# TBC1D32 variants disrupt retinal ciliogenesis and cause retinitis pigmentosa

Béatrice Bocquet, Caroline Borday, Nejla Erkilic, Daria Mamaeva, Alicia Donval, Christel Masson-Garcia, Karine Parain, Karolina Kaminska, Mathieu Quinodoz, Irene Perea-Romero, et al.

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Béatrice Bocquet, Caroline Borday, Nejla Erkilic, Daria Mamaeva, Alicia Donval, et al. TBC1D32 variants disrupt retinal ciliogenesis and cause retinitis pigmentosa. JCI Insight, 2023, 8 (21), pp.e169426. 10.1172/jci.insight.169426. hal-04229192

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# TBC1D32 variants disrupt retinal ciliogenesis and cause retinitis pigmentosa

Béatrice Bocquet, ..., Muriel Perron, Vasiliki Kalatzis

JCI Insight. 2023. https://doi.org/10.1172/jci.insight.169426.

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Ophthalmology

## Graphical abstract

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TBC1D32 variants disrupt retinal ciliogenesis and cause retinitis pigmentosa

Béatrice Bocquet<sup>1,2,#,*</sup>, Caroline Borday<sup>3,#</sup>, Nejla Erkilic<sup>1,2,#</sup>, Daria Mamaeva<sup>1,#</sup>, Alicia Donval<sup>3</sup>, Christel Masson-Garcia<sup>3</sup>, Karine Parain<sup>3</sup>, Karolina Kaminska<sup>4,5</sup>, Mathieu Quinodoz<sup>4,5,6</sup>, Irene Perea-Romero<sup>7,8</sup>, Gema Garcia-Garcia<sup>8,9,10</sup>, Carla Jimenez-Medina<sup>1</sup>, Hassan Boukhaddaoui<sup>1</sup>, Arthur Coget<sup>11,12</sup>, Nicolas Leboucq<sup>11</sup>, Giacomo Calzetti<sup>4,5,13</sup>, Stefano Gandolfi<sup>13</sup>, Antonio Percesepe<sup>13</sup>, Valeria Barili<sup>13</sup>, Vera Uliana<sup>14</sup>, Marco Delsante<sup>15</sup>, Francesca Bozzetti<sup>16</sup>, Hendrik PN Scholl<sup>4,5</sup>, Marta Corton<sup>7,8</sup>, Carmen Ayuso<sup>7,8</sup>, José M. Millan<sup>8,9,10</sup>, Carlo Rivolta<sup>4,5,6</sup>, Isabelle Meunier<sup>1,2,§</sup>, Muriel Perron<sup>3,§,*</sup> and Vasiliki Kalatzis<sup>1,§,*</sup>

<sup>1</sup>Institute for Neurosciences of Montpellier (INM), University of Montpellier, Inserm, Montpellier, France

<sup>2</sup>National Reference Centre for Inherited Sensory Diseases, University of Montpellier, CHU, Montpellier, France

<sup>3</sup>Université Paris-Saclay, CNRS, Institut des Neurosciences Paris-Saclay, Saclay, France

<sup>4</sup>Institute of Molecular and Clinical Ophthalmology Basel (IOB), Basel, Switzerland

<sup>5</sup>Department of Ophthalmology, University of Basel, Basel, Switzerland

<sup>6</sup>Department of Genetics and Genome Biology, University of Leicester, Leicester, UK

<sup>7</sup>Department of Genetics, Health Research Institute-Fundación Jiménez Díaz University Hospital, Universidad Autónoma de Madrid (IIS-FJD, UAM), Madrid, Spain

<sup>8</sup>Center for Biomedical Network Research on Rare Diseases (CIBERER), Instituto de Salud Carlos III, Madrid, Spain

<sup>9</sup>Molecular, Cellular and Genomics Biomedicine Research Group, Instituto de Investigación Sanitaria La Fe (IIS La Fe), Valencia, Spain

<sup>10</sup>Joint Unit of Rare Diseases, IIS La Fe-Centro de Investigación Príncipe Felipe, Valencia, Spain

<sup>11</sup>Department of Neuroradiology, University of Montpellier, CHU, Montpellier, France

<sup>12</sup>Institute for Human Functional Imaging (I2FH), University of Montpellier, CHU, Montpellier, France

<sup>13</sup>Department of Medicine and Surgery, University Hospital of Parma, Parma, Italy

<sup>14</sup>Department of Medical Genetics, University Hospital of Parma, Parma, Italy

<sup>15</sup>Department of Nephrology, University Hospital of Parma, Parma, Italy

<sup>16</sup>Neuroradiology Unit, Diagnostic Department, University Hospital of Parma, Parma, Italy

<sup>#</sup>These authors contributed equally to this work

<sup>§</sup>These authors contributed equally to this work

<sup>*</sup>Authors for correspondence:

Beatrice Bocquet
35 CRM R Maolya

36 Hôpital Gui de Chauliac

37 80 Ave Augustin Fliche

38 34295 Montpellier

39 France

40 Phone: +33 4 67 33 69 74

41 Email: beatrice.bocquet@inserm.fr

42

43 Muriel Perron

44 Paris-Saclay Institute of Neuroscience

45 Campus CEA Saclay

46 Bâtiment 151

47 91400 Saclay

48 France

49 Phone: +33 1 69 82 60 92

50 Email: muriel.perron@universite-paris-saclay.fr

51

52 Vasiliki Kalatzis

53 INM, Inserm U1298

54 Hôpital St Eloi, BP 74103

55 80 Ave Augustin Fliche

56 34091 Montpellier

57 France

58 Phone: +33 4 99 63 60 76

59 Email: vasiliki.kalatzis@inserm.fr

60

61
62 **Abstract (<200 words)**

63 Retinitis pigmentosa (RP) is the most common inherited retinal disease (IRD) and is characterized
64 by photoreceptor degeneration and progressive loss of vision. We report here four patients who presented
65 with RP from three unrelated families with variants in *TBC1D32*, which to date has never been associated
66 with an IRD. To validate *TBC1D32* as a putative RP causative gene, we combined *Xenopus* in vivo
67 approaches and human iPSC-derived retinal models. Our data showed that *TBC1D32* was expressed during
68 retinal development and that it played an important role in retinal pigment epithelium (RPE) differentiation.
69 Furthermore, we identified a role for TBC1D32 in ciliogenesis of the RPE. We demonstrated elongated
70 ciliary defects that resulted in disrupted apical tight junctions, loss of functionality (delayed retinoid cycling
71 and altered secretion balance), and the onset of an epithelial-mesenchymal transition-like phenotype. Lastly,
72 our results also suggested photoreceptor differentiation defects, including connecting cilium anomalies, that
73 resulted in impaired trafficking to the outer segment in both cones and rods in *TBC1D32* iPSC-derived
74 retinal organoids. Overall, our data not only highlight a critical role for *TBC1D32* in the retina but
75 demonstrate that *TBC1D32* mutations lead to RP. We thus identify *TBC1D32* as an IRD causative gene.

76

77 **Keywords**

78 *TBC1D32/Bromi*; retinitis pigmentosa; RPE; primary cilium; epithelial-mesenchymal transition (EMT);

79 *Xenopus*; induced-pluripotent stem cells (iPSCs); retinal organoids

80

81
## 82 Introduction

Inherited retinal diseases (IRDs) are a group of clinically and genetically heterogeneous disorders characterized by a dysfunction or degeneration of the light-sensing cells of the retina, the photoreceptors, and/or their underlying support tissue, the retinal pigment epithelium (RPE) (1). In the majority of cases, IRDs are associated with a progressive loss of vision that can have a variable age of onset and severity of symptoms. Although each disease is monogenic, there are >270 known causative genes (https://sph.uth.edu/retnet). The most common IRD is a rod-cone dystrophy known as retinitis pigmentosa (RP) (2), characterized by night blindness appearing in childhood and followed by a progressive loss of peripheral vision. There are >90 causative RP genes with a collective prevalence of 1/4000 (3) and with autosomal dominant, autosomal recessive or X-linked modes of transmission (4). Furthermore, mutations in the same gene can give rise to either isolated RP or RP associated with extra-retinal symptoms.

The clinical heterogeneity associated with RP is partly explained by the diverse localizations and roles of the proteins encoded by the causative genes within the retina (5). The photoreceptors have a unique morphology comprising an axon, nucleus-containing cell body, a mitochondria-rich inner segment (IS) and an outer segment (OS) filled with lipid disks where the phototransduction process is initiated following light interaction. The IS is connected to the OS by a highly modified primary cilium called the connecting cilium (CC). The genes responsible for isolated RP can be divided into groups that play key roles in the cell body (splicing, transcription, development), IS (transport, signaling) or OS (structure, phototransduction) (3, 5). The genes for syndromic RP are mostly localized at the level of the CC or the periciliary membrane complex (6). These diseases are also known as ciliopathies and, although they can give rise to isolated RP, retinal defects are often linked to defects in other ciliated tissues (7). Interestingly, the RPE also contains a primary cilium on its apical side, which is critical for RPE maturation and homeostasis maintenance (8), and RPE ciliary defects were recently associated with retinal ciliopathies (9).

Currently, the genetic causes in ~40% of RP patients remain unknown (4). In order to resolve such cases, we performed whole exome sequencing (WES) in the probands from three unrelated families presenting clinically with autosomal recessive RP and identified previously undescribed biallelic variants in *TBC1D32* (OMIM: 615867). *TBC1D32* belongs to a family of Rab GTPase-activating proteins (GAPs) containing a Tre-2, Bub-2, and Cdc16 (TBC) domain. *TBC1D32* has been shown to have a role in regulating the structure of the primary cilium in the neural tube (10). Zebrafish *tbc1d32* morphants showed a curvature of the body axis and hydrocephalus and a *Tbc1d32* knock-out mouse model exhibited neural patterning defects comprising, exencephaly, preaxial polydactyly as well as poorly developed eyes. Accordingly, recessive loss-of-function *TBC1D32* variants were reported as responsible for Oral-Facial-Digital (OFD) syndrome type IX (OMIM: 258865), a ciliopathy characterized by defects in the development of the oral cavity, face and digits, associated with microphthalmia/anophthalmia (11). However, OFD syndrome is not
classically associated with retinal anomalies, and the *TBC1D32* animal models do not have reported retinal defects.

Therefore, to validate *TBC1D32* as a putative RP causative gene, we combined animal and human disease modeling by (i) performing hypomorphic knockdowns in an in vivo *Xenopus* model and (ii) generating iPSC-derived RPE and retinal organoids from an RP patient carrying *TBC1D32* variants. Together, our data show that *TBC1D32* is expressed during retinal development and plays an important role in ciliogenesis. The disruption of *TBC1D32* results in abnormal RPE maturation and function, as well as defective photoreceptor differentiation and OS trafficking. Overall, we identify *TBC1D32* as an IRD causative gene, which has implications for alleviating current diagnostic deadlocks.

125
# Results

## Three unrelated families present with clinical features of RP

Four patients from 3 unrelated families (Patient 1, Family 1; Patient 2, Family 2; Patients 3 and 4, Family 3) were studied (Supplemental Figure 1) and their ocular data are summarized in Supplemental Table 1. Night blindness occurred in childhood (Patients 1 and 2) or in early adulthood (Patients 3 and 4). Refraction errors were highly variable, the two patients older than 50 years were legally blind. Peripheral visual field impairment occurred early in the second decade with a severe impairment after the age of 50 years. Beyond the classic signs of RP noted on multimodal imaging and on spectral-domain optical coherence tomography (SD-OCT), all patients shared abnormal visibility of large choroidal vessels in the entire peripheral retina suggesting RPE loss with photoreceptor degeneration (Figure 1). None of the patients had documented macular edema. On full-field electroretinogram recordings, none of the patients had discernable rod and cone responses (data not shown). The patients did not mention any systemic symptoms, and consistently a systemic check-up only revealed mild MRI anomalies with an atrophic aspect of the superior vermis in patients 1 and 2 (Supplemental Figure 2). In addition, Patient 2 was diagnosed at 27 years of age with non-proteinuric chronic kidney disease. Urinary protein-to-creatinine and albumin-to-creatinine ratios were 0.19 and 0.10 g/g, respectively, with no notable urinary sediment. Autoimmunity markers were negative and C3 and C4 complement fractions, and IgG, IgA and IgM plasmatic levels, were normal. Kidneys were bilaterally reduced (95 mm) and showed mild cortical hyperechogenicity with a few bilateral cortical and parapelvic cysts (max 9 mm). Negative genetic testing for *PKD1* and *PKD2* genes further ruled out polycystic kidney disease.

## *TBC1D32* pathogenic variants segregate with the RP in three families

Trio WES was performed on the Patient 1 and his healthy parents (Supplemental Figure 1A). No mutation in genes previously associated with syndromic and non-syndromic RP (https://sph.uth.edu/retnet/) was detected. We identified two variants in the *TBC1D32* gene (NM_001367760.1) in trans in the proband, one of which was carried by each parent. The maternal intron 2 variant c.317+5G>A was predicted to probably alter splicing according to Human Splicing Finder and described at a frequency of 0.000004092 in the gnomAD database (https://gnomad.broadinstitute.org/). To ascertain pathogenicity, we performed RT-PCR amplification of exons 1 to 5 on whole blood cells (WBCs) and detected two bands in the proband and maternal samples, the smaller of which corresponded to in-frame skipping of exon 2 (Supplemental Figure 1D). The paternal variant, c.846delTCCTA; p.(Asn282Lysfs*7) (exon 7), was absent from the gnomAD database, and was predicted to induce a frameshift and the introduction of a premature termination codon (PTC).
WES performed for Patient 2 (Supplemental Figure 1B) showed that he was compound heterozygous for the variants c.18_27del; p.(Ser6Argfs*8) in exon 1 (maternal allele) and c.1141-1G>A in intron 10 (paternal allele). Both variants were absent from databases. The second variant was predicted to strongly alter splicing according to an in silico prediction software (https://mobidetails.iurc.montp.inserm.fr/MD). To confirm its pathogenicity, we performed RT-PCR amplification of exons 9 to 13 on WBCs of the patient and his father, and detected as many as seven additional splicing events that bore PTCs and were not present in the control (Supplemental Figure 1E).

Trio WES was performed on Patients 3 and 4, and a healthy sister (Supplemental Figure 1C). The affected siblings were compound heterozygous for the variants c.1267G>T; p.(Glu423*) in exon 12, and c.3513G>T; p.(Trp1171Cys) in exon 31, whereas the unaffected sister was heterozygous for the c.3513G>T variant. The frequency in gnomAD of the c.1267G>T and c.3513G>T variants were 0.0000124 (described three times in the database) and 0.00000402 (described once), respectively. The missense variant is situated in the carboxy-terminal Rab GAP TBC domain, in a highly conserved position relative to the orthologous proteins (Supplemental Figure 1F). The in silico pathogenicity scores were ambiguous (https://varsome.com/). Structural analysis using Alphafold showed that the Tryptophan at position 1171 was located at the beginning of an alpha helix, and its substitution by a Cysteine residue was likely to create a steric hindrance with a Tyrosine at position 1019. However, with a CADD pathogenicity score of 31 (https://cadd.gs.washington.edu/), this variant was classed as of unknown significance according to the ACMG classification. Taken together, the probands of all three families carried biallelic *TBC1D32* variants, of which only one allele could be predicted as loss of function.

### *tbc1d32* is expressed in the neuroretina and RPE during *Xenopus* development

As *TBC1D32* variants were never previously linked to RP, we used a *Xenopus laevis* model, which is highly suited to functional development and gene knockdown studies (12), to test the expression and role of *tbc1d32* in the retina. *Xenopus tbc1d32* is located on chromosome 5L, displays an exon/intron structure identical to the human homologue (33 exons), and is predicted to encode a 1297 aa protein with ~60% identity to human TBC1D32 (accession # NM_001367759.1 or NM_001367760.1 or NM_152730.6). We cloned the *Xenopus tbc1d32* full-length coding sequence (Supplemental Figure 3) and analyzed its expression during development (Figure 2A). *tbc1d32* transcripts were first detected at stage 25 (13) in multiciliated epithelial cells in the epidermis and in the optic vesicle. From stage 27-28, expression was also detected in the otic vesicle, the brain, and the pronephric nephrostomes. We further analyzed the expression of *tbc1d32*, and *indian hedgehog* (*ihh*) as an RPE marker (14), on retinal sections using an RNAscope fluorescent multiplex assay (Figure 2B). At stage 25, *tbc1d32* was expressed in all progenitor cells of the optic vesicle. At stage 27-28, transcripts were abundantly detected in the presumptive neural
retina, but only a few molecules were detected in the presumptive RPE, labelled with *ihh*. At stages 32-33, *tbc1d32* transcripts were clearly visible in the developing RPE, with a peak at stage 35-36. From stage 37-38 onwards, *tbc1d32* expression decreased in the RPE, the inner nuclear, and ganglion cell layers but remained well expressed in the outer nuclear layer (ONL) containing the photoreceptors, where it decreased at later stages. Finally, we confirmed the presence of *tbc1d32* transcripts in RPE cells by qPCR on mRNAs from dissected RPE at stage 35-36 (Figure 2C). In conclusion, in the retina, *tbc1d32* is expressed in retinal progenitors, developing photoreceptors and RPE cells, consistent with a putative role in retinogenesis.

## Knockdown of *Xenopus tbc1d32* leads to developmental RPE and photoreceptor defects

To determine the impact of *tbc1d32* knockdown in retinal development, we designed two different morpholinos (Mo1 and Mo2) that we injected at the 4-cell stage in one blastomere giving rise to neural ectoderm (Figure 3, A and B, and Supplemental Figures 3 and 4). This led to a unilateral *tbc1d32* knockdown mainly limited to neural tissues (Supplemental Figure 4). A high dose (20 ng) of either Mo1 or Mo2 led to severe developmental defects, including curvature of the body axis and microphthalmia/anophthalmia of the injected embryos (data not shown). No discernable eye malformations were observed upon injection of a control Mo.

As we suspected hypomorphic pathogenicity in *TBC1D32* patients, we tested lower doses of *tbc1d32* Mo. At a dose of 10 or 15 ng of Mo1, *Xenopus* embryos no longer displayed a microphthalmic phenotype, and we did not detect retinal toxicity using a cell death assay (Supplemental Figure 5). By contrast, morphant embryos exhibited retinal pigmentation defects, comprising a decreased intensity and discontinuity in pigmentation on retinal sections compared to controls (Figure 3C). This phenotype was similar upon Mo2 injection (Supplemental Figure 6A) and was partially restored upon co-injection with Mo-resistant *tbc1d32* mRNAs (i.e. not recognized by the Mo), demonstrating its specificity (Figure 3C). We further analyzed actin cytoskeleton organization in RPE cells by phalloidin staining of dissected eyes injected with Mo1 or Mo2 (Figure 3D and Supplemental Figure 6B). In controls, phalloidin staining was uniform, outlining cell membranes and highlighting the typical hexagonal shape of RPE cells, whereas the staining was heterogeneous and irregular in *tbc1d32* morphants, highlighting an altered shape of RPE cells. We hence investigated the impact of *tbc1d32* knockdown on RPE differentiation using *mitf* and *ihh* as RPE markers (14). Using in situ hybridization, we found that their expression was significantly decreased in the retinas of embryos injected with Mo1 or Mo2 compared to controls (Figure 4, A-D and Supplemental Figure 7A and B). This decrease was confirmed by qPCR (Supplemental Figure 8). Importantly, this effect was partially rescued upon co-injection with Mo-resistant *tbc1d32* mRNAs (Supplemental Figure 9). Collectively, these data suggest a key role for *tbc1d32* in RPE differentiation.
We also explored whether photoreceptor development was affected upon *tbc1d32* knockdown. By immunofluorescence (IF) at stage 41, when photoreceptors are well-differentiated, we found that the vast majority of morphant retinas injected with 10 ng of Mo1 exhibited normal labeling in the ONL of both Rhodopsin and S/M cone Opsin (Figure 4D). By contrast, at a higher dose of 15 ng, around half of the morphant embryos showed reduced staining. To determine whether these defects arise early during photoreceptor differentiation, we assessed *rhodopsin* expression at stage 35-36, when *rhodopsin* RNA begins to be detected. We found significantly reduced staining in *tbc1d32* morphant retinas compared to controls (Figure 4E), which was confirmed following Mo2 injections (Supplemental Figure 7C). Taken together, *tbc1d32* knockdown results in RPE defects, while photoreceptors are affected in a dose-dependent manner, being either largely preserved or exhibiting obvious differentiation defects.

### Xenopus RPE ciliogenesis is affected upon *tbc1d32* knockdown

Defects in primary cilia morphology, namely curled cilia, were reported in both neural progenitor cells of *Tbc1d32*⁻/⁻ mouse embryos and in renal tubules of zebrafish *tbc1d32* morphants (10). Consistently, by staining with the widely used cilium markers Arl13b (ciliary shaft marker) and γ-Tubulin (basal body marker), we detected ciliogenesis defects in neural tube cells of *Xenopus tbc1d32* morphants, which displayed longer primary cilia than controls (Figure 5, A and B). We thus assayed if the RPE developmental defects in the *tbc1d32* morphants were also associated with ciliogenesis defects. As primary cilia are necessary for the maturation of mouse RPE but then disappear in fully differentiated cells (15), we first characterized the developmental window during *Xenopus* retinogenesis when the RPE cilia were present. We used both the RPE marker Otx2, to delineate cells, and acetylated α-Tubulin and γ-Tub, to label primary cilia. First, we verified that acetylated α-Tubulin co-localized with Arl13b along the cilium and with γ-Tub at the basal body level, confirming that it is a reliable marker of the *Xenopus* RPE cilium (Figure 5C). We detected ciliated RPE cells in the presumptive retina from stage 27-28. The percentage of ciliated cells increased and peaked at stage 35, and then decreased from stage 37-38 onwards (Figure 5, D and E). In *tbc1d32* morphant retinas of stage 35 embryos, the mean cilia length was significantly higher than in control retinas (Figure 5, F and G; Supplemental Figure 10, A and B). We then analyzed photoreceptor ciliogenesis and found a significant decrease in the number of photoreceptors showing Arl13b staining in *tbc1d32* morphant retinas compared to controls (Figure 5, H and I and Supplemental Figure 10, C and D). In conclusion, *tbc1d32* knockdown impacted ciliogenesis in both *Xenopus* RPE and photoreceptor cells.

### Human *TBC1D32* fibroblasts display a primary cilium defect

Human fibroblasts have long constituted a classical screening assay for ciliary defects (16). Therefore, we cultured fibroblasts from a skin biopsy of Patient 1 to better study the expression of both
mutant *TBC1D32* alleles and their impact on ciliogenesis. Using primers specific to exons 1 and 8 to assay the c.317+5G>A allele, we detected a 931-bp band in control cells and three bands in the patient cells (Supplemental Figure 11A): the first was similar in size to the control band but contained the last 18 bp of intron 2 (Supplemental Figure 11B) that introduced a PTC; the second and most predominant was 769 bp and corresponded to the in-frame skipping of exon 2 (Supplemental Figure 11C), as observed in patient WBCs; the third was 591 bp and corresponded to the skipping of exons 2 and 3 that also introduced a PTC (Supplemental Figure 11D). Thus, only transcript 2 was predicted to lead to translation, which corroborated our observations in WBCs, despite the less comprehensive analysis in this cell type. Using primers specific to exons 6 and 11 to assay the c.846delTCCTA allele, we detected an ~600-bp amplicon in control and patient cells (Supplemental Figure 11E), the latter of which contained a transcript with the 5-bp deletion in exon 7 (Supplemental Figure 11F) that introduced a PTC. These results suggested that the mutant transcript did not undergo nonsense-mediated decay, which was consistent with the qPCR results showing similar levels of *TBC1D32* expression in both control and patient fibroblasts (Supplemental Figure 11G).

We then assayed ciliogenesis by IF studies of ARL13B with the axoneme markers acetylated α-tubulin and polyglutamylated tubulin (GT335), and the basal body marker pericentrin (PCN), which is present in both the mother and daughter centrosomes (17). We detected a similar spatial distribution between control and TBC1D32 fibroblasts (Figure 6, A-F). By contrast, and consistent with the *Xenopus* data, we detected a significantly elongated ARL13B-labelled cilium in the patient cells compared to controls (Figure 6, A-G). We also assayed intraflagellar transport (IFT) by IF studies of IFT88 (18, 19), which correctly extended beyond acetylated α-tubulin at both ends of the cilium in control and patient cells (Figure 6, H and I). Lastly, consistent with the qPCR data, we clearly detected TBC1D32 using an antibody specific to an epitope encoded by exons 24 to 28 (Supplemental Figure 11G). TBC1D32 was located basal to acetylated α-tubulin in control and patient fibroblasts (Figure 6, J and K), where it overlapped with PCN (Figure 6, L-O). Taken together, TBC1D32 localized to the centrosomes of the primary cilium in human fibroblasts, and mutant TBC1D32 proteins led to elongation defects.

## TBC1D32 iPSC-derived RPE has an elongated cilium and disrupted tight junctions

To unravel the pathophysiology associated with *TBC1D32* mutations, we reprogrammed the fibroblasts of Patient 1 into induced pluripotent stem cells (iPSCs) to subsequently generate highly relevant human retinal models. Two *TBC1D32* iPSC lines were confirmed as pluripotent by the expression of the host pluripotency markers (Supplemental Figure 12, A and B) and germline markers following an embryoid body (EB) differentiation assay (Supplemental Figure 12C). In addition, we confirmed genomic stability by testing the copy number of the recombination hotspots in iPSCs (20) (Supplemental Figure 12D). Interestingly, in contrast to the fibroblast data, *TBC1D32* expression levels in the two patient iPSC lines
were lower than in control cells (Supplemental Figure 12E). Lastly, we confirmed the presence of the causative *TBC1D32* mutations c.317+5G>A in exon 2 and c.846delTCCTA in exon 7 in the patient iPSC lines by Sanger sequencing (Supplemental Figure 12F).

We then differentiated the patient and control lines into iPSC-derived RPE. Due to the different qPCR profiles observed between fibroblasts and iPSCs, we assayed *TBC1D32* expression in the iPSC-derived RPE and detected lower levels in the patient RPE compared to controls (Supplemental Figure 13A), further suggesting tissue-specific regulation. By IF analysis, TBC1D32 partially co-localized with PCN in control and patient iPSC-derived RPE, as in primary fibroblasts, but the staining intensity was lower in the patient RPE (Supplemental Figure 13, B and C), consistent with the qPCR results. We then assayed the morphology of the *TBC1D32* iPSC-derived RPE from 12 weeks post-seeding. Co-labelling of the apical tight junction marker ZO1 and ARL13B demonstrated a regular cobblestone morphology and primary cilium staining for the control RPE (Figure 7A). By contrast, the morphology of the *TBC1D32* patient iPSC-derived RPE was highly irregular with cilia significantly longer than controls (Figure 7, B and C). Furthermore, we detected gaps between cells, and ZO1 aggregates at the contact points of multiple cells, in the *TBC1D32* RPE that were absent from control RPE (Figure 7, D and E). The disrupted tight junctions were confirmed by trans-epithelial resistance (TER) measurements, which steadily decreased in the *TBC1D32* RPE from 6 weeks post-seeding to levels significantly lower than control RPE at 13 weeks (Figure 7F). In conclusion, the *TBC1D32* patient iPSC-derived RPE presented with a primary cilium defect and disrupted apical tight junctions.

## *TBC1D32* iPSC-derived RPE displays an EMT-like phenotype

Primary cilium defects in RPE reportedly provoke an EMT-like phenotype, characterized initially by a loss of tight junctions (21). We thus assayed the *TBC1D32* RPE for signs of cytoskeletal changes associated with EMT by testing the expression of vimentin and SMA (22) (Figure 8, A-D). A low and relatively regular intensity of vimentin expression was detected in the control iPSC-derived RPE, whereas islands of upregulated expression were detected in the patient RPE, which were associated with a loss of MERTK expression, further confirming the loss of RPE identity. Similarly, large and abundant SMA-positive islands were detected in the *TBC1D32* iPSC-derived RPE, compared to controls, which were associated with disrupted ZO1 expression (Supplemental Figure 13, D-F). Thus, the disrupted apical junctions in the *TBC1D32* iPSC-derived RPE were accompanied by a transition to a mesenchymal state.

These observations were further confirmed by gene expression levels of other EMT markers (Figure 8, E-I). The genes encoding the adherens junction markers E-cadherin (*CDH1*), P-cadherin (*CDH3*) and β-catenin (*CTNNB1*) showed reduced expression levels in the *TBC1D32* RPE, compared to controls, confirming the loss of epithelial cell-cell contacts. Furthermore, the transcription factor SNAIL (*SNAI1*),
328 which mediates EMT, and Fibronectin 1 (FN1), a marker of EMT, showed increased expression levels.
329 Consistently, western blot analysis of E- and P-cadherin showed significantly decreased levels in the patient
330 RPE compared to controls (Figure 8, J and K). In contrast to the qPCR data, β-catenin showed similar levels
331 in control and patient RPE (Figure 8L). We thus analyzed its subcellular distribution and showed that it was
332 predominately localized to the membrane fraction in control RPE and to the cytosolic fraction in the patient
333 RPE (Figure 8M). This difference was consistent with the release of β-catenin from the membrane, which
334 allows it to act as a transcriptional activator of EMT genes, such as SNAI1 (23). Taken together, the
335 TBC1D32 iPSC-derived RPE displays altered morphology and deregulated expression profiles that are
336 hallmarks of EMT activation.
337

338 **TBC1D32 iPSC-derived RPE shows altered functionality**
339

340 The loss of tight and adherens junctions reportedly affects the apical-basal polarity of RPE with an
341 impact on functionality (21). Hence, we assayed the expression of CRALBP, an RPE marker and actor of
342 the visual cycle, together with N-cadherin, an adherens junction and EMT marker (22) (Figure 9, A and B).
343 We detected a regular membrane staining of N-cadherin in control RPE compared to notable patches of
344 discontinuous N-cadherin staining in the patient RPE. Similarly, we detected reduced CRALBP staining at
345 the membrane and an increased signal in the cytosol in the patient RPE compared to controls. This shift
346 was confirmed by western blot analyses (Figure 9C). Furthermore, the most pronounced areas of CRALBP
347 staining showed reduced N-cadherin levels. Moreover, we detected a decrease in expression of the
348 corresponding gene, RLBP1, as well as other visual cycle genes, LRAT and RPE65 in TBC1D32 RPE
349 compared to controls (Supplemental Figure 13, G-I). ABCA4, which is involved in the visual cycle in
350 photoreceptors, was recently described to be also expressed in the RPE (24) and, accordingly, its expression
351 was also reduced in the patient RPE (Supplemental Figure 13J).

351 To determine whether the differential expression of these visual cycle actors in the patient RPE
352 impacted its function, we assayed for the accumulation of retinoids, detectable as lipid droplets termed
353 retinosomes (25) by transmission electron microscopy (TEM). We detected an accumulation of lipid
354 droplets in the TBCD1D32 RPE as compared to controls (Figure 9, D-F), which were identified as
355 retinosomes by the specific marker perilipin-2 (Figure 9G). The significant abundance of retinosomes
356 prompted us to assay the polarized secretion of the angiogenic factors pigment epithelium-derived factor
357 (PEDF) and VEGF, another key RPE function, by ELISA (26). We detected a significant increase in apical,
358 and a significant decrease in basal, PEDF and VEGF secretion in the TBC1D32 RPE, as compared to
359 controls (Figure 9H). Taken together, the cellular changes in the TBC1D32 RPE result in disrupted retinoid
360 cycling and altered secretion balance.
361
362 **TBC1D32** iPSC-derived retinal organoids show defective trafficking to outer segments
363
As photoreceptors possess a specialized cilium, the CC, we assayed whether *TBC1D32* defects also
364
led to photoreceptor anomalies in iPSC-derived retinal organoids (27). Bright-field imaging showed that
365
control and *TBC1D32* organoids had a similar morphology with a brush border (Figure 10, A and B)
366
containing IS and OS-like structures and a distinct lamina corresponding to the ONL (Figure 10, C and D).
367
This organization was confirmed by IF studies of CRX and recoverin (RCVRN), although staining was
368
visibly reduced in the patient organoids (Figure 10, E and F). Furthermore, we assayed the expression of
369
ZO1 and CRALBP, markers of the outer limiting membrane (OLM) formed by the end feet of Müller glial
370
cells. The OLM was clearly discernible by the co-localized CRALBP/ZO1 signals in the control organoids,
371
whereas we detected only a faint CRALBP signal and no ZO1 signal in the *TBC1D32* organoids (Figure
372
10, G and H). Moreover, co-staining of ZO1 and rhodopsin kinase (RK) showed that RK expression
373
extended beyond the OLM into the IS/OS layer in control organoids but not as far in the *TBC1D32*
374
organoids (Figure 10, I and J). We thus investigated the CC by assaying the expression of ARL13B in
375
parallel to rhodopsin (RHO). In control organoids, ARL13B was visible as punctate dots close to the ONL
376
with RHO extended beyond into the OS, whereas, in *TBC1D32* organoids, ARL13B was only faintly
377
discernible and RHO remained close to the ONL (Figure 10, K and L). The RK and RHO expression profiles
378
suggested that trafficking to the OS was impaired, thus we tested the expression of the cone and rod OS
379
marker ABCA4 and the rod-specific OS marker PDE6B. ABCA4 and PDE6B staining was clearly
380
detectable in the OS of the control organoids but visibly reduced in the *TBC1D32* organoids (Figure 10, M
381
and N). In addition, arrestin 3 (ARR3)-positive cones of varying lengths were identified in the control
382
organoids, a subset of which co-expressed the more mature R/G opsin cone marker (Figure 10O). By
383
comparison, the cones in the *TBC1D32* organoids appeared stunted with a distinctly visible co-localization
384
signal at the edge of the cells beyond the ONL (Figure 10P). In conclusion, in addition to affecting the RPE,
385
*TBC1D32* variants directly affect photoreceptors at the level of the CC and result in impaired trafficking to
386
the OS in both cones and rods.
387 **Discussion**

388 With the technological advancements of next-generation sequencing, the genetic diagnosis yield of 389 IRDs had considerably increased over the last ten years (28). However, recently, the slope of the 390 identification curve has reached a plateau (https://sph.uth.edu/retnet), raising challenges for genetic 391 counselling and therapeutic opportunities for patients. Furthermore, the genetic landscape for IRDs is 392 complexifying, as it becomes increasingly evident that mutations in the same gene can give rise to 393 differential phenotypes or even clinically distinct disorders (29). *TBC1D32* is now another perfect example 394 of the breakdown in borders between disorders. To date, *TBC1D32* variants were only associated with OFD 395 syndrome, a severe malformative disorder. However, here, we report pathogenic *TBC1D32* variants in four 396 patients from three independent families presenting clinically with RP. Furthermore, by combining an in 397 *Xenopus* model and human iPSC-derived retinal models, we describe a critical role for *TBC1D32* in 398 retinal differentiation and ciliogenesis, and demonstrate that reduced *TBC1D32* levels impact the RPE and 399 photoreceptors. We thus identify *TBC1D32* as a causative IRD gene.

400 OFD syndromes form a rare subgroup of ciliopathies characterized by oral cavity, facial and digital 401 anomalies with or without heart defects, polycystic kidney disease, or corpus callosum agenesis (30). There 402 is a high clinical heterogeneity linked to sixteen causative genes. The most frequent is the X-linked *OFD1*, 403 which encodes a centrosomal protein. Interestingly, rare *OFD1* patients have been described as presenting 404 with non-syndromic retinal degeneration (31), suggesting that variants in other *OFD* genes may also give 405 rise to IRDs. In 2014, loss-of-function *TBC1D32* variants were first described as associated with the OFD 406 syndrome type IX (11). Clinical symptoms ranged from midline cleft, anophthalmia, and polydactyly to 407 choanal atresia, agenesis of corpus callum, vermis and pituitary hypoplasia, and hydrocephalus. 408 Subsequently, seven patients from five families were reported as carrying different pathogenic *TBC1D32* 409 variants and presenting with syndromic ciliopathies (32). It is noteworthy that one patient from a Finnish 410 family with compound heterozygous indels in *TBC1D32* also exhibited a progressive retinal dystrophy but 411 this was not further explored (33).

412 Taking into account the clinical data in relation to age, none of the patients reported here showed 413 signs of *TBC1D32*-related OFD syndrome. It should be noted, though, that MRI revealed mild subclinical 414 alterations, which suggests that *TBC1D32* variants may give rise to a disease spectrum with a certain 415 threshold for the appearance of clinical signs. It is likely that type or severity of the associated variant 416 accounts for this spectrum. We now raise the number of identified pathogenic *TBC1D32* variants, identified 417 in eight unrelated families, to ten: one missense (c.3513G>T), two nonsense (c.1267G>T; c.3724C>T), 418 three splicing (c.317+5G>A; c.1141-1G>A; c.1372+1G>T) and four indels (c.18_27del; c.846delTCCT; 419 c.1165_1166dup; c.2151del) ((32-34) and this study). Although it is possible that nonsense and indel 420 *TBC1D32* variants result in loss of function, splicing and missense variants are more ambiguous.
The c.317+5G>A splicing variant, carried by Patient 1 in trans to a frameshift indel, only results in mild RP, which is likely explained by the resulting in-frame exon skipping that may result in a shorter partially functional protein. Similarly, the c.1141-1G>A variant, carried by Patient 2 in trans to a frameshift indel, which promotes the creation of multiple aberrant splicing variants also did not result in severe syndromic disease. However, this patient presented with additional renal defects, which were possibly of a ciliopathic nature. An association between altered ciliary function and the development of renal cysts has been linked to variants in the ciliary function-related genes *PKD1/PKD2* (35). Patient 2 was negative for pathogenic *PKD1/PKD2* variants, raising the possibility that the renal anomalies may be due to the *TBC1D32* variants. This is supported by the zebrafish *tbc1d32* morphant model that shows dose-dependent ciliary defects in the distal kidney tubules (10), and the association of renal anomalies with other ciliopathies (7). Along this line, a recent study described developmental cochlear defects in the knockout *Tbc1d32* mouse model (36), and we also detected *tbc1d32* in the *Xenopus* otic vesicle, suggesting that *TBC1D32* variants may also provoke ciliopathies with deafness associated. Finally, the mildest *TBC1D32*-associated phenotype reported to date is the later onset RP in Patients 3 and 4 carrying the missense c.3513G>T variant in trans to a nonsense variant. The predicted structural changes associated with this variant suggest the production of a mutant protein, which we hypothesise may be partially functional to account for the moderate phenotype. Taken together, and in consideration of the dose-dependent phenotypic severity in the *Xenopus tbc1d32* morphants, we propose that hypomorphic *TBC1D32* variants give rise to RP and loss-of-function variants to OFD syndrome.

Pertinently, none of the patients showed ocular malformations, whereas they all presented with features of retinal degeneration that were compatible with both RPE and photoreceptor defects. This clinical data is strongly supported by our functional data. Firstly, the phenotypes of the *Xenopus tbc1d32* morphants and the human *TBC1D32* iPSC-derived RPE models highlighted RPE differentiation defects that affected functionality. Most notably, the slowing down of retinoid recycling, as evidenced by retinosome accumulation, is reminiscent of the RPE defect associated with mutations in the visual cycle protein *RPE65*, which causes Leber congenital amaurosis (37). Visual cycle disruption reduces the availability of 11 *cis*-retinol, which subsequently impacts the phototransduction process causing visual impairment. Secondly, the *Xenopus* morphants and iPSC-derived retinal organoids showed photoreceptor differentiation anomalies and disturbed trafficking of photoreceptor proteins to the OS, which would affect their function. In addition, the disrupted apical VEGF secretion identified in the *TBC1D32* iPSC-derived RPE, would also impact neurodevelopment and photoreceptor protection throughout life (38), further impacting phototransduction and vision.

The disrupted ciliogenesis of the RPE and photoreceptors in both the *Xenopus* and human models leads us to propose that *TBC1D32*-associated RP is a retinal ciliopathy (7). In particular, we show that
TBC1D32 defects result in elongated primary cilia in fibroblasts, neural tube and RPE cells. Ciliary length is determined by an interplay between ciliary assembly and disassembly processes, which rely on an effective bidirectional IFT system (39). There have been examples in the past of gene variants associated with retinal degeneration that result in elongated cilia. For example, variants in the gene encoding IFT172, have been associated with non-syndromic or syndromic forms of RP (40, 41) and elongated cilia were detected on patient's fibroblasts (41). Similarly, variants in the gene encoding the transition zone protein CEP290 (42), associated with the early onset retinal dystrophy Leber congenital amaurosis (LCA), resulted in elongated primary cilia on patient fibroblasts (43) Furthermore, these fibroblasts showed normal IFT88 distribution, similar to our observations on *TBC1D32* cells, indicating that anterograde IFT was normal. Finally, variants in the gene encoding NPHP5/IQCB1 also associated with LCA were recently shown to provoke elongated primary cilia in patient-derived fibroblasts and iPSC-derived RPE (44), further confirming that elongated ciliary defects give rise to retinal ciliopathies.

Importantly, we show that the tight junction defects of the *TBC1D32* RPE were associated with an EMT-like phenotype both at the mRNA and protein levels. We did observe a slight discrepancy for β-catenin, as the decreased mRNA levels did not corroborate with the unchanged protein levels. The regulation of β-catenin in EMT, however, is highly complex due to the protein's dual roles in adhesion and transcription and the interplay between these pools has not been elucidated (45). Nonetheless, we showed that there was a release of β-catenin into the cytosol, which would increase the transcriptionally active pool consistent with EMT (23). We can thus only speculate that, at the time of analysis, the RPE was in a dynamic EMT state and that feedback regulations on mRNA or protein levels were in process. TBC1D32 is an actor of the Hedgehog (HH) pathway in primary cilia, which acts to regulate the GLI family of transcription factors (46). TBC1D32 controls the HH pathway by allowing GLI proteins to be properly localized in the primary cilia and correctly activated in response to high-level HH signaling (10). In cancer cells, it has been shown that activated GLI proteins bind and transcribe genes that control EMT (namely *SNAIL*) among other cell processes (47). Therefore, our data suggest that ciliary elongation in the RPE due to TBC1D32 disruption may cause HH signaling perturbations, which would activate EMT causing a loss of identity and function. Although beyond the scope of this manuscript, future studies should be aimed at precisely deciphering the link between TBC1D32, HH signaling, and EMT in the retina. Lastly, a similar link between primary ciliary defects and EMT in the RPE was also made in a murine model of the syndromic retinal ciliopathy Bardet-Biedl syndrome (*BBS8*^−/−), where an EMT-like phenotype was associated with defective cellular polarization and morphology (21), further validating our observations.

In general, the proteins encoded by OFD genes have been attributed to three functional modules in primary cilia: centriole elongation, transition zone, and intraflagellar transport (30). TBC1D32 was attributed to the elongation module as zebrafish *tbc1d32* interacts with cell cycle-related kinase to regulate
ciliary membrane and axonemal growth in the neural tube (10) and murine Tbc1d32 acts redundantly with the centrosomal protein Dzip11 to regulate ciliogenesis (48). We now demonstrate that TBC1D32 also regulates ciliogenesis in the retina in human and *Xenopus* models. Consistent with the fact that TBC1D32 is not associated with the intraflagellar transport module, we did not detect abnormal IFT88 distribution in *TBC1D32* fibroblasts. Interestingly though, IFT88 pooling was previously reported in the fibroblasts of the *TBC1D32*-associated OFD type IX patient (19) and in a *TBC1D32* knockout hTERT-RPE1 cell line (49). Furthermore, the murine *tbc1d32*−/− and zebrafish morphant models displayed curled axonemes enveloped by dilated ciliary membranes (10), in contrast to the elongated ciliary defect that we report here in the *Xenopus* knockdown model and in the *TBC1D32* patient cells. These differences could be due to a dose-dependent effect related to the amount of remaining functional TBC1D32 protein. Thus, taken together, we propose that the severity of TBC1D32 disruption will lead to phenotypic variability at the level of the primary cilium and an ensuing disease spectrum.

In conclusion, we identify *TBC1D32* as an IRD gene responsible for RP and broaden the clinical spectrum associated with *TBC1D32* variants. As we identified pathogenic variants in multiple unrelated families and, as one of the probands also presented with renal anomalies, we highly recommend that *TBC1D32* be added to the list of candidate genes to be screened for both isolated and syndromic RP.
## Methods

### Clinical investigations

Patients underwent a comprehensive ophthalmological examination (best-corrected visual acuity, kinetic visual field, SD-OCT, short-wavelength fundus autofluorescence (SW-FAF), color fundus photography and full-field electroretinography performed in accordance with the guidelines of the International Society for Electrophysiology of Vision). Age of onset, initial symptoms and extra-ocular manifestations were also reviewed. The systemic check-up included a cerebral MRI, hypothalamic-pituitary hormone dosages, skeletal investigations, ear, nose, throat heart and kidney evaluations.

### Whole exome sequencing and bioinformatic analyses

Targeted exome sequencing library preparation, exome capture, sequencing and data analysis for Patient 1 and his unaffected parents were performed by IntegraGen SA (Evry, France) using Sureselect All Exon V5 (Agilent Technologies) on an Illumina HiSeq4000, as previously described (50). WES for Patient 2 was performed using the Twist Comprehensive Exome Panel (Twist bioscience) and sequenced on a HiSeq 4000 instrument (Illumina) with an average coverage of 100–120X at each nucleotide position. WES for Patients 3 and 4 and a healthy sibling was conducted at CNAG-CRG (Centro Nacional de Análisis Genómico, Barcelona, Spain) using Sureselect All Exon V5 (Agilent Technologies) and captured libraries were paired-end (2x100 bp) sequenced on Illumina HiSeq2000 platform. For Patients 2 to 4, BWA (V0.7.17) was used for mapping raw reads to the human genome reference sequence (build hg19) and the bioinformatic analysis was performed as described (51). For Patients 3 and 4, the raw genomic data was processed in parallel through the RD-Connect analysis pipeline in the centralized RD-Connect database (52).

### Xenopus embryo collection, RNA extraction and cloning

*Xenopus laevis* embryos were obtained from TEFOR (Paris-Saclay) by conventional methods of hormone-induced egg laying and in vitro fertilization, staged according to Nieuwkoop and Faber's table of development (13). Total RNA from wild-type *Xenopus laevis* embryos was isolated using the Nucleospin RNA XS kit (Macherey Nagel). Following reverse transcription, PCR amplification was performed using specific primers (Supplemental Table 2) designed from the *tbc1d32* sequence in the National Center for Biology Information (NCBI) *Xenopus laevis* EST database (accession # XM_041562734.1). The amplified *tbc1d32* sequence was then cloned using the ligation-free cloning system in the XhoI-linearized pCS2+ plasmid (E001, ABM) and verified by sequencing (Supplemental Figure 3).

### Xenopus whole-mount in situ hybridization
Digoxigenin-labeled antisense RNA probes were generated according to the manufacturer's instruction (Roche) from gifted plasmids: pBS-*ihh* (Stephen Ekker, Mayo Clinic, Minnesota, USA); pGEM-Teasy-*mitf* (Andrea Viczian, Upstate Medical University, New York, USA); pGEM-T-*rhodopsin* (Thomas Hollemann, Martin-Luther-University Halle-Wittenberg, Halle, Germany). The *tbc1d32* probe was generated from the entire *tbc1d32* coding region cloned into pCS2+. Whole-mount *in situ* hybridizations were carried out as described (14) and analyzed on 50 µm vibratome transverse sections.

### RNAscope *in situ* hybridization

*Xenopus* embryos were fixed in 4% PFA at 4°C for 24 h, embedded in OCT and sectioned at 10 µm thickness on a cryostat (Leica). The RNAscope Multiplex Fluorescent 2.0 Assay was performed according to the manufacturer's protocols using the HybEZ oven. Opal 520 and 570 dyes were applied to slides along with DAPI counterstain.

### Morpholinos and Microinjection

Translation-blocking antisense Morpholino oligonucleotides (GeneTools) are listed in Supplemental Table 2. Five nL microinjections were performed at the four-cell stage into one of the two blastomeres giving rise to the neural ectoderm. 500 pg of *tbc1d32* mRNA (synthesized with mMessage mMachine kit, Life Technologies) and/or 10 to 20 ng of *tbc1d32* Mo1 or Mo2 were injected; a standard Mo (GeneTools) was injected as a control. For each condition, mRNA encoding GFP was co-injected as a lineage tracer. The efficacy of the two *tbc1d32* Mo was tested by analysing in vivo GFP fluorescence following co-injection of a chimeric GFP construct fused downstream of the Mo-complementary sequence (Supplemental Figure 4).

### Fibroblasts and iPSCs

Control fibroblast and iPSCs lines were previously reported (53). The skin biopsy of Patient 1 was performed at the National Reference Center for Inherited Sensory Diseases (Montpellier, France). Dermal fibroblasts were reprogrammed using the CytoTune-iPS 2.0 Sendai Reprogramming Kit (Thermo Fisher Scientific), and iPSC colonies cultured in Essential 8 Medium (Gibco) (54). Trilineage differentiation potential was determined by an EB differentiation assay as described (54). Genomic stability of iPSCs was analyzed using droplet digital qPCR analysis by Stem Genomics (Montpellier, France). iPSC genomic DNA was isolated, PCR amplified using *TBC1D32*-specific primers (Supplemental Table 3), and the amplicons sequenced on an Applied Biosystems 3130xL or SeqStudio genetic analyzer (54).

### iPSC-derived RPE and retinal organoids
iPSC-derived RPE was differentiated (53) and cryostored at passage 2 (26) as described. All experiments were performed on passage 3 RPE seeded on Matrigel-coated translucent Corning cell culture inserts with high-density 0.4 µM pores (BD Falcon). The TER of 4 independent inserts at each time point was measured using the Epithelial Volt/Ohm Meter EVOM2 (Word Precision Instruments) and expressed as an average value in Ω/cm² (53). The TEM analysis of iPSC-derived RPE grown on transwell inserts was performed as described (53). Retinal organoids were differentiated from adherent iPSCs and cultured as free-floating structures in DMEM/F12+GlutaMAX (Gibco) supplemented with taurine and retinoic acid, as described (27).

## RT-PCR and qPCR

Total *Xenopus* RNA from at least 12 optic vesicles at stage 30, 12 whole retinas, 40 dissected RPE, or 40 neural retinas at stage 35-36, was extracted using trizol and Nucleospin RNA Kit (Macherey Nagel). cDNA was synthesized from 290 ng of RNA with Superscript IV (Invitrogen). For qPCR experiments, 10 ng of cDNA was amplified in triplicate using SsoFast EvaGreen Supermix (BioRad) on a C1000 thermal cycler (CFX96 real-time system, BioRad). Quantification was performed using the ΔΔCt method, and levels were normalized to *odc*, *ef1a*, and *rpl8* and expressed relative to control Mo or neural retina. Total human RNA was extracted from WBCs, fibroblasts, iPSCs, and iPSC-derived RPE using the PAXgene RNA kit (BD Biosciences), Tempus Spin RNA Isolation kit (Applied Biosystems), or RNeasy Mini Kit (Qiagen). cDNA was synthesized from 500 ng of RNA with the SuperScript III First-Strand Synthesis System using random hexamers (Life Technologies) or from 2 µg of RNA with the High Capacity cDNA Reverse Transcription Kit (Applied Biosystems). RT-PCR amplification of Patient 1's WBCs and fibroblasts was performed using AmpliTaq Gold 360 Master Mix (Applied Biosystems) and Patient 2's WBCs using GoTaq G2 DNA Polymerase (Promega). Amplicons were either gel purified and directly sequenced (WBC Patient 1), or subcloned into pGEMTeasy vectors and then sequenced. For qPCR experiments, a 1/10 (iPSCs) or a 1/20 dilution (RPE) of the cDNA was amplified in triplicate using the FastStart SYBR Green Master mix on a LightCycler 480 II thermal cycler (Roche). Quantification was performed using the ΔΔCt method normalized to *GAPDH*, and levels expressed relative to control. Experiments were duplicated using a second housekeeping gene. Primer sequences are presented in Supplemental Table 4.

## Immunofluorescence studies

*Xenopus* embryos were fixed in 4% paraformaldehyde (PFA) and immunolabelling was performed on paraffin sections with antibodies listed in Supplemental Table 5 using standard procedures. Antigen retrieval was performed by boiling the sections in 10 mM sodium citrate and 0.05% Tween-20 for 9 min. For cilia analysis, sections were bleached in 10% H₂O₂ in PBS at 55°C. Nuclei were stained with Hoechst
(Sigma-Aldrich). Sections were imaged with a Zeiss M2 microscope, or a Zeiss LSM 710 confocal microscope for cilia analysis, and processed with Zen software (Zeiss). For phalloidin staining, dissected eyes were fixed in 4% PFA and labelled with Alexa Fluor 568-phalloidin (1/40, Molecular Probes). IF studies of human fibroblasts, iPSCs, EBs, RPE and retinal organoids were performed as previously described (53-55). Primary antibodies were incubated at 4°C overnight and secondary antibodies with 0.2 µg/ml Hoechst 33258 (Sigma-Aldrich) were incubated at room temperature for 1 h (Supplemental Table 6). Samples were imaged using a Zeiss ApoTome 2 Upright wide-field microscope or Confocal LSM700 microscope.

## Image quantification

All *Xenopus* experiments were performed at least in duplicate and results from one representative experiment are shown. To quantify activated caspase3⁺ labeled cells, 6 to 9 sections per retina were analyzed using Adobe Photoshop CS4 software. *In situ* hybridization staining integrated density in eyes or in retinal sections was measured in the delineated region of interest using Fiji software (https://fiji.sc/). Fiji software was also used to quantify *tbc1d32* expression upon RNAscope *in situ* hybridization. To image and measure cilia lengths, Z-stacks of 11 µm retina sections were taken with 0.3 µm steps and converted into single planes by maximum projection with Fiji software. For each condition, the length of 3 to 8 cilia per retina was quantified on image stacks using the ObjectJ plugin (https://sils.fnwi.uva.nl/bcb/objectj/). The quantification of the cilium lengths in human fibroblasts and iPSC-derived RPE were performed using Image J and Imaris Software (Bitplane), respectively. For the RPE, five random regions of a transwell filter were measured. The quantification of the SMA area was performed on a montage of 16 regions of a transwell filter using the Imaris Software. The quantification of the lipid vesicles in the TEM images was performed manually.

## Western blot analyses

Unless otherwise stated, hiPSC-derived RPE cells were seeded on inserts, scraped 12 weeks post-seeding, resuspended in Laemmli sample buffer containing 2-mercaptoethanol, and western blot analysis performed as previously described (53). The Mem-PERTM Plus Kit (Thermo Scientific) was used to separate the cytosolic and membrane fractions prior to mixing with Laemmli buffer. Primary and secondary antibodies (Supplemental Table 6) were incubated overnight at 4°C or 45 min at room temperature, respectively. Fluorescence was detected using the ODYSSEY CLx imaging system (LI-COR) and quantified with the Image Studio Lite software.

## Secretion assays
VEGF and PEDF secretion of human iPSC-derived RPE was assayed by ELISA (R&D Systems) on 641 642 24 h conditioned media collected from the apical and basolateral chambers of the cultured inserts as 643 described (26). Samples were collected from 16 to 19 inserts and assayed in triplicate. The mean 644 concentration values of the *TBC1D32* samples were expressed as a percentage of the control values. 645

Statistical analyses 646 Statistical analyses were performed using the two-tailed Mann and Whitney test, unpaired two-tailed Student's t-test, or the Fisher's exact test with GraphPad Prism 8.3 software. The number of samples 647 per experiment and condition is indicated in the corresponding figures and legends. For all analyses, a P 648 value less than 0.05 was considered significant. 649

Study approval 651 652 Clinical and genetic analyses were performed after signed informed consent using approved 653 protocols of the Montpellier University Hospital, France (ID n°: IRB-MTP_2021_11_202100959), the 654 Research Ethics Committee of the Fundación Jiménez Díaz University Hospital, Madrid, Spain (ID n°: CM 655 06-2016 FJD), the Research Ethics Committee of the University Hospital La Fe, Valencia, Spain (ID n°: 656 2019/0098) and the Institute of Molecular and Clinical Ophthalmology Basel (ID n°: 2019-01660), and in 657 agreement with the Declaration of Helsinki. Skin biopsies and iPSC reprogramming were approved by the 658 French National Agency for the Safety of Medicines and Health Products (ID n°: 2014-A00549-38). 659 *Xenopus* care followed institutional guidelines (licenses A91272108 and C 91-471-102) and study protocols 660 were approved by the institutional animal care committee CEEA #59 and the Direction Départementale de 661 la Protection des Populations (APAFIS #21474-2019071210549691v2 and #32589- 662 2021072719047904v4). 663

Data availability 664 665 All data are available in the main text or the supplementary materials. Data point values for all graphs are 666 found in the Supporting Data Values file. 667

668
669 **Author contributions**

670 The authorship order among co-first was determined by the chronological order that they began
671 working on the project. BB, CB, NE, DM, IM, MP and VK designed research studies. BB, CB, NE, DM,
672 AD, KP, CMG, KK, MQ, IPR and MC conducted experiments. CJM and HB analysed data. GGG, AC,
673 NL, GC, SG, AP, VB, VU, MD, FB, HPNS, CA, JMM, IM performed clinical investigations. CB, CA,
674 JMM, CR, IM, MP, VK performed supervision. BB, CB, IM, MP, VK wrote the manuscript. All authors
675 reviewed and edited the manuscript.

676

677 **Acknowledgements**

678 This work is dedicated to Pr. Christian Hamel who initiated the study, identified TBC1D32 as the
679 potential candidate gene, and passed away prematurely in 2017. We thank L. Surcouf, J. Guillot, T. Crouzet-
680 Deprost, M. Pegart and K. Damodar for technical assistance; M. Le Quintrec Donnette for helpful
681 discussions; L. Mansard for variant classification; C. Sanjurjo-Soriano, J. Roger and C.-M. Dhaenens for
682 critical reading of the manuscript; C. Fabre for EMT discussions; C. Cazevieille of the INM CoMET
683 electron microscopy facility; P. Clair of the high-debit qPCR platform (qPHD, Univ Montpellier); the MRI
684 Imaging platform, and the CNAG-CRG for assistance with WES. This work was funded by: Fondation de
685 France, Fondation Valentin Haüy and the associations Retina France (MP), France Choroideremia (VK),
686 AVIESAN-UNADEV (VK, MP) and IRRP (BB and IM); Swiss National Science Foundation (grant n°
687 176097 and 204285) (CR); BBMRI-LPC 2016 access call for Whole Exome Sequencing (FP7/2007-2013,
688 grant agreement n° 313010) (CA); Instituto de Salud Carlos III (Spain) (PI19/00321, PI19/00303 and
689 FI17/00192) (CA and JMM); IIS-FJD BioBank (PT13/0010/0012) (CA); Comunidad de Madrid (CAM,
690 RAREGenomics Project, B2017/BMD-3721) (CA); Organización Nacional de Ciegos Españoles (ONCE)
691 (CA); University Chair UAM-IIS-FJD of Genomic Medicine (CA); Centro de Investigación Biomédica en
692 Red Enfermedades Raras (CIBERER, 06/07/0036) (CA), and European Regional Development Fund
693 (FEDER) (CA). The RD-Connect Genome Phenome Analysis platform was developed under a FP7/2007-
694 2013-funded project (grant agreement n° 305444; CA). CMG is supported by Retina France. BB, CA, JM,
695 CR, IM and VK are members of the European Retinal Disease Consortium (ERDC).

696

697
699 **References**

700 1. Berger W, et al. The molecular basis of human retinal and vitreoretinal diseases. *Prog Retin Eye Res.* 2010;29(5):335-75.

701 2. Hanany M, et al. Worldwide carrier frequency and genetic prevalence of autosomal recessive inherited retinal diseases. *Proc Natl Acad Sci U S A.* 2020;117(5):2710-6.

702 3. Verbakel SK, et al. Non-syndromic retinitis pigmentosa. *Prog Retin Eye Res.* 2018;66:157-86.

703 4. Dias MF, et al. Molecular genetics and emerging therapies for retinitis pigmentosa: Basic research and clinical perspectives. *Prog Retin Eye Res.* 2018;63:107-31.

704 5. Hamel CP. Gene discovery and prevalence in inherited retinal dystrophies. *C R Biol.* 2014;337(3):160-6.

705 6. Bachmann-Gagescu R, Neuhauss SC. The photoreceptor cilium and its diseases. *Curr Opin Genet Dev.* 2019;56:22-33.

706 7. Bujakowska KM, et al. Photoreceptor Cilia and Retinal Ciliopathies. *Cold Spring Harb Perspect Biol.* 2017;9(10).

707 8. Sun C, et al. Primary cilia in retinal pigment epithelium development and diseases. *J Cell Mol Med.* 2021;25(19):9084-8.

708 9. May-Simera HL, et al. Primary Cilium-Mediated Retinal Pigment Epithelium Maturation Is Disrupted in Ciliopathy Patient Cells. *Cell Rep.* 2018;22(1):189-205.

709 10. Ko HW, et al. Broad-minded links cell cycle-related kinase to cilia assembly and hedgehog signal transduction. *Dev Cell.* 2010;18(2):237-47.

710 11. Adly N, et al. Ciliary genes TBC1D32/C6orf170 and SCLT1 are mutated in patients with OFD type IX. *Hum Mutat.* 2014;35(1):36-40.

711 12. Kostiuk V, Khokha MK. Xenopus as a platform for discovery of genes relevant to human disease. *Curr Top Dev Biol.* 2021;145:277-312.

712 13. Zahn N, et al. Normal Table of Xenopus development: a new graphical resource. *Development.* 2022;149(14).

713 14. Perron M, et al. A novel function for Hedgehog signalling in retinal pigment epithelium differentiation. *Development.* 2003;130(8):1565-77.

714 15. Patnaik SR, et al. Bardet-Biedl Syndrome proteins regulate cilia disassembly during tissue maturation. *Cellular and molecular life sciences : CMLS.* 2019;76(4):757-75.

715 16. Doornbos C, et al. Cell-based assay for ciliopathy patients to improve accurate diagnosis using ALPACA. *Eur J Hum Genet.* 2021;29(11):1677-89.

716 17. Muhlhans J, et al. The centrosomal protein pericentrin identified at the basal body complex of the connecting cilium in mouse photoreceptors. *PLoS One.* 2011;6(10):e26496.

717 18. Pazour GJ, et al. The intraflagellar transport protein, IFT88, is essential for vertebrate photoreceptor assembly and maintenance. *J Cell Biol.* 2002;157(1):103-13.

718 19. Shaheen R, et al. Bi-allelic Mutations in FAM149B1 Cause Abnormal Primary Cilium and a Range of Ciliopathy Phenotypes in Humans. *Am J Hum Genet.* 2019;104(4):731-7.

719 20. Assou S, et al. Recurrent Genetic Abnormalities in Human Pluripotent Stem Cells: Definition and Routine Detection in Culture Supernatant by Targeted Droplet Digital PCR. *Stem Cell Reports.* 2020;14(1):1-8.

720 21. Schneider S, et al. Loss of Ciliary Gene Bbs8 Results in Physiological Defects in the Retinal Pigment Epithelium. *Front Cell Dev Biol.* 2021;9:607121.

721 22. Zeisberg M, Neilson EG. Biomarkers for epithelial-mesenchymal transitions. *J Clin Invest.* 2009;119(6):1429-37.

722 23. Zhou M, et al. Role of Epithelial-Mesenchymal Transition in Retinal Pigment Epithelium Dysfunction. *Front Cell Dev Biol.* 2020;8:501.

723 24. Lenis TL, et al. Expression of ABCA4 in the retinal pigment epithelium and its implications for Stargardt macular degeneration. *Proc Natl Acad Sci U S A.* 2018;115(47):E11120-E7.
748 25. Imanishi Y, et al. Retinosomes: new insights into intracellular managing of hydrophobic substances in lipid bodies. *J Cell Biol.* 2004;166(4):447-53.

749 26. Mamaeva D, et al. Novel roles for voltage-gated T-type Ca(2+) and ClC-2 channels in phagocytosis and angiogenic factor balance identified in human iPSC-derived RPE. *FASEB J.* 2021;35(4):e21406.

751 27. Sanjurjo-Soriano C, et al. Retinoic acid delays initial photoreceptor differentiation and results in a highly structured mature retinal organoid. *Stem Cell Res Ther.* 2022;13:478.

754 28. Tatour Y, Ben-Yosef T. Syndromic Inherited Retinal Diseases: Genetic, Clinical and Diagnostic Aspects. *Diagnostics (Basel).* 2020;10(10).

756 29. Talib M, Boon CJF. Retinal Dystrophies and the Road to Treatment: Clinical Requirements and Considerations. *Asia Pac J Ophthalmol (Phila).* 2020;9(3):159-79.

758 30. Bruel AL, et al. Fifteen years of research on oral-facial-digital syndromes: from 1 to 16 causal genes. *J Med Genet.* 2017;54(6):371-80.

760 31. Pezzella N, et al. OFD1: One gene, several disorders. *Am J Med Genet C Semin Med Genet.* 2022;190(1):57-71.

762 32. Alsahan N, Alkuraya FS. Confirming TBC1D32-related ciliopathy in humans. *American journal of medical genetics Part A.* 2020;182(8):1985-7.

763 33. Hietamaki J, et al. Loss-of-Function Variants in TBC1D32 Underlie Syndromic Hypopituitarism. *J Clin Endocrinol Metab.* 2020;105(6).

766 34. Monies D, et al. Lessons Learned from Large-Scale, First-Tier Clinical Exome Sequencing in a Highly Consanguineous Population. *Am J Hum Genet.* 2019;104(6):1182-201.

768 35. Walker RV, et al. Ciliary exclusion of Polycystin-2 promotes kidney cystogenesis in an autosomal dominant polycystic kidney disease model. *Nature communications.* 2019;10(1):4072.

769 36. Moon KH, et al. Dysregulation of sonic hedgehog signaling causes hearing loss in ciliopathy mouse models. *Elife.* 2020;9.

771 37. Marlhens F, et al. Mutations in RPE65 cause Leber's congenital amaurosis. *Nat Genet.* 1997;17:139-41.

773 38. Klettner AK. In: Klettner AK, and Dithmar S eds. *Retinal Pigment Epithelium in Health and Disease*. Switzerland: Springer Nature; 2020:85-100.

776 39. Keeling J, et al. Cellular Mechanisms of Ciliary Length Control. *Cells.* 2016;5(1).

777 40. Bujakowska KM, et al. Mutations in IFT172 cause isolated retinal degeneration and Bardet-Biedl syndrome. *Hum Mol Genet.* 2015;24(1):230-42.

778 41. Halbritter J, et al. Defects in the IFT-B component IFT172 cause Jeune and Mainzer-Saldino syndromes in humans. *Am J Hum Genet.* 2013;93(5):915-25.

779 42. Rachel RA, et al. Photoreceptor sensory cilia and ciliopathies: focus on CEP290, RPGR and their interacting proteins. *Cilia.* 2012;1(1):22.

781 43. Barny I, et al. AON-Mediated Exon Skipping to Bypass Protein Truncation in Retinal Dystrophies Due to the Recurrent CEP290 c.4723A > T Mutation. *Fact or Fiction? Genes (Basel).* 2019;10(5).

783 44. Kruczek K, et al. In vitro modeling and rescue of ciliopathy associated with IQCB1/NPHP5 mutations using patient-derived cells. *Stem Cell Reports.* 2022;17(10):2172-86.

785 45. van der Wal T, van Amerongen R. Walking the tight wire between cell adhesion and WNT signalling: a balancing act for beta-catenin. *Open Biol.* 2020;10(12):200267.

788 46. Veland IR, et al. Primary cilia and signaling pathways in mammalian development, health and disease. *Nephron Physiol.* 2009;111(3):p39-53.

790 47. Liu H, et al. Ciliary signalling in cancer. *Nat Rev Cancer.* 2018;18(8):511-24.

792 48. Wang C, et al. Centrosomal protein Dzip11 binds Cby, promotes ciliary bud formation, and acts redundantly with Bromi to regulate ciliogenesis in the mouse. *Development.* 2018;145(6).

794 49. Satoda Y, et al. BROM1/TBC1D32 together with CCRK/CDK20 and FAM149B1/JBTS36 contributes to intraflagellar transport turnaround involving ICK/CILK1. *Mol Biol Cell.* 2022;33(9):ar79.

796 50. Piro-Megy C, et al. Dominant mutations in mtDNA maintenance gene SSBP1 cause optic atrophy and foveopathy. *J Clin Invest.* 2020;130(1):143-56.
798 51. Azhar Baig HM, et al. Genetic Analysis of Consanguineous Pakistani Families with Congenital
799 Stationary Night Blindness. *Ophthalmic research*. 2022;65(1):104-10.

800 52. Thompson R, et al. RD-Connect: an integrated platform connecting databases, registries, biobanks and
801 clinical bioinformatics for rare disease research. *J Gen Intern Med*. 2014;29 Suppl 3:S780-7.

802 53. Torriano S, et al. Pathogenicity of a novel missense variant associated with choroideremia and its
803 impact on gene replacement therapy. *Hum Mol Genet*. 2017;26(18):3573-84.

804 54. Erkilic N, et al. Generation of a human iPSC line, INMi004-A, with a point mutation in CRX
805 associated with autosomal dominant Leber congenital amaurosis. *Stem Cell Res*. 2019;38:101476.

806 55. Sanjurjo-Soriano C, et al. *USH2A* variants causing retinitis pigmentosa or Usher syndrome provoke
differential retinal phenotypes in disease-specific organoids. *HGG Adv*. 2023;4(4):100229.

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<figure><img src="image_5.png" /><figcaption>**Figure 1.** Multimodal imaging of the patients. Patient 1: **A)** Colour fundus photograph showing pallor of the optic nerve head and reduced caliber of the retinal vessels. Note the abnormal visibility of the choroidal vessels due to RPE damage in the midperiphery. **B)** SW-FAF showing a parafoveal ring characteristic of RP (white arrows). **C)** SD-OCT macular scan showing preserved segmentation in the foveal area with peripheral loss of the ellipsoidal line nasal to the fovea (white arrow). Patient 2: **D)** Color fundus photograph showing increased choroidal visibility beyond the temporal vascular arcades. Inset: pigment accumulation in the superior retinal periphery. **E)** SW-FAF showing a relatively preserved intensity within the macular region and loss of the signal beyond the temporal vascular arcades. An incomplete hyper-autofluorescent ring is visible in the temporal parafoveal region (yellow arrow). **F)** SD-OCT scan along the horizontal meridian through the fovea showing preservation of retinal layers in the central macula and gradual disappearance of photoreceptor layers with increasing eccentricity in the temporal direction. The white arrow indicates the beginning of the disappearance of the EZ line. Patient 3: **G, H)** Colour fundus imaging of both eyes showing peripheral retinal bone spicules (blue arrows), papillary pallor and evident atrophy of the RPE-choriocapillaris complex except in the macular area (delineated by white arrows). **I)** SD-OCT macular scan showing an ellipsoid line only visible beneath the fovea (between white arrows).</figcaption></figure>

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<figure><img src="image_6.png" /><figcaption>**Figure 2.** *tbc1d32* expression during *Xenopus* development and retinogenesis. A) Lateral views of *Xenopus* embryo heads (anterior to the right) following *tbc1d32* whole-mount *in situ* hybridization. Enlargement: at stage 25, epidermal cells are stained. *tbc1d32* expression is also detected in the eye (black arrows), and later in the pronephric nephrostomes (black dotted arrows), the brain (black arrowheads) and the otic vesicle (white arrowheads). B) RNAscope *in situ* hybridizations for *tbc1d32* and *ihh* on cryostat sections. A region of RPE cells, delineated by white dotted boxes in the upper panels, is enlarged in the lower panels where the RPE layer is outlined with white lines. Scatter plots with bars represent the</figcaption></figure>
quantification of *tbc1d32* expression in the RPE or in the photoreceptor layer at each stage (st). Data are represented as mean ± SEM. Each dot represents one retina. AU: arbitrary units, pR: presumptive retina, RPE: retinal pigment epithelium. PR: photoreceptors. Scale bars = 400 µm in A, 50 µm in B. C) qPCR analysis of *ihh* and *tbc1d32* expression in neural retina and RPE tissues, dissected at stage 35-36. *ihh* serves as a specific RPE marker. Data are represented as geometric mean with 95% CI; n = 3 technical replicates.
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<figure><img src="image_12.png" /><figcaption>**Figure 3. Xenopus RPE phenotype following tbc1d32 knockdown.** A) Mo1 and Mo2 target sequences located in the 5'UTR of *tbc1d32* mRNA. B) Diagram of the experimental design. C) Upper panels, lateral views of one control and two morphant embryo heads with moderate and severe phenotypes (anterior to the right). Lower panels, transverse retinal sections of control and morphant embryos (dorsal side up). The histogram represents the percentage of embryos with a severe decrease in pigmentation among control (control Mo), morphant (*tbc1d32* Mo1), *tbc1d32* mRNA-injected, and *tbc1d32* Mo1/*tbc1d32* mRNA co-injected, groups. The total number of embryos analyzed per condition is indicated in each bar. **p<0.01; ****p<0.0001; Fisher's exact test. D) Phalloidin staining of filamentous actin on dissected eyes of control or morphant *Xenopus* embryos. The histogram represents the proportion of eyes with an altered distribution</figcaption></figure>
850 of F-actin for each condition. The total number of eyes analyzed per condition is indicated in each bar.

851 \*p<0.05; \*\*\*p<0.0001; Fisher's exact test. Scale bars = 400 µm for whole mounts and 60 µm for sections

852 in C, and 20 µm in D.

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<figure><img src="image_13.png" /><figcaption>**Figure 4.** *Xenopus* RPE and photoreceptor marker expression following *tbc1d32* knockdown. **A)** Diagram of the experimental design. Whole mount (**B**) or retinal sections (**C**) following in *situ* hybridization against *mitf* or *ihh*, respectively, on embryos injected with control Mo or *tbc1d32* Mo1. Scatter plots represent the quantification of the integrated density of the staining per eye relative to control Mo; each dot corresponds to one eye or one section, respectively. **D)** Rho and S+M Opsin immunolabelling on retinal</figcaption></figure>

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sections of embryos injected with control Mo or embryos injected with 2 doses of *tbc1d32* Mo1 (10 or 15 ng). Lower panels, enlargement of the areas indicated by white dashed boxes in the upper panels. The histogram represents the proportion of eyes with altered staining of Rho and S+M Opsin for each condition.

The number of eyes analyzed per condition is indicated in each bar. E) Upper panels, whole mount *in situ* hybridization against *rhodopsin (rho)* in embryos injected with control Mo or *tbc1d32* Mo1. Lower panels, transverse retinal sections of control and morphant embryos. The scatter plots represent the quantification of the integrated density of *rho* staining relative to control Mo; each dot corresponds to one eye (left) or one section (right). For all scatter plots, data are represented as mean ± SEM. ns: non-significant; \*p\<0.05; \*\*p\<0.01; \*\*\*p\<0.001; \*\*\*\*p\<0.0001; Fisher's exact test (D); Mann and Whitney test (B, C, and E). Scale bars = 400 µm in B, 40 µm in C and E, 400 µm for whole mount and 40 µm for sections.

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<figure><img src="image_22.png" /><figcaption>**Figure 5.** *Xenopus* ciliogenesis following *tbc1d32* knockdown. A) Arl13b and γ-Tubulin (γ-Tub) immunolabelling on neural tube sections of stage 35-36 wild-type (WT), control Mo, or *tbc1d32* Mo1 embryos. B) Scatter plot shows the mean length of neural tube cilia. Each dot corresponds to the mean length for one embryo. C) Immunolabelling of cilia markers, acetylated α-Tubulin (α-Tub), Arl13b, and γ-</figcaption></figure>
Tub, on retinal sections of stage 35-36 WT embryo. The right panels show an enlargement of the area delineated by a white dotted box in the left panel. D) α-Tub immunolabelling of RPE cilia (arrowheads) at different stages. Sections are co-stained with Otx2 to identify RPE cells, which are enlarged in the lower panels. NR: neural retina, ONL: outer nuclear layer, OS: outer segment. E) Proportion of ciliated cells among RPE cells at different stages (st). F) α-Tub and Otx2 immunolabelling showing primary RPE cilia (brackets) on retinal sections of stage 35 WT, control Mo or *tbc1d32* Mo1 embryos. G) Scatter plot with bars showing the mean cilia length in RPE cells. Each dot corresponds to the mean length for one embryo. H) Immunolabelling of Arl13b showing the photoreceptor connective cilium on retinal sections of stage 35 WT, control Mo or *tbc1d32* Mo1 embryos. I) The scatter plot shows the mean number of Arl13⁺ ciliated photoreceptors in one field of the central retina. Each dot corresponds to one embryo. All data are represented as mean ± SEM. ns: non-significant, \*p\<0.05, \*\*p\<0.001, \*\*\*p\<0.0001; Mann and Whitney test. Scale bars = 2 µm in A, 50 µm and 2 µm for enlargements in C, 5 µm and 2 µm for enlargements in D, 25 µm in H.

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<figure><img src="image_23.png" /><figcaption>**Figure 6. Morphology of the primary cilium in human fibroblasts.** IF studies and single plane confocal imaging to assay ciliogenesis in control (A, C, E) and *TBC1D32* (B, D, E) fibroblasts with antibodies directed against ARL13B and acetylated α-tubulin (Ac αTUB; A, B); GT335 (C, D) and PCN (E, F). G) Quantification of cilium length in control and *TBC1D32* fibroblasts. Data are represented as mean ± SEM; n = 66 cells; ****p<0.0001; Student's t-test. IF studies to assay intraflagellar transport in control (H) and *TBC1D32* (I) fibroblasts with antibodies directed against IFT88 and Ac αTUB. IF studies in control (J) and *TBC1D32* (K) fibroblasts to assay the localization of TBC1D32 in relation to Ac αTUB. IF studies in control (L, N) and *TBC1D32* (M, O) fibroblasts to assay the localization of TBC1D32 in relation to PCN in a basal (L, M) and apical (N, O) plane. Insets show magnification of the labelled basal bodies/centrosomes in each panel. Scale bars = 5 µm.</figcaption></figure>
<figure><img src="image_24.png" /><figcaption>**Figure 7. Morphology of the human iPSC-derived RPE.** IF studies and maximum intensity projection (MIP) confocal imaging of control (A) and *TBC1D32* (B) iPSC-derived RPE using antibodies directed against ZO1 and ARL13. White arrows indicate elongated cilia. The disrupted *TBC1D32* RPE monolayer can be seen on the orthogonal planes as compared to control. Scale bars = 10 µm. C) Quantification of the cilium length in control (n= 229 cells) and *TBC1D32* iPSC-derived RPE (n= 416 cells). Data are represented as mean ± SEM; ****p<0.0001; Student's t-test. D) Higher magnification of control iPSC-derived RPE showing the regular cobblestone morphology and cilia length. E) Higher magnification of *TBC1D32* RPE showing irregularly-shaped cells, separated tight junctions (white arrows) and ZO1 aggregates. Scale bars = 5 µm. F) Weekly TER measurements expressed in Ω/cm² in control (black line) and *TBC1D32* (grey line) iPSC-derived RPE. Data are represented as mean ± SEM; *p<0.05; n = 4 inserts; Mann and Whitney test.</figcaption></figure>
<figure><img src="image_25.png" /><figcaption>**Figure 8.** EMT-like phenotype in human **TBC1D32** iPSC-derived RPE. IF studies and MIP confocal imaging of control (A) and **TBC1D32** (B) iPSC-derived RPE using antibodies directed against vimentin and MERTK. The boxed area in B indicates areas of upregulated vimentin and disrupted MERTK expression. Scale bars = 10 µm. IF studies and MIP confocal imaging of control (C) and **TBC1D32** (D) iPSC-derived RPE using antibodies directed against SMA and ZO1. Scale bars = 20 µm. qPCR analysis of</figcaption></figure>
CDH1 (E), CDH3 (F), CTNNB1 (G), SNAI1 (H) and FN1 (I) in control and TBC1D32 RPE. Data are represented as mean ± SEM; n = 3 technical replicates; \*p\<0.05; Mann and Whitney test. Representative western blot analysis of E-cadherin (J), P-cadherin (K) and β-catenin (L) expression and quantification relative to the β-actin loading control; the same membrane was hybridized with two different primary antibodies in panels J and L. Data represented as mean ± SEM; n = 3 blots; \*p\<0.05. M) Representative western blot analysis and quantification of two independent blots of β-catenin distribution in the cytosol versus membrane fractions of control and TBC1D32 iPSC-derived RPE relative to the β-actin loading control and displayed on a logarithmic scale. Data are represented as mean ± SEM.

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<figure><img src="image_26.png" /><figcaption>**Figure 9.** Retinosome accumulation and altered secretion in human *TBC1D32* iPSC-derived RPE. IF studies and MIP confocal imaging of control (A) and *TBC1D32* (B) iPSC-derived RPE using antibodies directed against N-cadherin and CRALBP. Redistribution of N-cadherin and CRALBP can be seen on the orthogonal planes. Scale bars = 10 µm. C) Representative western blot analysis and quantification of two independent blots of CRALBP distribution in the cytosol versus membrane fractions of control and *TBC1D32* iPSC-derived RPE relative to the β-actin loading control. Data represented as mean ± SEM. TEM of control (D) and *TBC1D32* (E) iPSC-derived RPE showing apical microvilli (MV), basal nuclei (N), pigmented melanosomes and vesicles corresponding to lipid droplets (black arrow). Scale bar = 2 µm. F) Quantification of the number of vesicles per cell in control (n = 38 cells) and *TBC1D32* (n = 39 cells) RPE. Data are represented as mean ± SEM; ****p<0.0001; Student's t-test. G) Representative western blot</figcaption></figure>
942 analysis of perilipin expression and quantification relative to the β-actin loading control. Data represented
943 as mean ± SEM. n = 3 blots; *p<0.05. H) ELISA of PEDF and VEGF secretion in apical and basal chambers
944 of control and TBC1D32 RPE cultured on transwell membrane inserts. Data are expressed as a percentage
945 of control; *p<0.05; **p<0.01; ***p<0.001; ****p<0.0001; the number of samples are indicated within
946 bars. Student's t-test.
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<figure><img src="image_27.png" /><figcaption>**Figure 10.** Morphology of human iPSC-derived retinal organoids. Bright-field microscopy of control (A) and *TBC1D32* (B) retinal organoids at day 225 of differentiation showing the brush border. Scale bars = 100 µm. Higher magnification of the ONL of control (C) and *TBC1D32* (D) organoids. Scale bars = 50 µm. IF studies and MIP confocal imaging of control (E,G,I) and *TBC1D32* (F,H,J) organoids with antibodies directed to CRX and RCVRN (E,F) to assay general photoreceptor morphology, CRALBP and ZO1 to assay the OLM (G,H), and RK and ZO1 to assay the prolongation of the IS and OS (I,J). IF studies of control (K,M,O) and *TBC1D32* (L,N,P) organoids with antibodies directed to ARL13B and RHO (K,L) to assay the CC and prolongation of the OS, ABCA4 and PDE6B (M,N) to assay the OS of rods and/or cones, and ARR3 and (in red) (O,P) to assay the cones. All nuclei are labelled with Hoechst. Scale bars = 20 µm.</figcaption></figure>