The main function of red blood cells is the delivery of O2 from the lung to the periphery and to transport CO2 from tissues to the lung. The precise control of regional blood flow is required to match the O2 demand, CO2 production, and blood flow. This is of particular importance when the metabolic activity is high, such as in an exercising skeletal muscle.
Microcirculation is adjusted by myogenic and metabolic controllers. One such controlling molecule is the vasodilator nitric oxide (NO), which is formed mainly in vascular endothelial cells. Red blood cells also produce bioactive NO equivalents in an O2 saturation and shear stress–dependent manner (3,11,23,29,40). Another stimulus for endothelium-dependent NO release seems to be adenosine triphosphate (ATP) in plasma (37). Its occurrence there depends on the presence of red blood cells (4,37). In skeletal muscle, red blood cells seem to function as an oxygen sensor and release ATP to cause an NO-dependent increase in blood flow (13). In vivo, plasma ATP has been found to be increased in the venous effluent from exercising forearm muscle (7,8). It was increased even further when exercise was performed in hypoxia (13). Forrester et al. (9) found that topically applied ATP leads to strong vasodilation. Taken together, these results indicate that ATP released from red blood cells or one of its breakdown products might induce local vasodilation.
The major stimulus for ATP release from red blood cells seems to be mechanical deformation (6,37). Also, in vitro exposure to hypoxia stimulates the release of ATP from red blood cells (1), which is in agreement with findings by Gonzalez-Alonso et al. (12), showing that local blood flow and ATP release depend on the oxygenation state of hemoglobin. ATP release is also stimulated by beta adrenergic stimulators and prostacyclin (22). Several release mechanisms for ATP have been described (for a review, see Praetorius and Leipziger ). In red blood cells, nonvesicular pannexin 1–dependent (27) and pannexin 1–independent (28) ATP release has been observed.
Exercise increases shear stress, causes local desaturation of hemoglobin, and increases the temperature in the working muscle, therefore resulting in a typical situation where ATP release from red blood cells might improve local blood flow (6,13,15). Rozier et al. (30) reported that elevated lactate at neutral pH reduced ATP release from red blood cells. However, heavy exercise is also associated with strong acidification from both muscle CO2 production and lactic acid formation, which decreases the oxygen saturation by decreasing the O2 affinity of hemoglobin, a mechanism expected to further facilitate ATP release (12,30). Another stimulus for ATP release might be the exercise-induced increase in muscle temperature (15).
Exercise induces pronounced intravascular hemolysis by the mechanical destruction of red blood cells as they pass through constricting skeletal muscles (33,35) and also in plantar blood vessels during running (47). Because the destruction of red blood cells also releases ATP contained in the cells, it is evident that exercise-induced intravascular hemolysis contributes to elevated ATP in plasma from venous blood leaving exercising muscles (13).
We tested the hypothesis that lactic acidosis as observed in working skeletal muscle, equivalent acidosis by HCl, and increased lactate at neutral pH enhance shear stress and hypoxia-induced release of ATP from human red blood cells. In these experiments, we also attempted to quantify to which extent hemolysis contributed to ATP appearing in the incubation media. It follows that any ATP not accounted for by hemolysis would have to be released by specific mechanisms. Red blood cells were exposed to shear stress and other experimental conditions in vitro in a Couette viscometer, where rotational shear stress induces the tank trading of the red cells as it is observed in vivo. This technique allows the quantification of release by hemolysis and by hemolysis-independent mechanisms. It also allows continued exposure to shear stress as it occurs during exercise. Our results show that in shear stress–exposed red blood cells in hypoxia, the appearance of most of the extracellular ATP was independent of hemolysis and may thus be explained by cellular release. We also found that lactic acidosis had no significant effect.
Heparinized blood was collected from antecubital veins from seven healthy, nonsmoking, male individuals (22–45 yr) after written informed consent during a medical checkup for inclusion into another study, which was approved by the local ethics committee. Twenty-gauge needles were used to minimize hemolysis during blood drawing. Red blood cells were washed in HEPES-buffered medium (145 mM NaCl, 4 mM KCl, 10 mM glucose, 2 mM MgCl2, 1 mM CaCl2, 10 mM HEPES, pH 7.4 at 37°C). Dextran 40 was added (20%) to obtain the appropriate shear stress in the viscometer (14). Washed red blood cells were suspended in the dextran-containing medium, and the hematocrit was adjusted to 10%.
Suspensions of red blood cells in buffer without dextran were equilibrated with nitrogen in a tonometer at 37°C. After equilibration, they were mixed with the dextran-containing medium, which was also preequilibrated with N2. The O2 saturation of the final cell suspension was approximately 20% as measured by oximetry (OSM 2a; Radiometer, Denmark). Normoxic cells were equilibrated with room air (O2 saturation > 90%).
To simulate exercise-induced changes in the microenvironment of red blood cells, hydrochloric acid (HCl; 15 mM, pH 7.0), lactic acid (15 mM, pH 7.0), and Na lactate (15 mM, pH 7.4) was added.
The cell suspension was transferred to a prewarmed (37°C) rotating concentric cylinder Couette viscometer (21). The rotational speed (70 and 210 rpm), which was adjusted to obtain shear rates in the low physiological range (21 and 65 dyn·cm−2, respectively), is up to approximately 15 times higher than the values reported for capillary shear stress at rest (46) but far below the hemolytic shear stress threshold of 1500 dyn·cm−2 (42) and also below the shear stress where a net K+ leak has been observed (21).
Cell suspensions (5 mL) were pipetted into the gap (0.63 mm) between the stationary center rod heated to 37°C with a circulating water bath and the rotating outer cylinder of the viscometer and were exposed to shear stress for 10 and 20 min. When working with hypoxic erythrocytes, the gap of the viscometer above the cell suspension was flushed with nitrogen to prevent reoxygenation of the cells. Control cells were kept static (nonsheared) at 37°C in polyethylene tubes. The high dextran content prevented sedimentation of red blood cells during nonsheared incubation. After exposure, aliquots of cell suspensions were mixed with one volume of dextran-free medium and centrifuged (60 s, 10,000 rpm). Aliquots of the supernatant and of the original cell suspension were denatured with 0.1 N perchloric acid for the measurement of supernatant and total ATP.
Hemoglobin in the cell suspension and in the supernatant was measured with a modified Drabkin assay. The detection limit was approximately 0.02% hemolysis. ATP in the supernatant was measured with a luciferin–luciferase chemiluminescence assay (Roche, Germany). The specific release of ATP by red blood cells was calculated from the total erythrocyte and supernatant ATP and the percentage of hemolysis assuming complete liberation of all ATP from the hemolyzed red blood cells.
To avoid problems with interpreting results due to changes in red cell volume upon exposure to isotonic and hypertonic media, supernatant ATP was calculated as the number of ATP molecules released from 500 μL of red cells at their original volume in isotonic buffer, pH 7.4, and in normoxia. Results shown are mean ± SD values. Differences were analyzed by one- and two-way ANOVA for repeated-measures and LSD post hoc tests. Group differences were also analyzed with paired t-tests when appropriate. The SigmaStat and SigmaPlot software (SYSTAT, Erkrath, Germany) was used for analysis. The level of significance was P < 0.05.
Figure 1A indicates that none of the treatments significantly affected total ATP in red blood cells. In normoxic cells, the SO2 was >90% (not shown). Exposure to hypoxia resulted in an SO2 of approximately 22% (Fig. 1B). SO2 was not affected by treatment with Na lactate but was significantly decreased by lactic acid and HCl, which is due to the decrease in Hb–O2 affinity by acidification.
We determined then whether treatments induce a cellular release of ATP to the supernatant and to which extent this was due to hemolysis. Figure 2A indicates that the nonsheared incubation of red blood cells in the dextran-containing buffer caused some increase in extracellular ATP. Ten minutes of shearing at 70 rpm did not cause a statistically significant increase in extracellular ATP, but ATP was increased significantly after 20 min of shearing (P < 0.05). Extracellular ATP was elevated after 10 and 20 min when cells were sheared at 210 rpm (P < 0.05). Figure 2B shows that there was some hemolysis even in nonsheared red blood cells, and that hemolysis increased further with increasing the shear rate (P < 0.05). Subsequent experiments were performed by exposure of cells for 20 min to a rotational speed of 210 rpm.
In normoxic cells, shear stress significantly (P < 0.01) increased ATP in the medium in control and in cells treated with Na lactate and hydrochloric acid (pH 7.0), but not when red blood cells were exposed to lactic acid (pH 7.0) (Fig. 3A). Supernatant ATP was significantly elevated in hypoxic red blood cells in all experimental conditions, even when cells were not exposed to shear stress (P < 0.001). The exposure of hypoxic cells to shear stress caused an additional increase in ATP (P < 0.001), which was less pronounced when cells were exposed to lactic acid.
Figure 3B shows that there was some hemolysis in red blood cells not exposed to shear stress in normoxia and hypoxia; there was no significant difference between control and chemical treatments. Shear stress induced a significant increase in hemolysis in normoxic red blood cells (P < 0.001). As in the nonsheared cells, shear stress–induced hemolysis did not differ between treatment groups. Shear stress–induced hemolysis was less pronounced in the hypoxic cells (P < 0.05).
From the supernatant ATP, the total red blood cell ATP, and the percentage of hemolysis, we calculated how much of the supernatant ATP can be explained by hemolysis. Figures 4A and 4B shows that in normoxia, shear stress increased hemolysis-related ATP about fivefold in normoxic control cells and in cells treated with Na lactate. There was only an about twofold increase in hemolysis-associated extracellular ATP in cells treated with lactic acid and HCl (in all cases P < 0.001). No increase in shear stress–induced, hemolysis-associated ATP was found in hypoxia control cells (P < 0.358), whereas a 30% increase was found after treatment with lactic acid (P < 0.001). A more than twofold increase was found in cells treated with Na lactate and HCl (P < 0.001) (Fig. 4A).
Under all conditions, non–hemolysis-associated ATP accounted for a large portion of supernatant ATP, indicating ATP release by specific mechanisms (Fig. 4C). Shear stress did not induce a significant specific release of ATP in normoxic cells (P < 0.659), whereas hypoxia induced a pronounced increase in released ATP (P < 0.001) even when cells were not exposed to shear stress (P < 0.001). Release was significantly more pronounced when cells were exposed to shear stress and hypoxia together (P < 0.001). In the hypoxic cells, shear stress–induced ATP release was higher in Na lactate and HCL than that in control and in cells treated with lactic acid (P < 0.05).
Extracellular ATP and its metabolites are potent local vasodilators. The cellular release of ATP from red blood cells contributes to elevated plasma ATP, and several specific release mechanisms have been postulated (for a review, see Ellsworth et al. ). Intravascular hemolysis also releases ATP into plasma. Here we show that in vitro shear stress of red blood cells in a Couette viscometer causes the release of ATP, and that at a high oxygen saturation of hemoglobin, a large proportion can be explained by hemolysis. We confirm that hypoxia of nonsheared cells also causes ATP release and shows that it is further enhanced by shear stress and mostly independent of hemolysis. Lactic acidosis did not alter shear stress and hypoxia-induced ATP release.
Total ATP release from red blood cells: concentrations achieved in blood capillaries
Our experiments were designed to measure accumulating amounts of extracellular ATP by exposure to the stressors for up to 20 min. Because cells were washed and suspended in Krebs buffer, the degradation of released ATP by exonucleotidases was avoided. Therefore, our results reflect the maximum capacity of ATP release from red blood cells. Because ATP release was linear over time, it is possible to calculate ATP release for short periods, such as capillary transit time (s), from the 20-min incubations.
Thus, from the measured supernatant ATP concentrations, we calculated that the 500 μL of red blood cells used in the assay (hematocrit 10%) released approximately 13 × 1014 molecules of ATP during the 20-min incubation in normoxia without shear stress. This value was nearly doubled by shear stress in normoxia (21 × 1014 molecules), whereas exposure to hypoxia and shear stress together increased release to 150 × 1014 ATP molecules per 20 min (Fig. 3A). For extrapolation to the situation in a blood capillary, we used the results by Kindig et al. (17), who reported a red blood cell velocity of 270 μm·s−1 and 428 μm·s−1 in resting and electrically stimulated, respectively, rat spinotrapezius muscle. Assuming an average length of muscle capillaries of approximately 1 mm (24), a red blood cell spends approximately 3.7 s in the muscle capillary at rest but only 2.3 s during exercise. Thus, it can be calculated that ATP release from a single red blood cell might generate a local ATP concentration of approximately 7 nM at the venous end of a capillary. This value might increase to approximately 13 nM during muscle contractions when the shear stress is increased as in our in vitro experiments and when the PO2 is high. Deoxygenation of hemoglobin, which occurs when muscle cell O2 consumption is increased, might raise the end capillary ATP concentration to almost 100 nM. The delayed onset of the ATP release of approximately 29 ms reported by Wan et al. (45) should not decrease these numbers significantly because this time is very short relative to the total capillary transit time. These calculated local ATP concentrations are in the range where arteriolar vasodilation was found upon intraluminal ATP application (19). In contrast, a concentration of 1 μM was estimated from in vitro ATP release from red blood cells by McCullough et al. (19). The differences might be in the shear stress applied, which was relatively low in our experiments (65 dyn·cm−2) relative to others (up to ∼2900 dyn·cm−2 ). Results from in vivo measurements indicate plasma ATP concentrations up to 2 μM in the venous blood from exercising muscle (8,13), but the compartment (vascular endothelium, muscle cells, and blood cells), where ATP was coming from, was unclear.
Contribution of hemolysis
Whereas several mechanisms for the specific release of ATP from red blood cells have been described (for a review, see Ellsworth et al. ), the potential contribution of ATP released by hemolysis has not been addressed experimentally. The hemolysis of red blood cells occurs at shear rates between 1000 and 4000 dyn·cm−2 (31,42). An increased flow in blood vessels, the shear stress while passing through blood capillaries, the mechanical pressure of the contracting muscle during physical exercise, and the mechanical strain by physical impact can cause intravascular hemolysis. It has been found to be related to the intensity and kind of exercise (20,47). Foot strike has been identified as a significant source of hemolysis in runners (43) that can be prevented by good shoe cushioning (5,47). Intravascular hemolysis after strength training was indicated by decreased haptoglobin (32) and by hemoglobinuria after karate (41). An increased concentration of hemoglobin in plasma (e.g., [32,43]) is a direct indicator of intravascular hemolysis. Telford et al. (43) reported plasma hemoglobin of approximately 30 mg·L−1 at rest and approximately 120 mg·L−1 after running exercise, indicating that approximately 0.04% of red blood cells were lyzed during exercise. In vitro, we found 0.05% of hemolysis in nonsheared cells and a fourfold increase by shear stress of normoxic cells. Hemolysis in our setting might be somewhat increased because of a possible effect of dextran and the absence of plasma (16). From the degree of hemolysis found in vivo, it can be estimated that every 500th to 2000th red blood cell would lyze when exposed to shear stress. If occurring during passage through a capillary, hemolysis would not result in a stable elevation of ATP at the venular end of a capillary but rather result in ATP pulses. If hemolysis is a consequence of traumatic impact, it occurs most likely not only in small but also in larger blood vessels. It is of question whether this is relevant to microcirculation.
Hypoxia of nonsheared cells did not affect the degree of hemolysis. Interestingly, hypoxia blunted the shear stress–related increase in hemolysis. This is in contrast to results from prolonged in vivo hypoxia showing a decreased deformability and increased susceptibility to red cell damage (18,34).
Specific ATP release
Several groups have measured directly ATP release after red blood cells had passed through a narrow capillary or porous filters to induce shear stress using the luciferase assay (26,37,39,45). In these systems, red blood cells are exposed to the stimulus for milliseconds only. In contrast, we exposed cells to the different stimuli for 20 min and calculated “specific ATP release” as the proportion of extracellular ATP that could not be explained by hemolysis. Also in this setting, we observed a hypoxia-induced increase in ATP release, thus confirming earlier findings (1). In contrast to others (6,37), we found no significant effect of shear stress on specific ATP release from normoxic cells.
Effects of lactic acidosis
Besides increased shear stress and deoxygenation, exercise causes a variety of other changes of the microenvironment, which red blood cells encounter while passing through capillaries in working muscle. The most prominent is lactic acidosis. We found that specific ATP release and hemolysis in nonsheared and sheared, normoxic, and hypoxic red blood cells were not affected by lactic acidosis. We also looked for the independent effects of acidosis (by adding HCl) and elevated lactate (15 mM at neutral pH). Interestingly, both treatments increased specific cellular ATP release from sheared red blood cells, which was even more pronounced in hypoxia. It appears as if the combination of both, elevated lactate plus acidosis abolishes these individual effects by mechanisms that require further clarification. This might be because the increase in cell volume by increased extracellular osmolality due to the elevated lactate, which widens the difference between intra- and extracellular lactate concentration (36), is balanced by cell swelling caused by acidosis (10,44). Individually, each of the effects, shrinkage and swelling of red cells, would decrease their deformability (2), which might also increase shear stress–related ATP release.
Our results on lactate effects are in contrast to a recent report showing that in nonsheared, hypoxic rabbit red blood cells, the addition of Na lactate (pH 7.4) significantly impaired ATP release. The discrepancy might be explained by the almost twofold higher concentration of lactate in the latter study (30) and possibly also by the different species used. Together, our results indicate that chemical changes induced by lactate, Cl (from HCl), and acidosis can modify the shear stress and hypoxia induced the release of ATP.
Besides increasing shear stress and decreasing oxygenation, exercise induces several additional strains on red cells that might affect ATP release. The most prominent is an increase in temperature in the working skeletal muscle and an elevated CO2 due to increased aerobic metabolism. Kalsi and Gonzalez-Alonso (15) have shown that an increase in temperature augments ATP release from red blood cells in vitro, which is not caused by hemolysis. It has not been studied whether altered temperature modifies ATP release induced by shear stress and by hypoxia. Ueda and Bookchin (44) have shown that an elevation of CO2 from 40 to 80 mm Hg at constant extracellular pH causes shrinkage of red blood cells by approximately 8%, which would completely abolish cell swelling by a decrease in pH from 7.4 to 7.1 induced by fixed acids. In vivo, elevated CO2 might therefore somewhat decrease deformability, which might stimulate ATP release. Together, elevated temperature and CO2 might enhance ATP release caused by shear stress and hypoxia.
In conclusion, data from in vivo and in vitro studies provide strong evidence that significant amounts of ATP are released from red blood cells upon exposure to hypoxia and shear stress at the same time. The release of ATP seems to be independent of the glycolysis because the cellular concentration of ATP remained unchanged during stress. Although in normoxia up to 50% of ATP release is due to hemolysis in our experimental setting, its contribution decreases less than 5% in hypoxia, which likely represents the situation in exercising muscle. The ATP concentration achieved by specific release from red blood cells appears to reach levels relevant to cause local vasodilation. Although probably all red blood cells exposed to the stressors contribute to specific release, only few red blood cells will lyze, causing only occasional ATP pulses. Therefore, ATP from specific release rather than from hemolysis contributes most to the regulation of blood flow in the microcirculation.
The authors thank Mrs. Christiane Herth and Mrs. Sonja Engelhardt for excellent technical assistance. They also thank Robert P. Hebbel, University of Minnesota, for providing us with the Couette viscometer and Joseph F. Hoffman, Yale School of Medicine, for fruitful conversation regarding the interpretation of the data and work on the manuscript. The results of the present study do not constitute endorsement by the American College of Sports Medicine.
The study was financed by departmental funds.
The authors declare no conflict of interest.
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