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GROWTH, DIFFERENTIATION, AND APOPTOSIS
-induced epithelial barrier destabilization and tissue repair1Division of Physiology, Department of Physiology and Medical Physics, and 2Department of Therapeutic Radiology and Oncology, Innsbruck Medical University, Innsbruck, Austria
Submitted 17 August 2007 ; accepted in final form 2 October 2007
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ABSTRACT |
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, a cytokine crucial for linking innate and adaptive immune responses. EGF was implicated in rescue mechanisms from renal injury. To study the interplay between the two processes, we determined if EGF can prevent IFN--induced barrier permeabilization. EGF did not counteract but even exacerbated the IFN--induced decrease of transepithelial electrical resistance in LLC-PK1 monolayers. For this effect Erk1/2 activation was necessary, linking barrier regulation to EGF-induced cell cycle progression. In contrast to its damage-intensifying effect, EGF also facilitated the regeneration of epithelial barrier function after the termination of IFN- treatment. This effect was not mediated by Erk1/2 activation or cell proliferation since U0126, an Erk1/2 inhibitor, did not prevent but ameliorated recovery. However, EGF accelerated the downregulation of caspase-3 in recovering cells. Similarly, a pan-caspase inhibitor was able to block caspase activity and, concomitantly, promote restoration of barrier function. Thus, barrier repair might be linked to an EGF-mediated antiapoptotic mechanism. EGF appears to sensitize epithelial cells to the detrimental effects of IFN- but also helps to restore barrier function in the healing phase. The observed dual effect of EGF might be explained by the different impact of proproliferative and antiapoptotic signaling pathways during and after cytokine treatment. The timing of epithelial exposure to damaging agents and repair factors was identified as a crucial parameter determining tissue fate. cytokine; proximal tubular cells; proliferation; type I interferon; apoptosis
A pivotal process in tissue repair following injury is the replacement of proximal tubular cells lost by cell death. Although the potential implications of bone marrow and/or renal stem cells in renal regenerative processes have been discussed (6), the extent to which stem cells may participate in renal repair processes has not yet been determined. It is clear, however, that resident differentiated proximal tubular cells are a major source of newly generated tubular cells (16, 19). Healthy cells have been shown to transiently dedifferentiate and increase their proliferative capacity after sublethal injury (5, 8, 27).
Cell cycle progression is induced by growth factors, which have been implicated as autocrine or paracrine regulators in proximal tubular recovery (18, 23). EGF receptor activation has been found to be required for proximal tubular cell proliferation and migration after mechanical damage in vitro (31). In addition, functional EGF and EGF receptor activity have been demonstrated to play a crucial role in recovery from acute nephrotoxic injury in vivo (29). Furthermore, EGF has been shown to reduce the extent of renal dysfunction, accelerate recovery of tubular cells, and decrease mortality when administered to animals after ischemia (10). However, long-term treatment with high levels of EGF is suspected to promote the development of renal carcinoma (26). Overexpression of growth factors, furthermore, has been associated with renal deterioration processes due to uncontrolled proliferation characterized by tubulointerstitial lesions (25). Therefore, further insights into the regulation of growth factor-induced renal tissue repair are needed.
A major intracellular signaling pathway induced by EGF is the MAPK pathway. EGF receptor tyrosine phosphorylation has been shown to create docking sites for the assembly of a signaling complex leading to the activation of small G proteins of the Ras family. This process activates a protein kinase cascade leading to the phosphorylation and thus activation of Erk1/2 by Mek1/2. This Mek1/2-Erk1/2 pathway has been implicated in the regulation of proliferation and cell survival (30).
While most studies have concentrated on the role of cell proliferation by EGF in response to damage, EGF might also be implicated in the preservation or regeneration of epithelial barrier function. Since EGF treatment of renal epithelial monolayers in vitro has been shown to cause an increase of transepithelial electrical resistance (TEER) (9, 22, 24), we hypothesized that barrier stabilization by EGF could counteract barrier permeabilization by damaging agents such as inflammatory cytokines.
We have previously identified IFN- as an epithelial barrier-destabilizing factor since it has been found to decrease TEER in proximal tubular monolayers (12, 13). IFN- is released by activated immune cells and plays a key role for linking innate and adaptive immune responses (4). Release of cytokines is thought to cause damage to the proximal tubular barrier, compromising kidney function. "Backleak" of glomerular filtrate into the renal interstitium might thus be induced, causing edema and a further exacerbation of the inflammatory process (14). Epithelial damage by inflammatory cytokines might also be relevant for ischemia-reperfusion injury and toxin-induced ARF due to locally activated inflammatory mechanisms (21).
The present study investigated the effect of EGF on IFN--mediated renal epithelial barrier destabilization. Specifically, we studied if EGF prevented IFN--induced barrier permeabilization or influenced recovery phases after the termination of IFN- treatment.
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MATERIALS AND METHODS |
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5 days followed by serum-free medium for at least 1 day before the start of the experiment. Well-established confluence was carefully controlled for by microscopic inspection, establishment of a stable TEER, and the ability to maintain a FITC-inulin (Sigma-Aldrich) apicobasolateral gradient. All experiments were performed in serum-free culture medium. Human recombinant EGF was purchased from Sigma-Aldrich. The IFN- preparation commonly used for the described experiments was recombinant human IFN-2b in phosphate buffer (Strathmann-Biotech, Hamburg, Germany). Alternatively, a clinical IFN-2b preparation (IntronA, Schering, Berlin, Germany) was also used, yielding the same results. U0126, a specific inhibitor of the Erk1/2-activating kinases Mek1/2, was from Promega (Mannheim, Germany). U0126 treatment was started 30 min before the addition of IFN- and/or EGF.
TEER measurements.
TEER was measured with fixed electrodes utilizing an Endohm (World Precision Instruments, Sarasota, FL) coupled to an Evom volt-ohm meter (Millipore, Vienna, Austria). TEER values (in 
·cm2) were calculated by multiplying raw values by the surface area of the filter and subtracting the TEER of empty filters. TEER is expressed either as the difference to time-matched controls (in ·cm2) or as a percentage of time-matched controls. The mean TEER value (±SD) of controls was 95.6 (±11.4) ·cm2.
Bromodeoxyuridine proliferation assay and cell cycle analysis by flow cytometry. Confluent serum-starved LLC-PK1 cells on 25-mm microporous growth supports were labeled with 20 µM bromodeoxyuridine (BrdU; Roche Diagnostics, Mannheim, Germany) in cell culture medium for 2 h. Adherent cells were harvested by trypsination and pooled with cells floating in the supernatant medium. Fixation was performed in 70% ethanol for 30 min on ice followed by storage at –20°C for at least 1 day. BrdU was detected by staining with an anti-BrdU Alexa fluor488-labeled antibody (Molecular Probes-Invitrogen, Paisley, UK) after treatment of the cells with 4 N HCl for 30 min at room temperature. DNA was stained with propidium iodide. Fluorescence was measured in a FACScan flow cytometer (Becton-Dickinson, Heidelberg, Germany) at 488-nm excitation, and emission was determined in fluorescence 1 (FL-1; 530/30-nm band pass) and fluorescence 3 (FL-3; 650-nm long pass) channels. At least 25,000 events in G0/G1, S and G2/M phases were analyzed in slow mode excluding debris and aggregates. Quantitative analysis of BrdU-labeled cells was performed on dot plots of FL-1 versus FL-3 using Cellquest software (Becton-Dickinson). Cell cycle phases were, in addition, calculated from histograms with linear FL-3 area recordings using Modfit software (Becton-Dickinson).
Western blot analysis. LLC-PK1 cells were cultured on 25-mm microporous growth supports. Whole cell extracts were prepared by rinsing monolayers with ice-cold PBS and scraping in lysis buffer (50 mM Tris·HCl, 100 mM NaCl, 50 mM NaF, 40 mM β-glycerophosphate, 5 mM EDTA, and 1% Triton-X 100) containing protease inhibitor cocktail (Sigma-Aldrich) and Na-orthovanadate. The protein content of LLC-PK1 cell homogenates was determined with the BCA protein determination reagent (Pierce, Rockford, IL), and samples were matched with lysis buffer. Homogenates of identical protein content were boiled for 3 min and applied to gels for SDS-PAGE. For immunoblot analysis, proteins were transferred from gels to an Immobilon P membrane (Millipore, Vienna, Austria), which was incubated with the primary antibody (anti-phospho-Erk1/2 or anti-β-actin antibody from Cell Signaling Biolabs, Frankfurt, Germany) after being blocked in 5% (wt/vol) nonfat dry milk or BSA according to the manufacturer's recommendations. Secondary antibodies labeled with horseradish peroxidase were used (Sigma-Aldrich or Cell Signaling Biolabs). Enhanced chemiluminescence (ECL or ECL plus, GE Healthcare, Vienna, Austria) or the Phototope-HRP Western blot detection system (Cell Signaling Biolabs) and Hyperfilm (GE Healthcare) were used for detection.
Caspase-3 enzymatic assay.
Caspase-3 activity was determined using the caspase-3 fluorescence-based enzymatic assay (Molecular Probes, Leiden, The Netherlands) according to the manufacturer's instructions. LLC-PK1 cells were grown on 10-mm microporous growth supports. Cellular extracts were obtained by incubating cells on ice for 30 min in lysis buffer (10 mM Tris, 1 mM EDTA, and 1% Triton X-100; pH 7.4). Caspase-3 activity was measured by cleavage of rhodamine 110-conjugated bis-N-acetyl-L-aspartyl-L-glutamyl-L-valyl-L-aspartic acid amide (Ac-DEVD-R110). Fluorescence was measured in a spectrofluorimeter (Tecan/Grödig, Austria) at excitation/emission wavelengths of 485/535 nm. Fluorescence signals were corrected by subtracting background fluorescence and also for protein content determined by a BCA assay (Pierce) according to the manufacturer's instructions. Caspase-3 activity is expressed as fold over control. For caspase inhibition, 5 µM Z-Val-Ala-Asp(OCH3)-fluoromethylketone (Z-VAD-FMK), a cell-permeable irreversible pan caspase inhibitor (Calbiochem-Merck Biosciences, Bad Soden, Germany), was used.
Statistical analysis. Values are expressed as means ± SD. At least three independent experiments were performed in triplicate or more. Statistical evaluation was performed using an unpaired two-sided Student's t-test or ANOVA and Bonferroni comparison of the means where appropriate, utilizing the "OriginPro7" software package. P values of <0.05 were deemed statistically significant.
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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.
Since EGF has been implicated as a protective factor in kidney dysfunction, we investigated the influence of EGF on cytokine-induced proximal tubular barrier destabilization. However, EGF pretreatment for 24 h, which caused a transient increase in TEER, did not prevent the TEER decrease by IFN- (Fig. 1A). Concomitant treatment of LLC-PK1 epithelial monolayers with EGF and IFN- even resulted in a further decrease of TEER compared with IFN- monotreatment. EGF alone, in contrast, caused persistently increased TEER (Fig. 1B). Interestingly, exposure of LLC-PK1 cultures to EGF after IFN- withdrawal did accelerate TEER recovery (Fig. 1B), implicating EGF as a rescue factor in epithelial barrier repair.
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Role of proliferation and Mek1/2-Erk1/2 signaling in barrier destabilization by IFN-.
Since EGF is a key regulator of epithelial cell proliferation, we tested if cell cycle progression was involved in epithelial barrier regulation. IFN- was able to induce LLC-PK1 cells to progress into the S phase, as shown by flow cytometric analysis of BrdU-labeled cells (Fig. 2A, light gray bars) (see Supplemental Fig. s1, for a representation of the original flow cytometric data).1 The addition of EGF during IFN- treatment increased the BrdU-labeling, resulting in a similar proliferation rate as induced by EGF alone. Since simultaneous EGF and IFN- treatment has also been shown to decrease TEER more than IFN- alone (Figs. 1A and 2B, light gray bars), a higher proliferation rate thus appears to render cells more susceptible to IFN--induced barrier destabilization. On the other hand, treatment with EGF alone was able to stabilize epithelial barrier function, as shown by a TEER increase, confirming previous findings (22, 24). This result indicates that proliferation does not per se cause barrier destabilization. Cell cycle progression might, however, render epithelial barriers more susceptible to agents that cause barrier destabilization.
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and/or EGF was dependent on Erk1/2 activation. By flow cytometry of BrdU-labeled cells, it was demonstrated that U0126 was able to significantly reduce the percentage of cells in the S phase under all treatment conditions (Fig. 2A, solid bars).
To analyze how Erk1/2 activation influenced barrier regulation, the TEER of LLC-PK1 monolayers treated with U0126 was determined (Fig. 2B, solid bars). As a result, the TEER of IFN--treated or IFN- and EGF-treated cultures was significantly higher in the presence of U0126 than in its absence. This result implies that IFN--induced barrier destabilization depends on Erk1/2 activation. Since proliferation was decreased by U0126 and TEER was concomitantly increased, these data also suggest that proliferating cultures might be more susceptible to IFN--induced barrier destabilization. U0126 was also effective in blocking the effects of EGF alone on proliferation as well as on TEER.
To demonstrate Erk1/2 activation, Western blot analysis was performed utilizing a phosphorylation state-specific Erk1/2 antibody that selectively binds to the active form of Erk1 and Erk2. EGF induced a transient immediate-early activation of Erk1/2 (Fig. 2C) (see Supplemental Fig. s2, for gel loading controls with a β-actin antibody). Remarkably, IFN- was also able to induce phosphorylation of Erk1/2, but with different activation kinetics compared with EGF. The Erk1/2 phosphorylation induced by IFN- occurred after hours and was then sustained. Cotreatment with IFN- and EGF resulted in a combination of high-level early and late activation. U0126 effectively reduced Erk1/2 phosphorylation.
Role of proliferation and the Mek1/2-Erk1/2 pathway in barrier repair.
Since EGF not only exacerbated barrier destabilization during IFN- treatment but also promoted barrier repair after IFN- withdrawal, we next determined if proliferation was responsible for this EGF effect. In the recovery phase after IFN- removal, however, EGF had no effect on proliferation (Fig. 3A, light gray bars) (see Supplemental Fig. s3, for a representation of the original flow cytometric data). This may be due to the fact that cells recovering from treatment in medium only were already at the maximum proliferative capacity. In line with this finding, Western blot analysis showed that a high level of Erk1/2 phosphorylation was maintained after the termination of IFN- treatment independent of EGF treatment (Fig. 3C) (see Supplemental Fig. s4, containing a gel loading control with a β-actin antibody).
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withdrawal, we first treated the monolayers with IFN-. After the removal of IFN-, monolayers were exposed to EGF in the absence or presence of U0126. U0126 was able to counteract the cell cycle progression (Fig. 3A, solid bars) as well as Erk1/2 phosphorylation (Fig. 3C), revealing the importance of the Mek1/2-Erk1/2 pathway for this proliferative response. U0126 did not, however, prevent the effect of EGF during TEER recovery (Fig. 3B, solid bars). In contrast, U0126 per se had a favorable effect on TEER reestablishment. The highest TEER was obtained when EGF and U0126 were concomitantly added to the medium after IFN- withdrawal. These results suggest that barrier restoration was not mediated but even hindered by Erk1/2 activation. Since the Erk1/2 pathway has been shown to be necessary for the induction of cell cycle progression, the favorable effect of EGF in TEER recovery thus cannot be mediated by proliferation. Thus, we attempted to discover which alternative mechanism could be activated by EGF resulting in the accelerated recovery of barrier function after cytokine challenge.
Influence of antiapoptotic pathways in barrier repair.
Since we have recently found that IFN- was able to induce programmed cell death of LLC-PK1 cells (J. Lechner, unpublished observations), we investigated the role of apoptosis on EGF-induced barrier alterations. We hypothesized that a downregulation of apoptotic pathways during recovery may be favorable to restoration of barrier function after IFN- withdrawal. To test this hypothesis, LLC-PK1 monolayers were exposed to a pan-caspase inhibitor (Z-VAD-FMK) following IFN- treatment. In fact, TEER recovery after IFN- withdrawal was accelerated by the presence of the pan-caspase inhibitor (Fig. 4A), similar to that observed for EGF (Fig. 1B). However, the pan-caspase inhibitor (as with EGF) did not prevent barrier destabilization when it was present before and during IFN- treatment (J. Lechner, unpublished observations).
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withdrawal appeared to be hindered by apoptotic cell death, we analyzed if apoptotic signaling was still detectable after IFN- withdrawal. Twenty-four hours after the termination of IFN- treatment, caspase-3 activity was found to be elevated (Fig. 4B). At 48 h of recovery, caspase-3 activity was similar to controls (data not shown). Remarkably, EGF was able to significantly accelerate the downregulation of active caspase-3 in recovering cell populations (Fig. 4B). These results establish a link between the beneficial effects of EGF during TEER recovery and EGF-induced antiapoptotic signaling.
However, in the IFN- induction phase, EGF did not prevent but exacerbated apoptosis (Fig. 4C). A similar effect was obtained by pretreating cells with EGF for 24 h before the start of IFN- incubation. IFN--induced caspase-3 activation was also increased in subconfluent proliferating cultures.
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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was applied, EGF did not protect but even exacerbated injury. EGF was beneficial in the recovery phase after IFN- removal. Thus, EGF has not only the potential to aid tissue repair but also to enhance injury processes. The aim of this study was to attempt to unravel the distinct pathways that may be responsible for these alternate effects of EGF.
Proliferation increases the susceptibility of epithelial barriers to IFN--induced destabilization.
Since EGF is a key regulator of epithelial cell proliferation, we analyzed if mitosis triggered by EGF had an impact on the barrier stability of LLC-PK1 monolayers. Cotreatment with EGF and IFN- resulted in an increased proliferation rate compared with IFN- monotreatment. Since cotreatment also decreased TEER more than IFN- alone, the higher proliferation rate correlated with an increased susceptibility to IFN--induced TEER decreases.
U0126 was utilized to specifically inhibit the Mek1/2-Erk1/2 signaling pathway. Blockade of Erk1/2 activation by U0126 not only attenuated cell cycle progression but also concurrently counteracted the TEER decrease. The lower proliferation rate, thus, was linked to a higher TEER, showing (again) an inverse correlation between proliferation and barrier stability. Therefore, exacerbation of IFN--induced epithelial barrier dysfunction by EGF may be the result of enhanced proliferation.
Mitotic cell division and epithelial barrier stability. The question remains as to by which mechanism cell proliferation might influence epithelial barrier function. Mitosis involves rounding up of cells during cytokinesis, an event that potentially affects epithelial barrier function by destabilizing cell-cell contacts. However, proliferation induced by EGF treatment alone was previously found to not increase but even decrease transepithelial permeability (22, 24). This apparent contradiction might be resolved by assuming that EGF (in addition to proliferation) was able to promote cellular counterreactions leading to epithelial barrier stabilization. In fact, EGF and other growth factors have been shown to induce a preferential expression of supposedly barrier tightness-conferring claudins (17, 24). These proteins are discussed as regulators of tight junctional (TJ) permeability characteristics (3, 7). Thus, concomitant restructuring of TJ strands leading to a TEER increase could counteract the destabilizing effects conferred by proliferation.
While EGF monotreatment resulted in overall barrier stabilization, IFN- cotreatment with EGF led to an exacerbation of the TEER decrease. Junctional complex disorganization by IFN- might potentially outweigh the TJ tightening effects of EGF. Thus, the barrier destabilizing component, potentially conferred by EGF-enhanced proliferation, might prevail under these treatment conditions. The result might be an augmented barrier-destabilizing effect by EGF and IFN- cotreatment compared with IFN- treatment alone, as was observed.
The Erk1/2 phosphorylation pattern: early short-term activation by EGF versus late sustained activation by IFN-.
Both EGF and IFN- were able to induce Erk1/2 phosphorylation, but with different kinetics. Erk1/2 phosphorylation was reached after minutes by EGF treatment compared with hours with IFN- treatment. Induction of immediate-early gene expression by EGF has been previously shown to be necessary for cell cycle progression (30). The delayed onset of Erk1/2 activation by IFN- exposure might induce those genes less efficiently, thereby causing a lower proliferation rate. IFN- and EGF cotreatment resulted in a combination of early and late activation accompanied by a proliferation rate equal to EGF monotreatment.
EGF and Erk1/2 signaling display prodeath and prosurvival functions.
Our results implicate that EGF and Erk1/2 signaling do not protect renal cells from barrier destabilization by IFN-. These findings do not support a general repair and survival function of EGF and Erk1/2. Similar observations have been reported in proximal tubular cells treated with other damaging agents. Inhibition of EGF receptor-Erk1/2 signaling has been shown to prevent cisplatin-induced cell death in mouse proximal tubular cells (1, 11). In addition, cisplatin cytotoxicity was reported to depend on Cdk2 activation, linking it to cell proliferation (20). A proapoptotic role of Erk1/2 was furthermore demonstrated in cellular damage by reactive oxygen species since concomitant Erk1/2 activation promoted hydrogen peroxide-induced apoptosis in renal epithelial cells (33). Another group, however, found that sustained activation of Erk1/2 via the canonical EGF receptor pathway favored cell survival after oxidant-mediated injury to proximal tubular cells (2). A detailed evaluation revealed that this rescue effect might result from Erk1/2 activation during recovery from injury. Thus, the outcome might be mainly determined by the timing of EGF receptor and Erk1/2 activation in relation to the damaging process. This explanation might resolve the apparent conflict in the literature describing the growth factor-Erk1/2 pathway as both a prodeath and prosurvival signal (28, 32). Our present report adds evidence to this hypothesis. EGF was shown to induce proliferation and, concomitantly, increase the susceptibility of cells to IFN--induced barrier destabilization and caspase activation. While replacing dead cells by mitosis is obviously necessary to eventually repair a damaged epithelium, dividing cells also appear to be more susceptible to influences inducing barrier destabilization and apoptosis. Thus, the kinetics of the injury process and the onset of Erk1/2 signaling appear to determine which of the dual roles prevails: a prosurvival effect or an intensification of damage.
Barrier restoration is not accelerated but hindered by the proliferative response following injury.
We observed a proliferative response of cells in recovery after challenge by IFN-. As demonstrated, this proliferative response was dependent on Erk1/2 activation and could not be further increased by supplementing the medium with EGF. To analyze if restoration of barrier tightness after IFN- withdrawal was dependent on this proliferative response, we blocked Erk1/2 activation by U0126, thereby inhibiting cell cycle progression. Blockade of the proliferation, however, did not prevent but rather accelerated reestablishment of barrier tightness after the termination of IFN- treatment.
Barrier repair is favored by EGF through an antiapoptotic mechanism.
Since proliferation was not favorable to barrier repair, we sought for another potentially responsible mechanism to explain the favorable effect of EGF in barrier restoration after IFN- withdrawal. To analyze if persistent apoptosis hindered reestablishment of barrier tightness after removal of IFN-, we made use of a pan-caspase inhibitor. This inhibitor effectively blocked apoptosis by downregulation of the caspase cascade. Confirming our hypothesis, the pan-caspase inhibitor had a favorable effect on the recovery of epithelial barrier function. Thus, apoptotic signaling significantly compromised the reestablishment of barrier tightness.
In addition, monolayers recovering from IFN--induced damage in medium alone showed a higher degree of caspase-3 activity than cultures treated with EGF after IFN- withdrawal. These findings suggest that the barrier-rescuing function of EGF might be linked to the downregulation of apoptotic pathways.
However, the pan-caspase inhibitor was not able to prevent barrier destabilization by IFN- if the cells were treated with the inhibitor before or concomitantly with IFN-. While the pan-caspase inhibitor efficiently reduced apoptotic cell death, the TEER decrease induced by IFN- was not affected (J. Lechner, unpublished observations). This previous finding indicates that barrier destabilization is not necessarily coupled to apoptotic cell death and may be mediated by apoptosis-independent mechanisms as well.
Furthermore, EGF was not able to prevent the onset of IFN--induced apoptosis since caspase-3 activity after concurrent treatment was not reduced but even increased compared with IFN- monotreatment. Enhanced apoptotic rates were also obtained by EGF pretreatment and using subconfluent growing LLC-PK1 cells. Since these treatment conditions involved increased cell cycle progression, the higher susceptibility of the cells to apoptosis may be due to actively cycling cells more easily entering an apoptotic pathway. Thus, EGF was found to display a dual, timing-dependent effect on IFN--induced apoptosis. These findings mirror the opposing effects of EGF on IFN--mediated barrier destabilization.
Summary and conclusions.
The data presented have shown that IFN--induced epithelial destabilization was not prevented but rather exacerbated by EGF. Erk1/2 signaling was necessary for this effect, linking it to enhanced cell proliferation. In addition to its damage-intensifying effect, EGF also positively influenced epithelial barrier recovery when administered after IFN- withdrawal. This action correlated with an antiapoptotic mechanism induced by EGF and was independent of proliferation.
The timing of EGF signaling and onset of proliferation were identified as crucial parameters determining tissue fate. Thus, EGF might play opposing roles in cytokine-induced epithelial tissue injury: exacerbating dysfunction of normal tissue during the presence of the damaging agent and favoring the recovery of barrier function in the healing phase. Seemingly contradicting results in the literature concerning the role of EGF as a mediator of restorative as well as deteriorative processes might potentially be explained if the timing of the EGF signaling versus damaging events is taken into account. Therefore, the possible dual nature of growth factors on epithelial tissue function and integrity should be considered whenever growth factor signaling is manipulated with a therapeutic goal.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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.
1 Supplemental material for this article is available online at the American Journal of Physiology-Cell Physiology website. 
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) or concomitantly with IFN-
) and compared with treatment with IFN-
). The 0-h time point denotes the addition of IFN-
). TEER reestablishment was compared with the recovery in serum-free medium without further additives (
). At least 8 independent treatments were analyzed. Values are represented as means ± SD. *Statistically significant difference (P < 0.05) vs. IFN-
