Chapter 16 Isolation and Analysis of Microtubule Motor Proteins

CHAPTER 16

Isolation and Analysis of Microtubule Motor Proteins William M. Saxton Department of Biology Indiana University Bloomington, Indiana 47405

I. Introduction 11. Isolation of Motor Proteins A. Preparing Embryos B. Homogenization C. Clarification D. Microtubule Polymerization and Kinesin Binding E. Differential Sedimentation of Microtubules and Kinesin F. Additional Fractionation 111. Characterization of Microtubule Motors A. Microscopic Assays for Motor Proteins B. A Simple Motility Assay IV. Conclusions and Summary References

I. Introduction Eukaryotes employ force producing mechanochemical enzymes (motors) to move cytoplasmic components along microtubules. A few years ago, only two cytoplasmic microtubule-based motors were known: kinesin, which moves toward the plus ends of microtubules, and cytoplasmic dynein, which moves toward the minus ends of microtubules. Now, through gene sequence comparisons and biochemical analyses, it has become apparent that there are many microtubule-based motors in a wide variety of organisms. Those for which the gene sequences are known fall into two groups: the kinesin and dynein METHODS IN CELL BIOLOGY. VOL 44 Copyright D 1994 by Academic Press. 1°C. All rights of reproduction

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superfamilies. In Drosophila alone, genes for 11 members of the kinesin superfamily (Stewart et a f . , 1991) and 9 members of the dynein superfamily have been characterized (Rasmussen et al., 1994). To understand microtubule-based motility, the functions of each motor need to be identified. Some motors, including kinesin, appear to be expressed in all cell types and at all stages of development (Saxton et al., 1988; Hollenbeck, 1989; Stewart et al., 1991). Others are either stage or tissue specific (Stewart et al., 1991). While it is likely that some motors have unique functions, it has been shown that some have overlapping or redundant functions (Roof et al., 1992; Saunders and Hoyt, 1992). Thus, determining which motor or motors are responsible for a given motility process will not always be straightforward. Sorting out the biological functions of individual and sets of microtubule motors will require a variety of approaches including genetics. Drosophila provides an excellent system for such an endeavor, especially if one wishes to learn how microtubule motors are used during development and in differentiated tissues. Biochemistry and molecular techniques can be used to identify, isolate, and characterize motors, and then genetics and physiological studies can be used to analyze motor functions in uiuo. The isolation of Drosophila kinesin is described here to provide a guide for those who wish to isolate kinesin, to point out possible avenues for the isolation of other microtubule motor proteins, and to illustrate some of the gains that can result from an initial biochemical approach to studying motor proteins. The methods have evolved from pioneering work on in uitro microtubule polymerization (Weisenberg et al., 1968; Weisenberg, 1972), the identification of nucleotide analogs that can induce rigor binding of motors to microtubules (Lasek and Brady, 1985), and the development of in uitro assays for microtubule-based motility (Brady et al., 1982; Vale et al., 1985a). The specific methods I use for isolating kinesin from Drosophila (Saxton et al., 1988) were derived from those developed by Vale et a f . (1985b) and Scholey et al. (1985) fL*i the isolation of kinesin from squid neural tissue and sea urchin eggs, respectively. The identification and characterization of Drosophila kinesin helped to establish the widespread distribution and similarity of this motor in metazoans and provided the background necessary for entry into molecular and genetic analyses. The Drosophila kinesin heavy-chain gene (khc) was the first microtubule motor gene to be cloned and characterized (Yang et al., 1988). Its sequence founded the kinesin superfamily of related genes and also allowed secondary structure predictions for kinesin heavy chain (KHC) (Yang et a f . ,1989). Another advance was the development of an Escherichia coli expression system that uses khc to produce functional KHC protein. This expression system has been used to identify fragments of KHC that are necessary and sufficient for the production of microtubule-based movement in uitro (Yang et al., 1990), to study the structure of the “stalk domain” (deCuevas et al., 1992), and to identify proteins that can bind KHC (Gauger and Goldstein, 1993). KHC produced in E. coli promises future contributions to our understanding of how kinesin

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generates force and movement by allowing biochemical studies of mutant KHCs and by providing material for high-resolution structural analyses. Genetic tests of Drosophila KHC are being used to study the contributions of kinesin to the development and function of differentiated cells. Mutations in Drosophila khc were isolated (Saxton et al., 1991) using reverse genetic methods such as those described by Wolfner and Goldberg and by Hamilton and Zinn in Chapters 3 and 4, respectively. The mutants have been used to demonstrate that KHC is essential and that it is likely to function as a motor for axonal transport in the nervous system (Saxton et al., 1991; Gho et al., 1992). Mutations in khc result in impaired action potential propagation and impaired neurotransmitter release. Based on these studies, it has been hypothesized that kinesin is involved in the delivery of ion channels to the axonal membrane (Gho et al., 1992). Additional studies of the neuronal physiology of khc mutants, identification of genetic loci that interact with khc, and clonal analysis (Perrimon; Chapter 34) of khc function in a variety of nonneuronal tissues should provide further insight into the in vivo functions of kinesin.

11. Isolation of Motor Proteins A. Preparing Embryos If one wishes to prepare Drosophila microtubule motors from large amounts of material, the simplest and least expensive source is a population cage. Descriptions of how to develop and maintain large populations of flies are provided by Schaffer et al. in Chapter 5 and by Mahowald in Chapter 7. I have prepared active kinesin from as little as 1 ml of packed embryos but normally use between 25 and 100 ml. Egg collection pads covered with a thick layer of yeast paste are placed in population cages for approximately 24 hr and then rinsed into a set of graded copper screens (40, 60, and 120 mesh) with a gentle stream of room-temperature water. Embryos trapped in the bottom screen are then rinsed three times with E-wash (0.4% NaC1/0.03% Triton X-loo), immersed in a 50% solution of commercial bleach to remove the chorions (about 3 min), rinsed with E-wash until the odor of bleach is completely gone, rinsed twice with distilled water, and rinsed once with extraction buffer consisting of 0.1 M Pipes, pH 6.9,0.9 M glycerol, 5 mM EGTA, 2.5 mM MgSO,, and protease inhibitors: 1 pg/ml leupeptin, 2pg/ml aprotinin, 1 pg/ml pepstatin, 1 p M phenylmethylsulfonyl fluoride (PMSF), 1 mg/ml p-tosyl-L-arginine methyl ester (TAME), and 0.1 mg/ml soybean trypsin inhibitor (STI). The quality of the protease inhibitors is important. The activities of some decline rapidly in aqueous solution, so it is best to add the inhibitors to cold extraction buffer just prior to use. We use stock solutions of leupeptin and aprotinin at 1 mg/ml in H,O (stored at 4"C), pepstatin at 1 mg/ml in ethanol (stored at 4'0, and PMSF at 0.1 M in DMSO (stored at -20°C). TAME and STI are added to the extraction buffer dry.

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During the preceding steps, it is important to remember that the embryos are alive and sensitive to stress. To optimize results, abnormal temperatures and poor oxygenation should be avoided. Embryos should be kept in a thin layer or actively suspended in a large volume of room-temperature liquid as much as possible. In addition, one should move through embryo preparation to the point of homogenization as quickly as possible. B. Homogenization After embryos have been rinsed with extraction buffer, they are placed in a chilled Wheaton glass homogenizer with an equal volume of ice-cold extraction buffer. Keeping the homogenizer in ice, five strokes with the loose pestle (“A”) followed by seven strokes with the tight pestle (“B”) should completely disrupt the embryos. The Wheaton homogenizer is cheap and simple but not ideal. Substantial force is required and it must be applied in a controlled manner to avoid spewing homogenate out of the tube and onto one’s face. We have tried Brinkman and Vertishear blade-driven tissue homogenizers as well as sonication with little success. Tim Karr’s lab uses a teflon pestle driven by a drill motor in a Dounce homogenizer tube with good results (Karr et al., 1982). C. Clarification After disruption, the homogenate is clarified by low-speed and high-speed centrifugation steps. The low-speed spin is done in open plastic tubes in a swinging bucket rotor at 15,OOOg for 40 min at 4°C.A hardened layer of lipid will be found overlying the sample after the spin. The lipid layer should be picked up with a cotton-tipped applicator and discarded. The supernatant should then be removed with a large volume pipet, avoiding the cloudy material that lies on top of the pellet. The supernatant is transferred to clean plastic tubes and centrifuged in a fixed angle rotor at 50,OOOg for 30 min at 4°C.The resulting high-speed supernatant (HSS) can be used imediately or can be placed at -70°C for long-term storage. I have had good results with HSS that has been frozen for 2 years. D. Microtubule Polymerization and Kinesin Binding Microtubule polymerization is induced in HSS by the addition of GTP (0.3 mM) and taxol(20 p M ) , followed by gentle agitation at room temperature for 20 min. At this point, if one is interested in identifying novel motors whose binding to and release from microtubules is nucleotide sensitive, two samples of HSS should be treated in parallel. In sample “A”, rigor binding of kinesin and other proteins to microtubules is induced by addition of the nonhydrolyzable ATP analog 5’-adenylyl imidodiphosphate (AMPPNP) to 2.5 mM, followed by an additional 10 min of incubation at room temperature. To sample “B”, ATP

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and MgSO, are added to 2.5 mM each in place of AMPPNP. Proteins like kinesin, which bind to microtubules in A but not in B, are candidate motor proteins. If one is simply interested in quantitative preparation of kinesin, then all of the HSS should be treated with AMPPNP as described for sample A. E. Differential Sedimentation of Microtubules and Kinesin

After the binding step, microtubules are sedimented through a sucrose cushion (20% sucrose and 10 p M taxol in extraction buffer) by centrifugation in a fixed angle rotor at 23,OOOgfor 30 min at 4°C. The microtubules are then washed by resuspension in extraction buffer containing 10 p M taxol and 75 mM NaCl, followed by sedimentation through another sucrose cushion. The pellet from this salt wash is then resuspended in extraction buffer with 20 ph4 taxol, 75 mM NaCl, 10 mM MgSO,, and 10 mM ATP. The volume of buffer used in this resuspension will determine the final concentration of kinesin. My lab aims for 2.5% of the initial HSS volume. Suspended microtubules are then sedimented in a fixed angle rotor at 120,OOOg for 20 min at 4°C. The supernatant (“ATP extract”) contains highly enriched kinesin as well as some less abundant proteins. The activity of the kinesin is greatest if used fresh, but it can be stored at -70°C for long periods if a slight decrease in activity is acceptable. F. Additional Fractionation Column chromatography can provide fractionation of the ATP extract if required. Good results can be obtained with a Bio-Gel A5M gel filtration column (2.5 x 35 cm, 15 ml/hr) equilibrated with extraction buffer at 4°C. Bio-Gel A1.5M resin has also been used and produces good results (Cole er al., 1993). We have not explored other chromatography methods, but ion-exchange columns and FPLC can undoubtedly be used for kinesin (Hackney, 1991;Urrutia and Kachar, 1992)and may be essential for purifying less abundant motors. Fractionation of the ATP extract can also be achieved by repetitive binding to and release from microtubules. Taxol-stabilized microtubules are added to the ATP extract (0.5-1 mg/ml tubulin), rigor binding of motors is induced by addition of AMPPNP (10 mM), the microtubules are pelleted, and the bound proteins are released with ATP as explained earlier. The cycle can then be repeated. If one wishes to reduce cost, the ATP in the ATP extract can be depleted with hexokinase/glucose (Cohn et al., 1987), which allows the use of less AMPPNP (1.5 mM). This microtubule bindinghelease method can provide an enrichment of proteins like kinesin that bind microtubules with relatively low k,s in the presence of AMPPNP and relatively high k,s in the presence of ATP. If one were attempting to purify novel motors, the use of different nucleotides, nucleotide analogs (Shimizu et al., 1991, 1993), and ionic strengths might be used to differentially enrich for them.

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111. Characterization of Microtubule Motors A. Microscopic Assays for Motor Proteins The development of microscope-based in vitro motility assays has had a huge impact on our understanding of intracellular motility (Sale and Satir, 1977; Brady et al., 1982; Sheetz and Spudich, 1983; Vale et al., 1985a). Any attempt to identify new microtubule-based motors will eventually go to the microscope. The most common approach is to coat a glass coverslip with the cytoplasmic fraction to be tested, add microtubules and ATP, and watch for gliding of the microtubules over the coated surface (Vale et al., 1985a; reviewed by Paschal and Vallee, 1993;Cohn et al., 1993).Another approach is to anchor microtubules to a coverslip, add ATP, add microscopic beads coated with a cytoplasmic fraction, and look for translocation of the beads along the microtubules (Vale et al., 1985a; Gelles et al., 1988). When a cytoplasmic fraction that can induce microtubule-based movements has been identified, microscopic assays can be used to test a variety of characteristics. These include the velocity of movement (Cohn et al., 1993), the kinetics of nucleotide use (Cohn et al., 1993), the polarity of movement (Vale et al., 1985c; Porter et al., 1987; Yang et al., 1990; Paschal and Vallee, 1993; Howard and Hyman, 1993), the effects of ATP analogs on movement (Porter et al., 1987; Shimizu et al., 1993),force production (Hall et al., 1993; Kuo and Sheetz, 1993), and step size (Gelles et al., 1988; Svoboda et al., 1993). Some of these tests require highly specialized and expensive equipment. All of them require a video microscope. The most common system utilizes a video camera attached to a microscope that is equipped for differential interference contrast with a high-intensity source of light (Cohn et al., 1993). However, with suitable video cameras, dark-field and fluorescence microscopy can also be used (Vale and Hotani, 1988; Sale et al., 1993; Howard and Hyman, 1993).

B. A Simple Motility Assay If one wishes to assay microtubule motors in vitro with minimal fuss and little or no video equipment, sea urchin sperm provide a useful substrate (Lye et al., 1989; Yang et al., 1990). Sperm can be demembranated and treated with salt to expose dynein-free axonemal bundles of microtubules. Many axonemes will remain attached to their sperm heads via the minus ends of the microtubules. When applied to a coverslip that is coated with a cytoplasmic fraction, the axonemes can indicate both :he presence of microtubule motors and the polarity of movement. An axoneme that pushes the sperm head in front is being moved by pius-end-directed motors and one that pulls the sperm head along behind is being moved by minus-end-directed motors. Although images of the dernembranated sperm will be improved with video-enhanced contrast and image processing, they can be seen with regular light microscopic techniques.

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This assay was developed by C. M. Pfarr and J. R. McIntosh (unpublished), building on methods described by Gibbons (1982). A male urchin injected with 0.5 M KCI is placed upside down on a small beaker that is resting on ice. After sperm are shed into the beaker, a 25-pi aliquot is mixed gently with 75 pl of extraction buffer containing 0.5 M KCl and 0.1% Triton X-100. After 10-20 min at room temperature, the suspension is diluted 50- to 100-fold with extraction buffer and placed on ice. During the preparation, vigorous mixing and pipetting should be avoided to prevent fragmentation of the axonemes. This stock solution of demembranated sperm can be used for several hours in place of microtubules in a standard gliding assay (Cohn et al., 1993).

IV. Conclusions and Summary Isolation of microtubule motor proteins is needed both for the discovery of new motors and for characterization of the products of motor-related genes. The sequences of motor-related genes cannot yet be used to predict the mechanochemical properties of the gene products. This was illustrated by the first kinesin-related gene product to be characterized. Protein expressed from the ncd gene moved toward the minus ends of microtubules (Walker er al., 1990; McDonald et al., 1990), while kinesin itself moves toward the plus ends. Until the relationship between mechanochemical function and amino acid sequence is more thoroughly understood, biochemical isolation and characterization of microtubule motor proteins will remain essential. Two approaches for getting useful quantities of microtubule motor proteins have been used: isolation from cytosol as described under Section I1 above and isolation from bacteria carrying cloned motor protein genes in expression vectors. Bacterial expression of functional microtubule motors has been successful to date in only a few cases (Yang et al., 1990; Walker et al., 1990, McDonald et al., 1990). Additional progress is expected with the expression of cloned genes from viral vectors in cultured eukaryotic cells, but broad success has not yet been reported. Biochemical isolation of motors from their natural cytosol has some distinct advantages. One can have confidence that a given motor will be folded properly and have normal post-translational modifications. In addition, if it exists in uiuo as a heteromultimer, a microtubule motor isolated from its native cytosol will carry with it a normal complement of associated proteins. Studies of such associated proteins will be important in learning how motors accomplish their tasks in uivo. Drosophila cytosol should be a rich source of microtubule motors. Drosophila carry at least 11 and perhaps as many as 30 genes that are related to kinesin (Stewart er al., 1991; Endow and Hatsumi, 1991). The work of Tom Hays’ lab indicates that Drosophila carry more than nine dynein related genes (Rasmussen et al., 1994). Relatively little effort to isolate the products of these genes from

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cytosol has been made. The only work that I am aware of has produced a kinesin-like microtubule motor (D. G. Cole, K. B. Sheehan, W. M. Saxton, and J. M. Scholey, in progress) that may be the Drosophila homolog ofXenopus eg5 (Sawin et al., 1992). This isolation was straightforward, and efforts to identify additional motors are almost assured of success. Acknowledgments This work was supported in part by a grant from the NIH (GM-46295) to W. M. Saxton. The author is an Established Investigator of the American Heart Association.

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