Force measurements on cargoes in living cells revealcollective dynamics of microtubule motorsAdam G. Hendricks, Erika L. F. Holzbaur, and Yale E. Goldman1

Pennsylvania Muscle Institute and Department of Physiology, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA 19104

Edited by Ronald D. Vale, University of California, San Francisco, CA, and approved September 28, 2012 (received for review September 10, 2012)

Many cellular cargoes move bidirectionally along microtubules,driven by teams of plus- and minus-end–directed motor proteins.To probe the forces exerted on cargoes during intracellular trans-port, we examined latex beads phagocytosed into livingmammalianmacrophages. These latex bead compartments (LBCs) are encased inmembrane and transported along the cytoskeleton by a comple-ment of endogenous kinesin-1, kinesin-2, and dynein motors. Thesize and refractive index of LBCs makes themwell-suited for manip-ulation with an optical trap. We developed methods that provide insitu calibration of the optical trap in the complex cellular environ-ment, taking into account any variations among cargoes and localviscoelastic properties of the cytoplasm.We found that centrally andperipherally directed forces exerted on LBCs are of similar magni-tude, with maximum forces of ∼20 pN. During force events greaterthan 10 pN, we often observe 8-nm steps in both directions, indicat-ing that the stepping of multiple motors is correlated. These obser-vations suggest bidirectional transport of LBCs is driven by opposingteams of stably bound motors that operate near force balance.

optical tweezers | microrheology | intracellular trafficking | tug-of-war |laser trap calibration

Active transport of vesicular cargoes is vital to the targeteddelivery of organelles, proteins, and signaling molecules in

cells. Accordingly, defects in transport are linked to developmental,neurodegenerative, pigmentation, immunological, and other dis-eases (1, 2). Many cargoes are transported by teams of plus- andminus-end–directed motor proteins along the microtubule cyto-skeleton. In mammals, members of the kinesin superfamily drivetransport in the plus-end direction toward the cell periphery,whereas cytoplasmic dynein drives transport toward microtubuleminus ends organized at the cell center.Themechanochemistry of isolatedmotor proteins has been well

characterized through biochemical and single-molecule techni-ques (3–8). In vivo, multiple motor proteins function on a singleorganelle, interact with binding partners and effectors, and oper-ate in a crowded, viscoelastic cellular environment (Fig. 1A). Theextent to which these factors modulate motor protein dynamics inthe cell is not well understood. To investigate the behavior ofmotor proteins in the complex cellular environment, we developedtechniques that allowed us to precisely manipulate vesicular car-goes and measure the forces exerted on them in living cells byusing an optical trap. Using these techniques, we calibrate theoptical trap in situ, directly measuring the position and forcesexerted on intracellular cargoes with high mechanical (<1 nm,<0.2 pN) and temporal (<100 μs) resolution.

ResultsBidirectional Motility of Phagocytosed Latex Beads. We examined1.0-μm latex beads that had been phagocytosed into mouse mac-rophage cells. When they have been internalized, these beads areenveloped in a native phagosome to form latex bead compartments(LBCs) (9), which are transported bidirectionally along micro-tubules by a complement of stably bound endogenous kinesin anddynein motors (10) (SI Appendix, Fig. S1). The motility of LBCswas examined using automated tracking analysis (Fig. 1B–D and SIAppendix, Fig. S2) (11, 12). To ensure we observed LBCs at similar

maturation states, we focused on the time period between 1 and 2 hafter internalization and on cell projections in which microtubulepolarity was well defined. Consistent with previous work (10), wefound that long-range LBC motility was unaffected by perturba-tions to the actin cytoskeleton, but was markedly suppressed whenmicrotubules were depolymerized (SI Appendix, Fig. S2 A–C). Byusing dominant-negative constructs, we found that LBCs aredriven by kinesin-1, kinesin-2, and cytoplasmic dynein (SI Appen-dix, Figs. S1 and S2 D–F). Although these results indicate that theLBC motility at this stage of maturation is primarily driven bymicrotubule motors, we cannot entirely exclude minor con-tributions frommicrotubule dynamics (SI Appendix, Fig. S2A) (13),actin-based motors (14), and cell shape changes.The motility of LBCs is characteristic of bidirectional cargoes.

LBCs in polarized projections exhibited 16% stationary, 73%diffusive, and 11% processive motility, and average velocities of730 nm/s and 880 nm/s in the anterograde (i.e., toward the cellperiphery) and retrograde (i.e., toward the cell center) directions,respectively (Fig. 1E). This motility was similar to the previouslycharacterized motility of bidirectional neuronal cargoes, in whichpurified axonal transport vesicles in vitro and late endosomes/lysosomes in cortical neurons exhibited ∼20% stationary, ∼70%diffusive, and ∼10% processive motility, and average velocitiesof±900 nm/s (12). At the stage of maturation observed (1–2 h afterinternalization), LBC movements were approximately equally di-vided between anterograde and retrograde motility (Fig. 1E).

Optical Trap Calibration in Viscoelastic Environments. We sought tomeasure the forces exerted by microtubule motors on LBCs in thecell by using an optical trap. To obtain accurate force and posi-tion data in living cells, new calibration methods were neededto transform the signals from the quadrant photodiode detectormeasuring optical trap deflection into LBC force and position,explicitly taking into account the viscoelastic nature of the cellularenvironment (15). Briefly, the viscoelastic response of the cyto-plasm is approximated with three components, a constant vis-cosity term (γ), a constant stiffness term (kcyt,0), and a frequency-dependent viscoelasticity [kcyt,1(jω)

α] characteristic of an entan-gled or cross-linked network of semiflexible polymers (16) (SIAppendix,Methods). In Newtonian fluids, the thermal fluctuationsof the bead are adequate to calibrate the optical trap, as theelastic response is entirely prescribed by the stiffness of the op-tical trap. In the cell, other viscoelastic components including thecytoskeleton, the cell membrane, and the motors themselves con-tribute to the elastic response (Fig. 2A), and thus more informationis needed to resolve the stiffness of the optical trap (17). To ad-dress this issue, the spontaneous fluctuations of the bead were

Author contributions: A.G.H., E.L.F.H., and Y.E.G. designed research; A.G.H. performedexperiments; A.G.H. and Y.E.G. analyzed data; and A.G.H., E.L.F.H., and Y.E.G. wrotethe paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.1To whom correspondence should be addressed. E-mail: goldmany@mail.med.upenn.edu.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1215462109/-/DCSupplemental.

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analyzed, as were the forced response to sinusoidal excitationsapplied over a wide range of frequencies through a piezoelectricstage or by positioning the laser via an acousto-optic device. Boththe forced response of the bead to sinusoidal perturbations andthe spontaneous (i.e., unforced) fluctuations of the bead were fitglobally to analytical spectra based on these viscoelastic com-ponents (Fig. 2B and SI Appendix, Fig. S3 and Methods). Incontrast to previous studies (18–21), this method allows in situcalibration of the cargo of interest, taking into account any dif-ferences among cargoes as well as local variations in the cellularenvironment (Fig. 2C). The method implemented here wasdesigned to mitigate the effects of nonequilibrium disturbances,and as such is particularly applicable in living cells. The relativecontributions of the cytoplasm and the optical trap are resolvedby using only the forced response, for which the effect of dis-turbances from biological processes is negligible. In addition,a simple approximation of the viscoelastic response relates theforced response and spontaneous fluctuations in different fre-quency ranges, allowing us to use only the frequency range of thespontaneous fluctuations that is free of apparent disturbanceswhile using the forced response over a large range of frequen-cies, unlike alternate methods (17) (SI Appendix, Fig. S4 shows acomparison with other methods).Consistent with microrheological measurements (22, 23), cali-

brations performed in living cells indicate that the cytoplasm ishighly viscoelastic, with storage and loss moduli of similar order inthe frequency range examined (1 Hz to 5 kHz), and viscosity twoorders of magnitude greater than that of water (Fig. 2 D and E).To validate the calibration method in viscoelastic fluids, we per-formed calibrations in solutions of 0%, 1%, and 2% (wt/vol)methylcellulose (molecular weight of 88,000 Da), which forms anentangled polymer network (Fig. 2 D and E). The viscoelasticparameters measured for methylcellulose solutions comparedwell to previous estimates (24). These measurements also provide

a useful comparison with the properties of the cytoplasm. Al-though 2% methylcellulose approximates the contribution of anentangled polymer network to the viscoelasticity of the cytoplasm,the storage modulus of the cytoplasm deviates from methylcellu-lose at low frequencies as a result of the presence of a significantfrequency-independent, constant stiffness component. Severalfactors may contribute to this constant stiffness, including cross-linking of the cytoskeleton, interactions with the cell membrane,or cross-linking of the LBC to the cytoskeleton by motor proteins.

LBCs in Living Cells. Calibrated optical trap recordings of LBCsin living cells showed forces as high as ∼20 pN generated by teamsof opposite-polarity motors in the anterograde and retrogradedirections (Fig. 3A and SI Appendix, Fig. S5). Frequent short, low-force events were interspersed with less common large-forceevents (Fig. 3 A and C). Interestingly, unlike optical trap record-ings of multiple motors in low-viscosity buffer (25, 19), LBCs in thecell do not tend to stall or dissociate and diffuse quickly (i.e., snapback) into the center of the trap. Instead, LBCs often advanced instepwise movements away and toward the trap center (Fig. 3B).Possible explanations are that the relaxation of LBCs back to thecenter of the trap is slowed in the viscoelastic cellular environ-ment, or that motors in the cell may step backward under load, ashas been observed under high loads in vitro (26, 5). Alternatively,stepwise movements away and toward the center of the trap sug-gest that motors of opposite polarity are simultaneously engaged.Previous studies suggest that bidirectional cargoes are often

driven by few kinesin motors with unitary stall forces of ∼5 to 6 pN(27, 28) andmany dynein motors with unitary stall forces of∼1 pN,nearly balancing the net forces in the plus andminus end directions(12, 19). Although there is some controversy over the unitary stallforce of mammalian dynein (29) likely arising from the sensitivityof dynein’s biophysical properties to buffer conditions (30, 31), ourlaboratory and others find that individual dynein motors isolated

Fig. 1. Latex bead-containing phagosomes exhibit bidirectional motility in the cell. (A) In the cell, motor proteins function collectively in a crowded, viscoelasticenvironment to transport vesicular cargoes. (B) Polystyrene beads 1 μm in diameter (example indicated by arrow) are phagocytosed by J774A.1 mouse mac-rophage cells. (C) These LBCs are transported bidirectionally in the cell, as shown by typical trajectories. The trajectories were projected onto the black line(drawn parallel to trajectories based on a maximum projection image) to quantify displacements in the retrograde and anterograde directions. (D) Anterogradedisplacements are plotted in the upward direction; retrograde displacements are directed downward. Black dots indicate reversals. (E) LBC motility is typical ofbidirectional transport, with short directed runs interspersed with apparent diffusion and pausing. At the stage of maturation used in these experiments (1–2 hafter internalization), the LBCs exhibit approximately equal fractions of plus- and minus-end–directed motility, with similar average velocities in the ante-rograde and retrograde directions. Error bars indicate SEM (n = 52 processive runs, n = 1,261 total runs between reversals, n = 153 trajectories from six cells).

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from mammalian tissue produce ∼1 pN (25, 32, 33). As clear stallevents are rare in the cellular data, we define a force event as anexcursion from the trap center greater than ±0.5 pN, includingboth events in which motors stall and ones in which motors detachbefore reaching stall. When considering only force events >1 s induration, the retrograde force histogram exhibited clear peaksspaced at multiples of 1.6 to 2.3 pN, consistent with events beingdriven by several dynein motors, each with a unitary stall force∼1.7 pN (Fig. 3D and SI Appendix, Table S2). In contrast, theanterograde force histogram showed a broad distribution (Fig. 3Dand SI Appendix, Table S1). A component at∼6 pN is indicative ofsingle kinesin motors reaching stall. Lower-force events are likelycaused by runs that terminate before reaching kinesin’s maximalstall force. Consistent with this interpretation, in vitro single-molecule studies of kinesin indicate that kinesin often dissociatesbefore reaching the maximal stall force (27, 34). Further, althoughkinesin-2 has a similar unitary stall force as kinesin-1, the de-tachment rate of kinesin-2 is very sensitive to force, increasing thefrequency of detachments before stall (35). According to themagnitude of the forces generated, most anterograde events ob-served in living cells are driven by one kinesin, with infrequentevents driven bymultiple motors. Similar to this observation, for invitro experiments in which two kinesin motors were attached toa bead via a DNA scaffold, motility was often driven by one en-gaged motor, and multiple-motor events were rare (34).To further understand how teams of kinesin and dynein motors

function collectively, we used the Kerssemakers step-finding

algorithm (36) to compute step size distributions for events inwhich the maximum force is greater than ±10 pN, as these eventscorrespond to transport by multiple motors. In addition, steps aremore readily observed at high loads as the effect of the compliancebetween the motors and the bead becomes less significant, as aresult of the nonlinear force–extension curve of motor proteins(37, 34). Multiple motors transporting a cargo are expected toproduce fractional steps (less that the step size of a single motor)unless the motors’ steps are synchronized. For example, if twomotors are attached to a cargo and only one of them takes a 8-nmstep, the cargo equilibrates the strain between the motors bymoving half of the step distance, or 4 nm (38, 39). Surprisingly, fornet anterograde and net retrograde runs, we observed frequent8-nm steps, suggesting that multiple motors correlate their step-ping when coupled through a common cargo, apparently steppingnearly simultaneously (Fig. 3 E and F). Independent analysis ofstepping via pairwise distances and Gaussian kernel density esti-mation confirmed that 8-nm steps are frequent at high loads (SIAppendix, Fig. S6); and analysis of simulated stepping tracesverifies that we can reliably detect 8-nm steps at the noise levelspresent in the recorded traces (SI Appendix, Fig. S7). Intracellularcargoes have been observed to advance with 8-nm steps previously(40, 18). However, 8-nm steps are expected if one motor is en-gaged. Here we found that 8-nm steps occurred during high-forceevents driven by multiple motors. This unexpected result providesstrong evidence that the stepping of coupled motors is correlatedat high loads. We also analyzed steps for low-force events (i.e., less

Fig. 2. The optical trap was calibrated in the viscoelastic cellular environment. (A) The diagram depicts the forces on the LBC that result from the optical trapand the viscoelastic cytoplasm. (B) The calibration uses a global fit to the response of the LBC to sinusoidal oscillations of the stage or optical trap and theportion of the power spectrum of spontaneous fluctuations of the LBC greater than 300 Hz assumed to be thermal motions (black line). At frequencies of lessthan 300 Hz, the power spectrum shows disturbances as a result of nonequilibrium, biological processes in the cell, and vibrations of the stage caused by thecoupling of the stage and the LBC in the viscoelastic cytoplasm. Motions of the beads in cells are subdiffusive, as the slope of the power spectrum is less than2 (red line indicates a slope of 2). Insets: Spectra from a bead in water. Note that for a purely viscous fluid like water (kcyt = 0), the magnitude of the forcedresponse continues to decrease at low frequencies, and the slope of the high-frequency fluctuations is near 2. (C) The calibration gives the optical trapstiffness (ktrap) and sensitivity (β), shown here for five LBCs in separate cells. (D and E ) The storage and elastic moduli of the cytoplasm for several cells (106

pN/nm2 = 1 Pa). Results for water, 1% methylcellulose, and 2% methylcellulose solutions are shown for comparison.

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than ±5 pN), and, there, we observed more frequent sub–8-nmsteps. However, interpretation of these values is complicated bytwo factors: the effect of compliance is more significant at lowforces and motors step faster at low loads, making steps moredifficult to detect.

Motility of Isolated LBCs in Vitro. We isolated LBCs from macro-phage cells and reconstituted their motility in vitro, thus decou-pling the dynamics intrinsic to the cargo and bound motors fromthe influence of the complex cellular environment. LBCs wereisolated from cell lysates through floatation on a sucrose step gra-dient and placed on paclitaxel-stabilized, polarity-marked micro-tubules (41, 12). In contrast to previous studies in which additionalcytosolic factors were added (10, 14), we observed motility causedsolely by a stably bound complement of endogenous motor proteins

(SI Appendix, Fig. S1). Isolated LBCs exerted forces bidirec-tionally (Fig. 4 A and B and SI Appendix, Fig. S8) and, as in livingcells, most events were short and low-force (Fig. 4C). When an-alyzing long events (>1 s), we again observed separate peaks in theretrograde force histogram corresponding to transport by multi-ple dynein motors (Fig. 4D). Also similarly to live cell data, theanterograde force histogram exhibited a broad distribution con-sistent with forces primarily caused by single kinesin motors, oftendetaching before reaching stall. The peaks in the force histogramsare strikingly similar to those observed in living cells for ante-rograde and retrograde forces, providing evidence that the cel-lular environment does not substantially modulate the forceproduced by individual motors.Although the unitary forces attributed to motors are similar in

living cells and in vitro, we observe several important differences

Fig. 3. Collective dynamics of kinesin and dynein motors drive LBC motility. (A) Forces are exerted bidirectionally on LBCs in living cells. The optical trapwas calibrated separately for each cargo in the same position within the cell where forces were recorded. Signals were acquired at 2 kHz (gray), thenmedian-filtered to 4 Hz (black). (B) Boxed portions in A are shown in detail, and black lines are filtered to 20 Hz. (C ) Similar forces are exerted by plus- andminus-end–directed motors. Force events are defined as excursions from the trap center greater than ±0.5 pN. The maximum force is recorded for eachevent. [Event criteria: abs(force) >0.5 pN, event duration > 250 ms; n = 2,165 events from 14 recordings.] (D) When only force events longer than 1 s areincluded, the retrograde forces show components at 1.6- to 2.3-pN intervals, whereas anterograde forces show a broader distribution consistent with∼6-pN stalls by single kinesin motors. Low-force events are likely the result of detachments before reaching kinesin’s stall force (n = 855 events).The number of fitted Gaussian components was chosen by using Bayes information criterion (SI Appendix, Fig. S10). (E and F ) Force events greaterthan ±10 pN were analyzed to select for transport driven by multiple motors. For net anterograde (E ) and net retrograde (F ) runs, step size distributionsare centered around ∼8 nm, with occasional back-steps also centered around ∼8 nm (SI Appendix, Figs. S6 and S7). Note that the position data (Inset) aretaken from the traces in B.

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in the collective dynamics of motors between LBCs in living cellsand isolated LBCs in vitro. Unlike force traces in living cells,isolated LBCs often quickly snap back to the center of the trapfollowing detachment, suggesting that fewer motors are engagedat any one time on single microtubules in vitro. In addition,maximum forces of ∼12 pN were observed in vitro, whereasforces as high as ∼20 pN were observed in cells. Although wecannot exclude the possibility that motors dissociate from theLBCs during fractionation, we found that characteristics of forceand motility are stable after isolation for several days on ice,indicating that motors remain active and stably bound to isolatedLBCs. These results suggest that, for the same cargo, largernumbers of motors are able to engage in the cell than on singlemicrotubules. These differences point to several ways in whichthe cellular environment influences bidirectional transport. First,the viscoelastic environment confines diffusion, increasing thetime that cargoes remain near the microtubule, thus promotingmotor binding. Additionally, the motors on a cargo may be ableto access multiple microtubules in the cell, allowing larger num-bers of motors to engage. Indeed, the microtubule cytoskeleton inmacrophage cells is quite dense (SI Appendix, Fig. S9), whereas invitro motility was performed along single microtubules. This ef-fect is likely to be particularly important for large cargoes such asmitochondria and autophagosomes.

DiscussionThe techniques for optical trapping in living cells presented hereenabled reliable, high-resolution measurements of the forces onintracellular cargoes and their motility. Through examining thebidirectional transport of cargoes in the cell, we found that theforces exerted by motor teams in the anterograde and retrogradedirections are nearly balanced, but the characteristics of forcesgenerated by teams of plus- and minus-end–directed motors dif-fer greatly. Many dynein motors (as many as 12, assuming forcesare additive), each producing ∼1.7 pN, exert forces collectively inthe retrograde direction. In contrast, a few kinesin motors (1–3),each exerting ∼5 pN, drive anterograde motility. As a result of the

stochastic nature of kinesin and dynein stepping, it is expectedthat multiple motors transporting a cargo would be synchronizedonly if the motors are strongly coupled to one another (42). Thecompliance of the motors themselves and the elasticity of thevesicular membrane might suggest weak coupling between motors.However, analysis of the stepping dynamics led to a strikingobservation: plus- and minus-end–directed motors seem to cor-relate their steps when functioning collectively at high loads.Further studies will be needed to determine how motors arecoupled on vesicular cargoes and the mechanisms leading to thesecollective motor dynamics. Scaffolding proteins may influence thecoupling between motor proteins to allow them to function ef-fectively in teams. Motor proteins may also display altered gatingwhen functioning collectively to promote correlated stepping.Optical trap measurements suggest that forces exerted by in-

dividual motors on LBCs in living cells are the same as those inisolated LBCs in a simple in vitro system with single microtubules,as we observe similar peaks in the force histograms. However, theviscoelastic environment and cytoskeletal networks present in thecell promote motor binding and allow more motors to engage,leading to simultaneous activity of opposite-polarity motors andhigher maximum forces (Fig. 1A and SI Appendix, Fig. S9).

Materials and MethodsCell Culture. J774A.1 mouse macrophage cells (American Type Culture Col-lection) were cultured as described (43, 41). Briefly, cells were grown in 10-cm dishes in DMEM supplemented with 10% FCS and 1% glutamine at 37 °Cin a 5% CO2 atmosphere. LBCs were formed by incubating the cells withBSA-coated polystyrene beads (1.0 μm diameter, carboxylated; Polysciences)for 10 min at 37 °C. Cells were washed with complete medium, and wereimaged after a 1-h chase. Imaging was performed at 37 °C in media sup-plemented with 10 mM Hepes.

In Vitro Motility of Isolated LBCs. LBCs were isolated as described (43), witha few modifications. After a 90-min chase, cells were lysed by using a Douncehomogenizer with a tight-fitting pestle in motility assay buffer (MAB; 10 mMPipes, 50 mM K-acetate, 4 mM MgCl2, 1 mM EGTA, pH 7.0) supplementedwith protease inhibitors, 1 mM DTT, 1 mM ATP, and 8.5% sucrose. All sucrose

Fig. 4. Bidirectional forces are exerted by motors stably bound to isolated LBCs along paclitaxel-stabilized microtubules in vitro. (A and B) Force traces wereacquired at 2 kHz (gray) and median-filtered to 20 Hz (black). (C) As in live cells, most events are short and low-force (n = 1,137 events from 44 recordings).(D) Retrograde force events >1 s exhibit components at ∼1.5-pN intervals, at peaks strikingly similar to those observed in living cells. Anterograde forceevents >1 s exhibit a component at ∼5 pN, corresponding approximately to the stall force of single kinesin motors and low-force components likely caused byearly detachments (n = 401 events).

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solutions were made in MAB and supplemented as described earlier. Afterisolation, the LBCs maintained activity for at least 2 d stored on ice.

Flow chambers were constructed with a silanized coverslip and a glassslide. Polarity-marked microtubules were bound to the coverslip by using atubulin antibody (anti–β-tubulin, clone Tub2.1; Sigma; diluted 1:20 in MAB).Chambers were blocked with Pluronic F-127 (Sigma) and then washed withtwo chamber volumes of MAB plus 20 μM paclitaxel. Purified LBCs werethen added to the chamber, diluted in MAB, and supplemented with 25 mMDTT, 100 μM MgATP, 20 μM paclitaxel, and an oxygen scavenging system(10 mg/mL glucose, 1 μg/mL glucose oxidase, and 0.5 μg/mL catalase). Thechamber was sealed with vacuum grease, mounted on the microscope, andimaged at ∼20 °C.

Optical Trap. The optical trap was built on a inverted microscope (EclipseTE-2000U; Nikon) with a 1.49 NA oil-immersion objective. The beam from a1,064 nm Nd:YVO4 laser (Spectra Physics) was expanded to overfill the backaperture of the objective. The light passing through the trapped object iscollected by using an oil-immersion condenser objective. A quadrant pho-todiode (Current Designs), positioned conjugate to the back focal plane ofthe objective, provided a measurement of the bead displacement from thetrap center, which correlated directly to force (44). A photodiode bias volt-age of 180 V was used to increase the time resolution of the photodiodeto >10 kHz. The laser position was modulated by using an acousto-opticdeflector (NEOS) and direct digital frequency synthesizer, controlled via afield-programmable gate array by using custom Labview routines (NationalInstruments). For live cell experiments, cells were plated on a 50-mm cov-erslip, then mounted in a customized FCS2 live cell chamber used in com-bination with an objective heater (Bioptechs) to maintain the sample at37 °C during imaging. The reflection of a low-intensity 532-nm laser beam

was positioned onto a quadrant photodiode, and the signal was sentthrough a feedback control system to the piezoelectric stage to autofocusthe distance between the coverslip and the objective during measurements.

Optical Trap Calibration in Living Cells. For each LBC examined, we first re-corded the forces exerted on the LBC (Fig. 3 and SI Appendix, Fig. S5). We nextrecorded the power spectrum of the spontaneous bead fluctuations (Fig. 2Band SI Appendix, Fig. S3). Biological processes in the cell and vibrations ofthe stage result in added noise in the frequency range <300 Hz, so these dataare not used for the calibration. The response at frequencies >300 Hz wasfree of apparent disturbances and assumed to be driven by thermal fluctu-ations. We then recorded the forced response to sinusoidal oscillations ofthe stage [for excitation frequencies (fexc) <50 Hz] or the trap position (fexc>50 Hz) over a large range of frequencies (1 Hz ≤ fexc ≤ 5 kHz; Fig. 2B and SIAppendix, Fig. S3). At each frequency, the amplitude of the excitation wastailored to ensure a linear response. To obtain trap and viscoelastic param-eters, the magnitude of the forced response and the power spectrum of thespontaneous bead fluctuations were fit globally to the analytical response.Derivation of the analytical response is described in SI Appendix, Methods.Force traces recorded before and after the calibration procedure showedsimilar results, indicating that motor proteins were not damaged by pro-longed exposure to the IR laser in the cell.

ACKNOWLEDGMENTS. The authors thank Mr. Pritish Agarwal and Ms.Mariko Tokito for technical assistance, Dr. Henry Shuman for helpful advice,and Gheorghe Chistol for providing the Gaussian kernel density algorithm.This work was supported by National Institutes of Health Grants GM087253(to E.L.F.H. and Y.E.G.) and GM089077 (to A.G.H.).

1. Chevalier-Larsen E, Holzbaur ELF (2006) Axonal transport and neurodegenerativedisease. Biochim Biophys Acta 1762(11-12):1094–1108.

2. Akhmanova A, Hammer JA, 3rd (2010) Linking molecular motors to membrane cargo.Curr Opin Cell Biol 22(4):479–487.

3. Hackney DD (1994) Evidence for alternating head catalysis by kinesin during micro-tubule-stimulated ATP hydrolysis. Proc Natl Acad Sci USA 91(15):6865–6869.

4. Visscher K, Schnitzer MJ, Block SM (1999) Single kinesin molecules studied witha molecular force clamp. Nature 400(6740):184–189.

5. Clancy BE, Behnke-Parks WM, Andreasson JOL, Rosenfeld SS, Block SM (2011) A uni-versal pathway for kinesin stepping. Nat Struct Mol Biol 18(9):1020–1027.

6. Ross JL, Wallace K, Shuman H, Goldman YE, Holzbaur ELF (2006) Processive bi-directional motion of dynein-dynactin complexes in vitro. Nat Cell Biol 8(6):562–570.

7. DeWitt MA, Chang AY, Combs PA, Yildiz A (2012) Cytoplasmic dynein moves throughuncoordinated stepping of the AAA+ ring domains. Science 335(6065):221–225.

8. Qiu W, et al. (2012) Dynein achieves processive motion using both stochastic andcoordinated stepping. Nat Struct Mol Biol 19(2):193–200.

9. Desjardins M, Griffiths G (2003) Phagocytosis: Latex leads the way. Curr Opin Cell Biol15(4):498–503.

10. Blocker A, et al. (1997) Molecular requirements for bi-directional movement ofphagosomes along microtubules. J Cell Biol 137(1):113–129.

11. Ruhnow F, Zwicker D, Diez S (2011) Tracking single particles and elongated filamentswith nanometer precision. Biophys J 100(11):2820–2828.

12. Hendricks AG, et al. (2010) Motor coordination via a tug-of-war mechanism drivesbidirectional vesicle transport. Curr Biol 20(8):697–702.

13. Blocker A, Griffiths G, Olivo JC, Hyman AA, Severin FF (1998) A role for microtubuledynamics in phagosome movement. J Cell Sci 111(Pt 3):303–312.

14. Al-Haddad A, et al. (2001) Myosin Va bound to phagosomes binds to F-actin anddelays microtubule-dependent motility. Mol Biol Cell 12(9):2742–2755.

15. Veigel C, Schmidt CF (2011) Moving into the cell: Single-molecule studies of molecularmotors in complex environments. Nat Rev Mol Cell Biol 12(3):163–176.

16. Gittes F, MacKintosh FC (1998) Dynamic shear modulus of a semiflexible polymernetwork. Phys Rev E Stat Phys Plasmas Fluids Relat Interdiscip Topics 58:R1241–R1244.

17. Fischer M, Richardson AC, Reihani SNS, Oddershede LB, Berg-Sørensen K (2010) Ac-tive-passive calibration of optical tweezers in viscoelastic media. Rev Sci Instrum 81(1):015103.

18. Sims PA, Xie XS (2009) Probing dynein and kinesin stepping with mechanical ma-nipulation in a living cell. ChemPhysChem 10(9-10):1511–1516.

19. Soppina V, Rai AK, Ramaiya AJ, Barak P, Mallik R (2009) Tug-of-war between dis-similar teams of microtubule motors regulates transport and fission of endosomes.Proc Natl Acad Sci USA 106(46):19381–19386.

20. Shubeita GT, et al. (2008) Consequences of motor copy number on the intracellulartransport of kinesin-1-driven lipid droplets. Cell 135(6):1098–1107.

21. Leidel C, Longoria RA, Gutierrez FM, Shubeita GT (2012) Measuring molecular motorforces in vivo: Implications for tug-of-war models of bidirectional transport. Biophys J103(3):492–500.

22. Hoffman BD, Massiera G, Van Citters KM, Crocker JC (2006) The consensus mechanicsof cultured mammalian cells. Proc Natl Acad Sci USA 103(27):10259–10264.

23. Jonas M, Huang H, Kamm RD, So PTC (2008) Fast fluorescence laser tracking micro-rheometry, II: Quantitative studies of cytoskeletal mechanotransduction. Biophys J 95(2):895–909.

24. Brau RR, et al. (2007) Passive and active microrheology with optical tweezers. J Opt A,

Pure Appl Opt 9:S103.25. Schroeder HW, 3rd, Mitchell C, Shuman H, Holzbaur ELF, Goldman YE (2010) Motor

number controls cargo switching at actin-microtubule intersections in vitro. Curr Biol

20(8):687–696.26. Gennerich A, Carter AP, Reck-Peterson SL, Vale RD (2007) Force-induced bidirectional

stepping of cytoplasmic dynein. Cell 131(5):952–965.27. Svoboda K, Block SM (1994) Force and velocity measured for single kinesin molecules.

Cell 77(5):773–784.28. Khalil AS, et al. (2008) Kinesin’s cover-neck bundle folds forward to generate force.

Proc Natl Acad Sci USA 105(49):19247–19252.29. Toba S, Watanabe TM, Yamaguchi-Okimoto L, Toyoshima YY, Higuchi H (2006)

Overlapping hand-over-hand mechanism of single molecular motility of cytoplasmic

dynein. Proc Natl Acad Sci USA 103(15):5741–5745.30. Walter WJ, Brenner B, Steffen W (2010) Cytoplasmic dynein is not a conventional

processive motor. J Struct Biol 170(2):266–269.31. Walter WJ, Koonce MP, Brenner B, Steffen W (2012) Two independent switches

regulate cytoplasmic dynein’s processivity and directionality. Proc Natl Acad Sci USA

109(14):5289–5293.32. Mallik R, Carter BC, Lex SA, King SJ, Gross SP (2004) Cytoplasmic dynein functions as

a gear in response to load. Nature 427(6975):649–652.33. McKenney RJ, Vershinin M, Kunwar A, Vallee RB, Gross SP (2010) LIS1 and NudE in-

duce a persistent dynein force-producing state. Cell 141(2):304–314.34. Jamison DK, Driver JW, Rogers AR, Constantinou PE, Diehl MR (2010) Two kinesins

transport cargo primarily via the action of one motor: implications for intracellular

transport. Biophys J 99(9):2967–2977.35. Schroeder HW, 3rd, et al. (2012) Force-dependent detachment of kinesin-2 biases

track switching at cytoskeletal filament intersections. Biophys J 103(1):48–58.36. Kerssemakers JW, et al. (2006) Assembly dynamics of microtubules at molecular res-

olution. Nature 442(7103):709–712.37. Svoboda K, Schmidt CF, Schnapp BJ, Block SM (1993) Direct observation of kinesin

stepping by optical trapping interferometry. Nature 365(6448):721–727.38. Holzbaur ELF, Goldman YE (2010) Coordination of molecular motors: From in vitro

assays to intracellular dynamics. Curr Opin Cell Biol 22(1):4–13.39. Leduc C, Ruhnow F, Howard J, Diez S (2007) Detection of fractional steps in cargo

movement by the collective operation of kinesin-1 motors. Proc Natl Acad Sci USA 104

(26):10847–10852.40. Nan X, Sims PA, Xie XS (2008) Organelle tracking in a living cell with microsecond time

resolution and nanometer spatial precision. ChemPhysChem 9:707–712.41. Desjardins M, et al. (1994) Molecular characterization of phagosomes. J Biol Chem 269

(51):32194–32200.42. Hendricks AG, Epureanu BI, Meyhöfer E (2009) Collective dynamics of kinesin. Phys

Rev E Stat Nonlin Soft Matter Phys 79(3 Pt 1):031929.43. Vinet AF, Descoteaux A (2009) Large scale phagosome preparation.Methods Mol Biol

531:329–346.44. Tolic-Norrelykke SF, et al. (2006) Calibration of optical tweezers with positional de-

tection in the back focal plane. Rev Sci Instrum 77:103101.

18452 | www.pnas.org/cgi/doi/10.1073/pnas.1215462109 Hendricks et al.

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