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Cellular microtubules are rigid in comparison to other cytoskeletal elements [1, 2]. To facilitate cytoplasmic remodeling and timely responses to cell signaling events, microtubules depolymerize and repolymerize rapidly at their ends . These dynamic properties are critically important for many cellular functions, such as spindle assembly, the capture and segregation of chromosomes during cell division and cell motility. Microtubule dynamics are spatially and temporally controlled in the cell by accessory proteins. Molecular motor proteins of the kinesin superfamily that act to destabilize microtubules play important roles in this regulation .
The kinesin superfamily of proteins has recently been classified into fourteen subfamilies based on primary protein structure . The proteins that comprise many of these subfamilies utilize the energy from ATP hydrolysis to transport along microtubules. However, at least two kinesin subfamilies, the kinesin-13s and kinesin-14s, contain motors with microtubule destabilizing activity [6–8]. Detailed mechanistic studies of the microtubule destabilizing activity exhibited by these motors have significantly contributed to our understanding of their cellular functions.
In this chapter, we discuss assays for measuring the microtubule destabilizing activity of kinesins in both a purified system and within the context of mammalian cells. We first describe a turbidity assay that can be used to determine whether a kinesin is a bona fide microtubule destabilizer and to quantitatively compare the activities of different kinesin preparations. In addition, we report a fluorescence microscopy assay for analyzing the effect of kinesin expression on microtubule stability in cells. We have applied these approaches in combination with site-directed mutagenesis to study the microtubule destabilizing activity of MCAK, one of the founding members of the kinesin-13 family [9–12]. For many of these assays MCAK protein or constructs provides an admirable positive control for microtubule destabilizing activity.
Assembled microtubules cause detectable light scattering at 350 nm. This characteristic turbidity can be utilized to assay kinesin-mediated microtubule disassembly . The turbidity assay is best performed using GMP-CPP stabilized microtubules. Longer taxol stabilized microtubules tend to be bundled by added protein, and bundling of microtubules will artificially contribute to light scattering. We describe here how to prepare GMP-CPP microtubules and carry out the turbidity assay. To facilitate accurate comparisons between different preparations or variants of depolymerizing kinesins using the turbidity assay, we also describe a method used to determine the concentration of active motors competent to release ADP and bind ATP.
Tubulin has a higher affinity for GTP than it does for GMP-CPP . Thus, in order to grow microtubules with an uninterrupted GMP-CPP lattice, a method is required for removing the GTP that is commonly present in solution with purchased tubulin. To accomplish this, we recommend cycling the tubulin once in GMPCPP as noted in steps 1–4.
In addition to measuring microtubule depolymerization in vitro, it is important to determine the ability of a kinesin to depolymerize microtubules in vivo. This in vivo depolymerization assay depends in large part on the tubulin autoregulatory system. When microtubules are depolymerized by nocodazole or transiently transfected MCAK, the decrease of microtubule polymer and increase of tubulin dimer causes the degradation of β-tubulin mRNA [10, 14, 15]. Over a period of 24 hours, this causes an overall reduction of tubulin in the cell, which can be detected by a decrease in tubulin immunofluorescence. To measure microtubule depolymerization of a known or suspected depolymerizing kinesin in vivo, tubulin immunofluorescence intensity can be measured after kinesin expression and compared to tubulin immunofluorescence in control cells . We utilize Chinese hamster ovarian (CHO) cells for this assay because they consistently express transfected DNA quickly (i.e. within 12–24 hours).
We gratefully acknowledge the excellent work of Dave Coy, Jo Howard, Andy Hunter, Yulia Ovechkina, and Todd Maney in the development of these assays. This work was supported by a National Science Foundation NCI IGERT grant to J. Cooper, a National Institutes of Health Predoctoral training grant (GM07270) to K. Rankin and a National Institutes of Health grant (GM69429) to L. Wordeman.
1We routinely use pEGFP-C1 (Clontech). This vector drives strong, quick expression of EGFP fusion proteins under the control of a CMV promoter. The EGFP coded in the vector folds quickly and bleaches slowly, which allows for quick expression and stable photographing, respectively. Additionally, this vector has an f1 single strand DNA origin for replication that is useful for fast production of multiple mutant transgenes. N-terminal versus C-terminal placement of the transgene relative to the fluorescent protein should be considered and tested to determine whether placement has an effect on expression and function.
2The original tubulin concentration prior to assembly can be measured by measuring the absorbance (A) at 280nm assuming an extinction coefficient for tubulin of 115,000 mol−1cm−1. Using this value, a 1 mM solution of tubulin would possess an A280=1.15. This extinction coefficient is noted in Hyman et al. 1991 .
3Coverslips are mounted inside the centrifuge tube. Depending on the type of centrifuge tubes used, it may be necessary to machine a small fitting for mounting the coverslips securely at the bottom of the tube. It is important that the coverslip fits securely into the tube to ensure all microtubules are pelleted onto the glass. We routinely spin microtubules onto 5 mm diameter coverslips in 8 mm (outer diameter) centrifuge tubes using a Beckman airfuge. As an alternative to a machined fitting for the centrifuge tubes, a small amount of epoxy can be placed in the bottom of the tube and then spun while the epoxy is curing. This will generate a flat angled surface on which to lay the coverslip.
4Measure the average number of microtubules per frame, multiply this number by the total coverslip surface area, and divide by the surface area of one frame. This gives the total number of microtubules on the coverslip. Next, divide by the initial starting volume to determine the initial microtubule concentration. Multiply this number by two to determine the microtubule end concentration.
5For a typical ATP binding assay, prepare three nitrocellulose filters per protein prep being assayed and three for the buffer blank control. Additionally, we routinely wash a few spare membranes in case some break during the procedure.
6α32P-ATP, 800 Ci/mmol, 10 mCi/ml or 12.5 µM α32P-ATP on reference date. Concentration on date of assay is A(t) = Ao * e−lt where Ao = concentration on reference date, t = number of days after reference date, and l = ln2(1/half-life) or 0.0485 day−1 for 32P.
7Note that the hot: cold ATP ratio in this solution is assumed to be 1:50. If the stock concentration of active protein much exceeds 1 µM, the cold ATP carried by the protein will be significant, and a dilution of protein to 1–2 µM in motor storage buffer + 5 µM ATP should be made prior to the assay
8Each step in this process should be done as quickly and consistently as possible. Process 3 filters for each assay mixture or buffer blank.
9Example of ATP-binding activity calculation:
10A typical test reaction would employ 300 nM GMP-CPP microtubules and 3 nM full-length MCAK dimmer, but optimal motor concentration of other kinesins should be determined empirically.
11Addition of CaCl2 to a final concentration of 5 mM will disassemble all GMP-CPP microtubule polymer . This is a useful control to determine the A350 of fully disassembled microtubule polymer. When modest concentrations of active motor are added to microtubules, destabilizers like MCAK will disassemble the microtubules to a new steady state polymer concentration. It is unwise to assume that the lowest A350 reading represents fully disassembled microtubules.
12Turbidity traces can be converted to tubulin polymer concentration using a standard curve in which the zero time point corresponds to the concentration of tubulin in the assembled microtubules (i.e. 300 nM) and the turbidity after complete disassembly in CaCl2 is equal to 0 nM assembled tubulin.
13Confluency refers to the density of the cells adhered to the plate. For example, 100% confluency means cells that have adhered to and spread out on the plate are dense enough that they are touching each other and taking up 100% of the plate surface.
14One day prior to transfection, we plate CHO cells in a 24-well plate containing one acid-washed 12 mm coverslip in each well. Under these conditions, seed cells at a density of 3.5 × 104 cells per well in 500 µl culturing media. If cells are seeded too densely, analysis of tubulin fluorescence will be difficult because cells will not spread properly (section 3.2.3).
15We fix and wash our cells in 250 ml plastic beakers containing 100 ml of fix solution or PBS. The coverslips are held vertically in a coverglass staining rack and the cell containing side of each coverslip is carefully tracked through the procedure.
16We block and stain cells by placing coverslips cell-side up on a piece of parafilm laid flat inside a 25 mm Petri dish. Damp paper towels are rolled and placed around the inside of the dish to provide humidity and prevent drying during incubation periods.
17Choose cells that do not touch any other cells for this analysis because fluorescence from contacting cells can complicate the quantification procedure. Also, choose cells that provide a good dynamic range with respect to fluorescence intensity and avoid cells that contain areas of fluorescence saturation (at or above the maximal intensity value).
18We routinely take separate images for microtubules and GFP, and number the files sequentially (e.g. Kif2C_01_GFP; Kif2C__01_MT, where “Kif2C” is the name of the kinesin being analyzed). We keep all the files for each construct in a single folder. This organization is optimal for importing the files as a sequence into Image J for analysis.
19All the photos in the folder will open as a stack, with the GFP channel first and the microtubule channel second. To go forward within the stack, use the period key (.), and to go backward use the comma key (,).
20In ImageJ, set parameters by opening Analyze-Set measurements. Click the boxes for “Area”, “Mean Gray Value”, “Min and Max Gray Value” and “Display Label.”
21In ImageJ, take measurements with the Analyze-Measure command. Toggle between the GFP and tubulin images to take measurements using the same cell outline.
22The average GFP intensities should be statistically the same for all constructs in the assay by a student’s t-test analysis. Cells transfected with GFP should give a microtubule fluorescence ratio of approximately one, while cells expressing a microtubule depolymerizing kinesin should give a tubulin fluorescence ratio that is less than one (for example see Figure 2).