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Centriolar remodeling underlies basal body maturation during ciliogenesis in Caenorhabditis elegans

  1. Inna V Nechipurenko  Is a corresponding author
  2. Cristina Berciu
  3. Piali Sengupta  Is a corresponding author
  4. Daniela Nicastro  Is a corresponding author
  1. Brandeis University, United States
  2. University of Texas Southwestern Medical Center, United States
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Cite this article as: eLife 2017;6:e25686 doi: 10.7554/eLife.25686

Abstract

The primary cilium is nucleated by the mother centriole-derived basal body (BB) via as yet poorly characterized mechanisms. BBs have been reported to degenerate following ciliogenesis in the C. elegans embryo, although neither BB architecture nor early ciliogenesis steps have been described in this organism. In a previous study (Doroquez et al., 2014), we described the three-dimensional morphologies of sensory neuron cilia in adult C. elegans hermaphrodites at high resolution. Here, we use serial section electron microscopy and tomography of staged C. elegans embryos to demonstrate that BBs remodel to support ciliogenesis in a subset of sensory neurons. We show that centriolar singlet microtubules are converted into BB doublets which subsequently grow asynchronously to template the ciliary axoneme, visualize degeneration of the centriole core, and define the developmental stage at which the transition zone is established. Our work provides a framework for future investigations into the mechanisms underlying BB remodeling.

https://doi.org/10.7554/eLife.25686.001

Introduction

Cilia are evolutionarily conserved microtubule (MT)-based organelles that play key roles in regulating embryonic development, sensory signaling, and motility among other cellular functions (Goetz and Anderson, 2010; Green and Mykytyn, 2010; Yildiz and Khanna, 2012; Falk et al., 2015). Both immotile primary and motile cilia are nucleated by a basal body (BB) that is generally derived from the mother centriole (Marshall, 2007; Kim and Dynlacht, 2013). BBs possess accessory structures such as transition fibers that associate with a ciliary vesicle or dock with the plasma membrane and provide a platform for assembly of intraflagellar transport (IFT) complexes that are essential for elongation of the ciliary axoneme (Kim and Dynlacht, 2013; Azimzadeh and Marshall, 2010; Reiter et al., 2012; Dawe et al., 2007). Although overall organization of centrioles/BBs, as well as many proteins required for their assembly and function are conserved, ultrastructural features of these cellular structures can differ among and within species. For instance, centrioles/BBs are cylindrical structures that can be comprised of a radially symmetric array of MT singlets, doublets, or triplets depending on the species and cellular context (Azimzadeh and Marshall, 2010; Winey and O'Toole, 2014; Carvalho-Santos et al., 2011; Gottardo et al., 2015; González et al., 1998; Jana et al., 2016). It remains unclear whether centrioles of distinct ultrastructural organization transition to BBs and nucleate cilia via similar or distinct mechanisms.

Ultrastructural analyses of centrioles in one-cell C. elegans embryos have shown that centrioles in this organism are structurally distinct from their mammalian counterparts (Pelletier et al., 2006). C. elegans centrioles are relatively small compared to those in mammals and are comprised of a central tube surrounded by nine singlet MTs (sMTs), as compared to the cartwheel structure surrounded by triplet MTs found in larger mammalian centrioles (Winey and O'Toole, 2014; Pelletier et al., 2006; Hilbert et al., 2013; Gönczy, 2012). Despite these differences, C. elegans and vertebrate centrioles are built using subsets of conserved proteins (Carvalho-Santos et al., 2011; Gönczy, 2012).

Primary cilia are present only on sensory neurons in C. elegans (Ward et al., 1975; Perkins et al., 1986). As in other organisms, these cilia are templated by BBs derived from centrioles (Perkins et al., 1986). However, BBs in C. elegans have been reported to degenerate following cilia assembly in the embryo, and no canonical BB structures or core centriolar components are detected in ciliated neurons in wild-type animals at postembryonic stages (Perkins et al., 1986; Dammermann et al., 2009; Schouteden et al., 2015; Doroquez et al., 2014). Intriguingly, despite this apparent degeneration, a subset of BB-associated proteins remains enriched at the cilia base in adult animals as shown via immunofluorescence (Dammermann et al., 2009; Mohan et al., 2013; Wei et al., 2013, 2016), suggesting the presence of centriolar/BB ‘remnants’. Since sensory neurons are born and differentiate at late embryonic stages (Sulston et al., 1983) that are technically challenging to analyze experimentally, key early steps in ciliogenesis including the centriole-to-BB transition, the precise timing of centriolar degeneration, and initiation of axoneme elongation, have yet to be examined in this organism.

In a recent report, we described the three-dimensional morphologies of sensory cilia in the nose of C. elegans hermaphrodites at high resolution using serial section transmission electron microscopy (ssTEM) and serial section electron tomography (ssET) of high pressure-frozen and freeze-substituted (HPF-FS) adult animals (Doroquez et al., 2014). Here, we use these imaging methods to describe early steps of ciliogenesis in the C. elegans embryo. We find that sMTs of centrioles in early embryos contain hook-like appendages that remodel to dMTs during BB maturation and prior to axoneme elongation, and template the dMTs of the ciliary axoneme. We show that these BB dMTs at the cilia base ‘flare’ at later embryonic stages, and that this flaring coincides with degeneration of the central tube of the centriole/BB. We also visualize formation of the transition zone (TZ), a compartment that acts as a diffusion barrier at the ciliary base, and the apical ring, a structure present at the distal TZ (Perkins et al., 1986; Doroquez et al., 2014; Blacque and Sanders, 2014). Our observations indicate that the centriole/BB does not fully degenerate, but that the outer centriole wall remodels to nucleate the axoneme and persists through adulthood in a subset of C. elegans sensory neurons. This work reports key early steps in BB maturation and ciliogenesis and extends our previous ultrastructural analyses of adult sensory cilia in this organism.

Results

Hook-like appendages of A-tubules close to form the B-tubules of the BB and axoneme in a subset of C. elegans ciliated sensory neurons

Twelve pairs of ciliated sensory neurons are found in the bilateral amphid sensory organs of the head in the C. elegans hermaphrodite (Ward et al., 1975; Perkins et al., 1986). Eight of these neurons extend their simple rod-like cilia through a channel created by glial cells (Ward et al., 1975; Perkins et al., 1986; Doroquez et al., 2014) (Figure 1—figure supplement 1). Since these channels, and neuronal endings contained therein, are readily identifiable in serial sections of the embryo, we focused our attention on this subset of ciliated cells. Amphid sensory neurons are born over a period of time from the end of ventral closure to the comma stage of embryogenesis (Sulston et al., 1983) (Figure 1A), and cilia of 3 μm or longer have been previously detected in these neurons starting at the three-fold stage using fluorescent reporters (Fujiwara et al., 1999). However, core centriole proteins such as SAS-4 are not detected at these later developmental times (Dammermann et al., 2009; Schouteden et al., 2015; Kirkham et al., 2003) (Figure 1A), suggesting that the centriole has at least partly degenerated, and that cilia have elongated by these late embryonic stages.

Figure 1 with 2 supplements see all
The centrioles of embryonic amphid sensory neurons remodel to initiate ciliogenesis.

(A) Timeline of embryonic development at 22°C (adapted from IntroFIG7 http://www.wormatlas.org/ver1/handbook/anatomyintro/anatomyintro.htm) showing select stages defined by morphology between fertilization (t = 0) and hatching (L1). The approximate time period during which amphid neurons are born is marked. The developmental stages during which core centriole components (e.g. SAS-4 and BB-associated protein HYLS-1 [Dammermann et al., 2009; Wei et al., 2016]) are detected by immunofluorescence (IF) in amphid neurons are marked. Dashed line indicates that the exact time of SAS-4 loss in amphid neurons is unknown. mpf – minutes post fertilization. (B–D) TEM images of the amphid channel in cross-section (Ci and Di), cross-section ET slices and schematics of a centriole in an unidentified cell of a 350 mpf embryo (Bi) and BBs in amphid neurons (Cii, Ciii, Dii, Diii), and ET slices showing centrioles/BBs in longitudinal orientation (Bii, Civ, Div) at the indicated stages of embryogenesis. Two different examples of bean-stage centrioles (#s 1 and 2) undergoing remodeling are shown in Cii and Ciii, respectively. Arrowheads: centrioles/BBs (white), dMTs (yellow), sMTs with hook appendages (red), central tube (light blue), putative nascent Y-links (purple). Scale bars: 100 nm.

https://doi.org/10.7554/eLife.25686.002

To visualize centrioles and cilia in the embryo, we examined wild-type C. elegans embryos at multiple developmental stages using ssTEM and ssET. Although a subset of amphid neurons has already been born by 350 min post-fertilization (mpf) (Figure 1A), we were unable to detect amphid channels formed by glial cells in embryos of this stage (Figure 1—figure supplement 2A), and thus, could not unambiguously distinguish amphid sensory neurons from other cell types. Nevertheless, consistent with previous reports (Pelletier et al., 2006; Delattre et al., 2004; Mikeladze-Dvali et al., 2012), we observed centrioles comprised of the central tube surrounded by sMTs with hook-like appendages in many cells (Figure 1B). The average diameter and length of centrioles at this developmental stage were 88.2 ± 4.2 nm and 100.7 ± 0.2 nm, respectively (Figure 3A).

We next examined serial sections of embryos frozen at the ‘bean’ stage (Figure 1A). In contrast to our observations at 350 mpf, bilateral amphid channels containing sensory neuron endings were readily visible at this and all subsequent stages (Figure 1—figure supplement 2B, Figure 2—figure supplement 1). Centrioles in this stage were not located in close proximity to the cell surface and dendritic tip but were instead found deep within the cell (Figure 1Ci and iv). Interestingly, at this stage, we observed a transition from sMTs with hooks to dMTs. Specifically, centrioles in several amphid neurons were comprised of the central tube surrounded by a mixture of dMTs and sMTs with hooks (Figure 1Cii–iii). The average length of these structures at the bean stage was similar to that at the 350 mpf stage (Figure 3B). We, therefore, infer that sMTs remodel to form dMTs by closure of the A tubule-associated hooks to generate B-tubules. As these centrioles contain both dMTs and sMTs with hooks within the same 70 nm section, the transition from sMTs to dMTs likely occurs asynchronously within a centriole. We henceforth refer to this remodeled structure as the BB, and conclude that this remodeling is initiated by the bean stage of embryonic development in a subset of amphid sensory neurons.

By the ‘comma’ stage (Figure 1A), BBs in a subset of amphid neurons were found in close proximity to the cell surface (Figure 1Di and iv). Although the average length of these structures was mildly increased, their mean diameter and length were not significantly different relative to those at the bean stage (Figure 3A–B). At the comma stage, dMTs surrounded the central tube in the most proximal BB regions; however, a mixture of dMTs and sMTs with hooks was present in more distal BB/axoneme regions in most examined cells (Figure 1Dii–iii). As BBs/axonemes elongate in later stage embryos (Figure 2A–C, Figure 3B), only dMTs were detected in proximal regions, whereas a mixture of dMTs and sMTs with hooks were present in more distal regions of axonemes in 1.5- and two-fold embryos (Figure 2A–C). These observations suggest that A- and B-tubules of BBs/axonemes grow asynchronously in examined amphid sensory neurons (summarized in Figure 3C).

Figure 2 with 3 supplements see all
Degeneration of the central tube and dMT flaring at the ciliary base are observed by the 1.5-fold stage of embryogenesis.

(A–C) TEM images (Ai and Bi) and ET slice (Ci) of the amphid channel in cross-section, cross-section ssET images and schematics of BBs in amphid neurons (Aiiv, Biiv, Ciiv), and ET slices (Avi, Avii, Cvi) and TEM image (Bvi) showing BBs/axonemes in longitudinal orientation at the indicated stages of embryogenesis. Ciiv show a subset of ET slices from the serial section tomogram. Examples of BBs/axonemes in 1.5-fold embryo with a largely degenerated central tube and an intact central tube are shown in Aiv and Av, respectively. Arrowheads: centrioles/BBs (large white), flared dMTs at cilia base (small white double), dMTs (yellow), sMTs with hook appendages (red), central tube (light blue), Y-links (purple), apical ring (pink), isMTs (orange), a vesicle (green). Each bracket delineates a single BB/axoneme with its proximal and distal regions marked accordingly. Scale bars: 100 nm.

https://doi.org/10.7554/eLife.25686.005
Quantification of BB/axoneme diameter and length in embryonic amphid sensory neurons.

(A) Quantification of the centriole/BB/axoneme diameter measured as the distance between centers of A-tubules. The first tomographic slice of a ssET sequence showing the entire proximal region of each BB/axoneme in cross-section was used for measurements. Each dot represents a measurement from an individual BB/axoneme in different neurons from the same embryo. Horizontal bars indicate mean. Errors are SD. ** indicate that marked data sets are different at p<0.01 (Kruskal-Wallis test with post-hoc correction for multiple comparisons). (B) Quantification of the centriole/BB/axoneme length at the indicated stages of embryonic development. Each dot represents a measurement from an individual BB/axoneme in different neurons from the same embryo. Horizontal bars indicate mean. Errors are SD. *, **, and *** indicate that marked data sets are different at p<0.05, 0.01, and 0.001, respectively (Kruskal-Wallis test with post-hoc correction for multiple comparisons). (C) Model summarizing key early ciliogenesis stages in the examined subset of C. elegans sensory neurons. BB – basal body; TZ – transition zone; AX – axoneme. Blue and pink circles indicate the central tube and apical ring, respectively. Model is based solely on our ability to visualize specific ciliary structures, and no assumptions are made regarding the presence or absence of proteins associated with these structures.

https://doi.org/10.7554/eLife.25686.009

Degeneration of the centriole core coincides with increased ciliary base diameter

We previously reported flaring of dMTs at the ciliary base in adult amphid neurons, and proposed that this flaring is a consequence of BB degeneration (Doroquez et al., 2014). We found that in embryos, the diameter of the proximal BB/axoneme region was variable at the 1.5-fold stage and significantly increased by the two-fold stage (Figure 3A). Previous observations have reported loss of core centriolar/BB markers such as SAS-4 by the two-fold stage by immunofluorescence (Dammermann et al., 2009; Schouteden et al., 2015), raising the possibility that the increased BB diameter is a consequence of degeneration of core centriolar structures.

All examined cross-sections of centrioles/BBs in 350 mpf embryos as well as bean and comma-stage amphid neurons contained the central tube with an average diameter of 60.9 ± 4.2 nm (Figure 1B–D). However, the central tube appeared to be present in only a subset of examined BBs in 1.5-fold embryos (compare Figure 2Aiv with Figure 2Av, Figure 2—figure supplement 2) and was absent from all examined BBs in two- and three-fold embryos (Figure 2B–C). The diameter of the proximal regions of BBs with seemingly degenerated central tubes was larger than that of BBs with intact central tubes (compare Figure 2Av with Figure 2Aiv, Figure 2Bv and Cv). Consistent with a significant increase in proximal BB/axoneme diameter evident from cross-sections, we observed flaring of dMTs at the ciliary base in longitudinal sections in a subset of amphid neurons at the 1.5-fold stage (Figure 2Ai and Avii), and in all examined amphid neurons at the two- and three-fold embryonic stages (Figure 2Bi, Bvi, 3Ci and Cvi). These results suggest that central tubes degenerate asynchronously in individual amphid neurons, and that degeneration of the central tube starting at the 1.5-fold stage likely accounts for the flaring of BB dMT arrays and increased BB diameters (Figure 3A and C). As we reported previously (Doroquez et al., 2014), we did not detect any obvious structures resembling transition fibers associated with BBs in C. elegans amphid neurons at any developmental stage by TEM of HPF-FS samples.

The transition zone is formed by the 1.5-fold stage of embryogenesis in a subset of sensory neurons

The TZ at the cilia base is the proximal-most compartment of the axoneme proper. This compartment is defined ultrastructurally by the presence of proteinacious Y-links that originate at the outer junction between A- and B-tubules of axonemal dMTs, and project toward and usually connect to the plasma membrane (Reiter et al., 2012; Blacque and Sanders, 2014; Czarnecki and Shah, 2012). We investigated when Y-link structures are first observed ultrastructurally during axoneme elongation. Analyses of cross-sections identified obvious Y-shaped fibers emanating from dMTs at the 1.5-, two-, and three-fold stages (Figure 2A–C). At the comma stage, we observed shorter fiber-like densities without fully formed Y-link endings extending from subsets of dMTs (Figure 1Diii), potentially representing nascent Y-links. These observations suggest that structural features of TZs of a subset of amphid sensory neurons are at least partly established by the 1.5-fold stage of embryogenesis.

A puzzling feature of amphid sensory cilia is the presence of a variable number of inner singlet MTs (isMTs) inside adult axonemes (Perkins et al., 1986; Doroquez et al., 2014). These isMTs are smaller in diameter compared to A-tubules of the centriole/BB/axoneme and contain 11 protofilaments similar to cytoplasmic MTs in C. elegans (Perkins et al., 1986; Doroquez et al., 2014; Chalfie and Thomson, 1979). We defined isMTs as fully closed MTs that were observed inside axonemes over multiple tomographic slices. Using these criteria, we observed only one isMT in one axoneme at the two-fold stage and multiple isMTs in all examined axonemes at the three-fold stage (Figure 2Biv, Ciii and Civ; Figure 2—figure supplement 3). However, we detected incompletely closed MTs and short MT-like structures inside multiple axonemes at the two-fold stage (Figure 2—figure supplement 3A); these structures may represent early stages of isMT assembly. We also noted a ring-like structure in distal regions of the TZs in two- and three-fold axonemes (Figure 2Bii–iii, Cii–iii). This structure is likely the apical ring hypothesized to provide an attachment site for isMTs (Perkins et al., 1986; Doroquez et al., 2014; Blacque and Sanders, 2014). The origin and function of the isMTs remain to be determined.

Discussion

Our observations suggest that in a subset of ciliated sensory neurons in the head amphid organs of C. elegans, the outer centriole wall is remodeled to initiate ciliogenesis and persists thereafter into adulthood, while the centriole core degenerates starting at the 1.5-fold embryonic stage (summarized in Figure 3C). This remodeling is consistent with the persistence of a subset of outer centriole wall- and centriole-associated proteins (eg. HYLS-1 and DYF-19/FBF1) through postembryonic stages (Dammermann et al., 2009; Mohan et al., 2013; Wei et al., 2013, 2016). Thus, amphid channel cilia in adult C. elegans hermaphrodites do not completely lack a BB, but instead possess a modified BB at their base. Proteins (e.g. kinesin-II, DYF-19/FBF1, and HYLS-1) localized to this region in adult cilia have previously been implicated in loading, ciliary import, and recycling of IFT machinery (Wei et al., 2013, 2016; Prevo et al., 2015), suggesting that this modified BB retains at least a subset of the functions of canonical BBs in other organisms.

The observed closure of sMT-associated hooks to form dMTs in C. elegans BBs is reminiscent of observations in Drosophila. Although mature centrioles generally contain nine dMTs in the Drosophila embryo, centrioles consisting of nine sMTs with lateral hooks have also been observed, and it has been speculated that these structures represent intermediates in the centriole assembly process (Gottardo et al., 2015). Similar to flagella in Chlamydomonas, where B-tubules are assembled by tubulin addition onto A-tubules from the outer to inner AB junction (Nicastro et al., 2011; Linck and Stephens, 2007), B-tubules in C. elegans amphid BBs appear to form via closure of A-tubule-associated hooks at the inner AB junction. Intriguingly, B-tubules open at the inner AB junction have been observed in C. elegans and mice mutant for the TZ component NPHP-4 and the small GTPase Arl13b, respectively, raising the possibility that these defects may be a consequence of failure to close this junction during development (Jauregui et al., 2008; Caspary et al., 2007). Similar to our findings, asynchronous growth of dMTs comprising the outer centriole wall has been reported in Paramecium (Dippell, 1968), Chlamydomonas (Preble et al., 2001), human lymphoblasts (Guichard et al., 2010), and Drosophila germline stem cells (Gottardo et al., 2015). The mechanisms that mediate centriole wall remodeling are unknown.

Similar to Drosophila, BBs in C. elegans appear to lack structurally visible appendages (Gottardo et al., 2015; González et al., 1998; Doroquez et al., 2014; Callaini and Riparbelli, 1990; Callaini et al., 1997). As proposed previously, the absence of transition fibers suggests that BBs in C. elegans dock to the cell membrane via alternate mechanisms possibly requiring interaction of the BB and/or TZ with cell adhesion molecules (Schouteden et al., 2015; Williams et al., 2011; Nechipurenko et al., 2016; Heiman and Shaham, 2009). However, we note that in HPF-FS specimens, mesh-like structures are often less clearly visible than in conventional chemically fixed specimens in which fibrous networks can collapse into dark electron-dense appearing structures (McEwen et al., 1998). Thus, it remains possible that in C. elegans, transition fiber-associated proteins are organized in a less compact structure that is distinct from that in vertebrates. We detect structures resembling putative nascent Y-links of the TZ associated with dMTs in the comma stage when the central tube has not yet degenerated. At the 1.5-fold stage, clear Y-links are associated with BBs that either lack or contain the central tube, suggesting that while initiation of Y-link formation may require the centriole core, the core is dispensable for axoneme elongation. We find that loss of the central tube coincides with widening of the cilia base starting at the 1.5-fold stage of embryogenesis, suggesting that dMT flaring at the ciliary base is a consequence of central tube loss. We note that the timing of early ciliogenesis events may be distinct in ciliated cell types that were not examined in this study. In the future, it will be important to correlate the presence (or absence) of distinct subciliary structures with that of their known molecular components in individual cell types across developmental stages in order to obtain a more complete description of early ciliogenic steps. We expect that the ability to visualize centrioles/BBs and cilia in single cells in vivo together with the genetic power of C. elegans will allow further characterization of the conserved and species-specific mechanisms that underlie biogenesis and maintenance of these important cellular organelles.

Materials and methods

Strains

Wild-type strain of C. elegans (Bristol N2) was obtained from the Caenorhabditis Genetics Center and cultured on standard nematode growth media plates seeded with E. coli OP50.

Specimen preparation

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Gravid hermaphrodites were cut open to release eggs, which were then allowed to develop to the desired stage at room temperature in M9 buffer. Bean, comma, 1.5-fold, two-fold, and three-fold embryos were identified based on morphology. To collect 350 mpf embryos, single-cell embryos were manually sorted and allowed to develop for ~350 mins in M9 at 22°C. Embryos of the desired developmental stage were suctioned into cellulose capillary tubes (200 µm diameter, Leica Microsystems) in M9 and sealed.

The HPF-FS preparation was performed as described previously (Doroquez et al., 2014). Briefly, embryos in sealed capillary tubes were placed in the cavity between two aluminum planchettes (type ‘A’ hat, 100 µm deep, and the flat surface of type ‘B’ hat, Wohlwend, Switzerland) that was filled with 20% bovine serum albumin (BSA) in M9. The quickly assembled planchette sandwich was rapidly high pressure-frozen using a Leica EM HPM100 HPF system (Leica Microsystems, Vienna, Austria). Freeze-substitution was performed at −90°C over 3–4 days in fixation solution [1% osmium tetroxide (19100, EMS), 0.5% glutaraldehyde (16530, EMS), 2% water in anhydrous acetone (AC32680-1000, Fisher)] using a Leica EM AFS2 FS system, before the temperature was progressively increased to 4°C (5°C/hr). After 1 hr at 4°C, samples were washed with anhydrous acetone (4 × 30 mins), infiltrated and flat embedded in Araldite 502/Embed-812 Resin [Araldite (10900, EMS), Embed-812 (14900, EMS), DDSA (13710, EMS)] at room temperature, and polymerized at 60°C for several days. Flat-embedded samples were subsequently re-embedded in order to obtain cross sections of embryos.

Serial section TEM and ET

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Serial sectioning, electron microscopy and tomography, and image processing were performed as described previously (Doroquez et al., 2014). Briefly, serial plastic sections (70-nm thick) were collected on slot grids covered with Formvar support film, post-stained with saturated solution of uranyl acetate (0379, Polysciences, Inc., Warrington, PA) for 15 min, and Reynold’s lead citrate (Lead nitrate - 17900, EMS, and Sodium citrate - S-279, Fisher) for 7 min, and imaged using a Tecnai F20 (200 keV) or F30 (300 keV) transmission electron microscope (FEI, Hillsboro, OR) equipped with a 2K ×2K charged-coupled device (CCD) camera. For large overviews of sections, we acquired montages of overlapping high-magnification images. For electron tomography, BSA-coated, 10 nm colloidal gold fiducials (Au - Sigma-Aldrich, St. Louis, MO; BSA - SC-2323, Santa Cruz Biotechnology, Inc.) (Iancu et al., 2006) were applied to the sections, before acquiring dual-axis tilt series with a tilt range of ±60° with 1° increments around each axis. Automated montage and tilt series acquisition was facilitated by the microscope control software SerialEM (Mastronarde, 2005). Image processing, such as blending montages and reconstructing tomograms, was performed using various tools from the IMOD software package (Kremer et al., 1996).

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Decision letter

  1. Oliver Hobert
    Reviewing Editor; Howard Hughes Medical Institute, Columbia University, United States

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "Centriolar remodeling underlies basal body maturation during ciliogenesis in Caenorhabditis elegans” for consideration by eLife. Your article has been reviewed by two peer reviewers, and the evaluation has been overseen by Oliver Hobert as Reviewing Editor and K VijayRaghavan as the Senior Editor.

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

The manuscript by Nechipurenko et al. is a logical follow up from authors' previous eLife paper defining an ultrastructural map of C. elegans sensory cilia and glia. Here, the authors again use ssTEM and ssET to capture early steps of ciliogenesis in rapidly forming cilia in C. elegans embryos, which is a difficult and heroic accomplishment in microscopy. The authors make a number of very interesting and unexpected findings that are worthy of publication pending some revisions.

1) Many of the inferred temporal events presented in the concluding model appear somewhat speculative in the absence of some type of complementary real-time visualization using light microscopy. The authors need to better emphasize the caveats associated with their deductions.

2) Some of the implications of the work seem somewhat overstated – e.g. in the Discussion section the authors conclude that "[…]cilia in adult C. elegans […] possess a modified BB[…]that[…]provide(s) a site for docking, assembly and turnaround of IFT proteins…" but what is the evidence that these 3 processes require the modified BB? the authors should consider discussing their speculations about the import of the IFT machinery in the context of the paper by Prevo et al., 2015, Nat Cell Biol, 17, 1536 which reports the specific role of heterotrimeric kinesin-2 in importing the IFT machinery through structures described in the current paper at the base of the cilium.

3) Some implications of the authors findings are not emphasized in this manuscript and the authors are encouraged to discuss implications of their findings that will appeal to a broader audience. Overstatements should be avoided and caveats be presented:a) For example the hook-like structures on A-tubules have been observed in degenerating ciliary axonemes in adult C. elegans(for example: arl-13, nphp-4, ccpp-1), mouse (for example, Arl13b/hennin mutant), and zebrafish (Fleer mutant). Findings in this report suggest that these hook-like structures observed in other systems may represent a defect in development and not degeneration, and that cilia employ an evolutionarily conserved mechanism to construct the B-tubule. This would be a major change in how we view, interpret, and seek treatment options for ciliopathies.b) Another example, the authors observe vesicles within the C. elegans TZ and conclude "that the ciliary gate function of C. elegans TZs may be partly distinct from that in vertebrate cilia." An alternative possibility is that this study has provided unprecedented insight to multiple ciliated cell types and that this is not a worm oddity. Rather, it is technically impossible to do the same experiment in vertebrates (ie survey 10-12 vertebrate ciliated cell types at the same time in a developing embryo and in the adult). Because of the neuroanatomy of the worm (all the cilia are in the head) and studies by the Nicastro and Sengupta labs, we now have an appreciation of the diversity and complexity of developing and adult C. elegans cilia – there is no reason to think that vertebrate cilia are less complex. We just can't see it yet.

https://doi.org/10.7554/eLife.25686.011

Author response

The manuscript by Nechipurenko et al. is a logical follow up from authors' previous eLife paper defining an ultrastructural map of C. elegans sensory cilia and glia. Here, the authors again use ssTEM and ssET to capture early steps of ciliogenesis in rapidly forming cilia in C. elegans embryos, which is a difficult and heroic accomplishment in microscopy. The authors make a number of very interesting and unexpected findings that are worthy of publication pending some revisions.

1) Many of the inferred temporal events presented in the concluding model appear somewhat speculative in the absence of some type of complementary real-time visualization using light microscopy. The authors need to better emphasize the caveats associated with their deductions.

We now emphasize this very valid caveat in the legend to Figure 2C. We also note this caveat in the Discussion.

2) Some of the implications of the work seem somewhat overstated – e.g. in the Discussion section the authors conclude that "[…]cilia in adult C. elegans […] possess a modified BB[…]that[…]provide(s) a site for docking, assembly and turnaround of IFT proteins…" but what is the evidence that these 3 processes require the modified BB? the authors should consider discussing their speculations about the import of the IFT machinery in the context of the paper by Prevo et al., 2015, Nat Cell Biol, 17, 1536 which reports the specific role of heterotrimeric kinesin-2 in importing the IFT machinery through structures described in the current paper at the base of the cilium.

Our statement was based on work from the Dammermann and Hu labs. These groups recently reported that in animals mutant for the centriolar outer wall-associated protein HYLS-1, the docking and import of IFT machinery, and formation of the transition zone (TZ) and axoneme are severely impaired (1). HYLS-1 is also required for the formation of transition fiber (TF)-like structures comprised of the DYF-19/FBF1 protein among others. In an earlier report, the Hu lab showed that IFT particle entry into the cilium is also compromised in dyf-19 mutants (2). Together, these papers suggest that at least some proteins localized to the modified BB region are important for distinct aspects of IFT. It is possible that the structural defects in hyls-1 and dyf-19 mutants compromise IFT particle entry by affecting kinesin-II-dependent transport of IFT trains through the ciliary base and TZ (3). We have now modified this and related sentences in the manuscript.

3) Some implications of the authors findings are not emphasized in this manuscript and the authors are encouraged to discuss implications of their findings that will appeal to a broader audience. Overstatements should be avoided and caveats be presented:a) For example the hook-like structures on A-tubules have been observed in degenerating ciliary axonemes in adult C. elegans (for example: arl-13, nphp-4, ccpp-1), mouse (for example, Arl13b/hennin mutant), and zebrafish (Fleer mutant). Findings in this report suggest that these hook-like structures observed in other systems may represent a defect in development and not degeneration, and that cilia employ an evolutionarily conserved mechanism to construct the B-tubule. This would be a major change in how we view, interpret, and seek treatment options for ciliopathies.

This is an intriguing suggestion. As noted in this manuscript, the inner AB junction is open initially forming A-tubules with the associated hooks originating at the outer AB junction. The inner junction closes later in development to form the AB doublets. Thus, B-tubules that are open at the inner AB junction such as in Arl13b hnn mutant mice (4) and nphp-4 mutant worms (5) could potentially represent a developmental defect in B-tubule formation. However, ccpp-1 tubulin deglutamylase mutants in C. elegans exhibit a range of B-tubule defects complicating interpretation (6), and the fleer mutant in zebrafish that exhibits reduced tubulin polyglutamylation appears to show defects in the outer AB junction (7). Compared to the inner AB junction, the outer junction has been reported to be more fragile and susceptible to breakage during biochemical experiments (8-10) suggesting that the hook-like structures observed in fleer mutants may arise due to weakening of the outer AB junction as a consequence of altered tubulin posttranslational modifications. We have now added speculation regarding the ultrastructural defects in hnn and nphp-4 mutants to the text.

b) Another example, the authors observe vesicles within the C. elegans TZ and conclude "that the ciliary gate function of C. elegans TZs may be partly distinct from that in vertebrate cilia." An alternative possibility is that this study has provided unprecedented insight to multiple ciliated cell types and that this is not a worm oddity. Rather, it is technically impossible to do the same experiment in vertebrates (ie survey 10-12 vertebrate ciliated cell types at the same time in a developing embryo and in the adult). Because of the neuroanatomy of the worm (all the cilia are in the head) and studies by the Nicastro and Sengupta labs, we now have an appreciation of the diversity and complexity of developing and adult C. elegans cilia – there is no reason to think that vertebrate cilia are less complex. We just can't see it yet.

The reviewer makes a good point. Given that this was pure speculation and somewhat tangential to the main points of the manuscript, we deleted this section of the Discussion.

References

1. Wei Q, Zhang Y, Schouteden C, Zhang Y, Zhang Q, Dong J, et al. The hydrolethalus syndrome protein HYLS-1 regulates formation of the ciliary gate. Nat Commun. 2016;7: 12437.

2. Wei Q, Xu Q, Zhang Y, Li Y, Zhang Q, Hu Z, et al. Transition fibre protein FBF1 is required for the ciliary entry of assembled intraflagellar transport complexes. Nat Commun. 2013;4: 2750.

3. Prevo B, Mangeol P, Oswald F, Scholey JM, Peterman EJ. Functional differentiation of cooperating kinesin-2 motors orchestrates cargo import and transport in C. elegans cilia. Nat Cell Biol. 2015;17: 1536-1545.

4. Caspary T, Larkins CE, Anderson KV. The graded response to Sonic Hedgehog depends on cilia architecture. Dev Cell. 2007;12: 767-778.

5. Jauregui AR, Nguyen KC, Hall DH, Barr MM. The Caenorhabditis elegans nephrocystins act as global modifiers of cilium structure. J Cell Biol. 2008;180: 973-988.

6. O'Hagan R, Piasecki BP, Silva M, Phirke P, Nguyen KC, Hall DH, et al. The tubulin deglutamylase CCPP-1 regulates the function and stability of sensory cilia in C. elegans. Curr Biol. 2011;21: 1685-1694.

7. Pathak N, Obara T, Mangos S, Liu Y, Drummond IA. The zebrafish fleer gene encodes an essential regulator of cilia tubulin polyglutamylation. Mol Biol Cell. 2007;18: 4353-4364.

8. Witman GB, Carlson K, Berliner J, Rosenbaum JL. Chlamydomonas flagella. I. Isolation and electrophoretic analysis of microtubules, matrix, membranes, and mastigonemes. J Cell Biol. 1972;54: 507-539.

9. Stephens RE. Thermal fractionation of outer fiber doublet microtubules into A- and B-subfiber components. A- and B-tubulin. J Mol Biol. 1970;47: 353-363.

10. Linck RW, Langevin GL. Reassembly of flagellar B (α β) tubulin into singlet microtubules: consequences for cytoplasmic microtubule structure and assembly. J Cell Biol. 1981;89: 323-337.

https://doi.org/10.7554/eLife.25686.012

Article and author information

Author details

  1. Inna V Nechipurenko

    Department of Biology and National Center for Behavioral Genomics, Brandeis University, Waltham, United States
    Contribution
    IVN, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing—original draft, Writing—review and editing
    Contributed equally with
    Cristina Berciu
    For correspondence
    ivn@brandeis.edu
    Competing interests
    No competing interests declared.
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-0249-6620
  2. Cristina Berciu

    Department of Biology and Rosenstiel Basic Medical Sciences Research Center, Brandeis University, Waltham, United States
    Present address
    Microscopy Core Facility McLean Hospital, Belmont, United States
    Contribution
    CB, Formal analysis, Validation, Investigation, Visualization, Methodology
    Contributed equally with
    Inna V Nechipurenko
    Competing interests
    No competing interests declared.
  3. Piali Sengupta

    Department of Biology and National Center for Behavioral Genomics, Brandeis University, Waltham, United States
    Contribution
    PS, Conceptualization, Supervision, Funding acquisition, Visualization, Writing—original draft, Project administration, Writing—review and editing
    For correspondence
    sengupta@brandeis.edu
    Competing interests
    PS: Reviewing editor, eLife
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7468-0035
  4. Daniela Nicastro

    1. Department of Biology and Rosenstiel Basic Medical Sciences Research Center, Brandeis University, Waltham, United States
    2. Departments of Cell Biology and Biophysics, University of Texas Southwestern Medical Center, Dallas, United States
    Contribution
    DN, Conceptualization, Formal analysis, Supervision, Funding acquisition, Visualization, Project administration, Writing—review and editing
    For correspondence
    daniela.nicastro@utsouthwestern.edu
    Competing interests
    No competing interests declared.
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0122-7173

Funding

National Institutes of Health (R37 GM56223)

  • Piali Sengupta

National Institutes of Health (R01 GM083122)

  • Daniela Nicastro

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

Acknowledgements

We are grateful to Alex Dammermann, Max Heiman and Shai Shaham for discussions and communication of data prior to publication. We thank Oliver Blacque for comments on the manuscript. We are grateful to Chen Xu for providing training and for management of the Brandeis EM facility. This work was supported in part by the NIH (R37 GM56223 – PS and R01 GM083122 – DN).

Reviewing Editor

  1. Oliver Hobert, Howard Hughes Medical Institute, Columbia University, United States

Publication history

  1. Received: February 3, 2017
  2. Accepted: March 15, 2017
  3. Version of Record published: April 15, 2017 (version 1)

Copyright

© 2017, Nechipurenko 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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