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. Author manuscript; available in PMC: 2018 Apr 3.
Published in final edited form as: Curr Biol. 2017 Mar 16;27(7):968–980. doi: 10.1016/j.cub.2017.02.039

Cell-specific α-tubulin isotype regulates ciliary microtubule ultrastructure, intraflagellar transport, and extracellular vesicle biology

Malan Silva 1,2, Natalia Morsci 2, Ken C Q Nguyen 3, Anza Rizvi 1, Christopher Rongo 1,2, David H Hall 3, Maureen M Barr 1,2,*
PMCID: PMC5688951  NIHMSID: NIHMS859000  PMID: 28318980

Summary

Cilia are found on most non-dividing cells in the human body and, when faulty, cause a wide range of pathologies called ciliopathies. Ciliary specialization in form and function is observed throughout the animal kingdom, yet mechanisms generating ciliary diversity are poorly understood. The “tubulin code” – a combination of tubulin isotypes and tubulin post-translational modifications – can generate microtubule diversity. Using C. elegans, we show that α-tubulin isotype TBA-6 sculpts 18 A- and B-tubule singlets from nine ciliary A-B doublet microtubules in cephalic male (CEM) neurons. In CEM cilia, tba-6 regulates velocities and cargoes of intraflagellar transport (IFT) kinesin-2 motors kinesin-II and OSM-3/KIF17 without affecting kinesin-3 KLP-6 motility. In addition to their unique ultrastructure and accessory kinesin-3 motor, CEM cilia are specialized to produce extracellular vesicles. tba-6 also influences several aspects of extracellular vesicle biology, including cargo sorting, release, and bioactivity. We conclude that this cell-specific α-tubulin isotype dictates the hallmarks of CEM cilia specialization. These findings provide insight into mechanisms generating ciliary diversity and lay a foundation for further understanding the tubulin code.

eTOC blurb

Silva et al. show that a cell-specific α-tubulin is required for ciliary specialization, motor coordination, and several aspects of extracellular vesicle biology.

Introduction

Microtubules are essential for many cytoskeletal processes including cell division, ciliogenesis, and intracellular trafficking. Microtubules are composed of α+β tubulin heterodimers arranged into protofilaments that form cylinders [1]. These cylinders can also form higher order microtubule structures like A-B tubule doublets and A-B-C triplets found in the axonemes and basal bodies of cilia and flagella [2]. Eukaryotes express multiple tubulin isotypes that may function redundantly or perform specialized functions [3, 4]. Mutations in certain α and β-tubulin isotypes cause a spectrum of neurological disorders [5]. The mechanisms by which tubulin isotypes contribute to higher order microtubule structural and functional diversity are not well understood.

Cilia are organelles with a microtubule core, called the axoneme, that exhibit a conserved architecture of nine outer doublet microtubules with a variable number of inner singlets [6, 7]. The axoneme has distinct structural regions: the basal body (microtubule triplets), transition zone (microtubule doublets attached to the surrounding membrane via “Y” links), middle segment (microtubule doublets), and distal segment (microtubule singlets). Microtubule doublets consist of complete A-tubules with 13-protofilaments and attached incomplete 10-protofilament B-tubules [8]. Axonemal structural variations may arise in the relative length of each region and the presence or absence of the distal segment, where A-tubule singlets extend and B-tubules terminate [7]. For example, the mammalian kidney displays a variant ciliary axoneme that is composed of a short proximal microtubule doublet and long distal microtubule singlet region [9] and is reminiscent of C. elegans amphid channel cilia [10, 11]. Ciliary specialization mechanisms including those regulating microtubule doublet to singlet conversion are unknown.

C. elegans possesses numerous ciliary specializations that can be visualized in vivo using fluorescent reporters and examined ultrastructurally using transmission electron microscopy and electron tomography [10–12]. Cilia are built by a conserved process called intraflagellar transport (IFT). IFT is a bidirectional microtubule-based motor driven cargo transport system that consist of biochemically separable and functionally distinct IFT-A and -B particles transported by anterograde heterotrimeric kinesin-II and retrograde dynein-2 motors [6, 13]. In some cilia, anterograde IFT transport is modulated by accessory kinesins. In C. elegans amphid neurons, homodimeric kinesin-2 OSM-3/KIF17 participates in IFT and is implicated in generating sensory cilia of diverse morphology and function [14]. In C. elegans cephalic male (CEM) neurons, kinesin-3 KLP-6 modulates velocities of both IFT kinesin-2 motors and also regulates release of ciliary extracellular vesicles (EVs) [15–17].

The “tubulin code” employs tubulin isotypes and tubulin post-translational modifications to orchestrate cytoskeletal functions [3, 18–20]. The tubulin code is “written” by tubulin isotypes and tubulin modifying enzymes such as tubulin tyrosine ligase-like enzymes (TTLLs) that glutamylate microtubules [21]. The tubulin code can be erased by tubulin modifying enzymes such as cytosolic carboxypeptidase (CCP) deglutamylases that remove glutamylates from microtubules [22]. The tubulin code is “read” or interpreted by cellular effectors such as molecular motors or microtubule associated proteins. We previously showed that the C. elegans ccpp-1 deglutamylase regulates stability of B-tubules and controls the velocity of kinesin-3 KLP-6 and kinesin-2 OSM-3/KIF17 without affecting the transport of kinesin-II cargo [23].

The C. elegans genome encodes nine α-tubulin and six β-tubulins [24, 25]. The CEM neurons express a cell-type specific α-tubulin isoform TBA-6 [26], suggesting that TBA-6 may contribute to CEM specialization. Using serial transmission electron microscopy and electron tomography of CEM cilia, we discovered a novel axonemal variation whereby doublet microtubules are splayed to form complete A- and B-tubule singlets in middle regions while remaining attached at their proximal and distal ends. α-tubulin TBA-6 was essential for B-tubule singlet formation. In tba-6 mutants, the CEM axoneme was comprised of a proximal doublet microtubule region followed by a distal A-tubule singlet microtubule region. The coordination of kinesin-2 motors with IFT-A and IFT-B complexes was disrupted in tba-6 mutant cilia. tba-6 was also required for extracellular vesicle (EV) cargo sorting, EV release, and EV bioactivity. Our findings demonstrate that the tubulin code via α-tubulin isotype TBA-6 influences axonemal microtubule architecture, IFT dynamics, and extracellular vesicle biology.

Results

CEM ciliary doublet microtubules splay to form A-tubule and B-tubule singlet microtubules

We characterized the ultrastructure of CEM cilia using transmission electron microscopy and electron tomography. In the male head, four quadrant cephalic sensilla contain the cilia of the male-specific CEM and non-sex specific CEP ciliated sensory neurons and associated glial socket and sheath cells that create a lumen surrounding the cilia (Figure 1A). The CEM ciliary axoneme can be subdivided into distinct segments based on microtubule ultrastructure (Figure 1B), described here in proximal-to-distal order.

Figure 1. Specialized ultrastructure of adult CEM cilia reveals a novel axonemal microtubule arrangement.

Figure 1

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A Left: Adult C. elegans male DIC image with region of interest boxed. Middle: Each of four quadrant cephalic sensilla contains the endings of CEM and CEP neurons and the socket and sheath support cells. Right: The cephalic sensillum contains cilia of the CEM and CEP neurons surrounded by socket (blue) and sheath (pink) cells. The CEM cilium (green) curves out and the tip protrudes to the environment via a cuticular pore (orange); while CEP cilium (gray) curves in and embeds in the cuticle. The cephalic lumen formed by the sheath and socket cells contains CEM-derived EVs [17].

B The CEM axoneme has five ultrastructurally distinct regions. Right: Serial electron tomogram model shows the relative positions of the five distinguishable regions. Sections labeled C-F correspond to subsequent panels. Left: TEM inserts show the representative microtubule organization. Scale bar, 250nm. dMT, doublet microtubule; sMT, singlet microtubule. Cartoon inserts show a stereotypical microtubule arrangement within each segment and do not directly reflect adjacent TEM sections.

C Tomogram model of A- and B- tubules of a microtubule doublet splitting into an A-tubule and B-tubule singlet. Left: Longitudinal section taken from a serial electron tomogram depicting a representative microtubule doublet splitting into A- and B- tubule singlets at the proximal doublet region. Middle: segmented tomogram model of the left panel depicting microtubules in orange, and ciliary membrane is green. Teal rings depict the boundary of 500–600nm long region where most microtubule doublets split into two singlets. Numbers correspond to the equivalent regions in the tomogram and model. Right: tomogram cross-sections of a representative doublet splitting into two singlets. Refer to Figure S1 for images of all nine microtubules.

D Some microtubules form bridges with the ciliary membrane. Left: flat-plane image taken from electron tomogram illustrating a microtubule singlet making three bridges with ciliary membrane. Right: density-based isosurface 3D model of this location. Middle: overlay. Scale bar, 50nm. Refer to Movie S2 for annotated 3D tomogram. In the 75nm cilia section examined, 5 out of 9 A-tubules, 6 out of 8 B-tubules, and 0 out of 2 inner singlets were connected the ciliary membrane at least at one point. Of those membrane-associated tubules, the number of connections was: 4.4±6.3 for an A-tubule and 11.16±3.9 for a B-tubule (mean±SD). The difference between A-tubule and B-tubule connections did not reach statistical significance

E Tomogram model of the sub-distal region depicting A- and B- tubule singlet fusion. Left: segmented electron tomogram model depicting fusion of A-tubule and B-tubule singlets (pink and teal false-colored), and the A-tubule singlet continuing to the distal segment (incomplete). Ciliary membrane is green. Right: false-colored longitudinal section from the electron tomogram used to generate the model on the left.

F B-tubules form transient C-shaped structures during the singlet-to-doublet transition in the sub-distal segment (refer to Figure 1B). Left panel: successive images from a tomogram. Scale bar, 50nm. Right panel: isosurface 3D models corresponding to images on the left. Middle panel: overlay.

See also Figure S1, Movies S1 and S2.

The transition zone contains nine doublet microtubules attached to the ciliary membrane by Y-links and arranged in a ring around one to four inner singlet microtubules. Doublet microtubules at the base of the transition zone flare out into the periciliary membrane compartment (PCMC) and terminate at varying lengths (Movie S1). At the anterior end of the transition zone, doublet microtubules became progressively devoid of Y-links. The doublet microtubule region is ~0.5μm and lacks Y-links. After the initial doublet segment, A- and B-tubules of the nine doublet microtubules splay to form discrete A-tubule and B-tubule singlets (Figure 1C). Incomplete B-subfibers separate at inner and outer seams from their partner A-tubules transiently display concave “C” shapes, and then seal to form complete B-tubule singlets (Figure 1F, Figure S1A–B). As a consequence of doublet splaying, the CEM axoneme contains 20±2 singlet microtubules along the length of its middle region (Figure 1B, quantified in 2D).

Some singlet microtubules closely associate with the ciliary membrane via electron-dense bridges (Figure 1D, Movie S2). By tracing A- and B-tubules along the cilia length, we found that A- and B- singlet microtubules eventually fuse to each other in the sub-distal region. The distal region possesses only A-tubule singlets (Figure 1B and E, Figure S1A). The distal-most ciliary region consists of tightly packed A-tubule singlets (Figure 1B also Figure S1B for an oblique view of the CEM cuticle pore). The number of singlets in this region ranged from nine immediately after the sub-distal region to as few as one at the tip. The CEM ciliary tip protrudes to the outside environment via an opening in the cuticle bulge (Figure 1A, Figure S1C). We also observed electron-opaque membrane structures continuous with CEM pore (Figure S1B). In sum, this is the first evidence of doublet microtubules splaying to generate discrete A-tubule and B-tubule singlets microtubules. B-tubule singlets exhibit a transient C-shape conformation during the doublet to singlet microtubule conversion.

α-tubulin isotype tba-6 is required for CEM ciliary ultrastructure

CEM cilia are approximately 3.5±0.4μm long and have a sinusoid curved shape visible by both transmission electron microscopy in fixed specimens and fluorescent reporters in living animals (Figure 1B, Figure 2A, 2D, 2F). In wild type (WT), 90% of CEM cilia curve outward from the central body axis of the worm (Figure 2A, 2F). Several genes regulate CEM cilia shape including α-tubulin tba-6, the IFT kinesin-II subunits klp-11, klp-20, kap-1, and the transition zone component nphp-4 [16, 26, 27]. tba-6 expression in the ciliated nervous system is restricted to the 27 ciliated extracellular vesicle (EV) releasing neurons including the four CEMs [26], while klp-11, klp-20, kap-1 and nphp-4 are pan-ciliary genes. A functional GFP-tagged TBA-6 reporter localizes along the length of the curved CEM axoneme (Figure S2C) [26]. We therefore focused on tba-6 as a potential CEM ciliary specialization factor.

Figure 2. tba-6 is required for CEM cilia curvature, microtubule architecture, and tubulin composition.

Figure 2

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A, B, B′ Serial TEM reconstructions of male cephalic sensilla of WT and tba-6 males. Pie charts (bottom) indicate the penetrance of the cilia curvature phenotype as assayed by TBB-4::tagRFP (WT N=83, tba-6 N=102 males).

A In WT, CEM (yellow) and CEP cilia (gray) share a lumen formed by the sheath (inner border in pink) and socket (teal) (refer to Figure 1A). The distal tip of the CEM cilium protrudes to the environment through a cuticular pore (brown) (also, Figure S1C). The CEP cilium curves in and embeds in the cuticle. EVs (green) are located in the proximal region of the cephalic lumen.

B 59% of tba-6 cilia curve-out but do not protrude through the cuticular pore. EVs were observed inside the cuticular pore (for TEM cross sections of tba-6, see Figure 4C).

B′ 41% of tba-6 cilia curve-in, follow the path of the CEP cilium, and embed in the cuticle. EVs were found in the lumen surrounding CEM and CEP cilia.

C In tba-6, microtubule doublets terminate and do not form A- and B- tubule singlets. Right: Serial TEM images of tba-6 cilia with A-tubules continuing as singlets and B-tubules terminating (black arrow). Left: Cross-sections of WT cilia from a serial tomogram showing a representative A- and B-tubule of a microtubule doublet splitting to form two singlets (white arrow) (refer to Figure 1F and Figure S1A, B). Scale bar, 50nm.

D Cartoon depicting microtubule architectures and cilia shapes.

E Cross sections of sub-distal CEM cilia in WT and tba-6 animals. Scale bar, 50nm.

Right: Microtubule singlet quantification: WT= 20±2, tba-6 curved-out = 14±2 tubules, mean±SD, p-value=0.0269 by Mann Whitney, N=3 animals, and n=12 cilia for each genotype.

F, F′, G, G′ β-tubulin TBB-4::GFP relative ciliary levels and ciliary localization. In WT, TBB-4::GFP localizes to CEM cilia in a stereotypical pattern; lines indicate two of four CEM cilia, quantified in G, n= 25 cilia. Ciliary TBB-4::GFP levels were normalized to levels in the CEM dendrite, mean fold intensity ± SD. This pattern is altered in tba-6 mutants, quantified in G′, n=20 cilia. In tba-6, TBB-4::GFP ciliary levels are significantly reduced and TBB-4::GFP accumulates at ciliary bases. F′, N=86 and 102 animals for WT and tba-6 respectively, p-value < 0.0001 by Mann-Whitney U test, Scale bar,10μm.

See also Table S1, and Figure S2.

We examined CEM cilia of tba-6(cxP4018) loss of function mutants (Figure S2) using fluorescent reporters. To visualize the CEM ciliary axoneme in vivo, we expressed GFP-tagged β-tubulin tbb-4 under the CEM-specific pkd-2 promoter (Figure 2F, Table S1). tbb-4 encodes a pan-ciliary β-tubulin isotype [28]. In WT CEM cilia, TBB-4::GFP localizes along the sinusoid curved-out axoneme, with reduced GFP intensity at the transition zone and sub-distal region (Figure 2F quantified in 2G). In tba-6 mutants, 41% of CEM cilia showed an abnormal curved-in morphology that deflected away from the cuticular pore and bent inward adjacent to the CEP cilium (Figure 2B′ and 2F). 59% of tba-6 CEM cilia curved-out and terminated before reaching the cuticular pore (Figure 2B).

Using serial transmission electron microscopy and electron tomography, we found that doublet microtubules do not splay into singlets in either curved-out or curved-in tba-6 CEM cilia (Figure 2C and 2D). Instead, the doublet microtubule region was defined by the abrupt termination of B-tubules, with A-tubule singlets extending distally (Figure 2C–D). As a consequence, the number of singlets was reduced to almost half in tba-6 males (Figure 2E). The doublet microtubule region was longer in curved-in tba-6 than curved-out tba-6 CEM cilia (Figure 2D). We conclude that α-tubulin TBA-6 is required for formation of B-tubule singlets and characteristic shape of CEM cilia.

TBA-6 regulates relative ciliary abundance of ciliary kinesin-2 and kinesin-3 motors but only affects ciliary distribution of accessory kinesin motors

To determine whether tba-6 regulates localization of ciliary proteins, we examined GFP-tagged ciliary kinesin motors and IFT polypeptides in tba-6 and WT CEM cilia (Figure 3A–C, Table S1). Ciliary fluorescence intensity of all ciliary motors were significantly increased in tba-6 mutants (Figure 3A–C, Table S1). However, only OSM-3::GFP and KLP-6::GFP ciliary distribution patterns were abnormal in tba-6 mutant cilia (Figure 3A, quantified in 3B and 3C). CEM ciliary distributions of IFT-A complex component CHE-11::GFP, IFT-B complex proteins OSM-5::GFP and OSM-6::GFP were similar in WT and tba-6 mutant males (Figure 3A and 3B), while ciliary fluorescence intensity levels of KAP-1::GFP and OSM-5::GFP were slightly elevated in tba-6 mutants. These results suggest that the loss of TBA-6 selectively affects ciliary distribution patterns of a subset of ciliary motors and IFT proteins, with the most dramatic changes observed for accessory kinesins OSM-3 and KLP-6.

Figure 3. Localization and velocity distributions of IFT motors and polypeptides in CEM cilia of wild type and tba-6 mutant animals.

Figure 3

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A Widefield fluorescence images of GFP tagged IFT polypeptides and motors. Lines in each image indicate two of the four CEM cilia. OSM-3::GFP and KLP-6 GFP are driven by the klp-6 promoter and expressed in both CEM (arrows) and IL2 cilia. All other reporters are driven by the pkd-2 promoter and expressed in CEM cilia. Phenotypes are summarized in Table S1. Scale bar is 10μm and applicable to all panels.

B Comparison of fluorescence levels in cilia of WT and tba-6 males normalized to cell body. Mean ± SD. ‘*’, ‘**’, ‘***’ indicate p-values of <0.01, <0.001 and, <0.0001 respectively by Mann-Whitney U test.

C Average CEM ciliary localization pattern of KAP-1::GFP and OSM-3::GFP in WT and tba-6 mutant backgrounds. ‘◆’ indicates feature used to align intensity profiles; center-line depicts the mean intensity and the shaded area represent the standard deviation.

D Velocity distributions of GFP-tagged IFT motors and polypeptides in CEM cilia of WT and tba-6 males. Distribution averages, standard deviations, and statistical analyses are summarized in Table 1.

See also Table S1.

tba-6 regulates anterograde velocity and coordination of IFT kinesin-2 motors but not kinesin-3 KLP-6

To determine whether tba-6 regulates IFT, we examined anterograde velocity of CHE-11::GFP (IFT-A), OSM-5::GFP (IFT-B), OSM-6::GFP (IFT-B), IFT kinesin-2 motors kinesin-II (KAP-1::GFP) and OSM-3::GFP in WT and tba-6 CEM cilia (Figure 3D, Table 1) [12]. In WT, the velocity ranges of CHE-11::GFP, OSM-5::GFP, OSM-6::GFP, and KAP-1::GFP overlap at ~0.4 μm/sec, while OSM-3::GFP moved at distinctly faster velocity range of ~0.6 μm/sec (Figure 3D, Table 1). These observations are consistent with our model in which heterotrimeric kinesin-II but not OSM-3 drives IFT in CEM cilia [16].

Table 1.

Velocity and frequency of anterograde IFT in CEM cilia.

Complex Reporter Genotype Velocity, μm/s p value n = # Cilia Frequency/30sec
Meana SD* Mean SD
Heterotrimeric kinesin II IFT motor KAP-1 WT 0.39 0.11 2.87E-12 25 5 2
tba-6 0.46 0.13 28 6 2
Homodimeric kinesin-2 IFT motor OSM-3 WT 0.62 0.18 3.02E-32 55 11 4
tba-6 0.74 0.24 57 14 4
Kinesin-3 ciliary motor KLP-6 WT 0.67 0.21 0.11 15 11 4
tba-6 0.64 0.21 28 11 3
IFT-A polypeptide CHE-11 WT 0.37 0.10 8.29E-44 26 9 3
tba-6 0.52 0.17 16 9 3
IFT-B polypeptide OSM-5 WT 0.43 0.15 8.64E-66 22 7 3
tba-6 0.69 0.24 21 13 3
IFT-B polypeptide OSM-6 WT 0.45 0.12 1.80E-53 16 9 3
tba-6 0.65 0.20 22 14 3
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Data obtained from in vivo imaging of GFP-tagged transgenic reporters. p-values indicate results of unpaired two-tailed t-test (with unequal variances) comparing wild type and tba-6 groups for a given reporter.

a

Mean and Standard Deviation were calculated based on the population of particles.

See Figure 3D for frequency histograms of these reporters.

In tba-6 mutants, IFT-A and IFT-B moved at different velocity ranges. IFT-A protein CHE-11::GFP and KAP-1::GFP moved at ~0.5 μm/sec. IFT-B proteins OSM-5::GFP and OSM-6::GFP velocities overlapped with OSM-3::GFP at ~0.7 μm/sec (Figure 3D, Table 1). Kinesin-3 KLP-6 velocity was unaffected (Table 1). These data suggest that, in tba-6 CEM cilia, heterotrimeric kinesin-II transports IFT-A and homodimeric kinesin-2 OSM-3 transports IFT-B.

tba-6 mutants retain ciliary extracellular vesicles (EVs) in the cephalic lumen

CEM neurons shed and release EVs [17, 29, 30]. GFP-tagged TRP polycystin PKD-2 and peripheral membrane protein CIL-7 localize to the CEM ciliary base, cilium, and EVs (Figure 4A–B, 5A) [31]. The ciliary base region includes the periciliary membrane compartment (PCMC) internal to the CEM neuron and the extracellular lumen surrounding the cilium. Ciliary base localization of PKD-2::GFP and CIL-7::GFP may reflect EVs that are shed into the lumen but not environmentally released [31]. In tba-6 mutants, PKD-2::GFP and CIL-7::GFP accumulated at ciliary base, suggesting an EV release defect (Figure 4A–B).

Figure 4. tba-6 regulates EV location, abundance, and size in the cephalic sensilla.

Figure 4

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A, B PKD-2::GFP and CIL-7::GFP abnormally accumulate in tba-6 mutant animals. Quantified on right. See Table S1 for statistical analysis. Lines indicate cilia. Scale bar, 10μm.

C Simplified serial TEM reconstructions depicting relative positions of lumenal EVs. Genotypes are indicated at the top and respective reconstructions and TEM images are arranged vertically. Scale bars in TEM images are 500nm.

C-top row The CEM cilium is red and distal dendrite is yellow. For simplicity, the cuticular pore is outlined using a dotted line and the CEP cilium is not shown. Dashed lines indicated by ‘Cu’(cuticular pore) or ‘P’ (proximal lumen) denote the relative position of the TEM cross sections.

C-middle row TEM cross section at the cuticular pore. The WT distal CEM cilium is present and contains microtubule singlets. The tba-6 curved out CEM cilium does not reach the cuticular pore. Instead, EVs are seen in the empty pore. tba-6 curved in cilia are deflected from the pore and embedded in the cuticle lumen next to the CEP cilium. EVs are seen in the cephalic lumen and cuticle pore.

C-bottom row TEM cross-sections from proximal sheath cell level. In WT, few EVs are observed in the cephalic lumen. In tba-6, EVs abnormally accumulate in the cephalic lumen.

D and E Comparison of average number of EVs and average EV diameter in the distal and proximal cephalic lumen. In WT, EVs were more abundant in the proximal lumen (distal=14±10 EVs, proximal=80±60 EVs, p-value=0.0087, ‘*’, by Mann-Whitney U test). When present, distally located EVs were significantly smaller than proximally located EVs (distal=36.67±0.63nm, proximal=105±29.14nm, mean±SD, n=8 cilia, p-value<0.0001 for size, ‘***’, by Mann-Whitney U test). tba-6 accumulated significantly more lumenal EVs than WT (distal=252±10 and proximal=753±77 EVs, mean±SD, p-value=0.007 for both with WT using Kruskal-Wallis with Dunn’s correction). tba-6 distally located EVs were larger than WT distally located EVs (diameters were 61.65±13.96nm and 54.86±9.76nm for curved out (n=6) and curved in (n= 2) sensilla, p-value<0.0001 compared with WT by Kruskal-Wallis with Dunn’s correction). tba-6 proximally located EVs were smaller than WT proximally located EVs (diameters were 84.57±35.21nm and 71.48±19.75nm, mean±SD, for curved-out and curved–in sensilla, p-values<0.0001 compared with wild type by Kruskal-Wallis with Dunn’s correction).

G and H Average histograms of EV diameter within the distal and proximal cephalic lumen in WT and tba-6. In tba-6 curved-in cilia 50–70nm sized EVs accumulate in the lumen (WT distal: 8 50–60nm EVs, tba-6 curved-in distal: 59 50–60nm EVs; WT proximal: 13 50–60nm EVs, tba-6 curved-in proximal: 346 50–60nm EVs n=8 sensilla for WT and n=2 sensilla for tba-6 curved-in, p-value<0.0001 by two-way ANOVA).

I Occlusion of the CEM cilia tip correlates with luminal EV accumulation. In WT, CEM cilia tips are environmentally exposed through the cuticular pore (see Figure S1C) and contain 47±53 lumenal EVs, n=7 sensilla. Tips of the tba-6 curved-out cilia do not reach pore (hence “partial” label), accumulate 116±170 EVs (n=6 sensilla). Tips of tba-6 curved-in cilia are not environmentally exposed and accumulate on 502±272 lumenal EVs (n=4 sensilla). R2 indicates ‘goodness of fit’ of the cubic polynomial line. Number of EVs reflect mean ± SD and n is number of cephalic sensilla analyzed via quantitative serial EM.

See also Table S1

Figure 5. tba-6 is required for ciliary EV cargo composition, release, and bioactivity.

Figure 5

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A WT and tba-6 males release PKD-2::GFP containing EVs. B Cartoon depicts the single-focal-plane fluorescence image acquired at the surface of the cover slip showing PKD-2::GFP-containing EVs in WT and tba-6 worms. In A, arrowheads point to EVs; yellow brackets mark PKD-2::GFP EV streaks. Abundance of EV streaks is quantified below. Environmental release of PKD-2::GFP-containing EVs is significantly reduced in tba-6 mutants (WT: ‘0’ = 0, ‘≤10’= 0.04±21, ‘≤50,’= 0.34±0.48, ‘>50’=0.60±0.49, fraction ± SD, N=88 worms; tba-6: ‘0’ = 0.14±35, ‘≤10’= 0.30±0.46, ‘≤50’= 0.52±0.50, ‘>50’=0.05±0.21, fraction ± SD, N=86 worms). WT and tba-6 mutants are significantly different in ‘≤10’, ‘≤50’ and ‘>50’catogories, ‘*’ and ‘***’ indicate p-values of <0.01 and <0.0001, respectively, using the Kruskal-Wallis test with Dunn’s correction for multiple comparisons. Error bars indicate SEM. Fraction of animals with PKD-2::GFP-positive EV streaks was significantly reduced in tba-6 background (WT = 0.37±0.49, tba-6 = 0.10±0.34) ‘***’ indicates p-value <0.0001 using Mann-Whitney test. Fraction ± SD, N = 88 (WT) and 86 (tba-6) animals. Error bars indicate SEM.

C In WT, KLP-6::GFP is excluded from EVs. In tba-6 mutants, KLP-6::GFP is ectopically shed and released into EVs. Arrowheads point to ectopic EVs (refer Figure 3). Quantified on right. Statistical details are summarized in Table S1. Scale bar is 10μm.

D TBA-6 is required for bioactivity of EVs.

Left: Young adult virgin WT males were individually placed on a freshly made EV-containing bacterial lawn and video recorded for 5 minutes. Inset Sample time series of tail-chasing behavior; in this case the male was tail-chasing from 117th to 124th second. Middle: WT males exhibit a basal level of tail chasing behavior (buffer control). WT-derived EVs increased the fraction of males exhibiting tail chasing (0.71±0.06 events/5 minutes vs buffer control 32±0.06 events/5 minutes, mean ± SEM, N=56 males and N=69 males for WT and buffer control, p-value<0.0001 with buffer control). In contrast, tail chasing events were similar with tba-6 derived EVs and buffer control (0.42±0.05 events/5 minutes, N= 69 tba-6 males, p-value>0.05 with buffer control and p-value<0.001 with WT EVs). Number within bars indicates number of animals. Letters indicate statistically distinct groups by Kruskal-Wallis test with Dunn’s correction for multiple comparisons. Error bars indicate SEM.

Right: Number of tail chasing episodes per assay increases when WT males are exposed to WT EVs but not tba-6-derived EVs.

See also Table S1

To resolve whether tba-6 regulates EV shedding into the lumen and/or EV environmental release, we used serial TEM. We examined the location of EVs, EV numbers, and EV diameters. The WT cephalic lumen contains two discernable EV populations based on location, abundance, and size. The distal lumen contained a few rare and smaller-sized EVs (39±10nm) (Figure 4C–F). The proximal lumen surrounding the PCMC and proximal cilium contained larger-sized EVs (105±29) (Figure 4C–E and, G). Proximally located EVs varied in size and appearance, with some being electron opaque and others translucent (Figure 4C; average distribution of EV sizes quantified in Figure 4G).

In tba-6 mutants with curved in cilia, EV numbers increased in distal and proximal luminal regions (distal 14±10 and proximal 80±60 EVs in WT versus distal=252±10 and proximal=753±77 EVs in tba-6 curved in cilia) (Figure 4C; quantified in 4D–G). This increase was primarily due to enrichment of 50–60nm diameter EVs (10–30 fold higher in the cephalic lumen of a curved in tba-6 cilium) (Figure 4F–G). Lumenal EV accumulation increased exponentially with decreasing environmental access of the CEM ciliary tip (Figure 5H). We conclude that TBA-6 is required for CEM-specific axoneme microtubule ultrastructure, cilia shape and EV release as manifested by abnormally high levels of lumenal EVs.

tba-6 regulates EV release and cargo composition

To confirm that lumenal EV accumulation correlates with an EV environmental release defect, we scored release of PKD-2::GFP-containing EVs in living animals. tba-6 mutants released fewer PKD-2::GFP-containing EVs and fewer PKD-2::GFP-labeled EV streaks from CEM cilia compared to WT (Figure 5A). In WT, KLP-6::GFP is not EV cargo [17]. Surprisingly, in tba-6 mutants, KLP-6::GFP localized to environmentally released EVs (Figure 5C), which may correlate with KLP-6::GFP abnormal ciliary localization patterns (Figure 3A). GFP-tagged kinesin-2 ciliary motors, IFT-A CHE-11, IFT-B OSM-5 and OSM-6, and TBB-4 were not released in EVs in WT or tba-6 animals. These results suggest that EV cargo composition is different between tba-6 and WT animals.

tba-6 is required for EV bioactivity

EVs isolated from WT animals stimulate male tail-chasing behavior, indicating a role in inter-animal communication [17]. EVs isolated from klp-6 animals do not contain PKD-2::GFP and do not stimulate male tail-chasing behavior, suggesting that EV bioactivity is affected by cargo content. We therefore examined whether TBA-6 regulates EV bioactivity.

We isolated EVs from mixed stage, male-enriched cultures of WT and tba-6 mutants as previously described [17]. We compared bioactivity of WT and tba-6-isolated EVs by measuring the ability of EV preps to evoke male tail chasing. When exposed to WT EV preparations, 71±0.05% of WT males displayed tail chasing behavior, as compared to 32 ±0.06% to buffer control (Figure 5D, middle panel). When exposed to EV preparations derived from tba-6 mutants, 42±0.06% of wild-type males displayed tail chasing behavior which is not significantly different from buffer control (Figure 5D, middle panel). For those males that displayed tail-chasing behavior, the frequency of repetitive tail chasing was more robust in response to WT EVs than to tba-6-derived EVs (Figure 5D, right panel). We conclude that α-tubulin TBA-6 is important for EV cargo content and EV bioactivity.

Discussion

α-tubulin isotype TBA-6 sculpts 18 singlet MTs from nine doublet MTs. Flagella of mammalian spermatozoa also display A- and B-tubule singlets extending from microtubule doublets [32, 33], hinting that a conserved but unexplored mechanism generates this ciliary ultrastructure. How are nine A-tubule and nine B-tubule singlets generated from an axoneme with nine doublets? In vitro studies showed that formation of B-tubule based singlets can be energetically stable and persist in partially unwound states [34, 35]. We observed B-tubules transiently forming C-shapes when separating from partner A tubules and before sealing to form B-tubule singlets (Figure 1C, 1F). In the absence of tba-6, the CEM axoneme is composed of proximal microtubule doublets followed by a distal A-tubule microtubule singlet region, resembling C. elegans amphid channel cilia [10, 11]. We conclude that α-tubulin TBA-6 is essential for architectural remodeling of axonemal MT-doublets into A- and B-tubule singlets (see model in Figure 6).

Figure 6. Model of a doublet microtubule, IFT motors and polypeptides in WT and tba-6 CEM cilia.

Figure 6

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In WT, microtubule doublets splay to form A- and B-tubule singlets. Heterotrimeric kinesin-II and IFT A- and B-polypeptides are co-transported at overlapping velocities. Homodimeric kinesin-2 OSM-3 travels at a higher velocity separate from IFT polypeptides, transporting unknown cargo. KLP-6 is not included in this model.

In tba-6, microtubule doublets do not splay to form A- and B- tubule singlets. The doublet region is elongated and terminates abruptly. Only A-tubule singlets extend. Axonemal abnormalities accompany changes in IFT. IFT-A and –B polypeptides travel at distinct velocities that overlap with heterotrimeric kinesin-II and OSM-3, respectively.

Red squares indicate possible TBA-6 sites of action. We propose a model where the separation of microtubule doublets into distinct A-tubule and B-tubule singlets acts as a physical substrate for the functional separation of these two conserved IFT kinesins. Alternatively, tubulin composition and post-translational modifications may also impact IFT-motor-cargo dynamics.

We propose three potential mechanisms by which TBA-6 and the tubulin code generate A- and B-tubule singlets from doublet microtubules. TBA-6 may directly or indirectly regulate protofilament composition necessary for stabilizing the intermediate C-tubule structure and the transition of B-tubules from doublets to singlets (Figure 1 and 6). Consistent with this hypothesis, β-tubulin β3 regulates protofilament number in flagellar microtubules of Drosophila spermatozoa [36] and α-tubulin MEC-12 and tubulin acetyltransferase MEC-17 generate 15-protofilament neuronal microtubules in non-ciliated C. elegans touch receptor neurons [37–39].

Alternatively, TBA-6 may act as a buffer to temper against hyperglutamylation and resultant B-tubule instability [23, 40]. The C-terminus of TBA-6 is an unusually long and positively charged compared to other α-tubulin isotypes (Figure S2B). The C-tail of α-tubulin TBA-6 lacks glutamate residues (Figure S2B) and cannot be glutamylated by a TTLL enzyme. We previously showed that microtubule hyperglutamylation causes B-tubule instability [23]. While TBA-6 itself may not be a direct substrate of the TTLL glutamylase enzymes, TTLL enzymes glutamylate tubulin in the context of the microtubule. Structural characterization of TTLL7 reveals that TTLL7 binds both α and β tails of tubulin dimers and that this binding is required for glutamylation of β-tubulin [41]. Hence α-tubulin TBA-6 may impact TTLL activity without itself being a substrate.

Finally, TBA-6 may function in CEM ciliary maturation. CEM neurons are embryonically-derived, yet tba-6 is not expressed until the L4 stage immediately preceding sexual maturation [26]. In this instance, tba-6 may be required for extending doublet microtubules, while other factors such as tubulin glutamylases, tubulin deglutamylases, or microtubule associated proteins may sculpt 18 singlets from nine doublet microtubules. None of these models are mutually exclusive.

tba-6 also regulates IFT. How ciliary motors walk along microtubule tracks is a fundamental and unresolved question in cell biology. In Chlamydomonas flagella, anterograde heterotrimeric kinesin-II walks on the B-tubule while retrograde dynein-2 moves on the A-tubule [42]. Metazoans such as C. elegans acquired an additional accessory IFT kinesin-2 motor, homodimeric KIF17/OSM-3, which depending on the specific cilium type, maybe functionally redundant with kinesin-II or seemingly not involved in IFT at all [16, 43–46]. How A- and B-tubule ultrastructure relates to microtubule motor coordination in cilia with multiple kinesins is not known. In C. elegans amphid and phasmid channel cilia that express two IFT kinesin-2 motors, kinesin-II transports IFT through the transition zone before handing its cargo over to OSM-3 in the proximal cilium [47]. Here, kinesin-II is restricted to the region of the cilium that contains microtubule doublets and excluded from the distal A-tubule singlet region [47, 48]. In the distal region OSM-3 alone moves the entire IFT-A/B complex [47], indicating that OSM-3 utilizes A-tubule singlets [49].

In CEM cilia, anterograde IFT is driven predominantly by kinesin-II with minimal modulation from accessory kinesins OSM-3 and KLP-6 [16]. Based on the above literature and our data, we propose that core IFT motor kinesin-II moves on B-tubules while accessory ciliary kinesins OSM-3 and KLP-6 are restricted to A-tubules (Figure 6). The loss of specialized ciliary ultrastructure in tba-6 mutants is accompanied by the loss of CEM-specific coordination of the IFT kinesins and IFT-A/B complex transport. In tba-6 mutants, the inherently faster OSM-3 motor is abnormally recruited into the IFT machinery, resulting in the formation of discrete populations of kinesin-II•IFT-A and OSM-3•IFT-B moving at different velocities. KLP-6 moves independently of the IFT machinery in WT and tba-6 mutant cilia, consistent with KLP-6 moving on A-tubules. To our knowledge, this is the first in vivo demonstration that a tubulin isotype directly affects IFT and ciliary microtubule-based transport.

α-tubulin isotype TBA-6 is required for the cilium’s ability to release bioactive EVs. CEM cilia shed and release EVs to the outside environment via the cuticle pore [17]. TBA-6 is required for EV environmental release, EV cargo selection, and EV bioactivity. In tba-6 mutants, environmental access of the ciliary tip inversely correlates with lumenal EV accumulation (Figure 4H). The abnormal inward curvature of the CEM cilia in tba-6 mutants precludes environmental access to the cuticle pore. This may result in lumenal accumulation of smaller sized 50–60 nm EVs. These data suggest that ciliary EVs may be shed from two sites – larger EVs that are shed at the ciliary base into the lumen and smaller EVs that are shed at the ciliary tip and released into the environment.

Our studies raise several new and interesting questions. What mechanisms drive axonemal remodeling? How do microtubule tracks and the tubulin code control ciliary motors and multiple aspects of EV biology? Our findings reveal the far-reaching effects of the tubulin code and how a specific tubulin isotype can be a defining regulator of the structural and functional identity of a cilium.

Supplementary Material

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Highlights.

  • The tubulin code specifies ciliary structure, motor transport, and function

  • α-tubulin isotype TBA-6 generates 18 singlet from nine doublet microtubules

  • tba-6 regulates IFT kinesin 2 motor coordination but not kinesin-3 KLP-6 velocity

  • tba-6 is required for extracellular vesicle cargo content, release, and bioactivity

Acknowledgments

We thank Leslie Gunther and Geoff Perumal for help in HPF-FS performed at Einstein, and William Rice and Ed Eng at the New York Structural Biology Center (NYSBC), for help in electron tomography; Daniella Nicastro, Antonina Roll-Mecak, Kristen Verhey for insights on the tubulin code; the Barr lab and Rutgers C. elegans community somewhere in the swamps of Jersey for discussion and constructive criticisms; Rutgers Human Genetics Institute and Genetics Department for critical bridge funding; WormBase and WormAtlas for online resources. Use of the NYSBC facilities was supported by the Albert Einstein College of Medicine. Some strains were provided by the National Bioresource Project and the Caenorhabditis Genetics Center (CGC), which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440). This work was funded by NIH DK059418 and DK074746 (to M.M.B.), NIH OD 010943 (to D.H.H.), NIH R01GM101972 and R01NS42023 (to C.R.), and Waksman Institute Charles and Johanna Busch Fellowship (to M.S).

Footnotes

Author contributions

Conceptualization: M.S and M.M.B.; Methodology, investigation, and analysis: M.S. N.M, K.Q.C.N, A.R., C.R., D.H.H., and M.M.B.; Writing: M.S, N.M., M.M.B.; Funding Acquisition M.S., C.R., D.H.H., and M.M.B.

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