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43kinesin illuminated

Chapter 2.2SINGLE-MOLECULE SPECTROSCOPY OF MOTOR PROTEINS

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44 single-molecule spectroscopy of motor proteins

2.2.1 Abstract

Single-molecule fluorescence spectroscopy has become a crucial tool to study the behavior of biomolecules and their roles in complex biological processes. Single- molecule methods have in particular contributed to our understanding of the mechanism of force generation and motility of motor proteins. Motor proteins are enzymes that convert the chemical free energy obtained from ATP hydrolysis in mechanical work, in order to drive processes such as muscle contraction and intracellular transport. In this chapter, I review how single-molecule fluorescence spectroscopy can be applied to motor proteins of the kinesin, myosin and dynein superfamilies and how such studies have advanced our knowledge of the molecu-lar basis of motor-protein action. 2.2


INTRODUCTION

Over the last two decades, single-molecule studies have become an indispensable tool for studying the properties of biomolecules (1). Single-molecule methods have several key advantages over bulk measurements: (i) they allow straightfor-ward resolution of heterogeneity, (ii) they allow correlating distinct properties of biomolecules of interest, (iii) they allow direct observation of stochastic changes of properties, without the need for synchronizing all the molecules in the sam-ple, (iv) they allow studying proteins that, in the cell, act on their own (2, 3, 4). Different single-molecule methods have been developed. Several of these techniques are well suited to measure forces acting on biomolecules and/or to apply forces to them: optical tweezers, magnetic tweezers and atomic force microscopy (5). These methods have been used extensively to determine the mechanical stability of proteins, DNA and RNA, and to determine the forces generated by enzymes such as DNA and RNA polymerase moving along a DNA track. Such measurements, in the biologically highly relevant picoNewton force range, were not possible before and have opened up a new perspective on the mechanics of biomolecules (5). Another very popular, but quite different approach is single- molecule fluorescence spectroscopy. Here the fluorescence emitted by a probe, part of or attached to the biomolecule of interest, is detected and processed to extract detailed information on behavior and properties of the biomolecule while no mechanical forces are involved. Fluorescence detection can provide real-time information with millisecond temporal resolution and nanometer spatial preci-sion, without substantial disturbance of the molecule’s behavior. Single-molecule fluorescence methods have been applied to a wide variety of biological and non-biological systems, shedding light on processes such as protein and RNA folding and conformational changes, the mechanism of enzymes and DNA- processing proteins, the diffusion of proteins in a cell membrane, a virus invading a cell and many others (1).

In this chapter the application of single-molecule fluorescence spectroscopy to a specific class of dynamic proteins, namely motor proteins, is discussed. In fact, the development of single-molecule methods, in particular optical tweezers and single-molecule fluorescence spectroscopy, has been closely linked with our in-creased knowledge about the mechanisms of this class of proteins (6). An over-view is provided of the biochemical assays and instrumentation required, and applications of single-molecule spectroscopy to motor proteins are reviewed. The

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chapter ends with an outlook of the potential future development. In the follow-ing, I will discuss the biological role of motor proteins, their general properties and focus on the three motor superfamilies: kinesin, myosin and dynein.

MOTOR PROTEINS INVOLVED IN INTRACELLULAR TRANSPORT

Eukaryotic cells are generally too big and too complex to fully rely on diffusion for the transport of macromolecules, proteins and organelles. To overcome this limitation, the cell makes use of sophisticated and ingenious transport systems where cargo is actively and efficiently transported in a directional way along the cytoskeleton (7). The cytoskeleton is a complex network that provides mechanical strength and structure to the cell (8). The cytoskeleton network consists of three classes of filaments: intermediate filaments (i.e. elastin, vimentin, lamin), microtubules and actin filaments (9). The latter two also function as tracks to the cell’s transport systems. Microtubules are tube-like structures built out of heterodimeric tubulin subunits. Microtubules are polarized structures: in inter-phase, the minus-end is usually located close to the nucleus, whereas the plus-ends extend to the periphery of the cell. Actin filaments are built out of actin-subunits and are found right underneath the cell cortex. Actin filaments are also polarized: they have a plus (“barbed”) and a minus (“pointed”) end. Many cell types have elongated protrusions such as axons, dendrites, cilia and flagella, ranging from several micrometers up to meters. These protrusions are generally rich of micro-tubules and are maintained by specialized directional transport mechanisms such as axonal transport (10) and intraflagellar transport (11, 12).

Active, directional transport of cargoes along actin filaments and microtubules is driven by motor proteins, using the hydrolysis of adenosine triphosphate (ATP) to adenosine diphosphate (ADP) as their source of free energy (13). Most motor proteins involved in intracellular transport are processive: this means that du-ring a single encounter with the track, they make hundreds of steps in a given direction along the track before releasing. This chapter is focused on processive motors, since interaction times and displacements of non-processive ones are often too short to be detected with single-molecule fluorescence spectroscopy. Eukaryotic cells contain three superfamilies of motor proteins: the myosins (which interact with actin filaments), dyneins and kinesins (which both interact with microtubules). In the next paragraphs the properties of these three super-families will be discussed in more detail.

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2.2.2 The kinesin superfamily

The kinesin superfamily consists of fourteen families of motor proteins that all use the microtubules as track and in most cases move towards the plus-end of the microtubule (14, 15). All members of the superfamily share sequence homology with Kinesin-1 (conventional kinesin), the founding member of the superfam-ily discovered in 1985 (16). Kinesin-1 is a tetrameric protein consisting of two identical light chains involved in cargo binding and two identical heavy chains (Fig. 1A). The heavy chains contain a long stalk (responsible for dimer formation) and a globular motor domain, that takes care of ATP hydrolysis, microtubule binding and undergoes nucleotide dependent conformational changes that drive motility (17). The biological function of Kinesin-1 is transport of membrane- enclosed organelles in axons (15).

A B C

Kinesin-1 Myosin VI Myosin V Dynein

D

Figure 1: Schematic representation of the structures of different motor proteins. (A) Kinesin-1 dimerizes through a coiled-coil structure that connects the motor domains (blue) to the cargo binding domain (brown). (B) Myosin V dimerizes through a coiled-coil structure that connects the motor domains (blue) to the cargo binding domain (brown). The myosin V lever arm, which connects the motor domain to the tail domain, contains six IQ motifs that all bind a single calmodulin light chain (red). (C) Myosin VI dimerizes through a coiled-coil structure that connects the motor domains (blue) to the cargo binding domain (brown). The myosin VI lever arm, which connects the motor domain to the tail domain, contains a unique insert that binds a calmodulin with a calcium ion bound to it (yellow), one conventional IQ motif that binds a single calmodulin light chain (red), and a three-helix bundle that can extend the lever arm. (D) The dynein AAA+ domains (blue) are organized in a hexameric ring from which a stalk extends to the microtubule binding domains (green). Dynein dimerizes through a tail domain interacting with multiple associated subunits (purple) and a cargo bin-ding domain (brown).

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The sequence homology between kinesin families lies in the motor domain. Several other kinesin families, for example Kinesin-2 and Kinesin-3 have over-all structures resembling that of Kinesin-1 (i.e. two globular motor domains connected with a long coiled coil tail) and have similar function in intracellular transport (although some Kinesin-3’s seem to be monomeric) (15). Others look similar, but have quite different functions: e.g. Kinesin-13 is involved in mi-crotubule depolymerization during mitosis (15). A special case is Kinesin-5, a homotetrameric kinesin with two motor domains on each end of a central stalk that is involved in crosslinking and sliding of microtubules in the mitotic spindle (15). Kinesin-14 members have a similar function but are dimeric. They are the only kinesins with their motor domains attached to the C-terminus of a coiled coil stalk (for other kinesins it is the N-terminus of the stalk). Surprisingly, this makes the motor move in the minus-end direction of microtubules, opposite to most other kinesins (15). The kinesins are small compared to the other motor families. This makes them easier to express and manipulate. Single-molecule flu-orescence methods have, as a consequence, been first developed for Kinesin-1 and later been applied to other motors. A central theme has been to relate differences in motility parameters to the differences in structures of distinct motor proteins.

2.2.3 The myosin superfamily

The myosin superfamily consists of eighteen families of motor proteins (8, 9, 18). The most well-known is myosin II, the motor protein that drives muscle con-traction by hydrolyzing ATP (18). In contrast to most kinesins, myosin II is not processive: a single myosin II interacts only very briefly with an actin filament, binding, generating a power stroke and releasing. Muscle contraction is driven by many mechanically connected myosin II motors working together (9). Mem-bers of the myosin superfamily consist of homologous N-terminal actin-binding motor domains, containing the active sites for ATP hydrolysis and force generation. Important parts of myosin motor domains are very similar to those of kinesins, very likely reflecting that these two motor superfamilies share a common evolutionary origin (19). Myosin polypeptides furthermore contain lever arms of varying length, in most cases, dimerization domains and C-terminal cargo- binding domains (9). The myosin families that are discussed in more detail in this review do not function in muscle action; they are involved in intracellular trans-port along the actin cytoskeleton. These motors, myosin V and VI, consist of two identical motor domains, are processive and, as a consequence, compatible with

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single-molecule fluorescence motility assays (Fig. 1B, C). Myosin V is involved in the intracellular transport of vesicles (20) and moves along actin filaments in the plus-end direction. Myosin VI is the only known minus-end directed myosin (21). It plays a key role in intracellular transport and is involved in processes as diverse as hearing, cancer metastasis and endocytosis (22).

2.2.4 The dynein superfamily

Dyneins are motor proteins that move to the minus-end of microtubules. In many ways, dyneins are different from kinesins and myosins. First of all, they are significantly larger (more than a mega Dalton) and contain many large, accessory subunits. Second, the core subunit of dynein is the Dynein Heavy Chain (DHC), which consists of six AAA+ (ATPase associated with diverse cellular activities) domains in a hexameric ring, and two domains protruding from this ring: a microtubule-binding domain, and a large tail domain (Fig. 1D). This tail domain is responsible for dimerization and is the site of binding for accessory subunits (23). These accessory subunits are called Intermediate Chains (ICs), Light Chains (LCs) and Light-Intermediate Chains (LICs) and are involved in motor regulation and cargo binding, but are not essential for motility in vitro (23, 24). In contrast to the other motor superfamilies, only a limited number of dis-tinct dynein motor classes have been identified (25). One class, axonemal dynein, slides and bends microtubules in axonemes and thus drives the beating of motile cilia (9, 24). The other class, cytoplasmic dynein, is involved in the transport of a wide range of cargoes for example in dendritic transport (26) and intraflagellar transport in cilia (12). Moreover, dynein plays an important role in cell division by performing functions such as the positioning of the mitotic spindle (24).

METHODS TO STUDY MOTOR PROTEINS

Understanding the biological function and mechanism of motor proteins re-quires precise knowledge of their properties and behavior. Determination of general motility parameters is often not sufficient, because of heterogeneity in the motor population: some of the motors might be bound to a microtubule, others not; furthermore, the motors generally do not work in the same pace or rhythm and it can be difficult to synchronize them. The last two decades single-molecule fluorescence microscopy has emerged as a crucial technique for measuring the characteristics and behavior of individual motor proteins, eliminating the need

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to synchronize motors, providing direct access to quantifying heterogeneous be-havior, and allowing these proteins to be measured in more natural conditions: in the cell, they often work on their own, or in small numbers (1). The parallel development of sophisticated motility assays was equally important (6). The ear-liest assays to probe motor function relied on measuring the energy consumption of the motor system as a whole. Advancement of motor assays allowed for in vitro reconstitution of motor transport systems using only a subset of components. Further developments of fluorescence microscopy allowed for detection of sin-gle fluorescently labeled proteins under ambient conditions. These developments were essential for accurate in vitro reconstitution of motor transport systems that allow for studying single motor activity. Below I will give an overview of key experiments and techniques that contributed to the evolution of motor assays.

2.2.5 ATPase assays of motor proteins

As discussed above, motor proteins use the free energy released from ATP hydrolysis to induce conformational changes that allow the motor to generate force. A catalytic site ensures efficient ATP turnover so that typically ~10-100 ATP mole-cules can be hydrolyzed every second. This allows kinesin, for example, to move along the microtubule track at 0.8 µm s-1, thereby hydrolyzing one ATP each 8 nm step. In order to prevent motors from consuming ATP unnecessarily their enzymatic activity is tightly coupled to track interactions. This ATP dependence of motor proteins was first described for myosin motors during muscle contrac-tion (27), a discovery that led to the development of ATPase assays. In such assays, the ATP consumption by myosin or actomyosin was monitored at specific time intervals (28). ATPase assays proved to be very insightful for kinetic analysis of motor proteins. As a result, different ATP and ADP nucleotide analogs were developed to probe the ATPase cycle (29, 30). One of these nucleotides was a non-hydrolyzable analog of ATP, called adenylyl imidodiphospate (AMP-PNP), which was shown to be a competitive inhibitor of the ATPase cycle of myosin (31). This specific nucleotide would fulfill a crucial role in the development of motility assays.

2.2.6 Multi-motor gliding and bead assays

In 1983, a breakthrough in the development of motor assays was made by Sheetz and Spudich who showed that myosin-coated beads move unidirec-

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tional along exposed actin filaments of the alga Nitella (32). This was a pivotal step as it demonstrated that the motility of motor proteins could be reconsti-tuted in vitro in cell-free conditions. Furthermore, it provided insight into the general intracellular transport activities that already had been observed for a long time using bright-field microscopy (33). Around the same time, microscopy was revolutionized through the development of video-enhanced differential inter-ference contrast (VE-DIC) microscopy, which allowed for the observation and recording of objects substantially smaller than the wavelength of light (34, 35). VE-DIC was used to show that small particles of about a tenth of a micron move in anterograde or retrograde direction parallel to linear elements inside the squid giant axon (36). These linear elements were later identified as microtubules capable of supporting transport in both directions (37). Amazingly, this microtubule transport system could be reconstituted in vitro using purified microtubules and axoplasmic (cytoplasm from axon) supernatant, yielding microtubules gliding along the glass coverslip (38). This was the first description of the microtubule gliding assay, which would later become a widely used experimental technique for studying behavior of groups of motor proteins (18).

In 1985, the work described above would lead to the discovery and purification of kinesin (16, 39, 40). For purification the researchers relied on blocking of the kinesin ATPase cycle by AMP-PNP. After addition of AMP-PNP kinesin get stuck to the microtubules, which permitted spinning down the motor proteins in the form of microtubule-motor complexes. Subsequently, ATP was added to re-lease the motor proteins. This microtubule-affinity purification procedure is still widely used for purification. Having access to purified motor proteins was a pre-requisite for setting up minimal motility assays to probe motor activity in a con-trolled in vitro environment, opening up a whole new world of studies (41). In gliding assays, microtubules are transported by motor proteins that are adsorbed to the glass surface (Fig. 2A). The movement of microtubules can be observed using VE-DIC or fluorescence microscopy and motility characteristics such as velocity can be readily obtained. Polarity-marked fluorescent microtubules can be used to determine the directionality of the motor proteins.

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A B

Gliding assay Bead assay

Optical trapping assay Single-motor assay

Sliding assay Multi-motor assembly assay

C D

E F

Figure 2: Schematic representations of different in vitro motility assays. Arrows indicate moving elements and direction of movement. (A) A gliding assay where dimeric motor proteins (blue) bound to a coverslip transport a microtubule (green) along the surface of the glass. (B) A bead assay where a bead (grey) covered with motor proteins (blue) moves along a microtubule (green). (C) An optical trapping assay, similar to the bead assay, but now with an optically trapped bead (grey). A motor protein (blue) exerts force on the bead, pulling it out of the center of the optical trap (red). The trap can be used to apply a force to the motor pro-tein and measure the force it generates. (D) A single-motor assay where single motor proteins (blue) temporarily interact and walk along a microtubule that is bound to a coverslip. (E) A sliding assay where homotetrameric motor proteins (blue) move along a microtubule (green) that is bound to the glass and another microtubule (green) that is sliding along the surface of the glass. (F) A multi-motor assembly assay where trains containing different types of kinesin motors (blue and red) move along microtubules (green) that are attached to the glass.

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The bead assay inverts the microtubule-motor geometry by adherence of the microtubules to the glass and using microspheres coated with motor proteins to reconstitute motility (Fig. 2B). In this assay, it is the bead that moves, while the tracks remain stationary. Bead movement can be readily tracked using VE-DIC, or other bright field or fluorescence microscopy methods. Multi-motor gliding and bead assays have been crucial for our understanding of motor-protein driven transport (18). It is important to realize that these assays probe the motility driven by an often unknown multiple of motor proteins and not by a single one.

2.2.7 Optical tweezers experiments with motor proteins

The biological function of motor proteins is to generate mechanical work, i.e. to drive displacement working against a force. The method of choice to measure forces generated by and to apply forces to motor proteins is optical tweezers (5, 42, 43). In optical-tweezers experiments on processive motors, the bead assay discussed above is applied. Instead of only tracking bead displacement using bright-field or fluorescence microscopy, optical tweezers are applied to hold the motor-coated bead and measure its displacement (Fig. 2C). An optical tweezers instrument is, in its most straightforward implementation, nothing more than a near-infrared laser beam focused using a high-numerical-aperture objective in a diffraction-limited spot. In this focus, a transparent micrometer-sized micro-sphere can be held in three dimensions, due to transfer of momentum from pho-tons to microsphere, caused by refraction of the light. By steering the laser beam, the location of the optical trap can be altered. In this way, forces up to ~100 pN can be applied to the microsphere. In the case of motor-protein experiments, the interaction between motor protein attached to the bead and microtubule or actin filament on the surface is probed. At the same time, the trapping light passing through the sample can be detected using back-focal plane interferometry in or-der to determine the displacement of the bead with respect to the trap, with sub- nanometer accuracy (44). Since the optical trap generates a harmonic potential well for the bead (in first order) this displacement can be converted into a force signal after careful calibration of the instrument. Optical tweezers were first ap-plied to Kinesin-1: beads coated with at most a single Kinesin-1 motor held in an optical trap were brought into contact with a microtubule attached to the cover-slip (45). When the motor protein started walking along the filament, it pulled the bead out of the center of the trap, such that it experienced an increasing counteracting force. Displacement of the bead was in steps of 8 nm, independent

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of load and ATP concentration, which corresponds to the length of the repetitive unit of microtubules, indicating that Kinesin-1 steps between consecutive tubu-lin dimers. Kinesin-1 velocity was shown to decrease with applied force, until the motor stalled at ~6 pN (46). The mechanical work produced by a motor is the product of step size and maximum force it can overcome. For Kinesin-1 this is about 48 pN nm, corresponding to 12 (Boltzmann’s constant times absolute temperature), which is approximately half of the free energy that can be released by the hydrolysis of a single ATP molecule to ADP under cellular conditions (~25 ) (18). Since these first experiments on single Kinesin-1, optical twee-zers experiments have been used to unravel many aspects of the mechanochem-ical cycle of Kinesin-1 (47, 48, 49). Furthermore, it has applied to many other motor proteins, including Kinesin-14 (50), myosin V (51) and myosin VI (52).

2.2.8 Detection of single-motor activity using fluorescence microscopy

In the gliding assays and bead assays described above VE-DIC and fluorescence microscopy are used to follow movement of microtubules or beads. One require-ment for using fluorescence microscopy was that the microtubules or beads had to be labeled with multiple fluorophores, because of limited sensitivity of the detection system. At that time it was not possible to detect the signal from a single fluorophore. However, technical advancements in the fluorescence microscopy field pushed this detection limit further and further. In 1989, Moerner and co-workers achieved single-molecule sensitivity when they were able to detect the fluorescence of a single dye under extremely low temperatures (53). One year later, researchers were able to detect the fluorescent signal from a single fluorescent molecule in water (54), progress essential for fluorescence-based single- molecule assays of motor proteins (55). In the following I will discuss in more detail how fluorescence experiments can be performed on single motor proteins.

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SINGLE-MOLECULE FLUORESCENCE MICROSCOPY ON MOTOR PROTEINS

Single-molecule fluorescence microscopy allows tracking the location of individual, fluorescent proteins and is thus an ideal tool for studying single-motor pro-tein activity. A prerequisite is that the motor proteins are labeled with a fluoro-phore, since motor proteins are not fluorescent by themselves. An instrumental setup needs to be used that is sensitive enough to resolve the fluorescence emitted by a single fluorophore over background signals. In this part fluorescence and the properties of fluorescent probes will be briefly discussed. Next, an overview of will be given of the microscopy techniques used for single-motor-protein detec-tion. Finally, a brief description will be given of how fluorescence signals obtained from single motor proteins can be used to obtain insight in their mobility.

2.2.9 Fluorescence and fluorophores

Fluorescence is the phenomenon of the emission of a photon by a molecule im-mediately after the absorption of another photon by the same molecule (56). In fluorescent molecules, absorption of light results in an electronic transition from electronic ground state to excited singlet state. On a timescale of typically nano-seconds (in the condensed phase at room temperature), the excess energy stored in the excited state can be released by emission of a fluorescence photon, while the molecule returns to its ground state. The wavelength of the emitted photon is typically longer than that of the absorbed photon, the so-called Stokes’ shift, caused by fast vibrational relaxation and solvent reorientation in the excited state. Apart from wavelengths of absorption and emission, several other properties of fluorophores are crucial for their use in single-molecule experiments. The extinc-tion coefficient of a fluorophore should be high, i.e. at a given excitation intensity, the molecule should be excited frequently. It should have a high fluorescence quantum yield, such that a large fraction of the excited states generated result in the emission of fluorescence photons and that non-radiative decay processes of the excited state are relatively infrequent. In addition, the fluorescence intensity should be constant and not fluctuate due to blinking on and off. Finally, the probability of photobleaching should be low, such that the fluorescence signal can be followed for a substantial amount of time (1, 57).

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Three classes of fluorophores have been used to label motor proteins in in vitro single-molecule motility assays. (i) Synthetic dyes, such as Rhodamins, Alexa Fluor and Cy dyes are relatively small organic molecules that are obtainable in forms that can easily be attached to thiols or amines in the biomolecule of in-terest (58). In general, these molecules have optimized optical properties that are well known. (ii) Fluorescent proteins, such as Green Fluorescent Protein (GFP) and variants can be genetically encoded allowing for stoichiometric specific la-beling of proteins by generating genetic fusions between protein of interest and fluorescent protein (59). Fluorescent proteins have a size (~5 nm) that is often comparable to the protein they are fused to. Care has to be taken to assess that protein function is not altered by labeling. In addition, the optical properties of fluorescent proteins (including photostability) are substantially less than the best synthetic dyes. (iii) Quantum dots (QDs) are nanometer-sized spheres (6-60 nm in diameter) made of a semiconductor material (58). They are very bright fluoro-phores, have very wide absorption spectra and narrowly peaked emission bands. QDs have a high quantum yield and extinction coefficient, but tend to blink (60). QDs are usually coated with streptavidin for attachment to biotinylated versions of the biomolecule of interest.

2.2.10 Fluorescence microscopy

Fluorescence microscopy is required to image the fluorescently labeled mo-tor proteins with high enough sensitivity and specificity to resolve them over background signals and with good enough spatial and temporal resolution to track them in time. A typical fluorescence microscope consists of four parts: a light source, an objective lens, optical filters, and a detector (2, 61). For single- molecule studies continuous-wave lasers are generally used, because of their spec-tral purity (which allows good rejection of scattered excitation light), stability, and nearly perfect Gaussian beams (which allows diffraction-limited focusing). For effective excitation, the laser wavelength needs to be chosen such that it is close to the fluorophore’s absorption maximum. An objective is used to project the excitation light in the sample and to collect the fluorescent light emitted by the sample. Crucial is that an objective with high numerical aperture is used (for efficient collection of the fluorescence signal) and that the objective is well corrected for optical aberrations. A combination of optical filters is used to sep-arate excitation from fluorescence light. In most fluorescence microscopes three optical filters are used. (i) An excitation filter (often not necessary when lasers are

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used) selects the desired part of the spectrum from the excitation lights source for fluorescence excitation. (ii) A dichroic mirror separates excitation light from the fluorescence by specifically reflecting the excitation wavelengths but transmitting fluorescence wavelengths. (iii) An emission filter transmits most of the fluores-cence wavelengths and blocks unwanted back-ground signals, such as scattered excitation light. Good blockage of excitation light with more than 6 orders of magnitude is crucial for obtaining high-quality images. In case several, spectrally distinct fluorophores are used, specific multiband filters and dichroic mirrors can be applied. It is also possible to separate emission light in distinct spectral paths detected by different detectors. (iv) For single-molecule detection it is crucial to use a detector with high sensitivity, low noise and reasonable speed. Nowadays, usually electron-multiplying charge coupled device (EMCCD) cameras or ava-lanche photodiodes (APDs) are used. These two detectors are used in different types of fluorescence microscopes.

APDs are point detectors that are frequently used in confocal fluorescence microscopy. In this imaging modality, the excitation laser light is focused by the objective in a diffraction-limited spot (Fig. 3A). Light emitted from this spot (with a size on the order of half the wavelength of the light) is collected by the same objective and imaged on a pinhole. The pinhole effectively rejects out-of-focus light, ensuring optical sectioning. The fluorescence light passing through the pinhole is detected by the APD. In order to make an image with a confocal microscope, the sample or the confocal spot has to be scanned (using a piezo stage or scanning mirrors, respectively). In single-molecule applications, scanning is typically slow, since enough photons need to be detected at each sample point. In many single-molecule applications of confocal fluorescence microscopy, no images are recorded, but a stationary confocal excitation / de-tection spot is used to detect fluorescence bursts from fluorescently labeled biomolecules diffusing freely in and out (62, 63).

EMCCD cameras are array detectors, consisting of ten-thousands of micrometer-sized light detectors. These cameras are used in wide-field fluorescence microscopy. In wide-field epi-fluorescence microscopy the excitation light illumi-nates a larger area of the sample, typically tens of micrometers and is not focused in a diffraction-limited spot (Fig. 3B). The fluorescence emitted by this illuminated area is imaged on the camera, making scanning unnecessary. A key drawback of this approach is that not only fluorophores in the focal plane are excited, but

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also out-of-focus fluorophores are excited with comparable efficiency. This can re-sult in detection of a substantial amount of out-of-focus fluorescence, appearing as an unstructured haze in the images, often overwhelming the signal of interest. Wide-field epi-fluorescence microscopy is thus best suited for samples where the majority of fluorophores is located in a relatively thin layer (i.e. less than seve-ral micrometers). A wide-field imaging modality that overcomes this problem is total-internal-reflection fluorescence microscopy (TIRF). In TIRF the parallel excitation beam is totally internally reflected on the cover glass – water (sample) interface, producing a so-called evanescent wave in the sample (Fig. 3C) (64). The evanescent wave is a bound, exponentially decaying electromagnetic wave that penetrates only ~100 nanometers in the sample. With TIRF, fluorophores within the first 100 nanometers from the coverslip can be excited exclusively, largely eliminating background fluorescence emerging from deeper in the sample (2, 57).

Figure 3: Single-motor assays based on different fluorescence microscopy techniques. (A) Confocal fluorescence microscopy. A confocal spot (orange) is positioned on top of a micro-tubule (green). Motor proteins walk through the confocal spot where the fluorophore (red) is excited by the incoming excitation light (orange). (B) Epi-widefield fluorescence microscopy. Collimated excitation light (orange) is exciting the fluorophores (red) of several motor pro-teins (blue). (C) Total internal reflection fluorescence microscopy (TIRF). The incident ex-citation light (orange) falls on the coverglass such that it totally internally reflected. Because of the specific boundary conditions an evanescent wave propagates through the medium exciting only the fluorophores (red) of the few motors (blue) that are within ~100 nm from the surface.

A B

C

Confocal �uorescence microscopy Epi-wide�eld �uorescence microscopy

TIRF microscopy

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2.2.11 Sample preparation

Fluorescence-based single-molecule motility assays are typically performed inside a sample chamber built from a glass slide and a functionalized glass coverslip that allows controlled exchange of different solutions. The glass needs to be cleaned thoroughly and subsequently functionalized in order to minimize background fluorescence and to attach actin filaments or microtubules efficiently to the sur-face. Microtubules are generally grown from tubulin seeds (stabilized with slowly hydrolyzable GTP analogues) using fluorescently labeled tubulin, followed by stabilization with taxol. Microtubules can be attached to the cover-slip surface via charge interactions (between amino-modified or poly-L-lysine coated cover glass and the negatively charged microtubules), tubulin antibodies or streptavidin (when biotinylated tubulin is used) (Fig. 2D). Actin filaments are often made with fluorescently labeled actin monomers, subsequently stabilized with phal-loidin. They can be attached to the surface using actin antibodies or, in case biotinylated actin is used, steptavidin. Care has to be taken to find the right balance between stable attachment and minimal deformation of microtubules or actin filaments due to too tight binding. In order to prevent unwanted, aspecific adsorption of motor proteins to the remaining exposed glass surface, resulting in loss of motors from solution and increased background fluorescence, the surface needs to be passivated (blocked) using proteins (casein, serum albumin) or poly-mers (pluronic, PEG). Next, fluorescently labeled motor proteins are flushed into the sample chamber, at low concentration, together with ATP, and other required cofactors such as Mg2+. In order to decrease photobleaching anti-bleach cocktails can be added that remove reactive oxygen species (e.g. glucose-oxidase, catalase and glucose oxygen scavenger mix) and oxidizers/reducers (e.g. trolox, ß-mercap-toethanol). For each protein, concentrations, salt and buffer conditions, incuba-tion times and surface-blocking methods need to be optimized for best results (57, 65). Small differences in buffer conditions can result in substantially differ-ent motor motility behavior (66). Care needs to be taken to control temperature, which can have a substantial effect on motility parameters. After preparation of the sample, single-motor activity can be detected in the fluorescence microscope by localizing fluorescent spots due to individual motor proteins walking over their track.

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2.2.12 Data acquisition and analysis

Once the sample is prepared, data can be acquired by recording a sequence of fluorescence images with a sensitive camera such as a back-illuminated elec-tron-multiplying CCD camera. Frame rate and excitation intensity need to be balanced and optimized in order to obtain a high enough signal-to-noise ratio to discriminate the single fluorophores over back-ground signals, to reduce pho-tobleaching and to reduce motion blurring in individual image frames (depend-ing on the distance covered by the motor protein studied within the duration of a single frame). Image sequences obtained can be analyzed in several ways. A straightforward approach is to convert the image stack into kymographs, using open source image processing software such as ImageJ (http://imagej.nih.gov/). To this end, a line is drawn along a microtubule, the fluorescence intensities along this line for each frame are plotted in a 2-dimensional (time versus position along line) graph, the kymograph. Directed motion at a constant speed shows up in the kymograph as a straight, slanted line, with a slope directly related to its velocity. Kymographs are a valuable means to get a quick overview of the motility behavior but can also be used to obtain detailed quantitative insight (for more information regarding their quantitative analysis see Chapter 3.2). To obtain deeper and more quantitative insight about the behavior of single motor proteins, their position can be determined as a function of time with sub-diffraction limit accuracy, by Gaussian fitting of the fluorescence-intensity profiles using a track-ing algorithm (67, 68). Several key parameters can be readily obtained. First of all, the fluorescence intensity of moving spots can be analyzed to determine the number of fluorophores and, as a consequence, the amount of motors. Usually, single-step bleaching behavior is a used as a signature for individual fluorophores (2). Second, from position-time traces, velocities can be determined and, in case motility is characterized by random components, diffusion constants (66). By labeling the two motor domains with two spectrally distinct fluorophores the relative distance of the two motor domains during a processive run could, in principle, be followed. A third crucial motility parameter that can be readily obtained from straight-forward fluorescence imaging of moving individual motor proteins is the run length, the average distance a motor protein travels over its track before releasing. For accurate determination of run lengths, the contribu-tion of photobleaching to run termination has to be taken into account. This can be readily done by measuring the run termination times as a function of fluores-cence excitation intensity and extrapolating to zero intensity (69).

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2.2.13 Advanced fluorescence methods applied to single-motor motility assays

More advanced fluorescence measurements can be performed to obtain more specific information. Distances or length changes on the sub-ten-nanometer scale can be obtained using Förster resonance energy transfer (FRET) (4, 56). FRET is the transfer of excitations from a donor fluorophore to an acceptor fluorophore. The efficiency of FRET depends on spectral overlap of, relative orientation of and distance between the fluorophores. The distance dependence is very strong, FRET efficiency scaling with the inverse sixth power of distance, making FRET an excellent tool to scrutinize conformational dynamics in biomolecules such as motor proteins. Fluorescence polarization spectroscopy on individual fluoro-phores can be applied to learn about the orientational dynamics of different parts of the motor protein (4, 56, 70). The way the electric field of excitation and emission light couples with the fluorophore is described by the so-called transi-tion dipole moments of the fluorophore, vectors within the molecular frame. The relative orientation of the electric field of the excitation light (its polarization) and the excitation transition dipole moment determines how effective excitation can be. The orientation of the emission transition dipole moment determines the polarization direction of the fluorescence light. By making use of excitation light with alternating polarization and/or polarization-selective detection of fluores-cence, information can be obtained in the orientation of the fluorophore (and consequently the labeled protein) and changes therein (70). For more detailed information about FRET and its application to study the stepping mechanism of Kinesin-1 see Chapter 2.3.

A conceptually different approach of detecting single-motor protein activity was developed on the basis of confocal fluorescence microscopy (71). In this assay (Fig. 3A), similar samples were used, but no fluorescence images were recorded. Instead, a laser was focused on a microtubule and fluorescence photons emitted by labeled motor proteins walking through the confocal spot were recorded on an APD. Intensity time traces were Gaussians, from which readily motor-protein velocity and stoichiometry could be obtained. Key advantage of this approach is its increased time resolution, which can be down to about 100 microseconds in case the time traces are analyzed using autocorrelation methods. This makes it very attractive for studying motor dynamics with sub-step kinetics, for example in combination with FRET (72).

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APPLICATIONS OF SINGLE-MOLECULE FLUORESCENCE SPEC-TROSCOPY TO MOTOR PROTEINS

Above, I have discussed the fundamentals of the approaches to study single motor proteins with fluorescence microscopy. In the remainder of the contribution an overview is presented of applications of these techniques to motor proteins for the dynein, kinesin and myosin super families. Note that obtaining motility pa-rameters from single non-processive motor proteins using fluorescence detection is very difficult and often not very informative. This is why the following discus-sion almost entirely focuses on processive motor proteins.

SINGLE-MOLECULE FLUORESCENCE EXPERIMENTS ON KINE-SINS

The field of single-motor-protein fluorescence studies emerged from first measurements on Kinesin-1, the prototypical processive motor protein (73). The suc-cess of these initial experiments inspired more complex studies on Kinesin-1 and applications first to other kinesins and later to dyneins and myosins. Before giving an overview of these studies I will start by briefly describing the history of single-molecule fluorescence studies on Kinesin-1.

2.2.14 Kinesin-1 is processive

Kinesin-1 was first purified and identified as a motor protein using surface- gliding experiments (16). Later surface-gliding assays performed at very low mo-tor densities showed intriguing behavior. Under these conditions, microtubules still attached to the surface moved forward. Their motion was, however, quite different: they appeared to be attached to the surface at only one point, around which they swivelled (74). This observation was interpreted as indicating that Kinesin-1 is processive: it makes a multitude of steps along the microtubule track before detaching. This notion was supported by experiments involving micro-spheres coated with a low density of motors being tracked using video micros-copy (75) or optical tweezers (76). Under conditions that statistically at most a single Kinesin-1 could interact with the microtubule, movement over more than one micrometer was still observed. The most direct way of showing that a motor is processive is by tracking the motion of a single motor with single-molecule fluorescence microscopy. In a landmark experiment performed by Vale, Yanagida

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and their coworkers, Cy3-labeled Kinesin-1 motors were tracked moving in the plus-end direction of microtubules (73). For many of the moving motors the fluorescence intensity disappeared in two steps due to photobleaching, indicating that Kinesin-1 dimers (with in total two Cy3-molecules attached) can drive pro-cessive motility. From the single-motor traces (position versus time), important motility parameters, such as run length (i.e. the average distance a motor tra-vels before detaching) and velocity can be readily determined. In this first study, a velocity of 0.3 µm s-1 and a run length of about half a micron were determined. Later studies, most likely under better controlled conditions, have reported high-er values, 0.9 µm s-1 and 1.1 µm respectively, more in agreement with surface- gliding and in vivo measurements (e.g. (71)). Nkin, a Kinesin-1 from the fun-gus Neurospora crassa, was shown to move substantially faster, at ~2 µm s-1 (77). It is not well understood what the molecular basis is of this velocity difference. A hint might be that the sequences of the motor domains of Neurospora crassa and human Kinesin-1 are quite similar, while those of the neck regions differ substan-tially. Processivity of human Kinesin-1 could be altered considerably by mutating the motors, particular in this neck coiled-coil regions (78). Addition of extra positively charged amino-acid residues increased processivity substantially (69).

2.2.15 Kinesin-1 walks hand-over-hand

Optical tweezers experiments have revealed that the step size of Kinesin-1 is 8 nm (45), corresponding to the length of tubulin dimers, the building blocks of microtubules (79). This important observation indicated that Kinesin-1 steps between consecutive tubulin dimers. Subsequent experiments performed at low ATP concentrations revealed that binding of an ATP molecule is the single rate-limiting step under these conditions, strongly indicating that an 8 nm step is the consequence of the hydrolysis of one ATP molecule (80). Further evidence that dimeric Kinesin-1 hydrolyzes one ATP molecule per step at all ATP con-centrations was provided by measuring the displacement of microspheres coated with single motors, in combination with ATPase rate determinations (81, 82). Such experiments do, however, not explain how Kinesin-1’s two motor domains, with identical microtubule binding sites and ATPase active sites, work togeth-er to drive processive motion. In a landmark study, super-resolution localiza-tion of single fluorophores (83) was used to determine the stepping mechanism of Kinesin-1 (84). Kinesin-1 labeled on one of the motor domains was imaged with wide-field fluorescence microscopy, while moving at a very low velocity by

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applying a low ATP concentration. Dwell times between subsequent steps lasted relatively long, such that acquisition times of 330 ms could be used, resulting in high numbers of detected fluorescence photons. In this way, the localization of the fluorescent probe could be determined with an astonishingly high accuracy of only a few nanometers (84). Using this approach, step sizes of 16 nm of individual motor domains were observed, as expected in a “hand-over-hand model”, in which the motor domains alternate leading and trailing roles (Fig. 4A). Waiting times between such 16 nm steps were not distributed exponentially (i.e. due to a single rate-limiting process, the binding of ATP), but by a convolution of two identical exponents. This was to be expected, since in a hand-over-hand model this waiting time is the consequence of binding and hydrolyzing ATP and step-ping of both motor domains subsequently, resulting in 16 nm steps of each motor domain, but 8 nm steps of the dimeric motor as a whole. Subsequent hydrolysis of ATP by both motor domains was further supported by confocal single-motor assays involving fluorescent ATP-analogs (85). Using FRET between a donor

Figure 4: Stepping and diffusion of kinesin along the microtubule track. Arrow indicates movement of the subdomain with the same color. (A) Kinesin steps hand-over-hand in an ATP-dependent manner, with its two motor domains (red and blue) alternating between being in leading or trailing position. Each motor head tightly docks the tubulin dimer, which consists of one -tubulin (dark green) and one -tubulin (light green). (B) Kinesin diffusing along the microtubule lattice with the two motor domains (red and blue) weakly interacting with the surface of the microtubule (green).

A B

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on one of the motor domains and acceptor-labeled ATP, fluctuations in donor fluorescence intensity as a consequence of ATP binding and ADP release could be measured. A quantitative description of the magnitude and timescale of these fluctuations as a function of ATP concentration could only be obtained using alternating-site catalysis models, consistent with hand-over-hand stepping.

2.2.16 Conformational changes during Kinesin-1 motility studied using single-molecule fluorescence polarization

The processivity of kinesin has intrigued many researchers, raising questions on how the two identical motor domains cooperate to avoid falling off the track during stepping and what conformational changes in the motor domains are connected to stepping. To shed light on these aspects, single-molecule approach-es have been applied addressing the conformation of the motor domains during stepping. Two general approaches have been taken: FRET and fluorescence po-larization. We will first focus on the latter. Early single-molecule fluorescence polarization measurements focused on the orientation of motor domains in mon-omeric (86) and dimeric Kinesin-1 constructs (87). To this end, the motor do-main was labeled with a bisfunctional fluorescent probe, attached to two cysteins, such that the orientation of the dye with respect to the motor domain is known. The orientation of the motor domains could be obtained with great accuracy in fixed, non-walking states using nucleotide analogs. Improved measurements could resolve the orientation of the motor domains during stepping and indi-cated that the motor switches between a state with both motor domains tightly bound to the microtubule in a fixed orientation (one with ATP bound, the other without nucleotide), to a state where one motor domain is attached in this same fixed orientation (with no nucleotide bound) and the other mobile but tethered (most likely with ADP bound) (88). In another set of single-molecule fluores-cence polarization experiments on Kinesin-1, the focus was on the orientation of the neck linker, the 12-residue peptide that connects motor domains with stalk (89). Before, it had been shown, by a combination of electron microscopy and electron paramagnetic resonance, that the neck linker in monomeric Kinesin-1 constructs undergoes a conformational change from an ordered conformation, in which the neck linker points in the forward direction of Kinesin-1 movement, to a less ordered one (90). The fluorescence polarization measurements on individu-al Kinesin1 dimers demonstrated that this conformational change is also present in processively stepping dimeric Kinesin-1 constructs (89).

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2.2.17 Conformational changes during Kinesin-1 motility studied with Förster Resonance Energy Transfer (FRET)

Conformational changes of dimeric Kinesin-1 have also been tracked using FRET detected with wide-field fluorescence microscopy. In a first study, the focus was on the conformational change of the neck linker (91). To this end, heterodimeric Kinesin1 constructs were generated with a donor and an acceptor fluorophore at-tached to specifically engineered cysteines in the motor domain and neck linker of one of the polypeptide chains. This study confirmed (in dimeric Kinesin-1) that the neck linker undergoes a nucleotide-state dependent conformational change that plays a key role in Kinesin-1’s hand-over-hand motility. In a next study, the same authors focused on the location of the motor domains during processive motion (92). To this end they used heterodimeric Kinesin-1 with a donor and an acceptor attached to each motor domain at different locations. They found that Kinesin-1, while moving at maximum velocity, spends most of its time in a configuration with both motor domains microtubule-bound. At low ATP con-centrations, the motor is slowed down substantially and spends most of its time in a configuration with one motor domain microtubule-bound and “waiting” for ATP to bind, while the other is tethered, with ADP bound, and relatively free to explore the next binding site on the microtubule. A better time resolution was achieved in FRET studies employing a confocal fluorescence based motility assay (72). This study employed homodimeric Kinesin-1 constructs with a donor and an acceptor attached to each motor domain on the same location. This approach revealed that at maximum velocity, the two motor domains of Kinesin-1 are most of the time bound to the microtubule, 8 nm apart, but a fraction of the time in a specific conformation with one motor domain microtubule bound and the other one placed specifically, possibly docked on the bound motor domain. This additional configuration could only be detected in Kinesin-1 constructs with the fluorescent labels in specific parts of the motor domains. At lower ATP con-centrations, an additional configuration was observed, with one motor domain microtubule bound and the other tethered, consistent with the “ADP-waiting” state discussed above.

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2.2.18 Kinesin-1 motility is regulated by conformational changes induced by cargo binding

In the early days of Kinesin-1 research, it was observed that the motility of surface- bound motors (in gliding or bead motility assays) was often higher than expected on basis of the ATPase activity of the same preparations. Later, most researchers started using substantially shorter Kinesin-1 constructs, only containing motor and dimerization domains, with higher ATPase activity, more consistent with motility measurements. On the basis of such observations it was proposed that, when no cargo is bound, Kinesin-1’s tail can fold back on the motor domains inactivating them (93, 94, 95). When cargo binds to the tail, back-folding is prevented and the motor domains can bind to microtubules and drive motility. This proposal was tested using single-molecule fluorescence assays by comparing the motility parameters of Kinesin-1 constructs with different tails (96). Full-length Kinesin-1, with and without light chain, hardly bound to microtubules and moved very irregularly, with bursts of rapid unidirectional motion inter-spersed by frequent pauses. Short Kinesin-1 constructs with minimal tails, in contrast, moved with constant, high velocity. Smooth and fast moving Kinesin-1 constructs with long tails were generated by prohibiting back folding by stabiliz-ing the coiled-coil regions in the tail or specific mutations in the neck region to which the back-folded tail docks (see Fig. 4G on page 114).

SINGLE-MOLECULE FLUORESCENCE EXPERIMENTS ON OTHER KINESINS

Following the successful and fascinating studies of Kinesin-1 also other members of the kinesin superfamily have been scrutinized with single-molecule fluores-cence methods. Some of these other kinesins showed motility behavior similar to Kinesin-1, while other behaved surprisingly different. In the following, sin-gle-molecule fluorescence motility experiments on these kinesin motors on basis of their motility behavior will be discussed.

2.2.19 Processive kinesins

Several other classes of kinesins studied with single-molecule fluorescence mo-tility assays showed velocities and run length similar to Kinesin-1. Examples are the homodimeric Kinesin-3 UNC-104 (from Caenorhabditis elegans) (velocity

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~2 µm s-1 and run length of several micrometers, depending on construct) (97) and the homodimeric Kinesin-2 OSM-3 also from C. elegans (velocity 1-1.5 µm s-1

and run length ~1.2 µm, depending on the exact construct used) (98). For the latter motor an inhibition mechanism by tail back folding, similar to Kinesin-1 was demonstrated to exist. Kinesins-1, -2, and -3 all function in intracellular transport. It thus appears that the cell has devised a similar working mechanism for kinesins with this function. A kinesin with a totally different function in the cell is Kip3p, a fungal Kinesin-8. This motor is involved in the length regula-tion of microtubules by stimulating depolymerization from the microtubule plus ends. In single-molecule fluorescence assays, this motor showed slow (~50 nm s-1) plus-end directed motility, with unusual processivity, run lengths of more than 10 µm (99). In a later study it was shown that this long run length, in conjunc-tion with Kip3p’s removal of a limited number of tubulin dimers in a cooperative mechanism, allows Kip3p to tightly control microtubule length.

2.2.20 Kinesin diffusing along microtubules

Other microtubule-depolymerizing kinesins employ other motility strategies to reach microtubule ends. In single-molecule fluorescence assays it was shown that human Kinesin-13 MCAK binds to microtubules but does not move unidirec-tionally, with a constant velocity (100). It moves in a completely random way along the microtubule lattice (Fig. 4B). Diffusive motion was only observed in the presence of ATP and ADP. With no nucleotide present or non-hydrolyzable ATP analogs MCAK was stuck to microtubules. It was argued that one-dimen-sional diffusion over the microtubule lattice is an efficient search mechanism for microtubule ends, faster than direct binding from solution.

Interesting behavior was also observed in monomeric Kinesin-3 constructs of human KIF1A. Although the monomers moved erratic and changed direction constantly, overall they moved in a directional way (101). Careful analysis of the mean squared displacement of KIF1A showed that its motility can be understood as a biased random walk: KIF1A can step backward or forward like MCAK dis-cussed above, but the probability for stepping forward is higher (unlike MCAK). A follow-up study showed that the KIF1A motor cycles between a weakly bound and a strongly bound state, in a nucleotide-state dependent way (102). In the weakly bound state, KIF1A can move freely along the microtubule, while posi-tively charged KIF1A residues interact with the flexible, negatively charged C-ter-

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minal region of microtubules (the “E-hook”). The bias of motion in the plus-end direction of the microtubule is provided by an ATP-dependent conformational change, the power stroke. It remains, however, to be seen whether such a mecha-nism is relevant in vivo, since the related Kinesin-3 UNC104 is, in dimeric form, a processive plus-end directed motor (97). It might very well be that KIF1A in vivo is dimeric and that it thus far has been impossible to isolate intact dimers. Nevertheless, biased diffusion is a very interesting mechanism from the perspec-tive of theoretical concepts of motor action.

2.2.21 Kinesin-5, a tetrameric microtubule-crosslinker

Kinesins of the Kinesin-5 family are homotetrameric, with four motor domains, two on each side of a stalk (103, 104). For these motors microtubules are both track and cargo: they can crosslink two microtubules and slide them apart by walking on both of them (Fig. 2E) (105). This specific function of Kinesin-5 is essential for proper formation of the mitotic spindle. Single-molecule fluores-cence assays on GFP-tagged Eg5 (Kinesin-5 from Xenopus leavis) revealed a mix-ture of diffusive and slow plus-end directed motion (106). Interestingly, motility on a single microtubule was completely diffusive in the presence of ADP and at relatively high salt concentrations (66). Between two microtubules, however, Eg5 was shown to switch to directional motion in the presence of ATP (also at relatively high salt concentrations). These results reflect a regulation mechanism in which the motor switches from passive diffusion when only bound to a sin-gle microtubule to active motion when crosslinking two microtubules (66). It is unknown what part of the motor mediates the diffusive microtubule interaction and the motility switch. It has become clear, however, that Eg5 has additional, non-motor microtubule binding sites that are essential for crosslinking and rela-tive sliding of microtubules (107).

Cin8 is a Kinesin-5 from budding yeast, with motility properties remarkably different from Eg5. When bound to single microtubules it moves processively in the minus-end direction of microtubules, while it can slide anti-parallel micro-tubules apart, the consequence of plus-end directed motility when sandwiched between two microtubules (108). In a later study switching of directionality was shown to be inducible by salt in vitro: at high salt Cin8 moves processively in the minus-end direction along a single microtubule; at low salt Cin8 continuously switches direction from minus-end to plus-end directed processive motion (109).

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At high salt, Cin8 almost exclusively moved in the minus-end direction between parallel microtubules. In contrast, between anti-parallel microtubules, Cin8 al-ternated between plus and minus-end directionality. These findings indicate that also Cin8 can switch its motility parameters in a cargo-dependent way. Further studies are required to unravel the molecular nature of this remarkably complex behavior.

2.2.22 Kinesin-14, a non-processive minus-end directed kinesin

The canonical minus-end kinesin is Kinesin-14. This motor family is unique among kinesins in that its motor domain is on the C-terminal end of the polypeptide chain. Ncd, a Kinesin-14 from Drosophila melanogaster, showed steady minus-end directed motion in multi-motor surface gliding experiments (110, 111). In single-molecule fluorescence assays using GFP-tagged, truncat-ed Ncd constructs, only short binding events could be observed, runs of hun-dreds of nanometers and less, consistent with Ncd being a non-processive motor (112). In a subsequent study, full-length GFP-tagged Ncd constructs containing additional non-motor microtubule-binding domains were studied (113). These constructs remained bound to microtubules for a substantial amount of time and diffused along them. This motility mode was mediated by the non-motor microtubule-binding domains, as evidenced by similar binding and diffusive mo-tion by constructs lacking the motor domains. Many full-length Ncd motors working together were able to crosslink and slide apart anti-parallel microtubules, by bridging the two microtubules by interactions with the motor domains on one side and the additional microtubule-binding domain on the other (113). By crosslinking and sliding microtubules in this way, Kinesin-14 functions in constructing, maintaining and transforming the mitotic spindle, in concert with Kinesin-5 and other non-motor microtubule cross-linking proteins from the MAP65/Ase1/PRC1 family (114).

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SINGLE-MOLECULE FLUORESCENCE EXPERIMENTS ON MYO-SINS

The myosin superfamily of motor proteins interacts with actin filaments. Many myosins are non-processive motors involved in muscle contraction and oth-er force-generation tasks. Several myosin families consist of processive motors, involved in vesicle transport. Below, the focus will be on myosin V and VI, motor proteins with opposite directionality: myosin V is plus-end directed, myosin VI minus-end directed. Different types of single-molecule fluorescence spectroscopy have been applied to unravel the molecular basis of the different motility parameters of these myosins.

2.2.23 Myosin V walks hand-over-hand

Myosin V is the best studied processive plus-end directed motor of the myosin superfamily. In addition to the overall features of myosin, the myosin V lever arm (also called “light chain domain” or LCD) contains six IQ motifs that each bind a single calmodulin (CaM) light chain (Fig. 1C). The lever arm is responsible for amplifying the conformational changes of the motor domain upon ATP con-version in order to generate a step. The first single-molecule fluorescence studies of myosin V were employed to dissect its stepping mechanism. The movement of the lever arm was tracked by fluorescent labeling of CaM. It was shown that myosin V walks hand-over-hand with an average step size of ~37 nm (68, 115, 116, 117). This step size agreed with optical trapping experiments on the same motor (51). Using fluorescence polarization microscopy it was shown that the lever arm adopts two different orientations depending on being in leading or trailing position (115). Final support for the hand-over-hand mode of stepping was obtained from studies in which the lever arms of both motor domains were labeled with different fluorophores. Using this approach, the spacing between myosin V’s two motor domains bound to actin filaments was found to be ~36 nm, with step sizes of individual motor domains of ~74 nm (118, 119). Fluo-rescent ATP analogs were applied to demonstrate that myosin V’s processivity is the result of the tight coupling of the mechanochemical cycles of both motor domains (120).

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2.2.24 Myosin V step size is determined by the length of the lever arm

The ~36 nm step size is ideal for myosin V to take advantage of the pseudo re-peat of actin in order to stay on top of the actin filament rather than spiralling around it. To determine whether the step size was solely controlled by the pseudo repeat or if the length of the lever arm also plays a role, Sakamoto et al. per-formed a series of experiments changing the lever arm length of myosin V (121). The movement of the lever arm was tracked by Cy3-labeling of a single CaM. By inserting or deleting IQ motifs into or from the lever arm they found that myosin V could take a wide variety of steps indicating that step size is most likely determined by the neck linker length and not necessarily by the pseudo repeat of the actin filament.

2.2.25 Myosin V can explore different actin binding sites

Some of the step sizes observed in the experiments above would require consid-erable azimuthal twisting of the motor domain. Indeed, azimuthal variation of the myosin V lever arm was observed using “Defocused Orientation and Position Imaging” (DOPI), a fluorescence polarization technique in which the fluores-cence pattern of an out-of-focus fluorophore is analyzed to determine position and orientation, which showed that the motor makes ~27 degrees sideways swings back-and-forth while stepping (122). Moreover, it was found that the myosin motor can use its flexibility to explore different actin binding sites pri-or to making a full step (123, 124). This would allow myosin to successfully navigate through the complex actin network, an idea further supported by in vitro motility assays using actin-actin or actin-microtubule intersections (125). The authors showed that myosin could change actin filament upon encountering an actin-actin intersection or diffuse along the microtubule lattice upon encoun-tering an actin-microtubule intersection (Fig. 5). It might very well be that the dynamics of the myosin V motor as described above are important for cargo exchange between microtubule-based motors and myosins.

2.2.26 Myosin VI walks hand-over-hand

Minus-end directed myosin VI has a lever arm that contains only one conven-tional IQ motif but two light chains (Fig. 1C). Furthermore, it has a 53-amino acid insertion between the motor-converter domain and the lever arm that caus-

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es the arm to swing in opposite direction as most other myosins, inverting the direction (21, 126). On basis of the short lever arm, the step size of myosin VI was predicted to be smaller than that of myosin V. Optical trapping and single- molecule fluorescence experiments showed, however, that myosin VI takes ~36 nm steps, similar to myosin V (52, 127). Like myosin V, myosin VI moves processively in a hand-over-hand fashion (68, 128, 129, 130). The distribution of myosin VI step sizes is broader than those of myosin V, indicating that myosin VI stepping includes a substantial diffusive search component or small substeps. In a landmark study, Nishikawa et al. showed that the broad stepsize distribu-tion of myosin VI actually consists of two underlying distributions, one caused by ~72 nm hand-over-hand steps, and another one caused by ~44 nm “inch-worm”-like steps (131). In the inchworm mechanism, one of the motor domains is always in the lead, and steps while the other one limps behind.

How is the myosin VI motor able to make such big steps while having such a small lever arm? The hypothesis was that the 53-amino acid insertion had to be very flexible to allow for extension of the lever arm. However, this was soon ruled out, since a CaM was found to bind the insertion in a structural permanent

Figure 5: Myosin V moving along actin filaments and microtubules. Myosin V steps hand-over-hand along an actin filament (red). Upon encounter of an intersecting microtubule the myosin V motor can switch tracks and diffuse along the microtubule lattice (green).

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manner resulting in stiffening of the lever arm (132). In later studies a three-helix bundle was identified preceding the myosin tail (Fig. 1C) (133). This three-helix bundle unfolds upon dimerization of the motor, extending the lever arm from 4 nm to 13 nm as is required for ~36 nm steps.

2.2.27 Myosin VI is regulated by autoinhibition

It was thought that monomeric myosin VI exists in a compact, inactive confor-mation that helps the motor to diffuse through the dense network of actin fila-ments. This idea was supported by measurements on single GFP-tagged myosin VI showing that in the monomeric form the motor is in an autoinhibited state that can be activated by cargo binding (134). Furthermore, it was shown that a high density of monomers on the actin filament is necessary for dimerization and that cargo binding influences the dimerization process (135). Altogether, these results indicate that the monomeric form of myosin VI might diffuse through the actin meshwork where cargo binding controls dimerization and relieves the autoinhibition of the dimeric myosin VI motor (136).

2.2.28 Myosin VI can explore different actin binding sites

Myosin VI, now moving its cargo from the cell periphery towards the center of the cell has to navigate a complex network of actin intersections. As with myo-sin V, one would predict that myosin VI might benefit from having the flexibility to explore different actin binding sites. Earlier data demonstrated that the lever arm is highly mobile in the ATP state, and rather immobile in the ADP state, in-dicating that the motor has a flexible reach (68). This idea was further supported by measurements using single-molecule polarized total internal reflection fluo-rescence microscopy to measure angular changes of a bisfunctional rhodamine labeled lever arm (137). A broad distribution of lever arm angles was found when the lever arm was in leading position, whereas, this distribution was narrower in the trailing position. This would allow variable tilting of the leading motor do-main in order to explore different actin binding sites. Contrary to these results are DOPI measurements that show that the fluorescent pattern remains unchanged after a myosin step, indicating that the lever arm rotates 180 degrees during the power stroke (138). A critical aspect of the interpretation of these measurements is to relate the measured angle to the correct hemisphere, because of the twofold symmetry of the dipole (139, 140).

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SINGLE-MOLECULE FLUORESCENCE EXPERIMENTS ON DYNEIN

The third major class of motor proteins is the dynein family. In intracellular transport, dynein is responsible for transport along microtubules towards the mi-nus end. Dynein is a very large protein (heavy chains > 500 kDa), consisting of a C-terminal domain responsible for ATP hydrolysis, an N-terminal tail domain involved in dimerization and cargo binding, and a microtubule-binding domain. The C-terminal domain contains 6 AAA+ ATPase domains arranged in a ring. Dynein can bind multiple accessory proteins regulating motor function. Because of its size and complexity, in vitro assays on dynein have become feasible only since the last decade.

2.2.29 Dynein step size is variable

Cytoplasmic dynein purified from porcine brain was used in single-molecule motility assays to unravel its motile properties. It was found that dynein walks hand-over-hand with predominantly ~8 nm steps. The step-size distribution was, however, broader than that of other motors, including ~16 nm forward and ~8 nm backward steps (141). The distribution of dwell times between adjacent steps could be fitted to a single exponential function indicating that a single step is coupled to the hydrolysis of a single ATP molecule, consistent with the idea that ATP hydrolysis by only one of the AAA+ domains (AAA1) is sufficient for dynein motor activity (142). A truncated monomeric form of cytoplasmic dynein purified from S. cerevisiae was shown to be non-processive. When, however, these monomers were linked by adding dimerization domains they displayed robust processive movement, suggesting that dimerization is a prerequisite for processiv-ity but that the N-terminal tail domain or additional subunits (such as Lis1) are not required for processivity in vitro (143). Remarkably, also this dynein prepara-tion showed a wide variety of step sizes and, in addition, off-axis stepping. More recent work focused on high-resolution colocalization of the two ring domains and revealed that the two subunits walk on different microtubule protofilaments (144). After the majority of steps (~83%) a subunit remained in the leading or trailing position, indicating that the stepping mechanism is more complex than Kinesin-1’s hand-over-hand motion and involves inchworm-like episodes of mo-tion. Moreover, the trailing subunit was shown to have a higher probability to step if the spacing between the subunits is increased. It thus appears that dynein moves processively by individual subunits stepping stochastically and relatively

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independently when they are close together and in a coordinated way when they are stretched further apart (144).

2.2.30 Dynein can walk in both directions

The flexibility of dynein to make off-axis and backward steps of different siz-es might be essential for navigating through complex cytoskeleton networks. In measurements on GFP-labeled dynein-dynactin complexes purified from trans-genic mice, about 30% of the processive runs was purely plus-end directed, while another 30% was bidirectional (145), suggesting that dynactin regulates dynein directionality. The biological relevance of this flexibility in directionality was fur-ther emphasized in in vitro assays employing intersecting and crossing microtubules. In these assays, dynein-dynactin complexes displayed a range of behav-iors upon reaching an intersection: pausing, passing, switching track, dissociating and reversing, all with more or less the same chance of occurrence (146). Similar behavior was observed when dynein-dynactin encountered patches of the micro-tubule-associated protein (MAP) tau on microtubules (147). Intriguingly, in the same study it was shown that the affinity of kinesin for microtubules decreases with increasing tau concentration, whereas that of dynein is not affected. Afore-mentioned data illustrate that dynein motility is more complex and variable than that of kinesin. Future experiments will shed further light on dynein regulation and potential differences between proteins isolated from different sources.

IN VITRO FLUORESCENCE MICROSCOPY STUDIES OF MULTI-MOTOR ASSEMBLIES

In the cell, in many cases a cargo is not transported by a single motor protein. Generally, several motors, of the same or different species, are mechanically cou-pled (148, 149). Optical tweezers experiments have unequivocally shown that force applied to motors influences their motile properties (velocity, run length) substantially (150). This force dependence plays in fact a critical role in the com-munication between the two motor domains that drive the processive motion of for example Kinesin-1 (47). It is widely considered that motor proteins mechani-cally coupled to the same cargo will apply forces on each other and thus influence each other’s motility parameters. This motor cooperativity is a current active field of research, in vivo and in vitro (148, 149). In the following section we will focus on fluorescence-based in vitro experiments performed on assemblies of motors.

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2.2.31 Bead assays

A very straightforward way to study motor cooperativity is to attach multiple motors to a single microsphere and track the motion of the microsphere along the motor’s tracks using fluorescence or bright-field microscopy (Fig. 2B). In a recent application of such an approach to the cooperativity of Kinesin-1 (151), it was shown that run length increases with the number of coupled motor proteins, while the velocity is hardly affected, in agreement with previous work (74, 76, 152). A key limitation of such approaches is that the actual number of motor proteins on a particular microsphere is not known. Only the average number of motors can be inferred, in separate experiments, for example using dynamic light scattering (151). Furthermore, given the geometry of the microspheres and mo-tor size, the number of motors sterically able to interact with their track depends on the distribution of the motors over the area of the sphere.

2.2.32 Engineered assemblies of Kinesin-1 motors

To overcome these issues, Diehl and coworkers engineered well-controlled as-semblies of multiple Kinesin-1 motors fused to leucine zippers, first using purely protein-based scaffolds (elastin with a complementary leucine zipper) (153), later using leucine zippers connected to DNA strands that can be linked together with a complementary DNA strand (154). Using this latter approach, well-defined and dependable complexes of two Kinesin-1 motors were generated and studied using optical tweezers by attaching them to a transparent microsphere or using fluorescence microscopy by attaching them to a quantum dot (154, 155). Track-ing the motility of these two-motor complexes showed that the coupled kinesins produce larger average run lengths than single motors. Other motility parame-ters, such as velocity and stall force were shown to be very similar for coupled and individual motors. These results indicate that, in these motor assemblies, the individual motors can adopt many different configurations, but that in the large majority of them only one of the motors is effectively engaged with the track, limiting cooperative effects of highly processive motors (155).

2.2.33 Engineered assemblies of multiple, different motors

More recently, two novel approaches have been presented to construct larger mo-tor assemblies, allowing inclusion of different motor protein species (Fig. 2F)

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(156, 157). In one of these approaches (156) DNA origami (158) was used as a stable scaffold to which up to seven motors can be attached in a controlled way, by hybridizing single-stranded DNA attached to the motor proteins to comple-mentary sequences in the DNA origami. Using this approach it was confirmed for processive motors like Kinesin-1 and cytoplasmic dynein that velocity is hard-ly affected by the number of coupled motors, while run length increases substan-tially. DNA origami was also used to couple cytoplasmic dynein and Kinesin-1 in one mechanically connected assembly in order to test what motor wins the “tug of war”. It was shown such assemblies could move in the plus-end direc-tion, the minus-end direction or did not move at all. The ratios of these three possibilities depended strongly on the number of Kinesin-1 compared to dynein motors. Overall, dynein appeared to be stronger in the tug of war, resulting in more minus-end directed motion than to be expected on basis of the ratio of these particular motor proteins. This result indicates that not only stall force, but also other parameters such as run length, microtubule affinity, detachment force and force-dependent velocity play important roles. Non-motile dynein/kinesin assemblies could be brought in motion by photo-cleaving one type of motor, resulting in motion opposite to the directionality of the dissociated motor, indi-cating that these assemblies were stalled in a tug of war.

In a distinct approach to make large motor assemblies of different motor proteins, a simpler, fluorescently labeled DNA scaffold, consisting of hybridized oligonucleotides was used (157). Some of the oligonucleotides were covalently con-nected to the motors using Snap-tag or Halo-tag attachments (159, 160). Also using this approach the effect of number of coupled Kinesin-1 motors on motility parameters was tested, confirming that velocity is hardly affected, but run length increases substantially with motor number. Similar experiments were performed using the non-processive Kinesin-14 Ncd. Remarkably, coupling two or more Ncd’s turned the assembly processive, with run length increasing with number of coupled Ncd’s. Tug-of-war events between Kinesin-1 and up to four Ncd’s were generally “won” by Kinesin-1. The velocity of such assemblies was, however, substantially lower, indicating that Ncd generates substantial friction due to its interaction with microtubules.

These data illustrate that advances in bioengineering have created possibilities to create motor assemblies in a well-controlled way, providing opportunities to test how multiple motors work together in driving intracellular transport. These

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approaches allow modifying number and species of motor protein, their order and the mechanic properties of the linkers. It is to be expected that these ap-proaches will be applied to different types of motors in order to learn more about motor cooperativity in vivo.

SINGLE-MOTOR FLUORESCENCE-MICROSCOPY STUDIES IN VIVO

In vitro single-molecule fluorescence microscopy studies on molecular motors allow for measuring individual properties of motor proteins with high spatio- temporal precision and a high degree of control over experimental conditions. Most of these assays are minimal assays, containing only purified motors, micro-tubules, and a motility buffer with ATP. Inside the cell, conditions are, however, far more complex: microtubules and actin can undergo post-translational modifi-cations and accessory proteins can regulate motor activity and cargo attachment. Furthermore, motors often cooperate to transport one cargo through a dense and complex cytoskeletal environment. An example is intraflagellar transport (IFT) in the chemosensory cilia of Caenorhabditis elegans, where two motors of the Kinesin-2 family cooperate in building and maintaining the cilium (161). Both motors (OSM-3 and kinesin-II) drive transport in the beginning of the cilium, walking along microtubule doublets. Remarkably, only OSM-3 drives transport along the rest of the cilium walking over microtubule singlets and is essential for its formation and maintenance (in Chapter 4.1 the IFT-system is quantitatively dissected revealing the precise roles of the two kinesin-2 motors). Dynein drives transport in the opposite direction, from cilium tip to base. Although only three motors are involved, IFT is complex, involving well-balanced cooperativity and regulation of the motors, which makes it hard to mimic in vitro. In order to un-derstand the full complexity of transport processes in the cell it is pivotal to also study the behavior of individual motor proteins in their natural environment.

The ultimate goal is to be able to follow individual motor proteins while they are driving intracellular transport in a living cell or organism (see Chapter 4.1). So far, studies have been limited in visualization of individual motors not connected to cargo moving along the cytoskeleton of living cells, using fluorescence microscopy. The central problem in such studies is to overcome the autofluorescence background caused by naturally occurring fluorescing molecules such as flavins, collagen, NADH and elastin. The emission spectrum of autofluorescence is

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generally broad and overlaps with that of commonly used fluorophores decreas-ing the signal-to-background ratio (162). One approach to overcome autoflu-orescence was to introduce purified and quantum-dot labeled Kinesin-1 using pinocytosis into HeLa cells (163). Kinesin-1 was truncated and thus unable to connect to cargo (other than the quantum dot). The high brightness of quan-tum dots allowed tracking of individual motors. In a different approach, the signal-to-background ratio was improved by expressing fusion proteins of Kine-sin-1 with three Citrine monomers (a variant of yellow fluorescent protein) in COS cells and visualizing individual motors using total internal reflection fluo-rescence microscopy (164). In both studies, similar motility parameters (velocity, run length) were observed in vivo compared to those obtained using in vitro mo-tility assays, indicating that in vitro assays are good models for in vivo Kinesin-1 functionality. Later, the triple Citrine labeling approach was used to show that Kinesin-1 only moves along ~10% of the microtubules, having a strong prefer-ence for stable, post-translationally modified microtubules (165). In contrast, Ki-nesin-2 and Kinesin-3 did not appear to have a preference for a subset of microtubules. Similar approaches were used to track quantum-dot labeled myosin V moving along actin filaments inside the cell, which also showed that motility parameters are comparable to those measured using in vitro assays (166, 167).

As indicated, single-motor in vivo experiments have so far been applied only to motor proteins lacking cargo-binding functionality. In this way, the motor’s activity could be investigated in the cellular context. These studies have laid the groundwork for in vivo studies of individual motor proteins involved in intracel-lular transport of cargo. Such studies will undoubtedly shed more light on motor regulation and cooperativity within the years to come.

OUTLOOK

From the previous paragraphs it is clear that application of single-molecule spec-troscopy has tremendously enhanced our understanding of motor proteins. I fore-see that this will continue in the future, thanks to advances in instrumentation and assays (for example in bioengineering of motor assemblies, or purification of new motor proteins). Progress can be expected in these four areas in particular:

(A) Study of newly isolated motor proteins. Not all motor proteins of the kinesin, myosin and dynein superfamilies have been isolated and studied using in vitro

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assays. For example, it is unclear how the different kinds of dynein (cytoplasmic, IFT and flagellar, from different organisms) differ in motility properties.

(B) Study of multi-motor assemblies. Only recently it has become possible to study, in a quantitative way, the motility of well-defined motor assemblies (153, 156, 157). In the future, such assays will be used to study the cooperative action of motor proteins in conditions that are more like what is found in cells. For example, it would be interesting to understand how the two Kinesin-2’s cooper-ating in IFT influence each other’s motility in well-controlled in vitro conditions.

(C) Study of individual motor proteins engaged in intracellular transport in cells or whole organisms. So far, first experiments have been performed on motor proteins in living cells using truncated motor proteins not involved in cargo transport. A logical next step is to study motors participating in cellular process-es. Such studies will undoubtedly shed new light on our understanding of the role, regulation and mechanism of motor proteins in intracellular transport (see Chapter 4.1).

(D) In vitro reconstitution of complete intracellular transport systems. In the spirit of Richard Feynman, who stated “What I cannot create, I do not under-stand”, we have only just begun to understand complex biological processes such as intracellular transport. In vivo measurements (see above) will be essential to advance our knowledge. One can argue, though, that full comprehension of in-tracellular transport is only reached when it can be reconstituted in vitro from its components, displaying the full complexity of behaviors observed in the cell. This will not only involve motor proteins, but also their cargo, tracks and regulatory proteins.

Studies of motor proteins and the development of single-molecule assays will thus remain closely intertwined (6), yielding important new insights in the func-tionality and mechanism of this fascinating and crucial class of proteins.

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2.2.34 Acknowledgements

I would like to thank my colleagues and collaborators for insightful discussions. Work in the laboratory of Erwin J. G. Peterman at the VU University Amsterdam is supported by a Vici fellowship from the Netherlands Organization for Scien-tific Research, Physics (NWO-N), by an ALW Open Programme grant from NWO, Earth and Life Sciences (NWO-ALW), an NWO Groot investment grant and is part of the “Barriers in the Brain” research program of the Foundation for Fundamental Research on Matter (FOM), which is financially supported by NWO.

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