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Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

The secreted thiol-activated cytolysin listeriolysin O (LLO) was responsible for L. monocytogenes-induced high-molecular glycoproteins (HMGs) exocytosis in cultured human mucosecreting HT29-MTX cells. By biochemical analysis we demonstrate that the majority of secreted HMGs in LLO-stimulated cells are of mucin origin. In parallel, analysis of the expression of MUCs genes showed that the transcription of the MUC3, MUC4 and MUC12 genes encoding for membrane-bound mucins was increased in LLO-stimulated cells. Upregulation of the MUC3 gene correlates with an increased expression of the membrane-bound MUC3 mucin. In contrast, increase in secretion of the gel-forming MUC5AC mucin develops without upregulation of the MUC5AC gene. Finally, results showed that NF-κB and AP-1 transcription factors were not involved in LLO-induced upregulation of MUCs genes in HT29-MTX cells, whereas L. monocytogenes infection was able to promote the degradation of IκB proteins in the cells.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

In the human small intestinal mucosa, the mucus layer is the first obstacle encountered by enteric pathogens. The mucus coating at the surface results from the secretion of mucins by the specialized goblet cell (Neutra and Forstner, 1987). Inoculation of L. monocytogenes into the rat ligated ileal loops was associated with the massive release of mucus by goblet cells (Pron et al., 1998). Consistent with this in vivo observation, we have recently reported that L. monocytogenes infection of cultured human mucin-secreting intestinal cells is followed by an increase in mucin secretion through an listeriolysin (LLO)-dependent mechanism (Coconnier et al., 1998; Coconnier et al., 2000). Apical infection of the cultured human polarized mucosecreting HT29-MTX cells by the Gram-positive facultative intracellular human pathogen Listeria monocytogenes is followed by the stimulation of mucus exocytosis without cell-entry (Coconnier et al., 1998; 2000). Using a set of isogenic mutants and purified LLO, the L. monocytogenes thiol-activated exotoxin LLO has been identified as agonist of the mucus secretion. Listeriolysin O (Geoffroy et al., 1987; Cossart et al., 1989), encoded by the gene hly (Mengaud et al., 1987;1988), is an essential virulence factor for the L. monocytogenes intracellular lifestyle: it mediates the lysis of the phagosomal membrane formed after bacterial entry, thus allowing bacterial access to the cytosol. Moreover, LLO in conjunction with phosphatidylinositol-specific phospholipase C and broad-range phospholipase C, encoded by plcA and plcB, respectively, allows lysis of the double-membrane vacuole formed during the bacterial cell-to-cell passage allowing propagation of infection (Goldfine et al., 1995). When examining the mechanism by which LLO induces mucin exocytosis, we have demonstrated that this cellular response develops without any relationship with the known pore forming activity of the toxin (Coconnier et al., 2000). The LLO-induced mucin exocytosis required a functional microtubules network, suggesting that LLO stimulates the steady vesicular constitutive pathway of mucin exocytosis. Moreover, the LLO-induced mucin exocytosis is related to the binding of the toxin to multiple membrane-associated lipid receptors allowing the oligomerization of the toxin monomers. Finally, demonstration has been provided that the membrane-bound LLO is internalized through detergent insoluble glycolipid microdomains (caveolae) containing VIP/21 caveolin. The low-efficient caveolae-dependent pathway of endocytosis, in comparison to the clathrin-dependent pathway of endocytosis, allows a small quantity of the membrane-bound LLO to enter the polarized epithelial intestinal cells.

The present study was conducted in order to gain further insights into the mechanism by which LLO promotes the increase in mucus exocytosis in human mucin-secreting cells. Inducibility of the genes encoding for the secreted and membrane-bound human mucins was examined. Because it has been recently established that LLO elicits a NF-κB-dependent expression of pro-inflammatory genes in intestinal epithelial cells, we examined whether or not a NF-κB-dependent mechanism is involved in the LLO-induced mucin genes activation.

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

The high-molecular glycoproteins secreted in LLO-stimulated human polarized intestinal mucin-secreting HT29-MTX cells are mucins

We examine the HMGs exocytosis in three cultured human polarized intestinal cells, the HT29-MTX, HT29-Cl.16E and Caco-2/TC7 cells, which have different intestinal cell types and produce mucins, proteoglycans (PG) or both. The homogenous mucin-secreting population HT29-MTX (Lesuffleur et al., 1990) secreted HMGs, which are both PGs or mucins (Huet et al., 1995; Molist et al., 1998). It has been recently established that the enterocyte-like Caco-2 cells secreted PGs (Salmivirta et al., 1998), but although expressing the MUC1, MUC3, MUC4 and MUC5AC genes, these cells failed to secrete mucins (van Klinken et al., 1996). As shown in Fig. 1, LLO stimulation of mucin- and PG-secreting HT29-MTX and HT29-Cl.16E cells results in an significant increase (3.1-fold and 2.7-fold respectively) in the secretion of metabolically cells radiolabelled HMGs. The LLO-stimulation in PG-secreting Caco-2/TC7 results in a moderate increase (1.6-fold) in HMG secretion.

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Figure 1. Apically secreted [3H]-HMGs in control unstimulated and LLO-stimulated mucin-secreting HT29-MTX and HT29-Cl.16E and enterocyte-like Caco-2/TC7 cells. Endogenous HMGs were metabolically radiolabelled with d-6-[3H]glucosamine (18 h of labelling at 37°C in a 10% CO2/90% air atmosphere). For LLO cell stimulation, cells were washed and incubated 2 h with no addition; or LLO (90 µg/well applied apically in DMEM without FCS). HMGs secretion was measured in the apical compartment during a period of 2 h. Secreted radiolabelled HMGs were determined as described in Experimental procedures. Each value shown is the mean of three experiments (three successive passages of cultured cells) ± standard error. Statistical analysis was performed with a Student's t-test. In LLO-stimulated HT29-MTX and HT29-Cl.16E cells, P <0.01 compared with control unstimulated cells. In LLO-stimulated Caco-2/TC7 cells, no significant difference compared with control unstimulated cells.

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The results reported above prompted us to conduct a biochemical characterization of the HMGs secreted upon LLO stimulation in HT29-MTX cells. The secreted and cellular HMGs in control unstimulated and LLO-stimulated cells were isolated by ultracentrifugation through caesium bromide (CsBr) gradient and were analysed for amino acid composition, sugar analysis and sulphate content. As reported in Table 1, no change in amino acid and sugar compositions, and sulphate contents was found when intracellular HMGs stores within control unstimulated and LLO-stimulated cells were compared. In contrast, compared with control unstimulated cells, the culture medium of LLO-stimulated cells had higher relative Ala, Gly and Pro contents and lower relative Phe, His, Ile, Leu, Ser and Tyr contents. However, no change in sugar composition and sulphate contents was found when culture medium of control unstimulated and LLO-stimulated cells were compared. Both the control unstimulated and LLO-stimulated HT29-MTX cells show a Thr/Ser ratio of 1.48 and 1.34, respectively, a typical feature of mucins (Huet et al., 1995). In contrast, both the culture medium of control unstimulated and LLO-stimulated HT29-MTX cells had a lower Thr/Ser ratio (0.66 and 0.81 respectively). Surprisingly, a high proportion of Gly was found in the culture medium of control cells and increase in LLO-stimulated cells. This amino acid is not characteristic of mucins but it was commonly present in another class of HMGs, the PGs. Altogether, results of the biochemical analysis of secreted and cellular HMGs in control unstimulated and LLO-stimulated cells suggest the presence of both mucins and PGs.

Table 1.   Amino acid and sugar composition, and sulphate content in control unstimulated and LLO-stimulated HT29-MTX cells
 Culture mediumCells
LLOControlLLO
  1. Amino acid are expressed as percentage; sugar and sulphate are expressed as molar ratios to GalNac. ND: not detectable.

Ala 6.1210.15 6.04 6.72
Asp + Asn 8.14 6.15 4.16 3.64
Glu + Gln13.46 9.89 9.4410.16
Phe 2.28 1.19 1.91 2.09
Gly17.5330.31 9.4711.39
His 1.76 1.041.92 1.84
Ile 3.28 1.81 2.91 2.82
Lys 4.15 2.91 3.2 3.57
Leu 6.16 2.84 4.4 4.57
MetND 0.35 0.4 0.67
Pro 5.4512.8512.6 9.89
Arg 4.27 4.06 4.16 3.64
Ser11.88 6.4811.7812.58
Thr 7.81 5.2517.416.9
Val 4.73 3.64 5.46 4.97
Tyr 2.98 1.07 2.13 1.36
Fucose 0 0 0 0
Mannose 0.2 0.2NDND
Galactose 0.9 1.2 0.9 0.9
Glucose 0.3 0.3 0.1 0.1
GalNac 1 1 1 1
GlucNac 0.4 0.6 0.3 0.3
NANA 0.5 0.7 0.6 0.6
Sulphates 1.3 0.8NDND

A biochemical analysis of the PGs was conducted by cellulose acetate electrophoresis followed by DMB staining. As shown in Fig. 2A, results reveal that PGs are present in the cell lysate and culture medium of both control unstimulated and LLO-stimulated HT29-MTX cells. Quantification of the bands by scanning reveals that there is no significant difference between control unstimulated and LLO-stimulated HT29-MTX cells (data not shown). Interestingly, the bands found in control and LLO-stimulated cells migrate as the heparan sulphate. Confirmation that heparan sulphate are present is provided by an experiment conducted with heparinase III treatment, which specifically hydrolyses the glycosidic linkage present in heparan sulphate. Both in control heparan sulphate and LLO-stimulated cells, the enzymatic treatment leads to the disappearance of the bands. In order to determine the level of PGs in the culture medium of LLO-stimulated cells, we conduct experiment with heparinase III and chondroitinase ABC, which specifically removes chondroitin sulphate and dermatan sulphate. As disclosed in Fig. 2B, these enzymatic treatments did not affect significantly the level of metabolically radiolabelled HMGs secreted by the LLO-stimulated HT29-MTX cells, suggesting that PGs are a minor part of the HGMs secreted.

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Figure 2. Proteoglycan (PG) analysis of the HMGs secreted in LLO-stimulated HT29-MTX cells.

A. PG analysis by cellulose acetate electrophoresis and DMB staining in cell lysate and culture medium of 2 h LLO-stimulated HT29-MTX cells. Heparan sulphate, dermatan sulphate and chondroitin sulphate migrated as controls. Heparinase III treatment: 0.5 IU per ml in PBS, 37°C, 45 min. A band migrating as heparan sulphate and sensitive to heparinase III treatment was found in cell lysate and supernatant medium of control and LLO-stimulated cells. Micrograph is representative of three independent experiments.

B. The effect of glycosaminoglycan lyase treatments on the [3H]-HMGs secreted into the culture medium of LLO-stimulated HT29-MTX cells. The 3H-labelled HMGs-containing medium of LLO-stimulated HT29-MTX cells was treated with heparinase III and chondroitinase ABC (0.5–1 IU per ml in PBS, 37°C, 45 min) and then analysed for HMGs (described in Experimental procedures). Each value shown is the mean of three experiments (three successive passages of cultured cells) ± standard error. Statistical analysis was performed with a Student's t-test. In heparinase- and chondroitinase-treated LLO-stimulated HT29-MTX cells, no significant difference was found compared with untreated LLO-stimulated HT29-MTX cells.

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The MUC5AC mucin was one of the secreted mucins found in HT29-MTX cells (Hennebicq-Reig et al., 1998). We determine by indirect immunolabelling and radioimmunoassay the level of the MUC5AC mucin secreted upon LLO stimulation in HT29-MTX and HT29-Cl.16E cells. Expression of MUC5AC protein in LLO-stimulated HT29-MTX cells was examined by indirect immunofluorescence labelling with the anti-MUC5AC 1-13M1 mAb, followed by a confocal laser scanning microscopy (CLSM) analysis. As observed in Fig. 3A, an increase in positive MUC5AC immunoreactivity was found at the cell surface of the LLO-stimulated cells as compared with control unstimulated cells. Determination of the level of secreted MUC5AC mucin by radioimmunoassay in the culture medium of control unstimulated and LLO-stimulated cells shows a significant increase in secretion of MUC5AC mucin in LLO-stimulated HT29-MTX and HT29-Cl.16E cells (Fig. 3B). Although the Caco-2/TC7 cells did not produce mucins, we analyse the presence of MUC5AC mucin in the culture medium of the cells stimulated by LLO and observe the absence of this mucin. Altogether, these results are in favour with a major presence of mucins in the culture medium of LLO-stimulated HT29-MTX cells.

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Figure 3. Expression of MUC5AC mucin in control unstimulated and LLO-stimulated mucin-secreting HT29-MTX and HT29-Cl.16E and enterocyte-like Caco-2/TC7 cells.

A. Indirect immunofluorescence labelling of MUC5AC using I-13M1 mAb in control (A) and LLO-stimulated (90 µg/well applied apically in DMEM without FCS, at 37°C) (B) HT29-MTX cells. Cells were fixed with 3.5% paraformaldehyde, washed, permeabilized with Triton X-100, and processed for indirect immunofluorescence labelling as described in the Experimental procedures. Reconstruction of successive En face micrographs obtained by CLSM analysis (horizontal x–y optical sections, one section every 1 µm) shows an increase in MUC5AC expression in LLO-stimulated cells, compared with control unstimulated cells. (Magnification ×  100). Micrographs are representative of three independent experiments.

B. Culture media of control unstimulated and LLO-stimulated cells (90 µg/well applied apically in DMEM without FCS, during 2 h at 37°C) were analysed for determination of MUC5AC mucin, quantitated by immunoradiometric assay using the anti-MUC5AC 1–13M1 and PM7 mAbs. Each value shown is the mean of three experiments (three successive passages of cultured cells) ± standard error. Statistical analysis was performed with a Student's t-test. In LLO-stimulated HT29-MTX and HT29-Cl.16E cells, p < 0.01 compared with control unstimulated cells. In LLO-stimulated Caco-2/TC7 cells, no significant difference compared with control unstimulated cells.

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Upregulation of mucin genes transcription in LLO-stimulated HT29-MTX cells

It has been recently reported that exoproducts of Pseudomonas aeruginosa are able to promote the upregulation of MUC2 and MUC5AC genes transcription (Dohrman et al., 1998). Previous results (Coconnier et al., 1998; 2000) and the results reported above led us to examine whether or not LLO promotes MUC genes activation in HT29-MTX cells.

We conducted an analysis of the expression of MUC1-4, MUC5AC, MUC5B, MUC6, MUC7, MUC11, and MUC12 genes in control and LLO-stimulated HT29-MTX cells at different time points post stimulation. Results reported in Fig. 4A show that the MUC1, MUC2, MUC3, MUC4, and MUC5AC mRNAs were present in HT29-MTX cells, as previously observed (Lesuffleur et al., 1993). In contrast, no expression of MUC6 and MUC7 mRNAs was observed. We now report that HT29-MTX cells highly expressed the MUC11 mRNA, whereas the MUC12 mRNA was present at a very low level. As shown in Fig. 4A, in cells stimulated during 2 h and 24 h by LLO the transcription of the MUC3, MUC4 and MUC12 genes was increased compared with control, unstimulated cells. In contrast, no change in the transcription of MUC1, MUC2, MUC5B, MUC5AC, MUC6, MUC7, and MUC11 genes upon LLO-stimulation was observed after both 2 h and 24 h of exposure (Fig. 4B). A long exposure of HT29-MTX cells to LLO during 3 and 7 days shows that the upregulation of the MUC3 gene transcription remained stable, whereas the upregulation of the MUC4 and MUC12 gene transcription was transient (data not shown).

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Figure 4. Analysis of mucin mRNAs expression in control and LLO-stimulated HT29-MTX.

A. For each mucin mRNA: Lane 1, control; Lane 2, 2 h LLO-stimulated HT29-MTX cells (90 µg/well applied apically in DMEM without FCS, at 37°C); Lane 3, 24 h LLO-stimulated HT29-MTX cells.

B. PCR products were quantified with the Kodak Digital Science 1D analysis software, relative to a GADPH cDNA amplification control. Each value shown is the mean of three experiments (three successive passages of cultured cells) ± standard error. Statistical analysis was performed with a Student's t-test. In LLO-stimulated HT29-MTX cells, p <0.01 for MUC3 at 2 h and 24 h of stimulation, compared with control unstimulated cells. For MUC4 and MUC12, p <0.01 at 2 h of stimulation, and no significant difference at 24 h of stimulation, compared with control unstimulated cells.

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Whether or not the upregulation of MUC genes encoding for membrane-bound mucins was accompanied by a change in apical expression of MUC proteins in LLO-stimulated HT29-MTX cells was examined. We chose to examine the apical expression of MUC3 protein using a rabbit polyclonal anti-MUC3 antibody. Examination of MUC3 distribution in permeabilized LLO-stimulated cells by indirect immunofluorescence labelling followed by a confocal laser scanning microscopy (CLSM) analysis reveals that stimulation is accompanied by a dramatic increase in the apical expression of the MUC3 mucin as compared to control unstimulated cells (Fig. 5).

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Figure 5. Confocal laser scanning microscopy analysis of MUC3 in control and LLO- stimulated HT29-MTX cells.

A, C and E, control unstimulated cells. B, D and F, LLO-stimulated HT29-MTX cells (90 µg/well applied apically in DMEM without FCS, at 37°C). Before MUC3 immunolabelling, mucins were deglycosylated (3 min, 37°C, neuraminidase 0.05 U). Cells were fixed with 3.5% methanol-acetone, washed, permeabilized, and processed for indirect immunofluorescence labelling as described in the Experimental procedures. In A, B, C and D, reconstruction of successive En face micrographs obtained by CLSM analysis (horizontal x–y optical sections, one section every 1 µm). In A and C, micrographs show the detection of membrane-bound MUC3 present within the apical cell membrane of control unstimulated cells. The labelling of MUC3 is non-homogeneous among the cells. In B and D, micrographs show an increase in expression of MUC3 in LLO-stimulated cells. (Magnifications × 40 in A and B and ×63 in C and D). In E and F, successive En face micrographs obtained in permeabilized control unstimulated and LLO-stimulated cells by CLSM analysis (horizontal x–y optical sections, one section every 1 µm) were reconstructed in lateral views (vertical x–z) by integration of images gathered at a step position of 1 on the x–y axis. Lateral views allowed for the observation of the distribution of MUC3 within the cells. In control unstimulated cells (E), the tight band of MUC3 labelling is localized at the apical domain of the cells. In LLO-stimulated cells (F), the MUC3 labelling is dramatically modified showing an enlargement of the band. Hatched line localizes the basal domain of the cell monolayer. (Magnification × 63 in E and F).

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LLO-induced mucin exocytosis is a NF-κΒ- and AP1-independent event

Analysing the mechanism by which P. aeruginosa promotes the upregulation of several MUC genes, Li et al. (1998) recently demonstrated that a signalling pathway leads to activation of the nuclear factor NF-κB. Moreover, it has been recently reported that LLO was able to promote the upregulation of chemokines in endothelial cells by activation of the nuclear factor NF-κB (Kayal et al., 1999; Rose et al., 2001). We examine whether or not the nuclear factor NF-κB plays a role in the LLO-induced mucin exocytosis. For this purpose, we chose to examine the chemokine IL-8 secretion, the IκB degradation and the level of c-Fos in HT29-MTX cells subjected to LLO (Fig. 6).

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Figure 6. Secretion of pro-inflammatory IL-8 chemokine, and immunoblot analysis of IκB proteins degradation and c-Fos synthesis in control unstimulated, LLO-stimulated, and L. monocytogenes- and Salmonella-infected HT29-MTX cells.

A. Secreted IL-8 determined by immunoassay in control cells, Salmonella-infected and LLO-stimulated HT29-MTX cells. Monolayers of HT29-MTX cells were stimulated with LLO in the culture medium (90 µg/well, 37°C in 10% CO2/90% air atmosphere, 6 h), or apically infected with S. enterica serovar Typhimurium (STM) strain SL1344 (5 × 107 CFU/well, at 37°C, 1 h infection followed by a subculture time of 5 h in DMEM). Each value shown is the mean of three experiments (three successive passages of cultured cells) ± standard error. Statistical analysis was performed with a Student's t-test.In B and C, determination of IκB and c-Fos in control cells, apically Salmonella- and L. monocytogenes EGD-infected, and LLO-stimulated HT29-MTX cells.

B. Cytoplasmic proteins were isolated and IκB proteins were detected with a mouse anti-IκBα mAb, as described in Experimental procedures.

C. The nuclear extracts were examined for the presence of c-Fos by Western blotting using polyclonal antiserum as described in Experimental procedures. MW: molecular weights. Micrographs are representative of three independent experiments.

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We first control the IL-8 chemokine production in HT29-MTX cells by infecting the cells with Salmonella serovar Typhimurium, which promotes an adhesion-dependent IL-8 secretion in intestinal Caco-2 cells through activation of the nuclear factors NF-κB and AP-1 (Eaves-Pyles et al., 1999). As shown in Fig. 6A, infection of the cells by Salmonella was followed by a sixfold increase in IL-8 secretion. In contrast, no increase in IL-8 secretion was found after LLO stimulation compared with control unstimulated cells.

The nuclear translocation of NF-κB is regulated by the cytoplasmic inhibitory factor IκB which blocks its translocation to the nucleus (Baeuerle, 1991). Degradation of IκB triggers activation of NF-κB (Baeuerle and Henkel, 1994). As for the above reported IL-8 production, Salmonella-infected HT29-MTX cells were analysed for positive control of IκB degradation. As shown in Fig. 6B, Salmonella infection results in a degradation of IκB in HT29-MTX cells consistent with a previous observation in Caco-2 cells (Eaves-Pyles et al., 1999). In LLO-stimulated HT29-MTX cells, the results clearly indicate that the IκB protein level in the cytoplasm of HT29-MTX cells was not altered. As a control, we infect the cells with L. monocytogenes EGD. Results show that a rapid but transient degradation of IκB protein develops in L. monocytogenes EGD-infected HT29-MTX cells.

Cellular responses involving inducible genes are regulated by the transcription nuclear factor AP-1 (May and Ghosh, 1998). As a measure of AP-1 activity, we measured the level of c-Fos by Western immunoblotting analysis with an anti-c-Fos polyclonal Ab, in the nucleus of HT29-MTX cells after stimulation by LLO. As shown in Fig. 6C, stimulation of HT29-MTX cells by LLO at different times resulted in no increase in the nuclear levels of c-Fos. As above reported for IL-8 production and degradation of IκB, Salmonella-infected HT29-MTX cells were analysed for positive control of c-Fos synthesis. The Salmonella infection results in a marked increase in the nuclear levels of c-Fos, which paralleled the increase in IL-8 secretion and the degradation of IκB reported above.

Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

The results reported here are in line with and extend our previous results (Coconnier et al., 1998; 2000). We demonstrate that the L. monocytogenes thiol-activated exotoxin LLO promotes the mucin exocytosis in a human mucin- and PGs-secreting cell line. In parallel, we report for the first time that LLO induced the upregulation of several MUC genes encoding for membrane-bound mucins. When examining the mechanism by which LLO induced this cellular response, we found that LLO did not activate the nuclear transcriptional factors NF-κB and AP-1.

LLO enhanced specifically the mucin exocytosis in PGs- and mucin-secreting human polarized intestinal HT29-MTX cells

Host cell heparan sulphate PGs (HSPG) are integral components of plasma membranes, ubiquitously distributed among cell populations of mammalian tissues (for a review, see Prydz and Dalen, 2000). As for mucins, regulated and constitutive secretion pathways of PGs have been characterized (Brion et al., 1992). Interestingly, it has been established that PGs acted as receptors for a large variety of virulent Gram-negative and Gram-positive bacteria (for a review, see Rostand and Esko, 1997), particularly in L. monocytogenes pathogenicity (Alvarez-Dominguez et al., 1997; Jonquieres et al., 2001). Results reported here clearly outlined that the HMGs secreted upon LLO stimulation are in majority of a mucin nature.

A high level of MUC5AC mucin was found in culture medium of LLO-stimulated HT29-MTX cells. It is important to note that physiologically, the MUC5AC is known to be expressed in the respiratory tract, gastric mucosa and reproductive mucosa. The high expression level of MUC5AC mucin in the human adenocarcinoma colonic HT29-MTX and HT29-Cl.16E cell lines is reminiscent of the differentiation of the human fetal colonic epithelium (for a review, see Zweibaum et al., 1991). It could be not excluded that other mucins such as intestinal mucins expressed by the cells are secreted upon LLO stimulation. However, the levels of those mucins could not be determined by radioimmunoassay, because antibodies directed against these mucins are not currently suitable for quantitative determination.

Surprisingly, increase in MUC5AC mucin secretion in LLO-stimulated HT29-MTX cells develops without upregulation of the corresponding MUC5AC gene. There are two secretory pathways in mucin-secreting polarized cells (for reviews, Forstner, 1995; Laboisse et al., 1995). One is a steady vesicular constitutive pathway of mucin exocytosis in which small vesicles could be continuously transported directly to the cell surface to undergo exocytosis of their mucin content. Mucins are also stored in large vesicles to form a granule mass which undergoes the so-called regulated pathway of mucin exocytosis. This mucin exocytosis pathway is regulated by specific stimuli involving signalling molecules. We have previously demonstrated that LLO-induced mucin exocytosis is insensitive to pharmacological blockers of neuroendocrine receptors and coupled signalling systems, and to inflammatory agents known to be involved in controlling the regulated pathway of mucin exocytosis (Coconnier et al., 1998). Moreover, on the basis of the observation that LLO-stimulated mucin exocytosis is dependent on microtubular organization (Coconnier et al., 2000), a characteristic of the constitutive vesicular pathway of mucin exocytosis (for a review, see Forstner, 1995), we have previously suggested that LLO may activate baseline mucin secretion. Results reported here indicate that MUC5AC mucin is one of the mucins transported and secreted through the vesicular pathway of mucin exocytosis stimulated by LLO. In the related HT29–18 N2 clone, all conventional large secretory granules have been shown stain with MUC5AC antibodies (Stanley and Phillips, 1999). The fact that the MUC5AC mucin secretion was increased upon LLO stimulation without upregulation of the corresponding MUC5AC gene indicates that the mucin is from a preformed origin. We have conducted an additional experiment in which mucins have been radiolabelled during a short pulse of 1 h and 2 h (data not shown). No increase in mucins exocytosis upon LLO stimulation was observed after a short pulse compared with the increased mucins exocytosis found after a 18 h of pulse. In consequence, this result disagrees with our previous hypothesis that LLO stimulate the constitutive, non-regulated pathway of mucins exocytosis ant in contrast suggests that the mucins liberated upon LLO stimulation are not from a newly synthesized origin but from a preformed origin. Interestingly, Forstner (1995) in discussing the properties and characteristics of the mucin secretion pathways in intestinal cells, has postulated that a population of small vesicles could be stored as lateral granules on the granule mass formed in majority by large granules, which is involved in the regulated pathway of mucin exocytosis. We have previously reported that the large mucin-containing granules, located beneath the apical surface of the mucin-secreting cells and characteristic of the regulated pathway of mucin exocytosis, remain unchanged in LLO-stimulated cells (Coconnier et al., 2000). In order to reconcile our previous (Coconnier et al., 1998; 2000) and the present results, one explanation is that LLO could stimulate a subpopulation of vesicles containing MUC5AC in the granule mass forming the storage of mucins in intestinal cells.

LLO elicits upregulation of MUC genes encoding for membrane-bound mucins without activation of the nuclear transcription factors NF-κΒ and AP-1

Epithelial mucins can be classified into two main groups: membrane-associated and secreted mucins. The secreted mucins can be subdivided in two groups: gel-forming mucins and non-gel-forming mucins. Currently, 12 human mucin genes (MUC1-4, MUC5B, MUC5AC, MUC6-7, MUC11, MUC12, MUC13, and MUC14) have been identified (for reviews, Gendler and Spicer, 1995;Seregni et al., 1997; Williams et al., 1999a; Yin and Lloyd, 2001). More than one mucin gene can be expressed in a given tissue and in cultured mucin-secreting cell lines. The complete primary amino acid sequence of only MUC1, MUC2, MUC3, MUC4, MUC5AC, MUC5B and MUC7 have been deduced (for a review, see Crawley et al., 1999). The secreted gel-forming mucins are encoded by a cluster of four mucin genes (MUC2, MUC6, MUC5B and MUC5AC) located on the chromosom 11p15 (Desseyn et al., 1997). Four genes,MUC1, MUC3, MUC4 and MUC12, encoding for membrane-associated mucins have been characterized. The MUC1 gene is located on chromosome 1q21–24, MUC2, MUC6, MUC4 on 3q29, MUC7 on 4q13–21, and MUC3, MUC11 and MUC12 on 7q22 (Gum et al., 1997; Williams et al., 1999a, b).

We reported here the expression of MUC1-4, MUC5B, MUC5AC, MUC11 and MUC12 genes, but not of MUC6 and MUC7 genes in HT29-MTX cells. The observation that MUC1-4, MUC5B and MUC5AC genes are expressed in HT29-MTX cells is consistent with previous reports (Lesuffleur et al., 1993; Debailleul et al., 1998). We observed that among the MUC genes investigated, LLO promotes specifically the upregulation of MUC3, MUC4 and MUC12 genes. The present work is the four report that describes MUC gene upregulation upon bacterial infection. Indeed, it has been recently reported that the Gram-negative P. aeruginosa and Haemophilus influ­enzae and Gram-positive Staphylococcus aureus, S. epidermis and Streptococcus pyogenes promoted the upregulation of MUC2 and MUC5AC genes in NCIH292 epithelial cells (Dohrman et al., 1998; Li et al., 1998; Wang et al., 2002). All the MUC genes found to be upregulated by LLO encoded for membrane-associated mucins (Nollet et al., 1998; Crawley et al., 1999; Moniaux et al., 1999; Williams et al., 1999b).

Many cellular responses involving inducible genes are regulated by the widely used transcription nuclear factors NF-κB and AP-1 (May and Ghosh, 1999). The pleiotropic mediator of induction and tissue-specific gene control NF-κB is a member of the Rel family of transcriptional activator proteins. In activated cells, degradation of IκB proteins allows the entry of free NF-κB into the nucleus, where it binds to its target sequences and activates transcription. AP-1 is a homo- and heterodimeric transcription factor composed of members of the Jun and Fos family of DNA-binding proteins (Angel and Karin, 1991). Several stimuli such as growth factors, cytokines, hormones and microbial infections are known to induce this transcription factor. Exoproducts of P. aeruginosa promoted upregulation of MUC2 gene in NCIH292 and HM3 epithelial cells, through the activation of the transcription factor NF-κB via a Src-dependent Ras-MEK1/2-ERK1/2-pp90rsk pathway (Li et al., 1998). When examining whether or not the transcription factors NF-κB and AP-1 are involved in LLO-induced upregulation of MUC3, MUC4 and MUC12 genes, we found that upregulation occurs without activation of NF-κB and c-Fos synthesis. Cellular responses through the nuclear translocation of NF-κB by L. monocytogenes virulence factors have been recently documented, but our results are not in agreement with those showing that LLO elicits cellular responses through a NF-κB-dependent mechanism. Interestingly, we observed that control infection of the HT29-MTX cells by S. enterica serovar Typhimurium and L. monocytogenes EGD is followed by degradation of IκB proteins. LLO and L. monocytogenes in the absence of cell invasion elicits NF-κB-dependent cellular responses (Kayal et al., 1999; Rose et al., 2001). However other reports suggest the possibility that the L. monocytogenes virulence factors synergistically act to promote NF-κB-dependent cellular responses in several cell types (Schwarzer et al., 1998; Hauf et al., 1997). Analysing the NF-κB activation mechanism, it appears that the cell response involves two phases of activation, a transient activation phase by lipoteichoic acid (LTA) followed by a second persistent activation phase by the two phospholipases encoded by the virulence genes plcA and plcB. Our results obtained with L. monocytogenes EGD infection and LLO stimulation are consistent with the hypothesis that L. monocytogenes virulence factors synergistically act to promote NF-κB-dependent cellular responses. Indeed, we observed that IκB protein degradation develops when the HT29-MTX cells are infected with L. monocytogenes EGD, whereas LLO is not able to promote this effect.

Pathophysiological consequences of the LLO-induced upregulation of MUCs genes

It has been established that during L. monocytogenes infection signalling events are elicited by virulence factors interacting with host receptor signal transducing molecules (Ireton et al., 1996; Shen et al., 2000). Overexpression of MUC3, MUC4 and MUC12 mucins upon LLO stimulation reported in the present work, might be a new mechanism by which L. monocytogenes promotes signal transduction and/or influences cell growth in human mucin-secreting intestinal cells. Indeed, recent data highlight that the C-termini of human/rat/mouse MUC3, human MUC4 and MUC12 contain two conserved cysteine-rich, EGF-like domains (Williams et al., 1999a,b). It was interesting to note that MUC12 possesses a cytoplasmic tail containing an amino acid sequence, which is similar to motifs recognized by SH2 domain-containing proteins (Songyang et al., 1994). Moreover, the first EGF-like domain in MUC12 shows homology to a number of EGF receptor-binding growth factors. Finally, the rat MUC4 isoform containing the EGF-like domains binds the c-erB-2 growth factor receptor and promotes signalling (McNeer et al., 1997; Williams et al., 1999a).

The LLO-upregulated genes encoding for membrane-associated MUC3 and MUC12, are localized to the human chromosome band 7q22 (Gum et al., 1997; Williams et al., 1999a, b). A recent report described evidence for a link between inflammatory bowel disease and markers on chromosome 7q22 (Satsangi et al., 1996). Moreover, MUC3 has been proposed as a candidate susceptibility gene for inflammatory bowel disease (Kyo et al., 1999). An increase in L. monocytogenes immunoreactivity has been observed in biopsies of patients with inflammatory bowel disease (Liu et al., 1995). Moreover, an association between listeriolysin O and the induction of severe inflammatory disease in rat has been documented (Warner et al., 1996). The present results add interest to the proposed putative link between L. monocytogenes intestinal infection and inflammatory bowel disease.

Experimental procedures

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Reagents and antibodies

d-[6-3H]-glucosamine hydrochloride (specific activity 20–40 Ci mmole−1) was from Amersham Pharmacia Biotech (Buckinghamshire, UK). AEBSF, EDTA, leupeptin, benzamidine, N-ethyl malemide, heparinase III (heparin lyase III: EC 4.2.2.8) and chondroitinase ABC (chondroitin lyase ABC: EC 4.2.2.4) were obtained from Sigma-Aldrich Chimie SARL (L′Isle d'Abeau Chesnes, France). The mouse IκBα mAb H-4 and the rabbit polyclonal anti-c-Fos antibody were from Santa Cruz Biotechnology (Santa Cruz, CA).The rabbit polyclonal anti-MUC3 antibody reactive against deglycosylated mucin was from Biomeda (Forster City, CA). The anti-MUC5AC 1–13 M1 mAb and PM7 mAbs (a mixture of seven different anti-M1 mAbs) were from J. Bara (INSERM Unit 482, Paris, France). Enhancing chemiluminescence reagents (ECL-plus kit) were from Amersham (Les Ulis, France). The IL-8 immunoassay kit was from Diaclone Research Biotest (Buc, France). Polystyrene stars were from Oris Industrie (Saint Quentin en Yvelines, France). The Rneasy Midi kit was from Quiagen (Courtaboeuf Les Ulis, France). The first-strand TM cDNA synthesis kit was from Clonetech (Basingstoke, Hampshire, UK).

Bacterial strains

L. monocytogenes EGD wild type strain was a gift of J.-L. Gaiilard (INSERM U411, Paris). Bacteria were routinely grown 18 h at 37°C in tryptic soy (TS). S. enterica serovar Typhimurium strain SL1344 was a gift of B. A. D. Stocker (Stanford, California). Bacteria were cultured in Luria broth (Difco Laboratories, Detroit, MI) at 37°C.

Purification of LLO

LLO was purified from the L. monocytogenes strain EGD-SmR, a streptomycin-resistant derivative of strain EGD, routinely grown 18 h at 37°C in brain heart infusion (BHI) broth with spectinomycin 60 µg ml−1, or plated on sheep blood or TS agar plates. Protocol of LLO purification has been adapted by J.-L. Beretti (INSERM U411, Paris, France) from Geoffroy et al., 1987. Briefly, 500 µl of an overnight bacterial culture in BHI broth was grown in 10 ml of P3 broth [Protease Peptone N°3 (Difco, Detroit, MI) (0.5%, w/v)], yeast extract (0.5%, w/v), Na2HPO4 (0.5 mM), KH2PO4 (0.5 mM), sterile charcoal (0.1%, w/v), sterile H2O, complemented with 1% (w/v) glucose, for 8 h at 37°C with continuous shaking. 10 ml of P3 culture was used to inoculate 1 litre of the same medium supplemented with charcoal (0.1%, w/v). After 12 h of incubation at 37°C without shaking, the bacteria were removed by centrifugation at 5000 g for 20 min at 4°C. The cell-free supernatant was centrifugated at 10 000 g for 20 min at 4°C followed by filtration through a 0.45-µm Stericup HV5 filter unit (Millipore, Bedford, MA). EGTA 1 mM and PMSF (1 mM) were added to the supernatant to block proteases. Ammonium sulphate was added to give a final concentration of 40%. After 30 min of stirring at 4°C, the precipitate was collected by centrifugation at 10 000 g at 4°C. The precipitate was suspended in deionised H2O, dialysed against H2O containing sodium azide (0.02%, w/v) overnight at 4°C. The concentrated crude supernatant was then applied to a Q-Sepharose column (Pharmacia, Uppsala, Sweden), and eluted with Tris-HCl (50 mM, pH 6.7, the pHi of LLO).

Cell lines and culture

We used the mucin-secreting HT29-MTX cell subpopulation (Lesuffleur et al., 1990) and the clone HT29-Cl.16E (Augeron and Laboisse, 1984) selected from the parental, mainly undifferentiated HT-29 cell line (Fogh et al., 1977). Cells were routinely grown in Dulbecco's modified Eagle's minimal essential medium (DMEM) 25 mM glucose (Invitrogen, Cergy, France), supplemented with 10% heat-inactivated (30 min, 56°C) fetal bovine serum (Invitrogen).

The Caco-2/TC-7 clone (Chantret et al., 1994), established from the cultured human colonic adenocarcinoma parental Caco-2 cell line (Fogh et al., 1977), which spontaneously differentiates in culture (Pinto et al., 1983), was used. Cells were routinely grown in DMEM supplemented with 15% heat-inactivated (30 min, 56°C) fetal calf serum (Invitrogen) and 1% nonessential amino acids (Invitrogen) as previously described (Peiffer et al., 2001).

For maintenance purposes, cells were passaged weekly using 0.02% trypsin in Ca2+Mg2± free PBS containing 3 mM EDTA. Experiments and maintenance of the cells were carried out at 37°C in a 10% CO2/90% air atmosphere. The culture medium was changed daily. Cultures were used at late post-confluence.

LLO stimulation of HMGs exocytosis

Stimulation of cultured human intestinal cells by LLO was conducted has previously reported (Coconnier et al., 1998; 2000). LLO in the culture medium was added in the apical reservoir of the culture plate insert (90 µg well-1, 37°C in 10% CO2/90% air atmosphere).

For determination of HMGs exocytosis, cells were grown in filters mounted in chamber culture (Costar culture plate inserts, 3 µm porosity, 5 × 105 cells per cm2) (Dutscher, Brumath, France), which delineates an apical (luminal) and a basolateral (serosal) reservoir. Secretory HMGs were quantified using the previously described specific and sensitive electrophoretic method (Bou-Hanna et al., 1994). Briefly, before processing, the cells were metabolically labelled with 10 µCi ml−1D- [6-3H]-glucosamine hydrochloride in DMEM for 18 h. Prior to LLO challenge, the labelled cells were washed three times with DMEM medium.

The secreted [3H]-HMGs in the apical compartment were measured. The apical medium was sampled and the monolayer rinsed with additional DMEM to remove adherent molecules. Sampled apical medium and rinse were pooled. The 3H-labelled HMGs-containing medium was dialysed at 4°C for 36–48 h against several changes of deionized water and subsequently freeze-dried. The 3H-labelled HMGs were separated by SDS/PAGE on 3% gels. The band at the stacking/running gel interface, which contains the 3H-HMGs, was cut out, placed in a vial, and incubated overnight at 40°C in the presence of 0.5 ml Soluene 350 (Packard, Orsay, France). Radioactivity was determined by liquid-scintillation counting. Results were expressed as counts per minute (CPM) per 106 cells.

Enzymatic treatments

The 3H-labelled HMGs-containing medium of LLO-stimulated HT29-MTX cells was subjected to PGs lyases, heparinase III and chondroitinase ABC (0.5–1 IU per ml in PBS, 37°C, 45 min) and then analysed for HMGs determination as described above.

Biochemical analysis of secreted HMGs

Mucins and PGs synthesized by control unstimulated and LLO-stimulated HT29-MTX cells were isolated and analysed for amino acid composition, sugar analysis and sulphate content as previously described (Huet et al., 1995). Culture media containing secreted HMGs were collected. The remaining cells were rinsed twice with PBS, scraped with a rubber policeman, and directly lysed by ultrasonication in PBS. Cell extracts were collected after centrifugation for 5 min at 1000 g. In sampled culture media and cell extracts, protease inhibitors were added (AEBSF, 10 mM; EDTA, 1 mM; leupeptin, 10 µg ml−1; benzamidine, 2 mM; N-ethyl maleimide, 2 mM).

Ultracentrifugation

After adding CsBr (0.42 g ml−1), culture medium and cell extracts were ultracentrifuged (200 000 g for 72 h) using a Beckman 70.1 Ti rotor. Fractions of 1 ml were collected, weighed to determine density, and analysed for absorbance at 280 nm. The fractions 1 and 2 of the 13 fractions sampled from the bottom of the CsBr gradient, contained mucins.

Compositional analyses

For amino acid analysis, samples were hydrolysed in 5.6 M HCl for 24 h under a vacuum and processed using a 7300 Beckman amino acid analyse (Palo Alto, CA) equipped with a high performance sodium column (4 mm × 120 mm) (Beckman). Sugar analysis was carried out by gas-liquid chromatography of trimethylsilyl derivatives of methyl glucosides formed by methanolysis in 1.5 M HCl in methanol at 80°C for 24 h. To determine sulphate content, samples were hydrolysed in 1 M HCl for 5 h at 100°C and sulphate was determined by AE-HPLC. All composition analyses were performed twice to assure the reproductibility of the results.

For proteoglycan analysis, cellulose acetate plates were immersed in 0.05 M barium acetate. Samples were spotted and migration was carried out for 20 min. The plates were then stained in 0.02% dimethylmethylene blue in 1% acetic acid for 10 min and destained in 10% acetic acid.

MUC5AC immunoradiometric assay

A solid-phase double-antibody-sandwich immunoradiometric assay (IRMA) was used as previously described (Bara et al., 1988; 1991; 1998). Polystyrene stars (Oris Industrie) were coated with the anti-MUC5AC 1–13 M1 mAb (10 µg ml−1 in PBS, pH 7.4) by incubating overnight at 37°C, rinsed three times with PBS-1% BSA-tween 20 1 overnight at 37°C. After several washings, stars were dried at 40°C and stored at 4°C until use. The M1/MUC5AC mucin standard (10 µg ml−1), as well as the culture media of control unstimulated and LLO-stimulated cells were serially diluted in PBS-0.1% Tween 20 plus 1 mM NaHCO3. A volume of 300 µl of each dilution was added to the 1–13 M1 mAb coated-stars and incubated overnight at 37°C. Stars were then washed with PBS-0.1% Tween 20 and incubated with PM7 mAbs previously radiolabelled with 125I (5 × 105 cpm ml−1) overnight at 37°C. Subsequently, the stars were washed and the radioactivity was measured in a gamma counter (Wizard Model 147 005). The concentration of mucin in each sample was estimated from the IRMA standard curve obtained with the standard M1/MUC5AC mucin.

Immunofluorescence

Monolayers of HT29-MTX cells were prepared on glass coverslips, which were placed in 24-well TPP tissue culture plates (ATGC, Marne la Vallée, France). After LLO stimulation, cell preparations were fixed for 5 min at room temperature in methanol-acetone or PBS-3% paraformaldehyde. They were washed three times with PBS. Examination of MUC3 and MUC5AC expression was conducted by indirect immunolabelling with the rabbit polyclonal anti-MUC3 antibody and 1–13 M1 mAb respectively. Before MUC3 immunolabelling, mucins were deglycosylated (3 min, 37°C, neuraminidase 0.05 U). For immunolabelling, the cells were permeabilized with methanol-acetone (1/1, v/v) or 0.2% Triton X-100. Fixed and permeabilized monolayers were incubated with the primary antibody for 1 h at room temperature. After three washes in PBS, incubation with an Alexa fluor 488 secondary antibody (Molecular Probes, Eugene, OR) was performed for 1 h at room temperature. After three washes, the cells were mounted with Vectashield (Biosys SA, Compiègne, France) for immunofluorescence examination. Specimens were examined using a confocal laser scanning microscope (CLSM) (model LSM 510 Zeiss, equipped with an air cooled argon ion laser 488 nm, and a helium neon laser 543 nm) configured with an Axiovert 100 M microscope using a Plan Apochromat 63×/1.40 oil objective. Optical sectioning was used to collect 50 En face images 1 µm apart. Lateral views were obtained by integration of the images gathered at a step position of 1 on the x–y axis using the accom­panying Zeiss software LSM510 2.5 on Windows NT4. Photographic images were resized, organized, and labelled using Adobe Photoshop software (San Jose, CA). The printed images (Kodak XLS 8600 PS, Eastman Kodak, Rochester, NY) are representative of the original data. All photographs were taken on Kodak Electronic Imaging Paper (Eastman Kodak).

Reverse transcription-PCR (RT-PCR) and amplification procedures

The mRNAs were extracted from control unstimulated and LLO-stimulated HT29-MTX cells with the Rneasy Midi kit (Quiagen). First-strand cDNA species were synthesized using the first-strand TM cDNA synthesis kit (Clonetech). Oligonucleotides primers (Table 2) used in amplification procedure were synthesized by Invitrogen. Amplifications were performed in a Perkin-Elmer Thermal Cycler 480 (Perkin Elmer, Paris, France). PCR amplification reactions were conducted in 50 µl reaction volumes containing 5 µl of 10× buffer (100 nM Tris-HCl/500 mM KCl, pH 8.3), 5 µl MgCl2 25 mM, 5 µl of 10 mM deoxynucleoside triphosphates, 5 µl of the first-strand cDNA, 2 µl of each primer (10 µM) and 2 units of Ampli Taq Gold DNA polymerase (Perkin Elmer). The mixture was denatured at 95°C for 6 min followed by the amplification procedure: 35 cycles at 96°C for 30 s, annealing at 60°C for 30 s, and extension at 72°C for 1 min. The final elongation step was extended for an additional 15 min. The amplified products were electrophoresed on 2% agarose gel. PCR products were quantitated relative to a GADPH cDNA amplification control using the Kodak Digital Science 1D analysis software.

Table 2.   Oligonucleotides used for MUCs RT-PCR
GeneSequenceBind nucleotideFragmentAccession number
MUC1
5′GAACTACGGGCAGCTGGACATC1295–1316448 bpL41589
3′GCTCTCTGGGCCAGTCCTCCTG1742– 1721  
MUC2
5′CTGCACCAAGACCGTCCTCATG15291–15312401 bpNM_002457
3′GCAAGGACTGAACAAAGACTCAGAC15691–15667  
MUC3
5′AGTCCACGTTGACCACCACTGC1505–1526522 bpAB038784
3′TGTTCACATCCTGGCTGGCG2026 −2007  
MUC4
5′CGCGGTGGTGGAGGCGTTCTT2994–3014597 bpAJ010901
3′GAAGAATCCTGACAGCCTTCA3590–3570  
MUC5AC
5′TGATCATCCAGCAGCAGGGCT2897–2917409 bpAJ001402
3′CCGAGCTCAGAGGACATATGGG3305–3284  
MUC5B
5′CTGCGAGACCGAGGTCAACATC879–900415 bpXM_039876
3′TGGGCAGCAGGAGCACGGAG1293–1274  
MUC6
5′GCATGGCGAACGTGACGGTAA1034–1054421 bpU97698
3′TAGTCTGAGCCCCTGCTTGGCA1454–1433  
MUC7
5′CCACACCTAATTCTTCCCCAACTAC1010–1034407 bpXM_050370
3′CTGGCTTGTGGGATAGAGGCATT1416–1394  
MUC11
5′CAGGCGTCAGTCAGGAATCTACAG47–70169 bpAF147791
3′GAGGCTGTGGTGTTGTCAGGTAAG215–192  
MUC12
5′TGAAGGGCGACAATCTTCCTC952–972511 bpAF147790
3′TACACGAGGCTCTTGGCGATGTTG1462–1439  
GAPDH
5′TGAAGGTCGGAGTCAACGGATTTGGT113–138983 bpXM_033259
3′CATGTGGGCCATGAGGTCCACCAC1095–1072  

IL-8 assay

For determination of IL-8, monolayers of HT29-MTX cells were prepared in 24-well tissue culture plates. Prior to LLO challenge, the cells were washed three times with DMEM medium. LLO in the culture medium was added in the apical compartment (90 µg/well, 37°C in 10% C02/90% air atmosphere, 6 h). As a control for IL-8 secretion the cells were apically infected with Salmonella serovar Typhimurium strain SL1344 (5 × 107 CFU/well) for 1 h at 37°C followed, after washing, by a subculture time of 5 h in DMEM-gentamicin (100 µg ml−1). In all conditions, the culture medium in which IL-8 was determined were first centrifuged for 20 min at 12 000 g to pellet residual bacteria and cells. The IL-8 concentration was determined with the human IL-8 immunoassay kit (Diaclone Research Biotest, Buc, France). In a preliminary experiment we determined that LLO did not interfere with the IL-8 immunoassay.

IκB proteins and c-Fos immunoblotting

HT29-MTX cells were stimulated with LLO (90 µg/well, 37°C in 10% C02/90% air atmosphere). As a control, the cells were apically infected with S. serovar Typhimurium strain SL1344 (5 × 107 CFU/well, 37°C in 10% C02/90% air atmosphere). At the indicated time points post stimulation, cytoplasmic protein extract and nuclear extracts were prepared (Hobbie et al., 1997).

To examine the presence of IκB proteins at different times after stimulation, the cells were washed with PBS containing NaVO3 (1 mM) and subsequently lysed in lysis buffer (10 mM Hepes, pH 7.5, 150 mM Nacl, 10% glycerol, 1 mM NaVO3, 0.6% Triton X-100, 5 µl aprotinine, 1 µM phenylmethylsulphonyl fluoride. The cell lysate was processed for detection of IκBα protein by Western blot analysis. The cytoplasmic protein extracts were separated on 10% SDS/PAGE, transferred to PVDF, probed with the mouse anti-IκBα mAb, and developed using an enhanced chemiluminescence (ECL)-plus kit.

To examine the presence of c-Fos levels in nuclear fractions at different times after stimulation, the cells were washed in PBS containing 1 mM NaVO4, and harvested in lysis buffer (150 mM NaCl, 1.5 mM MgCl2, 0.6% NP40, 10 mM Tris/HCl, pH 7.4). Nuclei were collected by centrifugation at 12 000 g at 4°C for 10 min and examined with Western immunobloting using a c-Fos Ab.

Statistics

Data are expressed in mean ± S.E.M. of several experiments, with at least three monolayers from three successive passages of cells per experiment. The statistical significance was assessed by a Student's t-test

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

We are grateful to R. Amsellem for her expert assistance in cell culture. We thank V. Nicolas (Imagerie Cellulaire – IFR75-ISIT) for his expert assistance on confocal laser scanning microscopy analysis. We thank J.-L. Beretti (INSERM U411) for his assistance in LLO purification that was essential for this study. We thank M. Métioui for critical reading of the manuscript.

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  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
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