# The Mn-motif protein MAP6d1 assembles ciliary doublet microtubules

Dharshini Gopal, Juliette Wu, Julie Delaroche, Christophe Bosc, Manon de Andrade, Eric Denarier, Gregory Effantin, Annie Andrieux, Sylvie Gory-Fauré, Laurence Serre, et al.

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Dharshini Gopal, Juliette Wu, Julie Delaroche, Christophe Bosc, Manon de Andrade, et al. The Mn-motif protein MAP6d1 assembles ciliary doublet microtubules. Nature Communications, 2025, 16 (1), pp.6210. 10.1038/s41467-025-61679-0. hal-05151424

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# The Mn-motif protein MAP6d1 assembles ciliary doublet microtubules

Received: 30 December 2024

Accepted: 27 June 2025

Published online: 05 July 2025

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Dharshini Gopal Ⓓ1,3, Juliette Wu Ⓓ1,3, Julie Delaroche<fnref n="1" />, Christophe Bosc Ⓓ1,
Manon De Andrade<fnref n="1" />, Eric Denarier Ⓓ1, Gregory Effantin Ⓓ2, Annie Andrieux<fnref n="1" />,
Sylvie Gory-Fauré Ⓓ1, Laurence Serre Ⓓ1,4 & Isabelle Arnal Ⓓ1,4

Most eukaryotic cells have cilia that serve vital functions in sensing, signalling, motility. The core architecture of cilia is an array of microtubule doublets, which consist of a complete A-tubule and an incomplete B-tubule. How these structures assemble remains poorly understood. Using total internal reflection fluorescence microscopy and cryo-electron tomography, we investigate the role of MAP6d1, a brain-specific protein containing microtubule lumen-targeting Mn-motifs. We show that MAP6d1 assembles stable microtubule doublets by recruiting tubulin dimers onto the A-tubule lattice to initiate B-tubule nucleation. MAP6d1 also promotes the formation of luminal proto-filaments in singlet and doublet microtubules, a previously undescribed phenomenon that likely enhances microtubule stability. In neurons, MAP6d1 localises to the proximal part of primary cilia via its Mn-motif, with its loss resulting in shortened cilia, a characteristic of ciliopathies. MAP6d1 is thus a neuronal Mn-motif protein with a specific role in assembling microtubule doublets and regulating ciliary length.

Microtubules are one of the major classes of cytoskeletal proteins that serve a wide range of cellular functions, providing tracks for cargo transport and structural support to maintain cellular architecture and enable motility<fnref n="1" />,². In order for cells to quickly reorganise their cytoskeleton in response to various triggers, dynamic microtubules alternate between phases of growth and shrinkage, known as dynamic instability³. In contrast, long-lived stable microtubules with reduced tubulin turnover predominate in terminally differentiated cells such as neurons, where they maintain cell morphology and support long-distance intracellular transport⁴. Particularly stable microtubules are also found in the axonemes of cilia and flagella, where they form complex arrays of doublets, ensuring the integrity and functions of these cellular appendages⁵. Motile cilia and flagella are responsible for moving cells (e.g. sperm) or a part of their environment (e.g. mucus along the trachea), while non-motile or primary cilia function as an antenna for sensory perception and

signal transduction on most vertebrate cells⁶,⁷. Defects in cilia assembly and structure can result in a number of diseases, collectively referred to as ciliopathies⁸,⁹.

Cilia share a core structure, the axoneme, composed of nine doublet microtubules arranged radially around two central singlet microtubules in motile cilia, or without a central pair in primary cilia. Each doublet microtubule typically comprises one complete 13 protofilament A-tubule and one incomplete 10 protofilament B-tubule. Studies of cilia and flagella in unicellular organisms such as *Chlamydomonas* and *Tetrahymena* revealed the presence of proteins located in the microtubule lumen known as Microtubule Inner Proteins (MIPs), which have since emerged as key contributors to the remarkable stability of axonemal doublet microtubules¹⁰⁻¹⁴. In recent years, more than 60 axonemal MIPs have been identified across species, and they exhibit a variety of luminal microtubule-binding modes¹⁵⁻¹⁷. To date, however, the function of most MIPs remains obscure. More generally,

<footnote n="1">¹Grenoble Institut Neurosciences (GIN), INSERM, U1216, CNRS, CEA, Université Grenoble Alpes, Grenoble, France. ²Institut de Biologie Structurale (IBS), CNRS, CEA, Université Grenoble Alpes, Grenoble, France. ³These authors contributed equally: Dharshini Gopal, Juliette Wu. ⁴These authors jointly supervised this work: Laurence Serre, Isabelle Arnal. ✉e-mail: sylvie.gory-faure@univ-grenoble-alpes.fr; laurence.serre@univ-grenoble-alpes.fr; isabelle.arnal@univ-grenoble-alpes.fr</footnote>
the mechanisms by which microtubule doublets are assembled and the role of MIPs in this process are still poorly understood.

MIPs are also present in neurons, where they have long been observed as intraluminal particles in both cytoplasmic and ciliary microtubules<sup>18-23</sup>. By analogy with axonalal MIPs, they are thought to contribute to neuronal microtubule stability, although their identity remains largely unknown. MAP6, a microtubule-stabilising factor linked to psychiatric disorders<sup>24,25</sup>, was the first neuronal MIP located in the microtubule lumen. Intraluminal MAP6 generates highly stable microtubules that grow in a helicoidal pattern<sup>26</sup>. MAP6 belongs to the SAXO (Stabiliser of AXOnemal microtubules) family of proteins that contain a small helical motif in their microtubule-binding domains known as the Mn-motif<sup>17,27-29</sup>. This structural element has recently been recognised as a universal microtubule luminal binding feature in axonal- nemal MIPs, highlighting its potential significance in stabilising both axonal and neuronal microtubules. Interestingly, only two SAXO family members, MAP6 and MAP6d1 (MAP6 domain containing 1, formerly known as SL21 for 21-kDa STOP-like protein), are currently known to be expressed in the brain.

Here we focus on MAP6d1, the sole SAXO family member known to be expressed exclusively in the postnatal brain<sup>30</sup>. MAP6d1 contains two Mn-motifs and can protect microtubules from drug- and cold- induced depolymerisation via its Mn-motif-containing microtubule- binding domains<sup>31</sup>. Beyond its cellular localisation to microtubules, mitochondria, Golgi and membranes following palmitoylation<sup>30</sup>, little is known about MAP6d1 functions. To investigate the effects of MAP6d1 on microtubules and related cellular roles, we used a combi- nation of total internal reflection fluorescence (TIRF) microscopy- based in vitro reconstitution assays, cryo-electron tomography (cryo- ET), and cultured hippocampal neurons. Our findings uncover a direct role for MAP6d1 in the assembly and stabilisation of doublet micro- tubules, shedding light on its involvement in maintaining the archi- tecture of neuronal primary cilia.

Results

MAP6d1 induces microtubule pausing

To gain insight into the mechanism underlying MAP6d1's ability to stabilise microtubules, we used TIRF microscopy and in vitro recon- stitution assays to study microtubule dynamics (Fig. 1a and Supple- mentary Movie 1). We found that MAP6d1 did not affect the frequency of catastrophes but did reduce both the growth and shrinkage rates while promoting rescues, at both plus and minus ends (Fig. 1b and Supplementary Fig. 1a). The net effect was the occurrence of pauses during which the microtubules neither grew nor shrank. Pauses became more frequent at MAP6d1 concentrations of 20 nM and lasted sig- nificantly longer at concentrations of 50 nM (Fig. 1c). The longest paus- ses, lasting up to 20 min, blocked microtubule dynamics, with a mean paused microtubule length of about 8 ± 4.2 µm (n = 45 microtubules).

To evaluate the stability of these MAP6d1-paused microtubules, we performed dilution-induced depolymerisation assays. Microtubules assembled with tubulin alone quickly depolymerised within seconds upon buffer perfusion (Fig. 1d, left), whereas those assembled in the presence of MAP6d1 resisted depolymerisation with three distinct behaviours (Fig. 1d, right). One population of microtubules (37.4%) remained in a paused state for the entire duration of the movie (30 min) following buffer perfusion; a second (29.3%) showed pauses interrupted by small catastrophes; and a third population (32.1%) continuously but slowly depolymerised after buffer perfusion (Fig. 1e). The shrinkage rate of these slowly depolymerising microtubules was 50-fold slower than that observed for microtubules assembled without MAP6d1 (Fig. 1f).

We next performed a two-step perfusion TIRF experiment, poly-
merising microtubules from red tubulin in the presence of MAP6d1,
then exchanging the solution with green-fluorescent tubulin before
recording. In this experiment, approximately 70% of the dynamic
green-fluorescent microtubules, polymerising from the extremities of

MAP6d1 co-assembled microtubules, depolymerised up to the jun-
cion with the MAP6d1-assembled microtubule lattice, which served as a
rescue point (Fig. 1g). Thus, microtubules assembled in the presence of
MAP6d1 resist buffer-induced depolymerisation and are stable enough
to stop the depolymerisation of a dynamic microtubule.

Overall, these results reveal that MAP6d1 is a strong microtubule
stabiliser.

MAP6d1 induces the assembly of doublet microtubules by
recruiting tubulin

To investigate how MAP6d1 influences microtubule architecture, we performed negative staining electron microscopy on microtubules polymerised in the presence or absence of MAP6d1 (Fig. 2a). To our surprise, MAP6d1 enabled the formation of doublet microtubules, which were not seen with microtubules polymerised with tubulin alone. Cryo-ET revealed distinct microtubule architectures, including doublet microtubules with varying B-tubules, as shown in tomograms in (Fig. 2b and Supplementary Fig. 2). The complete microtubules, i.e. the A-tubules of doublet microtubules in our dataset, were consistently composed of 14 protofilaments, whereas the z-slices of the example tomograms delineated eleven classes of B-tubule with different numbers of protofilaments. Most B-tubules had 7–11 protofilaments, with 10 protofilaments being the most abundant class (~40 % of the extracted particles). This heterogeneity in B-tubule architecture could represent different stages of doublet microtubule assembly. Notably, the B-tubule was highly flexible, as its extremity did not anchor back to the A-tubule but exhibited global curvature similar to that observed in vivo (13.15 nm versus 13.50 nm, respectively, Fig. 2c and see below). Using subtomogram averaging, we reconstructed a doublet microtubule from these data, showing an A-tubule with 14 protofilaments and a B-tubule with 10 protofilaments (Fig. 2d).

To better understand the underlying mechanism of doublet microtubule formation, we performed tubulin recruitment assays using TIRF microscopy (Fig. 2e). Red-fluorescent pre-polymerised microtubules were incubated with free green-fluorescent tubulin. In the absence of MAP6d1, there was no tubulin recruitment (Fig. 2e, left), but with MAP6d1 we observed recruitment of free tubulin onto 57% of the pre-polymerised microtubules (Fig. 2e, f). We also noticed elon- gation of this recruited tubulin along the pre-existing microtubules (Supplementary Movie 2). To investigate the impact of tubulin recruitment on microtubule structure, we next imaged these samples by negative staining electron microscopy. In the presence of MAP6d1, 40% of microtubules were doublets, but without MAP6d1, there appeared only singlet microtubules (Fig. 2e, f). This confirms that the ability of MAP6d1 to assemble doublet microtubules depends on its capacity to recruit tubulin dimers along the microtubule lattice.

The Mn-motif of MAP6d1 determines its microtubule-regulating activities

We analysed the domains previously identified as essential for prop-
erties of MAP6d1 (Supplementary Fig. 3). The mutant MAP6d1-ΔMn2,
which deletes the microtubule-binding domain encompassing the
second Mn-motif and its C-terminal flanking region, conserved in
MAP6<sup>30</sup>, abolished the microtubule-stabilising activity of MAP6d1
(Fig. 3a, b and Supplementary Fig. 1b). This deletion also eliminated the
ability of MAP6d1 to recruit tubulin dimers along the microtubule
lattice and to promote microtubule doublet assembly (Fig. 3c, d). To
assess whether these effects were directly linked to the Mn-motif, we
designed a mutant in which the seven residues of the second Mn-motif
were replaced by alanine (MAP6d1-Mn2-7A). This mutant was also
unable to stabilise microtubules, promote doublet assembly or recruit
tubulin onto the microtubule lattice (Fig. 3). We then assessed the role
of the first Mn-motif using a similar alanine substitution mutant
(MAP6d1-Mn1-7A). This mutant retained residual microtubule-
stabilising activity, as evidenced by low growth and shrinkage rates
<figure><img src="image_5.png" /><figcaption>**Fig. 1** | MAP6d1 induces microtubule pauses. a Representative kymographs of microtubules grown from seeds (S) and 12 µM tubulin with increasing MAP6d1 concentrations. Arrowheads indicate pauses. b Graphs showing the growth rate (n = 301, 427, 712, and 364 growth events for 0, 10, 20 and 50 nM MAP6d1, respectively), shrinkage rate (n = 168, 274, 334, and 264 shrinkage events), catastrophe and rescue frequencies (n = 36, 41, 54 and 45 frequencies) of microtubules assembled with or without MAP6d1. c Graphs showing the pause frequency (n = 41, 54, and 45 pause frequencies, for 10, 20, and 50 nM MAP6d1, respectively) and durations (n = 24, 121, and 245 pause events) in similar conditions. In the absence of MAP6d1, microtubules did not exhibit any pauses. d Representative kymographs of microtubules undergoing disassembly after tubulin dilution. Microtubules were grown from seeds (S) and 25 µM tubulin (control, left) or 12 µM tubulin and 50 nM MAP6d1 (right). After dilution, three populations were observed in the presence of MAP6d1: continuous pause after dilution, pauses interrupted by small catastrophes (arrowheads), and continuous slow depolymerisation. e Percentages of the three</figcaption></figure>

populations presented in (d). f Shrinkage rate of control and population 3 microtubules shown in (d) (n = 32 and 50 shrinkage events, respectively). g Kymograph showing rescue events of a microtubule grown with 12 µM GTP-tubulin (cyan) at the extremity of a microtubule assembled with 12 µM GTP-tubulin and 50 nM MAP6d1 (magenta). The experimental procedure is depicted above the kymograph. The kymograph shows the behaviour of a dynamic microtubule following GTP-tubulin perfusion. Arrows indicate rescues occurring at the junction (*) between the MAP6d1-assembled microtubule (magenta) and the newly growing dynamic microtubule (cyan). The graph represents the number of rescues occurring at or beyond the junction (n = 194 and 20, respectively). For graphs in (b), (c), (f), (g), bars represent mean ± SD from at least three independent experiments. Circles represent the mean of each individual experiment. p values are indicated (Kruskal-Wallis ANOVA followed by post hoc Dunn's multiple comparison test). Source data are provided as a Source Data file.

and a higher binding affinity for microtubules (Supplementary Fig. 4a–d); it did not, however, promote pause events, recruit tubulin to the microtubule lattice or assemble doublet microtubules (Supplementary Fig. 4e, f). Overall, these results indicate that both Mn motifs contribute to the observed microtubule-regulatory roles of MAP6d1, with the Mn2 motif playing a dominant role.

## The N-terminal domain of MAP6d1 is required for microtubule pausing and doublet formation

Next, we compared the microtubule-regulatory activity of MAP6d1 to that of a mutant lacking its N-terminal domain (MAP6d1-Δ2-35), which is conserved in MAP6 and involved in MAP6-mediated microtubule stabilisation and luminal particle formation²⁶,³⁰. Relative to the control
a

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b

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e

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**Fig. 2** | MAP6d1 induces the assembly of doublet microtubules by recruiting tubulin. **a** Negative staining electron microscopy images of microtubules assembled with 25 µM tubulin in absence or presence of 250 nM MAP6d1. White and blue arrowheads indicate singlet (SMT) and doublet (DMT) microtubules, respectively. The graph represents the percentage of SMT and DMT assembled without and with MAP6d1 (total measured lengths of 304 µm and 401 µm, respectively, from 3 experiments). **b** Examples of cryo-electron tomograms of doublet microtubules assembled with MAP6d1. The top panel shows longitudinal sections of tomograms along with a transverse section at the position of the blue dashed lines. A sub-tomogram average map is presented below each tomogram. **c** B-tubule traces of MAP6d1-induced doublet microtubules starting at the outer junction (blue square) and overlapping the curvature of the B-tubule of the *Chlamydomonas* flagellar doublet microtubules (EMD-40621) for comparison (n = 45 B-tubule curvatures). **d** Reconstruction of MAP6d1-induced doublet microtubules obtained by combining the maps of the doublet microtubule and the B-tubule. **e** Top: snapshots of TIRF microscopy recorded movies of 0.325 µM tubulin (cyan) mixed with GMPCPP-stabilised microtubule seeds (magenta) in the absence or presence of 100 nM MAP6d1 and 50 µM GMPCPP. White and blue arrowheads indicate tubulin recruitment and polymerisation at microtubule ends, respectively. Bottom: negative staining electron microscopy images of microtubules assembled in the same conditions as for TIRF microscopy. White and blue arrowheads indicate SMT and DMT, respectively. MT microtubule, Tub tubulin. **f** Left: percentage of microtubules that have or have not recruited tubulin (n = 57 and 282 microtubules from 3 and 14 independent experiments, respectively). Black dots represent the percentage of microtubules with recruitment for each experiment. **Right:** percentage of singlet and doublet microtubules in the control and with MAP6d1. The total measured lengths of microtubules were 225 µm and 543 µm for the control and with MAP6d1, respectively (three independent experiments). Black dots represent the percentage of doublet microtubules for each experiment. In (a, f), Fischer's exact contingency test (two-sided) was applied (p values are indicated). Source data are provided as a Source Data file.

c

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d

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f

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<figure><img src="image_14.png" /><figcaption>**Fig. 3** | The microtubule-regulating activities of MAP6d1 depend on its Mn-motif and N-terminus. **a** Kymographs depicting microtubules grown from seeds (S) and 12 µM tubulin alone or with 50 nM MAP6d1, 300 nM MAP6d1-ΔMn2, 300 nM MAP6d1-Mn2-7A or 50 nM MAP6d1-Δ2-35 (n = 105, 57, 81, 49 and 38 microtubules, respectively). Arrowheads indicate microtubule pauses.</figcaption></figure>

**b** Dynamical parameters of microtubules assembled without or with MAP6d1, MAP6d1-ΔMn2 MAP6d1-Mn2-7A and MAP6d1-Δ2-35: growth rate (n = 556, 437, 190, 397 and 286 growth events, respectively), shrinkage rate (n = 444, 284, 162, 348 and 197 shrinkage events, respectively), catastrophe and rescue frequencies (n = 105, 56, 38, 81 and 48 frequencies, respectively). Bars represent mean ± SD from at least three independent experiments. Circles represent the mean of each individual experiment. p values are indicated (Kruskal-Wallis ANOVA followed by post hoc Dunn's multiple comparison test). **c** Left: snapshots of TIRF movies of 0.325 µM tubulin (cyan) mixed with GMPCPP seeds (magenta) in the absence or presence of 100 nM MAP6d1, 300 nM MAP6d1-ΔMn2 300 nM MAP6d1-Mn2-7A and 100 nM MAP6d1-Δ2-35. White and blue arrowheads indicate tubulin

recruitment and polymerisation at the seed extremities, respectively. **Right:** Negative staining electron microscopy images of microtubules assembled in the same conditions as for TIRF microscopy. The blue arrowhead indicates a doublet microtubule (DMT). MT microtubule, Tub tubulin. **d Top:** percentage of microtubules with or without recruited tubulin (n = 57, 282, 63, 164 and 152 microtubules from 7, 17, 3, 6 and 3 independent experiments for the control and with MAP6d1, MAP6d1-ΔMn2, MAP6d1-Mn2-7A and MAP6d1-Δ2-35, respectively). Black dots represent the percentage of microtubules with recruitment for each experiment. **Bottom:** percentage of singlet (SMT) and doublet (DMT) microtubules in the control and with MAP6d1 and its mutants (total measured lengths of 361, 752, 149, 221 and 198 µm for MAP6d1, MAP6d1-ΔMn2, MAP6d1-Mn2-7A and MAP6d1-Δ2-35, respectively). Black dots represent the percentage of doublet microtubules for each experiment. Fischer's exact contingency test (two-sided) was applied. All statistical analyses were performed from at least three independent experiments and p values are indicated. Source data are provided as a Source Data file.

<figure><img src="image_15.png" /></figure>
condition, MAP6d1-Δ2-35 stabilised microtubules similarly to MAP6d1 by increasing the rescue frequency and reducing the shrinkage rate (Fig. 3a, b and Supplementary Fig. 1b) while also promoting the growth rate, leading to persistent microtubule growth. Unlike MAP6d1, however, MAP6d1-Δ2-35 did not induce microtubule pausing (Fig. 3a), and it suppressed doublet microtubule formation (only -1% vs. the -45% with MAP6d1; Fig. 3c, d). Consistent with these observations, this mutant weakly recruited free tubulin along stabilised microtubules (-10% versus -61% for MAP6d1) (Fig. 3c, d). Thus, although the N-terminal domain is not needed for stabilising microtubules, it is essential to induce pauses by decreasing the microtubule growth rate and to form doublets by recruiting tubulin.

## MAP6d1 localises to microtubule doublets in neuronal primary cilia and regulates ciliary length

Building on evidence of MAP6d1's ability to assemble microtubule doublets in vitro, we moved to studies in neurons, focusing on primary cilia. We compared primary cilia from cultured hippocampal neurons derived from Wild-Type (WT) and MAP6d1-KnockOut (KO) mice (Fig. 4a, b). The percentage of ciliated neurons was similar in both cases (Supplementary Fig. 5a). At 4 days in vitro (DIV), ciliary length (4.3 ± 1.861 µm) was also similar to that observed in WT (4.4 ± 1.746 µm) (Fig. 4a). Given that MAP6d1 expression begins around 7 DIV³⁰, we re-examined ciliary length at 9 DIV, and found that MAP6d1-KO cilia had become 15% shorter than those in WT neurons (5.38 ± 2.022 µm to 4.7 ± 1.695 µm) (Fig. 4b); this difference persisted through 23 DIV (Supplementary Fig. 5b).

To determine the localisation of MAP6d1 in neurons, we then expressed ectopic GFP-tagged MAP6d1 in the hippocampal cells. We observed that MAP6d1-GFP localised to the proximal part of the primary cilia in all transfected cells and led to cilia ~24% longer than those in control cells (Fig. 4c and Supplementary Fig. 5c). This asymmetric distribution contrasted with that of the ciliary marker Arl13b, which spans the entire length of the cilium. The proximal localisation of MAP6d1 corresponds to the microtubule doublet region identified by cryo-ET showing that doublet microtubules lose their B-tubule, eventually leaving only singlet A-tubules in the distal part of the cilia³²,³³. Expansion microscopy showed that MAP6d1-GFP co-localised with polyglutamylated tubulin—a post-translational modification (PTM) enriched on the B-tubules (and therefore doublet microtubules) in motile cilia³⁴⁻³⁶— in the central region of the cilia, positioned between the two distinct bands of the membrane-associated protein Adenylate Cyclase that delineate the borders of the cilia (Fig. 4d, e).

To further investigate MAP6d1's preferential proximal enrichment, we performed immunofluorescence co-staining with PTMs of tubulin known to accumulate in ciliary axonemes. We confirmed that MAP6d1-GFP co-localised with polyglutamylated tubulin but not with tyrosinated tubulin, a PTM associated with dynamic microtubules³⁷,³⁸ and enriched at the distal end of the cilia (Fig. 4f, g). This finding is consistent with previous observations that single dynamic microtubules are found predominantly at the ciliary tips³⁹,⁴⁰. To assess whether MAP6d1 influences the distribution of doublet or singlet MTs in cilia, we compared the distribution of tubulin PTMs associated with these microtubule architectures in wild-type and MAP6d1-KO neurons. Both polyglutamylated and tyrosinated tubulin exhibited similar patterns in the two types of neurons, despite MAP6d1-KO neurons bearing shorter cilia (Supplementary Fig. 5d).

To determine whether MAP6d1's Mn-motif or N-terminal influence ciliary localisation, we expressed GPF-tagged-ΔMn2 and Δ2-35 mutants in hippocampal neurons (Fig. 4c and Supplementary Fig. 5c). Unlike MAP6d1, MAP6d1-ΔMn2-GFP failed to localise to the cilia, which were similar in length to those in wild-type cells. In contrast, MAP6d1-Δ2-35-GFP was present all along the cilia, which were twice as long as those in control neurons. The distribution of tubulin PTMs remained

similar to that observed in MAP6d1-expressing neurons (Supplementary Fig. 5e).

MAP6d1's ciliary localisation therefore requires the Mn2 motif, and its targeting of the proximal cilia, associated with doublet microtubules, is linked to the N-terminal domain. Moreover, these results suggest that MAP6d1 plays a role in regulating ciliary length in neurons, likely by assembling and stabilising microtubule doublet architecture.

## MAP6d1 assembles protofilaments in the lumen of singlet and doublet microtubules

Tomograms of microtubules assembled in the presence of MAP6d1 revealed peculiar structures within the lumen of both doublet and singlet microtubules (Fig. 5a). These structures correspond to two adjacent tubulin protofilaments that would interact with the microtubule internal surface. Indeed, subtomogram averages applied to these MAP6d1-induced structural entities allowed to obtain reconstructions of doublet and singlet microtubules with two luminal protofilaments (Fig. 5b, c). These reconstructions indicate that the luminal protofilaments in the A-tubule of the doublet face protofilaments A10-A13, adjacent to the putative seam (Fig. 5b) which we positioned between protofilaments A9 and A10 in the doublet microtubule, based on previous structural studies¹¹,⁴¹. We did not detect any obvious lattice defects in proximity to these luminal protofilaments (in a total of 220 microtubules), suggesting that luminal protofilaments are formed during microtubule copolymerisation with MAP6d1 rather than through diffusion via breaks in the lattice.

We next compared the B-tubule curvatures of doublet microtubules with and without luminal protofilaments (Fig. 5d) and found that the mean radii of the B-tubules with luminal protofilaments increased by ~15% compared to those without (Fig. 5d). As luminal protofilaments are localised in the A-tubule right next to where the B-tubule begins, known as the outer junction, they may influence the conformation of the B-tubule.

Because protofilaments within the microtubule lumen have not been previously described, we used cryo-electron tomography to look for the presence of such microtubule architectures in mature neurons (Fig. 5e and Supplementary Fig. 6). We observed a few rare examples of singlet microtubule structures containing filamentous densities that resemble the luminal protofilaments induced in vitro by MAP6d1, in neuritic extensions (Fig. 5f and Supplementary Fig. 6). We propose that these luminal protofilaments could help explain the strong stability of neuronal microtubules.

## Discussion

These data reveal that MAP6d1 regulates microtubule architecture within neuronal primary cilia. As MAP6d1 is expressed in the brain postnatally, it likely contributes to ciliary elongation and maturation, a process that lasts for several weeks after birth¹². More specifically, MAP6d1 induces the assembly of doublet microtubules and is able to independently reconstitute these ciliary structures. We also observed a previously undescribed phenomenon of protofilaments within the lumen that we suspect lend extra strength to neuronal microtubules.

MAP6d1 forms doublet microtubules through tubulin recruitment on the A-tubule lattice, facilitating the nucleation of the B-tubule. Our data support a model in which MAP6d1 behaves both as a MAP, in that it binds to the surface of the microtubule lattice, and as a MIP, in that it contains two universal lumen-targeting Mn-motifs. These features, respectively, are necessary for its ability to recruit tubulin to the MT lattice and form intraluminal filaments. We propose that this 'dual binding' mode is key to MAP6d1's unique ability to assemble doublet microtubules by bridging the A-tubule to the B-tubule (Fig. 6). Given that the C-terminal tail of tubulin plays a crucial role in doublet microtubule assembly⁴³,⁴⁴, it may be that MAP6d1 binding to the A-tubule triggers a conformational change in the tubulin C-terminal
<figure><img src="image_16.png" /><figcaption>**Fig. 4** | MAP6d1 preferentially localises to microtubule doublets in neuronal primary cilia and regulates ciliary length. **a** Representative images of 4 days in vitro (DIV) wild-type (WT) and MAP6d1-knockout (KO) neurons stained against cilia marker Arl13b and nuclear dye Hoechst. The graph represents cilia length of WT (n = 253) and KO neurons (n = 262). **b** Same as (a) for 9 DIV neurons (WT: n = 407, KO: n = 398). In (a, b), bars represent mean ± SD from three independent experiments. Circles represent individual experiment means. *p* values are indicated (Mann-Whitney's test). **c** Localisation of ectopic GFP, MAP6d1-GFP, MAP6d1-ΔMn2-GFP and MAP6d1-Δ2-35-GFP compared to ciliary marker Arl13b, in 7 DIV neurons. The graph represents cilia length in neurons expressing GFP (n = 128 cilia), MAP6d1-GFP (n = 68 cilia), MAP6d1-ΔMn2-GFP (n = 59 cilia), MAP6d1-Δ2-35-GFP (n = 86 cilia) from at least three independent experiments. The graph presents individual data points with bars representing mean ± SD. Circles represent individual experiment means. *p* values are indicated (One-way ANOVA and pairwise comparisons). **d, e** 4X Expansion microscopy</figcaption></figure>

images of MAP6d1's ciliary localisation with PolyE tubulin (d) or the cilia membrane marker Adenylate cyclase (e). Plot profiles drawn perpendicular to the cilia within the white rectangles. Images are representative of 20 cilia from two independent experiments. f Representative images of primary cilia from 7 DIV neurons expressing ectopic MAP6d1-GFP and stained for acetylated (Ac tub) and polyglutamylated tubulin (PolyE tub). The graph represents the percentage of normalised maximum fluorescence of MAP6d1-GFP and polyglutamylated tubulin (PolyE tub) along the ciliary length determined by acetylated tubulin (Ac tub). Values represent mean ± SEM of n = 58 cilia from 3 independent experiments. g Representative images of primary cilia from 7 DIV neurons expressing ectopic MAP6d1-GFP stained for Arl13b and tyrosinated tubulin (Tyr tub). The graph represents the percentage of normalised maximum fluorescence of MAP6d1-GFP and tyrosinated tubulin along the ciliary length determined by Arl13b. Bars represent mean ± SEM of n = 46 cilia from 3 experiments. Source data are provided as a Source Data file.

tail, promoting tubulin recruitment and subsequent B-tubule nucleation. Although removal of the C-terminal tail of tubulin allows B-tubule nucleation from multiple sites on the A-tubule⁴³, MAP6d1 binding is sufficient to restrict B-tubule nucleation to a single A-tubule protofilament. MAP6d1 therefore appears to provide positional information

to recruit tubulin at a specific site on the A-tubule lattice, likely related to its binding near the seam region. Since deletion of the N-terminal part of MAP6d1 prevents both doublet formation and tubulin recruitment, we suspect the N-terminal participates in this local C-terminal tubulin conformational change.
a

<figure><img src="image_17.png" /></figure>

b

<figure><img src="image_18.png" /></figure>

e

<figure><img src="image_19.png" /></figure>

c

<figure><img src="image_20.png" /></figure>

d

<figure><img src="image_21.png" /></figure>

f

<figure><img src="image_22.png" /><figcaption>**Fig. 5** | MAP6d1 assembles protofilaments in the microtubule lumen.</figcaption></figure>

a Examples of tomograms of doublet (left) and singlet (right) microtubules containing luminal protofilaments. b Subtomogram averaged model of doublet microtubules containing luminal protofilaments (pf). The putative location of the microtubule seam is indicated by a dashed line. c Subtomogram averaged model of singlet microtubules containing luminal protofilament (left) and its superposition with the focused map of the luminal protofilaments (yellow, right). d B-tubule curvatures traced for doublet microtubules without (top) and with (bottom) luminal protofilaments. The graph shows the mean radii of the B-tubule curvatures (n = 36

and 27 for doublet microtubules without and with luminal protofilaments, respectively). Bars represent mean ± SD. Mann-Whitney test was applied (p values are indicated). e Example of a tomogram showing neuronal microtubules containing luminal protofilaments (blue arrowheads). f Comparison of microtubules containing luminal protofilaments assembled with MAP6d1 in vitro (left) with those observed in neurons (right) and their corresponding schemes. Pink and blue arrowheads show luminal protofilaments in microtubules assembled in vitro and in neurons, respectively. Source data are provided as a Source Data file.

MAP6d1 has one particular feature that may help organise both singlet and doublet microtubules in neurons: it is one of the few proteins, along with the neuronal kinesin-4 (KIF21B) and the ciliary MIP CSPP1, that can induce microtubule pausing⁴⁵,⁴⁶. It is worth noting in this context that deletion of CSPP1 in zebrafish reduces the length of primary cilia⁴⁷, similar to what we observed in MAP6d1-KO neurons. On the other hand, the MAP6d1-Δ2-35 mutant failed to induce pausing in vitro and doubled the length of primary cilia when overexpressed in neurons. It thus seems that microtubule pauses are important for maintaining

optimal ciliary length, which varies considerably in response to physiological and pathological changes⁴⁸. Understanding the mechanism by which MAP6d1 induces pauses will require further study, but it could result from a combination of a strong lattice-stabilising effect that prevents tubulin release, as indicated by MAP6d1's inhibition of microtubule shrinkage, and a modified tip conformation that hinders tubulin incorporation, as proposed for KIF21B⁴⁶.

MAP6d1 may also contribute to microtubule stability in a unique way by assembling protofilaments within the microtubule lumen. In
<figure><img src="image_23.png" /><figcaption>**Fig. 6 | Models of MAP6d1-microtubule interactions leading to the formation of the B-tubule and luminal protofilaments.** a MAP6d1 binds to the A-tubule lattice, recruits free tubulin dimers to initiate the B-tubule and forms a doublet</figcaption></figure>

microtubule. b During co-polymerisation with microtubules, MAP6d1 assembles protofilaments in the lumen.

doublet microtubules, luminal protofilaments in the A-tubule always localise near the outer junction, reinforcing the positional specificity of MAP6d1 binding at the B-tubule nucleation site, as discussed above. Such luminal protofilaments could constrain the architecture of the B-tubule or stabilise the doublet outer junction, similar to the organisation of MIPs in axonemal doublets¹⁰,¹⁷,⁴⁹.

MAP6d1 is part of the SAXO family, which is defined by the universal microtubule lumen-targeting Mn-motif. Most proteins in this family are found within the doublet microtubules of cilia and flagella, where they are thought to stabilise these structures¹¹,¹⁶,¹⁷,³¹,⁴⁹. Although structural studies have mapped the distribution of these proteins in doublets, the mechanisms by which they contribute to microtubule stabilisation and/or doublet formation remain largely unexplored. MAP6d1 stands out as a Mn-motif protein with a specific role in primary cilia and more generally, in the assembly of microtubule doublets. Together, these findings mark an important step toward uncovering the molecular mechanisms driving microtubule doublet formation and understanding the specialised functions of Mn-motif proteins across different cell types.

## Methods

### Ethical statement

All experiments involving animals were approved by the local animal welfare committee (Comité Local CEEA n°44 – APAFIS number 45896-2023111317017897) and in compliance with the European Community Council Directive (directive 2010/63/EU).

### Plasmid constructs

cDNAs encoding mouse MAP6d1 (full length, aa 1-191), MAP6d1-Δ2-35 (deletion of residues 2-35), MAP6d1-ΔMn2 (deletion of residues 123 to 145) and MAP6d1-Mn2-7A (amino acids 123 to 128 replaced by alanine) were amplified by PCR using Phusion High-Fidelity DNA polymerase (Thermo Scientific) and cloned into pET42a vector with In-Fusion HD cloning kit (Clontech). Primer sequencies are listed in Supplementary Table 1. All encoded proteins exhibited an 8-histidine tag at their C-terminal end. For eukaryotic expression, we used

plasmids of mouse MAP6d1-GFP, MAP6d1-Δ2-35-GFP and MAP6d1-ΔMn2-GFP³⁰.

### Tubulin preparation

Tubulin was purified from bovine brain and coupled with biotin, ATTO-565 or ATTO-496 as previously described⁵⁰,⁵¹. Briefly, brain tissue was homogenised in cold AB buffer (0.1 M Pipes, 0.5 mM MgCl₂, 2 mM EGTA, 0.1 mM EDTA, pH 6.8; 1 mL per gram of tissue) and centrifuged at 100,000 × g for 90 min at 4 °C. Tubulin in the supernatant was polymerised by adding 33% (v/v) glycerol, 0.2 mM GTP, 1.5 mM ATP, and 3 mM MgCl₂, followed by incubation at 37 °C for 60 min. The mixture was then centrifuged at 100,000 × g for 90 min at 35 °C. Pellets were resuspended in AB buffer and incubated for 30 min at 4 °C to induce microtubule depolymerisation, then centrifuged at 140,000 × g for 30 min at 4 °C. The supernatant was supplemented with 33% glycerol, 1 mM GTP, 0.5 mM ATP, and 3 mM MgCl₂, and subjected to a second polymerisation-depolymerisation cycle. Microtubule-associated proteins (MAPs) in the solution were separated from tubulin by cation-exchange chromatography using Fractogel EMD SO₃ resin (VWR) equilibrated in BRB80 buffer (80 mM Pipes, 1 mM EGTA, 1 mM MgCl₂, 1 mM DTT, pH 6.8) supplemented with 0.1 mM GTP. Fractions containing tubulin were pooled, supplemented with 33% glycerol and 1 mM GTP, and incubated for 1 h at 37 °C, before been poured onto a BRB80/60% glycerol cushion and centrifuged at 100,000 × g at 35 °C for 90 min. Pelleted microtubules were resuspended in cold BRB80 buffer and incubated for 30 min at 4 °C to induce complete depolymerisation. Tubulin aliquots (5-10 µL) were flash-frozen and stored in liquid nitrogen.

For tubulin labelling, sulfo-NHS-LC-biotin (Thermo Scientific) or NHS-ester-ATTO-565/-ATTO-488 (ATTO-TEC Gmbh, Germany) was dissolved in anhydrous DMSO at 100 mM and mixed (5% v/v) with microtubules polymerized from purified tubulin, in a 100 mM Na-Hepes buffer (pH 8.6) containing 40% glycerol, at 35 °C for 10 min. The labelling reaction was stopped by adding two volumes of freshly prepared 100 mM potassium glutamate solution. Labelled microtubules were layered onto a BRB80/60% glycerol cushion and
centrifuged at 100,000 × g for 30 min at 35 °C. Pellets were resus-
pended in cold BRB80, centrifuged at 100,000 × g for 15 min at 4 °C,
and subjected to an additional polymerisation–depolymerisation
cycle before aliquoting, flash-freezing, and storing in liquid nitrogen.

Recombinant protein expression and purification

MAP6d1 and its mutants were expressed in BL21 (DE3) Star *Escherichia coli* cells. After induction by 0.1 mM IPTG overnight at 18 °C, cells were sonicated in a lysis buffer (50 mM Hepes pH 7.4, 200 mM KCl, 0.1% Triton, and complete EDTA-free protease inhibitor cocktail tablets [GE, Healthcare]) before centrifugation at 160,000 × g for 30 min at 4 °C. The clarified lysate was incubated with a cobalt affinity resin (Talon, Clontech) and the protein was eluted by increasing concentrations of imidazole. Collected fractions were concentrated, dialysed against 50 mM Hepes pH 7.4, 200 mM KCl, 20 mM DTT, centrifuged at 100,000 × g for 5 min at 4 °C before freezing in liquid nitrogen. Protein concentrations were determined by comparing band intensities on SDS-PAGE gels to a standard curve generated by loading a range of known BSA concentrations on the same gels. The intensities of the bands were quantified with ImageJ software<sup>32</sup> (version 2.160/1.54p). Purified proteins used in this study are shown in Supplementary Figs. 3 and 4.

Preparation of microtubule seeds and Taxol-stabilised microtubules

GMPCPP-microtubule seeds (50% biotinylated, 50% ATTO-565 labelled tubulin) were prepared at a final concentration of 10 µM tubulin by incubating 5 µM biotin-labelled tubulin and 5 µM ATTO-565 labelled tubulin in BRB80 buffer supplemented with 1 mM GMPCPP for 1 h at 37 °C<sup>51</sup>. Microtubules were centrifuged at 85,000 × g for 5 min at 37 °C, resuspended in the same volume of BRB80 buffer supplemented with 1 mM GMPCPP, aliquoted, and stored in liquid nitrogen.

*Taxol-stabilised microtubules* were polymerised by incubating
70 µM tubulin (50% biotinylated tubulin and 50% ATTO-565 labelled
tubulin) in BRB80 buffer with 1 mM GTP for 45 min at 37 °C. 50 µM
Taxol was then added and microtubules were incubated for 30 min
before being centrifuged at 85,000 × g for 5 min at 37 °C and resus-
pended in BRB80 buffer supplemented with 10 µM Taxol.

Co-sedimentation assay

To assess the microtubule-binding properties of MAP6d1 and its
mutants, 1 µM Taxol-stabilised microtubules were incubated for
25 min at 35 °C with 300 nM MAP6d1, MAP6d1-Δ2-35, MAP6d1-
ΔMn2, MAP6d1-Mn2-7A or MAP6d1-Mn1-7A in BRB80 buffer sup-
plemented with 125 mM KCl. The mixtures were then centrifuged at
190,000 × g (Beckman Coulter rotor TLA-100) for 15 min at 35 °C.
Protein contents of the supernatants and pellets were analysed by
Western blotting using mouse monoclonal anti-tubulin (1:2000,
clone alpha3A1)⁵³ and Penta His HRP-conjugated (1:10000, Qiagen,
34460) antibodies (Supplementary Figs. 3 and 4). The intensity of
the bands corresponding to MAP6d1 and mutants was quantified
using ImageJ.

Microtubule dynamics using TIRF microscopy

Chambers made of glass slides and coverslips, functionalised with
polyethylene glycol (PEG)-silane (30 Kda, Creative PEGWorks) and
PEG-silane-biotin (3.4 Kda, Laysan BIO), respectively, were prepared
as previously described⁵¹. Briefly, glass slides and coverslips were
plasma-cleaned and immediately incubated overnight in the dark
under gentle agitation with solutions of mPEG-silane (1 mg/mL in 96%
ethanol, 0.2% HCl) and biotin-PEG-silane (1 mg/mL in 96% ethanol,
0.2% HCl), respectively. After incubation, slides and coverslips were
thoroughly rinsed with ethanol and deionized water, dried, and
assembled into flow chambers. The glass chamber was perfused with

neutravidin (25 µg/ml) (Pierce) in 1% BSA in BRB80, followed by
Poly(L-lysine) (PLL)-g-PEG (0.1 mg/ml) (JenKem) in 10 mM Hepes, pH
7.4 and flushed with 1% BSA in BRB80. GMPCPP-microtubule seeds
were then perfused and incubated for 5 min before being washed
twice with 1% BSA in BRB80 buffer. Microtubules were polymerised
from seeds by adding 12 µM tubulin (15% ATTO-565 labelled tubulin)
with different concentrations of MAP6d1 or its mutants in TIRF buffer
(BRB80 containing 1 mM GTP, 25 mM KCl, 3 mM dithiothreitol [DTT],
1% BSA, glucose [1 mg/ml], catalase [70 µg/ml], glucose oxidase
[600 µg/ml] and 0.1% methylcellulose [4000 cP]). For experiments
comparing MAP6d1 and mutants (Fig. 3a, b), the concentrations of
MAP6d1-ΔMn2 and MAP6d1-Mn2-7A were increased to 300 nM
because of their lower microtubule-binding activity relative to
MAP6d1 (Supplementary Fig. 3b, c). Flow chambers were sealed, and
movies were acquired on an inverted microscope (Eclipse TI, Nikon)
equipped with an iLas TIRF system (Roper Scientific), a CMOS camera
(Prime 95B, Photometrics) and a temperature-controlled stage
(LINKAM MC60), under MetaMorph software (version 7.7, Molecular
Devices). Samples were excited using a 561-nm laser and observed
with an Apochromat 100x oil-immersion objective (N.A., 1.49). Time-
lapse acquisitions were carried out for 30 min at 35 °C with a 100 ms
exposure and one frame every 2 s.

To assess the effect of dilution of microtubule stability, micro-
tubules were polymerised in glass chambers from ATTO-565-GMPCPP
seeds in the presence of 12 µM tubulin (15% ATTO-565 labelled tubulin)
and 50 nM MAP6d1 in TIRF buffer containing 1 mM GTP. For control
conditions, microtubules were polymerised from 25 µM tubulin in
TIRF buffer with 1 mM GTP. Unsealed chambers were then incubated
for 20 min at 35 °C in a humid environment. Chambers were then
placed on the warm stage of the TIRF microscope and microtubule
depolymerisation was induced by flowing warm TIRF buffer. Micro-
tubule behaviour was immediately recorded using 561-nm laser and
images were acquired every 2 s with a 100 ms exposure for
30 min at 35 °C.

To evaluate the dynamic behaviour of microtubules polymerised
at the extremity of MAP6d1-assembled-microtubules, microtubules
were grown from ATTO-565-GMPCPP seeds in the presence of 12 µM
tubulin (15 % ATTO-565 labelled tubulin) and 50 nM MAP6d1 in TIRF
buffer supplemented with 1 mM GTP. The unsealed chambers were
incubated for 20 min at 35 °C in a humid environment, before perfu-
sion of a solution containing 12 µM tubulin with 1 mM GTP in TIRF
buffer. Images were then acquired under the microscope every 2 s with
100 ms exposure for 30 min at 35 °C.

Tubulin recruitment assay

Mixtures containing 0.325 µM tubulin (15 % ATTO-491 labelled tubulin) and either MAP6d1 (100 nM), MAP6d1-Δ2-35 (100 nM), MAP6d1-ΔMn2 (300 nM), MAP6d1-Mn2-7A (300 nM) or MAP6d1-Mn1-7A (300 nM) in TIRF buffer with 50 µM GMPCPP and 50 mM KCl were incubated with ATTO-565-GMPCPP seeds in chambers. Samples were excited using 491- and 561-nm lasers, and time-lapse images were acquired on the TIRF microscope every 2 s with 100 ms exposure for 30 min at 35 °C.

TIRF analysis

Movies were analysed using ImageJ software to generate kymographs
of individual microtubules. The four parameters to assess microtubule
dynamics were extracted from kymographs using an in-house plugin³¹.
Growth and shrinkage rates were calculated from the slopes of the
corresponding growth and shrinkage events, respectively. Cata-
strophe and rescue frequencies were calculated for each microtubule
by dividing the number of events by their corresponding growth or
shrinkage event durations. Microtubule pauses were defined by
growth/shrinkage rates less than 0.24 µm/min at the plus end and
0.12 µm/min at the minus end.
## Negative staining electron microscopy

We used two protocols to evaluate microtubule doublet formation. In one, we co-polymerised 25 µM purified brain tubulin with 250 nM MAP6d1 in BRB80 buffer supplemented with 1 mM GTP and 125 nM KCl for 10 min at 35 °C (Fig. 2a). In the second approach, we incubated 10 µM stabilised GMPCPP-microtubule seeds with purified brain tubulin (0.325 µM) and either MAP6d1 (100 nM), MAP6d1-ΔA2-35 (100 nM), MAP6d1-ΔMn2 300 nM), MAP6d1-Mn2-7A (300 nM) or MAP6d1-Mn1-7A (300 nM) in BRB80 buffer supplemented with 50 µM GMPCPP and 50 mM KCl for 10 min at 35 °C (Fig. 2e and Supplementary Fig. 4e). We then loaded 4 µL of each sample on a 400-mesh copper carbon grid (Electron Microscopy Sciences), washed it briefly with warm BRB80 buffer and stained it with 2.5% uranyl acetate. Images were acquired using a transmission electron microscope (TEM) JEOL 1200 EX (Veleta). The length of microtubule doublets and singlets was measured on images using ImageJ software.

## Cryo-electron tomography

**In vitro sample preparation.** 25 µM tubulin was co-polymerised with 250 nM MAP6d1 supplemented with 1 mM GTP and 125 mM KCl in BRB80 buffer for 5–10 min at 35 °C. Four microliters of the solution was mixed to 1 µL of cationic BSA-coated gold beads (Aurion Gold Tracers, 210111) and then loaded on a glow-discharged Quantifoil R2/2 300 mesh copper grid (Electron Microscopy Sciences) and placed in the Leica EM-GP2 95 humidity chamber. The grid was blotted for 2 s and plunge-frozen in liquid ethane.

**Neuronal sample preparation.** Glow-discharged Quantifoil R2/2 200 mesh gold grids (Electron Microscopy Sciences) were placed in poly-L-lysine-coated sterile 35-mm glass-bottom dishes (Ibidi, FluoroDish) and hippocampal neurons grown on the above-mentioned grids for 25 days, following the protocol described for neuronal culture. The grid was blotted for 5 s using the vitrification robot Leica EM-GP2 and plunge-frozen in liquid ethane.

**Image acquisition.** The tilt series were acquired using K3 Titan Krios at the ESRF facility⁵⁴ operated at 300 keV, at a magnification of ×33,000 between −60° and +60° with a 3° increment under an electron dose of 2.87 e/Å² at a defocus of −3 to −6 µm, using Tomography 5 software (version 5.16, Thermo Fisher Scientific). The acquisition was done with a super-resolution pixel size of 1.35 Å/px and the data processing was carried out in binning 2 with a pixel size of 2.7 Å/px (see Supplementary Table 2).

**Image processing.** The 252 and 118 tilt series composed of 41 images were acquired for the in vitro samples and neurons, respectively. Raw tilt images were motion-corrected using MotionCor2 (version 1.4.0)⁵⁵ and then aligned for reconstructing the 3D tomograms by automatic alignment followed by contrast transfer function (CTF) estimation using the tomography pipeline in Eman2 software (version 2.99)⁵⁶ (Supplementary Fig. 7 and Supplementary Table 2).

For in vitro samples, we manually selected different microtubule architectures (singlet and doublet microtubules, with or without luminal protofilaments, see Supplementary Fig. 2a, b) using the filament tracing tool and particles were extracted with periodicities of 8 nm. The number of protofilaments in singlet microtubules and the A-tubule of doublet microtubules was determined on extracted particles by performing 3D classification in Eman2, using microtubules with 11–16 protofilaments as references. For the control, protofilament analysis was performed based on the microtubule fringe pattern on 2D cryo-EM images⁵⁷. In the presence of MAP6d1, singlet microtubules and the A-tubule of doublet MTs consistently contained 14 protofilaments (Supplementary Fig. 2c).

For doublet microtubule reconstruction, 4997 particles were selected. First, we built reference models of doublet microtubules consisting of an A-tubule with 14 protofilaments and a B-tubule with variable numbers of protofilaments (Fig. 2b). We then classified the particles based on the number of protofilaments in the B-tubule, using Eman2's multi-reference refinement. The distribution of particles with 7, 8, 9, 10, or 11 protofilaments in the B-tubule made up 21.27, 11.28, 9.85, 39.92 and 17.67%, respectively (Supplementary Fig. 2c). Subtomogram averaging refinement was performed on all extracted particles. Since the largest class (39.92%) corresponded to doublet microtubules with a 14-profilament A-tubule and a 10-profilament B-tubule, we used this configuration as the reference for the 3D-refinement step applying Eman2's 3D refinement pipeline that includes iterative steps of 3D particle alignment, sub-tilt translational and rotational refinements followed by defocus variation and using a mask including both A- and B-tubule. We then applied a local mask to the B-tubule to refine it independently of the A-tubule. The map displayed is the combination of both maps (Fig. 2d and Supplementary Fig. 2d).

We selected 5881 and 1680 particles, respectively, to reconstruct singlet and doublet microtubules with luminal protofilaments. We performed subtomogram averaging using Eman2's 3D refinement pipeline as described above. We verified that all singlet microtubules contained two luminal protofilaments through 3D classification using 14-profilament microtubules with either one or two luminal protofilaments as references. To improve the reconstruction of the luminal protofilaments in singlet microtubules, we applied a local mask to the two luminal protofilaments to refine them independently of the microtubule lattice (Fig. 5c).

To visualise luminal protofilaments on tomograms of mature neurons, we searched for structures similar to those identified in vitro, i.e., filamentous densities close to the microtubule wall and/or displaying curved extremities. Note that the observation of these events is limited by the low signal-to-noise ratio of tomograms due to the thickness of the sample and the presence of numerous MIPs in the microtubule lumen, which may obscure elongated luminal structures.

The IMOD package (version 4.11.25) was used for displaying tomograms³⁸. Electron density maps were visualised with Chimera (version 1.19) and ChimeraX (version 1.2.5)⁵⁹. FSC plots were extracted from Eman2 refinement output (Supplementary Fig. 2e).

## B-tubule curvature analysis

To measure the curvature radii of B-tubules (related to Figs. 2 and 5), we selected a set of four doublet microtubules without luminal protofilaments and three doublet microtubules with luminal protofilaments. To ensure that curvature measurements were not influenced by neighbouring structures, we chose only doublet microtubules far from the carbon support film and well isolated from each other. Curvature was measured at nine different z-sections along each doublet. For each z-section, about ten coordinates were selected to trace the B-tubule curve using the segmented line tool in Fiji software⁶⁰ (version 2.160/1.54p). To superimpose the resulting B-tubule traces, we defined two reference points: the centre of the A-tubule (O) and the outer junction where the B-tubule attaches (X). Each set of the z-section coordinates was then rotated and translated such that all O-X segments were aligned and all X points were centred at the same position. Mean curvature radii were calculated along each B-tubule trace using the curvature radius.bsh script, originally written by Olivier Burri (BIOP EPFL) and modified for our study (https://github.com/EricDenarier/MAPd1_MT_Doublets_2025), and the results were plotted on graphs.

The mean curvature radius of the B-tubule in in vivo doublet microtubules was determined using Fiji by drawing two circles on a z-stack of the *Chlamydomonas* flagellar doublet microtubule (EMD-40621)¹⁷, one corresponding to the external surface of the tubulin, the
other to its internal surface. The final radius corresponds to the average value calculated from the radii of these two circles.

### Animals used in this study

The *Map6d1* conditional KO mutant mouse line was established at the Institut Clinique de la Souris (ICS) - PHENOMIN. The targeting vector was constructed as follows: A 3 kb 3' homology arm fragment was amplified by PCR (from 129svpass genomic DNA) and subcloned in an ICS proprietary vector. This ICS vector bore a floxed and flipped Neomycin resistance cassette and a 5' LoxP site. A 1.9 kb fragment encompassing 628 bps of proximal promoter and *Map6d1* exon 1 (ENSMUSE00000268483) was cloned in a second step. Finally, a 4.2 kb fragment corresponding to the 5' homology arms was amplified by PCR and subcloned in step 2 plasmid to generate the final targeting construct. The linearised construct was electroporated in 129svpass (ICS derived P1 line) mouse embryonic stem (ES) cells. After G418 selection, targeted clones were identified by 5' and 3' Long 6range PCRs and further confirmed by Southern blot with a Neo probe (5' and 3' digests). Three positive ES clones were injected into C57BL/6J blastocysts. Chimeras were bred with Flp deleter females (in a C57BL/6J genetic background) in order to obtain the cKO allele with no selection cassette. Germline transmission was achieved.

To obtain the plain KO allele, chimeras were bred with Cre deleter females (also in a C57BL/6J genetic background) (Supplementary Fig. 8). Genotyping was performed by PCR on genomic DNA isolated from mouse tails using following oligonucleotides: 5'-GG GGCTTATGCCTGTGGCTATATG-3' plus 5'-CAGTCCTCCTAGGTGCAGA CTG-3' to detect the mutant allele and 5'-CATGGATTCCAGGGATCCA TCTCTC-3' to detect the wild-type allele. All experiment were performed on a C57BL6J genetic background. Mice were house in a temperature controlled (22 ± 1°C) under 12:12 light/dark cycle (light from 7 A.M. to 7 P.M.) with *ad libitum* access to food and water and humidity between 45 and 65%.

### Cultured hippocampal neurons

Hippocampal neurons were isolated from the brains of wild-type and *MAP6d1*-knockout (*MAP6d1*-KO) littermate embryos (of both sex) at embryonic day 17.5 (E17.5) and cultured as described by Gory-Fauré et al.⁶¹. Briefly, the brains were dissected, and the hippocampi were removed, dissociated, and plated on poly-L-lysine-coated coverslips in 35 mm dishes containing DMEM 10% horse serum. After 2 h 30 min, the culture medium was replaced with MACS medium.

In some experiments, neurons were transfected prior to plating using the Lonza Mouse Neuron Nucleofector 2b Kit (program 0-005, following the manufacturer's instructions). Neurons were electroporated with 4 µg of plasmid encoding control GFP, MAP6d1-GFP, or MAP6d1-GFP deletion mutants immediately following dissociation. To ensure efficient transfection, neurons from one hippocampus were plated per dish.

### Immunofluorescence microscopy and quantification

Cells were fixed in 4% paraformaldehyde, 4% sucrose in phosphate buffered saline (PBS) for 30 min at 37 °C, followed by permeabilisation in 0.2% Triton X-100 in PBS for 2 min at room temperature. Cells were then incubated with primary antibodies for 1 h at room temperature. Coverslips were washed 3 times in PBS - 0.1% Tween 20 and incubated with conjugated secondary antibodies for 1 h in the dark at room temperature. After three final PBS 0.1% Tween20 washes and one with PBS, nuclei were stained using Hoechst (Sigma) in the mounting medium (Dako).

Primary antibodies used included mouse monoclonal anti-Arl13b (1:1000, Addgene, 180085), mouse monoclonal anti-polyglutamylated tubulin (1:1000, Coger, AG-20B-0020), rabbit polyclonal anti-acetylated tubulin (1:5000, Millipore, ABT241), and rat monoclonal anti-tyrosinated tubulin (1:500, clone YL1/2⁶²). The secondary

antibodies were donkey anti-mouse Cy3 (715-165-151), donkey anti-rabbit AF647 (711-605-152) and donkey anti-rat AF488 (712-545-153) (1:500, Jackson Immuno-Research Laboratory).

Images of cilia were acquired on a Zeiss LSM 710 confocal microscope (equipped with a Zeiss AiryScan module), using a 63x oil-immersion NA 1.4 objective and Zen software (Zen 2.1 SP3 FP3 version 14.0.23.201, Carl Zeiss Microlmaging).

We measured ciliary length using Fiji on the maximal projection of confocal images of cilia, all combined in a single file to facilitate manipulation. Only cells with a visible GFP signal at the Golgi and neurites were included (Supplementary Fig. 5c). We traced the cilia manually to determine their lengths using the segmented line tool.

For fluorescence intensity measurements, we imported confocal images into Fiji and max-projected the z-planes spanning the cilium. The entire cilium was manually traced from the most proximal end to the distal end with a width of 0.17 µm. We then extracted the intensity profile of the selected region, normalising length and intensity to a 0-100 scale. To compare MAP6d1-GFP localisation to polyglutamylated tubulin, we drew a line along acetylated tubulin ciliary staining and compared the signal of both proteins along this line. Similarly, we drew a line along Arl13b ciliary staining to compare the respective protein signals of MAP6d1-GFP and tyrosinated tubulin localisation.

### Expansion microscopy

At DIV9, five days after transduction with a lentivirus expressing MAP6d1-EGFP, neurons were fixed in 4% paraformaldehyde and 4% sucrose in phosphate-buffered saline (PBS) for 30 min at 37 °C. Following fixation, neurons were stored in PBS for 4 days prior to processing for post-expansion staining, following the protocol from Gambarotto et al.⁶³. Briefly, coverslips were incubated for 5 h at 37 °C in a solution of 2% acrylamide and 1.4% formaldehyde in PBS. Polymerisation was then initiated by incubating the samples for 1 h at 37 °C in a monomer solution containing 19% (w/v) sodium acrylate, 10% acrylamide, 0.1% bis-acrylamide, 0.5% (v/v) TEMED, and 0.5% (w/v) ammonium persulfate in PBS. This was followed by a denaturation step at 95 °C for 1 h and 15 min.

Following the first round of expansion, gels were immunostained with the following primary antibodies: rabbit anti-GFP (1:500, Life Technologies, A11122) and mouse anti-adenylate cyclase (1:250, Life Technologies, MA5-47381) or mouse anti-polyglutamylated tubulin (1:250, Coger clone GT335). The secondary antibodies were donkey anti-mouse Cy3 and donkey anti-rabbit AF488 (711-545-152) (1:500, Jackson Immuno-Research Laboratory). Images were acquired using a Zeiss LSM900 confocal microscope equipped with a 63x, NA 1.3 objective in Airyscan mode. To isolate the ciliary structure and reduce adjacent labelling, cilia outlined from different z-planes were extracted using the BigTrace plugin for Fiji (https://github.com/ekatrukha/BigTrace). A maximal intensity projection is presented in Fig. 4d, e.

### Statistical analysis

All experiments were repeated at least three times. Statistical analysis was performed with Prism GraphPad version 10. Statistical tests and sample sizes are all indicated in the figure legends.

All in vitro experiments conducted with MAP6d1 and its mutants are summarised in Supplementary Table 3, including the concentrations used and the results obtained.

### Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

### Data availability

All data supporting the results of this study can be found in the article, Supplementary Information, and Source Data files. Materials can be
obtained from the corresponding authors under a material transfer agreement. The subtomogram average maps data in this study have been deposited in the EMDB under the following accession codes: EMD-52569 (singlet microtubule with luminal protofilaments), EMD-52572 (doublet microtubule with luminal protofilaments), EMD-52575 (doublet microtubule), EMD-53457 (combined map of singlet microtubule with luminal protofilaments, from focused refined maps EMD-53454 (singlet microtubule) and EMD-53452 (luminal protofilaments)), EMD-52623 (combined map of doublet microtubule from focused refined map EMD-52574 (B-tubule)). Source data are provided with this paper.

## Code availability

We have used the curvature radius.bsh script, originally developed by Olivier Burri (BIOP, EPFL) and modified for our study to analyze the curvature of B-tubule traces and calculate their mean curvature radii, which is available via GitHub [https://github.com/EricDenarier/MAPd1 MT_Doublets_2025].

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## Acknowledgements

We thank N. Chaumontel and C. Miscopein for helping protein purification and cell biology experiments, M. Benoît for helpful discussion and V.L. Brandt for editing the manuscript. This work used the Photonic Imaging Center of Grenoble Institute of Neuroscience (PIC-GIN, University Grenoble Alpes - INSERM U1216), which is part of the ISdV core facility and certified by the IBiSA label. We also used the two EM facilities in Grenoble: the European Synchrotron Radiation Facility (beam time on CM01) and the Grenoble Instruct-ERIC Center (ISBG; UMS 3518 CNRS CEA-UGA-EMBL) with support from the French Infrastructure for Integrated Structural Biology (FRISBI; ANR-10-INSB-05-02) and GRAL, a project of the University Grenoble Alpes graduate school (Ecoles Universitaires de Recherche) CBH-EUR-GS (ANR-17-EURE-0003) within the Grenoble Partnership for Structural Biology. The IBS Electron Microscope facility is supported by the Auvergne Rhône-Alpes Region, the Fonds Feder, the Fondation pour la Recherche Médicale and GIS-IBiSA. We thank the staff members of PIC-GIN and both EM facilities for their assistance. The mice used in this study were bred at the animal facility of IRIG-DRF-CEA (Grenoble, France), which is supported by funding from GRAL, a program of the University Grenoble Alpes graduate school (Ecoles Universitaires de Recherche) CBH-EUR-GS (ANR-17-EURE-0003). We thank the zootechnicians, S. Bama-Toupet and C. Magallon, for their assistance. This work was supported by the Institut National pour la Santé et la Recherche Médicale (INSERM), the Centre National de la Recherche Scientifique (CNRS) and l'Agence Nationale pour la Recherche (ANR-20-CE13-0005-01 to A.A., ANR-22-CE16-0018-01 to A.A., ANR-24-CE13-5131-01 to I.A.). D.G. was supported by ANR (20-CE13-0005-01), J.W. by the French MESR Ministry and M. de A. by ANR (ANR-22-CE16-0018-01).

## Author contributions

Conceptualisation: S.G.-F., L.S., I.A. Investigation: D.G., J.W., J.D., C.B., M. de A., G.E., E.D., S.G.-F., L.S. Formal analysis: D.G., J.W., J.D., M. de A., E.D., S.G.-F., L.S., I.A. Resources. G.E., E.D. Writing - Original draft: D.G, L.S., I.A. Writing - review and editing: A.A., D.G, J.W., E.D., G.E., C.B., S.G-F, L.S., I.A. Visualisation: D.G., J.W. Supervision: S.G-F, L.S., I.A. Funding acquisition: A.A., I.A.

## Competing interests

The authors declare no competing interests.

## Additional information

**Supplementary information** The online version contains supplementary material available at https://doi.org/10.1038/s41467-025-61679-0.

**Correspondence** and requests for materials should be addressed to Sylvie Gory-Fauré, Laurence Serre or Isabelle Arnal.

**Peer review information** *Nature Communications* thanks Khanh Huy Bui, who co-reviewed with Thibault Legal, and the other anonymous reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

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© The Author(s) 2025
**Supplementary Information**

**The Mn-motif protein MAP6d1 assembles ciliary doublet microtubules**

Dharshini Gopal<fnref n="1" />#, Juliette Wu<fnref n="1" />#, Julie Delaroche<fnref n="1" />, Christophe Bosc<fnref n="1" />, Manon De Andrade<fnref n="1" />,
Eric Denarier<fnref n="1" />, Gregory Effantin<fnref n="2" />, Annie Andrieux<fnref n="1" />, Sylvie Gory-Fauré<fnref n="1" />*, Laurence Serre<fnref n="1" />‡*,
Isabelle Arnal<fnref n="1" />‡‡

<footnote n="1">¹Université Grenoble Alpes, INSERM, U1216, CNRS, CEA, Grenoble Institut Neurosciences (GIN), Grenoble, France.</footnote>

<footnote n="2">²Institut de Biologie Structurale (IBS), Université Grenoble Alpes, CNRS, CEA, 71 Avenue des Martyrs, 38042 Grenoble, France.</footnote>
<figure><img src="image_24.png" /></figure>

<figure><img src="image_25.png" /><figcaption>Supplementary Figure 1. Microtubule minus end dynamic modulation by MAP6d1 and its mutants.</figcaption></figure>

a Graphs showing the growth rate (n = 158, 266, 361, and 217 events for 0, 10, 20 and 50 nM MAP6d1, respectively), shrinkage rate (n = 133, 182, 202 and 116 events), catastrophe and rescue frequencies (n =
35, 43, 72 and 51 MTs), the pause frequencies (n = 31, 41 and 39 MTs, for 10, 20 and 50 nM MAP6d1, respectively) and pause durations (n = 4, 34 and 49 events) of MTs assembled with increasing concentrations of MAP6d1, extracted from kymographs depicted in **Fig. 1a. b** Dynamical parameters of microtubules assembled with tubulin alone and in the presence of MAP6d1, MAP6d1-ΔMn2, MAP6d1-Mn2-7A and MAP6d1-Δ2-35: growth rate (n = 187, 175, 68, 239 and 184 growth events, respectively), shrinkage rate (n = 162, 98, 56, 205 and 117 shrinkage events, respectively), catastrophe and rescue frequencies (n = 39, 41, 18, 46 and 41 frequencies), extracted from kymographs depicted in **Fig. 3a**. Bars represent mean ± SD from at least three independent experiments. Circles with different colours represent the mean of each individual experiment. p values are indicated (Kruskal-Wallis ANOVA followed by post hoc Dunn's multiple comparison test. For clarity, only significant statistics are indicated for (b). Source data are provided as a Source Data file.
<figure><img src="image_26.png" /><figcaption>**Supplementary Figure 2. Analysis of MAP6d1-induced microtubule architectures.** a Distribution of microtubule architectures in the tomography data set. b Examples of different microtubule architectures induced by MAP6d1. Blue dashed lines indicate the location at which the z-sections are displayed. c Distribution of microtubules (MT) with different number of protofilaments (PF) measured along 265 µm of microtubule length for control, and for microtubules polymerised by MAP6d1 are 820 and 115 µm of singlet (SMT) and doublet (DMT) microtubule lengths, respectively. d Subtomogram averaged models of the initial doublet microtubule, B-tubule and combined map. e Fourier Shell Correlation (FSC) plots of</figcaption></figure>
reconstructions shown in this study obtained from Eman2 refinement output. Source data are provided as a Source Data file.
<figure><img src="image_27.png" /><figcaption>c</figcaption></figure>

<figure><img src="image_28.png" /><figcaption>b</figcaption></figure>

<figure><img src="image_29.png" /><figcaption>**Supplementary Figure 3. Purified recombinant proteins used in this study and their microtubule-binding properties.** a Purified MAP6d1 and the mutants (cyan asterisk) on Coomassie-blue-stained gels (10 % SDS PAGE). To the left are diagrams of full-length MAP6d1 and the mutants with the N-terminal in grey and the Mn-motifs with a pink asterisk. MW, molecular weight. b Representative immunoblots of co-sedimentation assays of Taxol-stabilised microtubules (Tx-MT) with MAP6d1, and the mutants detected with antibodies against tubulin and histidine-tag (to detect his-tag purified MAP6d1 and mutants). S-Supernatant; P-Pellet. c Analysis of blots presented in (b). Black dots represent the percentage of proteins in the pellet for each blot. Source data are provided as a Source Data file.</figcaption></figure>
<figure><img src="image_30.png" /></figure>

<figure><img src="image_31.png" /></figure>

<figure><img src="image_32.png" /></figure>

<figure><img src="image_33.png" /></figure>

<figure><img src="image_34.png" /></figure>

<figure><img src="image_35.png" /><figcaption>Supplementary Figure 4. Effects of the mutant MAP6d1-Mn1-7A on microtubule dynamics, tubulin recruitment and doublet microtubule formation. a Representative immunoblots of co-sedimentation assays of Taxol-stabilised microtubules (Tx-MT) with MAP6d1-Mn1-7A, detected with antibodies against tubulin and histidine-tag (to detect his-tag purified MAP6d1 mutant). S-Supernatant; P-Pellet. b Analysis of blots presented in (a). Black dots represent the percentage of proteins in the pellet for each blot. c Representative kymographs of microtubules grown from seeds (S) and 12 µM tubulin alone or with</figcaption></figure>
300 nM MAP6d1-Mn1-7A. **d** Dynamical parameters of microtubules assembled with tubulin alone or in the presence of MAP6d1-Mn1-7A. Plus end: growth rate (n = 323 and 319 events, respectively), shrinkage rate (n = 233 and 274 events, respectively), catastrophe and rescue frequencies (n = 60 and 62 microtubules, respectively). Minus end: growth rate (n = 243 and 152 events, respectively), shrinkage rate (n = 201 and 130 events, respectively), catastrophe and rescue frequencies (n = 52 and 38 microtubules, respectively). Bars represent mean ± SD from at least three independent experiments. Circles represent the mean of each individual experiment. p values are indicated (Kruskal-Wallis ANOVA followed by post hoc Dunn's multiple comparison test). **e Left:** snapshots of TIRF movies of 0.325 µM tubulin (cyan) mixed with GMPCPP seeds (magenta) in the absence (Control) or presence of 300 nM MAP6d1-Mn1-7A. **Right:** Negative staining electron microscopy images of microtubules assembled in the same conditions as for TIRF microscopy. MT, microtubule; Tub, tubulin. **f Left:** Percentage of microtubules with or without recruited tubulin in the control and with MAP6d1-Mn1-7A (87 and 117 microtubules for the control and with MAP6d1-Mn1-7A, respectively). Black dots represent the percentage of microtubules with recruitment for each experiment. **Right:** percentage of singlet (SMT) and doublet (DMT) microtubules in the control and with MAP6d1-Mn1-7A (total measured lengths of 195 and 409 µm, respectively). Black dots represent the percentage of doublet microtubules for each experiment. p values are indicated (two-sided Fischer's exact contingency test). Source data are provided as a Source Data file.
<figure><img src="image_36.png" /><figcaption>**Supplementary Figure 5.** Comparison of primary cilia from wildtype and MAP6d1 deficient neurons. **a** Quantification of ciliated cells in hippocampal neurons at 4 (left) and 9 days in vitro (DIV) (right). n ≥ 350 cells for each experiment. Bars represent mean ± SD from three independent experiments. Circles represent the ratio of ciliated cells of each experiment. p values are indicated (Mann-Whitney's test). **b** Representative images of wild-type (WT) and MAP6d1-knockout (KO) hippocampal neurons at 23 DIV stained for Arl13b and Hoechst. Quantification of cilium length from WT (n = 344 cilia) and KO neurons (n = 385 cilia) (3 experiments). Bars represent mean ± SD and circles, the mean of each experiment. p value is indicated (Mann-Whitney's test). **c** Hippocampal neuron at 7 DIV transfected with MAP6d1-GFP and stained for Arl13b. White arrow indicates the primary cilium. The graph represents the proportion of localisation to the cilia in neurons transfected with plasmids encoding GFP (n = 128 neurons), MAP6d1-GFP (n = 68 neurons), MAP6d1-ΔMn2-GFP (n = 59 neurons), MAP6d1-Δ2-35-GFP (n = 35 neurons).</figcaption></figure>
(n = 86 neurons) from at least three independent experiments. **d Left:** Normalised maximum fluorescence of tyrosinated tubulin along the ciliary length determined by Arl13b in 9 DIV WT (n = 42 cilia) and MAP6d1-KO (n = 47 cilia) hippocampal neurons. **Right:** Normalised maximum fluorescence of polyglutamylated tubulin along the ciliary length determined by acetylated tubulin in 9 DIV WT (n = 45 cilia) and MAP6d1-KO (n = 47 cilia) hippocampal neurons. Values represent mean ± SEM from 3 independent experiments. **e Top:** Representative images of primary cilia from 7 DIV hippocampal neurons expressing MAP6d1-Δ2-35-GFP stained for acetylated tubulin (Ac tub) and polyglutamylated tubulin (PolyE tub). Solid lines show normalised maximum fluorescence of MAP6d1-Δ2-35-GFP and polyglutamylated tubulin (PolyE tub) along the ciliary length determined by acetylated tubulin. Values represent mean ± SEM of n = 94 cilia from 3 independent experiments. Dotted lines represent normalised maximum fluorescence of MAP6d1-GFP and polyglutamylated tubulin (PolyE tub) along the ciliary length in 7 DIV hippocampal neurons expressing ectopic MAP6d1-GFP (data from **Fig. 4f**). **Bottom:** Representative images of primary cilia from 7 DIV hippocampal neurons expressing MAP6d1-Δ2-35-GFP stained for Arl13b and tyrosinated tubulin (Tyr tub). Plain lines show normalised maximum fluorescence of MAP6d1-Δ2-35-GFP and tyrosinated tubulin (Tyr tub) along the ciliary length determined by Arl13b. Values represent mean ± SEM of n = 35 cilia from 3 independent experiments. Dotted lines show normalised maximum fluorescence of MAP6d1-GFP and tyrosinated tubulin (Tyr tub) along the ciliary length in 7 DIV hippocampal neurons expressing ectopic MAP6d1-GFP (**Fig. 4g**). Source data are provided as a Source Data file.
<figure><img src="image_37.png" /><figcaption>**Supplementary Figure 6.** Cryo-electron tomography of microtubules in neurons. **a** Low-magnification images of mature neurons on EM grids acquired using Titan Krios cryo-electron microscope. White arrows indicate neurites. **b** High-magnification tomogram of neuritic extensions highlighting microtubules (MTs) with white arrows. **c** Zoomed-in image of the neuronal microtubule with luminal protofilaments indicated by cyan arrows. **d** Gallery of images showing filamentous densities resembling luminal protofilaments in singlet neuronal microtubules, and their corresponding schemes.</figcaption></figure>
<figure><img src="image_38.png" /></figure>

<caption>**Supplementary Figure 7. Cryo-EM processing workflow.** MT, microtubule; pf, protofilament.</caption>

<footnote>* All singlet MTs with luminal pfs were classified as 14-pf MTs with two luminal pfs.</footnote>
Map6d1

<figure><img src="image_39.png" /></figure>

**Supplementary Figure 8. MAP6D1-KO mouse generation and genotyping strategy.** Diagram of the targeting vector used and all the possible alleles for *MAP6D1*, and of the wild-type and knock-out alleles for *MAP6D1* (bottom). Orange bar: genomic DNA. Black box: exons with their corresponding number. Green and purple arrowheads: LoxP and Flp sequences, respectively. White bar: neo cassette, with the neomycin resistance gene (white box). Blue lines: zone of sequence homology for homologous recombination, with the corresponding size in kbp. Black arrowheads: primers used for the PCR genotyping.
<table><caption>**Supplementary Table 1. Sequence of the primers used to produce plasmids encoding MAP6d1 and its mutants.** For each construct, the sequence of the Forward (For) and Reverse (Rev) primers are indicated, along with the matrix used to produce the PCR fragments and the restriction enzymes used to linearize the cloning vector. All PCR matrices are from ref (30).</caption><thead><tr><th>Plasmid</th><th>Sequence</th><th>PCR Matrix</th><th>Cloning Vector</th></tr></thead><tbody><tr><td>MAP6d1</td><td>For 5'-AAGGAGATATACATATGGCGTGGCCCTGCATCAG-3'<br>Rev 5'-GGTGGTGGTGCTCGAGTGCGGCCGCTCAATGGTGATGGTGATGATGGTGATGGGTGGCGACCGGTGGATCC-3'</td><td>Map6d1-EGFP</td><td>pET42a (Nde I / Xho I) Novagen</td></tr><tr><td>MAP6d1 -Δ2-35</td><td>For 5'-AAGGAGATATACATATGGGGAGCGAGGAGTCGGGC-3'<br>Rev 5'-GGTGGTGGTGCTCGAGTGCGGCCGCTCAATGGTGATGGTGATGATGGTGATGGGTGGCGACCGGTGGATCC-3'</td><td>Map6d1-EGFP</td><td>pET42a (Nde I / Xho I) Novagen</td></tr><tr><td>MAP6d1 -ΔMn2</td><td>For 5'-AAGGAGATATACATATGGCGTGGCCCTGCATCAG-3'<br>Rev 5'-GGCGACCGGTGGATCCCCCACATTGAGAGTCTGAGGAGC-3'</td><td>MAP6d1 -ΔMn3</td><td>MAP6d1 + pET42a (Nde I / BamH I) This work</td></tr><tr><td>MAP6d1 -Mn2-7A</td><td>For 5'-AAGGAGATATACATATGGCGTGGCCCTGCATCAG-3'<br>Rev 5'-ACTCCAGTCCAAGCTTGCGCTGCCGCAGCCGCGGCCGCGGTCGCTACTACAGCTGCGTC-3'</td><td>Map6d1-EGFP</td><td>MAP6d1 + pET42a (Nde I / Hind III) This work</td></tr></tbody></table>
<table><caption>**Supplementary Table 2. Parameters for cryo-electron tomography data collection and processing.**</caption><thead><tr><th>Name</th><th>SMT with luminal pfs</th><th>DMT with luminal pfs</th><th>DMT</th><th>luminal pfs in SMT</th><th>B-tubule in DMT</th></tr></thead><tbody><tr><td>EMDB code</td><td>EMD-52569</td><td>EMD-52572</td><td>EMD-52575</td><td>EMD-53452</td><td>EMD-52574</td></tr><tr><td>Magnification</td><td colspan="5">33,000 X (K3 Gatan camera)</td></tr><tr><td>Voltage</td><td colspan="5">300 kV (Titan Krios)</td></tr><tr><td>Total electron dose</td><td colspan="5">117 electrons /Å<sup>2</sup></td></tr><tr><td>Defocus range</td><td colspan="5">-3 to -6 µm</td></tr><tr><td>Pixel size</td><td colspan="5">2.7 Å</td></tr><tr><td>Symmetry imposed</td><td colspan="5">C1</td></tr><tr><td>Subtomogram number</td><td>5881</td><td>1680</td><td>4997</td><td>5881</td><td>4997</td></tr><tr><td>Map resolution masked/unmasked at FSC 0.5</td><td>12.5 / 16.7 Å</td><td>28.6 / 37.4 Å</td><td>16.1 / 20 Å</td><td>21 / 24.7 Å</td><td>21.7 / 23.8 Å</td></tr></tbody></table>

FSC, Fourier Shell Correlation; DMT, doublet microtubule; SMT, singlet microtubule; pfs, protofilaments.
<table><thead><tr><th></th><th>Type of experiments</th><th>[Tubulin]</th><th>[MAP6d1]<br>[mutants]</th><th>Figure</th><th>Results</th></tr></thead><tbody><tr><td rowspan="4">MAP6d1</td><td>MT dynamics (TIRFm)</td><td>12 µM</td><td>10-50 nM</td><td>Fig 1a-c<br>Fig S1a</td><td>MT stabilization: reduced<br>growth &amp; shrinkage;<br>increased rescues &amp; pauses</td></tr><tr><td>MT architectures (EM)</td><td>25 µM</td><td>250 nM</td><td>Fig 2a-d<br>Fig S2</td><td>DMT</td></tr><tr><td>Tubulin recruitment on<br>GMPCPP stabilized MTs<br>(TIRFm &amp; EM)</td><td>0.325 µM<br>soluble<br>tubulin</td><td>100 nM</td><td>Fig 2e-f</td><td>Tubulin recruitment and<br>DMT</td></tr><tr><td>Co-sedimentation with<br>Taxol-stabilized MTs</td><td>1 µM</td><td>300 nM</td><td>Fig S3b-c</td><td>65% of MAP6d1 co-pelleted<br>with MT</td></tr><tr><td rowspan="3">Mn-motif<br>mutants</td><td>MT dynamics (TIRFm)</td><td>12 mM</td><td>300 nM</td><td>Fig 3a-b<br>Fig S1b<br>Fig S4c-d</td><td>No MT stabilization</td></tr><tr><td>Tubulin recruitment on<br>GMPCPP stabilized MTs<br>(TIRFm &amp; EM)</td><td>0.325 µM<br>soluble<br>tubulin</td><td>300 nM</td><td>Fig 3c-d<br>Fig S4e-f</td><td>No tubulin recruitment,<br>no DMT</td></tr><tr><td>Co-sedimentation with<br>Taxol-stabilized MTs</td><td>1 µM</td><td>300 nM</td><td>Fig S3b-c<br>Fig S4a-b</td><td>&lt;25% (Mn2-mutants) and<br>58% (Mn1 mutant) co-<br>pelleted with MT</td></tr><tr><td rowspan="3">Δ2-35<br>mutant</td><td>MT dynamics (TIRFm)</td><td>12 µM</td><td>50 nM</td><td>Fig 3a-b<br>Fig S1b</td><td>MT stabilization: Increased<br>growth rate and rescue<br>frequency</td></tr><tr><td>Tubulin recruitment on<br>GMPCPP stabilized MTs<br>(TIRFm &amp; EM)</td><td>0.325 µM<br>soluble<br>tubulin</td><td>100 nM</td><td>Fig 3c-d</td><td>No tubulin recruitment and<br>no DMT</td></tr><tr><td>Co-sedimentation with<br>Taxol-stabilized MTs</td><td>1 µM</td><td>300 nM</td><td>Fig S3</td><td>80% co-pelleted with MT</td></tr></tbody></table>

**Supplementary Table 3.** Overview of the *in vitro* techniques and protein concentrations used in this study and the main conclusions. DMT, doublet microtubule; MT, microtubule; EM, electron microscopy; TIRFm, TIRF microscopy.