Using intestinal Caco-2 cells, we previously showed that assembly of cytoskeleton is required for monolayer barrier function, but the underlying mechanisms remain poorly understood. Because the θ-isoform of PKC is present in wild-type (WT) intestinal cells, we hypothesized that PKC-θ is crucial for changes in cytoskeletal and barrier dynamics. We have created the first multiple sets of gastrointestinal cell clones transfected with varying levels of cDNA to stably inhibit native PKC-θ (antisense, AS; dominant negative, DN) or to express its activity (sense). We studied transfected and WT Caco-2 cells. First, relative to WT cells, AS clones underexpressing PKC-θ showed monolayer injury as indicated by decreased native PKC-θ activity, reduced tubulin phosphorylation, increased tubulin disassembly (decreased polymerized and increased monomeric pools), reduced architectural integrity of microtubules, reduced stability of occludin, and increased barrier hyperpermeability. In these AS clones, PKC-θ was substantially reduced in the particulate fractions, indicating its inactivation. In WT cells, 82-kDa PKC-θ was constitutively active and coassociated with 50-kDa tubulin, forming an endogenous PKC-θ/tubulin complex. Second, DN transfection to inhibit the endogenous PKC-θ led to similar destabilizing effects on monolayers, including cytoskeletal hypophosphorylation, depolymerization, and instability as well as barrier disruption. Third, stable overexpression of PKC-θ led to a mostly cytosolic distribution of θ-isoform (<10% in particulate fractions), indicating its inactivation. In these sense clones, we also found disruption of occludin and microtubule assembly and increased barrier dysfunction. In conclusion, 1) PKC-θ isoform is required for changes in the cytoskeletal assembly and barrier permeability in intestinal monolayers, and 2) the molecular event underlying this novel biological effect of PKC-θ involves changes in phosphorylation and/or assembly of the subunit components of the cytoskeleton. The ability to alter the cytoskeletal and barrier dynamics is a unique function not previously attributed to PKC-θ.
- epithelial barrier permeability
- protein kinase C isoform
the epithelial cells of the intestinal mucosa form a selective permeability barrier that separates the hostile luminal environment from the internal milieu. Barrier permeability permits the absorption from the lumen of needed nutrients, water, and electrolytes but prevents the passage of external proinflammatory and immunoreactive molecules (e.g., bacterial derivatives) into the mucosa and the systemic circulation. Indeed, loss of barrier permeability (hyperpermeability) causes the penetration of bacterial endotoxin and other harmful luminal antigens into the mucosa and promotes the initiation or continuation of inflammatory processes and injury (3, 23–25, 31). Not surprisingly, disruption of barrier permeability has been implicated in the pathogenesis of a wide variety of gastrointestinal (GI) and systemic disorders (24, 25, 28, 30–32, 42, 46). For example, intestinal hyperpermeability (“leaky gut”) has often been reported and implicated in the pathogenesis of inflammatory bowel disease (IBD) (see, e.g., Refs. 23–25, 28, 42, 46).
An important advance in our understanding of GI inflammation, especially IBD, has been the realization that a poorly maintained barrier permeability, the so-called leaky gut, can lead to intestinal inflammation and tissue injury. In animal models, loss of gut barrier permeability caused by the injection of bacterial products into the mucosa elicits IBD-like conditions (48). Similarly, transgenic animals with a hyperpermeable gut barrier exhibit symptoms of intestinal inflammation (23). One of the underlying difficulties in managing IBD disorders is due in large part to our limited understanding of the molecular processes involved in alterations of gut barrier.
The stability of intestinal epithelial barrier depends on a complex array of cytoskeletal protein filaments that includes occludin and microtubules (2–4, 7, 9, 11–13, 29, 47). Using monolayers of intestinal Caco-2 cells, we reported (2–4, 7–9, 11) that microtubule cytoskeletal assembly and stability are required for epithelial barrier integrity. Not surprisingly, microtubules play a central role in maintaining cellular integrity, structure, and shape (3, 15, 33, 34, 44). As such, microtubules govern cell membrane morphology and polarity, functions essential to the maintenance of epithelial monolayer (barrier) integrity (3, 12, 15, 19). Protein kinase C (PKC), in general, is known to affect epithelial barrier, but the roles of specific PKC isoforms in cytoskeletal and barrier changes remain largely unknown.
PKC consists of a family of serine- and threonine-specific kinases. This family, which includes at least 12 known PKC isoenzymes, can be classified into three subfamilies on the basis of differences in sequence homology and cofactor requirement (5–8, 10, 14, 18, 26, 35, 43). The conventional (or classical) PKC isoforms (α, β1, β2, γ) require calcium and phospholipid for their activation, whereas the novel PKC isoforms (δ, ε, θ, η, μ) are calcium independent but require phospholipid. Activation of the third group, atypical PKC isoforms (λ, τ, ζ), is independent of calcium and phorbol compounds (10). Intestinal cells (e.g., Caco-2) express at least 10 isoforms of PKC, including PKC-α, PKC-β1, PKC-β2, PKC-δ, PKC-ε, PKC-θ, PKC-η, PKC-ζ, PKC-λ, and PKC-τ (5–8, 10, 14, 35, 36). These isoenzymes are different in their intracellular distribution, expression, substrate type, and activation. It is therefore thought that each PKC isoform can perform unique biological functions (5, 6, 10, 37, 39, 41, 43). In our previous studies, we investigated both the damaging and protective pathways for gut barrier permeability. We showed that growth factor [epidermal growth factor (EGF) or transforming growth factor-α (TGF-α)] prevents oxidant-induced barrier disruption through the activation of the classical PKC-β1 and the atypical PKC-ζ isoforms, leading to the protection of the cytoskeleton (e.g., 7, 10, 14). Nonetheless, the role of specific PKC isoforms in alterations of intestinal cytoskeletal assembly and barrier permeability still remains largely unexplored.
In the current investigation, we have explored the role of the θ-isoform of PKC because this member of the novel subfamily of PKC is both present and abundant in our wild-type (WT; naive) Caco-2 cells and therefore could be a possible contributor to epithelial monolayer barrier integrity and cytoskeletal assembly. To this end, we studied the PKC-θ using targeted molecular interventions, transfecting cells to create the first multiple clones of intestinal cells for this isoform: in several clones, the endogenous (82 kDa) PKC-θ isoform was reliably inhibited by either antisense underexpression or dominant negative inactivation; in the other clones, native PKC-θ was ectopically increased by sense expression. Using these targeted approaches, we sought to determine whether PKC-θ isoform activity is required for changes in the dynamics of the cytoskeleton (e.g., microtubules and occludin) and barrier integrity in intestinal epithelial monolayers. In this article, we describe novel mechanisms dependent on the θ-isoform of PKC, namely, alterations of the cytoskeletal assembly and permeability clearance in cell monolayers.
MATERIALS AND METHODS
Caco-2 cells were obtained from American Type Culture Collection (Rockville, MD) at passage 15. This colonic cell line was chosen for our studies because the cells form monolayers that morphologically resemble intestinal cells, with defined apical brush borders and a highly organized microtubule network upon differentiation (12, 19, 38). Cells were maintained at 37°C in complete Dulbecco's modified Eagle's medium (DMEM) in an atmosphere of 5% CO2 and 100% relative humidity. WT cells or stably transfected cells (see Plasmids and transfection) were split at a ratio of 1:6 upon reaching confluency and were set up in either 6- or 24-well plates for experiments or in T-75 flasks for propagation. The utility and characterization of this cell line have been previously reported (2–14, 38).
Plasmids and transfection.
The sense, antisense, and dominant negative plasmids of PKC-θ were constructed as we previously described (5, 7). A unique tetracycline-responsive expression (TRE) system was used to overexpress the native PKC-θ. cDNA encoding the entire reading frame of PKC-θ plasmid was subcloned into the TRE vector, creating TRE-PKC-θ. The antisense or dominant negative PKC-θ plasmid was also subcloned to create AS-PKC-θ or DN-PKC-θ, respectively.
Cultures of Caco-2 cells grown to 50–60% confluence were cotransfected with hygromycin resistance plasmid (pβ-hygro) and with expression plasmids encoding sense PKC-θ, antisense PKC-θ, or dominant negative PKC-θ by using Lipofectin reagent (GIBCO BRL) as we described (5, 7). Control conditions included vector alone. Briefly, cells were incubated for 16 h at 37°C with the plasmid DNA in serum-free medium in the presence of Lipofectamine (25 μl per 25-cm2 flask). Subsequently, the DNA-containing solution was removed and replaced with fresh medium containing 10% FBS to relieve cells from the shock of exposure to serum-free medium. After transfection, cells were subjected to hygromycin selection (1 mg/ml). Resistant cells were maintained in DMEM-FBS and 0.2 mg/ml hygromycin (selection medium). For inducible expression of PKC-θ, Caco-2 cells were transfected with a plasmid expressing the tetracycline-responsive transactivator (tTA) along with a second plasmid conferring resistance to G-418. After selection in 0.6 mg/ml G-418 (selection medium), one such clone (i.e., parental tTA) was then itself transfected with the TRE-PKC-θ system. pβ-Hygro was included to confer resistance to hygromycin (selection marker, 1 mg/ml). Control conditions included vector alone (TRE-z). Multiple clones expressing PKC-θ or lacking PKC-θ activity were assessed by immunoblotting and activity assay and then used for experiments.
In the first series of experiments, postconfluent monolayers of WT (naive) cells were incubated with vehicle (isotonic saline) for 30 min and then assessed for baseline conditions. Experiments were then repeated by using multiple clones of stably transfected cells. In all experiments, we assessed microtubule cytoskeletal stability (cytoarchitecture, disassembly), tubulin assembly (polymerized and monomeric tubulin pools), tubulin phosphorylation, occludin stability (cytoarchitecture), PKC-θ subcellular distribution (membrane, cytoskeletal, and cytosolic fractions), PKC-θ activity (immunoprecipitation and in vitro kinase assay), and monolayer integrity/permeability (clearance of different-sized probes).
In the second series of experiments, transfected cell clones overexpressing PKC-θ were used. Multiple clones that were stably overexpressing PKC-θ (i.e., TRE-PKC-θ) were grown as monolayers and then exposed to vehicle. Outcomes were measured as described above. In these overexpression studies, cells were grown for 48 h in the absence of tetracycline before experiments were performed.
In the third series of experiments, monolayers of antisense-transfected cells lacking PKC-θ activity were treated with vehicle. In all experiments, PKC-θ activity was determined in immunoprecipitated samples (see Immunoprecipitation and PKC-θ activity assay). In a corollary series of experiments, we investigated the effects of PKC-θ dominant negative mutant on the state of monolayer integrity, tubulin assembly, and microtubule disassembly and cytoarchitecture. To this end, monomeric (S1) and polymerized (S2) fractions of tubulin (the structural protein subunit of microtubules) were isolated and then analyzed by immunoblotting (2, 3, 12). Microtubule integrity was assessed by 1) immunofluorescent labeling and fluorescence microscopy, to determine the percentage of cells with normal microtubules; 2) detailed analysis with the use of high-resolution laser scanning confocal microscopy; and 3) immunoblot analysis of monomeric and polymerized tubulin pools.
Fractionation and immunoblotting of PKC.
Cell monolayers grown in large 75-cm2 flasks were processed for isolation of the cytosolic, membrane, and cytoskeletal fractions (7, 14, 20). Protein content of these various fractions was assessed according to the Bradford method (16). Samples (5 μg protein/lane) were separated by PAGE. The immunoblotted proteins were visualized by enhanced chemiluminescence (ECL; Amersham, Arlington Heights, IL) and autoradiography (e.g., 1 h at −20°C). The exposure times were adjusted to ensure linear responses. Under these conditions, the chemiluminescence assay was linear between 1 and 10 μg of total protein. Standard (purified PKC-θ) loading controls (1 μg/lane) were also run concurrently. Blots were routinely stained with 0.1% india ink in Tris-buffered saline containing Tween 20 to further verify equal loading.
For isolation of the cell fractions, after treatments, postconfluent monolayers were scraped and ultrasonically homogenized (GE130; amplitude 50, 6 pulses/s, duration 20 s; GE Ultrasonic Processor) in Tris·HCl buffer (20 mM Tris·HCl, pH 7.5, 0.25 mM sucrose, 2 mM EDTA, 10 mM EGTA, 2 μg/ml aprotinin, 2 μg/ml pepstatin, 2 μg/ml leupeptin, and 2 μg/ml PMSF). The homogenates were then ultracentrifuged (100,000 g, 40 min, 4°C), and the supernatant was removed and used as the source of the cytosolic fraction. Next, pellets were washed with 0.2 ml of Tris·HCl buffer, resuspended in 0.8 ml of buffer containing 0.3% Triton X-100, and maintained on ice for 1 h. The samples were then centrifuged (100,000 g, 1 h, 4°C), and the supernatant was used as the source of the membrane fraction. To this remaining pellet, we added 0.3 ml of cold (4°C) lysis buffer (150 mM NaCl, 50 mM Tris·HCl, 1 mM EDTA, 1 mM EGTA, 1% NP-40, 0.1% sodium deoxycholate, 0.1% SDS, 2 μg/ml aprotinin, 2 μg/ml pepstatin, 2 μg/ml leupeptin, and 2 μg/ml PMSF). The samples were then placed on ice for 1 h and ultracentrifuged as described above. The remainder of the lysate or Triton-insoluble cytoskeletal fraction was then removed. For total extraction, which provides the fraction used to confirm total PKC-θ, scraped monolayers were placed directly into 1.5 ml of cold lysis buffer and subsequently ultracentrifuged as described above. The supernatant was used for bulk protein determination.
For immunoblotting, samples (5 μg protein/lane) were added to a standard SDS buffer, boiled, and then separated on 7.5% SDS-PAGE (7, 14). The immunoblotted proteins were incubated with a primary monoclonal antibody to PKC-θ [nPKC-θ (E7), no. sc-1680 (human reactive); Santa Cruz Biotechnology, Santa Cruz, CA] at 1:3,000 dilution. A horseradish peroxidase-conjugated antibody (Molecular Probes, Eugene, OR) was used as a secondary antibody at 1:4,000 dilution. Proteins were visualized by enhanced chemiluminescence and autoradiography and subsequently analyzed by densitometry. The identity of the PKC-θ band was assessed according to a procedure that we previously described (5, 7): 1) Use of a PKC-θ blocking peptide (no. sc-1680 P; Santa Cruz Biotechnology) in combination with the anti-PKC-θ antibody prevented the appearance of the corresponding “major” band in Western blots. 2) In addition, in the absence of the primary antibody to PKC-θ, no corresponding band for PKC-θ was observed. 3) The PKC-θ band ran at the expected molecular mass of 82 kDa, as confirmed by a known positive control for PKC-θ (from rat brain lysates). 4) Prestained molecular weight (Mr) markers (Mr 67,000 and 93,000) were run in adjacent lanes. In initial studies, using total PKC extracts, we confirmed that expression of PKC-θ or inhibition of PKC-θ did not affect the relative expression levels of other PKC isoforms.
Immunoprecipitation and PKC-θ activity assay.
Immunoprecipitated PKC-θ was collected and processed for its ability to phosphorylate a synthetic peptide (5, 10). Briefly, after treatments, confluent cell monolayers were lysed by incubation for 20 min in 500 μl of cold lysis buffer (20 mM Tris·HCl, pH 7.4, 150 mM NaCl, 10 μg/ml anti-protease cocktail, 10% glycerol, 1 mM sodium orthovanadate, 5 mM NaF, and 1% Triton X-100). The lysates were clarified by centrifugation at 14,000 g for 10 min at 4°C. For immunoprecipitation, the lysates were incubated for 90 min at 4°C with anti-PKC-θ (1:2,000 dilution, in excess). The extracts were then incubated with protein A/G-agarose for 1 h at 4°C. The immunocomplexes were collected by centrifugation (2,500 g, 5 min) in a microfuge tube and washed three times with immunoprecipitation buffer containing 5 mM Tris·HCl, pH 7.4, and 0.2% Triton X-100. They were then washed one time with kinase buffer (20 mM HEPES, pH 7.5, 10 mM MgCl2, 2 mM MnCl2, and 20 μM ATP), resuspended in 20 μl of kinase buffer and 5 μl of 5× reaction buffer (1 mg/ml histone H1 and 0.25 mg/ml l-α-phosphatidyl-l-serine) plus 5 μCi [γ-32P]ATP, and subsequently incubated for 5 min at 30°C. Reactions were then stopped by the addition of 8 μl of 5× sample buffer, and the samples were boiled at 95°C for 5 min before separation by 7.5% PAGE. The extent of histone H1 phosphorylation was determined by scintillation counting of excised Coomassie blue-stained histone polypeptide bands. Counts for blanks were subtracted from the sample activity. Sample activity was also corrected for protein concentration (Bradford method, Ref. 16), and PKC-θ activity was reported as picomoles per minute per milligram of protein.
Immunofluorescent staining and high-resolution laser scanning confocal microscopy of microtubule cytoskeleton and occludin.
Cells from monolayers were fixed in cytoskeletal stabilization buffer and then postfixed in 95% ethanol at −20°C as we described previously (2–4, 7–9, 11, 12). Cells were subsequently processed for incubation with a primary antibody (monoclonal mouse anti-β-tubulin antibody; Sigma, St. Louis, MO) and then with a secondary antibody (FITC-conjugated goat anti-mouse antibody; Sigma). For occludin staining, cells were processed for incubation with a primary monoclonal antibody (mouse anti-occludin antibody; Zymed, San Francisco, CA) and then with a secondary antibody (FITC-conjugated goat anti-mouse antibody; Zymed). After staining, cells were observed with an argon laser (λ = 488 nm) with a ×63 oil-immersion Plan-Apochromat objective (NA 1.4; Zeiss). The cytoskeletal elements were examined in a blinded fashion for their overall morphology, orientation, and disruption as we described previously (2–4, 7–9, 11, 12). The identity of the treatment groups for all slides was decoded only after examination was complete.
Microtubule (tubulin) fractionation and immunoblotting analysis of polymerized tubulin and monomeric tubulin (tubulin assembly/disassembly).
Polymerized (S2) and monomeric (S1) fractions of tubulin were isolated according to a method we described previously (3, 12). Cells were gently scraped and pelleted with centrifugation at low speed (700 rpm, 7 min, 4°C) and resuspended in microtubule stabilization-extraction buffer (0.1 M PIPES, pH 6.9, 30% glycerol, 5% DMSO, 1 mM MgSO4, 10 μg/ml anti-protease cocktail, 1 mM EGTA, and 1% Triton X-100) at room temperature for 20 min. Tubulin fractions were separated after a series of centrifugation and extraction steps. Specifically, cell lysates were centrifuged at 105,000 g for 45 min at 4°C, and the supernatant containing the soluble monomeric pool of tubulin (S1 fraction) was gently removed. The remaining pellet was then resuspended in 0.3 ml of Ca2+-containing depolymerization buffer (0.1 M PIPES, pH 6.9, 1 mM MgSO4, 10 μg/ml, and 10 mM CaCl2) and incubated on ice for 60 min. Subsequently, samples were centrifuged at 48,000 g for 15 min at 4°C, and the supernatant (S2 fraction or cold/Ca2+-soluble fraction) was removed. To ensure complete removal of the S2 fraction, we treated the remaining pellet with the Ca2+-containing depolymerization buffer twice more by resuspension and centrifugation. The “microtubules” were recovered by separate incubation (at 37°C for 30 min) of the S1 and S2 fractions with the stabilizing agents taxol (10 μM) and GTP (1 mM) in microtubule stabilization buffer (MSB: 0.1 M PIPES, pH 6.9, 30% glycerol, 5% DMSO, 10 μg/ml anti-protease cocktail, 1 mM EGTA, 1 mM MgCl2, and 1 mM GTP) to promote polymerization of tubulin. Tubulin was then recovered by centrifugation and resuspended in MSB. Fractionated S1 and S2 samples were then flash frozen in liquid N2 and stored at −70°C until immunoblotting. For immunoblotting, samples (5 μg protein/lane) were placed in a standard SDS sample buffer, boiled, and then subjected to PAGE on 7.5% gels. Standard (purified) tubulin controls (5 μg/lane) were run concurrently with each run. To quantify the relative levels of tubulin, we used a laser densitometer to measure the optical density of the bands corresponding to immunolabeled tubulin.
Immunoprecipitation and Western blot analysis of tubulin phosphorylation.
Immunoprecipitated tubulin was collected and then assessed for phosphorylation by PAGE (7). For immunoprecipitation, cell lysates were incubated for 4 h at 4°C with monoclonal anti-β-tubulin (1:50 dilution, in excess). The extracts were then incubated with protein G-Sepharose 4B (Zymed) for 2 h at 4°C. The immunocomplexes were collected by centrifugation (2,500 g, 5 min) in a microfuge tube and washed three times with immunoprecipitation buffer containing 5 mM Tris·HCl, pH 7.4, and 0.2% Triton X-100. The resultant pellets were resuspended in a standard SDS sample buffer and boiled at 95°C for 5 min before separation by PAGE. Gels were transferred to nitrocellulose membranes, blocked with 1% bovine serum albumin and 0.01% Tween 20 in phosphate-buffered saline for blotting by anti-phosphothreonine (1:3,000 dilution; Transduction Labs, Lexington, KY) and then, for detection of immune complexes by horseradish peroxidase-conjugated secondary antibody, incubated with chemiluminescent reagents (ECL) and autoradiographed.
Determination of monolayer barrier integrity by fluorometry.
Status of the monolayer barrier integrity was assessed by a widely used and validated technique that measures the apical-to-basolateral paracellular flux of fluorescent markers such as fluorescein sulfonic acid (FS; 200 μg/ml, 0.478 kDa) as we (2–5, 7–9) and others (27, 29, 45, 47) described. In select experiments, higher molecular mass fluorescein dextran (FD4; 1 mg/ml, 4 kDa) also was used. Briefly, fresh phenol-free DMEM (800 μl) was placed into the lower (basolateral) chamber, and phenol-free DMEM (300 μl) containing probe (FS or FD4) was placed in the upper (apical) chamber. Aliquots (50 μl) were obtained from the upper and lower chambers at time 0 and at subsequent time points and transferred into clear 96-well plates (clear bottom; Costar, Cambridge, MA). Fluorescent signals from samples were quantitated using a fluorescence multiplate reader (FL 600; Bio-Tek Instruments). The excitation and emission spectra for probes were as follows: excitation = 485 nm, emission = 530 nm. Clearance was calculated as Cl (nl·h−1·cm−2) = Fab/([FS or FD4]a × S), where Fab is the apical-to-basolateral flux of FS or FD4 (light units/h), [FS or FD4]a is the probe concentration at baseline (light units/nl), and S is the surface area (0.3 cm2). Simultaneous controls were performed with each experiment.
Data are presented as means ± SE. All experiments were carried out with a sample size of at least six observations per treatment group. Statistical analysis comparing treatment groups was performed with analysis of variance followed by Dunnett's multiple-range test (22). Correlational analyses were done using the Pearson test for parametric analysis or, when applicable, the Spearman test for nonparametric analysis. P values < 0.05 were deemed statistically significant.
Stable underexpression of novel PKC-θ isoform in multiple clones of intestinal cells.
Intestinal cells cotransfected with cDNA encoding both hygromycin resistance (for selection) and PKC-θ antisense (AS-PKC-θ) stably underexpressed the novel θ-isoform of PKC (Fig. 1). Cell lysates of confluent monolayers were prepared from these transfected cell clones and then analyzed by immunoblotting. Multiple clones of intestinal cells transfected with 1, 2, 3, 4, or 5 μg of AS-PKC-θ demonstrated a dose-dependent underexpression of the PKC-θ (82 kDa) isoform. The clone transfected with 4 μg of AS-PKC-θ provided maximum reduction (−99.5%) in the levels of PKC-θ protein. As might be expected, transfection of only the empty vector (vector alone) did not underexpress PKC-θ (Fig. 1). In empty vector-transfected cells, PKC-θ levels were comparable to those of the WT cells, which exhibited native, steady-state levels of θ-isoform. Underexpression of PKC-θ caused no cellular toxicity (0% cell death assessed by ethidium homodimer probe).
Reduction of native PKC-θ isoform levels and disruption of monolayer integrity.
Underexpression of native PKC-θ by AS-PKC-θ led to loss of monolayer integrity (barrier permeability) in transfected intestinal cells (Fig. 2, A and B). Assessment of monolayer integrity from multiple clones of intestinal cells transfected with 1–5 μg of AS-PKC-θ showed a dose-dependent disruption of monolayer permeability as determined by increased FS clearance (i.e., FS hyperpermeability) (Fig. 2A). For example, in 4 μg of clone underexpressing PKC-θ (and exposed to vehicle), monolayer permeability was disrupted as shown by increased FS clearance to ∼2,250 ± 51 nl·h−1·cm−2, paralleling findings on the reduction of native PKC-θ protein levels in this same clone. WT cells (those expressing native PKC-θ levels), on the other hand, showed normal permeability as demonstrated by basal, low FS clearance. As might be expected, transfection of the empty vector alone did not cause barrier disruption [FS Cl = 17 ± 4 nl·h−1·cm−2 for vector-transfected cells exposed to vehicle and 18 ± 7 nl·h−1·cm−2 for WT cells exposed to vehicle]. Indeed, both empty vector-transfected cells and WT cells responded in a similar fashion to vehicle, exhibiting normal barrier permeability.
Comparison of the destabilizing (hyperpermeabilizing) effects of PKC-θ underexpression on monolayers by using a larger (molecular mass) size probe, namely, FD4 (4 kDa) (Fig. 2B), is also consistent with the noted FS (0.478 kDa) findings. There was a dose-dependent loss of monolayer integrity (increased FD4 clearance) in these same transfected clones (1–5 μg of AS-PKC-θ clones). Not surprisingly, 4 μg of clone underexpressing endogenous PKC-θ led to a maximum increase in monolayer hyperpermeability. As expected, there was a molecular mass-dependent increase in the permeability clearance of these probes in order of decreasing size (4 kDa FD4 < 0.478 kDa FS).
Underexpression of endogenous PKC-θ isoform causes instability of cytoskeletal assembly and cytoarchitecture.
Native PKC-θ underexpression, also in a dose-dependent fashion, injured the microtubule cytoskeleton as demonstrated by a low percentage of intestinal cells displaying normal microtubules (Fig. 3A). In WT cells, similarly to barrier permeability, microtubules were normal. Furthermore, transfection of vector alone, as might be expected, did not affect the microtubules (%normal microtubules = 99 ± 1% for empty vector-transfected cells exposed to vehicle and 98 ± 2% for WT cells exposed to vehicle).
Laser scanning confocal microscopy of immunofluorescently stained microtubule cytoskeleton (Fig. 3B) showed that the 4 μg of clone underexpressing PKC-θ exhibited a loss of the normal cytoskeletal architecture (Fig. 3Bb). This instability was indicated by the intracellular appearance of a fragmented, disrupted, and collapsed microtubule network. WT cells (Fig. 3Ba), in contrast, displayed an intact microtubule network that was dispersed radially throughout the cytosol. This normal cytoarchitecture was indistinguishable from that of the empty vector-transfected cells (Fig. 3Bc), which were also exposed to vehicle. We also assessed occludin in our PKC-θ clones (Fig. 3C). Confocal microscopy revealed that in WT cells (Fig. 3Ca), occludin appeared as an intact structure on the inner side of the plasma membrane. This was demonstrated by a continuous and smooth distribution of the occludin ring (Fig. 3Ca, arrow). In PKC-θ-underexpressing (4 μg) clone (Fig. 3Cb), on the other hand, the occludin ring showed clear fragmentation, disorganization, and disruption. In empty vector clone (Fig. 3Cc), as expected, occludin architecture was highly preserved, resembling that of the WT cells. These findings parallel the destabilizing effects of underexpressing PKC-θ on monolayer integrity (FS or FD4 hyperpermeability).
We then determined the effects of native PKC-θ underexpression on the dynamic alterations in the polymerization and depolymerization states of the microtubule cytoskeleton by assessing the 50-kDa structural protein of microtubules, namely, tubulin. To this end, we isolated and analyzed the tubulin pools (S1 and S2 fractions) using a unique SDS-PAGE fractionation procedure. Analysis of tubulin fractions (Fig. 4A) corroborated the microtubule studies noted above. Only the PKC-θ-underexpressing clones (1–5 μg of AS-PKC-θ) showed an abnormal tubulin assembly as demonstrated by graded reductions in the polymerized S2 tubulin and increases in the monomeric S1 tubulin, indicating disruption of microtubules. As before, the 4-μg inhibitory clone was the most effective at inducing injurious alterations, in this case to the assembly of tubulin. In WT cells, in contrast, neither any decreases in polymerized S2 tubulin nor any increases in monomeric S1 tubulin were observed, indicating normal assembly of the microtubule cytoskeleton. In these WT cells, tubulin polymerization was comparable to the levels seen in the empty vector-transfected cells. Indeed, transfection of vector alone, similarly to its lack of effects on microtubules and barrier permeability, did not affect tubulin assembly (%tubulin assembly = 66% ± 0.4% for empty vector-transfected cells exposed to vehicle and 65% ± 0.3% for WT cells exposed to vehicle).
Figure 4B shows representative immunoblots of the alterations in tubulin assembly (S2 and S1 pools) demonstrating once again that PKC-θ underexpression (4-μg AS-PKC-θ clone shown) effectively decreased the stable polymerized tubulin band density (Fig. 4B, lane c, top) and increased the unstable monomeric tubulin band density (Fig. 4B, lane c, bottom) compared with those of either WT or vector cells (Fig. 4B, lanes a and b, respectively). These findings parallel the destabilizing effects of underexpressing native PKC-θ on intestinal microtubule and monolayer integrity. Because the clone transfected with 4 μg of AS-PKC-θ led to maximum levels of monolayer barrier and cytoskeletal instability, we used this clone for subsequent mechanistic experiments.
Antisense inhibition of PKC-θ induces alterations in phosphorylation state of microtubule (tubulin based) cytoskeleton in intestinal monolayers.
We next probed potential molecular mechanisms underlying the observed effects of PKC-θ on the monolayer cytoskeleton and barrier integrity. To this end, we subjected tubulin, fractionated from both transfected and WT monolayers and then immunoprecipitated, to Western immunoblotting to assess phosphorylation (Fig. 5). Tubulin was markedly phosphorylated in WT cells (Fig. 5, lane a), whereas antisense inhibition of native PKC-θ substantially reduced tubulin phosphorylation (Fig. 5, lane c). In the empty vector-transfected cells (expressing steady-state levels of PKC-θ), as might be expected, such suppression of tubulin phosphorylation was not seen (Fig. 5, lane b). Indeed, the level of tubulin phosphorylation in the empty vector clone was similar to that of WT cells (which also exhibited normal tubulin phosphorylation and native, steady-state levels of PKC-θ).
Endogenous PKC-θ is complexed with tubulin.
Using immunoprecipitation analysis (Figs. 6, A and B), we further investigated the mechanism underlying the apparent stabilizing effects of native PKC-θ on tubulin. In a first series of protocols, cells were immunoprecipitated with a monoclonal PKC-θ antibody, and the immunoprecipitates were then analyzed for the presence of tubulin to determine whether this PKC isoform physically associates with tubulin. Antisense-transfected cells underexpressing native PKC-θ (suppressing endogenous θ) did not show any association between these proteins (Fig. 6A, lane c). In contrast, the amount of tubulin coprecipitation was substantially increased in resting (WT) vehicle-treated cells (Fig. 6A, lane b), indicating the presence of a PKC-θ/tubulin complex under native conditions. Compared with the antisense-transfected cells, as might be expected, transfection of the empty vector alone was ineffective (Fig. 6A, lane d), exhibiting PKC-θ/tubulin coprecipitation comparable to that of WT cells. In a second (reverse) series of protocols (Fig. 6B), anti-tubulin antibody was used, and immune complexes were then analyzed for the presence of PKC-θ. These data further corroborate the above-noted findings. PKC-θ was not seen in the complex in antisense-transfected cells (i.e., no coprecipitation with tubulin) (Fig. 6B, lane c), whereas vehicle-treated WT cells showed an accumulation of native PKC-θ complexes (Fig. 6B, lane b). Similarly, empty vector-transfected cells showed formation of the endogenous PKC-θ/tubulin complexes (Fig. 6B, lane d). In a third series of protocols, we further examined the specificity of the inhibition of formation of the PKC-θ/tubulin complexes in our PKC-θ antisense clone. As expected, probing cell lysates from two other PKC isoform antisense clones (the classical PKC-β2 and the PKC-α antisense) did not prevent the formation of PKC-θ/tubulin complexes (not shown).
Subcellular distribution and activity levels of PKC-θ in intestinal cells: native PKC-θ isoform is constitutively active and found largely in particulate (membrane and cytoskeletal) pools.
Immunoblotting assessment of the cytosolic, membrane, and cytoskeletal fractions of WT cells showed that the native (82 kDa) θ-isoform of PKC is distributed mostly in the particulate fractions (particulate = membrane + cytoskeletal fractions) with a much smaller distribution in the cytosolic fractions (Fig. 7A), indicating the constitutive activity of the θ-isoform of PKC. PKC-θ isoform is “constitutively active” because it is found to be most active in the particulate fractions under native conditions (see Fig. 8 below). In antisense-transfected clone underexpressing PKC-θ (Fig. 7B), in contrast, we found an almost complete absence of PKC-θ in either the particulate fraction or the cytosolic fraction, indicating its inactivity.
Figure 8 shows activity levels of the PKC-θ isoform (in vitro kinase assay) from immunoprecipitated particulate and cytosolic fractions of intestinal cells of both transfected and WT origin. WT cells exposed to vehicle showed normal activation levels for native PKC-θ in the particulate and cytosolic fractions, with most of this activity in the particulate fractions, confirming findings in Fig. 7. In the AS-PKC-θ, there was an almost complete lack of native θ-isoform activity in any cell fractions compared with the vector alone-transfected cells. Vector alone-transfected cells were indistinguishable from WT cells with regard to PKC-θ activity. The above findings collectively indicate that endogenous, native PKC-θ isoform is constitutively active in the membrane and cytoskeletal fractions.
Changes in activity of PKC-θ in intestinal cells robustly correlate with several different indexes of monolayer barrier stability.
Using data from all experimental conditions, we found significant (P < 0.05) correlations (r = 0.97) between PKC-θ levels (in vitro kinase activity assay or optical density from the particulate fractions) and enhanced monolayer barrier integrity (decreased FS clearance). Similarly, we found other robust correlations when two other markers of stability, either microtubule integrity (i.e., %normal) or tubulin assembly (i.e., S2 pool), were correlated with the PKC-θ levels (r = 0.95 or 0.92, respectively, P < 0.05 for each). Still, when other markers of stability, such as enhanced tubulin phosphorylation or reduced tubulin disassembly (i.e., decreased S1 pool), were used against PKC-θ, additional supporting correlations were found (r = 0.93 or 0.96, respectively, P < 0.05).
Dominant negative targeted approach to inactivate endogenous PKC-θ isoform and its resultant loss of monolayer barrier integrity and cytoskeletal stability.
We then used a dominant negative approach for PKC-θ to stably decrease the activity of endogenous PKC-θ isoform. Cells were thus transfected with PKC-θ dominant negative cDNA (DN-PKC-θ) and with plasmid encoding hygromycin resistance. Using this dominant negative mutant model, we were able to nearly completely abrogate the steady-state activity levels for native PKC-θ isoform by ∼99.7% (Fig. 9, 3 μg of clone). By comparison, in WT cells exposed to vehicle, native PKC-θ activity was high in the particulate pools, exhibiting the endogenous, steady-state constitutive activity levels for this isoform. The empty vector-transfected cells exposed to vehicle also showed the steady-state constitutive activity of PKC-θ.
Table 1 demonstrates the dose-dependent effects of varying amounts (1, 2, 3, 4, or 5 μg) of DN-PKC-θ plasmid on the loss of monolayer integrity in intestinal cells. The cell clone that was stably transfected with 3 μg of DN-PKC-θ plasmid resulted in maximum loss of monolayer integrity (hyperpermeability). Thus this dominant negative mutant clone was used for other inhibition experiments.
Analysis of the percentage of dominant negative mutant cells with a normal microtubule cytoskeleton demonstrated that dominant negative inactivation of native PKC-θ causes microtubule disruption (Fig. 10A). Only a low percentage (∼20%) of mutant cells exhibited normal microtubules, indicating microtubule instability. In WT cells, on the other hand, a large percentage of cells, nearly 100%, had normal microtubules.
In parallel, analysis of the state of tubulin assembly from these dominant negative cells demonstrated (Fig. 10B) that inactivation of native PKC-θ attenuates the state of tubulin assembly, leading to tubulin disassembly. A representative tubulin immunoblot further corroborates these findings (Fig. 10C). Furthermore, assessment of tubulin phosphorylation from the same dominant negative mutant (Fig. 10D) indicated that inhibition of endogenous PKC-θ largely reduces tubulin phosphorylation. In WT cells, tubulin was markedly phosphorylated (exhibiting normal steady-state phosphorylation levels for tubulin).
Stable overexpression of PKC-θ isoform effects intestinal monolayer barrier integrity.
The aforementioned findings obtained with the use of two independent, targeted molecular approaches (antisense and dominant negative) suggest that PKC-θ could play a novel role in cytoskeletal and monolayer alterations. In a third independent approach, we further investigated the role of PKC-θ in these alterations. To this end, we cotransfected parental Caco-2 cells (tTA parental cells) with cDNA encoding both hygromycin resistance (for selection) and a TRE system for full-length native PKC-θ, namely, TRE-PKC-θ (Fig. 11). In this TRE system, overexpression of PKC-θ is achieved in the absence of tetracycline, whereas its presence reduces expression to the levels seen in the parental cell line. Cell lysates from these stably transfected cells were then analyzed by Western immunoblotting. Figure 11 shows overexpression of the 82-kDa PKC-θ isozyme in these transfected clones. Total PKC-θ levels were elevated 2.0-fold in these TRE-PKC-θ-overexpressing cells compared with parental cells. Overexpression of PKC-θ at these levels caused no cellular toxicity (0% cell death assessed by ethidium homodimer probe).
Overexpression of PKC-θ isoform destabilizes cell monolayers.
Multiple clones of intestinal cells transfected with 1, 2, 3, 4, or 5 μg of TRE-PKC-θ demonstrated (Table 2) a dose-dependent loss of monolayer integrity. The clone transfected with 4 μg of TRE-PKC-θ provided the highest increase in monolayer hyperpermeability, and thus it was used for subsequent expression experiments.
For example, in Caco-2 cells, PKC-θ overexpression deleteriously affected monolayer integrity (Fig. 12) and the microtubule cytoskeleton (Fig. 13). In cells stably overexpressing native PKC-θ (TRE-PKC-θ) and exposed to vehicle, monolayer integrity was disrupted as determined by increased FS clearance (Fig. 12). Incubation of these same clones with tetracycline (where tetracycline prevents overexpression of PKC-θ), as might be expected, maintained monolayer integrity at normal levels. Similarly, parental cells (those not overexpressing PKC-θ) exposed to vehicle (with or without tetracycline) also showed intact monolayer integrity. Furthermore, transfection of only the empty vector (TRE-z) by itself did not cause monolayer instability (FS Cl = 17 ± 8 nl·h−1·cm−2 for empty vector-transfected cells exposed to vehicle and 19 ± 5 nl·h−1·cm−2 for parental cells exposed to vehicle compared with 3,452 ± 103 and 17 ± 8 nl·h−1·cm−2 for TRE-PKC-θ-overexpressing cells exposed to vehicle). Both empty vector-transfected and parental cells responded in a similar fashion to vehicle, exhibiting normal monolayer integrity (permeability).
PKC-θ expression by itself also disrupted the microtubule cytoskeleton, as demonstrated by the low percentage of cells with normal microtubules (Fig. 13A). This overexpression-induced instability was prevented when tetracycline was present. In parental cells, as for monolayer integrity, microtubules were normal. As before, transfection of control vector alone was ineffective (%normal microtubules = 99 ± 1% for empty vector-transfected cells exposed to vehicle and 97 ± 2% for parental cells exposed to vehicle compared with 9 ± 4% for TRE-PKC-θ-overexpressing cells exposed to vehicle).
Scanning confocal microscopy of the intracellular organization of the microtubule cytoskeleton also showed (Fig. 13B) that clones overexpressing PKC-θ (i.e., TRE-PKC-θ exposed to vehicle) exhibit an abnormal arrangement of the cytoskeleton (Fig. 13Bc). This abnormality was shown by the appearance of a fragmented and collapsed microtubule network. In the presence of tetracycline (which prevents overexpression of PKC-θ, Fig. 13Bd), these same cells displayed a preserved microtubule network as shown by an intact (stellate) architecture of the cytoskeleton. This normal cytoarchitecture is indistinguishable from that of the parental cells exposed to vehicle (Fig. 13B, a and b, with or without tetracycline, respectively). We also assessed the effects of PKC-θ overexpression on occludin (Fig. 13C). Confocal microscopy demonstrated that in clones overexpressing PKC-θ (Fig. 13Cc, TRE-PKC-θ exposed to vehicle), the occludin appeared beaded, fragmented, and disorganized. In the presence of tetracycline (to inhibit PKC-θ overexpression, Fig. 13Cd) in this same clone, normal occludin ring architecture was preserved. Similarly, in the parental cells with or without tetracycline (Fig. 13C, a and b, respectively), occludin appeared as an intact and smooth ring at areas of cell-cell contact.
Analysis of tubulin assembly (Fig. 14A) further demonstrated that PKC-θ-overexpressing clone (TRE PKC-θ, vehicle treated) showed an unstable tubulin assembly as indicated by a reduction in polymerized tubulin. In tetracycline-incubated TRE PKC-θ cells, as might be expected, tubulin assembly was preserved, which was comparable to the normal tubulin polymerization seen in parental cells. Transfection of vector alone, similarly to its lack of effects on microtubules and monolayers, did not alter tubulin assembly (not shown).
Figure 14B shows representative Western blots of the changes in tubulin pools (S2 and S1), further demonstrating that PKC-θ overexpression decreased the stable polymerized tubulin (Fig. 14B, lane c, top), whereas it increased the unstable monomeric tubulin (Fig. 14B, lane c, bottom); these overexpression-induced disruptive effects were prevented by tetracycline (Fig. 14B, corresponding lane d).
PKC-θ expression also reduced tubulin phosphorylation, as determined by immunoblotting of immunoprecipitated tubulin (Fig. 14C). PKC-θ-overexpressing cells showed an abnormally low level of tubulin phosphorylation (Fig. 14C, lane c), which was prevented when tetracycline was present (Fig. 14C, lane d). In parental cells, tubulin phosphorylation was normal (Fig. 14C, lanes a and b). These findings on both tubulin assembly and phosphorylation parallel the destabilizing effects of PKC-θ overexpression on intestinal microtubule and monolayer integrity.
Expression of PKC-θ leads to its increased cytosolic activity and reduced particulate activity.
Overexpression of the native PKC-θ in intestinal cells led to the distribution of its activity (in vitro kinase assay, Fig. 15) into mostly the cytosolic fractions, with a much smaller distribution of activity in the particulate fractions, indicating a reduction in the constitutive activity of PKC-θ isoform. In contrast, parental cells exposed to vehicle (with or without tetracycline) showed constitutive activation of PKC-θ. PKC-θ is constitutively active in parental cells because it is found endogenously most active in the particulate fractions under native conditions. In the presence of tetracycline, as might be expected, PKC-θ-transfected cells also showed normal constitutive activity of PKC-θ.
Finally, we also found significant (P < 0.05) correlations (e.g., r = −0.90) between reduced PKC-θ levels (reduced constitutive activity in the particulate fractions) and increased monolayer instability. Similarly, we found other robust correlations when either microtubule instability (i.e., %disruption) or tubulin disassembly (i.e., increased S1 pool) were correlated with the PKC-θ activity (r = −0.89 or −0.91, respectively, P < 0.05 for each). When other markers of instability, either reduced tubulin assembly (i.e., decreased S2 pool) or decreased tubulin phosphorylation, were used against PKC-θ, additional robust correlations were found (r = −0.88 or −0.95, respectively, P < 0.05 for each).
Using monolayers of intestinal epithelium, we have shown in the current investigation that 1) the 82-kDa θ-isoform of PKC is required for alterations in the microtubule cytoskeleton, occludin, and barrier permeability in epithelial cell monolayers, and 2) the molecular mechanism underlying this unique effect of the PKC-θ isoform appears to involve changes in the phosphorylation and/or assembly of the subunits of the cytoskeleton. To our knowledge, this is the first report that PKC-θ can effect the dynamics of cytoskeletal assembly and permeability clearance in cells.
These two conclusions are supported by independent lines of evidence. First, antisense underexpression of native PKC-θ induces disruption of monolayers (increases in FS and FD4 clearance). This barrier disruption appears to require underexpression and inactivation of PKC-θ activity. In particular, monolayer barrier instability is dependent on inactivation of PKC-θ because of the reduced distribution of PKC-θ in the key particulate (cytoskeletal and membrane) fractions. Second, antisense underexpression of PKC-θ appears to induce instability of the cytoskeleton. For example, underexpression of endogenous PKC-θ reduces the phosphorylation of tubulin; promotes the instability of the molecular dynamics of the tubulin-based cytoskeleton (increases the unstable monomeric tubulin and reduces the stable polymerized tubulin), and decreases the percentage of Caco-2 cells displaying normal microtubules. In WT cells, on the other hand, endogenous PKC-θ not only was constitutively active in the particulate (e.g., cytoskeletal) fractions but also was complexed with the cytoskeleton. Third, dominant negative inactivation of endogenous PKC-θ induces an antisense-like disruption in intestinal cells, causing a destabilizing cascade of alterations:
Fourth, overexpression of PKC-θ, which also leads to a reduction in the amount of PKC-θ in the cytoskeletal and membrane fractions, promotes all measures of monolayer barrier disruption. In these sense-transfected clones, tubulin, microtubules, occludin, and monolayer barrier permeability were substantially disrupted. Fifth, the high strength of the correlations (e.g., PKC-θ vs. cytoskeletal assembly or monolayer barrier integrity, etc.), which explains 85–95% of the variance, indicates that PKC-θ is key for alterations in the dynamic assembly of the cytoskeleton and barrier permeability in intestinal cell monolayers.
Our findings with the use of molecular biology are consistent with the known biochemical properties of PKC isoforms in cells. All PKCs consist of the NH2-terminal regulatory domains that are separated by a flexible hinge region from the COOH-terminal catalytic domains (21). In resting cells, some isoforms of PKC can be found in an inactive form, because they are mostly distributed in the cytosolic (soluble) fractions and only loosely bound to membrane and cytoskeletal (particulate) components. In contrast, in the same resting cells, other PKC isoforms are found in an active form, because they are mainly distributed in the membrane and cytoskeletal fractions. Regulatory domains of PKC isoforms vary from one subfamily to the next as well as among individual isozymes within the same subfamily (5, 10, 18, 21, 40). Indeed, the effects of PKC in cells are complex and appear to vary widely with different experimental settings as well as tissue types. This is not surprising because not only do most cells express multiple, different isozymes of PKC from various subfamilies but also the differences among these PKC isotypes with respect to conditions of activation and intracellular distribution allow individual isoforms to perform distinct biological functions (1, 5, 6, 10, 14, 26, 37, 39). For example, an immunofluorescence study proposed that PKC-ε may be involved in TNF-α-induced damage in the intestinal (IEC-18) cells (17), whereas a pharmacological study suggested that bryostatin-1 attenuates TNF-α-induced barrier disruption via PKC-ε in colonic (T-84) cells (49). A recent study from our laboratory (5) that was based on both pharmacological and molecular biological approaches showed that PKC-δ is involved in oxidant-induced damage in the intestinal (Caco-2) cells. We also showed that the 78-kDa classical β1-isoform of PKC (PKC-β1) is required for both normalization of Ca2+ homeostasis and suppression of NF-κB activation and consequent protection of colonocytes against oxidant injury (6, 14). In other studies, we showed (10) that the 72-kDa atypical ζ-isoform of PKC (PKC-ζ) is required for the suppression of NF-κB and inducible nitric oxide synthase-driven oxidative stress and protection of colonocytes. We thus showed that both PKC-β1 and PKC-ζ isoforms are required for growth factor (EGF, TGF-α)-induced protection of the intestinal epithelium (6, 10, 14), indicating that these PKC isoforms perform unique tasks in protection of intestinal cells. Thus mimicking or activating different isoforms of PKC appears to mediate distinct biological effects (e.g., EGF protection, oxidant injury) on the epithelium. Nonetheless, our current findings on the 82-kDa θ-isoform of PKC, we believe, indicate a unique role among the novel subfamily of PKC isoforms, namely, effects on epithelial barrier permeability via alterations in the phosphorylation and/or assembly of the subunit components of cytoskeleton. We have thus discovered a novel biological mechanism among this subfamily of PKC isoforms.
In the current investigation, we chose to utilize well-established and widely used fluorescent probes such as FS and FD4 for assessing intestinal monolayer barrier stability for several reasons. Fluorescently labeled FD4 and FS compounds offer appropriate choices for determination of epithelial monolayer barrier integrity. First, these probes are convenient because they come in a range of sizes, are nontoxic to cells, are membrane impermeable, and are relatively inexpensive (e.g., 2–10, 14, 27, 29, 45, 47). Second, both in vivo and in vitro studies, including our own, have revealed that these probes move through the monolayer paracellular space and thus are appropriate for assessing monolayer stability (7, 27, 29, 45). Third, the probes are nontoxic lipophobic moieties and are considered to be cell membrane impermeable (7, 27, 45), so their permeability characteristics make them ideal for use in studying monolayer barrier integrity. Accordingly, these probes can be and have been used as tracers to examine many aspects of endothelial and epithelial monolayer barrier stability. For example, the FS and FD probes have been widely used by numerous groups studying GI inflammation (2–4, 7–9, 27, 29, 45, 47). In several studies, these fluorescent probes were used to assess intestinal Caco-2 monolayer barrier integrity (2–10, 14, 27, 29, 38, 47).
The ability of PKC-θ to affect cytoskeleton and monolayer barrier integrity could potentially lead to development of new therapeutics for inflammation, in general, and IBD, in particular, because disruption of mucosal barrier integrity has been documented under these conditions. Specifically, this concept is consistent with characterizations of the pathophysiology of IBD and of the disrupted nature of barrier in IBD (23–25, 30–32, 28, 42, 46). The manifestations of IBD, including ulcerative colitis and Crohn's disease, wax and wane between active (symptomatic) phases of disease, when barrier permeability (i.e., barrier disruption) is thought to be high, and inactive (asymptomatic) phases, when barrier permeability is low. Not surprisingly, we previously showed that the degree of mucosal cytoskeletal instability (an index of barrier stability) correlated with the degree of inflammation and disease severity score in patients with IBD flare-up (30). The presence of unstable cytoskeletal components, which indicates an abnormally leaky barrier, has also been shown in intestinal epithelial monolayers (e.g., Caco-2 cells) under IBD-like conditions (3, 5–7, 10, 14). Consistent with these studies, in other studies loss of gut barrier permeability elicited symptoms of intestinal inflammation and IBD-like conditions in animals (23, 48). Thus induction of barrier hyperpermeability (leaky gut) appears to be crucial to the perpetuation of the active, symptomatic phase of IBD, the so-called IBD flare-up, when intestinal inflammation creates a vicious cycle dependent on oxidative stress, cytoskeletal instability, and, ultimately, mucosal tissue damage. The cytoskeletal and monolayer effects by PKC-θ, as we have found in intestinal cells, could be pivotal in suppressing and/or attenuating the continuation of barrier leakiness and inflammatory processes. Accordingly, developing a means of maintaining mucosal barrier integrity during IBD by using agents that attenuate endogenous signaling pathways (e.g., PKC-θ) might be a simple and effective strategy for the treatment of IBD. PKC-θ mimetics might even synergize with the effects of agents triggering protective pathways (e.g., EGF, PKC-ζ) and/or the currently used antioxidants so that inflammatory processes would be more effectively attenuated via the alterations of key intracellular mechanisms.
In summary, our findings with the use of targeted molecular approaches to stably underexpress, inactivate, or overexpress PKC-θ demonstrate an original concept: this PKC isoform appears to be crucial for a substantial portion of the endogenous stability of the cytoskeleton and barrier permeability in intestinal epithelial monolayers. PKC-θ perhaps is also key in attenuating establishment of an uncontrolled, leaky gut-initiated inflammatory process that can be found under pathophysiological conditions in the GI tract.
This work was supported in part by a grant from Rush University Medical Center, Department of Internal Medicine, and by two National Institutes of Health (NIH) R01 grants, DK-60511 (to A. Banan) and AA-13745 (to A. Keshavarzian).
Portions of this work were presented in abstract form during the annual meeting of the American Gastroenterological Association (Digestive Disease Week), May 2004.
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.
- Copyright © 2004 the American Physiological Society