Advertisement
Research Article
Slp2-a controls renal epithelial cell size through regulation of Rap–ezrin signaling independently of Rab27
Takao Yasuda, Mitsunori Fukuda
Journal of Cell Science 2014 127: 557-570; doi: 10.1242/jcs.134056

ABSTRACT

Synaptotagmin-like protein 2 (Slp2-a/Sytl2) is a Rab27 effector protein that regulates transport of Rab27-bearing vesicles and organelles through its N-terminal Rab27-binding domain and a phospholipid-binding C2A domain. Here we demonstrate a Rab27-independent function of Slp2-a in the control of renal cell size through a previously uncharacterized C2B domain. We found that by recruiting Rap1 GAPs to the plasma membrane of MDCK II cells through the C2B domain, Slp2-a inactivates Rap signaling and modulates the size of the cells. Functional ablation of Slp2-a resulted in an increase in the size of MDCK II cells. Drosophila Slp Bitesize was found to compensate for the function of Slp2-a in MDCK II cells, thereby indicating that the mechanism of the cell size control by Slp proteins has been evolutionarily conserved. Interestingly, blockade of the activity of ezrin, a downstream target of Rap, with the glucosylceramide synthase inhibitor, miglustat, effectively inhibited cell spreading of Slp2-a-knockdown cells. We also discovered aberrant expression of Slp2-a and increased activity of ezrin in pcy (Nphp3pcy) mice, a model of polycystic kidney disease that is characterized by renal cell spreading. Our findings indicate that Slp2-a controls renal cell size through regulation of Rap–ezrin signaling independently of Rab27.

INTRODUCTION

Synaptotagmin-like protein 2 (Slp2-a; also known as Sytl2) is a member of the synaptotagmin-like protein family, whose members consist of an N-terminal Slp homology domain (SHD) and C-terminal tandem C2 domains (named the C2A domain and C2B domain) (Fukuda et al., 2001; Fukuda and Mikoshiba, 2001), and it was originally characterized as an effector molecule for the small GTPase Rab27 (Fukuda, 2006; Kuroda et al., 2002). In melanocytes, Slp2-a is localized to mature melanosomes and promotes their anchoring to the plasma membrane through interaction with Rab27 on the melanosomes through the SHD and with phospholipids in the plasma membrane through its C2A domain (Kuroda and Fukuda, 2004). Similarly, Slp2-a promotes docking of secretory vesicles to the plasma membrane in certain types of secretory cells (Holt et al., 2008; Ménasché et al., 2008; Saegusa et al., 2006). Slp2-a has recently been shown to be localized to the apical membrane of renal epithelial cells and to be involved in the regulation of cell signaling and tubulogenesis by promoting docking of Rab27-bearing podocalyxin-containing vesicles to the apical membrane (Gálvez-Santisteban et al., 2012; Yasuda et al., 2012). Based on these findings, Slp2-a is generally thought to be a key mediator of the docking of Rab27-bearing vesicles and organelles to the plasma membrane, and the docking process is thought to require both the Rab27 binding activity of the SHD and the phospholipid binding activity of the C2A domain. In contrast to the C2A domain, however, the physiological function of the C2B domain, which exhibits only weak phospholipid binding ability (Gálvez-Santisteban et al., 2012; Kuroda and Fukuda, 2004), has yet to be determined.

Although the Rab27 effector function of Slp2-a has been well established in the literature, several studies implicate a Rab27-independent role of Slp2-a in cell morphology. As an example, Slp2-a has been shown to be involved in generation of the elongated morphology of melanocytes, but the molecular mechanism by which it does so remain unknown (Kuroda and Fukuda, 2004). Another example is the involvement of the Drosophila Slp homologue Bitesize (Btsz), the only homologue of mammalian Slp that shows high sequence similarity to Slp2-a (Fukuda et al., 2001), in maintenance of epithelial cell morphology (Pilot et al., 2006; Serano and Rubin, 2003), although the precise mechanism here also remains unknown. Because Btsz protein lacks an N-terminal SHD, i.e. a Rab27-binding domain, Rab27 effector function must not have been retained during evolution, and we hypothesized the existence of a conserved function of Slp proteins in cell morphology.

In this study we investigated the possible involvement of Slp2-a in controlling the morphology of Madin–Darby canine kidney II (MDCK II) cells (a renal epithelial cell line) and showed that it regulates renal cell size through Rap–ezrin signaling by recruiting Rap1 GAPs to the plasma membrane through the C2B domain. Because Slp2-a is also expressed in the renal tubules of mice (Yasuda et al., 2012), we investigated expression of Slp2-a in pcy (Nphp3pcy) mice, a spontaneous model of polycystic kidney disease (PKD), that have a homozygous missense mutation (I614S) in the nephrocystin-3 (NPHP3) gene (Bergmann et al., 2008; Hildebrandt et al., 2009; Olbrich et al., 2003; Torres and Harris, 2009). Cystic kidney diseases, including autosomal dominant and autosomal recessive PKDs and nephronophthisis (NPHP), are a heterogeneous group of genetic disorders all of which are characterized by renal cell spreading, renal cyst growth, and progression to kidney failure (Hildebrandt et al., 2009; Torres and Harris, 2009; Wilson, 2004), but the precise mechanisms underlying the diseases are poorly understood. We discovered aberrant expression of Slp2-a and increased activity of ezrin in pcy mice. Our findings indicate that Slp2-a-mediated control of renal cell size is crucial for normal kidney function, and we discuss Slp2-a-mediated Rap–ezrin signaling as a potential target for the treatment of PKD.

RESULTS

Slp2-a regulates cell size in renal epithelial cells

First, we investigated the involvement of Slp2-a in renal cell proliferation by comparing the growth rates of control MDCK II cells and Slp2-a-knockdown (KD) MDCK II cells that we had established previously by using Slp2-a short hairpin RNA (shRNA) (Yasuda et al., 2012) (Fig. 1A). Although there was no difference in their growth rate (Fig. 1B), the Slp2-a-KD cells became confluent much faster than the control cells. We then investigated the morphology of Slp2-a-KD cells (Fig. 1C). Intriguingly, the Slp2-a-KD cells were significantly larger (approximately four times larger) than the control cells under subconfluent conditions. Even under polarized conditions Slp2-a-KD cells were larger (approximately 1.5 times larger than control cells) (see Yasuda et al., 2012), but since their phenotype was not so clearly distinguishable as the phenotype of the subconfluent cells, we decided to use subconfluent MDCK II cells to investigate the role of Slp2-a in cell size control in the subsequent analyses. The greater size of the Slp2-a-KD cells was unlikely to have been attributable to an off-target effect of the shRNA, because the same results were obtained by transient expression of three different Slp2-a small interfering RNAs (siRNAs) in MDCK II cells (supplementary material Fig. S1A,B) and because re-expression of an shRNA-resistant form of Slp2-a (Slp2-aSR) in the Slp2-a-KD cells restored them to normal cell size (Fig. 1D). Because Slp2-a was originally described as a Rab27 effector (Fukuda, 2006; Kuroda and Fukuda, 2004), we investigated whether the Rab27 binding ability of Slp2-a is required for control of cell size by using a Rab27-binding-deficient mutant, Slp2-a E11A/R32A (Kuroda and Fukuda, 2004). As shown in Fig. 1D, the Slp2-a mutant protein was localized to the plasma membrane, the same as the wild-type protein, and was able to restore the size of Slp2-a-KD cells. Consistent with these findings, functional ablation of Rab27A either by expression of a constitutively active mutant (Q78L) or negative mutant (T23N) of Rab27A (supplementary material Fig. S1D), or by knockdown of endogenous Rab27A expression (supplementary material Fig. S1E,F) had no effect on the size of the control MDCK II cells. Furthermore, the difference between the level of Rab27A expression in the control cells and the Slp2-a-KD cells was not observed (supplementary material Fig. S1C). Taken together, these results indicate that Slp2-a regulates renal epithelial cell size independently of Rab27.

Fig. 1.
Fig. 1.

Loss of Slp2-a leads to larger MDCK II cells. (A) Slp2-a expression in control and Slp2-a-KD cells. Note that the level of Slp2-a protein in the Slp2-a-KD cells was approximately 5% of the level in the control cells. GAPDH was used as an internal control. The positions of the molecular mass markers (in kilodaltons) are shown on the left. (B) Growth curve of control and Slp2-a-KD cells. 50,000 cells were seeded in individual 10-cm dishes at day 0, and trypsinized cells were counted at the times indicated. (C) Increased size of Slp2-a-KD cells. (Left) Control and Slp2-a-KD cells were cultured for 24 hours, fixed, permeabilized, and stained with Texas-Red-conjugated phalloidin (for actin staining; red). The size of each cell type was determined by measuring the area that stained positive for actin. Nuclei were stained with DAPI (blue). Scale bars: 10 µm. (Right) Quantification of the cell sizes shown in the images on the left. *P<0.001, Student's unpaired t-test (n = 50 from three independent experiments). (D) Rescue experiments using Slp2-a-KD cells. (Left) Slp2-a-KD cells were transfected with pEGFP-C1, pEGFP-C1-Slp2-aSR or pEGFP-C1-Slp2-aSR-E11A/R32A (green), and cultured for 48 hours. The cells were re-plated on 35-mm glass-bottomed dishes, cultured for 24 hours, and then stained with Texas-Red-conjugated phalloidin (red). Nuclei were stained with DAPI (blue). Scale bars: 10 µm. (Right) Quantification of the cell sizes shown in the images on the left. *P<0.001, Dunnett's test (n = 55 from three independent experiments). Values are means and s.e.m.

Slp2-a interacts with Rap1GAP2

We focused our exploration of the mechanism of the Slp2-a-mediated control of cell size on Rap1, because Rap1 is known to play a central role in basic cellular events, e.g. adhesion and spreading (Bos et al., 2001), and because Rap1GAP2, a GTPase-activating protein (GAP) for Rap1 (Schultess et al., 2005), has been shown to interact with Slp1 (Neumüller et al., 2009), the closest homologue of Slp2-a (Fukuda et al., 2001). First, we performed co-immunoprecipitation assays to test for a possible interaction between Slp2-a and Rap1GAP2. As expected, both Slp1 and Slp2-a interacted with Rap1GAP2 (Fig. 2A), and the Rap1GAP2-binding site in Slp2-a was mapped to the C2B domain (Fig. 2B, bottom, lane 5). To investigate the subcellular localization of Slp2-a and Rap1GAP2, we co-expressed C-terminal Venus (a variant of yellow fluorescent protein)-tagged Rap1GAP2 and monomeric red fluorescent protein (mRFP)-tagged Slp2-a in control MDCK II cells. Consistent with the results of the co-immunoprecipitation assays described above, Slp2-a and Rap1GAP2 were mainly colocalized to the plasma membrane (Fig. 2C). When Rap1GAP2 alone was expressed in control MDCK II cells, it was also localized to the plasma membrane, whereas it was dispersed in the cytoplasm of Slp2-a-KD cells (Fig. 2D), indicating that Slp2-a is required for the plasma membrane-localization of Rap1GAP2.

Fig. 2.
Fig. 2.

Slp2-a is required for the plasma membrane localization of Rap1GAP2. (A) Interaction between Slp1/2-a and Rap1GAP2. T7-tagged Slp1 (or Slp2-a) and FLAG-tagged Rap1GAP2 were co-expressed in COS-7 cells, and their associations were analyzed by co-immunoprecipitation assays with anti-FLAG tag antibody-conjugated agarose beads. The positions of the molecular mass markers (in kilodaltons) are shown on the left. (B) Mapping of the Rap1GAP2-binding site in Slp2-a. (Top) Schematic representation of the domain organization of Slp2-a. The truncated mutants of Slp2-a used in the co-immunoprecipitation assays are represented by solid lines. The Rap1GAP2 binding activity of each mutant is shown on the right. SHD, Slp homology domain; the C2A and C2B domains are also shown. (Bottom) T7-tagged Slp2-a truncated mutants and FLAG-tagged Rap1GAP2 were co-expressed in COS-7 cells, and their associations were analyzed as described in A. The asterisks indicate immunoglobulin G heavy and light chains. (C) Colocalization of Slp2-a and Rap1GAP2. MDCK II cells were co-transfected with pVenus-N1-Rap1GAP2 (green) and pmRFP-C1-Slp2-a (red) and cultured for 48 hours. The cells were re-plated on 35-mm glass-bottomed dishes for 4 hours and then fixed. Nuclei were stained with DAPI (blue). Fluorescence intensity along the broken white lines (left panels) is shown on the right. The colocalization rate of Rap1GAP2 with Slp2-a was 59.2±4% (n = 25). (D) Plasma membrane localization of Rap1GAP2. Control and Slp2-a-KD cells were transfected with pVenus-N1-Rap1GAP2 (green) and cultured for 48 hours. The cells were re-plated on 35-mm glass-bottomed dishes, cultured for 4 hours, fixed, permeabilized and stained with Texas-Red-conjugated phalloidin (red). Nuclei were stained with DAPI (blue). Fluorescence intensity along the broken white lines (upper panels) is shown at the bottom. Scale bars: 10 µm.

Disruption of the Slp2-a–Rap1GAP2 interaction affects cell size

Because Rap1GAP2 has been shown to interact with Slp1 through its C-terminal EVTKTT sequence (amino acids 537–542 of Rap1GAP2) (Neumüller et al., 2009), we investigated whether Slp2-a also interacts with this sequence. To this end, we generated an N-terminal truncated mutant of Rap1GAP2 (amino acids 518–687 of Rap1GAP2; designated as RGDN) and a deletion mutant of the EVTKTT sequence from the RGDN (RGDN-ΔEVTKTT; Fig. 3A, left), and then performed co-immunoprecipitation assays with the C2B domain of Slp2-a. As shown in Fig. 3A (right), the RGDN construct, but not the RGDN-ΔEVTKTT mutant, interacted with the C2B domain of Slp2-a, indicating that the EVTKTT sequence is also required for the interaction between Slp2-a and Rap1GAP2. If the interaction between Slp2-a and Rap1GAP2 is actually involved in the control of cell size, disruption of the interaction by a dominant-negative construct, e.g. the binding site alone, should affect cell size. As expected, overexpression of Venus-tagged RGDN in control MDCK II cells resulted in an increase in cell size, whereas overexpression of the Slp2-a-binding-deficient mutant (RGDN-ΔEVTKTT) did not (Fig. 3B). Similarly, GFP-tagged Slp2-a-C2B exhibited a dominant-negative effect, i.e. its overexpression in control MDCK II cells resulted in an increase in cell size (Fig. 3C), whereas GFP-tagged Slp2-a-C2A, which lacks Rap1GAP2 binding ability (Fig. 2B), had no effect on cell size (Fig. 3C). Taken together, these results indicate that Slp2-a regulates renal epithelial cell size through interaction with Rap1GAP2.

Fig. 3.
Fig. 3.

Functional disruption of the Slp2-a–Rap1GAP2 interaction leads to cell spreading. (A) The Slp2-a-binding site in Rap1GAP2. (Left) Schematic representation of the functional domains of Rap1GAP2 and its deletion mutants and point mutants used in this study. The Slp2-a-C2B binding activity of each mutant is shown on the right. The CAAX after the C-terminus of Rap1GAP2 represents the plasma membrane-targeting sequence, and the A within the GAP domain represents an N357A substitution, which impairs GAP activity. (Right) T7-tagged Slp2-a-C2B and FLAG-tagged RGDN or RGDN-ΔEVTKTT were co-expressed in COS-7 cells, and their associations were analyzed by co-immunoprecipitation assays with anti-T7 tag antibody-conjugated agarose beads. The positions of the molecular mass markers (in kilodaltons) are shown on the left. (B) Effect of the RGDN of Rap1GAP2 on cell size. (Left) MDCK II cells were transfected with pVenus-C1, pVenus-C1-RGDN or pVenus-C1-RGDN-ΔEVTKTT (green), and cultured for 48 hours. The cells were re-plated on 35-mm glass-bottomed dishes, cultured for 24 hours, and then stained with Texas-Red-conjugated phalloidin (red). The size of each cell type was determined by measuring the area that stained positive for actin. Note that both the RGDN construct and RGDN-ΔEVTKTT construct were dispersed in the cytoplasm. Nuclei were stained with DAPI (blue). Scale bars: 10 µm. (Right) Quantification of the cell sizes shown in the images on the left. *P<0.001, Dunnett's test (n = 50 from three independent experiments). (C) Effect of the C2B domain of Slp2-a on cell size. (Left) MDCK II cells were transfected with pEGFP-C1, pEGFP-C1-Slp2-a-C2A or pEGFP-C1-Slp2-a-C2B (green), and their cell sizes were measured as described above. Note that GFP-Slp2-a-C2A was localized to the plasma membrane, whereas GFP-Slp2-a-C2B was mainly present in the cytoplasm. Scale bars: 10 µm. (Right) Quantification of the cell sizes shown in the images on the left. *P<0.001, Dunnett's test (n = 45 from three independent experiments). Values are means and s.e.m.

Targeting of Rap1GAP2 to the plasma membrane is required for the control of cell size

Because Rap1GAP2 was dispersed in the cytoplasm of the Slp2-a-KD cells (Fig. 2D), we hypothesized that Rap1GAP2 functions at the plasma membrane during the control of cell size. To test our hypothesis, we generated a plasma membrane-targeted Rap1GAP2 construct by using a plasma membrane-targeting sequence (CAAX) of K-Ras (Fig. 3A, left) (Kobayashi and Fukuda, 2013). Unlike wild-type Rap1GAP2, expression of Rap1GAP2-CAAX in Slp2-a-KD cells restored the cells to the normal size (Fig. 4A). To determine whether the GAP activity of Rap1GAP2 is actually involved in the control of cell size, we generated an additional construct, Rap1GAP2-N357A-CAAX, in which the mutation has been shown to completely impair GAP activity (Schultess et al., 2005), and evaluated its effect on the size of Slp2-a-KD cells. As expected, the mutant construct had no effect on cell size, even though it was clearly localized to the plasma membrane (Fig. 4A). These results indicate that targeting of Rap1GAP2 to the plasma membrane and its subsequent activation is required for control of cell size.

Fig. 4.
Fig. 4.

Plasma membrane localization of Rap1GAP2 is required for the control of cell size. (A) Effect of forcible targeting of Rap1GAP2 to the plasma membrane on the size of Slp2-a-KD cells. (Left) Slp2-a-KD cells were transfected with pEGFP-C1, pVenus-N1-Rap1GAP2, pEGFP-C1-Rap1GAP2-CAAX or pEGFP-C1-Rap1GAP2-N357A-CAAX (green), and their size was measured as described in the legend of Fig. 1. In contrast to the cytoplasmic localization of Rap1GAP2, both Rap1GAP2-CAAX and Rap1GAP2-N357A-CAAX (see also Fig. 3A) were localized to the plasma membrane in the Slp2-a-KD cells. Scale bars: 10 µm. (Right) Quantification of the cell sizes shown in the images on the left. *P<0.001, Tukey–Kramer test (n = 40 from three independent experiments). (B) Interaction between Btsz and Rap1GAP2. T7-tagged Btsz (or Slp2-a) and FLAG-tagged Rap1GAP2 were co-expressed in COS-7 cells, and their associations were analyzed by co-immunoprecipitation assays with anti-FLAG tag antibody-conjugated agarose beads. The asterisks indicate immunoglobulin G heavy chain. The positions of the molecular mass markers (in kilodaltons) are shown on the left. (C) Effect of the Btsz on cell size. (Left) Control cells and Slp2-a-KD cells were transfected with pVenus-C1 or pVenus-C1-Btsz (green). Btsz was localized to the plasma membrane in both control and Slp2-a-KD cells. Scale bars: 10 µm. (Right) Quantification of the cell sizes shown in the images on the left. *P<0.001, Student's unpaired t-test (n = 45 from three independent experiments). Values are means and s.e.m.

Drosophila Btsz is the only known homologue of mammalian Slp that contains C-terminal tandem C2 domains but lacks an SHD (Fukuda et al., 2001; Pilot et al., 2006). Because Btsz mutations have been shown to affect growth and cell size in Drosophila (Serano and Rubin, 2003), we also investigated whether Btsz could compensate for the function of Slp2-a in MDCK II cells. Co-immunoprecipitation assays were performed to test for a possible interaction between a short form of Btsz that contains only the tandem C2 domains and Rap1GAP2, and, as expected, Btsz interacted with Rap1GAP2, in the same way as Slp2-a (Fig. 4B). Next, we expressed Venus-tagged Btsz in control and Slp2-a-KD cells and evaluated its effect on cell size. It is noteworthy that Btsz was localized to the plasma membrane and that expression of Btsz in the Slp2-a-KD cells restored normal cell size. Intriguingly, however, Btsz did not affect the size of control MDCK II cells (Fig. 4C). These results suggested that the Rap1GAP2 recruiting function of Slp in the control of cell size may have been conserved across species from Drosophila to humans.

Rap signaling is activated in Slp2-a-KD cells

Because Rap1GAP has the ability to inactivate both Rap1 and Rap2 (Gloerich and Bos, 2011), we measured the level of GTP-Rap1/2 in Slp2-a-KD cells as described in the Materials and Methods and found that both Rap1 and Rap2 activities (i.e. their GTP levels) were increased in the Slp2-a-KD cells. The amount of active Rap1 and Rap2 in the Slp2-a-KD cells expressed as a percentage of their amount in the control cells was 217±16% (n = 3) and 326±51% (n = 3), respectively (Fig. 5A; supplementary material Fig. S2A). Although both Rap1 and Rap2 modulate Rac1 signaling, they have been reported to regulate cell size by different mechanisms: active Rap1 has been reported to regulate cell size in HeLa cells by promoting cell spreading by localizing Rac1 guanine nucleotide exchange factors (GEFs) (Arthur et al., 2004), whereas active Rap2 has been reported to regulate cell size in intestinal epithelial cells by activating an upstream target of Rac1 (Gloerich et al., 2012) independently of Rap1. We demonstrated that Rac1 is also activated in Slp2-a-KD cells [the amount of active Rac1 in the Slp2-a-KD cells expressed as a percentage of its amount in the control cells was 160±20% (n = 5)] (Fig. 5A; supplementary material Fig. S2E), and then investigated whether activated Rap1 and Rap2 induce renal cell spreading by different mechanisms. We generated constitutively active and negative mutants (G12V and S17N, respectively) of Rap1A and Rap2A and expressed them in control MDCK II cells (Fig. 5B; supplementary material Fig. S2B). As expected, both active mutants induced cell spreading, but neither negative mutant did. Intriguingly, when active Rap1A and Rap2A were simultaneously expressed in control MDCK II cells, comparison with cells expressing each of the active mutants showed that their cell size had increased additively (Fig. 5B), suggesting that Rap1 and Rap2 regulate the size of renal epithelial cells through different pathways. We next investigated the possible involvement of Rac1 in cell size by using constitutively active and negative mutants of Rac1 (G12V and T17N, respectively; Fig. 5C; supplementary material Fig. S2C). Expression of the active mutant of Rac1, but not of its negative mutant, in the control MDCK II cells induced cell spreading, whereas all of the negative mutants of Rap1A, Rap2A and Rac1 decreased the size of the Slp2-a-KD cells (Fig. 5D; supplementary material Fig. S2D), indicating that the Slp2-a–Rap1GAP2 interaction regulates renal epithelial cell size by modulating Rap signaling.

Fig. 5.
Fig. 5.

Rap signaling underlying the control of cell size in renal epithelial cells. (A) Activation of Rap signaling in Slp2-a-KD cells. Active Rap1, Rap2 and Rac1 were analyzed by pulldown assays and immunoblotting as described in the Materials and Methods. The results of negative control experiments are shown in supplementary material Fig. S2A. GAPDH was used as an internal control. The top and middle panels show active and total small G proteins, respectively. The bottom panels show the GST fusion proteins that were used for the pulldown assays. Rap1, Rap2 and Rac1 were activated in the Slp2-a-KD cells. The positions of the molecular mass markers (in kilodaltons) are shown on the left. (B) Modulation of cell size by manipulating Rap signaling. (Left) MDCK II cells were transfected with pVenus-C1, pVenus-C1-Rap1A-G12V, pVenus-C1-Rap1A-S17N, pVenus-C1-Rap2A-G12V or pVenus-C1-Rap2A-S17N (green), and co-transfected with pVenus-C1-Rap1A-G12V (green) and pmStr-C1-Rap2A-G12V (red), and their size was measured as described in the legend of Fig. 1. The expression levels of the fluorescent proteins used here are shown in supplementary material Fig. S2B–D. Scale bars: 10 µm. (Right) Quantification of the cell sizes shown in the images on the left. *P<0.001, Tukey–Kramer test (n = 46 from three independent experiments). (C) Modulation of cell size by manipulating Rac1. (Left) MDCK II cells were transfected with pEGFP-C1, pEGFP-C1-Rac1-G12V or pEGFP-C1-Rac1-T17N (green). Scale bars: 10 µm. (Right) Quantification of the cell sizes shown in the images on the left. *P<0.001, Dunnett's test (n = 70 from three independent experiments). (D) Effect of inhibition of Rap signaling on the size of Slp2-a-KD cells. (Left) Slp2-a-KD cells were transfected with pEGFP-C1, pVenus-C1-Rap1A-S17N, pVenus-C1-Rap2A-S17N or pEGFP-C1-Rac1-T17N (green). Scale bars: 10 µm. (Right) Quantification of the cell sizes shown in the images on the left. *P<0.001 Dunnett's test (n = 50 from three independent experiments).

Ezrin is activated in Slp2-a-KD cells

Because Rap2 has been reported to activate ezrin (Gloerich et al., 2012), one of the ERM family proteins that bind F-actin and are involved in the regulation of cell spreading (Bretscher et al., 2002; Ross et al., 2011), and because increased phosphorylation of ezrin (active form) was shown in confluent Slp2-a-KD cells in our previous study (Yasuda et al., 2012), we hypothesized that the increased Rap2 activity in subconfluent Slp2-a-KD cells also activates ezrin. The results of the immunoblot analysis showed a clear increase in ezrin phosphorylation in the subconfluent Slp2-a-KD cells in comparison with the control MDCK II cells (Fig. 6A). Because activated ezrin is known to initiate activation of Rac1 through its ability to bind and sequester the Rho GDP dissociation inhibitor (Rho-GDI), which interacts with Rac1 and inhibits Rac1 activity (Hsu et al., 2010; Yang and Hinds, 2006), we further hypothesized that activated ezrin and Rho-GDI induce cell spreading. To test our hypothesis, we generated a constitutively active mutant of ezrin, a phosphomimetic form of ezrin (ezrin T567D), and a C-terminal truncated mutant of ezrin (ezrin-N) that possesses Rho-GDI-binding ability, and evaluated their effect on cell size in control MDCK II cells. Both mutants, but not the wild-type (WT) ezrin, increased the size of control MDCK II cells (Fig. 6B), suggesting that activated ezrin induces cell spreading through Rac1 activation. In an attempt to further support our hypothesis, we co-expressed a constitutively negative mutant of either Rap2A or Rac1 together with an active mutant of ezrin, ezrin T567D in control MDCK II cells (Fig. 6C). As expected, the Rac1 mutant (Rac1 T17N) antagonized the effect of ezrin T567D, i.e. it inhibited the cell spreading induced by ezrin T567D, whereas the Rap2A mutant, Rap2A S17N, had no effect on ezrin-T567D-mediated cell spreading. Taken together, these results strongly indicate that activated Rap2A facilitates activation of Rac1 through phosphorylation of ezrin, which in turn leads to cell spreading.

Fig. 6.
Fig. 6.

Ezrin signaling underlying the control of cell size in renal epithelial cells. (A) Activation of ezrin in Slp2-a-KD cells. The cell lysates were analyzed by immunoblotting with the antibodies indicated. The positions of the molecular mass markers (in kilodaltons) are shown on the left. (B) Modulation of cell size by manipulating ezrin. (Left) MDCK II cells were transfected with pVenus-C1, pVenus-N1-Ezrin-WT, pVenus-N1-Ezrin-T567D, or pEGFP-N1-Ezrin-N, and their size was measured as described in the legend of Fig. 1. Scale bars: 10 µm. (Right) Quantification of the cell sizes shown in the images on the left. *P<0.001, Dunnett's test (n = 30 from three independent experiments). (C) Activation of ezrin by Rap signaling. (Left) MDCK II cells were transfected with pEGFP-C1, pVenus-C1-Rap2A-S17N, or pEGFP-C1-Rac1-T17N together with pmStr-N1-Ezrin-T567D (or pEGFP-C1 together with pmStr-N1). (Right) Quantification of the cell sizes shown in the images on the left. *P<0.001 Dunnett's test (n = 30 from three independent experiments). (D) Inhibition of ezrin activity by miglustat. Slp2-a-KD cells were treated with miglustat (10 µM or 50 µM) for 24 hours and harvested. Cell lysates were analyzed by immunoblotting with the antibodies indicated. (E) Decreased size of Slp2-a-KD cells in response to miglustat. Slp2-a-KD cells were treated with miglustat (10 µM or 50 µM) for 48 hours and then re-plated on the 35-mm glass-bottomed dishes and cultured with miglustat (10 µM or 50 µM) for 24 hours. The cells were stained with Texas-Red-conjugated phalloidin and their sizes were measured (right). *P<0.001, Dunnett's test (n = 70 from three independent experiments). Values are means and s.e.m.

Increased ceramide levels have been shown to inhibit ezrin phosphorylation by activating protein phosphatases (PP1 and/or PP2A) (Canals et al., 2010; Canals et al., 2012; Zeidan et al., 2008) and to induce cell growth arrest (Shayman et al., 1991). The ceramide level of renal epithelial cells has been reported to be increased by glucosylceramide (GlcCer) synthase inhibitor treatment (Natoli et al., 2010; Shayman et al., 1991). We therefore investigated the effect of a GlcCer synthase inhibitor, miglustat (drug for the treatment of Niemannn-Pick C disease) (Venier and Igdoura, 2012), on the ezrin activity and size of Slp2-a-KD cells (Fig. 6D,E). Miglustat inactivated ezrin and reduced the size of the Slp2-a-KD cells.

Mislocalization of Slp2-a in the cytoplasm induces increased MDCK II cell size

To investigate whether mislocalization of Slp2-a from the plasma membrane affects cell size, we overexpressed GFP-tagged Slp2-a in control MDCK II cells (Fig. 7A). When the level of GFP–Slp2-a expression was relatively low (Fig. 7A, top right, open circles), it was predominantly localized to the plasma membrane (Fig. 7A, top left, open circles) and the size of the GFP–Slp2-a-expressing MDCK II cells was almost the same as that of the control GFP-expressing cells (Fig. 7A, left two panels in the lower images). As the GFP–Slp2-a expression level increased (Fig. 7A, top right, closed circles), however, the overexpressed GFP–Slp2-a was often dispersed in the cytoplasm and induced cell spreading (Fig. 7A, top left, closed circles; and bottom, right two panels). Because the interaction between Slp2-a and Rap1GAP2 was actually involved in the control of cell size (Figs 2–⇑4), mislocalization of Slp2-a was expected to affect the localization of Rap1GAP2. Just as expected, when Rap1GAP2 was co-expressed with Slp2-a in control MDCK II cells (Fig. 7B), the localization patterns of Rap1GAP2 were similar to that of Slp2-a: cytoplasmic localization of Rap1GAP2 was often observed in MDCK II cells that expressed a high level of Slp2-a (Fig. 7B, left graphs, closed circles), as opposed to the plasma membrane localization of Rap1GAP2 in MDCK II cells that expressed Rap1GAP2 alone (Fig. 7B, lower left graphs, red triangles). Because, as noted above, plasma membrane localization of Slp2-a and Rap1GAP2 is required for proper cell signaling, mislocalization of Slp2-a and Rap1GAP2 in the cytoplasm as a result of Slp2-a overexpression may lead to abnormal signaling in the control of cell size.

Fig. 7.
Fig. 7.

Mislocalization of Slp2-a and Rap1GAP2 in the cytoplasm leads to cell spreading. (A) Overexpression of Slp2-a in MDCK II cells. (Bottom) MDCK II cells were transfected with pEGFP-C1 or pEGFP-C1-Slp2-a (green), and cultured for 48 hours. The cells were re-plated on 35-mm glass-bottomed dishes, cultured for 24 hours, and then stained with Texas-Red-conjugated phalloidin (red). Nuclei were stained with DAPI (blue). Scale bars: 10 µm. (Top left) Localization of Slp2-a near the plasma membrane (within 1.5 µm from the plasma membrane) was quantified using ImageJ software. The fluorescence signals near the plasma membrane were quantified as the ratio of fluorescence intensity near the plasma membrane to total fluorescence intensity, and the ratios were plotted against cell size. (Top right) Total fluorescence intensity was plotted against cell size. Red triangles, control GFP-expressing MDCK II cells; open circles and closed circles, Slp2-a-overexpressing MDCK II cells, in which the proportion of plasma-membrane-localized Slp2-a was more than 30% and less than 30%, respectively (the plasma membrane localization of GFP was ∼30%). (B) Co-expression of Rap1GAP2 with Slp2-a. (Right) MDCK II cells were transfected with pEGFP-C1-Slp2-a (green) and pmStr-N1-Rap1GAP2 (red), and their locations and cell sizes were determined as described above. (Left) Slp2-a and Rap1GAP2 exhibited similar localization patterns in MDCK II cells. However, when Rap1GAP2 alone was expressed in control MDCK II cells (red triangles), it was mainly present at the plasma membrane and did not affect cell size.

Increased Slp2-a expression and ezrin activation in the renal cyst epithelium of the kidney of pcy mice

Because Slp2-a regulates the size of MDCK II cells (a renal epithelial cell line; Fig. 1) and is expressed in mouse renal proximal tubules (Fig. 8B, top two rows) (Yasuda et al., 2012), in the final set of experiments we investigated the possibility of a link between dysfunction of Slp2-a-mediated signaling and cell-spreading-related disorders in the kidney (Hildebrandt et al., 2009; Torres and Harris, 2009; Wilson, 2004). A spontaneous mouse model (pcy) of the late-onset (adolescent) type of human NPHP that is characterized by multicystic dysplastic kidneys, situs inversus and congenital heart defects (Bergmann et al., 2008; Olbrich et al., 2003), have very large kidneys (Fig. 8A). Intriguingly, Slp2-a was found to be expressed in the renal cyst epithelium of pcy mice (Fig. 8B, arrowheads in the bottom panel) in addition to the renal proximal tubules, where Slp2-a is normally expressed in the WT mice. When we then compared the Slp2-a expression level in the kidneys of WT and pcy mice (Fig. 8C), we found that the Slp2-a expression level was clearly higher in the kidneys of the pcy mice, but that the expression levels of other Slp family proteins were not. A reverse transcriptase (RT)-PCR analysis showed that Slp2-a expression in pcy mice was also increased at the mRNA expression level, but that of Slp1, Rap1GAP1 and Rap1GAP2 was unaltered (Fig. 8D). Because the correct amount of Slp2-a is required for the control of MDCK II cell size (i.e. both knockdown and overexpression of Slp2-a induce cell spreading; Fig. 1C; Fig. 7A), it is highly possible that the increased amount of Slp2-a in pcy mice causes abnormal signaling in their renal cyst epithelium. To investigate this possibility, we searched for the cell signaling responsible for cyst expansion in the kidneys of pcy mice (Fig. 8E). As previously reported (Omori et al., 2006; Torres and Harris, 2006), the mitogen-activated protein kinase (MAPK) pathway was activated because of the increased expression of epidermal growth factor receptor (EGFR) in the kidneys of pcy mice (Fig. 8E, upper three panels in the left blots), and the cAMP-dependent protein kinase A (PKA)–CREB pathway was activated by the increased level of circulating vasopressin in the kidneys of pcy mice (Gattone et al., 2003; Torres and Harris, 2009) (Fig. 8E, lower two panels in the left blots). Intriguingly, we found that phosphorylation of ezrin was clearly increased in the kidneys of pcy mice, and that the ezrin–phosphoinositide 3-kinase (PI3K)–AKT pathway (Gautreau et al., 1999) was activated as a result (Fig. 8E, right blots), suggesting that ezrin signaling is involved in the control of cell signaling in the kidneys of pcy mice. Because ezrin phosphorylation was increased in the kidneys of pcy mice, we proceeded to compare the localization of activated ezrin and Slp2-a in the kidneys of WT and pcy mice (Fig. 8F,G). The results showed that most of the ezrin in both the WT and pcy mice was activated and localized to the brush border (apical membrane) of the epithelial cells of the renal proximal tubules (Bretscher et al., 2002; Hatano et al., 2013), where Slp2-a was abundantly expressed (Yasuda et al., 2012). In the pcy mice, however, activated ezrin was also found to be localized to the basolateral membrane of the renal cyst epithelium (Fig. 8F,G, arrowheads in the bottom panels), where Slp2-a was also expressed. Because we and others had previously shown that excessively activated ezrin was mislocalized to the basolateral membrane (Pujuguet et al., 2003; Yasuda et al., 2012), this finding suggested that an excess of Slp2-a leads to excessive activation of ezrin in renal cyst epithelial cells, and consequently to cell spreading.

Fig. 8.
Fig. 8.

Increased Slp2-a expression and ezrin activity in the kidneys of pcy mice. (A) Kidneys of WT and pcy mice. Scale bar: 1 cm. (B) Immunohistochemical staining with anti-Slp2-a antibody (red) and anti-claudin-2 antibody (green) in the kidneys of WT and pcy mice. Scale bars: 50 µm. (C) Expression of the Slp family members in the kidneys of WT and pcy mice. The lysates of kidneys of WT and pcy mice were analyzed by immunoblotting with the antibodies indicated. The positions of the molecular mass markers (in kilodaltons) are shown on the left. (D) RT-PCR analysis of Slp1, Slp2-a, Rap1GAP1 and Rap1GAP2 mRNA in the kidneys of WT and pcy mice. The size of the molecular weight markers (kb) is shown on the left. (E) Activity of the EGFR–MAPK, cAMP–PKA and ezrin–PI3K/AKT pathways in the kidneys of WT and pcy mice. The asterisk in the right-hand blots indicates nonspecific signals. (F,G) Immunohistochemical staining with anti-P-ERM antibody (green) and anti-ezrin antibody (red; F), and with anti-Slp2-a antibody (green) and anti-P-ERM antibody (red; G) in the kidneys of WT and pcy mice. P-ezrin (P-ERM) was localized to the brush border (apical membrane) of the epithelial cells of the renal proximal tubules in both the WT (top) and pcy mice (middle). P-ezrin was also localized to the basolateral membrane of the renal cyst epithelium in the pcy mice (bottom). The asterisks and arrowheads indicate the cyst and ezrin localized to the basolateral membrane, respectively. Scale bars: 50 µm.

DISCUSSION

The Slp family consists of five members (Slp1–5) in humans and mice (Fukuda et al., 2001; Fukuda and Mikoshiba, 2001; Kuroda et al., 2002), and all members have an N-terminal SHD and C-terminal tandem C2 domains. The SHD specifically interacts with an active form of Rab27 (Kuroda et al., 2002), whereas the C2 domains, especially the C2A domain, interact with phospholipids in the plasma membrane (Catz et al., 2001; Fukuda, 2002; Kuroda and Fukuda, 2004; Gálvez-Santisteban et al., 2012). All of the Slp family members have been reported to regulate the movement of Rab27-bearing vesicles and organelles and/or their docking to the plasma membrane through the N-terminal SHD (Fukuda, 2006), but whether they have additional functions other than the Rab27 effector function had never been determined. In the present study we discovered a novel function of Slp2-a in the control of renal epithelial cell size, which is independent of Rab27 (Figs 1; supplementary material Fig. S1). Slp2-a, but not Slp1 (supplementary material Fig. S1C,G,H), controls renal epithelial cell size by recruiting Rap1GAP2 to the plasma membrane through its interaction with the C2B domain of Slp2-a (Figs 2, 3), and the Rap1GAP2 recruiting function (i.e. Slp-mediated Rap signaling) appears to have been conserved across species from Drosophila to humans (Fig. 4B,C).

Rap1 and Rap2 coordinately regulate cell spreading through Rac1 activation and through ezrin activation, respectively (Figs 5, 6). Rap signaling is controlled by specific guanine nucleotide exchange factors (GEFs) and GAPs (Gloerich and Bos, 2011), and we have succeeded in showing a functional link between Rap signaling and Slp2-a. We found that Slp2-a interacts with both Rap1GAP1 (supplementary material Fig. S3A) and Rap1GAP2 (Fig. 2A), both of which are expressed in mouse kidneys (Fig. 8D), and that it anchors them to the plasma membrane (Fig. 2C,D). Dysregulation of Slp2-a either by knockdown or by overexpression (i.e. mislocalization of Slp2-a and Rap1GAP2 in the cytoplasm) induced cell spreading in renal epithelial cells (Fig. 1C; Fig. 7; supplementary material Fig. S1B), indicating that the activity of Rap1 and Rap2 is regulated in close proximity to the plasma membrane by Rap1 GAPs. Because activation of Rap signaling, Rac signaling and/or ezrin signaling, has been shown to impair cell–cell adhesion by destabilizing E-cadherin at the cell–cell junctions (Lozano et al., 2008; Pujuguet et al., 2003; Tsygankova et al., 2010) and because reduced accumulation of E-cadherin at the cell–cell junctions was observed in the Slp2-a-KD cells (supplementary material Fig. S4A), it is highly possible that activation of these signals facilitates cell spreading by disrupting cell–cell adhesion.

We also present evidence for a link between impairment of Slp2-a-mediated Rap–ezrin signaling and PKD. PKD is the most common inherited renal cystic disease and a leading cause of end-stage renal disease. It occurs worldwide and in all races with a prevalence estimated to be between 1∶400 and 1∶1000. Dysfunctions of several cell signaling pathways, including the excess activation of the EGFR–MAPK pathway and excess activation of the cAMP–PKA pathway due to an increased level of circulating vasopressin, have been reported to contribute to the pathogenesis of PKD (Torres and Harris, 2006; Torres and Harris, 2009) (Fig. 8E). In addition to these activated signaling pathways, we found that Slp2-a expression is increased in the kidneys of pcy mice (Fig. 8C). Because a candidate CREB-binding site was identified in the mouse Slp2-a gene by a search of the CREB Target Gene Database (http://natural.salk.edu/CREB/), it is highly possible that Slp2-a mRNA transcription was activated by CREB through PKA phosphorylation. Actually, Slp2-a expression in the control MDCK II cells was dose-dependently increased in response to stimulation with a cAMP-elevating reagent, forskolin (supplementary material Fig. S3B), and the size of the cells increased in tandem (supplementary material Fig. S3C). cAMP also activates cAMP-dependent RapGEFs (cAMP-GEFs) that activate both Rap1 and Rap2, and cAMP-GEFs have been shown to be expressed in the kidney (Li et al., 2008). Because activation of cAMP-GEFs induced cell spreading in renal epithelial cells (supplementary material Fig. S3D) (Ross et al., 2011), Rap signaling must also be enhanced and contribute to the pathogenesis of PKD. Because membrane polarity proteins, e.g. E-cadherin, are known to be destabilized in the kidneys of PKD and NPHP patients (Charron et al., 2000; Marciano et al., 2011; Okada et al., 2000; Rocco et al., 1992; Wilson, 2011), and because we also observed a similar phenotype in pcy mice (supplementary material Fig. S4B), it is probable that both impairment of Slp2-a-mediated Rap–ezrin signaling and enhancement of cAMP-induced RapGEF signaling ultimately activate Rac1 and that the activated Rac1 causes cell spreading by destabilizing E-cadherin at the cell–cell junctions in the kidneys of pcy mice (supplementary material Fig. S5, right).

Preclinical studies of several agents that decrease the intracellular cAMP level have shown that they reduce cell growth in the cystic kidneys of PKD patients (Gattone et al., 2003; Torres and Harris, 2009; Torres et al., 2012). However, because the decrease in intracellular cAMP level has the undesirable effect of decreasing expression of the water channel aquaporin-2 and its apical membrane localization (Nielsen et al., 2002), the aquaretic side effects (e.g. thirst, polyurea, nocturia and polydipsia) of treatment with such agents in PKD patients are an important issue that needs to be resolved. Natoli et al. recently reported that GlcCer and ganglioside GM3 levels were increased in human and mouse PKD kidneys and that blockade of GlcCer accumulation with a GlcCer synthase inhibitor effectively inhibited cystogenesis in mouse models of PKD (Pkd1 conditional knockout mice) and NPHP (jck and pcy mice) by largely unknown mechanisms (Natoli et al., 2010). It should be noted that the ceramide level of renal epithelial cells is increased by GlcCer synthase inhibitor treatment (Natoli et al., 2010; Shayman et al., 1991) and that increased ceramide levels have been shown to inhibit ezrin phosphorylation by activating protein phosphatases (PP1 and/or PP2A) (Canals et al., 2010; Canals et al., 2012; Zeidan et al., 2008) and to induce cell growth arrest (Shayman et al., 1991). We also found that the GlcCer synthase inhibitor miglustat has the effect of inactivating the ezrin–PI3K/AKT pathway and of reducing the size of Slp2-a-KD cells (Fig. 6D,E). Taken together, these findings indicate that inactivating Rac1 as a means of slowing cell growth in renal epithelial cells (Fig. 5D; Fig. 6C) should be an effective approach to curing PKD patients, because ezrin regulates Rac1 activity through Rho-GDI (Hsu et al., 2010; Yang and Hinds, 2006) (supplementary material Fig. S5, right). Thus, ezrin (or glycosphingolipid metabolism) should be a promising therapeutic target for PKD, and it will be necessary to search for new drugs that antagonize ezrin signaling to treat PKD patients.

In summary, we have identified Rap1 GAPs as novel interactors for the Slp2-a-C2B domain and discovered that their interactions are crucial for the control of renal epithelial cell size independently of the known Rab27 effector function of Slp2-a. In the absence of the proper level of Slp2-a-mediated signaling, renal epithelial cells exhibit a cell spreading phenotype due to increased Rap–ezrin signaling, and the increased Rap–ezrin signaling is likely to be associated with the renal cell spreading that occurs in PKD. Hence, the Slp2-a-mediated Rap–ezrin signaling that controls renal epithelial cell size may provide a novel target for the treatment of PKD.

MATERIALS AND METHODS

Materials

Horseradish peroxidase (HRP)-conjugated anti-T7 tag mouse monoclonal antibody and anti-T7 tag mouse monoclonal antibody-conjugated agarose were purchased from Merck Biosciences Novagen (Darmstadt, Germany). HRP-conjugated anti-GAPDH mouse monoclonal antibody, HRP-conjugated anti-GFP rabbit polyclonal antibody, and HRP-conjugated anti-red fluorescent protein rabbit polyclonal antibody were obtained from MBL (Nagoya, Japan). HRP-conjugated anti-FLAG tag (M2) mouse monoclonal antibody and anti-FLAG tag mouse monoclonal antibody-conjugated agarose were from Sigma-Aldrich (St. Louis, MO). HRP-conjugated anti-GST rabbit polyclonal antibody was from Santa Cruz Biotechnology (Santa Cruz, CA). Anti-ezrin mouse monoclonal antibody, anti-CREB rabbit polyclonal antibody, and anti-CREB (phospho-Ser133) rabbit polyclonal antibody were from Abcam (Cambridge, MA). Anti-AKT mouse monoclonal antibody and anti-E-cadherin mouse monoclonal antibody were from BD Transduction Laboratories (Lexington, KY). Anti-phospho-ezrin(Thr567)/radixin(Thr564)/moesin(Thr558) (anti-phospho-ERM) rabbit polyclonal antibody, anti-p44/42 MAPK (ERK1/2) rabbit polyclonal antibody, anti-phospho-p44/42 (ERK1/2) (Thr202/Tyr204) mouse monoclonal antibody, and anti-phospho-AKT (Thr308) rabbit polyclonal antibody were from Cell Signaling Technology (Danvers, MA). Anti-claudin-2 rabbit polyclonal antibody and the Alexa-Fluor-488/594-conjugated goat secondary antibodies were from Invitrogen (Carlsbad, CA). Anti-EGFR sheep polyclonal antibody was from Fitzgerald (Concord, MA). Anti-Slp1, anti-Slp2-a, anti-Slp3-a, anti-Slp4-a, anti-Slp5 and anti-Rab27A rabbit polyclonal antibodies were prepared as described previously (Imai et al., 2004). Three stealth RNAiTM siRNAs against canine Slp2-a site 1 (target site no. 1: 5′-GAAGAAGCTTCAGGTAGCAGCTGAA-3′), site 2 (target site no. 2: 5′-GAGACTTTGGCAATCTGGAAGTAAA-3′), and site 3 (target site no. 3: 5′-CCCTGCCTCTCAGAATGCTGTTGAT-3′) were synthesized by Invitrogen. Three siRNAs against canine/human Rab27A site 1 (19-base target site no. 1: 5′-CCAGTGTACTTTACCAATA-3′), site 2 (19-base target site no. 2: 5′-GGAGAGGTTTCGTAGCTTA-3′) and site 3 (19-base target site no. 3: 5′-CGACAGCGTTCTTCAGAGA-3′) were synthesized by Nippon Gene Material Co., Ltd (Toyama, Japan). DBA/2JJcl mouse kidneys and pcy mouse kidneys were purchased from CLEA Japan Inc. (Tokyo, Japan) and KYUDO Co. (Saga, Japan), respectively.

Plasmid construction

pEGFP-C1 (Clontech-Takara Bio, Shiga, Japan), pmRFP-C1 and/or pEF-T7 tag expression vectors harboring cDNAs of mouse Slp2-a and Rab27A and their mutants were prepared essentially as described previously (Fukuda and Kuroda, 2002; Kuroda and Fukuda, 2004). cDNAs encoding human Rap1A, Rap2A, Rap1GAP1 and Rap1GAP2, and mouse ezrin were amplified from Marathon-Ready adult human or mouse brain and testis cDNA (Clontech-Takara Bio), and Drosophila Btsz was amplified from a Drosophila cDNA library (Merck Biosciences Novagen) by PCR with the following pairs of oligonucleotides having a restriction enzyme site (underlined) as described previously (Fukuda et al., 1999): 5′-GGATCCATGCGTGAGTACAAGCTAGT-3′ and 5′-GTCGACCTAGAGCAGCAGACATGATT-3′ for Rap1A, 5′-GGATCCATGCGCGAGTACAAAGTGGT-3′ and 5′-GTCGACCTATTGTATGTTACATGCAG-3′ for Rap2A, 5′-GGATCCATGAGCGGCCGGAAGCGCA-3′ and 5′-CTAACAGCCCAGCTGGGGCAT-3′ for Rap1GAP1, 5′-AGATCTATGTTTGGCCGGAAGCGCAG-3′ and 5′-GTCGACTTAGTGACCCGCACCAGAGC-3′ for Rap1GAP2, 5′-GGATCCTCGCACAGTGGGGGCATCCCTGGC-3′ and 5′-GTCGACCTAGCTGGGGCTCTCGCTGCTCGG-3′ for RGDN, 5′-AGATCTCCACCATGCCCAAGCCAATCAACG-3′ and 5′-GGTACCCGGGCCTGGGCCTTCATCTGCTG-3′ for the N-terminal domain of ezrin, 5′-AGATCTGCCACCATGAAGGCCCAGGCCCGG-3′ and 5′-GGTACCCATGGCCTCGAACTCGTCAATGCG-3′ for the C-terminal domain of ezrin, and 5′-GGATCCATGGAAGCCAAAGTGGCCGCT-3′ and 5′-GTCGACCTATGGCAAATTGGCTCGAATGC-3′ for Btsz. To obtain the full-length ezrin cDNA, each fragment was digested with the appropriate restriction enzymes and then the two fragments were ligated. The resulting full-length ezrin cDNA was subcloned into the pEF-FLAG tag expression vector (Fukuda et al., 1999). cDNAs encoding Rap1A G12V, Rap1A S17N, Rap2A G12V, Rap2A S17N, Rac1 G12V, Rac1 T17N, ezrin T567D, Rap1GAP2 N357A, and RGDN-ΔEVTKTT were produced by conventional or two-step PCR techniques as described previously (Tamura et al., 2011). shRNA targeted to canine Slp1 (21-base target site: 5′-GCGCAAGACCACAGTGAAGAA-3′) was constructed as described previously (Kuroda and Fukuda, 2004), by using the pSilencer-neo 2.0-U6 vector (Ambion, Austin, TX), which expresses shRNA.

Cell culture, transfection and drug treatment

MDCK II cells and COS-7 cells were cultured in Dulbecco's modified Eagle's medium (DMEM; Wako Pure Chemical Industries, Osaka, Japan) containing 10% fetal bovine serum, penicillin G (100 U/ml) and streptomycin (100 µg/ml). Plasmids were transfected into MDCK II cells or COS-7 cells by using Lipofectamine 2000 (Invitrogen) according to the manufacturer's instructions. MDCK II cells were cultured in serum-free DMEM for 16 hours and then treated with 100 nM, 500 nM or 1 µM forskolin (Wako Pure Chemical Industries) for 5 minutes (to detect CREB activity) or for 48 hours (to detect Slp2-a expression). Slp2-a-KD cells were also treated with 10 µM or 50 µM miglustat (Sigma-Aldrich) for 24 hours.

Establishment of Slp2-a-knockdown cell lines by RNA interference

Control MDCK II cells and stable Slp2-a-knockdown MDCK II cells were established as described previously [cell line no. 2 in Yasuda et al. (Yasuda et al., 2012)]. The cells were grown in DMEM containing 800 µg/ml G418 (Invitrogen). Stealth siRNAs against canine Slp2-a were transfected into MDCK II cells by using Lipofectamine RNAiMax (Invitrogen) according to the manufacturer's instructions.

Total RNA preparation and RT-PCR analysis

Total RNA preparation and the RT-PCR analyses were performed as described previously (Yasuda et al., 2012). The mouse Slp1, Slp2-a, Rap1GAP1, and Rap1GAP2 primer pairs were designed to amplify fragments of, 532, 447, 715 and 511 bp, respectively. The primers were used for Slp1 (forward primer, 5′-GCAGATCTTCTCCAGATGAGCTTCCCAG-3′ and reverse primer, 5′-GCGAATTCTGCCCTGGGGACCAGGTT-3′), Slp2-a (forward primer, 5′-GTTGCCCTCGAGACAGAAAAC-3′ and reverse primer, 5′-CATCTTCTCCCAGAGAGCAA-3′), Rap1GAP1 (forward primer, 5′-ATCAGAAGCTTGGGCAGACCT-3′ and reverse primer, 5′-CTCCGGATGACCCGCTTGAAA-3′), and Rap1GAP2 (forward primer, 5′-AGATGCTGGAGAAAATGCAG-3′ and reverse primer, 5′-GCATCATCACAGAAAGCCTT-3′). The mouse G3PDH (glyceraldehyde 3-phosphate dehydrogenase; GAPDH) was amplified using mouse G3PDH-5′ and G3PDH-3′ primers (BD Biosciences Clontech) as a reference. cDNAs were amplified by PCR using rTaq DNA polymerase (Toyobo) with 35 cycles of denaturation at 94°C for 1 minute, annealing at 52°C for 1 minute, and extension at 72°C for 1.5 minutes.

Immunofluorescence analysis

Stable and transfected MDCK II cells were cultured on glass-bottomed dishes (35-mm dish; MatTek, Ashland, MA) for 24 hours and then fixed with 4% paraformaldehyde for 10 minutes at room temperature. To visualize cell–cell adhesion, cells were permeabilized with 0.3% Triton X-100 in PBS for 2 minutes and blocked with the blocking buffer (1% BSA and 0.1% Triton X-100 in PBS) for 1 hour at room temperature. The cells were then incubated for 2 hours at room temperature in the same buffer containing anti-E-cadherin antibody (1/100 dilution) followed by incubation with secondary antibodies (Alexa-Fluor-488-conjugated antibodies; Invitrogen) and stained with DAPI. To measure cell size, cells were stained with Texas-Red-conjugated phalloidin (1/200 dilution; Invitrogen). The stained cells were examined for immunofluorescence signals with a confocal fluorescence microscope (Fluoview 1000; Olympus, Tokyo, Japan) and with MetaMorph software (Molecular Devices, Sunnyvale, CA) (for Fig. 2C,D). The immunofluorescence data are single plane images of cells. The immunofluorescence intensity was quantified with ImageJ software (version 1.44o; NIH; for Fig. 7; supplementary material Fig. S4A).

Immunoblot analysis

MDCK II cells (1×105 cells/6-cm dish) were cultured for 48 hours and harvested. After washing the cells with PBS and lysing them in a lysis buffer [50 mM HEPES-KOH, pH 7.2, 150 mM NaCl, 1 mM MgCl2, 1% Triton X-100, a protease inhibitor cocktail (Roche, Mannheim, Germany) and phosphatase inhibitor cocktail (Sigma-Aldrich)], the cell lysates were subjected to 10% SDS-PAGE followed by immunoblotting with specific primary antibodies, indicated in each figure. Immunoreactive bands were visualized with appropriate HRP-conjugated secondary antibodies (GE Healthcare Ltd, Little Chalfont, UK) and detected by enhanced chemiluminescence (ECL, GE Healthcare Ltd). The blots shown are representative of three or four independent experiments. Mouse kidneys were lysed in the same lysis buffer and analyzed by 10% SDS-PAGE as described above. The intensity of the immunoreactive bands was quantified with ImageJ software.

Co-immunoprecipitation assays

COS-7 cells (3×105 cells/6-cm dish, the day before transfection) were transfected with plasmids (pEF-T7-Slps and pEF-FLAG-Rap1GAPs), cultured for 2 days, and harvested. Co-immunoprecipitation assays with anti-FLAG tag or anti-T7 tag antibody-conjugated agarose were performed as described previously (Kuroda et al., 2002).

Assays for Rap activity and Rac activity

MDCK II cells were cultured for 48 hours and lysed in lysis buffer to assay for Rap activity (50 mM Tris-HCl pH 7.5, 200 mM NaCl, 2 mM MgCl2, 1% NP-40, 1 mM DTT, 10% glycerol, protease inhibitor cocktail and phosphatase inhibitor cocktail), or in lysis buffer to assay for Rac activity (50 mM Tris-HCl pH 7.5, 150 mM NaCl, 10 mM MgCl2, 1% Triton X-100, 500 µM DTT, 15% glycerol, protease inhibitor cocktail and phosphatase inhibitor cocktail). The cell lysates were centrifuged at 15,000 g for 10 minutes, and 1 mg samples were incubated at 4°C for 2 hours with glutathione–Sepharose 4B beads (GE Healthcare Ltd) coupled with 30 µg of GST alone as a negative control, GST–RalGDS (for Rap) or GST–PAK1-PBD (for Rac). After washing the beads three times with the lysis buffer, proteins trapped by the beads (active Rap or active Rac1) and total lysates (total Rap or total Rac1) were analyzed by immunoblotting with anti-Rap antibody or anti-Rac1 antibody.

Immunohistochemical analysis

Immunohistochemical staining was performed as described previously (Yasuda et al., 2012). The samples were stained with anti-E-cadherin antibody, anti-Slp2-a antibody, anti-claudin-2 antibody, anti-ezrin antibody, and anti-P-ERM antibody. To double stain for Slp2-a and claudin-2 or for Slp2-a and P-ERM, the Slp2-a antibody was directly labeled with Alexa Fluor 488 or 594 using a direct labeling kit (Zenon Alexa Fluor 488 or 594 rabbit IgG labeling kit; Invitrogen). The immunostained sections were examined with a confocal fluorescence microscope.

Statistical analyses

The data are expressed as means and s.e.m. Data were tested for statistically significant differences by using Student's unpaired t-test, Dunnett's test or the Tukey–Kramer test as indicated in the legend of each figure. A probability level of P<0.05 was considered statistically significant.

Acknowledgments

We thank Megumi Aizawa for technical assistance and members of the Fukuda Laboratory for valuable discussions.

Footnotes

  • Competing interests

    The authors declare no competing interests.

  • Author contributions

    T.Y. designed and performed the experiments and wrote the manuscript. M.F. supervised the project and wrote the manuscript.

  • Funding

    This work was supported in part by the Japan Society for the Promotion of Science KAKENHI [grant number 242766 to T.Y.]; and Grants-in-Aid for Scientific Research from the Ministry of Education, Culture, Sports, and Technology of Japan [grant numbers 24370077 and 24657125 to M.F.].

  • Supplementary material available online at http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.134056/-/DC1

  • Received April 27, 2013.
  • Accepted October 30, 2013.

References

  1. ↵
    1. Arthur, W. T.,
    2. Quilliam, L. A. and
    3. Cooper, J. A.
    (2004). Rap1 promotes cell spreading by localizing Rac guanine nucleotide exchange factors. J. Cell Biol. 167, 111–122. doi:10.1083/jcb.200404068
    OpenUrlAbstract/FREE Full Text
  2. ↵
    1. Bergmann, C.,
    2. Fliegauf, M.,
    3. Brüchle, N. O.,
    4. Frank, V.,
    5. Olbrich, H.,
    6. Kirschner, J.,
    7. Schermer, B.,
    8. Schmedding, I.,
    9. Kispert, A. and
    10. Kränzlin, B.
    et al.(2008). Loss of nephrocystin-3 function can cause embryonic lethality, Meckel-Gruber-like syndrome, situs inversus, and renal-hepatic-pancreatic dysplasia. Am. J. Hum. Genet. 82, 959–970. doi:10.1016/j.ajhg.2008.02.017
    OpenUrlCrossRefPubMedWeb of Science
  3. ↵
    1. Bos, J. L.,
    2. de Rooij, J. and
    3. Reedquist, K. A.
    (2001). Rap1 signalling: adhering to new models. Nat. Rev. Mol. Cell Biol. 2, 369–377. doi:10.1038/35073073
    OpenUrlCrossRefPubMedWeb of Science
  4. ↵
    1. Bretscher, A.,
    2. Edwards, K. and
    3. Fehon, R. G.
    (2002). ERM proteins and merlin: integrators at the cell cortex. Nat. Rev. Mol. Cell Biol. 3, 586–599. doi:10.1038/nrm882
    OpenUrlCrossRefPubMedWeb of Science
  5. ↵
    1. Canals, D.,
    2. Jenkins, R. W.,
    3. Roddy, P.,
    4. Hernández-Corbacho, M. J.,
    5. Obeid, L. M. and
    6. Hannun, Y. A.
    (2010). Differential effects of ceramide and sphingosine 1-phosphate on ERM phosphorylation: probing sphingolipid signaling at the outer plasma membrane. J. Biol. Chem. 285, 32476–32485. doi:10.1074/jbc.M110.141028
    OpenUrlAbstract/FREE Full Text
  6. ↵
    1. Canals, D.,
    2. Roddy, P. and
    3. Hannun, Y. A.
    (2012). Protein phosphatase 1α mediates ceramide-induced ERM protein dephosphorylation: a novel mechanism independent of phosphatidylinositol 4, 5-biphosphate (PIP2) and myosin/ERM phosphatase. J. Biol. Chem. 287, 10145–10155. doi:10.1074/jbc.M111.306456
    OpenUrlAbstract/FREE Full Text
  7. ↵
    1. Catz, S. D.,
    2. Johnson, J. L. and
    3. Babior, B. M.
    (2001). Characterization of the nucleotide-binding capacity and the ATPase activity of the PIP3-binding protein JFC1. Proc. Natl. Acad. Sci. USA 98, 11230–11235. doi:10.1073/pnas.191369598
    OpenUrlAbstract/FREE Full Text
  8. ↵
    1. Charron, A. J.,
    2. Nakamura, S.,
    3. Bacallao, R. and
    4. Wandinger-Ness, A.
    (2000). Compromised cytoarchitecture and polarized trafficking in autosomal dominant polycystic kidney disease cells. J. Cell Biol. 149, 111–124. doi:10.1083/jcb.149.1.111
    OpenUrlAbstract/FREE Full Text
  9. ↵
    1. Fukuda, M.
    (2002). The C2A domain of synaptotagmin-like protein 3 (Slp3) is an atypical calcium-dependent phospholipid-binding machine: comparison with the C2A domain of synaptotagmin I. Biochem. J. 366, 681–687. doi:10.1042/BJ20020484
    OpenUrlCrossRefPubMedWeb of Science
  10. ↵
    1. Fukuda, M.
    (2006). Rab27 and its effectors in secretory granule exocytosis: a novel docking machinery composed of a Rab27·effector complex. Biochem. Soc. Trans. 34, 691–695. doi:10.1042/BST0340691
    OpenUrlCrossRefPubMedWeb of Science
  11. ↵
    1. Fukuda, M. and
    2. Kuroda, T. S.
    (2002). Slac2-c (synaptotagmin-like protein homologue lacking C2 domains-c), a novel linker protein that interacts with Rab27, myosin Va/VIIa, and actin. J. Biol. Chem. 277, 43096–43103. doi:10.1074/jbc.M203862200
    OpenUrlAbstract/FREE Full Text
  12. ↵
    1. Fukuda, M. and
    2. Mikoshiba, K.
    (2001). Synaptotagmin-like protein 1-3: a novel family of C-terminal-type tandem C2 proteins. Biochem. Biophys. Res. Commun. 281, 1226–1233. doi:10.1006/bbrc.2001.4512
    OpenUrlCrossRefPubMedWeb of Science
  13. ↵
    1. Fukuda, M.,
    2. Kanno, E. and
    3. Mikoshiba, K.
    (1999). Conserved N-terminal cysteine motif is essential for homo- and heterodimer formation of synaptotagmins III, V, VI, and X. J. Biol. Chem. 274, 31421–31427. doi:10.1074/jbc.274.44.31421
    OpenUrlAbstract/FREE Full Text
  14. ↵
    1. Fukuda, M.,
    2. Saegusa, C. and
    3. Mikoshiba, K.
    (2001). Novel splicing isoforms of synaptotagmin-like proteins 2 and 3: identification of the Slp homology domain. Biochem. Biophys. Res. Commun. 283, 513–519. doi:10.1006/bbrc.2001.4803
    OpenUrlCrossRefPubMedWeb of Science
  15. ↵
    1. Gálvez-Santisteban, M.,
    2. Rodriguez-Fraticelli, A. E.,
    3. Bryant, D. M.,
    4. Vergarajauregui, S.,
    5. Yasuda, T.,
    6. Bañón-Rodríguez, I.,
    7. Bernascone, I.,
    8. Datta, A.,
    9. Spivak, N. and
    10. Young, K.
    et al.(2012). Synaptotagmin-like proteins control the formation of a single apical membrane domain in epithelial cells. Nat. Cell Biol. 14, 838–849. doi:10.1038/ncb2541
    OpenUrlCrossRefPubMed
  16. ↵
    1. Gattone, V. H. 2nd.,
    2. Wang, X.,
    3. Harris, P. C. and
    4. Torres, V. E.
    (2003). Inhibition of renal cystic disease development and progression by a vasopressin V2 receptor antagonist. Nat. Med. 9, 1323–1326. doi:10.1038/nm935
    OpenUrlCrossRefPubMedWeb of Science
  17. ↵
    1. Gautreau, A.,
    2. Poullet, P.,
    3. Louvard, D. and
    4. Arpin, M.
    (1999). Ezrin, a plasma membrane-microfilament linker, signals cell survival through the phosphatidylinositol 3-kinase/Akt pathway. Proc. Natl. Acad. Sci. USA 96, 7300–7305. doi:10.1073/pnas.96.13.7300
    OpenUrlAbstract/FREE Full Text
  18. ↵
    1. Gloerich, M. and
    2. Bos, J. L.
    (2011). Regulating Rap small G-proteins in time and space. Trends Cell Biol. 21, 615–623. doi:10.1016/j.tcb.2011.07.001
    OpenUrlCrossRefPubMedWeb of Science
  19. ↵
    1. Gloerich, M.,
    2. ten Klooster, J. P.,
    3. Vliem, M. J.,
    4. Koorman, T.,
    5. Zwartkruis, F. J.,
    6. Clevers, H. and
    7. Bos, J. L.
    (2012). Rap2A links intestinal cell polarity to brush border formation. Nat. Cell Biol. 14, 793–801. doi:10.1038/ncb2537
    OpenUrlCrossRefPubMed
  20. ↵
    1. Hatano, R.,
    2. Fujii, E.,
    3. Segawa, H.,
    4. Mukaisho, K.,
    5. Matsubara, M.,
    6. Miyamoto, K.,
    7. Hattori, T.,
    8. Sugihara, H. and
    9. Asano, S.
    (2013). Ezrin, a membrane cytoskeletal cross-linker, is essential for the regulation of phosphate and calcium homeostasis. Kidney Int. 83, 41–49. doi:10.1038/ki.2012.308
    OpenUrlCrossRefPubMed
  21. ↵
    1. Hildebrandt, F.,
    2. Attanasio, M. and
    3. Otto, E.
    (2009). Nephronophthisis: disease mechanisms of a ciliopathy. J. Am. Soc. Nephrol. 20, 23–35. doi:10.1681/ASN.2008050456
    OpenUrlAbstract/FREE Full Text
  22. ↵
    1. Holt, O.,
    2. Kanno, E.,
    3. Bossi, G.,
    4. Booth, S.,
    5. Daniele, T.,
    6. Santoro, A.,
    7. Arico, M.,
    8. Saegusa, C.,
    9. Fukuda, M. and
    10. Griffiths, G. M.
    (2008). Slp1 and Slp2-a localize to the plasma membrane of CTL and contribute to secretion from the immunological synapse. Traffic 9, 446–457. doi:10.1111/j.1600-0854.2008.00714.x
    OpenUrlCrossRefPubMedWeb of Science
  23. ↵
    1. Hsu, Y. H.,
    2. Lin, W. L.,
    3. Hou, Y. T.,
    4. Pu, Y. S.,
    5. Shun, C. T.,
    6. Chen, C. L.,
    7. Wu, Y. Y.,
    8. Chen, J. Y.,
    9. Chen, T. H. and
    10. Jou, T. S.
    (2010). Podocalyxin EBP50 ezrin molecular complex enhances the metastatic potential of renal cell carcinoma through recruiting Rac1 guanine nucleotide exchange factor ARHGEF7. Am. J. Pathol. 176, 3050–3061. doi:10.2353/ajpath.2010.090539
    OpenUrlCrossRefPubMedWeb of Science
  24. ↵
    1. Imai, A.,
    2. Yoshie, S.,
    3. Nashida, T.,
    4. Shimomura, H. and
    5. Fukuda, M.
    (2004). The small GTPase Rab27B regulates amylase release from rat parotid acinar cells. J. Cell Sci. 117, 1945–1953. doi:10.1242/jcs.01048
    OpenUrlAbstract/FREE Full Text
  25. ↵
    1. Kobayashi, H. and
    2. Fukuda, M.
    (2013). Rab35 establishes the EHD1-association site by coordinating two distinct effectors during neurite outgrowth. J. Cell Sci. 126, 2424–2435. doi:10.1242/jcs.117846
    OpenUrlAbstract/FREE Full Text
  26. ↵
    1. Kuroda, T. S. and
    2. Fukuda, M.
    (2004). Rab27A-binding protein Slp2-a is required for peripheral melanosome distribution and elongated cell shape in melanocytes. Nat. Cell Biol. 6, 1195–1203. doi:10.1038/ncb1197
    OpenUrlCrossRefPubMedWeb of Science
  27. ↵
    1. Kuroda, T. S.,
    2. Fukuda, M.,
    3. Ariga, H. and
    4. Mikoshiba, K.
    (2002). The Slp homology domain of synaptotagmin-like proteins 1-4 and Slac2 functions as a novel Rab27A binding domain. J. Biol. Chem. 277, 9212–9218. doi:10.1074/jbc.M112414200
    OpenUrlAbstract/FREE Full Text
  28. ↵
    1. Li, Y.,
    2. Konings, I. B. M.,
    3. Zhao, J.,
    4. Price, L. S.,
    5. de Heer, E. and
    6. Deen, P. M. T.
    (2008). Renal expression of exchange protein directly activated by cAMP (Epac) 1 and 2. Am. J. Physiol. 295, F525–F533. doi:10.1152/ajprenal.00448.2007
    OpenUrlCrossRef
  29. ↵
    1. Lozano, E.,
    2. Frasa, M. A. M.,
    3. Smolarczyk, K.,
    4. Knaus, U. G. and
    5. Braga, V. M. M.
    (2008). PAK is required for the disruption of E-cadherin adhesion by the small GTPase Rac. J. Cell Sci. 121, 933–938. doi:10.1242/jcs.016121
    OpenUrlAbstract/FREE Full Text
  30. ↵
    1. Marciano, D. K.,
    2. Brakeman, P. R.,
    3. Lee, C. Z.,
    4. Spivak, N.,
    5. Eastburn, D. J.,
    6. Bryant, D. M.,
    7. Beaudoin, G. M. 3rd.,
    8. Hofmann, I.,
    9. Mostov, K. E. and
    10. Reichardt, L. F.
    (2011). p120 catenin is required for normal renal tubulogenesis and glomerulogenesis. Development 138, 2099–2109. doi:10.1242/dev.056564
    OpenUrlAbstract/FREE Full Text
  31. ↵
    1. Ménasché, G.,
    2. Ménager, M. M.,
    3. Lefebvre, J. M.,
    4. Deutsch, E.,
    5. Athman, R.,
    6. Lambert, N.,
    7. Mahlaoui, N.,
    8. Court, M.,
    9. Garin, J. and
    10. Fischer, A.
    et al.(2008). A newly identified isoform of Slp2a associates with Rab27a in cytotoxic T cells and participates to cytotoxic granule secretion. Blood 112, 5052–5062. doi:10.1182/blood-2008-02-141069
    OpenUrlAbstract/FREE Full Text
  32. ↵
    1. Natoli, T. A.,
    2. Smith, L. A.,
    3. Rogers, K. A.,
    4. Wang, B.,
    5. Komarnitsky, S.,
    6. Budman, Y.,
    7. Belenky, A.,
    8. Bukanov, N. O.,
    9. Dackowski, W. R. and
    10. Husson, H.
    et al.(2010). Inhibition of glucosylceramide accumulation results in effective blockade of polycystic kidney disease in mouse models. Nat. Med. 16, 788–792. doi:10.1038/nm.2171
    OpenUrlCrossRefPubMedWeb of Science
  33. ↵
    1. Neumüller, O.,
    2. Hoffmeister, M.,
    3. Babica, J.,
    4. Prelle, C.,
    5. Gegenbauer, K. and
    6. Smolenski, A. P.
    (2009). Synaptotagmin-like protein 1 interacts with the GTPase-activating protein Rap1GAP2 and regulates dense granule secretion in platelets. Blood 114, 1396–1404. doi:10.1182/blood-2008-05-155234
    OpenUrlAbstract/FREE Full Text
  34. ↵
    1. Nielsen, S.,
    2. Frøkiaer, J.,
    3. Marples, D.,
    4. Kwon, T. H.,
    5. Agre, P. and
    6. Knepper, M. A.
    (2002). Aquaporins in the kidney: from molecules to medicine. Physiol. Rev. 82, 205–244.
    OpenUrlAbstract/FREE Full Text
  35. ↵
    1. Okada, H.,
    2. Ban, S.,
    3. Nagao, S.,
    4. Takahashi, H.,
    5. Suzuki, H. and
    6. Neilson, E. G.
    (2000). Progressive renal fibrosis in murine polycystic kidney disease: an immunohistochemical observation. Kidney Int. 58, 587–597. doi:10.1046/j.1523-1755.2000.00205.x
    OpenUrlCrossRefPubMedWeb of Science
  36. ↵
    1. Olbrich, H.,
    2. Fliegauf, M.,
    3. Hoefele, J.,
    4. Kispert, A.,
    5. Otto, E.,
    6. Volz, A.,
    7. Wolf, M. T.,
    8. Sasmaz, G.,
    9. Trauer, U. and
    10. Reinhardt, R.
    et al.(2003). Mutations in a novel gene, NPHP3, cause adolescent nephronophthisis, tapeto-retinal degeneration and hepatic fibrosis. Nat. Genet. 34, 455–459. doi:10.1038/ng1216
    OpenUrlCrossRefPubMedWeb of Science
  37. ↵
    1. Omori, S.,
    2. Hida, M.,
    3. Fujita, H.,
    4. Takahashi, H.,
    5. Tanimura, S.,
    6. Kohno, M. and
    7. Awazu, M.
    (2006). Extracellular signal-regulated kinase inhibition slows disease progression in mice with polycystic kidney disease. J. Am. Soc. Nephrol. 17, 1604–1614. doi:10.1681/ASN.2004090800
    OpenUrlAbstract/FREE Full Text
  38. ↵
    1. Pilot, F.,
    2. Philippe, J. M.,
    3. Lemmers, C. and
    4. Lecuit, T.
    (2006). Spatial control of actin organization at adherens junctions by a synaptotagmin-like protein Btsz. Nature 442, 580–584. doi:10.1038/nature04935
    OpenUrlCrossRefPubMedWeb of Science
  39. ↵
    1. Pujuguet, P.,
    2. Del Maestro, L.,
    3. Gautreau, A.,
    4. Louvard, D. and
    5. Arpin, M.
    (2003). Ezrin regulates E-cadherin-dependent adherens junction assembly through Rac1 activation. Mol. Biol. Cell 14, 2181–2191. doi:10.1091/mbc.E02-07-0410
    OpenUrlAbstract/FREE Full Text
  40. ↵
    1. Rocco, M. V.,
    2. Neilson, E. G.,
    3. Hoyer, J. R. and
    4. Ziyadeh, F. N.
    (1992). Attenuated expression of epithelial cell adhesion molecules in murine polycystic kidney disease. Am. J. Physiol. 262, F679–F686.
    OpenUrlPubMed
  41. ↵
    1. Ross, S. H.,
    2. Post, A.,
    3. Raaijmakers, J. H.,
    4. Verlaan, I.,
    5. Gloerich, M. and
    6. Bos, J. L.
    (2011). Ezrin is required for efficient Rap1-induced cell spreading. J. Cell Sci. 124, 1808–1818. doi:10.1242/jcs.079830
    OpenUrlAbstract/FREE Full Text
  42. ↵
    1. Saegusa, C.,
    2. Tanaka, T.,
    3. Tani, S.,
    4. Itohara, S.,
    5. Mikoshiba, K. and
    6. Fukuda, M.
    (2006). Decreased basal mucus secretion by Slp2-a-deficient gastric surface mucous cells. Genes Cells 11, 623–631. doi:10.1111/j.1365-2443.2006.00964.x
    OpenUrlCrossRefPubMedWeb of Science
  43. ↵
    1. Schultess, J.,
    2. Danielewski, O. and
    3. Smolenski, A. P.
    (2005). Rap1GAP2 is a new GTPase-activating protein of Rap1 expressed in human platelets. Blood 105, 3185–3192. doi:10.1182/blood-2004-09-3605
    OpenUrlAbstract/FREE Full Text
  44. ↵
    1. Serano, J. and
    2. Rubin, G. M.
    (2003). The Drosophila synaptotagmin-like protein bitesize is required for growth and has mRNA localization sequences within its open reading frame. Proc. Natl. Acad. Sci. USA 100, 13368–13373. doi:10.1073/pnas.1835727100
    OpenUrlAbstract/FREE Full Text
  45. ↵
    1. Shayman, J. A.,
    2. Deshmukh, G. D.,
    3. Mahdiyoun, S.,
    4. Thomas, T. P.,
    5. Wu, D.,
    6. Barcelon, F. S. and
    7. Radin, N. S.
    (1991). Modulation of renal epithelial cell growth by glucosylceramide. Association with protein kinase C, sphingosine, and diacylglycerol. J. Biol. Chem. 266, 22968–22974.
    OpenUrlAbstract/FREE Full Text
  46. ↵
    1. Tamura, K.,
    2. Ohbayashi, N.,
    3. Ishibashi, K. and
    4. Fukuda, M.
    (2011). Structure-function analysis of VPS9-ankyrin-repeat protein (Varp) in the trafficking of tyrosinase-related protein 1 in melanocytes. J. Biol. Chem. 286, 7507–7521. doi:10.1074/jbc.M110.191205
    OpenUrlAbstract/FREE Full Text
  47. ↵
    1. Torres, V. E. and
    2. Harris, P. C.
    (2006). Mechanisms of Disease: autosomal dominant and recessive polycystic kidney diseases. Nat. Clin. Pract. Nephrol. 2, 40–55, quiz 55. doi:10.1038/ncpneph0070
    OpenUrlCrossRefPubMedWeb of Science
  48. ↵
    1. Torres, V. E. and
    2. Harris, P. C.
    (2009). Autosomal dominant polycystic kidney disease: the last 3 years. Kidney Int. 76, 149–168. doi:10.1038/ki.2009.128
    OpenUrlCrossRefPubMedWeb of Science
  49. ↵
    1. Torres, V. E.,
    2. Chapman, A. B.,
    3. Devuyst, O.,
    4. Gansevoort, R. T.,
    5. Grantham, J. J.,
    6. Higashihara, E.,
    7. Perrone, R. D.,
    8. Krasa, H. B.,
    9. Ouyang, J. and
    10. Czerwiec, F. S.
    TEMPO 3:4 Trial Investigators(2012). Tolvaptan in patients with autosomal dominant polycystic kidney disease. N. Engl. J. Med. 367, 2407–2418. doi:10.1056/NEJMoa1205511
    OpenUrlCrossRefPubMedWeb of Science
  50. ↵
    1. Tsygankova, O. M.,
    2. Ma, C.,
    3. Tang, W.,
    4. Korch, C.,
    5. Feldman, M. D.,
    6. Lv, Y.,
    7. Brose, M. S. and
    8. Meinkoth, J. L.
    (2010). Downregulation of Rap1GAP in human tumor cells alters cell/matrix and cell/cell adhesion. Mol. Cell. Biol. 30, 3262–3274. doi:10.1128/MCB.01345-09
    OpenUrlAbstract/FREE Full Text
  51. ↵
    1. Venier, R. E. and
    2. Igdoura, S. A.
    (2012). Miglustat as a therapeutic agent: prospects and caveats. J. Med. Genet. 49, 591–597. doi:10.1136/jmedgenet-2012-101070
    OpenUrlAbstract/FREE Full Text
  52. ↵
    1. Wilson, P. D.
    (2004). Polycystic kidney disease. N. Engl. J. Med. 350, 151–164. doi:10.1056/NEJMra022161
    OpenUrlCrossRefPubMedWeb of Science
  53. ↵
    1. Wilson, P. D.
    (2011). Apico-basal polarity in polycystic kidney disease epithelia. Biochim. Biophys. Acta 1812, 1239–1248. doi:10.1016/j.bbadis.2011.05.008
    OpenUrlCrossRefPubMedWeb of Science
  54. ↵
    1. Yang, H. S. and
    2. Hinds, P. W.
    (2006). Phosphorylation of ezrin by cyclin-dependent kinase 5 induces the release of Rho GDP dissociation inhibitor to inhibit Rac1 activity in senescent cells. Cancer Res. 66, 2708–2715. doi:10.1158/0008-5472.CAN-05-3141
    OpenUrlAbstract/FREE Full Text
  55. ↵
    1. Yasuda, T.,
    2. Saegusa, C.,
    3. Kamakura, S.,
    4. Sumimoto, H. and
    5. Fukuda, M.
    (2012). Rab27 effector Slp2-a transports the apical signaling molecule podocalyxin to the apical surface of MDCK II cells and regulates claudin-2 expression. Mol. Biol. Cell 23, 3229–3239. doi:10.1091/mbc.E12-02-0104
    OpenUrlAbstract/FREE Full Text
  56. ↵
    1. Zeidan, Y. H.,
    2. Jenkins, R. W. and
    3. Hannun, Y. A.
    (2008). Remodeling of cellular cytoskeleton by the acid sphingomyelinase/ceramide pathway. J. Cell Biol. 181, 335–350. doi:10.1083/jcb.200705060
    OpenUrlAbstract/FREE Full Text
Back to top

Keywords

 Download PDF

Email
Research Article
Slp2-a controls renal epithelial cell size through regulation of Rap–ezrin signaling independently of Rab27
Takao Yasuda, Mitsunori Fukuda
Journal of Cell Science 2014 127: 557-570; doi: 10.1242/jcs.134056
del.icio.us logo Digg logo Reddit logo Twitter logo CiteULike logo Facebook logo Google logo Mendeley logo
Citation Tools
Alerts

Article navigation

Other journals from The Company of Biologists

Advertisement

Introducing FocalPlane’s new Community Manager, Esperanza Agullo-Pascual

We are pleased to welcome Esperanza to the Journal of Cell Science team. The new Community Manager for FocalPlane, Esperanza is joining us from the Microscopy Core at Mount Sinai School of Medicine. Find out more about Esperanza in her introductory post over on FocalPlane.

New funding scheme supports sustainable events

As part of our Sustainable Conferencing Initiative, we are pleased to announce funding for organisers that seek to reduce the environmental footprint of their event. The next deadline to apply for a Scientific Meeting grant is 26 March 2021.

Read & Publish participation continues to grow

"Alongside pre-printing for early documentation of work, such mechanisms are particularly helpful for early-career researchers like me.”

Dr Chris MacDonald (University of York) shares his experience of publishing Open Access as part of our growing Read & Publish initiative. We now have over 150 institutions in 15 countries and four library consortia taking part – find out more and view our full list of participating institutions.

Cell scientist to watch: Romain Levayer

In an interview, Romain Levayer talks about starting his own lab, his love for preprints and his experience of balancing parenting with his research goals.

Live lactating mammary tissue

In a stunning video, Stewart et al. demonstrate warping of the alveolar unit due to basal cell-generated force as part of their recent work investigating roles for mechanically activated ion channels in lactation and involution.

Visit our YouTube channel to watch more videos from JCS, our sister journals and the Company.

JCS and COVID-19

For more information on measures Journal of Cell Science is taking to support the community during the COVID-19 pandemic, please see here.

If you have any questions or concerns, please do not hestiate to contact the Editorial Office.