Bryostatin-1 attenuates TNF-induced epithelial barrier dysfunction: role of novel PKC isozymes

James Yoo1, Anthony Nichols2, Jaekyung C. Song2, Joshua Mammen2, Isabel Calvo2, Roger T. Worrell2, John Cuppoletti2, Karl Matlin2, and Jeffrey B. Matthews2

1 Department of Surgery, Beth Israel Deaconess Medical Center, Boston, Massachusetts 02215; and 2 Department of Surgery, University of Cincinnati Medical Center, Cincinnati, Ohio 45267


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Tumor necrosis factor (TNF) increases epithelial permeability in many model systems. Protein kinase C (PKC) isozymes regulate epithelial barrier function and alter ligand-receptor interactions. We sought to define the impact of PKC on TNF-induced barrier dysfunction in T84 intestinal epithelia. TNF induced a dose- and time-dependent fall in transepithelial electrical resistance (TER) and an increase in [3H]mannitol flux. The TNF-induced fall in TER was not PKC mediated but was prevented by pretreatment with bryostatin-1, a PKC agonist. As demonstrated by a pattern of sensitivity to pharmacological inhibitors of PKC, this epithelial barrier preservation was mediated by novel PKC isozymes. Bryostatin-1 reduced TNF receptor (TNF-R1) surface availability, as demonstrated by radiolabeled TNF binding and cell surface biotinylation assays, and increased TNF-R1 receptor shedding. The pattern of sensitivity to isozyme-selective PKC inhibitors suggested that these effects were mediated by activation of PKC-epsilon . In addition, after bryostatin-1 treatment, PKC-delta and TNF-R1 became associated, as determined by mutual coimmunoprecipitation assay, which has been shown to lead to receptor desensitization in neutrophils. TNF-induced barrier dysfunction occurs independently of PKC, but selective modulation of novel PKC isozymes may regulate TNF-R1 signaling.

protein kinase C; tumor necrosis factor; epithelial barrier function


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

TUMOR NECROSIS FACTOR (TNF) is a 17-kDa cytokine that has been implicated in the pathogenesis of numerous ischemic and inflammatory diseases such as Crohn's disease, Behcet's disease, and rheumatoid arthritis (19, 49). Elevated mucosal levels of TNF are thought to contribute to the chronic inflammation, diarrhea, and increased mucosal permeability seen in patients with inflammatory bowel disease and may be important in initiating and perpetuating general inflammatory and ischemic states. Two high-affinity TNF receptors, p55 (type I, TNF-R1) and p75 (type II, TNF-R2), have been identified and are located on the basolateral surface of intestinal epithelia (16). TNF-R1 appears to be the dominant receptor involved in most of the known actions of TNF (14, 21).

An important property of intestinal epithelia is its ability to form a selectively permeable barrier that separates the internal and external environments. Barrier function, which is thought to reside in the tight junctional complexes between adjacent epithelial cells, is known to be perturbed under a variety of inflammatory, infectious, and ischemic enteropathies. TNF is one of several cytokines that has been shown to disrupt epithelial barrier function in a number of in vivo and in vitro models (14, 16, 17, 24, 32, 34, 42). However, the mechanism of TNF-induced barrier dysfunction remains incompletely understood.

Protein kinase C (PKC) represents a family of serine-threonine kinases involved in the modulation of diverse cellular processes, including epithelial barrier function (10, 11, 43, 45, 46). PKC is thought to be involved in junction reformation after calcium switch (2) by a mechanism that involves the regulated assembly and subcellular localization of tight junction proteins (2, 28) as well as their phosphorylation states (2, 11). There are at least 11 different isozymes of PKC classified into three broad groups. The classic PKC isozymes (alpha , beta I, beta II, and gamma ) are both Ca2+ and diacylglycerol (DAG) dependent. The novel PKC isozymes (delta , epsilon , eta , and theta ) are Ca2+ independent but DAG dependent. The atypical PKC isozymes (zeta , iota , and lambda ) are neither Ca2+ nor DAG dependent. Specificity of individual PKC isozyme action is thought to be conferred by translocation to, and interaction with, membrane-bound target proteins that recognize the distinct isozymes. A number of instances have been identified in which specific isozymes affect a given cell function in similar ways as well as in opposite ways.

PKC has been linked to a number of TNF-mediated processes (15, 18, 27, 31, 33, 50). PKC activity is elevated in colonic samples of inflammatory bowel disease patients, where TNF levels are also known to be elevated (39). In various model systems, PKC isozymes have variously been found to be either protective against or to directly mediate the pathological effects of TNF (22, 33, 50), suggesting that a counterregulatory relationship among PKC isozymes could account for the divergent experimental results. Evidence exists that activation or inhibition of specific PKC isozymes may protect against inflammatory and ischemic injury (6, 9, 52). In the heart, the novel PKC-epsilon isozyme is activated by ischemia and has been shown to protect against ischemic insults through a process known as preconditioning (9); PKC-delta under these circumstances appears to exacerbate ischemic damage (12). PKC-epsilon also appears to dampen the Cl- secretory response in a model of intestinal epithelial ischemia (51).

In the present study, we used the T84 human intestinal epithelial cell line to study the relationship between TNF and PKC on epithelial barrier function. T84 cells have been widely adopted as a model for studying the role of various inflammatory cytokines and enteric bacteria on intestinal barrier function (7, 16, 24, 32), and we and others (10, 20, 43, 46, 51) have previously shown that activation of PKC regulates barrier function in T84 cells. To examine the effects of TNF on PKC, we studied whether PKC activation is related to TNF-induced barrier dysfunction and whether pharmacological modulation of PKC alters the effects of TNF. Our results suggest that the novel isozymes PKC-delta and -epsilon interact with TNF signaling at the level of its membrane receptor and may mitigate the pathological effects of TNF on barrier function in model T84 epithelia.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Cell culture. T84 cells obtained from the American Type Culture Collection (Manassas, VA) were grown in a 5% CO2 humidified incubator at 37° on 162-cm2 flasks (Corning Costar, Acton, MA) with media containing a 1:1 mixture of Ham's F-12 nutrient mixture and DMEM supplemented with 6% heat-inactivated fetal bovine serum, 15 mM HEPES, 14.3 mM NaHCO3, and antibiotics-antimycotics (100 U/ml aqueous penicillin G, 100 µg/ml streptomycin, 250 ng/ml amphotericin B) at a pH of 7.4. Cells were passaged weekly on reaching confluence. For experiments, cells were plated onto collagen-coated permeable supports, where they were fed every 3 days and were maintained until steady-state transepithelial resistance (TER) was achieved and were used from days 7 through 14.

Cytokine treatment and TER measurements. Dual voltage-current clamp and apical and basolateral Ag-AgCl and calomel electrodes interfaced with "chopstick" KCl-agar bridges were used to assess TER in confluent monolayers grown on collagen-coated 0.33-cm2 inserts as previously described (30, 43). TER measurements have been used as a measure of paracellular permeability and barrier function in confluent T84 monolayers (20, 43). Baseline levels of TER in confluent T84 monolayers generally exceed 1,000 Omega  · cm2.

Confluent T84 monolayers were equilibrated in HEPES-phosphate-buffered Ringer solution [HPBR; containing (in mM) 135 NaCl, 5 KCl, 3.33 NaHPO4, 1 CaCl2, 1 MgCl2, 10 glucose, and 5 HEPES at pH 7.4] for 30 min before further treatment. TNF was applied to the basolateral compartment of confluent T84 monolayers at varying concentrations (10-1,000 U/ml) over a 4-h time course. Bryostatin-1 (100 nM) was applied to the basolateral compartment before and simultaneous with TNF treatment, and TER was measured over various time periods. Selective PKC inhibitors (Gö-6850, Gö-6976, and röttlerin) were added 30 min before treatment with either TNF or bryostatin-1.

Surface biotinylation. Confluent T84 monolayers grown on 4.7-cm2 Transwell inserts were treated with bryostatin-1 for up to 1 h, followed by three washes with ice-cold PBS containing 0.1 mM CaCl2 and 1.0 mM MgCl2, then followed by two 20-min incubation periods with buffer containing N-hydroxysuccinimidyl (NHS)-biotin added to both apical and basolateral surfaces. This was followed by two washes and a subsequent 20-min incubation with PBS-Ca2+-Mg2+ supplemented with 100 mM glycine to serve as the NHS-biotin quenching solution. Samples were washed again with PBS-Ca2+-Mg2+ twice, and monolayers were solubilized with 1 ml lysis buffer added for 1 h on ice. Cells were scraped from the filter, lysates were centrifuged at 14,000 g for 10 min at 4°C, and then lysates were filtered. Protein concentrations were normalized by using the Bradford method. Samples were incubated with 100 µl streptavidin beads overnight at 4°C with end-over-end rotation. The beads were then washed three times with lysis buffer, followed by two washes with high-salt buffer and one wash with no-salt buffer. Proteins were eluted by the addition of 5× Laemmli sample buffer containing DTT. Supernatants were subjected to SDS-PAGE (8% gel) followed by blotting with monoclonal antibody to TNF-R1. Samples were also taken before the addition of streptavidin beads for parallel Western blotting experiments.

125I-labeled TNF binding. Confluent T84 monolayers grown on 1.13-cm2 Transwell inserts were equilibrated for 30 min in HPBR followed by treatment with bryostatin-1 (100 nM) for 60 min. Monolayers were washed three times with ice-cold PBS and subsequently incubated at 4°C in HPBR containing 125I-TNF (10 ng/ml) in the basolateral compartment in the presence or absence of 200-fold excess cold TNF. After a 1-h incubation, filters were cut out and placed in vials containing ScintiSafe 30% Advanced Safety LSC-Cocktail scintillation fluid (Fischer Scientific, Pittsburgh, PA), and samples were measured on a scintillation counter.

In vitro kinase assay. As previously described (58), T84 monolayers grown to confluence on 4.7-cm2 Transwell inserts were treated with TNF (250 U/ml) for 30 min followed by two washes in ice-cold PBS. Protein extraction occurred by a 30-min incubation with lysis buffer containing (in mM) 50 Tris · HCl, pH 7.5, 140 EGTA, 30 sodium pyrophosphate, and 50 NaF with 100 µM Na3VO4 and complete protease inhibitor cocktail tablets. Samples were normalized to a concentration of 1.25 mg/ml and were incubated overnight at 4°C with polyclonal antibodies to PKC-alpha (2 µg), PKC-delta (2 µg), and PKC-epsilon (4 µg). Immune complexes were precipitated by using protein A-agarose beads (2 h incubation) and were washed three times and resuspended in 20 µl kinase buffer containing 35 mM Tris · HCl, pH 7.5, 10 mM MgCl2, 0.5 mM EGTA, 10 µCi [gamma -32P]ATP, 60 µM cold ATP, and 1 mM Na3VO4 in the presence of 10 µg myelin basic protein at 30°C for 30 min. Sample buffer (5× Laemmli) was added to terminate the reaction, and samples were boiled for 5 min. Supernatants were subjected to SDS-PAGE (12% gels), and gels were dried and analyzed by autoradiography.

Immunoprecipitation. Confluent T84 monolayers on 4.7-cm2 Transwell inserts were treated with bryostatin-1 (100 nM) for up to 30 min followed by two washes in ice-cold PBS. Protein extraction occurred by a 30-min incubation with lysis buffer containing (in mM) 50 Tris · HCl, pH 7.5, 140 EGTA, 30 sodium pyrophosphate, and 50 NaF, with 100 µM Na3VO4 and complete protease inhibitor cocktail tablets. Samples were normalized to a concentration of 1.25 mg/ml and were incubated overnight at 4°C in the presence of monoclonal antibody to TNF-R1. Immune complexes were precipitated by using protein A-agarose beads (2-h incubation) and were washed three times, then Laemmli sample buffer with 10% beta -mercaptoethanol was added and samples were boiled for 5 min. Supernatants were subjected to SDS-PAGE (8% gels), and gels were blotted with polyclonal antibody to PKC-alpha , -delta , and -epsilon .

[3H]mannitol flux. Confluent T84 monolayers grown on 4.7-cm2 Transwell inserts were incubated in HPBR for 30 min followed by treatment with basolaterally applied TNF (250 U/ml) in the presence of [3H]mannitol (5 µCi/ml) in the basolateral compartment. Apical solution was sampled every 30 min (to detect the presence of [3H]mannitol) and exchanged with fresh HPBR buffer solution for a total of 4 h. The collected apical samples were placed in vials containing 4 ml of ScintiSafe 30% Advanced Safety LSC-Cocktail scintillation fluid and analyzed for the presence of [3H]mannitol by scintillation counter.

Detection of soluble TNF-R1. Confluent T84 monolayers grown on 1.13-cm2 Transwell inserts were equilibrated for 30 min in HPBR buffer solution followed by treatment with bryostatin-1 (100 nM) for 1-4 h. Basolateral supernatant was sampled at regular time points, and amounts of soluble TNF-R1 present were evaluated with ELISA by using a human soluble TNF-R1 detection kit supplied by R&D Systems (Minneapolis, MN).

Gel electrophoresis and Western blotting. Samples were loaded at equal concentrations as determined by the Bradford assay after addition of Laemmli sample buffer containing 10% beta -mercaptoethanol and after 5 min of boiling. Proteins were separated by electrophoresis on 8-12% gels and transblotted on nitrocellulose membranes, followed by a 1-h incubation at room temperature in blocking buffer [containing 20 mM Tris (pH 7.5), 500 mM NaCl, 5% nonfat dry milk, and 0.2% Tween 20], a 1-h incubation with blocking buffer containing primary antibody, a 30-min rinse in wash buffer (20 mM Tris, pH 7.5, 500 mM NaCl, and 0.2% Tween 20), a 1-h incubation with blocking buffer containing secondary antibody, and another 30-min rinse in wash buffer. Bands were detected by using enhanced chemiluminescence detection reagents.

Cytotoxicity assay. Confluent T84 monolayers grown on 1.13-cm2 Transwell inserts were treated with TNF (1,000 U/ml) for 4 h, then washed twice with warm (37°C) PBS and incubated with PBS containing methylthiazoletetrazolium (MTT; thiazolyl blue, 5 mg/ml) for 20 min. This was followed by two washes with ice-cold PBS. Filters were cut out and placed into an Eppendorf tube containing 1 ml of 0.1 N HCl/isopropanol and sonicated for 30 s. Supernatant was removed, spun at 14,000 g for 10 min, and again removed and measured at 570 nM.

Materials. TNF was purchased from Sigma (St. Louis, MO). Transwell inserts were purchased from Corning Costar. Polyclonal antibodies to PKC-alpha , -delta , and -epsilon and the monoclonal TNF-R1 antibody were obtained from Santa Cruz Biotechnologies (Santa Cruz, CA). Bryostatin-1 was obtained from Biomol (Plymouth Meeting, PA). Tissue culture reagents and agarose beads were purchased from Life Technologies (Gaithersburg, MD). Gel electrophoresis and gel blotting reagents were purchased from Bio-Rad (Hercules, CA). Enhanced chemiluminescence detection reagents were purchased from Amersham (Piscataway, NJ). Complete protease inhibitor tablets were purchased from Boehringer Mannheim (Indianapolis, IN). The PKC inhibitors Gö-6850, Gö-6976, and röttlerin were purchased from Calbiochem (San Diego, CA). [gamma -32P]ATP (specific activity = 3,000 Ci/mmol), [3H]mannitol (specific activity = 17 Ci/mmol), and 125I-TNF (specific activity = 780 Ci/mmol) were purchased from PerkinElmer Life Sciences (Boston, MA). A Quantikine sTNF R1 immunoassay detection kit was purchased from R&D Systems.

Statistical analysis. Data are expressed as means ± SE. Statistical analysis was performed by Student's t-test and by one-way ANOVA with the Bonferroni-Dunn post hoc test for comparison with control, with P < 0.05 considered statistically significant. For all means, n >=  3.


    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

TNF causes a dose- and time-dependent fall in TER in T84 cells. TNF was applied to the basolateral membrane of confluent T84 monolayers at varying concentrations (10-1,000 U/ml), and TER was measured (Fig. 1A). There is a decrease in TER after 4 h with increasing concentrations of TNF (10 units, 96 ± 1% control; 100 units, 81 ± 1% control; 500 units, 43 ± 3% control; 1,000 units, 37 ± 3% control). Basal short-circuit current is unchanged with TNF treatment. TNF also leads to a time-dependent fall in TER (Fig. 1B). With TNF treatment (1,000 U/ml), there is an initial rise in TER after 1 h (113 ± 2% control) followed by a steady decline that was maximal at 4 h (37 ± 3% control). With lower concentrations of TNF, the fall in TER is slightly delayed and less pronounced (at 2 h, 250 U/ml TNF = 77 ± 8% control vs. 1,000 U/ml TNF = 65 ± 3% control; at 4 h, 250 U/ml TNF = 46 ± 5% control vs. 1,000 U/ml TNF = 37 ± 3% control). An MTT assay was used to assess cell viability at 4 h, and there was no difference between untreated and TNF-treated cells at 1,000 U/ml TNF, suggesting that direct cytotoxicity did not account for the fall in TER.


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Fig. 1.   Effect of tumor necrosis factor (TNF) on transepithelial electrical resistance (TER) and epithelial permeability. A: confluent T84 monolayers were treated with increasing concentrations of TNF (10-1,000 U/ml) applied to the basolateral compartment. TER was measured after 4 h. Data are mean percentages of control ± SE; n>= 3 in triplicate. After 4 h, control TER ~ 500 Omega  · cm2. *Statistical significance (P < 0.05) in TER compared with control. B: confluent T84 monolayers were treated with TNF (1,000 U/ml), and TER was measured over the time course indicated. Data are mean percentages of control ± SE; n >=  3 in triplicate. *Statistical significance (P < 0.05) in TER compared with control. C: cumulative [3H]mannitol flux following TNF treatment. TNF (250 U/ml) was applied to T84 cells in the presence of 5 µCi/ml [3H]mannitol in the basolateral compartment. Apical buffer was sampled every 30 min over a 4-h time period. * Statistically significant difference in flux rates between untreated and TNF-treated monolayers at given time point. Data are means ± SE; n = 6.

The TNF-induced fall in TER was accompanied by a time-dependent increase in the transepithelial flux of [3H]mannitol (mol wt = 182) from the basolateral to the apical compartment (Fig. 1C). The difference in flux rates between untreated and TNF-treated monolayers increased over time ([3H]mannitol flux rate difference: at 1h, 0.015 ± 0.002 pmol · h-1 · cm2; at 4 h, 214 ± 0.019 pmol · h-1 · cm2). The divergence of flux rates between untreated and TNF-treated monolayers reached significance after 3 h (At 3 h, untreated = 0.336 ± 0.014 vs. TNF = 0.413 ± 0.016 pmol · h-1 · cm2, P = 0.0059; at 4 h, untreated = 0.498 ± 0.008 vs. TNF = 0.713 ± 0.027 pmol · h-1 · cm2, P < 0.0001).

TNF-induced barrier dysfunction did not appear to be related to gross disruption of monolayer integrity. For example, TNF treatment (1,000 U/ml for 4 h) did not alter the subcellular localization of several tight junction proteins [claudin-1, -2, -3, and -5; occludin; zonula occludens (ZO)-1 and -2] as defined biochemically by their partitioning into Triton X-100-soluble and -insoluble fractions (35), suggesting that TNF-induced barrier dysfunction involved more subtle signaling-based alterations of junctional regulation rather than a wholesale remodeling of tight junction structure.

Evidence that the TNF-induced fall in TER is not mediated by PKC. The effect of TNF (250 U/ml) on PKC isozyme activity was assessed by in vitro kinase assay. After 30 min, TNF led to an increase in PKC-alpha activity (178 ± 25% control, P = 0.0364) and a decrease in the activity of PKC-epsilon (59 ± 10% control, P = 0.0160) and PKC-delta (51 ± 14% control, P = 0.0259) (Fig. 2A).


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Fig. 2.   TNF affects protein kinase C (PKC) isoform activity in T84 cells. A: T84 cells were treated with TNF (250 U/ml) for 30 min, and PKC isoform activity was assessed by in vitro kinase assay. After 30 min, PKC-alpha activity increased, whereas PKC-delta and -epsilon demonstrate decreased activity. Data analyzed by densitometry are expressed as mean percentages of control ± SE; n = 3. *P < 0.05. B: confluent T84 monolayers were treated with TNF (250 U/ml) with or without a 30-min pretreatment with various PKC inhibitors. Both Gö-6850 and Gö-6976 had no effect on the TNF-associated fall in TER. Data are mean percentages of control ± SE; n = 3 in triplicate. After 4 h, control TER ~500 Omega  · cm2 and TNF-treated (250 U/ml) TER ~200 Omega  · cm2. *P < 0.05.

To assess whether the TNF-induced fall in TER was mediated by the activity of these PKC isozymes, T84 cells were pretreated with the PKC inhibitors Gö-6850 (5 µM) and Gö-6976 (5 µM) before TNF (250 U/ml). Gö-6850 (5 µM) is an inhibitor of conventional (alpha ) and novel (delta , epsilon ) PKC isozymes, whereas Gö-6976 (5 µM) inhibits only the conventional (alpha ) PKC isozymes. Both G ö-6850 and Gö-6976 failed to block the fall in TER after treatment with TNF (Fig. 2B), suggesting that the mechanism behind this fall in TER is not mediated by activation of PKC-alpha . Although we cannot exclude the possibility that deactivation of PKC-delta and/or PKC-epsilon contributes to the fall in TER, the failure of either of these PKC inhibitors to modulate the TNF effect makes this possibility unlikely.

Bryostatin-1 pretreatment prevents the TNF-induced fall in TER. Treatment of T84 monolayers with bryostatin-1 (100 nM) for at least 30 min before TNF exposure (250 U/ml) prevented the TNF-induced fall in TER (at 2 h, TNF alone = 77 ± 8 vs. TNF + bryostatin-1 = 94 ± 2% control; at 3 h, TNF alone = 46 ± 7 vs. TNF + bryostatin-1 = 83 ± 1% control, P = 0.0103; at 4 h, TNF alone = 46 ± 5 vs. TNF + bryostatin-1 = 81 ± 1% control, P = 0.0020) (Fig. 3). Bryostatin-1 alone increases TER in T84 cells (117 ± 4% control after 4 h, n = 3 in triplicate, P < 0.05) through a mechanism that may be related to the regulated assembly of tight junction proteins (51). To control for the independent effects of bryostatin-1 on TER, the TER measurements of the TNF and bryostatin-1-treated monolayers were normalized to monolayers treated with bryostatin-1 alone. Subsequent data are normalized to a bryostatin-1 control to account for the small but significant independent effects of this agent on TER. Bryostatin-1 did not attenuate the TNF effect on TER if administered simultaneously with (Fig. 4) or subsequent to (not shown) TNF.


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Fig. 3.   Treatment of T84 cells with bryostatin-1 before TNF exposure attenuates the TNF-associated fall in TER. T84 cells were treated with bryostatin-1 (100 nM) for 30 min, followed by treatment with TNF (250 U/ml), and TER was measured over 4 h. T84 monolayers pretreated with bryostatin-1 maintained TER compared with monolayers exposed to TNF alone, with statistical significance occurring at 3 and 4 h. Data are mean percentages of control ± SE; n >=  3 in triplicate. *P < 0.05 compared with control.



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Fig. 4.   Pretreatment with bryostatin-1 is necessary to block the TNF-induced fall in TER. T84 monolayers pretreated with bryostatin-1 (100 nM) for 30 min maintained TER after exposure to TNF (250 U/ml). When bryostatin-1 and TNF were given simultaneously, the TNF-induced fall in TER was not prevented. Data are mean percentages of control ± SE; n>= 3 in triplicate. After 4 h, control TER ~500 Omega  · cm2, TNF-treated (250 U/ml) TER ~200 Omega  · cm2, and TNF- and bryostatin-1-pretreated TER ~450 Omega  · cm2. *P < 0.05 comparing pretreatment monolayers with TNF alone.

The effect of bryostatin-1 is mediated by a novel PKC isozyme. We have previously shown by both subcellular fractionation and Western blot as well as by in vitro kinase assay that bryostatin-1 induces rapid and sustained activation of the novel PKC-delta and -epsilon isozymes in T84 cells (43). In contrast, activation of the conventional PKC-alpha isozyme occurs only after 2 h, followed by downregulation (43). Thus the most likely candidate isozymes to mediate the effects of bryostatin-1 on the TNF response are novel PKC-delta and -epsilon . To confirm this, T84 monolayers were pretreated for 30 min with two different PKC inhibitors before treatment with bryostatin-1 and TNF (250 U/ml). Gö-6850 (5 µM) inhibits conventional (alpha ) and novel (delta , epsilon ) PKC isozymes, whereas Gö-6976 (5 µM) inhibits only the conventional (alpha ) PKC isozymes. Treatment with Gö-6850 blocked the bryostatin-1 effect (at 4 h, TNF + bryostatin-1 = 81 ± 1, TNF + bryostatin-1 + Gö-6850 = 52 ± 3, and TNF alone = 46 ± 5% control), whereas Gö-6976 did not (TNF + bryostatin-1 + Gö-6976 = 83 ± 2% control) (Fig. 5). This suggests that a novel PKC isozyme (delta  or epsilon ) is responsible for the effects of bryostatin-1 on preventing the TNF-induced fall in TER. This data is also consistent with the finding that a 30-min pretreatment with bryostatin-1 is sufficient time to prevent the fall in TER by TNF, since only PKC-delta and -epsilon are activated by bryostatin-1 during this time period (43).


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Fig. 5.   The effect of bryostatin-1 is mediated by a novel PKC isozyme. Confluent T84 monolayers were treated with PKC inhibitors [either Gö-6850 (5 µM) or Gö-6976 (5 µM)] for 30 min followed by bryostatin-1 (100 nM) and TNF (250 U/ml). TER was measured after 4 h. Pretreatment with Gö-6850, which inhibits conventional (alpha ) and novel (delta , epsilon ) isozymes, blocked the effect of bryostatin-1, whereas Gö-6976, which inhibits only conventional PKC isozymes, had no effect. Data are mean percentages of control ± SE; n >=  3 in triplicate. *P < 0.05 compared with control.

Bryostatin-1 induces PKC-delta coprecipitation with TNF-R1. The association between TNF-R1 and PKC isozymes was assessed by treating T84 cells with bryostatin-1 (100 nM) for up to 30 min, followed by immunoprecipitation with TNF-R1 and subsequent Western blotting using antibodies to various PKC isozymes. After bryostatin-1 treatment, PKC-delta was found to coprecipitate with TNF-R1, whereas PKC-alpha and -epsilon did not (Fig. 6A). Coprecipitation of PKC-delta with TNF-R1 became statistically significant as early as 15 min after bryostatin-1 treatment and was sustained over 30 min (Fig. 6B).


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Fig. 6.   Bryostatin-1 treatment leads to selective coprecipitation of PKC-delta with TNF receptor type I (TNF-R1). A: T84 cells were treated with bryostatin-1 (100 nM) over 30 min. Cell lysates were immunoprecipitated (IP) by using TNF-R1 antibody, followed by Western blot (WB) with various PKC isozymes. Bryostatin-1 treatment led to the coprecipitation of PKC-delta with TNF-R1. PKC-alpha and -epsilon did not coprecipitate with TNF-R1 after bryostatin-1 treatment over this time course. B: densitometric analysis of PKC-delta coprecipitating with TNF-R1. There is an increase in the amount of PKC-delta that coprecipitates with TNF-R1 that is sustained for 30 min. Data are mean percentages of control densitometry units ± SE; n >=  3. *P < 0.05.

PKC-epsilon activation by bryostatin-1 decreases surface TNF-R1. Bryostatin-1 treatment led to a progressive decrease in the surface expression of TNF-R1 as assessed by surface biotinylation and Western blot (bryostatin-1 for 30 min = 61 ± 8 vs. for 60 min = 53 ± 27% control) (Fig. 7A). According to Western blot, there was no decrease in total TNF-R1 after bryostatin-1 treatment (Fig. 7B), suggesting that the decrease in surface expression of TNF-R1 was not due to a decrease in total TNF-R1 protein. This data is consistent with 125I-TNF studies that used radiolabeled TNF to assess cell surface binding. Bryostatin-1-treated monolayers had reduced 125I-TNF binding (at 1 h, 71 ± 6% control, P = 0.022) (Fig. 7C), which provides further evidence that there was a decrease in the number of surface TNF receptors. The discrepancy between the biotinylation data (53 ± 27% control) and the 125I-TNF data (71 ± 6% control) may be explained by nonspecific binding of 125I-TNF to the cell surface or by binding of 125I-TNF to the TNF-R2 receptor, whose surface expression we have not characterized and which may not be affected by bryostatin-1.


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Fig. 7.   Bryostatin-1 treatment decreases surface expression of TNF-R1. A: T84 cells were treated with bryostatin-1 (100 nM) for 30 and 60 min. This was followed by surface biotinylation and Western blot for TNF-R1. There was a time-dependent decrease in surface expression of TNF-R1 that was significant by 30 min. Data are mean percentages of control densitometry units ± SE; n = 3. *P < 0.05. B: Western blot analysis of TNF-R1 after 30 and 60 min of treatment with bryostatin-1 (100 nM). There was no difference in total TNF-R1 protein after treatment with bryostatin-1; n = 3. C: 125I-labeled TNF binding following bryostatin-1 treatment. T84 cells were treated with bryostatin-1 for 30 min followed by cold incubation for 1 h with 125I-labeled TNF (10 ng/ml) in the presence or absence of 200-fold excess cold TNF. Specific binding was assessed under these conditions. Data are mean percentages of control ± SE; n = 6. *P < 0.05.

PKC-epsilon activation by bryostatin-1 induces TNF receptor (TNF-R1) shedding. T84 cells were treated with bryostatin-1, and T84 cell culture supernatant was collected from the basolateral compartment after 1-4 h and analyzed by ELISA. Bryostatin-1 increased the amount of soluble TNF-R1 in the basolateral supernatant in a time-dependent fashion (Fig. 8).


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Fig. 8.   Bryostatin-1 treatment leads to a time-dependent increase in TNF-R1 shedding. T84 cells were treated with bryostatin-1 (100 nM) for 1-4 h. Basolateral supernatant was collected and analyzed by ELISA for the presence of soluble TNF-R1. Bryostatin-1 treatment increased the amount of soluble TNF-R1 present in the basolateral supernatant, with significant differences occurring as early as 1 h after treatment (in ng/ml: at 1 h, control = 0.00365 ± 0.001 vs. bryostatin-1 = 0.01006 ± 0.001, P = 0.0483; at 2 h, control = 0.0113 ± 0.001 vs. bryostatin-1 = 0.037 ± 0.007, P < 0.0001; at 3 h, control = 0.013 ± 0.0027 vs. bryostatin-1 = 0.0546 ± 0.005, P = 0.0023; at 4 h, control = 0.0143 ± 0.005 vs. bryostatin-1 = 0.079 ± 0.009, P = 0.0035) *P < 0.05 vs. control monolayers at the same time point. Data are means ± SE; n = 3.

Bryostatin-1-induced TNF-R1 shedding was assessed in the presence of various PKC inhibitors. As stated previously, Gö-6850 (5 µM) is an inhibitor of conventional (alpha ) and novel (delta , epsilon ) PKC isozymes, whereas Gö-6976 (5 µM) inhibits only the conventional PKC-alpha isozymes. Röttlerin (10 µM), which is a specific inhibitor of PKC-delta , was also used. Gö-6850 inhibited bryostatin-1-induced TNF-R1 shedding (at 3 h, bryostatin-1 alone = 0.055 ± 0.005 vs. Gö-6850 + bryostatin-1 = 0.014 ± 0.001 ng/ml), whereas Gö-6976 had no effect (Gö-6976 + bryostatin-1 = 0.069 ± 0.003 ng/ml) (Fig. 9B). Röttlerin also failed to block bryostatin-1-induced TNF-R1 shedding (at 2 h, bryostatin-1 alone = 0.035 ± 0.006 vs. röttlerin + bryostatin-1 = 0.032 ± 0.005 ng/ml) (Fig. 9A). Together, the data suggest that bryostatin-1-induced shedding of the TNF receptor (TNF-R1) is mediated through the specific activation of PKC-epsilon .


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Fig. 9.   The bryostatin-1-induced increase in TNF-R1 shedding is mediated by PKC-epsilon A: T84 cells were treated with bryostatin-1 for 2 h in the presence or absence of various PKC inhibitors. Pretreatment with Gö-6850 (5 µM), which inhibits conventional (alpha ) and novel (delta , epsilon ) isozymes, inhibited bryostatin-1-induced TNF-R1 shedding, whereas röttlerin (10 µM), which specifically inhibits PKC-delta , had no effect. *P < 0.05. B: T84 cells were treated with bryostatin-1 for 3 h in the presence of Gö-6850 and Gö-6976, which inhibits conventional (alpha ) PKC isozymes. Although Gö-6850 blocked TNF-R1 shedding, Gö-6976 had no effect. *P < 0.05.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The proinflammatory cytokine TNF increases intestinal mucosal permeability in vivo and in vitro. We used the T84 human intestinal cell line as a reductionist model to study the effects of TNF on epithelial permeability. TNF has been shown in various cell lines and natural tissue models to increase epithelial permeability, although the mechanism underlying this effect is not yet established. In pulmonary endothelial cells, TNF-induced barrier dysfunction appears to be mediated by PKC-alpha (15). Another study implicated the p38-MAPK pathway in TNF-induced endothelial permeability (14). In the human intestinal cell line HT-29/B6, TNF increased epithelial permeability but could be partially inhibited by the tyrosine kinase inhibitor genistein and the protein kinase A inhibitor H-8 (42).

Several signaling pathways are known to regulate epithelial permeability, and TNF signaling has been shown in a number of instances to interact with these pathways. These include p44/p42 ERKs (24), p38 MAPK (14), PLC (21), PKC (8, 15, 38), myosin light-chain kinase (MLCK) (47), phosphoinositol 3-kinase (PI3-kinase) (7), and tyrosine kinase (25, 42). It is unknown whether activation of these pathways in the T84 model contribute to the TNF-induced fall in TER; however, in our hands (unpublished data), neither the MEK inhibitor PD-98059 (50 µM), the p38 MAPK inhibitor SB-203580 (25 µM), the PLC inhibitor D-609 (10 µM), the MLCK inhibitor ML-7 (20 µM), the PI3-kinase inhibitor wortmannin (100 nM), nor the general tyrosine kinase inhibitor genistein (50 µM) attenuated the TNF-associated fall in TER. Although TNF induced minor changes in the activation state of various PKC isozymes, neither the general PKC inhibitor Gö-6850 (5 µM) nor the conventional PKC inhibitor Gö-6976 (5 µM) attenuated the fall in TER, making it unlikely that PKC directly mediates the effects of TNF on junctional permeability in the T84 model.

The mechanism of the TNF-induced increase in epithelial permeability did not appear to involve gross disruption of junctional protein integrity, based on biochemically-defined subcellular localization and on immunohistochemical analysis (Fig. 10). Specifically, TNF did not appear to alter the distribution of several key tight junction proteins (claudin-1, -2, -3, and -5; occludin; ZO-1 and -2) between Triton X-100-soluble and -insoluble fractions. Although TNF does induce apoptosis in the T84 line (unpublished data), this effect lags many hours behind the initial fall in TER and is unlikely to contribute significantly to the observed changes over the experimental time course examined.


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Fig. 10.   Immunohistochemical staining of T84 cells with zonula occludens (ZO)-1. T84 cells were treated with TNF (250 U/ml) for 4 h, followed by immunostaining with antibody to ZO-1. There was no difference in ZO-1 staining after TNF treatment. Staining of F-actin served as a control.

In the present study, we have demonstrated that bryostatin-1 attenuates TNF-associated barrier dysfunction, likely through activation of a novel PKC isozyme. The observation that pretreatment with bryostatin-1 appears to be required in this regard suggests that the underlying mechanism involves the regulation of an early signaling event, perhaps occurring at the level of the TNF receptor and the initiation of subsequent signaling events. Both the novel PKC-delta and -epsilon isozymes appeared to interact with TNF-R1, albeit in strikingly different ways (Fig. 11). It is not yet certain whether all of these effects are required to attenuate TNF-induced barrier dysfunction.


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Fig. 11.   Diagram depicting the early and late effects of bryostatin-1 on PKC isozymes and the TNF-R1. Bryostatin-1 activates the novel PKC isozymes PKC-delta and PKC-epsilon , which have early and late effects on the TNF-R1. PKC-delta is activated within 15 min and coassociates with TNF-R1 (1). PKC-epsilon is activated within 30 min and leads to TNF-R1 receptor internalization (2). PKC-epsilon activation also leads to TNF-R1 shedding, an effect that is most significant after 3 h (3).

We have shown that on bryostatin-1 treatment PKC-delta coprecipitates with TNF-R1 within 30 min (depicted schematically in Fig. 11, 1). TNF-R1 is a known substrate of PKC-delta , although this has not been previously demonstrated in an epithelial cell line. In neutrophils, TNF treatment leads to coprecipitation of PKC-delta with TNF-R1 and is associated with serine phosphorylation of the receptor and subsequent receptor desensitization (23).

Two major effects of PKC-epsilon on TNF-R1 were observed (depicted schematically in Fig. 11). The initial decrease in surface TNF-R1, as evidenced by both surface biotinylation studies and 125I-TNF binding studies, is likely due to internalization of surface receptors via endocytosis (Fig. 11, 2). Activation of PKC-epsilon has been shown to increase the rate of fluid-phase endocytosis at the basolateral aspect of T84 cells (44), and it is clear that endocytosis alters the surface expression of a number of ion transporters and is involved in cell surface receptor recycling (3, 13, 26). PKC activation has been shown in a number of cell lines to downregulate TNF receptor function either by decreasing receptor number or influencing receptor affinity (1, 4, 22, 41, 48).

PKC-epsilon activation also leads to a time-dependent increase in TNF-R1 shedding (Fig. 11, 3). The concept of TNF receptor shedding has been studied most extensively in neutrophils (1, 4, 37, 41, 48), although there has been at least one report of TNF receptor shedding in a colonic cell line (29). We confirm that TNF-R1 shedding does occur in T84 intestinal epithelial cells in response to bryostatin-1, as well as to other known activators of shedding such as TNF and PMA (data not shown). Similar to its effects in neutrophils, receptor shedding may be another important mechanism of downregulation of TNF responses in epithelia. Soluble TNF-R1 proteolytically released from the basolateral membrane is known to bind to exogenous TNF, thereby affecting TNF bioavailability (5, 36). TNF-R1 shedding may also decrease the number of TNF-R1 available on the cell surface, although the effect of PKC-epsilon on receptor synthesis or trafficking of intracellular stores in T84 epithelia is not known. Although PKC is known to be involved in TNF receptor shedding in multiple cell lines (1, 4, 41, 48), a specific isozyme has not been previously implicated. Our data suggest that PKC-epsilon may be the key isozyme involved in TNF-R1 shedding in T84 epithelia. Whether PKC-epsilon also affects receptor affinity in this cell line has not yet been examined.

Inhibition of TNF signaling at the receptor level has been used in the treatment of inflammatory gastrointestinal conditions associated with elevated levels of TNF. For example, monoclonal TNF antibody is used in the treatment of patients with refractory Crohn's disease (49). Soluble TNF receptor is currently in use in clinical trials for the treatment of Crohn's disease and rheumatoid arthritis (40). TNF receptor surface regulation, through either internalization or shedding of endogenous soluble receptor, appears to be important in mitigating the pathological actions of TNF on model T84 intestinal epithelia. The development of isozyme-selective PKC activators, which are currently being developed for the treatment of ischemic heart disease (12), may be an alternative approach to the treatment of inflammatory conditions characterized by elevated TNF.


    ACKNOWLEDGEMENTS

This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grants DK-48010, DK-51630, and T32 DK-007754 (to J. Yoo).


    FOOTNOTES

Address for reprint requests and other correspondence: J. B. Matthews, Univ. of Cincinnati Medical Center, P.O. Box 670558, 231 Albert Sabin Way, Cincinnati, OH 45267-0558 (E-mail: jeffrey.matthews{at}uc.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

First published December 27, 2002;10.1152/ajpgi.00214.2002

Received 6 June 2002; accepted in final form 16 December 2002.


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