Ultrastructural organization of NompC in themechanoreceptive organelle of Drosophilacampaniform mechanoreceptorsLandi Suna,b, Yuan Gaoc, Jianfeng Hea,b, Lihong Cuia,b, Jana Meissnerd, Jean-Marc Verbavatzd, Bo Lic, Xiqiao Fengc,and Xin Lianga,b,1

aTsinghua-Peking Joint Center for Life Sciences, School of Life Sciences, Tsinghua University, 100084 Beijing, China; bMax Planck Partner Group, School ofLife Sciences, Tsinghua University, 100084 Beijing, China; cSchool of Aerospace Engineering, Tsinghua University, 100084 Beijing, China; and dElectronMicroscopy Facility, Max Planck Institute of Molecular Cell Biology and Genetics, 01307 Dresden, Germany

Edited by Miriam B. Goodman, Stanford University, Stanford, CA, and accepted by Editorial Board Member Yuh Nung Jan February 27, 2019 (received forreview November 14, 2018)

Mechanoreceptive organelles (MOs) are specialized subcellularentities in mechanoreceptors that transform extracellular mechan-ical stimuli into intracellular signals. Their ultrastructures are keyto understanding the molecular nature and mechanics of mecha-notransduction. Campaniform sensilla detect cuticular straincaused by muscular activities or external stimuli in Drosophila.Each campaniform sensillum has an MO located at the distal tipof its dendrite. Here we analyzed the molecular architecture of theMOs in fly campaniform mechanoreceptors using electron micro-scopic tomography. We focused on the ultrastructural organizationof NompC (a force-sensitive channel) that is linked to the array ofmicrotubules in these MOs via membrane-microtubule connectors(MMCs). We found that NompC channels are arranged in a regularpattern, with their number increasing from the distal to the proximalend of the MO. Double-length MMCs in nompC29+29ARs confirm theankyrin-repeat domain of NompC (NompC-AR) as a structural com-ponent of MMCs. The unexpected finding of regularly spacedNompC-independent linkers in nompC3 suggests that MMCs maycontain non-NompC components. Localized laser ablation experi-ments on mechanoreceptor arrays in halteres suggest that MMCsbear tension, providing a possible mechanism for why the MMCsare longer when NompC-AR is duplicated or absent in mutants.Finally, mechanical modeling shows that upon cuticular deforma-tion, sensillar architecture imposes a rotational activating force,with the proximal end of the MO, where more NOMPC channelsare located, being subject to larger forces than the distal end. Ouranalysis reveals an ultrastructural pattern of NompC that is struc-turally and mechanically optimized for the sensory functions ofcampaniform mechanoreceptors.

NompC | microtubule | mechanoreceptive organelle |electron tomography | mechanoreceptor

The conversion of mechanical signals into electrical signals incells, known as mechanotransduction, is required for the

perception of sound, touch, and acceleration (1, 2). Mechano-transduction occurs much more rapidly than visual photo-transduction or olfactory transduction (2, 3). This suggests that inmechanotransduction mechanical stimuli are directly converted tointracellular signals rather than through a second messenger as invisual and olfactory transduction (1, 2, 4). Based on electrophysi-ological and mechanical measurements, it has been hypothesizedthat the mechanotransduction apparatus contains a transductionchannel coupled to a molecular spring (3, 5). Mechanical signals areconveyed to the transduction channel by the spring, and the channelresponds by changing the opening probability of its pore, throughwhich ion influx initiates electrical signals (1). The molecular springis a compliant structure and its compliance has two functions. First,it matches the mechanical impedance of rigid structures, such asintracellular cytoskeleton or extracellular matrix, to that of morecompliant structures, such as the channel’s gating apparatus. Sec-

ond, it allows the channel’s gate to fluctuate between open andclosed states, thereby encoding incoming stimuli into graded sig-nals. For these reasons, this compliant structure has been termedthe “gating spring” (1, 5). A key question is how the mechano-transduction apparatus operates in vivo: How are external forcesconveyed to the gating spring and how does the gating spring inturn couple these forces to the channel?In mechanosensory cells, mechanoreceptive organelles (MOs)

are specialized subcellular entities where the transduction ap-paratuses reside and function. The ultrastructural architecturesof MOs have been studied in various model cells, including thehair bundles of inner ear hair cells (4), the microtubule-baseddendrites of Caenorhabditis elegans touch cells (6, 7), and theciliated dendrites of fly type I mechanoreceptors (8–11). Thesestudies provide structural insights into the molecular basis ofmechanotransduction in these types of mechanoreceptors.Campaniform mechanoreceptors are type I insect mechanore-

ceptors whose dendrites contain a modified cilium (8, 12). Theyrespond to cuticular strain caused by muscular activities or externalstimuli to provide mechanosensory feedback during locomotion(13). In Drosophila, campaniform mechanoreceptors at differentlocations (wing, haltere, leg, etc.) vary in their cuticular, supporting,and neuronal structures (14, 15). This morphological diversity is

Significance

Mechanosensory cells convert environmental mechanicalstimuli into intracellular signals. This process, termed mecha-notransduction, occurs in specialized mechanoreceptive or-ganelles. Using electron tomography we discovered that themechanoreceptive organelle in fly campaniform mechanore-ceptors contains thousands of force-sensitive ion channels thatare arranged in a regular pattern, aligned to the intracellularmicrotubule cytoskeleton. A mechanical model suggests thatthe pattern is structurally and functionally optimized, becausemore force-sensitive channels are located at regions that aresubject to larger activating forces. We propose that such apattern enhances the sensitivity and broadens the dynamicrange of mechanosensation in this type of mechanoreceptor.

Author contributions: B.L., X.F. and X.L. designed research; L.S., Y.G., L.C., J.M., B.L., andX.L. performed research; J.-M.V. contributed new reagents/analytic tools; L.S., Y.G., J.H.,B.L., X.F., and X.L. analyzed data; and L.S. and X.L. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission. M.B.G. is a guest editor invited by theEditorial Board.

Published under the PNAS license.1To whom correspondence should be addressed. Email: xinliang@tsinghua.edu.cn.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1819371116/-/DCSupplemental.

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thought to be important for these receptors to detect differenttypes of cuticular strains caused by their natural stimuli. TheMOs of campaniform mechanoreceptors are located at the distaltips of the modified cilia (10, 12). In previous work, the MOs ofpedicellar campaniform mechanoreceptors of fly halteres werestudied by transmission electron microscopy (TEM) (10, 16),using thin sections and glutaraldehyde fixation. A set of seriallyconnected structures in the MO is thought to form a mechanicalsignaling pathway that links extracellular structures to the trans-duction channels (16). In particular, the ankyrin-repeat (AR) do-main of NompC (NompC-AR) was found to contribute structurallyto a membrane–microtubule connector (MMC). Based on struc-tural and mechanical analyses, we proposed a molecular model inwhich a transduction channel (i.e., NompC) is connected to amolecular spring (i.e., NompC-AR) (17, 18), as predicted by the“gating-spring” model (1). The finding of this NompC–microtubulecomplex is consistent with other findings on NompC, includingthose showing that NompC is a bona fide force-sensitive ionchannel (19, 20), that it is mechanically important in fly hearing (9,21), and that NompC-AR is required for microtubule binding andmechanosensory gating of NompC channels (22, 23). Most recently,Jin et al. (24) reported the atomic structure of NompC resolved bycryo-EM, which showed that NompC is a homotetrameric channeland, most strikingly, that four NompC-ARs form an AR bundle.The new NompC structure raises many questions (25), especiallyhow it relates to the architecture of the transduction apparatus inthe MO of fly mechanoreceptors.Earlier studies of the MOs of campaniform mechanoreceptors

by conventional TEM suffered from technical limitations. Forexample, the glutaraldehyde fixation increased the risk of tissuedisruption, in particular to membranes and fine filaments (SIAppendix, Fig. S1). In addition, the low z-resolution of thin-section-based TEM (50 to 100 nm) and the high likelihoodthat cutting oblique sections (SI Appendix, Fig. S2) made struc-tural measurements inaccurate. These limitations made it diffi-cult to compare the wild-type and mutant structures and therebyprecluded more detailed functional and mechanical studies.Therefore, new techniques are required to further determineMO ultrastructure.In the present study, we used high-pressure freezing (HPF)

and dual-axis electron tomography (ET) (26) to analyze the 3Dultrastructure of MOs in fly campaniform mechanoreceptors (SIAppendix, Supplementary Note 1). We found that NompCchannels are arranged in a regular pattern on the MO membraneand their number increases from the distal to the proximal end ofthe MO. Mechanical modeling showed that as a product of thesensillar architectures the MO is strained by a rotational acti-vating force. In this model, the proximal end of the MO, wheremore NompC are located, receives larger forces than the distalend. Therefore, the spatial pattern of NompC matches the dis-tribution of the activating forces on the MO, suggesting astructural and mechanical optimization for the sensory functionof the MOs in fly campaniform mechanoreceptors. Additionalstructural analysis on nompC mutants confirmed NompC-AR asa component of MMCs and unexpectedly revealed regularlyspaced NompC-independent linkers, suggesting that MMCs mayalso contain non-NompC components.

ResultsSensillar Structures of the Modified Cilium. Halteres are the fly’sgyroscopes. Forces produced by rotations of the fly body duringflight generate stresses at the bases of the rapidly oscillatinghalteres; the resulting strains are sensed by several arrays ofcampaniform mechanoreceptors in the pedicel and scabellumsegments of the haltere (SI Appendix, Fig. S3). To understandthe 3D ultrastructure of MOs in these mechanoreceptors, weapplied serial block-face imaging with an FIB/SEM (focusedion beam/scanning electron microscope). Campaniform mecha-

noreceptors in haltere pedicel and scabellum arrays showedsimilar morphological organizations but differed in their cutic-ular, supporting, and neuronal architectures (Fig. 1, SI Appendix,Fig. S4, and Movies S1–S4). These results agree with the pre-vious observations using conventional TEM and SEM but pro-vide 3D structures for the entire sensillum (14, 15).In the present study, we focused on campaniform mechano-

receptors in haltere pedicel arrays (Fig. 1). In this type ofmechanoreceptor, the cupola is connected to the cuticle via jointmembranes and overlies the septum, dendritic sheath, and MO(Fig. 1 A and B). The dendritic terminal of the sensory cell isspecialized into a modified cilium and abuts the cuticularstructures. The MO of campaniform mechanoreceptor is locatedat the distal tip of modified cilium (12). The entire sensillarstructure is better visualized in 3D models of the differentcomponents (Fig. 1C and Movie S2). The 3D structure of a

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Fig. 1. Sensillar organization of the modified cilium in a pedicellar cam-paniform mechanoreceptor of the fly haltere. (A and B) Two representativelateral views of a modified cilium in a campaniform mechanoreceptor takenfrom the FIB/SEM volume data. The structures are reproduced and presentedas cartoon schematics on the right. The colors indicate different structures.The numbers label different segments of the modified cilium (outlined inblack). The original volume data are shown in Movie S1. The axes are de-fined in the main text and shown in each panel. (Scale bars: 1 μm.) (C) Thestructural model segmented from the volume data. The 3D segmentation ispresented in Movie S2. Note that the volume data of a haltere scabellumcampaniform mechanoreceptor and the segmented model are also pre-sented (SI Appendix, Fig. S4 and Movies S3 and S4). TB, tubular body. (Scalebar: 1 μm.)

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campaniform mechanoreceptor from the haltere scabellum isshown for morphological comparison (SI Appendix, Fig. S4 andMovies S3 and S4). To better describe the 3D structures, wedefined three axes: The z axis (Fig. 1A) is perpendicular to thecuticle surface and is directed into the interior of the tissue; the xaxis (Fig. 1A) and y axis (Fig. 1B) are parallel to the longer andshorter axes of the MO. Based on these coordinates, the MOshowed a fan shape in the x–z plane (Fig. 1A), a finger shape inthe y–z plane (Fig. 1B), and a round-ended rectangular or el-liptical shape in the x–y plane (Fig. 2B and Movie S1).

Structures of Sheath–Membrane Connectors and MMCs. To recon-struct the intracellular structures of the modified cilium withhigher resolution, we used ET and HPF. We collected tilt-seriesET data on 250- or 300-nm tissue sections. Ten sections in serieswere stitched in 3D space to reconstruct the entire modifiedcilium (Fig. 2A and Movie S5). Taking advantage of the ETvolume data, we corrected the potential problem of obliquesections by adjusting the visualization plane for each structure ofinterest in 3D space (SI Appendix, Fig. S2).We focused on the ultrastructure of the MO (Fig. 2B), and in

particular the sheath–membrane connectors (SMCs) and MMCs,because these two linkers are likely essential elements in formingthe mechanical signaling pathway in the mechanotransductionapparatus of campaniform mechanoreceptors (10). In our ETdata, SMCs had a filamentous shape and inserted deeply into the

sheath matrix (SI Appendix, Fig. S5); these features differ fromthe “short column”-like shape observed in our previous study(10). The SMC filaments had a density of about 4,000 to5,000 μm−2, thereby forming a dense connective layer betweenthe membrane and sheath (SI Appendix, Fig. S5).In our ET data, MMCs appeared to be thin and filamentous

connectors that linked the membrane to the microtubules (Fig.2C; also see SI Appendix, Fig. S6). They were often branched atboth the membrane and microtubule ends (Figs. 2C and 3H andSI Appendix, Fig. S6). Most strikingly, we found that each MMCwas linked to a button-shaped structure on the MO membrane(Fig. 2 C and D). Collectively, these buttons were aligned,through MMCs, to the arrayed microtubules and formed a 2Darray on the membrane (Fig. 2D). The spatial periodicity be-tween adjacent buttons was ∼28 nm along the x axis, similar tothe intermicrotubule distance (10) and ∼18 nm along the lon-gitudinal axis of microtubules (Fig. 3F). The overall density ofthese buttons on the MO membrane was 2,032 ± 323 μm−2 inwild-type cells (n = 4). A segmented model for the local struc-tures is presented for better visualization (Fig. 2E and Movie S6).

NompC Channels Structurally Contribute to the Bulk of Button-Shaped Structures. Based on the observations described aboveand in the previous studies (10, 22), we wondered if the button-shaped structures that are linked to MMCs correspond to orcontain the channel domains of NompC. In wild-type, these

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buttons were arranged in an array (Fig. 3A). They had an av-erage diameter of 10.3 ± 1.6 nm (n = 364 buttons from n =4 cells) (Fig. 3B), similar to the size of NompC channel domain(Fig. 3C). Interestingly, these button-shaped structures werenearly absent in nompC3, a nompC null mutant (Fig. 3A). InnompC3, we observed small dots on the membrane (Fig. 3A).These dots were smaller in size compared with the buttons inwild-type flies (Fig. 3 A and B). We also studied nompC29+29ARs

flies in which the wild-type NompC is replaced with onecontaining two serially linked AR domains on the amino termi-nus but an unchanged channel domain (22). In this mutant, thebutton-shaped structures were present, similar to wild-type cells(Fig. 3A). Based on these results, we conclude that the channeldomain of NompC structurally contributes to the bulk ofmembrane-associated button. This observation shows thatNompC are arrayed on MO membrane by aligning to the mi-crotubules via MMCs.

NompC-Independent Linkers in nompC3 Are Regularly Spaced butHave Different Structures from Wild-Type MMCs. We then won-dered what the small dots in nompC3 are. Could they possiblycorrespond to the membrane-contacting structures of the irreg-ularly spaced NompC-independent linkers found in previousstudies (10, 22)? Therefore, we checked the linkers in nompC3 inthe x–y planes. Surprisingly, we found that in ET data, themembrane-microtubules linkers in nompC3 were regularlyspaced (Fig. 3D), different from previous observations in TEMdata (10).

To better understand these linkers, we analyzed their spatialdistribution and structure. In the x–y plane, nearly every micro-tubule was associated with an MMC in both wild-type andnompC3 (Fig. 3D). In addition, the spatial density of these linkersalong the z axis (Fig. 3 E and F) was similar in wild-type, nompC3,and nompC29+29ARs (Fig. 3G). We then compared the mor-phology of individual linkers in wild-type and nompC3 (Fig. 3 H–Jand SI Appendix, Fig. S6). Wild-type MMCs were morphologi-cally heterogeneous (Fig. 3H), so we classified them into fourtypes (Fig. 3J): Type 1 has no branched ends, type 2 has branchedends only on membrane side, type 3 has branched ends only onmicrotubule side, and type 4 has branched ends on both sides. Inwild-type, over 60% of MMCs were branched at both sides (i.e.,type 4) and more than 90% were branched at one end or theother (i.e., type 2, type 3, and type 4) (Fig. 3J). In a sharp con-trast, nearly 80% of the linkers in nompC3 fell into type 1 (i.e., nobranched ends on either side) (Fig. 3 I and J and SI Appendix,Fig. S6). Therefore, NompC-independent linkers have a spatialdistribution similar to wild-type MMCs, but they appear to havea different structure.We were curious why fewer linkers were observed in nompC3

in previous studies (10, 22). We suspected that these linkersmight be more susceptible to the glutaraldehyde-based samplepreparation. Therefore, we imaged the glutaraldehyde fixedwild-type and nompC3 samples by ET (SI Appendix, Fig. S7).Interestingly, we found that wild-type samples showed regularlyspaced MMCs (SI Appendix, Fig. S7). However, the linkers innompC3 were nearly absent (SI Appendix, Fig. S7), recapitulatingour previous results (10). These observations suggest that the

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Fig. 3. Button-shaped structures and MMCs in wild-type and nompC3. (A) The x–z plane views of MOmembrane inwild-type (Upper), nompC3 (Middle), andnompC29+29ARs (Lower). The genotype of nompC29+29ARs:nompC2, uas-nompC29+29ARs-GFP/nompC2; +/nompC-gal4.Both nompC3 and nompC2 are null alleles of the nompCgene. Red arrows indicate the button-shaped structuresin wild-type and nompC29+29ARs or the dot densities innompC3. (Scale bar: 100 nm.) (B) The size distributions ofbutton-shaped structures in wild-type and small dots innompC3. We used the diameters of the bounding circlesto quantify the sizes of these structures. (C) Size mea-surements of NompC transmembrane domains (NompC-TMs). Atomic model: Protein Data Bank ID code 5VKQ.(Left) The channel domain or linker domain of NompChas a size of 6 to 9 nm (edge length of the boundingsquares). (Right) The bounding circle of NompC channeldomain has a diameter of ∼11 nm. (D) Representativeimages of MOs in x–y planes show that nearly every mi-crotubule is associated with a MMC in wild-type andnompC3. (Scale bar: 100 nm.) (E) Representative imagesof the local regions in MOs in x–z planes (between themicrotubule plane and membrane plane) show thatMMCs (Lower) are aligned to microtubules (Upper). Bothwild-type and nompC3 data are presented for compari-son. (Scale bar: 100 nm.) (F ) The distributions of thespatial interval (d ) between adjacent MMCs alongmicrotubules in wild-type and nompC3. (G) The statisti-cal comparison of d in wild-type (black), nompC3 (blue),and nompC29+29ARs (red) strains. (H) The representativex–y plane images for several wild-type MMCs. The lastimage in each panel contains the segmentation ofa typical MMC (red). Note the branched ends of wild-type MMCs on both membrane and microtubule ends(red arrowheads). (Scale bar: 100 nm.) (I) Representativex–y plane images for several linkers in nompC3. Thesegmentation (red) of a linker is presented in thelast image. Note that most of the linkers have nobranched ends. (Scale bar: 100 nm.) (J) The morphological classifications of MMCs in wild-type and nompC3. Representative images of wild-type MMCs are shown inthe insets (1–4). In these insets, the membrane is on the left side and the microtubule is on the right side. Several other representative images of MMCs in wild-typeand nompC3 are presented in SI Appendix, Fig. S6.

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glutaraldehyde fixation-based sample preparation is likely a majorreason for the significant disappearance of linkers in nompC3. Thesusceptibility to the glutaraldehyde-based method suggests thatNompC-independent linkers are less stable in comparison withwild-type MMCs.

The Number and Length of MMCs in Wild-Type. Having confirmedthe presence of NompC-independent linkers, we wondered ifthis observation would argue against the previous conclusionthat NompC-AR structurally contributes to MMCs or, on thecontrary, suggest that MMCs contain non-NompC compo-nents in addition to NompC-AR. To address this issue, wemeasured the numbers and lengths of MMCs in the MO ofwild-type, nompC29+29ARs, and nompC3 cells (Fig. 4 A–C andSI Appendix, Fig. S8).The number of MMCs in the MOs varied with z-position,

reflecting the MO geometry. The distal region had the leastnumber of MMCs. Toward the proximal side, the number ofMMCs first increased then stayed relatively constant through thedepth of the MO and only slightly decreased near the proximalend (Fig. 4D). Both nompC29+29ARs and nompC3 had numbers ofMMCs similar to wild-type (Fig. 4D).The mean length of wild-type MMCs was 42.7 ± 13.5 nm (n =

416 MMCs from four cells). The length distribution was broad,ranging from 20 to 80 nm (Fig. 4E). The large variation wasprimarily because MMCs at different locations had differentlengths. There were two regular patterns. First, in x–y planes, thelengths of MMCs in the middle region of MO were longer thanthose on two sides (Fig. 4 A and F and SI Appendix, Fig. S8).Second, along the z axis, the average length of MMCs first in-

creased and then stayed relatively constant until the proximalend of the MO (Fig. 4G). We noticed that at all z-positionsMMCs showed a similar range of lengths (Fig. 4F and SI Ap-pendix, Fig. S8), suggesting that the increase in the mean lengthof MMCs is primarily due to the increase in the ratio oflonger MMCs.We also noticed a spatial correlation between the length of

MMCs and the local membrane–sheath contact. In the distalpart of the MO, the membrane only loosely connected to thesheath (Fig. 4A) and appeared to be closer to the microtubules.In these regions, the MMCs were mostly shorter. On the con-trary, in the proximal part of the MO, where most of the mem-brane was tightly attached to the sheath (Fig. 4A), many moreMMCs were longer. Similarly, in the x–y planes at all z-positionsthe membrane on two sides of the MO (x axis) only looselyconnected to or even detached from the sheath. In these regions,the MMCs were generally shorter than those in the middle re-gion, where the sheath had a tighter contact with the membrane(Fig. 4A). Thus, the proximity between the sheath and mem-brane varies with both the z- and x-positions.

MMCs Are Longer in nompC29+29ARs and nompC3. Having measuredthe length of wild-type MMCs, we then analyzed the MMCs innompC29+29ARs. If NompC-AR contributes to the MMCs, thendoubling the length of NompC-AR should increase the length ofthe MMCs. The length distribution of MMCs in nompC29+29ARs

showed a location-dependent pattern, similar to wild-typeMMCs (Fig. 4B and SI Appendix, Fig. S8). The mean lengthsof MMCs in nompC29+29ARs were twice as long as wild-typeMMCs at nearly all z-positions (Fig. 4G). The maximum width

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Fig. 4. The numbers and lengths of MMCs in wild-type and mutant MOs. (A–C) The x–y plane views of distal (Upper), middle (Middle), and proximal (Lower)regions of a representative MO in wild-type (A), nompC29+29ARs (B), and nompC3 (C). Note in the wild-type MO the membrane is loosely connected to thesheath in the distal region (Upper) and at two sides of MO (along the x axis) in all three planes. Another set of raw ET images for each genotype is provided inSI Appendix, Fig. S8. (Scale bars: 100 nm.) (D) The number of MMCs is lowest at the distal region and increases toward the proximal end in wild-type (n = 4,black), nompC3 (n = 4, blue), and nompC29+29ARs (n = 5, red). The data are presented as mean ± SD. (E) The length distribution of wild-type MMCs. (F) Thelengths of wild-type MMCs at different positions along x axis in the distal, middle, and proximal regions of the MO (blue circles: distal; red squares, middle;green triangles, proximal). The lines are fitting curves to show the location-dependent length change. (G) The average lengths of MMCs change with their z-positions inwild-type (n = 4, black), nompC3 (n = 4, blue), and nompC29+29ARs (n = 5, red). The data are presented as mean ± SD. Note that the average lengthsof MMCs in nompC29+29ARs and nompC3 in all z-positions are longer than wild-type MMCs.

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of MO increased from 268.3 ± 55.2 nm (n = 4 cells) in wild-typeto 406.6 ± 64.9 nm (n = 5 cells) in nompC29+29ARs. Two controlssupported this finding. First, to rule out the possibility that theexpansion of MOs might be an artifact caused by ET samplepreparation, we imaged live campaniform receptors in freshlydissected halteres using spinning-disk confocal microscopy (SIAppendix, Fig. S9). In confocal images, control MOs showed asolid stripe, while the nompC29+29ARs MOs showed an oval ringshape (SI Appendix, Fig. S9) in which the membranes of MOscould be resolved. In the confocal data, the maximum width ofMOs in nompC29+29ARs was 405.5 ± 54.4 nm (haltere pedicel, n =7 cells), consistent with the ET data. Second, to check if thewidening of MOs could be caused by the Gal4 expression system,we measured the MO membrane areas in both wild-type andnompC29+29ARs samples. They were nearly the same [wild type:1.13 ± 0.15 μm2 (n = 4 cells); nompC29+29ARs: 1.09 ± 0.14 μm2

(n = 5 cells)]. Given the similar density of MMCs in these twostrains (Fig. 3F), the total number of MMCs should be similar(also shown in Fig. 4D). Therefore, the Gal4 expression system innompC29+29ARs did not increase the MO membrane area or thenumber of MMCs. Finally, we do not believe that GFP fusion onthe carboxyl terminus of NompC29+29ARs causes the widening ofthe MOs. GFP molecules have a size of about 3∼4 nm andcannot account for the 40-nm increase in the mean length ofMMCs. Thus, the doubling of MMC lengths in nompC29+29ARs

reflects the doubling of NompC-AR in this mutant, consistentwith the previous conclusion that the NompC-AR structurallycontributes to MMCs.We also analyzed the lengths of MMCs in nompC3. If NompC-

AR is not a component of MMCs, then the absence of NompC isnot expected to alter the MMC. On the contrary, if MMCscontain both NompC-AR and non-NompC component, the lossof NompC may change the structure and mechanical propertiesof MMCs, in turn causing length changes. We found that thelength distribution of MMCs was location-dependent in nompC3,similar to wild-type MMCs (Fig. 4C and SI Appendix, Fig. S8).However, the mean lengths of NompC-independent linkers innompC3 were longer than wild-type MMCs at all z-positions (Fig.4G). The maximum width (y axis) of MOs in nompC3 [301.2 ±57.2 nm (n = 4 cells)] was also larger than that in wild-type cells.These results, together with the morphological analysis (Fig. 3J),showed that the NompC-independent linkers distribute in apattern similar to wild-type MMCs, but their structures appear tobe changed due to the loss of NompC.In summary, our observations in nompC29+29ARs confirm that

NompC-AR is a structural component of MMCs that spans thegap between membrane and microtubule. In addition, structuralanalysis on NompC-independent linkers in nompC3 suggests thatMMCs may contain a non-NompC component.

A Candidate Mechanism: MMCs May Bear Tensions. Previous studiesshowed that NompC, as a membrane protein, could bind tomicrotubules in the absence of other binding partners (24). Suchmicrotubule-binding ability relies on the NompC-AR domain(22, 23). Therefore, NompC-AR is expected to span the gapbetween the MO membrane (channel domain) and microtubules(NompC-AR). MMCs are the only structures in the cytoplasm ofMO that may morphologically fit or contain NompC-ARs. Fur-thermore, the double-length MMCs observed in nompC29+29ARs

suggest that NompC-AR is a component of MMCs. Finally,structural analysis of linkers in nompC3 suggests that MMCs maycontain NompC-AR and non-NompC components. Based onthese results, our observations raise a question: Why are MMCsin nompC29+29ARs and nompC3 longer than wild-type MMCs?Among several alternative possibilities (Discussion), a simple

mechanism to account for the length change of mutant MMCs isthat MMCs bear tension. Several observations in the presentstudy agree with this hypothesis. First, most wild-type MMCs

(404 of 416 MMCs) are longer than purified NompC-AR(∼20 nm) in vitro (24), consistent with NompC-AR’s being acomponent of MMCs and MMCs’ being stretched. Second, ifMMCs bear tension, then sheath–membrane contact would beimportant for the stability of this tension as it holds the mem-brane side of MMCs. In the case of loose sheath–membranecontact, MMCs are expected to be less stretched and appear tobe shorter. This agrees with the correlation between the MMClengths and the tightness of sheath–membrane contact (Fig. 4A).Third, if MMCs are considered as spring-like structures, doubledNompC-ARs in nompC29+29ARs can be thought of as two seriallylinked springs, with half of the stiffness of a single NompC-AR(κ2AR = 0.5κAR). Upon the same stretching force (fex), the lengthchanges of doubled NompC-ARs should also double (Δl2AR =2ΔlAR). Therefore, the MMC length is expected to be doubled innompC29+29ARs [l2AR+Δl2AR = 2 (lAR+ΔlAR)], consistent withour observations (Fig. 4G). Fourth, if MMCs are compoundstructures of NompC-AR and non-NompC component, the lossof NompC-AR is expected to reduce the stiffness of individualMMCs (κMMC = κNompC-AR + κnon-NompC). Given that thenumber of linkers is nearly unchanged in nompC3 (Fig. 4D), thecollective stiffness of NompC-independent linkers in the entireMO is expected to be reduced. In this case, the linkers areexpected to be stretched more (Δllinker = fex/κnon-NompC > fex/κMMC) and appear to be longer than wild-type MMCs. Thisagrees with our observations in nompC3.What is the source of tension in MMCs? We suspected that it

may be extracellular forces that stretch the MO membranethrough the sheath–membrane contact and then in turn stretchthe MMCs (Fig. 5A). Are such forces present? To test this hy-pothesis, we performed localized laser ablation experiments onfreshly dissected halteres. The mechanical response of the tissueto laser ablation should test whether the MOs are under tension.We first ablated a single MO in wild-type samples. The struc-tures (cuticle and the adjacent MOs) next to the ablated MOshowed a rapid expansion (Fig. 5B, Upper and Movie S7), re-capitulating a typical elastic relaxation process and therebysuggesting the presence of tension in the MO. Similar observa-tions were made in nompC29+29ARs (Fig. 5B, Middle and MovieS8), suggesting that tension is also present when the NompC-ARis doubled. We then reasoned that if such extracellular forces actin every receptor, there would be tissue tensions across the entirearray of campaniform mechanoreceptors (Fig. 5A). We testedthis by using a laser to ablate a bigger area (around two MOs).The surrounding tissue relaxed rapidly (Fig. 5B, Lower andMovie S9) after the ablation. Interestingly, we noticed that thesurrounding tissue expanded along both the x and y axes, sug-gesting the presence of tissue tension in both directions.

Modeling Analysis: The Moment Activation Mechanism of the MO. Tofurther understand the mechanics of the MO, we took a theo-retical approach, building a mechanical model using the finiteelement method (Fig. 6A). In this model, geometric informationof the large structures was based on the FIB/SEM volume data(Fig. 1). SMCs and MMCs were incorporated based on thestructural analysis on ET data (Fig. 2). The mechanical property(Young’s modulus or stiffness) of each component was primarilytaken from the literatures (SI Appendix, Table S1). The simula-tion results showed that the pedicellar campaniform sensillumconverted cuticle deformation (Fig. 6A) into rotational forces(i.e., a moment, M) on the MO membrane, with a positive stressindicating tension (Fig. 6B). This appears to be due to the ar-chitecture and mechanical properties of supporting structures.As a result, the proximal end of the MO received a greatermoment than the distal end and was strained to a greater extent(Fig. 6 B and C). We termed such a mechanical process “mo-ment activation mechanism.” The resulting forces on MMCswere linearly proportional to cuticular deformation (Fig. 6C).

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With a cuticular deformation at the nanometer scale, the forceon each MMC was at the piconewton level. Because of the un-certainty in estimating the mechanical properties, we simulateddifferent conditions to explore variation in several parameters(i.e., septum, joint membrane, MMC, and membrane). We foundthat the Young’s moduli of the supporting materials (i.e., jointmembrane and septum) and the stiffness of the MMCs wereimportant for determining the resulting forces on MMCs, whilethe membrane contributed little to the overall stiffness (SI Ap-pendix, Fig. S10). However, in all simulation conditions, theproximal region of MO received larger forces than the distalregion, in agreement with the moment activation mechanism (SIAppendix, Fig. S10).

Based on these numerical simulations, we derived a simplified“spring-beam” model to account for the mechanical mechanismunderlying the operation of the MO (Eq. 1) (Fig. 6D). Modeldetails are provided in SI Appendix. Briefly, in this model, yi isthe vertical position of the springs (i.e., MMCs) relative to theneutral axis, E is the Young’s modulus of the beam (cuticle, jointmembrane, and septum), M is the torque produced by the sup-porting structures due to cuticle deformation, l0 is the initiallength of MMCs, κMMC is the stiffness of MMC, w is the thick-ness of the beam, and fi is the resulting force on MMC. In thismodel, cuticular deformation is converted to a torque (M), whichcreates a larger force (fi) on the proximal spring and a smallerforce on the distal spring, consistent with the finite elementsimulation results and essentially accounting for the momentactivation mechanism. This equation shows that the geometry(a, b, w) and Young’s modulus (E) of the beam (i.e., the cuticle,joint membrane, and septum) (Eq. 1) are important for extra-cellular mechanics. It provides a possible explanation for thediversities in the cuticular structures of different campaniformmechanoreceptors, namely that the receptors with differentsupporting structures are designed for sensing different forces. Inaddition, the stiffness (κMMC), distribution (yi) and geometry (l0)of MMCs are important for intracellular mechanics (Eq. 1),demonstrating that the spatial distribution of NompC and themolecular composition of MMCs are both key factors in un-derstanding the gating mechanics of NompC in vivo.

fi =3MκMMCyil0

Ewða3 + b3Þ+ 3κMMCl0�y21 + y22

� [1]

DiscussionIn the present study, we reconstructed the 3D structures of MOsin fly campaniform mechanoreceptors. The MO comprises reg-ularly arranged mechanotransduction apparatuses, each of whichis composed of an MMC which terminates in a button-shapedstructure in the membrane. The AR domain of the NompCprotein contributes to the MMC and the channel component ofthe protein likely contributes to the button (Fig. 7). In addition,the MMC may contain a NompC-independent component.Mechanical modeling of the campaniform sensillum shows thatcuticular deformation is converted to a rotational force that actson the MO (Fig. 7). Our structural and mechanical analyses showthat the predicted force distribution on the MO matches wellwith the spatial distribution of NompC channels, suggesting thatthe MO may have an optimized structural–mechanical design.We now justify these conclusions and their potential implicationsfor understanding fly mechanotransduction.

Ultrastructural Distribution of NompC in the MO. We observed a 2Darray of button-shaped structures on the MO membrane (Figs.2D and 7). Based on the size comparison and mutant analysis, weconclude that the channel domains of NompC structurally con-tribute to the bulk of these buttons. This implies that the densityof NompC in MOs is about 2,000 μm−2 (Fig. 7). Such a highdensity of ion channels is similar to that of voltage-gated chan-nels at the axon initial segment and likely to be functionallyimportant in ensuring that the electrical responses are large andthat the signaling is robust.We found that, due to the MO geometry, the number of

NompC channels first increases from the distal end of the MOand then stays relatively constant from the middle region to theproximal end (Fig. 7). Modeling analysis shows that cuticulardeformation is converted to a mechanical moment that creates alarger force on the proximal end than on the distal end of theMO (Fig. 7). Therefore, our results suggest that more NompCchannels are placed in the regions that receive larger forces,

A fEX fEX

fEX

fEXfEXfEX

Bablated mergedcontrol

wild

-type

nom

pC29

+29A

Rs

nom

pC29

+29A

Rs

xy

zy

xy

y

x

Fig. 5. Tissue tension across the haltere pedicel receptor field. (A, Left) Acartoon schematic showing the potential extracellular tension (fEX) on theMO along the y axis. (A, Right) A y–z plane image showing the directions ofthe extracellular tension in the receptor array. (Scale bar: 1 μm.) (B) Laserablation experiments: both wild-type and nompC29+29ARs were used in thisexperiment. The observations on two strains are similar. Note that in wild-type we used the auto-fluorescence of haltere cuticle to locate the focusplane for visualization and laser ablation. In the nompC29+29ARs strain, thepedicellar MOs are visible due to the expression of NompC29+29ARs-GFP.(Upper) Single MO ablation on a wild-type haltere (Movie S7). (Middle)Single MO ablation on a nompC29+29AR haltere (Movie S8). (Lower) Two MOsablation on a nompC29+29ARs haltere (Movie S9). The cuticular structuressurrounding the ablated MOs in haltere receptor field show instantaneousexpansion in all three experiments. The white arrows indicate the movingdirections of adjacent MOs after the laser ablation. In the merged images,the red channels are the images of the MOs taken right before the ablationand the green channels are immediately taken after the ablation. The mis-match of red and green channels in the merged channel indicates thestructural movements after the laser ablation. Note that in the case of twoMOs ablation in nompC29+29ARs, the shift away of the adjacent MOs occursalong both x and y axes, suggesting the presence of extracellular tensionthroughout the receptor array in both directions. (Scale bars: 2 μm.)

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which we propose increases the sensitivity of the MO. However,the NompC channels on the distal region could make additionalresponses in case of stronger stimuli, thereby conferring a broaddynamic range to the MO. In this way, the MOs could makegraded responses to a wide range of stimuli.

MMCs Have both NompC-AR and Non-NompC Components. Our re-sults provide several lines of evidence to confirm a previouslyproposed argument that NompC-AR contributes structurally tothe MMCs (10, 22). First, the button-shaped structures on MOmembrane resemble the channel domain of NompC in size, areabsent in the nompC3, and are rescued in nompC29+29ARs, sug-gesting that the channel domains of NompC structurally con-tribute to the bulk of these buttons. These buttons are aligned tothe arrayed microtubules in MO cytoplasm through MMCs. Thisis consistent with the molecular picture in which the channeldomain and the AR domain of NompC contribute to the button-shaped structure and MMC, respectively (Fig. 7). Second, dou-bling the AR domain in nompC29+29ARs doubles the MMC lengths.In a previous study, the length of MMCs in nompC29+29ARs (18 ±5 nm) was found to be only slightly longer than wild-type MMCs(15 ± 5 nm) (22). This is likely due to the differences in samplepreparation and imaging methods. Double-length MMCs re-flect the doubling of NompC-AR and confirm that NompC-ARstructurally contributes to the entire length of MMCs. Third,most wild-type MMCs show filamentous shapes and havebranched ends. However, the length and morphology oflinkers are altered in nompC3. This correlates with the lossof NompC-ARs.

NompC-independent linkers are poorly preserved in glutaraldehyde-fixed samples, so in the previous work we were not sure if theywere random denatured protein filaments or real componentof MMCs (10). The observation of regularly spaced NompC-independent linkers in ET data suggests that they are not ran-dom denatured protein. It is formally possible that these linkersare non-NompC molecules that “invade” the MOs only in theabsence of NompC. However, their similar density to wild-typeMMCs suggests a similar binding pattern to the arrayed micro-tubules in theMO. Therefore, the presence of non-NompC linkerssuggests that MMCs may contain non-NompC components. Basedon our previous immunostaining data using an antibody againstthe amino terminus of NompC (12), we think that these linkers arenot fragments or other isoforms of NompC that may be expressedin nompC3.NompC molecules can bind to microtubules on their own and

this interaction depends on NompC-AR (23, 24). Interestingly,NompC-independent linkers show direct connections to bothmembrane and microtubule. If NompC-independent linkersrepresent the non-NompC component of MMCs, this wouldsuggest a molecular picture in which NompC-AR and the non-NompC linkers together form MMCs (Fig. 7). In this model,NompC, in addition to being a mechanotransduction channel,contributes to the mechanical coupling in the mechanical sig-naling pathway. In the absence of NompC, mechanical couplingbetween membrane and microtubule could still occur, but likelywould be less stiff. Therefore, if there are additional mechan-ically sensitive channels, as is the case in the fly’s auditory cells(27, 28), mechanosensitivity in nompC null mutants might beadditionally reduced due to effects on mechanical coupling.

MEκMMC

x

y

h

a

bO

l0

y1

y2

C D

stimulus (nm)

forc

e (p

N)

proximalmiddledistal 200

100

-200

-100

sheath

EDM

MMC

membrane

SMC

A (10-4×GPa)B6.8

-2.012.0

2.8

proximal MMC

Mstress (10-2×GPa)

u

distal MMC

Fig. 6. Mechanical modeling of the pedicellar campaniform sensillum. (A) Modeling schematics; all structural components are indicated (see SI Appendix formodeling details). The mechanical rigidities used in the initial simulations: cuticle, 2 GPa; sheath, 2 GPa; SMC, 2 GPa; membrane, 30 kT; MMC, 16 pN/nm;septum, 0.15 GPa; joint membrane, 0.5 GPa. The mechanical constraint from tubular body is modeled as elastic springs in three axes (only two of the threesprings are shown here). The deformation stimulus is indicated as u. The red arrow indicates the symmetry axis. (B) A representative simulation result in whichcolor encodes stress (gigapascals) on each point. The MO is strained by a mechanical moment (M, i.e., a rotational force). Color bars used for the completesensillum (Right) and the enlarged MO (Left) indicate different scales. A distal MMC and a proximal MMC (indicated by two dashed boxes in the enlarged MO)are further enlarged, showing that the proximal one withstands a larger force. Note that the MO can be strained by stretching or compressive forces. A typicalsimulation with a stretching force (u = 60 nm) is shown here as an example. (C) The deformation-force curves of the most distal (blue), the middle (red), andthe most proximal (black) MMCs in the MO are shown. Note that the proximal end withstands larger forces. The horizontal axis represents stimulus(nanometers) and the vertical axis represents the resulting forces on the MMC (piconewtons). (D) A simplified spring-beam model to account for the momentactivation mechanism. The x axis is the neutral axis and O is the origin of the neutral axis. Further modeling details are provided in SI Appendix. Note that thiscartoon is only 2D, so w (i.e., thickness of the beam) is not labeled in this schematic.

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Future work to identify the molecules that form NompC-independent linkers and to dissect the structural-mechanics ofthe MO in fly’s auditory cells is necessary.

MMCs May Be Under Tension.Alternative mechanisms.We found that wild-type MMCs in vivo aregenerally longer than NompC-AR in vitro (24), MMCs arelonger in nompC3 and nompC29+29ARs, and MMCs showlocation-dependent lengths. To understand these findings, weproposed a model in which MMCs in vivo are under tensionand thereby are stretched. We now discuss some alternativemechanisms.First, could it be possible that NompC-AR in purified NompC

is compressed due to the presence of nanodisc, while MMCs invivo are unstressed so that they appear to be longer? Reconsti-tution in nanodisc stabilizes the transmembrane domains ofmembrane proteins. However, NompC-ARs are cytoplasmicdomains and thereby not likely to be axially compressed. Fur-thermore, MMCs in vivo being unstressed would predict thatthey do not contribute to the mechanical feedback in keeping themembrane–microtubule distances in the MO. If this were thecase, one would not expect any changes in the MMC lengths andMO sizes in nompC3 and nompC29+29ARs. Therefore, MMCs’being unstressed in vivo is inconsistent with our observations.Second, could the increases in the lengths of MMCs in nompC

mutants be the secondary effects of dysfunctional mechano-transduction (e.g., disrupting NompC function)? Our observa-

tions in nompC3 and nompC29+29ARs do not support this hypothesis.NompC29+29ARs was reported to have a similar mechanosensoryresponse to mechanical stimuli in comparison with wild-typeNompC (22). We confirmed that the number of NompC29+29ARs

in the MO of nompC29+29ARs cells is similar to wild-type. Using abehavioral assay, we found that NompC29+29ARs rescues, to a largeextent, the phenotype observed in nompC3: nompC29+29ARs fliescould walk fairly well and showed moderate flying ability (SI Ap-pendix, Fig. S11). If MMCs’ being longer is a nonspecific conse-quence of disrupting NompC function, one would expect minorstructural changes in nompC29+29ARs (partial rescue) and greaterstructural changes in nompC3 (loss of function). However, this isinconsistent with our observations that structural changes in theMMCs and MOs in nompC29+29ARs are greater than in nompC3.Third, could the MMCs at different locations have different

molecular compositions, for example more or fewer repeatingunits, so they show different lengths? This would require MMCswith different compositions to be patterned in the tiny space ofthe MO (0.5 × 1.0 × 0.3 μm3) along both x and z axes. Preciselyarranged structural (e.g., cytoskeleton- or matrix-associated)cues might contribute to such differential molecular specifica-tion. However, membrane–microtubule distance is unlikely theunderlying cue as this distance can be changed when the com-ponents (e.g., NompC-AR) of MMCs are altered. We also donot think chemical cues (small diffusive molecules) can be thepatterning signals as free diffusion would rapidly eliminate thegradient in this small space.Structural and functional implications. Given the stiffness of eachNompC-AR to be at the order of 1 pN/nm, 20-nm extensionpredicts the stretching force to be around 20 pN. Such a force isable to straighten the 24 ankyrin repeats in human Ankyrin-R,while larger forces can partially unfold the ankyrin repeats (29).Our analysis raises a question of how the NompC-AR bundleresponds structurally to this force. One possibility is that the ARdomains of NompC in vivo are present in a straightened orpartially unstructured form and undergo stretching/relaxing cycleswhile halteres are rapidly beating. In this view, the NompC-ARwould behave similarly to other mechanosensitive proteins, liketalin in the focal adhesion complex (30) and titin in muscle (31).Another interesting issue is how tension may regulate the

gating of NompC. We consider two possible scenarios. First, ifthe NompC channel is sensitive to pulling or pushing forces fromMMCs along the axial direction, the resting tension maymechanically offset the channels’ sensitivity to gating forces.Second, if the NompC channel is sensitive to changes in mem-brane tension, the channels’ responses will be symmetric if thereis zero resting tension in the MMC because both pulling andpushing forces will lead to an increase in membrane tension. Thepresence of resting tension in MMCs will make the channels’responses asymmetric, so these channels can be both excitedand inhibited.

Moment Activation Mechanism. Our modeling studies, based onthe architecture and the mechanical properties of the supportingstructures, suggest that cuticular deformation will lead to rota-tion of the campaniform mechanoreceptor so that the strain onthe MO is not uniform but is larger at the proximal end. We termthis the moment activation mechanism. This conclusion holdsover a wide range of mechanical parameters. One caveat, how-ever, is that our numerical simulations and mechanical model areessentially static. Haltere campaniform mechanoreceptors,which detect forces during flight, sense highly dynamic signals.Therefore, our modeling analysis on MO mechanics cannot fullyreveal the complexity of dynamics at haltere, sensillar, and sub-cellular levels but provide an intuitive and qualitative descriptionon how the MO in fly haltere campaniform mechanoreceptorsmay operate.

y z

M

M

yx

distal

proximal

zx

zx

28 nm

18 n

m

NompC-AR

non-NompC

NompC array microtubule-MMC-NompC

NompC (tetrameric channel domain)

NompC

membrane

microtubule (lateral view) microtubule (top view)

electron dense materialnon-NompC component of MMC

Fig. 7. The spatial distribution of NompC matches the moment activationmechanism. NompC channels form an array on the MO membrane (LowerLeft). Due to the MO geometry, more NompC channels are located on theproximal end than on the distal end (Upper Left). Based on modelinganalysis, we propose a moment activation mechanism in which the proximalpart of the MO receives larger forces than the distal part (Upper Right).Therefore, the spatial distribution of NompC matches the force distributionon the MO. In addition, we also observed non-NompC linkers in the nompC3

mutant, suggesting that MMCs may contain non-NompC components inaddition to the NompC-AR (Lower Right).

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Materials and MethodsFlies.All flies were maintained on standardmedium at 23 to 25 °C.w1118wasused as wild-type strain. nompC3 strain was provided by Martin Göpfert,University of Göttingen, Göttingen, Germany. nompC2, nompC-gal4, andnompC29+29ARs strains were provided by Wei Zhang, Tsinghua University,Beijing, China, originally from the Jan laboratory, University of California,San Francisco (22).

Laser Ablation. Laser ablation experiment was performed using the Micro-point system (a 435-nm laser; Andor) added on the Andor spinning-diskconfocal microscope.

Structure Visualization. The atomic structure of NompC was visualized andanalyzed using Chimera (University of California, San Franciso) (32).

HPF and Freeze Substitution.HPF fixation of fly haltere and freeze substitutionmethods were developed based on a previous protocol (33). The detailedprotocols are described in SI Appendix, Supplementary Methods.

ET. The serial sections (250 or 300 nm) were prepared with a Leica UltracutUCT or Leica EM UC7 microtome (Leica) and collected on Formvar-coatedcopper slot grids. Poststaining was performed with 2% UA in 70% metha-nol, followed by 0.4% lead citrate (EMS 17900). Gold nanoparticles (15 nm)(EMGC15; BBI Solutions) were added to both sides of the sections as thefiducial markers. The dual-axis tilt series ranging from −60° to +60° wereacquired with a FEI Tecnai F30 or FEI Tecnai F20 electron microscope (ThermoFisher Scientific). FEI Tecnai F30 electron microscope was equipped with anAxial Gatan US1000 CCD camera and controlled with SerialEM automatedacquisition software (34). An FEI Tecnai F20 electron microscope wasequipped with a Gatan US4000 (895) CCD camera and controlled with FEIXplore 3D TEM tomography software.

Serial Block-Face Imaging Using FIB/SEM. The sample preparation for FIB/SEMwas similar to that used for ET imaging except for the embedding medium(DurcupanACM, 44610; Sigma). For serial FIBmilling and SEM imaging, a layerof block surface was milled by gallium ion beam and then the block surfacewas imaged using an electron beam with 2-kV acceleration voltages, 0.4-nAcurrent, and 8-μs dwell time on a FEI Helios NanoLab G3 UC FIB-SEM. Aftervolume data collection, the images were imported into Amira, aligned usingDualBeam 3D Wizard module, and exported as a stack of images in TIFformat. The image stacks were then used for structural segmentation.

Structure Reconstruction, Segmentation, and Measurement. Tomograms werereconstructed with the IMOD software package (v4.7) (35). To stitch theadjacent sections in the z axis, the microtubules were traced and used aslandmarks (SI Appendix, Fig. S12). These microtubules were first traced usingcylinder correlation and trace correlation lines modules in Amira (ThermoFisher Scientific) (SI Appendix, Fig. S12). The alignment and stitching wereperformed using the SerialSectionStack module in Amira (SI Appendix, Fig.S12). The structural segmentation and 3D surface generation for both ETand FIB/SEM data were performed in Amira. All structural measurements in3D space were performed in Amira.

ACKNOWLEDGMENTS.We thank Jonathon Howard for the initial support ofthis work and his comments on the manuscript; Tobias Fürstenhaupt fortechnical assistance in ET; Martin Göpfert for the nompC3 strain; Wei Zhang,Yuhnung Jan, and Lily Jan for the nompC2, nompC-gal4, and nompC29+29ARs

strains; and the electron microscopy facility of Tsinghua University and theelectron microscopy facility of MPI-CBG. This work was supported by Na-tional Key R&D Program of China Grant 2017YFA0503502; National NaturalSciences Foundation of China Grants 31671389, 31801129, and 11620101001;Qingdao National Laboratory for Marine Science and Technology GrantQNLM2016ORP0301; and startup funding from Tsinghua UniversityGrant 20151080423. X.L. was supported byMax Planck Partner Group Programand Tsinghua-Peking Joint Center for Life Sciences. L.S. was supported by apostdoctoral fellowship from Tsinghua-Peking Joint Center for Life Sciences.

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