Skip to main content
Advertisement

Main menu

  • Home
  • Articles
    • Accepted manuscripts
    • Issue in progress
    • Latest complete issue
    • Issue archive
    • Archive by article type
    • Interviews
    • Sign up for alerts
  • About us
    • About BiO
    • Editors and Board
    • Editor biographies
    • Grants and funding
    • Journal Meetings
    • Workshops
    • The Company of Biologists
    • Journal news
  • For authors
    • Submit a manuscript
    • Aims and scope
    • Presubmission enquiries
    • Article types
    • Manuscript preparation
    • Cover suggestions
    • Editorial process
    • Promoting your paper
    • Open Access
  • Journal info
    • Journal policies
    • Rights and permissions
    • Media policies
    • Reviewer guide
    • Sign up for alerts
  • Contact
    • Contact BiO
    • Advertising
    • Feedback
  • COB
    • About The Company of Biologists
    • Development
    • Journal of Cell Science
    • Journal of Experimental Biology
    • Disease Models & Mechanisms
    • Biology Open

User menu

  • Log in

Search

  • Advanced search
Biology Open
  • COB
    • About The Company of Biologists
    • Development
    • Journal of Cell Science
    • Journal of Experimental Biology
    • Disease Models & Mechanisms
    • Biology Open

supporting biologistsinspiring biology

Biology Open

Advanced search

RSS   Twitter   Facebook   YouTube

  • Home
  • Articles
    • Accepted manuscripts
    • Issue in progress
    • Latest complete issue
    • Issue archive
    • Archive by article type
    • Interviews
    • Sign up for alerts
  • About us
    • About BiO
    • Editors and Board
    • Editor biographies
    • Grants and funding
    • Journal Meetings
    • Workshops
    • The Company of Biologists
    • Journal news
  • For authors
    • Submit a manuscript
    • Aims and scope
    • Presubmission enquiries
    • Article types
    • Manuscript preparation
    • Cover suggestions
    • Editorial process
    • Promoting your paper
    • Open Access
  • Journal info
    • Journal policies
    • Rights and permissions
    • Media policies
    • Reviewer guide
    • Sign up for alerts
  • Contact
    • Contact BiO
    • Advertising
    • Feedback
Research Article
Mutations in the splicing regulator Prp31 lead to retinal degeneration in Drosophila
Sarita Hebbar, Malte Lehmann, Sarah Behrens, Catrin Hälsig, Weihua Leng, Michaela Yuan, Sylke Winkler, Elisabeth Knust
Biology Open 2021 10: bio052332 doi: 10.1242/bio.052332 Published 25 January 2021
Sarita Hebbar
Max-Planck-Institute of Molecular Cell Biology and Genetics, Pfotenhauerstrasse 108, 01307 Dresden, Germany
  • Find this author on Google Scholar
  • Find this author on PubMed
  • Search for this author on this site
  • ORCID record for Sarita Hebbar
Malte Lehmann
Max-Planck-Institute of Molecular Cell Biology and Genetics, Pfotenhauerstrasse 108, 01307 Dresden, Germany
  • Find this author on Google Scholar
  • Find this author on PubMed
  • Search for this author on this site
Sarah Behrens
Max-Planck-Institute of Molecular Cell Biology and Genetics, Pfotenhauerstrasse 108, 01307 Dresden, Germany
  • Find this author on Google Scholar
  • Find this author on PubMed
  • Search for this author on this site
Catrin Hälsig
Max-Planck-Institute of Molecular Cell Biology and Genetics, Pfotenhauerstrasse 108, 01307 Dresden, Germany
  • Find this author on Google Scholar
  • Find this author on PubMed
  • Search for this author on this site
Weihua Leng
Max-Planck-Institute of Molecular Cell Biology and Genetics, Pfotenhauerstrasse 108, 01307 Dresden, Germany
  • Find this author on Google Scholar
  • Find this author on PubMed
  • Search for this author on this site
Michaela Yuan
Max-Planck-Institute of Molecular Cell Biology and Genetics, Pfotenhauerstrasse 108, 01307 Dresden, Germany
  • Find this author on Google Scholar
  • Find this author on PubMed
  • Search for this author on this site
Sylke Winkler
Max-Planck-Institute of Molecular Cell Biology and Genetics, Pfotenhauerstrasse 108, 01307 Dresden, Germany
  • Find this author on Google Scholar
  • Find this author on PubMed
  • Search for this author on this site
Elisabeth Knust
Max-Planck-Institute of Molecular Cell Biology and Genetics, Pfotenhauerstrasse 108, 01307 Dresden, Germany
  • Find this author on Google Scholar
  • Find this author on PubMed
  • Search for this author on this site
  • ORCID record for Elisabeth Knust
  • For correspondence: knust@mpi-cbg.de
  • Article
  • Figures & tables
  • Supp info
  • Info & metrics
  • eLetters
  • PDF + SI
  • PDF
Loading

ABSTRACT

Retinitis pigmentosa (RP) is a clinically heterogeneous disease affecting 1.6 million people worldwide. The second-largest group of genes causing autosomal dominant RP in human encodes regulators of the splicing machinery. Yet, how defects in splicing factor genes are linked to the aetiology of the disease remains largely elusive. To explore possible mechanisms underlying retinal degeneration caused by mutations in regulators of the splicing machinery, we induced mutations in Drosophila Prp31, the orthologue of human PRPF31, mutations in which are associated with RP11. Flies heterozygous mutant for Prp31 are viable and develop normal eyes and retina. However, photoreceptors degenerate under light stress, thus resembling the human disease phenotype. Degeneration is associated with increased accumulation of the visual pigment rhodopsin 1 and increased mRNA levels of twinfilin, a gene associated with rhodopsin trafficking. Reducing rhodopsin levels by raising animals in a carotenoid-free medium not only attenuates rhodopsin accumulation, but also retinal degeneration. Given a similar importance of proper rhodopsin trafficking for photoreceptor homeostasis in human, results obtained in flies presented here will also contribute to further unravel molecular mechanisms underlying the human disease.

This paper has an associated First Person interview with the co-first authors of the article.

INTRODUCTION

Retinitis pigmentosa (RP; OMIM 268000) is a clinically heterogeneous group of retinal dystrophies, which affects more than one million people worldwide. It often starts with night blindness in early childhood, continues with the loss of the peripheral visual field (tunnel vision), and progresses to complete blindness in later life due to gradual degeneration of photoreceptor cells (PRCs). RP is a genetically heterogeneous disease and can be inherited as autosomal dominant (adRP), autosomal recessive (arRP) or X-linked (xlRP) disease. So far >90 genes have been identified that are causally related to non-syndromic RP (Ali et al., 2017; Verbakel et al., 2018). Affected genes are functionally diverse. Some of them are expressed specifically in PRCs and encode, among others, transcription factors (e. g. CRX, an otx-like photoreceptor homeobox gene), components of the light-induced signalling cascade, including the visual pigment rhodopsin (Rho/RHO in Drosophila/human), or genes controlling vitamin A metabolism (e.g. RLBP-1, encoding Retinaldehyde-binding protein). Other genes are associated with a more general control of cellular homeostasis, for example genes involved in trafficking or cell polarity (e.g. CRB1) [reviewed in (Daiger et al., 2014, 2013; Hollingsworth and Gross, 2012; Nemet et al., 2015)]. Interestingly, the second-largest group of genes causing adRP, comprising 7 of 25 genes known, encodes regulators of the splicing machinery. So far, mutations in five pre-mRNA processing factor (PRPF) genes, PRPF3, PRPF4, PRPF6, PRPF8 and PRPF31, have been linked to adRP, namely RP18, RP70, RP60, RP13 and RP11, respectively. Pim1-associated protein (PAP1) and small nuclear ribonuclearprotein-200 (SNRNP200), two genes also involved in splicing, have been suggested to be associated with RP9 and RP33, respectively (Maita et al., 2004; Zhao et al., 2009) [reviewed in (Liu and Zack, 2013; Mordes et al., 2006; Poulos et al., 2011; Ruzickova and Stanek, 2016)]. The five PRPF genes encode components regulating the assembly of the U4/U6.U5 tri-snRNP, a major module of the pre-mRNA spliceosome machinery (Nguyen et al., 2015; Patel and Bellini, 2008; Will and Luhrmann, 2011). Several hypotheses have been put forward to explain why mutations in ubiquitously expressed components of the general splicing machinery show a dominant phenotype only in the retina. One hypothesis suggests that PRCs with only half the copy number of a gene encoding a general splicing component cannot cope with the elevated demand of RNA-/protein synthesis required to maintain the exceptionally high metabolic rate of PRCs in comparison to other tissues. Hence, halving their gene dose eventually results in apoptosis. Although this model is currently favoured, other mechanisms, such as impaired splicing of PRC-specific mRNAs or toxic effects caused by accumulation of mutant proteins have been discussed and may contribute to the disease phenotype [discussed in (Mordes et al., 2006; Scotti and Swanson, 2016; Tanackovic et al., 2011)]. More recent data obtained from retinal organoids established from RP11 patients showed that removing one copy of PRPF31 affects the splicing machinery specifically in retinal and retinal pigment epithelial (RPE) cells, but not in patient-derived fibroblasts or iPS cells (Buskin et al., 2018).

The observation that all adRP-associated genes involved in splicing are highly conserved from yeast to human allows to use model organisms to unravel the genetic and cell biological functions of these genes in order to obtain mechanistic insight into the origin of the diseases. In the case of RP11, the disease caused by mutations in PRPF31, three mouse models have been generated by knock-in and knock-out approaches. Unexpectedly, mice heterozygous mutant for a null allele or a point mutation that recapitulates a mutation in the corresponding human gene did not show any sign of retinal degeneration in 12- and 18-month-old mice, respectively (Bujakowska et al., 2009). Further analyses revealed that the retinal pigment epithelium, rather than the PRCs, is the primary tissue affected in Prpf31 heterozygous mice (Farkas et al., 2014; Graziotto et al., 2011; Hamieh and Nandrot, 2019). Other data show that homozygous PRPF31 mice are not viable (Dickinson et al., 2016). Morpholino-induced knockdown of zebrafish Prpf31 results in strong defects in PRC morphogenesis and survival (Linder et al., 2011). Defects induced by retina-specific expression of zebrafish Prpf31 constructs that encode proteins with the same mutations as those mapped in RP11 patients (called AD5 and SP117) were explained to occur by either haplo-insufficiency or by a dominant-negative effect of the mutant protein (Yin et al., 2011). In Drosophila, no mutations in the orthologue Prp31 have been identified so far. RNAi-mediated knockdown of Prp31 in the Drosophila eye results in abnormal eye development, ranging from smaller eyes to complete absence of the eye, including loss of PRCs and pigment cells (Ray et al., 2010).

In order to get better insights into the mechanisms by which Prp31 prevents retinal degeneration we aimed to establish a meaningful Drosophila model for RP11-associated retinal degeneration. Therefore, we isolated two mutant alleles of Prp31, Prp31P17 and Prp31P18, which carry missense mutations affecting conserved amino acids. Flies heterozygous for either of these mutations are viable and develop normally. Strikingly, when exposed to constant light, these mutant flies undergo retinal degeneration, thus mimicking the disease phenotype of RP11 patients. Degeneration of mutant PRCs is associated with accumulation and abnormal distribution of the visual pigment rhodopsin, Rh1, in PRCs. Reduction of dietary vitamin A, a precursor of the chromophore 11-cis-3-hydroxyretinal, which binds to opsin to generate the functional rhodopsin, mitigates both aspects of the mutant phenotype, rhodopsin accumulation and retinal degeneration. From this we conclude that Rh1 accumulation and/or misdistribution reflect a degeneration-prone condition in the Prp31 mutant retina.

RESULTS

Two Prp31 alleles were discovered by TILLING

It was recently shown that RNAi-mediated knockdown of Drosophila Prp31 in the eye using eye-specific Gal4-lines [eyeless (ey)-Gal4 or GMR-Gal4] results in abnormal eye development, ranging from smaller eyes to complete absence of the eye, including loss of photoreceptor cells (PRCs) and pigment cells (Ray et al., 2010). Both Gal4-lines are expressed throughout eye development. Therefore, some of the defects observed could be the result of impaired early development of the eye, such as defective cell fate specification, which would only indirectly affect PRC development. Here, we aimed to establish a more meaningful Drosophila model for RP11-associated retinal degeneration, a human disease associated with mutations in the human orthologue PRPF31, which would allow a deeper insight into the role of this splicing factor in the origin and progression of the disease.

Therefore, we set out to isolate specific mutations in Drosophila Prp31 by targeting induced local lesions in genomes (TILLING), following a protocol described recently (Spannl et al., 2017). In total, 2.400 genomes of ethyl methanesulfonate (EMS)-mutagenised flies were screened for sequence variants in two different amplicons of Prp31. Four sequence variants were identified, which were predicted to result in potentially deleterious missense mutations. Two of the four lines, named Prp31P17 and Prp31P18, were recovered from the living fly library and crossed for three generations to control, white-eyed (w*) flies to reduce the number of accompanying sequence variations. We outcrossed the mutants with white-eyes flies (w*) rather than with wild-type, red-eyed flies to generate a sensitised background for light-dependent degeneration experiments, since the presence of the pigment granules surrounding each ommatidium contributes towards lower sensitivity to light (Stark and Carlson, 1984). Prp31P18 flies were viable as homozygotes and in trans over any of three deficiencies, which remove, among others, the Prp31 locus (Fig. 1A). In contrast, no homozygous Prp31P17 flies were obtained. However, Prp31P17 was viable in trans over Prp31P18 and over Df(3L)ED217. This suggests that the lethality was due to a second site mutation, which was not removed during outcrossing. We noticed that outcrossing Prp31P17 and Prp31P18 did not remove scarlet (st), one of the markers of the original, mutagenised chromosome (ru st e ca) mapping close to Prp31. Therefore, the correct genotypes of the two mutant lines are w*; Prp31P17, st1 and w*; Prp31P18, st1. For simplicity, we will refer to them as Prp31P17 and Prp31P18 throughout the text.

Fig. 1.
  • Download figure
  • Open in new tab
  • Download powerpoint
Fig. 1.

Eyes of Prp31 mutant flies have no gross morphological abnormalities at eclosion. (A) Schematic of chromosome arm 3L. Prp31 and scarlet (st) are situated 2 cM apart (3–42 and 3–44, respectively; cytological positions 71B6 and 73A3, respectively; www.flybase.org). In both Prp31 mutant alleles the marker st1 from the original mutagenised chromosome (ru st e ca) is retained. The three deficiencies used cover the Prp31 locus, but not the st locus. (B) Schematic overview of the Drosophila Prp31 protein. The figure is drawn to scale using IBS (Liu et al., 2015). Domains described here are indicated. The two Prp31 alleles studied here carry non-conservative missense mutations, G90R in Prp31P17 and P277L in Prp31P18. (C–F) Representative bright-field images of Toluidine-blue stained semi-thin sections of eyes of w* (C), w*;; st1/+ (D) Prp31P18 /+ (E) and Prp31P17 /+ (F). Complete genotypes can be found in Table S1. Upon eclosion, flies were kept for 2 days under regular light conditions. Note that the number and stereotypic arrangement of photoreceptor cells within the mutant ommatidia are not affected. Scale bar, 10 µm.

The molecular lesions in the two Prp31 alleles were mapped in the protein coding region. Drosophila PRP31 is a protein of 501 amino acids, which contains a NOSIC domain (named after the central domain of Nop56/SIK1-like protein), a Nucleolar protein (Nop) domain required for RNA binding, a PRP31_C-specific domain and a nuclear localization signal, NLS (Fig. 1B). Prp31P17 contained a point mutation that resulted in a non-conservative glutamine to arginine exchange (G90R) N-terminal to the NOSIC domain. Prp31P18 contained a non-conservative exchange of a proline to a leucine residue in the Nop domain (P277L) (Fig. 1B). Both mutations affect amino acids that are highly conserved in many metazoan species (Fig. S1A). Based on polymorphism phenotyping v2 (PolyPhen-2; http://genetics.bwh.harvard.edu/pph2/) (Adzhubei et al., 2013) analyses, both amino acid substitutions are predicted to be deleterious for the structure and/or function of the protein (Fig. S1B).

Flies hetero- or hemizygous for Prp31 undergo light-dependent retinal degeneration

Homo- and heterozygous Prp31P18 and heterozygous Prp31P17 animals raised and kept under regular light–dark cycles (12 h light; 12 h dark) have eyes of normal size. Histological sections revealed normal numbers of PRCs per ommatidium (distinguished by the number of rhabdomeres) and a normal stereotypic arrangement of PRCs (Fig. 1C–F and Fig. S2A). This indicates that the development of the retina was not affected by these mutations. However, PRCs of Prp31P17/+, Prp31P18/+ and Prp31P18/Prp31P18 flies showed clear signs of retinal degeneration when exposed to constant light for several days, manifested by a partial or complete loss of rhabdomeric integrity (Fig. 2C,D and Fig. S2B). Quantification of the number of surviving rhabdomeres in Prp31 mutant retinas revealed only about 48% of ommatidia with the full complement of seven PRCs (Fig. 2E), while w* mutant control flies exhibited 82% of all ommatidia displaying the full complement of rhabdomeres (Fig. 2A,E). The degree of degeneration observed in Prp31 alleles is less severe and more variable than that observed in the well-established RP12 disease model induced by mutations in the gene crumbs (crb) (Chartier et al., 2012; Johnson et al., 2002; Spannl et al., 2017). In the two crb alleles crb11A22 and crbp13A9 only 5 to 11% of all ommatidia displayed seven rhabdomeres upon exposure to constant light, respectively (Fig. 2E).

Fig. 2.
  • Download figure
  • Open in new tab
  • Download powerpoint
Fig. 2.

PRCs of heterozygous Prp31P17 and Prp31P18 flies undergo light-dependent degeneration. (A–D) Representative bright-field images of Toluidine-blue stained semi-thin sections of eyes of w* (A), w*;; st1/+ (B), Prp31P18/+ (C), and Prp31P17 /+ (D). Complete genotypes can be found in Table S1. Upon eclosion, flies were kept for 2 days under regular light conditions and then subjected to a degeneration paradigm of 7 days of continuous, high intensity light exposure. Whereas in w* (A) most ommatidia (red outlines) display seven rhabdomeres indicative of the seven PRCs, w*;; st1/+ and Prp31 mutant ommatidia (B–D, red outlines) display fewer rhabdomeres per ommatidium indicative of degeneration. Scale bar, 10 µm. (E) Quantification of retinal degeneration as indicated by the number of surviving rhabdomeres observed upon high intensity, continuous light exposure. Y-axis: percent frequency of ommatidia displaying one to seven rhabdomeres. Genotypes are indicated below. Number on top of each graph indicates the mean percentage of ommatidia displaying the full complement of seven rhabdomeres. Bars represent mean±s.e.m. (a minimum of n=60 ommatidia from eyes of three biological replicates). Statistical significance of differences in this parameter, between genotype pairs, is indicated in Table S2.

To further confirm that the degeneration phenotype observed in Prp31P18 and Prp31P17 heterozygous flies is due to mutations in Prp31, we used additional strategies to reduce/inactivate Prp31 function. First, we knocked down Prp31 by overexpressing Prp31 RNAi, mediated by Rh1-Gal4, which drives expression late in retinal development, from 70% pupal development into adulthood (Kumar and Ready, 1995). Thereby, we can rule out any early effects on PRC specification or morphogenesis induced by loss of Prp31. To make the data comparable to those obtained with Prp31 alleles (which are in a w background), we reduced the red-coloured screening pigments encoded by the w+-gene on the transgenes by expression of another transgene, GMR-wIR, which expresses white RNAi under the control of the GMR-promoter (Lee and Carthew, 2003). RNAi-mediated knockdown of Prp31 in PRCs (and concomitant ubiquitous knockdown of w) resulted in clear signs of degeneration upon light exposure, such as loss of rhabdomeres and accumulation of intensely stained structures reminiscent to apoptotic bodies (Fig. 3B). In fact, while 71% of control ommatidia revealed 7 identifiable rhabdomeres and no major morphological defects (Fig. 3A,C), the number of ommatidia with a full complement of rhabdomeres decreased to 48% upon induction of Prp31 RNAi (Fig. 3C).

Fig. 3.
  • Download figure
  • Open in new tab
  • Download powerpoint
Fig. 3.

RNAi-mediated knockdown of Prp31 results in light-dependent retinal degeneration. (A,B) Representative bright-field images of Toluidine-blue stained semi-thin sections of eyes of Rh1-Gal4> (A; control) and Rh1-Gal4>UAS Prp31RNAi (B). Complete genotypes can be found in Table S1. Upon eclosion, flies were kept for 2 days under regular light conditions and then subjected to a degeneration paradigm of 7 days of continuous, high-intensity light exposure. In case of Rh1-Gal4>UAS Prp31RNAi, fewer ommatidia with seven rhabdomeres are seen. Scale bar, 10 µm. (C) Quantification of retinal degeneration as indicated by the number of surviving rhabdomeres observed upon high intensity, continuous light exposure. Y-axis: percent frequency of ommatidia displaying one to seven rhabdomeres. Genotypes are indicated below. Number on top of each graph indicates the mean percentage of ommatidia displaying the full complement of seven rhabdomeres. Bars represent mean±s.e.m. (a minimum of n=60 ommatidia from eyes of three biological replicates). Whilst 71% of control ommatidia have seven rhabdomeres/ommatidium, this number is significantly reduced to 48% upon knocking-down Prp31 by RNAi (P<0.05, shown in Table S2).

As a second alternative strategy to confirm the role of Prp31 in retinal degeneration, we analysed the phenotype of three deficiency lines that remove the Prp31 locus (see Fig. 1A). For a proper comparison with the data obtained for the Prp31 alleles (which are in a w background), we removed the red pigments of the deficiency lines (caused by the presence of a w+-minigene) by studying their phenotype in a cn bw background, an alternative way to remove all screening pigments. Df(3L)Exel6262/+, Df(3L)ED217/+, and Df(3L)ED218/+ flies exhibited retinal degeneration similar as Prp31P17 or Prp31P18 heterozygous flies (Fig. 4), with only about 20% of their ommatidia showing seven rhabdomeres. These deficiency lines also had no obvious effects on retinal development (Fig. S2D–F). Degeneration was also observed in Prp31P18/Df (3L)217 and Prp31P17/Df (3L)217) flies (Fig. S2G,H).

Fig. 4.
  • Download figure
  • Open in new tab
  • Download powerpoint
Fig. 4.

Flies heterozygous for deficiencies that remove the Prp31, but not the scarlet locus, undergo light-dependent degeneration. (A–D) Representative bright-field images of Toluidine-blue stained semi-thin sections of eyes of males of cn bw (A), Df (3L) Exel 6262/+ (B), Df (3L) ED217/+ (C) and Df (3L) ED218/+ (D). Complete genotypes can be found in Table S1. Upon eclosion, flies were kept for 2 days under regular light conditions and then subjected to a degeneration paradigm of 7 days of continuous, high intensity light exposure. Red circles outline individual ommatidia. Scale bar, 10 µm. (E) Quantification of retinal degeneration as indicated by the number of surviving rhabdomeres observed upon high intensity, continuous light exposure. Y-axis: percent frequency of ommatidia displaying one to seven rhabdomeres. Genotypes are indicated below. Number on top of each graph indicates the mean percentage of ommatidia displaying the full complement of seven rhabdomeres. Bars represent mean±s.e.m. (a minimum of n=60 ommatidia from eyes of three biological replicates). Statistical significance of differences in this parameter between genotype pairs is indicated in Table S2.

We noticed that retinal degeneration in w*; st1/+ flies was enhanced compared to that of w* flies (Fig. 2A,B and E). To rule out that retinal degeneration observed in Prp31 mutant flies is influenced by the presence of a mutation in st mapping close by (Fig. 1A), we overexpressed st in the retina of Prp31P18 flies (simultaneously knocking-down w gene activity provided by the transgenes). Expression of st did not modify the degree of retinal degeneration of Prp31 mutants (Fig. 5, Table S2).

Fig. 5.
  • Download figure
  • Open in new tab
  • Download powerpoint
Fig. 5.

PRCs of heterozygous Prp31P18 flies undergo light-dependent degeneration in the presence of st overexpression. (A–C) Representative bright-field images of Toluidine-blue stained semi-thin sections of eyes of GMR-Gal4> (A; control) and GMR-Gal4>; Prp31P18 /+ (B; mutant control) and GMR-Gal4 >;UAS-st ; Prp31P18 /+ (C; mutant and st overexpression). Complete genotypes can be found in Table S1. Upon eclosion, flies were kept for 2 days under regular light conditions and then subjected to a degeneration paradigm of 7 days of continuous, high-intensity light exposure. In the presence of st overexpression, PRCs of heterozygous Prp31P18 flies undergo degeneration to the same extent (C) as without st overexpression (B). Scale bar, 10 µm. (D) Quantification of retinal degeneration as indicated by the number of surviving rhabdomeres observed upon high intensity, continuous light exposure. Y-axis: percent frequency of ommatidia displaying one to seven rhabdomeres. Genotypes are indicated below. Number on top of each graph indicates the percentage of the mean number of ommatidia displaying the full complement of seven rhabdomeres. Bars represent mean±s.e.m. (a minimum of n=60 ommatidia from eyes of three biological replicates). Statistical significance of differences in this parameter between genotype pairs is indicated in Table S2.

Taken together, these data support the conclusion that reducing the function of the Prp31 locus causes light-induced retinal degeneration.

Prp31 mutant photoreceptor cells show increased rhodopsin accumulation

A common cause of retinal degeneration, both in flies and in mammals, is abnormal localisation/levels of the visual pigment rhodopsin1 (Rh1) (Hollingsworth and Gross, 2012; Xiong and Bellen, 2013). Therefore, we asked if the degeneration observed in Prp31 mutant retinas is associated with altered Rh1 localisation/levels. Drosophila Rh1, encoded by ninaE, is the most abundant rhodopsin expressed in the outer PRCs R1-R6 (Harris et al., 1976; Ostroy et al., 1974). In control flies raised under regular light conditions (12 h light, 12 h dark), Rh1 is concentrated in the rhabdomeres. As reported previously, Rh1 either fills the entire rhabdomere, forms a crescent-shaped pattern, or is restricted to the base or the lateral edges of the rhabdomere (Chen et al., 2017; Chinchore et al., 2009; Mitra et al., 2011; Orem et al., 2006; Wang et al., 2014; Xiong et al., 2012). Differences in localisation have been attributed to inconsistencies in antibody penetration due to either the dense packing of microvilli in the rhabdomeres (Xiong et al., 2012) or to a light-induced staining artefact (Schopf et al., 2019). Rh1 staining is observed in rhabdomeres (red arrowheads in Fig. 6A–E) and outlines the rhabdomeric structure along its length (Fig. 6A′–C′). Rh1 could also be detected in cytoplasmic punctae (blue arrowheads in Fig. 6A–E). This intracellular pool of Rh1 presumably represents internalised Rh1 following light exposure (Satoh and Ready, 2005), since these flies were raised under 12 h light/12 h darkness. Strikingly, PRCs of adult flies heterozygous for Prp31 exhibited increased accumulation of Rh1 in the rhabdomeres in comparison to genetic controls (Fig. 6C,C′). Increased Rh1 immunostaining was observed in mutants independent of light conditions applied (Fig. S3). Prp3118 homozygotes exhibited a similar phenotype of enhanced Rh1 immunostaining intensity (Fig. 6E as compared to D). Further, all three deficiencies that remove the Prp31 locus exhibited increased Rh1 staining when heterozygous (Fig. S4B–D) in comparison to the genetic controls (Fig. S4A). Finally, RNAi-mediated knockdown of Prp31 also resulted in increased accumulation of Rh1 immunoreactivity (Fig. S4F) as compared to genetic control (Fig. S4E). Increased intensity of Rh1 immunostaining is due to increased levels of Rh1 as revealed by western blots of protein extracts isolated from adult heads of Prp3118 hetero- and homozygotes. On average, Rh1 levels were significantly increased by over 300% in heads from Prp31P18 heterozygous and by 140% in Prp31P18 homozygous flies as compared to heads of genetic controls (Fig. 6F,G). The variability in the magnitude of increased Rh1 levels (see biological replicates in Fig. 6G) parallels the variability in the degenerative phenotype in the Prp31 mutants.

Fig. 6.
  • Download figure
  • Open in new tab
  • Download powerpoint
Fig. 6.

Reduction of Prp31 results in increased Rhodopsin 1 accumulation. Representative confocal images of 1 µm optical sections from 12 µm cross-sections (A–E), or whole mounts (A′–C′) of eyes of adult males with the genotypes indicated, stained with anti-Rh1. Images were taken using the same settings for mutant conditions and their respective controls. Rhabdomeres are shown in cross sections (A–E) and along their length (A′–C′) with the distal end directed towards the top and the proximal end directed towards the bottom. Red arrowheads indicate Rh1 staining in the rhabdomere and blue arrowheads indicate intracellular Rh1. Rh1 staining is more intense in the rhabdomeric membrane of Prp31P18/+ (C,C′) as compared to controls, w* (A,A′) and w*;; st1/+ (B,B′). Increased Rh1 immunostaining intensity is also observed in Prp31P18/ Prp31P18 (E) as compared to its genetic control (D). Scale bars, 10 µm. (F) Representative western blot for β-Tubulin and Rh1 from head lysates of w*;;Prp31P18 /+ and its genetic control w*;;st1/+, and for w*;;Prp31P18 / Prp31P18 and its genetic control w*;;st1/ st1. Complete genotypes can be found in Table S1. (G) Quantification of Rh1 levels normalised to Tubulin. Graph displays mean±s.e.m. of Rh1 levels calculated from intensity measurements of blots after normalisation compared to that of loading control (Tubulin) with each dot representing one biological replicate. On average, Rh1 levels are increased by 320% (P<0.05, Student's Unpaired t-test) in w*;;Prp31P18/+ as compared to control, w*;;st1/+ and by 140% (P<0.05, Student's Unpaired t-test) in w*;;Prp31P18 / Prp31P18 as compared to its genetic control w*;;st1/ st1. Complete genotypes can be found in Table S1.

Impaired Prp31 function does not affect splicing or abundance of opsin mRNA, but results in increased twinfilin mRNA

To better understand the underlying cause of increased Rh1 in rhabdomeres, we aimed to find out whether ninaE/opsin1 mRNA levels were altered in these mutants. Using Real time qRT-PCR and primers targeting each of the exons (Table 1A) and the exon-intron junctions (Table 1B), no significant change in opsin1 mRNA levels was detected in heads of heterozygous and homozygous Prp31P18 flies. This implies that abundance and splicing of opsin1 mRNA is unaffected in these mutants. We next investigated whether trafficking of opsin/rhodopsin along its biosynthetic route is altered. Carotenoids are precursors of the chromophore 11-cis-3-hydroxyretinal, which binds to opsin to generate the functional visual pigment rhodopsin in flies (von Lintig et al., 2010). Reduction of the chromophore halts endoplasmic reticulum (ER) to Golgi transport and maturation of rhodopsin, resulting in the accumulation of an intermediate form in the perinuclear ER (Colley et al., 1991; Ozaki et al., 1993). Upon supplementation of retinal and induction of its isomerization by blue light, mature Rh1 is now trafficked to the rhabdomere (Satoh et al., 1997). An assay, called blue-light induced chromophore supply (BLICS) (Iwanami et al., 2016; Ozaki et al., 1993), allows to follow Rh1 trafficking along its biosynthetic route. Using the BLICS assay, no qualitative difference was observed in Rh1 reaching the rhabdomere in control and Prp3118 heterozygote flies (Fig. S5). There was no substantial increase in Rh1 reaching the rhabdomere upon its release from the ER. This suggests that the amount of Rh1 produced and trafficked to the rhabdomere (at least via Rab11) was not substantially altered in the Prp31 mutant.

View this table:
  • View inline
  • View popup
  • Download powerpoint
Table 1.

Summary of Real time qRT-PCR data

Finally, we evaluated mRNA levels of three genes (Table 2) that have been recently implicated in Rh1 trafficking (Laffafian and Tepass, 2019). Of these, mRNA levels of only twinfilin (twf), which encodes an actin monomer-binding protein, is increased in Prp31 mutants (heterozygous and homozygous alleles) as compared to those of the respective genetic background. Taken together, impaired Prp31 function is associated with increased rhodopsin protein levels and increased twinfilin mRNA, but does not affect the amount or splicing of opsin mRNA.

View this table:
  • View inline
  • View popup
  • Download powerpoint
Table 2.

Summary of Real time qRT-PCR data

Light-dependent photoreceptor degeneration in Prp31 mutants is suppressed upon elimination of Rh1 accumulation

There is evidence suggesting that increased levels of rhodopsin and/or mis-localised rhodopsin contributes to retinal degeneration, both in mammals and in flies (Hollingsworth and Gross, 2012). In crb mutant retinas, for example, the degree of light-dependent retinal degeneration could be strongly reduced in animals raised on carotenoid-depleted food (Johnson et al., 2002). To determine whether rhodopsin accumulation makes the retina of Prp31 mutant flies prone to light-induced degeneration, we experimentally reduced rhodopsin levels by raising animals in carotenoid-free diet from embryonic stages onward. In fact, in retinas of flies raised under this condition, Rh1 levels are reduced and rhabdomeric localisation of Rh1 is abolished. Instead, Rh1 can now be found perinuclear, both in control and in Prp31/+ retinas (Fig. 7B–C′). In contrast, in the retina of control flies raised on normal food, Rh1 is detected on the lateral edges of the rhabdomeres (Fig. 7A,A′, white arrowheads).

Fig. 7.
  • Download figure
  • Open in new tab
  • Download powerpoint
Fig. 7.

Carotenoid-depleted diet limits the extent of light-induced degeneration in heterozygous Prp31 mutants. (A–C′) Representative images of 1 µm confocal optical sections from 12 µm cryosections of male eyes. Genotypes indicated. Tissues were immunostained for Rh1 (white) and labelled with phalloidin (magenta) and DAPI (green), to stain the rhabdomeres and nuclei, respectively. (A–C) Overlay of two (A) and three (B,C) channels, (A′–C′) images showing the extracted channel (Rh1). Reduction in Rh1 levels and change in its localisation from the rhabdomeres to a peri-nuclear localisation is observed in control (B,B′) and mutant (C,C′) flies fed on a carotenoid-depleted diet (B–C′) as opposed to flies fed on standard food (A–A′). Arrowheads indicate Rh1 localisation in the rhabdomere as opposed to peri-nuclear localisation. Scale bar, 5 µm. (D–F) Representative bright-field images of Toluidine-blue stained semi-thin sections of eyes of w* (D), w*;Prp31P18/+ (E) and w*;;crb11A22 (F) adults. Complete genotypes can be found in Table S1. Animals were raised on a carotenoid-depleted diet. Upon eclosion, they were aged for 2 days under regular light conditions and then subjected to a degeneration paradigm of exposure for 7 days to continuous, high-intensity light. Scale bar, 10 µm. (G) Quantification of retinal degeneration as indicated by the number of surviving rhabdomeres observed upon high intensity, continuous light exposure. Y-axis: percent frequency of ommatidia displaying one to seven rhabdomeres. Genotypes are indicated below. Number on top of each graph indicates the mean percentage of ommatidia displaying the full complement of seven rhabdomeres. Bars represent mean±s.e.m. (a minimum of n=60 ommatidia from eyes of three biological replicates). Statistical significance of differences in this parameter between genotype pairs is indicated in Table S2.

After 7 days of constant light exposure, the overall appearance of the retinae of both the genetic control (w*) and Prp31/+ mutants appeared more damaged as revealed by fewer surviving rhabdomeres and more lacunae (compare the retina of w* in Figs 2 and 7). Note that rhabdomeres are smaller under this dietary condition, a result which is consistent with previous reports (Johnson et al., 2002; Sapp et al., 1991; Satoh et al., 1998). More importantly, the percentage of ommatidia with seven rhabdomeres was the same in heterozygous Prp31P18/+ and control (w*) retinas under carotenoid depletion (Fig. 7D,E and G). A similar result was obtained in crb mutant retinas prepared from flies raised under the same conditions (Fig. 7F,G) (Johnson et al., 2002), supporting the conclusion that carotenoid depletion prevents retinal degeneration.

To conclude, these results suggest that Rh1 accumulation in Prp31 mutant flies makes the retina more susceptible to light-induced degeneration.

DISCUSSION

Here we present a fly model for RP11, an autosomal-dominant human disease caused by mutations in the splicing regulator PRPF31, which leads to blindness in affected patients. Our results reveal that mutations in the Drosophila orthologue Prp31 induce PRC degeneration under light stress, thus mimicking features of RP11-associated symptoms. Similar to those in human, mutations in Drosophila Prp31 are haplo-insufficient and lead to retinal degeneration when heterozygous. This is in stark contrast to mice heterozygous for Prpf31, which did not show any signs of PRC degeneration (Bujakowska et al., 2009), but rather late-onset defects in the retinal pigment epithelium (Farkas et al., 2014; Graziotto et al., 2011).

By using three different genetic approaches we provide convincing evidence that the knockdown of Prp31 is the cause of the retinal degeneration observed. (1) The two Prp31 alleles induced by TILLING (Prp31P17 and Prp31P18) carry missense mutations in conserved amino acids of the coding region, which are predicted to be damaging. (2) Flies heterozygous for any of three deletions, which completely remove the Prp31 locus, exhibit comparable phenotypes as flies heterozygous for Prp31 point mutations. (3) RNAi-mediated knockdown of Prp31 results in light-induced retinal degeneration. The results obtained suggest that the two missense mutations mapped in Prp31P17 and Prp31P18 are strong hypomorphic alleles. First, the two Drosophila alleles characterised here are hemizygous (Prp31/deficiency) and homozygous (in the case of Prp31P18) viable and fertile. Second, mutations in the two established Prp31 fly lines are missense mutations, one located N-terminal to the NOSIC domain in Prp31P17 (G90R) and the other in the Nop domain in Prp31P18 (P277L) (see Fig. 1A), which most likely result in a reduced function of the respective proteins (Fig. S1B). Whether protein levels are also decreased cannot be answered due to the lack of specific antibodies. The mutated amino acid residue in Drosophila Prp31P18 (P277L) lies within the NOP domain. Interestingly, many point mutations in human PRPF31, which are linked to RP11, have been mapped to the Nop domain (Liu and Zack, 2013; Vithana et al., 2001; Wheway et al., 2020) (see ClinVar: https://www.ncbi.nlm.nih.gov/clinvar/?term=PRPF31%5Bgene%5D). Similar as in yeast (Hardin et al., 2015), the Nop domain in human PRPF31 is involved in an essential step in the formation of the U4/U6-U5 tri-snRNP by building a complex of the U4 snRNA and a 15.5K protein, thus stabilising the U4/U6 snRNA junction. The mutated proline in Drosophila Prp31P18 precedes a histidine (H278), which corresponds to amino acid H270 in the human protein (see Fig. S1A). Mutations in H270 in the Nop domain of human PRPF31 result in a reduced affinity of PRPF31 to the complex formed by a stem-loop structure of the U4 snRNA and the 15.5K protein (Liu et al., 2007; Schultz et al., 2006). Therefore, it is tempting to speculate that the Drosophila P277L mutation could similarly weaken, but not abolish the corresponding interaction of the mutant Prp31 protein with the U4/U6 complex. Further experiments are required to determine the functional consequences of the molecular lesions identified in Drosophila PRP31.

We noticed that the retinal phenotype observed upon reduction of Prp31 is more variable than that observed upon loss of crb (see, for example, Fig. 2E) (Johnson et al., 2002; Spannl et al., 2017). This could be due to the fact that all Prp31 conditions analysed represent hypomorphic conditions, possibly retaining some residual protein function(s). However, the expressivity of the mutant phenotype is not increased in Prp31/deficiency flies (carrying only one mutant copy) in comparison to that of Prp31/+ flies, which carry one mutant and one wild-type allele. Interestingly, human RP11 patients heterozygous for mutations in Prpf31 show an unusually high degree of phenotypic non-penetrance and can even be asymptomatic. Various causes have been uncovered to explain this feature (Wheway et al., 2020). These include a highly variable expression level of the remaining wild-type Prpf31 allele, possibly due to changes in the expression levels of trans-acting regulators (Rio Frio et al., 2008) (reviewed in Rose and Bhattacharya, 2016). In addition, mutant PRPF31 proteins can form cytoplasmic aggregates in RPE cells, thus reducing the amount of protein entering the nucleus (Valdes-Sanchez et al., 2019), or can impair overall transcription or splicing, as described in Prpf31 zebrafish models (Linder et al., 2011; Yin et al., 2011). Finally, mutations in unlinked genes have been suggested to modify the disease severity of patients (Venturini et al., 2012).

Not only in flies, but also in human, mutations in PRPF31 affect only the retina, despite the importance of this splicing regulator in all cells. Recently published data show that impaired PRPF31 function can affect the splicing of target genes in a cell-type specific manner. Strikingly, retinal cells isolated from RP11 patient-derived retinal organoids exhibit mis-splicing of genes that encode components of the splicing machinery itself. This was not observed in fibroblasts or iPS cells derived from the same patients (Buskin et al., 2018). These authors obtained similar results from the retina and the RPE of Prpf31/+ mice. Mutant RPE cells additionally revealed splicing defects in transcripts of genes with functions in ciliogenesis, cell polarity and cellular adhesion (Buskin et al., 2018), which could explain the previously described RPE defects in these mice (Farkas et al., 2014; Graziotto et al., 2011; Hamieh and Nandrot, 2019).

In the retina of flies lacking one functional copy of Prp31, PRCs showed increased levels of Rh1, both in the rhabdomeres and in the cytoplasm, as revealed by immunostaining and confirmed by western blot analysis. However, increased Rh1 levels did not affect rhabdomere size or structure. This is in contrast to results obtained in the mouse, where transgenic overexpression of either wild-type bovine or human rhodopsin induced an increase in outer segment volume of rod PRCs (Price et al., 2012; Wen et al., 2009). In several other Drosophila mutants, accumulation of Rh1 in endocytic compartments has been suggested to cause retinal degeneration due to its toxicity. For example, dominant mutations in Drosophila ninaE result in ER accumulation of misfolded Rh1 due to impaired protein maturation. This, in turn, causes an overproduction of ER cisternae and induces the unfolded protein response (UPR), which eventually leads to apoptosis of PRCs, both in flies and in mammals (Colley et al., 1995; Kroeger et al., 2018; Zhang et al., 2014).

Interestingly, mis-localisation of rhodopsin in human PRCs to sites other than the outer segment is a common characteristic of adRP induced by mutations in rhodopsin and is considered to contribute to the pathological severity (Hollingsworth and Gross, 2012). Our data suggest that increased accumulation of rhodopsin contributes to degeneration in Prp31 mutant retinas. Reduction of Rh1 by depletion of dietary carotenoid not only obliterated increased Rh1 immunoreactivity and opsin retention in perinuclear compartments in Prp31 mutants, but also reduced the degree of PRC degeneration. However, whether increased Rh1 accumulation in the rhabdomere or in the cytoplasm contributes to light-dependent PRC degeneration of Prp31 mutant flies needs to be explored in the future.

Our data further suggest that Prp31 regulates, directly or indirectly, Rh1 levels at a posttranscriptional level, since no increase of RNA levels was detected in heads of Prp31/+ flies. This result is different from that obtained in primary murine retinal cell cultures, where expression of a mutant Prpf31 gene reduced rhodopsin expression, as a result of impaired splicing of the rhodopsin pre-mRNA (Yuan et al., 2005). Similarly, siRNA-mediated knockdown of PRPF31 function in human organotypic retinal flat-mount cultures (HORFC) reduced mRNAs encoding genes involved in phototransduction and photoreceptor structure, including rhodopsin (Azizzadeh Pormehr et al., 2018). Interestingly, the Prp31 mutants described here show increased mRNA levels of an evolutionary conserved actin monomer binding protein called twinfilin (twf), which inhibits actin polymerisation. Knockdown of twf results in excessive cytoplasmic Rh1 staining, suggesting defects in its trafficking (Laffafian and Tepass, 2019). In Prp31 mutants, an increase in rhabdomeric Rh1 was observed as well as increased twf mRNA. From this correlation we hypothesise that upregulation of twf mRNA in Prp31 might be in part responsible for at least the rhabdomeric Rh1 accumulation. Rh1 also accumulates in the cytoplasm of Prp31 mutant PRCs. Our data exclude the role of Rab11-mediated targeting of Rh1 in this accumulation. Now, it remains to be determined if the deregulation of other trafficking routes or the upregulation of twf contributes to the increased Rh1 in the cytoplasm. In the future, it may be interesting to explore the link between increased Rh1 levels as observed in Drosophila Prp31 mutants, increased mRNA levels of twinfilin and impaired Rh1 trafficking. Additionally, a detailed transcriptome analysis should elucidate possible defects in transcription and/or splicing of target genes, thus also allowing a better understanding of the aetiology of the human disease.

MATERIALS AND METHODS

Fly strains and genetics

All phenotypic analyses were performed in age-matched males unless otherwise specified. Genotypes are summarised in Table S1. Flies were maintained at 25°C on standard yeast-cornmeal-agar food unless otherwise stated. To rule out differences in light sensitivity in the light-degeneration paradigm, we used white-eyed flies, bearing mutations in the white gene, both as general controls and in the respective mutant background. The white allele (w*) used here was tested by PCR and shown to carry a deletion that includes the transcription and translation start site of the white gene (data not provided). Loss of scarlet (st) function was rescued by Gal4-mediated expression of a scarlet transgene (Cunningham et al., 2018) in all cells of the retina using GMR-Gal4 (Hay et al., 1994). The RNAi line (ID: 35131) for the Prp31 gene was obtained from the Vienna Drosophila Resource Centre (VDRC, www.vdrc.at) (Dietzl et al., 2007). RNAi was induced using Rh1-Gal4 (Lee and Carthew, 2003) in combination with Dicer-2 expression and concomitant expression of white RNAi under the control of the GMR-promoter (GMR-wIR) (Lee and Carthew, 2003) allowing assay of degeneration in a non-pigmented background. Df(3L)Exel6262 with deleted segment 71B3;71C1 (Parks et al., 2004), Df(3L)ED217 with deleted segment 70F4;71E1 and Df(3L)ED218 with deleted segment 71B1–71E1 (Ryder et al., 2007) were obtained from the Bloomington Stock Centre. Since the deficiency lines carry a mini-white transgene due the way they were generated (Ryder et al., 2007), cn bw was recombined into these lines to obtain white-eyed flies and all phenotypes were compared with cn bw.

Isolation of Prp31 alleles by TILLING

To isolate point mutations in the Prp31 locus (FlyBase ID: FBgn0036487) a library of 2.400 fly lines with isogenised third chromosomes, which potentially carry point mutations caused by EMS treatment, was screened (Winkler et al., 2005). Our approach targeted exon 1–3 of the Prp31 locus containing two thirds (67%) of the coding sequence, which includes several predicted functional domains (the NOSIC (IPRO012976), the Nop (IPRO002687) and parts of the Prp31_C terminal (IPRO019175) domain), making use of two different PCR amplicons. A nested PCR approach was used, where the inner primers contain universal M13 tails that serve as primer binding sites of the Sanger sequencing reaction:

  • amplicon1 (covers exon 1 and 2), outer primer, forward: TTCAATGAACCGCATGG, reverse: GTCGATCTTTGCCTTCTCC, inner / nested primer, forward: TGTAAAACGA CGGCCAGT-AGCAACGGTCACTTCAATTC, reverse: AGGAAACAGCTATGACCAT-GAAAGGGAATGGGATTCAG);

  • amplicon 2 (covers exon 3), outer primer, forward: ATCGTGGGTGAAATCGAG, reverse: TGGTCTTCTCATCCACCTG, inner / nested primer, forward: TGTAAAACGA CGGCCAGT-AAGCTGCAGGCTATTCTCAC, reverse: AGGAAACAGCTATGACCAT-TAGGCATCCTCTTCGATCTG.

PCR-reactions were performed in 10 µl volume and with an annealing temperature of 57°C, in 384-well format, making use of automated liquid handling tools. PCR fragments were sequenced by Sanger sequencing optimised for amplicon re-sequencing in a large-scale format (Winkler et al., 2011, 2005). Primary hits, resembling sequence variants, which result in potential nonsense and missense mutations upon translation or affect a predicted splice site, were verified in independent PCR amplification and Sanger sequencing reactions. Two of the four lines, named Prp31P17 and Prp31P18, were recovered from the living fly library and crossed for three generations to control, w* flies to reduce the number of accompanying sequence variations. The removal of the markers of the original, mutagenised chromosome (ru st e ca) by the above outcrossing was verified as follows: the isolated alleles (males) were crossed to the original line (ru st e ca) and checked for the phenotypes associated with homozygous conditions of roughoid (ru; eye appearance), scarlet (st; eye colour), ebony (e; body colour), claret (ca; eye colour).

Experimental light conditions

Flies were reared in regular light conditions defined as 12 h of light (approximately 900–1300 lux)/12 h of darkness. For the light-induced degeneration setting, flies (2 days of age) were placed at 25°C for 7 days in an incubator dedicated for continuous, high intensity light exposure (Johnson et al., 2002). High intensity light was defined by 1200–1300 lux measured using an Extech 403125 Light ProbeMeter (Extech Instruments, USA) with the detector placed immediately adjacent to the vial and facing the nearest light source. At the end of 7 days, fly heads were processed for sectioning. For immunostaining and western blotting, flies (1 day) reared under regular light were processed as described below.

Vitamin A depletion, BLICS assay

For vitamin A depletion experiments, animals were raised and maintained from embryonic stages onward on carotenoid free food (10% dry yeast, 10% sucrose, 0.02% cholesterol, and 2% agar) as described (Pocha et al., 2011). The protocol for BLICS assays was as described in Hebbar et al., 2020 doi: https://doi.org/10.1083/jcb.201911100). Briefly, flies were raised on carotenoid free food and upon eclosion, they were placed in the dark containing carotenoid-free food or food supplemented with all-trans retinal. After 48 h, flies were exposed to blue light and after 180 min, fly heads were quickly dissected and put in fixative for cryosectioning and immunostaining. At least two regions of interest (comprising at least 6–8 ommatidia) from three biological replicates (fly eyes) were used for quantification.

Quantification of Degeneration

Toluidine blue stained semi-thin sections were imaged with a 63x Plan Apo oil objective (N.A. =1.4) on AxioImager.Z1 (Zeiss, Germany), fitted with AxioCamMRm camera, and analysed using the AxioVision software (Release 4.7). Quantification of degeneration was performed as described (Bulgakova et al., 2010). Briefly, the number of detectable rhabdomeres in each ommatidium was recorded from approximately 60–80 ommatidia per section and at least three eyes from different individuals were analysed. In case of degeneration, fewer ommatidia were counted since most of the tissue had degenerated.

Immunostaining of adult retina and confocal imaging

Adult eyes were dissected and fixed in 4% formaldehyde. They were then processed either directly for immunostaining of the whole eye after removal of the lens, or for cryosectioning. For sectioning, sucrose treatment and embedding of the tissues in Richard-Allan Scientific NEG-50TM (Thermo Fisher Scientific, UK) tissue embedding medium was done (Mishra and Knust, 2019). Eyes were cryosectioned at 12 µm thickness at -21°C. Sections were air-dried and then subjected to immunostaining, which was done as described previously (Spannl et al., 2017). Antibodies used were mouse anti-Rh 1 (1:50; 4C5) from Developmental Studies Hybridoma Bank (DSHB), University of Iowa, IA, USA. 4C5 [http://dshb.biology.uiowa.edu/4C5] was deposited to the DSHB by de Coet, H.G./Tanimura, T., and by Fambrough, D.M., and anti-Rab11 (1:1000; Satoh et al., 2005; kind gift of D. Ready). Alexa-Fluor conjugated secondary antibodies (1:200, Thermo Fisher Scientific, UK) were used. DAPI (4′,6-Diamidino-2-Phenylindole, Dihydrochloride; Thermo Fisher Scientific, UK) was used to label nuclei in tissue sections and Alexa-Fluor-555–phalloidin (Thermo Fisher Scientific, UK) was used to visualise F-actin enriched rhabdomeres. Sections and whole mounts were imaged with an Olympus Fluoview 1000 confocal microscope using an Olympus UPlanSApochromat 60x Oil objective (N.A. =1.35). They were subsequently visualised in Fiji (Schindelin et al., 2012) and corrected for brightness and contrast only.

Western blotting

Five fly heads from each genotype were homogenised in 10 µl of 4× SDS-PAGE sample buffer (200 mM Tris-HCl pH 6.8, 20% Glycerol, 8% SDS, 0.04% Bromophenol blue, 400 mM DTT). After dilution with RIPA buffer (150 mM sodium chloride, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, 50 mM Tris pH 8), lysates were heated at 37°C for 30 min. Lysates equivalent to 2.5 heads were loaded and run on a 15% acrylamide gel, and proteins were transferred onto a membrane (Nitrocellulose Blotting Membrane 10600002; GE Healthcare Life Sciences, PA, USA). Primary antibodies were incubated overnight at 4°C and included (1) anti-Rh1 [4C5 (http://dshb.biology.uiowa.edu/4C5); 1:500], from Developmental Studies Hybridoma Bank (DSHB), University of Iowa, IA, USA, deposited by de Coet, H.G./Tanimura, T. and (2) anti-α-Tubulin (T6074; 1:5000) from Sigma-Aldrich. As secondary antibody IRDye 800CW goat anti-Mouse IgG (1:15,000; LI-COR Biotechnology, NE, USA) was used for a 1 h incubation at room temperature. The fluorescent signal from the dry membrane was measured using a LI-COR Odyssey Sa Infrared Imaging System 9260-11P (LI-COR Biotechnology). The intensity of the bands was analysed using the Image Studio Ver 4.0 software. The reported value in Fig. 6 was obtained following normalisation of the intensity values for Rh1 with the corresponding Tubulin intensity values and the number of heads loaded onto the gel.

Real Time qRT-PCR analyses

RNA extraction, cDNA generation, qPCR analyses were performed as described in Hebbar et al. (2020) (doi: https://doi.org/10.1083/jcb.201911100). Primers are listed in Table S3.

Figure panel preparation

All figure panels were assembled using Adobe Photoshop CS5.1 or Adobe Illustrator CS3 (Adobe Systems, USA). Statistical analyses and graphs were generated using GraphPad Prism (GraphPad Software, Inc, USA) and Microsoft Excel. For protein sequence visualisation, Illustrator of Biological Sequences (IBS; Liu et al., 2015) software package was used.

Acknowledgements

We would like to thank the Bloomington Stock Centre for fly stocks and the Developmental Studies Hybridoma Bank (DSHB) for antibodies. This work was supported by the fly facility, the light and electron microscopy facility and the sequencing facility of MPI-CBG. We thank K. Kapp (University of Kassel, Germany) for technical advice on western blotting procedures.

Footnotes

  • Competing interests

    The authors declare no competing or financial interests.

  • Author contributions

    Conceptualization: S.H., M.L., E.K.; Methodology: S.H., M.L., S.B., C.H., W.L., M.Y., S.W., E.K.; Validation: S.H., S.W., E.K.; Formal analysis: S.H., M.L., S.W.; Investigation: S.H., S.B., C.H., W.L., M.Y.; Resources: E.K.; Data curation: S.H., S.W.; Writing - original draft: S.H., M.L., E.K.; Writing - review & editing: S.H., E.K.; Visualization: S.H.; Supervision: E.K.; Project administration: E.K.; Funding acquisition: E.K.

  • Funding

    This work was funded by the Max-Planck Society.

  • Supplementary information

    Supplementary information available online at https://bio.biologists.org/lookup/doi/10.1242/bio.052332.supplemental

  • Received May 26, 2020.
  • Accepted December 1, 2020.
  • © 2021. Published by The Company of Biologists Ltd
http://creativecommons.org/licenses/by/4.0

This is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution and reproduction in any medium provided that the original work is properly attributed.

References

  1. ↵
    1. Adzhubei, I.,
    2. Jordan, D. M. and
    3. Sunyaev, S. R.
    (2013). Predicting functional effect of human missense mutations using PolyPhen-2. Curr. Protoc. Hum. Genet. Chapter 7, Unit7 20. doi:10.1002/0471142905.hg0720s76
    OpenUrlCrossRef
  2. ↵
    1. Ali, M. U.,
    2. Rahman, M. S. U.,
    3. Cao, J. and
    4. Yuan, P. X.
    (2017). Genetic characterization and disease mechanism of retinitis pigmentosa; current scenario. 3 Biotech 7, 251. doi:10.1007/s13205-017-0878-3
    OpenUrlCrossRef
  3. ↵
    1. Azizzadeh Pormehr, L.,
    2. Daftarian, N.,
    3. Ahmadian, S.,
    4. Rezaei Kanavi, M.,
    5. Ahmadieh, H. and
    6. Shafiezadeh, M.
    (2018). Human organotypic retinal flat-mount culture (HORFC) as a model for retinitis pigmentosa11. J. Cell. Biochem. 119, 6775-6783. doi:10.1002/jcb.26871
    OpenUrlCrossRef
  4. ↵
    1. Bujakowska, K.,
    2. Maubaret, C.,
    3. Chakarova, C. F.,
    4. Tanimoto, N.,
    5. Beck, S. C.,
    6. Fahl, E.,
    7. Humphries, M. M.,
    8. Kenna, P. F.,
    9. Makarov, E.,
    10. Makarova, O. et al.
    (2009). Study of gene-targeted mouse models of splicing factor gene Prpf31 implicated in human autosomal dominant retinitis pigmentosa (RP). Invest. Ophthalmol. Vis. Sci. 50, 5927-5933. doi:10.1167/iovs.08-3275
    OpenUrlAbstract/FREE Full Text
  5. ↵
    1. Bulgakova, N. A.,
    2. Rentsch, M. and
    3. Knust, E.
    (2010). Antagonistic functions of two Stardust isoforms in Drosophila photoreceptor cells. Mol. Biol. Cell 21, 3915-3925. doi:10.1091/mbc.e09-10-0917
    OpenUrlAbstract/FREE Full Text
  6. ↵
    1. Buskin, A.,
    2. Zhu, L.,
    3. Chichagova, V.,
    4. Basu, B.,
    5. Mozaffari-Jovin, S.,
    6. Dolan, D.,
    7. Droop, A.,
    8. Collin, J.,
    9. Bronstein, R.,
    10. Mehrotra, S. et al.
    (2018). Disrupted alternative splicing for genes implicated in splicing and ciliogenesis causes PRPF31 retinitis pigmentosa. Nat. Commun. 9, 4234. doi:10.1038/s41467-018-06448-y
    OpenUrlCrossRefPubMed
  7. ↵
    1. Chartier, F. J.-M.,
    2. Hardy, E. J.-L. and
    3. Laprise, P.
    (2012). Crumbs limits oxidase-dependent signaling to maintain epithelial integrity and prevent photoreceptor cell death. J. Cell Biol. 198, 991-998. doi:10.1083/jcb.201203083
    OpenUrlAbstract/FREE Full Text
  8. ↵
    1. Chen, X.,
    2. Hall, H.,
    3. Simpson, J. P.,
    4. Leon-Salas, W. D.,
    5. Ready, D. F. and
    6. Weake, V. M.
    (2017). Cytochrome b5 protects photoreceptors from light stress-induced lipid peroxidation and retinal degeneration. NPJ Aging Mech. Dis. 3, 18. doi:10.1038/s41514-017-0019-6
    OpenUrlCrossRef
  9. ↵
    1. Chinchore, Y.,
    2. Mitra, A. and
    3. Dolph, P. J.
    (2009). Accumulation of rhodopsin in late endosomes triggers photoreceptor cell degeneration. PLoS Genet. 5, e1000377. doi:10.1371/journal.pgen.1000377
    OpenUrlCrossRefPubMed
  10. ↵
    1. Colley, N. J.,
    2. Baker, E. K.,
    3. Stamnes, M. A. and
    4. Zuker, C. S.
    (1991). The cyclophilin homolog ninaA is required in the secretory pathway. Cell 67, 255-263. doi:10.1016/0092-8674(91)90177-Z
    OpenUrlCrossRefPubMedWeb of Science
  11. ↵
    1. Colley, N. J.,
    2. Cassill, J. A.,
    3. Baker, E. K. and
    4. Zuker, C. S.
    (1995). Defective intracellular transport is the molecular basis of rhodopsin-dependent dominant retinal degeneration. Proc. Natl. Acad. Sci. USA 92, 3070-3074. doi:10.1073/pnas.92.7.3070
    OpenUrlAbstract/FREE Full Text
  12. ↵
    1. Cunningham, P. C.,
    2. Waldeck, K.,
    3. Ganetzky, B. and
    4. Babcock, D. T.
    (2018). Neurodegeneration and locomotor dysfunction in Drosophila scarlet mutants. J. Cell Sci. 131, jcs216697. doi:10.1242/jcs.216697
    OpenUrlAbstract/FREE Full Text
  13. ↵
    1. Daiger, S. P.,
    2. Sullivan, L. S. and
    3. Bowne, S. J.
    (2013). Genes and mutations causing retinitis pigmentosa. Clin. Genet. 84, 132-141. doi:10.1111/cge.12203
    OpenUrlCrossRefPubMed
  14. ↵
    1. Daiger, S. P.,
    2. Bowne, S. J. and
    3. Sullivan, L. S.
    (2014). Genes and mutations causing autosomal dominant retinitis pigmentosa. Cold Spring Harb. Perspect Med. 5, a017129. doi:10.1101/cshperspect.a017129
    OpenUrlAbstract/FREE Full Text
  15. ↵
    1. Dickinson, M. E.,
    2. Flenniken, A. M.,
    3. Ji, X.,
    4. Teboul, L.,
    5. Wong, M. D.,
    6. White, J. K.,
    7. Meehan, T. F.,
    8. Weninger, W. J.,
    9. Westerberg, H.,
    10. Adissu, H. et al.
    (2016). High-throughput discovery of novel developmental phenotypes. Nature 537, 508-514. doi:10.1038/nature19356
    OpenUrlCrossRefPubMed
  16. ↵
    1. Dietzl, G.,
    2. Chen, D.,
    3. Schnorrer, F.,
    4. Su, K. C.,
    5. Barinova, Y.,
    6. Fellner, M.,
    7. Gasser, B.,
    8. Kinsey, K.,
    9. Oppel, S.,
    10. Scheiblauer, S. et al.
    (2007). A genome-wide transgenic RNAi library for conditional gene inactivation in Drosophila. Nature 448, 151-156. doi:10.1038/nature05954
    OpenUrlCrossRefPubMedWeb of Science
  17. ↵
    1. Farkas, M. H.,
    2. Lew, D. S.,
    3. Sousa, M. E.,
    4. Bujakowska, K.,
    5. Chatagnon, J.,
    6. Bhattacharya, S. S.,
    7. Pierce, E. A. and
    8. Nandrot, E. F.
    (2014). Mutations in pre-mRNA processing factors 3, 8, and 31 cause dysfunction of the retinal pigment epithelium. Am. J. Pathol. 184, 2641-2652. doi:10.1016/j.ajpath.2014.06.026
    OpenUrlCrossRefPubMed
  18. ↵
    1. Graziotto, J. J.,
    2. Farkas, M. H.,
    3. Bujakowska, K.,
    4. Deramaudt, B. M.,
    5. Zhang, Q.,
    6. Nandrot, E. F.,
    7. Inglehearn, C. F.,
    8. Bhattacharya, S. S. and
    9. Pierce, E. A.
    (2011). Three gene-targeted mouse models of RNA splicing factor RP show late-onset RPE and retinal degeneration. Invest. Ophthalmol. Vis. Sci. 52, 190-198. doi:10.1167/iovs.10-5194
    OpenUrlAbstract/FREE Full Text
  19. ↵
    1. Hamieh, A. and
    2. Nandrot, E. F.
    (2019). Retinal pigment epithelial cells: the unveiled component in the etiology of Prpf splicing factor-associated retinitis pigmentosa. Adv. Exp. Med. Biol. 1185, 227-231. doi:10.1007/978-3-030-27378-1_37
    OpenUrlCrossRef
  20. ↵
    1. Hardin, J. W.,
    2. Warnasooriya, C.,
    3. Kondo, Y.,
    4. Nagai, K. and
    5. Rueda, D.
    (2015). Assembly and dynamics of the U4/U6 di-snRNP by single-molecule FRET. Nucleic Acids Res. 43, 10963-10974. doi:10.1093/nar/gkv1011
    OpenUrlCrossRefPubMed
  21. ↵
    1. Harris, W. A.,
    2. Stark, W. S. and
    3. Walker, J. A.
    (1976). Genetic dissection of the photoreceptor system in the compound eye of Drosophila melanogaster. J. Physiol. 256, 415-439. doi:10.1113/jphysiol.1976.sp011331
    OpenUrlCrossRefPubMedWeb of Science
  22. ↵
    1. Hay, B. A.,
    2. Wolff, T. and
    3. Rubin, G. M.
    (1994). Expression of baculovirus P35 prevents cell death in Drosophila. Development 120, 2121-2129.
    OpenUrlAbstract
  23. ↵
    1. Hebbar, S.,
    2. Schuhmann, K.,
    3. Shevchenko, A. and
    4. Knust, E.
    (2020). Hydroxylated sphingolipid biosynthesis regulates photoreceptor apical domain morphogenesis. J. Cell Biol. 219, e201911100. doi:10.1083/jcb.201911100
  24. ↵
    1. Hollingsworth, T. J. and
    2. Gross, A. K.
    (2012). Defective trafficking of rhodopsin and its role in retinal degenerations. Int. Rev. Cell Mol. Biol. 293, 1-44. doi:10.1016/B978-0-12-394304-0.00006-3
    OpenUrlCrossRefPubMed
  25. ↵
    1. Iwanami, N.,
    2. Nakamura, Y.,
    3. Satoh, T.,
    4. Liu, Z. and
    5. Satoh, A. K.
    (2016). Rab6 is required for multiple apical transport pathways but not the basolateral transport pathway in Drosophila photoreceptors. PLoS Genet. 12, e1005828. doi:10.1371/journal.pgen.1005828
    OpenUrlCrossRefPubMed
  26. ↵
    1. Johnson, K.,
    2. Grawe, F.,
    3. Grzeschik, N. and
    4. Knust, E.
    (2002). Drosophila Crumbs is required to inhibit light-induced photoreceptor degeneration. Curr. Biol. 12, 1675-1680. doi:10.1016/S0960-9822(02)01180-6
    OpenUrlCrossRefPubMedWeb of Science
  27. ↵
    1. Kroeger, H.,
    2. Chiang, W. C.,
    3. Felden, J.,
    4. Nguyen, A. and
    5. Lin, J. H.
    (2018). ER stress and unfolded protein response in ocular health and disease. FEBS J. 286, 399-412. doi:10.1111/febs.14522
    OpenUrlCrossRef
  28. ↵
    1. Kumar, J. P. and
    2. Ready, D. F.
    (1995). Rhodopsin plays an essential structural role in Drosophila photoreceptor development. Development 121, 4359-4370.
    OpenUrlAbstract
  29. ↵
    1. Laffafian, A. and
    2. Tepass, U.
    (2019). Identification of genes required for apical protein trafficking in Drosophila photoreceptor cells. G3 (Bethesda) 9, 4007-4017. doi:10.1534/g3.119.400635
    OpenUrlAbstract/FREE Full Text
  30. ↵
    1. Lee, Y. S. and
    2. Carthew, R. W.
    (2003). Making a better RNAi vector for Drosophila: use of intron spacers. Methods 30, 322-329. doi:10.1016/S1046-2023(03)00051-3
    OpenUrlCrossRefPubMedWeb of Science
  31. ↵
    1. Linder, B.,
    2. Dill, H.,
    3. Hirmer, A.,
    4. Brocher, J.,
    5. Lee, G. P.,
    6. Mathavan, S.,
    7. Bolz, H. J.,
    8. Winkler, C.,
    9. Laggerbauer, B. and
    10. Fischer, U.
    (2011). Systemic splicing factor deficiency causes tissue-specific defects: a zebrafish model for retinitis pigmentosa. Hum. Mol. Genet. 20, 368-377. doi:10.1093/hmg/ddq473
    OpenUrlCrossRefPubMedWeb of Science
  32. ↵
    1. Liu, M. M. and
    2. Zack, D. J.
    (2013). Alternative splicing and retinal degeneration. Clin. Genet. 84, 142-149. doi:10.1111/cge.12181
    OpenUrlCrossRefPubMed
  33. ↵
    1. Liu, S.,
    2. Li, P.,
    3. Dybkov, O.,
    4. Nottrott, S.,
    5. Hartmuth, K.,
    6. Luhrmann, R.,
    7. Carlomagno, T. and
    8. Wahl, M. C.
    (2007). Binding of the human Prp31 Nop domain to a composite RNA-protein platform in U4 snRNP. Science 316, 115-120. doi:10.1126/science.1137924
    OpenUrlAbstract/FREE Full Text
  34. ↵
    1. Liu, W.,
    2. Xie, Y.,
    3. Ma, J.,
    4. Luo, X.,
    5. Nie, P.,
    6. Zuo, Z.,
    7. Lahrmann, U.,
    8. Zhao, Q.,
    9. Zheng, Y.,
    10. Zhao, Y. et al.
    (2015). IBS: an illustrator for the presentation and visualization of biological sequences. Bioinformatics 31, 3359-3361. doi:10.1093/bioinformatics/btv362
    OpenUrlCrossRefPubMed
  35. ↵
    1. Maita, H.,
    2. Kitaura, H.,
    3. Keen, T. J.,
    4. Inglehearn, C. F.,
    5. Ariga, H. and
    6. Iguchi-Ariga, S. M.
    (2004). PAP-1, the mutated gene underlying the RP9 form of dominant retinitis pigmentosa, is a splicing factor. Exp. Cell Res. 300, 283-296. doi:10.1016/j.yexcr.2004.07.029
    OpenUrlCrossRefPubMedWeb of Science
  36. ↵
    1. Mishra, M. and
    2. Knust, E.
    (2019). Analysis of the Drosophila compound eye with light and electron microscopy. Methods Mol. Biol. 1834, 345-364. doi:10.1007/978-1-4939-8669-9_22
    OpenUrlCrossRef
  37. ↵
    1. Mitra, A.,
    2. Chinchore, Y.,
    3. Kinser, R. and
    4. Dolph, P. J.
    (2011). Characterization of two dominant alleles of the major rhodopsin-encoding gene ninaE in Drosophila. Mol. Vis. 17, 3224-3233.
    OpenUrlPubMed
  38. ↵
    1. Mordes, D.,
    2. Luo, X.,
    3. Kar, A.,
    4. Kuo, D.,
    5. Xu, L.,
    6. Fushimi, K.,
    7. Yu, G.,
    8. Sternberg, P., Jr.. and
    9. Wu, J. Y.
    (2006). Pre-mRNA splicing and retinitis pigmentosa. Mol. Vis. 12, 1259-1271.
    OpenUrlPubMedWeb of Science
  39. ↵
    1. Nemet, I.,
    2. Ropelewski, P. and
    3. Imanishi, Y.
    (2015). Rhodopsin Trafficking and Mistrafficking: Signals, Molecular Components, and Mechanisms. Prog. Mol. Biol. Transl. Sci. 132, 39-71. doi:10.1016/bs.pmbts.2015.02.007
    OpenUrlCrossRefPubMed
  40. ↵
    1. Nguyen, T. H.,
    2. Galej, W. P.,
    3. Bai, X. C.,
    4. Savva, C. G.,
    5. Newman, A. J.,
    6. Scheres, S. H. and
    7. Nagai, K.
    (2015). The architecture of the spliceosomal U4/U6.U5 tri-snRNP. Nature 523, 47-52. doi:10.1038/nature14548
    OpenUrlCrossRefPubMed
  41. ↵
    1. Orem, N. R.,
    2. Xia, L. and
    3. Dolph, P. J.
    (2006). An essential role for endocytosis of rhodopsin through interaction of visual arrestin with the AP-2 adaptor. J. Cell Sci. 119, 3141-3148. doi:10.1242/jcs.03052
    OpenUrlAbstract/FREE Full Text
  42. ↵
    1. Ostroy, S. E.,
    2. Wilson, M. and
    3. Pak, W. L.
    (1974). Drosophila rhodopsin: photochemistry, extraction and differences in the norp AP12 phototransduction mutant. Biochem. Biophys. Res. Commun. 59, 960-966. doi:10.1016/S0006-291X(74)80073-2
    OpenUrlCrossRefPubMedWeb of Science
  43. ↵
    1. Ozaki, K.,
    2. Nagatani, H.,
    3. Ozaki, M. and
    4. Tokunaga, F.
    (1993). Maturation of major Drosophila rhodopsin, ninaE, requires chromophore 3-hydroxyretinal. Neuron 10, 1113-1119. doi:10.1016/0896-6273(93)90059-Z
    OpenUrlCrossRefPubMedWeb of Science
  44. ↵
    1. Parks, A. L.,
    2. Cook, K. R.,
    3. Belvin, M.,
    4. Dompe, N. A.,
    5. Fawcett, R.,
    6. Huppert, K.,
    7. Tan, L. R.,
    8. Winter, C. G.,
    9. Bogart, K. P.,
    10. Deal, J. E. et al.
    (2004). Systematic generation of high-resolution deletion coverage of the Drosophila melanogaster genome. Nat. Genet. 36, 288-292. doi:10.1038/ng1312
    OpenUrlCrossRefPubMedWeb of Science
  45. ↵
    1. Patel, S. B. and
    2. Bellini, M.
    (2008). The assembly of a spliceosomal small nuclear ribonucleoprotein particle. Nucleic Acids Res. 36, 6482-6493. doi:10.1093/nar/gkn658
    OpenUrlCrossRefPubMedWeb of Science
  46. ↵
    1. Pocha, S. M.,
    2. Shevchenko, A. and
    3. Knust, E.
    (2011). Crumbs regulates rhodopsin transport by interacting with and stabilizing myosin V. J. Cell Biol. 195, 827-838. doi:10.1083/jcb.201105144
    OpenUrlAbstract/FREE Full Text
  47. ↵
    1. Poulos, M. G.,
    2. Batra, R.,
    3. Charizanis, K. and
    4. Swanson, M. S.
    (2011). Developments in RNA splicing and disease. Cold Spring Harb. Perspect Biol. 3, a000778. doi:10.1101/cshperspect.a000778
    OpenUrlAbstract/FREE Full Text
  48. ↵
    1. Price, B. A.,
    2. Sandoval, I. M.,
    3. Chan, F.,
    4. Nichols, R.,
    5. Roman-Sanchez, R.,
    6. Wensel, T. G. and
    7. Wilson, J. H.
    (2012). Rhodopsin gene expression determines rod outer segment size and rod cell resistance to a dominant-negative neurodegeneration mutant. PLoS ONE 7, e49889. doi:10.1371/journal.pone.0049889
    OpenUrlCrossRefPubMed
  49. ↵
    1. Ray, P.,
    2. Luo, X.,
    3. Rao, E. J.,
    4. Basha, A.,
    5. Woodruff, E. A., III. and
    6. Wu, J. Y.
    (2010). The splicing factor Prp31 is essential for photoreceptor development in Drosophila. Protein Cell 1, 267-274. doi:10.1007/s13238-010-0035-9
    OpenUrlCrossRefPubMed
  50. ↵
    1. Rio Frio, T.,
    2. Civic, N.,
    3. Ransijn, A.,
    4. Beckmann, J. S. and
    5. Rivolta, C.
    (2008). Two trans-acting eQTLs modulate the penetrance of PRPF31 mutations. Hum. Mol. Genet. 17, 3154-3165. doi:10.1093/hmg/ddn212
    OpenUrlCrossRefPubMedWeb of Science
  51. ↵
    1. Rose, A. M. and
    2. Bhattacharya, S. S.
    (2016). Variant haploinsufficiency and phenotypic non-penetrance in PRPF31-associated retinitis pigmentosa. Clin. Genet. 90, 118-126. doi:10.1111/cge.12758
    OpenUrlCrossRef
  52. ↵
    1. Ruzickova, S. and
    2. Stanek, D.
    (2016). Mutations in spliceosomal proteins and retina degeneration. RNA Biol. 14, 1-9. doi:10.1080/15476286.2016.1191735
    OpenUrlCrossRefPubMed
  53. ↵
    1. Ryder, E.,
    2. Ashburner, M.,
    3. Bautista-Llacer, R.,
    4. Drummond, J.,
    5. Webster, J.,
    6. Johnson, G.,
    7. Morley, T.,
    8. Chan, Y. S.,
    9. Blows, F.,
    10. Coulson, D. et al.
    (2007). The DrosDel deletion collection: a Drosophila genomewide chromosomal deficiency resource. Genetics 177, 615-629. doi:10.1534/genetics.107.076216
    OpenUrlAbstract/FREE Full Text
  54. ↵
    1. Sapp, R. J.,
    2. Christianson, J. S.,
    3. Maier, L.,
    4. Studer, K. and
    5. Stark, W. S.
    (1991). Carotenoid replacement therapy in Drosophila: recovery of membrane, opsin and visual pigment. Exp. Eye Res. 53, 73-79. doi:10.1016/0014-4835(91)90147-7
    OpenUrlCrossRefPubMed
  55. ↵
    1. Satoh, A. K. and
    2. Ready, D. F.
    (2005). Arrestin1 mediates light-dependent rhodopsin endocytosis and cell survival. Curr. Biol. 15, 1722-1733. doi:10.1016/j.cub.2005.08.064
    OpenUrlCrossRefPubMedWeb of Science
  56. ↵
    1. Satoh, A.,
    2. Tokunaga, F.,
    3. Kawamura, S. and
    4. Ozaki, K.
    (1997). In situ inhibition of vesicle transport and protein processing in the dominant negative Rab1 mutant of Drosophila. J. Cell Sci. 110, 2943-2953.
    OpenUrlAbstract/FREE Full Text
  57. ↵
    1. Satoh, A. K.,
    2. Nagatani, H.,
    3. Tokunaga, F.,
    4. Kawamura, S. and
    5. Ozaki, K.
    (1998). Rhodopsin transport and Rab expression in the carotenoid-deprived Drosophila melanogaster. Zoolog. Sci. 15, 651-659. doi:10.2108/zsj.15.651
    OpenUrlCrossRef
  58. ↵
    1. Satoh, A. K.,
    2. O'Tousa, J. E.,
    3. Ozaki, K. and
    4. Ready, D. F.
    (2005). Rab11 mediates post-Golgi trafficking of rhodopsin to the photosensitive apical membrane of Drosophila photoreceptors. Development 132, 1487-1497. doi:10.1242/dev.01704
    OpenUrlAbstract/FREE Full Text
  59. ↵
    1. Schindelin, J.,
    2. Arganda-Carreras, I.,
    3. Frise, E.,
    4. Kaynig, V.,
    5. Longair, M.,
    6. Pietzsch, T.,
    7. Preibisch, S.,
    8. Rueden, C.,
    9. Saalfeld, S.,
    10. Schmid, B. et al.
    (2012). Fiji: an open-source platform for biological-image analysis. Nat. Methods 9, 676-682. doi:10.1038/nmeth.2019
    OpenUrlCrossRefPubMedWeb of Science
  60. ↵
    1. Schopf, K.,
    2. Smylla, T. K. and
    3. Huber, A.
    (2019). Immunocytochemical Labeling of Rhabdomeric Proteins in Drosophila Photoreceptor Cells Is Compromised by a Light-dependent Technical Artifact. J. Histochem. Cytochem. 67, 745-757. doi:10.1369/0022155419859870
    OpenUrlCrossRef
  61. ↵
    1. Schultz, A.,
    2. Nottrott, S.,
    3. Hartmuth, K. and
    4. Luhrmann, R.
    (2006). RNA structural requirements for the association of the spliceosomal hPrp31 protein with the U4 and U4atac small nuclear ribonucleoproteins. J. Biol. Chem. 281, 28278-28286. doi:10.1074/jbc.M603350200
    OpenUrlAbstract/FREE Full Text
  62. ↵
    1. Scotti, M. M. and
    2. Swanson, M. S.
    (2016). RNA mis-splicing in disease. Nat. Rev. Genetics 17, 19-32. doi:10.1038/nrg.2015.3
    OpenUrlCrossRefPubMed
  63. ↵
    1. Spannl, S.,
    2. Kumichel, A.,
    3. Hebbar, S.,
    4. Kapp, K.,
    5. Gonzalez-Gaitan, M.,
    6. Winkler, S.,
    7. Blawid, R.,
    8. Jessberger, G. and
    9. Knust, E.
    (2017). The Crumbs_C isoform of Drosophila shows tissue- and stage-specific expression and prevents light-dependent retinal degeneration. Biol. Open 6, 165-175. doi:10.1242/bio.020040
    OpenUrlAbstract/FREE Full Text
  64. ↵
    1. Stark, W. S. and
    2. Carlson, S. D.
    (1984). Blue and ultraviolet light induced damage to the Drosophila retina: ultrastructure. Curr. Eye Res. 3, 1441-1454. doi:10.3109/02713688409000840
    OpenUrlCrossRefPubMed
  65. ↵
    1. Tanackovic, G.,
    2. Ransijn, A.,
    3. Thibault, P.,
    4. Abou Elela, S.,
    5. Klinck, R.,
    6. Berson, E. L.,
    7. Chabot, B. and
    8. Rivolta, C.
    (2011). PRPF mutations are associated with generalized defects in spliceosome formation and pre-mRNA splicing in patients with retinitis pigmentosa. Hum. Mol. Genet. 20, 2116-2130. doi:10.1093/hmg/ddr094
    OpenUrlCrossRefPubMedWeb of Science
  66. ↵
    1. Valdes-Sanchez, L.,
    2. Calado, S. M.,
    3. de la Cerda, B.,
    4. Aramburu, A.,
    5. Garcia-Delgado, A. B.,
    6. Massalini, S.,
    7. Montero-Sanchez, A.,
    8. Bhatia, V.,
    9. Rodriguez-Bocanegra, E.,
    10. Diez-Lloret, A. et al.
    (2019). Retinal pigment epithelium degeneration caused by aggregation of PRPF31 and the role of HSP70 family of proteins. Mol. Med. 26, 1. doi:10.1186/s10020-019-0124-z
    OpenUrlCrossRef
  67. ↵
    1. Venturini, G.,
    2. Rose, A. M.,
    3. Shah, A. Z.,
    4. Bhattacharya, S. S. and
    5. Rivolta, C.
    (2012). CNOT3 is a modifier of PRPF31 mutations in retinitis pigmentosa with incomplete penetrance. PLoS Genet. 8, e1003040. doi:10.1371/journal.pgen.1003040
    OpenUrlCrossRefPubMed
  68. ↵
    1. Verbakel, S. K.,
    2. van Huet, R. A. C.,
    3. Boon, C. J. F.,
    4. den Hollander, A. I.,
    5. Collin, R. W. J.,
    6. Klaver, C. C. W.,
    7. Hoyng, C. B.,
    8. Roepman, R. and
    9. Klevering, B. J.
    (2018). Non-syndromic retinitis pigmentosa. Prog. Retin. Eye Res. 66, 157-186. doi:10.1016/j.preteyeres.2018.03.005
    OpenUrlCrossRefPubMed
  69. ↵
    1. Vithana, E. N.,
    2. Abu-Safieh, L.,
    3. Allen, M. J.,
    4. Carey, A.,
    5. Papaioannou, M.,
    6. Chakarova, C.,
    7. Al-Maghtheh, M.,
    8. Ebenezer, N. D.,
    9. Willis, C.,
    10. Moore, A. T. et al.
    (2001). A human homolog of yeast pre-mRNA splicing gene, PRP31, underlies autosomal dominant retinitis pigmentosa on chromosome 19q13.4 (RP11). Mol. Cell 8, 375-381. doi:10.1016/S1097-2765(01)00305-7
    OpenUrlCrossRefPubMedWeb of Science
  70. ↵
    1. von Lintig, J.,
    2. Kiser, P. D.,
    3. Golczak, M. and
    4. Palczewski, K.
    (2010). The biochemical and structural basis for trans-to-cis isomerization of retinoids in the chemistry of vision. Trends Biochem. Sci. 35, 400-410. doi:10.1016/j.tibs.2010.01.005
    OpenUrlCrossRefPubMedWeb of Science
  71. ↵
    1. Wang, S.,
    2. Tan, K. L.,
    3. Agosto, M. A.,
    4. Xiong, B.,
    5. Yamamoto, S.,
    6. Sandoval, H.,
    7. Jaiswal, M.,
    8. Bayat, V.,
    9. Zhang, K.,
    10. Charng, W. L. et al.
    (2014). The retromer complex is required for rhodopsin recycling and its loss leads to photoreceptor degeneration. PLoS Biol. 12, e1001847. doi:10.1371/journal.pbio.1001847
    OpenUrlCrossRefPubMed
  72. ↵
    1. Wen, X. H.,
    2. Shen, L.,
    3. Brush, R. S.,
    4. Michaud, N.,
    5. Al-Ubaidi, M. R.,
    6. Gurevich, V. V.,
    7. Hamm, H. E.,
    8. Lem, J.,
    9. Dibenedetto, E.,
    10. Anderson, R. E. et al.
    (2009). Overexpression of rhodopsin alters the structure and photoresponse of rod photoreceptors. Biophys. J. 96, 939-950. doi:10.1016/j.bpj.2008.10.016
    OpenUrlCrossRefPubMedWeb of Science
  73. ↵
    1. Wheway, G.,
    2. Douglas, A.,
    3. Baralle, D. and
    4. Guillot, E.
    (2020). Mutation spectrum of PRPF31, genotype-phenotype correlation in retinitis pigmentosa, and opportunities for therapy. Exp. Eye Res. 192, 107950. doi:10.1016/j.exer.2020.107950
    OpenUrlCrossRef
  74. ↵
    1. Will, C. L. and
    2. Luhrmann, R.
    (2011). Spliceosome structure and function. Cold Spring Harb. Perspect Biol. 3, a003707. doi:10.1101/cshperspect.a003707
    OpenUrlAbstract/FREE Full Text
  75. ↵
    1. Winkler, S.,
    2. Schwabedissen, A.,
    3. Backasch, D.,
    4. Bokel, C.,
    5. Seidel, C.,
    6. Bonisch, S.,
    7. Furthauer, M.,
    8. Kuhrs, A.,
    9. Cobreros, L.,
    10. Brand, M. et al.
    (2005). Target-selected mutant screen by TILLING in Drosophila. Genome Res. 15, 718-723. doi:10.1101/gr.3721805
    OpenUrlAbstract/FREE Full Text
  76. ↵
    1. Winkler, S.,
    2. Gscheidel, N. and
    3. Brand, M.
    (2011). Mutant generation in vertebrate model organisms by TILLING. Methods Mol. Biol. 770, 475-504. doi:10.1007/978-1-61779-210-6_19
    OpenUrlCrossRefPubMed
  77. ↵
    1. Xiong, B. and
    2. Bellen, H. J.
    (2013). Rhodopsin homeostasis and retinal degeneration: lessons from the fly. Trends Neurosci. 36, 652-660. doi:10.1016/j.tins.2013.08.003
    OpenUrlCrossRefPubMedWeb of Science
  78. ↵
    1. Xiong, B.,
    2. Bayat, V.,
    3. Jaiswal, M.,
    4. Zhang, K.,
    5. Sandoval, H.,
    6. Charng, W. L.,
    7. Li, T.,
    8. David, G.,
    9. Duraine, L.,
    10. Lin, Y. Q. et al.
    (2012). Crag is a GEF for Rab11 required for rhodopsin trafficking and maintenance of adult photoreceptor cells. PLoS Biol. 10, e1001438. doi:10.1371/journal.pbio.1001438
    OpenUrlCrossRefPubMedWeb of Science
  79. ↵
    1. Yin, J.,
    2. Brocher, J.,
    3. Fischer, U. and
    4. Winkler, C.
    (2011). Mutant Prpf31 causes pre-mRNA splicing defects and rod photoreceptor cell degeneration in a zebrafish model for Retinitis pigmentosa. Mol. Neurodegener 6, 56. doi:10.1186/1750-1326-6-56
    OpenUrlCrossRefPubMed
  80. ↵
    1. Yuan, L.,
    2. Kawada, M.,
    3. Havlioglu, N.,
    4. Tang, H. and
    5. Wu, J. Y.
    (2005). Mutations in PRPF31 inhibit pre-mRNA splicing of rhodopsin gene and cause apoptosis of retinal cells. J. Neurosci. 25, 748-757. doi:10.1523/JNEUROSCI.2399-04.2005
    OpenUrlAbstract/FREE Full Text
  81. ↵
    1. Zhang, S. X.,
    2. Sanders, E.,
    3. Fliesler, S. J. and
    4. Wang, J. J.
    (2014). Endoplasmic reticulum stress and the unfolded protein responses in retinal degeneration. Exp. Eye Res. 125, 30-40. doi:10.1016/j.exer.2014.04.015
    OpenUrlCrossRefPubMedWeb of Science
  82. ↵
    1. Zhao, C.,
    2. Bellur, D. L.,
    3. Lu, S.,
    4. Zhao, F.,
    5. Grassi, M. A.,
    6. Bowne, S. J.,
    7. Sullivan, L. S.,
    8. Daiger, S. P.,
    9. Chen, L. J.,
    10. Pang, C. P. et al.
    (2009). Autosomal-dominant retinitis pigmentosa caused by a mutation in SNRNP200, a gene required for unwinding of U4/U6 snRNAs. Am. J. Hum. Genet. 85, 617-627. doi:10.1016/j.ajhg.2009.09.020
    OpenUrlCrossRefPubMedWeb of Science
Previous ArticleNext Article
Back to top
Previous ArticleNext Article

This Issue

RSSRSS

Keywords

  • Spliceosome
  • Photoreceptor cells
  • Rhodopsin
  • scarlet
  • twinfilin

 Download PDF

Email

Thank you for your interest in spreading the word on Biology Open.

NOTE: We only request your email address so that the person you are recommending the page to knows that you wanted them to see it, and that it is not junk mail. We do not capture any email address.

Enter multiple addresses on separate lines or separate them with commas.
Mutations in the splicing regulator Prp31 lead to retinal degeneration in Drosophila
(Your Name) has sent you a message from Biology Open
(Your Name) thought you would like to see the Biology Open web site.
CAPTCHA
This question is for testing whether or not you are a human visitor and to prevent automated spam submissions.
Share
Research Article
Mutations in the splicing regulator Prp31 lead to retinal degeneration in Drosophila
Sarita Hebbar, Malte Lehmann, Sarah Behrens, Catrin Hälsig, Weihua Leng, Michaela Yuan, Sylke Winkler, Elisabeth Knust
Biology Open 2021 10: bio052332 doi: 10.1242/bio.052332 Published 25 January 2021
del.icio.us logo Digg logo Reddit logo Twitter logo CiteULike logo Facebook logo Google logo Mendeley logo
Citation Tools
Research Article
Mutations in the splicing regulator Prp31 lead to retinal degeneration in Drosophila
Sarita Hebbar, Malte Lehmann, Sarah Behrens, Catrin Hälsig, Weihua Leng, Michaela Yuan, Sylke Winkler, Elisabeth Knust
Biology Open 2021 10: bio052332 doi: 10.1242/bio.052332 Published 25 January 2021

Citation Manager Formats

  • BibTeX
  • Bookends
  • EasyBib
  • EndNote (tagged)
  • EndNote 8 (xml)
  • Medlars
  • Mendeley
  • Papers
  • RefWorks Tagged
  • Ref Manager
  • RIS
  • Zotero
Alerts

Please log in to add an alert for this article.

Sign in to email alerts with your email address

Article Navigation

  • Top
  • Article
    • ABSTRACT
    • INTRODUCTION
    • RESULTS
    • DISCUSSION
    • MATERIALS AND METHODS
    • Acknowledgements
    • Footnotes
    • References
  • Figures & tables
  • Supp info
  • Info & metrics
  • eLetters
  • PDF + SI
  • PDF

Related articles

Cited by...

More in this TOC section

  • Unique histological features of the tail skin of cotton rat (Sigmodon hispidus) related to caudal autotomy
  • Sex differences in postprandial blood glucose and body surface temperature are contingent on flight in the fruit bat, Cynopterus sphinx
  • Interdomain interactions regulate the localization of a lipid transfer protein at ER-PM contact sites
Show more RESEARCH ARTICLE

Similar articles

Other journals from The Company of Biologists

Development

Journal of Cell Science

Journal of Experimental Biology

Disease Models & Mechanisms

Advertisement

Biology Open and COVID-19

We are aware that the COVID-19 pandemic is having an unprecedented impact on researchers worldwide. The Editors of all The Company of Biologists’ journals have been considering ways in which we can alleviate concerns that members of our community may have around publishing activities during this time. Read about the actions we are taking at this time.

Please don’t hesitate to contact the Editorial Office if you have any questions or concerns.


New funding scheme supports sustainable events

As part of our Sustainable Conferencing Initiative, we are pleased to announce funding for organisers that seek to reduce the environmental footprint of their event. The next deadline to apply for a Scientific Meeting grant is 26 March 2021.


Future Leader Review – early neurodegeneration of Alzheimer’s disease

A new Future Leader Review from Olayemi Olajide, Marcus Suvanto and Clifton Andrew Chapman evaluates the molecular mechanisms that may explain the vulnerability and susceptibility of the entorhinal cortex to early neurodegeneration during the pathogenesis of Alzheimer's disease.

Find out more about our Future Leader Reviews – they are an exclusive opportunity for early-career researchers who want to establish themselves in their field.


First author interviews

Catch up on our latest first author interviews to go behind the scenes of our latest research, find out more about the authors and hear from early-career researchers themselves how they’re finding life at the bench.


Retinal degeneration in Drosophila

Thank you to Elisabeth Knust and her team for their confocal image of a longitudinal section of an adult Drosophila retina, which brightens the cover of our latest issue. Read the research behind the cover.

Articles

  • Accepted manuscripts
  • Issue in progress
  • Latest complete issue
  • Issue archive
  • Archive by article type
  • Interviews
  • Sign up for alerts

About us

  • About BiO
  • Editors and Board
  • Editor biographies
  • Grants and funding
  • Journal Meetings
  • Workshops
  • The Company of Biologists

For Authors

  • Submit a manuscript
  • Aims and scope
  • Presubmission enquiries
  • Article types
  • Manuscript preparation
  • Cover suggestions
  • Editorial process
  • Promoting your paper
  • Open Access

Journal Info

  • Journal policies
  • Rights and permissions
  • Media policies
  • Reviewer guide
  • Sign up for alerts

Contact

  • Contact BiO
  • Advertising
  • Feedback

Twitter   YouTube   LinkedIn

© 2021   The Company of Biologists Ltd   Registered Charity 277992