We previously reported that disruption of EXTL3, the gene encoding a glycosyltransferase required for the biosynthesis of all HS chains, reduces LGR-independent RSPO3 signaling by ~80% (Lebensohn and Rohatgi, 2018). However, depleting either all SDCs (1-4) or PIGL, an enzyme required for the biogenesis of all GPCs, did not reduce LGR-independent signaling by RSPO3. These observations suggested functional redundancy between GPCs, SDCs and perhaps other HSPGs (Sarrazin et al., 2011) in mediating LGR-independent signaling, making loss-of-function genetic experiments aimed at characterizing this signaling mechanism challenging. Our previous experiments also left open the possibility that reducing the levels of all HSPGs by depleting EXTL3 may change the cell-surface abundance of a different co-receptor for RSPOs that mediates LGR-independent signaling. Therefore, we investigated the mechanism of LGR-independent signaling further by modifying the RSPO ligands themselves.

We previously showed that deleting the complete TSP/BR domain of RSPO3 eliminated its ability to potentiate WNT signaling in LGR4/5/6^KO cells (Lebensohn and Rohatgi, 2018). Deleting the entire TSP/BR domain is a drastic alteration that prevents binding to heparin (Nam et al., 2006), but could also disrupt interactions with other potential binding partners. To design more precise mutations that selectively disrupt the putative HS-binding surface of RSPO3, we created a structural model of the RSPO3 TSP/BR domain based on its homology to 26 of the most similar proteins in the Protein Data Bank (Figure 1A and Figure 1—video 1). Analogous to other TSP domains, the model shows an elongated three-stranded fold stabilized by disulfide bonds at both ends (Ayadi, 2008; Tan et al., 2002). Two positively charged grooves (Site-1 and Site-2 in Figure 1A and Figure 1—video 1) on one face of the model suggested potential sites of interaction with negatively charged HS polymers.

Based on this model, we mutated six (out of sixteen) lysine (K) and four (out of nine) arginine (R) residues in these grooves to charge-reversing glutamic acid (E) residues, thereby altering the electrostatic surface of the TSP/BR domain so that it should not be able to interact properly with HS chains. We refer to this mutant protein as RSPO3 TSP/BR (K/R→E) (Figure 1A and B, Figure 1—video 1, and Supplementary file 1). We produced a tagged version of RSPO3 TSP/BR (K/R→E) and purified it under physiological conditions using a one-step affinity purification strategy described in our previous eLife paper (see Figure 1—figure supplement 1A from Lebensohn and Rohatgi, 2018). We have demonstrated that tagged RSPO proteins produced and purified in this manner preserve the signaling properties of their untagged counterparts (see Figure 1—figure supplement 1C and D from Lebensohn and Rohatgi, 2018). A coomassie blue-stained gel of purified wild type (WT) RSPO3, RSPO3 TSP/BR (K/R→E) and RSPO3 ΔTSP/BR, a control construct lacking the TSP/BR domain entirely (Figure 1B), is shown in Figure 1C.

We tested the capacity of these recombinant RSPO3 proteins to potentiate signaling induced by a low dose of WNT3A conditioned medium (CM) in WT HAP1-7TGP or in LGR4/5/6^KO cells. HAP1-7TGP is a human haploid cell line harboring a fluorescent transcriptional reporter for WNT/β-catenin signaling that we have previously established as a valid and genetically tractable system to study both WNT- and RSPO-dependent signaling (Lebensohn et al., 2016). LGR4/5/6^KO cells are a clonal derivative of HAP1-7TGP in which LGRs 4, 5 and 6 were disrupted by CRISPR/Cas9-mediated genome editing (Lebensohn and Rohatgi, 2018). In LGR4/5/6^KO cells, the signaling response to low levels of WNT3A can be potentiated robustly by RSPO2 and RSPO3, but not by RSPO1 or RSPO4 (see Figure 1B and C from Lebensohn and Rohatgi, 2018), making them a convenient system to study LGR-independent RSPO signaling.

As we had observed previously (see Figure 2F and G from Lebensohn and Rohatgi, 2018), deleting the TSP/BR domain of RSPO3 reduced its potency by 82-fold in WT HAP1-7TGP cells (Figure 1D), and completely abolished signaling in LGR4/5/6^KO cells (Figure 1E). Similarly, the K/R→E mutations we introduced into the TSP/BR domain of RSPO3 reduced its potency by 13-fold in WT HAP1-7TGP cells (Figure 1D) and almost completely abolished its ability to potentiate WNT signaling in LGR4/5/6^KO cells (Figure 1E). Our data supports the model that interactions between basic K and R residues in the TSP/BR domain of RSPO3 and the HS chains of HSPGs contribute substantially to potentiating WNT signaling in the presence of LGRs. In the absence of LGRs, the RSPO3-HSPG interaction becomes essential to potentiate WNT signaling.

While the positively charged grooves we mutated in the TSP/BR domain (Figure 1A and Figure 1—video 1) are known to interact with HS chains in other TSP domain-containing proteins (Sarrazin et al., 2011; Tan et al., 2002), it remained possible that the K/R→E mutations disrupted the interaction of RSPO3 with a different, unidentified receptor. To resolve this issue, we sought to provide the HSPG-binding functionality of the TSP/BR domain through an entirely different protein. We engineered a synthetic RSPO3 protein in which we replaced the entire TSP/BR domain with HS20, an scFv antibody selected by phage display to bind to the HS chains of GPC3 (Gao et al., 2016; Gao et al., 2014; Geoghegan et al., 2017; Figure 2A and Supplementary file 1). We refer to this protein as RSPO3 ΔTSP/BR HS20. Due to the presence of the HS20 scFv in the RSPO3 ΔTSP/BR HS20 fusion protein, we excluded the Fc tag that we had used in other RSPO3 constructs to promote their solubility. We retained an N-terminal HA tag and a small C-terminal 1D4 tag, the latter of which was used for affinity purification of the fusion protein (Figure 2A and B).

Replacing the TSP/BR domain with HS20 in RSPO3 ΔTSP/BR HS20 did not disrupt the ability of RSPO3 to signal both in the presence and absence of LGRs (Figure 2C and D). Importantly, mutations in the antigen-binding third complementarity-determining region (CDR3) of HS20 (RSPO3 ΔTSP/BR HS20 (GS) and RSPO3 ΔTSP/BR HS20 (A) in Figure 2A and B) entirely eliminated the WNT-potentiating activity of RSPO3 ΔTSP/BR HS20, both in the presence and absence of LGRs (Figure 2E and F). The fact that HS20, a structurally unrelated protein specifically selected for HS-binding activity (Gao et al., 2014), can functionally substitute for the TSP/BR domain of RSPO3 provides strong support for our model that interactions with HSPGs are required to potentiate WNT signaling in cells lacking LGRs, and also contribute significantly in cells containing LGRs.

What are the relative contributions of ZNRF3/RNF43, LGRs and HSPGs to signaling by RSPO3 ΔTSP/BR HS20? In order to measure the contribution of FU1 domain interactions with ZNRF3/RNF43 and FU2 domain interactions with LGRs, we individually mutated the FU1 and FU2 domains of RSPO3 ΔTSP/BR HS20. Point mutations (R67A/Q72A) in the FU1 domain of RSPO3 ΔTSP/BR HS20 (RSPO3 ΔTSP/BR HS20 (R67A/Q72A) in Figure 2A and B) known to weaken the interaction with ZNRF3/RNF43 (Xie et al., 2013), reduced signaling potency by 10-fold in WT HAP1-7TGP cells (Figure 2G) and abolished signaling in LGR4/5/6^KO cells (Figure 2H). Point mutations (F106E/F110E) in the FU2 domain of RSPO3 ΔTSP/BR HS20 (RSPO3 ΔTSP/BR HS20 (F106E/F110E) in Figure 2A and B) that weaken interactions with LGRs (Xie et al., 2013) reduced signaling efficacy by 80% and potency by 3-fold in WT HAP1-7TGP cells (Figure 2G). As expected, disrupting the LGR-binding FU2 domain in RSPO3 ΔTSP/BR HS20 (F106E/F110E) did not reduce signaling in LGR4/5/6^KO cells (Figure 2H).

We conclude that interactions with ZNRF3/RNF43, LGRs and HSPGs all contribute to the potentiation of WNT signaling by RSPO3 ΔTSP/BR HS20 in the presence of LGRs. Interactions with ZNRF3/RNF43 and HSPGs are required for signaling in the absence of LGRs. The simplest model to explain these results is that RSPO3 can bind simultaneously to LGR and HSPG co-receptors, which together present the ligand to ZNRF3/RNF43, the key effectors that mediate endocytosis of the complex. Without LGRs, RSPO3 is presented by HSPGs alone, explaining the significantly lower potency of both WT RSPO3 and RSPO3 ΔTSP/BR HS20 in LGR4/5/6^KO compared to WT HAP1-7TGP cells (Figures 1D, E, 2C and D). Binding assays and structural studies with purified proteins will be required to test for the formation of a ternary RSPO3-LGR-HSPG complex.

We tested whether the mechanism whereby RSPO3 ΔTSP/BR HS20 potentiates WNT responses is equivalent to that of WT RSPO3. LGR-dependent RSPO signaling triggers internalization and lysosomal degradation of the E3 ubiquitin ligases ZNRF3 and RNF43, promoting the stabilization of WNT receptors (Hao et al., 2012; Koo et al., 2012). Whether LGR-independent signaling by RSPO3 proceeds through a similar mechanism is not known. We used a surface biotinylation assay (Koo et al., 2012) to measure RSPO3-mediated internalization and degradation of RNF43-Flag in WT HAP1-7TGP and in LGR4/5/6^KO cells. Treatment of either WT HAP1-7TGP or LGR4/5/6^KO cells with WT RSPO3 led to the depletion of RNF43-Flag from the cell surface and a reduction in its total cellular level (Figure 3A and B), suggesting similar signaling mechanisms for LGR-dependent and LGR-independent potentiation of WNT signaling. These results are consistent with our previous mutational analysis demonstrating that the ZNRF3/RNF43-interacting FU1 domain is essential for RSPO3 signaling in cells lacking LGRs (Lebensohn and Rohatgi, 2018).

Having established the mechanism of LGR-independent signaling by RSPO3, we tested whether RSPO3 ΔTSP/BR HS20 potentiates WNT responses through an equivalent mechanism. Deletion of the TSP/BR domain of RSPO3 (RSPO3 ΔTSP/BR in Figure 3A and B) prevented the internalization and degradation of RNF43-Flag in both WT HAP1-7TGP and LGR4/5/6^KO cells, consistent with the reduced capacity of RSPO3 ΔTSP/BR to promote LGR-dependent (Figure 1D) and LGR-independent (Figure 1E) signaling. In contrast, as with WT RSPO3, addition of RSPO3 ΔTSP/BR HS20 to either WT HAP1-7TGP (Figure 3A) or LGR4/5/6^KO (Figure 3B) cells resulted in the depletion of RNF43-Flag from the cell surface and a reduction in its total cellular level. The effects of RSPO3 ΔTSP/BR HS20 were dependent on the interaction between HS20 and HS chains, since the RSPO3 ΔTSP/BR HS20 (GS) mutant, which carries mutations in the HS-binding CDR3 loop of HS20, failed to promote internalization and degradation of RNF43-Flag (Figure 3A and B).

In summary, replacing the HS-binding TSP/BR domain of RSPO3 with HS20, an unrelated protein that binds HS chains, can rescue signaling in the absence of LGRs and substantially enhance signaling in their presence by promoting the internalization and degradation of RNF43. We propose that HSPGs represent the third major class of RSPO co-receptors, in addition to LGRs and ZNRF3/RNF43.

Are specific HSPGs required for RSPO signaling? Cell-surface HSPGs comprise six glypicans (GPC1-6), which are anchored to the plasma membrane by a glycophosphatidylinositol (GPI) linkage, and four syndecans (SDC1-4), which are single pass transmembrane proteins (Sarrazin et al., 2011). In our previous eLife paper (Lebensohn and Rohatgi, 2018), we presented evidence that RSPO3 can use the HS chains on multiple HSPGs to potentiate WNT signaling. However, HS20 is an scFv selected to bind to GPC3 (Gao et al., 2014), raising the possibility that the synthetic RSPO3 ΔTSP/BR HS20 ligand may have a narrower HSPG selectivity. In a binding assay with purified proteins, HS20 bound equally well to GPC3 and GPC4 (Figure 4A), the two GPCs expressed most highly in HAP1 cells (see Table 1 in Lebensohn and Rohatgi, 2018: average RPKM values for GPC3 and GPC4 were 157 and 220, respectively). HS20 failed to bind to mutant GPC3 and GPC4 proteins that cannot be modified by HS chains (GPC3ΔHS and GPC4ΔHS in Figure 4A). These results support a prior conclusion that HS20 recognizes the HS side-chain structure shared by multiple HSPGs, rather than binding to the protein core of a specific GPC (Gao et al., 2014).

To test the HSPG specificity of RSPO3 ΔTSP/BR HS20 more thoroughly in cells, we used CRISPR/Cas9 methods to generate a series of clonal cell lines (see Materials and methods and Supplementary files 2 and 3) with increasingly severe deficits in their HSPG composition: (1) GPC3^KO, (2) GPC4^KO, (3) PIGL^KO (lacking an enzyme required for GPI anchor biosynthesis and hence for the cell-surface expression of all GPCs), (4) SDC1/2/3/4^KO (lacking all four SDCs) and (5) EXTL3^KO (lacking a glycosyltransferase required for the synthesis of HS chains attached to all HSPGs, including all GPCs and SDCs, but not for the synthesis of other proteoglycans or glycosaminoglycans). Each of these mutations was introduced into LGR4/5/6^KO cells, in which the HS-binding activity of HS20 is required for signaling (Figure 2D), and hence the effect of eliminating different HSPGs can be measured most accurately. We also tested the specificity of WT RSPO3, containing the native HSPG-binding TSP/BR domain, in these cell lines.

Disrupting GPC3 in LGR4/5/6^KO cells had no significant effect on signaling induced by either WT RSPO3 or RSPO3 ΔTSP/BR HS20 (Figure 4B and C), consistent with the finding that HS20 can bind with equivalent affinity to the HS chains of other GPCs such as GPC4 (Figure 4A). Disrupting GPC4 or PIGL modestly reduced the potency (i.e. increased the EC₅₀) of both WT RSPO3 and RSPO3 ΔTSP/BR HS20 (Figure 4B and C). Combined disruption of all SDCs – SDC1, SDC2, SDC3 and SDC4 – surprisingly increased the potency (ie. lowered the EC₅₀) of both WT RSPO3 and RSPO3 ΔTSP/BR HS20 (Figure 4B and C). This increase in potency may be related to the previous observation that some SDCs inhibit WNT/β-catenin signaling through regulation of LRP6 and RSPO3 (Astudillo et al., 2014). Finally, disrupting EXTL3, predicted to globally block HS chain biosynthesis, impaired signaling by both proteins to the greatest extent (Figure 4B and C).

Overall, disrupting different HSPGs led to a qualitatively similar effect on signaling by WT RSPO3 and RSPO3 ΔTSP/BR HS20, with GPC3 disruption having no effect, GPC4 or PIGL disruption causing a modest reduction in signaling potency and EXTL3 disruption causing the greatest reduction in signaling potency. These results are most consistent with the model that RSPO3 ΔTSP/BR HS20, like WT RSPO3, can signal in a redundant manner via either GPCs, SDCs or another HSPG by engaging their HS chains rather than their protein cores.

Given the substantial contribution made by the interaction between the TSP/BR domain or HS20 and HSPGs to LGR-dependent signaling in haploid cells (Figures 1D and 2C), it was important to test whether this interaction also mediates RSPO signaling in a more physiological context. The growth of mouse small intestinal organoids is strictly dependent on exogenously supplied RSPOs (Sato et al., 2009) and on the expression of LGRs (de Lau et al., 2011). In addition, studies in mice have shown that depletion of HS chains in the intestinal epithelium compromise WNT-dependent crypt renewal and regeneration following radiation-induced injury (Yamamoto et al., 2013). We used this organoid culture system in combination with our mutant and engineered RSPO3 ligands to test the importance of RSPO-HSPG interactions for supporting the crypt-villus architecture of the intestine. WT RSPO3 promoted the growth of intestinal organoids with multiple budded structures at concentrations down to 0.5 nM (Figure 5A and B). The buds represent crypt domains, since they are enriched in markers of proliferating cells (Ki67) and of Paneth cells (lysozyme) (Figure 5C). RSPO3 ΔTSP/BR was markedly less potent in this assay, enabling only a few organoids to grow at a concentration of 10 nM (Figure 5B). While the FU1 and FU2 domains present in RSPO3 ΔTSP/BR were sufficient to promote organoid growth at the highest concentration tested, our titration experiment revealed that abolishing HSPG interactions by deleting the TSP/BR domain sharply reduced ligand potency.

To determine if the positive effect of the TSP/BR domain in promoting organoid growth is due to its interactions with HSPGs, we tested the organoid-supporting properties of RSPO3 ΔTSP/BR HS20. Not only was RSPO3 ΔTSP/BR HS20 able to fully support organoid growth, with budding of normal crypt domains, but it was more potent than WT RSPO3, supporting some organoid growth even at concentrations as low as 0.1 nM (Figure 5A and B). Organoids grown in the presence of RSPO3 ΔTSP/BR HS20 were indistinguishable from those grown in the presence of WT RSPO3, with crypt domains containing proliferating cells and Paneth cells (Figure 5C). The capacity of RSPO3 ΔTSP/BR HS20 to promote organoid growth required the interaction between HS20 and HS chains, since RSPO3 ΔTSP/BR HS20 (GS), carrying mutations in the HS-binding CDR3 loop (Figure 2A) failed to promote organoid growth at concentrations below 10 nM, similarly to RSPO3 ΔTSP/BR (Figure 5A and B).

We conclude from these experiments that the interaction between the TSP/BR domain, or HS20, and HSPGs contributes substantially to the capacity of RSPO3 to support the growth of mouse intestinal organoids. While additional work will be required to establish which specific HSPGs mediate RSPO reception in the intestine, we note that Gpc3 mRNA expression in mouse intestinal organoids is undetectable (Lindeboom et al., 2018), suggesting that, as in HAP1 cells (Figure 4B), RSPOs likely signal through other HSPGs. Multi-omics data shows that GPC4, SDC1 and SDC4 are expressed at detectable levels in mouse intestinal organoids (Lindeboom et al., 2018), making them candidate receptors for RSPO3 in this tissue.

While our results demonstrate that the interaction between RSPO3 and the HS chains of HSPGs is necessary to promote LGR-independent signaling in cells, and can significantly enhance LGR-dependent signaling in cells and intestinal organoids, they do not preclude the existence of another co-receptor capable of mediating these effects. We previously reported the results of a genome-wide, haploid genetic screen for regulators of RSPO1-potentiated, low-level WNT signaling in WT HAP1-7TGP cells (Lebensohn et al., 2016). That screen was focused on LGR-dependent signaling, since RSPO1 (unlike RSPO3) requires LGRs to potentiate WNT signaling (Lebensohn and Rohatgi, 2018). We attempted to identify any additional co-receptors required for LGR-independent signaling through a similar haploid genetic screen for regulators of RSPO3-potentiated, low-level WNT signaling in cells lacking LGRs 4, 5 and 6 (see Materials and methods). To distinguish genes required specifically for LGR-independent signaling from those required for LGR-dependent signaling, we compared the results of our new screen to those of our previous screen for RSPO1-potentiated signaling in WT HAP1-7TGP cells (Lebensohn et al., 2016).

The RSPO3 screen in LGR4/5/6^KO cells revealed a subset of the WNT signaling regulators previously identified in the RSPO1 screen in WT HAP1-7TGP cells (Figure 6A–C and Supplementary files 4 and 5), including the WNT co-receptor LRP6, members of the TCF/LEF transcriptional complex such as CTNNB1, CREBBP, BCL9 and DOT1L, the ubiquitin ligase RNF146, which ubiquitinates poly-ADP-ribosylated AXIN1 for proteasomal degradation, and a truncated form of AXIN2 that represses WNT signaling through an unknown mechanism (Lebensohn et al., 2016). LGR4, the top hit of the RSPO1 screen in WT HAP1-7TGP cells, was not a significant hit in the RSPO3 screen in LGR4/5/6^KO cells, in which LGR4 had been disrupted by CRISPR/Cas9-mediated genome editing (Figure 6A–C and Supplementary files 4 and 5).

The RSPO3 screen in LGR4/5/6^KO cells failed to reveal any unique significant hits compared to the RSPO1 screen in WT HAP1-7TGP cells (Figure 6A–C and Supplementary files 4 and 5), suggesting there were no genes selectively required for LGR-independent potentiation of WNT signaling by RSPO3. While several components of the GPI-anchor biosynthetic machinery, including PIGP, PIGS, PIGU, PIGL and DPM3, as well as GPC4 were hits in both screens, we have previously shown that their presence as significant hits in the RSPO1 screen in WT HAP1-7TGP cells was due to the requirement of GPI-anchored GPCs for low-level WNT signaling, independently of RSPOs (Lebensohn et al., 2016). In summary, haploid genetic screens did not reveal an additional co-receptor required for LGR-independent potentiation of WNT signaling by RSPO3.

Provided as Supplementary file 6.

Human haploid cells (HAP1) were validated using prodium iodide staining to ensure they have a haploid DNA content. All derivatives of HAP1 cells made using CRISPR/Cas9 methods were verified by DNA sequencing. HEK 293T cells were obtained from ATCC with a certificate of authentication and used at low passage (<10) without any additional STR profiling. Cell lines were confirmed to be mycoplasma-negative when cultures were started in the lab.

pHLsec-HA-hRSPO3-Tev-Fc-Avi-1D4 and pHLsec-HA-hRSPO3ΔTSP/BR-Tev-Fc-Avi-1D4 (encoding proteins containing an N-terminal HA tag and C-terminal Fc and 1D4 tags) were constructed as described previously (Lebensohn and Rohatgi, 2018).

pHLsec-HA-hRSPO3TSP/BR(K/R→E)-Tev-Fc-Avi-1D4 (Figure 1B and Supplementary file 1) was synthesized as a gBlock Gene Fragment (Integrated DNA Technologies (IDT)) flanked at the 5’ and 3’ ends, respectively, by 24 base pair (bp) overhangs overlapping the sequence upstream of the unique AgeI site and downstream of the unique KpnI site in the pHLsec-HA-Tev-Fc-Avi-1D4 vector. The gBlock was subcloned into pHLsec-HA-Tev-Fc-Avi-1D4, previously digested with AgeI and KpnI, using the NEBuilder HiFi DNA Assembly Master Mix (New England Biolabs (NEB) Cat. # E2621L).

pHLsec-HA-hRSPO3ΔTSP/BRHS20-Avi-1D4, pHLsec-HA-hRSPO3ΔTSP/BRHS20(GS)-Avi-1D4, and pHLsec-HA-hRSPO3ΔTSP/BRHS20(A)-Avi-1D4, all of which encode proteins containing an N-terminal HA tag and a C-terminal 1D4 tag but lacking the dimerizing Fc tag (Figure 2A and Supplementary file 1), were synthesized as gBlock Gene Fragments (IDT) flanked at the 5’ and 3’ ends, respectively, by 24 bp overhangs overlapping the sequence upstream of the unique AgeI site and downstream of the unique KpnI site in the pHLsec-HA-Avi-1D4 vector described previously (Lebensohn and Rohatgi, 2018). The gBlock was subcloned into pHLsec-HA-Avi-1D4, previously digested with AgeI and KpnI, using the NEBuilder HiFi DNA Assembly Master Mix (NEB Cat. # E2621L).

pHLsec-HA-hRSPO3ΔTSP/BRHS20(R67A/Q72A)-Avi-1D4 and pHLsec-HA-hRSPO3ΔTSP/BRHS20(F106E/F110E)-Avi-1D4 were made by replacing the FU1 and FU2 domains in pHLsec-HA-hRSPO3ΔTSP/BRHS20-Avi-1D4 with those from pHLsec-HA-hRSPO3(R67A/Q72A)-Tev-Fc-Avi-1D4 and pHLsec-HA-hRSPO3(F106E/F110E)-Tev-Fc-Avi-1D4 (Lebensohn and Rohatgi, 2018), respectively. A fragment containing the FU1 and FU2 domains was subcloned by double digestion with AgeI and MfeI followed by ligation.

RNF43-2xFlag-2xHA and N-terminal SNAP-tagged mouse FZD5 (Ala27-Val585) were described previously (Koo et al., 2012).

pVRC8400-GPC3ΔHS-hFc (pMH137) and pVRC8400-GPC4ΔHS-hFc (pMH373) were constructed as described previously (Feng et al., 2013). Briefly, serine to alanine substitutions were introduced at the HS attachment sites of GPC3 (S495A and S509A) or GPC4 (S494A, S498A and S500A) and the mutant constructs were PCR-amplified and inserted into the pVRC8400 expression vector (a gift from Dr. Gary J. Nabel, National Institute of Allergy and Infectious Diseases, Bethesda, MD). Sequences encoding the IL-2 signal peptide and a human Fc tag were added to the N-terminus and C-terminus of the mutant GPCs, respectively.

pX458-mCherry was made by replacing the coding sequence of GFP in pSpCas9(BB)−2A-GFP (pX458, a gift from Dr. Feng Zhang, Addgene plasmid # 48138) with that of a modified mCherry sequence. The WT mCherry sequence has a BbsI cleavage site that makes it incompatible with the BbsI restriction digestion required for cloning the single guide RNA (sgRNA) oligos into pX458. Therefore, a silent mutation was introduced into the mCherry sequence to eliminate the BbsI cleavage site. Terminal EcoRI sites and a T2A sequence were appended to the modified mCherry sequence to obtain an EcoRI-T2A-mCherry-EcoRI fragment. The EcoRI-T2A-EGFP-EcoRI fragment in px458 was replaced with the EcoRI-T2A-mCherry-EcoRI fragment by EcoRI double digestion and ligation to obtain pX458-mCherry.

Primary antibodies: Rho 1D4 purified monoclonal antibody (University of British Columbia, https://uilo.ubc.ca/rho-1d4-antibody), mouse anti-Flag M2 (MilliporeSigma Cat. # F3165), mouse anti-actin monoclonal (Clone C4, MP Biomedicals Cat. # 08691002), polyclonal rabbit anti-human lysozyme (Agilent Dako Cat. # A0099), rabbit anti-Ki67 (Abcam Cat. # ab15580).

Secondary antibodies: chicken anti-rabbit Alexa Fluor 488 (Thermo Fisher Scientific Cat. # A21441), goat anti-mouse Alexa Fluor 680 (Thermo Fisher Scientific Cat. # A21057), goat anti-mouse IRDye800 (Rockland Immunochemicals Cat. # 610-132-121), goat anti-human Ig kappa chain HRP conjugate (MilliporeSigma Cat. # AP502P).

Staining reagents: phalloidin-TRITC (MilliporeSigma Cat. # P1951), DAPI (MilliporeSigma Cat. # D9542).

A structural model of human RSPO3 (residues valine 146 - isoleucine 232; UniProtKB Q9BXY4) was built using the HHpred web server (Max Planck Institute Bioinformatics Toolkit) based on homology to the 26 most similar protein structures in the Protein Data Bank (Zimmermann et al., 2018). Polymers consisting of four disaccharide units of heparin (PDB ID 1HPN [Mulloy et al., 1993]) were docked to RSPO3 using AutoDock Vina (Trott and Olson, 2010). Heparin was kept rigid during docking. Positively charged side chains of residues 161, 162, 164, 169, 170, 198, 203–205, 208, 211 (Site-1) or 213, 214, 216, 218–222, 225, 229 (Site-2) were kept flexible during docking. The models depicted in Figure 1A and the animation shown as Figure 1—video 1 were rendered using PyMOL 2.3.2.

Tagged RSPO proteins were produced by transient transfection of 293T cells and immuno-affinity purification as described previously (Lebensohn and Rohatgi, 2018), with some modifications. The WT, mutant and truncated RSPO3 proteins have an N-terminal HA tag and C-terminal Fc and 1D4 tags; the engineered RSPO3 ΔTSP/BR HS20 fusion proteins, and mutant derivatives thereof, have an N-terminal HA tag and a C-terminal 1D4 tag (see Supplementary file 1 for nucleotide sequences). ~24 hr before transfection, 20 × 10⁶ 293T cells were seeded in a T-225 flask containing 55 ml of growth medium (Dulbecco’s Modified Eagle’s Medium (DMEM)/high glucose without L-glutamine, sodium pyruvate (HyClone, GE Healthcare Life Sciences Cat. # SH30081.FS); 10% fetal bovine serum (FBS, MilliporeSigma Cat. # S11150); 1 mM sodium pyruvate (Gibco, Thermo Fisher Scientific Cat. # 11360070); 1x MEM non-essential amino acids (Gibco, Thermo Fisher Scientific Cat. # 11140050); 2 mM L-glutamine (Gemini Bio Products Cat. # 400–106)). Once they had reached ~80% confluency, the cells in each flask were transfected with 1 ml of a transfection mix prepared as follows. 30 µg of DNA constructs encoding tagged WT, mutant or synthetic RSPO proteins were diluted in 910 µl of serum-free DMEM and 90 µl of polyethylenimine (PEI, linear, MW ~25,000, Polysciences, Inc Cat. # 23966) was added from a 1 mg/ml stock (prepared in sterile water, stored frozen and equilibrated to 37°C before use). The transfection mix was vortexed briefly, incubated for 15–20 min at room temperature (RT) and added to the cells without replacing the growth medium. ~24 hr post-transfection, the cells were washed with 55 ml PBS and the medium was replaced with 55 ml of CD 293 medium (Thermo Fisher Scientific Cat. # 11913019) supplemented with 1x L-glutamine solution and 2 mM valproic acid (MilliporeSigma Cat. # P4543, added from a 0.5 M stock prepared in water and sterilized by filtration through a 0.22 µm filter) to promote protein expression. ~96 hr post-transfection, the CM from each flask, containing secreted RSPO protein, was filtered through 0.45 µm filters (Nalgene aPES membrane, Thermo Fisher Scientific Cat. # 166–0045) to remove particulates and was reserved on ice. 300 µl of a ~ 50% slurry of Rho 1D4 resin (prepared as described in Lebensohn and Rohatgi, 2018) was added to a 50 ml conical tube containing the RSPO CM and the suspension was incubated overnight, rocking at 4°C. Following binding and during all subsequent washes, the resin was collected by centrifugation for 5 min at 400 x g in a swinging bucket rotor. The beads were washed three times at RT with 25 ml PBS by resuspending them in buffer and mixing by inversion for ~1 min. The resin was transferred to a 1.5 ml Eppendorf tube and washed three more times with 1 ml of PBS, 10% glycerol. Following the last wash, the buffer was aspirated and the resin was resuspended in 150 µl of 500 µM 1D4 peptide ((NH3)-T-E-T-S-Q-V-A-P-A-(COOH)) prepared in PBS, 10% glycerol, to obtain a ~ 50% slurry. Elution was carried out by rotating the tube horizontally overnight at 4°C. Following centrifugation of the resin, the eluate was recovered and reserved on ice. The resin was resuspended in another 150 µl of 500 µM 1D4 peptide in PBS, 10% glycerol, and a second round of elution was carried out for 2 hr at RT. Following centrifugation of the resin, the second eluate was recovered and pooled with the first. The final eluate was centrifuged again to remove residual resin and the supernatant containing tagged RSPO proteins was aliquoted, frozen in liquid nitrogen and stored at −80°C. Tagged RSPO proteins were quantified as described previously (Lebensohn and Rohatgi, 2018).

20,000 cells were plated per well of a 96-well plate and grown in complete IMDM growth medium (Iscove's Modified Dulbecco's Medium (IMDM) modified with L-glutamine, HEPES (GE Healthcare Life Sciences Cat. # SH30228.01); 2 mM L-glutamine; 40 units/ml penicillin:40 µg/ml Streptomycin (Gemini Bio-products Cat. # 400–109); 10% FBS). ~24 hr after plating, the cells were treated with complete IMDM growth medium containing 1.43% WNT3A CM and the indicated concentrations of RSPO proteins. After 24 hr of treatment, cells were trypsinized in 40 μl of Tryspin-EDTA (0.05%, Thermo Fisher Scientific Cat. # 25300054) and harvested in 160 μl of complete IMDM growth medium. WNT reporter (EGFP) fluorescence data for 2,500 singlet-gated cells was measured by flow cytometry on a BD Accuri RUO Special Order System (BD Biosciences). The median EGFP fluorescence from each well was depicted as circles in Figure 1D and E and Figure 2C–H. Dose-response curves were fitted using the nonlinear regression (curve fit) analysis tool in GraphPad Prism 8 using the [agonist] vs. response – variable slope (four parameters) equation. Results presented are representative of experiments repeated at least twice.

WT HAP1-7TGP or LGR4/5/6^KO cells were transfected with human RNF43-2xFlag-2xHA and SNAP-tagged mouse FZD5 using PEI (Koo et al., 2012). 24 hr after transfection, cells were washed with ice-cold PBS complete (containing Ca²⁺ and Mg²⁺) and kept on ice. 800 μM of EZ-Link Sulfo-NHS-SS-Biotin (Thermo Fisher Scientific Cat. # 21331) in PBS complete was added to each well and biotinylation was allowed to proceed for 30 min on ice. The medium was aspirated, cells were washed once with ice-cold PBS complete and unreacted biotin was quenched with complete IMDM growth medium containing 50 mM glycine for 5 min on ice. Cells were washed with ice-cold PBS complete and lysed for 30 min on ice in lysis buffer (20 mM Tris pH 7.5, 150 mM NaCl, 1% Triton X-100, 1 mM EDTA, 1 mM PMSF, 10 μg/ml Leupeptin, 10 μg/ml Aprotinin). Cell lysates were centrifuged for 5 min at 14,000 x g at 4°C. An aliquot of the supernatant (‘Total’ in Figure 3A and B) was mixed with SDS sample buffer and the remaining supernatant was incubated with 25 μl of a 50% slurry of Streptavidin agarose beads (Pierce, Thermo Fisher Scientific Cat. # 20349) for 60 min at 4°C. The beads were washed with 0.1x PBS complete, and biotinylated proteins (‘Surface’ in Figure 3A and B) were eluted in 20 μl 2x SDS sample buffer, 40 mM DTT for 30 min at 37°C and analyzed by SDS-PAGE and Western blot. Western blotting was performed using standard procedures. Samples were resolved by SDS-PAGE, transferred to Immobilon-FL PVDF membranes (MiliporeSigma Cat. # IPFL00005), blocked with Odyssey blocking buffer (Li-Cor Cat. # 927–40000), incubated with the indicated primary and secondary antibodies, and imaged using the Amersham Typhoon NIR laser scanner (GE Healthcare).

The GPC3 and GPC4 mutant proteins without HS chains (GPC3ΔHS and GPC4ΔHS in Figure 4A) were constructed by replacing serine with alanine residues at the HS attachment sites. pVRC8400-GPC3ΔHS-hFc or pVRC8400-GPC4ΔHS-hFc were transiently transfected into HEK-293T cells. The cell supernatant was harvested and the Fc-tagged GPC proteins were purified on a Protein A Hi-Trap column (GE Healthcare Cat. # 29048576) according to the manufacturer’s instructions. The quality and quantity of purified proteins were determined by SDS-PAGE and by measuring absorbance at 280 nm on a NanoDrop instrument (Thermo Fisher Scientific), respectively.

Increasing concentrations of HS20 (serial 2-fold dilutions starting from 1 μg/ml) were added into a 96-well ELISA plate coated with 5 μg/ml of glycosylated GPC3, GPC4 (R and D Systems Cat. # 2119-GP and 9195-GP, respectively), GPC3ΔHS, GPC4ΔHS, or an irrelevant human Fc-fusion protein (CD276-hFc, labeled ‘Control’ in Figure 4A) and incubated for 1 hr at RT. The plate was washed three times with PBST (PBS, 0.1% Tween-20) and incubated with 50 μl of goat anti-human kappa chain HRP conjugate (1:5,000 dilution) for 1 hr at RT. After washing six times with PBST, 50 μl/well of 3,3′,5,5′-tetramethylbenzidine detection reagent (Kirkegaard and Perry Laboratories Cat. # 95059–156) was added and incubated for 10 min at RT. Absorbance was read at 450 nm.

The LGR4/5/6^KO, LGR4/5/6^KO; PIGL^KO, LGR4/5/6^KO; SDC1/2/3/4^KO and LGR4/5/6^KO; EXTL3^KO cell lines used in this study have been described previously (Lebensohn and Rohatgi, 2018). The LGR4/5/6^KO; GPC3^KO and LGR4/5/6^KO; GPC4^KO cell lines were constructed as described previously (Lebensohn et al., 2016). Briefly, oligonucleotides encoding sgRNAs (Supplementary file 2) were selected from a published library (Doench et al., 2016) and cloned into pX458-mCherry according to a published protocol (Cong et al., 2013) (original version of ‘Target Sequence Cloning Protocol’ from http://www.genome-engineering.org/crispr/wp-content/uploads/2014/05/CRISPR-Reagent-Description-Rev20140509.pdf). Clonal cell lines were established as described previously (Lebensohn et al., 2016) by transient transfection of LGR4/5/6^KO cells with pX458-mCherry containing the desired sgRNAs (since we used pX458-mCherry, there was no need to co-transfect with pmCherry), followed by single-cell sorting of mCherry-positive cells. Genotyping was done as described previously (Lebensohn et al., 2016) using the primers indicated in Supplementary file 2, and the sequencing results are summarized in Supplementary file 3.

Small intestinal organoids were established and maintained as described (Sato et al., 2009), starting from isolated crypts collected from the entire length of the small intestine of B6 mice. The basic culture medium (advanced DMEM/F12, penicillin/streptomycin, 10 mM HEPES, 1x GlutaMAX, 1x N-2 supplement, 1x B-27 supplement (all from Gibco, Thermo Fisher Scientific Cat. # 12634010, 400–109, 15630080, 35050061, 17502001 and 17504044, respectively), 1 mM N-acetylcysteine (MilliporeSigma Cat. # 106425)), was supplemented with 50 ng/ml recombinant murine EGF (Peprotech Cat. # 315–09), and 1% v/v Noggin CM to obtain EN medium. Purified RSPO recombinant proteins were added to the basic culture medium as indicated. Organoids were cultured in matrigel droplets (Corning CellBIND, Corning Cat. # 3300, 3335, 3292). The medium was refreshed 3 and 6 days after splitting. Images were captured 8 days after splitting using an EVOS M5000 imaging system (Thermo Fisher Scientific).

Organoids were carefully harvested from matrigel, collected by centrifugation for 5 min at 100 x g at 4°C and washed with ice-cold medium. Organoids were transferred to a μ-slide 8 well (Ibidi Cat. # 80826) and fixed in paraformaldehyde (4%, diluted in 0.1 M sodium phosphate buffer pH 7.4) for 1 hr at RT. Paraformaldehyde was removed and 20 mM ammonium chloride (in PBS) was added for 10 min. Organoids were permeabilized in PBD 0.2 T buffer (1x PBS, 1% BSA, 1% DMSO, 0.2% TX-100) for 30 min at RT. Organoids were incubated with anti-human lysozyme or anti-Ki67 primary antibodies in PBD 0.2 T for 3 hr at RT, followed by anti-rabbit Alexa Fluor 488 secondary antibody containing 0.2 μg/ml DAPI and phalloidin-TRITC in PBD 0.2 T for 3 hr at RT. Organoids were mounted in Ibidi mounting medium (Ibidi Cat. # 50001) and images were acquired with a Zeiss LSM700 confocal microscope. Images were processed in ImageJ (National Institutes of Health).

HAP1-7TGP or LGR4/5/6^KO cells were mutagenized with a GT-containing retrovirus as described previously (Lebensohn et al., 2016). Six days following mutagenesis, the HAP1-7TGP or LGR4/5/6^KO cell populations were treated with a low dose (1.04%) of WNT3A CM in complete IMDM growth medium, combined with a saturating dose (10 ng/ml) of recombinant human RSPO1 (R and D Systems Cat. # 4645-RS) or RSPO3 (R and D Systems Cat. # 3500-RS), respectively. A total of 1.25 × 10⁸ singlet-gated cells were sorted by fluorescence-activated cell sorting (FACS), gating for the lowest ~10% WNT reporter (EGFP) fluorescence. A population of singlet-gated HAP1-7TGP or LGR4/5/6^KO cells (not gated based on EGFP fluorescence) was also sorted and carried along as a control to set FACS gates during the subsequent sort. Cells were expanded for 6 days to allow recovery and resetting of the WNT reporter, and a portion of the cells was used in a subsequent round of sorting, following the same treatment and FACS gating criteria. For both screens described, cells underwent two consecutive rounds of FACS sorting. 3 × 10⁷ cells expanded following the final sort for each screen were used to identify GT insertions in the sorted cell populations and an equivalent number of mutagenized but unsorted HAP1-7TGP or LGR4/5/6^KO cells were used to identify GT insertions in the respective control cell populations. Preparation of genomic GT insertion libraries for deep sequencing, mapping of sequencing reads and statistical analysis of GT insertion enrichment was conducted as described previously (Lebensohn et al., 2016), except that the libraries for the unsorted (control) and sorted cell populations from the RSPO3 screen in LGR4/5/6^KO cells were indexed and sequenced together in an Illumina NextSeq 500 system. FASTQ files containing sequencing data for the unsorted and sorted cell pupulations from the RSPO3 screen in LGR4/5/6^KO cells, as well as for the sorted cell population from the RSPO1 screen in HAP1-7TGP cells have been deposited in the National Center for Biotechnology Information (NCBI) Sequence Read Archive (SRA) with study accession number SRP260445. The FASTQ file containing the sequencing data for the unsorted cell pupulation from the RSPO1 screen in HAP1-7TGP cells had been deposited previously with study accession number SRP094861 (SRA accession number SRX2410629). We generated a ranked list of hits from each screen (Supplementary file 4) and used it to create circle plots depicting the data (Figure 6A and B). Note that the data presented here for the RSPO1 screen in HAP1-7TGP cells is different from that presented in our previous report (Lebensohn et al., 2016) because in order to maintain a comparable degree of phenotypic enrichment between this screen and the RSPO3 screen in LGR4/5/6^KO cells, we analyzed a cell population from our previous RSPO1 screen in HAP1-7TGP cells that had been sorted only twice (as opposed to four times) for the lowest ~10% WNT reporter activity.

We used two metrics to compare the results of the RSPO1 screen in HAP1-7TGP cells and the RSPO3 screen in LGR4/5/6^KO cells. The FDR-corrected p-value (Figure 6A and B, and Supplementary file 4) tests for the enrichment of inactivating GT insertions in the sorted cell population over total GT insertions in the control cell population (Lebensohn et al., 2016). While this is a good metric by which to compare the significance of hits in a given screen, it depends on many experimental variables affecting the identification of GT insertions in both the sorted and control cell populations, making it an equivocal metric to directly compare hits between screens. We therefore used the Intronic GT Insertion Orientation Bias (IGTIOB) score we developed previously (Lebensohn et al., 2016), which is based on the frequency of intronic sense vs. antisense GT insertions only in the sorted cell population from each screen, to directly compare significant hits between the RSPO1 screen in HAP1-7TGP cells and the RSPO3 screen in LGR4/5/6^KO cells. In order to avoid missing any potential hits unique to only one screen, we relaxed the GT enrichment FDR-corrected p-value threshold for inclusion in the comparative analysis from <10⁻⁴ (used in our previous studies [Lebensohn et al., 2016]) to <0.01. We generated a list of all genes with an FDR-corrected p-value<0.01 in at least one of the two screens (Supplementary file 5) and used the IGTIOB scores of those genes in both screens to build a heat map (Figure 6C) as described previously (Lebensohn et al., 2016).

Data analysis and visualization for the dose-response analysis of RSPO3 effects on WNT reporter fluorescence (Figure 1D and E, Figure 2C–H) were performed in GraphPad Prism 8. Dose-response curves were fitted with the non-linear regression curve fit using the [agonist] vs. response – variable slope (four parameters) equation. All reported EC₅₀ values were obtained from these curve fits. The statistical significance of the differences in EC₅₀ values for various RSPO variants and cell lines (Figure 4B and C) was determined by a two-tailed, unpaired t-test in GraphPad Prism 8. Statistical analysis of screen data was performed as described previously (Lebensohn et al., 2016).