Laser Trap Measurements of Flagellar Membrane Motility

CHAPTER FIVE

Laser Trap Measurements of Flagellar Membrane Motility William H. Guilford*,1, Robert A. Bloodgood†

*Department of Biomedical Engineering, University of Virginia, Charlottesville, Virginia, USA † Department of Cell Biology, University of Virginia, Charlottesville, Virginia, USA 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Laser Trap Overview 3. Measuring Flagellar Membrane Transport 3.1 Cells 3.2 Laser trap measurements of flagellar membrane transport 3.3 Pitfalls and limitations 3.4 Future directions Acknowledgments References

86 88 91 91 91 103 105 105 105

Abstract In addition to swimming motility, which is driven by propagation of bends along the flagellum, the unicellular green alga Chlamydomonas exhibits an unusual and alternative form of whole cell locomotion, called gliding motility. In gliding motility, a large flagellar membrane glycoprotein mediates flagellar membrane adhesion to solid substrates. This in turn activates a transmembrane signaling system that initiates the movement of a cross-linked cluster of glycoproteins within the plane of the flagellar membrane by activating and/or recruiting isoforms of the motor proteins kinesin and dynein. Flagellar membrane motility can be visualized through the bidirectional movement of microspheres adherent to the flagellar surface. This microsphere motility offers a unique, noninvasive experimental system for measuring the in vivo dynamics and regulation of microtubule-dependent molecular motors by using a laser trap transducer to capture and manipulate microspheres as they move along the flagellar surface. Detailed procedures for conducting such analyses are provided.

Methods in Enzymology, Volume 525 ISSN 0076-6879 http://dx.doi.org/10.1016/B978-0-12-397944-5.00005-5

#

2013 Elsevier Inc. All rights reserved.

85

86

William H. Guilford and Robert A. Bloodgood

1. INTRODUCTION Transport of cargo by molecular motors in cells is necessarily bidirectional; in general, material that is transported actively to the cell periphery (anterograde) will be subject to return transport (retrograde). In order to achieve this bidirectional transport, both plus- and minus-end directed motors bind cellular cargos. They engage together or operate separately to move the cargo, and at any point in time the motors can either be the engines driving cargo movement, or part of the cargo that is being moved. One of the most compelling questions in the field is how oppositely directed motors are coordinated in order to generate directional movement in a controlled manner. The unicellular green alga Chlamydomonas is a useful organism for studying the biophysics, cellular, and molecular biology underlying bidirectional, microtubule-based transport. Chlamydomonas can exhibit two very different forms of flagella-dependent whole cell locomotion: (1) swimming motility, dependent upon the propagation of bends along the axoneme due to sliding of outer doublet microtubules driven by axonemal dynein motors (Witman, 2009) and (2) whole cell gliding motility, in which the cell moves across a surface to which it is adherent, and which is dependent on force transduction occurring at the flagellar surface in the absence of any obvious deformation of the flagellum (Bloodgood, 1981; Lewin, 1952). Because most strains of Chlamydomonas are found within environments such as soil, sand, moss, glaciers, and the like, it is likely that gliding motility is the more physiologically relevant of the two flagella-dependent motilities in Chlamydomonas. Flagellar surface motility can also be dramatically visualized through the bidirectional movement of microspheres adherent to the flagellar surface (Bloodgood, 1977). A large flagellar membrane glycoprotein, termed FMG-1B, has been shown to be the protein that mediates flagella membrane adhesion to a solid substrate during gliding motility and adhesion to microspheres during microsphere motility (Bloodgood & Workman, 1984). Cross-linking of FMG-1B molecules initiates a transmembrane signaling system that initiates the movement of a cross-linked cluster of FMG-1B molecules within the plane of the flagellar membrane (Bloodgood, Woodward, & Salomonsky, 1986), presumably by activating and/or recruiting motor proteins (Bloodgood, 2009). FMG-1B is recruited to and concentrated at sites of contact with moving microspheres (Bloodgood & Salomonsky, 1998); microsphere movement and gliding motility are inhibited under experimental conditions in which

Laser Trap Measurements of Membrane Motility

87

movement of FMG-1B within the flagellar membrane is restricted (Bloodgood & Salomonsky, 1989). Intraflagellar Transport (IFT) is a form of motility that operates in the compartment located between the flagellar membrane and the outer doublet microtubules (Kozminski, Johnson, Forscher, & Rosenbaum, 1993). IFT is necessary for the assembly and maintenance of all cilia and flagella; anterograde IFT is associated with the kinesin-2 motor while retrograde IFT is associated with the cytoplasmic dynein-2 (also known as cytoplasmic dynein-1b) motor. It is not known whether IFT and gliding motility are reflective of the same underlying transport process, though it has been demonstrated that they both use kinesin-2 as the anterograde motor (Kozminski, Beech, & Rosenbaum, 1995; Laib, Marin, Bloodgood, & Guilford, 2009). Although less certain, they may both utilize cytoplasmic dynein-2 as the retrograde motor (Bloodgood, 2009; Pazour, Dickert, & Witman, 1999). There are differences between IFT and flagellar surface motility relating to velocities of movement, saltatory behavior, and calcium dependence (Bloodgood, Leffler, & Bojczuk, 1979; Bloodgood & Salomonsky, 1989). A small group of laboratories is currently examining the relationship among IFT, microsphere movement, and gliding motility. Our laboratories have recently taken advantage of the saltatory, bidirectional transport of polystyrene microspheres (and by inference the bidirectional movement of the FMG-1 membrane glycoproteins) to study the behavior of microtubule-dependent motors in the living cell (Laib et al., 2009). Microspheres were captured in a laser trap—a focused beam of laser light used to capture and hold particles in three dimensions. Trapped particles behave as Hookean springs over small displacements from trap center. The laser trap can therefore be used as both a micromanipulator and an ultrasensitive force and displacement transducer. With careful selection of wavelength, a laser trap can often be used on living cells with little or no ill effect. Trapped microspheres were brought into contact with the flagellar surface of paralyzed (nonswimming) Chlamydomonas cells (Fig. 5.1). We measured the average velocities and peak forces of transport in each direction and found that transport in either direction was driven by approximately 10 simultaneously engaged motors—a larger number than generally supposed. Quiescent periods between transport events (excursions) strongly support models of transport regulation where kinesin-2 and cytoplasmic dynein-2 are reciprocally regulated to engage in large numbers and for exclusive transport in a single direction. This is a unique, noninvasive method for understanding the functioning of molecular motors in cells

88

William H. Guilford and Robert A. Bloodgood

Trap

FMG ?

Cell body

ms

Kinesin Dynein MT

f

PLL-cover glass

Figure 5.1 Experimental approach. A paralyzed Chlamydomonas is adhered to a polyL-lysine (PLL)-coated cover glass. A microsphere (ms) is captured in a laser trap and brought into contact with opposite face of the flagellum (f), well away from the cell body. FMG-1B binds to the microsphere and clusters, activating flagellar membrane transport of the bead via unknown intermediate proteins (?); these proteins bridge FMG-1B to kinesin and dynein, which in turn move along the outer doublet microtubules (MTs).

and for understanding the importance and regulation of motor-driven processes in the assembly and maintenance of flagella (Mallik & Gross, 2009). An added advantage of the Chlamydomonas system is the availability of molecular genetic approaches and the wealth of mutant cell lines available through the Chlamydomonas Resource Center.

2. LASER TRAP OVERVIEW A basic laser trap consists of four components: a laser, a beam expander, a high NA microscope objective, and a means of aligning the expanded beam to the objective. A typical arrangement of the optics is shown in Fig. 5.2. A laser beam is expanded to fill the back aperture of a high numerical aperture microscope objective and carefully aligned to the optical axis, most often by a pair of moveable mirrors (not shown). The objective focuses the laser beam at the specimen plane where the trap is formed and the trapped object is imaged. Laser traps may be based on either upright or inverted microscopes. When doing high-resolution work, such as that described here, mechanical stability is of overriding importance, and inverted microscopes are preferred. There have been numerous articles and books published on the construction, use, and underlying physics of laser traps (Fa¨llman & Axner, 1997;

89

Laser Trap Measurements of Membrane Motility

cam D1

Laser L1

L2 O Stage C D2

QPD L3

ilum

Figure 5.2 Critical components of a laser trap for measuring flagellar membrane transport. The output of a near-infrared laser is expanded using two lenses (L1, L2) and reflected into the optical path by a dichroic mirror (D1) to fill the back aperture of a high numerical aperture microscope objective (O). This brings the beam to a focus where a microsphere (black dot) can be trapped. A condenser (C) serves its usual role, focusing light from an illuminator (ilum), but also collects the partially collimated laser light after it passes through the microsphere. This light is diverted by a dichroic mirror (D2), and the back aperture of the condenser is focused onto a quadrant photodiode (QPD) by lens L3. The QPD signal provides data on the displacement of the microsphere relative to trap center. The microscopic field is observed using a solid-state camera (cam).

Guilford, 2001; Sheetz, 1997; Svoboda & Block, 1994). Most of the design considerations are generalizable, so a detailed discussion is beyond the scope of this chapter. We instead focus on features that are particularly relevant to measurements of flagellar membrane transport. For trapping living cells or other biological materials, it is best to choose a wavelength at which the cell has very little extinction in order to minimize heating and photodamage. Near-infrared in the range of 800–1100 nm is generally best (Svoboda & Block, 1994) and appears to be innocuous to Chlamydomonas flagella. Refer to Section 3.3 for a detailed discussion of the effects of laser light on living cells. The laser should deliver at least 200 mW at the specimen plane to generate sufficient trapping forces for studies of flagellar membrane transport. However, the optics will significantly attenuate the laser beam, so lasers with continuously variable power >1 W are preferable. Of equal importance to trapping is detecting the position of the trapped object with sufficiently high-spatial resolution to observe the individual steps with which single molecular motors move, and preferably with sufficiently

90

William H. Guilford and Robert A. Bloodgood

high-temporal resolution to observe motor function and to enable straightforward calibration of the trap (see Section 3.2.6). A relatively simple approach that achieves both is back-focal-plane interferometry (Allersma, Gittes, deCastro, Stewart, & Schmidt, 1998). The trapping beam (or a second laser beam) passes through the microsphere and emerges partially collimated. The exit angle of this collimated beam depends on the lateral position of the microsphere relative to the focal point of the laser. To measure this exit angle, the back aperture of the condenser, which collects this light, is imaged through a lens onto a position-sensitive detector—a quadrant photodiode in our realization (Fig. 5.2, QPD). The lateral distribution of light is a direct measure of the lateral displacement of the microsphere. Sensitivities of <0.3 nm have been achieved (Abbondanzieri, Greenleaf, Shaevitz, Landick, & Block, 2005), as well as bandwidths of >150 kHz (Guilford, Tournas, Dascalu, & Watson, 2004); these are orders of magnitude better than image feature tracking algorithms. Laser traps can be (and have been) combined with virtually any type of light microscopic imaging. Simple bright field illumination is adequate for the experiments described here. However, combining the use of strains of Chlamydomonas with GFP tags on motor proteins with a laser trap equipped with fluorescence detection could provide useful information about motor loss or recruitment during directional changes. Such strains are currently available. For most of the experiments described here, a stationary laser trap is all that is needed. However, it is often desirable to be able to move the laser trap within the field of view. This is accomplished by changing the entry angle of the beam into the microscope objective. If only slow movements of the trap are needed, this is easily done by moving the first element of the beam expander (L1, Fig. 5.2) or by tilting one of a pair of mirrors. If high-speed displacement (<10 ms) is needed, one typically inserts an acousto-optic deflector (AOD) before the aforementioned mirrors. Depending on the aperture size, AODs can move a laser trap within a few microseconds. The microscope can be equipped with any suitable stage, provided that it is sufficiently stable. We use a three-dimensional, feedback-stabilized piezoelectric stage (nPoint) that affords us the ability to raise and lower the stage relative to the focal point. While a traditional two-dimensional rack-driven stage is arguably sufficient for these experiments, precision adjustment of the focal plane is vital to placing a microsphere atop a 300-nm-wide flagellum. Three-dimensional stages or piezoelectric focusing systems are vital in this regard.

Laser Trap Measurements of Membrane Motility

91

3. MEASURING FLAGELLAR MEMBRANE TRANSPORT 3.1. Cells A wide variety of strains of Chlamydomonas are available from the Chlamydomonas Resource Center (http://chlamycollection.org). Detailed information on maintenance and growth of Chlamydomonas can be found in Harris (2009). For the approaches described in this chapter, we have typically grown Chlamydomonas in liquid cultures utilizing Medium 1 of Sager and Granick (1953); cultures were bubbled with filtered air and their growth synchronized using a light/dark cycle of 14 h light/10 h dark. Cells were used during the middle of the light cycle when they were not dividing and possessed full-length flagella. It is vital to use paralyzed Chlamydomonas (nonbeating flagella) in these experiments, for the obvious reason that one cannot make nanometer and piconewton resolution measurements in a beating flagellum. Fortunately there are mutant strains of Chlamydomonas with full-length flagella that do not beat. Most extensively used by us is pf18, which is characterized by a complete absence of the axonemal central pair (Adams, Huang, Piperno, & Luck, 1981). We have also used pf1, which lacks the head of the radial spoke (Luck, Piperno, Ramanis, & Huang, 1977). Motile strains could presumably be used in association with small molecule inhibitors of beating. Indeed, beating motility can be inhibited in wild-type cells, without any effect on IFT (and presumably other motility systems), using 20 mM LiCl (Dentler, 2005; Dentler, Vanderwaal, & Porter, 2009).

3.2. Laser trap measurements of flagellar membrane transport 3.2.1 Flow cell assembly Flow cells are assembled from two microscope cover glasses, one slightly smaller in dimension than the other, separated by a pair of spacers and held together with a suitable adhesive. While we routinely use mylar shim stock as the spacers and a UV-curing adhesive, vacuum grease can also be used in place of adhesive, or double-sided tape used as a complete replacement for both the spacers and the adhesive. The selection is somewhat a matter of personal preference, but mylar shims and UV-curing adhesive provide for a reproducible and consistent flow cell depth without leaks.

92

William H. Guilford and Robert A. Bloodgood

The use of two different cover glass sizes is important. A small cover glass mounted atop a larger one provides a convenient “lip” against which one may load cell suspensions into the flow chamber. 3.2.1.1 Materials

1. 2. 3. 4. 5.

Norland optical adhesive. Mylar shim stock (0.004 in. thickness, typical. Practi-Shim). 22
22 mm cover glass. 18
18 mm cover glass. UV curing oven (ELC-500 Light Exposure system, Electro-Lite Corp, Bethel, CT).

3.2.1.2 Assembly

1. Put a dab of optical adhesive in a disposable Petri dish. 2. Mylar shim stock is cut into short strips 4 mm wide by 21 mm long. The length of the strips must be greater than the length of the smaller cover glass but shorter than the length of the larger cover glass. 3. Using forceps, grip the end of a mylar strip and drag it repeatedly through the adhesive to completely cover both sides of the strip. 4. Remove excess adhesive by dragging the mylar strip across a dry or lightly coated region of the Petri dish. The coating should be uniform but thin. 5. Put the mylar strip close to one edge of the 22
22 mm cover glass (see Fig. 5.3A). 6. Repeat steps #3–5 for a second mylar strip, placing it along the opposite edge of the cover glass from the first. A

18 x 18 cover glass

B

Shim 22 x 22 cover glass

Figure 5.3 Assembly of flow cells and humidors. (A) An 18
18 and a 22
22 cover glasses are separated by two mylar shims to create a  10
18
0.2 mm (36 mL) flow cell. The cover glasses and the shims are held together by UV-curing glue. (B) A humidor is made from a square plastic Petri dish with four wooden applicator sticks arranged on the bottom with tape—two stacked vertically and two more spaced horizontally (see text for measurements). Flow cells may be rested horizontally during poly-L-lysine coating or tilted against the stacked sticks to angle the flow cell to promote flagellum adhesion.

Laser Trap Measurements of Membrane Motility

93

7. Put the 18
18 mm cover glass atop the mylar strips. Push lightly on the edges of the cover glass to develop a continuous seal between the cover glasses and the mylar strips. 8. Put the flow cell in the ultraviolet curing oven for 2 min. 9. These flow cells keep indefinitely in a clean environment. 3.2.2 Preparing microspheres for laser trap experiments Conveniently for laser trap experiments, FMG-1B is nonspecifically adhesive to a number of substrates. It is not necessary to adsorb or cross-link ligands or antibodies to microspheres in order for adhesion to FMG-1B to occur. We routinely use polystyrene microspheres, prepared as follows. 3.2.2.1 Materials

1. Microspheres—typically 0.5–1.3 mm diameter polystyrene (Bangs Laboratories, Fishers, IN, or Polysciences, Inc., Warrington, PA), with 1 mm neutral polystyrene microspheres giving dependable results. 2. Growth Medium 1 of Sager and Granick (1953), as referenced above (Section 3.1). This is an inexpensive and very stable minimal medium lacking any carbon source, allowing for good synchronization of cell growth. 3.2.2.2 Preparation of microspheres

1. Add 1 mL of ddH2O to a 1.5-mL microcentrifuge tube. 2. Add 3.3 uL of polystyrene microspheres (from Bangs Laboratories) to the same centrifuge tube. 3. Spin for 5 min at maximum speed (13,000
g) in a desktop centrifuge (Eppendorf MiniSpin). Smaller microspheres (around 0.5 mm in diameter) need to be centrifuged longer—about 10 min. 4. Carefully remove the excess water from the tube without removing the microspheres. 5. Add 1 mL of ddH2O and vortex to suspend the microspheres. 6. Repeat steps 3–5 two more times. 7. At the end of the last centrifugation, remove the excess water and refill the tube with 1 mL of culture medium rather than water. This solution of microspheres can be kept for several days with refrigeration. Microsphere clumping can be minimized by briefly sonicating the suspension in an ultrasonic cleaning bath, or using an ultrasonic dismembranator— an immersible ultrasonic probe. The latter is particularly effective at breaking apart clumps of microspheres.

94

William H. Guilford and Robert A. Bloodgood

Others have reported using microspheres coated with anti-FMG-1B monoclonal antibody rather than bare polystyrene for similar studies (Shih, Kocabas, & Yildiz, 2011). 3.2.3 Preparing poly-L-lysine stock solution Cells must be adhered to the walls of the flow chamber in a manner that prevents them from gliding. This is accomplished by the use of poly-L-lysine. 1. Weigh a small amount of poly-L-lysine hydrobromide (30,000–70,000 molecular weight, Sigma–Aldrich). 2. Poly-L-lysine is sticky, so measuring a specific mass is very difficult. Instead use the mass from step 1 and add ddH2O to create a 10 mg/mL stock solution. 3. Vortex the mixture for at least 2 min. 4. Aliquot the solution into 0.5-mL centrifuge tubes. 5. This solution is stable at 20 for months. 3.2.4 Loading the flow cell 3.2.4.1 Constructing a humidor

Loading a flow cell entails two prolonged incubation steps during which one can see significant evaporation from the ends of the flow cell. A humidor is useful for slowing evaporation. 1. Four wood or plastic applicator stick supports are taped to the bottom of a Petri dish closer to one edge than the other, parallel to one another (Fig. 5.3B). 2. A paper wipe (such as a Kimwipe) is wound into a tight rope and placed along the opposite edge of the Petri dish from the supports. 3. Distilled water is sprayed onto the paper wipe until it is fully saturated. 4. The lid is applied, creating an environment with high-relative humidity. 3.2.4.2 Loading the flow cells

In our early experiments, it was difficult to locate cells whose flagella were well adherent to the surface of the flow cell. This difficulty was resolved when we realized that the flagella of paralyzed Chlamydomonas strains act like the fins of a bomb, causing the cell to fall cell body first onto the cover glass with the flagella often pointing “up.” Angling the cover glass as in step 7 (below) gets a significant fraction of the flagella close enough to the cover glass to adhere. 1. Vortex the poly-L-lysine solution for 15–30 s. 2. Set the flow cell at a  45 angle against a convenient object and atop a paper wipe to capture flow-through.

Laser Trap Measurements of Membrane Motility

95

3. Add about 30–35 mL of poly-L-lysine to a flow cell, or whatever volume is necessary to fill your particular flow cell from edge-to-edge. 4. Put the flow cell into the humidor, laying it flat on the supports, and incubate for 20 min. 5. Wash the flow cell 2
with ddH2O (i.e., flow about 30 mL of ddH2O through the flow cell twice), as in step 2. 6. Add about 30 mL of Chlamydomonas culture. 7. Put the flow cell into the humidor for 20 minutes, this time between the supports to angle it at 45 . The larger cover glass must face down. 8. Add about 30 mL of microsphere solution to the flow cell. 9. Seal the open ends of the flow cell with vacuum grease. This is easily done with a 5 mL syringe without a needle.

3.2.5 Laser trap data collection 1. Find a cell with flagella that meet the selection criteria. i. The cell body and both flagella should appear to be adherent to the poly-L-lysine coated surface, and completely immobile. ii. The flagellum to be used for the measurements should appear to be entirely in one focal plane, indicating that it is adherent along its entire length. 2. Go through a calibration procedure (see Section 3.2.6). Calibration values for the stiffness (pN/nm) and sensitivity (V/nm) of a laser trap tend to vary more between any two trapped microspheres than they do from experiment to experiment. It is therefore vital that each microsphere be independently calibrated. Since it is usually impossible to remove a microsphere from the flagellar surface once bound, calibration should be done immediately before each measurement. The best practice is to use more than one calibration procedure on each microsphere. i. Position the microsphere at a known distance above the surface of the cover glass, typically 1–3 mm, and near the flagellum of interest but not directly over it. Five to ten micrometers of lateral separation is appropriate. ii. Begin collecting data at a sampling rate no lower than 30 k samples/s, and record approximately 10 s of thermal motion of the microsphere within the trap. iii. If the trap is so equipped, use the AOD to rapidly displace the trap 100 nm in both directions (e.g., left and right) along the orthogonal axes (x and y).

96

William H. Guilford and Robert A. Bloodgood

iv. The combination of quiescent data and “stepped” data allow for two independent measures of calibration during analysis. 3. Position the microsphere over the flagellum and lower the microsphere until it is in contact with the flagellum—approximately 300 nm from the cover glass surface. i. If the trap is equipped to do back-focal-plane interferometry, contact can be judged through a shift in the “sum” signal—the total transmitted laser light passing through the microsphere. At wavelength distances from the cover glass, this is a sensitive measure of microsphere motion along the optical axis (z). 4. Recordings of 60 s to several minutes at sampling rates of 10–50 k samples/s are typical. Transport events typically appear as “sawtooth” displacements of the microsphere away from trap center, aligned with the flagellum (Fig. 5.4A, B, and E). 5. In the form of a crude drawing or a digital image, record the orientation of the flagella relative to the cell body for every measurement and the position of the microsphere along the length of the flagellum (Fig. 5.4D). This is important for identifying anterograde versus retrograde saltations.

80 40 0 -40 -80 100 80 60 40 20 0

C

X

D

E F F

1

0

2

3

4

5

Y

Cell body 0

1 Mag

2

3

4

Anterograde 40 0 Retrograde -40 -40

5

0

40

80

X Force (pN)

F

a

v p

a

r

80

Y Force (pN)

Force (pN) Force (pN)

B

80 40 0 -40 -80

Force (pN)

A

r a 0

1

2

3

s 4

5

Time (s)

Figure 5.4 Example data. (A and B) Force traces along the X and Y axes are combined to give (C), the absolute force magnitude relative to a given baseline (gray boxes). (D) An example sketch of the cell shows the position of the microsphere (black dot) on the flagellum (F) relative to the cell body for these same data. (E) Anterograde and retrograde transport events (a and r, respectively, on C) can be distinguished on a 2D plot of X versus Y; notice that the direction of transport is well aligned to the drawing of the flagellum in (D). (F) An enlarged section of the trace in (C) illustrates how velocity (v), peak force (p), and stall force (s) are estimated. Modified from Laib et al. (2009).

Laser Trap Measurements of Membrane Motility

97

3.2.6 Calibration There are two options for calibration that are particularly amenable to flagellar membrane measurements. The first is based on power spectral density (PSD), which describes how the variance of the signal (in units2/Hz) is distributed by frequency (Allersma et al., 1998; Guilford et al., 2004; Svoboda & Block, 1994). The microsphere is trapped at a known distance above the cover glass, in the vicinity of but not directly over the flagellum of interest. A quiescent record 10 s in length of thermal (Brownian) motion is collected from the microsphere. Either online or in postprocessing the PSD is computed and several spectra are averaged to reduce noise. The PSD should show a characteristic constant region at low frequencies followed by a linear decay above a corner frequency (f0). Trap stiffness and sensitivity can then be determined by fitting the Lorenzian (Eq. 5.1) to the PSD.

 ½5:1 Sð f Þ ¼ S0 f02 = f02 þ f 2 : Trap stiffness a may be determined from f0 by a ¼ 6p2 zd f0 ,

½5:2

where  is the viscosity of the medium (0.001 for water in SI units), and d is the diameter of the microsphere. z accounts for proximity to the planar surface of the flow cell, and is described in Eq. (5.5). The detector sensitivity is determined from qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi b ¼ 3p3 zdS0 f02 =kB T , ½5:3 where kB is the Boltzmann’s constant, and T is the absolute temperature (Allersma et al., 1998; Guilford et al., 2004). While computation of the PSD and subsequent curve fitting may be performed using any suitable software (e.g., Matlab), we use custom software developed in Delphi (Borland) based on standard algorithms (Press, Teukolsky, Vetterling, & Flannery, 1989), or in real time using LabView (National Instruments). Alternatively, if the instrument is equipped to rapidly displace the laser trap itself, trap stiffness and sensitivity can be calibrated by “step response” (Dupuis, Guilford, Wu, & Warshaw, 1997; Svoboda & Block, 1994). The trap is displaced by a known distance D in a single step to a new location. The movement of the microsphere back into the center of the trap is given by

98

William H. Guilford and Robert A. Bloodgood

  eðtÞ ¼ DV 1  eat=z ,

½5:4

where e is the distance from trap center, and the fitted parameters are DV, the peak of the transient from the detector, and a. Since the initial distance between the trap and the microsphere is known, the sensitivity is simply DV/D. This comes with an important caveat, however. The step must be “rapid,” and the detector must be fast. Otherwise, the first recorded separation between the trap and the microsphere will not represent the full displacement D, but rather something smaller. We use an AOD which can move the trap in  10 ms, and our detector has a bandwidth of 150 kHz, sampled at 50 kHz. This is adequate to measure calibration values within 20% of PSD. We recommend using both methods rather than choosing one over the other. In fact, a series of rapid back-and-forth steps <1 ms in duration will not significantly perturb the PSD, so both methods can be applied to the same “quiescent” period before engaging the flagellum. Regardless of the calibration approach, it is vital to correct for hydrodynamic interactions between the microsphere and the nearby flow cell wall. The correction factor is   9 a 1 a3 45 a4 1 a5 1 z¼ 1   , ½5:5 þ 16 h 8 h 256 h 16 h where a is the radius of the microsphere, and h is the distance from the center of the microsphere to the surface of the flow cell—the “depth” (Svoboda & Block, 1994). The effect mimics an increase in viscosity. Thus this factor should be applied wherever a viscous term applies, such as in Eqs. (5.2–5.4). Otherwise there will be a systematic underestimation of trap stiffness. 3.2.7 Identification and analysis of transport events 3.2.7.1 Event selection criteria

Flagellar membrane transport events are discriminated from nonspecific cell movement by the following selection criteria: 1. There must not be appreciable movement of either the flagellum or the cell body. 2. There must not be an appreciable difference in the level of the baseline before and after a transport event. 3. For events <6 pN in magnitude where the flagellum is oriented such that it should have x- and y-axis components (i.e., the flagellum is oriented diagonally relative to the detector), then there must be movement

Laser Trap Measurements of Membrane Motility

99

evident in both channels and in the same direction (i.e., both anterograde or both retrograde) aligned to the flagellum. 4. For events <6 pN which have the flagellum oriented along the x- or y-axis: there must be movement in the major axis channel aligned to the flagellum, but there does not need to be appreciable movement in the other channel. 5. For events >6 pN in magnitude aligned to the flagellum, one may still observe small amounts of motion along the axis orthogonal to the flagellum. We do not exclude these events since neither the camera nor the flagellum is often perfectly aligned to the detector, accounting for what appear to be lateral deflections. 3.2.7.2 Analysis

a. We routinely filter the trap data to 1 kHz after calibration, but prior to analysis, using a Gaussian digital filter. This reduces the influence of Brownian motion on parameter measurements. b. We calculate the root sum of squares of the microsphere displacement along both axes to determine an absolute magnitude m of displacement (Fig. 5.4C). Note that for this to be meaningful, one has to calculate the magnitude relative to an established baseline in each of the orthogonal axes. We do this by selecting a segment of the recording that we assume is quiescent (gray boxes in Fig. 5.4A and B) and subtract the averages over these segments ( x0 and y0 ) from their respective axes before they are squared and summed. Thus qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi ½5:6 mðtÞ ¼ ðxðtÞ  x0 Þ2 þ ðyðt Þ  y0 Þ2 : Because directional cues are lost in the magnitude trace, reference is made to the original along-axis recordings (Fig. 5.4A and B) and the hand-sketched cell (Fig. 5.4D) to determine whether an event is anterograde or retrograde. c. Event durations: There are several ways of measuring the duration of the sawtooth-shaped transport events, but we have found two approaches that work for most cases and give comparable results. i. One can manually measure event duration from the first obvious displacement of the microsphere above the baseline noise to the first evident data point in the return transient. ii. A threshold crossing technique can be used. A measurement is taken from the first time point beyond which the data remains above a

100

William H. Guilford and Robert A. Bloodgood

certain threshold for at least a defined minimum time (typically 5 ms) until the data returns below threshold. d. Velocity: Velocity can be measured on either an instantaneous basis, or the overall velocity of a transport event. Either way, we generally measure velocity from the magnitude (m(t)) plot to account for movement along both axes. i. The simplest approach to measuring the overall velocity of an event is to either fit by eye or, using linear regression, fit a line to the upward slope of the event (v in Fig. 5.4F). ii. An alternative approach is to measure the instantaneous velocity as it evolves over the course of an event by fitting a series of overlapping or nonoverlapping linear segments that include short time periods (10–50 ms, typical). This not only returns a histogram of velocities, but the mean of these instantaneous velocities is the same as the overall velocity of the event. This more data-rich approach is what we consider best practice. e. Force: Remembering that, in a laser trap, force increases in linear proportion with the displacement from trap center, the peak force (p in Fig. 5.4F) reached is simply measured as the product of the trap stiffness and the displacement of the highest-magnitude point in an event (after filtering). Once again, we generally measure this from the magnitude (m(t)) plot to account for movement along both axes. Stall force is more difficult to measure, in part because it depends on one’s criterion for “stall”—the force reached where no net motion of the cargo occurs (s in Fig. 5.4F). While we routinely use 100 ms in our definition of stall as the minimum time the microsphere must remain stationary, this choice is rather arbitrary. Likewise, the word “stationary” is subject to definition. It behooves practitioners to predefine these criteria and report them along with the data. 3.2.8 Force clamping and single motor steps The experiments reported thus far use a stationary laser trap. As molecular motors in the flagellum move the extracellular cargo, force increases in linear proportion to displacement. This is analogous to auxotonic contractions in muscle, where both the length (displacement) and the tension (force) change with time rather than one being held constant. Force clamps have been used in recent years to study the dynamics of single molecular motors under a constant applied load (Block, Asbury, Shaevitz, & Lang, 2003; Lang, Asbury, Shaevitz, & Block, 2002; Patlak,

Laser Trap Measurements of Membrane Motility

101

Kad, & Warshaw, 2004; Visscher, Schnitzer, & Block, 1999). Aside from the advantage of working under a defined load, force clamps remove the effects of series compliance between the microsphere and the molecular motors, leading to clearer records than are obtained using an auxotonic trap (Visscher et al., 1999). We recently used a force clamp in the study of flagellar membrane transport. While readers are referred to the above publications for technical details, the underlying sequence of events follows. 1. A microsphere is trapped and calibrated in the usual fashion, except that the calibration values of detector sensitivity and trap stiffness are needed immediately. Thus the calibration algorithms described in Section 3.2.6 must be executed in real time. 2. The microsphere is positioned atop a flagellum. 3. When a transport event displaces the microsphere such that the force acting on it reaches a target set by the user, the force clamp is activated. 4. The position of the laser trap is changed under feedback control to hold constant the displacement between the trap center and the microsphere, and therefore the force, at the preset level. 5. The position of the laser trap is recorded as a measure of displacement under constant load. Alternatively, the microscope stage can be moved, instead of the trap itself, to hold the force constant. An advantage to moving the stage is that microspheres can be force clamped over very long distances—say the entire length of a flagellum. The advantage of moving the laser trap itself is that higher bandwidths can generally be achieved than when moving the stage. An example of force clamp data from flagellar membrane transport in Chlamydomonas pf18 is shown in Fig. 5.5A. Force was held constant at 6 pN. We recorded a much richer history of microsphere movement that is typical of auxotonic measurements in which discrete transport events are separated by quiescent periods. In addition, there are visually evident periods of discrete steps (Fig. 5.5B). The gold standard for objectively and defensibly showing the existence of and measuring the size of steps is the pairwise-difference histogram (Svoboda, Schmidt, Schnapp, & Block, 1993). For a given segment of data, the absolute difference is taken between the displacement of the microsphere at time 1 and time 2 (|d(1)  d(2)|), and this value is binned in a histogram. This is repeated for the absolute difference between each data point and every other in the data set. If the microsphere dwells at multiples of some fundamental distance, then peaks will be evident in the histogram at each

102

Displacement (nm)

A

William H. Guilford and Robert A. Bloodgood

300

200

100

0 0.0

0.5

1.0

1.5

2.0 Time (s)

3.0

3.5

C 80

1000

70

800

Frequency

Displacement (nm)

B

2.5

60 50

600 400

40 200

30

0 0 0.02

0.06

0.1 Time (s)

0.14

0.18

0

5 10 15 Pairwise difference (nm)

20

Figure 5.5 An example of a 6 pN force clamp applied to membrane transport in a pf18 cell (A). These records typically exhibit constant motion of the microsphere along the flagellum, without the long quiescent periods reported previously (Laib et al., 2009). (B) An expanded view of the region outlined in (A) has visually evident dwells and steps. (C) The segment in (B) was subjected to pairwise difference analysis (see text), which revealed peaks at 6.2  0.1, 14.6  0.2 nm, and further harmonic peaks beyond.

of these multiples. An example of this is shown in Fig. 5.5C. Gaussian fits to the peaks, including a selfsimilar peak at d ¼ 0, reveal the size of the steps. Not surprisingly, the step sizes in this example were measured to be 6.2 and 14.6 nm—close to the 8 nm and multiples thereof expected for kinesin and dynein (Reck-Peterson et al., 2006; Svoboda et al., 1993). Others have reported 8 nm steps in flagellar membrane transport using nanometerresolution optical tracking (Shih et al., 2011). While single molecule steps are not always apparent in laser trap data, this method offers the exciting possibility of measuring single molecule motor dynamics in living cells under active loads.

Laser Trap Measurements of Membrane Motility

103

3.3. Pitfalls and limitations 3.3.1 Effects of laser light on cells While some trapping has been done in plants (Ashkin & Dziedzic, 1989; Sparkes, Ketelaar, De Ruijter, & Hawes, 2009) none to our knowledge have tested the effects of laser wavelength or power on plant or algal cells. However, there have been several studies of the effects of laser traps on mammalian cell physiology and viability. Adverse effects of trapping have been reported at NIR wavelengths ranging from 740 to 765 nm. These effects include formation of giant cells (Liang et al., 1997), cross-linking of mitotic chromosomes (Vorobjev, Liang, Wright, & Berns, 1993), and failure to multiply after plating (Liang et al., 1996; Liu, Sonek, Berns, & Tromberg, 1996). Cell viability is not immediately compromised at longer wavelengths, at least at low laser powers. However, extended irradiation at 1064 nm, to which cells are relatively transparent, increases temperature by 1  C/100 mW trapping power at 1064 nm, but does not alter cytoplasmic pH nor DNA structure; loss of cell viability was observed when trapping for >2 min at 300 mW (Liu et al., 1996). All the above studies targeted the body of the cell with the laser trap. In contrast, the study of flagellar membrane motility by definition avoids irradiating the cell body. We have discerned no ill effects of trapping on the flagellum at any laser power, though we have not made a concerted effort to discover the ultimate limits. In contrast, trapping of Chlamydomonas by the cell body (as in Hollm, Khan, Marongelli, & Guilford, 2009) at an incident power of 500 mW results in an instantaneous loss of viability. Thus it is advisable to work at the lowest possible laser powers. 3.3.2 Effect of the cell cycle When grown on a carbon-free medium, the cell cycle of Chlamydomonas can be synchronized using a light–dark cycle in the growth room; this typically involves 14 h of light followed 10 h of dark. Cells are in the G1 phase of the cell cycle during the light period and divide only in the dark. This allows one to conduct experiments with cells that are known to be in the G1 phase of the cell cycle. This is important because Chlamydomonas resorb their flagella late in the light period in preparation for cell division. For reasons that are not well understood, the flagella of pf18 cells (a useful cell line for studies of microsphere movement) begin to shorten their flagella earlier in the light period than wild-type cells (Tuxhorn, Daise, & Dentler, 1998). While it

104

William H. Guilford and Robert A. Bloodgood

is known that cells that are assembling or disassembling flagella continue to exhibit microsphere movement, it is not known whether any of the parameters we measure would be sensitive to the assembly state of the flagellum. We performed experiments both early (2.5–5 h) and late (8–10.5 h) in the light period of the cycle using unmodified polystyrene microspheres. Anterograde and retrograde transport events were compared separately. The data showed that average peak forces were higher early in the light period as compared to late in the light period of the light–dark cycle (Table 5.1). Average initial velocities were also higher early in the light period as compared to late in the light period. Given these data, it is obviously important to do experiments at consistent points in the cell cycle by using light/dark synchronized cells. Practitioners should avoid the first 2–3 h after the cells enter the light portion of the cycle, and likewise avoid the end of the light cycle when cells are most apt to be actively resorbing their flagella in preparation for cell division. 3.3.3 Effect of microsphere surface chemistry and size If force and velocity scale with the number of ligated FMG-1B molecules, then one might expect carboxylated polystyrene microspheres to yield lower forces and velocities than the unmodified polystyrene microspheres, as the negatively charged surface of the flagellar glycoproteins should repel the negatively charged carboxylated microspheres. Indeed, carboxylated microspheres show lower initial velocities and peak forces than neutral polystyrene microspheres (Table 5.2). Table 5.1 Effect of light cycle on forces and velocities measured from pf18 cells Velocity (nm/s) Velocity (nm/s) 2.5–5 h light Force (pN) 8–10.5 h light Force (pN)

Anterograde

957.5

29.4

768.1

24.4

Retrograde

995.9

34.2

847.0

17.1

Table 5.2 Effects of polystyrene microsphere surface chemistry, unmodified or carboxymodified, on forces and velocities measured from pf18 cells Velocity (nm/s) Velocity (nm/s) PS Force (pN) PS–COO Force (pN)

Anterograde

957.5

29.4

624.7

22.7

Retrograde

995.9

34.2

814.6

28.3

Laser Trap Measurements of Membrane Motility

105

Microsphere size also influences force measurements, with force increasing linearly with microsphere diameter (unpublished data). Again, this is congruent with the notion that the degree of FMG-1B ligation is a modulator of motor activity; larger microspheres are predicted to have a larger contact area with the flagellar surface, and therefore recruit more FMG-1B membrane proteins, and perhaps recruit or activate a larger number of motors.

3.4. Future directions Laser traps can and have been combined with a variety of microscopic imaging and micromanipulation techniques. The most basic of these, epifluorescence, is already found on most biological laser traps (including ours), and makes possible an exciting array of experiments using fluorescent probes, including fluorescent protein-expressing strains (e.g., GFP-labeled motors). These can be crossed with paralyzed strains to make them amenable to the types of measurements described here. In addition to studying the dynamics of molecular motors within the milieu of the living cell, combining laser trap measurements of flagellar membrane transport with small molecule inhibitors will enable a better understanding of how microtubule-based intracellular transport is regulated. This is vital to understanding gliding motility in Chlamydomonas specifically, but also the roles of intracellular transport in human physiology and pathology and the regulation of motor proteins in general.

ACKNOWLEDGMENTS The authors acknowledge the hard work and contributions of Ms. Jeneva Laib who generated the data on the effects of light cycle and microsphere surface chemistry on force and velocity as well as the force clamp data. Much of this work was funded by a grant from the National Science Foundation (MCB0718430).

REFERENCES Abbondanzieri, E. A., Greenleaf, W. J., Shaevitz, J. W., Landick, R., & Block, S. M. (2005). Direct observation of base-pair stepping by RNA polymerase. Nature, 438(7067), 460–465. http://dx.doi.org/10.1038/nature04268. Adams, G. M., Huang, B., Piperno, G., & Luck, D. J. (1981). Central-pair microtubular complex of Chlamydomonas flagella: Polypeptide composition as revealed by analysis of mutants. The Journal of Cell Biology, 91(1), 69–76. http://dx.doi.org/10.1083/ jcb.91.1.69. Allersma, M. W., Gittes, F., deCastro, M. J., Stewart, R. J., & Schmidt, C. F. (1998). Twodimensional tracking of ncd motility by back focal plane interferometry. Biophysical Journal, 74(2), 1074–1085. http://dx.doi.org/10.1016/S0006-3495(98)74031-7. Ashkin, A., & Dziedzic, J. M. (1989). Internal cell manipulation using infrared laser traps. Proceedings of the National Academy of Sciences of the United States of America, 86(20), 7914–7918.

106

William H. Guilford and Robert A. Bloodgood

Block, S. M., Asbury, C. L., Shaevitz, J. W., & Lang, M. J. (2003). Probing the kinesin reaction cycle with a 2D optical force clamp. Proceedings of the National Academy of Sciences of the United States of America, 100(5), 2351–2356. http://dx.doi.org/10.1073/ pnas.0436709100. Bloodgood, R. A. (1977). Motility occurring in association with the surface of the Chlamydomonas flagellum. The Journal of Cell Biology, 75(3), 983–989. Bloodgood, R. A. (1981). Flagella-dependent gliding motility in Chlamydomonas. Protoplasma, 106(3), 183–192. Bloodgood, R. A. (2009). The Chlamydomonas flagellar membrane and its dynamic properties. In G. B. Witman, et al. (Ed.) (2nd ed.). The Chlamydomonas sourcebook, Vol. 3, (pp. 309–368). San Diego: Academic Press. Bloodgood, R. A., Leffler, E. M., & Bojczuk, A. T. (1979). Reversible inhibition of Chlamydomonas flagellar surface motility. The Journal of Cell Biology, 82(3), 664–674. Bloodgood, R. A., & Salomonsky, N. L. (1989). Use of a novel Chlamydomonas mutant to demonstrate that flagellar glycoprotein movements are necessary for the expression of gliding motility. Cell Motility and the Cytoskeleton, 13(1), 1–8. Bloodgood, R. A., & Salomonsky, N. L. (1998). Microsphere attachment induces glycoprotein redistribution and transmembrane signaling in the Chlamydomonas flagellum. Protoplasma, 202(1–2), 76–83. Bloodgood, R. A., Woodward, M. P., & Salomonsky, N. L. (1986). Redistribution and shedding of flagellar membrane glycoproteins visualized using an anti-carbohydrate monoclonal antibody and concanavalin A. The Journal of Cell Biology, 102(5), 1797–1812. Bloodgood, R. A., & Workman, L. J. (1984). A flagellar surface glycoprotein mediating cellsubstrate interaction in Chlamydomonas. Cell Motility, 4(2), 77–87. Dentler, W. (2005). Intraflagellar transport (IFT) during assembly and disassembly of Chlamydomonas flagella. The Journal of Cell Biology, 170(4), 649–659. http://dx.doi. org/10.1083/jcb.200412021. Dentler, W., Vanderwaal, K., & Porter, M. E. (2009). Recording and analyzing IFT in Chlamydomonas flagella. Methods in Cell Biology, 93, 145–155. http://dx.doi.org/ 10.1016/S0091-679X(08)93008-9. Dupuis, D., Guilford, W., Wu, J., & Warshaw, D. (1997). Actin filament mechanics in the laser trap. Journal of Muscle Research and Cell Motility, 18, 17–30. Fa¨llman, E., & Axner, O. (1997). Design for fully steerable dual-trap optical tweezers. Applied Optics, 36(10), 2107–2113. Guilford, W. H. (2001). Laser traps in cell biology and biophysics. In Methods in cellular imaging, Vol. 1, (pp. 381–394). New York: Oxford University Press. Guilford, W. H., Tournas, J. A., Dascalu, D., & Watson, D. S. (2004). Creating multiple time-shared laser traps with simultaneous displacement detection using digital signal processing hardware. Analytical Biochemistry, 326(2), 153–166. Harris, E. H. (2009). The Chlamydomonas sourcebook: Introduction to Chlamydomonas and its laboratory use, Vol. 1. Academic Press. Hollm, J. A., Khan, R. P., Marongelli, E. N., & Guilford, W. H. (2009). Laser trap characterization and modeling of phototaxis in Chlamydomonas reinhardtii. Cellular and Molecular Bioengineering, 2(2), 244–254. Kozminski, K. G., Beech, P. L., & Rosenbaum, J. L. (1995). The Chlamydomonas kinesinlike protein fla10 is involved in motility associated with the flagellar membrane. The Journal of Cell Biology, 131(6), 1517–1527. Kozminski, K. G., Johnson, K. A., Forscher, P., & Rosenbaum, J. L. (1993). A motility in the eukaryotic flagellum unrelated to flagellar beating. Proceedings of the National Academy of Sciences, 90(12), 5519–5523. Laib, J., Marin, J., Bloodgood, R., & Guilford, W. (2009). The reciprocal coordination and mechanics of molecular motors in living cells. Proceedings of the National Academy of Sciences, 106(9), 3190–3195. http://dx.doi.org/10.1073/pnas.0809849106.

Laser Trap Measurements of Membrane Motility

107

Lang, M. J., Asbury, C. L., Shaevitz, J. W., & Block, S. M. (2002). An automated twodimensional optical force clamp for single molecule studies. Biophysical Journal, 83(1), 491–501. Lewin, R. A. (1952). Studies on the flagella of algae. I. General observations on Chlamydomonas moewusii Gerloff. The Biological Bulletin, 103(1), 74–79. Liang, H., Vu, K. T., Krishnan, P., Trang, T. C., Shin, D., Kimel, S., et al. (1996). Wavelength dependence of cell cloning efficiency after optical trapping. Biophysical Journal, 70(3), 1529–1533. http://dx.doi.org/10.1016/S0006-3495(96)79716-3. Liang, H., Vu, K. T., Trang, T. C., Shin, D., Lee, Y. E., Nguyen, D. C., et al. (1997). Giant cell formation in cells exposed to 740 nm and 760 nm optical traps. Lasers in Surgery and Medicine, 21(2), 159–165. Liu, Y., Sonek, G. J., Berns, M. W., & Tromberg, B. J. (1996). Physiological monitoring of optically trapped cells: Assessing the effects of confinement by 1064-nm laser tweezers using microfluorometry. Biophysical Journal, 71(4), 2158–2167. Luck, D., Piperno, G., Ramanis, Z., & Huang, B. (1977). Flagellar mutants of Chlamydomonas: Studies of radial spoke-defective strains by dikaryon and revertant analysis. Proceedings of the National Academy of Sciences, 74(8), 3456–3460. Mallik, R., & Gross, S. P. (2009). Intracellular transport: How do motors work together? Current Biology, 19(10), R416–R418. http://dx.doi.org/10.1016/j.cub.2009.04.007. Patlak, J. B., Kad, N. M., & Warshaw, D. M. (2004). Transient force clamp: A new catch and release program for single myosin molecules. Biophysical Journal, 86(1), 53a. Pazour, G. J., Dickert, B. L., & Witman, G. B. (1999). The DHC1b (DHC2) isoform of cytoplasmic dynein is required for flagellar assembly. The Journal of Cell Biology, 144(3), 473–481. Press, W. H., Teukolsky, S. A., Vetterling, W. T., & Flannery, B. P. (1989). Numerical recipes in Pascal. New York: Cambridge University Press. Reck-Peterson, S. L., Yildiz, A., Carter, A. P., Gennerich, A., Zhang, N., & Vale, R. D. (2006). Single-molecule analysis of dynein processivity and stepping behavior. Cell, 126(2), 335–348. http://dx.doi.org/10.1016/j.cell.2006.05.046. Sager, R., & Granick, S. (1953). Nutritional studies with Chlamydomonas reinhardi. Annals of the New York Academy of Sciences, 56(5), 831–838. http://dx.doi.org/10.1111/j.17496632.1953.tb30261.x. Sheetz, M. P. (Ed.), (1997). Laser tweezers in cell biology, In: Methods in Cell Biology Vol. 55. San Diego: Academic Press. Shih, S.-M., Kocabas, F., & Yildiz, A. (2011). Single-molecule analysis of intraflagellar transport in live Chlamydomonas cells. Biophysical Journal, 100(3 Suppl. 1), 11a. http://dx. doi.org/10.1016/j.bpj.2010.12.269. Sparkes, I. A., Ketelaar, T., De Ruijter, N. C. A., & Hawes, C. (2009). Grab a Golgi: Laser trapping of golgi bodies reveals in vivo interactions with the endoplasmic reticulum. Traffic, 10(5), 567–571. http://dx.doi.org/10.1111/j.1600-0854.2009.00891.x. Svoboda, K., & Block, S. M. (1994). Biological applications of optical forces. Annual Review of Biophysics and Biomolecular Structure, 23(1), 247–285. http://dx.doi.org/10.1146/ annurev.bb.23.060194.001335. Svoboda, K., Schmidt, C. F., Schnapp, B. J., & Block, S. M. (1993). Direct observation of kinesin stepping by optical trapping interferometry. Nature, 365(6448), 721–727. Tuxhorn, J., Daise, T., & Dentler, W. L. (1998). Regulation of flagellar length in Chlamydomonas. Cell Motility and the Cytoskeleton, 40(2), 133–146. http://dx.doi. org/10.1002/(SICI)1097-0169. Visscher, K., Schnitzer, M. J., & Block, S. M. (1999). Single kinesin molecules studied with a molecular force clamp. Nature, 400(6740), 184–189. Vorobjev, I. A., Liang, H., Wright, W. H., & Berns, M. W. (1993). Optical trapping for chromosome manipulation: A wavelength dependence of induced chromosome bridges. Biophysical Journal, 64(2), 533–538. http://dx.doi.org/10.1016/S0006-3495(93)81398-5. Witman, G. (Ed.), (2009). In: The Chlamydomonas sourcebook: Cell motility and behavior, Vol. 3. San Diego: Academic Press.