CHAPTER
Axonemal motility in Chlamydomonas
19
Ken-ichi Wakabayashi*, Ritsu Kamiyax, 1 *
x
Chemical Resources Laboratory, Tokyo Institute of Technology, Yokohama, Japan Department of Life Science, Faculty of Science, Gakushuin University, Tokyo, Japan 1
Corresponding author: E-mail: [email protected]
CHAPTER OUTLINE Introduction ............................................................................................................ 388 1. Methods ............................................................................................................ 389 1.1 Swimming Tracks and Average Swimming Velocities .............................. 389 1.1.1 Swimming velocities and swimming directions................................... 390 1.1.2 Estimation of net propulsive force ..................................................... 391 1.2 Waveform Analysis .............................................................................. 391 1.3 Beat Frequency Measurements ............................................................. 393 1.4 Reactivation of Demembranated Cell Models ......................................... 394 1.5 Reactivation of Isolated Flagellar Axonemes .......................................... 396 1.6 Sliding Disintegration of Axonemes....................................................... 397 Acknowledgments ................................................................................................... 400 References ............................................................................................................. 400
Abstract Motile cilia and flagella rapidly propagate bending waves and produce water flow over the cell surface. Their function is important for the physiology and development of various organisms including humans. The movement is based on the sliding between outer doublet microtubules driven by axonemal dyneins, and is regulated by various axonemal components and environmental factors. For studies aiming to elucidate the mechanism of cilia/flagella movement and regulation, Chlamydomonas is an invaluable model organism that offers a variety of mutants. This chapter introduces standard methods for studying Chlamydomonas flagellar motility including analysis of swimming paths, measurements of swimming speed and beat frequency, motility reactivation in demembranated cells (cell models), and observation of microtubule sliding in disintegrating axonemes. Most methods may be easily applied to other organisms with slight modifications of the medium conditions.
Methods in Cell Biology, Volume 127, ISSN 0091-679X, http://dx.doi.org/10.1016/bs.mcb.2014.12.002 © 2015 Elsevier Inc. All rights reserved.
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INTRODUCTION Motile cilia and flagella produce water flow over the cell surface by propagating bending waves. The movement is based on the function of axonemal dyneins that produce sliding between outer doublet microtubules (Satir, 1968; Summers & Gibbons, 1971). To meet the cell’s needs, the movement in most cases is regulated in beat frequency (the number of wave propagation event per second) and bending pattern (waveform). Previous studies have shown that Ca2þ and protein phosphorylation are common regulatory factors, which directly or indirectly act on dyneins and cause changes in the pattern and/or velocity of sliding between the nine outer doublet microtubules (Bannai, Yoshimura, Takahashi, & Shingyoji, 2000; Habermacher & Sale, 1997; Naitoh & Kaneko, 1972; Wargo & Smith, 2003). Beat frequency and waveform also vary depending on cell’s genetic background and physiological conditions such as the intracellular ATP concentration. Cilia and flagella movement has been studied in a variety of organisms ranging from protozoa to mammals depending on the purpose of study. For example, sea urchin sperm has been used for detailed analyses of waveform (Gibbons, 1986), force generation, and regulation in microtubule sliding (Kamimura & Takahashi, 1981; Morita & Shingyoji, 2004); and Paramecium for studies on the collective properties (Machemer, 1972) and ionic regulation of ciliary movements (Naitoh & Eckert, 1969). Mammalian trachea cilia have been studied in relation to human physiology (Lorenzo, Liedtke, Sanderson, & Valverde, 2008; Shah, Ben-Shahar, Moninger, Kline, & Welsh, 2009). Recently, however, Chlamydomonas has been by far the most extensively used for motility analysis, especially in studies aimed at an understanding of the function of particular axonemal proteins or assemblies in flagellar motility mechanism. Chlamydomonas is preferentially used because it has excellent properties not possessed by other organisms: it can be cultured in large quantities, displays flagellar movement that is easy to observe, and, in particular, offers a wide range of flagella-deficient mutants. With the establishment of genome database and flagellar proteome database, the majority of mutations can now be relatively easily identified at the gene level. Most importantly, the basic structure, as well as the major components, of Chlamydomonas flagella is common to cilia and flagella of other organisms including humans. This chapter describes several methods for the assessment of Chlamydomonas flagellar movements in live cells, demembranated and reactivated cells (cell models), and isolated axonemes. In addition, a simple method is introduced for velocity measurements of microtubule sliding in disintegrating axonemes. We used these methods for isolation and characterization of various mutants deficient in flagellar motility as well as for analysis of the effects of physical and chemical factors on flagellar motility (Kamiya, 2002). Measurements of beat frequency and waveforms demonstrated that outer arm dynein is particularly important for the flagella to beat with a high frequency, whereas inner arm dyneins are important for producing effective waveforms (Brokaw & Kamiya, 1987). Experiments with demembranated cells and axonemes revealed that the two flagellar axonemes on a
1. Methods
single cell are differentially regulated by submicromolar Ca2þ (Kamiya & Witman, 1984); cAMP-dependent protein phosphorylation inhibits flagellar motility (Hasegawa, Hayashi, Asakura, & Kamiya, 1987); reductioneoxidation balance affects flagellar beat frequency (Wakabayashi & King, 2006); and the axonemes of paralyzed-flagella mutants lacking the central pair and radial spokes can beat under some nonphysiological nucleotide conditions (Omoto, Yagi, Kurimoto, & Kamiya, 1996) and display Ca2þ-dependent waveform conversion (Wakabayashi, Yagi, & Kamiya, 1997). Measurements of microtubule sliding velocity showed that the outer arm dynein and inner arm dynein intrinsically differ in sliding velocity (Kurimoto & Kamiya, 1991; Okagaki & Kamiya, 1986), leading to the recognition that the axoneme is equipped with motors operating at different speeds. Although the details of the conditions described below are optimized for Chlamydomonas, most methods may be readily used in cilia and flagella of other organisms after slight modification of the conditions.
1. METHODS 1.1 SWIMMING TRACKS AND AVERAGE SWIMMING VELOCITIES Average swimming velocity of Chlamydomonas cells can be obtained from the cells’ swimming path lengths per a given time. Previously, we manually traced the swimming paths on a transparent sheet overlaid on the monitor screen and measured the length with a scale. Recently, it is easier to track each cell image in a personal computer. For example, an Image J plug-in, MTrack2 (http://valelab.ucsf.edu/ wnstuurman/IJplugins/MTrack2.html) enables automatic tracking (Figure 1). In addition to swimming velocities, swimming tracks provide useful information about
FIGURE 1 (A) Swimming tracks of Chlamydomonas cells taken under a dark-field microscope. Time interval, 0.1 s. Image brightness is increased with time, so that the swimming direction is readily determined. Bar: 0.1 mm. (B) Swimming paths of the cells in (A) auto-tracked by Image J and MTracks2 plug-in.
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the cell’s motility; the tracks are important for assessing cells’ phototactic activity and the tendency of directional change. The pitch of the helical swimming path reflects characteristic features of flagellar beating pattern of a particular cell type.
1.1.1 Swimming velocities and swimming directions 1. Observe and video record the swimming paths of Chlamydomonas cells under a microscope at low magnification (objective: 10). The record must be saved in Audio Video Interleave (AVI) format, uncompressed. Dark-field or phasecontrast microscope is recommended to obtain high-contrast images. Cells often accumulate in the field of observation due to phototaxis, which interferes with the observation of individual cells. In such a case, place a red filter (630 nm) on the light source. However, pay attention to the fact that the swimming velocity varies with illumination; cells swim slightly slower under red light than under white light because of a response to the light conditions (this phenomenon is called the photokinesis). 2. Playback the AVI file using Image J. Binarize the images so that the cells appear in black and background in white. Run MTrack2. Obtain swimming tracks for w50 cells each for 3e5 s. 3. Analyze the resultant tracking data using a spreadsheet program, such as Excel, to obtain the average swimming velocity, standard deviation, and if necessary, other parameters such as the frequency of directional changes in swimming paths. Notes For phototaxis analysis, place a red filter on the light source of the microscope and set a light emission diode (l ¼ 500e525 nm) to stimulate the cells from the side. Measure the angle between the light direction and the swimming path of each cell. The data can be shown as a polar histogram (Figure 2) or a phototactic
FIGURE 2 Polar histograms showing swimming directions of wild-type cells. Bars represent the percentage of cells moving in a particular direction relative to the stimulating light (l ¼ 525 nm) from the right (12 bins of 30 ; n ¼ 50 cells per conditions). Swimming tracks for 1.5 s were examined. (A) Without stimulating light, cells swim in random directions. (B) When stimulating light is illuminated from the right (0 ), most cells show positive phototaxis and swim toward the light source. Modified from Wakabayashi et al. (2011).
1. Methods
index, the average of jcos qj (Okita, Isogai, Hirono, Kamiya, & Yoshimura, 2005; Wakabayashi, Misawa, Mochiji, & Kamiya, 2011).
1.1.2 Estimation of net propulsive force As an application of the above method, we can estimate the net propulsive force of flagellar movement in viscous media from the swimming velocity. The force acting on the moving cell body is proportional to the product of the viscosity (h), cell size, and moving velocity (v); if we approximate the cell body as a sphere of radius r, then the viscous force acting on it is given by 6phrv (Stokes’ formula). Thus it is straightforward to compare the load dependence of net flagellar propulsive force between different strains. Using this method, we found that the outer arm dynein and inner arm dynein greatly differ in the load dependence (Minoura & Kamiya, 1995), and that a particular kind of single-headed inner arm dynein (dynein c) is important for propulsive force production at high viscosity (Yagi et al., 2005). 1. Prepare viscous solution by dissolving 0e16% (w/v) Ficoll (Type 400, Sigma; Mr ¼ 400,000) in appropriate medium or buffer. Other polymers can be also used, but Ficoll produces less local inhomogeneity in solution than other polymers such as methylcellulose and, therefore, is more appropriate (Berg & Turner, 1979). If the actual force value needs to be determined, measure the solution viscosity with an Ostwald or other types of viscometer. 2. Record the tracking of swimming cells in the viscous sample solutions. Care must be taken that the sample consists of cells with similar diameters, since the swimming velocity in viscous media sensitively varies with load, which is proportional to the cell diameter. To prepare cells with similar diameters, synchronize cell cultures on an appropriate lightedark cycle, and measure the motility at a constant phase of the cycle.
1.2 WAVEFORM ANALYSIS High-speed recording is necessary for observation of flagellar waveform. Flagellar beat frequency in wild-type Chlamydomonas is w60 Hz; therefore, one beat cycle can be recorded in w5 frames if a movie is taken at 300 fps. Cameras with this frame rate are available from several manufacturers at relatively low costs. Even some home-use digital still cameras are equipped with a high-speed recording mode. It is important to choose a fast shutter speed to obtain clear images of waveform. For waveform analysis, we must obtain a time series of flagellar images that are in focus for at least one beating cycle. We use uniflagellated cells that rotate within a small area by beating only the single flagellum and retain the flagellar image in focus for a long time. When observation of the two flagella on a single cell is necessary, use cells that are stuck between the cover slip and glass slide or those held with a micropipette. We use the following “uniflagella” method. 1. Produce uniflagellated cells by gently homogenizing the cell culture, or by producing cells with the background of a uniflagella mutation, uni1
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(Huang, Ramanis, Dutcher, & Luck, 1982). In the former case, we must choose by visual inspection an appropriate cell that lacks one of the two flagella while maintaining the other flagellum intact, each time a movie is taken. In the latter case, we must pay attention to the fact that the uni1 mutant retains only one particular flagellum (the trans-flagellum; the flagellum farthest from the eyespot) of the two flagella. Previous studies showed that the trans-flagellum tends to beat at higher frequency than the other, cis-flagellum (Kamiya & Hasegawa, 1987). 2. Take a high-speed movie of a uniflagellated cell rotating under a microscope with a 40 objective (Figure 3(A)).
FIGURE 3 (A) Sequential images of a uni1 cell rotating by beating its single flagellum. Taken at 300 frames/s. (B) The angle between the tangents to the waveform at the base (dotted line) and at point s (at distance s along the flagellar length from the base) represents the “shear angle” (q) at this position. (C) Typical shear curves of wild-type (left) and ida4 (right) flagella. ida4 displays a smaller shear amplitude. The slope of each line gives the curvature at a given position. Flagellar waveforms were manually traced three times and the average of these data was smoothed by the method of Brokaw (1983). Modified from Minoura and Kamiya (1995).
1. Methods
3. Analyze flagellar waveform from sequential images of flagellar beating. For most purposes, a series of flagellar beating images will give enough information even without any analysis. When quantitative data are required, each image must be traced and analyzed. A standard method is to measure the angle (shear angle, q) between the tangent to a small segment at the position s (measured from the base along the flagellum) and the tangent to the basal segment, and plot q against s for the entire length (Brokaw, 1979; Brokaw, Luck, & Huang, 1982) (Figure 3(B) and (C)). Because q is proportional to the sliding displacement between adjacent outer doublet microtubules at point s, the plots produce a single curve that represents the relative microtubule sliding along the flagellar length. Hence, the curve is called a shear curve. From several sets of plots for different beating phases over a complete beat cycle, we can determine the amplitude and velocity of back-and-forth microtubule sliding at each position along the flagellum. The curves also give parameters such as the bend angle and curvature in the principal and reverse bends. Several programs are available that generate sequential images from a movie file, such as iMovie (bundled with Mac OSX) and TMPGEnc (shareware, a free version is also available at http:// www.tmpgenc.net/). 4. Beat frequency is readily obtained from sequential images of flagellar beating. Choose a pair of images showing similar waveforms in n cycles (n ¼ 1e3) of beating strokes, count the number (m) of frames between the two similar waveforms, and calculate (n/m) (frame rate) to yield the beat frequency. For average beat frequency values, use data from 30 to 50 cells. Notes When uniflagellated cells are produced by homogenization of bi-flagellated cells, we must determine whether the flagellum being observed is cis or trans relative to the eyespot. Eyespot is more easily observed with an oil-immersion dark-filed condenser than with a dry condenser; it appears as a bright spot located on the side of a rotating cell.
1.3 BEAT FREQUENCY MEASUREMENTS A convenient method for estimating the average beat frequency in a population of cells has been developed that uses frequency analysis of the cell body vibration (Kamiya, 2000). This method relies on the fact that the Chlamydomonas cell body vibrates back-and-forth in synchrony with flagellar beating. The equipment for this measurement is relatively easily constructed from a photosensor, an A/D converter, and an Fast Fourier Transform (FFT) program run on a PC. The outline of the method is as follows. Images of 10e100 swimming cells in a dark-field microscope are projected on a graded filter, and the transmitted light is detected with the sensor. The graded filter converts the back-and-forth movements of cell images into a light intensity fluctuation (however, we found that this graded filter is practically not necessary, because the brightness of microscope image field is usually not uniform and the light
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intensity from the cell images fluctuates even without it). We are using a photodetector composed of an appropriate photodiode (such as Hamamatsu S1223), lownoise operational amplifiers (such as OPA111, Burr Brown, Tucson, AZ, USA), and 1-GU feedback resistors (circuit diagrams are available at the manufacturer’s Web site). The photodetector is attached to a dark-field microscope with an objective of 10 to 20 and a light source of a 100-W halogen lamp. FFT is performed using the SIGVIEW software package (http://www.sigview.com/). This method allows us to measure the average beat frequency within a minute. Unlike the high-speed video method using the uni1 mutant, this method allows measurements of average beat frequencies of the two flagella. In demembranated and reactivated cell models (see below), the two axonemes tend to beat independently, giving rise to two peak frequencies reflecting the two frequencies of the cell body vibration caused by both cis- and trans-flagellar axonemes (Figure 4).
1.4 REACTIVATION OF DEMEMBRANATED CELL MODELS In vitro reactivation of cell movement is a powerful method to study cell motility mechanism. Feasibility of such an experiment was first demonstrated by SzentGyo¨rgyi (1949) who used rabbit skeletal muscle fibers extracted with 50% glycerol. It is an excellent experimental system that allows us to easily change the composition of reactivating solution, which corresponds to the cytoplasm in vivo. In fact, it was used in experiments to establish that muscle contraction is dependent on ATP. This method has been used in various studies on muscle contraction, in order to investigate the effects of ATP concentration, Ca2þ concentration, protein phosphorylation, etc. Hoffman-Berling (1954) applied this method to sperm flagella and showed that flagellar movement is also energized by ATP. He called the extracted cells “cell models.” Glycerol was later replaced by nonionic detergent Triton X-100 by Gibbons and Gibbons (1972) for sea urchin sperm cell models and by Naitoh and Kaneko (1972) for Paramecium cell models. These extracted cells were called “Triton models.” In the case of Chlamydomonas cell models, Nonidet P-40 has been more frequently used than Triton X-100 (Witman, Plummer, & Sander, 1978), although both detergent give similarly good results (our unpublished observation). Currently we use the following protocols based on those used by Witman et al. (1978) for studying axonemal motility. Solutions (named after constituents) • •
• • •
HES: 10 mM Hepes, pH 7.4, 1 mM EGTA, 4% sucrose HMDEKP: 30 mM Hepes, pH 7.4, 5 mM MgSO4, 1 mM dithiothreitol (DTT), 1 mM EGTA, 50 mM K-acetate, 1% polyethylene glycol (Mr ¼ 20,000) (add an ATP-regeneration system consisting of 70 U/mL creatine phosphokinase and 5 mM creatine phosphate for stable reactivation at 105 M, axonemes tend to detach from the cell body because Chlamydomonas has a Ca2þ-dependent mechanism for flagellar autotomy. When observation of cell model movements at higher Ca2þ concentrations is necessary, use the mutant fa1 deficient in the autotomy (Lewin & Burrascano, 1983) (available from the Chlamydomonas Resource Center).
1.5 REACTIVATION OF ISOLATED FLAGELLAR AXONEMES Chlamydomonas flagella and axonemes can be readily isolated by deflagellation with dibucaine and differential centrifugation (Witman et al., 1978). The isolated flagella and axonemes can be used in various experiments, including motility reactivation, SDS-PAGE, immunofluorescence microscopy and electron microscopy. Motility reactivation in isolated axonemes has been used to examine the effect of different concentrations of Ca2þ (Bessen, Fay, & Witman, 1980) and cAMP (Hasegawa et al., 1987). Chlamydomonas axonemes change their beating patterns from an asymmetric, ciliary pattern at 2 min) tends to result in poor reactivation. Spin down the cell bodies by centrifugation at 600 g for 5 min at 4 C. Transfer the supernatant to a new tube (placed on ice), and centrifuge at 1600 g for 2.5 min at 4 C. Transfer the supernatant to a new tube (placed in ice), and subject to centrifugation at 14,000 g for 12 min at 4 C. Discard the supernatant, resuspend the precipitates in 200 mL 0.2% NP-40 in HMDEK. Mix 80 mL of HMDEKP, 10 mL of ATP solution, and 10 mL of axonemes in a 0.5-mL test tube. To examine the effect of Ca2þ, replace the EGTA with one of the Ca2þ buffers shown in Table 1.
1.6 SLIDING DISINTEGRATION OF AXONEMES Axonemes beat through microtubule sliding driven by various kinds of axonemal dyneins. Sliding movements can be directly observed in protease-treated axonemes, Table 1 Composition of Calcium Buffers for Reactivation Experiments Free [Ca2D] (M)* pCa4 pCa5 pCa6 pCa7 pCa8 *
4
0.96 10 1.01 105 1.04 106 1.06 107 1.00 108
At 1 mM ATP.
CaCl2 (M) 3
1.60 10 1.60 104 4.75 103 3.20 103 3.70 104
EGTA (M)
EDTA (M)
0 0 5.00 103 4.80 103 2.30 103
2.00 103 6.50 104 0 0 0
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CHAPTER 19 Axonemal motility in Chlamydomonas
FIGURE 5 Flow chamber for observation of sliding disintegration. A coverslip is held on a glass slide via two pieces of double-sided adhesive tape w80 mm thick. A reservoir for the perfusing solution is made with nail varnish by one of the two open sides. Perfusion is started by putting a drop of solution to the reservoir and placing a piece of filter paper on the other side.
as first shown by Summers and Gibbons (1971). The following protocols for Chlamydomonas axonemes are based on Okagaki and Kamiya (1986) and Kurimoto and Kamiya (1991) who measured the velocity of microtubule sliding during disintegration occurring upon perfusion with ATP and protease (Figure 6). Methods similar to the one outlined below have been used in studies that explored regulatory mechanism of axonemal motility (Habermacher & Sale, 1995; Smith & Sale, 1992). We tend to regard the sliding velocities measured in disintegrating axonemes as reflecting the movement under very little load. However, microtubule sliding in disintegrating axonemes does not reflect the maximal velocity that can be achieved by axonemal microtubules under minimal loads. This is because the sliding velocity increases if the axonemes are more extensively treated with protease (Kikushima, 2009; Yano & Miki-Noumura, 1981), which may well function
FIGURE 6 Sliding disintegration of a wild-type axoneme fragment. ATP, 1 mM. Interval: 1/15 s. Bar, 10 mm. This axoneme fragment is undergoing sliding at w13 mm/s.
1. Methods
to reduce interdoublet friction. Thus we must be careful in interpreting the sliding velocity data; it can be affected by the intrinsic sliding velocity and the force produced by dynein(s), and also by the friction between outer doublets. A higher sliding velocity does not necessarily mean a higher activity of axonemal dyneins. Solutions • • • • • 1.
2. 3. 4.
5. 6. 7. 8. 9. 10.
11.
12.
HMDEKP: 30 mM Hepes, pH 7.4, 5 mM MgSO4, 1 mM DTT, 1 mM EGTA, 50 mM K-acetate, 1% polyethylene glycol (Mr ¼ 20,000) HMDEK: 30 mM Hepes, pH 7.4, 5 mM MgSO4, 1 mM DTT, 1 mM EGTA, 50 mM K-acetate 2% Nonidet P-40 in HMDEKP 1 mM ATP in HMDEKP 2 mg/mL Nagarse (Type XXIV protease, Sigma), 1 mM ATP in HMDEKP Prepare a flow chamber from a glass slide and a coverslip with spacers (pieces of double-sided adhesive tape) (Figure 5). Use nail varnish to produce a reservoir for perfusion solution by one of the two open sides of the cover glass. Prepare flagella from an w200 mL cell culture as described above. Dilute the flagellar suspension with HMDEKP so that the total volume is w1 mL, in a 1.5 mL Eppendorf tube. Produce flagellar fragments by sonication with an appropriate sonicator. We use a Sonifier 250 (Branson) with a microtip and apply sonication for 4 s with the power set to “3” and the duty ratio to 30%. Collect flagellar fragments by centrifugation at 14,000 g for 12 min at 4 C. Carefully remove the supernatant and resuspend the precipitate in w50 mL HMDEKP. Mix 85 mL HMDEKP, 10 mL flagella fragments, and 5 mL 2% NP-40 in a test tube and place it in ice (this sample is referred to as “axoneme fragments”). Fill the flow chamber with 10 mL of axoneme fragments. Perfuse the chamber with 40 mL of ATP solution. Observe the sample with a dark-field microscope capable of visualizing single microtubules. We use an Olympus microscope equipped with an oilimmersion condenser (NA 1.2e1.4), a light source of 100-W mercury arc lamp and a 50 objective with an iris (NA 0.90). Images are recorded with a sensitive video camera. Find a field where many axoneme fragments are attached on the glass surface. Perfuse the sample with the Nagarse solution that contains both protease and ATP. Under optimal conditions, sliding disintegration will occur within w30 s after onset of perfusion. Measure the sliding velocity after tracing the moving microtubule images using Image J and an appropriate plug-in, as outlined above. Notes
1. Elastase can be used in place of “Nagarse.”
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ACKNOWLEDGMENTS We thank Toshiki Yagi (Prefectural University of Hiroshima) and Mikito Owa (University of Tokyo) for helpful advice, and Elina Nakamasu (Tokyo Institute of Technology) for her help in preparing figures. Our studies have been supported by grants-in-aid from Japan Society for Promotion of Sciences (#25113507, #25117506, #25291058, and #26650093 to KW, and #23570189 and #25117521 to RK).
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Summers, K. E., & Gibbons, I. R. (1971). Adenosine triphosphate-induced sliding of tubules in trypsin-treated flagella of sea-urchin sperm. Proceedings of the National Academy of Science of the United States of America, 68, 3092e3096. Szent-Gyo¨rgyi, A. (1949). Free-energy relations and contraction of actomyosin. Biological Bulletin, 96, 140e161. Wakabayashi, K., & King, S. M. (2006). Modulation of Chlamydomonas reinhardtii flagellar motility by redox poise. The Journal of Cell Biology, 173, 743e754. Wakabayashi, K., Misawa, Y., Mochiji, S., & Kamiya, R. (2011). Reduction-oxidation poise regulates the sign of phototaxis in Chlamydomonas reinhardtii. Proceedings of the National Academy of Sciences of the United States of America, 108, 11280e11284. Wakabayashi, K., Yagi, T., & Kamiya, R. (1997). Ca2þ-dependent waveform conversion in the flagellar axoneme of Chlamydomonas mutants lacking the central-pair/radial spoke system. Cell Motility and the Cytoskeleton, 38, 22e28. Wargo, M. J., & Smith, E. F. (2003). Asymmetry of the central apparatus defines the location of active microtubule sliding in Chlamydomonas flagella. Proceedings of the National Academy of Sciences of the United States of America, 100, 137e142. Witman, G. B., Plummer, J., & Sander, G. (1978). Chlamydomonas flagellar mutants lacking radial spokes and central tubules. Structure, composition, and function of specific axonemal components. The Journal of Cell Biology, 76, 729e747. Yagi, T., Minoura, I., Fujiwara, A., Saito, R., Yasunaga, T., Hirono, M., et al. (2005). An axonemal dynein particularly important for flagellar movement at high viscosity. Implications from a new Chlamydomonas mutant deficient in the dynein heavy chain gene DHC9. Journal of Biological Chemistry, 280, 41412e41420. Yano, Y., & Miki-Noumura, T. (1981). Recovery of sliding ability in arm-depleted flagellar axonemes after recombination with extracted dynein I. Journal of Cell Science, 48, 223e239.
Axonemal motility in Chlamydomonas
19
Ken-ichi Wakabayashi*, Ritsu Kamiyax, 1 *
x
Chemical Resources Laboratory, Tokyo Institute of Technology, Yokohama, Japan Department of Life Science, Faculty of Science, Gakushuin University, Tokyo, Japan 1
Corresponding author: E-mail: [email protected]
CHAPTER OUTLINE Introduction ............................................................................................................ 388 1. Methods ............................................................................................................ 389 1.1 Swimming Tracks and Average Swimming Velocities .............................. 389 1.1.1 Swimming velocities and swimming directions................................... 390 1.1.2 Estimation of net propulsive force ..................................................... 391 1.2 Waveform Analysis .............................................................................. 391 1.3 Beat Frequency Measurements ............................................................. 393 1.4 Reactivation of Demembranated Cell Models ......................................... 394 1.5 Reactivation of Isolated Flagellar Axonemes .......................................... 396 1.6 Sliding Disintegration of Axonemes....................................................... 397 Acknowledgments ................................................................................................... 400 References ............................................................................................................. 400
Abstract Motile cilia and flagella rapidly propagate bending waves and produce water flow over the cell surface. Their function is important for the physiology and development of various organisms including humans. The movement is based on the sliding between outer doublet microtubules driven by axonemal dyneins, and is regulated by various axonemal components and environmental factors. For studies aiming to elucidate the mechanism of cilia/flagella movement and regulation, Chlamydomonas is an invaluable model organism that offers a variety of mutants. This chapter introduces standard methods for studying Chlamydomonas flagellar motility including analysis of swimming paths, measurements of swimming speed and beat frequency, motility reactivation in demembranated cells (cell models), and observation of microtubule sliding in disintegrating axonemes. Most methods may be easily applied to other organisms with slight modifications of the medium conditions.
Methods in Cell Biology, Volume 127, ISSN 0091-679X, http://dx.doi.org/10.1016/bs.mcb.2014.12.002 © 2015 Elsevier Inc. All rights reserved.
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INTRODUCTION Motile cilia and flagella produce water flow over the cell surface by propagating bending waves. The movement is based on the function of axonemal dyneins that produce sliding between outer doublet microtubules (Satir, 1968; Summers & Gibbons, 1971). To meet the cell’s needs, the movement in most cases is regulated in beat frequency (the number of wave propagation event per second) and bending pattern (waveform). Previous studies have shown that Ca2þ and protein phosphorylation are common regulatory factors, which directly or indirectly act on dyneins and cause changes in the pattern and/or velocity of sliding between the nine outer doublet microtubules (Bannai, Yoshimura, Takahashi, & Shingyoji, 2000; Habermacher & Sale, 1997; Naitoh & Kaneko, 1972; Wargo & Smith, 2003). Beat frequency and waveform also vary depending on cell’s genetic background and physiological conditions such as the intracellular ATP concentration. Cilia and flagella movement has been studied in a variety of organisms ranging from protozoa to mammals depending on the purpose of study. For example, sea urchin sperm has been used for detailed analyses of waveform (Gibbons, 1986), force generation, and regulation in microtubule sliding (Kamimura & Takahashi, 1981; Morita & Shingyoji, 2004); and Paramecium for studies on the collective properties (Machemer, 1972) and ionic regulation of ciliary movements (Naitoh & Eckert, 1969). Mammalian trachea cilia have been studied in relation to human physiology (Lorenzo, Liedtke, Sanderson, & Valverde, 2008; Shah, Ben-Shahar, Moninger, Kline, & Welsh, 2009). Recently, however, Chlamydomonas has been by far the most extensively used for motility analysis, especially in studies aimed at an understanding of the function of particular axonemal proteins or assemblies in flagellar motility mechanism. Chlamydomonas is preferentially used because it has excellent properties not possessed by other organisms: it can be cultured in large quantities, displays flagellar movement that is easy to observe, and, in particular, offers a wide range of flagella-deficient mutants. With the establishment of genome database and flagellar proteome database, the majority of mutations can now be relatively easily identified at the gene level. Most importantly, the basic structure, as well as the major components, of Chlamydomonas flagella is common to cilia and flagella of other organisms including humans. This chapter describes several methods for the assessment of Chlamydomonas flagellar movements in live cells, demembranated and reactivated cells (cell models), and isolated axonemes. In addition, a simple method is introduced for velocity measurements of microtubule sliding in disintegrating axonemes. We used these methods for isolation and characterization of various mutants deficient in flagellar motility as well as for analysis of the effects of physical and chemical factors on flagellar motility (Kamiya, 2002). Measurements of beat frequency and waveforms demonstrated that outer arm dynein is particularly important for the flagella to beat with a high frequency, whereas inner arm dyneins are important for producing effective waveforms (Brokaw & Kamiya, 1987). Experiments with demembranated cells and axonemes revealed that the two flagellar axonemes on a
1. Methods
single cell are differentially regulated by submicromolar Ca2þ (Kamiya & Witman, 1984); cAMP-dependent protein phosphorylation inhibits flagellar motility (Hasegawa, Hayashi, Asakura, & Kamiya, 1987); reductioneoxidation balance affects flagellar beat frequency (Wakabayashi & King, 2006); and the axonemes of paralyzed-flagella mutants lacking the central pair and radial spokes can beat under some nonphysiological nucleotide conditions (Omoto, Yagi, Kurimoto, & Kamiya, 1996) and display Ca2þ-dependent waveform conversion (Wakabayashi, Yagi, & Kamiya, 1997). Measurements of microtubule sliding velocity showed that the outer arm dynein and inner arm dynein intrinsically differ in sliding velocity (Kurimoto & Kamiya, 1991; Okagaki & Kamiya, 1986), leading to the recognition that the axoneme is equipped with motors operating at different speeds. Although the details of the conditions described below are optimized for Chlamydomonas, most methods may be readily used in cilia and flagella of other organisms after slight modification of the conditions.
1. METHODS 1.1 SWIMMING TRACKS AND AVERAGE SWIMMING VELOCITIES Average swimming velocity of Chlamydomonas cells can be obtained from the cells’ swimming path lengths per a given time. Previously, we manually traced the swimming paths on a transparent sheet overlaid on the monitor screen and measured the length with a scale. Recently, it is easier to track each cell image in a personal computer. For example, an Image J plug-in, MTrack2 (http://valelab.ucsf.edu/ wnstuurman/IJplugins/MTrack2.html) enables automatic tracking (Figure 1). In addition to swimming velocities, swimming tracks provide useful information about
FIGURE 1 (A) Swimming tracks of Chlamydomonas cells taken under a dark-field microscope. Time interval, 0.1 s. Image brightness is increased with time, so that the swimming direction is readily determined. Bar: 0.1 mm. (B) Swimming paths of the cells in (A) auto-tracked by Image J and MTracks2 plug-in.
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the cell’s motility; the tracks are important for assessing cells’ phototactic activity and the tendency of directional change. The pitch of the helical swimming path reflects characteristic features of flagellar beating pattern of a particular cell type.
1.1.1 Swimming velocities and swimming directions 1. Observe and video record the swimming paths of Chlamydomonas cells under a microscope at low magnification (objective: 10). The record must be saved in Audio Video Interleave (AVI) format, uncompressed. Dark-field or phasecontrast microscope is recommended to obtain high-contrast images. Cells often accumulate in the field of observation due to phototaxis, which interferes with the observation of individual cells. In such a case, place a red filter (630 nm) on the light source. However, pay attention to the fact that the swimming velocity varies with illumination; cells swim slightly slower under red light than under white light because of a response to the light conditions (this phenomenon is called the photokinesis). 2. Playback the AVI file using Image J. Binarize the images so that the cells appear in black and background in white. Run MTrack2. Obtain swimming tracks for w50 cells each for 3e5 s. 3. Analyze the resultant tracking data using a spreadsheet program, such as Excel, to obtain the average swimming velocity, standard deviation, and if necessary, other parameters such as the frequency of directional changes in swimming paths. Notes For phototaxis analysis, place a red filter on the light source of the microscope and set a light emission diode (l ¼ 500e525 nm) to stimulate the cells from the side. Measure the angle between the light direction and the swimming path of each cell. The data can be shown as a polar histogram (Figure 2) or a phototactic
FIGURE 2 Polar histograms showing swimming directions of wild-type cells. Bars represent the percentage of cells moving in a particular direction relative to the stimulating light (l ¼ 525 nm) from the right (12 bins of 30 ; n ¼ 50 cells per conditions). Swimming tracks for 1.5 s were examined. (A) Without stimulating light, cells swim in random directions. (B) When stimulating light is illuminated from the right (0 ), most cells show positive phototaxis and swim toward the light source. Modified from Wakabayashi et al. (2011).
1. Methods
index, the average of jcos qj (Okita, Isogai, Hirono, Kamiya, & Yoshimura, 2005; Wakabayashi, Misawa, Mochiji, & Kamiya, 2011).
1.1.2 Estimation of net propulsive force As an application of the above method, we can estimate the net propulsive force of flagellar movement in viscous media from the swimming velocity. The force acting on the moving cell body is proportional to the product of the viscosity (h), cell size, and moving velocity (v); if we approximate the cell body as a sphere of radius r, then the viscous force acting on it is given by 6phrv (Stokes’ formula). Thus it is straightforward to compare the load dependence of net flagellar propulsive force between different strains. Using this method, we found that the outer arm dynein and inner arm dynein greatly differ in the load dependence (Minoura & Kamiya, 1995), and that a particular kind of single-headed inner arm dynein (dynein c) is important for propulsive force production at high viscosity (Yagi et al., 2005). 1. Prepare viscous solution by dissolving 0e16% (w/v) Ficoll (Type 400, Sigma; Mr ¼ 400,000) in appropriate medium or buffer. Other polymers can be also used, but Ficoll produces less local inhomogeneity in solution than other polymers such as methylcellulose and, therefore, is more appropriate (Berg & Turner, 1979). If the actual force value needs to be determined, measure the solution viscosity with an Ostwald or other types of viscometer. 2. Record the tracking of swimming cells in the viscous sample solutions. Care must be taken that the sample consists of cells with similar diameters, since the swimming velocity in viscous media sensitively varies with load, which is proportional to the cell diameter. To prepare cells with similar diameters, synchronize cell cultures on an appropriate lightedark cycle, and measure the motility at a constant phase of the cycle.
1.2 WAVEFORM ANALYSIS High-speed recording is necessary for observation of flagellar waveform. Flagellar beat frequency in wild-type Chlamydomonas is w60 Hz; therefore, one beat cycle can be recorded in w5 frames if a movie is taken at 300 fps. Cameras with this frame rate are available from several manufacturers at relatively low costs. Even some home-use digital still cameras are equipped with a high-speed recording mode. It is important to choose a fast shutter speed to obtain clear images of waveform. For waveform analysis, we must obtain a time series of flagellar images that are in focus for at least one beating cycle. We use uniflagellated cells that rotate within a small area by beating only the single flagellum and retain the flagellar image in focus for a long time. When observation of the two flagella on a single cell is necessary, use cells that are stuck between the cover slip and glass slide or those held with a micropipette. We use the following “uniflagella” method. 1. Produce uniflagellated cells by gently homogenizing the cell culture, or by producing cells with the background of a uniflagella mutation, uni1
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(Huang, Ramanis, Dutcher, & Luck, 1982). In the former case, we must choose by visual inspection an appropriate cell that lacks one of the two flagella while maintaining the other flagellum intact, each time a movie is taken. In the latter case, we must pay attention to the fact that the uni1 mutant retains only one particular flagellum (the trans-flagellum; the flagellum farthest from the eyespot) of the two flagella. Previous studies showed that the trans-flagellum tends to beat at higher frequency than the other, cis-flagellum (Kamiya & Hasegawa, 1987). 2. Take a high-speed movie of a uniflagellated cell rotating under a microscope with a 40 objective (Figure 3(A)).
FIGURE 3 (A) Sequential images of a uni1 cell rotating by beating its single flagellum. Taken at 300 frames/s. (B) The angle between the tangents to the waveform at the base (dotted line) and at point s (at distance s along the flagellar length from the base) represents the “shear angle” (q) at this position. (C) Typical shear curves of wild-type (left) and ida4 (right) flagella. ida4 displays a smaller shear amplitude. The slope of each line gives the curvature at a given position. Flagellar waveforms were manually traced three times and the average of these data was smoothed by the method of Brokaw (1983). Modified from Minoura and Kamiya (1995).
1. Methods
3. Analyze flagellar waveform from sequential images of flagellar beating. For most purposes, a series of flagellar beating images will give enough information even without any analysis. When quantitative data are required, each image must be traced and analyzed. A standard method is to measure the angle (shear angle, q) between the tangent to a small segment at the position s (measured from the base along the flagellum) and the tangent to the basal segment, and plot q against s for the entire length (Brokaw, 1979; Brokaw, Luck, & Huang, 1982) (Figure 3(B) and (C)). Because q is proportional to the sliding displacement between adjacent outer doublet microtubules at point s, the plots produce a single curve that represents the relative microtubule sliding along the flagellar length. Hence, the curve is called a shear curve. From several sets of plots for different beating phases over a complete beat cycle, we can determine the amplitude and velocity of back-and-forth microtubule sliding at each position along the flagellum. The curves also give parameters such as the bend angle and curvature in the principal and reverse bends. Several programs are available that generate sequential images from a movie file, such as iMovie (bundled with Mac OSX) and TMPGEnc (shareware, a free version is also available at http:// www.tmpgenc.net/). 4. Beat frequency is readily obtained from sequential images of flagellar beating. Choose a pair of images showing similar waveforms in n cycles (n ¼ 1e3) of beating strokes, count the number (m) of frames between the two similar waveforms, and calculate (n/m) (frame rate) to yield the beat frequency. For average beat frequency values, use data from 30 to 50 cells. Notes When uniflagellated cells are produced by homogenization of bi-flagellated cells, we must determine whether the flagellum being observed is cis or trans relative to the eyespot. Eyespot is more easily observed with an oil-immersion dark-filed condenser than with a dry condenser; it appears as a bright spot located on the side of a rotating cell.
1.3 BEAT FREQUENCY MEASUREMENTS A convenient method for estimating the average beat frequency in a population of cells has been developed that uses frequency analysis of the cell body vibration (Kamiya, 2000). This method relies on the fact that the Chlamydomonas cell body vibrates back-and-forth in synchrony with flagellar beating. The equipment for this measurement is relatively easily constructed from a photosensor, an A/D converter, and an Fast Fourier Transform (FFT) program run on a PC. The outline of the method is as follows. Images of 10e100 swimming cells in a dark-field microscope are projected on a graded filter, and the transmitted light is detected with the sensor. The graded filter converts the back-and-forth movements of cell images into a light intensity fluctuation (however, we found that this graded filter is practically not necessary, because the brightness of microscope image field is usually not uniform and the light
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CHAPTER 19 Axonemal motility in Chlamydomonas
intensity from the cell images fluctuates even without it). We are using a photodetector composed of an appropriate photodiode (such as Hamamatsu S1223), lownoise operational amplifiers (such as OPA111, Burr Brown, Tucson, AZ, USA), and 1-GU feedback resistors (circuit diagrams are available at the manufacturer’s Web site). The photodetector is attached to a dark-field microscope with an objective of 10 to 20 and a light source of a 100-W halogen lamp. FFT is performed using the SIGVIEW software package (http://www.sigview.com/). This method allows us to measure the average beat frequency within a minute. Unlike the high-speed video method using the uni1 mutant, this method allows measurements of average beat frequencies of the two flagella. In demembranated and reactivated cell models (see below), the two axonemes tend to beat independently, giving rise to two peak frequencies reflecting the two frequencies of the cell body vibration caused by both cis- and trans-flagellar axonemes (Figure 4).
1.4 REACTIVATION OF DEMEMBRANATED CELL MODELS In vitro reactivation of cell movement is a powerful method to study cell motility mechanism. Feasibility of such an experiment was first demonstrated by SzentGyo¨rgyi (1949) who used rabbit skeletal muscle fibers extracted with 50% glycerol. It is an excellent experimental system that allows us to easily change the composition of reactivating solution, which corresponds to the cytoplasm in vivo. In fact, it was used in experiments to establish that muscle contraction is dependent on ATP. This method has been used in various studies on muscle contraction, in order to investigate the effects of ATP concentration, Ca2þ concentration, protein phosphorylation, etc. Hoffman-Berling (1954) applied this method to sperm flagella and showed that flagellar movement is also energized by ATP. He called the extracted cells “cell models.” Glycerol was later replaced by nonionic detergent Triton X-100 by Gibbons and Gibbons (1972) for sea urchin sperm cell models and by Naitoh and Kaneko (1972) for Paramecium cell models. These extracted cells were called “Triton models.” In the case of Chlamydomonas cell models, Nonidet P-40 has been more frequently used than Triton X-100 (Witman, Plummer, & Sander, 1978), although both detergent give similarly good results (our unpublished observation). Currently we use the following protocols based on those used by Witman et al. (1978) for studying axonemal motility. Solutions (named after constituents) • •
• • •
HES: 10 mM Hepes, pH 7.4, 1 mM EGTA, 4% sucrose HMDEKP: 30 mM Hepes, pH 7.4, 5 mM MgSO4, 1 mM dithiothreitol (DTT), 1 mM EGTA, 50 mM K-acetate, 1% polyethylene glycol (Mr ¼ 20,000) (add an ATP-regeneration system consisting of 70 U/mL creatine phosphokinase and 5 mM creatine phosphate for stable reactivation at 105 M, axonemes tend to detach from the cell body because Chlamydomonas has a Ca2þ-dependent mechanism for flagellar autotomy. When observation of cell model movements at higher Ca2þ concentrations is necessary, use the mutant fa1 deficient in the autotomy (Lewin & Burrascano, 1983) (available from the Chlamydomonas Resource Center).
1.5 REACTIVATION OF ISOLATED FLAGELLAR AXONEMES Chlamydomonas flagella and axonemes can be readily isolated by deflagellation with dibucaine and differential centrifugation (Witman et al., 1978). The isolated flagella and axonemes can be used in various experiments, including motility reactivation, SDS-PAGE, immunofluorescence microscopy and electron microscopy. Motility reactivation in isolated axonemes has been used to examine the effect of different concentrations of Ca2þ (Bessen, Fay, & Witman, 1980) and cAMP (Hasegawa et al., 1987). Chlamydomonas axonemes change their beating patterns from an asymmetric, ciliary pattern at 2 min) tends to result in poor reactivation. Spin down the cell bodies by centrifugation at 600 g for 5 min at 4 C. Transfer the supernatant to a new tube (placed on ice), and centrifuge at 1600 g for 2.5 min at 4 C. Transfer the supernatant to a new tube (placed in ice), and subject to centrifugation at 14,000 g for 12 min at 4 C. Discard the supernatant, resuspend the precipitates in 200 mL 0.2% NP-40 in HMDEK. Mix 80 mL of HMDEKP, 10 mL of ATP solution, and 10 mL of axonemes in a 0.5-mL test tube. To examine the effect of Ca2þ, replace the EGTA with one of the Ca2þ buffers shown in Table 1.
1.6 SLIDING DISINTEGRATION OF AXONEMES Axonemes beat through microtubule sliding driven by various kinds of axonemal dyneins. Sliding movements can be directly observed in protease-treated axonemes, Table 1 Composition of Calcium Buffers for Reactivation Experiments Free [Ca2D] (M)* pCa4 pCa5 pCa6 pCa7 pCa8 *
4
0.96 10 1.01 105 1.04 106 1.06 107 1.00 108
At 1 mM ATP.
CaCl2 (M) 3
1.60 10 1.60 104 4.75 103 3.20 103 3.70 104
EGTA (M)
EDTA (M)
0 0 5.00 103 4.80 103 2.30 103
2.00 103 6.50 104 0 0 0
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CHAPTER 19 Axonemal motility in Chlamydomonas
FIGURE 5 Flow chamber for observation of sliding disintegration. A coverslip is held on a glass slide via two pieces of double-sided adhesive tape w80 mm thick. A reservoir for the perfusing solution is made with nail varnish by one of the two open sides. Perfusion is started by putting a drop of solution to the reservoir and placing a piece of filter paper on the other side.
as first shown by Summers and Gibbons (1971). The following protocols for Chlamydomonas axonemes are based on Okagaki and Kamiya (1986) and Kurimoto and Kamiya (1991) who measured the velocity of microtubule sliding during disintegration occurring upon perfusion with ATP and protease (Figure 6). Methods similar to the one outlined below have been used in studies that explored regulatory mechanism of axonemal motility (Habermacher & Sale, 1995; Smith & Sale, 1992). We tend to regard the sliding velocities measured in disintegrating axonemes as reflecting the movement under very little load. However, microtubule sliding in disintegrating axonemes does not reflect the maximal velocity that can be achieved by axonemal microtubules under minimal loads. This is because the sliding velocity increases if the axonemes are more extensively treated with protease (Kikushima, 2009; Yano & Miki-Noumura, 1981), which may well function
FIGURE 6 Sliding disintegration of a wild-type axoneme fragment. ATP, 1 mM. Interval: 1/15 s. Bar, 10 mm. This axoneme fragment is undergoing sliding at w13 mm/s.
1. Methods
to reduce interdoublet friction. Thus we must be careful in interpreting the sliding velocity data; it can be affected by the intrinsic sliding velocity and the force produced by dynein(s), and also by the friction between outer doublets. A higher sliding velocity does not necessarily mean a higher activity of axonemal dyneins. Solutions • • • • • 1.
2. 3. 4.
5. 6. 7. 8. 9. 10.
11.
12.
HMDEKP: 30 mM Hepes, pH 7.4, 5 mM MgSO4, 1 mM DTT, 1 mM EGTA, 50 mM K-acetate, 1% polyethylene glycol (Mr ¼ 20,000) HMDEK: 30 mM Hepes, pH 7.4, 5 mM MgSO4, 1 mM DTT, 1 mM EGTA, 50 mM K-acetate 2% Nonidet P-40 in HMDEKP 1 mM ATP in HMDEKP 2 mg/mL Nagarse (Type XXIV protease, Sigma), 1 mM ATP in HMDEKP Prepare a flow chamber from a glass slide and a coverslip with spacers (pieces of double-sided adhesive tape) (Figure 5). Use nail varnish to produce a reservoir for perfusion solution by one of the two open sides of the cover glass. Prepare flagella from an w200 mL cell culture as described above. Dilute the flagellar suspension with HMDEKP so that the total volume is w1 mL, in a 1.5 mL Eppendorf tube. Produce flagellar fragments by sonication with an appropriate sonicator. We use a Sonifier 250 (Branson) with a microtip and apply sonication for 4 s with the power set to “3” and the duty ratio to 30%. Collect flagellar fragments by centrifugation at 14,000 g for 12 min at 4 C. Carefully remove the supernatant and resuspend the precipitate in w50 mL HMDEKP. Mix 85 mL HMDEKP, 10 mL flagella fragments, and 5 mL 2% NP-40 in a test tube and place it in ice (this sample is referred to as “axoneme fragments”). Fill the flow chamber with 10 mL of axoneme fragments. Perfuse the chamber with 40 mL of ATP solution. Observe the sample with a dark-field microscope capable of visualizing single microtubules. We use an Olympus microscope equipped with an oilimmersion condenser (NA 1.2e1.4), a light source of 100-W mercury arc lamp and a 50 objective with an iris (NA 0.90). Images are recorded with a sensitive video camera. Find a field where many axoneme fragments are attached on the glass surface. Perfuse the sample with the Nagarse solution that contains both protease and ATP. Under optimal conditions, sliding disintegration will occur within w30 s after onset of perfusion. Measure the sliding velocity after tracing the moving microtubule images using Image J and an appropriate plug-in, as outlined above. Notes
1. Elastase can be used in place of “Nagarse.”
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ACKNOWLEDGMENTS We thank Toshiki Yagi (Prefectural University of Hiroshima) and Mikito Owa (University of Tokyo) for helpful advice, and Elina Nakamasu (Tokyo Institute of Technology) for her help in preparing figures. Our studies have been supported by grants-in-aid from Japan Society for Promotion of Sciences (#25113507, #25117506, #25291058, and #26650093 to KW, and #23570189 and #25117521 to RK).
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CHAPTER 19 Axonemal motility in Chlamydomonas
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