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Abstract

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Mutations in ATP8B1 cause familial intrahepatic cholestasis type 1, a spectrum of disorders characterized by intrahepatic cholestasis, reduced growth, deafness, and diarrhea. ATP8B1 belongs to the P4 P-type adenosine triphosphatase (ATPase) family of putative aminophospholipid translocases, and loss of aminophospholipid asymmetry in the canalicular membranes of ATP8B1-deficient liver cells has been proposed as the primary cause of impaired bile salt excretion. To explore the origin of the hepatic and extrahepatic symptoms associated with ATP8B1 deficiency, we investigated the impact of ATP8B1 depletion on the domain-specific aminophospholipid translocase activities and polarized organization of polarized epithelial Caco-2 cells. Caco-2 cells were stably transfected with short hairpin RNA constructs to block ATP8B1 expression. Aminophospholipid translocase activity was assessed using spin-labeled phospholipids. The polarized organization of these cells was determined by pulse-chase analysis, cell-fractionation, immunocytochemistry, and transmission electron microscopy. ATP8B1 was abundantly expressed in the apical membrane of Caco-2 cells, and its expression was markedly induced during differentiation and polarization. Blocking ATP8B1 expression by RNA interference (RNAi) affected neither aminophospholipid transport nor the asymmetrical distribution of aminophospholipids across the apical bilayer. Nonetheless, ATP8B1-depleted Caco-2 cells displayed profound perturbations in apical membrane organization, including a disorganized apical actin cytoskeleton, a loss in microvilli, and a posttranscriptional defect in apical protein expression. Conclusion: Our findings point to a critical role of ATP8B1 in apical membrane organization that is unrelated to its presumed aminophospholipid translocase activity, yet potentially relevant for the development of cholestasis and the manifestation of extrahepatic features associated with ATP8B1 deficiency. (HEPATOLOGY 2010)

Mutations in ATP8B1 cause familial intrahepaticcholestasis type 1 or ATP8B1 deficiency, asevere liver disease characterized primarily by impaired bile salt secretion from liver into bile.1 ATP8B1 deficiency was originally described as progressive intrahepatic cholestasis type 12 and benign recurrent intrahepatic cholestasis type 1.3 Clinically, this disorder is characterized by cholestasis, and it may cause progressive liver scarring. Patients with ATP8B1 deficiency also suffer from multiple extrahepatic symptoms, which include intractable diarrhea, malabsorption, pancreatitis, hearing loss, and growth retardation.4 Importantly, these extrahepatic symptoms are not fully resolved after orthotopic liver transplantation.4, 5 The expression of ATP8B1 is not restricted to the canalicular membrane of hepatocytes6, 7 but extends to many different epithelial cell types.7-9 This implies that, besides the role of ATP8B1 in bile salt secretion, the overall pathophysiology of ATP8B1 deficiency is the result of a more general cellular defect.

ATP8B1 is a member of the P4-subfamily of P-type adenosine triphosphatases (ATPases) (P4-ATPases). This subfamily comprises 14 human proteins, several of which have been linked to human disease.10 P4-ATPases have been implicated in the catalysis of inward aminophospholipid (APL) translocation. This activity is essential for the creation and maintenance of APL asymmetry in membranes of eukaryotic cells.11-16 Loss of the yeast Saccharomyces cerevisiae P4-ATPases Dnf1p and Dnf2p blocks the non-endocytic uptake of fluorescent 7-nitro-2-1,3-benzoxadiazol-4-yl–labeled APL analogs across the plasma membrane and causes aberrant exposure of APLs at the cell surface.17, 18 Similarly, heterologous expression of ATP8B1 has been shown to stimulate the uptake of NBD-labeled phosphatidylserine (PS) in a mutant Chinese hamster ovary cell line defective in PS transport.7, 19 A role of ATP8B1 as APL translocase (APLT) in the canalicular membrane of hepatocytes is further substantiated by the luminal accumulation of NBD-PS in ATP8B1-deficient hepatocytes20 and the enhanced recovery of PS in bile from Atp8b1G308V/G308V mutant mice after infusion of taurocholate.21 Based on these and other findings, it has been postulated that ATP8B1 dysfunction causes phospholipid randomization in the apical membrane of polarized cell types such as enterocytes and hepatocytes. In hepatocytes this would sensitize the canalicular membrane to enhanced extraction of cholesterol by hydrophobic bile salts, leading to intrahepatic cholestasis.21

Human intestinal epithelial Caco-2 cells, as a model of polarized epithelium in vivo, have been used extensively to investigate the mechanisms involved in maintaining cell surface polarity. To further explore the origin of the apical membrane defects associated with ATP8B1 deficiency, we investigated the impact of blocking ATP8B1 expression on the domain-specific APLT activities as well as on the functional organization of polarized Caco-2 cells.

Materials and Methods

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Cell Culture and RNAi.

Caco-2 cells were cultured in Dulbecco's modified Eagle's medium. Lipofectamine2000 was used to stably transfect Caco-2 cells with pSUPER.retro empty vector (Dr. R. Agami, The Netherlands Cancer Institute, Amsterdam, The Netherlands) or pSUPER.retro containing ATP8B1-specific RNAi targeting sequences (see Supporting Information Methods). Clonal selection of stably transfected cells was performed in medium containing 4.5 μg/mL puromycin dihydrochloride (Sigma-Aldrich). Cells were passaged maximally 10 to 15 times after clonal selection. Permanent knockdown of ATP8B1 expression was routinely verified by immunoblot analysis after each passage to exclude the possibility that the knockdown is transient. All experiments were replicated in at least three independently-generated ATP8B1 knockdown clones and in two independent empty vector control clones to exclude potential clonality artifacts.

Quantitative Reverse Transcription Polymerase Chain Reaction.

RNA was isolated from Caco-2 cells in biological quadruplicates. Quantitative reverse transcriptase PCR (qRT-PCR) reactions were performed as described in Supporting Information Methods.

Immunoblotting and Pulse-Chase Analysis.

Caco-2 cells were lysed in ristocetin-induced platelet aggregation buffer. Sodium dodecyl sulfate polyacrylamide gel electrophoresis, immunoblotting, pulse-chase analysis, and antibody incubation are described in Supporting Information Methods.

Confocal Microscopy.

Cells grown on glass coverslips for 14 days post-confluent (dpc) were fixed, labeled with primary antibodies, counterstained with Alexa-conjugated secondary antibodies, and imaged by a confocal microscope (D-eclipse C1; Nikon) as described in Supporting Information Methods.

Subcellular Membrane Fractionation.

Cells were seeded on permeable Transwell filter supports (0.4-μm pore size, Costar, the Netherlands) and grown 11 to 14 dpc. Cells were biotinylated on either the apical or basolateral side, and membranes were fractionated by sucrose-step gradient centrifugation as described in Supporting Information Methods. Fractions were subjected to sodium dodecyl sulfate polyacrylamide gel electrophoresis. Biotinylated proteins were stained using horseradish peroxidase-conjugated Neutravidin (Invitrogen).

Lipid Transport Assay on Suspension Cells and Filter-Grown Cells.

Lipid transport assays were performed essentially as described,22 with some modifications as detailed in Supporting Information Methods. Briefly, cells grown on Transwell filter supports for 11 dpc were labeled apically or basolaterally with spin-labeled phospholipid at 4°C. Filters were incubated at 10°C for 0 to 60 minutes, and back-extraction by 4% bovine serum albumin was performed for 10 minutes at 4°C. Lipid analogs were reoxidized and quantified as described.22

Drug Sensitivity Assay.

To determine PS or phosphatidylethanolamine (PE) exposure in the outer leaflet of the plasma membrane cells grown to 14 dpc in 96-well plates were incubated with papuamide B (Flintbox) or cinnamycin (Sigma-Aldrich) for 15 minutes, respectively. As a measure for membrane damage induced by papuamide B or cinnamycin, the lactate dehydrogenase release in the medium was determined. Control cells were either incubated with medium supplemented with dimethylsulfoxide (solvent for papuamide and cinnamycin) or in medium containing 1% Triton-X100. Experiments were performed in quadruplicate.

Electron Microscopy.

Cells were seeded on polytetrafluoroethylene membranes (Costar), differentiated for 7 or 14 dpc, fixed in glutaraldehyde, postfixed in 1% OsO4, and embedded in Epon. Ultrathin sections (80 nm) were stained with uranyl acetate and lead citrate and examined using a Jeol 1010 transmission electron microscope as described in Supporting Information Methods.

Results

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

ATP8B1 Is an Abundant P4 -ATPase in the Apical Membrane of Caco-2 Cells.

To validate Caco-2 cells as an appropriate model system for investigating ATP8B1 function at the apical membrane, we first demonstrated that ATP8B1 and CDC50A, an interacting protein required for plasma membrane localization of ATP8B1,19 are expressed in these cells (Supporting Information Fig. 1). Subsequent qRT-PCR experiments showed that ATP8B1 is among the most abundantly expressed P4-ATPases, regardless of the differentiation status of the cells (Fig. 1A). Immunoblotting showed that ATP8B1 protein levels gradually rise up to 13-fold during differentiation (Fig. 1B). When differentiated cells were treated with cycloheximide, ATP8B1 levels only declined 1.3-fold after 4 days of treatment. In contrast, COMMD1, a binding partner of the copper transporting P-type ATPase ATP7B,23 was completely turned over within 2 days (Fig. 1C). Confocal immunofluorescence microscopy showed that ATP8B1 colocalizes extensively with the apical membrane protein aminopeptidase N (CD13), but not with the lateral adherence marker E-cadherin (Fig. 1D). The apical localization of ATP8B1 was verified by subcellular fractionation on sucrose density gradients after side-specific biotinylation of filter-grown Caco-2 cells. Apical membranes (peak fractions 11, 12) segregated completely from basolateral membranes (peak fractions 2, 3; Fig. 1E). The fractionation profile of ATP8B1 corresponded to that of the biotinylated apical proteins and the apical protein intestinal alkaline phosphatase (AP). In contrast, fractions containing the bulk of basolateral proteins E-cadherin (E-cad) and Na+/K+-ATPase were devoid of ATP8B1 (Fig. 1F). Together, these results reveal that ATP8B1 is a stable and abundantly expressed P4-ATPase in the apical membrane of differentiated Caco-2 cells, consistent with its expression in apical membranes of epithelial cells in vivo.8

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Figure 1. ATP8B1 is an abundantly expressed P4-ATPase in the apical membrane of Caco-2 cells. (A) Relative mRNA expression levels of ATP8B1 and five most closely related P4-ATPases in undifferentiated (0 days postconfluent, 0 dpc) and differentiated (14 dpc) Caco-2 cells were quantified by qRT-PCR. Data were normalized relative to ATP8B1 expression at 0 dpc. Error bars: SD of four biological replicates. (B) Cell lysates harvested at 0, 5, 11, or 14 dpc were subjected to immunoblotting using antibodies against ATP8B1 and tubulin. ATP8B1 and tubulin protein levels were quantified by densitometry. ATP8B1 signal was corrected for the tubulin loading control (average of two experiments) and presented relative to 0 dpc. (C) Differentiated Caco-2 cells were incubated with cycloheximide for the indicated time periods and subjected to immunoblotting using antibodies against ATP8B1, tubulin, and COMMD1. Protein bands were quantified as in (B) (average of three experiments) and presented relative to 0 days of cycloheximide treatment. (D) Confocal microscopy images (X,Y versus Z-stacks) of cells stained with antibodies against ATP8B1, CD13, and E-cadherin (E-cad). Bar, 10 μm. (E,F) Filter-grown Caco-2 cells were biotinylated on the apical or basolateral side and analyzed by sucrose density gradient fractionation. Fractions were subjected to immunoblotting using horseradish peroxidase-conjugated Neutravidin (E) or antibodies against ATP8B1, intestinal alkaline phosphatase (AP), Na+/K+-ATPase, and E-cad (F).

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Aminophospholipid Transport and Asymmetry Is Unperturbed in ATP8B1-Depleted Caco-2 Cells.

To mimic ATP8B1 deficiency in a polarized epithelial cell model, we created Caco-2 cell lines stably transfected with one of two different ATP8B1 short hairpin RNA constructs (Fig. 2A). Several independently isolated clones (that is, clones A3, A5, B3) showed a marked 70% to 90% reduction in ATP8B1 protein levels relative to empty vector (EV) control cell lines (clones EV1, EV2; Fig. 2B). This reduction of ATP8B1 protein expression was maintained during at least 15 passages after clonal selection (data not shown). Furthermore, knockdown of ATP8B1 did not induce compensatory expression of the most closely related P4 ATPases as assessed by qRT-PCR, nor did it affect the messenger RNA (mRNA) expression of CDC50A (Fig. 2C). The clonal cell line A5 showed the most pronounced (approximately 90%) reduction in ATP8B1 levels. Therefore, most experiments presented in this work concern the A5 clone, but we excluded potential transfection and clonality artifacts by replication of all experiments in clones A3 and B3, using both EV1, EV2, and wild-type cells as controls. Quantitative RT-PCR analysis of intestinal differentiation markers c-myc and villin24 indicated that all clonal cell lines retained the ability to undergo normal epithelial differentiation (Fig. 2D).

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Figure 2. Establishment of ATP8B1-depleted Caco-2 cell lines. (A) Position of RNAi target sequences A and B in the coding region and 3′ UTR of the ATP8B1 mRNA, respectively. (B) Total protein extracts of clonal Caco-2 cell lines stably transfected with ATP8B1-targeting short hairpin RNAs (clones A3, A4, A5, B1, B2, B3) or empty vector (clones EV1, EV2) were subjected to immunoblotting using antibodies against ATP8B1 and actin. (C) Relative mRNA levels of ATP8B1 and the five most closely related P4 ATPases and of the P4 ATPase beta subunit CDC50A in differentiated Caco-2 cells (CDC50B is not expressed in Caco-2 cells). ATP8B1 qRT-PCR primers amplify a region in the 5′ end of the coding region, which explains the relative minor reduction in ATP8B1 mRNA level in the B3 clone (A). Data were normalized relative to ATP8B1 expression in EV1 cells. Error bars: SD of four biological replicates. (D) Relative mRNA levels of villin and c-myc in undifferentiated (0 dpc) and differentiated (14 dpc) cell lines were quantified by qRT-PCR. Data were normalized relative to expression in clone EV1 at 0 dpc. Error bars: SD of four biological replicates.

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Because ATP8B1 is believed to possess APL flippase activity,7, 19 we analyzed the impact of RNAi-mediated depletion of ATP8B1 on the movement of spin-labeled phospholipids from the outer to the inner leaflet of the plasma membrane by using back-exchange to serum albumin.22 Initial experiments were performed with cells in suspension at reduced temperatures to suppress hydrolysis of the spin-labeled phospholipid into lyso-derivatives and free fatty acids (consistently <10%) and to prevent endocytic uptake. When incubated at 20°C, EV1 cells displayed a fast inward movement of SL-phosphatidylserine (SL-PS) and SL-phosphatidylethanolamine. In contrast, SL-phosphatidylcholine was internalized at a considerably lower rate, whereas SL-sphingomyelin (SL-SM) was hardly taken up at all (Supporting Information Fig. 2A). The lipid transport kinetics in A5 cells were indistinguishable from those in EV1 cells, also when internalization rates were reduced by lowering the temperature to 10°C. Under these conditions, a Chinese hamster ovary-K1 mutant cell line defective in PS transport25 displayed a marked reduction in SL-PS uptake compared with its wild-type counterpart (Supporting Information Fig. 2B).

To discriminate between lipid transport activities in apical and basolateral membranes, we measured the kinetics of lipid uptake in differentiated Caco-2 cells grown on filters. Spin-labeled phospholipid analogs were never found in the bovine serum albumin medium added to the nonlabeled membrane domain, indicating the presence of intact tight junctions. As shown in Fig. 3A, SL-PS was efficiently internalized at the apical and basolateral membranes of EV1 cells, whereas the transbilayer movement of SL-SM was slow. The SL-PS uptake rates at the apical and basolateral membranes of A5 cells were essentially the same as those in EV1 cells. To verify the apparent lack of any perturbation in APLT kinetics, differentiated wild-type, EV1, and all ATP8B1-depleted cell lines were incubated with different concentrations of papuamide B and cinnamycin, two cytolytic peptides that require binding to cell surface-exposed PS or PE, respectively, to exert their cytotoxicity.17, 26 As shown in Supporting Information Fig. 3 and in Fig. 3B and 3C, all cell clones were equally sensitive to these apically administered, APL-binding drugs. Together, these results indicate that ATP8B1 is largely dispensable for maintenance of APL flippase activity and APL asymmetry in the apical membrane of differentiated Caco-2 cells, although we cannot exclude the possibility that residual levels of ATP8B1 may be sufficient to sustain normal APL transport and phospholipid asymmetry.

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Figure 3. Aminophospholipid transport and asymmetry in the apical membrane of ATP8B1-depleted Caco-2 cells is unperturbed. (A) Differentiated EV1 or A5 cells grown on filters were incubated with SL-PS or SL-SM on either the apical or the basolateral side at 10°C. The amount of SL-phospholipid present in the exoplasmic leaflet was assessed by back exchange to bovine serum albumin and expressed as the percentage of total label added at time zero. SL-PS, SL-phosphatidyl serine; SL-SM, SL-sphingo myelin. Differentiated Caco-2 cells were incubated with various concentrations of the PS-binding peptide papuamide B (B) or with various concentrations of the PE-binding peptide cinnamycin (C). Cytotoxic membrane damage was measured by determining the amount of lactate dehydrogenase release in the medium. Experiments were performed in quadruplicate, and one representative experiment out of four and one out of two are shown, respectively for papuamide B and cinnamycin. Error bars represent SD of quadruplicates.

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ATP8B1-Depleted Caco-2 Cells Display a Posttranscriptional Defect in Apical Protein Expression.

Because P4-ATPase dysfunction is often accompanied by perturbations in membrane trafficking from and to the plasma membrane,16, 18 we investigated whether loss of ATP8B1 affects the polarized delivery of membrane proteins in differentiated Caco-2 cells. As shown in Fig. 4, the bulk of basolateral proteins could be separated from apical proteins by density gradient centrifugation. Control (untransfected) cells and A5 cells essentially yielded the same profiles of apically and basolaterally biotinylated proteins. However, apical membranes isolated from A5 cells consistently migrated to a lower density area (compare Figs. 1E and 4A), suggesting that loss of ATP8B1 affects the global composition of these membranes. Immunofluorescence microscopy showed that, although the apical proteins AP, CD13, and sucrase-isomaltase (SI) retained their normal polarized distribution in A5 cells, their labeling intensity was greatly diminished compared with EV1 or untransfected Caco-2 cells (Fig. 5A/B; data not shown). A similar reduction in apical staining of these proteins was observed in A3 and B3 cells. Immunoblot analysis confirmed that AP and SI levels in differentiated A5, A3, and B3 cells were threefold to 10-fold lower than in EV1 or untransfected Caco-2 cells (Fig. 6A; data not shown). In contrast, the expression levels of the basolateral membrane proteins Na+/K+-ATPase and E-cadherin, or the tight junction protein ZO-1 were not affected. Hence, loss of ATP8B1 does not disrupt the polarized delivery of membrane proteins in differentiated Caco-2 cells, but rather causes a general defect in apical protein expression. This defect seems to occur at the posttranscriptional level, as ATP8B1-depleted and control cells contain similar amounts of AP-encoding mRNA (Fig. 6B). The reduced apical levels of AP, CD13, and SI were not accompanied by an increased intracellular staining (Fig. 5B) or enhanced recovery of these proteins from the apical medium (data not shown). This indicates that the defect in apical protein expression is unlikely because of perturbations in polarized membrane trafficking or increased shedding of the apical membrane.

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Figure 4. Apical and basolateral proteins retain their polarized distribution in ATP8B1-depleted Caco-2 cells. (A, B) Filter-grown A5 cells were biotinylated on the apical or basolateral side and analyzed by sucrose density gradient fractionation. Fractions were subjected to immunoblotting using horseradish peroxidase-conjugated Neutravidin (A) or antibodies against AP, Na+/K+-ATPase, and E-cad (B). Peak fractions from control gradients—indicated by asterisks in Fig. 1E, F—are marked as ‘control.’ (C) Refraction indices representing density profiles of the sucrose gradients used for fractionation of control (Fig. 1E, F) and A5 (Fig. 4A, B).

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Figure 5. ATP8B1-depleted Caco-2 cells display a reduced staining of apical membrane markers. (A) Confocal microscope cross-sections (Z-stacks) of differentiated EV1 and A5 cells stained with antibodies against E-cad and AP and aminopeptidase N (CD13). Confocal images of all samples in panel A were taken using the same confocal microscope settings. Bar, 10 μm. (B) Confocal microscopy projections of EV1 and ATP8B1-depleted cells (A3, A5, B3) stained with antibodies against ATP8B1, AP and CD13, E-cad, and ZO-1. (B) The confocal microscope settings were adjusted per sample for optimal detection. Bar, 20 μm.

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Figure 6. ATP8B1-depleted Caco-2 cells display a posttranscriptional defect in apical protein expression. (A) Differentiated EV1 and ATP8B1-depleted cells (A3, A5, B3) were subjected to immunoblotting using antibodies against ATP8B1, AP, sucrase-isomaltase (SI), Na+/K+-ATPase, E-cad, ZO-1, and actin. (B) Relative mRNA levels of AP and E-cad in undifferentiated (0 dpc) and differentiated (14 dpc) cell lines were quantified as in Fig. 1A. Data were normalized relative to AP and E-cad expression in clone EV1 at 0 dpc. Error bars: SD of four biological replicates.

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To further address the origin of this defect, we monitored the de novo synthesis and turnover of the apical proteins AP and SI by pulse-chase immunoprecipitation analysis. The level of AP biosynthesis was drastically (approximately 15-fold) reduced in differentiated A5 cells compared with EV1 cells (Fig. 7A; data not shown). A5 cells also displayed a substantial (approximately threefold) reduction in the level of SI biosynthesis. Similar reductions in apical protein biosynthesis were observed in other ATP8B1-depleted clones, with the degree of reduction showing a good correlation with the level of ATP8B1 depletion in these clones (Fig. 7B). In contrast, biosynthesis of E-cad was not affected. Whereas loss of ATP8B1 greatly reduced AP biosynthesis, the AP turnover rate appeared very similar between ATP8B1-depleted and control cells (Fig. 7A). Addition of the proteasome inhibitor MG132 had no effect on the levels of radiolabeled AP recovered from A5 cells (Fig. 7C). Next, we considered the possibility that this specific defect in apical protein expression was caused by activation of the unfolded protein response secondary to the depletion of ATP8B1. Using RT-PCR, we found no evidence of XBP1 splicing. Moreover, immunoblot analysis showed that these cells contained similar levels of the ER chaperones BiP and GRP94 as EV1 control cells (Supporting Information Fig. 4). Together, these findings indicate that loss of ATP8B1 in polarized Caco-2 cells causes a specific post-transcriptional defect in the expression of apical proteins, presumably at the level of mRNA translation.

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Figure 7. ATP8B1-depleted Caco-2 cells are defective in the de novo synthesis of apical proteins. (A) Differentiated EV1 and A5 cells were labeled using [35S]-methionine/cysteine for 20 minutes, and then chased for the indicated time periods. E-cad, SI, and AP were immunoprecipitated from cell extracts or the chase medium, as indicated. Immunoprecipitates were resolved by sodium dodecyl sulfate polyacrylamide gel electrophoresis, and radiolabeled proteins were visualized by phosphorimager screens. (B) Quantification of radiolabeled AP and E-cad levels in 20-minutes pulsed EV1 and ATP8B1-depleted cells (A3, A5, B3). (C) EV1 and A5 cells were incubated with the indicated concentrations of MG132 for 16 hours before pulse-chase analysis, as in (A). p, precursor; m, mature.

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ATP8B1 Is Required for Apical Brush Border Formation.

The perturbed expression of apical membrane proteins in ATP8B1-depleted cells suggested a possible defect in the assembly of the apical brush border.24, 27 Contrary to differentiated control cells (EV1, untransfected), ATP8B1-depleted clones (A5, A3, B3) generally lacked the patchy apical staining pattern indicative of microvilli (Fig. 8A). Microvilli formation correlates with the polymerization of F-actin at the core of the microvilli. Phalloidin staining of F-actin showed the presence of clearly visible actin bundles in the apical domain of differentiated EV1 or untransfected Caco-2 cells. However, these bundles were largely absent or seemed much shorter in all three differentiated ATP8B1-deficient clones (Fig. 8B; data not shown). We next analyzed the ultrastructural organization of the apical domain by transmission electron microscopy. Control cells (EV1, untransfected) cultured for 7 dpc on filters formed homogenous monolayers of cuboidal cells rich in microvilli (Fig. 8C, panels 1, 3, 5; data not shown). Their nuclei were invariably positioned in proximity to the basal membrane. In contrast, ATP8B1-depleted cells (A5, A3, B3) formed highly irregular layers of flat cells with far less and much shorter microvilli (Fig. 8C, panels 2, 4, 6; data not shown). Typically, less than 50% of the apical surface of these cells was covered with microvilli. However, like control cells, ATP8B1-depleted cells contained well-developed tight junctions and desmosomes (compare panels 5 and 6 in Fig. 8C), confirming that ATP8B1-depleted Caco-2 cells retain the ability to polarize. Extending the culture time to 14 dpc did not lead to any improvement in the number or length of the microvilli (Supporting Information Fig. 5). Together, this implies that loss of ATP8B1 results in a constitutive defect in apical brush border formation rather than a delay in this process.

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Figure 8. ATP8B1-depleted Caco-2 cells exhibit a disorganized apical actin cytoskeleton and are defective in microvilli formation. (A) Confocal microscopy projections of differentiated EV1 and ATP8B1-depleted cells (A5, B3) stained with antibodies against AP and CD13. Confocal microscope settings were adjusted per sample. (B) Phalloidin staining of F-actin and immunostaining of E-cad in EV1, A5, and B3 cells. Lower panels represent higher magnifications of the areas indicated by rectangles. Bars, 10 lm. (C) Transmission electron microscopy analysis of Eponembedded, 1-week filter-grown EV1 (panels 1, 3, 5) and A5 cells (panels 2, 4, 6). Tight junctions (arrowheads) and desmosomes (arrows) are indicated. MV, microvilli; N, nucleus; A, apical membrane; BL, basolateral membrane; M, mitochondrion. Bars, (1) 10 μm, (2) 5 μm, (3,4) 1 μm, (5,6) 500 nm

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Discussion

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Dysfunction of the putative APL transporter ATP8B1 causes ATP8B1 deficiency, a spectrum of disorders characterized by intrahepatic cholestasis and extrahepatic symptoms. Yet the pathophysiological mechanisms of ATP8B1 deficiency await elucidation. Here we show that blocking ATP8B1 expression in polarized Caco-2 cells results in a profound disorganization of the apical actin cytoskeleton, a substantial loss in microvilli, and a defect in the translation of mRNA species encoding apical membrane proteins. In contrast, APL transport and asymmetry across the apical bilayer are unperturbed. These findings provide evidence for a critical role of ATP8B1 in the apical membrane organization of polarized cells that is independent of its presumed APLT activity, yet potentially relevant for the clinical manifestations of ATP8B1 deficiency.

A function of ATP8B1 in APL transport is supported by the finding that expression of ATP8B1 restores the nonendocytic uptake of fluorescent PS-analogs in PS transport-defective Chinese hamster ovary mutant cells,7, 19 as well as by the enhanced recovery of PS in bile from Atp8b1G308V/G308V mutant mice21 and patients with ATP8B1 deficiency.28 Hence, it was unexpected that blocking ATP8B1 expression in differentiated Caco-2 cells had no measurable impact on the apically associated APLT activity, nor on the asymmetric distribution of APLs across the apical bilayer. How can these data be reconciled with the presumed flippase function of ATP8B1? Even though ATP8B1-related P4-ATPases are only poorly expressed in differentiated Caco-2 cells, it is feasible that these or other P4-ATPases can functionally compensate for the loss of ATP8B1. Alternatively, residual levels of ATP8B1 in RNAi-treated cells may be sufficient to sustain normal APL transport and asymmetry across the apical membrane.

Nonetheless, our data indicate that residual levels of ATP8B1 expression are clearly insufficient for normal apical membrane organization of ATP8B1-depleted Caco-2 cells. Whereas ATP8B1-depleted cells undergo epithelial differentiation and form fully developed tight junctions and desmosomes, less than 50% of their apical surface is covered with microvilli. Moreover, the microvilli are much shorter than in control cells. These changes are reminiscent of the morphological aberrations found in the canalicular membrane of liver cells from patients with ATP8B1 deficiency29 and Atp8b1G308V/G308V mutant mice when fed a cholate-supplemented diet.21 In addition, they provide a striking parallel with the loss of stereocilia from the apical surface of the inner hair cells in Atp8b1G308V/G308V mutant mice, which is associated with hearing loss.9 Collectively, these data point to a general function of ATP8B1 in the formation or stabilization of microvillar structures. The perturbed ultrastructural morphology of the apical domain in ATP8B1-depleted Caco-2 cells is not caused by a general defect in apical protein sorting or stability, but is accompanied by a global change in its molecular composition. This can be inferred from an altered fractionation profile on sucrose density gradients as well as from a marked reduction in the expression of multiple apical proteins.

Recent reports have proposed that the pathophysiology of ATP8B1 deficiency could be explained through decreased transcriptional activity of the nuclear Farnesoid X receptor.30 Our data indicate that loss of ATP8B1 causes a defect in the translation of apical mRNA rather than a block in gene transcription. This defect is specific, because expression of basolateral proteins is not affected. Interestingly, one ATP8B1-homolog in yeast, DRS2, was originally identified in a screen for mutants defective in ribosome assembly and function.31Drs2 null mutants are deficient in 40S ribosomal subunits, resulting in a reduced efficiency of translation. Whether ATP8B1 is also required for translational efficiency, or for mobilizing apical mRNAs from specific subcellular pools to which the translational machinery has no access,32 remains to be established.

A most striking result from our studies is that the critical function of ATP8B1 in apical membrane organization appears independent of its presumed APLT activity. This observation challenges a recently proposed model in which the morphological aberrancies and loss of ectoenzymes from the canalicular membrane in liver biopsies of Atp8b1G308V/G308V mutant mice are primarily ascribed to a disruption of APL asymmetry, which would render the canalicular membrane hypersensitive to extraction of membrane sterols and proteins by hydrophobic bile salts.21, 33 Our current findings therefore imply that ATP8B1 serves a dual role. Besides catalyzing APLT activity, we propose that ATP8B1 may provide a molecular scaffold in the apical membrane to recruit structural components or modulators of the actin cytoskeleton involved in microvilli formation. Consistent with a putative scaffold function, ATP8B1 expression on the apical membrane of Caco-2 cells increases dramatically during differentiation, concomitantly with cell polarization. Interestingly, loss of the P4-ATPase deficient for ribosomal subunits (Drs2p) disrupts the polarized organization of cortical actin filaments in yeast.34

ATP8B1 is not the only example of a P-type ATPase for which a transport-independent structural function has been proposed. Epithelial junction formation and tracheal tube-size control in Drosophila require a pump-independent function of the Na+/K+-ATPase.35 The catalytic α-subunits of this pump and the closely related H+/K+-ATPase interact with ankyrin, a cytoskeletal protein implicated in epithelial junction formation.36 Linking the cytoskeleton to specialized areas of the plasma membrane may thus be a feature shared among several members of the P-type ATPase superfamily. Disruption of cytoskeletal linkages between the actin bundle and the microvillar membrane results in a profound loss of SI.37 These and other data24 suggest that microvilli formation is tightly connected to apical protein biosynthesis. As microvilli dramatically increase the surface area of the apical membrane, this coupling may serve to prevent overloading of the apical membrane with protein. Thus, when formation of microvilli is perturbed, the translation of apical mRNAs is repressed. It appears likely that cross-talk between these two processes is mediated by proteins embedded in the apical membrane.19, 20, 36-39 The loss of microvilli associated with reduced ATP8B1 expression is reminiscent of morphological aberrations found in disorders such as celiac disease and microvillus inclusion disease, which are associated with a dramatic reduction of enterocyte apical membrane surface area and a concomitant reduction of absorptive capacity and diarrhea.37-39 Similarly, reduction of the apical surface area of hepatocytes would likely limit the bile salt excretory capacity of the liver in patients with ATP8B1 deficiency. Our findings thus provide a rationale for intrahepatic cholestasis as well as the intractable diarrhea and malabsorption as frequent symptoms associated with ATP8B1 deficiency.4 Collectively, these data point to a general function of ATP8B1 in the formation or stabilization of microvillar structures at the apical plasma membrane of polarized cell types and provide a novel framework for our understanding of the pathogenesis of familial intrahepatic cholestasis type 1.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

The authors thank Dr. J. Koenderink and Dr. J. Fransen, Radboud University Nijmegen, The Netherlands, and Dr. R. Agami, The Netherlands Cancer Institute, Amsterdam, The Netherlands, for sharing reagents. We acknowledge Rene Scriwanek and Marc van Peski for preparing the EM figures and Joep van den Dikkenberg for technical assistance.

References

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Supporting Information

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Additional Supporting Information may be found in the online version of this article.

FilenameFormatSizeDescription
HEP_23586_sm_suppinfo.doc74KSupporting Information
HEP_23586_sm_suppfig1.tif93KSupporting Figure 1
HEP_23586_sm_suppfig2.tif167KSupporting Figure 2
HEP_23586_sm_suppfig3.tif1776KSupporting Figure 3
HEP_23586_sm_suppfig4.tif269KSupporting Figure 4
HEP_23586_sm_suppfig5.tif4628KSupporting Figure 5

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