LUZP1, a novel regulator of primary cilia and the actin cytoskeleton, is a contributing factor in Townes-Brocks Syndrome

Primary cilia are sensory organelles crucial for cell signaling during development and organ homeostasis. Cilia arise from centrosomes and their formation and function is governed by numerous factors. Through our studies on Townes-Brocks Syndrome (TBS), a rare disease linked to abnormal cilia formation in human fibroblasts, we uncovered the leucine-zipper protein LUZP1 as an interactor of truncated SALL1, a dominantly-acting protein causing the disease. Using TurboID proximity labeling and pulldowns, we show that LUZP1 associates with factors linked to centrosome and actin filaments. Here, we show that LUZP1 is a cilia regulator. It localizes around the centrioles and to actin cytoskeleton. Loss of LUZP1 reduces F-actin levels, facilitates ciliogenesis and alters Sonic Hedgehog signaling, pointing to a key role in cytoskeleton-cilia interdependency. Truncated SALL1 increases the ubiquitin proteasome-mediated degradation of LUZP1. Together with other factors, alterations in LUZP1 may be contributing to TBS etiology.


Introduction
Townes-Brocks Syndrome (TBS1 [MIM: 107480]) is an autosomal dominant genetic disease, caused by mutations in a transcription factor called SALL1, characterized by the presence of imperforate anus, dysplastic ears, thumb malformations and often renal and heart impairment, among other symptoms (Botzenhart et al., 2007;Kohlhase et al., 1998). Some of these features overlap those in the ciliopathic spectrum. It has been recently demonstrated that primary cilia defects are contributing factors to TBS etiology (Bozal-Basterra et al., 2018). Truncated SALL1, either by itself or in complex with full length protein (SALL1 FL ), can interact with CCP110 and CEP97. As a consequence, those negative regulators are reduced at the mother centriole (MC) and ciliogenesis is promoted (Bozal-Basterra et al., 2018).
Primary cilia are sensory organelles that have a crucial role in cell signaling and protein trafficking during development and organ homeostasis. Although several key pathways are influenced by cilia function (Wnt, TGFbeta, PDGFRalpha, Notch), the best characterized is the Sonic Hedgehog (Shh) pathway (Goetz and Anderson, 2010). Briefly, Shh binds to its receptor PTCH1 and leads to ciliary enrichment of the transmembrane protein Smoothened (SMO), with concomitant conversion of the transcription factor GLI3 from a cleaved repressor to a full-length activator form, leading to activation of Shh target genes. Two such genes are PTCH1 and GLI1 (encoding the Shh receptor and a transcriptional activator, respectively), exemplifying the feedback and fine-tuning of the Shh pathway.
Cilia arise from the centrosome, a cellular organelle composed of two barrel-shaped microtubulebased structures called the centrioles. Primary cilia formation is very dynamic throughout the cell cycle. Cilia are nucleated from the MC at the membrane-anchored basal body upon entry into the G0 phase, and they reabsorb as cells progress from G1 to S phase, completely disassembling in mitosis (Rezabkova et al., 2016). Centrioles are surrounded by protein-based matrix, the pericentriolar material (PCM) (Conduit et al., 2015;Vertii et al., 2016). In eukaryotic cells, PCM proteins are concentrically arranged around a centriole in a highly organized manner (Fu and Glover, 2012;Lawo et al., 2012;Mennella et al., 2012;Sonnen et al., 2012). Based on this observation, proper positioning and organization of PCM proteins may be important for promoting different cellular processes in a spatially regulated way (Kim et al., 2019). Not surprisingly, aberrations in the function of PCM scaffolds are associated with several human diseases, including cancer and ciliopathies (Gö nczy, 2015;Nigg and Holland, 2018). Cilia assembly is regulated by diverse factors. Among them, CCP110 and CEP97 form a cilia suppressor complex that, when removed from the MC, allows eLife digest Primary cilia are the 'antennae' of animal cells: these small, flexible protrusions emerge from the surface of cells, where they help to sense and relay external signals. Cilia are assembled with the help of the cytoskeleton, a dynamic network of mesh-like filaments that spans the interior of the cell and controls many different biological processes. If cilia do not work properly, human diseases called ciliopathies can emerge.
Townes-Brocks Syndrome (TBS) is an incurable disease that presents a range of symptoms such as malformations of the toes or fingers, hearing impairment, and kidney or heart problems. It is caused by a change in the gene that codes for a protein called SALL1, and recent work has also showed that the cells of TBS patients have defective cilia. In addition, this prior research identified a second protein that interacted with the mutant version of SALL1; called LUZP1, this protein is already known to help maintain the cytoskeleton.
In this study, Bozal-Basterra et al. wanted to find out if LUZP1 caused the cilia defects in TBS. First, the protein was removed from mouse cells grown in the laboratory, which dramatically weakened the cytoskeleton. In keeping with this observation, both the number of cilia per cell and the length of the cilia were abnormal. Cells lacking LUZP1 also had defects in a signalling process that transmits signals received by cilia to different parts of the cell. All these defects were previously observed in cells isolated from TBS patients. In addition, LUZP1-deficient mouse cells showed the same problems with their cilia and cytoskeleton as the cells from individuals with TBS. Crucially, the cells from human TBS patients also had much lower levels of LUZP1 than normal, suggesting that the protein may contribute to the cilia defects present in this disease.
The work by Bozal-Basterra et al. sheds light on how primary cilia depend on the cytoskeleton, while also providing new insight into TBS. In the future, this knowledge could help researchers to develop therapies for this rare and currently untreatable disease. ciliogenesis to proceed (Spektor et al., 2007). The actin cytoskeleton is also emerging as key regulator of cilia formation and function, with both negative and positive roles (Copeland, 2020).
Ciliary dysfunction often results in early developmental problems including hydrocephalus, neural tube closure defects (NTD) and left-right anomalies (Fliegauf et al., 2007). These features are often reported in a variety of diseases, collectively known as ciliopathies, caused by failure of cilia formation and/or cilia-dependent signaling (Hildebrandt et al., 2011). In the adult, depending on the underlying mutation, ciliopathies present a broad spectrum of phenotypes comprising cystic kidneys, polydactyly, obesity or heart malformation.
Truncated SALL1 likely interferes with multiple factors to give rise to TBS phenotypes. Here we focus on LUZP1, a leucine-zipper motif containing protein that was identified by proximity proteomics as an interactor of truncated SALL1 (Bozal-Basterra et al., 2018). LUZP1 has been previously identified as an interactor of ACTR2 (ARP2 actin related protein two homologue) and filamin A (FLNA) and, recently, as an actin cross-linking protein (Hein et al., 2015;Wang and Nakamura, 2019). Furthermore, LUZP1 shows homology to FILIP1, a protein interactor of FLNA and actin (Gad et al., 2012;Nagano et al., 2004). Interestingly, mutations in Luzp1 resulted in cardiovascular defects and cranial NTD in mice (Hsu et al., 2008), phenotypes within the spectrum of those seen in TBS individuals and mouse models of dysfunctional cilia (Botzenhart et al., 2007;Botzenhart et al., 2005;Klena et al., 2016;Kohlhase et al., 1998;Surka et al., 2001;Toomer et al., 2019). Both the non-canonical Wnt/PCP (Wingless-Integrated/planar cell polarity) and the Shh pathways are influenced by the presence of functional cilia and regulate neural tube closure and patterning (Campbell, 2003;Copp, 2005;Fuccillo et al., 2006). Remarkably, ectopic Shh was observed in the dorsal lateral neuroepithelium of the Luzp1 -/mice (Hsu et al., 2008). However, in spite of the phenotypic overlaps, a link between LUZP1 and ciliogenesis has not been explored.
Here we demonstrate that LUZP1 is associated with centrosomal and actin cytoskeleton-related proteins. We show that LUZP1 localizes to the PCM, actin cytoskeleton and the midbody, and also provide evidence towards its regulatory role on actin dynamics and its subsequent impact on ciliogenesis. Notably, we demonstrate that Luzp1 -/cells exhibit reduced filamentous actin (F-actin), longer primary cilia, higher rates of ciliogenesis and increased Shh signaling. Furthermore, TBS-derived primary fibroblasts show a reduction in LUZP1 and actin filaments, possibly through SALL1-regulated LUZP1 degradation via the ubiquitin (Ub)-proteasome system (UPS). As a novel regulator of ciliogenesis and the actin cytoskeleton, LUZP1 might contribute to the aberrant cilia phenotype in TBS.

SALL1 interacts with LUZP1
Using proximity proteomics, we have previously shown that a truncated and mislocalized form of SALL1 present in TBS individuals (SALL1 275 ) can interact aberrantly with cytoplasmic proteins (Bozal-Basterra et al., 2018). LUZP1 was found among the most enriched proteins in the SALL1 275 proximal interactome. We confirmed this finding by independent BioID experiments analyzed by western blot using a LUZP1-specific antibody ( Figure 1A and Figure 1-figure supplement 1). To further characterize the interaction of LUZP1 with truncated SALL1, we performed pulldowns using tagged SALL1 275 -YFP in HEK 293FT cells. Our results showed that endogenous LUZP1 was able to interact with SALL1 275 , confirming our proximity proteomics data ( suggesting that heterodimerization of the truncated and FL forms does not inhibit the interaction with LUZP1. When expressed alone, we noted that SALL1 FL -YFP also interacts with LUZP1 in pulldown assays (Figure 1-figure supplement 2A and Figure 1-figure supplement 1). As these proteins have distinct localizations (nuclear and cytoplasmic, respectively), the interaction likely occurs in post-lysis cell extracts (more in Discussion). These results show that the truncated form of SALL1 expressed in TBS individuals, either by itself or in complex with the FL form, can interact with LUZP1.

LUZP1 interacts with centrosomal and actin cytoskeleton components
To gain insights into the function of LUZP1, we sought to identify its proximal interactome using the TurboID approach (Branon et al., 2018). We used RPE1 cells stably expressing low levels of FLAG- Figure 1. Proximity proteomics reveal LUZP1 interaction with truncated SALL1 and with centrosome-and actin cytoskeleton-associated proteins. (A) Western blot analysis of BioID, streptavidin pulldown (PD) of HEK 293FT cells transfected with Myc-tagged BirA*-SALL1 275 or BirA*-SALL1 FL . Specific antibodies (LUZP1, actin, Myc) were used as indicated. Anti-Myc antibody detected the self-biotinylated form of BirA*-SALL1 FL (asterisk) or BirA*-SALL1 275 (black arrowhead). (B) STRING core cluster network analysis of LUZP1 interactors (confidence 0.7 or higher), visualized using Cytoscape software. Color and size of the nodes indicate Log2 of the TurboID-LUZP1 (TbID-LUZP1) versus TbID alone ratio. The most highly interconnected clusters, centrosome and actin, are indicated separately. (C) Volcano plot representing the distribution of the candidates identified by proximity proteomics in three independent experiments. Proteins with more than 1-fold change in TbID-LUZP1 intensity with respect to the TbID (Log 2 ! 0) were considered as LUZP1-associated candidates (grey dots). Proteins associated with the actin cytoskeleton and the centrosome were colored in green and pink colors, respectively. (D) Graphical representation of the -Log 10 of the p-value for each of the represented GO terms of the TbID experiment performed on RPE1 stably expressing near endogenous levels of TbID-LUZP1 vs TbID. Pink dotted line represents the cutoff of p-value<0.01. Figure 1 continued on next page TurboID-LUZP1 (TbID-LUZP1) or FLAG-TurboID (TbID) as control. Transduced cells showed subendogenous expression levels of TbID-LUZP1 (Figure 1-figure supplement 2B, two asterisks; endogenous, two open arrowheads). Staining of transfected cells revealed that, proteins biotinylated by TbID are diffusely localized throughout the nucleus and cytoplasm, whereas those biotinylated by TbID-LUZP1 are localized primarily at the centrosome and actin cytoskeleton, as shown by fluorescent streptavidin (Figure 1-figure supplement 2C). Total lysates from TbID-LUZP1 or TbIDexpressing cells were subjected to streptavidin pulldown and isolated proteins were analyzed by liquid chromatography tandem mass spectrometry (LC-MS/MS). 234 high-confidence proximity LUZP1 interactors were enriched in the TbID-LUZP1 vs the TbID proteome in three replicates (Source Data 1). Proteins enriched among the identified LUZP1 proximal interactors were centrosomal and actin cytoskeleton-related proteins ( Figure 1B-C). With the purpose of obtaining a functional overview of the main pathways associated to LUZP1, a comparative Gene Ontology (GO) analysis was performed with all the hits enriched in TbID-LUZP1 versus TbID cells. In the Cellular Component domain, 'actin cytoskeleton', 'microtubule cytoskeleton', 'centrosome', 'cilium' and 'midbody' terms were highlighted among others ( Figure 1D and Source Data 1). In the category of Biological Process, LUZP1 proteome showed enrichment in the 'microtubule cytoskeleton organization', 'cell division', 'cilium assembly', 'vesicle-mediated transport', 'microtubule organizing center organization' and 'cell adhesion' categories among others ( Figure 1D and Source Data 1). With respect to Molecular Function, LUZP1 also showed enrichment in cytoskeleton-related proteins ('actin binding', 'actinin binding' and 'microtubule binding' terms; Figure 1D and Source Data 1). 37 or 96 of the verified or potential, respectively, centrosome/cilia gene products previously identified by proteomic studies (Alves-Cruzeiro et al., 2014;Gupta et al., 2015) were found as LUZP1 proximal interactors, supporting the enrichment of centrosome-related proteins among the potential interactors of LUZP1. In addition, 45 of LUZP1 proximal interactors were present among the actin-localized proteins identified by the Human Protein Atlas project based on subcellular localization to actin filaments (Uhlen et al., 2015).

LUZP1 localizes to the centrosome, actin cytoskeleton and midbody
We examined the subcellular localization of LUZP1 in diverse cell types. First, immunostainings showed that endogenous LUZP1 surrounds both centrioles, labelled by centrin-2 (CETN2) in human RPE1 cells ( Figure 2A) and gamma-tubulin in human dermal fibroblasts (Figure 2-video 1) (for specificity of LUZP1 antibody, check Figure 5). We examined LUZP1 localization at the centrosome in synchronized RPE1 cells. LUZP1 was reduced at the centrosome during G2/M and G0 phases (Figure 2-figure supplement 1). Interestingly, LUZP1 levels increased upon treatment with the proteasome inhibitor MG132 in G0 phase arrested-RPE1 cells. All together, these results indicate that LUZP1 levels are reduced at the centrosome in G2/M phase and upon starvation, and this reduction might be mediated by the UPS. The localization of LUZP1 at the centrosome was reproduced in U2OS cells expressing LUZP1-YFP ( Figure 2B-D). We did not observe colocalization of LUZP1 with the distal centriolar marker CCP110 ( Figure 2B), indicating that LUZP1 is likely found at the proximal end of both centrioles. We further imaged LUZP1 along with pericentrin (PCNT) and PCM1, markers of PCM. Interestingly, we observed that LUZP1 enveloped PCNT ( Figure 2C), with LUZP1 itself being surrounded by PCM1 ( Figure 2D). These results suggest that LUZP1 might be a novel PCM associated-protein, decorating the proximal end of both centrioles. In concordance with this localization, LUZP1 was associated to PCM1 in TurboID experiments (Source Data 1).
In addition to the localization at the centrosome/basal body, LUZP1 also localized to actin stress fibers ( Figure 2E) and to the midbody ( Figure 2F) in U2OS cells.  LUZP1 shows reduction at the centrosome in TBS fibroblasts and interacts with centrosome-associated proteins Based on the LUZP1 interaction with truncated SALL1, we checked its subcellular localization in fibroblasts derived from a TBS individual (TBS 275 ; see Materials and methods) as well as non-TBS control. We observed that LUZP1 was markedly decreased at the centrosome of TBS 275 cells . Schematic representation of LUZP1 localization at the centrosome according to their respective micrographs in (A-D). Scale bar, 1 mm. Imaging in (A-D) was performed using confocal microscopy (Leica SP8, 63x objective). Lightning software (Leica) was applied. (E, F) Immunofluorescence micrographs of U2OS cells stained with an antibody against endogenous LUZP1 (green), phalloidin to detect F-actin (magenta), and counterstained with DAPI (blue). Single green and magenta channels are shown in black and white. (F) LUZP1 at the midbody in dividing cells (yellow arrowhead). Scale bar, 10 mm (E, F). Imaging in (E, F) was performed using widefield fluorescence microscopy (Zeiss Axioimager D1, 63x objective). The online version of this article includes the following video and figure supplement(s) for figure 2:   . Beyond pulldowns, we found that immunoprecipitation of endogenous LUZP1 led to co-purification of endogenous CCP110 ( Figure 3E and Figure 3-figure supplement 1) and that anti-CEP97 antibodies immunoprecipitated endogenous LUZP1 ( Figure 3F and Figure 3-figure supplement 1). Both CCP110 and CEP97 were also identified as proximal interactors of TbID-LUZP1 (Source Data 1). Immunofluorescent colocalization on centrioles of LUZP1 (proximal) and CCP110 (distal) was not evident, suggesting that the interaction is indirect or occurs before proteins reach their destinations. However, these key regulators of ciliogenesis are just two of multiple centrosomal proteins associated with LUZP1, suggesting that LUZP1 may have a function at this dynamic organelle.

LUZP1 localizes to actin and is altered in TBS fibroblasts
In addition to the centrosome/basal body, LUZP1 also localized to actin stress fibers, as well as the midbody in dividing cells ( Figure 2). Intriguingly, when LUZP1 levels were examined in TBS 275 cells, a reduction in both actin-associated LUZP1 and phalloidin-labelled stress fibers was observed when compared to control cells ( Figure 4A-C). These results indicate that actin cytoskeleton might be altered in TBS cells. Using pulldown assays, we confirmed that LUZP1-YFP interacts with both actin and FLNA ( Figure 4D and

LUZP1 plays a role in primary cilia formation and shh signaling
Based on the localization of LUZP1 at the centrosome, its interaction with centrosomal proteins and the defects in ciliogenesis previously observed in TBS cells (Bozal-Basterra et al., 2018), we hypothesized that LUZP1 might have a role in cilia formation. To examine this, we analyzed ciliogenesis in Figure 3 continued starved TBS 275 compared to non-starved control fibroblasts. Imaging was performed using widefield fluorescence microscopy (Zeiss Axioimager D1, 63x objective). Scale bar, 4 mm. (C) Graphical representation of the LUZP1 mean intensities (left panel) or the centrosome area (right panel), corresponding to the experiments shown in (A-B); n ! 6 micrographs. Three independent experiments were pooled together. P-values were calculated using the unpaired two-tailed Student´s test or U-Mann-Whitney test. (D) Western blot of inputs (lanes 1 to 4) and GFP-Trap pulldowns (lines 5 to 8) performed in WT HEK 293FT cells or in 293 335 SALL1 mutant cells transfected with LUZP1-YFP (lanes 1, 3, 5 and 7) or YFP alone (lanes 2, 4, 6 and 8). Numbers under CCP110 and CEP97 panels result from dividing band intensities of each pulldown by their respective input levels. GAPDH was used as loading control. (E, F) Co-immunoprecipitation experiments show LUZP1-CCP110 (E) and CEP97-LUZP1 (F) interactions. Rabbit IgG was used for immunoprecipitation controls. GAPDH was used as loading and specificity control. In all panels, specific antibodies (LUZP1, GAPDH, CCP110, CEP97, GFP) were used as indicated. Blots shown here are representative of three independent experiments. Molecular weight markers are shown to the right. The online version of this article includes the following figure supplement(s) for figure 3:  . Reduction in LUZP1 coincides with decreased F-actin. (A) Immunofluorescence micrographs of Control and TBS 275 human fibroblasts stained with an antibody against endogenous LUZP1 (green), phalloidin to label F-actin (magenta), and counterstained with DAPI to label the nuclei (blue). Black and white images show the single green and magenta channels. Note the overall reduction in LUZP1 and F-actin levels in TBS 275 compared to control fibroblasts. Scale bar, 10 mm. Imaging was performed using widefield fluorescence microscopy (Zeiss Axioimager D1, 63x objective). (B, C) Graphical representation of the LUZP1 (B) and F-actin (C) mean intensities, corresponding to the experiments shown in (A); n ! 6 micrographs. Three independent experiments were pooled together. P-values were calculated using the unpaired two-tailed Student´s test or U-Mann-Whitney test. (D) Western blot of inputs or GFP-Trap pulldowns performed in HEK 293FT cells transfected with LUZP1-YFP or YFP alone. Anti-GFP antibody detected YFP alone (two black arrowheads) and LUZP1-YFP (two white arrowheads). Blots shown here are representative of three independent experiments. Molecular weight markers are shown to the right. Specific antibodies (LUZP1, GAPDH, CCP110, CEP97, GFP) were used as indicated. (E) Western blot of total cell lysates of HEK 293FT treated or not with cytochalasin D (CytoD) in a mild lysis buffer (TX-100 0.1%, lanes 1, 2) or a strong lysis buffer (WB5, lanes 3, 4). Note the increase in LUZP1 levels upon actin polymerization blockage with CytoD, exclusively when cells were lysed on 01% TX-100-based lysis buffer. GAPDH was used as loading control. In (D) and (E) panels, specific antibodies (LUZP1, GAPDH, actin, FLNA, GFP) were used as indicated. (F) Graphical representation of LUZP1 vs GAPDH band intensities in (E) normalized to lane 1. Graphs represent Mean and SEM of three independent experiments. P-value was calculated using two tailed unpaired Student´s t-test. Molecular weight markers in (D) and (E) are shown to the right. The online version of this article includes the following figure supplement(s) for figure 4:

Figure 4 continued on next page
Shh-LIGHT2 cells, a cell line derived from immortalized mouse NIH3T3 fibroblasts that can display primary cilia and report on Shh pathway status using integrated luciferase reporters (herein designated as WT) (Taipale et al., 2000). Using CRISPR/Cas9 gene editing directed to exon 1 of murine Luzp1, we generated Shh-LIGHT2 mouse embryonic fibroblasts null for Luzp1 (Luzp1 -/cells). For genetic rescue experiments, LUZP1 was restored to these cells by the expression of human LUZP1-YFP fusion (+LUZP1 cells). To examine the effect of the Luzp1 mutation and rescue strategies, we used anti-LUZP1 antibody and checked LUZP1 localization associated with the actin cytoskeleton and the centrosome by immunofluorescence ( Figure 5A,B), and its levels of expression by western blot ( Figure 5C and To analyze the role of LUZP1 in ciliation, WT, Luzp1 -/and +LUZP1 cells were plated at equal densities and induced to ciliate for 48 hr by serum withdrawal (starved; Figure 6A). We quantified ciliation rates and primary cilia length in non-starved and starved cells. Luzp1 -/fibroblasts displayed higher ciliation rate (60%) than WT (10.5%) and +LUZP1 (22.2%) when the cells were not subjected to starvation ( Figure 6B). However, Luzp1 -/cells were not significantly more ciliated than WT or    Figure 6B). In addition, primary cilia in Luzp1 -/cells were significantly longer than in non-starved WT cycling cells ( Figure 6A and C), while under starvation the differences were not significant ( Figure 6A and C). Note that, differently than the ciliation rate, cilia length was not rescued by adding human LUZP1. Taken together, these results confirm that Luzp1 -/cells display longer and more abundant primary cilia compared to WT cells in cycling conditions and indicate that LUZP1 might affect primary cilia dynamics.
One key event in ciliogenesis is the depletion of CCP110 and its partner CEP97 from the distal end of the MC, promoting the ciliary activating program in somatic cells (Goetz et al., 2012;Figure 6. Loss of Luzp1 causes aberrant cilia frequency and length and Shh signaling. (A) Micrographs of Shh-LIGHT2 cells (WT), Shh-LIGHT2 cells lacking Luzp1 (Luzp1 -/-) and Luzp1 -/cells rescued with human LUZP1-YFP (+LUZP1) analyzed in cycling conditions (non-starved), or during cilia assembly (48 hr starved). Cilia were visualized by acetylated alpha-tubulin (magenta), basal body by ODF2 (green) and nuclei by DAPI (blue). Scale bar 2.5 mm. (B, C) Graphical representation of percentage of ciliated cells per micrograph (B) and cilia length (C) measured in WT (blue circles, n > 34 micrographs), Luzp1 -/-(orange circles, n > 44 micrographs) or +LUZP1 cells (green circles, n > 30 micrographs) from three independent experiments. No starvation: WT 2.3 mm; Luzp1 -/cells 3.0 mm; +LUZP1 cells 2.9 mm; 48 hr starvation: WT 4.2 mm; Luzp1 -/cells 4.1 mm; +LUZP1 cells 4.8 mm; all average measures. (D) Immunofluorescence micrographs of WT and LUZP1 -/cells stained with antibodies against endogenous CCP110 (green), gamma-tubulin (gTub) to label the centrioles (magenta) and DAPI to label the nuclei (blue). Black and white images show the single green and magenta channels. Note the different distribution of CCP110 to the centrosome in LUZP1 -/compared to WT cells. Scale bar, 1 mm. (E) Graphical representation of the percentage of cells showing the presence of CCP110 to both centrioles per micrograph corresponding to the experiments in (D); n = 10 micrographs. Three independent experiments were pooled together. Kleylein-Sohn et al., 2007;Prosser and Morrison, 2015;Spektor et al., 2007;Tsang et al., 2008). We analyzed the centrosomal localization of CCP110 in WT and Luzp1 -/cells by immunofluorescence. Consistent with the higher ciliogenesis rate, CCP110 was present at two centrosomal spots at a lower proportion in Luzp1 -/cells (19%) compared to WT (84%; Figure 6D and E). This result suggests that the lack of LUZP1 might result in CCP110 reduction at the centrosome, leading to higher frequency of ciliogenesis in Luzp1 -/cells, and is reminiscent to the results obtained in TBS cells (Bozal-Basterra et al., 2018).
It is well-established that mammalian Shh signal transduction is dependent on functional primary cilia (Huangfu et al., 2003;Yin et al., 2009). Therefore, we examined whether Shh signaling is altered in Luzp1 -/cells. Cells were starved for 24 hr and incubated in the presence or absence of purmorphamine (a SMO agonist) for 24 hr to activate the Shh pathway. The mRNA expression of two Shh target genes (Gli1 and Ptch1) was quantified by qRT-PCR ( Figure 7A,B). We found that Gli1 and Ptch1 expression levels in non-treated Luzp1 -/cells were higher than in WT cells (Gli1 1.5 fold and Ptch1 2.3 fold increase in Luzp1 -/vs WT cells without purmorphamine) ( Figure 7A,B). To further study the role of LUZP1 in Shh signaling, we analyzed GLI3 processing by western blot using total lysates extracted from WT vs Luzp1 -/cells. Without purmorphamine induction, we found a significantly higher ratio of GLI3 activating form vs GLI3 repressive form (GLI3-A:GLI3-R) in Luzp1 -/cells compared to WT (2.9 fold increase in Luzp1 -/cells vs WT) ( Figure 7C and (n = 7) (B) obtained by qRT-PCR from wild-type Shh-LIGHT2 cells (WT; blue dots) or Shh-LIGHT2 cells lacking Luzp1 (Luzp1 -/-; orange dots), treated (+) or not (-) with purmorphamine for 24 hr. (C) Western blot analysis of lysates from WT and Luzp1 -/cells. Samples were probed against GLI3 using an antibody that detects both GLI3-activator form (GLI3-A) and GLI3-repressor form (GLI3-R); GAPDH was used as loading control. Numbers under the lanes are the Mean of 3 independent experiments, resulting of dividing the activator by the repressor intensities, taking WT non-induced value as 1. Molecular weight markers are shown to the right. (D, E) Graphical representation of fold-change in luciferase activation when WT (n > 7; blue dots), Luzp1 -/-(n > 7; orange dots) or +LUZP1 (n = 4; green dots) cells are treated for 6 and 24 hr or not (-) with purmorphamine. (E) Each treated condition was normalized against its respective non treated condition, taking the non-induced value as 1 (dashed line). All graphs represent the Mean and SEM. P-values were calculated using two-tailed unpaired Student´s t-test or One-way ANOVA and Bonferroni post-hoc test. The online version of this article includes the following figure supplement(s) for figure 7: supplement 1). After induction, the values were similar for Luzp1 -/and WT cells. We also examined the effects of lacking Luzp1 on Shh signaling by measuring the activity of the Shh-responsive Firefly luciferase reporter in the presence or absence of purmorphamine for 24 hr to activate the Shh pathway ( Figure 7D,E). Consistent with the Gli1 and Ptch1 qRT-PCR data, non-treated Luzp1 -/cells showed higher Shh activity compared to control or +LUZP1 cells, as observed in TBS-derived cells ( Figure 7D). However, the induction capacity of Luzp1 -/cells upon purmorphamine treatment was reduced compared to WT or +LUZP1 cells ( Figure 7E). Altogether, the observed defects in Ptch1 and Gli1 gene expression and Shh reporter misregulation point to a role for LUZP1 in Shh signaling.

LUZP1 and F-actin levels are correlated
Based on the localization of LUZP1 to actin stress fibers and interaction with cytoskeletal proteins, we hypothesized that LUZP1 might also affect F-actin cytoskeleton. We observed a reduction in F-actin (labelled by phalloidin) in the Luzp1 -/cells compared to WT, which was recovered in +LUZP1 cells ( Figure 8A). We also note the correlation between LUZP1 levels and actin filaments in nonstarved versus starved WT fibroblasts ( Figure 8B,C). These results suggest that LUZP1 can stabilize actin stress fibers and that starvation triggers both LUZP1 and F-actin reduction.
Truncated SALL1 promotes LUZP1 degradation via the ubiquitin proteasome system In concordance with immunofluorescence results in Figures 3 and 4, we confirmed a reduction in total LUZP1 levels in TBS 275 cells compared to controls by western blot ( Figure 9A,B and    1 and 3) or YFP alone (lanes 2 and 4) treated (+) or not (-) with MG132. Specific antibodies against LUZP1, GFP and GAPDH were used. One black arrowhead indicates SALL1 275 -YFP, two back arrowheads YFP alone. (F) Graphical representation of the fold changes of LUZP1/GAPDH ratios obtained in (E) for HEK 293FT cells transfected with SALL1 275 -YFP (orange dots) or YFP alone (blue dots) treated (+) or not (-) with MG132. Note that LUZP1 increases in the presence of MG132 when SALL1 275 -YFP was transfected. Data from at least three independent experiments pooled together are shown. P-values were calculated using two-tailed unpaired Student´s t-test. (G) Immunofluorescence micrographs of RPE1 cells treated (+MG132) or not (-MG132) with proteasome inhibitor showing LUZP1 associated with the cytoskeleton (upper panels) or in the centrosome (lower panels). Antibodies against endogenous LUZP1 (green), Centrin-2 (CETN2, blue) and CEP164 (magenta) were used. DAPI labelled the nuclei (blue). Single green channels are shown in black and white. Note the overall increase of LUZP1 upon MG132 treatment. Scale bar 10 mm (cytoskeleton panels) or 0.5 mm (centrosome panels). Imaging was performed using widefield fluorescence microscopy (Zeiss Axioimager D1, 63x objective).  control and TBS 275 samples (Figure 9-figure supplement 2), so we hypothesized that truncated SALL1 might lead to UPS-mediated LUZP1 degradation. We analyzed LUZP1 levels after treatment with the proteasome inhibitor MG132, both in control and TBS 275 cells, and LUZP1 levels were increased to a higher extent in TBS 275 compared to control cells (1.8 fold increase in control vs 2.4 fold increase in TBS 275 cells) ( Figure 9A,B). Moreover, we confirmed the reduction of LUZP1 levels in the CRISPR/Cas9 TBS model cell line (293 335 ), compared to its parental cell line (293 WT ) ( Figure 9C,D and Figure 9-figure supplement 1), and likewise in HEK 293FT cells stably overexpressing truncated SALL1 (SALL1 275 -YFP) compared to cells with YFP as control ( Figure 9E,F and Figure 9-figure supplement 1). A more prominent increase in LUZP1 accumulation upon MG132 treatment was also observed in 293 335 and HEK 293FT cells overexpressing SALL1 275 -YFP compared to controls ( Figure 9C,D and Figure 9E,F, respectively, and Figure 9-figure supplement 1). Additionally, we also observed LUZP1 accumulation upon MG132 treatment by immunofluorescence in RPE1 cells, both at the actin cytoskeleton ( Figure 9G, upper panels) and at the centrosome ( Figure 9G, lower panels). All together, these results show that LUZP1 levels are sensitive to degradation via the UPS pathway and suggest that truncated SALL1 may contribute to this process. Furthermore, we compared LUZP1 ubiquitination in 293 WT vs 293 335 cells using the BioUb strategy (see Materials and methods) (Pirone et al., 2017). In the pulldowns, we could observe a prominent band in presence of BioUb, possibly corresponding to a monoubiquitinated form of LUZP1 ( Figure 9H and

LUZP1 overexpression represses cilia formation and increases F-actin levels in TBS fibroblasts
Our results suggest that LUZP1 could be a mediator of TBS cilia phenotype and that this could be caused, at least in part, by the increased degradation of LUZP1 triggered by truncated SALL1. Therefore, increasing LUZP1 levels in TBS cells might affect the cilia and actin cytoskeleton phenotypes. To check whether an increase in LUZP1 levels is sufficient to repress ciliogenesis in primary human fibroblasts, Control and TBS 275 cells were transduced with YFP or LUZP1-YFP using lentivirus ( Figure 10A). Whereas most non-transduced surrounding cells, as well as 100% of the TBS 275 cells expressing YFP were ciliated, only 40% of the Control and TBS 275 cells transduced with LUZP1-YFP displayed cilia ( Figure 10B). Furthermore, we checked the actin cytoskeleton defects observed in TBS 275 cells to see the effect of overexpressing LUZP1-YFP. Immunostaining showed that LUZP1-YFP overexpression led to an increase in F-actin levels both in control and in TBS 275 cells compared to the surrounding non-transfected cells or TBS 275 cells overexpressing YFP ( Figure 10C).
F-actin has a suppressive effect on ciliogenesis, and CytoD-mediated actin depolymerization has been shown to be permissive to cilia formation in cultured cells (Kim et al., 2010). To corroborate the relationship between LUZP1, actin and ciliogenesis, we performed CytoD treatment experiments. We observed that LUZP1-YFP overexpression can counteract the positive effects of CytoD on cilia formation (Figure 10-figure supplement 1). All together, these results support the notion that LUZP1 is a negative regulator of cilia formation and an F-actin stabilizing protein.

Discussion
We conclude that LUZP1 might be a contributing factor to the TBS phenotype via its interaction with truncated SALL1 and its effect on ciliogenesis, based on several findings: i) LUZP1 levels are reduced in TBS-derived cells likely due to truncated SALL1-mediated degradation through the UPS; ii) LUZP1 interacts with proteins of the centrosome and of the actin cytoskeleton; iii) In the absence of LUZP1, the assembly and growth of primary cilia is enhanced in cycling cells, accompanied by an increase in basal Shh signaling, and the actin cytoskeleton is reduced; iv) Increase of LUZP1 reduces the levels of ciliogenesis in TBS-individuals derived cells. All these results do not rule out the possibility that, in addition to LUZP1, other factors might be contributing to TBS etiology.  Figure 10 continued on next page LUZP1 localizes to the centrosome and actin cytoskeleton LUZP1 was previously described as a nuclear protein, with expression limited to the mouse brain (Lee et al., 2001;Sun et al., 1996). We tested two different commercial antibodies against LUZP1 by immunofluorescence and, while nuclear localization was sporadic and weak, the most prominent localization of LUZP1 was observed in the actin cytoskeleton and centrosome, both in human and mouse cells. This localization is consistent with our TurboID analysis that showed an enrichment of factors associated with the actin cytoskeleton and/or centrosomes among the potential interactors of LUZP1. The localization of LUZP1 to the actin cytoskeleton, as well as its expression in tissues beyond the brain, is consistent with independent validation in cell lines by the Human Protein Atlas (HPA; proteinatlas.org) and other expression databases (e.g. EMBL EBI Expression Atlas ebi.ac.uk/ gxa). Two independent proximity labeling studies identified LUZP1 as a proximal interactor of centriole (Gupta et al., 2015) and centriolar satellite-related proteins (Gheiratmand et al., 2019). Moreover, LUZP1 has been recently reported as a centrosomal protein involved in cilia regulation (Gonç alves et al., 2019). These localizations are also consistent with fluorescent LUZP1 fusion proteins (this work; [Gupta et al., 2015; Gonç alves et al., 2019] #117). Discrepancies with the previously reported LUZP1 localization and distribution might be due to technical differences, such as epitope specificity for the antisera used in the immunohistochemistry.
Here, we report that LUZP1 surrounds the proximal end of both centrioles. Like LUZP1, a large number of centrosomal scaffold proteins contain coiled-coil regions, and the proteins are concentrically localized around a centriole in a highly organized fashion (e.g. Cep120, Cep57, Cep63, Cep152, CPAP, Cdk5Rap2, PCNT) (Fu and Glover, 2012;Lawo et al., 2012;Mennella et al., 2012). Furthermore, we show that LUZP1 interacts with centrosome and actin-related proteins (Figure 3 and Figure 4). LUZP1 is associated with CCP110 and CEP97, using pull down, immunoprecipitation and proximity proteomics approaches. This association is likely to be dynamic and potentially indirect, via other bridging factors, since our immunostainings show that LUZP1 and CCP110 are located in different areas of the centrioles. LUZP1 has also been identified as an interactor of ACTR2 (ARP2 actin related protein two homologue) and FLNA (Hein et al., 2015;Wang and Nakamura, 2019), and it has been recently described as an actin cross-linking protein (Wang and Nakamura, 2019). We found that LUZP1 localizes not only to centrioles and actin cytoskeleton, but also to the midbody in dividing cells, which was recently reported to influence ciliogenesis in polarized epithelial cells (Bernabé-Rubio et al., 2016). Our data suggest that the association of LUZP1 to centrosomes and actin filaments may contribute to its overall roles. Figure 10 continued arrow). AcTub: acetylated alpha-tubulin (magenta); CETN2: Centrin-2 (blue). (B) Graphical representation of the number of ciliated cells per micrograph in Control and TBS 275 cells overexpressing YFP or LUZP1-YFP (n > 10 micrographs). Graphs represent Mean and SEM of ciliation frequencies per micrograph in three independent experiments pulled together. P-values were calculated using One-way ANOVA and Bonferroni post-hoc test or two-tailed unpaired Student´s t-test. (C) Representative micrographs of Control and TBS 275 cells transduced with YFP (yellow arrowhead) or LUZP1-YFP (white asterisk) co-stained with phalloidin to label F-actin (magenta) and DAPI (blue). Note the increase in F-actin levels in cells transduced with LUZP1-YFP (white asterisk) compared to non-transfected cells (yellow arrow). Scale bar, 10 mm. Imaging was performed using widefield fluorescence microscopy (Zeiss Axioimager D1, 63x objective). (D) Schematic model representing how the presence of truncated SALL1 might cause cilia and actin malformations in TBS through LUZP1 interaction and UPS-mediated degradation. In control (or fed) cells (left), LUZP1 (in green) localizes to F-actin and to the proximal ends of the two centrioles, inhibiting cilia formation. LUZP1 proteostasis is maintained by the Ubiquitin Proteasome System (UPS). By contrast, in TBS (or starved) cells (right) the truncated form of SALL1 interacts with LUZP1 and promotes its UPS-mediated degradation. Likewise, others have shown ubiquitination and proteasome-mediated degradation of CCP110 at the mother centriole (MC) is permissive for ciliogenesis (Li et al., 2013). LUZP1 reduction at the centrosome and at the cytoskeleton favors the formation of the primary cilia. The online version of this article includes the following figure supplement(s) for figure 10: LUZP1 as an integrator of actin and primary-cilium dynamics Actin dynamics coordinate several processes that are crucial for ciliogenesis. For example, positioning the MC to the appropriate area at the cell cortex is an actin-dependent process (Boisvieux-Ulrich et al., 1990;Euteneuer and Schliwa, 1985). A reduction in cortical actin should promote ciliogenesis, since there is less physical restriction for docking of the MC. Supporting this hypothesis, several studies have found that disruptions in the actin cytoskeleton, induced either chemically or genetically, promote ciliogenesis or affect cilia length (Cao et al., 2012;Drummond et al., 2018;Hernandez-Hernandez et al., 2013;Kang et al., 2015;Kim et al., 2015;Kim et al., 2010). How actin regulates cilium length is not clear. Recently, a role for actin has been implicated in ectocytosis and cilium tip scission, preventing the axoneme from growing too long (Nager et al., 2017;Phua et al., 2017). Whether LUZP1 at the centrosome is complexed with filamin and actin is unknown, but if so, they could together serve to stabilize the basal body as the axoneme extends or is subjected to mechanical stress.

LUZP1 is altered in TBS-derived cells
TBS is caused by mutations in SALL1 gene, which give rise to truncated proteins that interfere with the normal function of the cell. We show that LUZP1interacts with truncated SALL1 and with SALL1 FL , suggesting that interaction occurs through an N-terminal domain shared by both. In control cells, LUZP1 and SALL1 FL likely have minimal or no interaction due to their respective localizations to the cytoplasm and nucleus. However, truncated SALL1, alone or together with SALL1 FL that is retained in the cytoplasm, can interact with cytoplasmic LUZP1, promoting its degradation and functional inhibition. Importantly, we detected an increase in LUZP1 levels upon treatment with the proteasome inhibitor MG132 (Figure 9), suggesting that LUZP1 degradation is proteasome-dependent. Next, we demonstrated that LUZP1 is ubiquitinated, and that truncated SALL1 both increases LUZP1 ubiquitination and decreases its stability. In agreement with our findings, LUZP1 ubiquitination has been detected in several proteomic screens for ubiquitinated proteins (Akimov et al., 2018;Mertins et al., 2013;Povlsen et al., 2012;Udeshi et al., 2013;Wagner et al., 2012).
Further experiments would be required to understand the precise mechanism by which truncated SALL1 can influence LUZP1 ubiquitination, but one possibility could be de novo complexes involving specific Ub E3 ligases or de-ubiquitinases which could influence LUZP1 stability. In fact, various E3s/ de-ubiquitinases, as well as other components of the UPS, were found as proximal interactors of truncated SALL1 and LUZP1. Furthermore, regulation by the UPS system has been reported for centrosomal factors, including the cilia regulator CCP110 (D'Angiolella et al., 2010;Hossain et al., 2017;Li et al., 2013).
Both Luzp1 -/and TBS cells showed a reduction in F-actin accompanied by an increase in ciliation. We suggest that the reduction in F-actin in TBS cells might contribute to their higher cilia abundance, longer cilia and increased Shh signaling. An increase in LUZP1 in control and TBS 275 cells is sufficient to increase F-actin levels and reduce cilia frequency, supporting that LUZP1 may have a contributing role in the TBS phenotype.

The role of LUZP1 in TBS phenotype
Cilia formation, Shh signaling, and the actin cytoskeleton is aberrant in TBS patient-derived fibroblasts (this work; [Bozal-Basterra et al., 2018]). Changes in Shh signaling have not been reported in mouse models for TBS, nor in any other cell types or tissues derived from TBS patients. Nevertheless, the phenotypes observed in TBS individuals fall within the spectrum of those observed in ciliopathies, characterized by malformations in digits, ears, heart, brain, kidneys and urogenital anomalies, phenotypes that are consistent with misregulated Shh signaling. Preaxial polydactyly has been associated with ectopic Shh expression in limbs (Dunn et al., 2011;Johnson et al., 2014;Lettice et al., 2003;Lettice et al., 2008;Liem et al., 2009;Zhulyn and Hui, 2015); Anal stenosis or imperforate anus have been related to misregulation of Shh pathway (Kang et al., 1997;Mo et al., 2001;Roberts et al., 1995), as well as deafness and dysplastic ears (Driver et al., 2008).
We observed primary cilia, Shh signaling and cytoskeletal defects in Luzp1 -/cells. Several studies have implicated defective primary cilia and Shh signaling in the etiology of neural tube closure defects, as well as crucial roles for the actin cytoskeleton (Wallingford, 2005). There are certain parallels between phenotypes observed in animal models of Luzp1 and Sall1. Exencephaly and neural tube defects were detected in mice and Xenopus Sall1 mutants (Bö hm et al., 2008;Exner et al., 2017;Kiefer et al., 2003). Luzp1 KO mice exhibit ectopic Shh expression in the hindbrain neuroepithelium and display NTDs, however the expression of Shh-responsive genes (such as Gli1 or Ptch1) was not reported (Hsu et al., 2008). Perhaps the role of LUZP1 in Shh signaling, in spatial control of the signal or the response (or both), contributes to the NTD phenotype. Exencephaly may also be caused by failure in bending at the dorsolateral hinge point of the neural folds, where cells undergo changes in apical actin architecture (Sadler et al., 1982). Luzp1 KO embryos exhibited dorsolateral neural folds that were convex instead of the concave morphology observed in WT embryos (Hsu et al., 2008), suggested that defective actin dynamics may contribute to the NTD phenotype. Taken together, defective actin dynamics, aberrant primary cilia and changes in Shh signaling might lead to NTDs observed in the LUZP1 mouse model, as well as other animal models of TBS and loss of Sall-related proteins.
In addition to NTDs, cardiac malformations are another feature found in human ciliopathies (Klena et al., 2017). Cardiac defects are observed in Luzp1 knockout mice (Hsu et al., 2008), as well as TBS patients (Kohlhase, 1993). TBS cardiac defects include atrial or ventricular septal defect, the latter of which is seen in Luzp1 knockout mice. Moreover, compound Sall1/Sall4 KO mutant mice exhibit both NTDs and cardiac problems (Bö hm et al., 2008). While Luzp1 and Sall1 may both contribute to neural tube and heart development, a novel crosstalk may arise in TBS due to dominantlyacting truncated SALL1 that could derail these processes and cause deformities.
In conclusion, our data indicate that LUZP1 functions as a cilia suppressor ( Figure 10D). It localizes to actin stress fibers and to the centrosome. In starved cells, likewise in TBS cells, overall LUZP1 levels are diminished in both structures, which facilitates the formation of the primary cilia. In TBS cells, the truncated form of SALL1 localizes to the cytoplasm, interacting with LUZP1 and other factors, leading to the degradation of LUZP1, simulating what happens when control cells undergo starvation. As a result, the frequency of cilia formation increases, and cilia are longer than in control cells.
Ciliogenesis requires communication between the actin cytoskeleton and the centrosome. Here, we propose that LUZP1 might contribute as a nexus in this complex intracellular network that is disrupted by truncated SALL1. Our findings point to the intriguing possibility that LUZP1 might be a key relay switch in this network that, together with other factors, might contribute to the phenotypes observed in TBS.
Cell synchronization and drug treatment RPE1 cells were arrested in G1 phase by treatment with mimosine (Sigma, 400 mM) for 24 hr. For S phase arrest, cells were subjected to thymidine treatment (Sigma, 2.5 mM) for 16 hr, followed by release for 8 hr, and subsequently blocked again for 16 hr. For G2/M phase arrest, cells were treated with RO-3306 (Sigma, 10 mM) for 20 hr. For G0 phase synchronization and inducing primary cilia formation, cells were starved for 48 hr (DMEM, 0% FBS, 1% penicillin and streptomycin). Treatments with the proteasome inhibitor MG132 (Calbiochem, 5 mM) were for 15 hr and with CytoD (Sigma, 50 nM) for 16 hr to stimulate actin depolymerization. HEK 293FT cells were transfected using calcium phosphate method and U2OS cells using Effectene Transfection Reagent (Qiagen).

Mass spectrometry
Analysis was done in RPE1 cells stably expressing TbID or TbID-LUZP1 at sub-endogenous levels. Three independent pulldown experiments (1.5 Â 10 8 cells per replicate) were analyzed by MS. Samples eluted from the NeutrAvidin beads were separated in SDS-PAGE (50% loaded) and stained with Sypro-Ruby (Biorad) according to manufacturer's instructions. Entire gel lanes were excised, divided into pieces and in-gel digested with trypsin. Recovered peptides were desalted using stage-tip C18 microcolumns (Zip-tip, Millipore) and resuspended in 0.1% FA prior to MS analysis. Samples were analyzed in a novel hybrid trapped ion mobility spectrometry -quadrupole time of flight mass spectrometer (timsTOF Pro with PASEF, Bruker Daltonics) coupled online to a nanoElute liquid chromatograph (Bruker). This mass spectrometer takes advantage of a novel scan mode termed parallel accumulation -serial fragmentation (PASEF), which multiplies the sequencing speed without any loss in sensitivity (Meier et al., 2015), providing outstanding analytical speed and sensibility for proteomics analyses (Meier et al., 2018). Sample (200 ng) was directly loaded in a 15 cm Bruker nanoelute FIFTEEN C18 analytical column (Bruker) and resolved at 400 nl/min with a 100 min gradient. Column was heated to 50˚C using an oven. Protein identification and quantification was carried out using PEAKS software (Bioinformatics solutions). Searches were carried out against a database consisting of human entries (Uniprot/Swissprot), with precursor and fragment tolerances of 20 ppm and 0.05 Da. Only proteins identified with at least two peptides at FDR < 5% were considered for further analysis. Data were loaded onto Perseus platform (Tyanova et al., 2016) and further processed (Log 2 transformation, selection of proteins with at least two valid values in at least one condition, imputation). A t-test was applied in order to determine the statistical significance of the differences detected, and heatmaps were generated using this tool. Protein IDs were ranked according to the number of peptides found and their corresponding intensities. Gene ontology (GO) term enrichment was analyzed using g:GOSt Profiler, a tool integrated in the g:Profiler web server (Reimand et al., 2016). GO enrichment was obtained by calculating -Log 10 of the P-value.
Network analysis of LUZP1 interactors was performed using the String app version 1.4.2 in Cytoscape version 3.7.2, with a high confidence interaction score (0.7). Transparency and width of the edges were continuously mapped to the String score (textmining, databases, coexpression, experiments, fusion, neighborhood and cooccurrence). Color, transparency and size of the nodes were discretely mapped to the Log2 enrichment value as described in Figure 1. The Molecular COmplex DEtection (MCODE) plug-in version 1.5.1 was used to identify highly connected subclusters of proteins (degree cutoff of 2; Cluster finding: Haircut; Node score cutoff of 0.5; K-Core of 2; Max. Depth of 100).

Immunoprecipitation
All steps were performed at 4˚C. Cells were collected and lysates were processed as described for GFP-trap pulldowns. After saving 40 ml of supernatant (input), the rest was incubated overnight with 1 mg of anti-CEP97 antibody (Proteintech), or anti-LUZP1 antibody (Sigma HPA028506) and for additional 4 hr with 40 ml of pre-washed Protein G Sepharose 4 Fast Flow beads (GE Healthcare) on a rotating wheel. Beads were washed 5 times for 5 min each with WB (10 mM Tris-HCl pH 7.5, 137 mM NaCl, 1 mM EDTA, 1% Triton X-100). Beads were centrifuged at 2000 x g for 2 min after each wash. For elution, samples were boiled for 5 min at 95˚C in 2x Laemmli buffer.

Luciferase assays
Shh-LIGHT2 cells were starved for 48 hr, and treated or not for the last 24 hr with purmorphamine (5 mM, ChemCruz) to induce Shh signaling pathway. Firefly luciferase expression was measured using the Dual-Luciferase Reporter Assay System (Promega) according to the manufacturer's instructions. Luminescence was measured and data were normalized to the Renilla luciferase readout. For each construct, luciferase activity upon purmorphamine treatment was divided by the activity of cells before treatment to obtain the fold change value. Experiments were performed with both biological (n = 3) and technical (n = 6) replicates.

Statistical analysis
Statistical analysis was performed using GraphPad 6.0 software. Data were analyzed by Shapiro-Wilk normality test and Levene´s test of variance. We used two-tailed unpaired Student´s t-test or Mann Whitney-U tests for comparing two groups, One-way ANOVA or Kruskall-Wallis and the corresponding post-hoc tests for more than two groups and two-way ANOVA to compare more than one variable in more than two groups. P values were represented by asterisks as follows: (*) p-value<0.05; (**) p-value<0.01; (***) p-value<0.001; (****) p-value<0.0001. Differences were considered significant when p<0.05. Values used for graphical representations and statistical analysis are available in Source Data 2.