Src-dependent, neutrophil-mediated vascular hyperpermeability and beta -catenin modification

John H. Tinsley, Elena E. Ustinova, Wenjuan Xu, and Sarah Y. Yuan

Departments of Surgery and Medical Physiology, Cardiovascular Research Institute, Texas A&M University System Health Science Center, Temple, Texas 76504


    ABSTRACT
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ABSTRACT
INTRODUCTION
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The hyperpermeability response of microvessels in inflammation involves complex signaling reactions and structural modifications in the endothelium. Our goal was to determine the role of Src-family kinases (Src) in neutrophil-mediated venular hyperpermeability and possible interactions between Src and endothelial barrier components. We found that inhibition of Src abolished the increases in albumin permeability caused by C5a-activated neutrophils in intact, perfused coronary venules, as well as in cultured endothelial monolayers. Activated neutrophils increased Src phosphorylation at Tyr416, which is located in the catalytic domain, and decreased phosphorylation at Tyr527 near the carboxyl terminus, events consistent with reports that phosphorylating and transforming activities of Src are upregulated by Tyr416 phosphorylation and negatively regulated by Tyr527 phosphorylation. Furthermore, neutrophil stimulation resulted in association of Src with the endothelial junction protein beta -catenin and beta -catenin tyrosine phosphorylation. These phenomena were abolished by blockage of Src activity. Taken together, our studies link for the first time neutrophil-induced hyperpermeability to a pathway involving Src kinase activation, Src/beta -catenin association, and beta -catenin tyrosine phosphorylation in the microvascular endothelium.

permeability; transfection; phosphorylation; endothelium; microvessels


    INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

VASCULAR ENDOTHELIUM serves as an effective barrier to control the transvascular passage of solutes, fluid, and blood cells. Alterations in the barrier function are involved in thrombogenesis, angiogenesis, inflammation, and ischemia-reperfusion injury. Binding of inflammatory agonists and cells to endothelial cells elicits a multitude of intracellular signaling events, which can result in an increase in endothelial permeability (12, 19, 30, 34). Actomyosin interaction generates contractile forces that pull tightly connected endothelial cells apart, leading to macromolecular efflux. A transmembrane adhesive protein called vascular endothelial (VE)-cadherin forms endothelial adherens junctions (AJ) that inhibit the paracellular leakage of macromolecules. When the equilibrium between these adhesive and contractile forces is altered, barrier dysfunction and leakage occur.

Previous studies have shown that a group of inflammatory cells, polymorphonuclear leukocytes (PMNs), adhere to and migrate through the endothelium into surrounding tissues at sites of injury or inflammation and that this process is associated with permeability increases (1, 2, 35). We have demonstrated that PMN-induced hyperpermeability occurs concomitantly with an increase in tyrosine phosphorylation of VE-cadherin and beta -catenin, an important member of a family of proteins that links the cadherin complex to the actin cytoskeleton (14, 16, 27). Recently, studies have shown an apparent link between beta -catenin and the Src-family tyrosine kinases (Src) (17). The Src kinases are known to play a role in signaling transduction of endothelial barrier dysfunction and angiogenesis (9, 13, 15, 24). Src activity is regulated by tyrosine phosphorylation at Tyr416, which upregulates the kinase, and Tyr527, which renders Src less active (24).

Although a copious amount of work has sought to clearly define the mechanisms responsible for PMN-induced hyperpermeability (1, 3, 27), the molecular targets of activated PMNs and consequential events in the venular endothelium where leakage occurs are still poorly understood. This study focuses on possible interactions among PMNs, tyrosine kinase pathways, and junctional proteins in the regulation of endothelial cell barrier function. Through transfection of a specific peptide (SRCi) or administration of PP1, a Src family kinase inhibitor, we were able to inhibit Src activity; this inhibition significantly attenuated PMN-induced hyperpermeability responses in both isolated coronary venules and cultured endothelial cells. A similarly negatively charged and phosphorylated peptide used as a control had no effect on hyperpermeability. We show that activated PMNs alter the phosphorylation status of Src with an increase in phosphorylation at Tyr416 and a decrease at Tyr527. Furthermore, these phosphorylation changes are blocked with the addition of SRCi. Immunoprecipitation shows that under PMN-stimulated conditions, the AJ protein beta -catenin associates with Src. This association occurs concomitantly with beta -catenin tyrosine phosphorylation and loss of beta -catenin at the cell periphery, and all three of these events are abolished upon Src inactivation. Taken together, our data demonstrate for the first time a link among PMNs, the Src tyrosine kinase pathway, and AJ in the regulation of endothelial cell barrier integrity.


    METHODS
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INTRODUCTION
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Chemicals and drugs. The chemicals used were human recombinant C5a and phenylarsine oxide (PAO; Sigma), SRCi and PP1 (Calbiochem), polyclonal anti-Src and polyclonal anti-beta -catenin (Santa Cruz), polyclonal antiphospho-Src (Tyr416) and (Tyr527) (Cell Signaling), and polyclonal antiphosphotyrosine (Transduction). SRCi [Ac-Tyr(PO3H2)-Tyr(PO3H2)-Tyr(PO3H2)-Ile-Glu-OH] competes for binding to SH2 domains (31). Negative control peptide was [Ac-Asp-Ser(PO3H2)-Thr(PO3H2)-Val-Ser(PO3H2)-OH].

Isolation and perfusion of microvessels. Pigs weighing 9-13 kg were sedated with ketamine (2.5 mg/kg im) and rompun (2.25 mg/kg im), anesthetized with pentobarbital sodium (25 mg/kg iv), and treated with heparin (250 U/kg iv). After a left thoracotomy, the heart was electrically fibrillated, excised, and placed in 4°C physiological saline. The left descending artery was cannulated, and 3 ml of India ink-gelatin-physiological salt solution were infused to clearly define microvessels. The methods for isolating and cannulating coronary microvessels have been described previously (28, 33, 34, 36). Briefly, venules (0.8-1.2 mm, diameter 20-50 µm) were dissected and transferred to a cannulating chamber mounted on a Zeiss Axiovert 135 inverted microscope. The vessel was cannulated with a micropipette on each end and secured with suture; a third, smaller pipette was inserted into the inflow pipette. The vessel was perfused with either albumin-physiological salt solution (APSS) through the outer inflow pipette or APSS containing FITC-albumin through the inner inflow pipette. The micropipettes were connected to a reservoir that allowed intraluminal pressure and flow velocity to be adjusted. The vessel image was displayed on a video monitor, diameter was measured with a video caliper, and flow velocity was measured with an optical Doppler velocimeter (Microcirculation Research Institute, Texas A&M University, College Station, TX). Permeability was quantified by measuring FITC-albumin fluorescence in the vessel and in the surrounding area (33, 34, 36). The apparent solute permeability coefficient of albumin (Pa) was calculated using the equation Pa = (1/Delta If)(dIf/dt)0(r/2), where Delta If is the initial step increase in fluorescent intensity, (dIf/dt)0 is the initial rate of gradual increase in intensity as solutes diffuse out of the vessel into the extravascular space, and r is the venular radius. The venule was perfused at a constant pressure of 10 cmH2O and a flow velocity of 7 mm/s, and samples were not used if leakage of FITC-albumin was detected.

Endothelial cell assays. For permeability studies, human umbilical vein endothelial cells (HUVEC; Clonetics) were grown on gelatin-coated Costar transwell membranes (VWR) as previously described (27). Cells were exposed to porcine PMN for 10 min, transfections were carried out for 1 h, and PP1 and DMSO treatments were 10 min. After these treatments and in the presence of transfection reagent, PMNs, and all other reagents, FITC-albumin was added to the luminal chamber for 30 min, and samples were collected from both the luminal and abluminal chambers for fluorometry analysis. Readings were converted with a standard curve to albumin concentrations, which were then used in the following equation to determine Pa: Pa = [A]V/tA[L]; where [A] is abluminal concentration, t is time in seconds, A is area of membrane in cm2, V is volume of abluminal chamber, and [L] is luminal concentration. For Western analysis, cells were grown on gelatin-coated 60-mm dishes, treated as appropriate, and lysed. Neutrophils were washed from the dishes before lysis. In each lane of a 6% polyacrylamide SDS-PAGE, 10 µg of total protein were electrophoresed, followed by transfer to nitrocellulose membrane. After exposure of primary antibody and secondary antibody conjugated to horseradish peroxidase (HRP), bands were detected by using enhanced chemiluminescence. Bands were quantitated by scanning densitometry. For immunoprecipitation, 100 µg of total protein were incubated with either anti-Src, anti-beta -catenin, or anti-phosphotyrosine antibodies followed by protein A/G conjugated to agarose beads to isolate the protein(s) of interest before subjection to PAGE. For beta -catenin localization, cells were grown to confluence on coverslips and treated in the same manner as that for Western analysis. After fixation and permeabilization, anti-beta -catenin antibody was applied, followed by secondary antibody conjugated to FITC. The coverslips were mounted on slides for microscopic observation.

Neutrophil isolation. Porcine neutrophils (PMN) were isolated as previously described (27). To activate, PMNs were exposed to human recombinant C5a (10-8 M). In the isolated venule preparations, PMNs were added to the suffusion bath and, in the case of HUVEC studies, were added directly on the monolayer, in both instances at a concentration of 106/ml. Previous studies have shown that C5a affects endothelial function via neutrophil-dependent pathways yet does not affect permeability by itself (27).

Protein transfection. To transfect venules with SRCi, vessels were perfused for 1 h with the peptide at 10 µg/ml in the presence of the polyamine transfection reagent TransIT-LT1 (PanVera) at 10 µl/ml. Previous transfection studies using green fluorescent protein and various inhibiting peptides had shown this to be a suitable way to introduce proteins/peptides into intact microvessels (28). Additionally, the TransIT-LT1 alone has no apparent effects on microvascular permeability or vasoreactivity (28). For HUVEC transfection, the SRCi and TransIT-LT1 were added to the cell medium at the same concentrations, and transfection was allowed to proceed for 1 h. Previous studies have shown successful protein transfection of endothelial cells using this technique (26).

Data analysis. In the immunoblot studies, a representative image of Western blots was selected to present. At least three repetitions were performed for each intervention, and optical densities of the protein bands were averaged. Analysis of variance was used to evaluate the significance of intergroup differences in the immunoblot analyses and permeability studies. A value of P < 0.05 was considered significant for the comparisons.


    RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

PMN-induced hyperpermeability in venules. To determine whether Src is involved in neutrophil-mediated hyperpermeability, coronary venules were transfected with SRCi or exposed to PP1, a cell-permeable Src inhibitor. As shown in Fig. 1c, when activated with C5a, PMNs induce a twofold increase in permeability over basal levels. This increase is not observed with the addition of unstimulated PMNs or C5a alone (Fig. 1, a and b). Interestingly, when the SRCi is transfected for 1 h before the addition of activated PMNs, the hyperpermeability response is attenuated to near basal levels (Fig. 1d). Transfection of a similarly negatively charged and phosphorylated control peptide (negative control peptide) had no significant effect on PMN-induced hyperpermeability (Fig. 1e). Furthermore, addition of the cell-permeable Src inhibitor PP1 blocked PMN-induced hyperpermeability (Fig. 1f), whereas the PP1 vehicle DMSO had no effect (Fig. 1g).


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Fig. 1.   Polymorphonuclear leukocytes (PMN)-induced venular hyperpermeability. Venules were perfused with albumin-physiological salt solution (APSS), and basal permeability coefficient of albumin (Pa) was determined over a 1-h period (first bar in each group). Results are expressed as a percentage of basal. Treatments were as follows: a: unactivated porcine PMN (106/ml), b: C5a (10-8 M), c: C5a activated PMN, d: SRCi transfected for 1 h followed by C5a activated PMN, e: negative control peptide transfected for 1 h followed by C5a activated PMN, f: PP1 (10-6 M) followed by C5a activated PMN, and g: DMSO (0.05%) followed by C5a activated PMN. Values are means ± SE. * Significant increase in basal permeability, P < 0.05; n = 4 for each treatment.

PMN-induced hyperpermeability in cultured endothelial cells. To determine that the permeability response of endothelial cell monolayers is in agreement with that of intact venules, HUVECs were exposed to activated PMNs after transfection with SRCi or negative control peptide or exposure to PP1. As shown in Fig. 2b, activated PMNs induced significant hyperpermeability responses above control levels. In addition, the supernatant obtained after activated PMNs were centrifuged had a similar effect on permeability (Fig. 2c). However, SRCi completely abolished these permeability increases (Fig. 2e). Transfection of the negative control peptide did not block PMN-induced hyperpermeability (Fig. 2g). In agreement with the effects of SRCi, PP1 also blocked PMN-induced hyperpermeability (Fig. 2i), whereas the vehicle DMSO had no effect (Fig. 2k).


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Fig. 2.   Hyperpermeability response in human umbilical vein endothelial cells (HUVEC). Treatments were as follows: a: control, b: C5a activated porcine PMN (106/ml), c: supernatant from activated PMN, d: SRCi, e: SRCi followed by activated PMN, f: negative control peptide, g: negative control peptide followed by activated PMN, h: PP1 (10-6 M), i: PP1 (10-6 M) followed by activated PMN, j: DMSO (0.05%), and k: DMSO followed by activated PMN. Pa was measured after 30 min, and results are expressed as a percentage of control. Values are means ± SE. * Significant increase in permeability, P < 0.05; n = 4 for each treatment.

Src tyrosine phosphorylation. As stated previously, Src is a tyrosine kinase that itself is regulated by tyrosine phosphorylation. Using HUVECs, we show that activated PMNs and PAO (a tyrosine phosphatase inhibitor) induce Src phosphorylation at Tyr416 (Fig. 3A, lanes 4 and 7), a condition known to activate Src. This Tyr416 phosphorylation was attenuated in cells that had been transfected with the SRCi (Fig. 3A, lanes 6 and 8). Figure 3B, lanes 4 and 7, shows that when Src is activated by PMNs or PAO, phosphorylation at Tyr527 is decreased. Band intensity from three different experiments was obtained by using scanning densitometry followed by quantitation using National Institutes of Health image software. This data showed that the phosphorylation changes of Src Tyr416 and Tyr527 in response to activated PMNs and PAO are significantly different than those of control levels (Fig. 3, D and E). The amount of Src present in the cells does not appear to vary significantly under any of the test conditions (Fig. 3C).


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Fig. 3.   Src phosphorylation in HUVECs. Treatments were as follows: 1: control, 2: C5a (10-8 M), 3: PMN (106/ml), 4: C5a activated PMN (106/ml), 5: SRCi transfected for 1 h, 6: SRCi followed by activated PMN (106/ml), 7: phenylarsine oxide (PAO) (10-8 M), and 8: SRCi followed by PAO (10-8 M). Treatments were for 10 min. After cell lysis, 10 µg of protein from each treatment were subjected to Western analysis on a 6% PAGE gel. Antibodies used were as follows: A: anti-phospho Src (Tyr416), B: anti-phospho Src (Tyr527), and C: anti-Src. D and E: bands from A and B, respectively, were quantitated by scanning densitometry, and Src Tyr416 (D) and Tyr527 (E) phosphorylation was expressed as percentage of control. These experiments were repeated 3 times. Values are means ± SE. *P < 0.05 vs. control.

Src activation and beta -catenin. Our previous studies (27) had shown a link between activated PMNs and associated AJ disorganization, whereas others had shown links between AJ proteins and Src (17). Therefore, we wanted to determine whether interaction among all three of these factors could be detected. First, we found that the AJ protein beta -catenin localizes to the cell periphery in confluent monolayers (Fig. 4A). When monolayers are exposed to activated PMNs, intercellular gaps are formed and beta -catenin staining is lost in areas where the cells no longer contact each other (Fig. 4B). Supernatant from activated PMNs also induced gap formation (Fig. 4C). Exposure to PP1 or transfection of SRCi attenuated PMN-induced gap formation (Fig. 4, D and F). We found that PMN-induced gap formation is reversible; that is, when the PMNs are washed away from the cells, the monolayer regains its cell-cell contacts (Fig. 4G). Finally, transfection of a negative control peptide did not prevent PMN-induced gap formation (Fig. 4H). Next, immunoprecipitation and immunoblotting studies showed that upon PMN stimulation of HUVECs, beta -catenin interacts with Src, and this interaction can be abolished by transfection of SRCi (Fig. 5A). Additionally, PMN-induced beta -catenin tyrosine phosphorylation is decreased when Src is inhibited (Fig. 5, C and D), whereas beta -catenin protein levels remain constant (Fig. 5E). These findings strongly suggest an interaction among activated PMNs, Src, and beta -catenin.


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Fig. 4.   beta -catenin localization in HUVEC. A: control cells; B: cells exposed to C5a-activated PMN; C: supernatant from C5a-activated PMN; D: PP1 (10-6 M) followed by activated PMN; E: SRCi transfected for 1 h; F: SRCi followed by C5a-activated PMN; G: C5a-activated PMN exposure, wash with PBS, return to complete media for 1 h; and H: negative control peptide followed by C5a-activated PMN. After fixation and permeabilization, cells were incubated with anti-beta -catenin primary antibody followed by FITC-conjugated secondary antibody. Note gap formation and loss of beta -catenin at the sites where cells have lost contact in B, C, and H.



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Fig. 5.   PMN-induced Src/beta -catenin association in HUVEC. For A and B, treatments were as follows: 1: control, 2: C5a activated PMN (106/ml), and 3: SRCi transfected for 1 h followed by C5a activated PMN (106/ml). After PMN treatment for 10 min, 100 µg of protein were used for Src immunoprecipitation. The immunoprecipitates were subjected to Western analysis and probed for either beta -catenin (A) or Src (B). For C, D, and E, treatments were as follows: 1: control, 2: C5a activated PMN (106/ml), 3: SRCi transfected for 1 h followed by C5a activated PMN (106/ml), and 4: SRCi transfection. After PMN treatment for 10 min, 100 µg of protein were used for beta -catenin immunoprecipitation followed by phosphotyrosine immunoblotting (C) or phosphotyrosine immunoprecipitation followed by beta -catenin immunoblotting (D). In E, 15 µg of total cellular protein were used for beta -catenin immunoblotting.


    DISCUSSION
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INTRODUCTION
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PMN-induced microvascular leakage has long been linked to the development of circulatory dysfunction (10). This paper reports on the effects of PMNs on isolated porcine coronary venules and cultured HUVECs. Porcine and human PMNs have very similar oxidative and chemotactic responses, and both show increased migration upon stimulation by porcine- or human-derived C5a (5, 22, 25). Although there is some debate as to whether or not human PMNs lead to HUVEC dysfunction, this study and others (4, 11, 32) have shown that activated human PMNs do elicit HUVEC retraction and transendothelial protein movement. Interestingly, we found that physical attachment between PMNs and endothelial cells is not required for gap formation and hyperpermeability responses. On the other hand, loss of the junctional protein beta -catenin apparently plays a role in endothelial barrier dysfunction, as seen by our immunofluorescence studies. These phenomena were observed when supernatant from activated PMNs was applied to endothelial monolayers, suggesting that the physical attachment of PMNs to the endothelial surface or transmigration is not a prerequisite for endothelial hyperpermeability or AJ disorganization. This result is in disagreement with another study suggesting that PMN adhesion is necessary for induction of endothelial hyperpermeability and that PMN-derived proteases do not modify AJs (7). An explanation for the discrepancy is the different cellular preparations and experimental interventions. In particular, PMN activation was accomplished by using different agents, in our case human recombinant C5a and PMA in the case of Del Maschio et al. (7). The two stimulators may cause different endothelial responses. Importantly, our recent study on isolated venules supports the importance of PMN release of endothelial activators, rather than physical adherence to the endothelium, in the induction of microvascular hyperpermeability, because in the venule study we found that application of C5a-activated PMNs to the abluminal side of vessels in the absence of adhesion and migration caused a significant increase in permeability (37). In agreement with this, others have determined that activated PMNs release factors such as proteases, cationic proteins, and oxygen radicals that can induce hyperpermeability responses and degradation of catenins in the endothelium (6, 8, 20, 29).

Our previous studies had shown definitive effects of PMNs on vascular endothelium using both intact microvessels and cultured cells, in which activated PMNs cause phosphorylation and conformational changes of AJ proteins in association with intercellular gap formation (27, 35). However, signaling events occurring between PMN adhesion and AJ alteration are not well understood. This study, for the first time, links the well-known Src signaling pathway to these processes. We know that regarding Src tyrosine kinases, which have six functional domains, phosphorylation of Tyr527 and interactions between the SH2 and SH3 domains stabilize the inactive form of Src (21). Conversely, phosphorylation of Tyr416 in the activating loop of the kinase domain activates Src (21). Others have shown a Src requirement for vascular permeability in response to vascular endothelial growth factor (9). Our results clearly show that PMN-induced hyperpermeability in both microvessels and endothelial cells can be greatly attenuated through Src inhibition. Activated PMNs increased Src Tyr416 phosphorylation and decreased Tyr527 phosphorylation, two events that are known to upregulate Src activity. These findings suggest that Src is a major component in PMN-mediated endothelial barrier dysfunction.

The precise molecular mechanisms that lead to microvascular leakage after PMN adhesion are not clearly understood. Previous studies with endothelial monolayers have shown that activated PMNs induce actin stress fiber formation, in contrast to unstimulated cells in which most of the filamentous actin is found at the cell periphery (27). Apparently, these fibers contact opposite sides of the cell membrane and induce cellular contraction, which breaks the contacts between cells and leads to gap formation. Our hypothesis is that AJ proteins interact with actin stress fibers and this interaction leads to a disorganization of the AJ and changes in cellular morphology. Phosphorylation of beta -catenin, known to link VE-cadherin to the actin cytoskeleton (14), may be a crucial signaling event directing such structural changes. The proposed mechanism of beta -catenin tyrosine phosphorylation in AJ disorganization is parallel to the Wnt/beta -catenin signaling pathway in which Wnt is found to stabilize beta -catenin by blocking its serine/threonine phosphorylation and subsequent targeting for degradation, leading to beta -catenin nuclear localization and transcriptional activation (23). Conversely, absence of Wnt leads to beta -catenin serine/threonine phosphorylation and proteasomal degradation (18).

One goal of this study was an attempt to correlate activated PMNs with a signaling pathway involving beta -catenin that leads to alteration of AJ components. This study showed that under PMN-stimulated conditions, beta -catenin coimmunoprecipitated with Src. This apparent Src/beta -catenin association was completely blocked when cells were transfected with SRCi. Additionally, we were able to demonstrate that PMN-induced beta -catenin tyrosine phosphorylation is blocked under conditions of Src inhibition in cultured endothelial cells. Taken together, these results suggest that Src and beta -catenin interaction and phosphorylation are necessary for PMN-induced hyperpermeability. Perhaps it is Src kinase that directly phosphorylates beta -catenin in response to activated PMNs; this event leads to the disorganization of the AJ and ultimately endothelial barrier dysfunction. It is apparent that Src and beta -catenin are involved in multiple cellular processes, and further studies will attempt to more completely understand the connection between these two proteins and their interactions with other components central to barrier integrity in the microvascular endothelium.


    ACKNOWLEDGEMENTS

This work was supported by National Heart, Lung, and Blood Institute Grants HL-61507 and HL-70752 (to S. Y. Yuan) and a VA VISN 17 grant (to J. H. Tinsley). S. Y. Yuan is a recipient of National Institutes of Health Research Career Award K02 HL-03606.


    FOOTNOTES

Address for reprint requests and other correspondence: J. H. Tinsley, Dept. of Medical Physiology, Texas A&M Univ. System Health Science Center, 702 SW HK Dodgen Loop, Rm. 206F, Temple, TX 76504 (E-mail: jht{at}tamu.edu).

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.

July 24, 2002;10.1152/ajpcell.00230.2002

Received 20 May 2002; accepted in final form 17 July 2002.


    REFERENCES
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
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Am J Physiol Cell Physiol 283(6):C1745-C1751
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