the Fas ligand in cells of T lineage. J Immunol. 1995;154:3806-3813[Abstract]. 9. Lynch DH, Ramsdell F, Alderson MR. Fas and FasL in homeostatic regulation of immune responses. Immunol Today. 1995;16:569-573[CrossRef][Medline] [Order article via Infotrieve].
10.
Hyde H, Borthwick NJ, Janossy G, Salmon M, Akbar AN.
Upregulation of intracellular glutathione by fibroblast-derived factor(s): enhanced survival of activated T cells in the presence of low bcl-2.
Blood.
1997;89:2453-2460
11.
Ayroldi E, Zollo O, Cannarile L, et al.
Interleukin 6 (IL-6) prevents activation-induced cell death: IL-2-independent inhibition of Fas/FasL expression and cell death.
Blood.
1998;92:4212-4219
12.
Vella AT, Dow S, Potter TA, Kappler J, Marrack P.
Cytokine-induced survival of activated T cells in vitro and in vivo.
Proc Natl Acad Sci U S A.
1998;95:3810-3815 13. Yakahashi T, Tanaka M, Brannan CI, et al. Generalized lymphoproliferative disease in mice caused by a point mutation in the Fas ligand. Cell. 1994;76:969-976[CrossRef][Medline] [Order article via Infotrieve].
14.
Rieux-Laucat F, Deist FL, Hivroz C, et al.
Mutations in Fas associated with human lymphoproliferative syndrome and autoimmunity.
Science.
1995;268:1347-1349 15. Fisher GH, Rosenberg FJ, Straus SR, et al. Dominant interfering Fas gene mutations impair apoptosis in a human autoimmune lymphoproliferative syndrome. Cell. 1995;81:935-946[CrossRef][Medline] [Order article via Infotrieve]. 16. Drappa J, Vainshaw AK, Sullivan KE, Chu J, Elkon KB. Fas gene mutation in the Canale-Smith syndrome, an inherited lymphoproliferative disorder associated with autoimmunity. N Engl J Med. 1996;35:1643-1649. 17. Laurence J, Mitra D, Steiner M, Lynch DH, Siegal FP, Staiano-Coico L. Apoptotic depletion of CD4+ T cells in idiopathic CD4+ T lymphopenia. J Clin Invest. 1996;97:672-680[Medline] [Order article via Infotrieve].
18.
Ozsahin H, Arredondo-Vega FX, Santisteban O, et al.
Adenosine deaminase deficiency in adults.
Blood.
1997;89:2849-2855 19. Sandstrom PA, Mannie MD, Buttke TM. Inhibition of activation-induced death in T cell hybridomas by thiol antioxidants: oxidative stress as a mediator of apoptosis. J Leukoc Biol. 1994;55:221-226[Abstract]. 20. Williams MS, Henkart PA. Role of reactive oxygen intermediates in TCR-induced death of T cell blasts and hybridomas. J Immunol. 1996;157:2395-2402[Abstract]. 21. Marchetti P, Hirsch T, Zamzami N, et al. Mitochondrial permeability transition triggers lymphocyte apoptosis. J Immunol. 1996;157:4830-4836[Abstract]. 22. Los M, Schenk H, Hexel K, Baeuerle PA, Droge W, Schulze-Osthoff K. IL-2 gene expression and NF-kB activation through CD28 requires reactive oxygen production by 5-lipoxygenase. EMBO J. 1995;14:3731-3740[Medline] [Order article via Infotrieve]. 23. Tatla S, Woodhead V, Foreman JC, Chain BM. The role of reactive oxygen species in triggering proliferation and IL-2 secretion in T cells. Free Radic Biol Med. 1999;26:14-24[CrossRef][Medline] [Order article via Infotrieve]. 24. Hildeman DA, Mitchell T, Teague TK, et al. Reactive oxygen species regulate activation-induced T cell apoptosis. Immunity. 1999;10:735-744[CrossRef][Medline] [Order article via Infotrieve]. 25. Westermann J, Bode U. Distribution of activated T cells migrating through the body: a matter of life and death. Immunol Today. 1999;20:302-306[CrossRef][Medline] [Order article via Infotrieve]. 26. Kessel GK. Basic Medical Histology: The Biology of Cells, Tissues and Organs. New York: Oxford University Press; 1998.
27.
Lynch RE, Fridovich I.
Permeation of the erythrocyte stroma by superoxide radical.
J Biol Chem.
1978;253:4697-4699 28. Winterbourn CC, Stern A. Human red cells scavenge extracellular hydrogen peroxide and inhibit formation of hypochlorous acid and hydroxyl radical. J Clin Invest. 1987;80:1486-1491.
29.
Denicola A, Souza JM, Radi R.
Diffusion of peroxynitrite across erythrocyte membranes.
Proc Natl Acad Sci U S A.
1998;95:3566-3571 30. Halliwell B, Gutteridge JMC. Free Radicals in Biology and Medicine. New York: Oxford University Press; 1999. 31. Tarnvik A. A role for red cells in phytohemagglutinin-induced lymphocyte stimulation. Acta Pathol Microbiol Scand (Sect B). 1970;78:733-740. 32. Johnson RA, Smith T, Kirkpatrick C. Augmentation of phytohemagglutinin mitogenic activity by erythrocyte membranes. Cell Immunol. 1972;3:186-197[CrossRef][Medline] [Order article via Infotrieve]. 33. Kalechman Y, Herman S, Gafter U, Sredni B. Enhancing effects of autologous erythrocytes on human or mouse cytokine secretion and IL-2R expression. Cell Immunol. 1993;148:114-129[CrossRef][Medline] [Order article via Infotrieve]. 34. Toth KM, Clifford DP, Berger EM, White CW, Repine JE. Intact human erythrocytes prevent hydrogen peroxide-mediated damage to isolated perfused rat lungs and cultured bovine pulmonary artery endothelial cells. J Clin Invest. 1984;74:292-295.
35.
Van Asbeck BS, Hoidal J, Vercellotti GM, Schwartz BA, Moldow CF, Jacob HS.
Protection against lethal hyperoxia by tracheal insufflation of erythrocytes: role of cell glutathione.
Science.
1985;227:756-759 36. Agar NS, Sadrzadeh SMH, Hallaway PE, Eaton JW. Erythrocyte catalase: a somatic oxidant defense? J Clin Invest. 1986;77:319-321.
37.
Brown JM, Grosso MA, Terada LS, et al.
Erythrocytes decrease myocardial H2O2 levels and reperfusion injury.
Am J Physiol.
1989;256:H584-H588 38. Hoffman RA, Kung PC, Hansen WP, Goldstein G. Simple and rapid measurement of human T lymphocytes and their subclasses in peripheral blood. Proc Natl Acad Sci U S A 77:4914-4919.
39.
Uchida K, Kanematsu M, Sakai K, et al.
Protein-bound acrolein: potential markers for oxidative stress.
Proc Natl Acad Sci U S A
1998;95:4882-4887 40. Bass DA, Parce JW, DeChatelet LR, Szejda P, Seeds MC, Thomas M. Flow cytometric studies of oxidative product formation by neutrophils: a graded response to membrane stimulation. J Immunol. 1983;130:1910-1917[Abstract]. 41. Gallop PM, Paz MA, Henson E, Latt SA. Dynamic approaches to the delivery of reporter reagents into living cells. BioTechniques. 1984;2:32-36.
42.
Arosa FA, de Jesus O, Porto G, Carmo AM, de Sousa M.
Calreticulin is expressed on the cell surface of activated human peripheral blood T lymphocytes in association with major histocompatibility complex class I molecules.
J Biol Chem.
1999;274:16917-16922
43.
Carmo AM, Castro MAA, Arosa FA.
CD2 and CD3 associate independently with CD5 and differentially regulate signaling through CD5 in Jurkat T cells.
J Immunol.
1999;163:4238-4245 44. Young IT. Proof without prejudice: use of the Kolmogorov-Smirnov test for the analysis of histograms from flow systems and other sources. J Histochem Cytochem. 1977;25:935-941[Abstract]. 45. Lampariello F. On the use of the Kolmogorov-Smirnov statistical test for immunofluorescence histogram comparison. Cytometry. 2000;39:179-188[CrossRef][Medline] [Order article via Infotrieve]. 46. Santos M, de Sousa M. In vitro modulation of T-cell surface molecules by iron. Cell Immunol. 1994;154:498-506[CrossRef][Medline] [Order article via Infotrieve]. 47. Freitas A, Rocha B. Peripheral T cell survival. Curr Opin Immunol. 1999;11:152-156[CrossRef][Medline] [Order article via Infotrieve].
48.
Mehal WZ, Juedes AE, Crispe IN.
Selective retention of activated CD8+ T cells by the normal liver.
J Immunol.
1999;163:3202-3210 49. Munn DH, Pressey J, Beall AC, Hudes R, Alderson MR. Selective activation-induced apoptosis of peripheral T cells imposed by macrophages: a potential mechanism of antigen-specific peripheral lymphocyte deletion. J Immunol. 1996;156:523-532[Abstract]. 50. Kay MBM, Marchalonis JJ, Schluter SF, Bosman G. Human erythrocyte aging: cellular and molecular biology. Transfus Med Rev. 1991;V:173-195. 51. Brittenham GM. The red cell cycle. In: Brock JH,Halliday JW,Pippard MJ,Powell LW, eds. Iron Metabolism in Health and Disease. London: WB Saunders; 1994:31-62. 52. Costa LMG, Moura EMF, Moura JJG, de Sousa M. Iron compounds after erythrophagocytosis: chemical characterization and immunomodulatory effects. Biochem Biophys Res Commun. 1998;247:159-165[CrossRef][Medline] [Order article via Infotrieve]. 53. Lahdenpohja N, Hurme M. CD28-mediated activation in CD45RA+ and CD45RO+ T cells: enhanced levels of reactive oxygen intermediates and c-Rel nuclear translocation in CD45RA+ cells. J Leukoc Biol. 1998;63:775-780[Abstract]. 54. Kagan VE, Fabisiak JP, Shvedova AA, et al. Oxidative signaling pathway for externalization of plasma membrane phosphatidylserine during apoptosis. FEBS Lett. 2000;477:1-7[CrossRef][Medline] [Order article via Infotrieve]. 55. Sagone AL, Husney R, Guter H, Clark L. Effect of catalase on the proliferation of human lymphocytes to phorbol myristate acetate. J Immunol. 1984;133:1488-1494[Abstract].
56.
Wesch D, Marx S, Kabelitz D.
Monocyte-dependent death of freshly isolated T lymphocytes: induction by phorbol ester and mitogens and differential effects of catalase.
J Immunol.
1998;161:1248-1256 57. Kumar S, Chakrabarti R. Amphotericin B both inhibits and enhances T-cell proliferation: inhibitory effect is mediated through H2O2 production via cyclooxygenase pathway by macrophages. J Cell Biochem. 2000;77:361-371[CrossRef][Medline] [Order article via Infotrieve]. 58. Virella G, Rugeles MT, Hyman B, et al. The interaction of CD2 with its LFA-3 ligand expressed by autologous erythrocytes results in enhancement of B cell responses. Cell Immunol. 1988;116:308-319[CrossRef][Medline] [Order article via Infotrieve]. 59. Ponka P, Lok CN. The transferrin receptor: role in health and disease. Int J Biochem Cell Biol. 1999;10:1111-1137.
60.
Pelosi E, Testa U, Louache F, et al.
Expression of transferrin receptors in phytohemagglutinin-stimulated human T-lymphocytes.
J Biol Chem
1986;261:3036-3042 61. Pantopoulos K, Hentze MW. Nitric oxide and oxidative stress (H2O2) control mammalian iron metabolism by different pathways. Mol Cell Biol. 1996;16:3781-3788[Abstract].
62.
Pantopoulos K, Mueller S, Atzberger A, Ansorge W, Stremmel W, Hentze MW.
Differences in the regulation of iron regulatory protein-1 (IRP-1) by extra- and intracellular oxidative stress.
J Biol Chem.
1997;272:9802-9808 63. Novogrodsky A, Suthanthiran M, Stenzel KH. Immune stimulatory properties of metalloporphyrins. J Immunol. 1989;143:3981-3987[Abstract]. 64. Novogrodsky A, Suthanthiran M, Stenzel KH. Ferro-mitogens: iron-containing compounds with lymphocyte-stimulatory properties. Cell Immunol. 1991;133:295-305[CrossRef][Medline] [Order article via Infotrieve]. 65. Saxena RK, Chandrasekhar B. A novel nonphagocytic mechanism of erythrocyte destruction involving direct cell-mediated cytotoxicity. Int J Hematol. 2000;71:227-237[Medline] [Order article via Infotrieve].
66.
Lipinski P, Drapier JC, Oliveira L, Retmanska H, Sochaniwicz B, Kruszewski M.
Intracellular iron status as a hallmark of mammalian cell susceptibility to oxidative stress: a study of L5178Y mouse lymphoma cell lines differentially sensitive to H2O2.
Blood.
2000;95:2960-2966 67. Uchida K. Current status of acrolein as a lipid peroxidation product. Trends Cardiovasc Med. 1999;9:109-113[CrossRef][Medline] [Order article via Infotrieve]. 68. Lowell MA, Xie C, Markesbery WR. Acrolein, a product of lipid peroxidation, inhibits glucose and glutamate uptake in primary neuronal cultures. Free Radic Biol Med. 2000;29:714-720[CrossRef][Medline] [Order article via Infotrieve].
69.
Aoshiba K, Nakajima Y, Yasui S, Tamaoki J, Nagai A.
Red blood cells inhibit apoptosis of human neutrophils.
Blood.
1999;93:4006-4010
70.
Liao JC, Hein TW, Vaughn MW, Huang K-T, Kuo L.
Intravascular flow decreases erythrocyte consumption of nitric oxide.
Proc Natl Acad Sci U S A.
1999;96:8757-8761 71. Munn LL, Melder RJ, Jain RK. Role of erythrocytes in leukocyte-endothelial interactions: mathematical model and experimental validation. Biophys J. 1996;71:466-478[Medline] [Order article via Infotrieve]. 72. Melder RJ, Yuan J, Munn LL, Jain RK. Erythrocytes enhance lymphocyte rolling and arrest in vivo. Microvasc Res. 2000;59:316-322[CrossRef][Medline] [Order article via Infotrieve]. 73. De Sousa M. Lymphocyte Circulation: Experimental and Clinical Aspects. London: John Wiley & Sons; 1981. 74. Shau H, Roth MD, Golub SH. Regulation of natural killer function by nonlymphoid cells. Nat Immunol. 1993;12:235-249. 75. Lachant NA, Tanaka KR. Impaired antioxidant defense in hemoglobin E-containing erythrocytes: a mechanism protective against malaria? Am J Hematol. 1987;26:211-219[Medline] [Order article via Infotrieve]. 76. Scott MD, Eaton JW. Thalassaemic erythrocytes: cellular suicide arising from iron and glutathione-dependent oxidation reactions? Br J Haematol. 1995;91:811-819[Medline] [Order article via Infotrieve]. 77. Wanachiwanawin W, Phucharoen J, Pattanapanyasat K, Fucharoen S, Webster HK. Lymphocytes in beta-thalassemia/HbE: subpopulations and mitogen responses. Eur J Haematol. 1996;56:153-157[Medline] [Order article via Infotrieve].
© 2001 by The American Society of Hematology.
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
 |
|
 |
|
  M. Minetti, L. Agati, and W. Malorni The microenvironment can shift erythrocytes from a friendly to a harmful behavior: Pathogenetic implications for vascular diseases Cardiovasc Res, July 1, 2007; 75(1): 21 - 28. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S. Corinti, L. Chiarantini, S. Dominici, M. E. Laguardia, M. Magnani, and G. Girolomoni Erythrocytes deliver Tat to interferon-{gamma}-treated human dendritic cells for efficient initiation of specific type 1 immune responses in vitro J. Leukoc. Biol., April 1, 2002; 71(4): 652 - 658. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
 |
 |
 |
|||||||
 |
 |
 |
 |
 |
 |
 |
 |
||
| Copyright © 2001 by American Society of Hematology Online ISSN: 1528-0020 | |||||||||
Top
Abstract
Introduction
receptor, with their agonists, CD95L (Fas ligand) and TNF-
) and without (
) RBCs was
determined by trypan blue exclusion, as indicated in "Material and
methods." Results show the mean ± SD of 3 independent
experiments. *P < .03.
