WNT-induced association of Frizzled and LRP6 is not sufficient for the initiation of WNT/β-catenin signaling

doi:10.21203/rs.3.rs-5238449/v1

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WNT-induced association of Frizzled and LRP6 is not sufficient for the initiation of WNT/β-catenin signaling

https://doi.org/10.21203/rs.3.rs-5238449/v1

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The Wingless/Int-1 (WNT) signaling network is essential to orchestrate central physiological processes such as embryonic development and tissue homeostasis. In the currently held tenet, WNT/β-catenin signaling is initiated by WNT-induced recruitment of Frizzleds (FZDs) and LRP5/6 followed by the formation of a multiprotein signalosome complex. Here, we use bioluminescence resonance energy transfer (BRET) to show that different WNT paralogs dynamically trigger FZD-LRP6 association. While WNT-induced receptor interaction was independent of C-terminal LRP6 phosphorylation, it was allosterically modulated by binding of the phosphoprotein Dishevelled (DVL) to FZD. WNT-16B emerged as a ligand of particular interest, as it efficiently promoted FZD-LRP6 association but, unlike WNT-3A, did not lead to WNT/β-catenin signaling. Transcriptomic analysis further revealed distinct transcriptional fingerprints of WNT-3A and WNT-16B stimulation in HEK293 cells. Additionally, single-molecule imaging demonstrated that, despite increasing FZD₅ and LRP6 confinement, WNT-16B stimulation did not result in formation of large receptor clusters, in contrast to WNT-3A. Our results suggest that FZD-WNT-LRP5/6 complex formation alone is not sufficient for the initiation of WNT/β-catenin signaling. Instead, we propose a two-step model, where initial ligand-induced FZD-LRP6 association must be followed by LRP6 phosphorylation and receptor clustering into higher-order complexes for efficient activation of WNT/β-catenin signaling.

Biological sciences/Cell biology/Cell signalling/Morphogen signalling

Biological sciences/Chemical biology/Pharmacology/Receptor pharmacology

Biological sciences/Chemical biology/Mechanism of action

WNT

signalosome

Frizzled

LRP5/6

BRET

GPCR

receptor cluster

single-molecule

RNA-seq

Wingless/Int-1 (WNT) lipoglycoproteins are morphogens of paramount importance in embryonic development, stem cell regulation, and tissue maintenance ^1,2. They initiate pleiotropic signaling cascades whose dysregulation results in pathologies such as diverse cancers and fibrosis, rendering components of the WNT signaling pathway attractive targets for therapeutic intervention ^1,3. WNT-dependent signaling cascades compose a tremendously complex network of pathways encompassing nineteen WNTs, their primary receptors (ten FZD paralogues, members of the class F of the G protein-coupled receptor superfamily), and diverse coreceptors (e.g., low-density lipoprotein receptor-related protein 5 and 6 (LRP5/6), receptor tyrosine kinase-like orphan receptor 1/2 (ROR1/2)) ⁴. On the intracellular side, the signal is propagated by two main downstream transducers, namely the phosphoprotein Disheveled (DVL) and heterotrimeric G proteins ^4–6. These components specify signaling that is characterized by the involvement of the transcriptional regulator β-catenin as either β-catenin-dependent (WNT/b-catenin signaling) or b-catenin-independent (e.g., planar-cell-polarity-like signaling). Signaling is initiated by WNT binding to the extracellular cysteine-rich domain (CRD) of FZDs. Simultaneous WNT binding to co-receptors contributes to signal specification and diversification, yet the molecular details remain poorly understood for most pathways. In WNT/β-catenin signaling, a signalosome model has emerged as a prevailing theory in the field ^1,3,7.

According to this model, WNT binding to FZD and the co-receptors LRP5/6 leads to the clustering of multiple FZD-WNT-LRP5/6 complexes, scaffolded by DVL, at the plasma membrane, forming the so-called signalosome ⁷. DVL condensates then serve as a docking platform for the multiprotein β-catenin destruction complex, that constitutively ubiquitinylates the transcriptional regulator β-catenin, dedicating it for proteasomal degradation ^3,8. In presence of WNTs, the destruction complex is disassembled and inactivated, and several components are recruited to the signalosome. As a result, β-catenin is stabilized and translocates to the nucleus, where it acts as a transcriptional regulator in concert with T-cell factor/lymphoid enhancer factor (TCF/LEF) transcription factors to induce the transcription of β-catenin target genes. This is accompanied by multiple biochemical changes, most prominently C-terminal phosphorylation of LRP6 and hyperphosphorylation of DVL.

More recently, synthetic activators of WNT/β-catenin signaling, known as WNT surrogates, were developed ^9–11. These surrogates mimic WNTs by binding simultaneously to FZD CRDs and LRP5/6, reinforcing the prevailing hypothesis that ligand-induced association of FZDs and LRP5/6 is not only necessary but also sufficient to initiate WNT/β-catenin signaling. Yet, due to the phylogenetic diversity of the 19 WNTs and ten FZD paralogs, it is reasonable to speculate that there are intricacies to the initiation of WNT/β-catenin signaling beyond purely crosslinking FZD and LRP5/6. In fact, FZD-WNT fusion proteins displayed FZD paralog-dependent differences in a WNT/β-catenin signaling output in vivo ¹². Further work unveiling essential conformational dynamics of the FZD core hinted at a more complex interplay of membrane proteins ^13–18.

In this work, we established a BRET assay between FZD and LRP5/6 to monitor the dynamic WNT-induced association between FZDs and LRP5/6. We demonstrated that FZD-LRP6 association was independent of LRP6 phosphorylation but was allosterically modulated by DVL. When probing WNT-FZD-LRP6 selectivity, we discovered that WNT-16B induced association of FZDs and LRP6 without triggering hallmarks of WNT/β-catenin signaling, such as LRP6 phosphorylation, signalosome formation, and TCF/LEF reporter gene activity. Based on these findings, we propose a model in which ligand-induced association of LRP6 with FZDs represents only the first step in signal initiation and is not necessarily followed by WNT/β-catenin, which additionally requires LRP6 phosphorylation and clustering of FZD and LRP6 into higher-order clusters.

A direct NanoBRET assay to capture dynamic association of FZD₅ and LRP6

To characterize the interactions between FZDs and LRP5/6 upon stimulation with WNTs, we established a direct NanoBRET assay using a C-terminally located Nanoluciferase (Nluc)/Venus system (Fig. 1A + B). For validation of the assay, we utilized FZD₅, which robustly activates WNT/β-catenin signaling ^14,19,20. Surface expression of all constructs used in BRET assays was analyzed by surface ELISA (Supp. Figure 1A + B). We could detect all FZD-Nluc/Venus and LRP-Nluc/Venus constructs at the cell surface with limited surface expression of LRP5 constructs and FZD₅-Venus, whose expression levels were below the detection threshold of the ELISA. LRP6 constructs required the co-expression of the chaperone MESD ²¹ for significant surface expression levels. Therefore, MESD was co-transfected in all experiments that included LRP5/6. Notably, co-expression of LRP6-Venus and FZD₅-Nluc decreased the surface expression of the latter, which nevertheless remained above the detection threshold (Supp. Figure 1C).

In acceptor titration NanoBRET experiments, we did not observe constitutive interactions between FZD₅-Nluc and LRP6-Venus (Supp. Figure 2A). Given that WNT-3A is known to elicit stabilization of β-catenin by recruiting both FZD₅ and LRP6, we next tested its effect in ligand stimulation experiments. Here, we observed a robust and dynamic BRET increase between FZD₅-Nluc and LRP6-Venus upon addition of recombinant purified WNT-3A in HEK293 cells, and a notably lower BRET increase between FZD₅-Nluc and LRP5-Venus, potentially due to lower surface expression of LRP5-Venus (Fig. 1C). The assay displayed similar results with an inverse probe setup using LRP5/6-Nluc and FZD₅-Venus (Supp. Figure 2B). We decided to continue with FZD-Nluc/LRP5/6-Venus probes since FZD₅-Venus showed low surface expression levels (Supp. Figure 1A). A dynamic BRET response was not observed upon stimulation of an unrelated GPCR (β₂-adrenoceptor) and a CRD-truncated FZD₅ construct (ΔCRD-FZD₅) in combination with LRP6, confirming assay specificity (Supp. Figure 2C). A potential impact of endogenously expressed LRP5/6 was addressed by performing the experiments in HEK293T ΔLRP5/6 cells (Fig. 1D). Kinetic traces obtained here closely resembled those obtained in regular HEK293 cells. Along these lines, stimulation with a WNT surrogate⁹ resulted in a strong increase in ΔBRET between FZD₅-Nluc and LRP6-Venus, and to a lesser extent between FZD₅ and LRP5-Venus, comparable to WNT-3A (Fig. 1E).

LRP5 and LRP6 transduce WNT signals with different efficacies, which was pinpointed to be a property of their C-terminal portions ²². As LRP5 displayed lower ΔBRET values, we reasoned that exchanging the LRP5/6 C-termini might change the magnitude of the response to that of the respective other paralog. However, the traces observed in BRET experiments using FZD₅-Nluc and an LRP6 chimera with an LRP5 C-tail (LRP6-5CT-Venus) or vice versa (LRP5-6CT-Venus) were practically unchanged compared to traces obtained with the respective wild-type LRP5/6-Venus (Fig. 1F). Notably, the cell surface expression of LRP5-Venus and LRP5-6CT-Venus did not surpass the detection threshold, while LRP6-5CT-Venus retained the approximate surface expression of LRP6-Venus (Supp. Figure 1). Our results suggest that the observed difference in ΔBRET between the traces of LRP5-Venus and LRP6-Venus did not result from a different conformational space obtained by their respective C-termini, but either from different expression levels or from their respective N-terminal/transmembrane domains.

Due to the lower surface expression of LRP5, we continued our experiments exclusively with LRP6. As WNT stimulation caused a dynamic BRET increase and constitutive interactions were not detectable, we conclude that our BRET assay measures WNT-induced association of FZD₅ and LRP6 rather than conformational dynamics of a pre-formed complex. Our findings contrast a study employing similar BRET probes that claimed a constitutive, but not dynamic interaction between mouse FZD₈ and LRP6 ²³.

LRP6 phosphorylation is not required for WNT-induced FZD-LRP6 association

WNT-induced association of LRP5/6 and FZD is a hallmark of WNT/β-catenin signaling. In the process of WNT/β-catenin signaling, five C-terminal PPP(S/T)P-motifs of LRP6 are phosphorylated by GSK3 and CK1 isoforms, components of the β-catenin destruction complex, presenting another proximal hallmark of the activation of the WNT/b-catenin pathway ²⁴. To determine whether LRP6 phosphorylation has an impact on receptor association as measured in the direct BRET assay, we generated an LRP6-Venus mutant lacking all phosphorylation sites in the C-tail (LRP6-5A-Venus; Fig. 1G) ²⁵. The mutant LRP6-5A-Venus expressed at the cell surface to a degree comparable to wild-type (WT) LRP6-Venus (Supp. Figure 1B). WNT-3A-stimulated BRET traces of FZD₅ and LRP6-5A-Venus behaved identical to WT LRP6-Venus (Fig. 1H), emphasizing that FZD₅-LRP6 association is independent of LRP6 phosphorylation.

To confirm signaling deficiency of the LRP6-5A mutant, we cloned a C-terminally untagged version of WT LRP6 and LRP6-5A. In agreement with the literature, LRP6-5A was in fact β-catenin signaling-incompetent both by overexpression and in response to WNTs, as assessed by TOPFlash reporter gene assays (Supp. Figure 3). In contrast to native LRP6, recombinant expression of LRP6-5A could not rescue TOPFlash reporter gene activity upon WNT-3A stimulation in ΔLRP5/6 cells and acted in a dominant-negative fashion when expressed in HEK293 cells, probably due to outcompeting endogenous LRP6. In summary, our data suggest that the phosphorylation status of the LRP6 C-terminus neither affected the WNT-3A-induced FZD₅-LRP6 association nor its C-terminal conformation, despite significantly different signaling outputs.

DVL modulates the conformation of the FZD-LRP6 complex

The phosphoprotein DVL is the main intracellular transducer of WNT-induced and FZD-mediated signaling and binds to FZDs with high affinity mainly via its Dishevelled, Egl-10 and Pleckstrin (DEP) domain ^17,20. DVL oligomers, polymerized by their Dishevelled/Axin (DIX) domains, are an integral part of the WNT receptor signalosome where they mediate interactions between the upstream WNT receptors and proteins of the β-catenin destruction complex. To assess the potential impact of DVL on the interaction between FZD₅-Nluc and LRP6-Venus, we have analyzed WNT-3A-induced BRET responses in mutational paradigms from two angles (Fig. 2A): (i) a receptor-based angle, where we have employed the FZD₅ R^6.32A mutant, which prefers G protein- over DVL-mediated signaling ¹⁶. (ii) a DVL-based angle employing HEK293 ΔDVL1-3 cells in combination with non-functional mutants of DVL2, i.e. the oligomerization-deficient M2/M4 mutant^26,27 as well as the L445E mutant incapable to bind to FZDs ^17,28.

In an initial experiment, we compared ΔBRET traces between LRP6-Venus and WT FZD₅-Nluc or FZD₅-Nluc R^6.32A upon WNT-3A stimulation and found no substantial difference between the traces (Fig. 2B). This suggested to us that DVL binding by FZD₅ would not modulate the WNT-3A-induced complex formation between FZD₅-Nluc and LRP6-Venus. However, when comparing the ΔBRET traces of WNT-3A-stimulated WT FZD₅-Nluc and LRP6-Venus between regular HEK293 cells and ΔDVL1-3 cells, the ΔBRET_max increased substantially when DVL was absent (Fig. 2C). As DVL supposedly serves as a platform allowing clustering of multiple FZD₅-WNT-LRP6 complexes into higher order complexes, the observed ΔBRET increase in ΔDVL1-3 cells came unexpected. We initially confirmed that DVL knockout and co-expression had no impact on FZD surface expression (Supp. Figure 4A). The increased ΔBRET in ΔDVL1-3 cells could, however, be lowered to a level similar to that observed in native HEK293 cells by co-transfection of DVL2. This suggested that the observed phenomenon is based on presence of DVL2 instead of a cell-line specific difference (Fig. 2C, Supp. Figure 4B). Transfection of the oligomerization-deficient DVL2 M2/M4 mutant into HEK293 ΔDVL1-3 cells showed the same effect as transfection of WT DVL2. DVL2 therefore modulates the interaction between FZD₅-Nluc and LRP6-Venus in a DIX polymerization-independent manner. Similarly, co-transfection of DVL1 or DVL3 led to similar reductions in ΔBRET_max (Fig. 2D, Supp. Figure 4C) indicating that there are no DVL paralog-specific effects.

Similar observations were made in a combined approach using HEK293 ΔDVL1-3 cells and the FZD₅ R^6.32A mutant, whereby ΔBRET between FZD₅ R^6.32A-Nluc and LRP6-Venus was increased in ΔDVL1-3 cells and could be lowered by recombinant DVL expression (Supp. Figure 4D + E). The ΔBRET_max observed for WT FZD₅ and the R^6.32A mutant in ΔDVL cells was not significantly different from each other, suggesting that WNT-induced FZD₅-LRP6 complex formation is similarly affected by the absence of DVL1-3 for both WT FZD₅ and FZD₅ R^6.32A (Supp. Figure 4F). Lastly, we utilized the DVL2 L445E mutant, whose DEP domain cannot interact with FZDs ^17,28, and a combined DVL2 M2/M4 L445E mutant. From this, we found that neither of the DVL2 L445E mutants could dampen the increased ΔBRET in ΔDVL1-3 cells, emphasizing that the FZD-DVL interaction is the main driver of the difference in WNT-3A-induced ΔBRET_max between HEK293 and ΔDVL cells (Fig. 2E, Supp. Figure 4G).

In conclusion, presence of FZD-binding DVL in the cell significantly decreases the dynamic BRET range between FZD₅-Nluc and LRP6-Venus upon WNT-3A stimulation, independent of DIX-dependent DVL polymerization and the DVL paralog. We interpret this data as evidence for a role of DVL in orchestrating the arrangement of FZD₅-Nluc and LRP6-Venus C-termini in a conformation disfavoring BRET (illustrated in Fig. 2F).

WNTs can stimulate FZD-LRP6 association without stabilizing β-catenin

To investigate FZD- and WNT-paralog selectivity, we transfected HEK293 cells with representative FZD-Nluc paralogs of the FZD homology clusters that are known to activate WNT/β-catenin signaling (FZD_4/5/7-Nluc) ^29,30 and LRP6-Venus, and stimulated them with diverse recombinant WNTs (WNT-3A, WNT-5A, WNT-10B, and WNT-16B) (Fig. 3A). FZD_4/5/7 displayed a robust BRET increase when stimulated with WNT-3A with differences in ΔBRET_max (FZD₅ > FZD_4/7). When stimulated with WNT-5A, most BRET traces showed a transient increase upon ligand addition, followed by a decrease to baseline. The signal remained slightly above the baseline for FZD₅ but displayed a markedly lower ΔBRET_max value compared to WNT-3A stimulation. WNT-10B stimulation yielded a weak signal for FZD₅ and FZD₇, but no reliable ΔBRET changes for FZD₄. WNT-16B stimulation resulted in a monophasic ΔBRET increase to a plateau similar to that of WNT-3A stimulation for all tested FZDs (with ΔBRET_max values FZD₅ > FZD₄ > FZD₇).

To determine whether the previously tested WNTs could activate WNT/β-catenin signaling, we investigated phosphorylation of LRP6 and DVL2 upon ligand stimulation in HEK293 cells by Western blotting (Fig. 3E-G). LRP6 phosphorylation was exclusively detected upon stimulation with WNT-3A or WNT surrogate, and not in response to WNT-5A, WNT-10B, or WNT-16B. Only WNT-3A, WNT-5A, and WNT-16B, however, led to a significant upward electrophoretic mobility shift of DVL2 indicative of WNT-induced DVL2 hyperphosphorylation (PS-DVL2) ³¹. In an orthogonal approach, we employed a TCF/LEF reporter gene assay (TOPFlash) as a measure of agonist-induced, β-catenin-dependent gene transcription ^32,33, yielding essentially the same results in HEK293 cells and HEK293 ΔFZD_1 − 10 cells specifically expressing FZD_4/5/7 (Fig. 3H; notably, the employed WNT surrogate does not bind to the FZD₄-CRD).

WNT-16B was of particular interest, as it induced association between FZD_4/5/7-Nluc and LRP6-Venus but neither LRP6 phosphorylation nor reporter gene activity. To exclude that quantitative FZD degradation upon WNT-16B treatment masks a potential TOPFlash signal, we treated samples with R-spondin 1 (RSPO1), which prevents FZD ubiquitinylation and dramatically amplifies WNT efficacy ^1,3. Co-treatment of HEK293 cells with WNT and RSPO1 greatly amplified the WNT-3A signal, but the WNT-16B signal remained indistinguishable from vehicle in TOPFlash reporter gene assays (Fig. 3I). Furthermore, we demonstrated that heat-inactivated WNT-16B (hi-WNT-16B) did not cause a ΔBRET increase between FZD₅-Nluc and LRP6-Venus (Supp. Figure 5A). Next, we set out to show that WNT-16B behaves similarly to WNT-3A in previously used BRET assay paradigms: Like WNT-3A, WNT-16B displayed no altered ΔBRET trace when using a phosphorylation-insensitive LRP6-5A-Venus mutant (Supp. Figure 5B), and it also displayed an increased ΔBRET in absence of DVL (Supp. Figure 5C).

In summary, we observed that WNT-16B does not induce β-catenin-dependent signaling in our cell model but leads to a detectable ΔBRET increase between FZD-Nluc and LRP6-Venus probes. Stimulation with WNT-16B is also not accompanied by LRP6 phosphorylation characteristic for active WNT/β-catenin signaling. Our observations infer that WNT-induced FZD-LRP6 association as measured by BRET is not sufficient to induce β-catenin-stabilization. Despite the inability of WNT-16B to elicit β-catenin-dependent signaling, the positive efficacy of WNT-16B in the BRET assay and the electrophoretic mobility shift of DVL2 emphasize the protein’s ability to interact with its receptors and elicit functional downstream events.

WNT-3A and WNT-16B induce different transcriptome changes in HEK293 cells

The signaling activity of WNT-16B has remained an enigma for quite some time ³⁴. To gain an unbiased perspective, we performed poly-A enriched bulk mRNA sequencing of HEK293 cells treated with vehicle control, WNT-3A, or WNT-16B. HEK293 cells were cultured in complete medium supplemented with 10 nM C59 to suppress endogenous WNT secretion for 24 h and were subsequently stimulated for another 24 h with recombinant WNTs. After variance stabilization transformation, the principal component analysis (PCA) of RNA-seq data demonstrated clear separation of cells by treatment (Supp. Figure 6). Untreated samples clustered quite closely to WNT-3A treated samples, which may hint at basal WNT-3A-like signaling in the sample cells. For differential gene expression analyses, thresholds were set to p-value < 0.05 and log2 fold-change of > 1.5. Volcano plots (Fig. 4A-C) depict significantly up- (red) and down-regulated (blue) transcripts. Stimulation with each WNT resulted in a unique mRNA transcriptome signature relative to vehicle-treated HEK293 cells (Fig. 4A,B; see Supp. Table 1 for datasets of differentially expressed genes). Direct comparison between WNT-3A and WNT-16B treatment also revealed a distinct mRNA signature (Fig. 4C). Genes regulated by one WNT sometimes correlated with genes regulated by the other WNT (Fig. 4D), indicating that some transcriptional responses were shared between the treatments, while the majority (1,206 out of 1,593 comparisons) did not meet significance criteria in both datasets (also see Fig. 4C). We conclude from this unbiased approach (i) that WNT-16B is transcriptionally active even though it was not inducing a TCF/LEF-dependent response in TOPFlash assays, and (ii) that despite some shared target genes, WNT-3A and WNT-16B mediate largely different transcriptional programs, reflecting the observed differences regarding the biophysical readouts and the TOPFlash data.

WNT-3A and WNT-16B induce FZD₅-LRP6 protein complexes with distinct architecture

WNT-3A and WNT-16B can interact with the same set of cell surface receptors but mediate different cellular signaling programs with respect to the activation of β-catenin-dependent signaling. How does WNT-receptor activation differ to accomplish distinct cellular responses? We hypothesized that the different signaling outputs of WNT-3A and WNT-16B originate from differences in the molecular interaction between FZD and LRP6, which could not be captured by ensemble methods such as BRET. To this end, we employed dual-color single-molecule fluorescence microscopy to track FZD₅ and LRP6 in live cells in response to stimulation with either WNT-3A or WNT-16B.

We transiently transfected N-terminally tagged SNAP-FZD₅ and Halo-LRP6 constructs into Chinese Hamster Ovary K1 (CHO-K1) cells to achieve low physiological molecule density (FZD₅: 0.42 ± 0.1, LRP6: 0.59 ± 0.15 molecules/µm²). Receptors were labeled with saturating concentrations of SNAP SiR-647 and Halo R110 fluorophores, respectively, and imaged with fast, multi-color total internal reflection fluorescence (TIRF) microscopy, combined with single-particle tracking (Fig. 5A + B). Data were acquired both under basal (Supp. Video 1 + 2 for raw movies and single-particle tracking with additional plotting of interactions) and after early (2–10 min) and late (11–25 min) stimulations with WNTs (Supp. Video 3–10). When analyzed by time-averaged mean squared displacement, FZD₅ and LRP6 molecules explored a range of diffusion profiles on the plasma membrane, alternating between confined and random Brownian diffusion (Supp. Figure 7A). Both WNT-3A and WNT-16B stimulations increased the proportion of confined FZD₅ and LRP6 molecules (Fig. 5C). To estimate the frequency and duration of FZD₅ and LRP6 interactions, we applied previously developed methods based on deconvolution of apparent colocalization times with those of random colocalizations ³⁵. Random colocalization times were estimated by imaging SNAP-β₂-adrenoceptors (β₂ARs), a prototypical GPCR with similar diffusion properties to that of FZD₅ (expressed at similar densities 0.4 ± 0.08 molecules/µm²), and Halo-LRP6. In the absence of WNTs, FZD₅ and LRP6 molecules did not colocalize for longer than what we measured for random colocalizations between SNAP-β₂AR and Halo-LRP6 confirming our data from BRET acceptor titration experiments (Supp. Figure 2A, Supp. Videos 11 + 12).

Following stimulation, both WNT-3A and WNT-16B caused a substantial increase in FZD₅ and LRP6 association rates at both early and late stimulation time-points. Notably, WNT-3A induced a ~ 1.5 and ~ 10-fold increase in association rate compared (k_on) to WNT-16B at early and late stimulation time-points, respectively (Fig. 5D, left). We estimated that following early and late WNT-16B stimulation, FZD₅ and LRP6 interactions lasted ~ 0.76 s and ~ 1.57 s, respectively (k_off early = 1.32 s^− 1, 95% confidence interval (CI): 0.81–1.84; k_off late = 0.64 s^− 1, 95% CI: 0.52–0.75). WNT-3A induced a ~ 2-fold (early and late) increase in FZD₅-LRP6 interaction times, compared to WNT-16B (k_off early = 0.65 s^− 1, 95% CI: 0.37–0.92; k_off late = 0.3 s^− 1, 95% CI: 0.24–0.36) (Fig. 5D, right). Importantly, WNT-3A interaction times at late stimulation time-points were reaching the limit of the observation window in our experiments, which indicated that the true length of interactions following WNT-3A late stimulation are in fact longer than measured. A spatial confinement analysis revealed that both ligands induced receptor interactions in a co-confined state, where FZD₅ and LRP6 are confined together, rather than a co-diffusing state, where both receptors move together across the plasma membrane. However, only WNT-3A showed statistically significant increase in receptor co-confinement. This shift from co-diffusion to co-confinement is reflected by the inverted pattern in the graph depicting the relative fraction of detected interactions (Fig. 5E). In addition, at the late stimulation time-point, we observed that WNT-3A induced aggregation of both FZD₅ and LRP6 into clustered, higher-order complexes, which were not observed when cells were stimulated with WNT-16B (Fig. 5F). Furthermore, we exploited the single-molecule data to analyze the size of such complexes. Monomeric, fluorescent particles are expected to photobleach in a single step, whereas higher order oligomers follow a stepwise photobleaching pattern (Supp. Figure 7B). In response to WNT-16B-stimulation, stepwise photobleaching analysis showed that co-confined FZD₅ and LRP6 molecules were largely monomeric (~ 60% and 80%, respectively). Conversely, WNT-3A-stimulated cells had a smaller fraction of monomeric co-confined FZD₅ and LRP6 molecules (~ 20%) and higher numbers of molecules that photobleached in multiple steps (up to 15), indicating the presence of higher-order clusters (Fig. 5G). Notably, by the time receptors had aggregated in higher-order clusters in the single-molecule fluorescence microscopy experiments, ΔBRET traces had already reached their plateau (> 10 min).

While stimulation with both WNTs increased receptor confinement, co-confinement, and interaction duration, the characteristic formation of FZD₅-LRP6 higher-order clusters was more prominent with WNT-3A than WNT-16B. These findings provide a mechanistic explanation for the lack of effective WNT/β-catenin signaling upon WNT-16B stimulation. Thus, ligand-induced receptor association and confinement, as seen with WNT-16B, is not sufficient for WNT/β-catenin signaling, which requires the formation of higher-order clusters.

In the present work, we use a combination of BRET assays, biochemical signaling readouts, transcriptomics, and single-molecule fluorescence microscopy to further delineate the signal initiation and specification in WNT/β-catenin signaling. We primarily base our work on a pharmacological comparison between WNT-3A and WNT-16B (see Fig. 6A), acting on FZD₅ and LRP6, which serve as model receptors throughout the study. The central finding of this work is that the addition of WNT-16B leads to association of FZD₅ and LRP6 but does not feed into WNT/β-catenin signaling. WNT-16B is an endogenous ligand that challenges the tenet that β-catenin signaling necessarily follows from ligand-induced interaction between FZDs and LRP6, as implied by the development of WNT surrogates that act as extracellular FZD-LRP6 crosslinkers to efficaciously activate WNT/β-catenin signaling ^9–11,36,37

The established direct NanoBRET assay between FZD₅ and LRP6 provides insight into the process of ligand-induced association between FZDs and LRP6. Firstly, we observed that FZD₅-LRP6 association is independent from LRP6 phosphorylation, inferring that receptor association is not modulated by downstream signaling events (see Fig. 1G + H, Supp. Figure 3). Secondly, the interaction between FZD₅ and LRP6 is exclusively ligand-induced. While this opposes a study claiming direct and constitutive interaction between FZD₈/LRP6 extracellular domains ²³, it is well in line with observations from single-molecule fluorescence microscopy ⁹.

Furthermore, we found that the orientation of the FZD₅ and LRP6 C-termini is allosterically modulated by DVL binding to FZD, where the presence of DVL results in a lowered WNT-induced ΔBRET_max (see Fig. 2, Supp. Figure 5C). This observation was independent of DVL polymerization and may speculatively be explained by DVL acting as a steric hindrance, impeding optimal Nluc-Venus energy transfer, or a scaffold for the FZD₅ C-terminus. DVL generally appears to be dispensable for the initial ligand-induced association of FZD and LRP6, yet the increased dynamic BRET range provides evidence that DVL already interacts with the FZD-LRP6 complex at this early stage of signal initiation. Combined with the recent observation that a WNT-induced change in the FZD-DEP interface was independent of LRP5/6 ^17,38, our data suggest that FZD-DVL interactions precede LRP6 association.

The observed ligand-induced association between FZDs and LRP6 as measured by BRET occurs in response to WNTs that do not elicit a TOPFlash signal (i.e., several WNT-FZD-LRP6 combinations result in a positive BRET shift, but not in a reporter gene signal, compare Fig. 3A-D and Fig. 3H). Thus, our experiments dissect receptor association from the WNT/b-catenin signaling output assessed by TOPFlash. Therefore, we suggest that BRET measurements do not exclusively capture the formation of signaling-competent receptor clusters, but especially the preceding FZD-LRP6 association and (co)-confinement (see also Fig. 5C-E), which does not necessarily provide a signaling-competent platform. The signal specification into WNT/β-catenin signaling must be taking place downstream of the initial association of FZD₅ and LRP6. Of note WNT-5A, a WNT that generally does not act via the WNT/β-catenin pathway ^39,40, resulted in a low positive ΔBRET at FZD₅. This aligns well with a report showing physical interaction between LRP6 and WNT-5A ⁴¹, but challenges the hypothesis that WNT signal specification is solely achieved by engagement of different co-receptors ^42,43.

Using single-molecule fluorescence microscopy we established that at comparable receptor expression levels, WNT-3A and WNT-16B behave differently with regards to the extent of receptor clustering over time (Fig. 5G), yet both ligands still confine FZD₅ and LRP6 (Fig. 5C, E), underlining their ability to interact with both receptors. It is of paramount importance to emphasize that WNT-16B stimulation still translated into substantial transcriptomic changes in HEK293 cells, despite being inactive in TOPFlash reporter gene assays (compare Fig. 3H + I with Fig. 4B + C), suggesting the activation of β-catenin-independent pathways. Whether LRP6 is relevant for the signal specification after WNT-16B stimulation or whether the signal is further specified by co-receptors other than LRP6 (or even in a DVL-independent manner) cannot be stated from results obtained here, and a further dissection of WNT-16B signaling mechanisms in HEK293 cells is beyond the scope of this work.

Lastly, we observed higher order clustering of receptors only after approx. 10 min of WNT-3A stimulation. Yet, receptor confinement and co-confinement had been observed prior to that, indicating that lower-order clustering or association of FZD₅ and LRP6 precede the formation of larger, signaling-competent signalosomes feeding into the WNT/b-catenin pathway. As a result, the term WNT/b-catenin signalosome describing a signaling-competent receptor agglomerate must be defined as a higher-order cluster rather than the minimal WNT/FZD/LRP/DVL complex as often depicted schematically ³. From a kinetic perspective, BRET traces reached their plateau within 10 min, again suggesting that they capture initial FZD-LRP5/6 co-confinement upon WNT stimulation and not higher order clustering. The later data points obviously monitor both association and clustering, however the formation of higher order structures does not further elevate the BRET signal.

How does signal initiation elicited by WNT-3A acting at FZD₅ and LRP6 differ from that in response to WNT-16B acting at the same receptors? We demonstrate that further mechanisms beyond ligand-induced FZD-LRP6 association are required to activate the WNT/β-catenin signaling pathway – specifically the phosphorylation of LRP6 and the formation of higher order FZD₅-LRP6 clusters are absent upon WNT-16B stimulation. These additional mechanisms are also activated by WNT surrogates, that fulfill all hallmarks of WNT/β-catenin signaling, even when engaging FZD₆, a receptor that typically does not signal via β-catenin ³⁶. Multivalency of WNT surrogates was described to significantly boost their efficacy, and one study found that a monovalent crosslinker (simultaneously binding 1 FZD and 1 LRP6 molecule) was inactive in reporter gene assays ¹¹. Yet WNTs themselves and first-generation surrogates supposedly engage FZDs in a 1:1 stoichiometry ^40,44, which is in accordance with our stepwise photobleaching analysis which suggests a similar average amount of both FZD₅ and LRP6 molecules in each cluster. Therefore, it is currently unclear which molecular properties of a ligand are decisive for whether a FZD-WNT-LRP6 complex passes the checkpoints to initiate WNT/β-catenin signaling, or not. We have previously shown that WNT-16B stabilizes a different conformation than WNT-3A on the intracellular side of FZD₅ ⁴⁵, suggesting that it could form a structurally distinct effector binding site. Yet, WNT-3A and WNT-16B show a rather similar dynamic effect on the orientation of the DVL2 DEP domain relative to FZD₅ ³⁸. There are, however, notable reports of WNT-16B/β-catenin signaling: Mouse WNT-16 was described to have a weak agonistic efficacy in WNT/β-catenin signaling in MC3T3 mouse osteocyte precursor cells ⁴⁶, and WNT-16B promoted chemotherapy resistance via activation of β-catenin signaling in human prostate cancer cells ⁴⁷. Speculatively, cell type-specific expression of a co-factor absent in HEK293/CHO cells might be required to initiate WNT/β-catenin signaling upon WNT-16B stimulation, or WNT-16B/β-catenin signaling might require activation of a specific FZD paralog. The same might apply for WNT-10B, a ligand commonly described to lead to WNT/β-catenin signaling, which was inactive in TOPFlash assays here (see Fig. 3) ⁴⁸.

In conclusion, we propose a two-step model for the efficient activation of WNT/β-catenin signaling, in which WNT stimulation initially leads to quick association and co-confinement of FZDs and LRP6 (signal initiation). In a second step, the signaling outcome of the respective ligand is specified in a checkpoint-like manner (Fig. 6B): If the complex shows hallmarks of WNT/β-catenin signaling such as higher order receptor clustering and LRP6 phosphorylation, it will activate WNT/β-catenin signaling (red WNT, top). Alternatively, WNT stimulation can still trigger transcriptional changes, presumably through other pathways (blue WNT, bottom). The molecular properties required for a FZD-WNT-LRP6 complex to feed into WNT/β-catenin signaling remain unclear and warrant further investigation. This model underscores that ligand-induced association of FZD and LRP5/6 is necessary but not sufficient for WNT/β-catenin signaling but also provides a framework for understanding why diverse WNTs, despite engaging the same receptors, result in distinct signaling outcomes.

Plasmids

Constructs for recombinant protein expression used in this study are listed in Table S1. All plasmids encode human receptor genes. Constructs generated in this study were created by standard seamless cloning techniques following the manufacturer’s protocol (New England Biolabs #E2611) ⁴⁹. This entails all LRP5/6 constructs and FZD-Venus constructs. The template sequences for LRP5 and LRP6 were obtained from Addgene (#115907 and #27282, respectively)^8,50. FZD constructs generally followed the following architecture: 5-HT_3A signal peptide (MRLCIPQVLLALFLSMLTGPGEGSR) – HA-Tag – BamH1 restriction site – FZD coding sequence (no signal peptide) – 10 residue linker – Nluc/mVenus. LRP5/6 constructs were constructed as follows: Influenza hemagglutinin signal peptide (MKTIIALSYIFCLVFA) – FLAG-tag – LRP5/6 coding sequence (no signal peptide) – QPVAT-linker – Nluc/mVenus. FZD₅ and LRP6 constructs used for single-molecule fluorescence microscopy followed the design 5HT3A signal peptide – BamH1 site (GS) – coding sequence, without further tags. Mutations, as described for DVL2, were introduced by site-directed mutagenesis (GeneArt™ Site-Directed Mutagenesis System, Thermo Fisher #A13282). LRP6-5A mutants were created with help of a gBlock (IDT) of the mutant LRP6 C-tail, which was cloned into linearized LRP6-Nluc/Venus lacking the respective part by Gibson assembly.

Cell culture and transient transfection

HEK293 cell lines were cultured in Dulbecco’s Modified Eagle Medium (DMEM; Gibco, Thermo Fisher Scientific, Waltham, MA), containing 10% FCS (Gibco), penicillin (100 U/ml, Gibco), and streptomycin (100 µg/ml, Gibco) at 37° C in a humidified atmosphere containing 5% CO₂. CHO-K1 cells were cultured in DMEM/F12 medium under the same conditions. When reaching approx. 80% confluency, cells were passaged by trypsinization. Cells were routinely confirmed to be free of mycoplasma contamination by a strip test kit (InVivoGen, Toulouse, France). HEK293 cells were obtained from Thermo Fisher Scientific (female origin, #R70507). HEK293 ΔLRP5/6 and HEK293 ΔFZD_1 − 10 cells were a gift from B. Vanhollebeke (Université Libre de Bruxelles, Belgium). HEK293 ΔDVL1-3 cells were a gift from M. Gammons (Cambridge Institute for Medical Research, UK).

For transient transfections, cells were trypsinized and the cell number was adjusted to 350,000/ml with complete medium. Each ml of cell suspension was transfected with 1 µg of cDNA, and all cDNA ratios will be given in % of the total amount of cDNA in the mixture. If the biosensor/protein of interest-encoding cDNA(s) did not add up to 100%, an empty pcDNA3.1 vector was used to adjust the cDNA amount. Transient transfection was performed using PEI MAX® (Polysciences, Warrington, PA) in a three-fold weight excess relative to DNA. The PEI-DNA mixture was incubated in a total volume of 100 µl DPBS per µg DNA for 15–45 min before addition of cells. The transfected cells were then directly added to the respective assay plate and incubated as detailed in the respective sections.

For single-molecule experiments, CHO-K1 cells were seeded onto ultraclean 25 mm round glass coverslips at a density of 3 x 10⁵ cells per well in a 6-well plate. On the next day, they were transfected with plasmids encoding SNAP-FZD₅ or a SNAP-β₂ adrenoceptor and HALO-LRP6 using Lipofectamine 2000, following the manufacturer’s protocol. Cells were labeled and imaged by single-molecule microscopy 3 hours after transfection to obtain low close-to physiological expression levels ^35,52.

Recombinant proteins

Recombinant WNTs and RSPO1 were purchased from Biotechne with the following catalogue numbers: human WNT-3A (5036-WN-010), human/mouse WNT-5A (645-WN-010/CF), recombinant human WNT-10B (7196-WN-010), human WNT-16B (7790-WN-025), RSPO1 (4645-RS-025). To heat-inactivate WNT-16B, it was subjected to 2 cycles of 60° C and − 20° C for 20 min each. Vehicle controls corrected for CHAPS, EDTA and BSA present in protein preparations WNT surrogate was purchased from U-protein express/IPA Therapeutics (WNT Surrogate-Fc Fusion Protein N001-0.5 mg).

Enzyme-linked immunosorbent assay (ELISA)

Transfected cells were seeded into a poly-D-lysine-coated clear 96 well plate at a density of 35,000 cells per well and incubated for 40 h. The medium was aspirated and cells were incubated in primary antibody solution (α-FLAG-M2, Sigma, F1804, or α-HA, abcam, ab9110; both 1:1000) in ELISA Buffer (DPBS supplemented with MgCl₂ and CaCl₂ (Gibco) + 1% bovine serum albumin (BSA)) for 1h at 4° C. The cells were washed five times for 5 min with ice-cold washing buffer (DPBS supplemented with MgCl₂ and CaCl₂ (Gibco) + 0.5% BSA) and then incubated with secondary HRP-conjugated anti-mouse (Thermo Fisher Scientific, 31430) or anti-rabbit (Thermo Fisher Scientific, 31460) secondary antibody for 1 h at 4° C (1:2500 in ELISA buffer), followed by five times washing performed as above. After the last washing step, 50 µl of 3,3′,5,5′-tetramethylbenzidine (Sigma Aldrich, T8665) were added to the cells and the plate was incubated in the dark for 30 min at room temperature. Then, the wells were acidified with 50 µl of 2 M HCl and the absorption was measured at 450 nm at a TECAN Spark multimode plate reader.

Bioluminescence resonance energy transfer assays

For BRET assays, 35,000 transfected cells were seeded into white 96-well plates and incubated for 40-48h before measurement. After washing the cells once with HBSS, 80 µl of HBSS + 0.1% BSA was added to each well. Luciferase substrate (10 µl furimazine (Promega), 1: 100 in assay buffer) was added to each well 6 min before measurement. For kinetic reads, basal Nluc and Venus signal was read 3 times with a 2 min interval before ligand stimulation. After ligand addition, signals were read for another 60 min in 2 min intervals. Data were double baseline-corrected for the baseline and for vehicle control. For acceptor titration experiments, fluorescence was read three times prior to substrate addition to determine the acceptor expression levels (excitation 485/20 nm; emission 535/25 nm), and 6 min after substrate addition, luminescence and acceptor fluorescence were read again thrice to measure the BRET ratio. Data were baseline corrected for background fluorescence and background BRET (at 0% acceptor expression) to obtain net BRET values. All measurements were performed with a Spark multimode plate reader (Tecan) at a temperature of 37° C. Nluc donor emission was detected at wavelengths of 445–485 nm, Venus acceptor emission was detected at 520–560 nm, each with an integration time of 100 ms.

Western Blotting

Samples for Western Blotting were obtained by seeding 300,000 cells in a 96-well plate for 40-48h in complete medium supplemented with 10 nM of the porcupine inhibitor C59. If indicated, cells were transfected as described above. Stimulation of the cells was performed 2 h before lysis by addition of 300 ng/ml WNT or 1 nM WNT surrogate. After the stimulation period, the medium was removed and cells were lysed by addition of 2x Laemmli buffer supplemented with 200 mM dithiothreitol and subsequent sonication. Lysates were heated to 60° C for 20 min before separation on a 7.5% Mini-Protean TGX precast gel (BioRad) with a constant voltage of 120 V. Protein transfer onto polyvinylidene difluoride membranes was performed with the TransBlot Turbo transfer system (BioRad) and the manufacturer’s “Mixed molecular weight” transfer protocol. Membranes were subsequently blocked in blocking buffer (TBS-T (25 mM Tris-HCl, 150 mM NaCl, 0.1% Tween 20, pH 7.6) + 5% dry milk powder) for 1 h and incubated in primary antibody (α-pLRP6, 1:1000, Cell Signaling Tech., Cat. No. 2568; α-DVL2, 1:1000, Cell Signaling Tech., Cat. No. 3216; α-GAPDH, 1:2500, Cell Signaling Tech., Cat. No. 2118; all diluted in blocking buffer). Membranes were washed 5x 5 min with TBS-T before addition of secondary HRP-coupled anti-rabbit antibody (Thermo Fisher Scientific, 31460; 1:5000 diluted in blocking buffer). After 1 h, membranes were washed again for 5x5 min with TBS-T. Blots were developed using Clarity Western ECL substrate (Bio-Rad) following the instructions of the manufacturer in a ChemiDoc chemiluminescence reader (Bio-Rad).

TOPFlash reporter gene assays

HEK293 cells or HEK293 ΔFZD_1 − 10 cells were transfected with 25% of an 8x SuperTOPFlash reporter plasmid (Addgene #12456), 5% of a pRL-TK control plasmid (Promega), 20% of HiBiT-FZD constructs (if indicated), and LRP6 constructs as annotated, and were seeded at a density of 35,000 and 45,000 cells per well, respectively, into a PDL-coated white 96-well flat bottom plate. After 16–20 h of incubation at 37° C/5% CO₂ in complete medium, the medium was removed and replaced with starvation medium (regular DMEM without supplements) + 10 nM of the porcupine inhibitor C59. At the same time, cells were stimulated with 300 ng/ml of the indicated WNT or 1 nM of WNT surrogate. Each stimulation condition was paired with an appropriate vehicle control. Cells were stimulated for 24 h, washed once with HBSS and lysed with 20 µl of 1x passive lysis buffer (Dual Luciferase Assay Kit, Promega, #E1910) for 20 min at room temperature. Then, 20 µl of LARII reagent was added and firefly luminescence was immediately read (550–620 nm, 2 s integration time) using a Spark multimode plate reader (Tecan). Subsequently, 20 µl of Stop-and-Glo reagent spiked with Renilla luciferase substrate were added, and Rluc luminescence was read (445–530 nm, 2 s integration time). The raw TOPFlash ratio was obtained by dividing Fluc counts by Rluc counts, which was afterwards normalized to vehicle control and/or a pcDNA-transfected control to calculate agonist-induced or protein expression-induced TOPFlash ratios.

Bulk mRNA sequencing

RNA was extracted from frozen tissue using the RNeasy Mini Kit (Qiagen). Subsequent mRNA library preparation with polyA-enrichment and NovaSeq X Plus Series Sequencing (paired end reads, ~ 70-100M reads per sample) were performed by Novogene. Reads were aligned to the Hg19 human genome using Rsubread ⁵³. Feature count was performed using featureCounts ⁵⁴, and differential expression and PCA analysis using DESeq2 ⁵⁵. Differential gene expression threshold was set to adjusted p-value < 0.05.

Single-molecule fluorescence microscopy

Live cell protein labeling for single-molecule microscopy was done by labeling with a combination of 2 µM SNAP SiR-647 (New England Biolabs) and 1 µM Halo R110 (Promega) in complete culture medium for 20 min at 37°C. Cells were then washed five times with complete culture medium, allowing 5 min incubation between washes.

Single-molecule microscopy experiments were performed using total internal reflection fluorescence (TIRF) illumination on a custom system (assembled by CAIRN Research) based on an Eclipse Ti2 microscope (Nikon, Japan) equipped with a 100x oil-immersion objective (SR HP APO TIRF NA 1.49, Nikon), 405, 488, 561, and 637 nm diode lasers (Coherent, Obis), an iLas2 TIRF illuminator (Gataca Systems), quadruple band excitation and dichroic filters, a quadruple beam splitter, 1.5x tube lens, four EMCCD cameras (iXon Ultra 897, Andor), hardware focus stabilization, and a temperature-controlled enclosure. The sample and objective were maintained at 37°C throughout the experiments. Coverslips were mounted in a microscopy chamber filled with Hank's balanced salt solution (HBSS) supplemented with 10 mM HEPES, pH 7.5. A reduced oxygen environment (2–4% O₂) was provided in the imaging chamber to decrease photobleaching without increasing cytotoxicity using a mixture of nitrogen and air and a home-built gas mixing and humidifying system, similar as previously described ⁵⁶. Multi-color single-molecule image sequences were acquired simultaneously at a rate of one image every 33 ms.

Automated single-particle detection and tracking were performed with the u-track software ⁵⁷ and the obtained trajectories were further analyzed using custom algorithms in MATLAB environment as previously described ^35,51. Image sequences from different channels were registered against each other, based on reference points taken with multi-color fluorescent beads (100 nm, TetraSpeck) ³⁵. The inter-channel localization precision after coordinate registration was ~ 20 nm. The time-averaged mean squared displacement (TAMSD) of individual trajectories as well as stepwise photobleaching were computed as previously described ^35,52. To obtain the diffusion coefficient (D), only trajectories lasting at least 100 frames were analyzed. For the stepwise photobleaching analysis, a step fitting of the intensity profile was calculated for each particle, setting the maximum step number as 15. Only co-confined were analyzed. A spatial confinement analysis was used to identify trajectory segments characterized by confinement ⁵⁸. The frequency and duration of FZD₅–LRP6 interactions were estimated using previously described methods based on deconvolution of the distribution of single-molecule colocalization times with the one expected for random colocalizations ³⁵. Trajectory segments were first linked to obtain continuous trajectories that are no longer interrupted by merging and splitting events. Then, for each particle in the FZD₅ channel at frame f, all particles in the LRP6 channel falling within a defined search radius (150 nm) was identified as colocalizing. If a colocalization was also present at frame f + 1, the colocalization was extended. The process was iterated until the last frame of the image sequence. These data were used to build a matrix containing information for each colocalization (involved particles as well as the start and end frames). The observed colocalization time corresponds to the duration of true interactions plus the duration of random colocalizations. Thus, the distribution of the observed colocalization times can be seen as a convolution of the distribution of true interaction times and random colocalization times. The distribution for random colocalizations was estimated in cells co-transfected with Halo-LRP6 and SNAP-β₂AR, a membrane receptor with similar diffusion characteristics to that of FZD₅. To obtain the true colocalization time, deconvolution with the Lucy − Richardson algorithm was performed. The separation of co-diffusion versus co-confinement was estimated as previously described, based on whether the interacting partners (FZD₅ and LRP6 molecules) were diffusing on the plasma membrane or were confined during their interaction time ⁵¹.

Acknowledgements

We thank Dr. Melissa Gammons (Cambridge Institute for Medical Research) for providing HEK293 ΔDVL1-3 cells, and Dr. Benoit Vanhollebeke (Université libre de Bruxelles) for providing HEK293 ΔFZD_1-10 and HEK293 ΔLRP5/6 cells. We thank S. Angers (University of Toronto) and V. Bryja (Brno University) for providing plasmids encoding MESD and DVL constructs, respectively.

The work was supported by grants from Karolinska Institutet, the Swedish Research Council (GS: 2019-01190), the Swedish Cancer Society (GS: 20 1102 PjF, 23 2825 Pj), the Novo Nordisk Foundation (GS: NNF22OC0078104), the German Research Foundation (DFG; LG: 504098926; JHV: 520506488), and Cancerfonden (GS: 23 2825 PJ). This project has received funding from the Innovative Medicines Initiative 2 Joint Undertaking (JU) under grant agreement No 875510. The JU receives support from the European Union’s Horizon 2020 research and innovation programme and EFPIA and Ontario Institute for Cancer Research, Royal Institution for the Advancement of Learning McGill University, Kungliga Tekniska Högskolan, Diamond Light Source Limited. This communication reflects the views of the authors and the JU is not liable for any use that may be made of the information contained herein. DC was supported by a Wellcome Trust Senior Research Fellowship (212313/Z/18/Z). JTL acknowledges the Swedish Research Council and the Strategic Research Program in Diabetes for funding.

Author contributions

Conceptualization: J.V., G.S.; methodology: all authors; data curation, formal analysis, and visualization: J.V., Z.K., E.S.; funding acquisition, project administration, and resources: G.S., D.C., J.T.L.; investigation: J.V., Z.K., Y.Y., E.S.; writing – original draft: J.V., Z.K., E.S., G.S.; writing – review & editing: J.V. and G.S. with contributions from all authors.

Materials & Correspondence

Correspondence and material requests should be addressed to G.S.

Competing interests

The authors declare no competing interests.

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WNT-induced association of Frizzled and LRP6 is not sufficient for the initiation of WNT/β-catenin signaling

Status:

Version 1

Abstract

Figures

Introduction

Results

A direct NanoBRET assay to capture dynamic association of FZD₅ and LRP6

LRP6 phosphorylation is not required for WNT-induced FZD-LRP6 association

DVL modulates the conformation of the FZD-LRP6 complex

WNTs can stimulate FZD-LRP6 association without stabilizing β-catenin

WNT-3A and WNT-16B induce different transcriptome changes in HEK293 cells

WNT-3A and WNT-16B induce FZD₅-LRP6 protein complexes with distinct architecture

Discussion

Materials and Methods

Plasmids

Cell culture and transient transfection

Recombinant proteins

Enzyme-linked immunosorbent assay (ELISA)

Bioluminescence resonance energy transfer assays

Western Blotting

TOPFlash reporter gene assays

Bulk mRNA sequencing

Single-molecule fluorescence microscopy

Declarations

References

Additional Declarations

Supplementary Files

Status:

Version 1

WNT-induced association of Frizzled and LRP6 is not sufficient for the initiation of WNT/β-catenin signaling

Status:

Version 1

Abstract

Figures

Introduction

Results

A direct NanoBRET assay to capture dynamic association of FZD5 and LRP6

LRP6 phosphorylation is not required for WNT-induced FZD-LRP6 association

DVL modulates the conformation of the FZD-LRP6 complex

WNTs can stimulate FZD-LRP6 association without stabilizing β-catenin

WNT-3A and WNT-16B induce different transcriptome changes in HEK293 cells

WNT-3A and WNT-16B induce FZD5-LRP6 protein complexes with distinct architecture

Discussion

Materials and Methods

Plasmids

Cell culture and transient transfection

Recombinant proteins

Enzyme-linked immunosorbent assay (ELISA)

Bioluminescence resonance energy transfer assays

Western Blotting

TOPFlash reporter gene assays

Bulk mRNA sequencing

Single-molecule fluorescence microscopy

Declarations

References

Additional Declarations

Supplementary Files

Status:

Version 1

A direct NanoBRET assay to capture dynamic association of FZD₅ and LRP6

WNT-3A and WNT-16B induce FZD₅-LRP6 protein complexes with distinct architecture