Control of cell membrane tension by myosin-IRajalakshmi Nambiar, Russell E. McConnell, and Matthew J. Tyska1

Department of Cell and Developmental Biology, Vanderbilt University Medical Center, Nashville, TN 37205

Edited by Edward D. Korn, National Heart, Lung and Blood Institute, Bethesda, MD, and approved May 29, 2009 (received for review February 13, 2009)

All cell functions that involve membrane deformation or a changein cell shape (e.g., endocytosis, exocytosis, cell motility, and cyto-kinesis) are regulated by membrane tension. While molecularcontacts between the plasma membrane and the underlying actincytoskeleton are known to make significant contributions to mem-brane tension, little is known about the molecules that mediatethese interactions. We used an optical trap to directly probe themolecular determinants of membrane tension in isolated or-ganelles and in living cells. Here, we show that class I myosins, afamily of membrane-binding, actin-based motor proteins, mediatemembrane/cytoskeleton adhesion and thus, make major contribu-tions to membrane tension. These studies show that class I myosinsdirectly control the mechanical properties of the cell membrane;they also position these motor proteins as master regulators ofcellular events involving membrane deformation.

actin � brush border � cytoskeleton � microvilli � optical trap

P lasma membrane tension in eukaryotic cells is a masterregulator of all cellular processes that involve membrane

deformation including endocytosis (1), exocytosis (2), mem-brane repair (3), cell motility (4), and cell spreading (5). Thetotal tension present in the plasma membrane (i.e., the ‘‘appar-ent’’ membrane tension, TApp) has a minor contribution from thesurface tension of the lipid bilayer (TM) and a substantialcontribution from the molecular contacts that afford adhesion tothe underlying actin cytoskeleton (�) (6). To prevent largechanges in tension, the plasma membrane must maintain con-tinuous interactions with the cytoskeleton, despite the fact thatthis structure is highly dynamic and continuously remodeling.However, little is known about the molecules that mediatedynamic interactions between membrane and cytoskeleton, orthe proteins that contribute directly to controlling membranetension in cells.

A striking biological example of the complex mechanicalinterplay between the plasma membrane and the supportingcytoskeleton is provided by the brush border found on the apexof intestinal epithelial cells (7). The brush border functions as theprimary site for nutrient absorption and consists of up to 1,000microvilli, protrusions that increase apical membrane surfacearea and release vesicles into the intestinal lumen (7, 8). Eachmicrovillus is supported by a parallel bundle of actin filamentsthat enables this structure to extend several microns from the cellsurface (7). To stabilize this convoluted morphology, epithelialcells must furnish the brush border with high levels of mem-brane-cytoskeleton adhesion energy. One candidate moleculefor carrying out this task is myosin-1a (Myo1a), a monomericactin-based motor protein that is present at high levels in themicrovillus and is known to bind directly to phospholipids byvirtue of its basic C-terminal tail homology 1 (TH1) domain (9,10). Myo1a KO mice exhibit large apical membrane herniationsthat are morphologically similar to ‘‘blebs’’ (11, 12). Becauseblebs represent complete delamination of membrane from theactin cytoskeleton, these results suggest that Myo1a makes amajor contribution to membrane-cytoskeleton adhesion in thebrush border. While previous studies have suggested a role forclass I myosins in the regulation of cortical stiffness (i.e., wholecell deformability) of Dictyostelium discoideum cells (13), therole of class I myosins in the control of membrane tension has

not been explored. Thus, the goal of the current study was todetermine whether class I myosins function in controlling themechanical properties of the plasma membrane.

ResultsProbing Membrane Tension with an Optical Trap. We sought toinvestigate the contribution of Myo1a and other class I myosinsto plasma membrane tension in isolated organelles and livingcells. To this end, we developed an optical trap assay that enabledus to measure the force exerted by a thin tubule or ‘‘tether’’extracted from a membrane (14). In a typical tether forceexperiment, a concanavalin-A-coated 2.0 �m diameter micro-sphere was captured in the optical trap and then brought incontact with an isolated brush border or intact cell, which wasfirmly attached to a glass coverslip surface. Membrane tetherswere then formed by translating the piezoelectric stage to movethe sample away from the trapped bead. Forces exerted bymembrane tethers on the bead were derived from microsphereposition data (15), acquired at video rate using a CCD camera;position data were converted to force using the stiffness of theoptical trap (kTrap), which for our studies ranged from 0.1–0.4pN/nm (Fig. S1).

Myo1a KO Brush Borders Demonstrate Defects in Membrane Force-Extension. As a first step, we examined the force-extensionproperties of the apical membrane associated with brush bordersisolated from WT or Myo1a KO mouse small intestine. Becauseisolated brush borders are prepared in the absence of ATP,tether formation in this case is an irreversible process. The firststep in these experiments was to capture a ConA-coated beadwith the optical trap and bring it in contact with a coverslip-adsorbed brush border. After waiting approximately 4 s to allowfor bead binding to the apical membrane, tethers were pulled bytranslating samples away from the trapped bead at 1.0 �m/s (Fig.1); recordings ended once the bead escaped from the trap. Underthese conditions we observed that the slope of the membraneforce-extension curve is significantly higher in WT brush bordersrelative to Myo1a KO samples (0.43 � 0.01 vs. 0.05 � 0.01pN/nm, respectively; Fig. 1E). This approach also revealed thatMyo1a limits the maximum length of membrane tether that wewere able to extract during these experiments (WT, 3.5 � 0.3 vs.KO, 7.8 � 2.0 �m; Fig. 1F). Because these measurements wereperformed in the absence of active membrane trafficking orother potentially confounding subcellular activities, they clearlyindicate that Myo1a makes a direct contribution to the mechan-ical stability of the brush border apical membrane.

Author contributions: R.N. and M.J.T. designed research; R.N. performed research; R.N.,R.E.M., and M.J.T. contributed new reagents/analytic tools; R.N. and M.J.T. analyzed data;and R.N. and M.J.T. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

1To whom correspondence should be addressed at: Department of Cell and DevelopmentalBiology, Vanderbilt University Medical Center, 3130 Medical Research Building III, 465 21stAvenue South, Nashville, TN 37232-8240. Email: matthew.tyska@vanderbilt.edu.

This article contains supporting information online at www.pnas.org/cgi/content/full/0901641106/DCSupplemental.

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Myo1a Controls TApp in Living Epithelial Cells. We next sought todetermine whether Myo1a plays a role in regulating apicalmembrane mechanics in the context of living, polarized epithe-lial cells. In intact cells, the force exerted by a membrane tetherheld at constant length is directly related to the level of apparentmembrane tension (6). Importantly, tether formation in this caseis a reversible process; after forming a tether, release of thetrapped microsphere allows the cell to resorb the extractedmembrane. We carried out tether force measurements using thecolonic adenocarcinoma cell line, NGI3 (16). Upon differenti-ation, NGI3 cells express endogenous Myo1a and build anelaborate brush border with densely packed microvilli. For theseexperiments, an optical trap was used to capture a ConA-coatedbead and bring it into contact with a surface-adsorbed NGI3 cellto enable binding as described above. To form a membranetether, the cell was translated away from the trapped bead at aconstant rate of 1 �m/s. Tethers were pulled to a length of 5 �mas our initial experiments with NGI3 cells revealed that tetherforce is independent of length in this regime of extension (Fig.S2), in a manner similar to previous results with other cell lines(17). To probe the contribution of Myo1a to apparent membranetension, we perturbed the endogenous population of this motor

using 2 methods: (1) expression of an EGFP-tagged Myo1a-TH1dominant negative construct, which disrupts the targeting ofendogenous Myo1a and gives rise to cellular phenotypes similarto those observed in Myo1a KO mice (11, 18), and (2) expressionof an EGFP-tagged full length Myo1a construct to supplementthe population of endogenous Myo1a (19) (Fig. S3). Strikingly,expression of the EGFP-Myo1a-TH1 dominant negative inNGI3 cells dramatically reduced the force observed during thetethered phase of individual records, relative to cells expressingEGFP as a negative control (17.9 � 5.1 vs. 32.1 � 4.6 pN; Figs.2A and B). In contrast, expressing EGFP-Myo1a in NGI3 cellsgave rise to an increase in tether force relative to control cells(42.5 � 4.2 pN; Fig. 2 A and B). Thus, in the context of liveepithelial cells, Myo1a enables the apical membrane to resistdeformation (i.e., tether formation) and makes a substantialcontribution to apparent membrane tension (TApp).

Myo1a Contributes to Membrane-Cytoskeleton Adhesion (�) in LivingEpithelial Cells and Fibroblasts. The results observed in NGI3 cellscould be explained in 1 of 2 ways: (1) Myo1a could have a directimpact on membrane surface tension (TM), or (2) Myo1a couldmake a significant contribution to membrane-cytoskeleton ad-

Fig. 1. (A) Confocal micrograph of a membrane tether pulled from a single isolated brush border labeled with Alexa488-Concanavalin A. Image is invertedand contrast enhanced to enable visualization of the extremely dim membrane tether. (B) Phalloidin signal from the isolated brush border shown in a revealsthat the tether is devoid of F-actin. The position of the trapped bead and membrane tether are indicated; bars in A and B are 2 �m. (C) Merge of images fromA and B (shown without contrast enhancement) demonstrates the colocalization of membrane (green) and F-actin (red) signals; panel on the right shows theorientation of microvillar actin bundles in this structure. (D) Cartoon depicting the polarity of microvillar actin bundles of the brush border imaged in A–C. (E)Force-extension records for membrane tethers extracted from WT (green) and Myo1a KO (red) brush borders. Linear fitting of raw data over the first �m ofextension yields spring constants of 0.43 � 0.01 pN/nm (n � 6, R2 � 0.99) and 0.05 � 0.01 pN/nm (n � 7, R2 � 0.92) for WT and KO brush borders, respectively.(F) Box-plots of maximum tether lengths demonstrate that Myo1a KO brush borders released significantly longer tethers (7.8 � 2.0 �m; red) relative to WTcontrols (3.50 � 0.34 �m; green). *P � 0.05.

Fig. 2. (A) Representative tether force records from NGI3 cells transfected with EGFP (green), EGFP-Myo1a-TH1 (red), or EGFP-Myo1a (blue). The region of therecord marked as ‘‘Tethered’’ corresponds to the quasi-stable equilibrium force achieved after piezo-stage translation has stopped and tether formation iscomplete. (B) Bar graphs of the average force observed during the tethered phase of individual records reveal that EGFP-Myo1a-TH1 dominant-negativeexpression lowers tether force (17.9 � 5.1 pN, n � 14), while expression of EGFP-Myo1a increases tether force (42.5 � 4.2 pN, n � 11) relative to EGFP-expressingcontrol cells (32.1 � 4.6 pN, n � 15). *P � 0.05, **P � 0.005. (C) Model representation of the correspondence between tether force measurements and themolecular level perturbations induced by expressing EGFP-Myo1a or EGFP-Myo1a-TH1, relative to the EGFP negative control.

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hesion energy (�) by physically linking the membrane to the actincytoskeleton. With its combined membrane- and actin-bindingactivities, Myo1a is ideally suited for mediating to membrane-cytoskeleton adhesion; decreasing or increasing the number offunctional Myo1a molecules per unit area of membrane with theexpression of EGFP-Myo1a-TH1 or EGFP-Myo1a, respectively,would give rise to a corresponding reduction or elevation inadhesion energy, and thus apparent membrane tension (Fig. 2C).To further test this hypothesis, we performed tether forcemeasurements under conditions that favored the formation ofmultiple membrane tethers (20, 21). When multiple membranetethers are simultaneously pulled from the same local region ofmembrane, they demonstrate a tendency to coalesce into a singletether in the absence of ‘‘pinning’’ forces (22), for example,forces provided by molecular links to the underlying cytoskele-ton. Thus, assaying the number of tethers formed from a singlemicrosphere/cell encounter provides a read-out on the density ofmolecular contacts between the membrane and cytoskeleton.Increasing the contact area between the microsphere and cell, orincreasing the loading rate (by raising trap stiffness) biasedevents toward the formation of multiple tethers (23) (Fig. S4Aand B). Multiple tether formation was confirmed using DICmicroscopy (Fig. S5A). The presence of multiple tethers was alsoindicated by the appearance of ‘‘staircases’’ during the tetheredphase of force records (Fig. 3A). The rapid drops or ‘‘steps’’between discrete plateaus in force are the result of adjacent

membrane tether coalescence (22, 24). Similar to previousstudies of multiple tether mechanics (20, 25), observed forcesteps appear as integer multiples of the force measured for singletethers (Fig. S5B and C). Intriguingly, visual inspection of forcerecords revealed that expression of the EGFP-Myo1a-TH1dominant negative significantly impaired the cell’s ability tostabilize multiple tethers. This was indicated by 2 importantobservations. First, the tethered phase of records from TH1-expressing cells appeared to start at a lower level of force (Fig.3A). We confirmed this by calculating force-time integrals acrossthe tethered phase of individual records. EGFP-Myo1a-TH1expression significantly reduced the mean force-time integral,suggesting a reduction in the total number of tethers thatcontribute to force during the tethered phase (Fig. 3D). Second,when the number of observable steps (i.e., coalescence events)was tallied from force records, TH1-expressing cells demon-strated a significantly reduced average number of events perrecord (Fig. 3B and C).

As discussed above, the force steps observed during multipletether events are integer multiples of a unitary tether force (�30pN in the case of NGI3 cells, Fig. 2 and Fig. S5B), suggesting thateach step represents the same underlying physical process, thatis, tether coalescence. Because the coalescence of membranetethers is due to the failure of bonds that link the membrane andunderlying cytoskeleton, we expect the irreversible transitionfrom n tethers to n-1 tethers to proceed as a first-order process

Fig. 3. (A) Representative multiple tether force records from NGI3 cells transfected with EGFP (green) or EGFP-Myo1a-TH1 (red). Arrows indicate the beginningof the tethered phase in each case. Rapid drops in force during the tethered phase correspond to tether coalescence events. (B) Multiple representative examplesof the tethered phases from records of NGI3 cells expressing EGFP (green) and EGFP-Myo1a-TH1 (red). Data were plotted with a fixed offset of 25 pN betweenall records. Visual inspection reveals that EGFP-Myo1a-TH1 records contain fewer observable coalescence events relative to EGFP control records of the sameduration. (C) Box plots show the mean number of coalescence events or ‘‘steps’’ observed in EGFP or EGFP-Myo1a-TH1 records; cells expressing the TH1 dominantnegative demonstrate significantly fewer coalescence events, relative to control cells (*P � 0.05). (D) Box plots of the force-time integrals calculated from thetethered phase of individual records; EGFP-Myo1a-TH1 expressing cells demonstrate significantly lower force-time integrals relative to EGFP-expressing controls(**P � 0.001). (E) Ensemble averages of tethered phase data from NGI3 cells transfected with EGFP (green) and EGFP-Myo1a-TH1 (red); open circles show averagevalues at 0.3-s intervals; solid lines are single exponential fits to averaged data. Similar force decay kinetics were observed for EGFP (0.12 � 0.01 s�1, n � 28, R2 �0.99) and EGFP-Myo1a-TH1 (0.13 � 0.01 s�1, n � 30, R2 � 0.98), respectively. (F) Model of the mechanism underlying the formation of multiple tethers in cellsexpressing EGFP (negative control) or EGFP-Myo1a-TH1 (dominant negative). These cartoons represent ‘‘snap shots’’ in the records shown in A, taken at thebeginning of the tethered phases at the time point indicated by the black arrows.

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with a rate equal to approximately 1/lifetime of a single tether(26). Indeed, ensemble averaging of the tethered phases fromindividual EGFP and EGFP-Myo1a-TH1 records revealed sin-gle-exponential decays in force for both data sets (Fig. 3E). Fitsto these data revealed comparable kinetics (EGFP, 0.12 � 0.01s�1; EGFP-Myo1a-TH1, 0.13 � 0.01 s�1) indicating that a similarmolecular process controls the rate of tether coalescence inEGFP and EGFP-Myo1a-TH1-expressing cells. We proposethat Myo1a plays a role in stabilizing multiple membrane tethersin both cases. However, in cells expressing the dominant nega-tive, a large fraction of the endogenous Myo1a population isdisplaced from the plasma membrane; this reduces its effectivecontribution to membrane-cytoskeleton adhesion and ultimatelyimpairs the cell’s ability to form and stabilize multiple tethers(Fig. 3F).

Based on the model outlined above, one prediction is that theexpression of EGFP-Myo1a in NGI3 cells should enhance thecell’s capacity for stabilizing multiple tethers. While we at-tempted multiple tether experiments with NGI3 cells expressingEGFP-Myo1a, our ability to extract tethers from these cells wasdramatically decreased. If Myo1a does contribute to membrane-cytoskeleton adhesion (�), then this observation might reflectthe expected outcome. Because NGI3 cells express endogenousMyo1a, transfection with EGFP-Myo1a creates an over-expression scenario that could give rise to exaggerated mem-brane-cytoskeleton adhesion. We suspect that the extraction ofmultiple tethers is precluded in this case, as the forces requiredare beyond the upper limits of our optical trap. In light of thispossibility, we examined the impact of EGFP-Myo1a expressionon multiple tether formation from cells that express low levels ofendogenous Myo1a. For these experiments, we used NIH 3T3fibroblasts, which do not express detectable Myo1a and exhibita low apparent membrane tension (�25% of TApp for NGI3, Fig.S4C) (5). Multiple tether force measurements in NIH 3T3 cellsproduced step-containing records as observed in NGI3 cellsdescribed above. Inspection of raw data and analysis of force-time integrals revealed that expression of EGFP-Myo1a in NIH3T3 cells produces a modest increase in the force observedduring the tethered phase (Fig. 4, royal blue data). These resultsshow that Myo1a expression is capable of increasing membrane-cytoskeleton adhesion outside the context of the polarizedcytoskeleton found in NGI3 cells.

Interestingly, the force decay kinetics in NIH 3T3 cells ex-pressing EGFP-Myo1a were comparable to those observed inexperiments with NGI3 cells (Fig. 4C vs. Fig. 3E; 0.17 � 0.01 s�1

vs. 0.12 � 0.01 s�1, respectively), which normally express signif-icant levels of endogenous Myo1a. This kinetic similarity sug-gests that our tether force measurements are in fact probing themechanical contributions made by Myo1a in the case of bothexperiments.

Control of Membrane Mechanics May Be a Class-Wide Function forClass I Myosins. The results summarized to this point indicate thatMyo1a controls apparent membrane tension (TApp) by contrib-uting to membrane-cytoskeleton adhesion (�). Given that all 8vertebrate class I myosins contain a basic TH1 domain (27),which mediates interactions with cellular membranes (28), wesought to determine whether other myosin-I isoforms could alsocontribute to membrane-cytoskeleton adhesion. To this end, weexpressed EGFP-tagged versions of 3 other short-tailed (Myo1b,Myo1c, and Myo1d) and 1 long-tailed (Myo1e) class I myosinsin NIH 3T3 cells (Fig. S6) and probed their impact on multipletether formation as described above. Inspection of raw datarecords qualitatively revealed that regardless of the isoform,class I myosin expression increased the force measured duringthe tethered phase relative to EGFP-expressing controls (Fig.4A). This was confirmed through quantitative analysis of mul-tiple tether records by calculating average force-time integrals(Fig. 4B) and producing ensemble averages of data (Fig. 4C) foreach construct as outlined above (see Fig. 3). Of note here is thefact that Myo1e, the only long-tailed isoform included in ouranalysis, had the most dramatic impact on multiple tetherformation. The uniform increase in multiple tether formationobserved with the expression of different myosin I isoforms,suggests that membrane-cytoskeleton adhesion and the controlof plasma membrane tension may be general functions for allvertebrate class I myosins.

DiscussionExperiments performed in multiple eukaryotic model systemshave implicated class I myosins in various aspects of membrane-related events including phagocytosis (29–33), endocytosis (34–37), exocytosis (38, 39), and membrane recycling (40). Althoughour current data set does not allow us to rule out the possibilitythat perturbations in membrane trafficking may contribute tothe changes in membrane tension observed in our experiments,our results do provide strong support for a model where class Imyosins play a direct role in the control of membrane tension,by contributing to adhesion between the plasma membrane andunderlying actin cytoskeleton. Indeed, mechanical measure-

Fig. 4. (A) Representative tether force records for NIH 3T3 fibroblasts expressing EGFP (green) or 1 of 5 different EGFP-tagged class I myosins (as labeled onthe plot). (B) Box plots of force-time integrals calculated over the tethered phases of individual records from NIH 3T3 cells expressing the various class I myosinconstructs (*P � 0.05, **P � 0.001). Cells expressing myosin-I constructs uniformly demonstrate higher force-time integrals relative to EGFP expressing controls.(C) Ensemble averages of tethered phases from records of NIH 3T3 cells transfected with various class I myosin constructs (open circles); solid lines show singleexponential fits to the data. Rate constants obtained from exponential fits are as follows: EGFP, 0.04 � 0.01 s�1

, n � 23, R2 � 0.97; EGFP-Myo1a, 0.17 � 0.01 s�1,

n � 21, R2 � 0.99; EGFP-Myo1b, 0.06 � 0.01 s�1, n � 15, R2 � 0.97; EGFP-Myo1c, 0.13 � 0.01 s�1

, n � 10, R2 � 0.91; EGFP-Myo1d, 0.20 � 0.01 s�1, n � 9, R2 � 0.96;

EGFP-Myo1e, 0.28 � 0.01 s�1, n � 5, R2 � 0.98.

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ments with isolated brush borders revealed that Myo1a increasesapical membrane force-extension stiffness approximately 10-fold(Fig. 1E). Because these experiments were performed in theabsence of ATP, potentially confounding processes such asmembrane trafficking were not active during these in vitromeasurements and had little impact on the observed mechanicalresponses. This model finds additional support when we considerthat disrupting Myo1a function in polarized epithelial (NGI3)cells reduced the mean tether force by approximately 50% (Fig.2A and B). Apparent membrane tension (TApp), considered tobe the sum of in-plane tension (Tm) and membrane/cytoskeletonadhesion (�), is directly related to the tether force (FTether): TApp� Tm � � � (FTether)2/(8B�2), where B is the membrane bendingstiffness (41). Thus, perturbation of Myo1a reduced apparentmembrane tension by approximately 70%. This value ap-proaches previously published estimates that attribute over 75%of apparent membrane tension to membrane-cytoskeleton ad-hesion (6). Finally, analysis of multiple tether formation providessome of the most direct support for this model. Expression of theMyo1a TH1 dominant negative decreased the ability of NGI3cells to support and stabilize multiple membrane tethers,whereas over-expression of Myo1a or other class I myosinsstabilized multiple membrane tethers (Figs. 3 and 4). Becausethe ability to form multiple tethers is directly linked to thedensity of molecular contacts between the membrane andcytoskeleton, these results tell us that class I myosins areimportant players in mediating these interactions. Thus, theresults presented here strongly support a model where class Imyosins play a direct role in the control of membrane tension,by contributing to adhesion between the plasma membrane andunderlying actin cytoskeleton.

The mechanical measurements presented here provide aphysical explanation for the phenotypes observed in the Myo1aKO mouse (11). Among the most striking defects observed inthis model are herniations of apical membrane that extend fromthe apical surface of KO enterocytes. In most cell types cytosolicf luid pressure, created by myosin-II powered contractility in thecell cortex, exerts a positive (i.e., outward) force on the plasmamembrane (42). In the enterocyte, the high levels of membrane-cytoskeleton adhesion provided by the microvillar population ofMyo1a function to counter cytosolic pressure so that the brushborder can stabilize the enormous quantity of plasma membranepacked into this domain.

In addition to providing access to information about mem-brane-cytoskeleton adhesion, the multiple tether experimentsdescribed here may provide important mechanistic informa-tion on the formation of ‘‘tethers’’ under normal physiologicalconditions. As an example, leukocytes rolling along endothe-lium extrude multiple membrane tethers to stabilize theirrolling velocities, ultimately enabling arrest and extravasation(25). Thus, one goal for future studies will be to determinewhether the class I myosins expressed in leukocytes, play a rolein the formation and stabilization of these important mem-brane structures.

While the importance of the actin cytoskeleton in shaping theplasma membrane and its mechanical properties is well estab-lished (14), the results described here show that actin-basedmotors, and specifically class I myosins, play a role in controllingthe mechanical interactions between these 2 systems. Class Imolecules are ideal candidates for fulfilling this function withincells. Cryo-electron microscopy studies have established thatMyo1a is an extended molecule arranged with the actin-bindingmotor domain at 1 end, and the putative lipid interacting domainat the other (43). This domain arrangement is well suited for thebivalent crosslinking of plasma membrane to actin filaments,while maintaining an approximate 15-nm (projected length ofMyo1a) gap between these 2 compartments. In addition todomain organization, specific mechanochemical properties ap-

pear to be tuned to enable these motors to contribute tomembrane tension (44). Recent single molecule studies indicatethat the activity of myosin-1b is exquisitely sensitive to opposingexternal load (45). These studies show that the rate of ADPrelease and thus detachment from actin slows down approxi-mately 100-fold in response to loads as small as 2 pN (45).Load-dependent kinetics may enable class I myosins to functionin membrane/cytoskeleton adhesion by allowing them to remainstrongly bound to F-actin for long periods, without hydrolyzingATP.

Although the detailed kinetics of myosin-I/actin interactionsare in many cases well-characterized (44), less is known aboutthe mechanism underlying myosin-I membrane binding. Whileit was established many years ago that Myo1a binds to acidicphospholipids via its basic TH1 domain (10), more recentstudies have revealed that vertebrate and Acanthamoeba classI myosin TH1 domains contain a PH motif able to bind tightlyto highly charged, acidic phosphoinositides such as PIP2 (46,47). These studies may help explain earlier biophysical dataimplicating PIP2 in membrane/cytoskeleton adhesion (48).However, PIP2 is estimated to comprise �1% of total phos-pholipid found in the inner leaf let of the plasma membrane(49). Thus, in domains such as the brush border, wheremembrane/cytoskeleton adhesion must be high to maintain acomplex morphology, higher abundance lipids (e.g., phospha-tidylserine) and alternate, unexplored lipid binding motifswithin the TH1 domain are likely to play a role. Finally,because TH1 domains have been identified in myosin-I genesfrom the earliest eukaryotes (27), the control of membranetension may represent an ancient and conserved function forthese molecular motors.

Materials and MethodsOptical Trap Instrumentation. Our optical trap is built around a single-modediode-pumped solid-state Nd:YVO4 laser (LG Laser Technologies; TEM00, 3 W,� � 1064 nm) that is coupled to a Nikon TE-2000-U inverted light microscopevia optics that are housed in central unit from Molecular Machines & Indus-tries. In addition to the laser head, the central unit contains beam-conditioning optics, z axis focusing lenses, and 2 galvanometer-mountedmirrors for the control of beam position. We used a Nikon PlanFluor 100x/1.3lens (72% transmission in the IR) to focus the laser and form a trap at the focalplace. A motorized X-Y scanning stage (Marzhauser) was used for coursecontrol of sample position. A piezoelectric stage insert (Mad City Labs) pro-vided high-resolution position control with subnanometer accuracy. Scanningstage position, laser power, and laser focal depth (z axis trap position) were allunder computer via software provided by MMI. Images of trapped particleswere captured with a Sony Exwave HAD color CCD using a National Instru-ments PCI-1410 image acquisition card with custom software written in Lab-VIEW 8.5. Time-lapse images acquired at video rate were used to obtain theposition of trapped particles with software developed by Carter et al. (15).Trap stiffness calibration was performed by measuring the excursion of atrapped bead in response to different viscous drag forces applied by movingthe flow chamber with the piezoelectric stage. Stokes’ law (6��rv � kTrap x;where v is solution velocity, � is coefficient of solution viscosity, r is the radiusof the microsphere, and x is microsphere displacement from trap center) wasused to calculate the value of kTrapfor our experiments (Fig. S1).

Brush Border Isolation and Manipulation. Brush borders were isolated usingpreviously described protocol (50). All procedures involving animals wereperformed under the protocols prescribed by the Vanderbilt University Med-ical Center Institutional Animal Care and Use Committee. For membranetether studies, brush borders were typically transferred into a flow cell assem-bled with a glass slide, a coverslip and 2 pieces of doubled-sided tape. Severalvolumes of buffer were applied to the flow cell to flush out brush borders thatwere only loosely anchored to the glass; tether experiments were then carriedout in 75 mM KCl, 10 mM imidazole, 1 mM EGTA, 2.5 mM MgCl2, and 0.01%Na-Azide, pH 7.2.

Cell Culture and Transfections. NGI3 and NIH 3T3 cells were cultured oncoverslips in 6-well plates at 37°C, with 5% CO2. Culture medium consistedof DMEM (Invitrogen) supplemented with 10% fetal bovine serum (Hy-

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Clone) and 2 mM glutamine (GibcoBRL). Transfections were carried out in6-well plates with 6 – 8 �g DNA per 2.5 mL plating media using the reagentLipofectamine2000 (Invitrogen). Cells were assayed 2–3 days post transfec-tion. In a typical experiment, cell-coated coverslips were assembled into aflow chamber using double-sided tape and a glass slide. For all experi-ments, cells were incubated in CO2-independent medium (Invitrogen) at37°C with an objective heater controlled by a TC-124 temperature control-ler (Warner Instruments).

ACKNOWLEDGMENTS. We thank members of the Tyska Laboratory forhelpful suggestions and Dr. Stefan Niehren of Molecular Machines &Industries for outstanding support. This work was supported in part bythe Vanderbilt Digestive Diseases Research Center P30 DK-058404, theVanderbilt University Medical Center Training Program in DevelopmentalBiology (R.E.M.), a predoctoral fellowship from the American Heart Asso-ciation (R.E.M.), a postdoctoral fellowship from the American Heart Asso-ciation (R.N.), and a National Institutes of Health Grant R01-DK075555 (toM.J.T.).

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