Cell–cell junction formation requires actin cytoskeletal remodeling. Here, we show that PLEKHG4B, a Rho-guanine nucleotide exchange factor (Rho-GEF), plays a crucial role in epithelial cell–cell junction formation. Knockdown of PLEKHG4B decreased Cdc42 activity and tended to increase RhoA activity in A549 cells. A549 monolayer cells showed ‘closed junctions’ with closely packed actin bundles along the cell–cell contacts, but PLEKHG4B knockdown suppressed closed junction formation, and PLEKHG4B-knockdown cells exhibited ‘open junctions’ with split actin bundles located away from the cell–cell boundary. In Ca2+-switch assays, PLEKHG4B knockdown delayed the conversion of open junctions to closed junctions and β-catenin accumulation at cell–cell junctions. Furthermore, PLEKHG4B knockdown abrogated the reduction in myosin activity normally seen in the later stage of junction formation. The aberrant myosin activation and impairments in closed junction formation in PLEKHG4B-knockdown cells were reverted by ROCK inhibition or LARG/PDZ-RhoGEF knockdown. These results suggest that PLEKHG4B enables actin remodeling during epithelial cell–cell junction maturation, probably by reducing myosin activity in the later stage of junction formation, through suppressing LARG/PDZ-RhoGEF and RhoA–ROCK pathway activities. We also showed that annexin A2 participates in PLEKHG4B localization to cell–cell junctions.
Cell–cell junctions regulate various structural and functional characteristics of cells, including morphology, polarity, motility, mechanical integrity, barrier function, proliferation and differentiation (Garcia et al., 2018), and thus play essential roles in maintaining correct tissue architecture, development and homeostasis. Abnormal control of cell–cell junction formation is implicated in various diseases, including cancer. Epithelial cells adhere to each other through junctional complexes consisting of tight junctions, adherens junctions (AJs) and desmosomes, which are typically aligned in the apical-to-basal direction on the lateral plasma membrane in vertebrate epithelial tissues (Garcia et al., 2018). AJs connect adjacent cells principally via homophilic interactions of cadherins, a family of Ca2+-dependent transmembrane cell–cell adhesion proteins. The cytoplasmic tails of cadherins are anchored to the actin filament bundles through AJ-associated proteins, including β-catenin and α-catenin (Harris and Tepass, 2010; Mège and Ishiyama, 2017; Takeichi, 2014). AJs integrate cadherin-based adhesion and actomyosin-based contractility, and their assembly and disassembly enable dynamic cell rearrangements – such as convergent extension, tissue invagination and collective migration – during epithelial morphogenesis and homeostasis (Lecuit and Yap, 2015). Thus, cytoskeletal reorganization is dependent on AJ formation and stability, which are themselves regulated by actin cytoskeletal dynamics and actomyosin-based contractility (Takeichi, 2014).
Epithelial cell–cell junction formation is a multistep process that is coupled with actin cytoskeletal reorganization (Cavey and Lecuit, 2009; Vaezi et al., 2002; Vasioukhin et al., 2000; Yonemura et al., 1995). Typically, at the onset of cell–cell contact, adjacent cells form ‘spot-like’ AJs (also termed punctum adherens), in which cadherin-containing puncta are concentrated at the tips of filopodial or lamellipodial membrane protrusions at the cell–cell contact sites. As the cell–cell contact region expands, adjacent cells become connected by the ‘zipper-like’ AJs, in which radial actin bundles link the cadherin puncta to the split actin bundles located away from the cell–cell boundary. Then, the split actin bundles in adjacent cells are reorganized to the closely packed actin bundles along the cell–cell contacts, eventually leading to the formation of mature cell–cell junctions with apicobasal cell polarity (Cavey and Lecuit, 2009; Yonemura et al., 1995; Zhang et al., 2005). Thus, proper regulation of actin cytoskeletal remodeling is required for cell–cell junction maturation.
The Rho family of small GTPases are key regulators of actin cytoskeletal remodeling (Hodge and Ridley, 2016). The three best-characterized members of the family – RhoA, Rac1 and Cdc42 – induce the formation of stress fibers, lamellipodia and filopodia, respectively. Rho GTPases are regulated by the opposing actions of Rho-guanine nucleotide exchange factors (Rho-GEFs) and Rho GTPase-activating proteins (Rho-GAPs). Rho-GEFs catalyze the conversion of Rho proteins from their inactive, GDP-bound, forms to their active, GTP-bound, forms, while Rho-GAPs stimulate the intrinsic GTPase activity of Rho GTPases to facilitate the return to the inactive conformation (Hodge and Ridley, 2016). Previous studies showed that Rho GTPases, Rho-GEFs [such as Tiam1, Trio and Tuba (also known as DNMBP)] and Rho-GAPs [including p190RhoGAP and FILGAP (also known as ARHGAP24)] regulate the formation of epithelial cell–cell adhesions (Arnold et al., 2017; Braga, 2018; Citi et al., 2014; Fukata and Kaibuchi, 2001; McCormack et al., 2013), while cadherin-based cell–cell adhesions control the activities of Rho GTPases. Thus, Rho GTPases and cadherins mutually regulate their actions during cell–cell adhesion formation and remodeling (Citi et al., 2014; Fukata and Kaibuchi, 2001). However, the precise mechanisms underlying actin reorganization during cell–cell adhesion formation and the role of Rho-GEFs in this process are not completely understood (McCormack et al., 2013; Takeichi, 2014).
There are ∼70 Dbl-like Rho-GEF genes in the human genome (Cook et al., 2014; Rossman et al., 2005). Dbl-like Rho-GEFs share a Dbl homology (DH) domain, which is responsible for their guanine nucleotide exchange activity, and a pleckstrin homology (PH) domain, which modulates the catalytic activity of the DH domain or mediates membrane association. Pleckstrin homology domain-containing family G member 4B (PLEKHG4B) is a Dbl-like Rho-GEF, with a structure closely related to that of Solo (also known as ARHGEF40) and PLEKHG4 (also known as puratrophin-1) (Cook et al., 2014). These three Rho-GEFs share a common domain structure with an N-terminal Solo domain, a CRAL/TRIO domain and spectrin repeats in the medial region, and a C-terminal DH-PH domain, except that PLEKHG4 lacks the N-terminal Solo domain (Fig. 1A). Solo activates RhoA and RhoC; PLEKHG4 activates RhoA, Rac1 and Cdc42; and PLEKHG4B activates Cdc42 (Abiko et al., 2015; Curtis et al., 2004; Gupta et al., 2013; Müller et al., 2020). We recently showed that Solo is required for tensional force-induced RhoA activation and stress fiber reinforcement in epithelial cells (Abiko et al., 2015; Fujiwara et al., 2016), as well as being involved in cyclic stretch-induced reorientation of vascular endothelial cells and collective migration and tubule morphogenesis of epithelial cells (Abiko et al., 2015; Isozaki et al., 2020; Nishimura et al., 2018). Although our understanding of the cellular roles of Solo has recently advanced, the function of PLEKHG4B remains to be investigated. Gene expression datasets (Expression Atlas, https://www.ebi.ac.uk) show that PLEKHG4B mRNA is widely expressed in mammalian tissues and cell types, with higher expression in testis, ovary, lung, kidney, glandular tissues and epithelial cell lines.
In this study, we provide evidence that PLEKHG4B enables actin cytoskeletal remodeling during epithelial cell–cell junction formation. Our results suggest that PLEKHG4B is involved in cell–cell junction maturation by reducing myosin activity at the late stage of cell–cell junction formation. We also demonstrate that annexin A2 (ANXA2) is involved in the localization of PLEKHG4B to cell–cell junctions.
PLEKHG4B overexpression induces F-actin-rich membrane protrusions toward adjacent cells
To investigate the cellular function of PLEKHG4B, we analyzed the effect of PLEKHG4B overexpression on the actin cytoskeletal organization and cell morphology. Human lung carcinoma-derived A549 epithelial cells were transfected with yellow fluorescent protein (YFP)-tagged PLEKHG4B or control YFP, and their actin filament organization was evaluated using rhodamine-labeled phalloidin staining. F-actin-rich filopodium-like membrane protrusions were observed in most YFP-PLEKHG4B-expressing cells, but not in control cells (Fig. 1B). Intriguingly, the protrusions extended only toward adjacent cells, and not toward the cell-free space (Fig. 1C). The x-z image analysis showed that the protrusions extended underneath the neighboring cell (Fig. S1A). In addition, the protrusions were hardly detectable in solitary cultured PLEKHG4B-overexpressing cells (Fig. S1B). These observations suggest that PLEKHG4B induces F-actin-rich membrane protrusions in a cell–cell-contact-dependent manner. It is conceivable that PLEKHG4B may play some role in actin cytoskeletal reorganization at cell–cell junctions. Staining with antibodies against E-cadherin (also known as CDH1) or β-catenin showed that both proteins were localized to the PLEKHG4B-induced protrusions in addition to cell–cell junctions (Fig. 1B,C). We also examined the effect of PLEKHG4B overexpression on actin filament organization and cell morphology in Madin-Darby canine kidney (MDCK) epithelial cells. In line with the findings in A549 cells, overexpression of YFP-PLEKHG4B induced F-actin-rich and E-cadherin-positive protrusions toward adjacent cells, although their shape was rounder than that of the protrusions in A549 cells (Fig. S1C).
PLEKHG4B-induced protrusion formation requires Rac1 and Cdc42 activity, but not RhoA activity
To elucidate which Rho GTPases are involved in the formation of PLEKHG4B-induced membrane protrusions, we evaluated the potential dominant-negative (DN) effects of Rho GTPase mutants. A549 cells were co-transfected with YFP-PLEKHG4B and cyan fluorescent protein (CFP)-tagged DN mutant of RhoA, Rac1 or Cdc42, and stained with rhodamine-phalloidin. DN Rac1 and Cdc42 markedly suppressed PLEKHG4B-induced protrusions, whereas DN RhoA had no apparent effect (Fig. 2A,B). We also examined the effects of Rac1, Cdc42 and Rho-associated kinase (ROCK; a downstream effector of RhoA) inhibition on PLEKHG4B-induced protrusion formation. Congruent with the results obtained using DN Rho GTPases, EHT-1864 (Rac1 inhibitor) and ML-141 (Cdc42 inhibitor), but not Y-27632 (ROCK inhibitor), significantly impaired PLEKHG4B-induced protrusion formation (Fig. 2C–F). These results suggest that the formation of PLEKHG4B-induced F-actin-rich membrane protrusions requires activation of Rac1 and Cdc42, but not RhoA.
Effects of PLEKHG4B knockdown on Rho GTPase activity
We next analyzed the effects of PLEKHG4B knockdown on RhoA, Rac1 and Cdc42 activity in cultured A549 monolayer cells. Cells were treated with control or PLEKHG4B-specific small interfering RNAs (siRNAs) and cultured for 48 h, and the cell lysates were subjected to glutathione S-transferase (GST) pull-down assays using the GST-fused RhoA-binding domain (RBD) of rhotekin and p21 (Rac1/Cdc42)-binding domain (PBD) of PAK1 to bind the active forms of RhoA and Rac1/Cdc42, respectively. Two independent PLEKHG4B-targeting siRNAs effectively suppressed the expression of PLEKHG4B transcripts in A549 cells (Fig. 3A). Knockdown efficiency of these siRNAs was also confirmed by immunoblot and immunostaining analyses of the expression levels of V5-tagged PLEKHG4B in MDCK cell lines stably expressing this protein (Fig. S2). GST pull-down assays, followed by quantitative analysis of the relative activity of each Rho GTPase (ratio of active-to-total Rho GTPase), revealed that PLEKHG4B knockdown significantly decreased the level of active Cdc42, but had no apparent effect on the level of active Rac1 (Fig. 3B,C). PLEKHG4B knockdown tended to increase the level of active RhoA, albeit not statistically significantly (Fig. 3B,C). These results suggest that Cdc42 is the primary target of PLEKHG4B, in accord with the recently reported results (Müller et al., 2020), and that PLEKHG4B knockdown may have the potential to increase RhoA activity in cells.
PLEKHG4B knockdown alters cell–cell junction formation
We next examined the effects of PLEKHG4B knockdown on actin filament organization and cell–cell adhesions. A549 cells were treated with PLEKHG4B-targeting siRNAs, cultured for 48 h to reach a subconfluent state, and then stained with rhodamine-phalloidin and anti-β-catenin antibody. Fluorescence microscopy revealed that PLEKHG4B knockdown markedly altered the organization of actin filaments and β-catenin. In cells treated with control siRNA, both actin filaments and β-catenin were mostly localized to cell–cell contact sites and aligned along the cell–cell boundary (Fig. 4A,B). This type of cell–cell junction is hereafter referred to as a ‘closed junction’ (Ozawa et al., 2020), in which actin bundles were closely packed together at the cell–cell contact sites (Fig. 4B). In contrast, in PLEKHG4B-silenced cells, actin bundles were split away from the cell–cell interface, and they were cross-bridged by β-catenin and thin actin fibers running perpendicularly to the cell–cell boundary (Fig. 4A,B). This type of cell–cell junction is hereafter referred to as an ‘open junction’ (Ozawa et al., 2020) (Fig. 4B). The actin and β-catenin organization observed in the open junction in PLEKHG4B-deficient cells was reminiscent of the zipper-like adhesions found in nascent cell–cell junctions (Vasioukhin et al., 2000; Vaezi et al., 2002; Yonemura et al., 1995). Indeed, the density of the open junction (measured as the length of the open junction as a percentage of the total length of the cell–cell contact) was significantly higher in PLEKHG4B knockdown cells than in control cells (Fig. 4C). These results suggest that PLEKHG4B is involved in the conversion of the open junction with split actin bundles to the closed junction with packed actin bundles during cell–cell adhesion formation.
PLEKHG4B knockdown delays cell–cell junction formation in Ca2+-switch assay
To further investigate the role of PLEKHG4B during cell–cell junction formation, we conducted a Ca2+-switch assay. We analyzed the effects of PLEKHG4B knockdown on the time-dependent changes in actin filaments and β-catenin organization during cell–cell adhesion formation. A549 cells were transfected with control or PLEKHG4B-targeting siRNA and cultured for 48 h until they formed a confluent monolayer. Then, the cells were incubated with EGTA to chelate Ca2+ and, therefore, disrupt cell–cell junctions. After 8 h of incubation, the medium was switched to a normal Ca2+-containing one to restore cell–cell adhesion. At 0, 2, 4, 8 and 24 h after Ca2+ replenishment, the cells were fixed and stained with rhodamine-phalloidin and anti-β-catenin antibody. Ca2+ depletion abrogated cell–cell adhesion, caused disappearance of β-catenin signals, and induced cell rounding in both control and PLEKHG4B-knockdown cells (Fig. 5A,B, 0 h). In control cells, actin bundles were detected along the cell periphery 2 h after Ca2+ restoration. At 4–8 h, there appeared the closed junction, where actin bundles of adjacent cells were closely packed to form linear actin bundles at the cell–cell boundary, while β-catenin signals became more intense in the cytoplasm and occasionally colocalized with, and accumulated on, the packed actin bundles of the closed junctions (Fig. 5A). At 24 h, β-catenin was concentrated at cell–cell junctions in most cells (Fig. 5A,C). In contrast, in PLEKHG4B-knockdown cells, actin bundles of adjacent cells were located away from the cell–cell boundary and remained split on the open junctions, and β-catenin signals remained weak, even 4–8 h after Ca2+ addition (Fig. 5B,C). Magnified images at 4 h clearly showed the difference in the mode of actin bundles at the cell–cell boundary between control and PLEKHG4B-knockdown cells (Fig. 5A,B, rightmost columns).
Further, the x-z images showed that control cells altered their morphology from round to squamous 4 h after Ca2+ replenishment (Fig. 5A), whereas PLEKHG4B-knockdown cells retained round cell shape, and their lateral adhesion was incomplete, even at the 8-h mark (Fig. 5B). To quantify the progression of junction formation, we analyzed the temporal changes in the total and lateral cell height by measuring the height of actin localization in whole cells and cell–cell adhesive regions in the x-z view (Kalaji et al., 2012). The total cell height did not significantly change during Ca2+-switch assay in control and PLEKHG4B-knockdown cells. In contrast, the lateral cell height was markedly increased 4–8 h after Ca2+ restoration in control cells, but not in PLEKHG4B-knockdown cells (Fig. 5D). Thus, PLEKHG4B knockdown appears to delay restoration of cell–cell junctions. These data imply that PLEKHG4B plays a crucial role in the maturation of cell–cell adhesions by promoting the conversion from the open junction with split actin bundles to the closed junction with packed actin bundles, and accumulation of β-catenin at cell–cell junctions.
We also examined the effects of PLEKHG4B knockdown on cell–cell adhesion formation in MDCK cells. Two independent siRNAs targeting dog PLEKHG4B effectively suppressed the expression of PLEKHG4B transcripts in MDCK cells (Fig. S3A). In steady-state MDCK cell monolayers, PLEKHG4B knockdown significantly decreased the intensity of E-cadherin staining at cell–cell junctions, but had no apparent effect on ZO-1 (also known as TJP1) staining (Fig. S3B,C), indicating that PLEKHG4B is involved in the regulation of the integrity of AJs, but not tight junctions. We also conducted a Ca2+-switch assay in MDCK cells. MDCK cells were transfected with control or PLEKHG4B-targeting siRNAs, cultured for 8 h in a low-Ca2+ medium to disrupt cell–cell adhesions, and then transferred to a normal-Ca2+ medium to restore them. MDCK cells restored cell–cell adhesions more rapidly than did A549 cells. In control MDCK cells, the closed junctions with linearly packed actin bundles and E-cadherin at the cell–cell boundary were detected as early as 15 min after Ca2+ addition, and further progressed at 30–60 min (Fig. S4A). In PLEKHG4B-knockdown cells, the closed junction was barely detected and E-cadherin was not apparently accumulated at the cell–cell boundary 15 min after Ca2+ restoration (Fig. S4B). Additionally, the x-z images showed that control cells started to form a squamous monolayer within 30 min, while PLEKHG4B-knockdown cells retained round shape even 1 h after Ca2+ replenishment (Fig. S4A,B). Quantitative analysis of the lateral cell height revealed that PLEKHG4B knockdown delays restoration of cell–cell junctions (Fig. S4C). Localization of ZO-1 to cell–cell adhesions was slightly delayed by PLEKHG4B knockdown. Collectively, these results are in agreement with those in A549 cells, and support that PLEKHG4B is involved in cell–cell adhesion maturation.
PLEKHG4B knockdown abrogates the decrease in ppMLC level in the late stage of cell–cell adhesion formation
Cell–cell adhesion formation has been shown to be regulated by myosin activity (Ivanov et al., 2007; Kishikawa et al., 2008; Miyake et al., 2006; Shewan et al., 2005; Zhang et al., 2005). Thus, we analyzed the effects of PLEKHG4B knockdown on the levels of Ser19-phosphorylated and Thr18/Ser19-diphosphorylated myosin regulatory light chain 2 (pMLC and ppMLC), which are widely used markers of active myosin II (Goeckeler and Wysolmerski, 1995; Mizutani et al., 2006). A549 and MDCK cells were transfected with either control or PLEKHG4B siRNAs, cultured for 48 h, and co-stained with rhodamine-phalloidin and anti-pMLC and anti-ppMLC antibodies. In PLEKHG4B-knockdown cells, actin bundles were split away from the cell–cell boundary to form the open junctions, and pMLC and ppMLC signals were observed on and near the split actin bundles, with the signal intensities markedly higher than those in control cells (Fig. S5A,B), indicating that PLEKHG4B depletion increases myosin activity in epithelial monolayer cells.
To further investigate the effects of PLEKHG4B knockdown on myosin activity during junction formation, A549 cells were transfected with either control or PLEKHG4B siRNAs and subjected to a Ca2+-switch assay, as described above. At 0, 2, 4 and 8 h after Ca2+ restoration, the cells were fixed and co-stained with rhodamine-phalloidin and anti-ppMLC antibody. In control cells, ppMLC signals on actin bundles were intensified at the 2-h mark before becoming weaker 4–8 h after Ca2+ replenishment (Fig. 6A). Concomitant with the decrease in ppMLC level, the open junctions with split actin bundles in adjacent cells were converted to the closed junctions with closely packed actin bundles at 4 h. In contrast, in PLEKHG4B-knockdown cells, ppMLC signals on split actin bundles were not decreased at 4–8 h, and the closed junction with packed actin cables had not been observed even 8 h after Ca2+ restoration (Fig. 6B). Immunoblotting also showed that the protein level of ppMLC was upregulated 2 h after Ca2+ addition in both control and PLEKHG4B-deficient cells. However, whereas in control cells the ppMLC level gradually decreased over the following 6 h, in PLEKHG4B-knockdown cells it remained elevated even 4–8 h after Ca2+ replenishment (Fig. 6C,D), indicating that PLEKHG4B is involved in the decrease in myosin activity in the late stage of cell–cell adhesion formation.
To investigate the role of the decrease in myosin activity in junctional maturation, we analyzed the effect of blebbistatin (a myosin II inhibitor) on junction formation in PLEKHG4B-knockdown cells in Ca2+-switch assay. A549 cells were treated with blebbistatin 2 h after Ca2+ restoration and stained for F-actin and ppMLC at 4 h. Treatment with blebbistatin decreased the level of ppMLC and partially induced the closed junction phenotype in PLEKHG4B-knockdown cells (Fig. 6E,F). Taken together, these results suggest that the decrease in myosin activity is correlated with the conversion of the open junction with split actin bundles to the closed junction with packed actin bundles, and that PLEKHG4B is likely involved in cell–cell junction maturation, at least in part, by decreasing myosin activity in the late stage of cell–cell adhesion formation.
A recent study reported that myosin IIA and IIB localize to distinct actin networks and differentially regulate AJ formation (Heuzé et al., 2019). However, myosin IIA and IIB mostly colocalized on the cell peripheral actin filaments and stress fibers in both A549 and MDCK cells under our experimental conditions (Fig. S5C,D). Their staining intensities appeared to increase in PLEKHG4B-knockdown cells.
ROCK and LARG/PDZ-RhoGEF are involved in aberrant myosin activation in PLEKHG4B-knockdown cells
ROCK is a key regulator of myosin II activity (Jaffe and Hall, 2005). To understand the mechanism underlying the aberrant myosin activation in PLEKHG4B-knockdown cells, we analyzed the effect of Y-27632 on the ppMLC level in PLEKHG4B-knockdown cells. A549 cells were treated with PLEKHG4B siRNAs and cultured for 48 h to form monolayers. Immunoblot and immunostaining analyses revealed that the level of ppMLC was increased in PLEKHG4B-knockdown cells, compared to those in control cells, but treatment with Y-27632 attenuated the PLEKHG4B-knockdown-induced increase in ppMLC (Fig. 6G,H). Immunostaining also showed that Y-27632 treatment recovered the defect in junction formation in PLEKHG4B-depleted cells, resulting in the formation of closed junctions (Fig. 6H). These results suggest that PLEKHG4B knockdown causes aberrant myosin activation and junctional defects through improper activation of the RhoA–ROCK pathway.
A recent study showed that PLEKHG4B interacts with two RhoA-targeting GEFs, LARG and PDZ-RhoGEF (also known as ARHGEF12 and ARHGEF11, respectively), and impairs their activities toward RhoA (Müller et al., 2020). We therefore examined the effects of knockdown of LARG and/or PDZ-RhoGEF on the increased level of ppMLC in PLEKHG4B-depleted cells. Immunoblot and immunostaining revealed that single or double knockdown of LARG and PDZ-RhoGEF attenuated the PLEKHG4B-depletion-induced elevation in ppMLC levels (Fig. 6I,J). Immunostaining also showed that double knockdown of LARG and PDZ-RhoGEF partially recovered the defect in junction formation in PLEKHG4B-depleted cells (Fig. 6J). Thus, it is likely that PLEKHG4B knockdown results in aberrant myosin activation and junctional defects through activation of LARG and PDZ-RhoGEF. Taken together, these results suggest that PLEKHG4B is involved in junction maturation by decreasing myosin activity in the late stage of junction formation through suppressing LARG and PDZ-RhoGEF activity and their downstream RhoA–ROCK pathway.
PLEKHG4B localizes to cell–cell adhesions
Next, we investigated the subcellular localization of PLEKHG4B using MDCK cell lines stably expressing V5-PLEKHG4B (MDCK/V5-PLEKHG4B cells). Immunoblotting confirmed that V5-PLEKHG4B was expressed at the predicted molecular size (Fig. 7A). The parental MDCK cells and MDCK/V5-PLEKHG4B cells were cultured until they formed a confluent monolayer, then stained with rhodamine-phalloidin and anti-V5 antibody to visualize the actin filaments and V5-PLEKHG4B, respectively. Confocal microscopic analyses revealed that V5-PLEKHG4B is non-uniformly but definitely located at the basal plane of cell–cell junctions, and z-stack images showed that it is also diffusely distributed in the nucleus and cytoplasm (Fig. 7B). The x-z images showed that V5-PLEKHG4B had a spot-like localization at the basal side and faintly at the apical side of cell–cell adhesions (Fig. 7B, arrowheads in x-z view), as well as diffuse distribution in the nucleus and cytoplasm. The localization of V5-PLEKHG4B at the basal plane of cell–cell junctions was reproducibly observed and partially colocalized with α-catenin (Fig. 7C), but not ZO-1 (Fig. S6A). V5-PLEKHG4B signals were also detected at the basal plane of cell–cell junctions throughout junction formation in Ca2+-switch assay (Fig. S6B). The localization of PLEKHG4B to the cell–cell junction and the nucleus was also observed for green fluorescent protein (GFP)-tagged PLEKHG4B in MDCK cells stably expressing it (Fig. S6C). The localization of PLEKHG4B to cell–cell adhesions further suggests the role of PLEKHG4B at cell–cell adhesions. In addition, compared with parental cells, MDCK/V5-PLEKHG4B cells exhibited wider F-actin organization at the basal side of cell–cell junctions, but did not form membrane protrusions toward adjacent cells (Fig. 7C), suggesting that the F-actin-rich protrusion formation observed in Fig. 1 is the result of overexpression of PLEKHG4B.
ANXA2 is required for PLEKHG4B localization to cell–cell adhesions
To gain insights into the mechanisms governing the localization and function of PLEKHG4B, we searched for its binding partners using proteomics analysis of co-precipitated proteins. A549 cells stably expressing Flag- and BirA-tagged PLEKHG4B were established. BirA (biotin ligase from Escherichia coli) was conjugated to PLEKHG4B for use in future proximity-dependent biotinylated protein identification (BioID) assays (Li et al., 2017), which were not a part of the current study. Flag-PLEKHG4B-BirA-expressing A549 cells and the parental cells were lysed and the cell lysates were immunoprecipitated with anti-Flag antibody. The co-precipitated proteins were separated by SDS–PAGE, stained with silver, and subjected to mass spectrometric analysis (Fig. 8A). Several proteins that were co-precipitated with Flag-PLEKHG4B-BirA were identified (Fig. 8B). Among these proteins, we focused on ANXA2, a phospholipid- and actin-binding protein that mediates the organization of membrane microdomains and regulates actin dynamics at specific membrane sites (Bharadwaj et al., 2013; Gerke et al., 2005). To confirm the interaction between ANXA2 and PLEKHG4B, CFP-tagged ANXA2 and Flag-PLEKHG4B-BirA were co-transfected into HEK293T cells. The cell lysates were immunoprecipitated with anti-Flag antibody, and the precipitates were analyzed by immunoblotting with anti-GFP and anti-Flag antibodies. CFP-ANXA2 was efficiently co-precipitated with Flag-PLEKHG4B-BirA, but not with control Flag-BirA (Fig. 8C), indicating that ANXA2 specifically interacts with PLEKHG4B in cultured cells. ANXA2 consists of the N-terminal cytosolic domain (amino acids 1–36) and the C-terminal membrane-bound domain (amino acids 37–339) (Gerke et al., 2005). To map the PLEKHG4B-binding region of ANXA2, we individually co-expressed CFP-ANXA2 or its N- or C-terminal fragments with Flag-PLEKHG4B-BirA and analyzed their PLEKHG4B-binding ability by co-precipitation assays. Both the N-terminal (1–36) and the C-terminal (37–339) fragments were co-precipitated with PLEKHG4B, but their binding abilities were weaker than that of full-length ANXA2 (Fig. 8D), indicating that both the N- and C-terminal domains of ANXA2 are required for its full PLEKHG4B-binding activity.
We next investigated the effect of ANXA2 knockdown on PLEKHG4B localization. Two ANXA2-targeting siRNAs efficiently suppressed the expression of endogenous ANXA2 protein in MDCK/V5-PLEKHG4B cells (Fig. 8E). ANXA2 knockdown decreased the localization of V5-PLEKHG4B to cell–cell adhesions, but not in the nucleus (Fig. 8F), suggesting the role of ANXA2 in the localization of PLEKHG4B to cell–cell adhesions. We also examined the effect of ANXA2 knockdown on epithelial junction integrity in MDCK cell monolayers. Similar to the effect of PLEKHG4B knockdown (Fig. S3B,C), ANXA2 knockdown significantly decreased the intensity of E-cadherin signals at cell–cell junctions (Fig. 8G,H). A previous study also showed that depletion of ANXA2 suppresses E-cadherin-dependent AJ formation (Yamada et al., 2005). These results suggest that ANXA2 is involved in epithelial junction integrity and that PLEKHG4B may participate, at least in part, in the function of ANXA2 in epithelial junction formation.
In this study, we showed that overexpression of PLEKHG4B induces F-actin-rich membrane protrusions. The protrusions were predominantly directed toward adjacent cells rather than the cell-free space, and were hardly detectable in solitary cultured cells, suggesting that the protrusion formation is cell–cell contact dependent. The PLEKHG4B-induced membrane protrusions were suppressed by blocking the activity of Rac1 or Cdc42, but not that of RhoA or ROCK, indicating that PLEKHG4B overexpression promotes membrane protrusions at cell–cell adhesions in a manner dependent on Rac1 and Cdc42 activity, but not the RhoA–ROCK pathway. GST pull-down assays revealed that PLEKHG4B knockdown significantly reduces Cdc42 activity, but not Rac1 activity, indicating that Cdc42 is the primary target of PLEKHG4B. This is consistent with the recently published data on a comprehensive analysis of Rho-GEF family target specificity (Müller et al., 2020). Rac1 inhibition interfered with PLEKHG4B-induced protrusion formation, probably because the basal level of Rac1 activity is required for membrane protrusion formation. Previous studies reported that cadherin-based cell–cell adhesions promote actin cytoskeletal remodeling through Rac1 and Cdc42 activation, and that several GEFs – such as Tuba, Tiam1 and Trio – are involved in this process (Braga, 2018; Citi et al., 2014; Takeichi, 2014). It is likely that PLEKHG4B cooperates with these GEFs to stimulate cell–cell contact-induced actin remodeling.
Epithelial cell–cell junction formation is a multistep process that involves actin cytoskeletal reorganization. We demonstrated that, in subconfluent A549 epithelial cells, knockdown of PLEKHG4B causes immature junction structure with distinct F-actin and β-catenin architecture. In control cells, the closed junctions with closely packed actin bundles and β-catenin were observed at the cell–cell boundary. In contrast, in PLEKHG4B-depleted cells, the open junctions with split actin bundles located away from the cell–cell boundary were observed, and β-catenin and thin actin fibers were located in between the split actin bundles in a zipper-like pattern, running perpendicularly to the cell–cell boundary. These results suggest that PLEKHG4B is required for the conversion of the open junction with split actin bundles to the closed junction with closely packed actin bundles during the maturation of cell–cell adhesions. Using Ca2+-switch assays, we demonstrated that PLEKHG4B depletion delays cell–cell adhesion formation. In control A549 cells, the closed junctions with non-split actin bundles were mostly formed 4 h after Ca2+ restoration. In contrast, in PLEKHG4B-depleted cells, they were hardly detectable at the same time point, and the open junctions with split actin bundles were predominantly observed instead. Localization of β-catenin to cell–cell adhesions and transition from round to squamous cell morphology were also markedly delayed in PLEKHG4B-deficient cells. Similar results were obtained in MDCK cells. Collectively, these results strongly suggest that PLEKHG4B is involved in actin cytoskeletal remodeling during the maturation of cell–cell junctions.
Further, in control cells, myosin activity (measured by the levels of ppMLC) increased 2 h after Ca2+ replenishment, then gradually decreased until the 8-h mark. The fall in ppMLC levels in the later stages of Ca2+ restoration was abrogated by PLEKHG4B knockdown. Between 2–4 h after Ca2+ addition, the open junctions were converted to the closed junctions in control cells, but such conversion was not achieved in PLEKHG4B-depleted cells. Thus, the reduction in myosin activity at 2–4 h correlates well with the conversion of the open to closed junctions, and PLEKHG4B is likely involved in this conversion via suppressing myosin activity at this stage. Treatment with blebbistatin 2 h after Ca2+ restoration induced closed junction formation in PLEKHG4B-knockdown cells at 4 h, supporting the notion that the decrease in myosin activity at this stage is involved in the conversion of open to closed junctions.
Myosin-mediated contractility has been shown to be required for the recruitment of E-cadherin to cell–cell contacts and the formation and stability of cell–cell junctions (Engl et al., 2014; Ivanov et al., 2007; Miyake et al., 2006; Shewan et al., 2005; Vicente-Manzanares et al., 2009). In these studies, inhibition of myosin activity suppressed cadherin-mediated junction formation. In contrast, we have shown here that inhibition of myosin activity promotes the conversion from the open junction to the closed junction during junction maturation. Similar results were observed previously, although the authors did not claim the conversion of junctional morphology (Zhang et al., 2005). In support of our results, Kishikawa et al. (2008) showed that aPKC-mediated antagonization against myosin-mediated contractile force is required for the process of conversion of open to closed junctions. In addition, the level of ppMLC in the split actin cables in the open junctions was shown to be higher than that in the non-split actin cables in the closed junctions in α-catenin-knockout cells (Ozawa et al., 2020). Intriguingly, we showed that myosin activity increases in the early stage of cell–cell junction formation, and thereafter decreases in the later stage in Ca2+-switch assays. Similar time-dependent changes in pMLC and ppMLC levels were also observed previously (Ivanov et al., 2005; Kishikawa et al., 2008; Zhang et al., 2005). Thus, it is plausible that both initial myosin activation and its following inactivation are required for the proper development of cell–cell junctions. It is likely that myosin plays both positive and negative roles in cell–cell junction formation, depending on the time and space of its action. Further studies are required to understand the mechanisms of how myosin activity is regulated time dependently and how myosin activation and inactivation drive the individual steps of cell–cell junction formation. It is also important to investigate whether the temporal changes in myosin activity generally occur and regulate cell–cell junction formation in other cell types.
To understand the mechanism via which PLEKHG4B knockdown increases myosin activity, we examined the effect of Y-27632. Treatment with this reagent attenuated the PLEKHG4B-knockdown-induced increase in ppMLC and restored the defect in junction formation in PLEKHG4B-depleted cells, suggesting that PLEKHG4B knockdown induces aberrant myosin activation and open junction phenotypes through activation of the RhoA–ROCK pathway. GST pull-down assays showed that PLEKHG4B knockdown had a tendency to increase RhoA activity in cells, which further supports this notion. Based on a recent report showing that PLEKHG4B suppresses activation of two RhoA-targeting GEFs, LARG and PDZ-RhoGEF (Müller et al., 2020), we tested the effects of their knockdown. Single or double knockdown of these Rho-GEFs decreased ppMLC levels and partially recovered closed junction phenotypes in PLEKHG4B-depleted cells. Collectively, these results suggest that PLEKHG4B is involved in the junction maturation probably by decreasing myosin activity at the late stage of cell–cell junction formation, through inhibition of LARG and PDZ-RhoGEF and their downstream RhoA–ROCK pathway.
In addition to its role in the late stage of cell–cell adhesions, PLEKHG4B might be involved in the membrane expansion step in the early stage of cell–cell adhesion formation via Cdc42 activation. Although PLEKHG4B overexpression induced membrane protrusions at cell–cell contact sites, its knockdown had no apparent effect on the early step of cell–cell adhesion formation, i.e. from initial cell–cell contacts to formation of open junctions (see Fig. 5C). Several Rho-GEFs can activate Rac1 and Cdc42 at cell–cell contact sites to promote membrane expansion in developing cell–cell adhesions (Arnold et al., 2017; Braga, 2018). Thus, the functional redundancy between PLEKHG4B and other Rho-GEFs might conceal the role of PLEKHG4B in the membrane expansion step.
Multiple Rho-GEFs and GAPs regulate the formation of cell–cell adhesions (Arnold et al., 2017; Braga, 2018). For example, Tuba, a Cdc42-targeting GEF, is concentrated in the apical region of cell–cell junctions, where it stimulates cell–cell adhesion formation (Otani et al., 2006). Tiam1, a Rac-targeting GEF, also promotes cell–cell adhesions; its activity is differentially regulated along the apicobasal axis of developing cell–cell junctions to facilitate the proper assembly of AJs and tight junctions and establishment of apicobasal cell polarity (Mack et al., 2012). The RhoA-targeting GAP p190RhoGAP is translocated to AJs and where it associates with p120-catenin upon Rac activation, and contributes to AJ formation by locally inhibiting RhoA activity (Wildenberg et al., 2006). Thus, it is conceivable that PLEKHG4B cooperates with various Rho-GEFs and Rho-GAPs to regulate actin rearrangements during cell–cell adhesion formation. Further research into the spatiotemporal control of Rho family regulators, and their crosstalk, is needed to delineate the mechanisms governing the dynamic control of cell–cell adhesion.
PLEKHG4B was localized to restricted areas of the apical and basal regions of cell–cell junctions in the epithelial cell sheet. Considering that AJs are formed at the apical sides of cell–cell contacts, the presence of PLEKHG4B at their basal interface suggests that the protein has a unique role in cell–cell adhesion. Thus, in addition to its involvement in apical cell–cell junctions, PLEKHG4B may regulate the formation and stability of the yet uncharacterized adhesive structure located at the basal ends of cell–cell junctions. It is also possible that PLEKHG4B acts to regulate basal actin fibers and indirectly affects apical junctional organization. Notably, a substantial amount of PLEKHG4B was also detected in the nuclei of epithelial cells. It would be interesting to investigate the significance and functional role of nuclear PLEKHG4B in a future study.
Finally, we revealed that ANXA2 co-precipitates with PLEKHG4B. ANXA2 knockdown suppressed the localization of PLEKHG4B to cell–cell adhesions, suggesting the role of ANXA2 in junctional localization of PLEKHG4B. Knockdown of ANXA2 decreased E-cadherin at cell–cell junctions in MDCK cells, as previously reported (Yamada et al., 2005). Similarly, knockdown of PLEKHG4B reduced E-cadherin at junctions. Thus, it is possible that ANXA2 is involved in the localization of E-cadherin to the cell–cell junction, at least in part, by recruiting PLEKHG4B to this site. ANXA2 is a multifunctional protein that binds to membrane phospholipids, such as phosphatidylinositol-4,5-bisphosphate, in a Ca2+-dependent manner (Bharadwaj et al., 2013; Gerke et al., 2005), participates in the dynamic organization of membrane microdomains, and recruits its interactors to the confined area of the plasma membrane. ANXA2 can also bind to actin filaments and their regulators, Rac1 and Cdc42, and thereby acts as a scaffold that connects the actin cytoskeleton to target sites on the plasma membrane (Bryant et al., 2010; Hansen et al., 2002; Ikebuchi and Waisman, 1990; Martin-Belmonte et al., 2007). Thus, ANXA2 and PLEKHG4B may act co-operatively to promote cell–cell junction formation by regulating actin dynamics at cell–cell contacts.
In conclusion, our results highlight a crucial role of PLEKHG4B in the reorganization of the actin cytoskeleton and formation of cell–cell adhesions. Further studies on the spatiotemporal regulation of Rho-GEFs and Rho-GAPs, and their interplay with the components of the actin cytoskeleton, cell adhesion molecules and cell polarity complexes, will advance our understanding of the mechanisms underlying the formation and remodeling of cell–cell adhesions, and their roles in epithelial development and homeostasis, as well as in cancer invasion and metastasis.
MATERIALS AND METHODS
Reagents and antibodies
Rhodamine-labeled phalloidin (181-02921) and Y-27632 (257-00511) were purchased from Wako Pure Chemical (Osaka, Japan). ML-141 (217708) and blebbistatin (203391) were obtained from Merck. EHT-1864 (CAY17258) was obtained from Funakoshi. Rabbit polyclonal antibodies against the following proteins were used: GFP [A6455, Life Technologies, 1:1000 for immunoblotting (IB)], E-cadherin [ab15148, Abcam, 1:500 for immunofluorescence (IF)], ppMLC (3674, Cell Signaling Technology, 1:500 for IB, 1:200 for IF), myosin IIB (909901, Biolegend, 1:200 for IF), GAPDH (2275-PC-100, R&D Systems, 1:1000 for IB), LARG (sc25638, Santa Cruz Biotechnology, 1:1000 for IB), PDZ-RhoGEF (ab11059, Abcam, 1:1000 for IB), ANXA2 (ab41803, Abcam, 1:2000 for IB) and α-catenin (C2081, Sigma-Aldrich, 1:500 for IF). Mouse monoclonal antibodies against the following proteins were used: RhoA (26C4, Santa Cruz Biotechnology, 1:250 for IB), Rac1 (05-389, Millipore, 1:1000 for IB), Cdc42 (610928, BD Transduction Laboratories, 1:250 for IB), β-catenin (610154, BD Transduction Laboratories, 1:500 for IF), E-cadherin (610181, BD Transduction Laboratories, 1:500 for IF), pMLC (3675, Cell Signaling Technology, 1:200 for IF), myosin IIA (ab55456, Abcam, 1:200 for IF), Flag M2 [F1804, Sigma-Aldrich, 1:1000 for IB, 1 μg/ml for immunoprecipitation (IP)], V5 (R960-25, Invitrogen, 1:1000 for IB, 1:500 for IF) and β-actin (ab6276, Abcam, 1:1000 for IB). Rat monoclonal antibody against ZO-1 (sc-33725, Santa Cruz Biotechnology, 1:750 for IF) was purchased from Santa Cruz Biotechnology. Alexa Fluor 488, Alexa Fluor 568, Alexa Fluor 633 and Alexa Fluor 647-conjugated goat anti-mouse IgG (A11029, A11031, A21052 and A21240, respectively), Alexa Fluor 488-conjugated goat anti-rabbit IgG (A11034) and Alexa Fluor 568-conjugated goat anti-rat IgG (A11077) were purchased from Thermo Fisher Scientific. Alexa Fluor 633-conjugated phalloidin (A22284) was purchased from Invitrogen. Horseradish peroxidase (HRP)-conjugated sheep anti-mouse IgG (NA931) and donkey anti-rabbit IgG (NA934) were purchased from GE Healthcare.
The siRNAs targeting human and dog PLEKHG4B, dog ANXA2, human LARG and PDZ-RhoGEF were purchased from Sigma-Aldrich. The target sequences are as follows: human PLEKHG4B #1 (5′-CUGCAAACCUGCUUGCUGA-3′) and #2 (5′-GCACCUUCCUUAUCCAGAA-3′); dog PLEKHG4B #1 (5′-GGUAUGUCAUUGACCAUUA-3′) and #2 (5′-GAAUAUUCACAGAACUCUA-3′); dog ANXA2 #1 (5′-GGAGUGAAGAGGAAAGGAA-3′) and #2 (5′-GGUCUGAAUUCAAGAGAAA-3′); human LARG #1 (5′-GATCGAATCATCAAGGTGA-3′); human PDZ-RhoGEF #1 (5′-GAGAUGAAACGGUCUCGAA-3′). MISSION siRNA Universal Negative Control (Sigma-Aldrich, SIC-001) was used as the negative control.
The cDNA of human PLEKHG4B (isoform 2, 0A1B0GW72-1 in UniProt), consisting of 1627 amino acids, was amplified by PCR using the Megaman Human Transcriptome Library (Biocompare, San Francisco, CA). Expression plasmids encoding YFP-tagged PLEKHG4B were constructed by inserting the PCR-amplified cDNA into the pEYFP-C1 vector (Clontech, Mountain View, CA). Expression plasmids encoding Flag-BirA-, V5- and GFP-tagged PLEKHG4B were constructed by inserting the PLEKHG4B cDNA and the BirA, V5 and GFP sequence, respectively, into the pCAG-GS-puro vector. BirA cDNA was a kind gift from Dr Hiroshi Masumoto (Nagasaki University, Nagasaki, Japan). The pCAG-GS-puro vector was generated by subcloning of the expression cassette of pPGK-puro into the SspI-SalI site of the pCAG-GS vector (Niwa et al., 1991). The cDNAs of DN mutants of Rho GTPases were provided by Dr Kozo Kaibuchi (Nagoya University, Nagoya, Japan). Expression plasmids for CFP-tagged DN mutants of Rho GTPases, and ANXA2 and its fragments, were constructed by inserting the PCR-amplified cDNAs into the pECFP-C1 vector (Clontech).
Cell culture and transfection
A549, MDCK and HEK293T cells were cultured in Dulbecco's modified Eagle medium (DMEM; 044-29765, Wako Pure Chemical) supplemented with 10% fetal calf serum (FCS; Cosmo Bio) at 37°C in 5% CO2. MDCK cells stably expressing V5- or GFP-PLEKHG4B are populational lines that were selected by adding puromycin (2 µg/ml) to the medium. In Fig. 7B, the cells were cultured on Transwell permeable support (3460, Corning). Plasmid transfection was performed using Lipofectamine LTX and Plus reagent (Thermo Fisher Scientific) for A549 and MDCK cells, and jetPEI (Polyplus) for HEK293T cells, according to the manufacturer’s protocols. Transfection of siRNAs was carried out using Lipofectamine RNAiMAX (Thermo Fisher Scientific). Briefly, A549 cells (2.3×105 cells/well) were seeded on coverslips in a 12-well culture plate and incubated with the siRNA transfection complex for at least 48 h. To efficiently introduce the siRNAs, MDCK cells were transfected twice: first they were seeded at 2.0×105 cells/well in a 12-well culture plate and incubated with the siRNA transfection complex for 24 h, then re-seeded at 3.5×105 cells/well and incubated with the siRNA transfection complex for an additional 48 h.
PLEKHG4B knockdown was confirmed by semi-quantitative RT-PCR. In brief, total RNA was extracted from the siRNA-transfected cells using the RNeasy mini kit and QIAshredder (Qiagen), and 1 µg of it was used as a template for first-strand cDNA synthesis using the cDNA Reverse Transcription Kit (Toyobo) or PrimeScript RT Reagent Kit (Takara), according to the manufacturer's protocol. The cDNA fragments were amplified by PCR using Ex Taq polymerase (Takara), subjected to agarose gel electrophoresis, imaged, and quantified by ethidium bromide staining. The expression level of GAPDH was used as an internal control. The following primers purchased from Eurofins Genomics were used for PCR primers: human PLEKHG4B (forward, 5′-TCCACAAGACCTGGGCATCCCGA-3′; reverse, 5′-TCTGGAGCTTCTGGGTGCCCTTTC-3′), human GAPDH (forward, 5′-GGATTTGGTCGTATTGGG-3′; reverse, 5′-GGAAGATGGTGATGGGATT-3′), dog PLEKHG4B (forward, 5′-CATGTGTTCCTCTTCGAAGACCTC-3′; reverse, 5′-CTTGTTGCCAATCCCCATTGACAC-3′) and dog GAPDH (forward, 5′-TGTCCCCACCCCCAATGTATC-3′; reverse, 5′-CTCCGATGCCTGCTTCACTACCTT-3′).
GST-RBD/PBD pull-down assay
To assess the effects of PLEKHG4B knockdown on RhoA and Rac1/Cdc42 activity, A549 cells were transfected with control or PLEKHG4B-targeting siRNAs, and the cell lysates were subjected to a pull-down assay using GST-fused RBD of rhotekin or PBD of PAK1, respectively, as described previously (Nishita et al., 2002). The precipitates were separated by SDS–PAGE, transferred to PVDF membranes, and immunoblotted for RhoA, Rac1 and Cdc42. The relative amount of the active form of each Rho GTPase was normalized to the total content of the corresponding Rho GTPase. The relative activity of each Rho GTPase was normalized to that in control cells.
For analyzing the interaction between PLEKHG4B and ANXA2, HEK293T cells were co-transfected with Flag-PLEKHG4B-BirA and CFP-ANXA2 or its mutants, cultured for 24 h, and lysed with ice-cold lysis buffer (25 mM Tris-HCl, pH 7.4, 1% Triton X-100, 140 mM NaCl, 2.5 mM MgCl2, 1 mM EGTA, 50 mM NaF, 1 mM Na3VO4, 1 mM PMSF, 10 µg/ml leupeptin and 2 µg/ml pepstatin A). Cell lysates were clarified by centrifugation at 18,000 g for 10 min and the supernatants were incubated with anti-Flag antibody and Protein G-Sepharose Fast Flow (GE Healthcare) for 2 h at 4°C. Immunoprecipitates were separated by SDS–PAGE and immunoblotted with anti-FLAG and anti-GFP antibodies.
Y-27632, EHT-1864 and ML-141 were used to inhibit ROCK, Rac1 and Cdc42 activity, respectively. A549 cells were transfected with YFP or YFP-PLEKHG4B expression plasmids, cultured for 3 days, then exposed to 10 µM Y-27632 for 30 min, 50 nM EHT-1864 for 1 h, or 20 µM ML-141 for 1 h. Water or dimethyl sulfoxide (DMSO) were used as vehicle controls for Y-27632 and EHT-1864/ML-141 treatment, respectively. Following incubation, the cells were subjected to immunostaining or immunoblot analysis. Blebbistatin was used to inhibit myosin activity during a Ca2+-switch assay. The cells treated with PLEKHG4B-targeting siRNA were subjected to a Ca2+-switch assay, treated with 20 µM blebbistatin 2 h after Ca2+ restoration, cultured for an additional 2 h, and then subjected to immunofluorescence and immunoblot analyses.
A549 cells were incubated in DMEM supplemented with 10% FBS and 2.2 mM EGTA to chelate Ca2+. After over 8 h of incubation, the cells were washed twice with PBS, and the culture medium was switched to standard DMEM containing 10% FBS. MDCK cells were incubated in DMEM without glutamine or Ca2+ (21068028, Gibco) supplemented with 5% Ca2+-depleted FBS and 3 µM Ca2+ for over 8 h. Ca2+-depleted FBS was made by mixing Chelex® 100 resin (142-2832, Bio-Rad) for 12 h, followed by filtration to remove chelated ions. Both cell lines were subjected to immunofluorescence analysis after the Ca2+ switch.
Immunofluorescence staining and fluorescence imaging
Cells were fixed with phosphate-buffered saline (PBS) containing 4% paraformaldehyde (PFA) for 20 min, and then permeabilized with 0.5% Triton X-100 in PBS for 10 min. After washing with PBS, cells were incubated with 2% fetal bovine serum (FBS) to reduce non-specific antibody binding, before they were probed with the primary antibodies overnight at 4°C. The next day, the cells were washed with PBS, incubated with the secondary antibodies and rhodamine-labeled phalloidin for 60 min, then washed with PBS. Fluorescence images were obtained using an LSM 510 and LSM 710 laser-scanning confocal microscopies (Carl Zeiss, Jena, Germany) equipped with a PL Apo 63× or 100× oil objective lens (1.4 NA). The z-section was acquired at a resolution of 0.5 µm z-stacks. The images were analyzed in ImageJ software. As for the analysis of V5-PLEKHG4B localization in MDCK monolayer cells (Fig. 7B), cells were loaded on the membrane of Transwell chamber (3460, Corning) and cultured for 48 h to accomplish well-polarized and cuboidal morphology, and then fixed and stained as above. Cells were co-stained with anti-V5 antibody, followed by Alexa Fluor 488-conjugated anti-mouse IgG, and Alexa Fluor 633-conjugated phalloidin. In other cases, cells were cultured on a cover glass.
A549 cells stably expressing Flag-PLEKHG4B-BirA and the parental cells were lysed with a lysis buffer [25 mM Tris-HCl, pH 7.4, 140 mM NaCl, 1 mM EGTA, 0.5% Triton X-100, PhosSTOP (04906837001, Roche), protease inhibitor cocktail (G6521, Promega) and 10 µg/ml RNase A]. After homogenization, the cell lysates were centrifuged at 100,000 g for 20 min, and the supernatants were subjected to immunoprecipitation using anti-Flag M2 antibody and Protein G-Sepharose (17-0618-01, GE Healthcare) at 4°C for 3 h. Sepharose was washed six times with a wash buffer (25 mM Tris-HCl, pH 7.4, 140 mM NaCl, 1 mM EGTA, 0.1% Triton X-100, PhosSTOP, protease inhibitor cocktail and 2 µg/ml RNase A). Bound proteins were eluted with an elution buffer (25 mM Tris-HCl, pH 7.4, 50 mM NaCl, 0.1% Triton X-100) containing 200 µg/ml 3× Flag peptide (CS-7068, ChemScene). The eluted proteins were separated by SDS–PAGE and trypsinized, before the recovered peptides were analyzed by matrix-assisted laser desorption/ionization time-of-flight tandem mass spectrometry (TOF/TOF 5800 system; AB Sciex, Framingham, MA). The proteins were identified using the MS-Fit software (http://prospector.ucsf.edu/prospector/mshome.htm).
Statistical data are expressed as the mean±s.e.m. of at least three independent experiments. All statistical analyses were carried out in Prism 6 (GraphPad Software, La Jolla, CA). P-values were calculated using the one-way ANOVA followed by Dunnett's test for multiple data set comparisons. In all cases, P<0.05 was considered statistically significant.
We thank Mr Shomu Komatsu for technical assistance and Drs Hiroshi Masumoto (Nagasaki University) and Kozo Kaibuchi (Nagoya University) for providing the cDNA constructs for BirA and Rho GTPases, respectively.
Conceptualization: K.N., K.M., K. Ohashi; Methodology: K.N.; Validation: K.N., K.Y.; Formal analysis: K.N., K. Ohta, K.Y.; Investigation: K.N., K. Ohta, K.Y.; Resources: K.N.; Data curation: K.N., K.Y.; Writing - original draft: K.N., K.M.; Writing - review & editing: K.N., K.M.; Visualization: K.N.; Supervision: K.M., K. Ohashi; Project administration: K.M., K. Ohashi; Funding acquisition: K.N., K.M., K. Ohashi.
This work was supported by Grants-in aid for Scientific Research from the Japan Society for the Promotion of Science (JSPS), KAKENHI (24370051 and 18K19280 to K.M.; 23112005, 20H03248 and 16K07335 to K.Ohashi; 20J12425 to K.N.), and the Japan Agency for Medical Research and Development (AMED) (19gm5810015h0004 to K.Ohashi).
Peer review history
The peer review history is available online at https://jcs.biologists.org/lookup/doi/10.1242/jcs.249078.reviewer-comments.pdf
The authors declare no competing or financial interests.