The polymeric IgR (pIgR) is a central component in the transport of IgA across enterocytes and thereby plays a crucial role in the defense against enteropathogens and in the regulation of circulating IgA levels. The present study was performed to address the novel regulation of pIgR expression in intestinal epithelia undergoing ribosome inactivation. Insults to mucosa that led to ribosome inactivation attenuated pIgR expression in enterocytes. However, IFN regulatory factor-1 (IRF-1) as a central transcription factor of pIgR induction was superinduced by ribosome inactivation in the presence of IFN-γ as a result of mRNA stabilization by the RNA-binding protein HuR. Another important transcription factor for pIgR expression, NF-κB, was marginally involved in suppression of pIgR by ribosome inactivation. In contrast to a positive contribution of HuR in early induction of IRF-1 expression, extended exposure to ribosome inactivation caused nuclear entrapment of HuR, resulting in destabilization of late-phase–induced pIgR mRNA. These HuR-linked differential regulations of pIgR and of IRF-1 led to a reduced mucosal secretion of IgA and, paradoxically, an induction of IRF-1–activated target genes, including colitis-associated IL-7. Therefore, these events can account for ribosome inactivation–related mucosal disorders and provide new insight into interventions for HuR-linked pathogenesis in diverse mucosa-associated diseases, including inflammatory bowel disease and IgA nephritis.

The mucosal epithelial immune system is the first line of defense against various environmental insults, including toxic chemicals and luminal pathogens (1, 2). To protect from environmental Ags, the primary defense molecule produced by lamina propria lymphocytes, IgA, is transported across the epithelium in a basolateral-to-apical direction via the polymeric IgR (pIgR) (3). The pIgR is a transmembrane glycoprotein selectively expressed in the basolateral membrane of the mucosal epithelial cells of the intestinal tract and lung (4, 5). Decreases in epithelial pIgR expression limit the luminal secretion of IgA, which may enhance the risk of luminal microbial infection and subsequent inflammation. IgA deficiency can thus be a predisposing factor to intestinal inflammatory diseases, including inflammatory bowel disease (IBD) (6, 7). Unrestrained commensal or pathogenic bacteria in IgA-deficient individuals are able to easily bind to mucosal epithelial cells and translocate into submucosal regions, leading to the initiation of inflammatory responses. In an animal model of dextran sodium sulfate–mediated colitis, pIgR expression is significantly decreased, and subsequently, the IgA-mediated humoral defense in the mucosa is disrupted (8). Furthermore, genetic ablation of pIgR is associated with decreased mucosal secretion of IgA as well as simultaneous elevations in serum IgA (9, 10). High levels of serum IgA in age-dependent hypo-pIgR, or high IgA-producing models, lead to IgA deposition in the glomerular mesangium, which is a typical symptom of human IgA nephritis (11).

Proinflammatory cytokines, such as IFN-γ, TNF-α, IL-1, and IL-4, play cardinal roles in pIgR gene expression during various bacterial and viral infections (12, 13). In terms of gene regulation, several transcription factor binding sites in the vicinity of the human pIgR gene have been identified that regulate transcriptional activity in response to proinflammatory cytokines (14, 15). IFN-γ or TNF-α enhance de novo synthesis of the IFN regulatory factor-1 (IRF-1), which binds to the IFN-stimulated response element (ISRE) in exon 1 of the pIgR gene in gut epithelial cells (15). Moreover, pIgR gene expression is affected by the NF-κB signaling cascade (16). NF-κB activation by IFN-γ promotes pIgR transcription synergistically with IRF-1 because the NF-κB binding site in intron 1 cooperates with the ISRE in the proximal promoter region of pIgR (16). In addition, posttranscriptional modulation also contributes to pIgR induction by proinflammatory cytokines (17). Chronic proinflammatory stimulation, or oncogenic insults, cause a marked increase in pIgR mRNA stability, mechanistically acting on the 1.8-kb 3′-untranslated region (3′-UTR) of human pIgR mRNA that contains multiple repetitive elements important for the processing and stability of pIgR transcripts (18). In particular, adenylate-uridylate–rich elements (AREs) located in the 3′-UTR of unstable genes mediate selective mRNA stabilization in response to diverse intra- and extracellular signaling molecules (19). Although most RNA-binding proteins destabilize targeted mRNAs, a few of these, including human antigen R (HuR), bind to 3′-UTRs with AREs and stabilize the transcripts, such as those encoding growth factors or proinflammatory cytokines (19).

Among the mechanisms through which environmental factors promote IBD-related pathogenesis, inactivation of mammalian ribosomes has been extensively studied (20). Ribosome inactivation causes acute and chronic mucosal inflammation, and has been investigated as a potent etiological factor of human gut epithelial inflammatory disorders, including IBD (21, 22). Ribosome-inactivating xenobiotics belong to a large family of ribonucleolytic agents. A number of these xenobiotics including deoxynivalenol (DON) and anisomycin (ANS) irreversibly cleave 28S rRNA at a single phosphodiester bond within a universally conserved sequence known as the sarcin-ricin loop (2325). This cleavage leads to peptidyltransferase dysfunction and subsequent global translational arrest (26). Moreover, the ribosomal inactivation elicits ribosome-derived stress responses that stimulate intracellular sentinel signaling pathways, which results in the expression of genes important for cellular homeostasis as well as ones integral for a variety of pathogenic processes involved in cell survival, proliferation, and stress responses (25, 27). For instance, the cellular stress signals from the ribosomal dysfunction evoke the production of proinflammatory cytokines in epithelial and other immune-related cells (28), which leads to both intestinal and systemic inflammation in human mucosal epithelial disorders, including intestinal ulcerative colitis (21, 22, 29). In addition to proinflammatory cytokines, chemical ribosome inactivation can enhance IgA levels in circulation and chronically lead to IgA deposition in the renal glomerular matrix (30, 31).

On the basis of the assumption that ribosome inactivation can alter IgA transport in the mucosa, the current study assessed the effects of ribosome inactivation on pIgR levels in gut epithelia and enterocytes. Alteration of epithelial pIgR levels would be expected to have significant effects on mucosal defenses and total circulating IgA levels in the body. Deficiencies or excesses of luminal IgA or circulating IgA may mechanistically account for ribosome inactivation-related disorders, including intestinal epithelial inflammation. Indeed, the regulatory mechanism of epithelial IgA transport by ribosome inactivation would provide new insights into interventions in human mucosa–associated diseases.

HCT-8, HT-29, and Intestine407 human epithelial cell lines were purchased from the American Type Culture Collection (Manassas, VA). They were maintained in RPMI 1640 medium (Welgene, Daegu, South Korea) supplemented with 10% (v/v) heat-inactivated FBS (Welgene), 50 U/ml penicillin (Welgene), and 50 mg/ml streptomycin (Welgene) in a 5% CO2 humidified incubator at 37°C. Cell number was assessed by trypan blue (Sigma-Aldrich, St. Louis, MO) dye exclusion using a hemocytometer. DON with a purity of 97.6% was isolated from Fusarium graminearum (Sigma-Aldrich). Additional ribosome-inactivating agents, such as ANS, nivalenol (NIV), and 15-acetyl DON, were purchased from Sigma-Aldrich. Recombinant human IFN-γ (PeproTech, Rocky Hill, NJ) was dissolved in distilled water containing 0.1% BSA. Bay 11-7082 was purchased from Calbiochem (San Diego, CA).

Six-week-old female B6C3F1 (C57B1/6J × C3H/HeJ) and C57BL/6J mice were purchased from SamTako Bio Korea (Osan, South Korea), and pIgR KO mice were purchased from Korea Research Institute of Bioscience and Biotechnology (Ochang, South Korea). Animal care and experimental procedures were reviewed and approved by the Institutional Animal Care and Use Committee (PNU-2013-0291). The mice were divided into five groups: vehicle, low-dose ANS (2.5 mg/kg bodyweight), middle-dose ANS (10 mg/kg), high-dose ANS (25 mg/kg), and DON (25 mg/kg). ANS or DON was administered once via oral gavage in 0.2 ml PBS. At 48 h after chemical treatment, the mice were sacrificed by ether anesthesia in a closed container. Intestines were rapidly surgically removed.

For enteropathogenic Escherichia coli (EPEC; E2348/69) infection, C57BL/6J mice were allowed to acclimate for 7 d. All mice were individually housed in ventilated cages with free access to food and water. EPEC was grown to stationary phase in Luria–Bertani broth. Aliquots of the broth culture (1 ml) were centrifuged, and the bacterial pellet was suspended in 1.25 ml PBS. Streptomycin pretreated mice were orally gavaged with PBS or 25mg/kg ANS. After 24 h, a suspension containing ∼1 × 109 E2348/69 cells in 200 μl PBS was introduced into the mice by gavage with a curved needle 4 cm in length with a steel ball at the tip. Control animals received 200 μl sterile PBS. Over the course of infection, the mice were observed daily to assess activity levels and water intake, and body weight was measured. On the fourth day following infection, the animals were sacrificed after anesthetization by isoflurane inhalation and intestinal tissues were processed for further analysis.

C57BL/6J mice were anesthetized and the small intestine was opened longitudinally and then washed in 20 ml HBSS several times until luminal content was completely removed. The intestine was cut into 0.5-mm-long fragments and incubated with 15 ml prewarmed 0.05% trypsin and EDTA (1 mM) in HBSS medium for 20 min at 37°C. After incubation, the detached enterocytes were passed through the cell strainer and collected in RPMI 1640 medium. Cells were separated by centrifugation in the 25/40% discontinuous Percoll (Sigma-Aldrich) gradient at 600 × g for 10 min. Small-intestinal enterocytes were collected from the interface between two different Percoll gradients and resuspended in RPMI 1640 medium.

A CMV-driven short hairpin RNA (shRNA) expression plasmid was constructed by inserting shRNA template into the pSilencer 4.1-CMV-neo vector (Ambion, Austin, TX, USA). Human IRF-1 shRNA targeted the sequence 5′-AGA CCA GAG CAG GAA CAA G-3′. Human sense HuR gene (HuR SC) and antisense HuR (HuR AS) gene construct were provided by Dr. G. Myriam (National Institutes of Health, Baltimore, MD) (32, 33). pGL3, containing a pIgR promoter construct (pSC1), as well as mutated forms pSC53, pSC55, pSC57, were donated by Dr. F.-E. Johansen (Rikshospitalet University Hospital, Oslo, Norway). Plasmid pGL2, containing the −563/+29 region of the pIgR promoter construct, was obtained from Dr. C.S. Kaetzel (University of Kentucky, Lexington, KY). The cDNA containing the entire coding region of human activating transcription factor 3 (ATF3) was cloned into the expression plasmid pcDNA3.1 Zeo(+) (Invitrogen) using T4 DNA ligase (New England Biolabs, Beverly, MA). This vector was named ATF3-SC.

Protein expression was analyzed by Western immunoblot analysis using rabbit polyclonal anti-human actin Ab, rabbit polyclonal anti-pIgR Ab (Santa Cruz Biotechnology, Santa Cruz, CA), and anti-rabbit IgG secondary Ab (Enzo Life Science, Plymouth Meeting, PA). Cells were washed with ice-cold PBS, lysed in boiling lysis buffer (1% [w/v] SDS, 1 mM sodium orthovanadate and 10 mM Tris [pH 7.4]), and sonicated for 5 s. Protein content of the lysates was quantified using a BCA protein assay kit (Pierce, Rockford, IL). Fifty micrograms of protein were separated using a Bio-Rad gel mini electrophoresis system (Bio-Rad, Hercules, CA). Proteins were transferred onto a polyvinylidene difluoride membrane (Amersham Biosciences, Piscataway, NJ), and the blots were blocked for 1 h with 5% skim milk in TBST and probed with each Ab overnight at 4°C. After washing three times with TBST, blots were incubated with HRP-conjugated secondary Ab for 2 h and washed with TBST an additional three times. Bound Abs were detected using an ECL substrate (ELPIS Biotech, Taejon, South Korea).

To isolate the nuclear and cytosolic protein, cells were harvested using cold PBS containing protease inhibitor mixture (Sigma-Aldrich). The cell pellet was suspended in protein lysis buffer containing 10 mM HEPES, 10 mM KCl, 1.5 mM MgCl2, 0.5 mM DTT, 0.5 mM PMSF, 0.1% Nonidet P-40, and protease inhibitor mixture; incubated for 10 min on ice; and centrifuged. Supernatant (the cytosolic fraction) was collected while the pellet was suspended in the buffer containing 20 mM HEPES, 1.5 mM MgCl2, 420 mM NaCl, 0.2 mM EDTA, 0.5 mM DTT, 0.2 mM PMSF, 25% glycerol, and a protease inhibitor mixture. After incubating for 10 min on ice, the samples were centrifuged, and the supernatants (nuclear proteins) were collected, aliquoted, and stored at −80°C before analysis.

RNA was extracted with Ribo EX (Gene All, Seoul, Korea) according to the manufacturer’s instructions. RNA (100 ng) from each sample was transcribed to cDNA by Prime Moloney murine leukemia virus reverse transcriptase (Genetbio, Nonsan, South Korea). The amplification was performed with HS Prime Taq polymerase (Genetbio) in a MyCycler thermal cycler (Bio-Rad) using the following manufacturer’s instructions. An aliquot of each PCR product was subjected to 1% (w/v) agarose gel electrophoresis and visualized by staining with ethidium bromide. The forward and reverse complement PCR primers for amplification of each gene were human pIgR (5′-TGC TAC TAC CCA CCC ACC TC-3′ and 5′-AAC CAC TGG AGT CGA TGA CC-3′), mouse pIgR (5′-GGT GAC TCT CGC TGG AGA AC-3′ and 5′-TGC ATC TGT TGG GTT GAC AT-3′), human IRF-1 (5′-ACC CTG GCT AGA GAT GCA GA-3′ and 5′-TAG CTG CTG TGG TCA TCA GG-3′), human IRF-2 (5′-TCT CCT GAG TAT GCG GTC CT-3′ and 5′-CCC CAT GTT GCT GAG GTA CT-3′, human IL-7 (5′-TTT TCC TGC GGT GAT TCG GA-3′ and 5′-CTC TCA CCG CCC ATA GTC AC-3′), mouse IL-7 (5′-GGG AGT GAT TAT GGG TGG TG-3′ and 5′-TGG TTC ATT ATT CGG GCA AT-3′), human GAPDH (5′-TCA ACG GAT TTG GTC GTA TT-3′ and 5′-CTG TGG TCA TGA GTC CTT CC-3′), and mouse GAPDH (5′-TCA ACG GAT TTG GCC GTA TT-3′ and 5′-CTG TGG TCA TGA GCC CTT CC-3′). For real-time PCR, FAM was used as the fluorescent reporter dye and was conjugated to the 5′-ends of the probes to detect amplified cDNA in an iCycler thermal cycler (Bio-Rad) using the following manufacturer’s instructions. Each sample was tested in triplicate to ensure statistical significance. The relative quantification of gene expression was performed using the comparative threshold cycle method. In all experiments, GAPDH was used as the endogenous control. Results were analyzed in a relative quantitation study with the vehicle control.

Cells were washed with cold PBS, lysed with passive lysis buffer (Promega), and then centrifuged at 12,000 × g for 4 min. Luciferase activity was measured with a Model TD-20/20 dual-mode luminometer (Turner Designs, Sunnyvale, CA). Supernatant (10 μl) was mixed with 40 μl firefly luciferase assay substrate solution, and the reaction was stopped with 40 μl Renilla luciferase stop solution (Promega). The firefly luciferase activity was normalized against Renilla luciferase activity by dividing firefly luciferase activity by Renilla luciferase activity. The calculated relative luciferase units were compared.

Trypsinized cells (5 × 105) were prepared and suspended in 1 ml culture medium. The cells were immediately fixed using 4% paraformaldehyde and incubated at room temperature for 10 min. After washing three times with PBS, cells were permeabilized by adding permeabilization buffer (0.1 mM EDTA and 0.1% Triton X-100 in PBS) at BD Biosciences for 20 min. After washing three times with PBS, cells were blocked for 30 min with 10% FBS in PBS and incubated in buffer (10% FBS in PBS) containing a 1:100 dilution of rabbit polyclonal anti-human pIgR primary Ab (Santa Cruz Biotechnology) at BD Biosciences for 2 h and then repeatedly washed using PBS. Cells were then incubated with Alexa Fluor 488–conjugated anti-rabbit IgG (H+L; Invitrogen) for 2 h at room temperature, followed by repeated PBS washes. The fluorescence associated with single cells was measured and analyzed using FACSCanto II (BD Biosciences, San Jose, CA). Data from 5000 cells were collected in the list mode.

Cells were seeded and incubated for 48 h in a glass-bottom culture dish (SPL Life Sciences, Pocheon, South Korea). After various experimental treatments, cells were fixed with 4% paraformaldehyde for 10 min, permeabilized with 0.2% Triton X-100 for 10 min, blocked with 3% BSA for 2 h, and incubated with a 1:200 dilution of mouse polyclonal anti-HuR Ab (Santa Cruz Biotechnology) in 3% BSA at room temperature for 2 h. The cells were then washed with PBS, incubated with Alexa Fluor 546–conjugated goat anti-mouse IgG (H+L; Invitrogen) for 2 h at room temperature, repeatedly washed with PBS, and stained with 100 ng/ml DAPI for 10 min. Confocal images were obtained by Olympus FV1000 confocal microscope (Olympus, Tokyo, Japan) with single-line excitation (546 nm) or multitrack sequential excitation (546 and 633 nm). Images were acquired and processed by using FV10-ASW software (Olympus). The intensity of signals from individual cells was measured by Multi Gauge software (Fujifilm, Tokyo, Japan).

Formalin-fixed and paraffin-embedded tissue sections were deparaffinized in xylene and epitope-retrieved in retrieval solution (DakoCytomation, Carpinteria, CA). To prevent nonspecific binding, deparaffinized sections were preincubated with 3% BSA in PBS for 1 h and then incubated with rabbit anti-human pIgR (Santa Cruz Biotechnology) primary Ab (diluted 1:200) overnight at 4°C. The slides were washed five times with TBST for 5 min and incubated with anti-rabbit IgG:HRP secondary Ab (Enzo Life Science) for 3 h at room temperature. Ab binding was detected using diaminobenzidine (DakoCytomation), and images were acquired using a Zeiss Axio Imager M2 (Carl Zeiss, Oberkochen, Germany). To analyze the proportion of positive stained cells, at least four representative areas were measured by computer-assisted analysis using Histo-quest software 4.0 (TissueGnostics, Vienna, Austria).

Mouse fecal IgA–coated bacterial analysis was performed using a modified protocol for human fecal IgA–coated bacterial analysis (34). Eighty milligrams of feces were diluted in 750 μl sterile PBS (pH 7.2) and homogenized with a Vortex mixer for 1 min. To separate fecal particles, homogenized feces were centrifuged at low speed (35 × g) for 20 min. One hundred microliters of each supernatant were washed once in 1 ml sterile PBS and centrifuged at 8,000 × g for 10 min. The pellet was suspended in 60 μl BSA/PBS containing a 1:100 dilution of FITC-labeled goat anti-mouse IgA Ab (Santa Cruz Biotechnology) and incubated for 1 h at room temperature. Samples were washed twice with 1 ml sterile PBS. To label total bacteria prior to flow cytometry detection, the pellet was suspended in PBS, and 20 μl propidium iodine (4 mg/ml) was added. The IgA-coated bacterial fluorescence was measured and analyzed using FACSCanto II (BD Biosciences). Data from 5,000 bacteria were collected in the list mode.

HCT-8 cells were seeded at 2.5 × 106 cells per 100-mm dish in complete RPMI 1640 and grown for 48 h. After treatment, protein and RNA were cross-linked with 1% formaldehyde for 10 min at room temperature. The cytoplasmic extract was incubated overnight at 4°C with 5 mg of either goat anti-mouse IgG (nonspecific control) or anti-HuR Ab. The Ab-bound complexes were precipitated with protein G–Sepharose beads, and then sequentially washed in low-salt, high-salt, LiCl, and Tris-EDTA buffers (5 -min per wash). The protein–RNA complexes were eluted from the protein G–Sepharose beads with 250 ml elution buffer at 37°C for 15 min. RNA in the immunoprecipitated complexes was released by reversing the cross-linkage by incubating at 65°C for 4–5 h in 200 mM NaCl and 20 mg proteinase K. RNA was then extracted with Ribo EX reagent (Gene All) and subjected to real-time PCR.

Results shown are representative of three independent experiments. Statistics were calculated using the Student two-tailed t test. Densitometric analyses were performed using SigmaPlot.

On the basis of the assumption that altered IgA production in experimental animals with ribosomal dysfunction can be related to intestinal pIgR expression, we assessed the effect of ribosome inactivation on epithelial pIgR expression in the murine intestine. Although normal gut epithelia expressed relatively high levels of pIgR, exposure of the gastrointestinal tract to chemical insults that cause ribosome inactivation (ANS or DON) led to suppression of levels of the pIgR protein (Fig. 1A). Moreover, ribosome-inactivating ANS decreased epithelial pIgR mRNA expression in a dose-dependent manner (Fig. 1B). In addition, the length of villus was also slightly reduced by the ribosomal inactivation (Fig. 1C). To address which part of the small intestine is the most sensitive to ribosome inactivation, pIgR expressions in duodenum, jejunum, and ileum were compared. Of note, ileal pIgR was significantly downregulated by chemical ribosomal inactivation (Fig. 1D). Although significantly changes were not detected, similar patterns of reduction were observed in duodenum and jejunum (Fig. 1D). Moreover, pIgR mRNA expression in isolated mouse enterocytes was also significantly suppressed by ribosome inactivation (Fig. 1E). Because pIgR plays an important role in IgA transport from the lamina propria to the lumen, pIgR suppression by genetic knockout or ribosome inactivation caused IgA accumulation in the lamina propria of intestinal villi (Fig. 2A). Subsequently, a decrease in luminal secretion of IgA was noninvasively detected by observing a reduction of fecal IgA-coated bacteria (Fig. 2B). Because the secreted IgA is a critical defense molecule against pathogenic bacteria, pIgR suppression and reduced release of IgA as a result of ribosome inactivation can allow for greater adherence or invasion of mucosal pathogens. In the current study, the distribution of GFP-labeled EPEC was monitored in murine intestinal epithelium in mice with pIgR deficiency or ribosomal dysfunction. Animal models have been used to investigate the epithelial response to EPEC homologs (35, 36); these models include rabbits infected with rabbit-specific EPEC and mice infected with Citrobacter rodentium. Although diseases because of infection with rabbit-specific EPEC or C. rodentium are similar to other EPEC-induced diseases, these homolog models are limited because of differences in many pathophysiological patterns observed in the EPEC-triggered infection. In particular, IgA-mediated defense plays critical roles in EPEC infection, whereas it is marginal in C. rodentium model (37, 38). The present study used an established mouse model of EPEC infection (3941) to assess the role of IgA secretion via pIgR. In this study, C57BL/6J mice are particularly susceptible to infection by EPEC and serve as a suitable in vivo model for studying epithelial response to EPEC infection without severe diarrhea. EPEC primarily causes inflammatory gastrointestinal illness by colonizing the epithelial lining of the small intestine. A fraction of the introduced EPEC established colonies on the epithelial surface, and their intestinal adherence persisted during the period of experimental observation. Of note, bacterial adherence to the colon epithelial surface was enhanced by pIgR deficiency or by ribosome inactivation (Fig. 2C). It is reported that the intestinal colonization by EPEC notably decreases from the third day of postinfection in the same model as that used in the current study (42). The persistent surface attachment of EPEC in mice with pIgR deficiency or ribosomal dysfunction was observed, compared with marginally remaining bacterial colonization in the control mice on the fourth day of postinfection (Fig. 2C). This suggested that movement of the bacteria toward the gut epithelium was facilitated by the breakdown of the IgA secretion–mediated defense mechanism.

FIGURE 1.

Effect of ribosome inactivation on pIgR expression in the mouse intestine. Mice were orally treated with the vehicle, 25 mg/kg ANS or DON for 24 h. (A) Sections of small intestinal tissues were stained with anti-pIgR Ab (original magnification ×100) and measured relative to the amount of pIgR expression using Histo-quest tissue analysis software (right panel). *p < 0.05, significant difference from the control. (B) The mRNA from whole small intestines was analyzed using real-time RT-PCR. *p < 0.05, significant difference from the control. (C) The length of villus of the proximal and distal small intestine were measured and compared. *p < 0.05, significant difference from the vehicle control group. (D) The mRNA from duodenum, jejunum, and ileum were analyzed using real-time RT-PCR. *p < 0.05, significant difference from each control. (E) Isolated mice enterocytes were treated with 500 ng/ml DON or 50 ng/ml ANS for 48 h. The mRNA levels were measured using real-time RT-PCR. *p < 0.05, significant difference from the control. Amounts of mouse mRNA were normalized to those of mouse GAPDH mRNA.

FIGURE 1.

Effect of ribosome inactivation on pIgR expression in the mouse intestine. Mice were orally treated with the vehicle, 25 mg/kg ANS or DON for 24 h. (A) Sections of small intestinal tissues were stained with anti-pIgR Ab (original magnification ×100) and measured relative to the amount of pIgR expression using Histo-quest tissue analysis software (right panel). *p < 0.05, significant difference from the control. (B) The mRNA from whole small intestines was analyzed using real-time RT-PCR. *p < 0.05, significant difference from the control. (C) The length of villus of the proximal and distal small intestine were measured and compared. *p < 0.05, significant difference from the vehicle control group. (D) The mRNA from duodenum, jejunum, and ileum were analyzed using real-time RT-PCR. *p < 0.05, significant difference from each control. (E) Isolated mice enterocytes were treated with 500 ng/ml DON or 50 ng/ml ANS for 48 h. The mRNA levels were measured using real-time RT-PCR. *p < 0.05, significant difference from the control. Amounts of mouse mRNA were normalized to those of mouse GAPDH mRNA.

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FIGURE 2.

Effect of ribosome inactivation on IgA secretion. (A) Mice were orally treated with the vehicle or ANS for 24 h. Sections of small intestinal tissues were stained with anti–IgA-FITC Ab (green) and DAPI (blue) (original magnification ×200). *p < 0.05, significant difference from the control. (B) Mice were orally treated with the vehicle or 25 mg/kg ANS for 24 or 48 h, and fecal bacteria coated with IgA were quantified using flow cytometry analysis. *p < 0.05, significant difference from the control. (C) Mice pretreated with streptomycin for 24 h were orally gavaged with the vehicle or 25 mg/kg ANS. After 24 h treatment, mice were infected with GFP-labeled EPEC (1 × 109 CFU/200 μl) for 4 d. Sections of mice intestinal tissues were stained with DAPI (blue) (original magnification ×200). *p < 0.05, significant difference from the control.

FIGURE 2.

Effect of ribosome inactivation on IgA secretion. (A) Mice were orally treated with the vehicle or ANS for 24 h. Sections of small intestinal tissues were stained with anti–IgA-FITC Ab (green) and DAPI (blue) (original magnification ×200). *p < 0.05, significant difference from the control. (B) Mice were orally treated with the vehicle or 25 mg/kg ANS for 24 or 48 h, and fecal bacteria coated with IgA were quantified using flow cytometry analysis. *p < 0.05, significant difference from the control. (C) Mice pretreated with streptomycin for 24 h were orally gavaged with the vehicle or 25 mg/kg ANS. After 24 h treatment, mice were infected with GFP-labeled EPEC (1 × 109 CFU/200 μl) for 4 d. Sections of mice intestinal tissues were stained with DAPI (blue) (original magnification ×200). *p < 0.05, significant difference from the control.

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The effect of ribosome inactivation on pIgR expression was also assessed in HCT-8 human intestinal epithelial cells. The HCT-8 cell line is widely used as an enterocyte model for inflammatory diseases and microbial infection (43, 44). In addition, the ileocecum of the small intestine, the source of HCT-8 cells, is one of the regions most susceptible to pathogenesis associated with ribosome inactivation (45, 46). However, isolated intestinal epithelial cells generally show marginal levels of pIgR expression; thus, additional pIgR-enhancing endogenous factors, such as IFN-γ, are added to simulate a physiologically inflamed gut epithelium environment. In the current study, IFN-γ was added to activate intestinal epithelial cells to augment pIgR expression. The expression of IFN-γ–induced pIgR protein in the presence of ribosome-inactivating stress was assessed via flow cytometry (Fig. 3). Similar to the in vivo data, pIgR expression was significantly suppressed by ribosome-inactivating chemicals, such as DON or ANS, in a dose-dependent manner (Fig. 3A, 3B). Moreover, other ribosome-insulting chemicals, including NIV and 15-acetyl-deoxynivalenol, also suppressed pIgR protein expression in different intestinal epithelial cell lines, including HCT-8, HT-29, and Intestine 407 (Fig. 3C–E). Taken together, the in vivo and in vitro data indicated that ribosome inactivation attenuated pIgR expression in gut epithelial cells, which accounted for the reduced luminal secretion of IgA and subsequent disruption of intestinal epithelial defense against enteropathogenic bacteria.

FIGURE 3.

Effect of ribosome inactivation on pIgR expression in human intestinal epithelial cells. (A and B) HCT-8 cells were treated with different doses of ribosome-inactivating agents (DON and ANS) in the presence of 20 ng/ml IFN-γ for 48 h. (C) HCT-8 cells were treated with 500 ng/ml of each ribosome-inactivating agent (DON, ANS, NIV, or 15-acetyl DON [15 AcD]) in the presence of 20 ng/ml IFN-γ for 48 h. (D and E) Other human intestinal epithelial cells (HT-29 and Intestine 407) were treated with 1000 ng/ml of each ribosome-inactivating agent (DON, ANS, NIV, or 15 AcD) in the presence of 20 ng/ml IFN-γ for 48 h. In (A–E), cellular fluorescence from binding of anti-pIgR Ab was measured using flow cytometry analysis. *p < 0.05, significant difference from only the IFN-γ–treated group. P1 gating was set for the pIgR expression of cells without the isotype Ig–responsiveness. Mean fluorescent intensity (MFI) ratio was calculated by dividing by MFI of the isotype Ig treatment group.

FIGURE 3.

Effect of ribosome inactivation on pIgR expression in human intestinal epithelial cells. (A and B) HCT-8 cells were treated with different doses of ribosome-inactivating agents (DON and ANS) in the presence of 20 ng/ml IFN-γ for 48 h. (C) HCT-8 cells were treated with 500 ng/ml of each ribosome-inactivating agent (DON, ANS, NIV, or 15-acetyl DON [15 AcD]) in the presence of 20 ng/ml IFN-γ for 48 h. (D and E) Other human intestinal epithelial cells (HT-29 and Intestine 407) were treated with 1000 ng/ml of each ribosome-inactivating agent (DON, ANS, NIV, or 15 AcD) in the presence of 20 ng/ml IFN-γ for 48 h. In (A–E), cellular fluorescence from binding of anti-pIgR Ab was measured using flow cytometry analysis. *p < 0.05, significant difference from only the IFN-γ–treated group. P1 gating was set for the pIgR expression of cells without the isotype Ig–responsiveness. Mean fluorescent intensity (MFI) ratio was calculated by dividing by MFI of the isotype Ig treatment group.

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In terms of pIgR induction, IRF-1 is a predominant transcription factor that binds to the IFN-stimulated response element in the pIgR gene (15). We therefore assessed the effect of ribosome inactivation on IRF-1 expression in human enterocytes. Genetic knockdown of IRF-1 using shRNA significantly suppressed IFN-γ–induced pIgR mRNA expression, indicating that pIgR induction that results from IFN-γ treatment is IRF-1 dependent (Fig. 4A), which confirms the previous study (15). IRF-1 regulates expression of target genes by binding to ISRE in their promoters. To test the effect of IFN-γ and ribosomal inactivation in pIgR promoter with ISRE, we assessed transcriptional activity by using the reporter plasmid for human pIgR promoter (−563 to +29), which includes three ISRE motifs (Fig. 4B). IFN-γ increased the pIgR transcriptional activity, which was significantly enhanced by ribosome inactivation (Fig. 4C). Moreover, to assess the effects of ribosome inactivation on IRF-1 expression, IFN-γ–induced IRF-1 mRNA expression was measured in enterocytes exposed to chemical ribosome inactivators, including DON and ANS. Surprisingly, ribosome inactivation in the presence of IFN-γ potently superinduced IRF-1 mRNA expression to levels greater than those in enterocytes exposed to only IFN-γ (Fig. 4D). In addition to the in vitro observation, the murine intestinal epithelia exposed to the ribosomal insult also showed significantly higher levels of IRF-1 expression than the control (Fig. 4E). Moreover, IL-7, which is induced by IFN-γ and is a representative target of IRF-1 (47), was potently upregulated by ribosome inactivation under IFN-γ treatment but was suppressed by genetic knockdown of IRF-1 (Fig. 4F). Similarly, the murine intestine exposed to the ribosomal insult also showed significantly higher levels of IL-7 expression than the control (Fig. 4G). To understand the mechanism of the ribosome inactivation–mediated superinduction of IRF-1 in the presence of IFN-γ, posttranscriptional regulation mechanism was assessed in enterocytes in the presence of IFN-γ. Ribosome inactivation had marginal effects on the stability of IFN-γ–induced IRF-1 mRNA at the late time points (6–24 h) after the transcriptional arrest (Fig. 4H). However, IRF-1 mRNA stability at the earlier time points (0–2 h) was significant enhanced by the ribosomal inactivation under IFN-γ treatment (Fig. 4I, 4J). To understand the molecular mechanism of ribosome inactivation–induced posttranscriptional regulation of IRF-1 mRNA stability, we examined the effects of altering the levels of the HuR protein, which is known to bind to and thereby stabilize chemokine mRNAs that contain AREs in the 3′-UTR region (48). The IRF-1 transcript has three such AREs in its 3′-UTR and thus can be potently regulated by the HuR protein. The genetic suppression of HuR expression using an antisense expression plasmid decreased IRF-1 mRNA induction by ribosome inactivation in the presence of IFN-γ (Fig. 4K). Moreover, IFN-γ–induced binding of HuR proteins to 3′-UTR of IRF-1 transcript, which was significantly enhanced by ribosomal inactivation (Fig. 4L). Taken together, ribosomal stress enhanced IFN-γ–induced IRF-1 expression via HuR protein. This does not, however, account for pIgR’s suppression by ribosomal stress, even though IRF-1 was a positive modulator of pIgR induction by IFN-γ. Therefore, other pIgR regulatory factors involved in pIgR expression need to be addressed in enterocytes under ribosomal stress.

FIGURE 4.

Involvement of IRF-1 in the regulation of pIgR expression in human enterocytes. (A) HCT-8 cells expressing the empty vector, or IRF-1–specific shRNA-containing plasmid, were treated with 20 ng/ml IFN-γ, 500 ng/ml DON, 50 ng/ml ANS, or combinations (IFN+DON or IFN+ANS) for 48 h. Each mRNA was measured using real-time RT-PCR. *p < 0.05, significant difference from the IFN-γ–treated group. Blots in the box indicate the representative data for cDNA from IFN-γ plus DON–treated cells using conventional RT-PCR. (B) Maps of reporter constructs for human pIgR promoter containing three ISRE binding sites from −563 to +29 bp without NF-κB binding site. (C) HCT-8 cells transfected with pIgR promoter–linked reporter plasmid in (B) were treated with vehicle, 20 ng/ml IFN-γ, 500 ng/ml DON, or combination for 24 h. *p < 0.05, significant difference from the group treated with IFN-γ alone (D) HCT-8 cells were treated with 20 ng/ml IFN-γ, 500 ng/ml DON, 50 ng/ml ANS, or combinations (IFN+DON or IFN+ANS) for 24 h. Each mRNA was measured using real-time RT-PCR. *p < 0.05, significant difference from the IFN-γ–treated group. (E) Mice were orally gavaged with vehicle or 25 mg/kg DON and intestinal tissues were collected at 24 h after treatment. Expression of IRF-1 was visualized with anti–IRF-1 Ab (brown), which was counterstained with hematoxylin (blue) (original magnification ×200). Percentage of IRF-1 expression was measured by using Histo-quest tissue analysis software. *p < 0.05, significant difference from the control. (F) HCT-8 cells expressing the control empty vector, or IRF-1 shRNA-expression plasmid, were treated with vehicle, 20 ng/ml IFNγ, 500 ng/ml DON, or combination for 4 h. The mRNA levels were measured using real-time RT-PCR. *p < 0.05, significant difference from the control group. Blots in the box indicate the representative data for cDNA from IFN-γ plus DON–treated cells using conventional RT-PCR. (G) Mice were orally treated with the vehicle or 10 mg/kg or 25 mg/kg ANS for 24 h. The mRNA from whole small intestines was analyzed using real-time RT-PCR. *p < 0.05, significant difference from the vehicle control. (HJ) HCT-8 cells were treated with 20 ng/ml IFN-γ for 45 min to reach the maximum level, and then, the transcription was arrested by adding 5 μM actinomycin D for the indicated times. The mRNA was analyzed by real-time RT-PCR. *p < 0.05, significant difference from the group treated with IFN-γ alone. (K) HCT-8 cells transfected with empty vector, or antisense HuR expression plasmid, were treated with vehicle, 20 ng/ml IFN-γ, 500 ng/ml DON, or combination for 30 min. mRNA was analyzed by real-time RT-PCR. *p < 0.05, significant difference in empty vector–transfected group. (L) RNA immunoprecipitation assays were performed to measure HuR protein bound to the IRF-1 transcript in cells. HCT-8 cells were treated with 20 ng/ml IFNγ (I) in the presence or absence of 500 ng/ml DON (D) for 1 h. Immunoprecipitated transcript was measured using real-time RT-PCR. *p < 0.05, significant difference from the control group.

FIGURE 4.

Involvement of IRF-1 in the regulation of pIgR expression in human enterocytes. (A) HCT-8 cells expressing the empty vector, or IRF-1–specific shRNA-containing plasmid, were treated with 20 ng/ml IFN-γ, 500 ng/ml DON, 50 ng/ml ANS, or combinations (IFN+DON or IFN+ANS) for 48 h. Each mRNA was measured using real-time RT-PCR. *p < 0.05, significant difference from the IFN-γ–treated group. Blots in the box indicate the representative data for cDNA from IFN-γ plus DON–treated cells using conventional RT-PCR. (B) Maps of reporter constructs for human pIgR promoter containing three ISRE binding sites from −563 to +29 bp without NF-κB binding site. (C) HCT-8 cells transfected with pIgR promoter–linked reporter plasmid in (B) were treated with vehicle, 20 ng/ml IFN-γ, 500 ng/ml DON, or combination for 24 h. *p < 0.05, significant difference from the group treated with IFN-γ alone (D) HCT-8 cells were treated with 20 ng/ml IFN-γ, 500 ng/ml DON, 50 ng/ml ANS, or combinations (IFN+DON or IFN+ANS) for 24 h. Each mRNA was measured using real-time RT-PCR. *p < 0.05, significant difference from the IFN-γ–treated group. (E) Mice were orally gavaged with vehicle or 25 mg/kg DON and intestinal tissues were collected at 24 h after treatment. Expression of IRF-1 was visualized with anti–IRF-1 Ab (brown), which was counterstained with hematoxylin (blue) (original magnification ×200). Percentage of IRF-1 expression was measured by using Histo-quest tissue analysis software. *p < 0.05, significant difference from the control. (F) HCT-8 cells expressing the control empty vector, or IRF-1 shRNA-expression plasmid, were treated with vehicle, 20 ng/ml IFNγ, 500 ng/ml DON, or combination for 4 h. The mRNA levels were measured using real-time RT-PCR. *p < 0.05, significant difference from the control group. Blots in the box indicate the representative data for cDNA from IFN-γ plus DON–treated cells using conventional RT-PCR. (G) Mice were orally treated with the vehicle or 10 mg/kg or 25 mg/kg ANS for 24 h. The mRNA from whole small intestines was analyzed using real-time RT-PCR. *p < 0.05, significant difference from the vehicle control. (HJ) HCT-8 cells were treated with 20 ng/ml IFN-γ for 45 min to reach the maximum level, and then, the transcription was arrested by adding 5 μM actinomycin D for the indicated times. The mRNA was analyzed by real-time RT-PCR. *p < 0.05, significant difference from the group treated with IFN-γ alone. (K) HCT-8 cells transfected with empty vector, or antisense HuR expression plasmid, were treated with vehicle, 20 ng/ml IFN-γ, 500 ng/ml DON, or combination for 30 min. mRNA was analyzed by real-time RT-PCR. *p < 0.05, significant difference in empty vector–transfected group. (L) RNA immunoprecipitation assays were performed to measure HuR protein bound to the IRF-1 transcript in cells. HCT-8 cells were treated with 20 ng/ml IFNγ (I) in the presence or absence of 500 ng/ml DON (D) for 1 h. Immunoprecipitated transcript was measured using real-time RT-PCR. *p < 0.05, significant difference from the control group.

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As previously mentioned, NF-κB is considered to be another crucial transcription factor for transcriptional activation of the pIgR promoter (49). We therefore first assessed the effects of ribosomal stress on IFN-γ–activated NF-κB in enterocytes. IFN-γ elevated p65 phosphorylation, which was suppressed by the chemical ribosomal-stress agents (Fig. 5A). The negative regulation of NF-κB responsiveness by ribosomal stress was confirmed by observing a reduction in nuclear translocation of NF-κB in enterocytes exposed to the ribosomal stress (Fig. 5B). This pattern of negative regulation of IFN-γ–activated NF-κB signaling by ribosomal stress was similar to findings from our previous investigations on the effects of ribosome inactivation on bacterial product–activated NF-κB signals in enterocytes (50). To assess the involvement of NF-κB in IFN-γ–triggered pIgR transcriptional activation, we examined pIgR promoter–dependent transcriptional activities in response to IFN-γ using an NF-κB binding site mutation construct. The luciferase reporter plasmid pSC1 is composed of 2.7 kb of upstream promoter sequences, ISRE-containing exon 1, 5.7 kb of intron 1, and exon 2. IRF-1 binds to an ISRE in exon 1 and NF-κB binding elements are located in the upstream promoter and intron 1 (Fig. 5C). Because previous results indicated that IRF-1 was not involved in pIgR suppression in response to ribosome inactivation, the effects of NF-κB binding site mutations on pIgR transcriptional activities were assessed in IFN-γ–treated enterocytes (Fig. 5 C). Although NF-κB levels were enhanced by IFN-γ, mutations of the NF-κB–binding cis elements did not significantly influence the pIgR transcriptional activity (Fig. 5D). This indicated that altered NF-κB signals did not affect IFN-γ–induced pIgR expression in enterocytes. To test the possibility that other NF-κB sites may play important roles in pIgR transcript, effects of blocking of NF-κB trans-elements, with broad-spectrum IKK inhibitor, BAY 11-7082, on pIgR transcriptional activities in response to IFN-γ were assessed. This blocking also failed to influence IFN-γ–induced pIgR transcriptional activity and mRNA expression (Fig. 5E, 5F, respectively). In addition, the activity of BAY 11-7082 was also confirmed by measuring NF-κB activation in response to IFN-γ (Fig. 5G). Therefore, it was concluded that attenuation of pIgR expression by ribosome inactivation was not due to suppression of NF-κB.

FIGURE 5.

Involvement of IFN-γ–induced NF-κB in pIgR expression in human enterocytes. (A) HCT-8 cells were treated with vehicle, 500 ng/ml DON, or 50 ng/ml ANS in the presence of 20 ng/ml IFN-γ for 2 h. Cellular lysate was subjected to Western blot analysis. *p < 0.05, significant difference from the only IFN-γ–treated group. (B) HCT-8 cells were treated with vehicle, 500 ng/ml DON, or 50 ng/ml ANS in the presence of 20 ng/ml IFN-γ for 2 h. The cells were fixed, immunostained, and visualized with a confocal microscope (original magnification ×1800). The graphs on the right represent the relative density of the nuclear localization of p65 visualized with the confocal microscopic. *p < 0.05, significant difference from the only IFN-γ–treated group. (C) Maps of reporter constructs containing the wild-type and mutant human pIgR promoters from −2.7 kb to +5.9 kb. (D) HCT-8 cells transfected with pIgR promoter–linked reporter plasmid were treated with vehicle or 20 ng/ml IFN-γ 24 h prior to the luciferase assay. *p < 0.05, significant difference from each vehicle control group. (E) HCT-8 cells transfected with pSC1 pIgR reporter plasmid and then treated with vehicle, 10 μM BAY11-7082 (BAY), or 20 ng/ml IFN-γ 24 h prior to the luciferase assay. *p < 0.05, significant difference from the only vehicle or BAY–treated group. (F and G) HCT-8 cells were pretreated with or without 10 μM BAY11-7082 for 2 h and added with 20 ng/ml IFN-γ for 48 h (F) or 2 h (G). The mRNA was analyzed by real-time RT-PCR (F) and cellular lysate was subjected to Western blot analysis (G). *p < 0.05, significant difference from the only vehicle or BAY11-7082–treated group.

FIGURE 5.

Involvement of IFN-γ–induced NF-κB in pIgR expression in human enterocytes. (A) HCT-8 cells were treated with vehicle, 500 ng/ml DON, or 50 ng/ml ANS in the presence of 20 ng/ml IFN-γ for 2 h. Cellular lysate was subjected to Western blot analysis. *p < 0.05, significant difference from the only IFN-γ–treated group. (B) HCT-8 cells were treated with vehicle, 500 ng/ml DON, or 50 ng/ml ANS in the presence of 20 ng/ml IFN-γ for 2 h. The cells were fixed, immunostained, and visualized with a confocal microscope (original magnification ×1800). The graphs on the right represent the relative density of the nuclear localization of p65 visualized with the confocal microscopic. *p < 0.05, significant difference from the only IFN-γ–treated group. (C) Maps of reporter constructs containing the wild-type and mutant human pIgR promoters from −2.7 kb to +5.9 kb. (D) HCT-8 cells transfected with pIgR promoter–linked reporter plasmid were treated with vehicle or 20 ng/ml IFN-γ 24 h prior to the luciferase assay. *p < 0.05, significant difference from each vehicle control group. (E) HCT-8 cells transfected with pSC1 pIgR reporter plasmid and then treated with vehicle, 10 μM BAY11-7082 (BAY), or 20 ng/ml IFN-γ 24 h prior to the luciferase assay. *p < 0.05, significant difference from the only vehicle or BAY–treated group. (F and G) HCT-8 cells were pretreated with or without 10 μM BAY11-7082 for 2 h and added with 20 ng/ml IFN-γ for 48 h (F) or 2 h (G). The mRNA was analyzed by real-time RT-PCR (F) and cellular lysate was subjected to Western blot analysis (G). *p < 0.05, significant difference from the only vehicle or BAY11-7082–treated group.

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Because transcriptional regulation via IRF-1 or NF-κB cannot account for suppression of pIgR expression by ribosome inactivation, we assessed the effects of ribosome inactivation on posttranscriptional regulation of pIgR mRNA stability. In contrast with the enhancing effects of ribosome inactivation on IRF-1 mRNA stability at the early time points (Fig. 4H–J), ribosome inactivation significantly decreased pIgR mRNA stability under IFN-γ–stimulated conditions at the late time points (Fig. 6A). Moreover, the RNA-binding factor HuR, which was positively involved in stabilizing IRF-1 transcripts, was also assessed for its role in stability of pIgR mRNA in enterocytes exposed to ribosome inactivation. Overexpressed HuR significantly reversed ribosome inactivation–mediated suppression of pIgR expression in HCT-8 human enterocytes (Fig. 6B). However, the translocation of HuR to the cytoplasm that occurred in response to ribosomal stress, and that stabilized IRF-1 mRNA, would not account for destabilization of pIgR transcripts by the same stress. To provide evidence of possible differential regulation of HuR translocation by ribosome inactivation at different times of stress exposure, we examined the cellular localization of the HuR protein with different times of exposure to chemical ribosome inactivators. Whereas the short time (1 h) exposure enhanced IFN-γ–triggered cytosolic translocation of HuR protein, extended exposure for 12 h led to nuclear entrapment of HuR (Fig. 6C, 6D). This restriction of translocation of HuR to the cytoplasm upon prolonged exposure to ribosomal stress could account for the reduced stability of pIgR mRNA. Moreover, IFN-γ significantly increased binding of HuR proteins to pIgR transcript, which were completely inhibited by ribosome inactivation (Fig. 6E). Taken together, ribosomal inactivation suppressed IFN-γ–induced pIgR expression because of the limited availability of cytoplasmic HuR protein. In contrast, HuR released immediately after ribosome inactivation contributed to superinduction of IRF-1 and subsequent expression of proinflammatory mediators that would result in ribosomal stress-induced epithelial pathogenesis (Fig. 7).

FIGURE 6.

Effect of ribosome inactivation on pIgR mRNA stability. (A) HCT-8 cells pre-exposed to vehicle, or 500 ng/ml DON for 12 h, were treated with 20 ng/ml IFN-γ for 48 h. Cellular transcription was terminated by adding 5 μM actinomycin D, and cellular RNA was extracted at each indicated time. Each mRNA was measured using RT-PCR. *p < 0.05, significant difference for the IFN-γ treatment group. (B) Empty vector or sense HuR expression plasmid–transfected HCT-8 cells were treated with vehicle, 500 ng/ml DON, or 50 ng/ml ANS in the presence of 20 ng/ml IFN-γ for 48 h. *p < 0.05, significant difference from the empty vector–transfected group. Blots in the dashed box represent data for proteins. (C) HCT-8 cells were treated with the vehicle, 500 ng/ml DON, or 50 ng/ml ANS in the presence of 20 ng/ml IFN-γ for 1 or 12 h. Cytosolic or nuclear fractions of cellular lysates were subjected to Western blot analysis. (D) HCT-8 cells were treated with the vehicle, 500 ng/ml DON, or 50 ng/ml ANS in the presence of 20 ng/ml IFN-γ for 1 or 12 h. Cells were fixed, immunostained, and visualized under the confocal microscope (original magnification ×1800). Red dots indicate the nuclear border identified by DAPI staining to quantify relative amounts of cytosolic HuR. *p < 0.05, significant difference from the only IFN-γ–treated group. (E) RNA immunoprecipitation assays were performed to measure HuR protein bound to the pIgR transcript in cells. HCT-8 cells were treated with 20 ng/ml IFN-γ (I) in the presence or absence of 500 ng/ml DON (D) for 48 h. Immunoprecipitated transcript was measured using real-time RT-PCR. *p < 0.05, significant difference from the control group.

FIGURE 6.

Effect of ribosome inactivation on pIgR mRNA stability. (A) HCT-8 cells pre-exposed to vehicle, or 500 ng/ml DON for 12 h, were treated with 20 ng/ml IFN-γ for 48 h. Cellular transcription was terminated by adding 5 μM actinomycin D, and cellular RNA was extracted at each indicated time. Each mRNA was measured using RT-PCR. *p < 0.05, significant difference for the IFN-γ treatment group. (B) Empty vector or sense HuR expression plasmid–transfected HCT-8 cells were treated with vehicle, 500 ng/ml DON, or 50 ng/ml ANS in the presence of 20 ng/ml IFN-γ for 48 h. *p < 0.05, significant difference from the empty vector–transfected group. Blots in the dashed box represent data for proteins. (C) HCT-8 cells were treated with the vehicle, 500 ng/ml DON, or 50 ng/ml ANS in the presence of 20 ng/ml IFN-γ for 1 or 12 h. Cytosolic or nuclear fractions of cellular lysates were subjected to Western blot analysis. (D) HCT-8 cells were treated with the vehicle, 500 ng/ml DON, or 50 ng/ml ANS in the presence of 20 ng/ml IFN-γ for 1 or 12 h. Cells were fixed, immunostained, and visualized under the confocal microscope (original magnification ×1800). Red dots indicate the nuclear border identified by DAPI staining to quantify relative amounts of cytosolic HuR. *p < 0.05, significant difference from the only IFN-γ–treated group. (E) RNA immunoprecipitation assays were performed to measure HuR protein bound to the pIgR transcript in cells. HCT-8 cells were treated with 20 ng/ml IFN-γ (I) in the presence or absence of 500 ng/ml DON (D) for 48 h. Immunoprecipitated transcript was measured using real-time RT-PCR. *p < 0.05, significant difference from the control group.

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FIGURE 7.

A putative scheme for the mechanism of ribosome inactivation–mediated mucosal immune disorders in the human mucosal epithelium. Ribosome inactivation alters HuR protein translocation, which can lead to serious mucosal disorder. In the early phase, ribosome inactivation triggers cytosolic translocation of HuR protein. Enhanced cytosolic HuR mediates IRF-1 mRNA stabilization and subsequent induction of target genes, such as IL-7, that establish excessive mucosal inflammation. In late phase, ribosome inactivation causes the entrapment of HuR protein in the nucleus. Deficiency in the pool of cytosolic HuR protein leads to decreased stabilization of pIgR mRNA and subsequent failure of IgA translocation to the intestinal lumen, which is associated with disruption of initial immune defenses against luminal Ags, including enteropathogens. Disruption of IgA mucosal secretion and excessive expression of proinflammatory responses via IRF-1 are thus expected to be critical steps for ribosome inactivation–mediated mucosal immune disorders, such as IBD.

FIGURE 7.

A putative scheme for the mechanism of ribosome inactivation–mediated mucosal immune disorders in the human mucosal epithelium. Ribosome inactivation alters HuR protein translocation, which can lead to serious mucosal disorder. In the early phase, ribosome inactivation triggers cytosolic translocation of HuR protein. Enhanced cytosolic HuR mediates IRF-1 mRNA stabilization and subsequent induction of target genes, such as IL-7, that establish excessive mucosal inflammation. In late phase, ribosome inactivation causes the entrapment of HuR protein in the nucleus. Deficiency in the pool of cytosolic HuR protein leads to decreased stabilization of pIgR mRNA and subsequent failure of IgA translocation to the intestinal lumen, which is associated with disruption of initial immune defenses against luminal Ags, including enteropathogens. Disruption of IgA mucosal secretion and excessive expression of proinflammatory responses via IRF-1 are thus expected to be critical steps for ribosome inactivation–mediated mucosal immune disorders, such as IBD.

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Epithelial pIgR plays a critical role in the barrier defense of the intestinal epithelium. Ribosome inactivation suppressed pIgR expression and caused defects in mucosal IgA transport. This led to IgA accumulation in the villus lamina propria and subsequent disruption of mucosal defense against enteropathogens, such as EPEC. However, ribosome inactivation also triggered superinduction of IRF-1, a key positive regulator of pIgR, by enhancing stability of IRF-1 mRNA and destabilizing pIgR mRNA. The biphasic localization of HuR in response to ribosome inactivation resulted in paradoxical patterns of expression of the two genes. Although early-response cytoplasm-translocated HuR contributed to IRF superinduction, suppression of pIgR expression was mostly a result of eventual entrapment of HuR protein in the nucleus, which prevented stabilization of the pIgR transcripts via binding to AREs in 3′-UTR. Although IRF-1 did not contribute to regulation of pIgR by ribosome inactivation, IRF-1 has been known to extensively facilitate the progression of intestinal inflammatory diseases (51, 52). In particular, IRF-1–induced IL-7 has been shown to activate the mucosal immune responses that would lead to the development of chronic intestinal inflammation (53, 54). As a result, the epithelial stresses that cause ribosome inactivation would exacerbate inflammatory responses via activation of IRF-1–linked signals and disruption of microbial defenses that result from pIgR depletion. This might mechanistically account for mucosal pathogenesis and implicate ribosome inactivation as an etiological factor of environmental IBD. Increasing evidence for a link between pIgR deficiency and colitis is being reported (55). Epithelial pIgR is involved in maintaining the barrier defense against luminal Ags and microbes by preventing excessive activation of innate immunity and subsequent tissue injuries in murine experimental colitis models (8, 55). In these models, a deficiency in pIgR triggers macrophage infiltration in response to greater penetration of luminal bacteria to the lamina propria. Therefore, attenuated pIgR expression resulting from ribosome inactivation may lead to similar patterns of barrier-disrupted inflammation and microbial translocation as are seen in mice with pIgR deficiency.

IRF-1 is a potent positive modulator in the pathogenesis of IBD because of its ability to upregulate proinflammatory mediators, including IL-6, IL-15, and IL-1β–converting enzyme (56). Moreover, IRF-1 facilitates the differentiation of naive CD4+ T cells into Th1 cells that play a role in Crohn’s disease (52). Of the IRF-1–dependent genes, IL-7 is critical for maintenance of chronic colitis by enhancing colitogenic CD4+ effector memory cells (57) and promoting chronic colitis in humans with IBD-linked histopathological characteristics (58). In the current study, ribosome inactivation potentiated IFN-γ–induced IRF-1 and subsequent IL-7 expression, which would facilitate ribosome inactivation-triggered IBD-like pathogenesis. Mechanistically, it is well known that IRF-1 is an important regulator of epithelial pIgR because of its ability to bind to ISREs in the pIgR promoter following stimulation with IFN-γ (14). However, IRF-1 was not involved in ribosome inactivation–mediated pIgR suppression in the current study. Instead, we hypothesized that ribosome inactivation limited pIgR expression via the IRF-1 transcriptional repression at ISREs. We thus evaluated IRF-2, a representative ISRE-binding transcriptional repressor of IRF-1 (59). Although IRF-2 was enhanced by ribosome inactivation in the presence of IFN-γ, suppression of IRF-2 failed to restore the pIgR suppression resulting from ribosome inactivation (data not shown). As another IRF-1 partner, NF-κB also plays important roles in pIgR transcription (60), and NF-κB binding to intron 1 can cooperate with IRF-1 binding to exon 1 in the pIgR gene transcript (16). Because ribosome inactivation attenuated IFN-γ–mediated NF-κB phosphorylation and nuclear translocation, it was expected that NF-κB suppression by ribosome inactivation would lead to pIgR suppression in the current study. However, NF-κB and IRF-1 were marginally involved in transcriptional suppression of pIgR by ribosomal inactivation. Rather than transcriptional regulation mediated by IRF-1 or NF-κB, posttranscriptional regulation via HuR played a critical role in pIgR expression in enterocytes exposed to ribosome inactivators. According to our previous studies (32, 50), early exposure to the ribosomal inactivation also enhanced cytosolic translocation of HuR protein which was involved in productions of chemokines and BAFF in enterocytes. Because the induction of IRF-1 expression required a time span as short as 1–2 h, ribosome inactivation–triggered HuR protein in the cytoplasm contributed to the stabilization of IRF-1 mRNA. This was consistent with our previous reports that showed that ribosome inactivation enhances early-inducible proinflammatory cytokines via the HuR protein, which was translocated to the cytoplasm via protein kinase C (32, 48). However, long-term mucosal ribosome inactivation leads to the nuclear entrapment of HuR. To speculate the molecular mechanism of nuclear entrapment of HuR under ribosomal inactivation, we confirmed the known interaction between HuR and nuclear ATF3 in the present model (Supplemental Fig. 1), based on our previous report (50). ATF3 as a transcription factor was induced and translocated into nuclei by ribosomal inactivation (Supplemental Fig. 1A). Exogenously introduced or ribosomal stress–induced ATF3 strongly interacted with HuR protein in the nuclei, which led to nuclear entrapment of HuR protein (Supplemental Fig. 1B, 1C) and partly contributed to the suppression of pIgR expression (Supplemental Fig. 1D). Therefore, it can be implicated that ATF3 may downregulate pIgR expression to some degree by limiting cytosolic availability of HuR protein as a pIgR mRNA–stabilizing protein in the current study. This mechanistic implication is in agreement with our previous report on regulation of Nod2-activated proinflammatory gene expression via association between ATF3 and HuR (50).

Because epithelial pIgR protects against initial infections by transporting dimeric IgA and pentameric IgM (3), IgA-deficient hosts are more susceptible to infection and subsequent stimulation of the innate immune system (61). Moreover, pIgR expression is also commonly increased by proinflammatory cytokines in response to viral or bacterial infections, thus linking innate and adaptive immunity. However, abnormal high levels of epithelial pIgR or the cleaved extracellular domain of pIgR in the sera have been associated with bad prognosis in patients with hepatocellular carcinoma and colonic carcinoma with liver metastasis (62). Mechanistically, overexpression of pIgR plays a role in induction of the epithelial–mesenchymal transition and metastasis through activation of Smad signaling (62). Therefore, interfering with the action or expression of pIgR via targeting IRF-1– and Smad-linked signals could be beneficial for pIgR-high cancer patients. In this way, ribosome inactivation–mediated pIgR suppression potentially provides promising insights into epithelial cancer treatment. However, because ribosomal inactivation can trigger the acute mucosal inflammatory responses, its chronic insults may exacerbate the inflammation-associated cancers, and thus, the specific intervention with cancer pIgR without activation of inflammatory signals needs to be developed in the future study. Moreover, although downregulation of pIgR could be effective against some epithelial cancers, it is hard to conclude that the chronic suppression of pIgR is beneficial against all malignant diseases. For instance, a prolonged reduction in the pool of epithelial pIgR restricts mucosal secretion of IgA, which subsequently result in an elevation of circulatory IgA and subsequent deposition of IgA immune complexes in the renal mesangium, which could be a crucial etiological factor of IgA nephritis. Several mycotoxins that trigger ribosome inactivation induce murine IgA nephritis, which is very similar to human Berger’s disease, the most prevalent type of human chronic nephritis in the world (31, 63). Therefore, a restoration of pIgR levels and normal secretion of mucosal IgA would be important for intervening with pathogenesis of IgA nephritis although prolonged overexpression of pIgR has been associated with tumor progression.

Taken in sum, the results of this study account for the bimodal effects of ribosome inactivation on intestinal epithelium integrity. Early cytosolic translocation of HuR in response to ribosome inactivation facilitated stabilization of IRF-1 transcripts and induction of IRF-1–targeted immune modulators including proinflammatory IL-7 in enterocytes, even though IRF-1 was not involved in pIgR suppression by ribosome inactivation. However, prolonged epithelial insult by ribosome inactivation suppressed mucosal IgA transport via attenuating expression of pIgR, which was due to the hindered cytosolic translocation of HuR and subsequently decreased stabilization of pIgR transcript in the enterocytes.

This work was supported by Basic Science Research Program through National Research Foundation of Korea Grant NRF-2015R1A2A1A15056056 funded by the Ministry of Science, Information & Communication Technology and Future Planning.

The online version of this article contains supplemental material.

Abbreviations used in this article:

ANS

anisomycin

ARE

adenylate-uridylate–rich element

ATF3

activating transcription factor 3

DON

deoxynivalenol

EPEC

enteropathogenic Escherichia coli

HuR

human antigen R

IBD

inflammatory bowel disease

IRF-1

IFN regulatory factor-1

ISRE

IFN-stimulated response element

NIV

nivalenol

pIgR

polymeric IgR

shRNA

short hairpin RNA.

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The authors have no financial conflicts of interest.

Supplementary data