Cilia and flagella protrude like bristles from the cell surface. They share the same basic ‘9+2’ axoneme structure, being made up of nine microtubule doublets that surround a central pair of singlet microtubules. Flagella are generally involved in cell propulsion, whereas motile cilia help to move fluids over cell surfaces.

Maintaining cilia and flagella is a challenge for cells, which must find a way to send new proteins all the way along the axoneme to the site of assembly at the flagellar tip. Cells achieve this via a process called intraflagellar transport, in which proteins are carried back and forth by kinesin and dynein motors along the axonemal doublet microtubules. Intraflagellar transport has been proposed to influence other functions of cilia and flagella, including the propulsion of cells over surfaces. However, the details of these interactions are unclear.

Through a combination of biophysical and microscopy approaches, Shih et al. describe the mechanism that the green alga Chalmydomonas uses to power flagellar gliding over surfaces. By tracking single fluorescently tagged molecules, Shih et al. observed that flagellar membrane glycoproteins are carried along the axoneme by the intraflagellar transport machinery. During transport, flagellar membrane glycoproteins transiently adhere to the surface, and dynein motors that were previously engaged in carrying these glycoproteins now transmit force that moves the axonemal microtubules. This process, which is dependent on the concentration of calcium ions in the extracellular environment, generates the force that propels the alga's flagella along the surface.

Gliding motility is thought to have been one of the initial driving forces for the evolution of cilia and flagella. How the intricate mechanism of flagellar beat motility could have evolved has been the subject of much discussion, as it would require the flagellum to have evolved first. In demonstrating that gliding motility is powered by the same intraflagellar transport mechanism that is required for flagellar assembly, Shih et al. provide strong evidence for the evolution of primitive flagella before the evolution of flagellar beating. Furthermore, since algal flagella have essentially the same structure as the cilia of human cells, these findings could ultimately aid in the development of treatments for diseases that result from defects in intraflagellar transport, including polycystic kidney disease and retinal degeneration.

Cilia and flagella are microtubule-based organelles that power the locomotion of many organisms, generate fluid flow over multiciliated surfaces, and mediate cell signaling (Liem et al., 2012). In order to assemble and maintain cilia, ciliary proteins are transported from cytoplasm to the tip by IFT along axonemes (Kozminski et al., 1993). In IFT, linear arrays of multiprotein complexes (IFT trains) are transported by kinesin-2 and dynein-1b in anterograde and retrograde directions, respectively (Cole et al., 1998; Porter et al., 1999). IFT is a universal mechanism for nearly all eukaryotic cilia and flagella, and defects in this process are linked to a wide range of human diseases, including polycystic kidney disease, retinal degeneration (Rosenbaum and Witman, 2002; Ishikawa and Marshall, 2011), and Bardet-Biedl syndrome (Ou et al., 2005; Lechtreck et al., 2009, 2013; Wei et al., 2012).

Several studies have suggested that IFT not only transports material between the cell body and the flagellar tip, but also interacts dynamically with the flagellar membrane (Kozminski et al., 1993) to regulate diverse ciliary functions including motility, mating, sensing extracellular signals and influencing developmental decisions (Huangfu et al., 2003; Snell et al., 2004; Pedersen and Rosenbaum, 2008; Ishikawa and Marshall, 2011). However, it has remained unclear how transport of IFT trains underneath the flagellar membrane transmits force to components at the exterior of the flagellar membrane.

In order to investigate interactions between IFT and the ciliary surface, we used Chlamydomonas reinhardtii gliding motility as a model system. In Chlamydomonas, the flagellar surface is highly dynamic; polystyrene microspheres and other inanimate small objects adhere to and are moved bidirectionally along the flagellar surface, and cells glide over solid surfaces via adhesion of their flagella (Lewin, 1952; Bloodgood, 1981). Gliding motility is central to understanding the function and evolution of cilia, as it may have existed in early cilia before the establishment of axonemal beating. There are indications that gliding and FMG1-B motility are driven by the same process (Bloodgood, 2009), because they move at comparable speeds and both require ligation of the major flagellar surface protein, FMG1-B, into large clusters (Bloodgood and Workman, 1984), accompanied by a Ca²⁺-dependent signaling pathway (Bloodgood and Salomonsky, 1994). Microsphere movement is considered a more easily assayed and quantitated surrogate for the force transduction system that drives whole cell gliding motility. However, gliding is driven by the pulling motion of the leading flagellum, whereas FMG1-B movement is bidirectional (Bloodgood, 2009), implying that the two motilities could employ different motors.

It has been proposed that IFT provides the force for gliding motility (Bloodgood, 2009). IFT trains make multiple connections with the flagellar membrane (Pigino et al., 2009) and carry several ciliary and flagellar membrane proteins (Qin et al., 2005; Huang et al., 2007; Lechtreck et al., 2009, 2013). Inactivation of kinesin-2 in the temperature-sensitive (ts) mutant fla10^ts stops both IFT and gliding motility (Kozminski et al., 1995). While these results suggest that kinesin-2 serves as the anterograde motor responsible for both microsphere movement and gliding motility (Kozminski et al., 1995; Laib et al., 2009), the retrograde motor for these motilities has not been clearly identified. Mutations in the LC8 subunit of dynein do not abolish FMG1-B movement (Pazour et al., 1998), and other flagellar motors, such as the minus-end directed kinesin KCBP (Dymek et al., 2006), have been proposed to drive FSM (Bloodgood, 2009).

Several studies have raised arguments against this model. IFT motility differs significantly from FSM in that trains move faster and more processively along the length of the flagellum (Kozminski et al., 1993; Bloodgood, 2009). FSM requires micromolar levels of free calcium, whereas IFT is Ca²⁺-independent (Kozminski et al., 1993; Bloodgood, 2009). Sexual agglutinins were observed to migrate from the cell body into flagella in the absence of IFT (Pan and Snell, 2002). Therefore, evidence supporting the role of IFT in gliding motility is indirect and the precise functions of IFT, molecular motors, FMG1-B and Ca²⁺ in FSM remain unclear.

To dissect the mechanism of FSM, we directly observed IFT, FMG1-B and gliding motilities using single-molecule imaging techniques. We monitored the movement of individual IFT trains by using total internal reflection fluorescence (TIRF) illumination to image paralyzed-flagella (pf) mutant cells that had adhered to the glass surface with both flagella. To establish a link between FMG1-B transport and IFT, we simultaneously tracked the movement of FMG1-B antibody-coated fluorescent beads (200 nm diameter, dark red) and IFT27-GFP in the pf18 strain. The beads performed short processive runs with reversals of direction whereas IFT trains moved in a regular and unidirectional manner. Multicolor kymography analysis shows that the beads transiently dissociated from one IFT train, diffused for a period of time and then bound to another IFT train (Figure 1A, Video 1) similar to the movement of extraflagellar particles observed along the flagellar membrane (Dentler, 2005). Diffraction-limited images of beads and IFT trains were fitted to a two-dimensional Gaussian to achieve a higher localization precision (Yildiz et al., 2003). Figure 1B shows the colocalization of a membrane-attached bead with an individual IFT train as it moves unidirectionally for >500 nm. The movements of the bead and the colocalized IFT trains correlate strongly (>0.99) with each other during the processive run (N = 30, see Figure 1—figure supplement 1 for more examples), excluding the possibility that microspheres coincidentally moved together with IFT trains. We measured the distance between microspheres and the nearest IFT trains moving in the same direction for both the experimental data and randomly generated kymographs. The Kolmogorov-Smirnov test (p=7 × 10⁻⁹, Figure 1C) demonstrates that IFT trains colocalize with FMG1-B along the flagellar membrane (Figure 1D).

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We next investigated whether IFT plays a direct role in gliding motility by monitoring individual IFT trains in IFT20-GFP and IFT27-GFP pf18 cells as they glided on coverslips. A small fraction of IFT trains displayed rare pauses (0.125 s⁻¹ per cell, N_cells = 23, N_pauses = 247) as they moved either in the retrograde or anterograde direction. Remarkably, the pausing of IFT trains was required for the initiation of whole-cell movements (Figure 2A, Figure 2—figure supplements 1 and 2). We did not observe the start of a gliding event without a paused IFT train. We were able to determine the directionality of 60% of the paused IFT trains, and all of the trains correspondent to the initiation of gliding motility stopped moving during retrograde transport (N = 148). To rule out the possibility that IFT pausing events and initiation of gliding motility are simply coincidental, we quantified the lag time between the pausing of the last retrograde IFT train and the initiation of gliding motility. The average lag time was 0.48 ± 0.05 s. In comparison, we observed individual IFT pausing events (including both anterograde and retrograde pauses) every 8.25 ± 0.96 s per cell (Figure 2—figure supplement 3). The Student’s t-test excludes the null hypothesis that the timing of retrograde IFT pausing and gliding motility are independent of each other (p<0.0001). Anterograde trains also displayed rare pauses (∼0.02 s⁻¹ per cell, N = 27), but we never observed pausing of an anterograde train before the initiation of gliding motility.

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In 24% of all cases (N = 50), a single paused train appeared sufficient to pull the entire cell (Figure 2A, Video 2). In other cases, multiple trains paused before the start of gliding motility (Figure 2B, Video 2). Pauses occurred in the leading flagellum, and cells moved in the opposite direction relative to the transport trajectory of the paused trains. In uniflagellate IFT27-GFP pf18 cells, we also always observed pausing of single or multiple IFT trains before the initiation of gliding motility (Figure 2—figure supplement 4). Gliding motility either stopped when paused IFT trains resumed movement (Figure 2A), or continued until the cell bodies reached the paused trains (Figure 2B). We never observed the cell body gliding further than the position of the paused IFT train(s).

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We found compelling evidence that dynein-1b is the primary motor responsible for gliding motility. First, uniflagellate pf18 cells always glided with the flagellum leading the cell body at 1.49 ± 0.10 µm/s (mean ± SEM) (Figure 2C–E, Video 3) (Bloodgood, 2009), suggesting that gliding motility is driven by pulling forces generated through a minus end directed microtubule motor. In contrast to pf18 cells, uniflagellate dynein-1b ts cells (dhc1b-3^ts) (Engel et al., 2012) were unable to display robust gliding at restrictive temperatures (Figure 2C, Video 3). We observed only short-range (<500 nm) gliding-like motion in 54% of these cells, compared to robust unidirectional gliding motility observed over 5 µm in pf18 cells. These short-range motions were bidirectional (Figure 2D) and significantly slower (forward: 0.35 ± 0.09 µm/s, backward: 0.47 ± 0.16 µm/s) than the gliding speed of pf18 cells (Figure 2E). Second, inactivation of dynein-1b in biflagellate dhc1b-3^ts cells decreased the fraction of gliding cells from 100% to 52% and reduced the gliding speed from 0.86 ± 0.08 µm/s to 0.18 ± 0.03 µm/s. In contrast, inactivation of kinesin-2 in fla10^ts cells resulted in a twofold increase in gliding speed (Figure 2E), indicating that kinesin plays an inhibitory role during dynein-1b driven gliding motility. These results agree with our observations that the pausing of anterograde trains does not initiate gliding motility.

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To further test the role of dynein-1b in gliding motility, we treated the cells with a small molecule inhibitor of dynein, ciliobrevin D (Firestone et al., 2012). We varied ciliobrevin D concentration between 0–150 µM and monitored IFT in cells adhered both of their flagella to surface 2 min after drug treatment (Figure 3A, Figure 3—figure supplement 1, Video 4). Both anterograde and retrograde IFT train frequencies dropped with increasing concentrations of ciliobrevin D (Figure 3B), and at >100 µM ciliobrevin D we observed accumulation of IFT trains at the flagellar tip (see example kymograph in Figure 3A). At 150 µM ciliobrevin D, retrograde IFT frequency was reduced by 92% compared to the 70% decrease observed after 6 hr of heat inactivation of dhc1b-3^ts cells (Engel et al., 2012). The velocities of retrograde and anterograde trains also decreased by 60% and 36%, respectively (Figure 3C). Importantly, inhibition of dynein-1b resulted in a significant reduction in the speed (79%) and frequency (79%) of gliding motility (N = 50, Figure 3D). These results further support our conclusion that dynein-1b motors produce the force for gliding motility.

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Based on these results, we propose a model to describe the functions of IFT motors, IFT trains and FMG1-B transport in gliding motility (Figure 2F). Surface adhesion of the FMG1-B cargo through its large extracellular carbohydrate domain (Bloodgood, 2009) stops the retrograde IFT train. Dynein motors previously engaged in transporting the paused IFT train exert force towards the microtubule minus end, causing the whole cell to move toward the plus-end flagellar tip. Thus, gliding motility works similarly to microtubule gliding assays, in which surface-immobilized dyneins glide microtubules with their plus-end tips in the lead.

To investigate how cells reverse gliding direction, we developed a kymography method for monitoring IFT trains as the cells reorient their flagella during gliding (Figure 4A–B, Video 5). During 60% of reversals (N = 40), the cell lifted one of its flagella and the paused IFT trains on the surface-adhered flagellum drove the motility (Bloodgood, 2009). In other cases, single or multiple paused IFT trains accumulated in one flagellum, and the cell body glided toward this cluster until the paused trains either detached from the glass surface or reached the flagellar base. We also observed cases with paused IFT trains in both flagella where the cell remained immotile, likely due to the balance of forces between bound dynein-1b motors (Figure 4C–D). There were no indications of coordination of the pausing events between the two flagella. Different modes of reversals in gliding motility may allow cells to search through the environment by a random walk when they adhere both of their flagella, and to move in unidirectional manner by lifting one of the flagella.

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Gliding motility in C. reinhardtii is controlled by a Ca²⁺-calmodulin regulated kinase and requires micromolar levels of Ca²⁺ in the media (Bloodgood and Spano, 2002). We tested whether the pausing of IFT trains depends on Ca²⁺ (Figure 5A–B) by analyzing the kymographs of IFT at different Ca²⁺ concentrations. To quantify pausing in flagella, we used Fourier space direction analysis (FSDA, see Figure 2—figure supplement 2) to separate the traces of paused IFT trains and moving IFT trains into two kymographs. Kymographs of paused IFT trains in Figure 5A,B show that pausing along the length of the flagellum was significantly reduced in Ca²⁺-deprived cells (see additional examples in Figure 5—figure supplement 1). After Ca²⁺ depletion, IFT trains rarely paused in the middle regions of flagella and accumulated at the base (Figure 5B, middle). Figure 5C plots the total fluorescence intensity of paused IFT trains (N_cells = 30 for each case) along the lengths of flagella at different Ca²⁺ levels. The frequency of pausing in Ca²⁺-deprived cells was significantly lower than IFT pausing in cells at a normal Ca²⁺ concentration. In regular TAP media (free [Ca²⁺] = 0.34 mM), 95% of all surface-adhered cells displayed gliding motility and individual pausing events were observed every ∼8 s per flagellum, on average. In contrast, both the fraction of gliding cells and IFT pausing frequency were reduced by ∼10-fold at <1 µM Ca²⁺ (Figure 5D). Next, we compared the IFT pausing frequencies in gliding and non-gliding cells at different Ca²⁺ levels. The pausing frequency in non-gliding cells was low and independent of Ca²⁺ concentration. In contrast, the pausing frequency in gliding cells was high at normal Ca²⁺ levels and gradually declined to match that of non-gliding cells as the Ca²⁺ concentration decreased below 1 nM (Figure 5E).

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In Ca²⁺-deprived cells, the beads freely diffused on the flagellar membrane but did not move processively by IFT trains (not shown). Thus, we propose that Ca²⁺ is required for attachment of FMG1-B to IFT trains, not for the activation of a motor protein that provides force for gliding, as previously suggested (Bloodgood, 2009). As the flagellar membrane is enriched with a PKD2-like Ca²⁺ channel (Pazour et al., 2005; Huang et al., 2007), Ca²⁺ signaling at flagellar adhesion sites may play a role in controlling the directionality and timing of gliding movement.

To measure the forces exerted by motors bound to individual IFT trains, we tracked the movement of FMG1-B-antibody coated bead motility using an optical trap. To rule out the possibility that loads exerted by the trap might disrupt the linkage between IFT and FMG1-B, we performed TIRF imaging of IFT27-GFP and optical trapping of beads simultaneously (Figure 6A). We observed that IFT trains remained colocalized with trapped beads when subjected to external forces (Figure 6B; see Figure 6—figure supplement 1 for additional examples). The average offset between the positions of the trapped beads and colocalized IFT trains was 280 ± 10 nm (N = 11). This is due to the fact that the beads (920 nm diameter) and IFT trains (200–1000 nm in length) (Pigino et al., 2009) are large objects relative to the resolution of conventional fluorescence imaging (approximately 200–250 nm) and the forces that stretch the bead-FMG1-B-IFT linkage displace the center of the bead from the IFT train (see the schematics in Figure 6—figure supplement 2). The trap measurements correspond to forces generated by IFT motors and provide direct evidence that IFT transports FMG1-B. Since beads are outside the cell, but are physically linked to the action of IFT trains, the assay combines the advantages of precise in vitro trapping with the ability to manipulate IFT motility under load.

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In a fixed trap assay, ∼50% of processive bead movements terminated with a stall before returning to the trap center (Figure 6C, Figure 6—figure supplement 3). We did not observe a significant difference between the force values of stall and release events. This suggests that motors attached to an IFT train may not be able to reach their maximum stall force, defined as the stall force of a single motor multiplied by the number of bound motors (Shubeita et al., 2008). Histograms of peak forces (Figure 6D) show that anterograde and retrograde IFT trains moved against 21.4 ± 0.7 pN and 25.2 ± 1.3 pN (SEM), respectively. Forces exerted on IFT trains well exceed the force-generation capability of a single motor. Previous studies demonstrated that multiple motors produce larger forces, move with higher velocities under load and carry cargos further than a single kinesin or dynein motor (Mallik et al., 2005; Vershinin et al., 2007). It remains controversial whether motor stall forces are additive at low motor copy numbers (Vershinin et al., 2007) or whether multiple motors tend to transport their cargo using only one load-bearing motor at a time (Jamison et al., 2010). Therefore, we believe that peak forces in our experiment represent a lower boundary for estimates of motor copy number. Assuming that IFT motors produce 6–7 pN forces (Gennerich et al., 2007; Brunnbauer et al., 2010), we estimate that at least four motors transport IFT trains at a time, in agreement with previously reported values (Engel et al., 2009b). It is possible that multiple motor engagements enhance the run length of individual IFT trains, allowing them to traverse the length of a flagellum. In addition, the fast transport of IFT trains in a viscous cellular environment and IFT’s role in FSM may require forces larger than single motor stall forces.

The measured forces in both directions only marginally changed (<20%) under different bead antibody-coating conditions (Figure 6—figure supplement 4). This result argues against the possibility that antibody-coating of the beads leads to the crosslinking of more than one IFT trains, which would be expected to multiply the average peak force. 20% of retrograde runs escaped the trap by producing more than 80 pN of force, presumably via clustering of FMG1-B (Bloodgood, 2009), while <1% escape events were observed in the anterograde direction.

We next investigated why the speed of gliding motility (1.49 ± 0.10 µm/s, SEM) is significantly slower than that of retrograde IFT (2.96 ± 0.14 µm/s), even though both processes are powered by dynein-1b motors actively transporting IFT trains. IFT trains that carry 1-μm beads moved ∼30% slower than IFT trains that were not associated with the beads (two-tailed Student’s t-test, p = 2.1 × 10⁻⁹ (anterograde) and 1.8 × 10⁻¹¹ (retrograde), Figure 7A). We reasoned that the viscous drag of the membrane may slow down IFT trains transporting FMG1-B clusters and led to the observed differences in speed between IFT and FSM. To estimate the drag constant of the bead-IFT complex, we analyzed individual stalling events in the optical trap assay (Figure 7B) and measured the recoil time of the bead after a stall (Figure 7C). The average drag constant of the bead-IFT complex was found to be 8.0 ± 2.8 pN s/μm (SEM, N = 30).

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We hypothesized that the large drag constants we measured were due to the interaction between the bead and the membrane. Our results rule out the possibility that microtubule motors step backwards during the recoiling of the bead, as the average bead velocity was 20 µm/s (an order of magnitude faster than that of dynein and kinesin) and there were no detectable backward steps. To rule out other possible interactions between IFT trains and axonemes, we oscillated individual beads on a flagellar membrane surface in a square wave pattern and measured the recoiling time when the beads are decoupled from IFT (Figure 7—figure supplement 1). The average viscous drag constant was 8.4 ± 4.2 pN.s/µm, which is similar to the drag that beads experience when they interact with IFT trains. Anterograde and retrograde trains moving at 2–3 µm/s would experience 16–24 pN resistive forces, comparable to the total motor force exerted on a single IFT train (Figure 7D). Therefore, IFT is subjected to a high drag force when it carries a large bead along the flagellar surface, which leads to the reduction of transport velocity.

We next investigated how kinesin-2 activity reduces the speed of gliding motility (Figure 2E). This could theoretically be explained by a tug-of-war between active kinesin and dynein motors that are both present on the same IFT train. Alternatively, kinesin-2 and dynein-1b motors may be exclusively active on anterograde and retrograde cargos, respectively, and pausing of anterograde trains could produce forces that oppose the gliding forces of paused retrograde trains. To distinguish between tug-of-war and coordinated transport mechanisms, we examined how inhibition of one class of motors affected the forces exerted on IFT trains traveling in the opposite direction (Laib et al., 2009). At permissive temperatures, the peak forces of IFT in fla10-1^ts and dhc1b-3^ts were in close agreement with wild-type (WT) cells. Heat-inactivation of kinesin-2 reduced the frequency of anterograde runs by 66%, but did not alter the peak forces on retrograde runs (Laib et al., 2009) (Figure 8A). Similarly, heat inactivation of dynein-1b significantly reduced the ratio of retrograde to anterograde transport events (0.15), but had minimal effect on the peak forces of anterograde runs (Figure 8B). These results exclude a tug-of-war mechanism in IFT, which would lead to an increase of force from motors walking in one direction upon inactivation of the motors walking in the opposite direction. Instead, only one type of a motor remains active on IFT trains at a time, which is consistent with the result that retrograde speed of fla10-1^ts does not change after kinesin-2 is inactivated. We propose that kinesin-2 on paused anterograde trains slows down gliding motility by exerting forces in the opposite direction to that of dynein-1b on paused retrograde trains.

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We present direct evidence for the mechanism of IFT-mediated cell-surface interactions using Chlamydomonas gliding motility and FMG1-B transport as a model system. In Chlamydomonas, IFT trains carry FMG1-B as a cargo. The interaction between IFT trains and FMG1-B clusters is transient, as FMG1-B boards moving trains but usually unloads and diffuses away before the trains arrive at the flagellar base or tip. During gliding motility, IFT trains pause due to attachment of the FMG1-B cargo to the surface, and the dynein-1b motors engaged to the paused IFT trains generate pulling forces along the microtubule long axis. While the majority of organismal motilities rely on the actin cytoskeleton or axonemal beating, gliding motility in C. reinhardtii is distinct in that it is powered by intracellular transport machinery along microtubules. By providing surface adhesion points, FMG1-B performs an analogous function to integrins in mammalian cell motility (Bloodgood, 2009; Lecuit et al., 2011).

Our results also demonstrate how different types of flagellar motility in Chlamydomonas have distinct characteristics despite being driven by the same forces. First, we showed that viscous drag of the membrane slows down the motility of IFT trains that carry FMG1-B-conjugated beads, causing gliding motility and FMG1-B transport to proceed slower than IFT. Second, previous work showed that IFT is unaffected by changes in environmental Ca²⁺ concentration (Bloodgood and Salomonsky, 1990; Kozminski et al., 1993), whereas gliding motility is Ca²⁺-dependent (Bloodgood and Salomonsky, 1990; Kozminski et al., 1993). This raised the possibility that Ca²⁺ may be required to activate a specific motor protein that provides force for gliding (Bloodgood, 2009). Our results are inconsistent with this hypothesis. We found that the presence of free Ca²⁺ leads to frequent pauses in IFT motility at flagellar adhesion sites. In the absence of Ca²⁺, IFT moves unidirectionally without interruptions, suggesting that attachment of FMG1-B to IFT trains is regulated by a Ca²⁺-dependent signaling pathway. The signaling pathway responsible for linking FMG1-B to IFT remains unclear. There is evidence that surface adhesion or crosslinking of FMG1-B into large clusters induces dephosphorylation of a transmembrane protein that co-immunoprecipitates with FMG1-B (Bloodgood and Salomonsky, 1994, 1998). The complex may be regulated by a calcium-calmodulin dependent gliding associated kinase (GAK), which is required for the gliding motility (Bloodgood and Spano, 2002). Further investigation is required to identify the rest of the signaling pathway regulating IFT-flagellar surface interactions.

Our force measurements show a number of similarities and differences with earlier published work. In a previous trapping assay, beads nonspecifically adsorbed to the flagellar membrane were found to exert 60 pN average peak forces (Laib et al., 2009). In contrast, our measured peaked forces are significantly lower (20–30 pN) than these values. Laib et al. could not determine whether their force measurements reflect the forces produced by motors attached to IFT trains or by motors that are directly bound to FMG1-B clusters. Our experiments directly link optical trap recordings of bead motility to forces generated by flagellar motors attached to IFT trains (Figure 1), enabling us to propose physical models for the coordination of IFT and its role in FSM. In agreement with Laib et al., we observe reciprocal coordination of motors during anterograde transport. Additionally, our measurements of dynein-1b inactivation verify that motors do not engage in a tug-of-war while travelling in the retrograde direction.

The mechanism we described for IFT pausing and force generation has broad implications for the traffic of ciliary sensory proteins as well as cell signaling at ciliary membrane adhesion points (Wang et al., 2006). For example, IFT is required for signal transduction during the mating of ciliated organisms. The initial interaction between gametes of two mating types in Chlamydomonas can occur at any point along their flagellar membranes, but the tips of their flagella must be aligned and locked before activating gametic fusion (Homan et al., 1987). Since IFT trains transport sexual agglutinins within the flagellar membrane of gametic cells (Ferris et al., 2005), adhesion between sexual agglutinins of opposite cell types may stall retrograde IFT movement. As a result, forces produced by IFT trains would move the flagellar contact points until a balance of forces is achieved when the flagella are properly aligned with respect to each other. Indeed, microspheres have been observed to move and accumulate at the flagellar tips during mating (Hoffman and Goodenough, 1980). We propose that the absence of retrograde bead movement may be related to the formation of adhesion contacts between the flagella and pausing of retrograde IFT trains.

Cilia in the retina, liver, and kidney cells were recently observed to make direct physical contacts, which may serve as ‘bridges’ for signaling networks between many cells (Ott et al., 2012). These contacts are tight adhesions between the ciliary membranes, mediated by N-linked glycoproteins. It is possible that IFT exerts force on cell-cell adhesion sites and determines the positioning of these adhesion sites by moving them along the length of the cilium. Ca²⁺ signaling at flagellar adhesion sites may play a major role in regulating attachment to IFT trains and controlling the direction of force generation. We believe that the assay we developed will be a starting point for deciphering the role of IFT in the signaling, mating and development of ciliated cells.

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Author details

1. Sheng Min Shih

   Department of Physics, University of California, Berkeley, Berkeley, United States

   Contribution

   SMS, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

2. Benjamin D Engel

   Department of Biochemistry, University of California, San Francisco, San Francisco, United States

   Contribution

   BDE, Analysis and interpretation of data, Drafting or revising the article, Contributed unpublished essential data or reagents

   Competing interests

   The authors declare that no competing interests exist.

3. Fatih Kocabas

   Department of Internal Medicine, University of Texas Southwestern Medical Center, Dallas, United States

   Present address

   Texas Institute of Biotechnology, North American College, Houston, United States

   Contribution

   FK, Acquisition of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

4. Thomas Bilyard

   Department of Physics, University of California, Berkeley, Berkeley, United States

   Contribution

   TB, Conception and design, Acquisition of data

   Competing interests

   The authors declare that no competing interests exist.

5. Arne Gennerich

   Anatomy and Structural Biology, Albert Einstein College of Medicine, Bronx, United States

   Contribution

   AG, Acquisition of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

6. Wallace F Marshall

   Department of Biochemistry, University of California, San Francisco, San Francisco, United States

   Contribution

   WFM, Conception and design, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

7. Ahmet Yildiz

   1. Department of Physics, University of California, Berkeley, Berkeley, United States
   2. Department of Molecular Cell Biology, University of California, Berkeley, Berkeley, United States

   Contribution

   AY, Conception and design, Analysis and interpretation of data, Drafting or revising the article

   For correspondence

   yildiz@berkeley.edu

   Competing interests

   The authors declare that no competing interests exist.

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