Intracellular and extracellular forces drive primarycilia movementChristopher Battlea,1, Carolyn M. Ottb,1, Dylan T. Burnettec, Jennifer Lippincott-Schwartzb,2, and Christoph F. Schmidta,2

aDrittes Physikalisches Institut, Georg-August-Universität, 37077 Göttingen, Germany; bEunice Kennedy Shriver National Institute of Child Health and HumanDevelopment, National Institutes of Health, Bethesda, MD 20892; and cDepartment of Cell and Developmental Biology, Vanderbilt Medical Center, Nashville,TN 37232

Contributed by Jennifer Lippincott-Schwartz, December 16, 2014 (sent for review August 29, 2014)

Primary cilia are ubiquitous, microtubule-based organelles that playdiverse roles in sensory transduction in many eukaryotic cells. Theyinterrogate the cellular environment through chemosensing, osmo-sensing, and mechanosensing using receptors and ion channels inthe ciliary membrane. Little is known about the mechanical andstructural properties of the cilium and how these propertiescontribute to ciliary perception. We probed the mechanical re-sponses of primary cilia from kidney epithelial cells [Madin–Darbycanine kidney-II (MDCK-II)], which sense fluid flow in renal ducts.We found that, on manipulation with an optical trap, cilia deflectby bending along their length and pivoting around an effectivehinge located below the basal body. The calculated bending rigid-ity indicates weak microtubule doublet coupling. Primary cilia ofMDCK cells lack interdoublet dynein motors. Nevertheless, wefound that the organelles display active motility. 3D trackingshowed correlated fluctuations of the cilium and basal body. Theseangular movements seemed random but were dependent on ATPand cytoplasmic myosin-II in the cell cortex. We conclude that forcegeneration by the actin cytoskeleton surrounding the basal bodyresults in active ciliary movement. We speculate that actin-drivenciliary movement might tune and calibrate ciliary sensory functions.

primary cilium | mechanosensing | ciliary motility | flexural rigidity |active fluctuations

Aliving cell is a dynamic, nonequilibrium system dependenton chemical and mechanical communication with its envi-

ronment. Communication is mediated in many mammalian cellsby the primary cilium, a specialized antenna that typically extendsfrom a cell’s apical surface. The primary cilium, with its specializedand segregated membrane compartment, has emerged as a keysignaling center that transduces mechanical and chemical extra-cellular cues (1–4). Flow detection in the kidney epithelium pro-motes cell homeostasis, whereas defects in sensing or signaling canlead to polycystic kidney disease (5). Forces generated by fluidflow are thought to lead to Ca2+ influx through a transient re-ceptor potential (TRP) family Ca2+ channel in the ciliary mem-brane, polycystin-2 (PC2) (6–9) (Fig. 1A). It has also been shownthat ciliary mechanosensation regulates kidney epithelial cell sizethrough the mammalian target-of-rapamycin pathway independentof Ca2+ transients (10). How do the mechanical properties of thecilium facilitate these cellular responses? Both bending andpivoting could trigger membrane channels. Previous studies haveused fluid flow to estimate primary cilium bending stiffness basedon deformation profiles (11–14), but the precise mechanical prop-erties of the cilium remain unknown. It is likely that mecha-nosensation needs subtle coordination and calibration of theintracellular machinery involving adaptation and feedback mech-anisms reacting to external stimuli, such as is the case in mam-malian inner-ear cells, where active mechanical processes arecrucial for hearing acuity (15, 16). We here applied micromanip-ulation and imaging methods to measure the mechanical proper-ties of primary cilia, their cellular anchoring, and their fluctuations.The core of the primary cilium is an axoneme made up of

microtubules. Nine microtubule doublets extend from the cylindrical

basal body formed by the cell’s mother centriole and provide themechanical backbone of the cilium. Unlike beating cilia and fla-gella (9 + 2 cilia), primary cilia (9 + 0 cilia) lack a central mi-crotubule doublet and cross-linking supports. As a consequence,the doublet microtubules can lose their ninefold symmetry as theyextend away from the basal body. Electron micrographs haverevealed so-called 9v arrangements in distal portions of cilia, inwhich the number of microtubule doublets varies from the usualnine down to a single doublet (17). These fundamental structuraldifferences are expected to make the elastic properties of 9 + 0 ciliadistinctly different from those of 9 + 2 cilia.Most 9 + 0 primary cilia are presumed to be passive sensors

because they lack the dynein motors that drive beating of activecilia (18), with one exception: motile 9 + 0 cilia in the organizernode of early embryos possess dynein arms and drive fluid flow(19). Cilia in the epithelium lining renal ducts are thought topassively respond to flow in the ducts. Surprisingly, we foundactive ciliary fluctuations. Even in the absence of dyneins, in-ternal activity in the cell can cause cilia to move, just as a sky-scraper can sway because of both wind and an earthquake. Themechanical properties of the cilium and its anchoring inside thecell determine the response of the assembly to both external andinternal forces. The distinct nonthermal component of ciliummotion that we document here challenges the view that the cil-ium is a strictly immotile sensor and raises the possibility thatendogenous cilium fluctuations may have a functional role.

ResultsFlexural Rigidity of Primary Cilia. As an essential step toward un-derstanding ciliary mechanosensation, we set out to characterizethe elastic properties of both the cilium and its basal anchor inthe cell. We used an optical trap to exert controlled forces on

Significance

A single primary cilium extends from the surface of many mam-malian cells—often into an aqueous lumen, such as a kidney duct.In kidney epithelial cells, primary cilia are believed to sense fluidflow. This mechanosensory function is critical for proper organfunction. Fluid flow is assumed to deflect cilia, leading to activa-tion of transmembrane ion channels. This study defines the me-chanical contributions of both bending and pivoting at the base tociliary deflection. In addition, we report that active intracellularforces drive ciliary pivoting. This cell-directed cilia movement maybe important for tuning ciliary mechanosensitivity.

Author contributions: C.B., C.M.O., J.L.-S., and C.F.S. designed research; C.B., C.M.O., andD.T.B. performed research; C.B. and C.M.O. contributed new reagents/analytic tools; C.B.analyzed data; and C.B., C.M.O., J.L.-S., and C.F.S. wrote the paper.

The authors declare no conflict of interest.

Freely available online through the PNAS open access option.1C.B. and C.M.O. contributed equally to this work.2To whom correspondence may be addressed. Email: lippincj@mail.nih.gov or cfs@physik3.gwdg.de.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1421845112/-/DCSupplemental.

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individual primary cilia of Madin–Darby canine kidney (MDCK)cells while imaging them with differential interference contrastmicroscopy (Fig. 1 B and C). Cilia develop in nondividing, po-larized MDCK cells in culture after confluence is reached. Cellswere grown to confluence on elastic polycarbonate membranes(PCMs) that were then folded and mounted in a microscopechamber, such that the cilia extended parallel to the specimenplane of the microscope (Materials and Methods). The orienta-tion of cilia in MDCK cells is typically close to normal to theapical cell surface. This orientation is established and maintainedby the anchoring of the basal body in the cell cortex and againstthe nucleus by distal appendages and striated rootlets (20). Themicrotubule doublets of the ciliary axoneme are likely to be themain contributors to the cilium’s flexural rigidity (Fig. 1A). Toassess how the microtubule doublets are mechanically coupled inthe cilium, we first measured its flexural rigidity. For a columnarbundle of rod-like filaments, the flexural rigidity depends stronglyon the degree of lateral cross-linking and varies with the individualfilament flexural rigidity as (21)

EIB ∝NαEIf ; [1]

where E is the Young’s modulus of the (assumed to be) homo-geneous material composing the filaments (here microtubuledoublets), IB and If are geometrical factors, EIB is the flexuralrigidity of the bundle, EIf is the flexural rigidity of an individualfilament (microtubule doublet), N is the number of filaments,and α is an exponent that varies between one and three depend-ing on the cross-linking within the bundle (assumed to be a hol-low cylinder with a wall strength of one filament and a circularcross-section) (21). In the limit of weak cross-linking (α= 1), thebundle rigidity is the sum of the flexural rigidities of the N in-dividual filaments. In the strongly cross-linked regime (α= 3),the bundle rigidity corresponds to the flexural rigidity of a solid,thin shell of circular cross-section. Because n = 9, Eq. 1 predictsthat the degree of interdoublet cross-linking in the ciliumstrongly affects its flexural rigidity. The conspicuous absence ofinterdoublet structures in 9 + 0 cilia suggests that they might bemore flexible than 9 + 2 cilia (22). To directly measure bendingstiffness, we used two different approaches (Table 1). First, in

the bend and relax experiments (Movie S1), we laterally dis-placed the tips of cilia with an optical trap and watched the timecourse of the tip relaxation to its equilibrium position (Fig. 2A).At low Reynolds number (here ∼ 10−6), the relaxation time con-stant depends on the balance between the elastic restoring forceacting on the cilium and the viscous drag produced by the exter-nal medium, leading to a simple exponential relaxation curve(23, 24). For small bending angles, the time constant is relatedto the bending stiffness EIB approximately as

yðtÞ≈ yð0Þe−rt;   where  r=EIk41ζL4 ; [2]

where yðtÞ designates the tip position parallel to the cell mono-layer at time t, L is the length of the cilium, ζ= 2:3× 10 mPa · s isthe drag on a cylinder with the cilium’s dimensions (10 μm inlength and 200 nm in diameter), and k1 ≈ 1:89 is the coefficientassociated with the lowest-order mode in the family of solutionsto the equations of motion for a slender rod with one endclamped and one end free (24). Second, in the bead-attachedbending experiments, we used the optical trap to deflect a ciliumwith the help of an attached silica bead of 1.5 μm in diameter andmeasured the force needed to bend the cilium by a given amount(SI Materials and Methods, Fig. S1, and Movie S2). The disad-vantages of the latter method were (i) that the proximity of therefractile cell monolayer and PCM typically perturbed the forcesignal and (ii) that cell debris accumulated in the trap from theculture medium, which added to the force measurement error.The values that we measured from the two methods agreedwithin a factor of two, with the bead-attached bending experi-ments yielding EIB = 2.5 ± 1.5 × 10−23 N·m2 and the bend andrelax experiments yielding EIB = 3.6 ± 0.8 × 10−23 N·m2.By using optical traps, we avoided uncertainties in the fluid

velocity field that can affect deformation measurements. Valuesmeasured here and those reported in the literature from flowexperiments (Table 1) (11–13), with the exception those in thework by Downs et al. (12), consistently fall in the range of 1–5 ×10−23 N·m2. Reported bending stiffness values for individ-ual microtubules are 0.1–3.4 × 10−23 N·m2 (23, 25–30) (1–10times lower). Given the number of microtubules (n = 9; ex-pression 1) and the reported bending stiffness of microtubules,we, therefore, conclude that α≈ 1. In other words, a weak-couplingmodel best explains the measured values. Although we do notaccount for it, the nine microtubule doublets might each besomewhat more rigid than a single microtubule. However, strongcoupling would lead to an about 10-fold stiffer response, whichshould be easily distinguishable. Weak coupling is consistent withthe lack of interdoublet structural proteins in the primary ciliumas well as reports showing the gradual loss of columnar order inthe primary cilium along its length (17). Assuming weak coupling(α= 1 in Eq. 1), we estimate the stiffness of a single microtubuledoublet to be 0.3–0.4 ± 0.2 × 10−23 N·m2

—approximately one-third of the only other estimate of doublet stiffness of which weare aware [1.4 × 10−23 N·m2 measured from the deformation ofdemembranated sperm flagella doublets under flow in thepresence of 0.1 mM ATP (31)]. The stiffness values that wemeasured for the primary cilium are 100–1,000 times lower thanthose of sperm flagella, consistent with the highly cross-linkedstructure of the latter (31, 32).

Viscoelastic Anchoring of Primary Cilia. When we closely examinedthe relaxation curves of cilia from the bend and relax experiments,

Fig. 1. Mechanical structure and anchoring of the primary cilium. (A)Schematic sketch of a primary cilium with PC2 Ca2+ channels and its in-tracellular anchoring. (B) Differential interference contrast (DIC) micrographof a primary cilium deflected in buffer flow of ∼4.8 μm/s. (C) DIC micrographof a primary cilium with an attached bead deflected by an optical trap.

Table 1. Bending stiffness (EIB) of primary cilia (values from this work and referenced published work)

Source Bend and relax Bead-attached bending Schwartz et al. (11) Young et al. (13) Downs et al. (12)

EI value (10 N·m) 3.6 ± 0.8 2.5 ± 1.5 3.1 ± 0.8, 1.4–1.6 8.4 (70%; 1 ≤ EI ≤ 5) 21 ± 4.5, 31 ± 7.0

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we found systematic deviations from single-exponential time de-pendence. Tip relaxation curves could typically be better fit bya sum of two exponentials (Fig. 2B). Initial fast relaxation occurson the order of hundreds of milliseconds followed by a slowerreorientation of the cilium to its resting position normal to the cellsurface (Movie S1). This evidence points to a viscoelasticallyhinged rather than a rigidly clamped boundary condition (BC) forciliary deflection. [By clamped BC, we mean y(z = 0) = 0 anddy/dzjx = 0 = 0, where y measures displacement parallel to the cellsurface and z measures position along the cilium, with z = 0 cor-responding to the cilium attachment point at the cell. In the caseof a limited hinge, the first condition still holds, but the seconddoes not. Rather, deviations in slope away from zero (i.e., normalto the cell surface) are penalized by an elastic energy cost.] Twocharacteristic relaxation times can arise from the effective dragcoefficients for ciliary hinging and bending and their respectiveelastic restoring forces. We can draw a rough conclusion about theratio between internal and external drag coefficients from the factthat we observed two clearly distinct relaxation times. The sloweroverall reorientation dynamics mean that cell-internal viscous dragmust dominate over external fluid drag on the cilium. This fact canbe seen as follows. The amplitudes of the two deflections, ciliumbending and hinge pivoting, were roughly equal. This observationimplies that the effective elastic constants were roughly equal aswell, with the bending elastic constant, of course, depending oncilium length. Although the bend relaxation is only determined bythe cell-external drag, the pivoting relaxation feels both internaland external drag. If the internal drag was small and the relaxationwas dominated by external drag, then the two relaxation timesshould be close to equal. Because they are not, the cell-internaldrag must dominate over the pivoting relaxation.Based on these results, we conclude that cilium deflection

under external force involves both axoneme bending and ciliumbase pivoting. We extracted the bending stiffness values quotedin Table 1 by fitting EIB to the fast decay time constant, becauseit is evident from Movie S1 that the backbone curvature relaxeson the fast timescale. However, the existence of a hinge at thecilium base modifies the coefficient k1 in Eq. 2 and leads toa systematic underestimate of the values given in Table 1 by anamount dependent on the ratio of the cilium bending stiffness tothe compliance of the hinge at its base. From our estimates ofthis ratio, we conclude that the values for the bend and relaxexperiments in Table 1 represent an underestimate by 10%or less.

Spontaneous Angular Fluctuations of Primary Cilia. Even in theabsence of external forcing by fluid flow or the optical trap,cilia exhibit angular fluctuations and, if long enough, detectablebending fluctuations. Given the micrometer-length scale of cilia,these fluctuations could be purely thermal. We recorded thesespontaneous fluctuations of cilia with high temporal resolution.We then reconstructed the backbone of the cilium for eachmovie frame using a custom-written MATLAB algorithm (Fig.3A). An overlay of 6,500 backbones from one such movie is

shown in Fig. 3B. The cilia that we analyzed moved withina limited range of angles over time, sweeping out a circularsector and giving the appearance of a stiff rod rotating ona limited hinge. To determine whether a hinged rigid rod is anaccurate description of the spontaneous cilium motion as op-posed to a bending rod with a fixed attachment angle, we plottedthe angle between each point on the cilium and a reference linenormal to the apical cell membrane in each frame. For the caseof a hinged stiff rod, this angle would be constant along the rod,whereas a flexing rod would exhibit a distribution of angles alongits length because of the nonzero curvature. The angles con-vincingly collapsed onto a single value (SI Materials and Methodsand Figs. S2–S4), which leads us to conclude that the maincomponent of the spontaneous motion of short cilia (<10 μm) isa rigid rod pivoting around a hinge point inside the cell. The factthat internal flexing of the cilium was not detectable in the ab-sence of strong external perturbations makes sense given theabsence of dynein and the stiffness values reported above: purelythermal bending for a 10-μm-long cilium with a flexural rigidityvalue of 3 × 10−23 N·m2 and a clamped end at the cell surfacewould only result in average tip displacements on the order of200 nm (hd2i= kBTl3=3EI, where d2 is the mean squared dis-placement) (33), much less than the observed overall tip dis-placements (∼2 μm in Fig. 3B).

Actively Driven Nonthermal Fluctuations. The evidence that ciliapivot around a hinge within the cell in the absence of an externalforce suggests that the orientation and the motion of primarycilia may be coupled to the mechanical activity of the cell cortex.To test for regular patterns in the motion of cilia, we establishedan additional assay that enabled us to track ciliary motion in 3D.We imaged confluent layers of MDCK cells with fluorescentlylabeled cilia grown on a coverslip with a spinning disk confocalmicroscope in an en face geometry. To maximize temporalresolution, we undersampled focal planes in the z dimension.

Fig. 2. Biexponential relaxation curves of cilia deflection. (A) Relaxationcurves of cilia of different lengths from the bend and relax experiments. Ciliaof different lengths relax with different time constants, which was expectedfrom Eq. 2. (B) Relaxation curves of cilia between 9 and 10 μm in length. Thered line is a double-exponential fit to these curves (τ1 = 120ms; τ2 = 1,500 ms).

Fig. 3. Spontaneous fluctuations of primary cilia. (A) Differential in-terference contrast micrographs of a primary cilium viewed sideways withthe tracked backbone superimposed. (B) The 6,500 superimposed backbonesfrom one cilium tracking experiment recorded at 25 Hz. (C) Maximum in-tensity projection of an en face confocal stack containing several smooth-ened yellow fluorescent protein (YFP)-labeled cilia recorded at 2 Hz. Tomaximize temporal resolution, widely spaced z planes were imaged, corre-sponding to the bright dots in the projection (Movie S3). (Inset) Distributionof eccentricities for 34 cilia in the xy plane 4 μm above the cell surface. (D) 3Dreconstruction of a cilium from the confocal stack in C at higher spatialresolution. The reconstruction consists of 17 planes 1 μm apart. Axes aremarked in micrometers (Movie S4).

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As a result, each cilium looks like beads on a string in a maximumintensity projection of a single stack (Fig. 3C and Movie S3). Weused the SpotTracker ImageJ plugin (34) to track time trajectoriesin each confocal plane, then reconstructed the cilium backbonealong three dimensions with MATLAB, and examined positionfluctuations over time (Fig. 3D and Movie S4). To compare the3D data to the 2D data, we projected the movement of the ciliumalong one axis (x or y) and analyzed the variance along the lengthof the cilium. Plots of the transverse variance along the ciliumlength revealed no significant difference between the side-viewexperiments and the en face experiments (Fig. 4A) and con-firmed the hinged pivoting motions. We performed confocalimaging experiments at room temperature as well as 37 °C andfound similar behavior (Fig. 4A).The en face experiments allowed us to look for anisotropy of

the 9 + 0 ciliary motion, which could reflect constraining an-choring conditions or anisotropic active driving. It is known that9 + 2 motile cilia perform strongly anisotropic motions, withtheir beating stroke occurring in a plane defined by their singularbasal foot, which extends from the basal body (35). Basal feet inprimary cilia vary in number from two to five (36) and have beenproposed to stabilize the cilium (Fig. 1A). We conjectured thata particular arrangement of basal feet could lead to anisotropicmotion in the primary cilium as well. To test for anisotropy, weanalyzed thresholded maximum intensity projections from con-focal movies in a plane 4 μm above the apical cell surface. Weobserved no obvious regular pattern, such as motion on a circleor arc. We next determined the eccentricity « of the best-fit el-lipse to the maximum intensity pattern in the intersecting planeand plotted a histogram of the eccentricity values (Fig. 3C, Inset).For perfectly isotropic motion, « = 1, whereas for motion entirelyalong one dimension, « = 0. We see a broad distribution ofvalues with an average « = 0.79 ± 0.12, indicating, at most, weakanisotropy in the cilium motion (eccentricity is a biased in-dicator, and any noise leads to values of « < 1). The embeddingof the cilium in the cell through its basal feet, thus, does notcreate a favored plane of motion, or there is enough variation inthe number and arrangement of feet among cilia that no cleartrend emerges.Because cilia are microscopic in size, thermal Brownian mo-

tion will be at least part of their observed fluctuations. However,it is possible that cilium motion has an actively driven compo-nent, despite the fact that they contain no dynein motors. Cellsare nonequilibrium systems with vigorous cytoskeletal dynamics.Aggregate random forces generated by stochastic motor activityhave recently been documented to stir the cytoplasm (37) andenhance the transport of small proteins and organelles in thecytoplasm (38). The basal body is embedded in the actin cortex(Figs. 1A and 5), and recent work showed that focal adhesionproteins localize near the basal body and indicates a link betweencilia and the myosin-containing actin meshwork in the cell cortex(39). Because we have established that pivoting around a cell-

internal hinge is a key aspect of primary cilia movement, corticalactin–myosin fluctuations could exert forces at or above the ef-fective hinge, which could then be translated into angular motionof the primary cilium.To test this possibility, we analyzed cilium motion in the pres-

ence of biochemical perturbations. We quantified cilia movementin the side-view geometry by calculating the angular mean-squareddisplacement (MSD). We found that, in unperturbed control cells,the angular MSD grew subdiffusively with time, with a power lawexponent of ∼ 0:66 (Fig. 4B). Subdiffusive exponents < 1 areexpected generically in thermally driven, viscoelastically confinedsystems, where Brownian motion is influenced by the elasticity ofthe surrounding matrix (40). In the cell cytoskeleton, one wouldexpect to see thermal motions only at short times of 100 ms or less,whereas at longer times, nonthermal, actively driven cytoskeletalfluctuations typically take over (41). Direct methods to test foractive cytoskeletal driving are to either deplete the energy supplyor disable specific motor proteins. Intriguingly, both depletion ofATP and treatment of ciliated MDCK cells with blebbistatin,a nonmuscle myosin II motor inhibitor (42), led to a significantdrop in the overall angular MSD amplitude of ciliary motion.Furthermore, both treatments resulted in a much lower power lawscaling exponent of ∼ 0:21 (Fig. 4B). The large drop in fluctuationamplitudes is indicative of a strong active component in themotion of primary cilia. We found a comparable drop in ciliaryfluctuations on ATP depletion in the confocal en face experimentsat 37 °C (Fig. 4A), indicating that the activity is not dependent ontemperature or geometry. Because ATP depletion and blebbista-tin treatment led to identical suppression of spontaneous fluctu-ations, we surmise that actin–myosin activity—acting on the ciliumbase and the basal body, which is embedded in the actin cortex—translates cellular fluctuations into ciliary movements.

Cilium and Centrosome Fluctuations Are Positively Correlated. Thecilium grows from the mother centriole, which forms the basalbody, and is, in turn, connected to the daughter centriole and thenucleus through bundles of microtubules, the striated rootlets,and the striated connector (Fig. 1A) (36). This whole complexis embedded in the actin cortex during ciliogenesis (Fig. 5) (20).To localize the pivot point in this complex, we tracked bothcilium and centriole positions simultaneously in cells expressingemerald–melanin-concentrating hormone receptor-1 (a ciliarymembrane protein) and monomeric red fluorescent proteinfused to the pericentrin-AKAP450 centrosomal targeting domain(mRFP-PACT; a centrosome marker). If the hinge point preciselycoincided with the basal body, we would expect no correlation be-tween cilium and basal body fluctuations. However, if the hingepoint was below or above the centrioles, the fluctuations would be

Fig. 4. Analysis of ciliary fluctuations. (A) Variance of the cilium’s transverseposition along its length for side-viewed cells imaged in differential inter-ference contrast (blue circles, n = 14) and from confocal stacks of fluorescentlylabeled cells at room temperature (RT; 21 °C) and 37 °C and ATP-depletedcells at 37 °C (yellow, n = 10; red, n = 6; and green, n = 18, respectively). (B)Angular MSD vs. lag time for control (blue, n = 14), ATP-depleted (green, n = 9),and blebbistatin-treated (Blebb; 50 μM; red, n = 9) cells.

Fig. 5. EM of a membrane-extracted apical domain of an MDCK cell. 3Dstereo image shows the base of a single primary cilium (asterisk) embeddedin a network of actin filaments (red arrows). (Scale bar: 500 nm.)

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correlated or anticorrelated, respectively. PACT labeling was visibleon both the daughter centriole and the basal body; however, tomaximize temporal resolution, we recorded only the focal planecontaining the basal body. Fig. 6A shows a time series of fluctua-tions for a cilium–centrosome pair, and Fig. 6B shows the associatedcorrelation plot. In the majority of cases, cilium and centrosomefluctuations were well-correlated (rx,avg = 0.60 ± 0.43 and ry,avg =0.78 ± 0.09); their Pearson correlation coefficients are listed inTable 2. Because the correlations in all but one case are positive, weconclude that the hinge point of the ciliary motion must be locatedbelow the centrosome, somewhere closer to the nucleus, such asdepicted in Fig. 6B, Inset.

DiscussionDeflection of the primary cilium in MDCK kidney epithelial cellsoccurs through a combination of bending and pivoting. Wecharacterized the flexural rigidity of cilium bending and de-termined that the pivot point for the limited hinge is locatedbelow the basal body. In contrast to the current paradigm thatprimary cilia are immotile, our experiments revealed that pri-mary cilia perform cell-directed active movements. Fluctuationsin the actin–myosin network seem to act above the pivot point tocause correlated centrosome and cilia movements.The primary cilium is soft compared with motile axonemes.

The bending stiffness of 9 + 0 cilia is only on the order of 1–10times that of an individual microtubule, whereas 9 + 2 axonemesare 100–1,000 times stiffer than primary cilia. The bendingstiffness of the cilium and the mechanical nature of its attach-ment to the cell body will affect functional aspects of mecha-nosensing, such as cell membrane tension or shear strain in thewhole organelle. These parameters, in turn, should affect thelocation and gating of mechanosensitive channels, about whichthere has been some debate (6, 7, 43, 44). Our results allow us tospeculate further about plausible sensory mechanisms. Becausethe system has a hinged BC, it has an additional rotational de-gree of freedom compared with a system with a clamped BC, andtherefore, tension in the ciliary membrane caused by externallyimposed curvature should be lower at a given tip deflection thanin the clamped case. The additional rotational degree of freedomallows the cilium to deflect farther in a given flow field beforebending. Whereas mechanosensitive channels at the base couldbe triggered when the cilium pivots, greater flow forces would berequired to trigger channels along the length of the cilium. Thus,we speculate that channels located at the base might be moreresponsive to weak flow than channels located along the ciliumlength. In addition, channels might be gated by cytoskeletalattachments through shear between the plasma membrane, theECM, and the axoneme.Transmission of force from the actin cortex to the cilium might

occur through simple steric interactions or specific connections

to the cytoskeleton through striated rootlets (20) or basal body-associated focal adhesion proteins (39, 45). The hinging degreeof freedom could be important for other cellular processes, suchas ciliary orientation in migrating fibroblasts, which have beenfound to align their cilia in their direction of motion (4). Thepreviously unidentified actively driven cilium motion quantifiedhere has an origin that is very different from the dynein motorsdriving beating in 9 + 2 cilia. The correlation between ciliaryfluctuations and both myosin II activity and basal body move-ments shows that primary cilia motions report underlying corticaldynamics. One could compare the primary cilium with an an-tenna that has both receiving and transmitting functions. Thequestion remains what the physiological role of such activitymight be. A very basic function might be to establish andmaintain the overall orientation of the cilium normal to theapical cell surface. The existence of cell-directed pivoting raisesthe intriguing possibility that active fluctuations could be in-tegrated into feedback loops that have a functional role in tuningthe response to extracellular forces, possibly similar to what isknown to occur in inner-ear hair cells (15, 16). Speculatively,cell-directed ciliary pivoting could gate base-located ion chan-nels, leading to a rise in ciliary calcium levels, which could bringsignaling pathways closer to the threshold for activation, effec-tively calibrating and sensitizing the response to flow. Interest-ingly, previous reports have shown that myosin II motorinhibition with the Rho-kinase inhibitor Y27632 abolishes theflow-induced calcium response of murine epithelial kidney cells(46). Furthermore, there is evidence that the Lkb1 pathway inmammalian target-of-rapamycin signaling is locally activated atthe cilia basal body through ciliary flow sensing, highlightinganother possible physiological role for ciliary fluctuations (10).The connection between the actin cytoskeleton and primary

cilia is just beginning to be appreciated. A screen for proteinsthat effect ciliary length revealed that depletion of actin regu-latory proteins significantly reduces cilium formation (47). Inaddition, that study showed that inhibiting actin polymerizationwith cytochalasin D promotes cilia formation (47). The challengenow is to understand the integrated function of the primarycilium and the dynamic actin–myosin cell cortex.

Materials and MethodsCell Culture. MDCK-II cells (a gift from Andreas Janshoff, Georg-AugustUniversität, Göttingen, Germany) were cultured in MEM with 2 mM L-glu-tamine, 1% penicillin-streptomycin, and 10% (vol/vol) FBS. Cells for theedge-viewing experiments were imaged 1–2 d after full confluence (5–7 d afterseeding) on poly-L-lysine–coated PCMs. For confocal experiments, cells werecultured on LabTek cover glass chambers in MEM with 3 mM L-glutamine and960 μg/mL Geneticin (G-418) for selection of transfected cells.

Transfection. Cells for the confocal experiments were stably transfected withsmoothened YFP (48), Smoothened Tomato (49), or emerald–melanin-con-centrating hormone receptor-1 (created by Michael Davidson, National HighMagnetic Field Laboratory, Florida State University, Tallahassee, FL), and thoseused in the centrosome-tracking experiments were additionally transfectedwith mRFP PACT (created by Alex Ritter, National Institutes of Health,Bethesda). Transfection was done using the FuGene transfection reagent.

Imaging Chambers for Side-View Geometry. Imaging chambers were con-structed out of a microscope slide and a coverslip sealed together on all sideswith extrathin double-stick tape, creating a chamber ∼80 μm in height. Cellson PCMs were folded over for side viewing as described in ref. 50. PCMswere mounted in the chamber with ∼50 μL medium.

Fig. 6. Correlations between cilium and centrosome fluctuations from con-focal data. (A) Low pass-filtered (LP filtered; 0.33-Hz cutoff) cilium fluctuations(blue) in a plane ∼1 μm above the apical membrane and centrosome (Centro.)fluctuations (red) recorded at a 2-Hz frame rate. (B) Correlation plot of thecilium–centrosome fluctuations plotted in A. Cilium and centrosome fluctua-tions are correlated with a Pearson correlation coefficient value of r = 0.89.(Inset) Schematic of the proposed location of the hinge point and cilium–

centrosome motion. Blue denotes the tracked point at the cilium base (1 μmabove the membrane), and red denotes the centrosome label.

Table 2. Pearson correlation coefficient (PCC) values for cilium–

centrosome fluctuations in x and y directions

Cilium 1 2 3 4 5 6 7 8 9

x PCC 0.84 0.85 0.72 0.06 0.37 −0.22 0.94 0.92 0.88y PCC 0.89 0.86 0.75 0.72 0.68 0.68 0.71 0.80 0.92

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Blebbistatin Treatment. Confluent cells were incubated in 50 μM activeblebbistatin (−) enantiomer (100 μM racemic solution; 203389; CalBiochem)at 37 °C for 30 min and then mounted with blebbistatin-containing mediumfor video-tracking experiments.

ATP Depletion. Cells were incubated in glucose-free DMEM solution for 3 hat 37 °C. Then, Antimycin A and 2-Deoxy-D-glucose were added to finalconcentrations of 10 μM and 10 mM, respectively. The cells were incubated10–15 min at 37 °C and mounted in the glucose-free Antimycin A- and2-Deoxy-D-glucose–containing solution for video-tracking experiments.

Microscopy. Edge-viewing experiments were performed using the custom-built differential interference contrast/opticaltrapping setup described in ref.51. Images were acquired with an MTI analog camera at a rate of 25 Hz andread out through a frame grabber card and a custom-written LabView VI.Confocal experiments were performed with a 63× objective on a MarianasIntelligent Imaging Innovations spinning disk confocal, and frames wereacquired at ∼2 Hz.

EM.MDCK-II cells were cultured on Nunc glass transwells (Thermo Scientific) 2 dafter full confluence. Membranes were extracted, and samples were preparedfor EM by rotary shadowing as previously described (52). Metal replicas were

then separated from the glass by floating them on 10% bleach to removeorganic materials and mounted on EM grids. Images were acquired at 80 kV.

Data Analysis. Tracking of cilia in the confocal data was done using the ImageJplugin SpotTracker (34). Reconstruction and analysis were done in MATLAB.Edge-viewing tracking and analysis were done using custom-writtenMATLAB routines.

ACKNOWLEDGMENTS. We thank Hiroaki Ishikawa, Claus Heussinger,Giovanni Marelli, and Andrej Vilfan for helpful discussion and AndreasJanshoff for providing us with the Madin–Darby canine kidney-II cell line.We acknowledge discussions at the Woods Hole Marine Biology Laboratory.We also thank Monica Bettencourt-Dias, Tristan Ursell, and Kimberly Yasutisfor help with early experiments monitoring cilia movement. Rafael Vilasmilfrom the National Eye Institute Flow Cytometry Core sorted cells for thisstudy. This research was supported by the Intramural Program of the Na-tional Institutes of Health, National Institute of Child Health and HumanDevelopment; the Deutsche Forschungsgemeinschaft through Sonderfor-schungsbereich 803 Project A7 and 937 Project A2; the Center for NanoscaleMicroscopy and Molecular Physiology of the Brain; and European ResearchCouncil Grant PF7 ERC-2013-AdG, Project 340528 (to C.F.S.). Fellowship fund-ing was provided by the Göttingen Graduate School for Neurosciences, Bio-physics, and Molecular Biosciences.

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