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Inner lumen proteins stabilize doublet microtubulesin cilia and ﬂagella

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Mikito Owa1, Takayuki Uchihashi 2, Haru-aki Yanagisawa1, Takashi Yamano3, Hiro Iguchi3, Hideya Fukuzawa3,
Ken-ichi Wakabayashi4, Toshio Ando 5 & Masahide Kikkawa 1

Motile cilia are microtubule-based organelles that play important roles in most eukaryotes.
Although axonemal microtubules are sufﬁciently stable to withstand their beating motion, it
remains unknown how they are stabilized while serving as tracks for axonemal dyneins. To
address this question, we have identiﬁed two uncharacterized proteins, FAP45 and FAP52, as
microtubule inner proteins (MIPs) in Chlamydomonas. These proteins are conserved among
eukaryotes with motile cilia. Using cryo-electron tomography (cryo-ET) and high-speed
atomic force microscopy (HS-AFM), we show that lack of these proteins leads to a loss of
inner protrusions in B-tubules and less stable microtubules. These protrusions are locatednear the inner junctions of doublet microtubules and lack of both FAP52 and a known innerjunction protein FAP20 results in detachment of the B-tubule from the A-tubule, as well as
ﬂagellar shortening. These results demonstrate that FAP45 and FAP52 bind to the inside of
microtubules and stabilize ciliary axonemes.

1 Department of Cell Biology and Anatomy, Graduate School of Medicine, The University of Tokyo, 7-3-1 Hongo Bunkyo-ku, Tokyo 113-0033, Japan.
2 Department of Physics and Structural Biology Research Center, Chikusa-ku, Nagoya University, Nagoya 464-8602, Japan. 3 Graduate School of Biostudies,
Kyoto University, Kyoto 606-8502, Japan. 4 Laboratory for Chemistry and Life Science, Institute of Innovative Research, Tokyo Institute of Technology,
Yokohama 226-8503, Japan. 5 WPI Nano Life Science Institute, Kanazawa University, Kakuma, Kanazawa 920-1192, Japan. Correspondence and requests formaterials should be addressed to M.K. (email: mkikkawa@m.u-tokyo.ac.jp)

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Cilia and ﬂagella are microtubule-based organelles that
operate as both antennae and propellers in eukaryotic
cells. Cilia are classiﬁed as either non-motile or motile.
Non-motile cilia, or primary cilia, function as antennae and are
involved in signal transduction through the hedgehog, Wnt, andCa2+ signaling1. Motile cilia and ﬂagella beat at 20–60 Hz2,3 anddrive cellular motility and ﬂuid ﬂow. Since the cells in most tissues
are ciliated, ciliary dysfunction leads to various types of humandiseases termed ciliopathies, which include hydrocephalus, situs
inversus, retinal degeneration, and nephronophthisis4. Furthermore,
recent studies revealed that the assembly of cilia is signiﬁcantly
decreased in several types of tumors5,6, implying some correlation
between ciliogenesis and tumorigenesis.
The structure and protein composition of motile cilia and ﬂagella are well conserved among eukaryotes. The axoneme, the core
structure of cilia and ﬂagella, is composed of a central pair of
microtubules cylindrically surrounded by nine doublet microtubules (DMTs). This arrangement is often referred to as the 9+2 structure (Fig. 1a). In DMTs, 10 protoﬁlaments of the B-tubule
are attached to 13 protoﬁlaments of the A-tubule at the inner andouter junction (Fig. 1a). In motile cilia and ﬂagella, structures
essential for motility, such as axonemal dyneins, radial spokes, andthe nexin–dynein regulatory complex (N-DRC), are arranged on
DMTs with a 96-nm repeating unit7–9. Axonemal dyneins boundon A-tubules slide on the neighboring B-tubules and this sliding
propagates along the axoneme, generating a bending force. The
activity of dyneins is regulated by the interaction between theradial spokes and the central pair of singlet MTs10–12. The N-DRC
forms a crossbridge between neighboring DMTs and is requiredfor orchestrating dynein activity and axonemal integrity13–16.
Recent advances in cryo-electron tomography (cryo-ET)
techniques have revealed structural details of DMTs, yet a signiﬁcant question remains—what stabilizes DMTs? Cytoplasmic
MTs frequently bend with various degrees of curvature17,18 andthis bending occasionally induces MT breakage19. Furthermore, arecent study demonstrated that cytoplasmic MTs were damaged
even after bending only several times20. Therefore, to ensure ahigh beat frequency, the DMTs of motile cilia and ﬂagella should
be structurally robust compared with cytoplasmic MTs. However,
the mechanism that provides DMTs with such robustness
remains to be clariﬁed.
Nicastro and colleagues have provided insights into thestructural basis of DMTs using cryo-ET of Chlamydomonas ﬂagella. They reported periodic high densities on the inner surfacesof A-tubules and B-tubules, which they named microtubule innerproteins (MIPs, Fig. 1a)7,21. To date, two MIPs in the A-tubule
have been identiﬁed by recent studies22,23. MIPs have also been
observed in the axonemes of higher organisms7,21,24–26, implyingthat these inner structures are essential for the integrity of DMTs
in motile cilia and ﬂagella.
In this paper, we explore the mechanisms stabilizing DMTs using
Chlamydomonas mutants and various techniques, including cryoET and high-speed atomic force microscopy (HS-AFM). We ﬁndthat two uncharacterized ﬂagellar-associated proteins (FAP), FAP45
and FAP52, are essential for the stability of B-tubules. Lack of both
FAP45 and FAP52 leads to the loss of MIPs in B-tubules, resulting
in disruption of the MT walls. Furthermore, the B-tubule wall
detaches from the A-tubule at the inner junction, and ﬂagellar
shortening is observed when both FAP52 and FAP20 are absent.
These results indicate that the B-tubule is reinforced by MIPs andits stability is essential for maintaining the structure of the axoneme.
Results
Identiﬁcation of proteins essential for DMT stabilization. To
identify proteins that stabilize DMTs, we searched the2Chlamydomonas ﬂagellar proteome database27. We postulated
that those proteins are (1) abundant in the axoneme fraction, (2)tightly bound on the axoneme even in the presence of high salt,and (3) highly conserved among ciliated organisms. The proteinsthat satisfy these criteria are listed in Supplementary Table 1. We
focused on FAP45, an uncharacterized protein whose peptides
were most frequently found in the proteomic analysis (Supplementary Table 1, total unique peptide and Axo columns). FAP45
is a coiled-coil protein composed of 501 amino acids, has a
predicted molecular weight of ∼59 kDa, and is conserved amongorganisms with motile cilia. The human ortholog of FAP45 iscoiled-coil domain-containing protein 19 (CCDC19), also known
as NESG1. The transcript of this protein is enriched in thenasopharyngeal epithelium and trachea28. However, the functionsof FAP45/CCDC19 are unclear. Thus, we chose FAP45 as a
candidate of proteins stabilizing DMTs.
We ﬁrst looked for partner(s) that interacts with FAP45 on theaxoneme using chemical crosslinking. We treated isolated wild
type axonemes with 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC, zero-length crosslinker). The crosslinked products
were solubilized and immunoprecipitated with a polyclonal antiFAP45 antibody. Tubulin was identiﬁed as the major crosslinked
partner of FAP45 using western blot (Fig. 1c, open arrowhead),suggesting a direct interaction between FAP45 and tubulin. In
addition, a mass spectrometric analysis revealed that the ∼130kDa product is composed of FAP45 (~59 kDa) and FAP52 (~66
kDa), probably in a 1:1 ratio based on the molecular weight(Fig. 1c and d, ﬁlled arrowhead; Supplementary Fig. 1d, arrowhead; Supplementary Table 2).
FAP52 is the Chlamydomonas ortholog of human WD40
repeat domain 16 (WDR16) and is listed in Supplementary
Table 1. It was previously shown that WDR16 knockdown inzebraﬁsh led to severe hydrocephalus29. Furthermore, a recent
study reported that homozygous deletion of the WDR16 gene inhuman leads to situs anomalies, which are typical phenotypes ofciliopathy30. However, the function of the FAP52/WDR16
protein in ﬂagellar motility and axonemal structure remains
unclear. We hypothesized that FAP45 and FAP52 proteins play arole in stabilizing DMTs.
Characterization of Chlamydomonas FAP45 and FAP52
mutants. We used Chlamydomonas mutants lacking FAP45 and
FAP52 to investigate their function. The mutants were isolated
from ∼10,000 clones in an insertionally mutagenized Chlamydomonas library31. In these mutant strains, the ~1.8 kbp aphVIII
fragment was inserted into the 5′UTR of the FAP45 gene or theﬁrst exon of the FAP52 gene (Supplementary Fig. 1b). By asouthern blot, we conﬁrmed that each mutant has one aphVIII
fragment inserted only at the targeted locus (Supplementary
Fig. 1c). These strains did not express detectable FAP45 or FAP52
protein as analyzed by a western blot of axonemal fractions andimmunoﬂuorescence of the axonemes (Fig. 1b; Supplementary
Fig. 2b and c). On the other hand, fap45 and fap52 axonemes
retained known major axonemal components (Fig. 1b). Also,
FAP45 and FAP52 were incorporated into the axoneme independently of each other (Fig. 1b). The ~130 kDa crosslinked
product of FAP45 and FAP52 was detected in the wild typeaxoneme treated with EDC but not in the crosslinked fap45 or
fap52 axoneme (Fig. 1c and d, ﬁlled arrowheads), consistent withthe mass spectroscopic analysis.
Next, we examined the motility phenotype of fap45 and fap52.
The swimming velocity and beat frequency of fap45 cells wereslightly reduced (Fig. 1e; Supplementary Movie 1), whereas thephenotype of fap52 was quite similar to that of wild type (Fig. 1f;
Supplementary Movie 1). Intriguingly, the double mutant fap45

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c

B-tubule

A-tubuleRS

IDA: IC140(110 kDa)

IDA: p28(22 kDa)RSP1(88 kDa)

DRC2(65 kDa)

DRC4

(kDa)

(kDa)

250

250

150

150

(55 kDa)

100

100

FAP20

(22 kDa)

75

75

FAP45

(59 kDa)

FAP52(66 kDa)

50

50

e

f
fap45

200

DRC

ODA

g

250

Wild type250

Wild type
fap52

200
150

150

100

100

100

50

50

50
0

0
0

25

50

75

100

Wild typefap45fap52

200

150

0
IDA

W
ild

(64 kDa)

Swimming
velocity (μm/s)

OJ

ty
pe
fa
p4
5
fa
p5
2

ODA: IC2250
IJ

WB : FAP52
1.5 mM EDC

W
ild

DMT

d

WB : FAP45
1.5 mM EDC

ty
pe
fa
p4
5
fa
p5
2

ty
p
fa
p4 e
5
fa
p5
fa 2
p4
5f
ap
52

b

W
ild

a

0

25

50

75

100

0

25

50

75

100

Beat frequency (Hz)Fig. 1 Characteristics of the FAP45 and FAP52 null mutants. a Top: a transverse view of the 9+2 structure of the Chlamydomonas axoneme. Scale bar =50 nm. Bottom: a magniﬁed DMT. Arrowheads indicate MIPs. Major structures are colored; Outer dynein arm (ODA, magenta); Inner dynein arm (IDA,
green); Dynein regulatory complex (DRC, purple); Radial spoke (RS, blue). IJ inner junction, OJ outer junction. Scale bar = 25 nm. b Western blot analyses
of wild type, fap45, fap52, and fap45fap52 double mutant axonemes stained with various antibodies. FAP45 and FAP52 proteins were not detected in fap45
and fap52, respectively. Proteins essential for ﬂagellar motility (ODA-IC2: outer dynein arm-intermediate chain 2; IDA-IC140: inner dynein armintermediate chain 140; IDA-p28: inner dynein arm-light chain p28; RSP1: radial spoke protein 1; DRC2 and 4: dynein regulatory complex 2 and 4; FAP20:
inner junction protein of DMT) were not reduced in the mutants. c, d Axonemes from wild type and mutant Chlamydomonas crosslinked using EDC (zerolength crosslinker) were immunoblotted with anti-FAP45 (c) and anti-FAP52 antibodies (d). Filled arrowheads indicate the crosslinked product of FAP45
and FAP52 in a 1:1 ratio. The open arrowhead indicates the crosslinked product of FAP45 and tubulin. e–g Motility phenotypes of the mutants were assayedusing the CLONA system. Swimming velocity and beat frequency were slightly reduced in fap45, whereas no signiﬁcant reduction was observed in fap52.
fap45fap52 showed a more severe phenotype than did fap45

and fap52 (fap45fap52) swam signiﬁcantly more slowly than wild
type, with a low beat frequency (Fig. 1g; Supplementary Movie 1).
Flagellar beating of swimming cells in these mutants appeared
normal (Supplementary Movie 2). Furthermore, there was anaccumulation of non-motile fap45fap52 cells under conﬂuent
culture conditions (Supplementary Movie 3). We also observed
the swimming behavior of wild type and fap45fap52 log phasecells in the viscous medium. Most of the wild type cells swam
slowly but smoothly (Supplementary Movie 4) whereas many of
the fap45fap52 cells stopped swimming (67 of 78 cells) or
struggled to swim against the viscous load (Supplementary Fig. 3;
Supplementary Movie 4), indicating that fap45fap52 ﬂagellacannot beat properly under this condition. These results indicatethat the lack of both FAP45 and FAP52 led to synergistic effects
on motility phenotypes.

FAP45 and FAP52 are luminal proteins in B-tubules. We
biochemically localized FAP45 and FAP52 in DMTs by fractionating the axonemal proteins using increasing concentrations ofsarkosyl32 (Supplementary Fig. 4a). FAP45 began to be extracted
from the pellet at 0.2% sarkosyl, half of the FAP45 was extracted

at 0.3%, and the protein was completely extracted from thepellet at 0.7% (Supplementary Fig. 4a). FAP52 started to besolubilized at 0.3% sarkosyl and almost all the protein was solubilized at 0.7%, although a small amount of protein still remainedin the pellet. These behaviors correlate well with solubilization of
the B-tubule.
To distinguish whether FAP45 and FAP52 are located outside
or inside the B-tubule, we prepared axonemes containing
biotinylated FAPs and tested whether or not the proteins areaccessible to streptavidin. These axonemes were puriﬁed fromrescue strains expressing FAP proteins whose N-terminus or Cterminus was fused to biotin carboxyl carrier protein tag (BCCPtag)33,34. These rescued strains expressing BCCP-tagged proteins(Supplementary Fig. 4b and c) swam like wild type. However,essentially no signal of biotinylation was detected on thoseaxonemes (Supplementary Fig. 4d), suggesting that streptavidin
access to BCCP tag was prevented, presumably by the B-tubulewall. Therefore, we treated axonemes with 0.15% sarkosyl, which
partially broke the B-tubule wall and, as expected, resulted in
detection of the tagged proteins (Supplementary Fig. 4e). Taken
together with the results that the loss of FAP45 and FAP52 did
not affect the incorporation of major axonemal proteins bound

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4

ODA

Wild type

FAP45 and FAP52 form MIPs in B-tubules. To reveal the defectcaused by the loss of FAP45 and FAP52, we observed the mutant
axonemes by cryo-ET. We found that the density of dynein b, a
species of inner arm dyneins was decreased in fap45 and fap52(Fig. 2a–c). This result suggests that in the mutants the number ofdynein b may be reduced or dynein b cannot bind to the B-tubule
properly for some reason.
For further visualization of missing structures in fap45 andfap52, we applied Student’s t-tests to compare wild type andmutant density maps33 (Fig. 3e and f, right). In agreement withthe above results, the mutants also showed structural defects
inside of the B-tubule (Fig. 3).
A comparison between wild type and fap45 DMTs shows thatthe luminal surface of wild type B-tubule is smoother than that of
fap45 (Fig. 3d and e). In the fap45 DMTs, the grooves between Btubule protoﬁlaments B7-B9 are prominent (Fig. 3b and e;
Supplementary Fig. 5b). The grooves seem to be partially ﬁlledwith ﬁlamentous densities as shown in the t-map (Fig. 3e right,
highlighted in green). These densities are arranged in a 48-nm
repeat, and reminiscent of a lateral ﬁlament connecting MIP3,
fMIP B8B9, and fMIP B7B8 in Tetrahymena DMTs35. Therefore,
we named the missing structures as MIP3c (Fig. 3e right, circled
in red).
A comparison between the wild type and fap52 DMT revealed
that an arch-like density is missing in the B-tubule of the fap52
mutant. In the wild type DMT, there are a large arch-like density
and a small spike-shaped density in a 16-nm repeat inside of the
B-tubule on the protoﬁlament B9 and B10, called MIP3a and
MIP3b, respectively (Fig. 3a and d, highlighted in yellow andorange; Supplementary Fig. 5a)21. The DMTs of fap52 completely
lacked the densities of MIP3a, whereas MIP3b appear to beunchanged (Fig. 3c and f; Supplementary Fig. 5a).
Next, we tested whether the predicted atomic structures of
FAP45 and FAP52 can ﬁt into these densities. Chlamydomonas
FAP52 contains 11 WD40 domains (Supplementary Fig. 1a).
Since WD40 repeat domains typically form a seven-bladed βpropeller, FAP52 probably contains two β-propellers. Indeed, amodel comprising two β-propellers ﬁt well into the Chlamydomonas density map (Supplementary Fig. 5d), as well as therecently published sub-nanometer map of Tetrahymena35(Supplementary Fig. 5e), suggesting that the density of MIP3acorresponds to one molecule of FAP52. For MIP3c, predicted
coiled-coils of FAP45 protein were ﬁt into the density(Supplementary Fig. 5c). FAP45 is mainly composed of helices
and coiled-coils that account for 85% of the total sequence(Supplementary Fig. 1a). The total length of helices and coiled
coils in FAP45 is ~60 nm. Therefore, the circled densities in
Supplementary Fig. 5c are probable one unit of MIP3c. Thus,
MIP3c probably consists of FAP45.
To investigate the reasons why the double mutant fap45fap52
cells swim more slowly than single mutants, we observed thefap45fap52 axoneme using cryo-ET. In this case, we could not
apply 3D sub-tomographic averaging because many of theaxonemes were frayed and the B-tubules were partially missing(Fig. 4a and b). Since cryo-ET is not suitable for observing theshapes of individual DMTs due to the missing wedge effects, thinsections of the fap45fap52 axoneme were observed by conventional EM and revealed that the B-tubules were missing in∼33.8% of the outer DMTs (Fig. 4c). Among them, 23% lacked allof the protoﬁlaments in the B-tubules, whereas 77% retained
several protoﬁlaments. These data suggest that the loss of both

a

DRC

IDA
RS

b

fap45

on the outer surface of the DMT (Fig. 1b), these data indicate that
FAP45 and FAP52 are most likely luminal proteins enclosed by
the B-tubule.

c

fap52

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Fig. 2 Dynein b density is reduced in the fap45 and fap52 DMT. Isosurface
renderings of averaged DMT repeats: wild type (a), fap45 (b), and fap52(c). The density of dynein b is decreased in fap45 and fap52 (arrowheads).
Other species of IDAs, ODAs, RS, and N-DRC were properly arranged on
the DMT in fap45 and fap52. Scale bar = 25 nm

FAP45 and FAP52 decreases structural stability between the Btubule protoﬁlaments.
Direct observation of B-tubule depolymerization by HS-AFM.
The above static DMT structures suggested that the B-tubules offap45fap52 are less stable than those of wild type. We directly
observed the stability of the B-tubule using HS-AFM36, in which asmall cantilever tip (tip diameter ∼1–2 nm) intermittently taps
the DMT surface ﬁxed on a mica stage. This intermittent contact
can eliminate frictional force during lateral scanning and damage
to the DMT caused by scanning because the feedback operation
maintains a constant tip–sample interaction force and the force isapplied for a short time (<100 ns).
We ﬁrst observed in vitro polymerized MTs in the absence oftaxol, a stabilizing agent. It was previously reported25 that MTs
depolymerize spontaneously within two or three frames (1.0–1.5 s),suggesting that depolymerization requires <50 nm/s. Actual depolymerization could be faster26 but our HS-AFM has insufﬁcient
temporal resolution to verify this. Compared with in vitro
polymerized MTs, DMTs are more stable and do not depolymerize
spontaneously under normal HS-AFM conditions and theirstructures can be observed in detail.
When DMTs are adsorbed on the mica surface, AFM images
were classiﬁed into three types as shown in Supplementary Fig. 7;
Class 1: The 24 nm periodicity of ODAs was visualized on the topof the DMT, suggesting that the B-tubule and the radial spokes were
immobilized on the mica; Class 2: The heads of the radial spokes in
96 nm periodicity were visualized on the top of the DMT,

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a

B5
B6

Wild typed

B-tubule90°

B4 B3 B2

B7

MIP3c

A-tubule

B1

B6
B7
B8
B9
B10

A12

B8
B9

A13
B10

A1

MIP3a MIP3b

IJ

b

e

fap45

c

f

fap52

MIP3c (t-map) on fap45
B6
B7
B8
B9
B10

B6
B7
B8
B9
B10

MIP3a (t-map) on fap52

Fig. 3 fap45 and fap52 mutant have structural defects in B-tubules. Isosurface renderings of averaged axonemal 96-nm repeats from wild type (a, d), fap45(b, e), and fap52 (c, f). (a–c) are cross-sectional and oblique, and (d–f) are longitudinal views. The black dashed lines in (a–c) indicate the slicing plane of(d–f). Protoﬁlament numbers are labeled in (a) and (d–f). Missing structures in fap45 and fap52 were determined by Student’s t-tests as previously
described33. The t-value maps acquired by the t-tests were overlaid on the renderings of fap45 (e, light green) and fap52 (f, light yellow). In this ﬁgure,
MIP3a and b were colored by yellow and orange, respectively. MIP3c (the missing structure in fap45) was colored by green, according to the t-value map
shown in (e). Scale bar = 25 nmab

ProximalA
B
A
B

Distal

ODA

A
A

c

B
A

A

B

AB

A

B

Fig. 4 B-tubules are partially depolymerized in fap45fap52. a, b A typical tomogram of a fap45fap52 axoneme in cross-sectional (a) and longitudinal view(b). A dashed line in (a) indicates the slicing plane of (b). Arrowheads in (b) show break points in the B-tubules. c Typical thin section TEM images offap45fap52 axonemes. Broken B-tubules were observed in 33.8% of the DMTs (n = 552). The 9+2 structure was disorganized and the axonemes were
frayed in regions containing broken B-tubules. Scale bar = 100 nm
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a

d
z

z

0s

z

Doublet MT
Cantilever

y

y

x

x

y

B
A

x

Mica stage

b
–1 s

0s

1s

Wild type
fap45

0 s∼

Length of depolymerized
B-tubule (nm)

∼0 s

fap52

400

fap45fap52

300
200
100
0
0

20

40

60

Time (s)Wild type
10 s

20 s

60 s

0s

1s

20 s

40 s

c
–1 s

fap45fap52
10 sFig. 5 Direct observation of B-tubule depolymerization by HS-AFM. a Schematic of high speed-AFM observation of the DMT. The tip of the cantilever (tipdiameter ~1–2 nm) intermittently tapped the DMT surface on a mica stage. At time 0 s, the tip of the cantilever was thrust into the B-tubule and made asmall hole (circled in red in b and c), immediately followed by normal HS-AFM observation. b, c Representative time-lapse images of wild type (b) andfap45fap52 (c) DMTs acquired by HS-AFM. In the wild type, enlargement of the hole usually stopped within a few seconds, whereas most of the B-tubulesin the ﬁeld were broken in fap45fap52 by 40 s. Scale bar = 100 nm. d The average length of the depolymerized B-tubules was plotted against time (wildtype: n = 11, total length = 7.25 μm; fap45: n = 8, total length = 4.92 μm; fap52: n = 7, total length = 3.83 μm; fap45fap52: n = 10, total length = 4.95 μm)

suggesting that both of the A and B-tubule were immobilized on themica (Supplementary Fig. 7b); Class 3: The radial spokes are
periodically and horizontally projected from the A-tubule, suggesting that the B-tubule was visualized on the top while the A-tubule
was immobilized on the mica (Supplementary Fig. 7c).
To determine how a defect inﬂuences the stability of DMTs, weselected DMTs in the class 2 and 3 and made a small hole bythrusting the tip of the cantilever into the B-tubule (Time = 0,
Fig. 5a) and monitored whether or not the damage propagates. In
wild type, the hole rarely grew during the observation period(Fig. 5b, Supplementary Movie 5). On the other hand, in the
mutants, the hole gradually expanded in both the proximal anddistal directions (Fig. 5c, Supplementary Movie 6–8). We did notobserve signiﬁcant difference between depolymerization toward
the plus end and the minus end of the DMTs. Time-course
analyses of the depolymerized B-tubule length demonstrated thatthe tendency for depolymerization followed the order as
fap45fap52 > fap45 > fap52 ≥ wild type (Fig. 5d). These datastrongly suggest that FAP45 and FAP52 stabilize the doublet Btubule and prevent depolymerization induced by damage to the
MT wall. Of note, the DMT of fap45fap52 are still more stable
than cytoplasmic microtubules, suggesting that B-tubules are alsostabilized by other mechanisms, such as post-translational
modiﬁcations of tubulin37,38 (Supplementary Fig. 8) and fMIPs
that bind inside the B-tubule along its length35.

investigated whether FAP52 is involved in stabilizing the junction. We previously reported that FAP20 is a component of theinner junction and a null mutant of FAP20 (fap20) showed anabnormal swimming phenotype, although the structure of the
DMTs appeared to be normal39. Based on the hypothesis that
MIP3a anchors the B-tubule to the A-tubule together with
FAP20, we constructed the double and triple mutants fap20fap45,
fap20fap52, and fap20fap45fap52. Most fap20fap45 cells weretrembling and their motility appeared slightly worse than that of
fap20 cells (Supplementary Movie 9). In contrast, fap20fap52 andfap20fap45fap52 cells were essentially paralyzed (Supplementary
Movie 9). Furthermore, the ﬂagellar lengths of fap20fap52 andfap20fap45fap52 deviated greatly from wild type, fap20, andfap20fap45 (Fig. 6a and b), with a far higher ratio of short ﬂagella.
To identify the cause of these defects, we observed the axonemesof the mutants by thin section TEM. The axonemes and DMTs of
fap20fap45 appeared similar to those of fap20 (Fig. 6c and d).
Interestingly, several doublet B-tubules in fap20fap52 andfap20fap45fap52 were detached from the A-tubules (Fig. 6carrowheads; Fig. 6d) whereas no detachment of B-tubules from
A-tubules was observed in fap20fap45 axonemes. Given that such
defects were rarely observed in fap20 axonemes39, these results
demonstrate that MIP3a is required for anchoring the B-tubule to
the A-tubule at the inner junction together with FAP20, and thejunction is important for stabilizing the axonemal structure.

MIP3a is important for the crossbridge of the inner junction.
Since MIP3a is located near the inner junction between the Atubule and B-tubule and appeared to attach to the A-tubule35, we

Discussion
Ciliary/ﬂagellar MTs are remarkably stable compared with cytoplasmic MTs. Of the 9+2 MTs, DMTs are directly stressed by the

6

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a

fap20
fap45fap52

Wild type

fap20

fap20fap45

fap20fap52

c

fap20

fap20fap45

fap20fap52

fap20
fap45fap52

Flagellar length (μm)

b
16
12
8
4

d
fap20

fap20
fap45fap52

p5

2

52

fap20fap52

fa
45

0f

ap

ap

p2
fa

p2

0f

fa

0f

ap

45

0
p2
fa

p2

ty
ild
W

fa

pe

0

fap20fap45

Fig. 6 MIP3a is important for B-tubule anchoring to the A-tubule at inner junctions. a Differential interference contrast (DIC) images of wild type, fap20,
fap20fap45, fap20fap52, and fap20fap45fap52 cells. Scale bar = 10 μm. b Box and whisker plots of ﬂagellar length in wild type, fap20, fap20fap45, andfap20fap52. fap20fap52 and fap20fap45fap52 ﬂagella were signiﬁcantly shorter than in wild type, fap20, and fap20fap45. The whiskers represent 95%
conﬁdence intervals. The box represents the interquartile range and the center line shows the median. c Cross-sectional EM images of fap20, fap20fap45,
fap20fap52, and fap20fap45fap52 axonemes. Arrowheads indicate disconnections between the A-tubule and B-tubule. Scale bar = 100 nm. d Crosssectional images of fap20, fap20fap45, fap20fap52, and fap20fap45fap52 DMTs. Scale bar = 50 nm

axonemal dynein-generated force and constantly bent andstraightened in Chlamydomonas ﬂagella or trachea cilia, yet theirstructures remain intact. The mechanism that stabilizes axonemal
MTs has not been clariﬁed. Here, we identiﬁed FAP45 and FAP52
as proteins that stabilize DMTs by binding to the inner lumen of
the B-tubule. In this report we identify factors essential for thestabilization of ciliary/ﬂagellar MTs.
Based on our studies and existing data, we propose a schematic
model of FAP45 and FAP52 (Fig. 7b). Our cryo-ET observations
show that MIP3a and MIP3c are composed of FAP52 and FAP45,
respectively (Fig. 7a). FAP52/MIP3a anchors the protoﬁlaments
A13 and B10, whereas FAP45/MIP3c is bound inside of B7, B8,
and B9 protoﬁlaments. Besides, biochemical crosslinking showsthat FAP45 and FAP52 directly interact with each other. Thus,
these two proteins bundle B7–B10 protoﬁlaments to reinforce the
B-tubule. Consistent with this model, the B-tubules of fap45 weremore vulnerable to physical stress than those of fap52 in the HSAFM observation (Fig. 5d, Supplementary Movie 7). The MIP3astructure was observed also in vertebrates7,24, but MIP3c was notclearly described probably due to the limitation of resolution.
However, the amino acid sequence of FAP45 is highly conserved
from protists to mammals (Supplementary Fig. 6), suggesting thatthe functions of FAP45 are conserved also in higher organisms as
in Chlamydomonas.
Our data also suggest that DMTs are stabilized by fail-safe
mechanisms. The single mutants of FAP45 and FAP52 did notshow signiﬁcant decreases in swimming speed, and only the
double mutant fap45fap52 showed slow swimming and defects of
the B-tubules. Although the B-tubules of fap45fap52 were moreeasily depolymerized than wild type B-tubules, the speed ofdepolymerization was slower than that of in vitro polymerized
pure MTs.
The inner junctions of DMTs are also stabilized by fail-safe
mechanisms involving multiple proteins. We previously demonstrated that FAP20 is a constituent of the inner juction39. In
addition to the true inner junction, our data clariﬁed that MIP3aalso connects B-tubules to A-tubules (Fig. 6c and d). Furthermore, some inner junctions appeared to be attached in fap20fap52
and fap20fap45fap52 mutants (Fig. 6c). In the triple mutant,
additional proteins, such as tektin and PACRG, may contribute tostabilizing the inner junction, given our previous observation thatthese two proteins were partially retained in fap2039.
Ciliopathy caused by FAP45/CCDC19 mutation has not beenreported to date. On the other hand, patients lacking FAP52/

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a

90°
B7

B6

B8

IJ

MIP3c:FAP45

B tubule

B5
B4

B9

B3

B10

B2
B1

OJ

B6
B7
B8
B9
B10

A tubuleb

MIP3a:FAP52+FAP45, FAP52
B7–FAP45, FAP52

B6

B8
B9

FAP45

B10
IJ A13
A1

FAP52

Fig. 7 A schematic model of MIP3. a Isosurface rendering of the wild type density map. MIP3a-c is highlighted as shown in Fig. 2 (MIP3a: yellow; MIP3b:orange; MIP3c: green). The right map is the longitudinal view sliced by the line in the left map. A possible unit of MIP3c is circled in red. Scale bar = 25 nm.
b A model of DMT stabilization. FAP45 (green) and FAP52 (yellow) are localized on the inner lumen of the B-tubule and stabilize the MT wall. In particular,
FAP45 ties protoﬁlaments B7–B9. FAP52 anchors B9 and B10 to A13 and binds to FAP45. Thus, these proteins strengthen the MT wall against DMTbending. Loss of both proteins leads to destabilization of the B-tubule and the B-tubule is depolymerized by bending

WDR16 were reported to have abdominal situs inversus and situs
inversus totalis30. Despite abnormal laterality, those patients did
not have typical symptoms of primary ciliary dyskinesia, such asrecurrent bronchiolitis, hydrocephalus, or low nasal nitric oxidelevels. These ﬁndings are consistent with the fap52 phenotype inChlamydomonas, where we could not detect abnormal phenotypes other than the lack of MIP3a structure. Given that thephenotype of fap45 was more severe than that of fap52 in Chlamydomonas, we predict that the lack of FAP45/CCDC19 inhuman causes ciliopathy.
The structures of MIPs in the A-tubule are more complex thanin the B-tubule. It is previously shown that hyper-stable ribbons
were composed of four protoﬁlaments to which MIP4 is bound40,
suggesting MIP4 stabilizes the ribbon structure. A recent paper
reported that FAP85 is a MIP in the A-tubule and stabilizes
cytoplasmic microtubules in vitro22. Besides, a study using Tetrahymena revealed that Rib72A and B are essential for assemblyof MIP6, which is important for proper ﬂagellar beating23.
Although how those MIPs contribute to the stability of the Atubule is still unclear, it is possible that MIPs in A-tubules also
have fail-safe mechanisms, similar to MIP3a and c.
Methods
Strains and culture conditions. The Chlamydomonas strains used in this study arelisted in Supplementary Table 3. The double and triple mutants were constructedby standard methods3. All cells were cultured on Tris-acetate-phosphate (TAP)
plates with 1.5% agar or in TAP medium. fap45 and fap52 mutants were isolated
from a library of mutants generated by aphVIII gene insertion31. The mutants were
backcrossed with wild type several times before use. For the PCR analyses, following primers were used; FAP45-F: 5′-GTGGCTTGAGCACCCTACTCACTT-3′;
FAP45-R: 5′-CCGCTTGCTACCTCCAAATAAAGA-3′; FAP52-F: 5′-CGCCGACCTCTACATTTCTGAGTT-3′; FAP52-R: 5′-CTTTGACTCCAGGTCCCAGATG
AT-3′; aphVIII-F: 5′-GTCGACTTGGAGGATCTGGACGA-3′. For the probe of
the southern blot, the aphVIII coding sequence was ampliﬁed with following primers; 5′-ATGGACGATGCGTTGCGTGC-3′; 5′-TCAGAAGAACTCGTCCAACAGCCG-3′. The probe was PCR-labeled with biotin-16-dCTP and detected by
Streptavidin-HRP.
8

Motility assay using the CLONA system. To assay the motilities of Chlamydomonas cells in TAP medium or TAP with ﬁcoll, videos were recorded using a highspeed camera (EX-F1; Casio) attached on a dark-ﬁeld light microscope (BX51;
Olympus) at 600 fps. The videos were analyzed using the CLONA system41 and theresults were plotted using R Project software.

Antibodies. The antibodies used in this study are described in Supplementary
Table 4. Anti-FAP45 and anti-FAP52 antibodies were raised against full-length
FAP45 and FAP52 proteins, respectively (Supplementary Fig. 2a). The full-length
cDNA of FAP45 or FAP52 was cloned between the EcoRI and BamHI sites ofpMAL-c2x (New England Biolabs). In both constructs, a 6×His tag was inserted
into the HindIII site of pMAL-c2x to enable puriﬁcation in two steps. For afﬁnity
puriﬁcation, each cDNA was also cloned between the NdeI and EcoRI sites ofpColdI (Takara). Expression of the recombinant proteins was induced in Escherichia coli BL21(DE3) (TAKARA BIO Inc.) with 0.3 mM IPTG, and almost all of theexpressed protein was solubilized from each construct. MBP-FAP45-6×His and
MBP-FAP52-6×His were puriﬁed with amylose resin (New England Biolabs) andthen further puriﬁed with Ni-NTA agarose (Qiagen). These puriﬁed proteins wereused as antigens. Each antibody was afﬁnity puriﬁed using polyvinylidene
diﬂuoride membranes blotted with FAP45-6×His or FAP52-6×His. For western
blot analyses, we loaded 5 μg of isolated axonemes in each lane. Full blot images areshown in Supplementary Fig. 9.

Chemical crosslinking of axonemes. Isolated axonemes of wild type, fap45, andfap52 were treated with EDC (Thermo Scientiﬁc) in HMEK (30 mM Hepes–KOH,
pH 7.4, 5 mM MgSO4, 1 mM EGTA, 50 mM K-acetate) for 60 min at room temperature. After quenching the reaction, crosslinked axonemes were analyzed by
SDS–PAGE and western blotting with anti-FAP45 and anti-FAP52 antibodies. The
crosslinked products were immunoprecipitated with anti-FAP45 antibody following a previously described method42. Mass spectrometry analysis (LC tandem MS)of the precipitates was performed at the Proteomics and Mass Spectrometry
Facility (University of Massachusetts Medical School).

Immunoﬂuorescence microscopy. Nucleoﬂagellar apparatus (NFA) was prepared
as previously described43. After ﬁxation with 2% formaldehyde for 10 min at room
temperature, NFAs were treated with cold methanol (−20 °C). The samples were
immunostained as previously described44. Images were taken with a CCD camera(ORCA-R2; Hamamatsu Photonics) linked to a ﬂuorescence microscope (IX70;
Olympus).

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Generation of BCCP-tagged strains. Fragments from the start codon to immediately before the stop codon in the FAP45 and FAP52 genes were ampliﬁed bygenomic PCR and inserted into pIC2 plasmids16. In the FAP52 construct, a 3×HA
tag was inserted into the C terminus of FAP52. Each construct was linearized andtransformed into fap45 or fap52 cells by electroporation. Streptavidin-Alexa546
staining of the axonemes was performed as previously described39.

2.
3.
4.
5.

Thin-section TEM. Samples for thin-section TEM were prepared as previously
described45 except that we used 50 mM Na-phosphate, pH 7.0 instead of cacodylate
buffer. The samples were observed using a transmission electron microscope (JEM3100FEF; JEOL) equipped with a 4096 × 4096-pixel CMOS camera (TemCamF416; TVIPS). All images were taken at 300 keV, with ~3 μm defocus, at a magniﬁcation of 40,000 and a pixel size of 2.5 Å.

6.
7.
8.

Cryo-sample preparation. Puriﬁed axonemes in HMDEK buffer (30 mM
Hepes–KOH (pH7.4), 5 mM MgSO4, 1 mM dithiothreitol, 1 mM EGTA, 50 mM
potassium acetate, protease inhibitor cocktail (Nacalai Tesque)) were incubatedwith anti-beta-tubulin antibody (1:10,000 ﬁnal; T0198, SIGMA) for 15 min on ice,followed by incubation with goat anti-mouse IgG (H+L) 15 nm gold (1:20 ﬁnal; BBInternational) and 15 nm colloidal gold conjugated with BSA. The mixtures were
loaded onto home-made holey carbon grids and plunged into liquid ethane at−180 °C using an automated plunge-freezing device (EM GP; Leica).

9.

10.

11.

Image acquisition. Grids were transferred into the JEM-3100FEF using a high-tilt
liquid nitrogen cryo-transfer holder (914; Gatan Inc.). Tilt series images wererecorded at −180 °C using a K2 summit direct detector (Gatan) and the serialEM46.
The angular range was ±60° with 2.0° increments. The total electron dose waslimited to 100 e−/Å2 and the nominal magniﬁcation was ×6000. An in-column
energy ﬁlter was used with a slit width of 20 eV and a pixel size of 7.1 Å.

12.

Image processing. Image processing for subtomogram averaging of DMT was
carried out as previously described33,34. Tilt series images were aligned and backprojected to reconstruct 3D tomograms using IMOD47. Alignment and averagingof the subtomograms were performed using custom Ruby-Helix scripts48 and
PEET software7 to average the 96-nm repeats of DMTs. UCSF Chimera was used
for isosurface renderings49. FSC curves were shown in Supplementary Fig. 5f.

15.

13.

14.

16.

17.
HS-AFM observation. Flagella were demembranated with 0.5% Nonidet P-40 in
HMDEK (30 mM Hepes–KOH, 30 mM Hepes–KOH, pH 7.4, 5 mM MgSO4, 1 mM
DTT, 1 mM EGTA, 50 mM K-acetate, protease inhibitor cocktail (Nacalai Tesque)), followed by centrifugation for 2 min. The axoneme pellets were suspended
with ATP buffer (HMDEK containing 0.1 mM ATP, 1 mM ADP, and 0.5% polyethylene glycol (20,000) and incubated for 1 min at room temperature. Frayedaxonemes were then diluted with HMDEK and placed on a mica stage. After 10
min incubation, residuals are washed with the HMDEK buffer. The AFM images
were recorded using a home-built high-speed atomic force microscope based ontapping mode50–52 at frame rates of 1–2 fps. All observations were performed in
HMDEK buffer at room temperature. We selected DMTs of which the angle to thefast-scan axis is <45° to minimize accidental damage on the DMTs. A miniature
AFM cantilever (Olympus) with a spring constant of ~0.2 N/m, a quality factor of~2, and a resonant frequency of ~800 kHz was used for the HS-AFM imaging. For
imaging DMTs, a free oscillation amplitude was set to be ~2 nm and the set-point
amplitude for the feedback control was 80% of the free oscillation amplitude,
corresponding to the tapping force of ~120 pN53 (used the Eq. (10)). To create
defects on the DMTs, the set-point amplitude was decreased to <5% of the freeoscillation amplitude and the tapping force was increased to >200 pN at the speciﬁed area.

18.
19.

20.
21.
22.
23.

24.

25.
26.

Data availability
Data supporting the ﬁndings of this manuscript are available from the corresponding
author upon reasonable request. A reporting summary for this article is available as a
Supplementary Information ﬁle. The source data underlying Figs. 1e–g, 5d and 6b areprovided as a Source Data ﬁle. Cryo-ET reconstructions are available from the Electron
Microscopy Data Bank (EMDB) with following accession numbers: fap45 (EMD-9766),
fap52 (EMD-9767), and wild type (EMD-9768).

Received: 27 August 2018 Accepted: 10 February 2019

27.
28.
29.

30.
31.

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Acknowledgements
We thank Dr. John Leszyk (UMMS) for mass spec analysis and Dr. Brian Dynlacht(NYU) for helpful discussion. M.O. is a recipient of the fellowship from Japan Society for
Promotion of Sciences for young scientists. This work was supported by CREST, theJapan Science and Technology Agency to M.K. and T.A., the Japan Society for the
Promotion of Science (JSPS) KAKENHI (18H01837) to T.U., JSPS KAKENHI (25117506
and 15H01206) to K.W., and JSPS KAKENHI (16H04805) and JST Advanced Low
Carbon Technology Research and Development Program (ALCA, JPMJAL1105) to H.F.

Author contributions
M.O. and M.K. designed the experiments. M.O. performed most of the experiments
and analyzed the data. M.O., T.U. and M.K. wrote the paper. M.O. and K.W. performed
chemical crosslinking experiments. M.O. and T.U. performed A.F.M. analyses. M.O.
and H.Y. performed cryo-ET and E.M. experiments. T.Y., H.I., and H.F. provided theChlamydomonas mutant library. T.A. devised the AFM method for sample manipulation.

Additional information
Supplementary Information accompanies this paper at https://doi.org/10.1038/s41467019-09051-x.
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