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Am J Physiol Cell Physiol 292: C1759-C1767, 2007. First published January 24, 2007; doi:10.1152/ajpcell.00158.2006
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GROWTH, DIFFERENTIATION, AND APOPTOSIS

Enhanced suicidal death of erythrocytes from gene-targeted mice lacking the Cl–/HCO3– exchanger AE1

Ahmad Akel,1 Carsten A. Wagner,2 Jana Kovacikova,2 Ravi. S. Kasinathan,1 Valentin Kiedaisch,1 Saisudha Koka,1 Seth L. Alper,3 Ingolf Bernhardt,4 Thomas Wieder,1 Stephan M. Huber,1 and Florian Lang1

1Department of Physiology, University of Tübingen, Tübingen, Germany; 2Institute of Physiology and Center for Integrative Human Physiology, University of Zürich, Zürich, Switzerland; 3Molecular and Vascular Medicine Unit and Renal Division, Beth Israel Deaconess Medical Center and Harvard Medical School, Boston, Massachusetts; and 4Zentrales Isotopenlabor/AG Biophysik, Saarland University, Saarbrücken, Germany

Submitted 16 April 2006 ; accepted in final form 19 January 2007


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Genetic defects of anion exchanger 1 (AE1) may lead to spherocytic erythrocyte morphology, severe hemolytic anemia, and/or cation leak. In normal erythrocytes, osmotic shock, Cl– removal, and energy depletion activate Ca2+-permeable cation channels with Ca2+-induced suicidal erythrocyte death, i.e., surface exposure of phosphatidylserine, cell shrinkage, and membrane blebbing, all features typical for apoptosis of nucleated cells. The present experiments explored whether AE1 deficiency favors suicidal erythrocyte death. Peripheral blood erythrocyte numbers were significantly smaller in gene-targeted mice lacking AE1 (AE1–/– mice) than in their wild-type littermates (AE1+/+ mice) despite increased percentages of reticulocytes (AE1–/–: 49%, AE1+/+: 2%), an indicator of enhanced erythropoiesis. Annexin binding, reflecting phosphatidylserine exposure, was significantly larger in AE1–/–erythrocytes/reticulocytes (~10%) than in AE1+/+ erythrocytes (~1%). Osmotic shock (addition of 400 mM sucrose), Cl– removal (replacement with gluconate), or energy depletion (removal of glucose) led to significantly stronger annexin binding in AE1–/– erythrocytes/reticulocytes than in AE1+/+ erythrocytes. The increase of annexin binding following exposure to the Ca2+ ionophore ionomycin (1 µM) was, however, similar in AE1–/– and in AE1+/+ erythrocytes. Fluo3 fluorescence revealed markedly increased cytosolic Ca2+ permeability in AE1–/– erythrocytes/reticulocytes. Clearance of carboxyfluorescein diacetate succinimidyl ester-labeled erythrocytes/reticulocytes from circulating blood was more rapid in AE1–/– mice than in AE1+/+ mice and was accelerated by ionomycin treatment in both genotypes. In conclusion, lack of AE1 is associated with enhanced Ca2+ entry and subsequent scrambling of cell membrane phospholipids.

annexin; cell volume; osmolarity; phosphatidylserine; energy depletion

ANION EXCHANGER 1 (AE1), a Cl–/HCO3– exchanger, is the most abundant intrinsic protein of the erythrocyte cell membrane (27). AE1 mediates the rapid exchange of Cl– with HCO3– and thus allows the erythrocyte to release HCO3– into the systemic capillaries that perfuse acidic, CO2-producing peripheral tissues. In pulmonary capillaries, erythrocyte AE1 takes up plasma HCO3– for intracellular conversion by carbonic anhydrase to CO2, which can then diffuse from the cell back into the plasma and across the alveolar epithelium for exhalation (27). Humans with defective AE1 may suffer from hereditary spherocytic anemia, ovalocytosis (1, 25, 33, 47), or stomatocytosis (11, 43). Animals completely lacking AE1 exhibit severe hemolytic anemia, often in the setting of hydrops fetalis (11, 42, 48). Most recently, several distinct missense mutations of AE1 have been associated with enhanced erythroid cation leak (11).

In normal erythrocytes, cation channels are activated by cell injury, such as oxidative stress, osmotic shock, or energy depletion (17, 24), leading to entry of Ca2+ and subsequent erythrocyte shrinkage, membrane blebbing, and breakdown of cell membrane phosphatidylserine (PS) asymmetry (28). The channels are also activated and PS exposure is triggered by removal of Cl– (16). PS exposure, cell shrinkage, and membrane blebbing could be similarly elicited by treatment of erythrocytes with the Ca2+ ionophore ionomycin (6, 9, 13, 28). Accounting for the similarities with and differences from apoptosis of nucleated cells (20, 21), the term "eryptosis" has been coined to describe this type of erythrocyte death (29).

To explore whether the machinery leading to suicidal erythrocyte death is modified by lack of AE1, the effects of osmotic shock, Cl– removal, and energy depletion were studied in erythrocytes from AE1 knockout (AE1–/–) mice and their wild-type (AE1+/+) littermates. Furthermore, we tested whether AE1–/– and AE1+/+ erythrocytes are more rapidly cleared from circulating blood in vivo and whether an increased Ca2+ concentration influences the removal of AE1–/– and AE1+/+ erythrocytes.


   METHODS
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 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Mice. Experiments were performed in erythrocytes from gene-targeted mice lacking AE1 (AE1–/–) and their AE1+/+ littermates (age: 3–5 mo). The generation and initial characterization of AE1–/– mice were as described by Peters et al. (42). The genotype was determined by PCR. For detection of the wild-type allele, oligonucleotides 5'-AGGTACACGGACAAGGTTTCTTGAT-3' and 5'-AGGCCAGAGGG TTAGAGGTGAATGTT-3' were used as forward and reverse primers, respectively, which amplified a 400-bp fragment. The primer pair 5'-CTTGGGTGGAGAGGCTATTC-3' and 5'-AGGTGAGATGACAGGAGATC-3' served to detect the mutant allele and amplified a 280-bp fragment. PCR results were detected on a 1.8% agarose gel. Animal experiments were conducted according to the guidelines of the American Physiological Society as well as the German law for the welfare of animals and were approved by local authorities. To obtain erythrocytes, animals were lightly anesthetized with isofluran (Abbott, Wiesbaden, Germany), and ~200 µl of blood were withdrawn into heparinized capillaries by puncturing the retroorbital plexus.

Solutions. Experiments were performed at 37°C in Ringer solution containing 125 mM NaCl, 5 mM KCl, 1 mM MgSO4, 32 mM HEPES-NaOH, 5 mM glucose, and 1 mM CaCl2 (pH 7.4). Where indicated, cells were exposed to osmotic shock (700 mosM by the addition of 400 mM sucrose), to Cl– removal (isosmotic replacement by gluconate), to energy depletion (glucose removal with or without the addition of 2-deoxyglucose), or to 1 µM of the Ca2+ ionophore ionomycin (Sigma, Taufkirchen, Germany). In further experiments, the HEPES-NaOH buffer of the Cl–-free solution was replaced by 25 mM NaHCO3, and cells were incubated in a 5% CO2 atmosphere. Where indicated, DIDS was added at a concentration of 100 µM.

Blood count. Erythrocyte number, packed cell volume (hematocrit), platelet number, and hemoglobin concentration were determined in blood from the mice using an electronic hematology particle counter (type MDM 905, Medical Diagnostics Marx, Butzbach, Germany) equipped with a photometric unit for hemoglobin determination and with volume settings adjusted for use with mouse erythrocytes.

Determination of GSH. Freshly drawn mouse erythrocytes (3% hematocrit) were washed once with glucose-free Ringer solution and incubated for 7 h at 37°C in Ringer solution, in glucose-free Ringer solution, or in glucose-free Ringer supplemented with 10 mM 2-deoxyglucose. After incubation, the supernatant was removed, and the pellet washed once with ice-cold PBS. Samples were then deproteinated by the addition of 200 µl of ice-cold sulfosalicylic acid (1%) and centrifuged for 5 min at 15,000 g. Lysates were diluted in distilled water (1:10), and the GSH content was determined by the addition of 100 µl of assay cocktail mix [92.3 µl of assay buffer (0.1 M NaPi and 1 mM EDTA; pH 7.5), 4 µl of 10 mM NADPH, 3 µl of 10 mM 5,5'-dithio-bis-(2-nitrobenzoic acid), and 0.7 µl of glutathione reductase (2 U/µl)] to 10 µl of the diluted sample. Finally, the absorbance of the samples was measured at 405 nm and compared with known concentrations of standard samples.

PS exposure. PS exposure was determined by annexin binding in FACS analysis as previously described (2, 28). After incubation, cells were washed in annexin binding buffer containing (in mM) 125 NaCl, 10 HEPES-NaOH (pH 7.4), and 5 CaCl2. Erythrocytes were stained with Annexin-Fluos (Roche Diagnostics, Mannheim, Germany) at a 1:100 dilution. After 15 min, samples were diluted 1:5 and measured by flow cytometric analysis on a FACS-Calibur from Becton Dickinson (Heidelberg, Germany). Annexin fluorescence intensity was measured in the fluorescence channel FL-1 with an excitation wavelength of 488 nm and an emission wavelength of 515–545 nm.

Double staining. Cells were exposed to osmotic shock (700 mosM by the addition of 400 mM sucrose to Ringer solution) for 4 h and to Cl– removal (isosmotic replacement by gluconate) for 7 h. Thereafter, cells were washed in annexin binding buffer and incubated (15 min) in annexin binding buffer containing annexin V-568 (1:50 dilution, Roche Diagnostics) and the DNA/RNA-specific dye Syto16 (30 nM, Molecular Probes, Leiden, The Netherlands). Samples were diluted 1:5, and Syto16- and annexin V-568-specific fluorescence was analyzed by flow cytometry in fluorescence channel FL-1 and FL-2 (emission wavelength of 564–606 nm), respectively.

Measurements of intracellular Ca2+ activity. Intracellular Ca2+ measurements were performed as previously described (4). Briefly, erythrocytes were loaded with Fluo-3 AM (Calbiochem, Bad Soden, Germany) by the addition of 2 µl of a Fluo-3 AM stock solution (1 mM in DMSO) to 1 ml of the erythrocyte suspension [0.3% hematocrit in Fluo-3 AM buffer containing 123 mM NaCl, 5 mM KCl, 1 mM MgSO4, 25 mM HEPES-NaOH (pH 7.4), 10 mM glucose, 10 mM pyruvate, and 2 mM CaCl2 (or 0 mM CaCl2 and 0.5 mM EGTA)]. Cells were incubated at 37°C for 15 min under shaking and with protection from light. An additional 2 µl of Fluo-3 AM (1 mM) were added with incubation carried out for 25 min without shaking. Fluo-3 AM-loaded erythrocytes were centrifuged at 1,800 rpm for 5 min at 22°C and washed with Ringer solution (or Ca2+-free Ringer solution additionally containing 0.5 mM EGTA). For flow cytometry, Fluo-3 AM-loaded erythrocytes were resuspended in 1 ml Ringer solution (0.3% hematocrit) containing the appropriate experimental solution with the Ca2+ ionophore ionomycin (1 µM) or vehicle alone (0.1% DMSO) and incubated for different time periods at 37°C. Ca2+-dependent fluorescence intensity was then measured in fluorescence channel FL-1 with an excitation wavelength of 488 nm and an emission wavelength of 530 nm.

Measurement of the half-life of fluorescence-labeled erythrocytes. Fluorescence-labeled erythrocytes were obtained by staining the cells with carboxyfluorescein diacetate succinimidyl ester (CFSE; Molecular Probes). The labeling solution was prepared by the addition of adequate amounts of a CFSE stock solution (10 mM in DMSO) to PBS to yield a final concentration of 5 µM. Cells were then incubated with labeling solution for 30 min at 37°C under light protection. Cells were pelleted at 2,000 rpm for 5 min, washed twice in PBS containing 1% FCS, and pelleted at 2,000 rpm for 5 min. The pellet was then resuspended in fresh, prewarmed Ringer solution. Fluorescence-labeled erythrocytes were injected in a volume of 100 µl into the tail veins of healthy C57BL/6 mice (female, 4 mo old). After the respective time periods, blood was taken from the injected mice, and the CFSE-dependent fluorescence intensity of the erythrocytes was measured in the fluorescence channel FL-1 as described above. Percentages of CFSE-positive erythrocytes were calculated as percentages of the total erythrocyte number. Control experiments revealed that the labeling procedure induced <10% lysis of AE1–/– or AE1+/+ erythrocytes (data not shown).

Statistics. Data are expressed as arithmetic means ± SE, and statistical analysis was made by paired or unpaired t-test or ANOVA using Dunnett's, Bonferroni, or Tukey's test as post hoc tests where appropriate.


   RESULTS
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 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Blood counts revealed severe anemia in AE1–/– mice. As shown in Table 1, erythrocyte number, packed cell volume, and plasma hemoglobin concentration in AE1–/– mice were only a fraction of the respective values in AE1+/+ mice. In contrast, AE1–/– mice had nearly 50% reticulocytosis compared with ~2% reticulocytes in AE1+/+ mice. Thus, as shown previously by Peters et al. (42) and Southgate et al. (48), the severe anemia was not due to impaired erythropoiesis but rather to hemolysis and/or accelerated clearance of circulating erythrocytes. Although slightly elevated, platelet counts in AE1–/– mice were not statistically different from those of AE1+/+ mice (P < 0.07).


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  		Table 1. Blood counts of AE1–/– and AE1+/+ mice

 
The loss of mature erythrocytes could have been due to cell membrane phospholipid scrambling leading to PS exposure at the cell membrane (28). Annexin binding, reflecting breakdown of PS asymmetry with exposure of PS at the cell surface, was indeed significantly larger in erythrocytes from AE1–/– mice than in erythrocytes from AE1+/+ mice (Fig. 1), a finding in accordance with earlier studies (22, 23). These data suggest a less stable PS asymmetry of the AE1–/– erythrocyte cell membrane.


Figure 1
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  		Fig. 1. Annexin binding to normal and anion exchanger 1 (AE1)-deficient erythrocytes: effect of osmotic shock. Annexin V binding was determined by FACS analysis of erythrocytes from AE1-deficient (AE1–/–) mice and their wild-type (AE1+/+) littermates after 4 h of incubation in isotonic solution (Ringer solution) or in 700 mosM hypertonic solution (addition to an isotonic solution of 400 mM sucrose). A: representative histograms illustrating the percentages of annexin V-binding erythrocytes. B: arithmetic means ± SE (n = 5) of the percentages of annexin V-binding erythrocytes. *Significant difference between control conditions and treatment with 700 mosM; #significant difference between AE1–/– and AE1+/+ mice.

 
Osmotic shock is known to stimulate PS exposure (28) in the absence of substantial hemolysis (30). As illustrated in Fig. 1, osmotic shock indeed led to breakdown of PS asymmetry of the cell membrane in erythrocytes from both genotypes. However, the increase in annexin binding following osmotic shock was significantly larger in AE1–/– erythrocytes than in AE1+/+ erythrocytes (Fig. 1).

Similar to osmotic shock, Cl– removal (isosmotic replacement with gluconate) from the extracellular fluid triggered breakdown of PS asymmetry in erythrocytes from both AE1–/– and AE1+/+ mice (Fig. 2). Exposure to Cl–-free extracellular solution significantly increased the number of annexin binding cells in both AE1+/+ and AE1–/– erythrocytes. Again, percentages of annexin-binding erythrocytes were significantly higher in AE1–/– blood than in AE1+/+ blood (Fig. 2). However, the absolute increment in the proportion of annexin-binding erythrocytes in Cl–-free medium was the same for both genotypes.


Figure 2
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  		Fig. 2. Effect of Cl– removal on erythrocyte annexin binding. Annexin V binding was determined by FACS analysis of erythrocytes from AE1–/– and AE1+/+ mice after 7 h of incubation in control Ringer solution or in Cl–-free Ringer solution (0 Cl–), i.e., Cl– replaced with gluconate. A: representative histograms illustrating the percentages of annexin V-binding erythrocytes. B: arithmetic means ± SE (n = 5 each) of the percentages of annexin V-binding erythrocytes. *Significant difference between control conditions and Cl– replacement; #significant difference between AE1–/– and AE1+/+ mice.

 
To explore whether the accelerated suicidal erythrocyte death in AE1–/– mice was due to high reticulocyte numbers in the AE1–/– blood, annexin binding was determined separately for erythrocytes and reticulocytes. Cells from AE1–/– and AE1+/+ mice were exposed for 7 h to Cl–-free extracellular solution or for 4 h to hyperosmotic shock and subsequently double stained with the DNA/RNA-specific dye Syto16 and fluorescence-labeled annexin (Fig. 3, A and B). As a result, under control conditions (Ringer solution) and following both removal of extracellular Cl– (Fig. 3C) and hyperosmotic shock (Fig. 3D), mature AE1–/– erythrocytes (i.e., cells with low Syto16 fluorescence) bound significantly more annexin than AE1+/+ erythrocytes. Annexin binding of reticulocytes (i.e., cells with high Syto16 fluorescence) did not differ between AE1–/– and AE1+/+ mice under control conditions (Fig. 3, C and D) and was significantly higher in AE1–/– than AE1+/+ mice following Cl– removal (Fig. 3C) but similar in AE1–/– and AE1+/+ mice following hyperosmotic shock (Fig. 3D). Taken together, these experiments indicated that the observed higher susceptibility of AE1–/– blood cells to suicidal death stimuli does not simply reflect the higher number of reticulocytes in AE1–/– blood.


Figure 3
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  		Fig. 3. Phosphatidylserine (PS) exposure following removal of extracellular Cl– or osmotic shock in reticulocytes and erythrocytes. A and B: dot blots of cells double stained with fluorescence-labeled annexin and the DNA/RNA-specific dye Syto16 depicting the annexin binding of erythrocytes (i.e., low Syto16 fluorescence) and reticulocytes (i.e., high Syto16 fluorescence) from AE1+/+ (A) and AE1–/– (B) mice. C and D: percentages of annexin-binding cells in erythrocytes and reticulocytes from AE1+/+ and AE1–/– mice after 7 h (C) or 4 h (D) of incubation in Ringer solution (Ringer solution and 290 mosm/kg, respectively) and in Cl–-free Ringer solution (C) or Ringer solution containing an additional 400 mM sucrose (700 mosm/kg; D), respectively. Data are arithmetic means ± SE (n = 3–4). ***P ≤ 0.001 ( by ANOVA).

 
In an additional series of experiments, Cl– removal experiments were repeated in 25 mM HCO3–/5% CO2-buffered saline. Furthermore, annexin binding following Cl– removal was studied in standard HEPES-buffered solutions in the presence of the AE1 inhibitor DIDS (100 µM). As shown in Fig. 4, A and B, respectively, neither the use of the HCO3–/CO2 buffer nor of DIDS blunted the increase in annexin binding elicited by extracellular Cl– removal. Thus, the enhanced sensitivity of AE1–/– cells to removal of extracellular Cl– was not due to the missing anion exchanger function of AE1.


Figure 4
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  		Fig. 4. Effect of AE1 activity on PS exposure of AE1+/+ and AE1–/– erythrocytes in Cl–-free medium. A: mean percentages of annexin-binding cells (±SE; n = 4) in AE1+/+ and AE1–/– erythrocytes incubated for 7 h in Ringer solution, in Cl–-free Ringer solution, or in Cl–-free solution buffered with 25 mM HCO3–/5% CO2. ***P ≤ 0.001 (by ANOVA). B: mean percentages of annexin-binding cells (±SE; n = 4) in AE1+/+ and AE1–/– erythrocytes incubated in the presence of the AE1 inhibitor DIDS (100 µM) for 7 h in Ringer solution or in Cl–-free Ringer solution. **P ≤ 0.01 and ***P ≤ 0.001 (by ANOVA).

 
Energy depletion, i.e., removal of glucose from the extracellular fluid, a maneuver that reduced the cellular ATP content by ~45% during 7 h of incubation (26), similarly led to sustained breakdown of PS asymmetry in erythrocytes from both AE1–/– and AE1+/+ mice (Fig. 5). Exposure to glucose-free extracellular fluid significantly increased the number of annexin binding cells in both genotypes, but the increment in annexin binding associated with energy depletion was again significantly larger in erythrocytes from AE1–/– mice (Fig. 5). Glucose depletion decreased the concentration of GSH in AE1–/– and AE1+/+ cells similarly (Fig. 6A), suggesting similar oxidation of AE1–/– and AE1+/+ cells upon glucose depletion. Even 2 h of glucose depletion or incubation with 2-deoxyglucose (10 mM) elicited annexin binding of >20% of AE1–/– cells while having no effect on AE1+/+ erythrocytes (Fig. 6B). Since 2-deoxyglucose reportedly fuels the pentosephosphate cycle leading to partial NADPH replenishment and glutathione regeneration (49), the difference in glucose depletion-stimulated annexin binding between AE1–/– and AE1+/+ cells was probably not due to a difference in redox state.


Figure 5
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  		Fig. 5. Effect of glucose depletion on erythrocyte annexin binding. Annexin binding was determined by FACS analysis of erythrocytes from AE1–/– and AE1+/+ mice following 7 h of incubation in control Ringer solution or glucose-free Ringer solution [0 glucose]. A: representative histograms illustrating the percentages of annexin V-binding erythrocytes. B: arithmetic means ± SE (n = 5) of the percentages of annexin V-binding erythrocytes. *Significant difference between control conditions and glucose removal; #significant difference between AE1–/– and AE1+/+ mice.

 

Figure 6
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  		Fig. 6. Energy depletion-induced changes in redox state and PS exposure of AE1+/+ and AE1–/– erythrocytes. A: normalized concentration (means ± SE; n = 3–4) of GSH in AE1+/+ and AE1–/– erythrocytes incubated for 7 h in Ringer solution and in glucose-free Ringer solution. B: mean percentage of annexin-binding cells (±SE; n = 4) in AE1+/+ and AE1–/– erythrocytes incubated for 2 h (top) and 7 h (bottom) in Ringer solution, in glucose-free Ringer solution, or in glucose-free Ringer solution supplemented with 10 mM 2-deoxyglucose (Deoxy-Glc). ***P ≤ 0.001 (by ANOVA).

 
The Ca2+ ionophore ionomycin (1 µM), which increases cytosolic Ca2+ activity, triggered breakdown of PS asymmetry in erythrocytes from both AE1–/– and AE1+/+ mice (Fig. 7). In the presence of ionomycin, elevated percentages of annexin binding erythrocytes did not differ between AE1+/+ and AE1–/– mice. Hence, the sensitivity of erythrocytes to elevated intracellular Ca2+ and subsequent suicidal erythrocyte death did not depend on the presence of AE1.


Figure 7
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  		Fig. 7. Effect of the Ca2+ ionophore ionomycin (Iono) on erythrocyte annexin binding. Annexin binding was determined by FACS analysis of erythrocytes from AE1–/– and AE1+/+ mice incubated for 30 min in control Ringer solution with (+) or without (–) the Ca2+ ionophore Iono (1 µM). A: representative histograms illustrating the percentages of annexin V-binding erythrocytes. B: arithmetic means ± SE (n = 3) of the percentages of annexin V-binding erythrocytes. *Significant difference between control conditions and treatment with ionomycin; #significant difference between AE1–/– and AE1+/+ mice.

 
From the experiments described above, we concluded that the enhanced basal level of PS exposure in AE1–/– erythrocytes could be due to enhanced Ca2+ entry (28). The increase of Fluo 3-dependent fluorescence during osmotic shock, as a measure of cytosolic free Ca2+ concentration, was indeed significantly steeper in AE1–/– mice than in AE1+/+ mice (Fig. 8, A–C). Moreover, the difference in annexin binding between AE1+/+ and AE1–/– cells following exposure to Cl–-free extracellular solution between AE1+/+ and AE1–/– cells was significantly blunted when experiments were performed in Ca2+-free solution (Fig. 8D). The observations are consistent with enhanced Ca2+ permeability of AE1–/– cells during cellular stress.


Figure 8
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  		Fig. 8. Increased Ca2+ entry into erythrocytes from AE1–/– mice following hyperosmotic or isosmotic cell shrinkage. A: overlay histograms showing Fluo 3 fluorescence of erythrocytes from AE1+/+ and AE1–/– mice at 2 and 25 min after the addition of Ca2+ and 400 mM sucrose to the medium. B: time course of mean Fluo 3 fluorescence changes (±SE; n = 4) in AE1+/+ and AE1–/– erythrocytes during hypertonic shock (addition of Ca2+ and 400 mM sucrose to the medium at time = 0 min). C: mean slope of the Fluo 3 fluorescence increase (±SE; n = 4) in AE1+/+ and AE1–/– erythrocytes as calculated from B between 2 and 10 min of recording. **P ≤ 0.01 (by a two-tailed Welch-corrected t-test). D: annexin binding of AE1+/+ and AE1–/– erythrocytes following 7 h of incubation in Ringer solution, in Cl–-free Ringer solution (0 Cl), and in Cl–-free/Ca2+-free Ringer solution (0 Cl/0 Ca). Data are arithmetic means ± SE (n = 5). ***P ≤ 0.001 (by ANOVA).

 
To explore the physiological significance of PS exposure, the clearance of erythrocytes from circulating blood was determined in vivo. To this end, erythrocytes were labeled with CFSE, leading to nearly complete labeling of the cell population (not shown). Additionally, a portion of erythrocytes was treated ex vivo with 1 µM ionomycin before being labeled, which leads to PS exposure in the majority of erythrocytes of both genotypes (~65%; see also Fig. 7). Given a lifetime of ~40 days, 10% of wild-type mouse erythrocytes are expected to be cleared within 4 days. In the present study, ~15% of CFSE-labeled wild-type erythrocytes were cleared during this time period (Fig. 9, open circles), indicating that CFSE labeling per se is not a strong trigger of erythrocyte clearance. The comparison of the results shown in Fig. 9 indicated that nontreated, CFSE-labeled erythrocytes from AE1–/– mice (closed circles) disappeared more rapidly from circulating blood than those from AE1+/+ mice (open circles) during the first 10 h of the experiment. In contrast to the nontreated cells, the vast majority of ionomycin-treated, CFSE-labeled erythrocytes from either genotype disappeared at a rapid clearance rate (Fig. 9, closed and open triangles). After 20 min, ~85% of the cells were already cleared. This rapid clearance was not due to cellular lysis during the labeling procedure, which was below 10% (data not shown).


Figure 9
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  		Fig. 9. Clearance of normal and AE1-deficient erythrocytes from circulating blood in C57BL/6 mice. Shown is the time-dependent decay of carboxyfluorescein diacetate succinimidyl ester (CFSE)-labeled circulating erythrocytes originally taken from AE1–/– or AE1+/+ mice, labeled with CFSE, and reinjected into C57BL/6 mice either without [control (C)] or with prior treatment with ionomycin (Io; 1 µM, 30 min). The percentage of CFSE-labeled cells is plotted against time after injection. Values are arithmetic means ± SE (n = 4). Some SEs are smaller than the respective symbols.

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The present results clearly demonstrate that the stress-stimulated Ca2+ permeability is higher in erythrocytes from AE1–/– mice than in erythrocytes from AE1+/+ littermates. Human and murine erythrocytes express Ca2+-permeable cation channels, which are opened by osmotic shock, Cl– removal, or energy depletion (16, 17, 24). An increased cytosolic Ca2+ concentration triggers Ca2+-sensitive scrambling of cell membrane lipids (15, 54) with subsequent PS exposure at the cell surface (6, 9, 28, 31, 32).

PS in the external leaflet of the plasma membrane of apoptotic cells reportedly is an "eat me signal" contributing to the recognition and phagocytosis of apoptotic cells by macrophages that are equipped with PS receptors (18, 19). PS exposure might also contribute to the clearance of erythrocytes. Accordingly, PS exposure parallels the time-dependent clearance of biotin-labeled rabbit erythrocytes in vivo (7). Moreover, erythrocytes are engulfed rapidly by macrophages in vitro (51) and cleared in vivo from peripheral blood after the insertion of a synthetic fluorescent PS analog into the erythrocyte membrane (45). Thus, the observed increase in Ca2+ permeability and the subsequent PS exposure are instrumental in the clearance of affected erythrocytes.

The present observations thus provide an explanation for the severe anemia of AE1 knockout mice (42, 48) and, potentially, also for the anemia of humans with AE1 mutations (1, 25, 33, 47). As noted previously by Peters et al. (42) and Southgate et al. (48), the dramatic increase of reticulocyte number in the AE1–/– mouse reflects profound stimulation of erythropoiesis that, however, is unable to prevent the severe anemia. No attempts have been made to discriminate between reticulocytes and mature erythrocytes during the determination of erythrocyte clearance. Thus, the relative large number of reticulocytes in AE1–/– mice may have influenced the magnitude of rapidity of erythrocyte clearance. Nevertheless, accelerated erythrocyte death accounts for, but does not result from, increased reticulocyte numbers.

Despite the absence of AE1, which normally constitutes 50% of intrinsic membrane protein, along with the secondary absence of other membrane proteins (10), AE1–/– erythrocytes contain a morphologically intact cytoskeleton (42). Therefore, the cytoskeletal consequences of loss of AE1 might not be expected directly to result in destabilization and randomization and, possibly, reorganization of the membrane lipids.

The enhanced PS exposure of AE1-deficient erythrocytes is reminiscent of the enhanced PS exposure of thrombocytes leading to a hypercoagulable state (22, 23). PS-exposing erythrocytes may contribute to vasoocclusion by binding to receptors in the vascular wall, thereby leading to obstruction of the microcirculation (3, 12). The correlation between PS exposure and thrombosis may, however, not hold in hereditary spherocytosis. All tested recessive hereditary spherocytosis diseases of the mouse (53) have been characterized by erythrocytes with elevated annexin V binding, whereas dominant hereditary spherocytosis in humans did not share this property (14). Moreover, the severity of thrombosis in spherocytic mice deficient in beta-spectrin or ankyrin (15–22% incidence of systemic thrombosis) or deficient in {alpha}-spectrin (85–100% incidence of thrombosis) did not correlate with the proportion or magnitude of erythrocyte annexin V binding (53).

The premature death of erythrocytes from AE1–/– mice is in seeming contrast to the putative role of AE1 in the machinery leading to physiological erythrocyte senescence (37–39, 46). Senescent erythrocytes are thought to increasingly bind naturally occurring anti-band 3 antibodies with subsequent activation of the classical complement pathway eventually leading to opsonization (36–40). Accordingly, the clearance of transfused red blood cells (RBCs) is delayed when blood to be banked undergoes the removal of complement (50). Oxidative stress damages hemoglobin and leads to the formation of hemichromes, which associate with the cytoplasmic domain of AE1 (34, 41, 52, 55), resulting in binding of endogenous circulating anti-band 3 antibody, activation of the complement system, and deposition of complement C3 molecules on the RBC surface (for reviews, see Refs. 35, 36, and 46). Although the survival of erythrocytes from AE1–/– mice was reduced despite the absence of the above AE1-dependent mechanism of physiological senescence, the present observations do not necessarily challenge the view that AE1-dependent complement activation leads to senescence and subsequent clearance of normal erythrocytes. Instead, AE1-dependent senescence and (stress-induced) suicidal erythrocyte death may be two distinct mechanisms limiting erythrocyte survival. While AE1-dependent senescence selects aged erythrocytes for removal from the circulation, suicidal erythrocyte death affects injured or stressed erythrocytes of all ages. In addition to senescence and suicidal erythrocyte death as mechanisms of erythroid demise, neocytolysis preferentially affects young erythrocytes, at least in humans (44). Several distinct pathways (5, 8, 29, 44, 46) may thus lead to premature or timely death of erythrocytes.


   GRANTS
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by Deutsche Forschungsgemeinschaft Grants La 315/4-3 and La 315/13-1; Bundesministerium für Bildung, Wissenschaft, Forschung und Technologie (Center for Interdisciplinary Clinical Research) Grant 01 KS 9602; IZKF-Promotionskolleg "Molekulare Medizin Grant 1547;" Biomed Program of the European Union Grant BMH4-CT96-0602; and National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-43495.


   ACKNOWLEDGMENTS
 
The authors acknowledge the meticulous preparation of the manuscript by Lejla Subasic and Natalie Eckberger.


   FOOTNOTES
 

Address for reprint requests and other correspondence: F. Lang, Physiologisches Institut, Universität Tübingen, Gmelinstrasse 5, Tübingen D-72076, Germany (e-mail: florian.lang{at}uni-tuebingen.de)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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