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Lipid raft‐dependent adhesion of Giardia intestinalis trophozoites to a cultured human enterocyte‐like Caco‐2/TC7 cell monolayer leads to cytoskeleton‐dependent functional injuries

Gardia intestinalis, the aetiological agent of giardiasis, one of the most common intestinal diseases in both developing and developed countries, induces a loss of epithelial barrier function and functional injuries of the enterocyte by mechanisms that remain unknown. Three possible mechanisms have been proposed: (i) Giardia may directly alter the epithelial barrier after a close interaction between the trophozoite and polarized intestinal cells, (ii) intestinal functions may be altered by factors secreted by Giardia including an ‘enterotoxin’, proteinases and lectins, and (iii) based on mouse studies, a mechanism involving the intervention of activated T lymphocytes. We used fully differentiated cultured human intestinal Caco‐2/TC7 cells forming a monolayer and expressing several polarized functions of enterocytes of small intestine to investigate the mechanisms by which G. intestinalis induces structural and functional alterations in the host intestinal epithelium. We first report that adhesion of G. intestinalis at the brush border of enterocyte‐like cells involves the lipid raft membrane microdomains of the trophozoite. We report an adhesion‐dependent disorganization of the apical F‐actin cytoskeleton that, in turn, results in a dramatic loss of distribution of functional brush border‐associated proteins, including sucrase‐isomaltase (SI), dipeptidylpeptidase IV (DPP IV) and fructose transporter, GLUT5, and a decrease in sucrose enzyme activity in G. intestinalis‐infected enterocyte‐like cells. We observed that the G. intestinalis trophozoite promotes an adhesion‐dependent decrease in transepithelial electrical resistance (TER) accompanied by a rearrangement of functional tight junction (TJ)‐associated occludin, and delocalization of claudin‐1. Finally, we found that whereas the occludin rearrangement induced by G. intestinalis was related to apical F‐actin disorganization, the delocalization of claudin‐1 was not.


Introduction
Giardia intestinalis (synonyms: G. lamblia and G. duodenalis), the aetiological agent of giardiasis, is a flagellated protozoan that specifically colonizes the small intestine of various vertebrates, including human beings. Giardiasis is one of the most common human intestinal diseases in both developing and developed countries (Müller and von Allmen, 2005;Lujan, 2006;Buret, 2007). G. intestinalis contributes to an estimated 280 million symptomatic human infections per year. However, most human Giardia infections result in no overt symptom, and are resolved spontaneously. In addition, G. intestinalis infection may facilitate the development of chronic enteric disorders, including inflammatory bowel disease, irritable bowel syndrome and various allergies (Buret, 2008).
Symptomatic giardiasis is characterized by severe and protracted watery diarrhoea, abdominal cramps, nausea, vomiting, malabsorption and weight loss. How the noninvasive Giardia spp. cause intestinal disease is not completely understood. The life cycle of Gardia spp. includes two main phases: the proliferative trophozoite phase and the non-proliferative, infectious cyst phase (Ankarklev et al., 2010). The infection begins with the ingestion of cysts that lead to the release of excyzoites into the proximal region of the intestine, each of which immediately divides into four trophozoites (Bernander et al., 2001). During the initial proliferative stage, Giardia trophozoites form an adhesive ventral disk that adheres to the microvillous brush border of the small intestine, consisting of enterocytes . It is possible that the ventral disk and the brush border interact via receptorligand interaction(s) (Müller and von Allmen, 2005). Giardia induces a loss of epithelial barrier function and enterocyte injuries (Müller and von Allmen, 2005;Lujan, 2006;Buret, 2007;Ankarklev et al., 2010). Insights into whether G. intestinalis promotes structural and functional injuries in small intestinal epithelium have been obtained using rodent infection models (Belosevic et al., 1989;Buret et al., 1990;Daniels and Belosevic, 1992;Gorowara et al., 1992;Cevallos et al., 1995;Humen et al., 2005), cultured non-human and human intestinal cells (Chavez et al., 1986;Favennec et al., 1991;Teoh et al., 2000;Sousa et al., 2001;Scott et al., 2002) and human biopsies (Oberhuber et al., 1997;Singh et al., 2000;Teoh et al., 2000). In addition, although Giardia does not induce the production of pro-inflammatory cytokines (Jung et al., 1995), infected human intestinal cells display a particular profile of chemokines, substances that are involved in the recruitment of dendritic cells and can interfere with host's innate immunity, thus allowing the parasite to control the host's response (Roxstrom-Lindquist et al., 2005;Kamda and Singer, 2009). The mechanisms used by G. intestinalis to induce structural and functional alterations in the host intestinal epithelia remain unknown. It has been postulated that most of the Giardia-induced intestinal functional disorders result from brush border damage induced by Giardia (Müller and von Allmen, 2005;Buret, 2007). Three mechanisms have been proposed. According to the first mechanism, unidentified Giardia virulence factors directly alter the epithelial barrier after close interaction has occurred between the trophozoites and polarized intestinal cells. The second mechanism involves the intervention of activated T lymphocytes. The third mechanism involves factors secreted by Giardia, including an 'enterotoxin', proteinases and lectins. In the present study, we focused our investigation on the mechanism by which G. intestinalis promotes structural and functional cellular injuries after interaction has occurred between trophozoites and human intestinal cells. As our cellular model, we used human intestinal Caco-2/TC7 cells (Chantret et al., 1994), which spontaneously differentiate in culture to produce to a confluent cell monolayer that mimics the intestinal epithelial barrier (Pinto et al., 1983). Importantly, fully differentiated and polarized Caco-2/TC7 cells, although they are of colonic origin, express apical and basolateral proteins displaying the specific functions of enterocytes of the small intestine (Zweibaum et al., 1991). Many of the cellular and molecular mechanisms of enterovirulent microorganisms identified in non-polarized epithelial cells have been validated for the intestinal situation using this cell line (Lencer, 2001;Boyle and Finlay, 2003). Moreover, structural and functional alterations induced by enterovirulent bacteria and enteric viruses have recently been investigated using fully differentiated Caco-2 cells or clones. In order to evaluate the pathological mechanisms that lead to G. intestinalis-induced alterations in the host's epithelial structure and intestinal function, we assessed cell integrity, cell polarization, the cellular brush border cytoskeleton, the distribution of brush border enzymes, the integrity of the epithelial barrier and the distribution of functional tight junction (TJ) proteins. We provide evidence that both the structural and functional injuries promoted by G. intestinalis strains at the brush border of Caco-2/TC7 enterocyte-like cells and at the TJ of Caco-2/TC7 monolayers are adhesiondependent. Moreover, we showed that adhesiondependent induced disassembly in the apical F-actin cytoskeleton controls the loss of distribution of brush border-associated functional proteins including sucraseisomaltase (SI), dipeptidylpeptidase IV (DPP IV) and the fructose transporter, GLUT5, and causes a decrease in sucrase enzyme activity. We demonstrate that G. intestinalis delocalizes the TJ-associated functional proteins, occluding and claudin-1. Finally, we observed that the disassembly of apical F-actin cytoskeleton plays a pivotal role in the rearrangement of the TJ-associated protein occludin, but not in the rearrangement of claudin-1.

Results
Trophozoite lipid rafts control the adhesion of G. intestinalis to human enterocyte-like Caco-2/TC7 cells We examined the attachment capacity of G. intestinalis strains GS/H7, WB/C6 and WB/1267 to apically infected, fully differentiated Caco-2/TC7 cells. The number of trophozoites attached was determined 3 h post infection (p.i.), after infection with multiplicity of infection (moi) of 0.5, 1, 2 and 8 trophozoites per cell. Concentrationdependent adhesion was observed for all the strains examined (Fig. 1A). Strains GS/H7 and WB/C6 were highly adhesive, whereas strain WB/1267 displayed lower adhesion capability. Both the motility and ability to grow of adhering trophozoites were normal (not shown). A timecourse infection with strain GS/H7 showed that the trophozoites adhered rapidly, and that adhesion levels did not evolve any further during the time-course of the infection (Fig. 1B).
Lipid raft microdomains at the cell plasma membrane play a pivotal role in infectious processes (van der Goot and Harder, 2001). It has recently been reported that lipid rafts are involved in adhesion of Entamoeba histolytica trophozoites (Laughlin et al., 2004;Mittal et al., 2008). We investigated whether methyl-b-cyclodextrin (MBCD) (10 mM), a lipid raft disorganizing agent, affects the adhesion of G. intestinalis trophozoites to Caco-2/TC7 cells (Fig. 2). When cells were apically infected with cultures of G. intestinalis strains GS/H7, WB/C6 or WB/1267 in the presence of MBCD, we found that adhesion of trophozoites was abolished compared with untreated, infected cells ( Fig. 2A). Since lipid rafts were present both in the cells and in the trophozoite membranes, we pre-treated either the Caco-2/TC7 cells or the G. intestinalis strain before the adhesion experiment. As shown in Fig. 2B, when the Caco-2/TC7 cells had been pre-treated with MBCD there was no inhibition of trophozoite adhesion compared with that of untreated cells. In contrast, when GS/H7, WB/C6 or WB/1267 trophozoites were pre-treated with MBCD before being used to infect Caco-2/TC7 cells, there was a complete inhibition of the adhesion (Fig. 2C). These findings constitute the first demonstration that G. intestinalis membrane rafts play a role in the adhesion of this organism to human, enterocyte-like cells.
The adhesion of G. intestinalis disrupts the apical F-actin network in human enterocyte-like Caco-2/TC7 cells We investigated differences in the distribution of F-actin at the apical brush border of fully differentiated control Caco-2/TC7 cells and of Caco-2/TC7 cells infected with cultures of G. intestinalis GS/H7, WB/C6 and WB/1267 (moi 8, 24 h infection). Confocal laser microscopy scanning (CLMS) x-y images of F-actin labelled with fluorescein-labelled phalloidin in non-permeabilized cells (Fig. 3A) show that control cells displayed the typical mosaic pattern of microvillus-associated F-actin due to the different morphologies of microvilli in fully differentiated Caco-2 cells (Peterson and Mooseker, 1992;. In contrast, Caco-2/TC7 cells apically infected with G. intestinalis GS/H7 and WB/C6 showed a dramatic alteration of the distribution of F-actin (Fig. 3A). F-actin immunolabelling revealed central translucent zones, with the accumulation of F-actin at cell-tocell contact points. In addition, vacuole-like structures were observed in infected cells. In WB/1267-infected cells, mildly impaired apical distribution of F-actin was observed (Fig. 3A). Quantification of the F-actin labelling confirms that F-actin distribution was less affected in Fig. 1. Adhesion of G. intestinalis trophozoites to human enterocyte-like Caco-2/TC7 cell monolayers. After incubating for 3 h, the adhering trophozoites were counted as described in Experimental procedures. A. Concentration-dependent adhesion of GS/H7, WB/C6 and WB/1267 trophozoites onto Caco-2/TC7 cells. Different amounts of trophozoites were added per well: 5 ¥ 10 5 , 1 ¥ 10 6 , 2 ¥ 10 6 or 8 ¥ 10 6 trophozoites ml -1 , with a multiplicity of infection (moi) of 0.5, 1, 2 and 8 respectively. B. The time-course of GS/H7 trophozoite adhesion (moi, 8). In (A), the asterisk (*) indicates a significant difference (P < 0.05) versus GS/H7 and GS/W6 at the same moi.
These findings prompted us to examine in greater detail the localization of F-actin within fully differentiated control Caco-2/TC7 cells and in G. intestinalis-infected Caco-2/ TC7 cells. To do this, cells were infected with G. intestinalis strain GS/H7 (moi 8, 24 h infection). The F-actin network was examined by direct labelling of Triton-permeabilized cells with fluorescein-labelled phalloidin and CLMS analysis. As shown in Fig. 3C, the control cell monolayers displayed the typical regular organization of fully differentiated Caco-2/TC7 cells, with F-actin localized apically at the brush border and laterally at cell-to-cell contact points in polarized cells (Peterson and Mooseker, 1992;. The cell-to-cell contacts expressing F-actin were still present in cells apically infected with G. intestinalis GS/H7 (Fig. 3C), indicating that the polarized organization of the cells had not been altered. Consistent with this, no signs of any disruption of cell integrity were observed when assessed by determining LDH activity in the supernatant, cell detachment and necrosis (data not shown). However, the GS/H7-infected cell monolayers appeared to have been dramatically altered, with irregular cell sizes compared with the control cells. In particular, in some infected cells the apical domain was rounded, and F-actin labelling at the brush border was irregular compared with that in control cells. Comparison of the sizes of the apical F-actin bands by CLMS and image analysis shows that, in control cells, the F-actin labelling was distributed as a homogeneous band measuring 4.8 Ϯ 0.2 mm. In contrast, in GS/H7-infected cells the apical the F-actin labelling was irregular, forming a band of~2.3 Ϯ 0.7 mm (P-value < 0.01).
F-actin immunolabelling was examined in fully differentiated cells infected with G. intestinalis strain GS/H7infected (moi 8), during a time-course of infection. Surprisingly, we observed no change in the distribution of brush border-associated F-actin in non-permeabilized, infected cells 3 h or 12 h p.i. (Fig. 3D) compared with that in infected cells at 24 h p.i., despite the fact that similar trophozoite adhesion to Caco-2/TC7 cells was seen 3 h, 12 h and 24 h p.i. (Fig. 1B). Fig. 2. Treatment of G. intestinalis GS/H7 trophozoites with the lipid raft-destabilizing agent, methyl-b-cyclodextrin (MBCD) inhibits their adhesion to Caco-2/TC7 cells. A. Adhesion of GS/H7 trophozoites with or without MBCD in the incubation medium. B. Adhesion of GS/H7 trophozoites to Caco-2/TC7 cells with or without MBCD pre-treatment of cells. C. Adhesion of untreated and MBCD-treated (10 mM) GS/H7 trophozoites to Caco-2/TC7 cells. All data are the means (SEM) of three independent experiments. In (B) and (C), The asterisks indicate a significant difference (P < 0.01) versus control cells without MBCD treatment. Fig. 3. Distribution of F-actin in human enterocyte-like Caco-2/TC7 cell monolayers infected with G. intestinalis strains. After infection, the cells were fixed, permeabilized with Triton X-100 (or not), and processed for direct immunofluorescence labelling of F-actin with fluorescein-phalloidin as described in Experimental procedures, and observed by CLMS microscopy. A. F-actin immunofluorescence in non-permeabilized control cells, and cells apically infected G. intestinalis trophozoites strains GS/H7, WB/C6 and WB/1267 (moi = 8) (horizontal x-y optical sections). B. Quantification of F-actin immunofluorescence by CLMS analysis in non-permeabilized control cells and in G. intestinalis-infected cells. The adhesion of G. intestinalis dramatically modified the distribution of the brush border enzymes, of SI and of the fructose transporter GLUT5 Functional proteins with specific intestinal functions are known to be localized at the brush border of cultured human enterocytes (Peterson and Mooseker, 1992;. Since modification of the brush border cytoskeleton could lead to functional deficiencies, we analysed the distribution of brush border enzymes and transporters in GS/H7-infected cells. The enzymes and transporters investigated included SI, an a-glycosidase that hydrolyses maltose, sucrose and maltotriose, DPP IV, a widely distributed membrane glycoprotein essential for the transport of proline-containing peptides, and the transporter GLUT5, which is required for the active transport of fructose (Fig. 4). The immunofluorescent labelling of SI in non-permeabilized GS/H7-infected cells at 24 h p.i. shows that a high proportion of the infected cells had lost the mosaic pattern of SI immunofluorescence, which had been replaced by small, SI-positive puncta ( Fig. 4A and B). In the same way, GS/H7-infected cells showed a dramatic reduction in the brush border-associated transporter GLUT5 as compared with uninfected cells ( Fig. 4A and B). In contrast, the distribution of DPP IV was modified after infection to a lesser extent than that of SI or GLUT5 ( Fig. 4A and B). Correlating with this modification of the SI distribution, we observed a dramatic decrease in the brush border membrane-associated sucrase activity in G. intestinalis GS/H7-infected Caco-2/TC7 cells compared with that in control cells (Table 1). We further examined SI and GLUT5 immunolabelling at the brush border of non-permeabilized cells during the time-course of infection with strain GS/H7 (moi 8). As observed above for F-actin ( Fig. 3D), we saw no change in the distribution of SI or GLUT5 at the brush border of the infected GS/H7infected cells 3 h and 12 h p.i. (Fig. 4C) compared with infected cells at 24 h p.i.
Since several authors have variously linked giardiasis to iron, haemoglobin and transferrin deficiencies (Sackey et al., 2003;Monajemzadeh and Monajemzadeh, 2008), we investigated whether G. intestinalis strain GS/H7 induces any modification in the transferrin receptor distribution. There was no difference between infected and control cells with regard to the distribution of the transferrin receptor (not shown).

Inhibition of adhesion and stabilization of the cytoskeleton antagonize the G. intestinalis GS/H7-induced impairment of SI distribution and activity
We investigated whether the G. intestinalis-induced structural and functional changes observed above in Caco-2/ TC7 cell monolayers are related to trophozoite adhesion or not. We checked whether the above-observed effects on F-actin, SI and GLUT5 distribution were related to substances secreted by G. intestinalis. When Caco-2/ TC7 cells were incubated for 24 h in the presence of a cell-free, spent culture supernatant of strain GS/H7, the distributions of F-actin, SI, DPP IV and GLUT5 were the same as those following incubation in the presence of a GS/H7 culture (Fig. 5A). In order to check that the adhesion of GS/H7 trophozoites determined the structural and functional changes observed above with GS/H7 culture, we used MBCD to inhibit the adhesion of G. intestinalis trophozoites. When Caco-2/TC7 cells were apically infected for 24 h with MBCD-treated G. intestinalis GS/H7 trophozoites (moi = 8), CLMS analysis of infected cells showed that the distribution of apical F-actin cytoskeleton, SI and GLUT5 was the same as that in cells infected with untreated GS/H7 ( Fig. 5B and C). We also found no difference between the sucrase activity in the brush border membrane of Caco-2/TC7 cells infected with MBCDtreated GS/H7 and that of cells infected with untreated GS/H7 (Table 1).
It has previously been reported that the functionality of brush border-associated proteins is dependent on the establishment and maintenance of the polarized organization of the cells, which is controlled by the cytoskeleton . In order to demonstrate whether the G. intestinalisinduced functional injuries reported above are directly related to cytoskeleton disorganization, we conducted experiments in which the cytoskeleton was stabilized before infection. To do this, we used jasplakinolide (JAS) (1 mM), an F-actin-stabilizing agent to protect the cell cytoskeleton in Caco-2/TC7 cells infected with an enterovirulent Escherichia coli (Peiffer et al., 2001). In a preliminary experiment, we checked that the JAS treatment had no effect on the adhesion of G. intestinalis GS/H7 to Caco-2/TC7 cells (not shown) and as expected, found that it did not modify the apical distribution of F-actin and SI in uninfected cells (Fig. 6A). We observed that the apical distribution of F-actin, SI and GLUT5 was the same in JAS-treated G. intestinalis GS/H7-infected cells as in Table 1. Adhesion-and lipid raft-dependent reduction of sucrase activity in the brush border of G. intestinalis GS/H7-infected enterocyte-like Caco-2/TC7 cells. untreated G. intestinalis GS/H7-infected cells ( Fig. 6A and B). Consistently with the CLMS examination, we observed that sucrase activity in the brush border membrane of JAS-treated, GS/H7-infected Caco-2/TC7 cells was the same as that of control uninfected cells and non-JAStreated GS/H7-infected cells (Table 1). Taken together, these results showed that the GS/H7-induced structural and functional brush border injuries are both adhesion- dependent. Our findings also demonstrate that GS/H7induced changes in the distribution and activity of brush border-associated SI hydrolase and GLUT5 transporter are dependent on the GS/H7-induced disassembly of the F-actin cytoskeleton.

G. intestinalis induces alterations in the distribution of the TJ-associated functional proteins, occludin and claudin-1
Previous reports have shown that decreased TER results from alterations of the TJ-associated proteins, ZO-1 and claudin-1, after Giardia infection of cell monolayers Scott et al., 2002) and in human biopsy specimens from patients with chronic giardiasis (Troeger et al., 2007). We confirmed that a decrease in TER develops in Caco-2/TC7 cell monolayers infected with G. intestinalis strains GS/H7, WB/C6 or WB/1267 (moi = 8, 24 h of infection) ( . We conducted an in-depth analysis of the distribution of two functional TJ-associated proteins: occludin (Furuse et al., 1993) and claudin-1 (Furuse et al., , the asterisk (*) indicates a significant difference (P < 0.01) versus control, and the double asterisk (**) indicates a significant difference (P < 0.01) versus the GS/H7 culture. In (C), the asterisk (*) indicates a significant difference (P < 0.01) versus control, and the double asterisk (**) indicates a significant difference (P < 0.01) versus GS/H7.

1998) in G. intestinalis-infected
Caco-2/TC7 cell monolayers (moi = 8, 24 h of infection). Indirect immunolabelling of occludin and CLMS analysis showed that the protein was localized at cell-to-cell contact points in uninfected Caco-2/TC7 cells, and the typical honeycomb-like pattern can be observed (Fig. 7A). In contrast, apical infection of Caco-2/TC7 cell monolayers with G. intestinalis strains GS/H7, WB/C6 or WB/1267 induced modification of the cell-to-cell organization characterized by an obvious loss of the typical honeycomb structure (Fig. 7A). Occludin distribution was modified, now showing a brightly stained continuous band lining each cell-to-cell contact.
Phosphorylated and non-phosphorylated forms of occludin have both been described (Farshori and Kachar, 1999). It has been established that non-phosphorylated occludin has a cytoplasmic localization, and that it is easily extracted using Triton X-100. In contrast, phosphorylated occludin, located at cell-to-cell contact points, is resistant to extraction with Triton X-100. Occludin distribution was analysed by Western blotting of Triton X-100-soluble and Triton X-100-resistant fractions of G. intestinalis GS/H7-infected Caco-2/TC7 cells. In Triton X-100-soluble fractions (Fig. 7B), a 65 kDa band corresponding to the non-phosphorylated form of occludin was observed. A densitometric analysis of the Triton X-100soluble fraction of G. intestinalis GS/H7-infected cells revealed that non-phosphorylated occludin was significantly lower (P < 0.05) in these cells than in the control cells (Fig. 7C). In the Triton X-100-resistant fraction (Fig. 7B), two bands were observed corresponding to the non-phosphorylated (65 kDa) and phosphorylated (72 kDa) forms of occludin respectively. Quantification of non-phosphorylated occludin in the Triton X-100-resistant fraction of G. intestinalis GS/H7-infected cells did not reveal any statistical difference from control cells (Fig. 7C). In contrast, the Triton X-100-resistant fraction of   G. intestinalis GS/H7-infected cells contained more of the phosphorylated form of occludin than the control cells (P < 0.05) ( Fig. 7B and D). The differential expression of claudins 1, 2, 3, 4, 5, 6, 7 and 8 had previously been reported depending on the Caco-2 cell clones used. Depletion of Caco-2 cell cholesterol with MBCD resulted in the displacement of claudins 3, 4 and 7, but not of claudin 1, from the cholesterol-rich domains (Lambert et al., 2007). Since MBCD treatment was used in our experiments, we decided to examine claudin-1 distribution in Caco-2/TC7 cells infected for 24 h with G. intestinalis strain GS/H7, WB/C6 or WB/1267 trophozoites. In the control cells, claudin-1 was seen at the cell-to-cell contact points (Fig. 8A). In contrast, there was a total loss of the claudin-1 immunolabelling at cell-to-cell contacts between infected cells, and claudin-1 was localized within small aggregates in the central region (Fig. 8A). The GS/H7-, WB/C6-and WB/1267-infected cells subjected to Triton X-100 treatment showed that the delocalized, centrally expressed, claudin-1 aggregates were resistant to extraction with the detergent (Fig. 8B).
Next to the TJs lies the adherens junction that orchestrates the intercellular junction, thereby playing a pivotal role in establishing and maintaining the epithelial architecture. Examining the distribution of the adherens junction-associated protein, E-cadherin, we observed that the Giardia infection did not modify the distribution of this protein (not shown), suggesting that the polarized organization of the Caco-2/TC7 cell monolayer was not modified after Giardia infection.
We then investigated whether inhibition of the adhesion of G. intestinalis GS/H7 trophozoites by MBCD and/or stabilization of the F-actin cytoskeleton by JAS modified the GS/H7-induced changes in TER, or the distribution of occludin and claudin-1. The results shown in Table 2 show that no G. intestinalis GS/H7-induced decrease in TER occurred when the GS/H7 trophozoites were treated before infection with MBCD or when the Caco-2/TC7 cells were pre-treated with JAS before infection. As shown in Fig. 9, both MBCD treatment (Fig. 9A) and JAS treatment (Fig. 9B) abolished the GS/H7-induced rearrangement of occludin. MBCD treatment of cells also abolished the GS/H7-induced delocalization of claudin-1. Surprisingly, JAS treatment of cells did not modify the GS/H7-induced delocalization of claudin-1. Overall, these results show that both the GS/H7-induced rearrangement of occludin and the delocalization of claudin-1 are adhesiondependent. Interestingly, we found that the adhesiondependent GS/H7-induced rearrangement of occludin resulted of GS/H7-induced disassembly of the F-actin cytoskeleton, whereas the GS/H7-induced delocalization of claudin-1 did not.

Discussion
The intestinal polarized cells that make up the epithelium provide physical and chemical barriers that protect the Fig. 8. Changes in the distribution of the tight junction-associated protein, claudin-1, in G. intestinalis-infected human enterocyte-like Caco-2/TC7 cell monolayers. Cells were apically infected with trophozoites of G. intestinalis strains GS/H7, WB/C6 and WB/1267 for 24 h (8·10 6 trophozoites ml -1 , moi = 8). Cells were processed for indirect immunofluorescence labelling of claudin-1, as described in Experimental procedures. A. Control cells and cells infected apically with trophozoites of G. intestinalis strains GS/H7, WB/C6 and WB/1267. B. Infected cells were processed for the indirect immunofluorescence labelling of claudin-1 after treatment with Triton X-100. CLMS analysis was carried out to obtain en face micrographs (horizontal x-y optical sections). The micrographs are representative of three independent experiments. host against the unwelcome intrusion of microorganisms (Lievin-Le Moal and Servin, 2006). The adhesion of G. intestinalis trophozoites to epithelial intestinal cells is the first step in the pathogenesis of intestinal disease. There is evidence showing that this adhesion step is a multi-factorial process that involves flagella motility, mechanical forces of the ventral disk and of the ventrolateral border structures, and surface molecules (Ankarklev et al., 2010). Previous studies have demonstrated the role of G. intestinalis surface lectins in the attachment of trophozoites to epithelial host cells (Pegado and de Souza, 1994;Katelaris et al., 1995;Sousa et al., 2001). We describe here for the first time that disruption of G. intestinalis trophozoites lipid rafts by MBCD treatment leads to the inhibition of adhesion of trophozoites at the brush border, and from the abolition of the G. intestinalis-induced structural and functional injuries in cultured enterocyte-like cells. Subdomains in cell membranes created by lipid-lipid and protein-lipid interactions are referred to as membrane lipid rafts rich in phospholipids, glycosphingolipids and cholesterol. Lipid rafts contain glycosylphosphatidylinositol-anchored proteins, and several studies have shown that myristoylation and palmitoylation play a role in the association of transmembrane proteins with raft membranes. The presence of membrane lipid rafts is suggested in Giardia by the observation of the presence of lipid raft-associated molecules including the invariant glycosylphosphatidylinositol (GPI)-anchored surface protein GP49 (Das et al., 1991), which alters electrolyte fluxes in cultured human colonic Fig. 9. Inhibition of G. intestinalis GS/H7 adhesion to human enterocyte-like Caco-2/TC7 cell monolayers by MBCD treatment of trophozoites abolishes both the trophozoite-induced rearrangements in the distributions of occludin and claudin-1, and stabilization of the F-actin cytoskeleton by jasplakinolide treatment of cells had no effect on claudin-1 rearrangement, but inhibited the rearrangement of occludin. Cells were apically infected with trophozoites of G. intestinalis strains GS/H7, WB/C6 and WB/1267 for 24 h (8·10 6 trophozoites ml -1 , moi = 8). Cells were processed for the indirect immunofluorescence labelling of occludin and claudin-1, as described in Experimental procedures. A. Occludin distribution in non-infected, control cells and in cells infected with trophozoites, which had or had not been treated with MBCD (10 mM). B. Distribution of occludin in control cells and in GS/H7-infected cells treated with JAS (1 mM). C. Claudin-1 distribution in uninfected control cells and cells infected with trophozoites, which had or had not been treated with MBCD (10 mM). D. Claudin-1 distribution in control cells and in GS/H7-infected cells treated with JAS (1 mM). CLMS analysis was carried out to obtain en face micrographs (horizontal x-y optical sections). The micrographs are representative of three independent experiments. epithelial T84 cells (Das et al., 1994), and palmitoylated and/or myristoylated giardins (Jenkins et al., 2009;Saric et al., 2009) that could contribute to the attachment of G. lamblia trophozoites to the intestinal epithelium . Moreover, many variant-specific surface proteins (VSPs) that cover the entire surface of Giardia, and which are antigenically modified when Giardia undergoes antigenic variation are thought to play an important role for evading the host's immune system (Nash, 2001), are palmitoylated (Papanastasiou et al., 1997;Hiltpold et al., 2000;Touz et al., 2005). GP49, an invariant protein, has been identified in strain WB/C6 (Das et al., 1991) and palmitoylated variant surface protein H7 in strain GS/H7 (Touz et al., 2005). Delta-giardin has been identified in the ventral disk of ATCC strain 30957 WB (Jenkins et al., 2009), palmitoylated and myristoylated a-19 giardin in the ventral flagella of the WB/C6 strain (Saric et al., 2009), and alpha8-giardin on the plasma membrane and flagella, but not on the ventral disk (Wei et al., 2010). It is interesting to note that, as we found here for G. intestinalis, disruption of raft-like membrane domains in E. histolytica by the cholesterolbinding agents, filipin and MBCD results in the inhibition of several important virulence functions, fluid-phase pinocytosis and adhesion to host cell monolayers (Laughlin et al., 2004). A raft-resident adhesin, galactose/ N-acetylgalactosamine-inhibitable lectin, has been found to be involved in the interaction of E. histolytica with the host extracellular matrix by binding to the galactose or N-acetylgalactosamine moieties of collagen receptor (Laughlin et al., 2004;Mittal et al., 2008). Pre-treatment of G. lamblia trophozoites with anti-delta-giardin sera caused morphological changes in the parasite and inhibited trophozoite binding to the surface of cell culture slides (Jenkins et al., 2009). Since the strong attachment of G. intestinalis trophozoites is produced by the adhesive disk and flagella (Müller and von Allmen, 2005;Ankarklev et al., 2010), it remains to be determined which lipid raftassociated VSPs and/or giardins associated with ventral disk and/or flagella trigger the lipid raft-dependent adhesion of G. intestinalis strains GS/H7, WB/C6 or WB/1267 to human enterocyte-like Caco-2/TC7 cells.
Electron microscopy studies indicate that the close adhesion of G. intestinalis trophozoites to intestinal cells by means of their ventral disc directly induces cell damage (Chavez et al., 1986;Favennec et al., 1991;Teoh et al., 2000;Sousa et al., 2001). Microvillous shortening, villous flattening or atrophy are all hallmarks of giardiasis. It has been reported that infecting intestinal cells by G. intestinalis trophozoites disrupts the cytoskeleton network (Chavez et al., 1986;Buret et al., 1990;Favennec et al., 1991;Scott et al., 2000;Teoh et al., 2000). Consistently with these findings, we observed a dramatic disorganization of the apical F-actin in enterocyte-like Caco-2/TC7 cells infected with all the G. intestinalis strains examined here. It is worth noting that no modification of apical F-actin developed when the cells were exposed to spent culture supernatants of G. intestinalis strains GS/H7, WB/C6 or WB/1267 cultures, or when trophozoite adhesion was inhibited by MBCD treatment. This is consistent with the hypothesis that the cytoskeleton damage is directly associated with the adhesion of the parasite to host cells. However, there is a discrepancy between this finding and a previous observation by Teoh et al. (2000), indicating that cultured, non-transformed human duodenal epithelial SCBN and enterocyte-like Caco-2 cells infected with spent culture medium of G. lamblia strain S2 showed F-actin rearrangements similar to those promoted by live or lysed G. lamblia. This discrepancy could be attributable to the fact that we used different strains in our study, and suggests that active molecule(s) are specifically secreted by strain S2.
Giardia-induced intestinal cell abnormalities lead to the impairment of the digestive process, and this in turn promotes malabsorption and diarrhoea. These abnormalities have been observed in animal models and human biopsies. Buret et al. (1990) observed that mice infected with Giardia muris developed atrophied intestinal villi and reduced disaccharidase activity. Humen et al. (2005), using a gerbil model infected with G. intestinalis, have reported the same effect. Interestingly, Daniels and Belosevic (1992; have reported that C3H/HeN and C57BL/6 mice infected with G. muris displayed reduced activities of intestinal lactase, sucrase, trehalase and maltase. Belosevic et al. (1989) found that infecting gerbils with G. lamblia caused a dramatic decrease in disaccharidase activity. Buret et al. (1991; have reported that in G. duodenalis-infected gerbils, the loss of microvillous border surface area is correlated with the reduction in mucosal sucrase and maltase activities, and with that of the jejunal glucosestimulated absorption of electrolytes, water, and 3-Omethyl-D-glucose. In human duodenal biopsy specimens from patients with chronic giardiasis, Troeger et al. (2007) observed that sodium-dependent glucose absorption is impaired and active electrogenic anion secretion is activated. Giardia-induced disturbance of intestinal functions has been attributed, in part, to shortening of brush border microvilli. In the present article, results obtained after the inhibition of trophozoite adhesion by pre-treating the trophozoites with MBCD or stabilizing the F-actin cytoskeleton by JAS treatment before infection provide evidence that the change in the distribution of brush border-associated SI, DPP IV hydrolases and GLUT5 transporter, and the decrease in sucrose enzyme activity result from the G. intestinalis-induced apical F-actin rearrangements.

Giardia intestinalis-induced cytoskeleton-dependent intestinal functional injuries 1695
Structural lesions of the brush border produced by adhering G. intestinalis trophozoites have been explained as resulting from the mechanical-tension extraction of microvilli mediated by suction between the ventral disk of the trophozoite and the epithelial intestinal cells forming the brush border (Müller and von Allmen, 2005;Ankarklev et al., 2010). The observation here that, even though G. intestinalis had adhered to enterocytelike Caco-2/TC7 cells at 3 h and 12 h p.i., the cells did not display any change in the distribution in the brush border of F-actin, SI and GLUT5 did not fit well with the presence of mechanical tension lesions at the brush border. We observed that the dramatic structural and functional brush border changes in cells infected with the GS/H7 culture seen at 24 h p.i., but not at 3 h and 12 h p.i. despite the fact that the same levels of G. intestinalis GS/H7 trophozoites were found at adhering 3 h, 12 h and 24 h p.i. These findings suggest that G. intestinalis produces harmful substances in response to adhesion. Analysis of the interaction between G. intestinalis and epithelial intestinal cells by transcriptional profiling of interacting trophozoites using Giardia microarrays has shown that a total of 200 transcripts were significantly up-or downregulated as a result of the interaction, lasting up to 18 h p.i. . Giardia is known to contain and/or release a variety of harmful substances. A sonicate of G. lamblia trophozoites leads to an increase in the transepithelial flux and ZO-1 rearrangement . Proteases (Hare et al., 1989;Parenti, 1989) and excreted-secreted substances (Chen et al., 1995;Kaur et al., 2001;Jimenez et al., 2004;Shant et al., 2004;Rodriguez-Fuentes et al., 2006) have been identified in G. intestinalis. Contact between G. lamblia and epithelial cells triggers the release of proteases (Rodriguez-Fuentes et al., 2006), metabolic enzymes including arginine deiminase, ornithine carbamoyl transferase and enolase, which might facilitate the effective colonization of the human small intestine (Ringqvist et al., 2008) and elongation factor 1-alpha (Skarin et al., 2011).
The integrity of the intestinal layer of epithelial cells is maintained by intercellular junctional complexes composed of TJs, adherent junctions and desmosomes, whereas gap junctions allow intercellular communication to occur (Schneeberger and Lynch, 2004). TJs, the most apical components of the junctional complex, create a semipermeable diffusion barrier between individual cells that can be regulated. Moreover, TJs constitute a functional limit between apical and basolateral cell membrane domains, and also help to maintain cell polarity. It is worth noting that many pathogenic enteric bacteria target and exploit the TJs in their pathogenic strategies (Berkes et al., 2003;Guttman and Finlay, 2008a;O'Hara and Buret, 2008). Many studies have shown that there is an increase in intestinal permeability, and a reduction in TER in giardiasis. Cultured SCBN, Caco-2 and MDCK cell monolayers infected with G. intestinalis trophozoites displayed a loss of TER and increased paracellular permeability (Chavez et al., 1986;Hardin et al., 1997;Teoh et al., 2000;Buret et al., 2002;Chin et al., 2002;Dagci et al., 2002). G. intestinalis or G. muris trophozoites disrupted the distribution of ZO-1, which is delocalized from the TJs to the cytosol in SCBN and Caco-2 cells Chin et al., 2002;Scott et al., 2002). In human duodenal biopsy specimens from patients with chronic giardiasis, Troeger et al. (2007) have observed moderate down expression of the TJ-associated claudin-1 and -7, but not of occludin and claudin-4, accompanied by a decrease in TER and an increase in paracellular permeability. We show here that G. intestinalis promotes the phosphorylation of occludin. Phosphorylation events have been reported in enterocytes infected with G. intestinalis sonicates, where myosin light chain kinase (MLCK) phosphorylates the myosin light chain at the actomyosin ring, thus altering the distribution of F-actin and the ZO-1 . Taking into consideration the fact that F-actin cytoskeleton disruption caused by G. intestinalis trophozoites develops in parallel with the delocalization of the structural TJ-associated protein ZO-1 Buret et al., 2002;Chin et al., 2002;Scott et al., 2002), it has been suggested that the Giardia-induced disruption of TJs results from disorganization of the F-actin cytoskeleton. In the present study we inhibited G. intestinalis adhesion with MBCD and demonstrated that the G. intestinalis-induced changes in TER and distribution of TJ-associated proteins depend directly on the adhesion of trophozoites to the brush border. Consistent with the role of structural TJ-associated ZO proteins and their connection with the F-actin cytoskeleton (Fanning et al., 1998), and the relevance of structural TJ proteins for the establishment and the maintenance of the TER (Gonzalez-Mariscal et al., 2000), we report that JAS treatment of Caco-2/ TC7 cells results in inhibition of the G. intestinalisinduced loss of TER. The association of occludin with the underlying F-actin cytoskeleton occurs via ZO-1 (Furuse et al., 1994;Fanning et al., 1998). Accordingly, we observe here that stabilization of the F-actin cytoskeleton of Caco-2/TC7 cells by JAS treatment before infection, abrogates the modification of occludin distribution thus indicating that the G. intestinalis-induced rearrangement of occludin at TJs, and the G. intestinalis-induced disassembly of apical F-actin cytoskeleton are related events. Alteration of the intestinal barrier as a result of the modification of structural ZO proteins and structural/functional occludin has been reported for both bacterial (Peiffer et al., 2000a;Simonovic et al., 2000;Sakaguchi et al., 2002;Guignot et al., 2007) and viral (Obert et al., 2000;Beau et al., 2007) enterovirulent pathogens infecting cultured human intestinal T84 or Caco-2 cells forming monolayers.
It has previously been reported that a decrease in the distribution of claudins develops in the intestine of patients with chronic giardiasis (Troeger et al., 2007). Claudins, which colocalizes with occludin at the cell membrane, form the backbone of the protein filaments in TJs (Anderson and Van Itallie, 2009;Tepass, 2003). There is evidence that enterovirulent bacteria are able to alter the distribution and properties of claudins, and thus to affect the intestinal barrier (Sakaguchi et al., 2002;Muza-Moons et al., 2004). Here, we report that in G. intestinalisinfected Caco-2/TC7 cells, claudin-1 disappeared from the TJs at the cell-to-cell contact points and relocalized to aggregates located in the central regions of the cells. As for occludin, this rearrangement of claudin-1 results directly from the adhesion of trophozoites, since inhibiting the adhesion by pre-treatment with MBCD, abolishes the delocalization of claudin-1. In contrast to that of occludin, we found that the G. intestinalis-induced delocalization of claudin-1 from TJs did not depend on the G. intestinalisinduced disassembly of apical F-actin cytoskeleton, since JAS treatment did not prevent the formation of claudin-1 aggregates. Aggresomes are cellular structures formed in response to misfolded proteins (Johnston et al., 1998). Aggresome-like structures have been observed in Salmonella enterica serovar Typhimurium-infected epithelial cells and macrophages with dramatic remodelling of the networks of cytoplasmic intermediate filament proteins, vimentin and cytokeratin (Guignot and Servin, 2008), and aggresome-like structures in Herpes simplex virus-2infected Vero cells (Nozawa et al., 2004). It remains to be determined: (i) whether G. intestinalis-induced claudin-1 aggregates are aggresomes or not, and (ii) whether other claudins are delocalized or not in response to G. intestinalis infection.
To summarize, our results show that the adhesion of G. intestinalis trophozoites to cultured human enterocytelike Caco-2/TC7 cells promotes the profound disassembly of the brush border-associated F-actin cell cytoskeleton that in turn leads to a dramatic modification of the distribution of several intestinal brush border-associated enzymes and transporters. In addition, adhesion-and cytoskeleton-dependent changes in TER and occludin distribution are observed in G. intestinalis-infected enterocyte-like cells, whereas the change in claudin-1 distribution was adhesion-dependent but cytoskeletonindependent. Similar losses of intestinal microvilli, cytoskeleton-dependent alterations in the distribution and activity of brush border-associated functional proteins, and disruption of epithelial TJs have been observed triggered by bacterial enterovirulent pathogens including attaching-effacing enterohaemorrhagic or enteropatho-genic E. coli and diffusely adhering E. coli, and Shigella flexneri (Guttman and Finlay, 2008b). Our in vitro observations do not rule out that structural or functional lesions at the brush border of enterocytes can take place in vivo following an immune response against G. intestinalis infection or the development of cellular caspasedependent apoptosis as reported by elsewhere (Müller and von Allmen, 2005;Buret, 2007;2008).

Culture of trophozoites
Giardia intestinalis strain GS/H7 (ATCC 50581) was purchased from the American Type Culture Collection. Clones 1267 and C6 of the WB strain of G. intestinalis were kindly provided by Hugo Luján (University of Córdoba, Argentina). Trophozoites were grown in Keister's modified TYI-S-33 medium (Keister, 1983) supplemented with 15 ml per litre of a solution containing 1000 IU ml -1 penicillin and 1000 g ml -1 streptomycin (Gibco-BRL/Life Technologies, Cergy, France). pH of cultures was adjusted to 6.9 before filter sterilization (0.22 mm). Parasites were cultured in 25 cm 2 cell culture flasks, and harvested as previously described .

Treatments
Jasplakinolide (1 mM) was added to the culture medium of the Caco-2/TC7 cells 30 min before trophozoite infection and maintained during the infection (Peiffer et al., 2001). At the concentrations used, JAS and MBCD did not affect the integrity or viability of the Caco-2/TC7 cells (not shown). In order to investigate the role of lipid rafts in the co-culture of trophozoites and Caco-2/TC7 cells, the cells were pre-treated with MBCD (10 mM) 1 h before trophozoite infection, and MBCD was maintained throughout the 3 h infection period. In order to investigate the role of the lipid rafts of Caco-2/TC7 cells, the cells were pre-treated with MBCD (10 mM) 1 h before trophozoite infection, washed twice with DMEM and then infected. In order to investigate the role of the lipid rafts of G. intestinalis, the trophozoites were pre-treated with MBCD (10 mM) for 1 h and washed twice with PBS before being infected. There were no changes in the morphology, motility or proliferation of the trophozoites after MBCD treatment (not shown).

Attachment assay
The attachment assay was carried out as previously described (Sousa et al., 2001). Postconfluence cell monolayers were washed twice with sterile PBS before being infected. Cultures of G. intestinalis trophozoites were decanted, and any remaining attached trophozoites were detached by chilling for 10 min in ice-cold PBS, at pH 7.2. Trophozoite suspensions were centrifuged at 1000 g for 10 min. The supernatant was discarded, and the pellet was washed once with ice-cold PBS. The pellet was then suspended in DMEM without fetal calf serum (DMEMadhesion medium). Trophozoite counts were done using a haemocytometer. Suspensions of trophozoites, containing 5 ¥ 10 5 , 1 ¥ 10 6 , 2 ¥ 10 6 or 8 ¥ 10 6 trophozoites ml -1 (moi 0.5, 1, 2 and 8 respectively), were then co-incubated with cultured cells. Plates were incubated at 37°C in a 10% CO2-90% air atmosphere. After incubating for 3 h, unattached trophozoites were removed by gently rinsing the culture plates three times with DMEM-adhesion medium at 37°C. Adhering trophozoites were then recovered by incubating at 4°C for 10 min in cold DMEMadhesion medium. Trophozoite suspensions were counted in a haemocytometer. Assays were conducted in triplicate, with three successive passages of Caco-2/TC7 cells.

Measurement of cell integrity
Cell detachment, necrosis and the release of LDH activity were assessed as reported elsewhere  to evaluate cell integrity and viability.

Measurement of TER
Monolayers of Caco-2/TC7 cells were grown on filters (0.4 mm polyester, tissue culture-treated, Transwell Costar). After apical infection, the integrity of the confluent polarized monolayer was assessed by measuring the TER with a volt-ohmmeter (Millicell-ERS; Millipore, Saint Quentin, France). The background reading for a control filter was subtracted, and results were expressed as percentages of the value for uninfected control cells.

Immunofluorescence
Labelling was conducted as previously reported (Peiffer et al., 2000b) on cells cultured on round glass coverslips (Karl Hecht, 97647 Sondheim, Germany). Briefly, infected cells were washed twice with PBS buffer and fixed for 15 min at room temperature in PBS-3% (w/v) in paraformaldehyde. Monolayers were washed three times with PBS, and then exposed to 50 mM NH4Cl for 10 min. For cell permeabilization, the coverslips were incubated for 4 min with PBS-0.2% Triton X-100. F-actin labelling was carried out by incubating with FITC-phalloidin (Molecular Probes, Eugene, OR) for 45 min at room temperature. Indirect immunofluorescence labelling of fixed cells was conducted for brush border-associated enzymes (SI, DPPIV and GLUT5), for the transferrin receptor, TJ proteins (occludin and claudin-1) and the adherens junction protein (E-cadherin). Anti-SI, anti-DPPIV and anti-claudin-1 antibodies were diluted 1:50. The anti-GLUT5 antibody was diluted 1:100. Anti-Rc transferin and anti-occludin antibodies were diluted 1:100; and the anti-E-cadherin antibody was diluted 1:300. All dilutions were performed in 0.2% (v/v) gelatin-PBS. The cells were incubated for 45 min at room temperature with the primary antibody. After three washes with PBS, the cells were incubated for another 45 min in the dark and at room temperature with the secondary antibody conjugated with FITC or TRITC diluted 1:200 in 0.2% gelatin-PBS. After exhaustive washing, samples were mounted in DakoCytomation fluorescent mounting medium for immunofluorescence examination. The coverslips were examined by conventional epifluorescence microscopy using a Leitz Aristoplan microscope. No fluorescence was detected when the primary antibody was omitted. Confocal analysis was performed using a CLMS (model LSM 510 Zeiss, equipped with an air-cooled, 488 nm argon ion laser, and a 543 nm helium neon laser) configured with an Axiovert 100M microscope using a Plan Apochromat 63 ¥/1.40 oil objective. Images obtained by CLMS were analysed using Imaris software (version 6.21) (Bitplane, Zurich).

Western blot analysis
SDS-PAGE gels were transferred onto a PVDF membrane (Immobilon-P; Millipore). Mouse anti-occludin (1/500) and rabbit anti-claudin-1 (1/250) antibodies were used to detect the TJ proteins. A secondary anti-rabbit or anti-mouse horseradish peroxidase-conjugated antibody was also used. A chemiluminescence analysis was then performed (ECL system Millipore). Quantification was carried out using the Scion Image processor software.

Enzyme assay
The cells were washed in ice-cold PBS, scraped off, suspended in PBS and homogenized. Sucrase activity was measured in the enriched brush border membrane fraction obtained after centrifuging the cell homogenates for 1 h at 100 000 g and at 4°C as previously described (Peiffer et al., 2001). Briefly, sucrase activity was assayed using a glucose oxidase/peroxidase reagent that contains 4-amino-antipyrine. Sucrase activity was determined by measuring the amount of glucose liberated from the sucrose. One unit is defined as the amount of enzyme that hydrolyses 1 mM of substrate min -1 at 37°C. Enzyme-specific activity is expressed as milliunits (mU) per mg of protein. Proteins were determined by the BCA assay.

Statistical analysis
All experiments were conducted in triplicate. The results are expressed as the mean Ϯ standard deviation of the mean (SD). Student's t-test was used for statistical comparisons.