June 2005
Volume 46, Issue 6
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Cornea  |   June 2005
LPA and S1P Increase Corneal Epithelial and Endothelial Cell Transcellular Resistance
Author Affiliations
  • Fei Yin
    From the Department of Physiology, University of Tennessee Health Science Center, Memphis, Tennessee.
  • Mitchell A. Watsky
    From the Department of Physiology, University of Tennessee Health Science Center, Memphis, Tennessee.
Investigative Ophthalmology & Visual Science June 2005, Vol.46, 1927-1933. doi:10.1167/iovs.04-1256
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      Fei Yin, Mitchell A. Watsky; LPA and S1P Increase Corneal Epithelial and Endothelial Cell Transcellular Resistance. Invest. Ophthalmol. Vis. Sci. 2005;46(6):1927-1933. doi: 10.1167/iovs.04-1256.

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      © ARVO (1962-2015); The Authors (2016-present)

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Abstract

purpose. To determine whether lysophosphatidic acid (LPA) or sphingosine-1-phosphate (S1P) affects transcellular resistance across cultured rabbit corneal epithelial and endothelial cells.

methods. Electric cell–substrate impedance sensing (ECIS) was used to measure electrical resistance across cultured rabbit corneal epithelial and endothelial monolayers. After a 1-hour equilibration period, different concentrations of LPA or S1P were added to each well, and the effect observed for 4 hours. For cells significantly affected by LPA or S1P, pertussis toxin (PTX) or dioctyl-glycerol pyrophosphate (DGPP 8:0) was added along with LPA or S1P in separate experiments. Cells were also treated with phorbol 12-myristate 13-acetate (PMA) in the presence of LPA or S1P in different tests. The influence of LPA and S1P on epithelial and endothelial cell F-actin was determined with immunohistochemistry.

results. LPA significantly increased the resistance of both the epithelial and endothelial monolayers, whereas S1P increased the resistance in only the endothelial cells. PTX blocked both the LPA- and S1P-induced increases in resistance, and DGPP (8:0) inhibited LPA-induced transcellular resistance in both the epithelium and endothelium. LPA and S1P prevented PMA-induced resistance decreases across epithelial and endothelial cells. F-actin staining around cell borders was more intense in both LPA- and S1P-treated cells.

conclusions. LPA increases transcellular resistance across cultured rabbit corneal epithelial and endothelial cell monolayers, and the effect is mediated through the LPA1 receptor and signaled through Gαi/o. S1P-stimulated increases in endothelial resistance are also signaled through Gαi/o. Both LPA and S1P prevented increased transcellular permeabilities induced by PMA, and increased actin stress fiber formation in epithelial and endothelial cells.

The corneal epithelium forms a tight barrier between the environment and the stroma of the cornea. 1 This barrier protects the cornea and intraocular structures from the external environment. In contrast, the corneal endothelium establishes a leaky barrier on the posterior surface of the cornea. The endothelial barrier allows aqueous humor to leak into the stroma to provide for the nutritional needs of cells of the stroma and basal epithelium. The fluid that leaks across the endothelial barrier is subsequently “pumped” back out of the stroma via the activity of endothelial Na,K-adenosine triphosphatase (ATPase) and secondary ion transport mechanisms coupled to the ion gradients created by Na,K-ATPase. Thus, the epithelial and endothelial tight junctional barriers play a key role in the homeostasis and host defense of the cornea. 1 2  
Tight junctions are located in the apical region of superficial corneal epithelial cells, as well as on the apical region of corneal endothelial cells. The interactions of tight junction proteins between adjacent cells form a barrier to the paracellular diffusion of water and solutes and maintain gradients generated by transcellular transport mechanisms. Whereas the tight junctions of the corneal epithelium form a high resistance barrier to solutes, the endothelial tight junctions form a relatively leaky barrier. Corneal tight junctions are not passive structures, but rather dynamic functional entities that can be regulated by a variety of signaling mechanisms to meet the changing needs and responses of the cornea. The resistance barrier of the cornea is regulated by cytokines, protein kinase C (PKC), phospholipase A (PLA), and mitogen-activated protein kinases (MAPKs). 2 3 4 Findings in work of Watsky et al. 3 4 and Riley et al. 5 showed that corneal endothelial permeability is controlled by PLC and PLA activation. 3 4 5 In addition, Wang et al. 2 found that activation of PKC by phorbol 12-myristate 13-acetate (PMA) results in an increase in paracellular permeability, as evidenced by a decrease in transepithelial electrical resistance. 
Both platelets and lipoproteins generate sphingosine 1-phosphate (S1P) and lysophosphatidic acid (LPA), two members of the phospholipids growth factor (PLGF) family. 6 To date, there have been four LPA receptors and five S1P receptors identified in mammals. The LPA receptors consist of LPA1 to -4, and S1P receptors consist of S1P1 to -5. 7 LPA and S1P are potent signaling molecules involved in a variety of physiological and pathologic processes, including cell differentiation and proliferation, 8 9 cytoskeletal rearrangement, 10 actin changes, 11 cell–cell communication, 6 and wound healing. 9 In addition, PLGFs have been shown to influence cell permeability. S1P has been reported to decrease the permeability of endothelial cells via Gαi, 12 whereas the influence of LPA on endothelial cell permeability is variable. For example, Alexander et al. 13 and Minnear et al. 14 reported that LPA decreases the permeability of endothelial monolayers in vitro, whereas Schulze et al. 15 reported that permeability in cultured brain endothelial cells is increased by LPA. 
The cytoskeleton, which has a high F-actin content, supports the cell membrane and gives it shape, 16 assists in cell motility, 17 and is an important factor in maintaining the integrity of tight junctions. 18 In the corneal epithelium and endothelium, F-actin forms a peripheral band around the apical region of the cells. 19 20 S1P-induced increases in F-actin density around the peripheral border of pulmonary endothelial cells has been associated with cell barrier enhancement. 21 In addition, LPA has been reported to have strong effects on the cortical actin stress fibers and focal adhesions of lens epithelial cells, in association with activation of Rho GTPase. 22  
LPA, along with several other PLGF family members, are present in aqueous humor and lacrimal gland fluid, and corneal injury results in a significant increase in the concentration of these lipids. 23 It has been established that many physiological actions induced by LPA and S1P are transduced through receptors coupled to the PTX-sensitive Gαi/o protein. 7 24 25 Watsky et al. 9 and Liliom et al. 23 have demonstrated that LPA stimulates proliferation in the three major cell types of the cornea in a dose-dependent manner and that the LPA-stimulated proliferation of corneal epithelial cells and keratocytes is PTX sensitive. In addition to the proliferation response, keratocytes from wounded corneas express an LPA-activated Cl current that is also activated by an increase in cell volume and serum, 26 and wound-healing affects the expression of PLGF receptor mRNA in corneal cells. 8 All these data suggest that LPA, S1P, and other PLGF family members are involved in maintaining the integrity of the normal cornea. Although PLGFs are present and active in the cornea and PLGFs have been shown to influence barrier function in vascular and pulmonary epithelial and endothelial cells, it remains unclear whether LPA and S1P have any direct effect on corneal epithelial or endothelial permeability, and if so, whether this effect is mediated by Gαi/o
Electric cell–substrate impedance sensing (ECIS) electrically monitors cell resistance across cultured cell layers noninvasively in real time. Increases in ion flow across the cell layers (decreased electrical resistance) indicate a disruption in the barrier integrity. Among many studies using ECIS to study barrier function, ECIS studies demonstrated differential pulmonary endothelial and alveolar epithelial barrier function regulation via cytoskeletal interactions with tight junction complexes. 27  
In this study, we used ECIS to determine whether LPA and S1P increase corneal transepithelial and endothelial resistance, and whether this increase is receptor-activated and signaled through Gαi/o. We found LPA increases the transcellular resistance in both epithelium and endothelium through Gαi/o via the LPA1 receptor, whereas S1P only increases resistance across the endothelium, also through Gαi/o. In addition, both LPA and S1P prevented the phorbol 12-myristate 13-acetate (PMA)-induced decreases in resistance across corneal epithelial and endothelial monolayers. LPA and S1P also increased actin stress fiber formation, particularly along the cell borders, in corneal epithelial and endothelial cells. 
Methods and Materials
Chemicals
Unless otherwise stated, chemicals were obtained from Sigma-Aldrich (St. Louis, MO). LPA (18:1), S1P, and DGPP (8:0) were purchased from Avanti Polar Lipids (Alabaster, AL). Rhodamine-phalloidin was purchased from Molecular Probes (Eugene, OR). 
Electric Cell–Substrate Impedance Sensing
The ECIS (model 1600R) system used in this study was from Applied BioPhysics (Applied BioPhysics, Troy, NY). In this system, gold-film electrodes are set into the bottom of eight-well tissue culture dishes. An AC current (40 kHz) is applied between the small active electrodes and a large counter electrode, with culture medium being used as the electrolyte. The instrument monitors both the voltage across the electrodes and its phase relative to the applied current. In addition to reporting total impedance, these data are converted to resistance and capacitance values, treating the cell–electrode system as a series resistance-capacitance (RC) circuit. When cells attach and spread over the small electrodes, the impedance increases as the insulating cell membranes force current into the narrow spaces beneath the cell and through the intercellular junctions. The microampere current and the resultant voltage decrease of a few millivolts have no measurable effect on the cells. Thus, the monitoring of cell behavior is noninvasive. 
Cell Culture
Corneal epithelial and endothelial cells were isolated from the corneas of 1- to 2-kg New Zealand White rabbits, as previously described. 28 29 Cells were grown using DMEM (Cellgro; Mediatech, Inc., Herndon, VA) and Ham’s F12 (Gibco, Rockville, MD) plus 20% fetal calf serum (FCS). Cells were passaged at approximately 80% confluence. Cells of passages 2 to 8 were used in all experiments. Rabbits were treated in accordance with the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research. 
Preparation and Inoculation of Electrodes
ECIS electrode arrays (8W10E) were obtained from Applied BioPhysics. Each array slide consists of eight individually addressable wells with surfaces treated for cell culture. Cells were harvested from 100-mm dishes and plated at 5 × 105 cells/mL in ECIS chambers to obtain a confluent layer of cells covering each electrode. Complete single-layer electrode coverage was confirmed microscopically before data analysis. 
Preparation of LPA and S1P
LPA and S1P were solubilized according to the manufacturer’s instructions. LPA was diluted to the desired concentrations using a stock 1 mM BSA/EGTA solution, and S1P was diluted in a BSA (4 mg/mL) solution. 
Resistance Measurements with ECIS
To determine whether LPA and S1P affect transcellular resistance across cultured rabbit corneal epithelial and endothelial cells, we used the ECIS to examine the effects of LPA and S1P on resistances across corneal epithelial and endothelial cell monolayers. 
Resistance was monitored for up to 5 hours, with measurements being collected every few seconds by the ECIS device. The corneal epithelial and endothelial cells were serum starved 15 hours before the ECIS assay. A 1-hour equilibration time was allowed in the ECIS incubator for all cultures, and then different concentrations of LPA (100 nM, 1 μM, or 10 μM) or S1P (10 nM, 100 nM, or 1 μM) were added to each well to determine whether LPA and S1P affect the resistance across cultured corneal epithelial and endothelial cells and whether there were dose-dependent effects of LPA and S1P on the resistance. BSA/EGTA (1 mM) and BSA (4 mg/mL) were used as LPA and S1P controls, respectively. Cell resistances were then recorded for an additional 4 hours. For cells significantly affected by LPA or S1P, pertussis toxin (PTX, 100 ng/mL) plus LPA or S1P were added to the serum-free DMEM in separate experiments, to determine whether Gαi/o is involved in the response. An additional study was performed to determine whether LPA increases the transcellular resistance in epithelial and endothelial cells through a specific LPA receptor. Dioctyl-glycerol pyrophosphate (1 or 10 μM; DGPP 8:0) plus LPA (10 μM) were added to the ECIS wells. DGPP (8:0) has been shown to block LPA3 and LPA1 receptors with K i values of 106 nM and 6.6 μM, respectively, 30 and is ineffective at blocking LPA2. To determine whether LPA and/or S1P prevented PMA-induced decreases in epithelial and/or endothelial resistance, we also examined the effects of LPA and S1P on resistances across corneal epithelial and endothelial cell monolayers treated with PMA. PMA (20 nM) plus LPA (10 μM) or S1P (100 nM) for epithelium, and 50 nM PMA plus the same concentrations of LPA or S1P for endothelium, were added to the ECIS wells to incubate in ECIS chamber for 4 hours. Two PMA controls (20 and 50 nM) were also examined. 
To analyze ECIS results, resistance values were normalized (divided by their values at time 0), and data points were analyzed at 20-minute intervals. 
Actin Staining and Immunofluorescence
Corneal epithelial and endothelial cells were serum starved for 15 hours after they reached confluence. LPA (10 μM) or S1P (100 nM) were then added to the media for 2 hours. A BSA (4 mg/mL) control was also measured. Cells were rinsed in PBS, fixed in 4% paraformaldehyde for 15 minutes, permeabilized with Triton-X-100, blocked with 6% FBS, and incubated with rhodamine-phalloidin for 15 minutes. Fluorescence was examined with a confocal laser scanning microscope (LSM 5 Pascal Laser Scanning Microscope; Carl Zeiss Meditec, Jena, Germany). 
Statistics
A two-factor ANOVA with replication was used to compare all results. Data were considered significant when P < 0.05. 
Results
Resistance Measurements
The effects of LPA on corneal epithelial and endothelial transcellular resistance were measured. In addition, the possible Gαi/o protein signaling pathway coupling LPA and S1P to changes in barrier function was explored with the specific Gαi/o inhibitor, PTX. Figure 1shows the effects of different concentrations of LPA on corneal epithelial and endothelial transcellular resistances. LPA significantly increased both corneal epithelial (Fig. 1A)and endothelial (Fig. 1B)transcellular resistances in a dose-dependent manner (P < 0.01). Pretreatment with PTX prevented the LPA-induced resistance increase across corneal epithelial and endothelial cell monolayers (Fig. 2) . PTX (100 ng/mL) alone had no effects on resistance across epithelial and endothelial cells (data not shown). In addition, the LPA-induced transcellular resistance increases of both epithelium and endothelium were inhibited by 10 μM DGPP (8:0), but not 1 μM DGPP (8:0) (Fig. 3) . These results suggest that LPA affects resistance through the Gi/o-protein-coupled LPA1 receptor. 
PMA has been shown to disrupt corneal epithelial and endothelial barrier function. 2 4 To determine whether LPA and/or S1P could counteract the barrier disrupting effects of PMA, we added each lipid along with PMA to corneal epithelial and endothelial cells grown in ECIS chambers. Figure 4Ashows that LPA and S1P inhibited PMA-induced decreases in endothelial transcellular resistance. LPA and S1P also prevented the PMA-induced decrease in transepithelial cell monolayer resistance, although this effect became insignificant after 4 hours in the S1P group (Fig. 4B) . For the epithelial S1P group, overall significance equaled 0.08 using all time points and 0.03 if the last two time points were omitted. The ANOVA results showed significance (P < 0.01) if time was not included in the statistical analysis. DGPP (1 μM, 10 μM) alone had no effect on epithelial or endothelial transcellular resistance (data not shown). 
Figure 5shows the effects of different concentrations of S1P on both corneal epithelial and endothelial transcellular resistances. S1P increased the resistance only in endothelial cells. Not only did S1P have no effect on corneal epithelial transcellular resistance, but higher concentrations of S1P (>1 μM) appeared to be toxic to the epithelial cells (data not shown). Figure 6shows that PTX blocked the S1P-induced increase in corneal endothelial transcellular resistance. This demonstrates that S1P affects endothelial resistance through the Gαi/o pathway. 
Immunohistochemistry
Figure 7shows the influence of the LPA and S1P on the f-actin organization in corneal epithelial and endothelial cell monolayers. Figure 7A 7B 7Cshows control, LPA, and S1P treated epithelial cells (respectively), whereas Figures 7D and 7Eshow control, LPA-, and S1P-treated endothelial cells (respectively). In all cases, LPA and S1P treatment for 2 hours resulted in more intense F-actin stress fiber staining, along with increased f-actin staining along the cell borders. 
Discussion
In this study, we demonstrate that LPA increases transcellular resistance in both rabbit corneal epithelium and endothelium, whereas S1P only increases resistance across corneal endothelial tight junctions. The LPA (epithelium and endothelium) and S1P (endothelium) effects were dose dependent and acted through the Gαi/o-signaling pathway. In addition, LPA (epithelium and endothelium) and S1P (endothelium) prevented PMA-induced decreases in transcellular resistances across cell monolayers. LPA and S1P increased actin stress fiber formation, particularly along the cell borders of both type cells. 
ECIS, a technique first described in 1984 by Giaever and Keese, 31 electrically monitors cell resistance noninvasively in real time. Impedance changes measured using ECIS have also been used to monitor cell attachment, spreading, and micromotion. 32 33 Because it is relatively simple to culture cells on the electrodes, and the measurements are not time consuming, we used the ECIS technique as a simple and sensitive method to study corneal epithelial and endothelial permeability. 
Dynamic regulation of transcellular permeability is fundamental to many physiological processes; therefore, it is important to understand the regulatory mechanisms of cell permeability. We previously demonstrated that tumor necrosis factor (TNF)-α increases corneal endothelial permeability. 3 Wang et al. 2 group reported that activation of the ERK1/2 MAPK pathway induces increased permeability in human corneal epithelial cells. At the same time, actin cytoskeleton changes also affect the control of epithelial and endothelial barriers. 34  
LPA and S1P are important bioactive lipid growth factors present in many cell types and several organ compartments. It has been shown that several endothelial cell types reduce their transcellular permeability in response to S1P. This response has been shown to be dependent on activation of Rho kinase and Gαi and is signaled through the Edg1 and -3 receptors. 12 S1P and FTY720, a high-affinity agonist for S1P receptors, potently inhibited VEGF-induced transcellular permeability in murine embryonic endothelial cells. 35 Peng et al. 36 also demonstrated that S1P decreased pulmonary and renal vascular leakage, in a murine model of LPS-mediated acute lung injury. 
The influences of LPA on the transcellular permeability of different cell types are diverse. LPA increases the permeability of brain endothelial cells as measured by increases in the flux of sucrose across confluent monolayers. 15 In contrast, LPA can effectively stabilize vascular endothelial monolayer barrier function, 37 and the permeability of cultured bovine fetal aortic endothelial monolayers can be decreased by LPA. 11 Pertussis toxin has been shown to attenuate LPA-induced decreases in endothelial permeability. 13 38  
We found that LPA increased the transcellular resistance of corneal endothelial and epithelial cells in a dose-dependent manner, whereas S1P only increased endothelial cell resistance. S1P did not affect corneal epithelial transcellular resistance, even at high concentrations. Concentrations of S1P greater than 1 μM appeared to be toxic to the epithelial cells, resulting in shrinkage and death of some cells (data not shown). 
Previous studies have established that LPA and S1P receptors coupled with PTX-sensitive G proteins are related to the multiple effects of LPA and S1P. 7 24 25 In the present study, we observed that the LPA-induced increase in endothelial and epithelial cell resistances, and the S1P-induced increase in endothelial cell resistance, were all suppressed by PTX treatment. This inhibition by PTX suggests that the effects of LPA and S1P on corneal cells are at least partially mediated through Gαi/o-signaling pathways. 
It is now clear that most of the responses documented for extracellular LPA are attributable to the activation of G-protein-coupled receptors. Exogenous overexpression of the LPA1 receptor in rat Schwann cells significantly reduced apoptosis on serum withdrawal, indicating that LPA promotion of Schwann cell survival was at least partially mediated by the LPA1 receptor. 39 We previously identified the different PLGF receptor subtypes present in the three cell types of the rabbit cornea and found the LPA1, LPA3, and S1P1 receptors. 8 We also discovered that activation of a Cl current in cultured corneal keratocytes is activated by LPA via a receptor-mediated signaling pathway. The LPA1 receptor was implicated in this response, because 1 μM DGPP was ineffective at preventing LPA-induced Cl current activity, whereas 10 μM DGPP prevented activation of the current. 40 In the present study, 1 μM DGPP (8:0) did not block the LPA-induced resistance increases across epithelial and endothelial cells, whereas 10 μM DGPP (8:0) significantly blocked this LPA-induced resistance increase. This suggests that LPA increases epithelial and endothelial transcellular resistance through the LPA1 receptor. S1P receptor specificity was not examined in this study. 
PMA, a PKC activator, leads to disruption of epithelial and endothelial cell monolayer integrity in corneal cells and cell types. 2 4 41 42 Our laboratory has demonstrated that PMA increases corneal endothelial permeability, likely through a PKC-mediated pathway. 4 In the present study, we found that PMA increased permeability of corneal epithelial and endothelial tight junctions, whereas LPA and S1P inhibited the effects of PMA on the resistances. This result suggests that the effects of LPA and S1P on corneal cell permeability may involve regulation of PKC activity. 
LPA and S1P are two modulators of the actin cytoskeleton. S1P organizes actin into a strong cortical ring and strengthens both intercellular and cell–matrix adherence to decrease vascular permeability to fluid. 43 LPA also causes Schwann cell actin changes. 11 In this study, LPA and S1P increased F-actin stress fiber formation, particularly along the cell borders, when corneal cells were exposed to LPA or S1P for 2 hours. This was the time frame in which we noted significant changes in transcellular resistance (Figs. 1 5) . Thus, LPA and S1P may increase the resistance across corneal epithelial and endothelial cell monolayers, at least in part, via regulation of the actin cytoskeleton. 
In conclusion, this study provided detailed evidence that LPA stimulates increases in transcellular epithelial and endothelial resistance, whereas S1P increases transcellular resistance only in endothelial cells. Both lipids signaled this response through Gαi/o, and the LPA-induced increase in corneal transepithelial and -endothelial resistance was mediated through the LPA1 receptor. LPA and S1P also prevent PMA-induced decreases in the transcellular resistance of corneal epithelial and endothelial tight junctions. Both lipids also increased the actin stress fiber formation in the two types of cells. This study also demonstrates that transcellular impedance measurement using the ECIS technique is a simple and sensitive method for studying corneal epithelial and endothelial permeability. 
 
Figure 1.
 
(A) Epithelial and (B) endothelial resistance values (normalized) measured with ECIS. After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 100 nM, 1 μM, or 10 μM LPA, along with a vehicle (1 mM BSA/EGTA) control. LPA significantly increased both corneal epithelial and endothelial transcellular resistance in a dose-dependent manner (n = 6, epithelium; n = 10, endothelium; P < 0.01).
Figure 1.
 
(A) Epithelial and (B) endothelial resistance values (normalized) measured with ECIS. After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 100 nM, 1 μM, or 10 μM LPA, along with a vehicle (1 mM BSA/EGTA) control. LPA significantly increased both corneal epithelial and endothelial transcellular resistance in a dose-dependent manner (n = 6, epithelium; n = 10, endothelium; P < 0.01).
Figure 2.
 
(A) Epithelial and (B) endothelial cells were pretreated with PTX (100 ng/mL) for 15 hours. After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing LPA (10 μM), along with a vehicle (1 mM BSA/EGTA) control. PTX prevented the LPA-induced increase in resistance in both cell types (vehicle versus LPA+PTX, n = 6; P > 0.05).
Figure 2.
 
(A) Epithelial and (B) endothelial cells were pretreated with PTX (100 ng/mL) for 15 hours. After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing LPA (10 μM), along with a vehicle (1 mM BSA/EGTA) control. PTX prevented the LPA-induced increase in resistance in both cell types (vehicle versus LPA+PTX, n = 6; P > 0.05).
Figure 3.
 
(A) Epithelial and (B) endothelial resistance values (normalized). After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 1 or 10 μM DGPP (8:0) plus 10 μM LPA, along with a 10-μM LPA control. The results showed that 10 μM, but not 1 μM DGPP, prevented the LPA-induced resistance increase in both cell types (1 μM DGPP (8:0)+LPA versus LPA, n = 6, P > 0.05; 10 μM DGPP (8:0)+LPA versus LPA, n = 6, P < 0.01).
Figure 3.
 
(A) Epithelial and (B) endothelial resistance values (normalized). After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 1 or 10 μM DGPP (8:0) plus 10 μM LPA, along with a 10-μM LPA control. The results showed that 10 μM, but not 1 μM DGPP, prevented the LPA-induced resistance increase in both cell types (1 μM DGPP (8:0)+LPA versus LPA, n = 6, P > 0.05; 10 μM DGPP (8:0)+LPA versus LPA, n = 6, P < 0.01).
Figure 4.
 
LPA and S1P prevent the PMA-induced decrease in transendothelial and epithelial resistance. (A) For endothelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing LPA (10 μM)+PMA (50 nM) or S1P (100 nM)+PMA (50 nM), along with a PMA (50 nM) and BSA (4 mg/mL) control. LPA and S1P prevented the PMA-induced decrease in resistance (LPA+PMA versus PMA+BSA; P < 0.01. S1P+PMA versus PMA+BSA; P < 0.01). (B) For epithelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing LPA (10 μM)+PMA (20 nM) or S1P (100 nM)+PMA (20 nM), along with a PMA (20 nM) and BSA (4 mg) control. LPA and S1P prevented the PMA-induced decrease in resistance (LPA+PMA versus PMA+BSA, n = 6; P < 0.01), although S1P was less effective (S1P+PMA versus PMA+BSA, n = 3; P = 0.08; with the last two time points omitted, P < 0.05; also P < 0.01 with the time component omitted from the analysis). Experiments were repeated two times. Representative results are shown.
Figure 4.
 
LPA and S1P prevent the PMA-induced decrease in transendothelial and epithelial resistance. (A) For endothelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing LPA (10 μM)+PMA (50 nM) or S1P (100 nM)+PMA (50 nM), along with a PMA (50 nM) and BSA (4 mg/mL) control. LPA and S1P prevented the PMA-induced decrease in resistance (LPA+PMA versus PMA+BSA; P < 0.01. S1P+PMA versus PMA+BSA; P < 0.01). (B) For epithelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing LPA (10 μM)+PMA (20 nM) or S1P (100 nM)+PMA (20 nM), along with a PMA (20 nM) and BSA (4 mg) control. LPA and S1P prevented the PMA-induced decrease in resistance (LPA+PMA versus PMA+BSA, n = 6; P < 0.01), although S1P was less effective (S1P+PMA versus PMA+BSA, n = 3; P = 0.08; with the last two time points omitted, P < 0.05; also P < 0.01 with the time component omitted from the analysis). Experiments were repeated two times. Representative results are shown.
Figure 5.
 
Epithelial and endothelial resistance levels (normalized). (A) In epithelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 100 nM or 1 μM S1P, along with a BSA (4 mg/mL) control. (B) For endothelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 10 or 100 nM S1P, along with a BSA (4 mg/mL) control. Whereas S1P had no effect on epithelial resistance (n = 10, P > 0.05), S1P significantly increased the endothelium transcellular resistance in a dose-dependent manner (n = 8, P < 0.01).
Figure 5.
 
Epithelial and endothelial resistance levels (normalized). (A) In epithelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 100 nM or 1 μM S1P, along with a BSA (4 mg/mL) control. (B) For endothelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 10 or 100 nM S1P, along with a BSA (4 mg/mL) control. Whereas S1P had no effect on epithelial resistance (n = 10, P > 0.05), S1P significantly increased the endothelium transcellular resistance in a dose-dependent manner (n = 8, P < 0.01).
Figure 6.
 
Endothelial cells were pretreated with PTX (100 ng/mL) for 15 hours. After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 100 nM S1P, along with a BSA (4 mg/mL) control. PTX pretreatment prevented the S1P-induced increase in resistance (BSA versus S1P+PTX pretreatment, n = 6, P > 0.05).
Figure 6.
 
Endothelial cells were pretreated with PTX (100 ng/mL) for 15 hours. After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 100 nM S1P, along with a BSA (4 mg/mL) control. PTX pretreatment prevented the S1P-induced increase in resistance (BSA versus S1P+PTX pretreatment, n = 6, P > 0.05).
Figure 7.
 
Effects of LPA and SIP on the actin cytoskeleton. (B) LPA- and (C) S1P-treated epithelial cells (2 hour exposure) showed increased F-actin staining with rhodamine phalloidin, particularly along the cell borders, compared with the untreated control (A). In a similar fashion, (E) LPA and (F) S1P endothelial cells treated for 2 hours also showed increased F-actin staining, particularly along the cell borders, compared to the untreated control (D).
Figure 7.
 
Effects of LPA and SIP on the actin cytoskeleton. (B) LPA- and (C) S1P-treated epithelial cells (2 hour exposure) showed increased F-actin staining with rhodamine phalloidin, particularly along the cell borders, compared with the untreated control (A). In a similar fashion, (E) LPA and (F) S1P endothelial cells treated for 2 hours also showed increased F-actin staining, particularly along the cell borders, compared to the untreated control (D).
The authors thank Victorina Pintea for technical assistance. 
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Figure 1.
 
(A) Epithelial and (B) endothelial resistance values (normalized) measured with ECIS. After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 100 nM, 1 μM, or 10 μM LPA, along with a vehicle (1 mM BSA/EGTA) control. LPA significantly increased both corneal epithelial and endothelial transcellular resistance in a dose-dependent manner (n = 6, epithelium; n = 10, endothelium; P < 0.01).
Figure 1.
 
(A) Epithelial and (B) endothelial resistance values (normalized) measured with ECIS. After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 100 nM, 1 μM, or 10 μM LPA, along with a vehicle (1 mM BSA/EGTA) control. LPA significantly increased both corneal epithelial and endothelial transcellular resistance in a dose-dependent manner (n = 6, epithelium; n = 10, endothelium; P < 0.01).
Figure 2.
 
(A) Epithelial and (B) endothelial cells were pretreated with PTX (100 ng/mL) for 15 hours. After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing LPA (10 μM), along with a vehicle (1 mM BSA/EGTA) control. PTX prevented the LPA-induced increase in resistance in both cell types (vehicle versus LPA+PTX, n = 6; P > 0.05).
Figure 2.
 
(A) Epithelial and (B) endothelial cells were pretreated with PTX (100 ng/mL) for 15 hours. After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing LPA (10 μM), along with a vehicle (1 mM BSA/EGTA) control. PTX prevented the LPA-induced increase in resistance in both cell types (vehicle versus LPA+PTX, n = 6; P > 0.05).
Figure 3.
 
(A) Epithelial and (B) endothelial resistance values (normalized). After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 1 or 10 μM DGPP (8:0) plus 10 μM LPA, along with a 10-μM LPA control. The results showed that 10 μM, but not 1 μM DGPP, prevented the LPA-induced resistance increase in both cell types (1 μM DGPP (8:0)+LPA versus LPA, n = 6, P > 0.05; 10 μM DGPP (8:0)+LPA versus LPA, n = 6, P < 0.01).
Figure 3.
 
(A) Epithelial and (B) endothelial resistance values (normalized). After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 1 or 10 μM DGPP (8:0) plus 10 μM LPA, along with a 10-μM LPA control. The results showed that 10 μM, but not 1 μM DGPP, prevented the LPA-induced resistance increase in both cell types (1 μM DGPP (8:0)+LPA versus LPA, n = 6, P > 0.05; 10 μM DGPP (8:0)+LPA versus LPA, n = 6, P < 0.01).
Figure 4.
 
LPA and S1P prevent the PMA-induced decrease in transendothelial and epithelial resistance. (A) For endothelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing LPA (10 μM)+PMA (50 nM) or S1P (100 nM)+PMA (50 nM), along with a PMA (50 nM) and BSA (4 mg/mL) control. LPA and S1P prevented the PMA-induced decrease in resistance (LPA+PMA versus PMA+BSA; P < 0.01. S1P+PMA versus PMA+BSA; P < 0.01). (B) For epithelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing LPA (10 μM)+PMA (20 nM) or S1P (100 nM)+PMA (20 nM), along with a PMA (20 nM) and BSA (4 mg) control. LPA and S1P prevented the PMA-induced decrease in resistance (LPA+PMA versus PMA+BSA, n = 6; P < 0.01), although S1P was less effective (S1P+PMA versus PMA+BSA, n = 3; P = 0.08; with the last two time points omitted, P < 0.05; also P < 0.01 with the time component omitted from the analysis). Experiments were repeated two times. Representative results are shown.
Figure 4.
 
LPA and S1P prevent the PMA-induced decrease in transendothelial and epithelial resistance. (A) For endothelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing LPA (10 μM)+PMA (50 nM) or S1P (100 nM)+PMA (50 nM), along with a PMA (50 nM) and BSA (4 mg/mL) control. LPA and S1P prevented the PMA-induced decrease in resistance (LPA+PMA versus PMA+BSA; P < 0.01. S1P+PMA versus PMA+BSA; P < 0.01). (B) For epithelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing LPA (10 μM)+PMA (20 nM) or S1P (100 nM)+PMA (20 nM), along with a PMA (20 nM) and BSA (4 mg) control. LPA and S1P prevented the PMA-induced decrease in resistance (LPA+PMA versus PMA+BSA, n = 6; P < 0.01), although S1P was less effective (S1P+PMA versus PMA+BSA, n = 3; P = 0.08; with the last two time points omitted, P < 0.05; also P < 0.01 with the time component omitted from the analysis). Experiments were repeated two times. Representative results are shown.
Figure 5.
 
Epithelial and endothelial resistance levels (normalized). (A) In epithelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 100 nM or 1 μM S1P, along with a BSA (4 mg/mL) control. (B) For endothelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 10 or 100 nM S1P, along with a BSA (4 mg/mL) control. Whereas S1P had no effect on epithelial resistance (n = 10, P > 0.05), S1P significantly increased the endothelium transcellular resistance in a dose-dependent manner (n = 8, P < 0.01).
Figure 5.
 
Epithelial and endothelial resistance levels (normalized). (A) In epithelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 100 nM or 1 μM S1P, along with a BSA (4 mg/mL) control. (B) For endothelial cells, after a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 10 or 100 nM S1P, along with a BSA (4 mg/mL) control. Whereas S1P had no effect on epithelial resistance (n = 10, P > 0.05), S1P significantly increased the endothelium transcellular resistance in a dose-dependent manner (n = 8, P < 0.01).
Figure 6.
 
Endothelial cells were pretreated with PTX (100 ng/mL) for 15 hours. After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 100 nM S1P, along with a BSA (4 mg/mL) control. PTX pretreatment prevented the S1P-induced increase in resistance (BSA versus S1P+PTX pretreatment, n = 6, P > 0.05).
Figure 6.
 
Endothelial cells were pretreated with PTX (100 ng/mL) for 15 hours. After a 1-hour equilibration (arrow), medium was replaced with serum-free DMEM containing 100 nM S1P, along with a BSA (4 mg/mL) control. PTX pretreatment prevented the S1P-induced increase in resistance (BSA versus S1P+PTX pretreatment, n = 6, P > 0.05).
Figure 7.
 
Effects of LPA and SIP on the actin cytoskeleton. (B) LPA- and (C) S1P-treated epithelial cells (2 hour exposure) showed increased F-actin staining with rhodamine phalloidin, particularly along the cell borders, compared with the untreated control (A). In a similar fashion, (E) LPA and (F) S1P endothelial cells treated for 2 hours also showed increased F-actin staining, particularly along the cell borders, compared to the untreated control (D).
Figure 7.
 
Effects of LPA and SIP on the actin cytoskeleton. (B) LPA- and (C) S1P-treated epithelial cells (2 hour exposure) showed increased F-actin staining with rhodamine phalloidin, particularly along the cell borders, compared with the untreated control (A). In a similar fashion, (E) LPA and (F) S1P endothelial cells treated for 2 hours also showed increased F-actin staining, particularly along the cell borders, compared to the untreated control (D).
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