1. Cell Biology

Evolutionary conservation of centriole rotational asymmetry in the human centrosome

  1. Noémie Gaudin
  2. Paula Martin Gil
  3. Meriem Boumendjel
  4. Dmitry Ershov
  5. Catherine Pioche-Durieu
  6. Manon Bouix
  7. Quentin Delobelle
  8. Lucia Maniscalco
  9. Than Bich Ngan Phan
  10. Vincent Heyer
  11. Bernardo Reina-San-Martin
  12. Juliette Azimzadeh  Is a corresponding author
  1. Université Paris Cité, CNRS, Institut Jacques Monod, France
  2. Image Analysis Hub, C2RT, Institut Pasteur, France
  3. Hub de Bioinformatique et Biostatistique – Département Biologie Computationnelle, Institut Pasteur, France
  4. Institut de Génétique et de Biologie Moléculaire et Cellulaire (IGBMC), France
  5. Institut National de la Santé et de la Recherche Médicale (INSERM), France
  6. Centre National de la Recherche Scientifique (CNRS), France
  7. Université de Strasbourg, France
https://doi.org/10.7554/eLife.72382
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Cite this article
  1. Noémie Gaudin
  2. Paula Martin Gil
  3. Meriem Boumendjel
  4. Dmitry Ershov
  5. Catherine Pioche-Durieu
  6. Manon Bouix
  7. Quentin Delobelle
  8. Lucia Maniscalco
  9. Than Bich Ngan Phan
  10. Vincent Heyer
  11. Bernardo Reina-San-Martin
  12. Juliette Azimzadeh
(2022)
Evolutionary conservation of centriole rotational asymmetry in the human centrosome
eLife 11:e72382.
https://doi.org/10.7554/eLife.72382
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Abstract

Centrioles are formed by microtubule triplets in a ninefold symmetric arrangement. In flagellated protists and animal multiciliated cells, accessory structures tethered to specific triplets render the centrioles rotationally asymmetric, a property that is key to cytoskeletal and cellular organization in these contexts. In contrast, centrioles within the centrosome of animal cells display no conspicuous rotational asymmetry. Here, we uncover rotationally asymmetric molecular features in human centrioles. Using ultrastructure expansion microscopy, we show that LRRCC1, the ortholog of a protein originally characterized in flagellate green algae, associates preferentially to two consecutive triplets in the distal lumen of human centrioles. LRRCC1 partially co-localizes and affects the recruitment of another distal component, C2CD3, which also has an asymmetric localization pattern in the centriole lumen. Together, LRRCC1 and C2CD3 delineate a structure reminiscent of a filamentous density observed by electron microscopy in flagellates, termed the ‘acorn.’ Functionally, the depletion of LRRCC1 in human cells induced defects in centriole structure, ciliary assembly, and ciliary signaling, supporting that LRRCC1 cooperates with C2CD3 to organizing the distal region of centrioles. Since a mutation in the LRRCC1 gene has been identified in Joubert syndrome patients, this finding is relevant in the context of human ciliopathies. Taken together, our results demonstrate that rotational asymmetry is an ancient property of centrioles that is broadly conserved in human cells. Our work also reveals that asymmetrically localized proteins are key for primary ciliogenesis and ciliary signaling in human cells.

Editor's evaluation

This work shows that, contrary to a widely accepted view, centrioles of the human centrosome are rotationally asymmetric, a feature previously known only from centrioles in flagellated protists and multiciliated cells. The authors identify LRRCC1, implicated in ciliary disease, as an asymmetrically localized protein of the centriole lumen and show that it contributes to proper centriole structure, ciliary assembly, and ciliary signaling.

https://doi.org/10.7554/eLife.72382.sa0

Introduction

Centrioles are cylindrical structures with a characteristic ninefold symmetry, which results from the arrangement of their constituent microtubule triplets (LeGuennec et al., 2021). In animal cells, centrioles are essential for the assembly of centrosomes and cilia. The centrosome, composed of two centrioles embedded in a pericentriolar material (PCM), is a major organizer of the microtubule cytoskeleton. In addition, most vertebrate cells possess a primary cilium, a sensory organelle that assembles from the oldest centriole within the centrosome, called mother centriole (Kumar and Reiter, 2021).

Centrioles within the centrosome show no apparent rotational asymmetry, that is, no structural asymmetry of the microtubule triplets. In vertebrates, the mother centriole carries distal appendages (DAs) and subdistal appendages arranged in a symmetric manner around the centriole cylinder (Kumar and Reiter, 2021). In contrast, the centriole/basal body complex of flagellates, to which the animal centrosome is evolutionary related, is characterized by marked rotational asymmetries (Azimzadeh, 2021; Yubuki and Leander, 2013). In flagellates, an array of fibers and microtubules anchored asymmetrically at centrioles controls the spatial organization of the cell (Feldman et al., 2007; Yubuki and Leander, 2013). The asymmetric attachment of cytoskeletal elements appears to rely on molecular differences between microtubule triplets. In the green alga Chlamydomonas reinhardtii, Vfl1p (variable flagella number 1 protein) localizes principally at two triplets near the attachment site of a striated fiber connecting the centrioles (Silflow et al., 2001). This fiber is absent or mispositioned in the vfl1 mutant, leading to defects in centriole position and number, and overall cytoskeleton disorganization (Adams et al., 1985; Feldman et al., 2007). In the same region, a rotationally asymmetric structure termed the ‘acorn’ was observed in the centriole lumen by transmission electron microscopy. The acorn appears as a filament connecting five successive triplets and is in part colocalized with Vfl1p (Geimer and Melkonian, 2005; Geimer and Melkonian, 2004).

We recently established that Vfl1p function is conserved in the multiciliated cells (MCCs) of planarian flatworms, which was recently confirmed in Xenopus (Basquin et al., 2019; Nommick et al., 2022). MCCs assemble large numbers of centrioles that are polarized in the plane of the plasma membrane to enable the directional beating of cilia (Meunier and Azimzadeh, 2016), like in C. reinhardtii. The planarian ortholog of Vfl1p is required for the assembly of two appendages that decorate MCC centrioles asymmetrically, the basal foot and the ciliary rootlet (Basquin et al., 2019). Depleting Vfl1p orthologs in planarian or xenopus MCCs alters centriole rotational polarity, reminiscent of the vfl1 phenotype in C. reinhardtii (Adams et al., 1985; Basquin et al., 2019; Nommick et al., 2022). Intriguingly, the human ortholog of Vfl1p, called LRRCC1 (leucine rich repeat and coiled coil containing 1) localizes at the centrosome despite the lack of rotationally asymmetric appendage in this organelle (Andersen et al., 2003; Muto et al., 2008). Furthermore, a homozygous mutation in the LRRCC1 gene was identified in two siblings affected by a ciliopathy called Joubert syndrome (JBTS), suggesting that LRRCC1 might somehow affect the function of non-motile cilia (Shaheen et al., 2016).

Here, we show that LRRCC1 localizes in a rotationally asymmetric manner in the centrioles of the human centrosome. We further establish that LRRCC1 is required for proper ciliary assembly and signaling, which likely explains its implication in JBTS. LRRCC1 affects the recruitment at centrioles of another ciliopathy protein called C2CD3 (C2 domain containing 3), which we found to also localize in a rotationally asymmetric manner, forming a pattern partly reminiscent of the acorn described in flagellates. Our findings uncover the unanticipated rotational asymmetry of centrioles in the human centrosome and show that this property is connected to the assembly and function of primary cilia.

Results

LRRCC1 localizes asymmetrically at the distal end of centrioles

To investigate a potential role of LRRCC1 at the centrosome, we first sought to determine its precise localization. We raised antibodies against two different fragments within the long C-terminal coiled-coil domain of LRRCC1 (Ab1, 2), which both stained the centrosome region in human retinal pigmented epithelial (RPE1) cells (Figure 1a, Figure 1—figure supplement 1a), as previously reported (Muto et al., 2008). Labeling intensity was decreased in LRRCC1-depleted cells for both antibodies, supporting their specificity (Figure 4a and b, Figure 1—figure supplement 1). LRRCC1 punctate labeling in the centrosomal region indicated that it is present within centriolar satellites, confirming a previous finding that LRRCC1 interacts with the satellite component PCM1 (Gupta et al., 2015). After nocodazole depolymerization of microtubules to disperse satellites, a fraction of LRRCC1 was retained at centrioles (Figure 1a, Figure 1—figure supplement 1a), providing evidence that LRRCC1 is also a core component of centrioles. To determine LRRCC1 localization more precisely within the centriolar structure, we used ultrastructure expansion microscopy (U-ExM) (Gambarotto et al., 2019) combined with imaging on a Zeiss Airyscan 2 confocal microscope, thereby increasing the resolution by a factor of ~8 compared to conventional confocal microscopy. We found that LRRCC1 localizes at the distal end of centrioles as well as of procentrioles (Figure 1b). Strikingly, and unlike other known centrosome components, LRRCC1 decorated the distal end of centrioles in a rotationally asymmetric manner. Indeed, LRRCC1 was detected close to the triplet blades and towards the lumen of the centriole (Figure 1c). The staining was often associated with two or more consecutive triplets, one of them being usually more brightly labeled than the others. In addition, a fainter staining was consistently detected along the entire length of all triplets (Figure 1b, brighter exposure). This pattern was observed in both RPE1 and HEK 293 cells and was obtained with both anti-LRRCC1 antibodies (Figure 1—figure supplement 1h), supporting its specificity. We verified that LRRCC1 asymmetric localization was also observed in unexpanded cells by directly analyzing immunofluorescence samples by Airyscan microscopy (Figure 1d). We measured the lateral distribution of LRRCC1 signal intensity peak relative to the long axis of the centriole. The distance between peaks was greater for LRRCC1 than for hPOC5, a marker that localizes symmetrically in the centriole (Azimzadeh et al., 2009; Le Guennec et al., 2020; Schweizer et al., 2021), confirming the asymmetry of LRRCC1 staining. The distal pattern obtained by U-ExM showed some variability, especially in the distance between LRRCC1 and the centriole wall (Figure 1c), which could result from the fact that centrioles were not perfectly orthogonal to the imaging plan. To obtain a more accurate picture of LRRCC1 localization, we generated 3D reconstructions that we realigned, first along the vertical axis, then with respect to one another using the most intense region of the LRRCC1 labeling as a reference point (Figure 1e, Figure 1—figure supplement 2a and b). An average 3D reconstruction was then generated (Figure 1f) and revealed that LRRCC1 was mainly associated to one triplet, and to a lesser extent to its direct neighbor counterclockwise, on their luminal side. A longitudinal view confirmed that LRRCC1 is principally located at the distal end of centrioles.

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LRRCC1 is localized in a rotationally asymmetric manner at the distal end of centrioles in the human centrosome.

(a) LRRCC1 localization in non-treated RPE1 cells (left) or in cells treated with nocodazole to disperse the pericentriolar satellites (right). LRRCC1 (Ab2, yellow), γ-tubulin (PCM, magenta), and DNA (cyan). Bar, 5 µm (insets, 2 µm). (b) Longitudinal view of centrioles and procentrioles in the duplicating centrosome of an RPE1 cell analyzed by ultrastructure expansion microscopy (U-ExM). LRRCC1 (Ab2, yellow), acetylated tubulin (magenta). Bar, 0.5 µm. (c) Centrioles from WT RPE1 cells as seen from the distal end. LRRCC1 (Ab2, yellow), acetylated tubulin (magenta). Images are maximum intensity projections of individual z-sections encompassing the LRRCC1 signal. Note that an apparent shift between channels occurs when centrioles are slightly angled with respect to the imaging axis. Bar, 0.2 µm. (d) Lateral distance between LRRCC1 (left, yellow) or hPOC5 (middle, cyan) signal intensity peaks and the centriole center (given by the position of acetylated tubulin intensity peak, magenta) in ciliated RPE1 cells. Individual intensity profiles were measured along the green lines. The approximate position of the centriole is shown (white cylinders). Note that LRRCC1 and hPOC5 were also detected at the periphery of the centriole, towards the proximal end for LRRCC1 and in the appendage region for hPOC5. Bar, 0.2 µm. Right: interpeak distance (d). Bars, mean ± SD, 31 cells from two different experiments (Kolmogorov–Smirnov test). (e) Workflow for calculating the average staining from 3D-reconstructed individual centrioles generated from confocal z-stacks. The brightest part of LRRCC1 signal was used as a reference point to align the centrioles. (f) Average LRRCC1 staining obtained from 34 individual centrioles viewed from the distal end, in transverse and longitudinal views. A diagram representing the average pattern in transverse view is also shown.

Together, our results show that LRRCC1 is localized asymmetrically within the distal centriole lumen, establishing that centrioles within the human centrosome are rotationally asymmetric.

The localization pattern of LRRCC1 is similar at the centrosome and in mouse MCCs

LRRCC1 orthologs are required for establishing centriole rotational polarity in planarian and xenopus MCCs, like in C. reinhardtii (Basquin et al., 2019; Nommick et al., 2022; Silflow et al., 2001). It is therefore plausible that LRRCC1-related proteins localize asymmetrically in MCC centrioles, and indeed, Lrrcc1 was recently found associated to the ciliary rootlet in xenopus MCCs (Nommick et al., 2022). To determine whether LRRCC1 also localizes at the distal end of MCC centrioles in addition to its rootlet localization, and if so, whether LRRCC1 localization pattern resembles that observed at the centrosome, we analyzed mouse ependymal and tracheal cells by U-ExM. In in vitro differentiated ependymal cells, the labeling generated by the anti-LRRCC1 antibody was consistent with our observations in human culture cells. Mouse Lrrcc1 localized asymmetrically at the distal end of centrioles, opposite to the side where the basal foot is attached (Figure 2a), as determined by co-staining with the basal foot marker γ-tubulin (Clare et al., 2014). Lrrcc1 was also present at the distal end of procentrioles forming via either the centriolar or acentriolar pathways (i.e., around parent centrioles or deuterosomes, respectively) (Figure 2b). We also examined tracheal explants, in which centrioles were docked and polarized at the apical membrane in higher proportions (Figure 2c). We obtained an average image of Lrrcc1 labeling from 35 individual centrioles aligned using the position of the basal foot as a reference point. This revealed that Lrrcc1 is principally located in the vicinity of three triplets opposite to the basal foot, to the right of basal foot main axis (triplet numbers 9, 1, and 2 on the diagram in Figure 2d). Lrrcc1 was located farther away from the triplet wall than in centrioles of the centrosome, but this was likely an effect of a deformation of the centrioles (Figure 2c and d) caused by the incomplete expansion of the underlying cartilage layer in tracheal explants. In agreement, Lrrcc1 was close to the triplets in ependymal cell monolayers, which expand isometrically. Besides the distal centriole staining, we found no evidence that Lrrcc1 is associated to the ciliary rootlet in mouse MCCs, unlike in xenopus. The Lrrcc1 pattern in mouse MCCs was thus similar to the pattern observed at the human centrosome.

Figure 2
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The LRRCC1 rotationally asymmetric pattern is conserved in mouse multiciliated cells (MCCs).

(a) Centrioles in the cytoplasm of mouse ependymal cells differentiating in vitro analyzed by ultrastructure expansion microscopy (U-ExM), in longitudinal and transverse view. Lrrcc1 (Ab2, yellow), γ-tubulin (basal foot cap, cyan), and acetylated tubulin (magenta). Of note, γ-tubulin was also detected in the proximal lumen of centrioles. Bar, 0.2 µm. (b) Procentrioles assembling via the centriolar (right) or the deuterosome pathway (left and center) in ependymal cells. Lrrcc1 (Ab2, yellow), acetylated tubulin (magenta). Bar, 0.2 µm. (c) Transverse view of centrioles docked at the apical membrane in fully differentiated mouse tracheal cells, viewed from the distal end. Lrrcc1 (Ab2, yellow), γ-tubulin (cyan), and acetylated tubulin (magenta). Bar, 0.2 µm. (d) Average image generated from 35 individual centrioles from mouse trachea, viewed from the distal end, shown in transverse and longitudinal views. The position of the basal foot (cyan dotted line) stained with γ-tubulin was used as a reference point to align the centrioles. A diagram of the average pattern in transverse view is shown, in which the direction of ciliary beat (Schneiter et al., 2021) is represented by a dotted arrow and the basal foot axis by a green line. Triplets are numbered counterclockwise from the LRRCC1 signal.

Together, these results show that LRRCC1 asymmetric localization is a conserved feature of mammalian centrioles, presumably linked to the control of centriole rotational polarity and ciliary beat direction in MCCs.

Procentriole assembly site is partly correlated with centriole rotational polarity

In C. reinhardtii, cytoskeleton organization and flagellar beat direction depend on the position and orientation at which new centrioles arise during cell division. Reflecting the stereotypical organization of centrioles and procentrioles in this species, Vfl1p is recruited early and at a fixed position at the distal end of procentrioles (Figure 3a; Geimer and Melkonian, 2004; Silflow et al., 2001). We therefore wondered whether this mechanism might be to some extent conserved at the centrosome, which could explain the persistence of centriole rotational asymmetry despite the absence of asymmetric appendages or ciliary motility in most animal cell types. We first analyzed the timing of LRRCC1 incorporation into procentrioles. LRRCC1 was already present at an early stage of centriole assembly when the procentrioles stained with acetylated tubulin and the cartwheel component SAS-6 were only about 100 nm in length (Figure 3b). LRRCC1 was then detected during successive stages of procentriole elongation, always localizing asymmetrically and distally (Figure 3c), like in C. reinhardtii. We then examined LRRCC1 localization in duplicating centrosomes by generating 3D reconstructions of diplosomes (i.e., orthogonal centriole pairs) from RPE1 and HEK 293 cells processed by U-ExM (Figure 3d). We analyzed two parameters: the angle between LRRCC1 in the procentriole and the long axis of the parent centriole used as reference (Figure 3d, LRRCC1 localization in procentrioles), and the angle between procentriole position and LRRCC1 in the parent centriole (Figure 3d, procentriole position with respect to centriolar LRRCC1). We found that LRRCC1 localization in procentrioles was more often aligned with the long axis of the parent centriole in RPE1 cells (Figure 3d, top-left panel, quadrants Q1 and Q3, respectively), but less so in HEK 293 cells (top-right panel), in which the distribution was closer to a random distribution. Thus, human procentrioles do not arise in a fixed orientation, although there appears to be a bias toward alignment of LRRCC1 with the main axis of the parent centriole in RPE1 cells. Next, we analyzed the position of procentrioles with respect to centriolar LRRCC1 (bottom panels). Based on current models, procentriole assembly is expected to occur at a random position around parent centrioles in animal cells (Takao et al., 2019). Identification of LRRCC1 provided the first opportunity to directly test this model. In diplosomes from both RPE1 and HEK 293 cells, the position of procentrioles with respect to LRRCC1 location in the parent centriole was variable, confirming that the position at which procentrioles assemble is not strictly controlled in human cells. Interestingly, however, the procentrioles were not distributed in a completely random fashion either. Procentrioles were found in quadrant Q2 (45–135° clockwise from LRRCC1 centroid) on average four times less often than in the other quadrants, both in RPE1 and HEK 293 cells, suggesting that rotational polarity of the parent centriole somehow impacts procentriole assembly.

Figure 3
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Procentriole assembly site is partly correlated with centriole rotational polarity.

(a) Diagram showing the localization of Vfl1p (cyan) in the centrioles/basal bodies (gray) and procentrioles/probasal bodies (pink) of C. reinhardtii. The microtubule roots are also shown. (b) Early stage of procentriole assembly stained for LRRCC1 (Ab2, cyan), SAS-6 (yellow), and acetylated tubulin (magenta) in a HEK 293 cell. The brightness of the acetylated tubulin labeling was increased in the insets. Bar, 0.1 µm. (c) Successive stages of centriole elongation in HEK 293 cells stained for LRRCC1 (Ab2, cyan) and acetylated tubulin (magenta). Bar, 0.1 µm. (d) Location of LRRCC1 in the procentrioles (top panels) and position of the procentriole relative to its parent centriole polarity (bottom panels), in RPE1 and HEK 293 centrioles analyzed by ultrastructure expansion microscopy (U-ExM). For each diplosome, the angle between LRRCC1 in the procentriole and the centriole long axis (top panels), or between the procentriole and LRRCC1 in the centriole (bottom panels) was measured. The number of diplosomes analyzed is indicated. p-Values are indicated when statistically different from a random distribution (χ2 test).

Overall, these results suggest that centriole rotational polarity influences centriole duplication, limiting procentriole assembly within a particular region of centriole periphery. Nevertheless, procentrioles are not formed at a strictly determined position, suggesting that the mechanisms involving the LRRCC1 ortholog Vfl1p in centriole duplication in C. reinhardtii are not or not completely conserved at the centrosome.

LRRCC1 is required for primary cilium assembly and ciliary signaling

A previous report identified a homozygous mutation in a splice acceptor site of the LRRCC1 gene in two siblings diagnosed with JBTS (Shaheen et al., 2016), but how disruption of LRRCC1 expression affects ciliary assembly and signaling has never been investigated. To address this, we generated RPE1 cell lines deficient in LRRCC1 using two different CRISPR/Cas9 strategies and targeting two different regions of the LRRCC1 locus. We could not recover null clones despite repeated attempts in RPE1 – both wildtype and p53-/- (Izquierdo et al., 2014), HEK 293 and U2-OS cells, suggesting that a complete lack of LRRCC1 is possibly deleterious. Nevertheless, we obtained partially depleted mutant clones, including three RPE1 clones targeted in either exons 8–9 (clone 1.1) or exons 11–12 (clones 1.2 and 1.9). Clone 1.1 carries deletions in both copies of the LRRCC1 gene (Figure 4—figure supplement 1a). However, long in-frame transcripts are expressed at reduced levels through alternative splicing (Figure 1—figure supplement 1c). These transcripts are expected to generate mutant protein isoforms carrying deletions in the beginning of the coiled-coil region (Figure 4—figure supplement 1). In contrast, only wildtype transcripts were detected in clones 1.2 and 1.9, which were present at approximately 30% of wildtype levels, as determined by quantitative RT-PCR (Figure 1—figure supplement 1c). We could not evaluate the overall decrease in LRRCC1 levels since the endogenous LRRCC1 protein was not detected by Western blot (Figure 1—figure supplement 1b). However, we confirmed the decrease in centrosomal LRRCC1 levels by immunofluorescence using the two different anti-LRRCC1 antibodies (Figure 4a, Figure 1—figure supplement 1d and e). The downregulation of LRRCC1 in CRISPR clones was overall of the same order as that achieved by RNAi, although treatment of CRISPR clones with the more efficient siRNA (si LRRCC1-1) could further reduce LRRCC1 levels (Figure 4a). Using Airyscan microscopy, we showed that LRRCC1 amounts were decreased not only at centriolar satellites, but also at the centrioles themselves in CRISPR clones (Figure 4b). Interestingly, the decrease in centriolar LRRCC1 was less for clone 1.1 than for the other clones, suggesting that the mutant isoforms produced in this clone have different dynamics than wildtype LRRCC1. Following induction of ciliogenesis, the proportion of ciliated cells was decreased in all three mutant clones compared to control cells (Figure 4c). We were unable to obtain stable RPE1 cell lines expressing tagged versions of LRRCC1, and transient overexpression of LRRCC1 in wildtype cells led to a decrease in the proportion of ciliated cells, making phenotype rescue experiments difficult to interpret. However, we used RNAi as an independent method to verify the specificity of ciliary defects observed in CRISPR clones. The proportion of ciliated cells was decreased by RNAi to a similar extent than in CRISPR clones (Figure 4c, Figure 1—figure supplement 1f). RNAi treatment of CRISPR clones did not lead to a greater decrease in ciliary frequency, suggesting that loss of LRRCC1 only partially inhibits ciliogenesis (Figure 4c). Sensory ciliopathies like JBTS result to a large extent from defective Hedgehog signaling (Romani et al., 2013). We determined the effect of LRRCC1-depletion on Hedgehog signaling by measuring the ciliary accumulation of the activator SMOOTHENED (SMO) upon induction of the pathway (Rohatgi et al., 2007). Depletion of LRRCC1 by either CRISPR editing or RNAi led to a drastic decrease in SMO accumulation at the primary cilium following induction of the Hedgehog pathway by SMO-agonist (SAG) (Figure 4d and e), and reduced expression of the target gene PTCH1 (Figure 1—figure supplement 1i; Goodrich et al., 1996). Taken together, our results demonstrate that LRRCC1 is required for proper ciliary assembly and signaling in human cells, further establishing its implication in JBTS.

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LRRCC1 is required for ciliary assembly and signaling.

(a) Left: LRRCC1 staining (Ab2) of WT or LRRCC1-defficient RPE1 cells obtained by CRISPR/Cas9 editing (clones 1.1, 1.2, and 1.9). Bar, 2 µm. Right: quantification of fluorescence intensity in WT or CRISPR clones treated with control or LRRCC1 siRNAs. Bars, mean ± SD, three independent experiments. p-Values are provided when statistically significant from the corresponding control (one-way ANOVA). (b) Quantification of LRRCC1 distal pool at the mother centriole of ciliated WT or CRISPR cells. Left: Airyscan images showing the region of interest (circled). LRRCC1 (yellow), acetylated tubulin (magenta). Bar: 0.5 µm. Right: quantification of the corresponding signal. Bars, mean ± SD, ≥47 cells from two independent experiments. p-Values are provided when statistically significant from the corresponding control (one-way ANOVA). (c) Percentage of ciliated cells in WT or CRISPR cells treated with control or LRRCC1 siRNAs and serum-deprived during 24 hr. Bars, mean ± SD, ≥204 cells from three independent experiments for each condition. p-Values are provided when statistically significant from the corresponding control (one-way ANOVA). (d) Left: SMO (yellow) accumulation at primary cilia (ARL13B, magenta) following SMO-agonist (SAG)-induction of the Hedgehog pathway, in WT or CRISPR cells. Bar, 2 µm. Right: quantification of ciliary SMO expressed as a percentage of the SAG-treated WT mean. Bars, mean ± SD, three independent experiments. p-Values are provided when statistically significant from the corresponding control (one-way ANOVA). (e) Ciliary SMO expressed as a percentage of the SAG-induced control mean in RPE1 cells treated with control or LRRCC1 siRNAs. Bars, mean ± SD, three independent experiments. p-Values are provided when statistically significant from the corresponding control (one-way ANOVA).

Depletion of LRRCC1 induces defects in centriole structure

Mutations in distal centriole components can alter centriole length regulation or the assembly of DAs, which both result in defective ciliogenesis (Reiter and Leroux, 2017; Sharma et al., 2021). We used U-ExM to search for possible defects in centriole structure in LRRCC1-depleted RPE1 cells. We measured centriole length in CRISPR clone 1.9, which has the lowest levels of centriolar LRRCC1 (Figure 4b), and in clone 1.1, which expresses mutant isoforms of LRRCC1. Centrioles were co-stained with anti-acetylated tubulin and an antibody against the DA component CEP164 to differentiate mother and daughter centrioles. We observed an increase in centriole length in clone 1.9 (Figure 5a) compared to control cells (483 ± 53 nm for mother and 372 ± 55 nm for daughter centrioles in clone 1.9; 427 ± 56 nm for mother and 320 ± 46 nm for daughter centrioles in control cells; mean ± SD). Although on a limited sample size, we also observed abnormally long centrioles by transmission electron microscopy in this clone (494 ± 73 nm in clone 1.9, N = 9; 429 ± 52 nm in control cells, N = 3; mean ± SD) (Figure 5c). The increase in centriole length was not due to mitotic delay as previously observed (Kong et al., 2020) since the duration of mitosis in clone 1.9 was similar as in control cells (Figure 1—figure supplement 1k). In addition, although centriole length was not modified in clone 1.1, further reduction of LRRCC1 levels by RNAi resulted in a significant increase in centriole length compared to control cells (Figure 5b). Next, we analyzed DA organization by labeling CEP164, which localizes to the outer part of DAs (Figure 5d; Yang et al., 2018). In RPE1 control cells, 80% ± 14% of mother centriole had nine properly shaped DAs, but this proportion fell to 57% ± 16% and 44% ± 17% (mean ± SD) in clones 1.1 and 1.9, respectively (Figure 5e). Mutant clones exhibited an increased proportion of centrioles with one or more abnormally shaped DAs (29% ± 17% and 42% ± 18% in clones 1.1 and 1.9, respectively, compared to 11% ± 11% in control cells; mean ± SD). We obtained similar results in a HEK 293 CRISPR clone expressing half the control levels of LRRCC1 (Figure 5f, Figure 1—figure supplement 1g). LRRCC1 depletion did not affect overall CEP164 levels at mother centrioles in the CRISPR clones (Figure 5—figure supplement 1a and d), consistent with the relatively mild defect in DA morphology observed by U-ExM. We also analyzed the distribution of CEP83, a DA component that localizes closer to the centriole wall (Yang et al., 2018). The proportion of centrioles with abnormal CEP83 labeling was not significantly different between control cells and CRISPR clones. However, this proportion became significantly lower than in the control after treating CRISPR clones with RNAi (41% ± 18% and 48% ± 4% in clones 1.1 and 1.9 treated with RNAi, respectively, compared to 77% ± 9% in control cells; mean ± SD; Figure 5g and h). Beyond these anomalies in centriolar structure, LRRCC1-depleted cells showed no defect in centriole number, supporting that centriole assembly is not affected by LRRCC1 downregulation (Figure 1—figure supplement 1j).

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Depleting LRRCC1 induces defects in centriole structure.

(a) Centriole length in mother (MC) and daughter (DC) centrioles analyzed by ultrastructure expansion microscopy (U-ExM) in WT or LRRCC1-deficient clones (1.1 and 1.9). Left: centrioles were stained for acetylated tubulin (magenta) and CEP164 (yellow) to measure centriole length (arrows). Bar, 0.5 µm. Right: quantification. Bars, mean ± SD, ≥38 centrioles from three independent experiments. p-Values are provided when statistically significant from the corresponding control (one-way ANOVA). (b) Centriole length in control cells or CRISPR cells treated with LRRCC1 siRNA-1 and stained for acetylated tubulin and CEP83. Bars, mean ± SD, ≥43 centrioles from three independent experiments. p-Values are provided when statistically significant from the corresponding control (one-way ANOVA). (c) Transmission electron microscopy view of centrioles in WT and CRISPR (clone 1.9) RPE1 cells. Note that the 1.9 centrioles are from the same cell. N = 9 centrioles from eight different cells for clone 1.9, 3 centrioles from two different cells for WT. Bar, 0.5 µm. (d) Examples of normal distal appendages (DAs), DAs with abnormal morphology (white arrowhead: abnormal spacing between consecutive DAs; cyan arrowhead: abnormal DA shape) or missing DAs (gray arrowhead) in RPE1 cells stained with CEP164 (yellow) and analyzed by U-ExM. Images are maximum intensity projections of individual z-sections encompassing the CEP164 signal. Note that an apparent shift between channels occurs when centrioles are slightly angled with respect to the imaging axis. Bar, 1 µm. (e) Percentages of centrioles presenting anomalies in CEP164 staining in WT or CRISPR RPE1 cells. ≥87 centrioles from eight independent experiments for each condition. p-Values are provided when statistically significant from the corresponding control (two-way ANOVA). (f) Percentages of centrioles presenting anomalies in CEP164 staining in WT or CRISPR HEK 293 (clone 25) cells. ≥40 centrioles from four independent experiments for each condition. p-Values are provided when statistically significant from the corresponding control (two-way ANOVA). (g) Examples of normal DAs, DAs with abnormal morphology (white arrowhead) or missing DAs (gray arrowhead) in RPE1 cells stained with CEP83 (yellow) and analyzed by U-ExM. Images are maximum intensity projections of individual z-sections encompassing the CEP83 signal. Note that apparent shift between channels and decreased circularity occurs when centrioles are slightly angled with respect to the imaging axis. Bar, 1 µm. (h) Percentages of centrioles presenting anomalies in CEP83 staining in WT RPE1 cells and CRISPR clones with or without RNAi treatment. ≥56 centrioles from three independent experiments for each condition. p-Values are provided when statistically significant from the corresponding control (two-way ANOVA).

Together, these results show that downregulation of LRRCC1 affects the formation of centriole distal structures, leading to centriole over-elongation and abnormal DA morphology.

LRRCC1 and C2CD3 delineate a rotationally asymmetric structure in human centrioles

We next wanted to determine whether LRRCC1 cooperates with other distal centriole components. Proteins shown to be recruited early at procentriole distal end include CEP290 (Kim et al., 2008), OFD1 (Singla et al., 2010), and C2CD3 (Thauvin-Robinet et al., 2014). Of particular interest, OFD1 and C2CD3 are required for DA assembly and centriole length control, and mutations in these proteins have been implicated in sensory ciliopathies (Singla et al., 2010; Thauvin-Robinet et al., 2014; Tsai et al., 2019; Wang et al., 2018). We first determined whether depletion of LRRCC1 either by CRISPR editing or by RNAi led to modifications in the recruitment of these proteins within centrioles. We found no major differences in the centrosomal levels of OFD1 and CEP290 compared to control cells (Figure 5—figure supplement 1b, c, e, and f). In contrast, C2CD3 levels were moderately increased in cells depleted from LRRCC1 either by CRISPR editing (clones 1.1 and 1.9) or by RNAi (Figure 6a and b). We thus analyzed C2CD3 further by U-ExM. As described previously, C2CD3 localized principally at the distal extremity of centrioles (Figure 6c; Tsai et al., 2019; Yang et al., 2018). Strikingly, the C2CD3 labeling was also asymmetric, often adopting a C-shape (Figure 6d). After correcting the vertical alignment of centrioles as previously, we generated an average 3D reconstruction of the C2CD3 pattern. To do this, we used one end of the C as a reference point in the xy-plane to superimpose individual centriole views. The resulting image supported that the C2CD3 labeling forms a C-shaped pattern positioned symmetrically in the centriole lumen (Figure 6e). To determine whether the C2CD3 localization pattern is affected by LRRCC1-depletion, we next analyzed C2CD3 in LRRCC1 CRISPR clones 1.1 and 1.9. The C2CD3 pattern was more variable than in control RPE1 cells, and often appeared abnormal in shape, position, or both (Figure 6f). Indeed, averaging the signal from multiple LRRCC1-depleted centrioles produced aberrant patterns, most strikingly for clone 1.9 (Figure 6g). Furthermore, the phenotype of clone 1.1 was enhanced by further reducing LRRCC1 levels using RNAi (Figure 6g). Thus, LRRCC1 is required for the proper assembly of the C2CD3-containing distal structure.

Figure 6
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C2CD3 localizes asymmetrically at the distal end of centrioles and is affected by LRRCC1 depletion.

(a) C2CD3 levels at the centrosome of WT or CRISPR RPE1 cells. Bars, mean ± SD, three independent experiments. p-Values are provided when statistically significant from the corresponding control (one-way ANOVA). (b) C2CD3 levels at the centrosome in RPE1 cells treated with control or LRRCC1 siRNAs. Bars, mean ± SD, three independent experiments. p-Values are provided when statistically significant from the corresponding control (one-way ANOVA). (c) Longitudinal view of a centriole analyzed by ultrastructure expansion microscopy (U-ExM) and stained for C2CD3 (yellow) and acetylated tubulin (magenta). Bar, 0.2 µm. (d) Centrioles from WT RPE1 cells as viewed from the distal end. C2CD3 (yellow), acetylated tubulin (magenta). Images are maximum intensity projections of individual z-sections encompassing the C2CD3 signal. Note that an apparent shift between channels occurs when centrioles are slightly angled with respect to the imaging axis. Bar, 0.2 µm. (e) Average C2CD3 images obtained from 33 individual centrioles from WT RPE1 cells viewed from the distal end, in transverse views. One end of the C-pattern was used as a reference point to align individual centrioles. (f) Centrioles from untreated CRISPR cells or CRISPR cells treated with LRRCC1 RNAi in transverse section as viewed from the distal end. C2CD3 (yellow), acetylated tubulin (magenta). Images are maximum intensity projections of individual z-sections encompassing the C2CD3 signal. Note that an apparent shift between channels occurs when centrioles are slightly angled with respect to the imaging axis. Bar, 0.2 µm. (g) Average C2CD3 images obtained from untreated or RNAi-treated CRISPR cells viewed from the distal end, in transverse views. The number or individual centrioles used for generating each average is indicated.

To determine whether LRRCC1 and C2CD3 might belong to a common structure, we next examined their respective positions within the centriole. We co-stained centrioles with our anti-LRRCC1 antibody and a second anti-C2CD3 antibody produced in sheep (Table 1). We confirmed that LRRCC1 and C2CD3 are present in the same distal region of the centriole (Figure 7a). In transverse views, the two proteins were usually not perfectly colocalized but found in close vicinity of one another near the microtubule wall. However, C2CD3 distal staining was consistently fainter than with the previous antibody, and we either could not observe a full C-shaped pattern or we could not image it due to fluorescence bleaching. Neither anti-C2CD3 antibodies worked in mouse, so we were not able to compare C2cd3 and Lrrcc1 localization in MCCs. Nevertheless, the results obtained by individually labeling LRRCC1 and C2CD3 at the centrosome (Figures 1f and 6e) together with the co-localization data (Figure 7a) are consistent with the hypothesis that LRRCC1 is located along the C2CD3-containing, C-shaped structure (Figure 7b). C2CD3 was not co-immunoprecipitated with a GFP-LRRCC1 fusion protein, however, suggesting that LRRCC1 and C2CD3 do not directly interact (Figure 7—figure supplement 1).

Figure 7 with 1 supplement see all
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C2CD3 and LRRCC1 partially colocalize at the distal end of centrioles.

(a) RPE1 centrioles processed for ultrastructure expansion microscopy (U-ExM) and stained for LRRCC1 (Ab2, yellow), C2CD3 (cyan), and acetylated tubulin (magenta). Bar, 0.1 µm. (b) Model showing the possible location of LRRCC1 and C2CD3 relative to each other within human centrioles. Right panel: diagram showing the respective positions of the acorn (Geimer and Melkonian, 2004) and Vfl1p (Silflow et al., 2001) in C. reinhardtii. The direction of the flagellar beat is indicated by a dotted arrow, and the distal striated fiber is in gray. (c) Evolution of the roles played by Vfl1p/LRRCC1 proteins and associated rotationally asymmetric centriolar substructures. In C. reinhardtii, Vfl1p is required for proper ciliary assembly (1), as well as for the formation of fibers and microtubular roots (2) that control the position of centrioles and procentrioles (3), and overall cellular organization (Adams et al., 1985; Silflow et al., 2001). In human cells, LRRCC1 and C2CD3 are required for primary cilium assembly (1) – this study and Thauvin-Robinet et al., 2014; Ye et al., 2014 – and a role in asymmetric anchoring of cytoskeletal elements to the centriole may also be conserved (2), which could indirectly affect the determination of procentriole assembly site.

Table 1
Antibodies used in this study.
AntibodyDilutionIFDilutionU-ExMDilutionWBRRID identifierSourceReference
Primary antibodies
Goat anti-ARL13B1:100//RRID:AB_2058502Santa Cruz Biotechnologysc-102318
Guinea pig anti-alpha tubulin AA344 monobody/1:500/ Geneva Antibody FacilityscFv-S11B
Guinea pig anti-beta tubulin AA345 monobody/1:500/ Geneva Antibody FacilityscFv-F2C
Mouse anti-acetylated tubulin (6-11B-1)1:10001:500/RRID:AB_628409Santa Cruz Biotechnologysc-23950
Mouse anti-Centrin (20H5)1:500//RRID:AB_10563501Sigma-Aldrich04–1624
Mouse anti-CEP290 (B-7)1:500//RRID:AB_2890036Santa Cruz Biotechnologysc-390462
Mouse anti-gamma tubulin (GTU88)1:20001:200/RRID:AB_532292Sigma-AldrichT5326
Mouse anti-SAS-6/1:100/RRID:AB_1128357Santa Cruz Biotechnologysc-81431
Mouse anti-Smoothened1:200//RRID:AB_1270802Abcamab72130
Rabbit anti-ARL13B1:500//RRID:AB_2060867Proteintech17711–1-AP
Rabbit anti-C2CD31:5001:500/RRID:AB_10669542Sigma-AldrichHPA038552
Rabbit anti-C2CD3//1:1000RRID:AB_2718714Thermo Fisher ScientificPA5-72860
Rabbit anti-CEP83/1:500/RRID:AB_10674547Sigma-AldrichHPA038161
Rabbit anti-CEP1641:5001:300/RRID:AB_2651175Proteintech22227–1-AP
Rabbit anti-GFP//1:1000RRID:AB_591816MBL International598
Rabbit anti-HA//1:1000RRID:AB_631618Santa Cruz Biotechnologysc-805
Rabbit anti-hPOC51:500// Azimzadeh et al., 2009
Rabbit anti-KI671:1000//RRID:AB_443209Abcamab15580
Rabbit anti-LRRCC1 Ab11:5001:200/ This study
Rabbit anti-LRRCC1 Ab21:5001:3001:1000 This study
Rabbit anti-OFD11:500//RRID:AB_2890033.Sigma-AldrichABC961
Sheep anti-C2CD31:2001:100/RRID:AB_10997138R&D SystemsAF7348
Secondary antibodies
Donkey anti-goat IgG H&L (Alexa Fluor 488)1:5001:500/RRID:AB_2687506Abcamab150129
Donkey anti-goat IgG H&L (Alexa Fluor 568)1:5001:500/RRID:AB_2636995Abcamab175474
Donkey anti-goat IgG H&L (Alexa Fluor 647)1:5001:100/RRID:AB_2732857Abcamab150131
Donkey anti-mouse IgG H&L (Alexa Fluor 488)1:5001:500/RRID:AB_2732856Abcamab150105
Donkey Anti-Mouse IgG H&L (Alexa Fluor 568)1:5001:500/RRID:AB_2636996Abcamab175472
Donkey anti-mouse IgG H&L (Alexa Fluor 647)1:5001:100/RRID:AB_2890037Abcamab150107
Donkey anti-rabbit IgG H&L (Alexa Fluor 488)1:5001:500/RRID:AB_2636877Abcamab150073
Donkey anti-rabbit IgG H&L (Alexa Fluor 647)1:5001:500/RRID:AB_2752244Abcamab150075
Donkey anti-sheep IgG H&L (Alexa Fluor 647)/1:100/RRID:AB_2884038Abcamab150179
Goat anti-guinea pig IgG (H+L) (Alexa Fluor 568)/1:100/RRID:AB_141954Thermo Fisher ScientificA-11075
Goat anti-rabbit IgG (H+L) horseradish peroxidase conjugate//1:1000RRID:AB_2536530Thermo Fisher ScientificG-21234

  1. IF: immunofluorescence; U-ExM: ultrastructure expansion microscopy WB: Western blot.

Taken together, our results support that C2CD3 localizes asymmetrically in the distal lumen of human centrioles, a pattern that depends in part on LRRCC1.

Discussion

Here, we show that centrioles within the human centrosome are rotationally asymmetric despite the apparent ninefold symmetry of their ultrastructure. This asymmetry is manifested by a specific enrichment in LRRCC1 near two consecutive triplets, and the C-shaped pattern of C2CD3. Depletion of LRRCC1 perturbed the recruitment of C2CD3 and induced defects in centriole structure, ciliogenesis, and ciliary signaling, supporting that LRRCC1 contributes to organizing the distal centriole region together with C2CD3. LRRCC1 localizes like its C. reinhardtii ortholog Vfl1p, and C2CD3 delineates a filamentous structure reminiscent of the acorn first described in C. reinhardtii (Geimer and Melkonian, 2005; Geimer and Melkonian, 2004) and later found in a wide variety of eukaryotic species (Cavalier-Smith, 2021; Vaughan and Gull, 2015). Collectively, our results support that rotational asymmetry is a conserved property of centrioles linked to ciliary assembly and signaling in humans.

LRRCC1 and C2CD3 belong to a conserved rotationally asymmetric structure

Our work identifies two proteins located asymmetrically in the distal centriole lumen of the human centrosome, each with a specific pattern. LRRCC1 localizes principally near two consecutive triplets, with the first triplet counterclockwise labeled approximately 50% more than the next one. This pattern is highly reminiscent of the LRRCC1 ortholog Vfl1p, which localizes predominantly to the triplet facing the second centriole (referred to as triplet 1), and to a lesser extent to its immediate neighbor counterclockwise (triplet 2; Figure 7b; Silflow et al., 2001). In C. reinhardtii, triplets 1 and 2 are positioned directly opposite to the direction of flagellar beat, which is directed towards triplet 6 (Figure 7b; Lin et al., 2012). In mouse MCCs, Lrrcc1 is associated to triplets located not exactly opposite to the basal foot but with a clockwise shift of at least 20° from the basal foot axis. However, the beating direction was shown to be also shifted approximately 20° clockwise relative to the position of the basal foot in bovine tracheal MCCs (Schneiter et al., 2021; Figure 2d). The position of Lrrcc1/Vfl1p-labeled triplets with respect to ciliary beat direction might thus be similar in C. reinhardtii and in animal MCCs. Overall, the specific localization pattern of Vfl1p-related proteins at the distal end of centrioles, and their requirement for centriole positioning and ciliary beat orientation when motile cilia are present, appears to be conserved between flagellates and animals.

The second protein conferring rotational asymmetry to human centrioles, C2CD3, delineates a C-shape in the distal lumen. Strikingly, this staining is reminiscent of a filament observed by electron microscopy, which is said to form an ‘incomplete circle’ in the distal lumen of human centrioles (Vorobjev and Chentsov, 1980). Several lines of evidence favor the hypothesis that the C2CD3-containing structure is homologous to the acorn, a conserved filamentous structure that in C. reinhardtii connects five consecutive triplets along the centriole wall and across the lumen (Figure 7b; Cavalier-Smith, 2021; Geimer and Melkonian, 2004; Vaughan and Gull, 2015). First, the C2CD3 labeling is consistent with a circular filament. Second, C2CD3 is partially co-localized with LRRCC1 near the microtubule wall. Lastly, C2CD3 orthologs are found in a variety of flagellated unicellular eukaryotes, including the green algae Micromonas pusilla (Zhang and Aravind, 2012) and Chlamydomonas eustigma (Uniprot_A0A250XH15), suggesting an ancestral association to centrioles and cilia. The partial co-localization of Vfl1p and the acorn in C. reinhardtii, and the observation that both are already present at the distal end of procentrioles, led to propose that Vfl1p might also be a component of the acorn (Geimer and Melkonian, 2004). Consistent with this idea, both LRRCC1 and C2CD3 are recruited early to the distal end of human procentrioles, and LRRCC1 is required for proper assembly of the C2CD3-containing structure. C2CD3 recruitment at the centrioles also depends on the proteins CEP120 and Talpid3 (Tsai et al., 2019). Future work will help deciphering the relationships between these different proteins and characterize in more detail the architecture of the rotationally asymmetric structure at the distal end of mammalian centrioles.

Rotationally asymmetric centriole components are required for ciliogenesis

Our results uncover a link between centriole rotational asymmetry and primary ciliogenesis in human cells. Mutations in C2CD3 have been involved in several sensory ciliopathies, including JBTS (Boczek et al., 2018; Cortés et al., 2016; Ooi, 2015; Thauvin-Robinet et al., 2014). The associated ciliary defects are likely caused by anomalies in the structure of centrioles since depleting C2CD3 inhibits centriole elongation and DA assembly, whereas C2CD3 overexpression leads to centriole hyper-elongation (Thauvin-Robinet et al., 2014; Wang et al., 2018; Ye et al., 2014). We observed similar defects in LRRCC1-depleted cells, but of comparatively lesser extent. DA morphology was altered and centriole length was slightly increased in cells depleted from LRRCC1. The fact that LRRCC1 depletion has a more limited impact on centriole assembly than perturbation of C2CD3 levels suggests that LRRCC1 might not be directly involved in centriole length control or DA formation, however. The defects observed in LRRCC1-depleted cells could instead result indirectly from the abnormal localization of C2CD3. Besides the defects in centriole structure, it is plausible that LRRCC1 depletion also perturbs the organization of the ciliary gate as LRRCC1-depleted cells exhibited a drastic reduction in Hedgehog signaling. Loss of ciliary gate integrity interferes with the accumulation of SMO in the cilium upon activation of the Hedgehog pathway and is a frequent consequence of ciliopathic mutations (Garcia-Gonzalo and Reiter, 2017). The ciliary gate consists of the (Transition zone) TZ and the DA region, which both contribute to regulating protein trafficking in and out of the cilium (Garcia-Gonzalo and Reiter, 2017; Nachury, 2018). The anomalies in DA morphology observed in LRRCC1-depleted cells could disrupt the organization of the so-called DA matrix (Yang et al., 2018), thus preventing SMO accumulation in the cilium. Another, nonmutually exclusive possibility is that the architecture of the TZ, which forms directly in contact with the distal end of the centriole, is altered by LRRCC1 depletion. In either case, our observations in RPE1 cells are consistent with the JBTS diagnosis in two siblings carrying a mutation in the LRRCC1 gene (Shaheen et al., 2016), further establishing that LRRCC1 is a novel ciliopathy gene. Besides signaling, ciliary gate integrity is required for axoneme extension, and indeed, LRRCC1-depleted cells formed cilia at lower frequency than control cells – a defect that might also result from perturbed DA architecture. In the vfl1 mutant of C. reinhardtii, both unanchored centrioles and centriole docked at the plasma membrane but lacking a flagellum were observed (Adams et al., 1985). This supports that LRRCC1/Vfl1p requirement for properly assembling the ciliary gate is a conserved functional aspect of this family of proteins (Figure 7c).

Why is there a rotationally asymmetric structure at the base of primary cilia, and how does this structure form and contribute to the assembly of the DAs and the cilium remain open questions. In C. reinhardtii and in MCCs, LRRCC1 function is linked to the assembly of asymmetric appendages, which must be correctly positioned in relation to ciliary beat direction (Figure 7c). An asymmetric structure present early during centriole assembly and ultimately located near the cilium appears well suited for this task. The conservation of such a structure at the base of the primary cilium could perhaps indicate that primary cilia also possess rotationally asymmetric features, which would open interesting new perspectives on ciliary roles in heath and disease.

Other roles for centriole rotational asymmetry in human cells

Our finding that procentrioles do not form completely at random with respect to LRRCC1 location in the parent centriole suggests that centriole rotational polarity can influence centriole duplication in human cells. In C. reinhartdtii, procentrioles are formed at fixed positions with respect to the parent centrioles, to which they are bound by a complex array of fibrous and microtubular roots (Figure 7c; Geimer and Melkonian, 2004; Yubuki and Leander, 2013). The process is likely different at the centrosome since the roots typical of flagellates are not conserved in animal cells (Azimzadeh, 2021; Yubuki and Leander, 2013). In mammalian cells, procentrioles form near the wall of the parent centriole following the recruitment of early centriole proteins directly to the PCM components CEP152 and CEP192 (Yamamoto and Kitagawa, 2021). It is nonetheless conceivable that an asymmetry in triplet composition could result in local changes in PCM composition, which in turn could negatively impact PLK4 activation in this region. For instance, our analyses in planarian MCCs led us to postulate that linkers might be tethered to one side of the centrioles in a VFL1-dependent manner and independently of centriole appendages (Basquin et al., 2019). Future work will allow deciphering how centriole rotational asymmetry influences centriole duplication, and whether it affects other aspects of centriole positioning and cellular organization.

Materials and methods

Cell culture

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RPE1 cells (hTERT-RPE1, RRID:CVCL_4388; American Type Culture Collection, authenticated by STP profiling, no mycoplasma detected) were cultured in DMEM/F-12 medium (Thermo Fisher Scientific) supplemented with 10% fetal calf serum (Thermo Fisher Scientific), 100 U/mL penicillin and 100 μg/mL streptomycin (Thermo Fisher Scientific). Ciliogenesis was induced by culturing RPE1 cells in medium without serum during 48 hr. HEK 293 cells (kind gift from F. Causeret, Institut Imagine, Paris) were cultured in DMEM medium (Thermo Fisher Scientific) supplemented with 10% fetal calf serum and antibiotics as previously. All cells were kept at 37°C in the presence of 5% CO2.

Mouse ependymal cells and tracheal tissue

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All experiments were performed in accordance with the French Agricultural Ministry and European guidelines for the care and use of laboratory animals. In vitro differentiated ependymal cells were a kind gift from A.R. Boudjema and A. Meunier (IBENS, Paris). They were prepared as described previously (Delgehyr et al., 2015; Mercey et al., 2019) from Cen2GFP mice (CB6-Tg(CAG-EGFP/CETN2)3-4Jgg/J, The Jackson Laboratory). The fragment of trachea was obtained from a wildtype mouse of the Swiss background (kindly provided by I. Le Parco, Institut Jacques Monod).

CRISPR/Cas9 editing

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LRRCC1 mutant clones were obtained by two different CRISPR/Cas9 strategies. First, RPE1 cells were co-transfected with plasmid px154-1 (U6p-gRNA#1_U6p-gRNA#2_CMVpnCas9-EGFP_SV40p-PuroR-pA with gRNA#1: 5′-AGA ATT CTA CCC TAC CTG-3′ and gRNA#2: 5′-TAA GGT AGT GCT TCC TAC-3′) targeting the LRRCC1 locus in exon 8, and px155-24 (U6p-gRNA#3_U6p-gRNA#4_CMVpnCas9-mCherry_SV40p-PuroR-pA; gRNA#1: 5′-ATC TAC TCG GAA AGC TGA-3′ and 5′-GCT TGA GGG CTC AAA TAC-3′) targeting exon 9. Both constructs express the nickase mutant of Cas9 fused to either EGFP or mCherry. Two days after transfection, EGFP- and mCherry-positive cells were sorted by flow cytometry and grown at low concentration. Individual clones were picked after 2 weeks and analyzed by PCR to detect short insertions/deletions. A single clone was obtained (clone 1.1), which was further characterized by genomic sequencing. Both alleles of LRRCC1 contained deletions (~0.6 kb deletion of exon 9 and an ~1.5 kb deletion of exon 8; Figure 4—figure supplement 1a), leading to frameshifts. In a second approach, cells were co-transfected using a mix of three CRISPR/Cas9 Knockout Plasmids (sc-413781; Santa Cruz Biotechnology) targeting exons 11 (5′-CTT GTT CTC TTT CTC GAT GA-3′ and 5′-ACT TCT TGC ATT GAA AGA AC-3′) or 12 (5′-CGT GTT AAG CCA GCA GTA TA-3′) of LRRCC1, together with the corresponding homology-directed repair plasmids carrying a puromycin-resistance cassette (sc-413781-HDR; Santa Cruz Biotechnology), following the recommendations of the manufacturer. Mutant clones were selected by addition of 2 µg/mL puromycin in the culture medium and further screened by immunofluorescence, allowing to identify two clones with decreased LRRCC1 levels (clones 1.2 and 1.9). Genomic insertion of the HDR cassette could not be detected in these clones by PCR, and no sequence anomalies were identified in PCR fragments corresponding to exons 10–13. This suggests that one copy of the LRRCC1 gene is intact, while the second copy may have undergone more extensive modifications via large deletions/insertions. For sequencing of LRRCC1 transcripts, total RNA extracts were obtained using the Nucleospin RNA kit (Macherey-Nagel) and cDNAs were synthesized using SuperScript III reverse transcriptase (Thermo Fisher Scientific). PCR primers specific to exons 4 and 8, 4 and 9, 8 and 19, or 9 and 19 were used to amplify cDNAs from clone 1.1; primers specific to exons 4 and 17 were used for clones 1.2 and 1.9. The resulting fragments were analyzed by sequencing.

Inducible HEK 293 cell lines

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LRRCC1 full-length coding sequence was amplified from cDNA clone IMAGE:5272572 (GenBank accession: BC070092.1), corresponding to the longest isoform of LRRCC1 (NM_033402.5), after correction of a frameshift error by PCR mutagenesis. As N- and C-terminal GFP fusions were not targeted to the centrosome, we inserted the GFP tag within the LRRCC1 sequence in disordered regions present between the leucine-rich repeat and coiled-coil domains, either after amino acid 251 or 402. The fusions were cloned into the pCDNA-5FRT (Thermo Fisher Scientific) vector using the Gibson assembly method (Gibson et al., 2009) and then integrated into the Flp-In-293 cell line using the Flp-In system (Thermo Fisher Scientific). Expression of the GFP-LRRCC1 fusions was induced by culturing the Flp-In-293 cell lines overnight in medium supplemented with 1 µg/mL doxycycline (Thermo Fisher Scientific).

RNAi

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Ready to use double-stranded siRNA LRRCC1-si1 (target sequence: 5′-AAG GAG AAA GAT GGA GAC GAT-3′) (Muto et al., 2008), LRRCC1-si2 (target sequence: 5′-TTA GAT GAC CAA ATT CTA CAA-3′), and control siRNA (AllStars Negative Control) were purchased from QIAGEN. siRNAs were delivered into cells using Lipofectamine RNAiMAX diluted in OptiMEM medium (Thermo Fisher Scientific). Cells were fixed after 48 hr and processed for immunofluorescence. For RNAi depletion of ciliated cells, RPE1 cells grown in complete culture medium were treated by RNAi, incubated for 2 days, then submitted to a second round of RNAi. After 8 hr, cells were washed 3× in PBS then cultured during 24 hr in serum-free medium to induce ciliogenesis.

qRT-PCR

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Total RNA extracts were obtained using the Nucleospin RNA kit (Macherey-Nagel) and cDNAs were synthetized using SuperScript III reverse transcriptase (Thermo Fisher Scientific). qPCR was performed in triplicate with the GoTaq qPCR Master Mix (Promega) in a LightCycler 480 instrument (Roche) using the primers listed in Table 2. Quantification of relative mRNA levels was performed using CHMP2A and EMC7 as reference genes following the MIQE guidelines (Bustin et al., 2009).

Table 2
Primers used for quantitative RT-PCR.
PrimerSequence
LRRCC1-1_FwCAA CAA GGA TCT TCT CTA GCC CA
LRRCC1-1_RvAGT TTG GTC GTC TAT GAT TTT GCA
LRRCC1-2_FwGCA CAA CAA GGA TCT TCT CTA GC
LRRCC1-2_RvTCG CAG ACA TTC ATT CTC TCT AGA
PTCH1_FwCCC CTG TAC GAA GTG GAC ACT CTC
PTCH1_RvAAG GAA GAT CAC CAC TAC CTT GGC T
CHMP2A_FwATG GGC ACC ATG AAC AGA CAG
CHMP2A_RvTCT CCT CTT CAT CTT CCT CAT CAC
EMC7_FwGTC AGA CTG CCC TAT CCT CTC C
EMC7_RvCAT GTC AGG ATC ACT TGT GTT GAC

Antibodies

Fragments encoding either aa 671–805 (Ab1) or aa 961–1032 (Ab2) of LRRCC1 (NP_208325.3) were cloned in pGST-Parallel1 and expressed in Escherichia coli. The GST-fusion proteins were purified under native conditions using glutathione agarose (Thermo Fisher Scientific), and the LRRCC1 fragments were recovered by Tev protease cleavage and dialyzed before rabbit immunization (Covalab). Antibodies were affinity-purified over the corresponding GST-LRRCC1 fusion bound to Affi-Gel 10 resin (Bio-Rad). Other primary and secondary antibodies used in this study are listed in Table 1.

Western blot

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For whole-cell extracts, Flp-In-293 cell lines expressing the GFP-LRRCC1 fusions were induced overnight with doxycycline, collected by centrifugation, and resuspended in Western blot sample buffer prior to incubation at 95°C for 5 min. For immunoprecipitation experiments, doxycycline-induced cells expressing LRRCC1 with a GFP inserted after aa 402 were resuspended in lysis buffer (50 mM Tris pH 8, 150 mM NaCl, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS) supplemented with 1 mM MgCl2, 20 µg/mL DNAse I (Roche), and a protease inhibitor cocktail (cOmplete Mini, EDTA-free, Roche). After 30 min on ice, the lysates were centrifuged at 15,000×g for 10 min at 4°C. The supernatants were then incubated with Dynabeads M-280 sheep-anti rabbit magnetic beads (Thermo Fisher Scientific) previously incubated with rabbit anti-IgGs, either anti-GFP or anti-HA tag for the control IP (Table 1), and rotated for 3 hr at 4°C. After 3 washes with lysis buffer, immunoprecipitated proteins were recovered by resuspending the beads in sample buffer and heating at 95°C for 5 min. The samples were then run on 4–20% Mini-Protean TGX precast protein gels (Bio-Rad) and transferred onto PVDF membrane using the iBlot 2 blot system (Thermo Fisher Scientific). The membranes were blocked and incubated with antibodies following standard procedures, then visualized using Pierce ECL plus chemiluminescence reagents (Thermo Fisher Scientific) on a ChemiDoc imaging system (Bio-Rad).

Immunofluorescence

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Cells were fixed in cold methanol for 5 min at – 20°C, blocked 10 min with 3% BSA (Sigma-Aldrich) in PBS containing 0.05% Tween-20 (PBST-0.05%), then incubated with primary antibodies diluted in PBST-0.05% containing 3% BSA for 1 hr. After washing 3 × 1 min in PBST-0.05%, cells were incubated 2 hr with secondary antibodies in PBST-0.05% containing 3% BSA and 5 µg/mL Hoechst 33342 (Thermo Fisher Scientific), washed in PBST-0.05% as previously, and mounted using Fluorescence Mounting Medium (Agilent). For staining of primary cilia with anti-acetylated tubulin, cells were incubated 2 hr on ice prior to methanol fixation. For quantification of SMO accumulation within cilia, confluent cells cultured during 24 hr in serum-free medium were supplemented with 200 nM SAG (Sigma) diluted in DMSO, or DMSO alone for 24 hr. Cells were then co-stained for SMO and ARL13B to determine the position of the primary cilium. For all experiments involving induction of ciliogenesis by serum deprivation, we verified that cells were arrested in G0 by immunofluorescence staining of Ki67. To visualize centriolar LRRCC1 and quantify CEP290 centrosomal levels, cells were treated during 1 hr with 5 µM nocodazole prior to fixation. Images were acquired using an Axio Observer Z.1 microscope (Zeiss) equipped with a sCMOS Orca Flash4 LT camera (Hamamatsu) and a ×63 objective (Plan Apo, N.A. 1.4). The structured illumination microscopy (SIM) image was acquired on an ELYRA PS.1 (Zeiss) equipped with an EMCCD iXon 885 camera (Andor) and a ×63 objective (Plan Apo, N.A. 1.4).

Ultrastructure expansion microscopy

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We used the U-ExM protocol described in Gambarotto et al., 2019 with slight modifications. Cells grown on glass coverslips were incubated in a fresh solution of 1% acrylamide and 0.7% formaldehyde diluted in PBS. After incubating 5 hr to overnight at 37°C, the coverslips were washed with PBS and placed cells down on a drop of 35 μL monomer solution (19.3% sodium acrylate, 10% acrylamide, 0.1% bis-acrylamide in PBS) to which 0.5% TEMED and 0.1% ammonium persulfate were added just before use. The coverslips were incubated 5 min on ice then 1 hr at 37°C, then transferred to denaturation buffer (200 mM SDS, 200 mM NaCl, 50 mM Tris pH9) for 15 min with agitation to detach the gels from the coverslips. The gels were then incubated in denaturation buffer 1.5 hr at 95°C, washed 2 × 30 min in deionized water, then incubated overnight in water at room temperature to allow expansion of the gel. The gels were measured at this step to determine the coefficient of expansion. After 2 × 10 min in PBS, the gels were cut into smaller pieces then incubated 3 hr at 37°C with primary antibodies diluted in saturation buffer (3% BSA, 0.05% Tween-20 in PBS). The gel fragments were then washed 3 × 10 min in PBST-0.1%, incubated 3 hr with secondary antibodies, and washed in PBST-0.1% as described previously. Finally, the gels were incubated 2 × 30 min in deionized water, then left to expand overnight in deionized water to regain their maximum size. For U-ExM of mouse tracheal cells, a fragment of WT mouse trachea (kind gift from I. Le Parco, IJM, Paris) was adhered on a poly-lysine-coated coverslip, then processed as described above with the following modifications: for the first step, the fragment of trachea was incubated overnight to 48 hr in 1% acrylamide and 0.7% formaldehyde in PBS; they were placed 15 min on ice prior to the 1 hr incubation at 37°C and the transfer to denaturation buffer. Note that GFP fluorescence was quenched during U-ExM processing, so the GFP-Cen2 construct expressed in ependymal cells was not detectable in final samples. Gels were imaged on Lab-Tek chamber slides (0.15 mm) coated with poly-lysine (Thermo Fisher Scientific). Images were acquired at room temperature using either a LSM780 confocal microscope (Zeiss) equipped with an oil ×63 objective (Plan Apo, N.A. 1.4) or an LSM980 confocal microscope with Airyscan 2 (Zeiss) equipped with an oil ×63 objective (Plan Apo, N.A. 1.4).

Image analysis

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Protein levels were determined using ImageJ software (Schneider et al., 2012) by measuring the fluorescence intensity in the centrosome or cilium area and subtracting the cytoplasmic background in z-series taken at 0.5 μm interval. Images of individual centrioles in U-ExM are maximum intensity projections of all z-sections comprising the signal of interest. Note that centrioles are presented as they are in the sample (i.e., without correcting their orientation), which leads to an apparent shift between channels or decreased circularity in the projections when centrioles are not parallel to the imaging axis. Analysis of DA morphology defects was performed on z-stacks and not on projected images. Daughter centriole length was determined by U-ExM using the acetylated tubulin staining. For mother centrioles, which could be associated with a primary cilium, the length was measured between the proximal end of the acetylated tubulin staining and DAs labeled by anti-CEP164 or CEP83. To generate average images of LRRCC1 and C2CD3, only centrioles that were nearly perpendicular to the imaging plane were acquired on the Airyscan microscope in order to maximize the resolution in transverse views. Calculating the average image consisted of several steps: cropping out individual centrioles, aligning them, providing reference points, standardizing centrioles using the reference points, and averaging (Figure 1—figure supplement 2). The cropping was done in ImageJ, and for aligning and providing the reference points a graphical user interface was developed based on Napari (Sofroniew et al., 2020). Centriole alignment: the direction of centriole long axis was selected manually and used to position the centriole vertically. Providing the reference points: reference points were manually selected to outline the circle of microtubules triplets and the location of the protein of interest. The centriole was also framed in Z dimension with a rectangle. Standardization: the reference points were used to calculate all necessary transformations (rotation, scaling, and translation) to map the original image of a centriole to the standard image. Averaging: an average image was calculated for all the successive XY planes of the standardized image stacks. For alignment of tracheal cell centrioles, since the current version of the graphical user interface can only accommodate two channels, the position of the basal foot provided by the γ-tubulin channel was reported manually in the acetylated tubulin channel using ImageJ. The images were then processed as before using the manual annotation as a reference point for the basal foot.

For analysis of procentriole position and LRRCC1 location in procentrioles, 3D reconstructions of diplosomes processed for U-ExM were obtained using Imaris software (Oxford Instruments).

Electron microscopy

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RPE1 cells were grown at confluence before induction of ciliogenesis for 72 hr by serum deprivation. Cells were fixed 30 min in 2.5% glutaraldehyde (Electron Microscopy Sciences), 2% paraformaldehyde (Electron Microscopy Sciences), 1 mM CaCl2 in PBS, then washed 3 × 5 min in PBS. Samples were then post-fixed during 30 min in 1% osmium tetroxide (Electron Microscopy Sciences), then washed 3 × 5 min in water. Dehydration was performed using graded series of ethanol in water for 5 min 30, 50, 70, 90, 100, and 100%. Resin infiltration was performed by incubating 30 min in an Agar low-viscosity resin (Agar Scientific Ltd) and EtOH (1:2) mix, then 30 min in a resin and EtOH (2:1) mix followed by overnight incubation in pure resin. The resin was then changed and the samples further incubated during 1.5 hr prior to inclusion in gelatin capsules and overnight polymerization at 60°C. 70 nm sections were obtained using an EM UC6 ultramicrotome (Leica), post-stained in 4% aqueous uranyl acetate and lead citrate, and observed at 80 kV with a Tecnai12 transmission electron microscope (Thermo Fisher Scientific) equipped with a 1K × 1K Keen View camera (OSIS).

Videomicroscopy

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To determine the duration of mitosis, individual frames of cells growing under normal culture conditions were acquired every 5 min for 24 hr using an IncuCyte ZOOM live-cell analysis system (Sartorius) equipped with a ×20 objective.

Statistical analysis

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All statistical analyses were performed using the Prism 9 for Mac OS X software (GraphPad Software, Inc). All values are provided as mean ± SD. The number of experimental replicates and the statistical test used are indicated in the figure legends, and the p-values are included when statistically different.

Data availability

All data generated or analyzed during this study are included in the manuscript and supporting files. Source data files are available from the Dryad database (doi:https://doi.org/10.5061/dryad.95x69p8m5).

The following data sets were generated
    1. Azimzadeh J
    (2022) Dryad Digital Repository
    Data from: Evolutionary conservation of centriole rotational asymmetry in the human centrosome.
    https://doi.org/10.5061/dryad.95x69p8m5

References

Decision letter

  1. Jens Lüders
    Reviewing Editor; Institute for Research in Biomedicine, Spain
  2. Anna Akhmanova
    Senior Editor; Utrecht University, Netherlands

Our editorial process produces two outputs: i) public reviews designed to be posted alongside the preprint for the benefit of readers; ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "Evolutionary conservation of centriole rotational asymmetry in the human centrosome" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Anna Akhmanova as the Senior Editor. The reviewers have opted to remain anonymous.

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

1) The most important finding is the demonstration of rotational asymmetry of LRRCC1 localization. The reviewers felt that this finding needs to be confirmed by an independent imaging method. As indicated by the reviewers, there are multiple options, but 3D-SIM may be the most accessible.

2) A careful analysis of centriole numbers in LRRCC1 deficient cells (e.g. at the end of the duplication cycle in mitotic cells) should be performed to determine whether correct duplication occurs.

3) All reviewers had concerns regarding the functional data in the partially depleted KO line. Could the authors use RNAi in the KO cell lines to achieve further depletion and more robust phenotypes? Related to this, the authors should use super-resolution imaging to demonstrate reduced levels at centrioles (not only at centrosomes or pericentriosomal region/satellites) in the KO lines.

Reviewer #1 (Recommendations for the authors):

The authors convincingly demonstrate rotational asymmetry in human centrosome centrioles and a contribution of LRRCC1 to cilium assembly and ciliary signaling. Overall the data is of high quality and mostly support the conclusions. There are a few points, however, that need to be addressed:

1) Does LRRCC1 affect centriole assembly? Is there a change in centriole number in the partial KO lines or after RNAi? This is an obvious question when analyzing the function of a centriole component and is also an important control when claiming cilium assembly defects. The authors should quantify centriole number in mitotic cells.

2) Considering the mild phenotypes after partial depletion of LRRCC1, could the authors use RNAi in the KO cell lines to achieve stronger depletion and more robust phenotypes?

3) Figure 5. Changes in centriole length: were these measurements done at specific cell cycle stages? Can the authors exclude that cell cycle effects in cells with reduced LRRCC1 levels may cause differences in measured centriole length? Related: is mitotic progression normal in these cells? Delayed mitotic progression was shown to also cause centriole elongation (e.g. doi: 10.1083/jcb.201910019). Since full knockout seem to prevent cell growth, it is possible that partial KO may also cause proliferation defects. FACS analysis and monitoring the timing of mitotic progression would address this.Reviewer #2 (Recommendations for the authors):

The data appear too preliminary for immediate publication in eLife. Nevertheless, the subject is intriguing and if authors could provide a cleaner evidence of LRRCC1 and C2CD3 localization, show how depletion of LRRCC1 affects centriole, appendage and cilia structure, the manuscript would be suitable for publication.

Point 1. Expansion microscopy is a powerful imaging approach. However, it is used here to determine localization of a relatively uncharacterized protein, using uncharacterized antibodies. This is risky. For instance: In figure 5 c-e, the authors judge the arrangement of Cep164 to classify centrosomes into 'normal, abnormal, and missing'. This characterization is made based on uneven distribution of Cep164 signals. However, if this criterion is to be extended to acetylated tubulin signal across all expansion images, many of control centrioles (Figure 1c, centriole2 and 3, Figure 3, centriole in b, 1.1 and 1.9 centrioles in Figure 6 f) must also be deemed 'aberrant', due to distortions, uneven spacings between MT blades etc. Clearly, expansion microscopy has its pitfalls.

There are also huge inconsistencies in the pattern and a great variability in the intensity of the LRRCC1 signals between figures and figure panels. For instance, in non-averaged images of expanded centrioles, LRRCC1 can be found in centriole's lumen, in association with MT wall, and in association with variable number of MT blades. In Figure 2b, and 1 b, LLRRCC1 signal is even associated with procentriole's proximal end. How much of the LLRRCC1 signal is indeed specific? The situation seems even more dramatic with C2CD3 data, where some of the signals lie entirely outside the centriole (for instance in 1.9 clone, Figure 6F). Can authors comment on how LRRCC1 and C2CD3 signals can protrude and be localized beyond centriole wall? Previous work (PMID: 30988386, PMID: 29789620) shows C2CD3 to be centered within centriole distal end, and not asymmetric.

It would be critical to use an additional super resolution microscopy technique to re-analyze the localization of LRRCC1 and C2CD3 in non-expanded samples and not to rely solely on expansion microscopy.

Point 2. It is not stated in the manuscript how and whether axial and lateral shifts between different channels were corrected? Was such correction done before image averaging or before image assembly?

Instead of showing averaged data, panels showing more individual examples of horizontally and vertically oriented centrioles would be more instructive. Additional supplemental figures could be assembled for this purpose, at least for the purpose of evaluation.

Point 3. Quantification of LRRCC1 levels in CRISPR clones (Figure 4 a) does not reflect centriole-associated levels of LRRCC1. It reflects the loss of pericentrosomal (satellite) population of LRRCC1, which is the most abundant. Quantification of the centriole bound LCCRR1 needs to be provided instead. Are centrosomal levels of LRRCC1 in 1.1, 1.2, and 1.9 CRISPR clones (Figure 4 a) decreased in comparison to the ones from nocodazole -treated cells (Figure 1A)? In addition, higher resolution images of LRRCC1 in CRISPR clones have not been provided to illustrate where the leftover protein is localized in CRISPR 1.1, 1.2, and 1.9 clones.

How can authors exclude the possibility that loss of the pericentrosomal population of LRRCC1 affects ciliation and ciliary signaling?

Point 4. Between 8 independent experiments conducted in wt RPE-1 cells, the percentage of centrioles with 'abnormal or missing" arrangement of Cep164 varies from ~0% – 40% (Figure 5C). This is a large variability since almost all wt RPE-1 cells can form a cilium (Figure 4b). In my view, this means either that the variability in Cep164 signal is introduced experimentally, or that it is physiological and cannot be used as a measure for ciliation. Indeed, previous super-resolution analysis has demonstrated that, although distal appendage proteins display characteristic pattern around centrioles, their levels on individual appendages can vary under physiological conditions (PMID: 30824690, PMID: 29789620). Therefore, significance of the data from Figure 5 c-e is unclear. Besides, additional appendage proteins (such as Cep83 and SCLT1, for instance) should be analyzed to reach a conclusion about distal appendage organization and number.

Point 5. The authors propose that LRRCC1 cooperates with C2CD3 in organization of distal centriole ends. However, partial reduction in LRRCC1 leads to elongated centrioles only in 1.9 CRISPR clone and not in other two clones, but it is unclear why, as differences between the centrosomal levels of LRRCC1 in CRISPR clones do not seem large enough to explain this difference. Increased centriole length in clone 1.9 could also be non-specific due to clonal variation, and it is unclear whether it is relevant. Especially, because both clones 1.1 and 1.9 seem to show similar defects in ciliation and Cep164 assembly.

To prove and understand what type of centriolar structural defects occur after LRRCC1 depletion, extensive ultrastructural analysis would need to be conducted. Perhaps LRRCC1 could be further depleted by siRNA in LRRCC1- CRISPR clones to generate more penetrant phenotypes?

Point 6. Figure 7. More evidence of LRRCC1/C2CD3 colocalization needs to be provided. Why is majority of C2CD3 signal colocalized with centriole MTs?

Reviewer #3 (Recommendations for the authors):

Critical to the central points of the paper are the asymmetric localization of LRRCC1 and C2CD3 at the distal centriole. It is not clear whether the expansion in expansion microscopy is always isotropic and anisotropic expansion could conceivably skew protein localization to appear asymmetric. Could an orthogonal superresolution method, such as 3D-SIM/STED/STORM be used to confirm asymmetry of LRRCC1 or C2CD3? Additionally, could performing 3D-reconstruction image analysis of LRRCC1/C2CD3 with a distal appendage component or another symmetric sub-component of the distal centriole is critical rule out potential artifacts that could be due to anisotropic expansion during U-ExM? The authors imaged OFD1 or CEP290 using U-ExM (fluorescent intensity quantification data are presented in Supplementary Figure S3, but no images are shown). Further confidence in the LRRCC1 conclusions would be garnered by data indicating that these proteins are symmetrically localized at the distal centriole.

The authors report that their results "demonstrate that rotational asymmetry is a conserved ancient property of centrioles." However, the basal foot is a previously described rotational asymmetry present in animal cells, no? Indeed, the basal foot seems to be anticorrelated with the localization of LRRCC1. The authors examine distal appendage formation in some depth. Is basal foot formation or location compromised in LRRCC1 depleted cells?

Quantitation of depletion of LRRCC1 is done by light microscopy. The remaining population, although quantitated, is not characterized spatially with any high degree of resolution. Should one subcellular population (centriolar versus satellite) be preferentially depleted by the reduction in LRRCC1 levels, one population might be implicated in the biological functions of LRRCC1. Where is the residual LRRCC1 staining in the CRISPR/Cas9 generated clones? For example, figure 4a would benefit from U-ExM and quantification of fluorescence intensity specifically at centrioles to determine if the centriolar pool of LRRCC1 is disrupted in the hypomorphs, as surmised by the authors.

It is curious that the authors have been unable to generate null mutations in LRRCC1. RPE1 cells fail to proliferate in the presence of centrinone, possibly hinting at a critical role for LRRCC1 in centriole duplication. Is under duplication or delayed duplication of centrioles observed in LRRCC1 depleted cells?

Relatedly, were null mutations in LRRCC1 generated in HEK 293 cells?

In the discussion, the authors speculate at some length about a role of LRRCC1 in the formation of the ciliary gate, given that LRRCC1 depletion inhibits the ciliary localization of Smoothened. However, one component of the ciliary gate, CEP290, is not reported to be altered in LRRCC1 knockdown. Can the authors examine the dependence of ciliary gate components required for Smoothened localization to cilia in the LRRCC1 knockdown cells?

I recognize that the rescue experiments were difficult. Could an inducible approach (e.g., TetOne system) be attempted? Rescue may help delineate which of the variably penetrant phenotypes are due to downregulation of LRRCC1 and which to other factors. For example, the authors observe a mild increase in centriole length in clone 1.9, but not in clone 1.1. The authors conclude that the slightly lower levels of LRRCC1 in clone 1.9 account for the difference. However, an alternative possibility is that clone 1.9 has another genetic or epigenetic variation that accounts for this incompletely penetrant phenotype.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "Evolutionary conservation of centriole rotational asymmetry in the human centrosome" for further consideration by eLife. Your revised article has been evaluated by Anna Akhmanova (Senior Editor) and a Reviewing Editor.

The manuscript has been much improved and you have addressed all concerns. There are a couple of items that I would like you to address before formal acceptance:

1) Figure 1d: The Airyscan localization experiment uses POC5 labelling as an example of a symmetrical localization. I suggest that you show the channels also separately so that one can appreciate the luminal POC5 signal better. In the legend you commented on the fact that the POC5 label is also on the outside – in fact the outside label seems stronger than the luminal signal. This is a bit confusing for the purpose of this experiment, but also considering previous POC5 localization studies by ExM (e.g. Le Guennec et al., Sci Adv, 2020; Schweizer et al., Nat Comms, 2021). Is this outside signal specific? It may be good to discuss/explain this a bit more.

2) It is somewhat surprising that you can reliably count individual centrioles using POC5 staining, which is proposed to mark the central lumen. Considering the unexpected distribution of the POC5 label in Fig, 1d, perhaps the more distal, outside label helps? In any case, it would be good to include example cells with POC5 staining of centriole pairs at spindle poles with the quantifications in Figure S1j.

https://doi.org/10.7554/eLife.72382.sa1

Author response

Essential revisions:

1) The most important finding is the demonstration of rotational asymmetry of LRRCC1 localization. The reviewers felt that this finding needs to be confirmed by an independent imaging method. As indicated by the reviewers, there are multiple options, but 3D-SIM may be the most accessible.

In the revised version of the manuscript, we now verify the asymmetry of LRRCC1 localization by a method that does not involve expansion. For this, we imaged conventional immunofluorescence samples using a Zeiss Airyscan 2 confocal microscope. The Airyscan 2 module increases the resolution by a factor of 1.8 compared to conventional confocal microscopy, which is close to the performance of 3D-SIM. The images we obtained are indeed comparable to those obtained using a Zeiss Elyra PS1 microscope (compare the Airyscan images now included in Figure 1d of the revised manuscript with the SIM image in Author response image 1).

Author response image 1
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To verify the asymmetry of LRRCC1 labeling, we measured the lateral distance between LRRCC1 intensity peak and the long axis of the centriole labeled with acetylated tubulin. To determine centriole orientation with greater accuracy, we measured only mother centrioles associated with a cilium. A comparison between the distribution of LRRCC1 and that of hPOC5, a protein localized symmetrically within centrioles (Le Guennec et al., 2020), confirms the asymmetric localization of LRRCC1 in samples without expansion. These results are now presented in Figure 1d. and discussed in the Results section (p7, lines 122-128).

2) A careful analysis of centriole numbers in LRRCC1 deficient cells (e.g. at the end of the duplication cycle in mitotic cells) should be performed to determine whether correct duplication occurs.

We never observed anomalies in centriole number in cells depleted in LRRCC1, but we had indeed neglected to formally quantify this. We have now included quantification of centriole number in LRRCC1-depleted cells in Supplemental Figure S1j of the revised manuscript. As suggested by the reviewers, we analyzed the number of centrioles at mitotic spindle poles using the hPOC5 protein as a centriole marker. We detected no difference between cells depleted in LRRCC1, either by CRISPR or RNAi, and control cells. We conclude that LRRCC1-depletion does not affect centriole duplication.

3) All reviewers had concerns regarding the functional data in the partially depleted KO line. Could the authors use RNAi in the KO cell lines to achieve further depletion and more robust phenotypes?

As suggested, we treated the CRISPR clones with RNAi to further reduce LRRCC1 levels. Overall LRRCC1 levels in the centrosome area were decreased after RNAi compared to CRISPR clones (Figure 4a of the revised manuscript). We then analyzed several aspects of the LRRCC1 phenotype in CRISPR cells treated with RNAi.

– We first show that further reducing LRRCC1 levels in CRISPR clones does not enhance the ciliogenesis defect, as the proportion of cells forming a primary cilium in these conditions is comparable to what is observed in CRISPR clones treated with a control siRNA. We conclude that depletion of LRRCC1 only partially inhibits ciliogenesis. These results are now presented in Figure 4c of the revised manuscript and discussed in the Results section (p11-12, lines 246-248).

– We show that RNAi treatment leads to a significant increase in mother centriole length in CRISPR clone 1.1 compared to the control. We conclude that depletion of LRRCC1 increases centriole length. These results are now presented in Figure 5b of the revised manuscript and discussed in the Results section (p12-13, lines 271-275).

– We analyzed the effect of RNAi treatment on the recruitment of a second component of distal appendages, CEP83. We show that the proportion of centrioles with abnormal CEP83 labelling is significantly less in RNAi-treated CRISPR clones 1.1 and 1.9 than in control cells, confirming that LRRCC1-depletion leads to abnormalities in distal appendages. These results are now presented in Figure 5g, h of the revised manuscript and discussed in the Results section (p13, lines 284-294).

– We determined how further reducing LRRCC1 levels affects C2CD3 localization pattern. The average images obtained after RNAi treatment of CRISPR clones show that while the treatment does not alter further the C2CD3 pattern in clone 1.9, which has lower levels of centriolar LRRCC1 (see below), it does so in clone 1.1. These results confirm that LRRCC1 depletion affects C2CD3 localization and are now presented in Figure 6g of the revised manuscript (see also Results section, p14-15, lines 321-322). Please note that we generated new average images of the C2CD3 staining for all conditions (Figure 6d and 6g; see the response to point 2 of reviewer #2 for details).

Altogether, our results confirm that LRRCC1 depletion interferes with ciliogenesis, increases centriole length, and leads to abnormal distal appendage morphology and C2CD3 recruitment.

Related to this, the authors should use super-resolution imaging to demonstrate reduced levels at centrioles (not only at centrosomes or pericentriosomal region/satellites) in the KO lines.

In the revised manuscript, we now quantify LRRCC1 levels at the distal end of centrioles by Airyscan microscopy. To identify the distal centriole pool with greater accuracy, we only analyzed mother centrioles associated with a primary cilium. Our results confirmed a decrease in the amounts of centriolar LRRCC1 in all 3 CRISPR clones, with lower levels in clone 1.9, for which we observed the strongest phenotypes. These results are now presented in Figure 4b of the revised manuscript (see also Results section, p11, lines 235-239).

Unexpectedly, the amount of distal protein detected in clone 1.1 was comparatively higher than in the other clones (on the order of 80% of the WT level, compared with < 50% for clone 1.9). Clone 1.1 has deletions in both copies of the LRRCC1 gene, which is expected to cause a loss of the reading frame in both cases (now in Supplemental Figure S3a of the revised manuscript). To determine what the proteins detected in clone 1.1 correspond to, we performed an analysis of the transcripts present in this clone. As now shown in Supplemental Figure S3b, we identified two long in-frame transcripts. In both cases, an exon adjacent to the exon deleted in the genomic sequence was eliminated by splicing, allowing frame recovery. These splicing profiles are not found in the databases, which supports that the corresponding proteins are mutant isoforms. In contrast, WT transcripts are detected, but at reduced levels compared to WT, in clones 1.2 and 1.9. Together, these analyses show that all 3 clones exhibit reduced levels of centriolar LRRCC1, and that in the case of clone 1.1 only mutant isoforms are present.

Reviewer #1 (Recommendations for the authors):

The authors convincingly demonstrate rotational asymmetry in human centrosome centrioles and a contribution of LRRCC1 to cilium assembly and ciliary signaling. Overall the data is of high quality and mostly support the conclusions. There are a few points, however, that need to be addressed:

1) Does LRRCC1 affect centriole assembly? Is there a change in centriole number in the partial KO lines or after RNAi? This is an obvious question when analyzing the function of a centriole component and is also an important control when claiming cilium assembly defects. The authors should quantify centriole number in mitotic cells.

Please see the response to point 2 of the Essential Revisions.

2) Considering the mild phenotypes after partial depletion of LRRCC1, could the authors use RNAi in the KO cell lines to achieve stronger depletion and more robust phenotypes?

Please see the response to point 3 of the Essential Revisions.

3) Figure 5. Changes in centriole length: were these measurements done at specific cell cycle stages? Can the authors exclude that cell cycle effects in cells with reduced LRRCC1 levels may cause differences in measured centriole length? Related: is mitotic progression normal in these cells? Delayed mitotic progression was shown to also cause centriole elongation (e.g. doi: 10.1083/jcb.201910019). Since full knockout seem to prevent cell growth, it is possible that partial KO may also cause proliferation defects. FACS analysis and monitoring the timing of mitotic progression would address this.

Centriole length measurements were not taken at a specific time in the cell cycle. However, we measured the length of mother and daughter centrioles separately (Figure 5a) and found both to be increased in clone 1.9. Since the length of the mother centriole does not vary over time, it should not be affected by the cell cycle stage at which it is measured. We have now also included an analysis of centriole length in RNAi-treated clones 1.1 and 1.9 showing that in both cases, mother centriole length is increased compared to the control (Figure 5b; see also point 3 of the Essential Revisions).

As suggested by the reviewer, we have measured the duration of mitosis in CRISPR clones to determine whether centriole elongation might be caused by extended mitosis. As shown in Supplemental Figure S1k of the revised manuscript, we found no difference in mitosis duration between clone 1.9 and WT cells (clones 1.1 and 1.2 show an increase of approximately 3 min on average compared to control cells). We conclude that centriole elongation is not driven by delayed mitosis in clone 1.9.

Reviewer #2 (Recommendations for the authors):

The data appear too preliminary for immediate publication in eLife. Nevertheless, the subject is intriguing and if authors could provide a cleaner evidence of LRRCC1 and C2CD3 localization, show how depletion of LRRCC1 affects centriole, appendage and cilia structure, the manuscript would be suitable for publication.

Point 1. Expansion microscopy is a powerful imaging approach. However, it is used here to determine localization of a relatively uncharacterized protein, using uncharacterized antibodies. This is risky. For instance: In figure 5 c-e, the authors judge the arrangement of Cep164 to classify centrosomes into 'normal, abnormal, and missing'. This characterization is made based on uneven distribution of Cep164 signals. However, if this criterion is to be extended to acetylated tubulin signal across all expansion images, many of control centrioles (Figure 1c, centriole2 and 3, Figure 3, centriole in b, 1.1 and 1.9 centrioles in Figure 6 f) must also be deemed 'aberrant', due to distortions, uneven spacings between MT blades etc. Clearly, expansion microscopy has its pitfalls.

There are also huge inconsistencies in the pattern and a great variability in the intensity of the LRRCC1 signals between figures and figure panels. For instance, in non-averaged images of expanded centrioles, LRRCC1 can be found in centriole's lumen, in association with MT wall, and in association with variable number of MT blades. In Figure 2b, and 1 b, LLRRCC1 signal is even associated with procentriole's proximal end. How much of the LLRRCC1 signal is indeed specific? The situation seems even more dramatic with C2CD3 data, where some of the signals lie entirely outside the centriole (for instance in 1.9 clone, Figure 6F). Can authors comment on how LRRCC1 and C2CD3 signals can protrude and be localized beyond centriole wall? Previous work (PMID: 30988386, PMID: 29789620) shows C2CD3 to be centered within centriole distal end, and not asymmetric.

It would be critical to use an additional super resolution microscopy technique to re-analyze the localization of LRRCC1 and C2CD3 in non-expanded samples and not to rely solely on expansion microscopy.

We agree with the reviewer that there are certain points to be careful about when analyzing U-ExM data. Although we systematically inspected our U-ExM samples to ensure that the geometry of the centrioles was preserved, clearly the expansion step can generate additional staining heterogeneity.

We addressed this problem by quantifying our analyses in two ways.

– In the case of distal appendage defects, we categorized centrioles into normal, abnormal and missing DAs, and indeed, we found 'abnormal' centrioles even in control cells. This proportion of abnormal centrioles may result from heterogeneity introduced during sample processing, variability in centriole structure and composition, or a combination of both. Nevertheless, by systematically analyzing multiple series of control and LRRCC1-depleted samples, we found in a highly reproducible manner that distal appendage defects are more frequent in the latter. We have now also added analyses using a second distal appendage marker, CEP83. In this case, it was more difficult to detect morphological variations, as the CEP83 staining is more compact than the CEP164 staining. Nevertheless, we observed a significant decrease in the proportion of centrioles with normal CEP83 labeling after RNAi treatment of CRISPR clones (now in Figure 5g, h). Together, our results support that LRRCC1-depletion affects the morphology of distal appendages.

– For LRRCC1 and C2CD3 labeling, we generated average images to synthesize the information present in the individual images. In our experience, the main issue with U-ExM is that is sometimes lead to suboptimal labeling, even in samples that expanded in an isometric fashion. In contrast, we have no evidence that this method leads to non-specific labeling, unlike SIM for which the reconstruction step can generate artifacts. Generating an average image from individual images is an approach that also has limitations, but which we believe is relevant to our analyses because it allows us to highlight the most salient aspects of LRRCC1 and C2CD3 localization patterns in WT cells. In LRRCC1-depleted cells, in which the variability of the C2CD3 staining is much greater, the average images allow us to visualize a divergence from the control pattern.

Regarding the aberrant patterns of C2CD3 such as the one mentioned by the reviewer, we would like to point out that these were not observed in control samples. Also, the individual centriole images shown in the figures have not been reoriented with respect to the observation axis. The images included in Figures 1c, 5d and g, 6d and f, are maximum intensity projections of individual z-sections encompassing the signal of interest in centrioles as they are in the sample. When the centrioles are tilted, the image projection generates an apparent shift between the two channels, which is due to the fact that the acetylated tubulin staining is fainter in the most distal part of the centriole where the different markers localize (see the lateral view of the average centriole in Figure 1f). This particular point is now addressed in the Material and Methods section (p 28, lines 638-642), as well as in the corresponding figure legends. To address this problem, we included in our image analysis tool a step allowing to reorient the centrioles before generating an average image. Unfortunately, this pipeline in its current state allows us to visually inspect individual centrioles after reorientation, but not to save them (only the average image can be saved). We hope in the future to further develop the potentialities of this tool.

This being said, the channels were not properly aligned in some of the images presented in the previous version of the manuscript. In some cases, it accentuated the shift between channels, including in the image the reviewer is referring to (now Figure 6g, bottom right panel). As explained in more detail in the response to the following comment (Point 2), this problem has now been addressed in the revised version of the manuscript.

Finally, we have now included data showing that LRRCC1 asymmetric localization can be detected in non-expanded samples using Airyscan microscopy (see response to point 3 of the Essential Revisions).

Point 2. It is not stated in the manuscript how and whether axial and lateral shifts between different channels were corrected? Was such correction done before image averaging or before image assembly?

Instead of showing averaged data, panels showing more individual examples of horizontally and vertically oriented centrioles would be more instructive. Additional supplemental figures could be assembled for this purpose, at least for the purpose of evaluation.

The alignment of the different channels was verified on series of individual images before generating average images. For this, we took advantage of the fact that the LRRCC1 and C2CD3 antibodies weakly label the centriole wall (see Figure 1b in the case of LRRCC1 labeling), which allows us to verify the overlap with acetylated tubulin labeling. We systematically checked the superposition of the channels in the first and last few images of each series. This was to verify that the U-ExM gel was not drifting, as sometimes happens, and that the microscope was correctly set up for acquisition. However, we have now checked each individual image and found that despite these precautions, the channels were incorrectly aligned in some of our C2CD3 images. No such misalignment was detected in LRRCC1 images. We therefore realigned the C2CD3 images using the weak centriole wall staining generated by the anti-C2CD3 antibody. Images in which such staining was absent or ambiguous were removed and replaced with new images.

In order to remain neutral to the hypothesis of an asymmetric vs. symmetric positioning of C2CD3 within the centriole, we now generated a new average using the shape of the staining pattern to superimpose images of individual centrioles. The resulting image shown in Figure 6e of the revised manuscript confirms the C-shape of C2CD3 labeling but does not show clear positional asymmetry within the centriole. This result is more consistent with previously published results (Yang et al., 2018; Tsai et al., 2019), as pointed above by the reviewer. Interestingly, a filament forming an ‘incomplete circle’ and lining the inner centriole wall was observed in the same region by electron microscopy (Vorobjev and Chentsov, 1980), which may correspond to the C2CD3-pattern we describe (see Discussion section p 17, line 374-376).

Individual images of C2CD3 in CRISPR clones were all reanalyzed and average images were generated in the same manner as for the WT (Figure 6f, g of the revised manuscript). We now also include images of CRISPR cells treated by RNAi to further reduce LRRCC1 levels. These results confirm that LRRCC1 depletion disrupts the C2CD3 localization pattern.

Point 3. Quantification of LRRCC1 levels in CRISPR clones (Figure 4 a) does not reflect centriole-associated levels of LRRCC1. It reflects the loss of pericentrosomal (satellite) population of LRRCC1, which is the most abundant. Quantification of the centriole bound LCCRR1 needs to be provided instead. Are centrosomal levels of LRRCC1 in 1.1, 1.2, and 1.9 CRISPR clones (Figure 4 a) decreased in comparison to the ones from nocodazole -treated cells (Figure 1A)? In addition, higher resolution images of LRRCC1 in CRISPR clones have not been provided to illustrate where the leftover protein is localized in CRISPR 1.1, 1.2, and 1.9 clones.

How can authors exclude the possibility that loss of the pericentrosomal population of LRRCC1 affects ciliation and ciliary signaling?

This issue is addressed in the response to point 3 of the Essential Revisions.

Point 4. Between 8 independent experiments conducted in wt RPE-1 cells, the percentage of centrioles with 'abnormal or missing" arrangement of Cep164 varies from ~0% – 40% (Figure 5C). This is a large variability since almost all wt RPE-1 cells can form a cilium (Figure 4b). In my view, this means either that the variability in Cep164 signal is introduced experimentally, or that it is physiological and cannot be used as a measure for ciliation. Indeed, previous super-resolution analysis has demonstrated that, although distal appendage proteins display characteristic pattern around centrioles, their levels on individual appendages can vary under physiological conditions (PMID: 30824690, PMID: 29789620). Therefore, significance of the data from Figure 5 c-e is unclear. Besides, additional appendage proteins (such as Cep83 and SCLT1, for instance) should be analyzed to reach a conclusion about distal appendage organization and number.

This issue is addressed in the response to the reviewer’s point 1.

Point 5. The authors propose that LRRCC1 cooperates with C2CD3 in organization of distal centriole ends. However, partial reduction in LRRCC1 leads to elongated centrioles only in 1.9 CRISPR clone and not in other two clones, but it is unclear why, as differences between the centrosomal levels of LRRCC1 in CRISPR clones do not seem large enough to explain this difference. Increased centriole length in clone 1.9 could also be non-specific due to clonal variation, and it is unclear whether it is relevant. Especially, because both clones 1.1 and 1.9 seem to show similar defects in ciliation and Cep164 assembly.

To prove and understand what type of centriolar structural defects occur after LRRCC1 depletion, extensive ultrastructural analysis would need to be conducted. Perhaps LRRCC1 could be further depleted by siRNA in LRRCC1- CRISPR clones to generate more penetrant phenotypes?

This issue is addressed in the response to point 3 of the Essential Revisions.

Point 6. Figure 7. More evidence of LRRCC1/C2CD3 colocalization needs to be provided. Why is majority of C2CD3 signal colocalized with centriole MTs?

We agree with the reviewer that the co-localization data are not entirely satisfactory, but we have tried extensively to get better images using various combinations of antibodies, so far without success. The main difficulty is that far red dyes are not stable under U-ExM conditions – i.e. in pure water, which is especially a problem for low intensity signals such as those generated by LRRCC1 or C2CD3 antibodies. This is really an issue we expect to solve in the future. However, although we cannot precisely map the respective locations of LRRCC1 and C2CD3, our images already show that C2CD3 and LRRCC1 localize to the same region of the centriole.

Reviewer #3 (Recommendations for the authors):

Critical to the central points of the paper are the asymmetric localization of LRRCC1 and C2CD3 at the distal centriole. It is not clear whether the expansion in expansion microscopy is always isotropic and anisotropic expansion could conceivably skew protein localization to appear asymmetric. Could an orthogonal superresolution method, such as 3D-SIM/STED/STORM be used to confirm asymmetry of LRRCC1 or C2CD3? Additionally, could performing 3D-reconstruction image analysis of LRRCC1/C2CD3 with a distal appendage component or another symmetric sub-component of the distal centriole is critical rule out potential artifacts that could be due to anisotropic expansion during U-ExM? The authors imaged OFD1 or CEP290 using U-ExM (fluorescent intensity quantification data are presented in Supplementary Figure S3, but no images are shown). Further confidence in the LRRCC1 conclusions would be garnered by data indicating that these proteins are symmetrically localized at the distal centriole.

As detailed in the response to point 2 of the Essential Revisions, we have now confirmed that LRRCC1 is asymmetrically localized by a method that does not involve expansion. With respect to C2CD3, as explained in more details in the response to reviewer #2 (point 2), we found that in a subset of C2CD3 images, the two channels were not properly aligned. After systematically correcting this anomaly and generating a new average image based on the shape of the staining pattern, we confirm the C-shape of the C2CD3 pattern but not its asymmetric position in the lumen (Figure 6e and Results section p14, lines 311-316).

Regarding the quantification of OFD1 and CEP290 proteins, it was performed on non-expanded immunofluorescence samples. However, we analyzed CEP164 and now CEP83 localization, and found that these were symmetrically distributed around the centrioles in the majority of WT cells. We made preliminary observations of other markers, including OFD1, CEP290, ODF2 and Talpid3 (see for example the image below, noting that the centriole is tilted, hence the apparent shift between the two channels). We did not observe any obvious asymmetry in these patterns, in sharp contrast to what we observed for LRRCC1. We should also mention that we use the same protocol as in the studies by Le Guennec et al. (2020; doi:10.1126/sciadv.aaz4137), Steib et al. (2020; doi: 10.7554/eLife.57205) or Le Borgne et al. (2021, doi.org/10.1101/2021.07.13.452210), which also show symmetric labeling for different markers localized in the centriolar lumen or at distal appendages.

The authors report that their results "demonstrate that rotational asymmetry is a conserved ancient property of centrioles." However, the basal foot is a previously described rotational asymmetry present in animal cells, no? Indeed, the basal foot seems to be anticorrelated with the localization of LRRCC1. The authors examine distal appendage formation in some depth. Is basal foot formation or location compromised in LRRCC1 depleted cells?

Regarding the first point, we meant to say that centriole rotational asymmetry is conserved beyond centrioles that carry asymmetric appendages like in multicellular cells. We have modified this sentence as follows: “Taken together, our results demonstrate that rotational asymmetry is an ancient property of centrioles that is broadly conserved in human cells” (p2, lines 41-43).

Regarding the role of LRRCC1 on basal foot assembly or position, we do not have the expertise for culture and targeted gene inactivation in mammalian multiciliated cells, unfortunately. However, we have shown that in planarian multiciliated cells, inactivation of the LRRCC1 ortholog (SMED-VFL1) indeed leads to defects in basal foot assembly and position (Basquin et al., 2019). Because of this observation, we also tried to determine whether the subdistal appendages of the mother centriole could be impacted at the centrosome of human cells. Unfortunately, subdistal appendages are not properly labelled in WT cells processed for U-ExM with the markers we have tested so far (CEP170, centriolin, ninein). But it is indeed an important point that we want to continue exploring in the future.

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Quantitation of depletion of LRRCC1 is done by light microscopy. The remaining population, although quantitated, is not characterized spatially with any high degree of resolution. Should one subcellular population (centriolar versus satellite) be preferentially depleted by the reduction in LRRCC1 levels, one population might be implicated in the biological functions of LRRCC1. Where is the residual LRRCC1 staining in the CRISPR/Cas9 generated clones? For example, figure 4a would benefit from U-ExM and quantification of fluorescence intensity specifically at centrioles to determine if the centriolar pool of LRRCC1 is disrupted in the hypomorphs, as surmised by the authors.

This is addressed in the response to point 3 of the Essential Revisions.

It is curious that the authors have been unable to generate null mutations in LRRCC1. RPE1 cells fail to proliferate in the presence of centrinone, possibly hinting at a critical role for LRRCC1 in centriole duplication. Is under duplication or delayed duplication of centrioles observed in LRRCC1 depleted cells?

Relatedly, were null mutations in LRRCC1 generated in HEK 293 cells?

We have now included data showing that centriole duplication is not affected in LRRCC1-depleted cells (Supplemental Figure S1j; see also response to point 2 of the Essential Revisions).

With respect to the lack of LRRCC1 null clones, we also initially thought that this might be related to the fact that RPE1 cells arrest or exhibit slower growth in a p53-dependent manner after a range of centrosome perturbations. However, we also failed to isolate null clones for HEK 293, U2-OS, or p53-/- RPE1 cells (generously donated by Brian Tsou). Thus, we have no explanation at this stage for the fact that null clones have not been isolated, while we have managed to isolate such clones for other genes without difficulty (e.g. CCDC61 and C-Nap1; doi: 10.1111/boc.201900038).

In the discussion, the authors speculate at some length about a role of LRRCC1 in the formation of the ciliary gate, given that LRRCC1 depletion inhibits the ciliary localization of Smoothened. However, one component of the ciliary gate, CEP290, is not reported to be altered in LRRCC1 knockdown. Can the authors examine the dependence of ciliary gate components required for Smoothened localization to cilia in the LRRCC1 knockdown cells?

We agree with the reviewer that this is an important point, but so far we have not been successful with the different antibodies against ciliary gate proteins we tested. We expect to find the right markers and conditions for future studies.

I recognize that the rescue experiments were difficult. Could an inducible approach (e.g., TetOne system) be attempted? Rescue may help delineate which of the variably penetrant phenotypes are due to downregulation of LRRCC1 and which to other factors. For example, the authors observe a mild increase in centriole length in clone 1.9, but not in clone 1.1. The authors conclude that the slightly lower levels of LRRCC1 in clone 1.9 account for the difference. However, an alternative possibility is that clone 1.9 has another genetic or epigenetic variation that accounts for this incompletely penetrant phenotype.

We have tried different approaches for performing rescue experiments, including attempts to generate stable lines expressing inducible constructs, without success. One difficulty we have faced is that the presence of larger tags such as GFP disrupts LRRCC1 localization. Fusions at the N- or C-terminus do not localize at all to the centrosome or the satellites, and fusions with a GFP inserted within the protein sequence (after aa. 251 or 402) localize to the centrosome area and satellites, but we did not detect them at centriole distal ends. We therefore attempted to establish stable lines using a myc-tagged construct since the myc tag is much smaller. Screening non-fluorescent cells is extremely tedious when the proportion of stably transformed cells is low however, and we abandoned this approach after several unsuccessful attempts.

Following the reviewer's recommendation, we have now included data showing the effect of further reducing LRRCC1 levels by RNAi. As also detailed in the response to point 3 of the Essential Revisions, under these conditions we observed a significant difference between clone 1.1 and control cells with respect to centriole length.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

The manuscript has been much improved and you have addressed all concerns. There are a couple of items that I would like you to address before formal acceptance:

1) Figure 1d: The Airyscan localization experiment uses POC5 labelling as an example of a symmetrical localization. I suggest that you show the channels also separately so that one can appreciate the luminal POC5 signal better. In the legend you commented on the fact that the POC5 label is also on the outside – in fact the outside label seems stronger than the luminal signal. This is a bit confusing for the purpose of this experiment, but also considering previous POC5 localization studies by ExM (e.g. Le Guennec et al., Sci Adv, 2020; Schweizer et al., Nat Comms, 2021). Is this outside signal specific? It may be good to discuss/explain this a bit more.

We have now included images showing the individual channels in Figure 1d. We also added a comment in the corresponding legend to specify that the anti-hPOC5 antibody labels a region of the mother centriole that broadly corresponds to the region of distal/subdistal appendages (highlighted in green).

Initially, we considered using OFD1, which localizes distally and near the triplets, as a marker. However, OFD1 labeling is wider and tends to separate into 2 peaks when observed by Airyscan microscopy, which made comparison with LRRCC1 difficult. Similar results were obtained using antibodies against ODF2 and Talpid-3. We therefore opted for hPOC5, which localizes symmetrically in the centriole lumen (Le Guennec et al; 2020; and Schweizer et al., 2021). The antibody we used was described in the initial characterization of the hPOC5 protein, the specificity of which was established via RNAi experiments (Azimzadeh et al., 2009; doi: 10.1083/jcb.200808082). We noted at the time that the immunofluorescence labeling was stronger at the mother centriole than at the daughter centriole, which we interpreted as resulting from a progressive recruitment of hPOC5 into the centriole lumen. The pattern observed by Airyscan microscopy rather suggests that hPOC5 localizes near mother centriole appendages in addition to being present in the centriole lumen. In U-ExM, our antibody only labels the centriole wall however, very similar to what was described by Le Guennec et al. and Schweizer et al. As mentioned in the response to Reviewer #3, none of the antibodies against subdistal appendage components that we tested worked, suggesting that these appendages may not be properly preserved by the U-ExM protocol. If hPOC5 localizes near subdistal appendages, this could explain why this staining is not visible by U-ExM.

In any case, by Airyscan microscopy, hPOC5 localizes as expected in the middle region of centrioles, which is the region in which we performed our measurements. In this region, hPOC5 is clearly localized more symmetrically than LRRCC1, confirming our finding that LRRCC1 localizes asymmetrically in the centriole lumen.

2) It is somewhat surprising that you can reliably count individual centrioles using POC5 staining, which is proposed to mark the central lumen. Considering the unexpected distribution of the POC5 label in Fig, 1d, perhaps the more distal, outside label helps? In any case, it would be good to include example cells with POC5 staining of centriole pairs at spindle poles with the quantifications in Figure S1j.

In the previous version of the manuscript, we forgot to indicate that centrioles were labeled for centrin in addition to hPOC5. An image has now been included in Figure 1—figure supplement 1j, and the figure legend has been modified accordingly (as well as the antibody list in Table 1; highlighted in green). As also shown in our previous study (Azimzadeh et al., 2009), hPOC5 labeling allows to visualize individual centrioles in mitotic cells in a manner similar than centrin.

https://doi.org/10.7554/eLife.72382.sa2

Article and author information

Author details

  1. Noémie Gaudin

    Université Paris Cité, CNRS, Institut Jacques Monod, Paris, France
    Contribution
    Conceptualization, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Resources, Supervision, Validation, Visualization, Writing – original draft
    Competing interests
    No competing interests declared

  2. Paula Martin Gil

    Université Paris Cité, CNRS, Institut Jacques Monod, Paris, France
    Contribution
    Data curation, Formal analysis, Investigation, Supervision, Validation, Visualization, Writing – original draft
    Competing interests
    No competing interests declared

  3. Meriem Boumendjel

    Université Paris Cité, CNRS, Institut Jacques Monod, Paris, France
    Contribution
    Formal analysis, Investigation, Visualization
    Competing interests
    No competing interests declared

  4. Dmitry Ershov

    1. Image Analysis Hub, C2RT, Institut Pasteur, Paris, France
    2. Hub de Bioinformatique et Biostatistique – Département Biologie Computationnelle, Institut Pasteur, Paris, France
    Contribution
    Methodology, Resources, Software, Visualization
    Competing interests
    No competing interests declared

  5. Catherine Pioche-Durieu

    Université Paris Cité, CNRS, Institut Jacques Monod, Paris, France
    Contribution
    Investigation, Visualization
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-0988-1169

  6. Manon Bouix

    Université Paris Cité, CNRS, Institut Jacques Monod, Paris, France
    Contribution
    Formal analysis, Investigation, Visualization
    Competing interests
    No competing interests declared

  7. Quentin Delobelle

    Université Paris Cité, CNRS, Institut Jacques Monod, Paris, France
    Contribution
    Formal analysis, Investigation, Visualization
    Competing interests
    No competing interests declared

  8. Lucia Maniscalco

    Université Paris Cité, CNRS, Institut Jacques Monod, Paris, France
    Contribution
    Formal analysis, Investigation, Visualization
    Competing interests
    No competing interests declared

  9. Than Bich Ngan Phan

    Université Paris Cité, CNRS, Institut Jacques Monod, Paris, France
    Contribution
    Formal analysis, Investigation, Validation
    Competing interests
    No competing interests declared

  10. Vincent Heyer

    1. Institut de Génétique et de Biologie Moléculaire et Cellulaire (IGBMC), Ilkirch, France
    2. Institut National de la Santé et de la Recherche Médicale (INSERM), Illkirch, France
    3. Centre National de la Recherche Scientifique (CNRS), Illkirch, France
    4. Université de Strasbourg, Illkirch, France
    Contribution
    Resources
    Competing interests
    No competing interests declared

  11. Bernardo Reina-San-Martin

    1. Institut de Génétique et de Biologie Moléculaire et Cellulaire (IGBMC), Ilkirch, France
    2. Institut National de la Santé et de la Recherche Médicale (INSERM), Illkirch, France
    3. Centre National de la Recherche Scientifique (CNRS), Illkirch, France
    4. Université de Strasbourg, Illkirch, France
    Contribution
    Resources
    Competing interests
    No competing interests declared

  12. Juliette Azimzadeh

    Université Paris Cité, CNRS, Institut Jacques Monod, Paris, France
    Contribution
    Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Project administration, Supervision, Validation, Writing – original draft
    For correspondence
    juliette.azimzadeh@cnrs.fr
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-7292-9973

Funding

Agence Nationale de la Recherche (ANR-21-CE13-008)

  • Juliette Azimzadeh

Fondation pour la Recherche Médicale (Graduate Student Fellowship)

  • Noémie Gaudin

Fondation ARC pour la Recherche sur le Cancer (ARCPJA32020060002055)

  • Juliette Azimzadeh

Ligue Contre le Cancer (RS16/75-105)

  • Juliette Azimzadeh

Labex Who Am I? (Thematic Program)

  • Juliette Azimzadeh

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We are deeply grateful to Marine Laporte, Virginie Hamel, Paul Guichard, and Davide Gambarotto for teaching us the U-ExM procedure and for sharing antibodies; Arnaud Echard and Takashi Ochi for critical reading of the manuscript; Amélie-Rose Boudjema and Alice Meunier for providing mouse ependymal cells and Isabelle Le Parco for the tracheal tissue; Juliane Da Graça and Simon Herman for technical help; Rémi Le Borgne for help with transmission electron microscopy and for critical reading of the manuscript. We acknowledge the core imaging facility of Institut Jacques Monod (ImagoSeine facility, member of the France BioImaging infrastructure supported by grant ANR-10-INBS-04 from the French National Research Agency). This work was supported by funding from La Ligue Contre le Cancer, Fondation ARC pour la recherche sur le cancer, Labex Who Am I? (supported by grants ANR-11-LABX-0071 and ANR-11-IDEX-0005-02) and ANR-21-CE13-008 to JA. NG was a recipient of a MESRI PhD fellowship from the French government and a 4th year PhD fellowship from the Fondation pour la Recherche Médicale.

Senior Editor

  1. Anna Akhmanova, Utrecht University, Netherlands

Reviewing Editor

  1. Jens Lüders, Institute for Research in Biomedicine, Spain

Publication history

  1. Received: July 21, 2021
  2. Preprint posted: July 22, 2021 (view preprint)
  3. Accepted: March 22, 2022
  4. Accepted Manuscript published: March 23, 2022 (version 1)
  5. Version of Record published: April 5, 2022 (version 2)

Copyright

© 2022, Gaudin et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Noémie Gaudin
  2. Paula Martin Gil
  3. Meriem Boumendjel
  4. Dmitry Ershov
  5. Catherine Pioche-Durieu
  6. Manon Bouix
  7. Quentin Delobelle
  8. Lucia Maniscalco
  9. Than Bich Ngan Phan
  10. Vincent Heyer
  11. Bernardo Reina-San-Martin
  12. Juliette Azimzadeh
(2022)
Evolutionary conservation of centriole rotational asymmetry in the human centrosome
eLife 11:e72382.
https://doi.org/10.7554/eLife.72382

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    2. Stem Cells and Regenerative Medicine

    Atf3 defines a population of pulmonary endothelial cells essential for lung regeneration

    Terren K Niethamer, Lillian I Levin ... Edward E Morrisey
    Research Article

    Following acute injury, the capillary vascular bed in the lung must be repaired to reestablish gas exchange with the external environment. Little is known about the transcriptional and signaling factors that drive pulmonary endothelial cell (EC) proliferation and subsequent regeneration of pulmonary capillaries, as well as their response to stress. Here, we show that the transcription factor Atf3 is essential for the regenerative response of the mouse pulmonary endothelium after influenza infection. Atf3 expression defines a subpopulation of capillary ECs enriched in genes involved in endothelial development, differentiation, and migration. During lung alveolar regeneration, this EC population expands and increases the expression of genes involved in angiogenesis, blood vessel development, and cellular response to stress. Importantly, endothelial cell-specific loss of Atf3 results in defective alveolar regeneration, in part through increased apoptosis and decreased proliferation in the endothelium. This leads to the general loss of alveolar endothelium and persistent morphological changes to the alveolar niche, including an emphysema-like phenotype with enlarged alveolar airspaces lined with regions that lack vascular investment. Taken together, these data implicate Atf3 as an essential component of the vascular response to acute lung injury that is required for successful lung alveolar regeneration.

    1. Cell Biology
    2. Microbiology and Infectious Disease

    Profiling the bloodstream form and procyclic form Trypanosoma brucei cell cycle using single-cell transcriptomics

    Emma M Briggs, Catarina A Marques ... Keith R Matthews
    Research Article Updated

    African trypanosomes proliferate as bloodstream forms (BSFs) and procyclic forms in the mammal and tsetse fly midgut, respectively. This allows them to colonise the host environment upon infection and ensure life cycle progression. Yet, understanding of the mechanisms that regulate and drive the cell replication cycle of these forms is limited. Using single-cell transcriptomics on unsynchronised cell populations, we have obtained high resolution cell cycle regulated (CCR) transcriptomes of both procyclic and slender BSF Trypanosoma brucei without prior cell sorting or synchronisation. Additionally, we describe an efficient freeze–thawing protocol that allows single-cell transcriptomic analysis of cryopreserved T. brucei. Computational reconstruction of the cell cycle using periodic pseudotime inference allowed the dynamic expression patterns of cycling genes to be profiled for both life cycle forms. Comparative analyses identify a core cycling transcriptome highly conserved between forms, as well as several genes where transcript levels dynamics are form specific. Comparing transcript expression patterns with protein abundance revealed that the majority of genes with periodic cycling transcript and protein levels exhibit a relative delay between peak transcript and protein expression. This work reveals novel detail of the CCR transcriptomes of both forms, which are available for further interrogation via an interactive webtool.