MINIREVIEW
Biomechanics, One Molecule at a Time*

Amit D. Mehta, Matthias Rief, and James A. SpudichDagger

From the Department of Biochemistry, Stanford University Medical Center, Stanford, California 94305

    INTRODUCTION
TOP
INTRODUCTION
Processive Motors
Non-processive Motors
Proteins Experiencing...
Conclusion
REFERENCES

Single-molecule observation has come of age. Parallel developments of sensitive mechanical probes and single fluorophore detection now fuse into unique combinations, allowing investigators to examine shape and chemical transitions of single molecules with ever increasing precision and finesse. Optical trapping, using focused laser beams to constrain dielectric particles in solution (for reviews see Ref. 1), has emerged as a widely used and versatile tool to examine mechanically interesting proteins and DNA. The associated forces of light on matter can be rendered sufficiently weak that single molecules compete with them. In most applications to date, molecules of interest are attached to uniform dielectric beads, which are trapped and used as handles to configure an appropriate experimental geometry. One can detect bead position with high precision, monitoring biological activity by tracking probe displacement. Such methods allow accurate, quantitative characterization of force and displacement transients driven or experienced by single molecules, providing a unique edge in deciphering the underlying mechanisms and reaction schemes. Several biomolecules have met variants on this theme. Here, we focus attention on three classes: processive motors, nonprocessive motors, and proteins experiencing significant strain.

    Processive Motors
TOP
INTRODUCTION
Processive Motors
Non-processive Motors
Proteins Experiencing...
Conclusion
REFERENCES

A processive enzyme undergoes multiple productive catalytic cycles per diffusional encounter with its binding partner. Illustrating this, a processive motor protein binds its polymer track and advances along it through several unitary cycles before dissociating (for reviews, see Refs. 2 and 3). Two widely studied examples are kinesin, involved in vesicle transport, and RNA polymerase (RNAP), 1 involved in DNA transcription.

Under in vitro conditions with purified proteins, a single kinesin molecule can move along its microtubule track for several microns before dissociating (4-6). To further characterize this movement, Svoboda et al. (7) attached kinesin at low density to silica beads, captured such a bead with an optical trap, and moved it to close proximity of microtubules fixed on a microscope coverslip (Fig. 1a). They observed discrete advances between dwell positions spaced 8 nm apart (Fig. 1b). Kinesin continued to advance the optically trapped and thus elastically loaded bead until it no longer could, a point at which resistive load is termed the "stall force," measured at 5 pN (7) to 7 pN (8). The bead eventually detached and fell back to the trap center before kinesin rebound the microtubule and began again to pull the bead.


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Fig. 1.   Single-molecule kinesin measurements. a, schematic illustration of the Svoboda et al. (7) kinesin experiment. The motor protein is linked to an optically trapped bead, and the bead is tracked with nanometer and millisecond resolution. The motor moves along its polymer track, pulling the bead behind it. b, sample record, reprinted from Schnitzer and Block (10). The bead advances in 8-nm increments.

These pioneering experiments uncovered the step character and distance but left open the question of correspondence between ATP turnover and the 8-nm advances. To address this, others have approached the kinetic scheme underlying mechanical activity by examining the distribution of dwell periods between unitary advances. Unfortunately, fast stepping or noisy data preclude unambiguous identification of all such transitions. Investigators circumvented this by examining the staircase-like data records at the ensemble level, examining the mean and variance of bead position across the ensemble as a function of time after the records begin (9, 10).

The ensemble average should increase with time, the slope being the average kinesin velocity. The ensemble variance, however, contains information regarding the dwell period distribution. If kinesin stepping proceeds with clocklike regularity, meaning a large number of processes are comparably rate-limiting for each mechanical step, then all kinesin encounters with its track should proceed with the same time course. The variance of bead position across the data record ensemble should remain constant, equaling zero if the records are synchronized. If instead the stepping events are stochastic, individual data records need not follow the same course and the ensemble variance will increase with time. As Svoboda et al. (9) calculate, if a single chemical transition limits the rate of each mechanical step, the variance rises with a slope of velocity times step size. If two kinetically comparable and rate-limiting chemical transitions precede each mechanical step, the variance rises with half that slope. If two mechanical steps follow each rate-limiting chemical transition, the variance rises with twice that slope.

Svoboda et al. (9) used such measurements to demonstrate that under saturating ATP conditions, each mechanical advance is rate-limited by two kinetically comparable processes. Schnitzer et al. (10) performed like experiments under a broad range of ATP concentrations and showed ensemble variance under limited ATP was consistent with a single rate-limiting process, ATP binding, per mechanical advance. This demonstrates that single ATP binding events separate all or nearly all 8-nm advances at all ATP concentrations. Kojima et al. (8) and Hua et al. (11) have reached similar conclusions by fitting the distribution of dwell times separating detected step transitions.

A similar experimental geometry has been used to examine the behavior of kinesin under load either along or against its direction of movement (12). Visscher et al. (13) developed an instrument capable of maintaining a fixed trap-bead separation and thus fixed system tension. One expects that single-molecule experiments using such a technique should clarify the nature of chemomechanical coupling as a function of load, shedding more light on observed slippage and stalling phenomenon. Despite the relative maturity of single-molecule kinesin experiments, more surprises seem likely. Investigators continue to puzzle over how so small a molecule with no visible means of 8-16-nm extension can behave as the above step measurements suggest (for review, see Ref. 14).

In contrast to kinesin, RNAP does not strike one as a molecular motor, as its biological function involves DNA transcription as opposed to moving cargo or generating tension. However, RNA synthesis requires using free energy released from nucleotide condensation for generating force to advance along the DNA template. Because RNAP usually does not dissociate from its track before finishing the transcript, it is a processive molecular motor (for review, see Ref. 15).

RNAP must thread through and presumably rotate a helical DNA strand, precluding the scheme of Svoboda et al. (7), fixing the motor upon a trapped bead and the polymer track upon the surface. Instead, Yin et al. (16) adapted an experiment designed by Shafer et al. (17), in which a surface-mounted RNAP binds and pulls on a solution DNA duplex, attached to a bead on its transcriptionally downstream end. As RNAP advances upon its template, the bead is drawn closer to the surface. Shafer et al. (17) monitored this single transcription reaction by observing the diffusive range of the tethered bead as a function of time. Yin et al. (16) extended this experiment by trapping the bead.


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Fig. 2.   Single-molecule myosin measurements. a, schematic illustration of the Finer et al. (24) myosin single-molecule experiment. An actin filament is attached at either end to a trapped bead. The filament is stretched taut and moved near surface-mounted silica spheres decorated sparsely with myosin molecules. The trapped beads are then tracked for movements parallel to the long actin filament axis. b, simultaneous measurement of myosin binding ATP and actin (29). Top trace shows bead position parallel to the long filament axis. Middle trace shows the stiffness constraining bead diffusion, which rises when a stiff myosin molecule binds the actin filament. Bottom trace shows the fluorescent emission count (F.I.), which rises by 1000 photons/s upon each ATP binding event. ATP binding corresponds to actin release and vice versa. (Reprinted with permission from Ishijima et al. (29).)

In early work, Yin et al. (16) measured the "stall force" against which the molecule ceased movement, analogous to that described above for kinesin. At high optical trap strength, they observed many transcribing RNAP molecules to stall against loads of 12.3 ± 3.5 pN, likely an underestimate because some complexes did not stall and some likely suffered damage from laser light exposure.

Wang et al. (18) used a feedback scheme to increase effective trap strength while reducing light exposure; they measured stall forces of 21-27 pN, considerably above the 5-7 pN required to stall single kinesin motors. RNAP may need such high forces to untangle DNA secondary structure during transcription. Once the feedback system was turned off following stall, RNAP recovered its transcription activity only after a 0-30-s delay. Wang et al. (18) observed intermittent stalls for comparable times when the enzyme moved against low load, suggesting these times reflect transition rates out of a nonproductive state, a phenomenon with no analogue in the kinesin data records.

Finally, Wang et al. (19) corrected for DNA and other elasticity as well as geometry to estimate protein movement along DNA from detected bead displacement (18). They computed RNAP velocity as a function of resistive force, a measurement of central importance in elucidating the reaction steps underlying movement for this and other motor enzymes (20, 21). The load dependence of velocity provides information regarding enzyme behavior in the absence of load. Wang et al. (18) showed velocity remained fairly constant against variable load until it fell off sharply just below the stall force, suggesting that the rate-limiting process under low load does not involve movement along the DNA long axis. From fitting the sharp velocity drop under loads approaching stall, Wang et al. (18) estimated that the force affects the underlying unitary RNAP reaction cycle over a distance spanning at least 5-10 base pairs. This may indicate that optical load induces stall through a conformational strain 5-10 times larger than the distance traversed in a given unitary cycle. Alternatively, load may engage a more complex pathway, perhaps favoring an inactive state that follows slippage by 5-10 base pairs in the transcriptionally upstream direction. Transient pauses in observed RNAP movement records lend further support to the long suggested existence of such a state.

One expects future experiments of like character to examine the influence of specific DNA sequences or protein cofactors on RNAP activity. More precise position detectors will track the enzyme with better than single base pair resolution. Moreover, once the problem of photoinduced enzyme damage is solved and instrumental drift is suitably reduced, the study of slow RNAP movement under limiting nucleotide conditions should allow step and chemomechanical coupling measurements analogous to those described for kinesin. Through these and other experiments, RNAP seems likely to usher in single-molecule study of a broad array of DNA-based proteins.

    Non-processive Motors
TOP
INTRODUCTION
Processive Motors
Non-processive Motors
Proteins Experiencing...
Conclusion
REFERENCES

A non-processive enzyme undergoes at most one productive catalytic cycle per diffusional encounter with its binding partner (for reviews, see Refs. 2 and 22). Although one or a few molecules of non-processive motors cannot transport cargo over long distances, large ensembles of them can move along their tracks at faster rates than comparable ensembles of processive ones. Several members of the myosin superfamily, motors that act on the filamentous polymer actin, can be so classified. Not many but only one unitary advance follows each binding event, demanding a different experimental geometry to capture this advance in detectable probe motion. Simple attachment of binding partners to bead and surface, respectively, would not suffice, because small protein movement would likely cause bead rotation, which is not detectable. Using a dual beam optical trap (23), Finer et al. (24) approached this problem by attaching each end of an actin filament to a polystyrene bead and using the trapped beads as handles. They stretched the filament to tension and moved it near silica spheres, mounted on a coverslip surface and decorated sparsely with skeletal muscle myosin II molecules (Fig. 2a). Myosin bound the filament, often pulled the bead from trap center, and maintained the deflected position for a variable dwell period before releasing (Fig. 2b, top). Because skeletal myosin II detaches from actin only after binding ATP from solution, Finer et al. (24) reduced ATP concentrations to extend the dwell period and clarify bead deflections. Such measured deflections ranged from 10 to 15 nm, more consistent with a tightly coupled mechanical ratchet than some predictions of much larger displacement values (cf. Ref. 25).

This pioneering experiment left open issues regarding probe thermal diffusion, detection of all binding events, protein orientation, myosin mounting on the surface, artifacts of geometry, and compliant linkages in the bead-actin system, all addressed in the work that followed.

All optical trap measurements to date indicate a unitary displacement of 5-15 nm (24, 26-30), though microneedle experiments place the estimate nearer 20 nm (31). In the above trapping experiments, the myosin molecule was oriented randomly with respect to the actin filament. Tanaka et al. (32) have used synthetic muscle-like filaments with very few intact motors to measure displacement as a function of alignment between the long axes of actin and myosin. They report bead displacements near 10 nm when the two are optimally aligned, around 5 nm when the axes are offset around 30°, near zero when the filaments are orthogonal, and, surprisingly, around 5 nm in the same actin filament direction when the proteins are oppositely aligned. Such findings indicate that myosin binds and moves actin in the same direction, albeit by lesser amounts, when constrained geometrically. Moreover, they suggest measurements of randomly oriented myosin may underestimate the unitary step distance.

Ishijima et al. (29) engineered a powerful synthesis of the Finer et al. (24) optical trapping geometry with total internal reflection microscopy, a technique used by Funatsu et al. (33) to image single fluorophores. They tracked simultaneously myosin mechanical activity and the diffusion of fluorescent Cy3-ATP into the focal plane. Working at the low (100 nM) Cy3-ATP concentrations required to reduce background fluorescence and render single fluorophores visible, Ishijima et al. (29) observed data records as shown in Fig. 2b. Myosin binding actin, as detected by an increase in stiffness constraining bead motion (Fig. 2b, middle), occurred coincident with a loss of 1000 detected Cy3 emission photons per second (Fig. 2b, bottom), indicating myosin released a single Cy3-nucleotide. Myosin releasing actin, detected by a drop in system stiffness, corresponded with the arrival of a single Cy3-ATP in the focal plane and presumably the binding of myosin by Cy3-ATP. By observing at once this mechanical and chemical activity of a single molecule, Ishijima et al. (29) sought to examine directly the issue of coupling between myosin displacement and ATP turnover.

The measured 15-nm deflections corresponded consistently with catching and releasing a single ATP. Ishijima et al. (29) left open the question of whether these deflections are driven by one or more cyclical conformational changes in the myosin head. Although unitary displacement estimates near 5 nm have seemed most consistent with structure-based predictions of movement from a single conformational change (34), those around 10-15 nm are not impossible to reconcile with them.

Moreover, Ishijima et al. (29) observed that in a significant minority of binding events, apparent ATP release preceded binding and moving the actin by several hundred milliseconds. They also observed that in the absence of ATP, myosin bound but did not move the actin filament. Based on this, they suggested a "hysteretic state," in which myosin preserves the memory of a recent ATP hydrolysis, using residual energy to move the actin filament. Such a state likely does not affect interaction cycles under physiological conditions, where actin binding occurs at a faster rate than nucleotide release. However, it does suggest myosin is capable of maintaining a long lived, energized chemical state following release of hydrolysis products in the absence of actin. Although dye photobleaching could also explain these apparently premature nucleotide dissociations, the authors argue against it.

This combination of optical trapping with single fluorophore detection is likely to inspire followers. For instance, Suzuki et al. (35) have recently observed conformational changes in the myosin head by tracking resonant energy transfer between fluorophores attached to the termini of the motor domain. Warshaw et al. (36) have used a spot confocal microscope to monitor changes in fluorescence polarization from a 6'-iodoacetamidotetramethylrhodamine probe linked to the neck region of single myosin molecules. In combination with optical trapping, such techniques should allow simultaneous measurement of myosin shape changes and the displacements they produce or perhaps even myosin shape changes induced by stress applied.

The future should see extension of these techniques to different forms and mutants of myosin. Along these lines, Guilford et al. (27) have observed that a single smooth muscle myosin molecule holds actin, after pulling it, for much longer than skeletal muscle myosin, explaining its slower speed of contraction and the higher forces it generates. One also expects increasingly precise measurements of myosin moving against load. Although some have attempted to measure the force produced by myosin under isometric conditions, such experiments have faced limits from feedback system performance (37) and compliant linkages separating the actin from the optically trapped bead (28, 30, 38). Once such hurdles are overcome investigators will measure force generated by myosin throughout its putative conformational change and observe its mechanical response to a sudden shift in load after binding the actin, analogous to seminal experiments with whole muscle fibers (39).

    Proteins Experiencing Significant Strain
TOP
INTRODUCTION
Processive Motors
Non-processive Motors
Proteins Experiencing...
Conclusion
REFERENCES

To observe displacement driven by single molecules, one must use probes sufficiently compliant that they do not perturb the structure or chemistry. In other experiments one can impose high loads by pulling a stiff probe to disrupt biomolecular structure in controlled ways (40-43). Smith et al. (40) and Cluzel et al. (41) have used relatively strong optical tweezers or microneedles to pull double-stranded B-DNA, inducing a cooperative transition to a structurally distorted conformation termed "overstretched" or "S-DNA." Others have pulled apart single ligand-receptor complexes, for instance antibody-antigen (44, 45), biotin-avidin (46, 47), or actomyosin (48). Tension dependence of detachment rates provides information regarding the binding chemistry (49, 50).

Three research groups (51-53) employing like schemes have induced reversible unfolding of isolated domains in the 3-MDa muscle protein titin (54), which consists largely of a series of 200 structurally similar immunoglobulin (Ig) and fibronectin III (Fn3) domains (55). In striated muscle, titin provides an elastic link that maintains structural integrity of the sarcomere under tension. Much of this elasticity derives from the PEVK region, a putative random coil segment of the molecule (56). However, investigators have long speculated that domains in titin may unfold reversibly to effect the large length changes required in passive muscle stretching. To explore such behavior at the single-molecule level, experimentalists fixed one end of an isolated titin molecule and pulled on the other.

Two groups pulled titin with optical tweezers (Fig. 3a) (51, 53), compliant probes offering sub-pN force resolution. A third group (52) used 100-1000-fold more stiff AFM cantilevers. The stiffness of probes used determined the type of data one could extract, with different methods yielding complementary results.


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Fig. 3.   Single-molecule titin measurements. a, schematic illustration of pulling titin with an optically trapped bead. b, force extension curve of native titin generated by Kellermeyer et al. (51), using a compliant optical trap (reprinted with permission from Kellermayer et al. (51)). c, force extension curve of native titin generated by Tskhovrebova et al. (53), using a less compliant trap to apply sudden force jumps and then tracking stepwise relaxation (reprinted with permission from Tskhovrebova et al. (53)). d, force extension curve of truncated titin construct generated by Rief et al. (52), using a stiff AFM cantilever (reprinted with permission from Rief et al. (52)).

Fig. 3b shows a force-extension curve for a single titin molecule through a stretch and relax cycle, induced and measured using a compliant optical trap (51). Until the molecule extends to its fully folded length (1-2 µm), entropic restoring forces (51, 53) dominate the response of titin to imposed stress. Further extension leads to a steep rise in resistive force up to around 30 pN, beyond which the protein offers less resistance to continued expansion. Kellermayer et al. (51) argue this reflects domain unfolding in the molecule. However, they could not visualize discrete domain unfolding events, because the compliant traps do not offer required spatial discrimination. This approach differed from that of Tskhovrebova et al. (53), who used a stiffer trap to apply rapidly a force over 100 pN to a single titin molecule. They observed the time course of the mechanical response of titin to such force transients (Fig. 3c). The molecule expanded in discrete steps of 19 ± 10 nm, the added contour length expected to follow a single domain unfolding. Such discrete unfolding events were most readily observed in the AFM records generated by Rief et al. (52) (Fig. 3d). Pulling on titin with a stiff cantilever and tracking its resistance, they observed a sawtooth pattern with edges presumed to reflect single domain unfolding. Rief et al. (57) demonstrated that the spatial resolution of a stiff cantilever allows them to map the measured length changes to unfolded polypeptide stretches with single amino acid resolution.

The three groups observed titin domains unfolding under different forces, ranging from 30 to 300 pN. In part, this reflects that not all of the 200 domains have the same fold stability. However, the widely used term "unfolding force" is a misnomer; domains unfold eventually under the slightest force, or even no force, if one waits long enough. The rate constant reflecting the stochastic transition from folded to unfolded depends exponentially on the applied force and induced strain. Discrepancy in "unfolding force" observed by Tskhovrebova et al. (53) and Rief et al. (52) therefore mainly derives from the different pulling speeds and thus time windows used in the experiments (58).

Similar experiments have shown recently that all-beta structures, like Ig domains in titin and Fn3 domains in titin and tenascin, resist unfolding to a higher force (57, 59) than the all-alpha domains of spectrin (60), when pulled at the same speed.

    Conclusion
TOP
INTRODUCTION
Processive Motors
Non-processive Motors
Proteins Experiencing...
Conclusion
REFERENCES

A broad variety of biologically interesting molecules have undergone study using optical traps to position, constrain, deform, or otherwise manipulate them. Targets for such study include proteins that exert force, motors, or those biomolecules that respond to it in interesting ways. The experiments described above represent a new class of measurements, which share common concepts, level of detail, and dimensions of movement and force. Future experiments in every class described here will likely draw upon fluorescent methods to observe at once protein shape changes and the associated binding events and physical displacements.

Investigators in these areas will continue to face similar problems, among them imperfect protein mounting, unwanted system compliances, limits in detector or actuator resolution or bandwidth, effects of probe thermal diffusion, biases in data analysis, instrumental drift, mechanically or optically induced protein damage, and equipment costs. As these and other problems find solutions, one expects that the kind of measurements described here should grow increasingly precise and the resulting mechanistic understanding increasingly detailed.

    FOOTNOTES

* This minireview will be reprinted in the 1999 Minireview Compendium, which will be available in December, 1999. This is the first article of four in the "Biochemistry at the Single-molecule Level Minireview Series."

Dagger To whom correspondence should be addressed. Tel.: 650-723-7634; Fax: 650-725-6044; E-mail: jspudich{at}cmgm.stanford.edu.

    ABBREVIATIONS

The abbreviations used are: RNAP, RNA polymerase; pN, piconewton(s); AFM, atomic force microscope.

    REFERENCES
TOP
INTRODUCTION
Processive Motors
Non-processive Motors
Proteins Experiencing...
Conclusion
REFERENCES
  1. Sheetz, M. (ed) (1998) Laser Tweezers in Cell Biology, Methods in Cell Biology, Vol. 55, Academic Press, Orlando, FL
  2. Howard, J. (1997) Nature 389, 561-567[CrossRef][Medline] [Order article via Infotrieve]
  3. Lohman, T. M., Thorn, K., and Vale, R. D. (1998) Cell 93, 9-12[Medline] [Order article via Infotrieve]
  4. Howard, J., Hudspeth, A. J., and Vale, R. D. (1989) Nature 342, 154-158[CrossRef][Medline] [Order article via Infotrieve]
  5. Block, S. M., Goldstein, L. S., and Schnapp, B. J. (1990) Nature 348, 348-352[CrossRef][Medline] [Order article via Infotrieve]
  6. Vale, R. D., Funatsu, T., Pierce, D. W., Romberg, L., Harada, Y., and Yanagida, T. (1996) Nature 380, 451-453[CrossRef][Medline] [Order article via Infotrieve]
  7. Svoboda, K., Schmidt, C. F., Schnapp, B. J., and Block, S. M. (1993) Nature 365, 721-727[CrossRef][Medline] [Order article via Infotrieve]
  8. Kojima, H., Muto, E., Higuchi, H., and Yanagida, T. (1997) Biophys. J. 73, 2012-2022[Abstract]
  9. Svoboda, K., Mitra, P. P., and Block, S. M. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 11782-11786[Abstract/Free Full Text]
  10. Schnitzer, M. J., and Block, S. M. (1997) Nature 388, 386-390[CrossRef][Medline] [Order article via Infotrieve]
  11. Hua, W., Young, E. C., Fleming, M. L., and Gelles, J. (1997) Nature 388, 390-393[CrossRef][Medline] [Order article via Infotrieve]
  12. Coppin, C. M., Pierce, D. W., Hsu, L., and Vale, R. D. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 8539-8544[Abstract/Free Full Text]
  13. Visscher, K., and Block, S. M. (1998) Methods Enzymol. 298, 460-489[Medline] [Order article via Infotrieve]
  14. Block, S. M. (1998) Cell 93, 5-8[Medline] [Order article via Infotrieve]
  15. Gelles, J., and Landick, R. (1998) Cell 93, 13-16[Medline] [Order article via Infotrieve]
  16. Yin, H., Wang, M. D., Svoboda, K., Landick, R., Block, S. M., and Gelles, J. (1995) Science 270, 1653-1657[Abstract]
  17. Schafer, D. A., Gelles, J., Sheetz, M. P., and Landick, R. (1991) Nature 352, 444-448[CrossRef][Medline] [Order article via Infotrieve]
  18. Wang, M. D., Schnitzer, M. J., Yin, H., Landick, R., Gelles, J., and Block, S. M. (1998) Science 282, 902-907[Abstract/Free Full Text]
  19. Wang, M. D., Yin, H., Landick, R., Gelles, J., and Block, S. M. (1997) Biophys. J. 72, 1335-1346[Abstract]
  20. Svoboda, K., and Block, S. M. (1994) Cell 77, 773-784[Medline] [Order article via Infotrieve]
  21. Hunt, A. J., Gittes, F., and Howard, J. (1994) Biophys. J. 67, 766-781[Abstract]
  22. Spudich, J. A. (1994) Nature 372, 515-518[Medline] [Order article via Infotrieve]
  23. Simmons, R. M., Finer, J. T., Chu, S., and Spudich, J. A. (1996) Biophys. J. 70, 1813-1822[Abstract]
  24. Finer, J. T., Simmons, R. M., and Spudich, J. A. (1994) Nature 368, 113-119[CrossRef][Medline] [Order article via Infotrieve]
  25. Harada, Y., Sakurada, K., Aoki, T., Thomas, D. D., and Yanagida, T. (1990) J. Mol. Biol. 216, 49-68[Medline] [Order article via Infotrieve]
  26. Molloy, J. E., Burns, J. E., Kendrick-Jones, J., Tregear, R. T., and White, D. C. (1995) Nature 378, 209-212[CrossRef][Medline] [Order article via Infotrieve]
  27. Guilford, W. H., Dupuis, D. E., Kennedy, G., Wu, J., Patlak, J. B., and Warshaw, D. M. (1997) Biophys. J. 72, 1006-1021[Abstract]
  28. Mehta, A. D., Finer, J. T., and Spudich, J. A. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 7927-7931[Abstract/Free Full Text]
  29. Ishijima, A., Kojima, H., Funatsu, T., Tokunaga, M., Higuchi, H., Tanaka, H., and Yanagida, T. (1998) Cell 92, 161-171[Medline] [Order article via Infotrieve]
  30. Veigel, C., Bartoo, M. L., White, D. C., Sparrow, J. C., and Molloy, J. E. (1998) Biophys. J. 75, 1424-1438[Abstract/Free Full Text]
  31. Ishijima, A., Kojima, H., Higuchi, H., Harada, Y., Funatsu, T., and Yanagida, T. (1996) Biophys. J. 70, 383-400[Abstract]
  32. Tanaka, H., Ishijima, A., Honda, M., Saito, K., and Yanagida, T. (1998) Biophys. J. 75, 1886-1894[Abstract/Free Full Text]
  33. Funatsu, T., Harada, Y., Tokunaga, M., Saito, K., and Yanagida, T. (1995) Nature 374, 555-559[Medline] [Order article via Infotrieve]
  34. Rayment, I., Rypniewski, W. R., Schmidt-Base, K., Smith, R., Tomchick, D. R., Benning, M. M., Winkelmann, D. A., Wesenberg, G., and Holden, H. M. (1993) Science 261, 50-58[Medline] [Order article via Infotrieve]
  35. Suzuki, Y., Yasunaga, T., Ohkura, R., Wakabayashi, T., and Sutoh, K. (1998) Nature 396, 380-383[CrossRef][Medline] [Order article via Infotrieve]
  36. Warshaw, D. M., Hayes, E., Gaffney, D., Lauzon, A. M., Wu, J., Kennedy, G., Trybus, K., Lowey, S., and Berger, C. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 8034-8039[Abstract/Free Full Text]
  37. Mehta, A. D., Finer, J. T., and Spudich, J. A. (1998) Methods Cell Biol. 55, 47-69[Medline] [Order article via Infotrieve]
  38. Dupuis, D. E., Guilford, W. H., Wu, J., and Warshaw, D. M. (1997) J. Muscle Res. Cell Motil. 18, 17-30[CrossRef][Medline] [Order article via Infotrieve]
  39. Huxley, A. F., and Simmons, R. M. (1971) Nature 233, 533-538[Medline] [Order article via Infotrieve]
  40. Smith, S. B., Cui, Y., and Bustamante, C. (1996) Science 271, 795-798[Abstract]
  41. Cluzel, P., Lebrun, A., Heller, C., Lavery, R., Viovy, J.-L., Chateny, D., and Caron, F. (1996) Science 271, 792-794[Abstract]
  42. Essevaz-Roulet, B., Bockelmann, U., and Heslot, F. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 11935-11940[Abstract/Free Full Text]
  43. Rief, M., Oesterhelt, F., Heymann, B., and Gaub, H. E. (1997) Science 275, 1295-1297[Abstract/Free Full Text]
  44. Dammer, U., Hegner, M., Anselmetti, D., Wagner, P., Dreier, M., Huber, W., and Güntherodt, H.-J. (1995) Biophys. J. 70, 2437-2441[Abstract]
  45. Hinterdorfer, P., Baumgartner, W., Gruber, H. J., Schilcher, K., and Schindler, H. (1996) Proc. Natl. Acad. Sci. U. S. A. 93, 3477-3481[Abstract/Free Full Text]
  46. Florin, E.-L., Moy, V. T., and Gaub, H. E. (1994) Science 264, 415-417[Medline] [Order article via Infotrieve]
  47. Lee, G. U., Kidwell, D. A., and Colton, R. J. (1994) Langmuir 10, 354-357
  48. Nishizaka, T., Miyata, H., Yoshikawa, H., Ishiwata, S., and Kinosita, K., Jr. (1995) Nature 377, 251-254[CrossRef][Medline] [Order article via Infotrieve]
  49. Bell, G. I. (1978) Science 200, 618-627[Medline] [Order article via Infotrieve]
  50. Evans, E., and Ritchie, K. (1997) Biophys. J. 72, 1541-1555[Abstract]
  51. Kellermayer, M. S., Smith, S. B., Granzier, H. L., and Bustamante, C. (1997) Science 276, 1112-1116[Abstract/Free Full Text]
  52. Rief, M., Gautel, M., Oesterhelt, F., Fernandez, J. M., and Gaub, H. E. (1997) Science 276, 1109-1112[Abstract/Free Full Text]
  53. Tskhovrebova, L., Trinick, J., Sleep, J. A., and Simmons, R. M. (1997) Nature 387, 308-312[CrossRef][Medline] [Order article via Infotrieve]
  54. Maruyama, K., Matsubara, S., Natori, R., Nonomura, Y., Kimura, S., Ohashi, K., Murakami, F., Handa, S., and Eguchi, G. (1977) J. Biochem. (Tokyo) 82, 317-337[Medline] [Order article via Infotrieve]
  55. Labeit, S., and Kolmerer, B. (1995) Science 270, 293-296[Abstract]
  56. Gautel, M., and Goulding, D. (1996) FEBS Lett. 385, 11-14[CrossRef][Medline] [Order article via Infotrieve]
  57. Rief, M., Gautel, M., Schemmel, A., and Gaub, H. E. (1998) Biophys. J. 75, 3008-3014[Abstract/Free Full Text]
  58. Rief, M., Fernandez, J. M., and Gaub, H. E. (1998) Phys. Rev. Lett. 81, 4764-4767[CrossRef]
  59. Oberhauser, A. F., Marszalek, P. E., Erickson, H. P., and Fernandez, J. M. (1998) Nature 393, 181-185[CrossRef][Medline] [Order article via Infotrieve]
  60. Rief, M., Pascual, J., Saraste, M., and Gaub, H. E. (1999) J. Mol. Biol. 286, 553-561[CrossRef][Medline] [Order article via Infotrieve]


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