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Microtubule disassembly breaks down the barrier integrity in a number of epithelial and endothelial monolayers. This study has investigated effects of TNF-α, which is implicated in corneal allograft rejection, on microtubules and barrier integrity in cultured bovine corneal endothelial cells. Exposure to TNF- α led to disassembly of the microtubules, and also caused disruption of the perijunctional actomyosin ring (PAMR). As a measure of barrier integrity, trans-endothelial electrical resistance (TER) was determined based on electrical cell-substrate impedance sensing in realtime. Exposure to TNF- α caused a slow decline in TER for > 20 h, and a similar exposure to cells grown on porous culture inserts led to a significant increase in permeability to FITC dextran. These changes, indicating a loss of barrier integrity, were also reflected by dislocation of ZO-1 at the cell border and disassembly of cadherins. These effects of TNF- α were inhibited upon stabilization of microtubules by pre-treatment with paclitaxel or epothilone B. Microtubule stabilization may be a useful strategy to overcome (TNF-α)-induced loss of the barrier integrity of corneal endothelium during inflammation associated with transplant rejection and uveitis.
Dysfunction of the corneal endothelium secondary to iatrogenic injury, inflammation, and genetic disorders (e.g., Fuchs’ dystrophy) can lead to a loss of corneal transparency (Joyce 2003; Edelhauser 2006). Currently, there are no pharmacological strategies to treat endothelial dysfunction, and the alternatives include endothelial or full-thickness corneal transplantation. In USA alone, there are nearly 40,000 corneal transplants performed annually (George and Larkin 2004). Even after transplantation, survival of the endothelium is a major concern because of inflammatory response secondary to allograft rejection (Armitage et al. 2003; Edelhauser 2006; Niederkorn 2007). Among the cytokines found in significant levels in the cornea and aqueous humor, TNF-α is also assumed to play a major role in corneal allograft rejection (George and Larkin 2004; Niederkorn 2007). Thus, it is important to investigate the influence of the cytokine on the barrier integrity of corneal endothelium, and to assess strategies to oppose its adverse influence.
TNF-α is known to increase permeability of epithelial and endothelial monolayers by mechanisms that are independent of apoptosis (Goldblum et al. 1993; Watsky et al. 1996; Bruewer et al. 2003; Ma et al. 2005; Ye et al. 2006; McKenzie and Ridley 2007; Kimura et al. 2008). Accordingly, the involvement of the cytokine in the breakdown of the barrier integrity is well known in many disorders including pulmonary edema (Petrache et al. 2003) and Crohn’s disease (Ma et al. 2004). Its influence on corneal endothelium has also been demonstrated in an in vitro model of rabbit eyes (Watsky et al. 1996). This study by Watsky et al., showed that exposure of the endothelium to the cytokine results in an increase in permeability to the hydrophilic carboxyfluorescein. In parallel with the loss of the barrier integrity, the actin cytoskeleton was disrupted. Furthermore, the study showed that elevated cAMP opposed the influence of TNF-α. Despite these early observations, molecular mechanisms involved in the breakdown of the actin cytoskeleton as well as the barrier integrity remain unknown.
Several recent studies on the (TNF-α)-induced loss of barrier integrity in vascular endothelium have implicated microtubule disassembly (Petrache et al. 2003; Kakiashvili et al. 2009). The microtubules, as part of the cytoskeleton, are essential for structural integrity of cells. Although their involvement in trafficking of membrane proteins, migration, polarity, mitosis, and maintenance of cell shape (Musch 2004) are relatively well understood, the mechanism(s) by which microtubule disassembly induces actomyosin contraction is not well established. One mechanism that is shown in vascular endothelium involves activation of RhoA (Birukova et al. 2005) through a release of RhoA–specific GEFs (guanine nucleotide exchange factors), which are anchored to the microtubules and released in response to its disassembly (van Horck et al. 2001; Birukova et al. 2006). In a recent study from our laboratory, we have shown that microtubule disassembly by exposure to nocodazole results in disruption of the actin cytoskeleton and thereby induces a loss of barrier integrity in bovine corneal endothelial monolayers (Jalimarada et al. 2009). In current study, we have investigated the role of microtubules in the (TNF-α)-induced loss of barrier integrity in corneal endothelium.
In studies involving pulmonary artery endothelial cells, it has been shown that microtubule stabilization effectively opposes loss of barrier integrity induced by TNF-α (Petrache et al. 2003). Among the agents known to stabilize microtubules, the efficacy of paclitaxel is well established (Amos and Lowe 1999). Epothilones are a new class of antineoplastic agents known to induce tubulin polymerization, and enhance microtubule stability (Reichenbach and Hofle 2008). Here, we have examined the ability of both paclitaxel and epothilone B to oppose the loss of barrier integrity induced by TNF-α. Our results show that (TNF-α)-induced microtubule disassembly contributes to barrier dysfunction of corneal endothelium, and that the stabilization of microtubules effectively opposes the TNF-α response.
TNF-α (biological activity of 2 × 107 U/mg; free of endotoxin), paclitaxel, α-tubulin antibody (anti-mouse), monoclonal pan-cadherin antibody (anti-mouse) and FITC dextran (10 kDa) were purchased from Sigma Aldrich (St. Louis, MO). Epothilone B was obtained from EMD Biosciences (La Jolla, CA). Texas red conjugated phalloidin, goat anti-mouse Alexa-488 and anti-fade agent were purchased from Molecular Probes (Eugene, OR). ZO-1 antibody (anti-mouse) was from Zymed (Long Island, NY). Electrodes (8W10E+) for TER measurements were from Applied Biophysics, Inc. (Troy, NY).
Primary cultures of BCEC from fresh eyes were established as previously described.(Satpathy et al. 2004; Srinivas et al. 2004) The growth medium contained Dulbecco’s Modified Eagle’s Medium (DMEM), supplemented with 10% bovine calf serum and an antibiotic-antimycotic mixture (Penicillin 100 U/ml, Streptomycin 100 μg/ml and amphotericin-B 0.25 μg/ml). Cells were cultured at 37°C in a humidified atmosphere containing 5% CO2. The medium was replaced every 2–3 days. Cells of the first and second passages were harvested and seeded onto glass coverslips and porous filters, then allowed to grow to confluence for 3–4 days before use. Cell culture supplies were from Invitrogen (Long Island, NY). For all the experiments, TNF-α (20 ng/ml), paclitaxel (10 μM), and epothilone B (0.1 μM) were used to treat the cells. Paclitaxel or epothilone B was pretreated for 1 h before the addition of TNF-α.
Cells grown on coverslips were washed with PBS after desired drug treatment, fixed with 3.7% paraformaldehyde and permeabilized using 0.2% Triton X-100 for 5 min. This was followed by staining for F-actin by phalloidin conjugated to Texas Red (1:1000) for 45–60 min at room temperature. Cells were stained for microtubule, ZO-1 and cadherins by fixing the cells followed by permeabilization with 0.01% saponin in PBS. The cells were exposed to blocking buffer for 45 min at RT, and then incubated with the antibody for α-tubulin (1:1000)/ZO-1 (1:25)/pan-cadherin (1:1000) in a mixture of 0.01% saponin in PBS-goat serum (1:1) overnight at 4°C. This was followed by washing with 0.01% saponin in PBS and incubation with secondary antibody (goat anti-mouse IgG Alexa Fluor 488 at 1:1000) for 1 h at RT. Stained cells were mounted using anti-fade medium with DAPI and visualized using an epifluorescence microscope equipped with a 60/100x oil immersion objective and 1.2 NA (Nikon, Tokyo, Japan).
TER was measured by ECIS (electric cell substrate impedance sensing) device (ECIS 1600R, Applied Biophysics, Troy, NY). The electrodes (8W10E+, Applied Biophysics, Inc.) were stabilized with aqueous solution of L-cysteine (10 mM) for 15 min in the incubator at 37°C, followed by washing with regular media two times before seeding of cells. Cells were seeded at a density of 5×105 cells/ml on electrodes and grown to confluence. The approach to confluence was monitored through spreading data acquired overnight. After the monolayers reached steady resistance values, cells were washed twice with serum-free media and allowed to equilibrate for 1 h before drug treatment. The resistive portion of impedance (TER) normalized to its initial value at time zero was monitored continuously and taken as a measure of barrier integrity.
Cells were grown on 0.2 μm pore-size collagen IV (1 mg/ml)-coated tissue culture inserts (Nunc, Fisher Scientific, Pittsburgh, PA) to confluence. The monolayers were then serum-starved for 1 h and either left untreated or exposed in triplicate with esired agents. Cells were pretreated with paclitaxel or epothilone B for 1 h before exposure to TNF-α. Following treatment, FITC-dextran (10 kDa) dissolved in the Ringer solution was placed in the apical compartment at 0.4 μg/mL and the cells were placed back in the incubator for 2 h. Samples were then taken from the basolateral chamber for fluorescence measurements. Fluorescence was measured by excitation at 492 nm and the emission was collected at 520 nm.
Statistical comparisons were made by one-way analysis of variance (ANOVA) with Bonferroni’s post test analysis using GraphPad Prism software (GraphPad Software, Inc., San Diego, CA; Version 5.0). A value of p<0.05 was considered statistically significant. Data are expressed as mean ± SE. “n” represents number of independent experiments.
Fig. 1 shows the influence of TNF-α on microtubules with and without paclitaxel or epothilone B pre-treatment. In untreated cells (Fig. 1A), microtubules extend from around the nucleus as fibrillary structures towards the cell periphery. Treatment with TNF-α for 6 h led to loss of the fibrillary extensions, and condensation of microtubules noticeable around the nucleus (Fig. 1B). Treatment with paclitaxel or epothilone B stabilized microtubule assembly (Figs. 1C and 1E, respectively). This loss is not apparent when cells were pre-treated with paclitaxel or epothilone B (Figs. 1D and 1F, respectively). These results show the ability of TNF-α to induce microtubule disassembly, and that the response is masked upon microtubule stabilization.
In order to establish the effect of (TNF-α)-induced microtubule disassembly on barrier integrity, we first examined its influence on permeability to FITC dextran (10 KDa) across cells grown to form a monolayer on porous culture inserts. After growing the cells to confluence, as established previously (Jalimarada et al. 2009), for 7 days, we challenged the apical chamber to TNF-α for 20 h with or without pre-treatment with paclitaxel or epothilone B. We then assessed flux of FITC dextran to the basolateral chamber. TNF-α significantly increased flux of FITC dextran when compared to untreated cells (Fig. 2). Furthermore, cells that were pre-treated with paclitaxel or epothilone B showed significantly attenuated flux in response to TNF-α compared to those treated with the cytokine alone. In addition, it may be noted that the opposition of epothilone B to the TNF-α response is significantly greater than that of paclitaxel.
In order to gain further insights into the loss of barrier integrity and its protection by microtubule stabilization, we examined trans-endothelial electrical resistance (TER). As shown by us (Jalimarada et al. 2009) and Watsky et al (Yin and Watsky 2005) for corneal endothelial cells, we employed electrical cell substrate impedance sensing (ECIS) approach, which is especially sensitive and therefore highly suitable for leaky epithelia. This method, which is widely employed with vascular endothelial cells,(Petrache et al. 2003; Birukova et al. 2004) is especially useful for the corneal endothelium. As shown in Fig. 3A, exposure to TNF-α led to a slow but a persistent decline in TER that continued beyond 20 h. Pre-treatment with paclitaxel attenuated the decline in TER induced by TNF-α. However, it did not alone influence TER significantly. Data from experiments similar to that shown in Fig. 3A are summarized in terms of % reduction in TER (Fig. 3B) assessed at 4 h intervals. It is clear that paclitaxel significantly opposes the TNF-α response beyond 12 h.
As an alternative microtubule stabilizing agent, we tested the efficacy of epothilone B. Similar to paclitaxel, pretreatment with epothilone B also opposed the TNF-α response (Fig. 4A). Furthermore, epothilone B alone led to a small increase in TER. Histogram analysis as before is shown in Fig. 4B. It is evident that epothilone B significantly opposes TNF-α response starting from 4 h itself. Thus, the effect of epothilone B on the (TNF-α)-induced decline in TER is pronounced compared to paclitaxel. But, however, it is evident that both microtubule agents elicit similar influence to the TNF-α response, which induces disassembly of microtubules. Therefore, (TNF-α)-induced microtubule disassembly underlies the loss in barrier integrity.
In order to confirm that the disassembly of microtubules contributes to the observed loss in barrier integrity, we also investigated the apical junctional complex by immunofluorescence. Specifically, we examined disposition of peri-junctional actomyosin ring (PAMR), ZO-1, and cadherins in response to TNF-α with and without the microtubule stabilizing agents.
ZO-1, a cytoplasmic marker of tight junctions and localized contiguous with that of the neighboring cells, stains uniformly along the cell borders when the monolayer exhibits significant barrier integrity (Fig 5A). This disposition was disturbed upon treatment with TNF-α for 6 h (Fig. 5B). However, upon pre-treatment with paclitaxel or epothilone B, the distribution of ZO-1 was largely unaltered in response to TNF-α (Figs. 5D and 5F respectively). This finding is consistent with the efficacy of both agents in opposing TNF-α response in terms of reduced TER and increase in FITC dextran flux. In contrast with ZO-1, which is closely associated with the tight junctions, cadherins represent transmembrane domains of the adherens junctions. Hence, to examine the influence of TNF-α on the adherens junctions, we examined localization of cadherins by immunofluorescence using a pan-cadherin antibody. In untreated cells, similar to ZO-1, cadherins distribution is contiguous with that of the neighboring cells, and accordingly densely stained at the intercellular borders (Fig. 6A). Exposure to TNF-α for 6 h led to a considerable dispersion in the distribution of cadherins at the intercellular regions leading to a compromise in cell-cell contacts (Fig. 6B). On treatment with paclitaxel or epothilone B, the cadherins distribution was largely unaltered and similar to that of untreated cells (Fig. 6C and 6E, respectively). Pre-treatment with paclitaxel or epothilone B (Fig. 6D and 6F, respectively) opposed the TNF-α response. These results are also in concordance with the observation that the microtubule disassembly underlies the loss of barrier integrity in response to TNF-α.
A significant component of the apical junctions in the polarized epithelia, such as the corneal endothelium, is the presence of a dense of band of actin cytoskeleton proximal to tight and adherens junctions, which has been referred to as the peri-junctional actomyosin ring (PAMR). When challenged by agents that enhance actomyosin contraction, the structural integrity of the PAMR is disturbed. The disruption of the PAMR, which is structurally linked to the proteins of the TJs and AJs, is known to reorganize the apical junction. This instability presumably is responsible for the breakdown of the barrier integrity in addition to loss of cell-cell tethering secondary to centripetal forces generated in the PAMR following actomyosin contraction. Thus, in order to determine if the (TNF-α)-induced redistribution of ZO-1 and cadherins could be attributed to the disruption of the PAMR, we characterized its reorganization following exposure to the cytokine. As shown in Fig. 7B, exposure to TNF-α resulted in significant disruption of the PAMR. Treatment with paclitaxel or epothilone B did not alter the organization of PAMR to a significant extent (Figs. 7C and 7E respectively). However, TNF-α failed to show a similar disruption of the PAMR upon pre-treatment with paclitaxel or epothilone B (Fig. 7D and 7F, respectively). Taken together, these findings suggest that TNF-α response includes significant alteration of the apical junctional complex concomitant with the disassembly of the microtubules.
The corneal endothelium, as a monolayer at the posterior surface of the cornea, is responsible for keeping the cornea transparent. The maintenance of transparency requires that hydration of the corneal stroma to be held constant. The endothelium is able to actively pump fluid that leaks into the stroma from the aqueous humor which bathes its apical surface. The leakage of fluid into the stroma is dependent on the barrier function of the endothelium, which is the focus of this study. When the barrier integrity fails, cornea becomes edematous since the fluid pump, which drives fluid flow in the direction opposite to that of the leak, is overwhelmed. In several recent studies, we have investigated the mechanisms by which inflammatory mediators elicit breakdown in barrier integrity and intercellular communication through altered cytoskeleton (Satpathy et al. 2004; Satpathy et al. 2005; Srinivas et al. 2006; D’Hondt et al. 2007; D’Hondt et al. 2007; Ponsaerts et al. 2008). In this study, our major goal was to investigate how TNF-α breaks down the barrier integrity in a commonly used cell culture model of the corneal endothelium. In this study, for the first time, we have demonstrated that (TNF-α)-induced loss of barrier integrity in the corneal endothelium occurs largely through disassembly of microtubules and accordingly, we have also shown that microtubule stabilization effectively opposes the response to the cytokine.
A number of studies have examined the influence of TNF-α on barrier integrity in a variety of epithelial and endothelial monolayers (Goldblum et al. 1993; Nwariaku et al. 2002; Petrache et al. 2003; Ma et al. 2004; McKenzie and Ridley 2007). As a pleiotropic cytokine, it is known to induce barrier dysfunction through a variety of mechanisms including disruption of actin cytoskeleton (Goldblum et al. 1993; Watsky et al. 1996), activation of ROS (reactive oxygen species) (Natarajan et al. 1998; Usatyuk et al. 2003), RhoA GTPase (McKenzie and Ridley 2007), MAPKs (Nwariaku et al. 2002; Petrache et al. 2003), transcriptional activation of MLCK (myosin light chain kinase) (Petrache et al. 2001; Ma et al. 2005; McKenzie and Ridley 2007) and/or Hsp27 (heat shock protein 27) (Kiemer et al. 2002), and microtubule disassembly (Petrache et al. 2003). Although Watsky et al documented the disruption of the actin cytoskeleton following exposure to TNF-α in the rabbit corneal endothelium, their study did not consider a role for microtubules (Watsky et al. 1996). In this study, we focused on microtubule disassembly since it has been recently reported that cytokines can induce the disassembly leading to disruption of actin cytoskeleton (Petrache et al. 2003; Xu et al. 2008).
In consistence with our main goal, we began this study by examining whether TNF-α can induce microtubule disassembly. As expected, we consistently observed loss of microtubule staining in the cellular periphery, which indicates its depolymerization. The extent to which depolymerization occurs in response to TNF-α is not evident when cells are pretreated with paclitaxel and epothilone B (Fig. 1D and 1F, respectively). It is possible that excessive polymerization induced by these drugs may have masked the TNF-α response, but we note that the net level of microtubule polymerization is significantly greater in cells pretreated with paclitaxel and epothilone B. Thus, we conclude from Fig. 1 that TNF-α induces microtubule disassembly and that the response to the cytokine is not noticeable in cells pre-treated with microtubule-stabilizing agents. Similar observations have been noted in pulmonary artery endothelial cells (Petrache et al. 2003).
Having established the influence of TNF-α on microtubules, we next examined the barrier integrity by two distinct methods. We first confirmed the TNF-α response on barrier integrity by examining the flux of FITC dextran in cells treated with the cytokine and compared with those pre-treated with paclitaxel and epothilone B. While TNF-α treatment alone increased the permeability significantly, microtubule stabilization masked the response to the cytokine (Fig. 2). This is our first indication to suggest that microtubule disassembly at the cellular periphery is responsible for loss of the barrier integrity. Despite this clear indication, the temporal response to TNF-α with and without microtubule stabilization is not evident. Hence, we took recourse to the ECIS approach, which we had employed to investigate the loss of barrier function upon deliberate microtubule disassembly by nocodazole (Jalimarada et al. 2009). As demonstrated in Figs 3 and and4,4, it is clear that exposure to TNF-α results in a sustained decline in TER, which is attenuated by both paclitaxel and epothilone B (Figs. 3 and and4,4, respectively). It may be noted that the opposition to TNF-α response by epothilone B is significantly more profound than paclitaxel. Although we did not perform a dose response study, it is important to note that epothilone B is known to have higher potency in stabilizing the microtubules compared to paclitaxel (Kowalski et al. 1997; Altmann et al. 2000).
In order to gain further insights into the (TNF-α)-induced loss of barrier integrity, we investigated structural instability at the apical junction by immunofluorescence. In addition to ZO-1 and cadherins as markers of tight and adherence junctions, we examined PAMR, which lies proximal to the tight and adherens junctions. Furthermore, PAMR is structurally linked to both tight and adherens junctions and hence its disruption results in their remodeling which may underlie the loss in barrier integrity. In Figs. 5 and and6,6, we have demonstrated that TNF-α induces dislocation of ZO-1 and cadherin, respectively, in consistence with the loss of barrier integrity induced by the cytokine. In agreement with masking of the effect of TNF-α on barrier integrity by paclitaxel and epothilone B, we also show that the cytokine-induced dispersions of ZO-1 and cadherins are also attenuated by microtubule stabilization (Figs. 5D, 5F, 6D, and 6F, respectively). These results reaffirm that the cytokine-induced loss in barrier integrity is brought about by microtubule disassembly. Of special significance is the finding in Fig. 7 in which we demonstrate that microtubule disassembly by the cytokine also disrupts the PAMR. This is in consistence with the dispersion of ZO-1 and cadherins from their normal locus as shown in Fig. 5 and Fig. 6.
The disruption of the PAMR secondary to microtubule disassembly caused by the cytokine is reminiscent of our recent work in which we demonstrated a similar effect in response to nocodazole (Jalimarada et al. 2009). As reported with pulmonary artery endothelial cells, we had found increased MLC phosphorylation secondary to microtubule disassembly. This was attributed to activation of small GTPase RhoA. While it is tempting to speculate similar downstream mechanisms secondary to microtubule disassembly induced by TNF-α, the pleiotropic nature of the cytokine and lack of response to Rho kinase inhibitors (data not shown) to subdue its response prevented us from further emphasis on RhoA-Rho kinase pathway. Similar to uncertainty in the downstream targets of the (TNF-α)-induced microtubule disassembly, the upstream mechanism(s) are also not well established. It has been reported that increased oxidative stress can increase iNOS activity, leading to decreased tubulin polymerization and consequent disruption of the microtubules (Banan et al. 2003). A study also suggests that ROS (reactive oxygen species) activation is mediated by TNF-α signaling (Rajashekhar et al. 2007). Hence, the TNF-α-induced microtubule disassembly may also involve upregulation of ROS and iNOS. Recent studies suggest that p38 MAP kinase is involved in TNF-α-mediated increase in endothelial permeability (Petrache et al. 2003). It has also been shown in human and bovine pulmonary artery endothelial cells that pharmacological inhibition of p38 MAP kinase attenuates nocodazole-induced microtubule depolymerization, actin remodeling, and barrier dysfunction (Birukova et al. 2005). Studies also suggest that p38 MAP kinase activation can induce phosphorylation of microtubule associated proteins such as tau, thereby reducing its ability to promote microtubule assembly (Goedert et al. 1997). It remains to be seen whether activation of p38 MAP kinase by TNF-α is responsible for microtubule disassembly and thereby the downstream events associated with the loss of corneal endothelial barrier function.
In conclusion, for the first time, we have shown that microtubule disassembly by TNF-α contributes to the barrier dysfunction in corneal endothelium. Hence, inhibition of microtubule disassembly through its stabilization may be effective in reducing corneal edema occurring as a result of acute graft rejection or anterior uveitis. However, further studies are necessary to delineate the signaling pathways activated by TNF-α, leading to microtubule disassembly and investigate potential adverse effects of microtubule stabilizing agents.
Supported by NIH grant R21-EY019119 (SPS) and Faculty Research Grant, VP of Research, IU Bloomington, IN (SPS).
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