From the Department of Biochemistry, Stanford University Medical Center, Stanford, California 94305
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.
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.
INTRODUCTION
TOP
INTRODUCTION
Processive Motors
Non-processive Motors
Proteins Experiencing...
Conclusion
REFERENCES
Processive Motors
TOP
INTRODUCTION
Processive Motors
Non-processive Motors
Proteins Experiencing...
Conclusion
REFERENCES
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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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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.
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Non-processive Motors |
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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).
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Proteins Experiencing Significant Strain |
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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. 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- structures, like
Ig domains in titin and Fn3 domains in titin and tenascin, resist
unfolding to a higher force (57, 59) than the all-
domains of
spectrin (60), when pulled at the same speed.
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Conclusion |
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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.
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FOOTNOTES |
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* 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."
To whom correspondence should be addressed. Tel.: 650-723-7634;
Fax: 650-725-6044; E-mail: jspudich{at}cmgm.stanford.edu.
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ABBREVIATIONS |
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The abbreviations used are: RNAP, RNA polymerase; pN, piconewton(s); AFM, atomic force microscope.
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