The canonical Wnt/β-catenin signaling pathway regulates cell proliferation in development and adult tissue homeostasis. Dysregulated signaling contributes to human diseases, in particular cancer. Growing evidence suggests a role for clathrin and/or endocytosis in the regulation of this pathway, but conflicting results exist and demand a deeper mechanistic understanding. We investigated the consequences of clathrin depletion on Wnt/β-catenin signaling in cell lines and found a pronounced reduction in β-catenin protein levels, which affects the amount of nuclear β-catenin and β-catenin target gene expression. Although we found no evidence that clathrin affects β-catenin levels via endocytosis or multivesicular endosome formation, an inhibition of protein transport through the biosynthetic pathway led to reduced levels of a Wnt co-receptor, low-density lipoprotein receptor-related protein 6 (LRP6), and cell adhesion molecules of the cadherin family, thereby affecting steady-state levels of β-catenin. We conclude that clathrin impacts on Wnt/β-catenin signaling by controlling exocytosis of transmembrane proteins, including cadherins and Wnt co-receptors that together control the membrane-bound and soluble pools of β-catenin.
The Wnt/β-catenin signaling pathway is an evolutionarily conserved pathway important for embryonal development and adult tissue homeostasis because it orchestrates cell proliferation and cell fate determination (Nusse and Clevers, 2017). Dysregulation of this pathway has been implicated in various human diseases and in cancer development in particular (MacDonald et al., 2009; Ng et al., 2019; Nusse and Clevers, 2017). β-Catenin is the central molecule of the canonical Wnt/β-catenin signaling pathway. It can act as a transcriptional co-activator together with transcription factors of the LEF/TCF family to drive expression of target genes that control cellular proliferation and differentiation programs. Cytosolic protein levels of β-catenin are usually kept low by a constant turnover through proteasomal degradation (Aberle et al., 1997). This process requires the so-called β-catenin destruction complex (also called degradasome), which consists of adenomatous polyposis coli proteins (APC/APC2), AXIN1/2, casein kinase 1 (CK1) and glycogen synthase kinase 3 (GSK3) and serves to phosphorylate β-catenin (Stamos and Weis, 2013). These phosphorylations function as a signal for subsequent ubiquitylation and degradation of β-catenin in the proteasome.
Wnt signaling can be activated by Wnt ligands, which upon binding to members of the Frizzled (FZD) family crosslink with their co-receptors, low-density lipoprotein receptor-related proteins 5/6 (LRP5/6). This leads to the phosphorylation of the cytosolic tails of LRP5/6, which in turn recruit components of the destruction complex (GKS3, AXIN1/2 and APC/APC2) and Dishevelled proteins (DVL1,2,3) to the plasma membrane, leading to the stabilization of β-catenin. The mechanism of β-catenin stabilization following ligand binding is still incompletely understood (Tortelote et al., 2017), but probably involves inhibition of the destruction complex through recruitment of degradasome components to the plasma membrane, forming so-called signalosomes (Bienz, 2014; Bilic et al., 2007). Alternatively, it has been suggested that ligand binding leads to LRP5/6 and GKS3 internalization into multivesicular endosomes, depleting the enzyme GSK3 from the cytosol and thus allowing the substrate β-catenin to accumulate (Taelman et al., 2010). Interestingly, both processes appear to require clathrin: in signalosome formation, clathrin-coated pits at the plasma membrane are thought to serve as sites of signalosome assembly (Gammons et al., 2016; Kim et al., 2013). For multivesicular endosome formation, the ESCRT (endosomal sorting complex required for transport) machinery cooperates with clathrin to form intraluminal vesicles (Raiborg et al., 2006; Wenzel et al., 2018).
The role of clathrin in canonical Wnt/β-catenin signaling remains poorly understood and even controversial (Brunt and Scholpp, 2018; Feng and Gao, 2015; Gagliardi et al., 2008). Although some Wnt components were shown to be directly regulated by vesicular trafficking (Feng and Gao, 2015), the net outcome of interfering with endocytosis or vesicle transport (dynamin inhibition) has shown partially conflicting results and requires mechanistic understanding (Brunt and Scholpp, 2018; Gagliardi et al., 2014, 2008).
We therefore set out to investigate the impact of clathrin depletion on Wnt/β-catenin signaling and the associated molecular mechanisms in several cell lines, focusing on HeLa cells and mouse L cells, which do not have mutations in Wnt/β-catenin signaling components.
Clathrin depletion reduces β-catenin protein levels
To investigate the role of clathrin in Wnt/β-catenin signaling, we depleted the clathrin heavy chain (CHC) in HeLaK cells with three different siRNA oligonucleotides (Fig. 1A) and observed a reduction in β-catenin protein levels with all three siRNAs to about 50% of endogenous β-catenin levels. To investigate whether clathrin also affects Wnt-3a stabilized β-catenin pools, we stimulated HeLaK cells with Wnt-3a conditioned medium for 2 and 5 h. Wnt-3a stimulation counteracted the reduction in β-catenin levels upon CHC depletion to a certain extent (Fig. 1B); however, clathrin depletion induced a reduction in β-catenin levels that was visible in both unstimulated basal conditions and upon Wnt-3a stimulation. Depleting different cell lines (MDA-MB-231, SW480 and RKO) for CHC resulted in a tendency for reduced β-catenin levels in all of them, but the extent of reduction was variable and less clear in comparison with HeLaK cells (Fig. S1). These marginal effects might be explained by cell line-specific factors (as discussed later) or by lower knockdown (KD) efficiencies in these cell lines. Indeed, a time course experiment in HeLaK cells indicated that complete depletion of CHC is necessary in order to affect β-catenin protein levels (Fig. 1C).
The clathrin-mediated decrease in β-catenin stability occurs at the protein level
Next, we wondered whether β-catenin levels are affected at the protein or mRNA level. Quantitative real-time PCR experiments showed that mRNA levels of β-catenin were not reduced in HeLaK cells upon siCHC treatment and even showed a tendency to increase (Fig. 2A). The same was true for MDA-MB-231 cells (Fig. S2A), which may indicate a compensatory upregulation of β-catenin mRNA expression, but clearly not a reduction upon clathrin depletion.
β-Catenin is constantly turned over via proteasomal degradation (Aberle et al., 1997). Inhibiting the proteasome with MG132 abrogated the reduction in β-catenin protein levels following clathrin depletion to a large extent (Fig. 2B). To verify the proteasome inhibitor experiments, we used HCT116 cells. HCT116 cells carry a mutation in β-catenin (ΔSer45), making it non-degradable by the proteasome (Morin et al., 1997). Despite a very good CHC KD efficiency, β-catenin protein levels were unaffected in HCT116 cells (Fig. S2B). Taken together, these findings clearly point to clathrin affecting β-catenin at the protein level.
We next addressed the role of the β-catenin destruction complex, which phosphorylates and marks β-catenin for ubiquitylation and subsequent proteasomal degradation: depleting APC, one of the key components of this enzyme complex, led to an increase in β-catenin protein levels as expected. Co-depletion of clathrin counteracted the effect of APC depletion (Fig. 2C). To manipulate the function of the destruction complex by another means, we inhibited GSK3 with indirubin or LiCl. In line with the results from the APC depletion experiment, GSK3 inhibition also resulted in an increase in β-catenin protein levels, which was reverted by clathrin depletion (Fig. 2D). Thus, disruption of the β-catenin destruction complex by APC depletion or GSK3 inhibition counteracts the clathrin depletion-induced reduction in β-catenin protein levels, but less efficiently than proteasome inhibition. In other words, inhibiting the destruction complex appeared to abrogate the effect of clathrin on β-catenin only partially. These results suggest that the canonical β-catenin turnover pathway via the destruction complex is not the sole mechanism by which clathrin affects β-catenin stability.
Clathrin depletion does not affect β-catenin protein levels via endocytosis inhibition or multivesicular endosome formation
Clathrin has several cellular functions, but is best known for mediating endocytosis, whereby clathrin triskelia form an endocytic coat on nascent endocytic vesicles. To assess whether interference with endocytosis affects β-catenin protein levels, we depleted the clathrin adaptor protein AP-2 by targeting two of its subunits (AP-2α and AP-2µ2) (Johannessen et al., 2006; Motley et al., 2003). We verified that KD of CHC and both AP-2 subunits were equally efficient in blocking endocytosis by testing transferrin uptake (transferrin is a typical cargo for clathrin-mediated endocytosis). Quantitative fluorescence microscopy analysis showed complete inhibition of endocytic uptake in all three conditions (Fig. S3). In contrast to clathrin, depletion of AP-2 did not reduce β-catenin protein levels (Fig. 3A), indicating that endocytosis does not play a role in affecting β-catenin levels in HeLaK cells. These results are in line with recent studies using KD and knockout model systems to abrogate endocytosis, which also found no effect on Wnt/β-catenin signaling (Rim et al., 2020).
Clathrin has another function on the limiting membrane of endosomes, where it cooperates with the ESCRT machinery to sort transmembrane proteins into intraluminal vesicles, thereby generating multivesicular endosomes (Raiborg et al., 2006; Wenzel et al., 2018). Intraluminal vesicle formation is a prerequisite for the degradation and downregulation of transmembrane cargo, such as growth factor receptors, but is also suggested to play a role in transmitting Wnt ligand-mediated signaling into the cell. Taelman et al. (2010) proposed that ESCRT-dependent multivesicular endosomes might sequester the destruction complex, and in particular GSK3, into intraluminal vesicles, thereby separating the enzyme from its substrate β-catenin and thus leading to β-catenin stabilization. To test whether the reduced β-catenin levels observed in clathrin-depleted cells were a result of the role of clathrin in formation of intraluminal vesicles, we depleted the ESCRT component HRS (hepatocyte growth factor receptor substrate). HRS is crucial for cargo sorting and downstream recruitment of the ESCRT machinery, but also for the recruitment of clathrin to the endosome membrane (Raiborg et al., 2001; Wenzel et al., 2018). HRS KD did not reduce β-catenin protein levels, neither in the steady state nor in Wnt-3a-stimulated cells (Fig. 3A). We conclude that, based on our findings with AP-2 and HRS depletion, clathrin does not affect β-catenin levels via clathrin-mediated endocytosis or multivesicular endosome formation.
Dynamin is a GTPase that functions in different clathrin-dependent cellular processes to mediate membrane fission (Henley et al., 1999). It has been shown previously that inhibiting dynamin with Dyngo-4a or Dynasore reduced Armadillo/β-Catenin protein levels in Drosophila and RKO cells, with a particularly strong effect in cells activated via Wnt signaling (Gagliardi et al., 2014). Inhibiting dynamin with Dyngo-4a in HeLaK cells reproduced the findings of Gagliardi et al. (2014), showing a weak reduction in β-catenin protein levels in steady-state and a clear reduction in Wnt signaling-activated cells (Wnt-3a-stimulated or GSK3-inhibited cells) (Fig. 3B).
The apparently contradictory findings with AP-2 depletion versus dynamin inhibition can probably be explained by the broader function of dynamin compared to AP-2 and possibly help to identify the mechanism of the effect of clathrin on β-catenin levels. Although the adaptor protein AP-2 functions specifically in clathrin-mediated endocytosis at the plasma membrane (Park and Guo, 2014), dynamin mediates membrane scission at multiple cellular compartments and may cooperate with clathrin not only during clathrin-mediated endocytosis, but also in membrane trafficking processes in the biosynthetic pathway (Henley et al., 1999). We therefore hypothesized that depletion of clathrin or dynamin affects intracellular trafficking of transmembrane proteins important for Wnt/β-catenin signaling.
LRP6 receptor shows reduced abundance and mislocalization in clathrin-depleted cells
We wondered whether the Wnt receptor and transmembrane protein LRP6 is affected by clathrin depletion. Western blot (WB) analysis showed that LRP6 was expressed in HeLaK cells, but at a low expression level (Fig. 4A). Therefore, we generated a stable cell line expressing fluorescently tagged LRP6 (LRP6-mNeongreen, ‘LRP6-mNG’) and quantified expression levels of endogenous and exogenous LRP6 protein in CHC KD cells. We saw a tendency for reduced protein levels of endogenous LRP6 in clathrin-depleted cells, which was more pronounced for the exogenous LRP6-mNG construct (Fig. 4A). To investigate the subcellular distribution and abundance of LRP6, we performed confocal fluorescence microscopy experiments. We could not detect endogenous LRP6 by immunostaining of parental HeLaK cells. Exogenous LRP6-mNG localized correctly at the plasma membrane of control cells, but total fluorescence intensity and plasma membrane localization were strongly reduced in CHC-depleted cells (Fig. 4B), which is in line with the findings from WB quantification (Fig. 4A).
To assess whether β-catenin stability might be affected via autocrine LRP5/6-mediated Wnt signaling in HeLaK cells, and whether LRP5/6 mis-trafficking in the absence of clathrin accounts for the destabilization of β-catenin, we generated LRP5/6 double-knockout (DKO) cells using CRISPR/Cas9. We identified several clones lacking protein expression of both LRP5 and LRP6, as judged by WB (Fig. 4C). In addition, we verified that the knockout was functionally complete, as the DKO clones failed to respond to Wnt-3a stimulation by stabilizing β-catenin (Fig. 4C). Importantly, the DKO clones appeared to have reduced levels of β-catenin, possibly mimicking the clathrin-depletion effect on β-catenin (Fig. 4D). However, KD of clathrin in the DKO clones clearly reduced β-catenin levels further (Fig. 4E), demanding an additional mechanistic explanation.
Clathrin depletion reduces the cell adhesion pool of β-catenin by impairing cadherin trafficking
β-Catenin resides in several cellular pools, either freely in the cytosol or bound to cell adhesion molecules of the cadherin family. We therefore set out to elucidate which cellular pool of β-catenin is affected by clathrin depletion. Fractionation experiments showed that the membrane-associated pool of β-catenin was reduced in clathrin-depleted cells (Fig. 5A) and we could verify this by immunofluorescence stainings, which showed a reduced fluorescence signal for β-catenin at cell–cell contact sites in siCHC-treated cells (Fig. 5B). To investigate the protein levels of cell adhesion molecules in the presence and absence of clathrin, we used a Pan-cadherin antibody for WB analysis and saw a reduction in a Pan-cadherin band upon clathrin KD (Fig. 5C). Unfortunately, we could not detect E-cadherin, N-cadherin or Pan-cadherin by immunofluorescence staining in HeLaK cells. Instead, we turned to HeLa-CCL2 cells, which are thought to be close to the original HeLa cell line but differ substantially on the genomic, proteomic and functional level compared to the HeLaK variant, which has a more uniform morphology and is particularly suitable for imaging (Liu et al., 2019). In addition, HeLa-CCL2 cells are known to express N-cadherin (Nakamura et al., 2008). HeLa-CCL2 cells also showed a reduction in Pan-cadherin levels upon clathrin depletion, detectable by both WB (Fig. 5D) and confocal microscopy (Fig. 5E). An antibody specific for N-cadherin detected a band with the same electrophoretic mobility, which was equally strongly reduced upon clathrin depletion (Fig. 5F), probably detecting the same protein as the Pan-cadherin antibody. Importantly, HeLa-CCL2 cells showed an efficient reduction in both β-catenin and N-cadherin upon CHC depletion (Fig. 5F,G) and the levels of reduction appeared to correlate with each other (Fig. S4A). Testing various cell lines showed typically a good correlation between Pan-cadherin and β-catenin levels following clathrin depletion (Fig. S4B). We conclude that impaired biogenesis of cadherin family members in clathrin-depleted cells leads to loss of the cell adhesion molecule-associated pool of β-catenin.
The cell adhesion pool and the soluble pool of β-catenin are controlled by clathrin
Next, we wondered whether we could manipulate the sensitivity of β-catenin to clathrin depletion by altering the expression of cadherin molecules. For this purpose, we used HeLaP cells and depleted them for N-cadherin. As expected, β-catenin levels followed the reduction of N-cadherin (Fig. 6A). An additional depletion of clathrin did not lead to a further reduction, making cadherin expression the main determinator of β-catenin stability in this cell line. However, the double KD appeared to impair the KD efficiency of N-cadherin, which may preclude a potential additional effect. We therefore employed another cell system: mouse L cells do not express cadherin molecules and we generated stable cell lines expressing cadherins (mouse E-cadherin–GFP, mouse N-cadherin–GFP and human E-cadherin–GFP) in these cells. Although parental L cells had barely detectable β-catenin levels, ectopic expression of all three cadherin variants stabilized β-catenin (Fig. 6B). Importantly, CHC KD indeed reduced the levels of β-catenin and of cadherin family members in the overexpression cell lines (Fig. 6B). Immunofluorescence staining for β-catenin and cadherin showed a strongly reduced signal in clathrin-depleted cells and the levels of β-catenin correlated well with the levels of cadherin family members (Fig. 6C). Interestingly, some cells showed an accumulation of cadherin molecules and β-catenin in a perinuclear region, which we identified as the trans-Golgi network (Fig. 6D). We interpret this accumulation as a sign of impaired anterograde trafficking in the absence of clathrin, whereby trans-Golgi-derived vesicles destined for the plasma membrane fail to be formed. This leads to an accumulation of both β-catenin and clathrin in the Golgi, which appears to be resolved in most cells, as indicated by the reduction of the overall fluorescent signal in most cells.
Next, we wanted to address the soluble β-catenin pool. For this purpose, we used L Wnt-3a cells, which stably express Wnt-3a ligand, leading to autocrine stimulation and elevated β-catenin levels compared with parental L cells (Fig. 6E). Importantly, we could also see a reduction in β-catenin levels upon clathrin depletion in these cells (Fig. 6E). Immunofluorescence staining (Fig. 6F) and fractionation experiments (Fig. 6G) showed that clathrin depletion can clearly affect both the membrane pool and the soluble pool of β-catenin.
Clathrin depletion leads to reduction in nuclear β-catenin levels and β-catenin target gene expression
Our data show that clathrin affects β-catenin protein levels by two means: it affects LRP5/6-mediated basal Wnt/β-catenin activity by regulating LRP5/6 biosynthesis and it affects the cell adhesion pool of β-catenin. Does the reduction in β-catenin levels upon clathrin depletion affect the transcriptional pool? The steady-state levels of β-catenin in the cytosol of HeLaK cells were close to our detection limit in WB, which made any further reduction hard to judge (Fig. 5A). However, fractionation experiments in which we purified the nuclear pool from HeLaK, HeLa-CCL2 or L cells clearly showed a reduction in nuclear β-catenin upon clathrin depletion in all three cell lines (Fig. 7A-C). Interestingly, this reduction was seen in cells where the cytosolic pool of β-catenin was high [Wnt-3a-stimulated HeLaK cells (Fig. 7A) and Wnt-3a-expressing L cells (Fig. 7B)] and in cells where the membrane pool of β-catenin dominates (HeLaP cells, N-cadherin expressing L cells) (Fig. 7B,C). Moreover, we investigated the consequences of clathrin depletion on Wnt/β-catenin target gene expression. KD of clathrin led to a slightly reduced expression of AXIN2 and cMyc mRNAs (Fig. 7D), indicating that the transcriptionally active pool of β-catenin can also be affected by clathrin.
The requirement for endocytosis in Wnt/β-catenin signaling has been controversial, and two lines of evidence point to an involvement of dynamin in Wnt signaling: a dominant-negative form of dynamin has been shown to affect Wnt target genes in Drosophila model systems (Piddini et al., 2005; Seto and Bellen, 2006) and inhibitors of dynamin have been used in cell culture model systems (Gagliardi et al., 2014). Effects on Wnt signaling were interpreted as being the result of inhibition of dynamin-dependent endocytosis (Gagliardi et al., 2014). In this study, we offer an alternative mechanistic explanation of how clathrin and dynamin affect Wnt/β-catenin signaling (Fig. 8).
We found no indications that clathrin-mediated endocytosis affects stimulated or unstimulated β-catenin levels and this is in accordance with recently published work using clathrin and AP-2 knockdown and knockout approaches (Rim et al., 2020). In line with Kim et al. (2013) and Gagliardi et al. (2014), we did not see any indications that LRP5/6 internalization is required for Wnt/β-catenin signaling. Clathrin depletion would in this case have led to an accumulation of LRP5/6 on the plasma membrane instead of reduced expression levels of LRP5/6. Instead, our data point to an inhibition in the biosynthetic pathway as the main mechanism, affecting both cell adhesion molecules of the cadherin family and LRP receptors.
How does clathrin affect the biosynthesis of cadherins and LRP receptors? Clathrin mediates vesicle budding from the trans-Golgi network during anterograde trafficking of newly synthesized transmembrane proteins from the endoplasmic reticulum (ER) to the plasma membrane (Bonifacino and Glick, 2004). Depletion of clathrin thus block anterograde protein transport, leading to protein accumulation in the Golgi and possibly ER, which could lead to ER stress, similar to that seen for Brefeldin A treatment (Moon et al., 2012). ER stress triggers the unfolded protein response (UPR), which alleviates the biosynthetic burden on the ER by downregulating protein synthesis and upregulating ER-associated degradation (ERAD) via the ubiquitin/proteasome system (Shaheen, 2018; Sun and Brodsky, 2019). This can reduce the protein level of transmembrane proteins such as LRP5/6 and cadherin molecules and could explain why inhibition of proteasomal degradation partially rescues the clathrin depletion-induced degradation of β-catenin (Fig. 2B). The consequence is a two-pronged mechanism of β-catenin inhibition. We show that HeLa cells appear to have a moderate basal autocrine Wnt-3a/β-catenin signaling level, which can be abrogated by knockout of the Wnt co-receptors LRP5/6, leading to a reduction in steady-state β-catenin levels (Fig. 4D). In L cells stably expressing Wnt-3a, this autocrine stimulation is even more prominent and we saw a strong reduction in soluble β-catenin levels upon clathrin depletion in these cells. In addition to the Wnt receptor-mediated effect, clathrin depletion leads to reduced levels of cell adhesion molecules of the cadherin family, which in turn reduces the cell adhesion pool of β-catenin. Importantly, in both cases, the reduced β-catenin protein levels resulted in reduced levels of nuclear β-catenin and reduced levels of β-catenin target genes.
The relative contributions of both mechanistic arms are highly cell line dependent, which can help explain confounding results from previous studies. Dependent on the proportion of cytosolic and adhesion molecule-associated β-catenin pools, various cell lines are expected to show variable sensitivities towards clathrin depletion. Although HeLaK cells have high levels of cell adhesion-associated β-catenin (Fig. 5A), SW480 cells have a high cytosolic pool and less β-catenin at contact sites (Wang et al., 2014), which can explain the marginal effect in CHC-depleted SW480 cells. In addition, SW480 cells are not expected to react to reduced LRP5/6 levels because their cytosolic β-catenin level is already maximal due to an APC mutation (Nishisho et al., 1991). Other cell lines that were investigated in this study (HeLaK, HeLa-CCL2, LentiX, RKO, MDA-MB-231 and L cells) do not have mutations in Wnt signaling components and are therefore able to keep cytosolic β-catenin levels low, making the cell adhesion pool the main contributor to total β-catenin levels. In these cell lines, we found a good correlation between Pan-cadherin levels and β-catenin, as expected.
The reduction in β-catenin protein levels induced by clathrin depletion was visible in unstimulated and Wnt-3a-stimulated conditions in HeLaK cells (Fig. 1B). By contrast, dynamin inhibition with Dyngo-4a led to a pronounced reduction in β-catenin levels, particularly in Wnt-3a-stimulated HeLaK cells (Fig. 3B) (Gagliardi et al., 2008). Although additional off-target effects of dynamin inhibitors cannot be ruled out (Preta et al., 2015), inhibition of dynamin can affect the biosynthetic pathway, but also dynamin-dependent endocytosis pathways such as caveolin-mediated endocytosis. Caveolins were found to be required for Wnt-3a stimulation-induced β-catenin stabilization (Yamamoto et al., 2006), offering a possible explanation for the additional effect of dynamin inhibition compared with clathrin depletion, particularly in Wnt-3a-stimulated HeLaK cells. The involvement of caveolins, clathrin and dynamin in canonical Wnt signaling has been controversial and the subject of numerous studies (reviewed by Brunt and Scholpp, 2018; Feng and Gao, 2015; Gagliardi et al., 2008). Our study now adds another layer of complexity by highlighting the role of the biosynthetic pathway in Wnt signaling, adding exocytosis to the debate on endocytosis in Wnt signaling activation.
MATERIALS AND METHODS
Cell culture and generation of stable cell lines
HeLaK (‘Kyoto’ were obtained from D. Gerlich, Institute of Molecular Biotechnology, Wien, Austria), HeLa-CCL2 (obtained from Institute Curie, Paris, France), MDA-MB-231 (HTB-26, ATCC), SW480 (CCL-228, ATCC), HCT116 (CCL-247, ATCC), L cells and L cells Wnt-3a (kindly provided by Mariann Bienz, Cambridge, UK) and RKO (CRL-2577, ATCC) cells were grown in DMEM high glucose (for HeLaK, HeLa-CCL2 and L cells; D0819 from Sigma-Aldrich) or RPMI (for MDA-MB-231 and SW480 cells; R2405 from Sigma Aldrich) or DMEM/F12 (for HCT116 and RKO; 31331-093 from Gibco) supplemented with 10% fetal calf serum, 100 U ml−1 penicillin and 100 µg ml−1 streptomycin and maintained at 37°C under 5% CO2. L cells were additionally supplemented with 1 mM sodium pyruvate. Cell lines were authenticated by genotyping and regularly tested for mycoplasma contamination.
HeLaK and mouse L cells stably expressing LRP6-mNG, TNKS1-mNG, mouse N-cadherin–GFP, mouse E-cadherin–GFP or human E-cadherin–GFP were generated using lentiviral transduction as described (Wenzel et al., 2018). Plasmids were generated by standard molecular biology methods; details and constructs can be obtained upon request to the authors. Human E-cadherin–GFP was Addgene plasmid #28009 (deposited by Jennifer Stow; Miranda et al., 2001), mouse E-cadherin-GFP was Addgene plasmid #67937 (deposited by Alpha Yap; Truffi et al., 2014) and mouse N-cadherin-EGFP was Addgene plasmid #18870 (deposited by Valeri Vasioukhin; Nechiporuk et al., 2007). To generate LRP5/6 double-knockout cell lines, HeLaK cells were transfected with plasmids encoding Cas9 and guide RNAs (kindly provided by Marianne Bienz and Melissa Gammons, Cambridge, UK) (Agajanian et al., 2019) targeting genomic loci of LRP5 (5′-CGTCCACCAGCCGTACGTCC) and LRP6 (5′-CGATTGGTTGATGCTACAAA). Clones were screened by immunoblotting.
siRNAs were purchased from Thermo Fisher Scientific (Ambion Silencer Select) or Dharmacon. Cells at 50% confluency were transfected using Lipofectamine RNAiMax transfection reagent (Life Technologies) following the manufacturer's instructions. Cells were transfected with siRNAs targeting the following: CHC, 5′-AUCCAAUUCGAAGACCAAU-3′ (Motley et al., 2003; Wenzel et al., 2018); CHC1, 5′-UAAUCCAAUUCGAAGACCAAU-3′ (Skånland et al., 2009); CHC2, 5′-GCAAUGAGCUGUUUGAAGA-3′ (Skånland et al., 2009); HRS 5′-GCACGUCUUUCCA GAAUUC-3′ (Wenzel et al., 2018); β-catenin, 5′-AGCTGATATTGATGGACAG-3′; APC, 5′-GACGUUGCGAGAAGUUGGA-3′ (Schneikert et al., 2013); AP-2α subunit, 5′-AAGAGCAUGUGCACGCUGGCCA-3′ (Johannessen et al., 2006; Motley et al., 2003); AP-2µ2 subunit, 5′-AAGUGGAUGCCUUUCGGGUCA-3′ (Johannessen et al., 2006; Motley et al., 2003); and N-cadherin/CDH2 (Ambion Pre-Designed oligos against CDH2, cat. no. s2771 and s2773). Non-targeting control siRNA from Thermo Fisher Scientific (Ambion Silencer Select, catalog number 4390844) or from Dharmacon (D-001810-01) were used as control.
Antibodies and reagents
We used the following antibodies at the dilutions stated for WB or immunofluorescence (IF): mouse anti-β-actin (A5316, Sigma-Aldrich; WB 1:10,000); mouse anti-β-catenin (610154, BD Transduction Laboratories; WB 1:1000, IF 1:400); rabbit anti-clathrin heavy chain (ab21679, Abcam; WB 1:1000); rabbit anti-HRS (Raiborg et al., 2001) (WB 1:1000); mouse-anti-mNG (32F6, Chromotek; IF 1:500); mouse anti-AP-2µ2 (612620, BD Transduction Laboratories; WB 1:500); mouse anti-AP-2α (sc-17771, Santa Cruz Biotechnologies; WB 1:1000); rabbit anti-Pan-cadherin (ab6529, Abcam; IF 1:100, WB 1:1000); mouse anti-N-cadherin (3B9, Invitrogen; IF 1:100), mouse anti-N-cadherin (610920, BD Biosciences; WB 1:1000); rabbit-anti-LRP5 (5731, Cell Signaling Technology; WB 1:1000); rabbit-anti-LRP6 (C5C7, Cell Signaling Technology; WB 1:1000), rabbit-anti-LaminA/C (EP4520, Epitomics; WB 1:10,000).
All secondary antibodies used for immunofluorescence studies were obtained from Jacksons ImmunoResearch Laboratories or from Molecular Probes (Life Technologies). Secondary antibodies used for western blotting were obtained from LI-COR Biosciences.
Wnt-3a- and control-conditioned media were generated from L-cells (ATCC CRL-2648) or Wnt-3a-producing L-cells (ATCC CRL-2647) according to specifications from ATCC. MG132 (C2211), dimethyl sulfoxide (DMSO) and indirubin-3′-oxime (I0404) were from Sigma-Aldrich. Dyngo-4a (ab120689) was from Abcam. G007-LK was a gift from Stefan Krauss and Jo Waaler, Oslo, Norway (Lau et al., 2013).
Immunoblotting and cell fractionations
Cells were washed with ice-cold PBS and lysed in 2× sample buffer (125 mM Tris-HCl pH 6.8, 4% SDS, 20% glycerol, 200 mM DTT and 0.004% bromophenol blue). Fractionations were done using the Mem-PER Plus Membrane Protein Extraction Kit (Thermo Fisher Scientific, #89842) according to manufacturer's specifications. To isolate detergent-insoluble proteins, we resuspended the last pellet in 2× sample buffer. To isolate nuclear proteins, we incubated the detergent-insoluble pellet in high salt buffer (300 mM NaCl, 50 mM TrisHCl pH 7.5, 0.5% TritonX-100) for 15 min at 4°C. Lysates were subjected to SDS-PAGE on 4-20% gradient gels (mini-PROTEAN TGX; Bio-Rad). Proteins were transferred to PVDF membranes (TransBlot Turbo LF PVDF, Bio-Rad) followed by antibody incubation in 2% BSA in Tris-buffered saline containing 0.1% Tween20. Membranes incubated with fluorescent secondary antibodies (IRDye680 or IRDye800; LI-COR) were developed with an Odyssey infrared scanner (LI-COR). Quantification of immunoblots was done using the Odyssey Software (version 3.0.30).
Transferrin uptake assay
Cells seeded on coverslips were starved for 45 min in serum-free medium before incubation with 25 µg ml−1 transferrin-AlexaFluor488 (Thermo Fisher T13342) in serum-free medium on ice for 30 min to allow ligand binding. The multiwell plates containing the coverslips with cells were then transferred to 37°C for 10 min to induce transferrin uptake; control coverslips were kept at 4°C to assess binding of transferrin-AlexaFluor488. Cells were then rinsed once in ice-cold PBS and washed for 40 s with ice-cold acidic stripping solution (500 mM NaCl and 100 mM glacial acetic acid in distilled water) to remove surface-bound transferrin. Cells were washed twice with ice-cold PBS and fixed with 3% formaldehyde at 4°C for 15 min. After PBS washes, coverslips were mounted in Mowiol containing 1 µg ml−1 Hoechst 33342 (Sigma-Aldrich) and imaged on a confocal microscope.
Immunostaining, confocal fluorescence microscopy and image analysis
Cells grown on coverslips were fixed in 3% formaldehyde for 10 min at room temperature or for 15 min on ice. Cells were washed twice in PBS and then permeabilized with 0.5% TritonX-100 for 5 min. Coverslips were then washed once in PBS containing 0.05% saponin before staining with the indicated primary antibodies for 1 h. After washing three times in 0.05% saponin in PBS, cells were stained with secondary antibodies for 1 h and washed three times in PBS. The cells were mounted in Mowiol containing 1 µg ml−1 Hoechst 33342 (Sigma-Aldrich).
Confocal fluorescence microscopy was carried out using a Zeiss LSM 710 or 780 microscope (Carl Zeiss MicroImaging) using standard filter sets and laser lines and a Plan Apo 63×1.4 N.A. oil lens. All images within one dataset were taken at fixed intensity settings below saturation.
Images were quantified using the Nikon NIS Elements software (version 5.11) using the General Analysis 3 package. The Hoechst channel was used to segment nuclei and to define ‘zones of influence’ corresponding to cell areas. For quantification of LRP intensity, the sum of intensities per cell was measured. For transferrin intensity quantification, transferrin spots were segmented using a suitable fixed threshold and the sum of intensities of the segmented transferrin spots per cell calculated.
Quantitative real-time PCR
Isolation of RNA and cDNA synthesis were performed using the Cell-to-Ct kit (Thermo Fisher Scientific, #AM1728) following the manufacturer's protocols. qPCR was performed with the TaqMan primers CTNNB1 #Hs00355049_m1, AXIN2 Hs00610344_m1, MYC #Hs99999003_m1 and GAPDH #Hs02786624_g1 (all from Thermo Fisher Scientific) and master mix (Thermo Fisher Scientific, #4369016), using cDNA from 100-500 cells per reaction. Relative quantifications were calculated using the 2-ΔΔCt method, with GAPDH as the endogenous control gene and negative control siRNA-treated sample as the calibrator.
Statistical analysis and considerations
Statistical analysis was done in GraphPad Prism Version 5.01. An unpaired t-test was used to test two samples with equal variance. For normalized datasets, a one-sample t-test was used. All error bars denote mean±s.d. or s.e.m., as indicated in the figure legends. n.s., not significant, *P<0.05, **P<0.01, ***P<0.001.
We thank Maria Lyngaas Torgersen for helpful discussions, Laura Sobotta for excellent technical help and Chema Bassols for IT assistance. The Core Facility for Advanced Light Microscopy at Oslo University Hospital is acknowledged for providing access to relevant microscopes.
Conceptualization: E.M.W.; Methodology: E.M., E.M.W.; Formal analysis: E.M., C.R., E.M.W.; Investigation: E.M., C.R., E.M.W.; Resources: C.R., H.S.; Writing - original draft: E.M.W.; Writing - review & editing: E.M., C.R., H.S., E.M.W.; Supervision: E.M.W.; Project administration: E.M.W.; Funding acquisition: E.M.W.
This work was supported by the South-Eastern Norway Regional Health Authority (Helse Sør-Øst RHF, grants 2015014 and 2016105 to E.M.W.).
Peer review history
The peer review history is available online at https://jcs.biologists.org/lookup/doi/10.1242/jcs.244467.reviewer-comments.pdf
The authors declare no competing or financial interests.