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Melanophores for Microtubule Dynamics and Motility Assays
CHAPTER 21
Melanophores for Microtubule Dynamics and Motility Assays Kazuho Ikeda, Irina Semenova, Olga Zhapparova, and Vladimir Rodionov Department of Cell Biology, R. D. Berlin Center for Cell Analysis and Modeling, University of Connecticut Health Center, Farmington, Connecticut 06032-1507
Abstract I. Introduction II. Experimental Procedures A. Cultivation of Melanophores B. Fluorescent Labeling of Tubulin C. Microinjection and Microsurgery D. Live Cell Imaging and Data Analysis E. Quantification of Aggregation and Dispersion of Pigment Granules III. Discussion
Acknowledgments
References
Abstract Microtubules (MTs) are cytoskeletal structures essential for cell division, locomotion, intracellular transport, and spatial organization of the cytoplasm. In most interphase cells, MTs are organized into a polarized radial array with minus-ends clustered at the centrosome and plus-ends extended to the cell periphery. This array directs transport of organelles driven by MT-based motor proteins that specifically move either to plus- or to minus-ends. Along with using MTs as tracks for cargo, motor proteins can organize MTs into a radial array in the absence of the centrosome. Transport of organelles and motor-dependent radial organization of MTs require MT dynamics, continuous addi tion and loss of tubulin subunits at minus- and plus-ends. A unique experimental system for studying the role of MT dynamics in these processes is the melanophore, METHODS IN CELL BIOLOGY, VOL. 97 Copyright � 2010 Elsevier Inc. All rights reserved.
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which provides a useful tool for imaging of both dynamic MTs and moving membrane organelles. Melanophores are filled with pigment granules that are synchronously transported by motor proteins in response to hormonal stimuli. The flat shape of the cell and the radial organization of MTs facilitate imaging of dynamic MT plusends and monitoring of their interaction with membrane organelles. Microsurgically produced cytoplasmic fragments of melanophores are used to study the centrosomeindependent rearrangement of MTs into a radial array. Here we describe the experi mental approaches to study the role of MT dynamics in intracellular transport and centrosome-independent MT organization in melanophores. We focus on the prepara tion of cell cultures, microsurgery and microinjection, fluorescence labeling, and live imaging of MTs.
I. Introduction Microtubules (MTs) are highly dynamic structures that continuously grow and shorten by addition and loss of tubulin subunits (Cassimeris et al., 1987; Desai and Mitchison, 1997; Howard and Hyman, 2009). Polarized radial array of MTs growing from the centrosome defines spatial organization of the cytoplasm and supports transport of organelles driven by MT-based motor proteins (Cole and LippincottSchwartz, 1995; Gross, 2004; Lane and Allan, 1998; Welte, 2004). In the absence of the centrosome, motor proteins establish radial organization of MTs (Borisy and Rodionov, 1999; Compton, 1998; Hyman and Karsenti, 1996; Sharp et al., 2000). MT dynamics are critical for both transport of membrane organelles and motordependent organization of a radial MT array. The major experimental approach to study the role of MT dynamics in these processes is live imaging of cells with fluorescently labeled MTs. The cells used for these assays should be suitable for both the observation of dynamic MTs and the moving organelles; these features are combined in melanophores—pigment cells of lower vertebrates. The main function of melanophores is fast and synchronous redistribution of numerous pigment granules, which aggregate at the cell center or disperse uniformly throughout the cytoplasm in response to hormones (Nascimento et al., 2003). The rapid and highly coordinated redistribution of pigment granules changes the color of animal skin and helps to elude predators (Nascimento et al., 2003). Pigment granules are transported along MTs by motor proteins—kinesins move them to the MT plusends during dispersion and cytoplasmic dynein to the minus-ends during aggregation (Nilsson and Wallin, 1997; Rodionov et al., 1991; Tuma et al., 1998). We have used Xenopus melanophores to test whether MT dynamics facilitates the interaction of pigment granules with MTs (Lomakin et al., 2009). Live cell imaging of melanophores with fluorescently labeled MTs revealed that the initiation of minus-end transport involved capturing of pigment granules by the growing MT plus-ends. We also found that stabilization of MTs with taxol dramatically inhibited pigment aggregation. These data demonstrate that dynamic MTs are required for the initiation of minus-end transport of pigment granules (Lomakin et al., 2009).
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MTs in melanophores are organized into a polarized radial array as a result of nucleation and anchoring at the centrosome. Remarkably, cytoplasmic fragments of melanophores form a radial MT array in the absence of the centrosome after the activation of dynein motors bound to pigment granules (McNiven et al., 1984; Rodionov and Borisy, 1997). Radial organization of MTs might result from minus end-directed transport of preassembled MTs by dynein motors, seen in mitotic cell extracts (Hyman and Karsenti, 1996; Verde et al., 1991) or from disassembly and reassembly of MTs. To determine the mechanism of radial MT organization, we injected cells with fluorescent tubulin taken at a low concentration (Waterman-Storer and Salmon, 1998). The labeled tubulin unevenly incorporated along the MTs and the resulting bright speckles allowed us to follow the position of reference points on individual MTs and determine whether they were moved in the cytoplasm by motor proteins and discriminate between the two mechanisms of MT reorganization. We found that fluorescent speckles did not move after the activation of dynein motors bound to pigment granules (Vorobjev et al., 2001), which indicated that MTs remained immotile. These data demonstrate that in centrosome-free fragments of melanophores, radial organization of MTs results from disassembly and reassembly of MTs, rather than motor-driven movement in the cytoplasm. Therefore, MT dynamics are important for the formation of a radial array in the absence of the centrosome. Here we report the methods for studying the role of dynamic MTs in melanophores and describe the details of cell culture preparation, microsurgery and microinjection, fluorescence labeling, and live imaging of MTs.
II. Experimental Procedures A. Cultivation of Melanophores Melanophores are obtained either from fish (Gymnocorymbus ternetzi) scales or from frog (Xenopus laevis) tadpoles. Primary cultures of fish melanophores are prepared from black tetra scales prior to each experiment. Cells are separated from the scales by collagenase treatment, plated on carbon-coated coverslips, and incubated overnight in tissue culture medium. Permanent cell lines of X. laevis melanophores are obtained according to the modified method of Daniolos et al. (1990) that includes the following steps: trituration of tadpoles, purification of melanophores by Percoll density centrifugation, and generation of stable cell lines by limiting dilution. Xenopus melanophores become spontaneously immortalized during the first 2–3 weeks of growth in primary culture, and therefore melanophore cell lines can be continually maintained in culture. Xenopus melanophores containing pigment do not survive freezing, and therefore cells should be depleted of melanin by treatment with PTU (N-phenylthiocarbamide) that inhibits tyrosinase, an enzyme responsible for melanin synthesis.
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1. Primary Culture of Fish Melanophores 1. Remove 10–20 scales from black tetra (G. ternetzi) under a dissection microscope using Dumond #5 or #7 tweezers. 2. Place scales in fish Ringer’s solution (5 mM Tris–HCl, 103 mM NaCl, 1.8 mM KCl, 0.8 mM NaHCO3, 2 mM CaCl2, pH 7.2) supplemented with 5 mg/ml BSA and 1 mg/ml collagenase (Worthington, type 3). 3. Incubate scales at 30°C for about 1 h. 4. Under a dissection microscope, hold down scales (pin them) using tweezers and detach cells by pipetting using a 2–20 µl automatic pipettor. 5. Wash off the excess collagenase by transferring individual cells with a pipettor from one 35 mm cell culture dish containing Ringer’s solution into another dish; repeat three to five times. To prevent attachment of cells to the bottom of the dish, treat dishes with 1% BSA solution in PBS for 10–15 min, and rinse with water prior to use. 6. Place cell suspension onto 22 22 mm carbon-coated coverslips. Carbon coating is applied using the DV-502 evaporator (Denton Vacuum, Inc.) and coverslips are mounted with silicon vacuum grease over a hole drilled in a 35 mm cell culture dish. Instead of carbon, coverslips can be coated with laminin (40 µg/ml) or poly-L-lysine (0.1 mg/ml), although melanophores spread more readily on carbon. 7. Add 3 ml of tissue culture medium into each dish (DMEM supplemented with 20 mM HEPES, 20% fetal bovine serum, 200 µg/ml streptomycin, and 200 units/ml penicillin) and incubate overnight at 30°C for the complete spreading of melanophores.
2. Generation of Immortalized Cell Lines of X. laevis Melanophores 1. Rinse 20 X. laevis tadpoles (stage 30–35; Nieuwkoop and Faber, 1967) three times with amphibian Ringer’s solution (9 mM HEPES, 115 mM NaCl, 3 mM KCl, 2 mM CaCl2, pH 7.3). 2. Rinse with 70% ethanol for 5 s. 3. Rinse three times with amphibian Ringer’s solution. 4. Grind tadpoles in 10 ml of Xenopus tissue culture medium (XTCM; 0.7 L-15 medium (Sigma-Aldrich, St. Louis, MO), supplemented with 10% heatinactivated fetal bovine serum (Gibco, Invitrogen Corp., Carlsbad, CA), 200 units/ml penicillin, 200 µg/ml streptomycin, and 5 µm insulin, pH 7.3–7.4). 5. Plate the resulting suspension of cells onto three 60 mm cell culture dishes. 6. Incubate at 27°C for 1 week. 7. Rinse with XTCM to remove unattached cells. 8. Cultivate for 1–2 months changing XTCM twice a week until black colonies of melanophores appear. 9. Detach cells by trypsinization (briefly incubate in trypsin solution containing 2 mg/ml trypsin, 0.2 mg/ml EDTA in PBS) and collect by centrifugation. 10. Purify melanophores by centrifugation through Percoll cushion. Resuspend cell pellet in 20% Percoll solution, load onto 12 ml of 33% Percoll solution (both
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Percoll solutions prepared on XTCM), and pellet cells by centrifugation at 500 g at for 5 min at a room temperature. 11. Resuspend the pellet, which contains 80–90% melanophores, in 5 ml of XTCM, plate into a 25 cm2 flask, and cultivate at 27°C. Change the medium twice a week. 12. When melanophores become confluent, repeat steps 9–11 until unpigmented cells are completely removed. Usually three rounds of Percoll density gradient purification yield pure primary culture of melanophores. 13. Prepare feeder layer of Xenopus fibroblasts by plating them into a 48-well plate and cultivate until cells form a monolayer. Treat cells with 10 µg/ml mitomycin C (M0440, Sigma-Aldrich) for 3 h at 27°C to arrest cell growth. Wash three times with XTCM to remove mitomycin C. 14. Detach melanophores by trypsinization, resuspend in 10 ml of XTCM, count using a hemocytometer (Bright-line Counting Chamber, #3100 Haussen Scientific, Horsham, PA), and dilute the suspension to concentration of one cell per 400 µl. 15. Add 200 µl of cell suspension to each well of the 48-well plate containing a monolayer of Xenopus fibroblasts (step 13). 16. Cultivate for about 1 month; colonies of melanophores will appear in the wells. Be sure that wells containing two or more colonies are not used for further steps. Change the medium twice a week. 17. Select colonies of cells that grow fast and respond well to hormones by completely aggregating or dispersing pigment granules within 20 min after the treatment with 10 nM melatonin or 10 nM melanocyte stimulating hormone (MSH), respectively. 18. Grow cells in the selected wells until they form a monolayer, detach by trypsinization, and transfer into a 24-well plate. 19. Repeat step 18, and replate cells consecutively into 12- and 6-well plates, and, finally, onto the 25 cm2 flask. Always maintain melanophores in a semi-confluent culture since at low density cells stop dividing and responding to hormones. 20. If required, freeze melanophores as follows. Incubate cells in the XTCM supplemented with 1 mM PTU (P7629, Sigma) for at least 3 weeks to deplete melanophores of melanin. Trypsinize cells and freeze cell suspension in 20% DMSO in XTCM. After thawing in PTU-free XTCM melanophores produce pigment within 5–6 days.
B. Fluorescent Labeling of Tubulin Purified tubulin is obtained from porcine brain by a standard procedure involving two cycles of MT temperature-dependent assembly-disassembly (Borisy et al., 1975). MT-associated proteins (MAPs) are removed by an additional cycle of disassemblyreassembly of MTs in a high-salt buffer (HS buffer). Conjugation with Cy3 involves incubation of assembled MTs with the dye followed by the two cycles of MT assembly-disassembly and purification of labeled MTs by centrifugation through a glycerol cushion.
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1. Experimental Procedure 1. Thaw 1 ml aliquot of twice cycled tubulin (8–10 mg/ml) in PM buffer (0.1 M PIPES, 1.0 mM MgCl2, pH 6.9) in water bath at 37°C. 2. Add guanosine triphosphatase (GTP) to a final concentration of 1 mM. 3. Polymerize MTs by incubating at 37°C for 10 min. The solution should become viscous and turbid. 4. Pellet MTs at 100,000 xg (50,000 rpm, Beckman TLA 100.3 rotor) at 37°C for 5 min; discard the supernatant. 5. Measure MT pellet volume and resuspend it in equal volume of HS buffer (0.5 M PIPES, 2 mM MgCl2, 1 mM EGTA, pH 6.9). 6. Add DMSO to a final concentration 10% and GTP to 1 mM. 7. Polymerize MTs and pellet them by centrifugation as described in steps 2–3. At this step, the most MAPs are removed. 8. Resuspend the MT pellet in a twofold excess volume of cold PEM buffer (0.1 M PIPES, 5 mM EGTA, 2 mM MgCl2, pH 6.9). 9. Depolymerize MTs by incubating on ice for 10 min. 10. Add DMSO to a final concentration of 10% and GTP to 1 mM. 11. Polymerize MTs by incubating at 37°C for 10 min. 12. Dissolve one vial of commercial aliquot of Cy3 bis-reactive dye (GE Healthcare, PA23000) in 20 µl of anhydrous DMSO. Each commercial vial of Cy3 dye is sufficient for conjugation with 8–10 mg of tubulin. 13. Add the Cy3 solution to polymerized MTs and vortex immediately. Incubate at 37°C for 30 min. 14. Pellet MTs at 100,000 xg at 37°C for 5 min; discard the supernatant. 15. Resuspend the MT pellet in a fivefold excess volume of cold PEM buffer. 16. Add GTP to a final concentration of 1 mM. 17. Depolymerize MTs by incubating on ice for 10 min. 18. Centrifuge tubulin solution at 100,000g at 4°C for 5 min. Transfer the supernatant into a new cold tube. 19. Measure the volume of supernatant and adjust DMSO and GTP concentrations to 10% and 1 mM, respectively. 20. Polymerize and centrifuge MTs as in steps 2–3. 21. Repeat steps 14–18. 22. Polymerize MTs by incubating at 37°C for 10 min. 23. Load MTs onto 900 µl warm 33% glycerol (37°C) cushion prepared on PEM buffer containing 1 mM GTP. Pellet MTs by centrifugation in TLS55 swinging bucket rotor (Beckman, Fullerton, CA; #343778 tubes) at 135,000g (40,000 rpm) at 37°C for 20 min. Aspirate the supernatant and glycerol cushion and resuspend MT pellet in 200 µl of PEM buffer with 1 mM GTP. 24. Depolymerize MTs on ice for 10 min. Centrifuge tubulin solution at 100,000g at 4°C for 5 min. Transfer the supernatant into a new cold tube. The concentration of tubulin solution will be about 6–8 mg/ml. 25. Freeze 10 µl aliquots of supernatant containing Cy3-labeled tubulin and store in liquid nitrogen.
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C. Microinjection and Microsurgery For fluorescent labeling of cytoplasmic MTs in melanophores, cells are injected with Cy3-labeled tubulin. Immediately before microinjection, Cy3-labeled tubulin is clarified by high-speed centrifugation to avoid clogging of microneedles. Injected cells are incubated for about 1 h to allow for the incorporation of Cy3-labeled tubulin into MTs. To produce fluorescent speckles along MTs, cells are injected with Cy3-tubulin solution at a low (0.5 mg/ml) needle concentration. Centrosome-free fragments of melanophores with fluorescently labeled MTs are obtained by microsurgical dissection of the microinjected cells with a sharp glass microneedle.
1. Microinjection 1. Plate cells onto photo-etched carbon-coated coverslips (24 24 mm; Bellco Biotechnology, Vineland, NJ) mounted over a hole drilled in a 35 mm cell culture dish; photo-etched coverslips have grids that enable relocation of the microinjected cells. 2. Thaw a 10 µl aliquot of Cy3-labeled tubulin and clarify by centrifugation at 135,000g (40,000 rpm) for 5 min at 4°C. 3. Transfer 1 µl of Cy3-labeled tubulin solution into a glass microneedle with tip diameter of about 0.1 µm produced using micropipette puller, such as Narishige PB-7 (Narishige, East Meadow, NY), from thin-wall borosilicate glass capillaries with filament (an outer capillary diameter is 1.5 mm and inner diameter—1.12 mm; WPI Inc., Sarasota, FL). 4. Place the glass microneedle into a micromanipulator with pipette holder and inject cells with Cy3-labeled tubulin under a pressure of 40 hPa applied by a microinjector (model 5242, Eppendorf, Hauppauge, NY). 5. Incubate for at least 1 h at 27°C for Xenopus or at 30°C for fish melanophores to allow the incorporation of labeled tubulin into MTs.
2. Microsurgery 1. Plate fish melanophores on photo-etched carbon-coated coverslips (24 24 mm; Bellco Biotechnology) and grow in tissue culture medium overnight. 2. Prepare microneedles as described above. 3. For the dissection of a cytoplasmic fragment, place the tip of an empty glass microneedle close to the cell surface; the fragment is separated by capillary suction. Try to select cells with processes and perform dissection at the base of the process.
D. Live Cell Imaging and Data Analysis Cells injected with Cy3-tubulin are observed on an inverted fluorescence microscope. Images are acquired with a sensitive cooled charge-coupled device (CCD) camera. To prevent photodamage of cells caused by light exposure during the observation, the
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oxygen scavenger Oxyrase is added to the medium and mineral oil is overlaid on the culture medium to retard gas exchange. 1. Take a dish with microinjected cells and add Oxyrase (Oxyrase, Ashland, OH) and its substrate lactic acid (L4263 Sigma) to final concentrations of 2% and 20 mM, respectively; check the pH and adjust, if necessary, to 7.3–7.4. 2. Overlay 2 ml of mineral oil (purchased in a local drug store) to retard gas exchange and incubate for 10 min at a room temperature for oxygen depletion. 3. Acquire fluorescence images of cells with an inverted fluorescence microscope, such as a Diaphot 300 (Nikon, Melville, NY) equipped with a Plan 100 1.25 numerical aperture objective lens and 100 W mercury arc lamp, and a narrow band rhodamine filter set, which is compatible with the excitation of Cy3 fluorescence. For image acquisition, use a CCD camera, such as Andor iXon EM-CCD sensor (Andor, South Windsor, CT), Photometrics series 300 cooled CCD camera (Photometrics, Tucson, AZ), or other cameras which have high quantum efficiency at the emission wavelengths of Cy3 and rhodamine. Collect time series of images of labeled MTs with the exposure time 100–500 ms and 2–3 s intervals between the frames using Metamorph software (MDC, Downingtown, PA). 4. Determine parameters of MT dynamics by tracking the positions of MT plus-ends and recording X, Y coordinates using Metamorph. Analyze the data with an appropriate computer software such as the program based on the Multiscale Trend Analysis algorithm (Zaliapin et al., 2005) to determine the following parameters: the duration, length and velocity of MT growth and shortening, and the frequency of catastrophes (transition from growth or pause to shortening) and rescues (the transition from shortening or pause to growth). Alternatively, determine these parameters manually by decomposing life history (distance vs time) plots of MTs into periods of growth and shortening and pauses assuming the changes of MT length over 0.5 µm as growth or shortening events, and others changes as pauses. The parameters of MT dynamics in control and Taxol-treated melanophores with stabilized MTs are represented in Table I.
Table I Parameters of MT Dynamic Instability in Control and Taxol-treated Xenopus Melanophores
Growth distance (µm) Growth rate (µm/s) Shortening distance (µm) Shortening rate (µm/s) Catastrophe frequency (s–1) Rescue frequency (s–1) Duration of pauses (s) Number of analyzed MTs Number of analyzed cells
Control
Taxol
2.96 ± 0.16 0.17 ± 0.03 3.17 ± 0.18 0.18 ± 0.04 0.026 ± 0.001 0.029 ± 0.001 3.99 ± 0.31 30 5
0.23 ± 0.01 0.06 ± 0.01 0.23 ± 0.01 0.06 ± 0.001 0.026 ± 0.002 0.042 ± 0.002 11.51 ± 0.57 30 6
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E. Quantification of Aggregation and Dispersion of Pigment Granules Responses of melanophores to hormones that induce aggregation or dispersion of pigment granules are quantified by estimating the fractions of cells with aggregated, partially aggregated, or dispersed pigment granules at a fixed time interval after induction. Cells in each category are manually counted using phase contrast micro scopy. Alternatively, the kinetics of aggregation or dispersion of pigment granules are determined by measuring pixel gray values within cell outlines using Metamorph region measurement tool. This parameter reflects the degree of homogenous pigment distribution throughout the cytoplasm.
1. Quantification of Cells with Aggregated, Partially Aggregated, or Dispersed Pigment Granules 1. Induce aggregation or dispersion of pigment granules. In the case of fish melanophores, treat cells with adrenaline taken at a final concentration 500 nM to induce aggregation. Induce dispersion by the addition of 5 mM caffeine. To induce pigment aggregation or dispersion in Xenopus melanophores, place cells in a serumfree medium, incubate for 1 h at 27°C, add melatonin or MSH, respectively, to the final concentration 10 nM. 2. Fix cells with 4% formaldehyde solution 5 or 10 min after the application of adrenaline or caffeine (fish melanophores) or 10 or 20 min after the stimulation with melatonin or MSH (Xenopus melanophores). 3. Count the cells with aggregated, partially aggregated, dispersed pigment granules by counting cells in each category using phase contrast microscopy. Completely aggregated cells have a compact aggregate of pigment granules in the center and no granules at the periphery. Fully dispersed cells contain granules randomly distributed throughout the cytoplasm. Partially aggregated cells have no granules at the margins and less compact pigment aggregate in the center.
2. Quantification of Kinetics of Aggregation and Dispersion of Pigment Granules in Xenopus Melanophores 1. Induce aggregation or dispersion of pigment granules in fish or Xenopus melanophores as described above. 2. Acquire time series of bright-field images of melanophores with 10 s time intervals using Metamorph time-lapse acquisition mode. 3. Measure the gray levels as integrated pixel values within cell outlines in each of the acquired images in the time-lapse series using Metamorph region measurement tool. The values in the fully dispersed state are taken as 100%. Percentage of gray levels is calculated for each image using the following equation: A ¼ ðIb It Þ=ðIb Id Þ100 where Ib is averaged background levels measured outside cell outlines, It is integrated pixel value within a cell outline at a given moment t, and Id is integrated pixel value
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within a cell outline in the fully dispersed state. Percentages of gray levels for each time point are averaged across the recorded cells and plotted as a function of time (Lomakin et al., 2009).
III. Discussion We have described the experimental approaches used to study the role of MT dynamics in intracellular transport and radial organization of MTs in melanophores. Fish or Xenopus melanophores provide a unique experimental system for imaging both dynamic MTs and moving membrane organelles. Dynamic MT plus-ends are easily followed in large and flat lamellae of melanophores. Synchronous movement of pigment granules driven by motor proteins in response to distinct hormonal stimuli can be observed using conventional light microscopy (Fig. 1). Large and flat fish Frog melanophores
Aggregated pigment
Dispersed pigment
Fish melanophore
Fig. 1 Phase contrast images of melanophores with dispersed or aggregated pigment granules. Fish (G. ternetzi, left images) and frog (X. laevis, right images) melanophores with dispersed (top) or aggregated pigment granules (bottom). Bar, 25 µm.
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melanophores are particularly good for micromanipulation, including microsurgical dissection of cytoplasmic fragments. However, they cannot be grown in large quan tities, and each experiment requires a laborious procedure of cell isolation. Therefore immortalized Xenopus melanophores are recommended for the experiments that do not involve microsurgery. Another advantage of Xenopus cells is that they are easy to transfect and are suitable for various biochemical experiments. For live cell imaging of MTs, melanophores are microinjected with fluorescently labeled tubulin. An alternative way of MT labeling involves the expression of tubulin fused with a fluorescent protein. This method is less laborious but involves transfec tion that is inefficient in cells grown in primary culture, such as fish melanophores.
(A)
0
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(B)
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Fig. 2 MT dynamics are required for aggregation of pigment granules in Xenopus melanophores. (A) Time series of images of fluorescently labeled MTs during pigment granule aggregation. The arrow and arrowhead indicate a growing MT tip and a pigment granule, respectively. Numbers indicate time in seconds. Bar, 2 µm. (B) Stabilization of MTs reduces pigment granule transport. Responses to aggregation (melatonin, 10 min) or dispersion (MSH, 15 min) in control melanophores or melanophores treated with taxol (1 µM) are quantified. The data are expressed as the percentages of cells with aggregated (white bars), partially dispersed (gray bars), or dispersed (black bars) pigment granules. (C) Kinetics of pigment granule aggregation (left) or dispersion (right) in control (white squares) or taxol-treated (black squares) melanophores.
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Besides, the level of exogenous tubulin expression varies from cell to cell being sometimes insufficient for real-time observation, or oppositely, excessive expression generates high background. Therefore, microinjection is a preferable approach for MT labeling in melanophores. The amount of fluorescent tubulin introduced into the cytoplasm by microinjection can be adjusted according to the aim of an experiment. High tubulin concentration results in even labeling of the entire MT and enables observation of dynamic plus-ends, whereas low concentration generates fluorescent speckles that allow studying MT movement and distinguishing between two types of translocation—growth and shortening at the ends or transport of the entire MT (Vorobjev et al., 2001). The described methods were used to define the role of MT dynamics in intracellular transport and in MT organization, driven by motor proteins. We hypothesized that constantly growing and shortening plus-ends searched the cytoplasm for cargo and that such dynamic behavior increased the probability of organelle capturing. Live cell imaging of melanophores stimulated to aggregate pigment granules revealed the events of granule capturing by the growing plus-ends leading to the initiation of transport (Fig. 2). We further confirmed the role of MT dynamics in this process by treating cells with a MT-stabilizing agent and demonstrated that aggregation of pigment granules was inhibited under these conditions (Lomakin et al., 2009). In the absence of the centrosome MTs are organized by motor proteins (Borisy and Rodionov, 1999; Compton, 1998; Hyman and Karsenti, 1996; Sharp et al., 2000). We suggested that MT dynamics were important for this organization. The rearrangement could result either from the transport of previously assembled MTs by motor proteins or from the nucleation of new MTs and their rapid reorganization due to active growth and shortening. Imaging of MTs labeled with fluorescent speckles (Fig. 3) revealed
(B)
After adrenaline
Before adrenaline
(A)
Fig. 3
Organization of a polarized radial array of MTs in cytoplasmic fragments of fish melanophores. (A) Fluorescence images of MTs in a centrosome-free fragment of fish melanophore before (top) and after (bottom) stimulation of aggregation with adrenaline. (B) Fluorescence image of MTs with speckles at low magnification (left) and time sequences of speckles on MTs in the regions indicated on the left panel (right). Numbers indicate the time in seconds. Scale bars, 10 µm (left) and 2 µm (right).
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that MTs were not transported in the cytoplasm but were nucleated de novo, formed small local asters, which merged into a single array as a result of MT growth and shortening (Rodionov and Borisy, 1997). Therefore, live cell imaging of MTs in melanophores elucidated the role of dynamics in intracellular transport and centrosome-independent organization of MTs.
Acknowledgments This work was supported by NIH grant GM62290 to V.I.R.
References Borisy, G. G., Marcum, J. M., Olmsted, J. B., Murphy, D. B., and Johnson, K. A. (1975). Purification of tubulin and associated high molecular weight proteins from porcine brain and characterization of microtubule assembly in vitro. Ann. N. Y. Acad. Sci. 253, 107–132. Borisy, G. G., and Rodionov, V. I. (1999). Lessons from the melanophore. FASEB J. 13(Suppl. 2), S221–S224. Cassimeris, L. U., Walker, R. A., Pryer, N. K., and Salmon, E. D. (1987). Dynamic instability of micro tubules. Bioessays 7, 149–154. Cole, N. B., and Lippincott-Schwartz, J. (1995). Organization of organelles and membrane traffic by microtubules. Curr. Opin. Cell Biol. 7, 55–64. Compton, D. A. (1998). Focusing on spindle poles. J. Cell Sci. 111(Pt 11), 1477–1481. Daniolos, A., Lerner, A. B., and Lerner, M. R. (1990). Action of light on frog pigment cells in culture. Pigment Cell Res. 3, 38–43. Desai, A., and Mitchison, T. J. (1997). Microtubule polymerization dynamics. Annu. Rev. Cell Dev. Biol. 13, 83–117. Gross, S. P. (2004). Hither and yon: A review of bi-directional microtubule-based transport. Phys. Biol. 1, R1–R11. Howard, J., and Hyman, A. A. (2009). Growth, fluctuation and switching at microtubule plus ends. Nat. Rev. Mol. Cell Biol. 10, 569–574. Hyman, A. A., and Karsenti, E. (1996). Morphogenetic properties of microtubules and mitotic spindle assembly. Cell 84, 401–410. Lane, J., and Allan, V. (1998). Microtubule-based membrane movement. Biochim. Biophys. Acta 1376, 27–55. Lomakin, A. J., Semenova, I., Zaliapin, I., Kraikivski, P., Nadezhdina, E., Slepchenko, B. M., Akhmanova, A., and Rodionov, V. (2009). CLIP-170-dependent capture of membrane organelles by microtubules initiates minus-end directed transport. Dev. Cell 17, 323–333. McNiven, M. A., Wang, M., and Porter, K. R. (1984). Microtubule polarity and the direction of pigment transport reverse simultaneously in surgically severed melanophore arms. Cell 37, 753–765. Nascimento, A. A., Roland, J. T., and Gelfand, V. I. (2003). Pigment cells: A model for the study of organelle transport. Annu. Rev. Cell Dev. Biol. 19, 469–491. Nieuwkoop, P. D., and Faber, J. (1967). “A Normal Table of Xenopus laevis (Daudin).” North Holland Publishing Co, Amsterdam. Nilsson, H., and Wallin, M. (1997). Evidence for several roles of dynein in pigment transport in melano phores. Cell Motil. Cytoskeleton 38, 397–409. Rodionov, V. I., and Borisy, G. G. (1997). Self-centring activity of cytoplasm. Nature 386, 170–173. Rodionov, V. I., Gyoeva, F. K., and Gelfand, V. I. (1991). Kinesin is responsible for centrifugal movement of pigment granules in melanophores. Proc. Natl. Acad. Sci. U. S.A. 88, 4956–4960. Sharp, D. J., Rogers, G. C., and Scholey, J. M. (2000). Roles of motor proteins in building microtubule-based structures: A basic principle of cellular design. Biochim. Biophys. Acta 1496, 128–141.
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Melanophores for Microtubule Dynamics and Motility Assays Kazuho Ikeda, Irina Semenova, Olga Zhapparova, and Vladimir Rodionov Department of Cell Biology, R. D. Berlin Center for Cell Analysis and Modeling, University of Connecticut Health Center, Farmington, Connecticut 06032-1507
Abstract I. Introduction II. Experimental Procedures A. Cultivation of Melanophores B. Fluorescent Labeling of Tubulin C. Microinjection and Microsurgery D. Live Cell Imaging and Data Analysis E. Quantification of Aggregation and Dispersion of Pigment Granules III. Discussion
Acknowledgments
References
Abstract Microtubules (MTs) are cytoskeletal structures essential for cell division, locomotion, intracellular transport, and spatial organization of the cytoplasm. In most interphase cells, MTs are organized into a polarized radial array with minus-ends clustered at the centrosome and plus-ends extended to the cell periphery. This array directs transport of organelles driven by MT-based motor proteins that specifically move either to plus- or to minus-ends. Along with using MTs as tracks for cargo, motor proteins can organize MTs into a radial array in the absence of the centrosome. Transport of organelles and motor-dependent radial organization of MTs require MT dynamics, continuous addi tion and loss of tubulin subunits at minus- and plus-ends. A unique experimental system for studying the role of MT dynamics in these processes is the melanophore, METHODS IN CELL BIOLOGY, VOL. 97 Copyright � 2010 Elsevier Inc. All rights reserved.
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which provides a useful tool for imaging of both dynamic MTs and moving membrane organelles. Melanophores are filled with pigment granules that are synchronously transported by motor proteins in response to hormonal stimuli. The flat shape of the cell and the radial organization of MTs facilitate imaging of dynamic MT plusends and monitoring of their interaction with membrane organelles. Microsurgically produced cytoplasmic fragments of melanophores are used to study the centrosomeindependent rearrangement of MTs into a radial array. Here we describe the experi mental approaches to study the role of MT dynamics in intracellular transport and centrosome-independent MT organization in melanophores. We focus on the prepara tion of cell cultures, microsurgery and microinjection, fluorescence labeling, and live imaging of MTs.
I. Introduction Microtubules (MTs) are highly dynamic structures that continuously grow and shorten by addition and loss of tubulin subunits (Cassimeris et al., 1987; Desai and Mitchison, 1997; Howard and Hyman, 2009). Polarized radial array of MTs growing from the centrosome defines spatial organization of the cytoplasm and supports transport of organelles driven by MT-based motor proteins (Cole and LippincottSchwartz, 1995; Gross, 2004; Lane and Allan, 1998; Welte, 2004). In the absence of the centrosome, motor proteins establish radial organization of MTs (Borisy and Rodionov, 1999; Compton, 1998; Hyman and Karsenti, 1996; Sharp et al., 2000). MT dynamics are critical for both transport of membrane organelles and motordependent organization of a radial MT array. The major experimental approach to study the role of MT dynamics in these processes is live imaging of cells with fluorescently labeled MTs. The cells used for these assays should be suitable for both the observation of dynamic MTs and the moving organelles; these features are combined in melanophores—pigment cells of lower vertebrates. The main function of melanophores is fast and synchronous redistribution of numerous pigment granules, which aggregate at the cell center or disperse uniformly throughout the cytoplasm in response to hormones (Nascimento et al., 2003). The rapid and highly coordinated redistribution of pigment granules changes the color of animal skin and helps to elude predators (Nascimento et al., 2003). Pigment granules are transported along MTs by motor proteins—kinesins move them to the MT plusends during dispersion and cytoplasmic dynein to the minus-ends during aggregation (Nilsson and Wallin, 1997; Rodionov et al., 1991; Tuma et al., 1998). We have used Xenopus melanophores to test whether MT dynamics facilitates the interaction of pigment granules with MTs (Lomakin et al., 2009). Live cell imaging of melanophores with fluorescently labeled MTs revealed that the initiation of minus-end transport involved capturing of pigment granules by the growing MT plus-ends. We also found that stabilization of MTs with taxol dramatically inhibited pigment aggregation. These data demonstrate that dynamic MTs are required for the initiation of minus-end transport of pigment granules (Lomakin et al., 2009).
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MTs in melanophores are organized into a polarized radial array as a result of nucleation and anchoring at the centrosome. Remarkably, cytoplasmic fragments of melanophores form a radial MT array in the absence of the centrosome after the activation of dynein motors bound to pigment granules (McNiven et al., 1984; Rodionov and Borisy, 1997). Radial organization of MTs might result from minus end-directed transport of preassembled MTs by dynein motors, seen in mitotic cell extracts (Hyman and Karsenti, 1996; Verde et al., 1991) or from disassembly and reassembly of MTs. To determine the mechanism of radial MT organization, we injected cells with fluorescent tubulin taken at a low concentration (Waterman-Storer and Salmon, 1998). The labeled tubulin unevenly incorporated along the MTs and the resulting bright speckles allowed us to follow the position of reference points on individual MTs and determine whether they were moved in the cytoplasm by motor proteins and discriminate between the two mechanisms of MT reorganization. We found that fluorescent speckles did not move after the activation of dynein motors bound to pigment granules (Vorobjev et al., 2001), which indicated that MTs remained immotile. These data demonstrate that in centrosome-free fragments of melanophores, radial organization of MTs results from disassembly and reassembly of MTs, rather than motor-driven movement in the cytoplasm. Therefore, MT dynamics are important for the formation of a radial array in the absence of the centrosome. Here we report the methods for studying the role of dynamic MTs in melanophores and describe the details of cell culture preparation, microsurgery and microinjection, fluorescence labeling, and live imaging of MTs.
II. Experimental Procedures A. Cultivation of Melanophores Melanophores are obtained either from fish (Gymnocorymbus ternetzi) scales or from frog (Xenopus laevis) tadpoles. Primary cultures of fish melanophores are prepared from black tetra scales prior to each experiment. Cells are separated from the scales by collagenase treatment, plated on carbon-coated coverslips, and incubated overnight in tissue culture medium. Permanent cell lines of X. laevis melanophores are obtained according to the modified method of Daniolos et al. (1990) that includes the following steps: trituration of tadpoles, purification of melanophores by Percoll density centrifugation, and generation of stable cell lines by limiting dilution. Xenopus melanophores become spontaneously immortalized during the first 2–3 weeks of growth in primary culture, and therefore melanophore cell lines can be continually maintained in culture. Xenopus melanophores containing pigment do not survive freezing, and therefore cells should be depleted of melanin by treatment with PTU (N-phenylthiocarbamide) that inhibits tyrosinase, an enzyme responsible for melanin synthesis.
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1. Primary Culture of Fish Melanophores 1. Remove 10–20 scales from black tetra (G. ternetzi) under a dissection microscope using Dumond #5 or #7 tweezers. 2. Place scales in fish Ringer’s solution (5 mM Tris–HCl, 103 mM NaCl, 1.8 mM KCl, 0.8 mM NaHCO3, 2 mM CaCl2, pH 7.2) supplemented with 5 mg/ml BSA and 1 mg/ml collagenase (Worthington, type 3). 3. Incubate scales at 30°C for about 1 h. 4. Under a dissection microscope, hold down scales (pin them) using tweezers and detach cells by pipetting using a 2–20 µl automatic pipettor. 5. Wash off the excess collagenase by transferring individual cells with a pipettor from one 35 mm cell culture dish containing Ringer’s solution into another dish; repeat three to five times. To prevent attachment of cells to the bottom of the dish, treat dishes with 1% BSA solution in PBS for 10–15 min, and rinse with water prior to use. 6. Place cell suspension onto 22 22 mm carbon-coated coverslips. Carbon coating is applied using the DV-502 evaporator (Denton Vacuum, Inc.) and coverslips are mounted with silicon vacuum grease over a hole drilled in a 35 mm cell culture dish. Instead of carbon, coverslips can be coated with laminin (40 µg/ml) or poly-L-lysine (0.1 mg/ml), although melanophores spread more readily on carbon. 7. Add 3 ml of tissue culture medium into each dish (DMEM supplemented with 20 mM HEPES, 20% fetal bovine serum, 200 µg/ml streptomycin, and 200 units/ml penicillin) and incubate overnight at 30°C for the complete spreading of melanophores.
2. Generation of Immortalized Cell Lines of X. laevis Melanophores 1. Rinse 20 X. laevis tadpoles (stage 30–35; Nieuwkoop and Faber, 1967) three times with amphibian Ringer’s solution (9 mM HEPES, 115 mM NaCl, 3 mM KCl, 2 mM CaCl2, pH 7.3). 2. Rinse with 70% ethanol for 5 s. 3. Rinse three times with amphibian Ringer’s solution. 4. Grind tadpoles in 10 ml of Xenopus tissue culture medium (XTCM; 0.7 L-15 medium (Sigma-Aldrich, St. Louis, MO), supplemented with 10% heatinactivated fetal bovine serum (Gibco, Invitrogen Corp., Carlsbad, CA), 200 units/ml penicillin, 200 µg/ml streptomycin, and 5 µm insulin, pH 7.3–7.4). 5. Plate the resulting suspension of cells onto three 60 mm cell culture dishes. 6. Incubate at 27°C for 1 week. 7. Rinse with XTCM to remove unattached cells. 8. Cultivate for 1–2 months changing XTCM twice a week until black colonies of melanophores appear. 9. Detach cells by trypsinization (briefly incubate in trypsin solution containing 2 mg/ml trypsin, 0.2 mg/ml EDTA in PBS) and collect by centrifugation. 10. Purify melanophores by centrifugation through Percoll cushion. Resuspend cell pellet in 20% Percoll solution, load onto 12 ml of 33% Percoll solution (both
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Percoll solutions prepared on XTCM), and pellet cells by centrifugation at 500 g at for 5 min at a room temperature. 11. Resuspend the pellet, which contains 80–90% melanophores, in 5 ml of XTCM, plate into a 25 cm2 flask, and cultivate at 27°C. Change the medium twice a week. 12. When melanophores become confluent, repeat steps 9–11 until unpigmented cells are completely removed. Usually three rounds of Percoll density gradient purification yield pure primary culture of melanophores. 13. Prepare feeder layer of Xenopus fibroblasts by plating them into a 48-well plate and cultivate until cells form a monolayer. Treat cells with 10 µg/ml mitomycin C (M0440, Sigma-Aldrich) for 3 h at 27°C to arrest cell growth. Wash three times with XTCM to remove mitomycin C. 14. Detach melanophores by trypsinization, resuspend in 10 ml of XTCM, count using a hemocytometer (Bright-line Counting Chamber, #3100 Haussen Scientific, Horsham, PA), and dilute the suspension to concentration of one cell per 400 µl. 15. Add 200 µl of cell suspension to each well of the 48-well plate containing a monolayer of Xenopus fibroblasts (step 13). 16. Cultivate for about 1 month; colonies of melanophores will appear in the wells. Be sure that wells containing two or more colonies are not used for further steps. Change the medium twice a week. 17. Select colonies of cells that grow fast and respond well to hormones by completely aggregating or dispersing pigment granules within 20 min after the treatment with 10 nM melatonin or 10 nM melanocyte stimulating hormone (MSH), respectively. 18. Grow cells in the selected wells until they form a monolayer, detach by trypsinization, and transfer into a 24-well plate. 19. Repeat step 18, and replate cells consecutively into 12- and 6-well plates, and, finally, onto the 25 cm2 flask. Always maintain melanophores in a semi-confluent culture since at low density cells stop dividing and responding to hormones. 20. If required, freeze melanophores as follows. Incubate cells in the XTCM supplemented with 1 mM PTU (P7629, Sigma) for at least 3 weeks to deplete melanophores of melanin. Trypsinize cells and freeze cell suspension in 20% DMSO in XTCM. After thawing in PTU-free XTCM melanophores produce pigment within 5–6 days.
B. Fluorescent Labeling of Tubulin Purified tubulin is obtained from porcine brain by a standard procedure involving two cycles of MT temperature-dependent assembly-disassembly (Borisy et al., 1975). MT-associated proteins (MAPs) are removed by an additional cycle of disassemblyreassembly of MTs in a high-salt buffer (HS buffer). Conjugation with Cy3 involves incubation of assembled MTs with the dye followed by the two cycles of MT assembly-disassembly and purification of labeled MTs by centrifugation through a glycerol cushion.
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1. Experimental Procedure 1. Thaw 1 ml aliquot of twice cycled tubulin (8–10 mg/ml) in PM buffer (0.1 M PIPES, 1.0 mM MgCl2, pH 6.9) in water bath at 37°C. 2. Add guanosine triphosphatase (GTP) to a final concentration of 1 mM. 3. Polymerize MTs by incubating at 37°C for 10 min. The solution should become viscous and turbid. 4. Pellet MTs at 100,000 xg (50,000 rpm, Beckman TLA 100.3 rotor) at 37°C for 5 min; discard the supernatant. 5. Measure MT pellet volume and resuspend it in equal volume of HS buffer (0.5 M PIPES, 2 mM MgCl2, 1 mM EGTA, pH 6.9). 6. Add DMSO to a final concentration 10% and GTP to 1 mM. 7. Polymerize MTs and pellet them by centrifugation as described in steps 2–3. At this step, the most MAPs are removed. 8. Resuspend the MT pellet in a twofold excess volume of cold PEM buffer (0.1 M PIPES, 5 mM EGTA, 2 mM MgCl2, pH 6.9). 9. Depolymerize MTs by incubating on ice for 10 min. 10. Add DMSO to a final concentration of 10% and GTP to 1 mM. 11. Polymerize MTs by incubating at 37°C for 10 min. 12. Dissolve one vial of commercial aliquot of Cy3 bis-reactive dye (GE Healthcare, PA23000) in 20 µl of anhydrous DMSO. Each commercial vial of Cy3 dye is sufficient for conjugation with 8–10 mg of tubulin. 13. Add the Cy3 solution to polymerized MTs and vortex immediately. Incubate at 37°C for 30 min. 14. Pellet MTs at 100,000 xg at 37°C for 5 min; discard the supernatant. 15. Resuspend the MT pellet in a fivefold excess volume of cold PEM buffer. 16. Add GTP to a final concentration of 1 mM. 17. Depolymerize MTs by incubating on ice for 10 min. 18. Centrifuge tubulin solution at 100,000g at 4°C for 5 min. Transfer the supernatant into a new cold tube. 19. Measure the volume of supernatant and adjust DMSO and GTP concentrations to 10% and 1 mM, respectively. 20. Polymerize and centrifuge MTs as in steps 2–3. 21. Repeat steps 14–18. 22. Polymerize MTs by incubating at 37°C for 10 min. 23. Load MTs onto 900 µl warm 33% glycerol (37°C) cushion prepared on PEM buffer containing 1 mM GTP. Pellet MTs by centrifugation in TLS55 swinging bucket rotor (Beckman, Fullerton, CA; #343778 tubes) at 135,000g (40,000 rpm) at 37°C for 20 min. Aspirate the supernatant and glycerol cushion and resuspend MT pellet in 200 µl of PEM buffer with 1 mM GTP. 24. Depolymerize MTs on ice for 10 min. Centrifuge tubulin solution at 100,000g at 4°C for 5 min. Transfer the supernatant into a new cold tube. The concentration of tubulin solution will be about 6–8 mg/ml. 25. Freeze 10 µl aliquots of supernatant containing Cy3-labeled tubulin and store in liquid nitrogen.
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C. Microinjection and Microsurgery For fluorescent labeling of cytoplasmic MTs in melanophores, cells are injected with Cy3-labeled tubulin. Immediately before microinjection, Cy3-labeled tubulin is clarified by high-speed centrifugation to avoid clogging of microneedles. Injected cells are incubated for about 1 h to allow for the incorporation of Cy3-labeled tubulin into MTs. To produce fluorescent speckles along MTs, cells are injected with Cy3-tubulin solution at a low (0.5 mg/ml) needle concentration. Centrosome-free fragments of melanophores with fluorescently labeled MTs are obtained by microsurgical dissection of the microinjected cells with a sharp glass microneedle.
1. Microinjection 1. Plate cells onto photo-etched carbon-coated coverslips (24 24 mm; Bellco Biotechnology, Vineland, NJ) mounted over a hole drilled in a 35 mm cell culture dish; photo-etched coverslips have grids that enable relocation of the microinjected cells. 2. Thaw a 10 µl aliquot of Cy3-labeled tubulin and clarify by centrifugation at 135,000g (40,000 rpm) for 5 min at 4°C. 3. Transfer 1 µl of Cy3-labeled tubulin solution into a glass microneedle with tip diameter of about 0.1 µm produced using micropipette puller, such as Narishige PB-7 (Narishige, East Meadow, NY), from thin-wall borosilicate glass capillaries with filament (an outer capillary diameter is 1.5 mm and inner diameter—1.12 mm; WPI Inc., Sarasota, FL). 4. Place the glass microneedle into a micromanipulator with pipette holder and inject cells with Cy3-labeled tubulin under a pressure of 40 hPa applied by a microinjector (model 5242, Eppendorf, Hauppauge, NY). 5. Incubate for at least 1 h at 27°C for Xenopus or at 30°C for fish melanophores to allow the incorporation of labeled tubulin into MTs.
2. Microsurgery 1. Plate fish melanophores on photo-etched carbon-coated coverslips (24 24 mm; Bellco Biotechnology) and grow in tissue culture medium overnight. 2. Prepare microneedles as described above. 3. For the dissection of a cytoplasmic fragment, place the tip of an empty glass microneedle close to the cell surface; the fragment is separated by capillary suction. Try to select cells with processes and perform dissection at the base of the process.
D. Live Cell Imaging and Data Analysis Cells injected with Cy3-tubulin are observed on an inverted fluorescence microscope. Images are acquired with a sensitive cooled charge-coupled device (CCD) camera. To prevent photodamage of cells caused by light exposure during the observation, the
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oxygen scavenger Oxyrase is added to the medium and mineral oil is overlaid on the culture medium to retard gas exchange. 1. Take a dish with microinjected cells and add Oxyrase (Oxyrase, Ashland, OH) and its substrate lactic acid (L4263 Sigma) to final concentrations of 2% and 20 mM, respectively; check the pH and adjust, if necessary, to 7.3–7.4. 2. Overlay 2 ml of mineral oil (purchased in a local drug store) to retard gas exchange and incubate for 10 min at a room temperature for oxygen depletion. 3. Acquire fluorescence images of cells with an inverted fluorescence microscope, such as a Diaphot 300 (Nikon, Melville, NY) equipped with a Plan 100 1.25 numerical aperture objective lens and 100 W mercury arc lamp, and a narrow band rhodamine filter set, which is compatible with the excitation of Cy3 fluorescence. For image acquisition, use a CCD camera, such as Andor iXon EM-CCD sensor (Andor, South Windsor, CT), Photometrics series 300 cooled CCD camera (Photometrics, Tucson, AZ), or other cameras which have high quantum efficiency at the emission wavelengths of Cy3 and rhodamine. Collect time series of images of labeled MTs with the exposure time 100–500 ms and 2–3 s intervals between the frames using Metamorph software (MDC, Downingtown, PA). 4. Determine parameters of MT dynamics by tracking the positions of MT plus-ends and recording X, Y coordinates using Metamorph. Analyze the data with an appropriate computer software such as the program based on the Multiscale Trend Analysis algorithm (Zaliapin et al., 2005) to determine the following parameters: the duration, length and velocity of MT growth and shortening, and the frequency of catastrophes (transition from growth or pause to shortening) and rescues (the transition from shortening or pause to growth). Alternatively, determine these parameters manually by decomposing life history (distance vs time) plots of MTs into periods of growth and shortening and pauses assuming the changes of MT length over 0.5 µm as growth or shortening events, and others changes as pauses. The parameters of MT dynamics in control and Taxol-treated melanophores with stabilized MTs are represented in Table I.
Table I Parameters of MT Dynamic Instability in Control and Taxol-treated Xenopus Melanophores
Growth distance (µm) Growth rate (µm/s) Shortening distance (µm) Shortening rate (µm/s) Catastrophe frequency (s–1) Rescue frequency (s–1) Duration of pauses (s) Number of analyzed MTs Number of analyzed cells
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2.96 ± 0.16 0.17 ± 0.03 3.17 ± 0.18 0.18 ± 0.04 0.026 ± 0.001 0.029 ± 0.001 3.99 ± 0.31 30 5
0.23 ± 0.01 0.06 ± 0.01 0.23 ± 0.01 0.06 ± 0.001 0.026 ± 0.002 0.042 ± 0.002 11.51 ± 0.57 30 6
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E. Quantification of Aggregation and Dispersion of Pigment Granules Responses of melanophores to hormones that induce aggregation or dispersion of pigment granules are quantified by estimating the fractions of cells with aggregated, partially aggregated, or dispersed pigment granules at a fixed time interval after induction. Cells in each category are manually counted using phase contrast micro scopy. Alternatively, the kinetics of aggregation or dispersion of pigment granules are determined by measuring pixel gray values within cell outlines using Metamorph region measurement tool. This parameter reflects the degree of homogenous pigment distribution throughout the cytoplasm.
1. Quantification of Cells with Aggregated, Partially Aggregated, or Dispersed Pigment Granules 1. Induce aggregation or dispersion of pigment granules. In the case of fish melanophores, treat cells with adrenaline taken at a final concentration 500 nM to induce aggregation. Induce dispersion by the addition of 5 mM caffeine. To induce pigment aggregation or dispersion in Xenopus melanophores, place cells in a serumfree medium, incubate for 1 h at 27°C, add melatonin or MSH, respectively, to the final concentration 10 nM. 2. Fix cells with 4% formaldehyde solution 5 or 10 min after the application of adrenaline or caffeine (fish melanophores) or 10 or 20 min after the stimulation with melatonin or MSH (Xenopus melanophores). 3. Count the cells with aggregated, partially aggregated, dispersed pigment granules by counting cells in each category using phase contrast microscopy. Completely aggregated cells have a compact aggregate of pigment granules in the center and no granules at the periphery. Fully dispersed cells contain granules randomly distributed throughout the cytoplasm. Partially aggregated cells have no granules at the margins and less compact pigment aggregate in the center.
2. Quantification of Kinetics of Aggregation and Dispersion of Pigment Granules in Xenopus Melanophores 1. Induce aggregation or dispersion of pigment granules in fish or Xenopus melanophores as described above. 2. Acquire time series of bright-field images of melanophores with 10 s time intervals using Metamorph time-lapse acquisition mode. 3. Measure the gray levels as integrated pixel values within cell outlines in each of the acquired images in the time-lapse series using Metamorph region measurement tool. The values in the fully dispersed state are taken as 100%. Percentage of gray levels is calculated for each image using the following equation: A ¼ ðIb It Þ=ðIb Id Þ100 where Ib is averaged background levels measured outside cell outlines, It is integrated pixel value within a cell outline at a given moment t, and Id is integrated pixel value
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within a cell outline in the fully dispersed state. Percentages of gray levels for each time point are averaged across the recorded cells and plotted as a function of time (Lomakin et al., 2009).
III. Discussion We have described the experimental approaches used to study the role of MT dynamics in intracellular transport and radial organization of MTs in melanophores. Fish or Xenopus melanophores provide a unique experimental system for imaging both dynamic MTs and moving membrane organelles. Dynamic MT plus-ends are easily followed in large and flat lamellae of melanophores. Synchronous movement of pigment granules driven by motor proteins in response to distinct hormonal stimuli can be observed using conventional light microscopy (Fig. 1). Large and flat fish Frog melanophores
Aggregated pigment
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Fig. 1 Phase contrast images of melanophores with dispersed or aggregated pigment granules. Fish (G. ternetzi, left images) and frog (X. laevis, right images) melanophores with dispersed (top) or aggregated pigment granules (bottom). Bar, 25 µm.
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melanophores are particularly good for micromanipulation, including microsurgical dissection of cytoplasmic fragments. However, they cannot be grown in large quan tities, and each experiment requires a laborious procedure of cell isolation. Therefore immortalized Xenopus melanophores are recommended for the experiments that do not involve microsurgery. Another advantage of Xenopus cells is that they are easy to transfect and are suitable for various biochemical experiments. For live cell imaging of MTs, melanophores are microinjected with fluorescently labeled tubulin. An alternative way of MT labeling involves the expression of tubulin fused with a fluorescent protein. This method is less laborious but involves transfec tion that is inefficient in cells grown in primary culture, such as fish melanophores.
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Fig. 2 MT dynamics are required for aggregation of pigment granules in Xenopus melanophores. (A) Time series of images of fluorescently labeled MTs during pigment granule aggregation. The arrow and arrowhead indicate a growing MT tip and a pigment granule, respectively. Numbers indicate time in seconds. Bar, 2 µm. (B) Stabilization of MTs reduces pigment granule transport. Responses to aggregation (melatonin, 10 min) or dispersion (MSH, 15 min) in control melanophores or melanophores treated with taxol (1 µM) are quantified. The data are expressed as the percentages of cells with aggregated (white bars), partially dispersed (gray bars), or dispersed (black bars) pigment granules. (C) Kinetics of pigment granule aggregation (left) or dispersion (right) in control (white squares) or taxol-treated (black squares) melanophores.
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Besides, the level of exogenous tubulin expression varies from cell to cell being sometimes insufficient for real-time observation, or oppositely, excessive expression generates high background. Therefore, microinjection is a preferable approach for MT labeling in melanophores. The amount of fluorescent tubulin introduced into the cytoplasm by microinjection can be adjusted according to the aim of an experiment. High tubulin concentration results in even labeling of the entire MT and enables observation of dynamic plus-ends, whereas low concentration generates fluorescent speckles that allow studying MT movement and distinguishing between two types of translocation—growth and shortening at the ends or transport of the entire MT (Vorobjev et al., 2001). The described methods were used to define the role of MT dynamics in intracellular transport and in MT organization, driven by motor proteins. We hypothesized that constantly growing and shortening plus-ends searched the cytoplasm for cargo and that such dynamic behavior increased the probability of organelle capturing. Live cell imaging of melanophores stimulated to aggregate pigment granules revealed the events of granule capturing by the growing plus-ends leading to the initiation of transport (Fig. 2). We further confirmed the role of MT dynamics in this process by treating cells with a MT-stabilizing agent and demonstrated that aggregation of pigment granules was inhibited under these conditions (Lomakin et al., 2009). In the absence of the centrosome MTs are organized by motor proteins (Borisy and Rodionov, 1999; Compton, 1998; Hyman and Karsenti, 1996; Sharp et al., 2000). We suggested that MT dynamics were important for this organization. The rearrangement could result either from the transport of previously assembled MTs by motor proteins or from the nucleation of new MTs and their rapid reorganization due to active growth and shortening. Imaging of MTs labeled with fluorescent speckles (Fig. 3) revealed
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(A)
Fig. 3
Organization of a polarized radial array of MTs in cytoplasmic fragments of fish melanophores. (A) Fluorescence images of MTs in a centrosome-free fragment of fish melanophore before (top) and after (bottom) stimulation of aggregation with adrenaline. (B) Fluorescence image of MTs with speckles at low magnification (left) and time sequences of speckles on MTs in the regions indicated on the left panel (right). Numbers indicate the time in seconds. Scale bars, 10 µm (left) and 2 µm (right).
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that MTs were not transported in the cytoplasm but were nucleated de novo, formed small local asters, which merged into a single array as a result of MT growth and shortening (Rodionov and Borisy, 1997). Therefore, live cell imaging of MTs in melanophores elucidated the role of dynamics in intracellular transport and centrosome-independent organization of MTs.
Acknowledgments This work was supported by NIH grant GM62290 to V.I.R.
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