Platelets and a platelet-released enhance endothelial barrier F. R. HASELTON Department
AND J. S. ALEXANDER
of Biomedical
Engineering,
Vanderbilt
Haselton, F. R., and J. S. Alexander. Platelets and a platelet-released factor enhance endothelial barrier. Am. J. Physiol. 263 (Lung Cell. Mol. Physiol. 7): L670-L678, 1992.The role of platelets in the maintenance of endothelial barrier is examined in an in vitro model of the microvasculature. Human platelets (6,000/~1) perfused through a cell column of endothelial-covered microcarriers decrease paracellular permeability of sodium fluorescein (mol wt 342) to 63% of baseline values. This effect is reversible and a second application and removal of platelets produces a similar response. This effect occurs within 5 min and reverses within 10 min after platelet removal. The reduction in permeability is not due to mechanical obstruction of endothelial junctions, since the number of recirculating platelets is not reduced and releasate from unstimulated 2-h platelet incubations also decreases permeability. Releasate from platelets stimulated with 0.1 U/ml of thrombin for 15 min have the same permeability reducing effect. In this system, the platelet factors serotonin ( 10e3 M) and ADP ( 10S4 M) have no effect on permeability. However, the platelet factors adenosine ( 10v4 M), ATP ( 10b5M), and ,&agonists decrease permeability. None of these appear to account for platelet permeability activity, since activity is not blocked by agents directed against these mediators (adenosine deaminase, apyrase, S-phenyltheophylline, or propranolol). The active factor(s) is stable at -2O”C, heat stable, sensitive to trypsin, and has an apparent molecular weight X00. We conclude that unstimulated platelets release a factor(s) that enhances endothelial barrier in vitro and may be important in maintenance of the normal in vivo barrier. paracellular permeability; serotonin; adenosine; adenosine 5’triphosphate; adenosine 5’-diphosphate
of studies have used cultured endothelial monolayers as a model system to study transvascular capillary exchange (1, 11,15,15a, 19,30,34). It has been observed in these systems that some physiological mediators, e.g., ,&agonists, bradykinin, thrombin, and histamine have direct effects on endothelial monolayer permeability in vitro (5, 11, 15, 21). Although not completely analogous, the responsiveness of these pure endothelial models continues to make this approach valuable in investigations of endothelial barrier. One consistent difference between in vitro and in vivo systems is the finding that in vitro endothelial monolayers have permeabilities that are from IO-1,000 times higher than the transvascular permeabilities values measured in vivo (1, 15). By definition, in vitro systems are incomplete, and it has been suggested that the permeability differences between these systems may be the lack of other vessel wall layers (26), differences in the properties of the in vitro extracellular matrix (7), or use of large vessel endothelial cells to model microvascular phenomena (4). Another possibility is that the in vitro tissue culture systems lack some essential circulating component of the vascular system that normally contributes to vessel integrity. Platelets are an important vascular component that
A NUMBER
L670
factor
1040-0605/92
$2.00
Copyright
University,
Nashville,
Tennessee
37235
have been shown to be involved in thrombosis, hemostasis, and the maintenance of vascular integrity. Experimental or pathological loss of circulating platelets (thrombocytopenia) is associated with increased vascular fragility and erythrocyte extravasation (2, 12, 16, 23, 28,32,33). However, it is unclear whether platelets contribute to the endothelial barrier in vivo. One possibility is that vasoactive compounds stored in platelets may promote barrier. Platelet contents are normally released by either slow leakage, e.g., 5-hydroxytryptamine (5HT), or stimulation that results in rapid release of CYand dense granules storage pools (17, 18). Recently, platelets have been shown to decrease the permeability of albumin across an endothelial monolayer separating two chambers (23, 31). In the endothelial model employed in that study (3l), the effect was not due to physical blockade, and a similar permeability effect was observed with either platelet lysate supernatant or thrombin stimulated platelets. The unknown active factor was not a P-agonist, serotonin, or a cyclooxygenase product (3 1). We have recently described cell-column chromatography as another in vitro model useful in studies of transvascular exchange (14,15). One of the unique properties of our approach is that it utilizes continuously flowing systems both in the culture system and in the design of the cell columns used to measure endothelial permeability. This method also finds permeabilities that are 10 times higher than measured in vivo values (15). In this study, we explore the possibility that circulating platelets are an important barrier enhancing component missing from in vitro systems. We measure the change in endothelial monolayer permeability produced by adding platelets and platelet releasate to our flowing in vitro system and partially characterize the responsible platelet factor(s). METHODS CeZl culture methods. Well-established techniques were used to isolate and culture bovine aortic endothelial cells (BAEC) and fetal aortic endothelial cells (BFAE); detailed procedures used for primary cell isolation, cell identification, monolayer culture subcultivation, and the determination of in vitro lifespan have been described previously (15, 27). Briefly, cultures of adult and fetal aortic endothelial cells were obtained from thoracic aortae by means of 0.1% collagenase treatment. Monolayer cultures were established in Dulbecco’s modified Eagle’s medium containing 10% fetal bovine serum and were subcultured using 0.1% trypsin-EDTA. Endothelial cell identity was verified by indirect immunofluorescent assay for factor VIII-related antigen and diI-low-density lipoprotein (LDL). Cell lines were used during the vigorous proliferative phase of their in vitro lifespan. The fetal bovine cell lines (AG-7680 and AG7681) are available from the Cell Repository at the Coriell Institute for Medical Research (Camden, NJ).
0 1992 The
American
Physiological
Society
Downloaded from www.physiology.org/journal/ajplung by ${individualUser.givenNames} ${individualUser.surname} (130.070.008.131) on January 16, 2019.
PLATELET-RELEASED
FACTOR
ENHANCES
Cells were cultured on microcarrier beads as previously described (15). Cells were seeded on Cytodex-3 microcarrier beads at a density of 2 x IO4 cells/cm2. Cell attachment was achieved by intermittent stirring overnight. Microcarrier cultures were maintained at 60 rpm continuously and fed three times a week. Cultures were used for these assays between 9 and 30 days postseeding. Unless otherwise noted, all biological and tissue culture supplies were purchased from Sigma Chemical (St. Louis, MO). Platelet isolation methods. Human platelets were obtained from normal volunteers and isolated by the two methods described below (8, 35). Subjects were nonsmokers, drug and aspirin free for at least 3 days before platelet donation. Sixty milliliters of blood was withdrawn through an extension tube using a 20-gauge needle into a syringe containing sodium citrate. In one method (method A) whole blood was spun for 30 min at 300 g over a Ficoll-hypaque density gradient (Flow Labs). The platelet-rich plasma was resuspended in Ca2+ free N2-hydroxyethylpiperazine-N’-2-ethanesulfonic acid (HEPES) buffer containing 0.35% bovine serum albumin. This platelet suspension was spun 15 min at 100 g to remove leukocytes and red blood cells. A second method (method B) developed by Timmons and Hawiger (35) was also used to remove adventitiously adherent serum proteins. Whole blood was spun for 15 min at 250 g. The platelet-rich plasma was layered over a lo-25% albumin gradient and spun 15 min at 100 g. Platelets were removed and resuspended in Ca 2+-free HEPES buffer and passed over a Sepharose 2B gel column (35). Platelet releasates were prepared by incubating platelet suspensions at room temperature for 120 min in Hanks’ balanced salt solution (HBSS) containing 0.5% bovine serum albumin and 25 mM HEPES. Platelets were removed from suspension by centrifugation at 100 g for 15 min and 0.5-ml aliquots were frozen at -20°C in polypropylene Eppendorf tubes. PLateLet activation assay. Activation of platelets was assessed by their ability to release radiolabeled serotonin on stimulation with 10m6 M phorbol myristate acetate (PMA). Platelets were loaded with 1.5 &i/ml of [14C]serotonin (Amersham). Serotonin release was measured by comparing the releasate from spun platelets with the total activity of Triton X-100 (1%) treated platelets. Platelet activation was achieved with 10V6 M PMA for 10 min, and the release was compared with that found in unstimulated conditions. Cell-column methods. We used a previously reported assay of endothelial monolayer permeability, with modifications, to measure changes in barrier produced by stimulation of endothelial monolayers with experimental treatments (15). The method uses a model of the vasculature consisting of a chromatographic cell-column filled with cell-covered microcarrier beads. The permeability of the endothelial monolayers covering the beads is determined from a comparison of the elution curves of tracers injected into the flow at the top of the column. The details of this method have been previously described (15), and this approach, with modifications, is briefly described below. Chromatographic cell columns were made from water-jacketed glass columns (0.65-cm diam, Rainin). Cell-covered beads were poured to a column height of -2 cm, which provided 130 cm2 of endothelial cell culture surface, or -1 X IO7 cells. The column was washed and equilibrated with HBSS containing 0.5% albumin. Perfusion through the column was maintained by a Gilson peristaltic pump, at 0.9 ml/min. This flow was chosen to approximate the gravity flow rate observed when the pump was not connected. A tracer bolus described below was applied by a rotary injection valve (Rainin) using a 50-~1 loop. The cell column and all perfusate solutions were kept at 37°C.
ENDOTHELIAL
BARRIER
L671
Multiple tracer indicator-dilution analysis was used to obtain cell layer permeability from the relative shapes of the elution profiles of tracers simultaneously applied to the top of a cell column. One of the applied tracers (Blue Dextran, 10 mg/ml, 2,000 kDa) is impermeant and follows the mobile phase, i.e., a flow tracer. Two other tracers, initially sodium fluorescein (mol wt 342), and in later experiments sodium fluorescein and cyanocobalamin (mol wt 1,355), were used as permeant tracers, able to permeate beneath the cell layer and diffuse into the bead matrix beneath the cells. Using only optical tracers allows us to collect and measure the concentration of three optical tracers simultaneously without having to rely on radioactive materials previously employed and also has the advantage of using a single instrument to quantify the tracer concentrations contained within each sample. Elution profiles were constructed from 66 samples of the column eluant using a modification of previously reported methods (15). A Gilson fraction collector (model 203) equipped with a drop counter was used to collect two drops of eluant per well for 66 wells of a 96-well microtiter plate. The absorbance of each of the 96 wells was read at 620,540, and 492 nm on a plate reader (Titertek MCC 340) and stored on a computer for analysis. The optical absorbances were used to calculate the fractional recovery per sample of each of the optically absorbing dyes. Estimates of permeability based on the column-elution patterns of multiple tracers is an adaptation and extension of techniques used in vivo to assess capillary permeability (13,29). To apply these techniques to these experiments, a mathematical model of tracer motion has been developed based on the physical picture of tracer behavior described above. In this previously described model (15), it is assumed that the elution profiles of cyanocobalamin and sodium fluorescein depend on the properties of the mobile phase of the column plus the paracellular permeability properties of the endothelial monolayer and the diffusive motion of the tracer within the microcarrier beads. In contrast to these permeant tracers, the elution of Blue Dextran depends only on the flow (mobile) phase properties of the column. A modified Marquardt iteration scheme was used to estimate monolayer permeability which best approximated the experimental data. Best fit was determined by the minimization of the coefficient of variation between a computer-generated prediction of the permeant tracer’s elution profile and the experimentally observed elution profile. A permeability value for any one column and time point was computed as the mean of two consecutive measurements (spanning 5-7 min). Significance among three or more column conditions was judged using one-way analysis of variance and Tukey’s modified t test. A P < 0.05 was judged significant. Treatment protocols. Consecutive permeability measurements on the same population of cells were made in duplicate during a number of consecutive experimental treatments. All treatment protocols followed this same basic design. In all protocols, cell columns were initially perfused with a baseline perfusate consisting of HBSS (pH 7.4), 25 mM HEPES, and 0.5% albumin. After a column equilibration period of 15 min, duplicate measurements of cell-column permeability were made. The column perfusate was then switched to HBSS containing the test agent(s) as described in Hatelets and pLatelet rezeasate. PlateZets and platelet releasate. Freshly prepared platelet suspensions containing 6,000 platelets/p1 in HBSS were applied to the column. To conserve platelets in these experiments the column effluent was returned to a lo-ml platelet reservoir. The effect of platelets on permeability was measured by injecting tracers over the column in the presence of platelets, as described in CelLcolumn methods. Platelet contents of the recirculating
Downloaded from www.physiology.org/journal/ajplung by ${individualUser.givenNames} ${individualUser.surname} (130.070.008.131) on January 16, 2019.
L672
PLATELET-RELEASED
FACTOR
ENHANCES
perfusate was monitored by removal of a 50 ~1 sample of perfusate and counting platelets with a Coulter counter (Coulter Instruments). During the measurement of permeability the effluent was switched to the fraction collector. In six experiments, the platelet containing perfusate was removed and replaced by baseline perfusate and permeability reassessed. In two experiments, this addition and removal of platelet containing solution was repeated again. Frozen aliquots of platelet releasates were thawed and diluted to the equivalent volume of releasate from 6,000 platelets/pi. These dilute solutions were applied to the cell-column in the same manner as the platelet suspensions. Platelet releasates were not recirculated to the column reservoir. Serum. Since serum contains significant quantities of platelet factors, we tested the permeability effect of switching from baseline column perfusate (HBSS) containing 10% serum to baseline column perfusate containing 0.5% bovine serum albumin. Measurements of permeability were recorded under perfusion with HBSS plus 10% bovine serum albumin, the perfusate was switched to our normal baseline perfusate containing HBSS and 0.5% bovine serum albumin and measurements of permeability were repeated. Thrombin-stimulated platelet releasate. Releasate from thrombin-stimulated platelets was prepared by adding 0.1 U of thrombin to each ml of platelet suspension. Thrombin stimulation was carried out for 30 min and PPACK (10e5 M) added to inactivate thrombin. Releasate was collected after spinning at 200 g for 15 min. Adenosine, ADP, ATP, serotonin. After baseline measurements of permeability, adenosine (10V4 M), ADP (lOA M), ATP ( 10m5M), and serotonin ( 10e3 M) were tested for permeability effects by adding them to the cell-column perfusates for 15 min and remeasuring the permeability of the endothelial monolayers. These doses were chosen to exceed the maximum published release levels achievable with 6000 platelets/p1 (18). Receptor antagonists/degradation enzymes. These protocols were designed to inactivate specific factors that might be active in platelet releasates. The effect of adenosine deaminase (2.5 U/ml for 30 min on previously frozen platelet releasate) on platelet releasate activity was tested by measuring permeability under the following four consecutive perfusate conditions: baseline, platelet releasate, adenosine deaminase alone, and releasate treated with adenosine deaminase. The effect of 8phenyltheophylline @PT, 10m5 M) on adenosine receptor blockade was tested by measuring permeability under the following five consecutive perfusate conditions: baseline, platelets, baseline, 8PT alone, 8PT plus platelets. The effect of degradation of ATP and ADP by apyrase (20) (10 U/ml; 30 min incubation with previously frozen releasate) was tested by measuring permeability under the following four consecutive conditions: baseline, platelet releasate, baseline, apyrase-treated platelet releasate. The importance of ,&receptor activation was tested by measuring permeability under the following four consecutive conditions: baseline, platelets, baseline, and platelets plus low5 M propran0101. Effects of heat, trypsin, and filtration on releasate activity. Previously frozen releasates were boiled, trypsin treated, and size-filtered to further characterize some of the properties of the active factor found in platelet releasates. Concentrated platelet releasate was heated at 100°C for 30 min to inactivate heat sensitive factors. Heated releasates were centrifuged at 10,000 g for 2 min and the releasate decanted. Activity of these treated releasates was tested by measuring permeability under baseline and heated platelet releasate conditions. Concentrated frozen releasates were treated with trypsin to degrade polypeptide factors (6). One milliliter of releasate was treated with trypsin (0.4 mg/ml) for 30 min at 37°C. Trypsin was neutralized with soybean inhibitor at a final concentration of (0.1 U/ml). The treated sample was then diluted to a con-
ENDOTHELIAL
BARRIER
centration equivalent to 6,000 platelets/p1 and activity tested by measuring permeability under the following four consecutive conditions: baseline, platelet releasate, baseline, and trypsintreated platelet releasate. Partial size characterization was done using microconcentrator- 100 spin columns according to manufacturers instructions (Amicon). Previously frozen platelet releasates were applied to the upper chamber of a C-100 column and spun at 1,000 g for 60 min. The filtrate and retentate were diluted by the same amount to reach a dilution equivalent to 6,000 platelet@. The activity of these fractions was tested by measuring permeability under the following six consecutive conditions: baseline, platelet releasate, baseline, X00-kDa releasate fraction, baseline, and
AND J. S. ALEXANDER
of Biomedical
Engineering,
Vanderbilt
Haselton, F. R., and J. S. Alexander. Platelets and a platelet-released factor enhance endothelial barrier. Am. J. Physiol. 263 (Lung Cell. Mol. Physiol. 7): L670-L678, 1992.The role of platelets in the maintenance of endothelial barrier is examined in an in vitro model of the microvasculature. Human platelets (6,000/~1) perfused through a cell column of endothelial-covered microcarriers decrease paracellular permeability of sodium fluorescein (mol wt 342) to 63% of baseline values. This effect is reversible and a second application and removal of platelets produces a similar response. This effect occurs within 5 min and reverses within 10 min after platelet removal. The reduction in permeability is not due to mechanical obstruction of endothelial junctions, since the number of recirculating platelets is not reduced and releasate from unstimulated 2-h platelet incubations also decreases permeability. Releasate from platelets stimulated with 0.1 U/ml of thrombin for 15 min have the same permeability reducing effect. In this system, the platelet factors serotonin ( 10e3 M) and ADP ( 10S4 M) have no effect on permeability. However, the platelet factors adenosine ( 10v4 M), ATP ( 10b5M), and ,&agonists decrease permeability. None of these appear to account for platelet permeability activity, since activity is not blocked by agents directed against these mediators (adenosine deaminase, apyrase, S-phenyltheophylline, or propranolol). The active factor(s) is stable at -2O”C, heat stable, sensitive to trypsin, and has an apparent molecular weight X00. We conclude that unstimulated platelets release a factor(s) that enhances endothelial barrier in vitro and may be important in maintenance of the normal in vivo barrier. paracellular permeability; serotonin; adenosine; adenosine 5’triphosphate; adenosine 5’-diphosphate
of studies have used cultured endothelial monolayers as a model system to study transvascular capillary exchange (1, 11,15,15a, 19,30,34). It has been observed in these systems that some physiological mediators, e.g., ,&agonists, bradykinin, thrombin, and histamine have direct effects on endothelial monolayer permeability in vitro (5, 11, 15, 21). Although not completely analogous, the responsiveness of these pure endothelial models continues to make this approach valuable in investigations of endothelial barrier. One consistent difference between in vitro and in vivo systems is the finding that in vitro endothelial monolayers have permeabilities that are from IO-1,000 times higher than the transvascular permeabilities values measured in vivo (1, 15). By definition, in vitro systems are incomplete, and it has been suggested that the permeability differences between these systems may be the lack of other vessel wall layers (26), differences in the properties of the in vitro extracellular matrix (7), or use of large vessel endothelial cells to model microvascular phenomena (4). Another possibility is that the in vitro tissue culture systems lack some essential circulating component of the vascular system that normally contributes to vessel integrity. Platelets are an important vascular component that
A NUMBER
L670
factor
1040-0605/92
$2.00
Copyright
University,
Nashville,
Tennessee
37235
have been shown to be involved in thrombosis, hemostasis, and the maintenance of vascular integrity. Experimental or pathological loss of circulating platelets (thrombocytopenia) is associated with increased vascular fragility and erythrocyte extravasation (2, 12, 16, 23, 28,32,33). However, it is unclear whether platelets contribute to the endothelial barrier in vivo. One possibility is that vasoactive compounds stored in platelets may promote barrier. Platelet contents are normally released by either slow leakage, e.g., 5-hydroxytryptamine (5HT), or stimulation that results in rapid release of CYand dense granules storage pools (17, 18). Recently, platelets have been shown to decrease the permeability of albumin across an endothelial monolayer separating two chambers (23, 31). In the endothelial model employed in that study (3l), the effect was not due to physical blockade, and a similar permeability effect was observed with either platelet lysate supernatant or thrombin stimulated platelets. The unknown active factor was not a P-agonist, serotonin, or a cyclooxygenase product (3 1). We have recently described cell-column chromatography as another in vitro model useful in studies of transvascular exchange (14,15). One of the unique properties of our approach is that it utilizes continuously flowing systems both in the culture system and in the design of the cell columns used to measure endothelial permeability. This method also finds permeabilities that are 10 times higher than measured in vivo values (15). In this study, we explore the possibility that circulating platelets are an important barrier enhancing component missing from in vitro systems. We measure the change in endothelial monolayer permeability produced by adding platelets and platelet releasate to our flowing in vitro system and partially characterize the responsible platelet factor(s). METHODS CeZl culture methods. Well-established techniques were used to isolate and culture bovine aortic endothelial cells (BAEC) and fetal aortic endothelial cells (BFAE); detailed procedures used for primary cell isolation, cell identification, monolayer culture subcultivation, and the determination of in vitro lifespan have been described previously (15, 27). Briefly, cultures of adult and fetal aortic endothelial cells were obtained from thoracic aortae by means of 0.1% collagenase treatment. Monolayer cultures were established in Dulbecco’s modified Eagle’s medium containing 10% fetal bovine serum and were subcultured using 0.1% trypsin-EDTA. Endothelial cell identity was verified by indirect immunofluorescent assay for factor VIII-related antigen and diI-low-density lipoprotein (LDL). Cell lines were used during the vigorous proliferative phase of their in vitro lifespan. The fetal bovine cell lines (AG-7680 and AG7681) are available from the Cell Repository at the Coriell Institute for Medical Research (Camden, NJ).
0 1992 The
American
Physiological
Society
Downloaded from www.physiology.org/journal/ajplung by ${individualUser.givenNames} ${individualUser.surname} (130.070.008.131) on January 16, 2019.
PLATELET-RELEASED
FACTOR
ENHANCES
Cells were cultured on microcarrier beads as previously described (15). Cells were seeded on Cytodex-3 microcarrier beads at a density of 2 x IO4 cells/cm2. Cell attachment was achieved by intermittent stirring overnight. Microcarrier cultures were maintained at 60 rpm continuously and fed three times a week. Cultures were used for these assays between 9 and 30 days postseeding. Unless otherwise noted, all biological and tissue culture supplies were purchased from Sigma Chemical (St. Louis, MO). Platelet isolation methods. Human platelets were obtained from normal volunteers and isolated by the two methods described below (8, 35). Subjects were nonsmokers, drug and aspirin free for at least 3 days before platelet donation. Sixty milliliters of blood was withdrawn through an extension tube using a 20-gauge needle into a syringe containing sodium citrate. In one method (method A) whole blood was spun for 30 min at 300 g over a Ficoll-hypaque density gradient (Flow Labs). The platelet-rich plasma was resuspended in Ca2+ free N2-hydroxyethylpiperazine-N’-2-ethanesulfonic acid (HEPES) buffer containing 0.35% bovine serum albumin. This platelet suspension was spun 15 min at 100 g to remove leukocytes and red blood cells. A second method (method B) developed by Timmons and Hawiger (35) was also used to remove adventitiously adherent serum proteins. Whole blood was spun for 15 min at 250 g. The platelet-rich plasma was layered over a lo-25% albumin gradient and spun 15 min at 100 g. Platelets were removed and resuspended in Ca 2+-free HEPES buffer and passed over a Sepharose 2B gel column (35). Platelet releasates were prepared by incubating platelet suspensions at room temperature for 120 min in Hanks’ balanced salt solution (HBSS) containing 0.5% bovine serum albumin and 25 mM HEPES. Platelets were removed from suspension by centrifugation at 100 g for 15 min and 0.5-ml aliquots were frozen at -20°C in polypropylene Eppendorf tubes. PLateLet activation assay. Activation of platelets was assessed by their ability to release radiolabeled serotonin on stimulation with 10m6 M phorbol myristate acetate (PMA). Platelets were loaded with 1.5 &i/ml of [14C]serotonin (Amersham). Serotonin release was measured by comparing the releasate from spun platelets with the total activity of Triton X-100 (1%) treated platelets. Platelet activation was achieved with 10V6 M PMA for 10 min, and the release was compared with that found in unstimulated conditions. Cell-column methods. We used a previously reported assay of endothelial monolayer permeability, with modifications, to measure changes in barrier produced by stimulation of endothelial monolayers with experimental treatments (15). The method uses a model of the vasculature consisting of a chromatographic cell-column filled with cell-covered microcarrier beads. The permeability of the endothelial monolayers covering the beads is determined from a comparison of the elution curves of tracers injected into the flow at the top of the column. The details of this method have been previously described (15), and this approach, with modifications, is briefly described below. Chromatographic cell columns were made from water-jacketed glass columns (0.65-cm diam, Rainin). Cell-covered beads were poured to a column height of -2 cm, which provided 130 cm2 of endothelial cell culture surface, or -1 X IO7 cells. The column was washed and equilibrated with HBSS containing 0.5% albumin. Perfusion through the column was maintained by a Gilson peristaltic pump, at 0.9 ml/min. This flow was chosen to approximate the gravity flow rate observed when the pump was not connected. A tracer bolus described below was applied by a rotary injection valve (Rainin) using a 50-~1 loop. The cell column and all perfusate solutions were kept at 37°C.
ENDOTHELIAL
BARRIER
L671
Multiple tracer indicator-dilution analysis was used to obtain cell layer permeability from the relative shapes of the elution profiles of tracers simultaneously applied to the top of a cell column. One of the applied tracers (Blue Dextran, 10 mg/ml, 2,000 kDa) is impermeant and follows the mobile phase, i.e., a flow tracer. Two other tracers, initially sodium fluorescein (mol wt 342), and in later experiments sodium fluorescein and cyanocobalamin (mol wt 1,355), were used as permeant tracers, able to permeate beneath the cell layer and diffuse into the bead matrix beneath the cells. Using only optical tracers allows us to collect and measure the concentration of three optical tracers simultaneously without having to rely on radioactive materials previously employed and also has the advantage of using a single instrument to quantify the tracer concentrations contained within each sample. Elution profiles were constructed from 66 samples of the column eluant using a modification of previously reported methods (15). A Gilson fraction collector (model 203) equipped with a drop counter was used to collect two drops of eluant per well for 66 wells of a 96-well microtiter plate. The absorbance of each of the 96 wells was read at 620,540, and 492 nm on a plate reader (Titertek MCC 340) and stored on a computer for analysis. The optical absorbances were used to calculate the fractional recovery per sample of each of the optically absorbing dyes. Estimates of permeability based on the column-elution patterns of multiple tracers is an adaptation and extension of techniques used in vivo to assess capillary permeability (13,29). To apply these techniques to these experiments, a mathematical model of tracer motion has been developed based on the physical picture of tracer behavior described above. In this previously described model (15), it is assumed that the elution profiles of cyanocobalamin and sodium fluorescein depend on the properties of the mobile phase of the column plus the paracellular permeability properties of the endothelial monolayer and the diffusive motion of the tracer within the microcarrier beads. In contrast to these permeant tracers, the elution of Blue Dextran depends only on the flow (mobile) phase properties of the column. A modified Marquardt iteration scheme was used to estimate monolayer permeability which best approximated the experimental data. Best fit was determined by the minimization of the coefficient of variation between a computer-generated prediction of the permeant tracer’s elution profile and the experimentally observed elution profile. A permeability value for any one column and time point was computed as the mean of two consecutive measurements (spanning 5-7 min). Significance among three or more column conditions was judged using one-way analysis of variance and Tukey’s modified t test. A P < 0.05 was judged significant. Treatment protocols. Consecutive permeability measurements on the same population of cells were made in duplicate during a number of consecutive experimental treatments. All treatment protocols followed this same basic design. In all protocols, cell columns were initially perfused with a baseline perfusate consisting of HBSS (pH 7.4), 25 mM HEPES, and 0.5% albumin. After a column equilibration period of 15 min, duplicate measurements of cell-column permeability were made. The column perfusate was then switched to HBSS containing the test agent(s) as described in Hatelets and pLatelet rezeasate. PlateZets and platelet releasate. Freshly prepared platelet suspensions containing 6,000 platelets/p1 in HBSS were applied to the column. To conserve platelets in these experiments the column effluent was returned to a lo-ml platelet reservoir. The effect of platelets on permeability was measured by injecting tracers over the column in the presence of platelets, as described in CelLcolumn methods. Platelet contents of the recirculating
Downloaded from www.physiology.org/journal/ajplung by ${individualUser.givenNames} ${individualUser.surname} (130.070.008.131) on January 16, 2019.
L672
PLATELET-RELEASED
FACTOR
ENHANCES
perfusate was monitored by removal of a 50 ~1 sample of perfusate and counting platelets with a Coulter counter (Coulter Instruments). During the measurement of permeability the effluent was switched to the fraction collector. In six experiments, the platelet containing perfusate was removed and replaced by baseline perfusate and permeability reassessed. In two experiments, this addition and removal of platelet containing solution was repeated again. Frozen aliquots of platelet releasates were thawed and diluted to the equivalent volume of releasate from 6,000 platelets/pi. These dilute solutions were applied to the cell-column in the same manner as the platelet suspensions. Platelet releasates were not recirculated to the column reservoir. Serum. Since serum contains significant quantities of platelet factors, we tested the permeability effect of switching from baseline column perfusate (HBSS) containing 10% serum to baseline column perfusate containing 0.5% bovine serum albumin. Measurements of permeability were recorded under perfusion with HBSS plus 10% bovine serum albumin, the perfusate was switched to our normal baseline perfusate containing HBSS and 0.5% bovine serum albumin and measurements of permeability were repeated. Thrombin-stimulated platelet releasate. Releasate from thrombin-stimulated platelets was prepared by adding 0.1 U of thrombin to each ml of platelet suspension. Thrombin stimulation was carried out for 30 min and PPACK (10e5 M) added to inactivate thrombin. Releasate was collected after spinning at 200 g for 15 min. Adenosine, ADP, ATP, serotonin. After baseline measurements of permeability, adenosine (10V4 M), ADP (lOA M), ATP ( 10m5M), and serotonin ( 10e3 M) were tested for permeability effects by adding them to the cell-column perfusates for 15 min and remeasuring the permeability of the endothelial monolayers. These doses were chosen to exceed the maximum published release levels achievable with 6000 platelets/p1 (18). Receptor antagonists/degradation enzymes. These protocols were designed to inactivate specific factors that might be active in platelet releasates. The effect of adenosine deaminase (2.5 U/ml for 30 min on previously frozen platelet releasate) on platelet releasate activity was tested by measuring permeability under the following four consecutive perfusate conditions: baseline, platelet releasate, adenosine deaminase alone, and releasate treated with adenosine deaminase. The effect of 8phenyltheophylline @PT, 10m5 M) on adenosine receptor blockade was tested by measuring permeability under the following five consecutive perfusate conditions: baseline, platelets, baseline, 8PT alone, 8PT plus platelets. The effect of degradation of ATP and ADP by apyrase (20) (10 U/ml; 30 min incubation with previously frozen releasate) was tested by measuring permeability under the following four consecutive conditions: baseline, platelet releasate, baseline, apyrase-treated platelet releasate. The importance of ,&receptor activation was tested by measuring permeability under the following four consecutive conditions: baseline, platelets, baseline, and platelets plus low5 M propran0101. Effects of heat, trypsin, and filtration on releasate activity. Previously frozen releasates were boiled, trypsin treated, and size-filtered to further characterize some of the properties of the active factor found in platelet releasates. Concentrated platelet releasate was heated at 100°C for 30 min to inactivate heat sensitive factors. Heated releasates were centrifuged at 10,000 g for 2 min and the releasate decanted. Activity of these treated releasates was tested by measuring permeability under baseline and heated platelet releasate conditions. Concentrated frozen releasates were treated with trypsin to degrade polypeptide factors (6). One milliliter of releasate was treated with trypsin (0.4 mg/ml) for 30 min at 37°C. Trypsin was neutralized with soybean inhibitor at a final concentration of (0.1 U/ml). The treated sample was then diluted to a con-
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centration equivalent to 6,000 platelets/p1 and activity tested by measuring permeability under the following four consecutive conditions: baseline, platelet releasate, baseline, and trypsintreated platelet releasate. Partial size characterization was done using microconcentrator- 100 spin columns according to manufacturers instructions (Amicon). Previously frozen platelet releasates were applied to the upper chamber of a C-100 column and spun at 1,000 g for 60 min. The filtrate and retentate were diluted by the same amount to reach a dilution equivalent to 6,000 platelet@. The activity of these fractions was tested by measuring permeability under the following six consecutive conditions: baseline, platelet releasate, baseline, X00-kDa releasate fraction, baseline, and
