User:Andy Maloney/Kinesin & Microtubule Page

Purpose

This page describes the various procedures needed in order to produce gliding motility assays. Some insight and information into the chemicals, tools, and procedures used are given as well as failures I have encountered while trying to run these experiments.

This is the first chapter in my completely open notebook science dissertation. If you would like to post questions, comments, or concerns, please join the wiki and post comments to the talk page. If you do not want to join the wiki and would still like to comment, feel free to email me by using the provided link below.

• Click here to join the wiki.
• Click here to email me.
• Click here to post general comments about the open dissertation.
• Click here to post comments to this chapter's talk page.

Introduction

Figure 1: Image of the birthday cake my wife made me for my 30th birthday.

Experiments require certain tools in order for them to work. These tools can be chemicals, proteins, equipment, and/or procedures. Chemicals and proteins can easily be though of as tools since they are nothing more than the ingredients for an experiment, much like flour and eggs are ingredients for a cake. I also believe that equipment are tools just like chemicals and proteins. It does not matter if the equipment is an oven to bake a cake in or, a 2 W infrared laser used in an optical tweezers, they are both things used to make something.

My definition of a tool can even be extended to include procedures. In science, much like in baking, we can follow recipes to produce either proteins or the carrot cake found in Figure 1. The end product of the baking or procedure to produce a protein is what I consider to be the platform where scientists or bakers can begin to become creative. As popular TV chefs of this era who decorate cakes will understand⁽¹⁾, the cake can be thought of as the tool or rather, a vehicle, for creative arts. I should state, however, that I'm not belittling the mastery of baking because it is tough and not everyone can follow a recipe properly. Just like the preparation of a cake can be the platform for an artist to create a piece of art, so too is the preparation of an experiment the platform for a scientist to be creative while investigating nature.

I feel that the preparation of experiments in a reproducible manner should be paramount to experimentalists. Paying close attention to the design and preparation of an experiment allows the researcher to prepare the tools consistently. If the tools are produced the same every time, then the scientist has the leisure of not having to worry about an outcome changing simply because the time of day changes or other black box scenarios. The same thing applies to the baker, without the consistency of the cake recipe and ingredients, customers are not guaranteed a tasty cake on Wednesday as opposed to Thursday. Wednesdays are special days since they are the day that the Baron found out that "we don't silence other people's cannons"⁽²⁾. Without careful planning, experiments can be just as strange as the quote just mentioned. But again, sometimes the sweet success of doing something aspartame-esque in the lab is good. So, my stringent belief that experiments should be conducted in a precise manner does not hold all the time. However, for the gliding motility assay to work properly I have found it necessary to acquire a certain level of strictness.

A great number of tools are used in gliding motility assays. These tools come in the form of chemicals, proteins, equipment, and procedures. As with any biological experiment, there are numerous variables and any number of those variables can cause problems in the experiment. By understanding the chemicals and proteins used and observing my previous failures, I can now pin point where a problem exists that caused the assay to awry. Thankfully I have been able to collect enough data to show how this assay can go awry and, I hope that my failures can be of some use to others who do these types of experiments.

Methods and materials

There are several chemicals used for gliding motility assays. The fun part of these experiments are when changes to the chemicals are made to try and fish out physics about the interactions of kinesin and microtubules. Before I talk about changing chemicals, I will describe my procedure on how to make the basic buffer solution used for nearly all experiments; the PEM buffer. I will then go into how to make the other buffers and solutions necessary to run the most basic gliding motility assay.

The PEM buffer

PEM buffer overview

PEM, also known as BRB80, is the basic buffer used in all the gliding motility assays. It was found that tubulin polymerized quite nicely in this buffer with maximum effectiveness occurring at similar concentrations to what we use today. Olmsted and Borisy⁽⁸⁾ showed that microtubules polymerized very well in a solution that contained approximately 100 mM PIPES, 1 mM EGTA, and stociometric ratios of MgCl₂ to tubulin. Since microtubules polymerize so well in this buffer, it seems only natural to keep it in everything. This is why PEM is used in this assay and why I make sure to put all my proteins in this buffer.

Before I discuss the recipe of PEM that I have used for nearly all my experiments, I will first discuss the ingredients to the buffer. I will also discuss the chemicals, how I store them, and link to the exact products used in the subsequent experiments.

PEM stands for:

• PIPES
• EGTA
• MgCl₂

I have adopted this terminology since it is the easiest for me to remember and, it goes well with the naming of the other solutions necessary for the experiment. One of the difficulties with working with many different chemical solutions is agreeing on a common language and a naming convention. Unfortunately there is no common naming convention in the literature and as such, it can be difficult for a beginning researcher in this field to understand what buffers one group or another uses. In fact, when I was first attempting to learn the assay, I was presented with a buffer solution called BRB80 that had no explanation as to why it was called this. Apparently, no one else knows what the initials stand for either as the link would suggest. I hope that the naming convention I have proposed is straight forward and easy to use for other researchers.

PIPES is an acid and is the pH buffer used in PEM. It has a pK_a of 6.76 which is why I pH PEM very close to this value. Unfortunately, there is no consensus in the literature about which form of PIPES to use in PEM, i.e. K₂•PIPES or Na₂•PIPES. Neither is there a consensus on using sodium or potassium in solution and how one may be better than the other. See for instance Ray et.al.⁽³⁾ for an example of a buffer pH-ed with KOH and Woehlke et. al.⁽⁴⁾ for an example using NaOH. After discussing this with Dr. Koch, we chose to use the acid form (no K⁺ or Na⁺ attached to it) of PIPES. Since this form of PIPES did not have any sodium or potassium ions on it, we could actively choose which counter ion we wanted in solution. I decided to use NaOH in my version of PEM which means I probably should call it Na-PEM but, I've dropped the Na for simplicity. If I were to use KOH, then I would probably call it K-PEM. The stock chemical of PIPES is stored at room temperature in the desiccator in its original bottle.

EGTA chelates both calcium and magnesium from solution and is also an acid. EGTA has a higher affinity for calcium than it does for magnesium which is good since for motility to work, magnesium must be in solution⁽⁵⁾. I have yet to find an article that describes why EGTA is used in PEM. I believe that EGTA is used in PEM because I think it will chelate calcium phosphate found in casein micelles. Casein is used to passivate glass and is discussed in detail below. The chelation of calcium phosphate breaks apart casein micelles and I believe that breaking up the micelles aids in surface passivation. In their book, Fox and McSweeney⁽⁶⁾ state that EDTA disintegrates casein micelles but, they do not talk about EGTA disintegrating casein micelles. EDTA and EGTA are very similar compounds as both are magnesium and calcium chelators. Holt et. al.⁽⁷⁾ talked about the similarities of EDTA and EGTA used to investigate the ratio of calcium to phosphate in milk. These two references makes me inclined to believe that EGTA will disintegrate casein micelles similarly to how EDTA does it. In fact, I believe that EGTA may do a better job of breaking up the micelles since EGTA has a higher affinity for calcium than EDTA does. EGTA is stored in the desiccator at room temperature in its original bottle.

Magnesium is essential for both the polymerization of microtubules⁽⁸⁾ and for the motility of kinesin⁽⁵⁾. This is why magnesium chloride is included in the PEM buffer. MgCl₂ is purchased in solution at a concentration of 1 M in water. I do not purchase the salt form of MgCl₂ because it is extraordinarily hygroscopic. So much so that if you leave a pellet of it out, it will suck up so much water from the atmosphere that it will basically put itself into solution. Even in the dry New Mexico air. Since MgCl₂ is in solution already, it does not need to be stored in the desiccator and it can be stored at room temperature in its original bottle.

Since the pH buffer PIPES and the divalent cation chelator EGTA are acid forms of the chemicals, counter ions must be in solution in order for them to dissolve. As previously discussed, those ions come in the flavor of NaOH.

• NaOH

Getting the correct amount of NaOH for a solution of PEM was tricky at first, however, now I know the approximate amount to use because of trial and error. Scientists typically will not state how much of a pH-ing chemical is added to a buffer. I'm not sure why this is the case. Especially since it is known that changing the ionic strength of the motility assay does affect gliding speeds⁽⁹⁾. NaOH comes in pellet form and should always be dessicated due to its hygroscopic nature. It is also a very strong base so care must be taken when handling it. When using this chemical one must work quickly, otherwise it will pull water from the atmosphere and throw off weight measurement. NaOH should be stored at room temperature in the desiccator at all times.

Ultimately these chemicals, PIPES, EGTA, MgCl₂, and NaOH must be put into an aqueous solution. In order to create a buffer that is known to have only those chemicals one puts in it, one must use pure water. We have a reverse osmosis deionizing water system that produces very pure 18.2 MΩ-cm H₂O.

• 18.2 MΩ-cm H₂O

Water is very important in the experiments that I conduct. I will discuss in greater detail some properties of water and its isotopes in cite Ch3.

PEM buffer recipe & procedure

This PEM buffer is not unique and neither is it the standard buffer used for gliding motility assays. See here for a few other labs' "PEM" buffers.

I prepare a 10x concentrated version of the PEM buffer used in assays. I do this for two main reasons; the first is when I make TSB which is discussed below and second is because 10x PEM is used in another assay that contains H₂¹⁸O water. Also, I can make a smaller volume of 10x PEM and store it more easily than I can a 1x PEM solution. The 10x PEM buffer contains the following concentrations of chemicals.

• 800 mM PIPES
• 10 mM EGTA
• 10 mM MgCl₂
• [math]\displaystyle{ \approx }[/math] 1.25 M NaOH
• 18.2 MΩ-cm H₂O

The procedure I use to make this buffer is as follows:

1. Weigh out the appropriate amount of PIPES, EGTA, and NaOH to make a 25 mL solution. This comes out to:
   • 6.0474 g PIPES
   • 0.0951 g EGTA
   • 250 µL MgCl₂
   • ≈ 1.2 g NaOH
     • I like to keep the amount of NaOH below the 1.25 M upper limit. That way I don't over shoot the amount of NaOH needed. If I do end up going past the pH goal of 6.89, I will restart this procedure since I do not want to have HCl in the buffer.
   • All components are placed in a 50 mL centrifuge tube and vortexed with just enough 18.2 MΩ-cm H₂O water such that the chemicals will completely dissolve. I never put more than 15 mL of water in the tube at this step.
2. Once all the chemicals are in solution, I add more 18.2 MΩ-cm H₂O to the tube till the total volume is about 22 mL. I don't add the total amount of volume needed (25 mL) since I know that I may have to pH the buffer. Having less than the total volume needed ensures that I don't add too much water to the solution thus diluting the chemicals if I need to pH it.
3. Prepare a solution of 10 N NaOH, typically about 10 mL worth in 18.2 MΩ-cm H₂O just in case pH-ing is needed.
4. Determine the pH of the solution. Add small amounts of the 10 N NaOH prepared in the previous step if needed in order to reach the 6.89 pH. If I over shoot this value, I scrap the buffer and try again.
5. Once the correct pH is achieved, add the appropriate amount of water to reach the 25 mL total volume mark.
6. The PEM solution is then syringe filtered using a 0.2 µm filter and aliquoted into 1 mL screw top vials that are then labeled and stored in the 4˚C fridge in a convenient fridge box.

Surface passivation chemicals

Casein overview

Figure 2: Graphical representation of a bovine casein micelle made by Anthony Salvagno. Since whole casein is not crystalizable, this image may or may not be accurate but it is one theory to how whole casein looks. Alpha and beta casein form a globular "sub-micelle" complex (yellow balls) that are stabilized by kappa casein (black lines). The casein sequesters calcium phosphate from solution (red and blue markers).

In the gliding motility assay, motility is sustained by first passivating the glass microscope slides. Passivation is done to prevent kinesin’s motor domains from becoming inactive when interacting with untreated glass. It is not understood how or why kinesin motor domains become inactive on untreated glass but they do. Passivation of glass can be done with bovine serum albumin (BSA)^(1-3), bovine casein^(4-9), a lot of kinesin⁽¹⁰⁾, or other chemical compositions⁽¹¹⁾. Bovine casein is the surface blocker of choice by many experimenters mainly because it works well at passivation and is inexpensive. 500 g of whole bovine casein costs about $30 at the time of this writing. Typical assays will use 10 - 50 μg of casein at a time. This means that the 500 g stock of bovine casein will outlast a graduate student's career and if stored properly, possibly a PI's.

Casein is a globular protein that does not have a known crystal structure⁽¹⁷⁾, see Figure 2. Bovine casein is comprised of four major subgroups: α_s1, α_s2, β, and κ. Depending on the mammal the caseins come from, there exists different ratios of these globular constituents. For instance, bovine casein contains α_s1 + α_s2 > β > κ and human casein contains β > κ with only trace amounts of α_s1 casein^(19,20). Each species has a finely tuned milk for their neonates as casein is a vehicle in milk for delivering calcium phosphate and amino acids to them.

Figure 2 also shows a theory about how the casein micelle looks, for a discussion of the other theories, see cite Punungath. Since whole casein can not be crystallized, no one really knows for sure what the micelle looks like inside. What this model suggests is that kappa casein is more than likely on the outside of the micelle and it is there to stabilize its overall size. Alpha and beta caseins form complexes that are inside the micelle and help to sequester calcium phosphate.

How casein passivates glass surfaces in order to support kinesin for the gliding motility assay is still not very well understood but, some work has been done to try and understand it. Ozeki et al. showed that two layers of casein form on the glass surface to help support kinesin for motility^(cite). Verma et al.^(cite) showed that the number of microtubules that landed on the kinesin surface was affected by the casein passivation. Hancock and Howard also showed that the number of microtubules that landed on the kinesin surface was dependent on the number of motor proteins adhered to the glass slide^(cite).

With all these studies showing how the kinesin and microtubule system can be affected by the passivation substrate, one can assume that the system will be affected by the individual components of bovine casein. I will discuss this line of thinking more in detail in cite ch2. For now, I will outline how to prepare a basic solution of whole casein in PEM which is what most researchers use as the passivation chemical for conducting gliding motility assays.

Whole casein does not dissolve very easily without the addition of heat. Thankfully, whole casein does not have a secondary and thus it does not denature due to heatingcite fox. This is why milk can be subjected to ultra high temperature (UHT) pasteurization with minimal ill effects. I have never tried low temperature pasteurized milk but I hear that the milk tastes better. UHT heats milk above 135°C for very short periods of time in order to kill bacteria. Low temperature pasteurization uses much lower temperatures for much longer periods of time to obtain the same results.

The type of whole casein I use in experiments is from Sigma.

• Whole casein

I should point out that I also tried a Vitamin free version of whole casein as a passivator. I could not get this casein to dissolve in PEM very easily and thus I decided to use the "technical grade" whole casein from the link above.

W-PEM recipe

Figure 3: Condenser unit attached to a solution of whole casein being mixed and heated in PEM.

When I prepare whole casein solutions, I use a condenser since heating up the solution will cause evaporation and the use of a condenser eliminates that problem, see Figure 3. If I didn't use a condenser, then I would have to replace the water that evaporated back into the flask. This is cumbersome since one never really knows for certain how much of the water evaporated.

I will prepare whole casein at a concentration 1.0 mg/mL in PEM. I call this solution W-PEM to differentiate it from the other solutions of casein that are prepared for other studies, cite ch2.

1. Setup the condenser, stir/hot plate and a temperature probe.
2. I typically make 25 mL of W-PEM so I weigh out 25 mg.
3. I add the whole casein to the flask and then 25 mL of PEM. PEM is made from the 10x concentrated PEM stock solution by dilution.
4. Stir the solution at the highest setting on the stir plate. If my stir/hot plated allowed me to take the stirring up to 11, I would have.
5. Turn on the hot plate to a medium low setting, 3 for my case.
6. Monitor the temperature of the solution with a temperature probe. Once it reaches between 60 - 80°C, turn off the heat and continue to stir. The heating time is about 15 minutes for my setup. If there is still casein in solution after the 15 minutes, I will keep it on the hot plate till I no longer see visible precipitates.
7. Let the hot plate cool back to room temperature while still stirring the W-PEM.
8. Once the hot plate is cool, I will remove the flask from the condenser and cover it with Parafilm. There should be a foam on top of the solution. This foam indicates that the casein has been dissolved.
9. I will then put the W-PEM in the 4°C fridge over night to ensure that the foam has incorporated itself back into solution.
10. After the foam has disappeared, I will aliquot the W-PEM into convenient screw top vials and store them at 4°C.

If the casein solutions are kept at 4°C, they will last for up to 6 months. I do not filter this solution. The reason why I do not filter it is because I do not want to loose any of the casein in the filter. Not filtering it ensures that I have a 1.0 mg/mL solution of whole casein in PEM.

Antifade

Antifade overview

I use a very common antifade system that consists of the following components.

• Glucose Oxidase (GOD).
• Catalase (CAT)
• [math]\displaystyle{ \beta }[/math]-mercaptoethanol (BME).
• D-glucose.

The antifade system is vital for observing a good gliding motility assay. It is used because it prolongs the observation time of fluorescent microtubules. Since I observe my microtubules with fluorescence, elongating the time it takes before the microtubule fades is crucial for taking good data with a high signal to noise ratio. I rather dislike using this antifade system because the BME has a very strong sulfer odor. There are other recipes for antifade systems that exist⁽¹⁰⁾, however, I have not preformed any experiments with them.

Glucose oxidase requires D-glucose in solution as this is its fuel source. GOD oxidizes D-glucose to gluconic acid while using up oxygen in the solution. This is a good thing since photobleaching is caused from highly reactive oxygen species. Unfortunately when GOD oxidizes D-glucose, it also produces hydrogen peroxide which can damage the kinesin microtubule system. Catalase is added to the mix in order to decompose the hydrogen peroxide. BME, β-mercaptoethanol, or 2-mercaptoethanol is used to prevent blinking of the fluorophore and to quench triplet states^(10,11). Glucose oxidase and catalase should be stored in the -20°C freezer and BME can be stored in the 4°C fridge all in their original containers.

BME is nasty stuff and smells quite terrible. If any of this stuff spills anywhere, the stench will permeate through the lab for days. I know this because it has happend in my lab. Even if it's just a microliter, it will stink up the place very badly. Anything that touches BME needs to be handled carefully and disposed of, or cleaned properly.

Antifade components

The antifade chemicals are mixed in PEM and stored in the -80°C freezer. Below I outline the stock solutions necessary to prepare the antifade cocktail. In the recipe section, I describe the recipe that I use to prepare what I call antifade.

PEM-GOD

Weighing out the amount of glucose oxidase needed to prepare a stock solution of PEM-GOD can be difficult since such a small quantity is needed. I typically opt to make a 1000x more concentrated solution than what is needed in the motility assay since it is much easier to weigh out at this concentration. I will then dilute the 1000x solution to a 100x solution as this is what is needed for the antifade cocktail. 1000x PEM-GOD contains:

• 20 mg/mL Glucose oxidase in PEM

I only make 1 mL of PEM-GOD at a time. This solution will keep for 6 months in the -80°C freezer in a screw top vial.

PEM-CAT

Again, weighing the amount of catalase needed is difficult. I again opt for making a 1000x concentrated solution of CAT in PEM and then dilute it by a factor of 10 to obtain the 100x solutions needed for the cocktail recipe. 1000x PEM-CAT contains:

• 8 mg/mL Catalase in PEM

I only make 1 mL of PEM-GOD at a time. This solution will keep for 6 months in the -80°C freezer in a screw top vial.

BME

BME does not have to be diluted in PEM. It is used as is from the stock solution purchased from Sigma.

PEM-Glu

Along with the above two solutions, I make up what I call PEM-Glu. PEM-Glu is nothing more than a 2M solution of D-glucose in PEM. Yes. I want to make a 2 M solution of glucose, which means that there is a lot of sugar to put into a very small volume of liquid. This is an extreme case where the volume of the solute has to be taken into consideration. If I measure out the total volume of liquid I want to make a 2 M solution of glucose with and just add the glucose to it, I will end up with a solution that is a larger volume than I anticipated. To prevent this from happening, I weigh out the glucose and add it to a smaller volume of liquid than my final target total volume. I vortex it so that a considerable portion of the glucose goes into solution and then add more PEM till I reach my target volume. I only make 1 mL total volume of PEM-Glu and aliquot it into 20 µL aliquots and store them in the -80˚C freezer. D-glucose should be stored at room temperature in its original container. I should note that the PEM-Glu is not added to the antifade system aliquots, it's added to the motility solution before observations.

Antifade recipe

The recipe and procedure for this antifade system is as follows.

• More importantly than the exact volumes listed below is the ratio of the chemicals used, i.e. PEM-GOD:PEM-CAT:BME is 2:2:1.
  1. 12 µL of 100x GOD is added to a microcentrifuge tube.
  2. 12 µL of 100x CAT is added to the same microcentrifuge tube.
  3. In the hood with the fan on, 6 µL of BME is added.
  4. Vortex the solution and spin it down.
  5. Aliquot the antifade into 5 µL aliquots stored in microcentrifuge tubes.

Antifade solutions stay viable for only one week if stored in the -20˚C freezer.

Tubulin suspension in TSB

The tubulin I use comes from bovine brains and is purchased from Cytoskeleton. Thankfully there is a company in which I can purchase tubulin from since I'm not too keen on having to harvest tubulin myself which involves liquefying brains⁽¹²⁾. I have three different types of tubulin in the lab:

• un-labeled bovine tubulin
• rhodamine-labeled bovine tubulin
• fluorescein-labeled bovine tubulin.

The tubulin is lyophilized (flash frozen) and comes from Cytoskeleton in 1 mg, 20 µg, and 20 µg aliquots respectively. Aliquots are stored in the -80˚C freezer at all times upon arrival. My gliding motility studies use rhodamine labeled tubulin exclusively, as it gives a much better signal to noise ratio than does the fluorescein labeled tubulin.

To prepare the tubulin for easy polymerization, I store it in what I call TSB or Tubulin Storage Buffer. The limited number of studies I have looked at show that tubulin polymerizes into microtubules best in PEM⁽⁶⁾. I believe this is why we use PEM in everything and TSB is no exception as TSB is just PEM with extra stuff in it. I use the following recipe to prepare TSB.

• 1.06x PEM
• 1 mM MgCl₂. This is an extra 1 mM above what is already in the PEM.
• 1 mM GTP.
• 6% (v/v) Glycerol

The extra 1 mM MgCl₂ is added since microtubules will not polymerize effectively without it in solution⁽¹²⁾. The lab has chosen to add an extra 1 mM in the TSB since EGTA chelates Mg²⁺ from solution and I'd much rather have too much MgCl₂ in solution than not enough.

Figure 2: Image of GTP bottle and its insert.

GTP comes packaged nicely in 10 mg jars so I do not have to weigh it out. It is highly toxic so I use caution when opening the jar. Preparing this suspension in the hood is highly recommended in order to prevent any inadvertent inhalation of GTP dust, as it definitely burns a lot when inhaled. I add the 1.06x PEM to the GTP bottle such that the GTP is at 100 mM. In order to get all the GTP from the bottle into solution, I have to slosh the contents around after adding the PEM. The GTP bottle has an insert in it that I pull out for easier mixing.

The reason as to why I put 100 mM GTP in a 1.06x concentrated solution of PEM and not the 1x PEM is because the GTP will be used exclusively in the TSB. TSB gets diluted by 6% (v/v) when adding the glycerol hence the need for GTP to be in a higher concentration of PEM. I transfer the GTP + 1.06x PEM mix into a screw top vial and flash freeze it with LN₂. I store it in the -80˚C freezer. I have used both a diluted PEM solution and one that takes into consideration the dilution of 6% when adding the glycerol. Both work fine in polymerizing microtubules.

Glycerol is extraordinarily viscous and is a terrible pain to try and measure out. The way I measure it out is with the 1000 µL pipettor. The 1000 µL pipet tip is large enough such that I can get the glycerol to go into it and I've found that glycerol won't go up a 100 µL pipet tip at all. Getting all the glycerol needed in the pipettor takes some zen motivation as it takes patience to wait till all the glycerol is in the tip. As Heinz ketchup would say back in the 1980's "Good things come to those who wait."

Glycerol is used to speed up microtubule polymerization⁽¹³⁾. Other chemicals can be used in polymerization from DMSO to excess Taxol. These three polymerization techniques result in three different types of microtubules being polymerized with the major difference in the microtubules being the number of protofilaments⁽³⁾.

GTP suspension

To prepare the GTP in 1.06x PEM I follow the below recipe.

• 1.06x PEM:
  • 2820 μL 18.2 MΩ-cm H₂O.
  • 180 μL 10x PEM

• GTP suspension:
  • 191.14 μL 1.06x PEM.
  • Add to the GTP bottle and mix. Place in a screw top vial and store in the -80°C freezer.

TSB recipe and procedure

I prepare a 2 mL solutions of TSB. I do this mainly so that I am in the lower range of the 1000 mL pipettor for glycerol measurements. If stored properly in the -80˚C freezer, TSB will last up to a year. Unfortunately, there is no way anyone will use all the 2 mL TSB solution before it goes bad. My recipe is below.

• 1858 μL 1.06x PEM
• 120 μL Glycerol
• 20 μL 100 mM GTP in 1.06x PEM
• 2 μL MgCl₂

Tubulin dimers are highly unstable, so care should be taken to keep them as cold as possible and to minimize steps that keep the tubulin from being frozen. Once I have the TSB prepared, I am ready to prepare aliquots of tubulin that can be used for polymerization into microtubules.

Un-labeled tubulin suspension

Un-labeled tubulin comes packed in vials containing 1 mg of tubulin. I suspend this tubulin to a final concentration of 5 mg/mL in TSB in convenient aliquots.

1. I remove a vial of tubulin from the -80˚C freezer and put it in the e•IceBucket to defrost. If necessary, I spin the vial to get all the tubulin to settle at the bottom. Be careful though since tubulin is very labile and may be destroyed during this step.
2. I then suspend the tubulin in 200 µL of TSB. I mix the solution by gently drawing the tubulin + TSB mixture back into the pipettor and blow it out again into the vial. I will do this three time.
3. I then aliquot into 5 µL aliquots in the 200 μL microcentrifuge tubes, flash freeze in LN₂ and store in the -80˚C freezer.

Labeled tubulin suspension

Rhodamine labeled tubulin and fluorescein labeled tubulin come packed in vials containing 20 µg of tubulin. I will suspend either of these tubulins to a final concentration of 5 mg/mL.

1. I remove a vial of tubulin from the -80˚C freezer and put it in the e•IceBucket to defrost. If necessary, I spin the vial to get all the tubulin to settle at the bottom. Be careful though since tubulin is very labile and may be destroyed during this step.
2. I then suspend the tubulin in 4 µL of TSB. I mix the solution by gently drawing the tubulin + TSB mixture back into the pipettor and blow it out again into the vial. I will do this three time.
3. I then aliquot into 2 µL aliquots in the 200 μL microcentrifuge tubes, flash freeze in LN₂ and store in the -80˚C freezer.

29% Labeled tubulin suspension

Using 100% rhodamine labeled tubulin in an experiment is not ideal. I have found an acceptable ratio of labeled tubulin to unlabeled tubulin using the 100W Hg lamp with 6% illumination. 29% rhodamine labeled tubulin to 61% unlabeled tubulin gives a great signal to noise ratio and the microtubules can be tracked easily. Since both my unlabeled and labeled tubulin are now in convenient aliquots, I will use them to make up a 29% rhodamine labeled suspension. I do this by:

1. I will dethaw one aliquot of the rhodamine labeled tubulin and unlabeled tubulin each in the electronic ice bucket.
2. I add the unlabeled tubulin to the labeled tubulin.
3. I mix gently with the pipettor to ensure that the two solutions are mixed properly.
4. I then aliquot into 1 μL aliquots and store in the -80°C freezer in the 500 μL microcentrifuge tubes. I will typically use a 25 mL centrifuge tube to store my tubulin aliquots in.

These 29% labeled tubulin suspension are what I will use in experiments. See below for a description of how I polymerize the tubulin into microtubules.

Taxol suspension

Taxol is purchased from Cytoskeleton and is stored in the -80˚C freezer in a container filled with desiccant. Taxol is used to stabilize the microtubules as they are quite labile and will break apart easily⁽¹⁴⁾. Breaking apart easily is a spectacular characteristic of microtubules as they are essentially roads, for motor proteins, under constant construction and deconstruction in the cell. The cell can determine if a road is no longer needed or if it needs to be repaired or elongated thus allowing the motors that travel along them to deliver items to different parts of the cell. This "instability" is essential for cell function and in mitosis. A good example of this instability is shown in the below video.

<html> <object width="480" height="385"><param name="movie" value="http://www.youtube.com/v/ZL3_BwrB6AM?fs=1&amp;hl=en_US"></param><param name="allowFullScreen" value="true"></param><param name="allowscriptaccess" value="always"></param><embed src="http://www.youtube.com/v/ZL3_BwrB6AM?fs=1&amp;hl=en_US" type="application/x-shockwave-flash" allowscriptaccess="always" allowfullscreen="true" width="480" height="385"></embed></object> </html>

Movie 1: Dynamic instabilities of microtubules.

The above movie is from the Yamada Lab at UC Davis. This video shows that the ends of microtubules, labeled with GFP, grow and shrink dynamically. While this is good for cells, it is not good for gliding motility experiments as I am measuring speeds at which the microtubules float across a sea of kinesin. If the microtubule under observation suddenly shrinks, this will not allow me to get a good measure of its speed. To prevent the breakup of microtubules (depolymerization) I use Taxol.

Taxol is an anti-cancer drug⁽¹⁵⁾ that stabilizes microtubules. Since cancer cells are fast growing cells, Taxol helps to slow down the spread of the tumor by inhibiting microtubule dynamics and thus cellular replication. The stabilizing effect of Taxol is a good thing for my experiments and is why it is used.

Taxol is hydrophobic so it will not go into an aqueous environment unless the solution has DMSO in it. DMSO is an amazing chemical that solubilizes Taxol such that I can add it to a PEM solution. DMSO is hygroscopic and reacts with just about everything. An interesting side note about DMSO is that if it gets on your skin, you will taste garlic almost immediately. Sigma packs DMSO in ampules and the liquid needs to be transferred into a different screw top container for easier access. Since DMSO likes just about everything, it must be stored in something that it will not react with. DMSO can be stored in HDPE, LDPE, and PP without any problems. The DMSO from one ampule can be stored in 3 cryo vials. These cryo vials are then stored in a secondary HDPE container filled with desiccant in a nitrogen environment and stored in the desiccator.

10 mM stock Taxol suspension procedure

The Taxol I get from Cytoskeleton has approximately 170 μg of Taxol in the vials. Adding 20 μL of DMSO will make a 10 mM Taxol in DMSO solution.

• One vial of Taxol from Cytoskeleton.
• 20 μL of DMSO.

I vortex the vial and spin it down for use. I always store the 10 mM Taxol solution in the e•IceBucket when not in use since it is set to 3˚C. DMSO freezes at 3˚C so this is a nice way to see if the solution is still good or not. If the solution does not freeze, this means that the DMSO has absorbed enough water from the atmosphere to render it unreliable for experiments.

If there is too much Taxol in solution or, if the Taxol stock has gone bad, I will get Taxol spindles as depicted in the movie I took below. For an explanation on how Taxol forms these spindles see^(16-18).

<html> <object width="480" height="385"><param name="movie" value="http://www.youtube.com/v/2Tn7P957iXU?fs=1&amp;hl=en_US"></param><param name="allowFullScreen" value="true"></param><param name="allowscriptaccess" value="always"></param><embed src="http://www.youtube.com/v/2Tn7P957iXU?fs=1&amp;hl=en_US" type="application/x-shockwave-flash" allowscriptaccess="always" allowfullscreen="true" width="480" height="385"></embed></object> </html>

Movie 2: Taxol crystals found in one of my assays. Note that the crystal does not photobleach.

If this occurs, I make a new stock solution of Taxol. The worst case scenario when this does happen is that the pipettor has gone out of calibration and I have put too much Taxol in solution. Thankfully, that doesn't happen too often and Taxol crystals typically only occur for me when my Taxol solution has stopped freezing at 3˚C.

There are some things to take note of about this spindle. The first thing is that the fibers are rigid. If I observe what I think is a microtubule but it doesn't bend at all, then it is more than likely a Taxol crystal. Also, the spindle does not photobleach very much. The above movie was observing the spindle with no ND filters in front of the Hg lamp. If this was a bunch of microtubules and not a Taxol crystal, the microtubules would have depolymerized quickly leaving nothing to observe.

PEM-A

ATP is the fuel for kinesin. It is what kinesin uses in its motor domains to produce a step on a microtubule. To prepare PEM-A, I use:

• PEM
• ATP

Since I have a salt form of ATP, it goes into solution very easily. ATP should be stored in a secondary container that is filled with desiccant and under a nitrogen environment in the -80˚C freezer.

The book, Molecular Cloning⁽²¹⁾, says to suspend ATP in a Tris buffer at pH 8.0 since ATP auto-hydrolyzes less in alkaline buffers as opposed to acidic ones. Alberty⁽²²⁾ shows a graph that the auto-hydrolysis of ATP doesn't change very much for ATP stored in buffers at pH 7. Since PEM is pH-ed to 6.89, I figured it would be just fine to store the ATP in it. I have not had any problems using ATP stored in PEM and I believe that not having to introduce another chemical, namely the Tris, into the assay is beneficial. I will note that after storage, the PEM-A solution will start to look cloudy. I will mix the solution as best as I can and spin the precipitate to the bottom of the tube using the mini centrifuge. I'll decant the fluid such that I do not get any of the precipitate in my assays.

Recipe and procedure

• 100 mM ATP in PEM
• I'll aliquot into 10 μL vials and store them in the -80°C freezer.

Flow cell

The above link shows an old method to how I prepared my flow cells. Rather than get rid of it entirely, I have opted to include the link so that it is easier to show the evolution of the flow cell preparation. I put some notes by the video pointing out where I did things incorrectly.

Below, you will find my updated version of how I create my flow cells.

<html> <object width="640" height="385"><param name="movie" value="http://www.youtube.com/v/rnp9qKNep8U?fs=1&amp;hl=en_US"></param><param name="allowFullScreen" value="true"></param><param name="allowscriptaccess" value="always"></param><embed src="http://www.youtube.com/v/rnp9qKNep8U?fs=1&amp;hl=en_US" type="application/x-shockwave-flash" allowscriptaccess="always" allowfullscreen="true" width="640" height="385"></embed></object> </html>

Microscope and software

In this section, I will describe the microscope that I use and briefly introduce the software that has been designed in Dr. Koch's lab, mainly by Larry Herskowitz for the observation, tracking, and automated data analysis of the gliding motility assays.

Programs and equipment

I use an Olympus IX71 inverted microscope setup for epifluorescence using a 100W Hg lamp from Olympus. The objective is an Olympus PlanApo 60x 1.42NA. The filter cube is a TRITC filter cube with Chroma filters in it. The microscope also has ultraviolet neutral density filters and I typically will illuminate my microtubules with 6% of the 100W Hg lamp. I have made a cover for my microscope out of blackout material and PVC pipe. The cover is really cheap and it blocks out stray light as well as keeping dust and strong air currents from affecting measurements. My objective has extra stuff attached to it that I will talk more about below.
I use an Andor Luca-S camera with custom LabVIEW acquisition software written by Larry Herskowitz. The acquisition software captures .png files from the camera (grey background program) and are time stamped and stored for later analysis. Another custom LabVIEW program written by Larry is used to convert the series of .pngs captured by the camera to .avis (pink background program). The camera is attached to a 3 axis stage as well as a rotation stage and not directly attached to the microscope. This allows me to center the camera in the field of view easily and reduces noise in the acquisition due to the camera's fan.
After the images are captured, another custom LabVIEW software written by Larry is used to automatically track microtubules as they glide over kinesin in the assays. This program searches a parent directory of pngs and tracks the minus and plus ends of the microtubules through pattern matching algorithms.
Once the microtubules have been tracked, yet another custom LabVIEW program (again, thanks to Larry) is used to analyze the data. This program loads data from the tracked microtubules and performs data smoothing and kernel density estimation analysis on the speeds of individual microtubules. It can do so in an automated manner and has the capability of analyzing thousands of microtubule tracks relatively quickly.
The current computer used to capture and analyze data has the following specs.

Objective heater

Temperature stabilization is crucial for obtaining stable speed measurements. I will talk more about why in a following chapter. I have followed the work done by Mahamdeh and Schaeffer⁽²⁰⁾ pretty closely with my objective heater build.

Materials

Most of the materials can be purchased from TeTech. The rest can be purchased from Mouser.

• Temperature controller.
• 15k thermistor.
• Another 15k thermistor.
• Objective thermal spacer.
• Power supply.
• Box and Box cover.
• LEDs.
• Switches.
• Flexible heaters.
• Copper tape.

Build

The temperature controller I purchased came in its own aluminum box with exposed screw terminals. I didn't like the idea of having exposed wiring from the control box so I made a break away box with banana connectors on it. It isn't pretty but, it gets the job done. I also connected an RS232 connector to the box in order to communicate with the computer. There are several connectors going to this box. 2 for power, 2 for power to the heater, and 4 connectors for 2 different thermistors. The control thermistor and the sensor thermistor.
I taped the larger thermistor to the base of the objective and placed the flexible heating element above it. The heating element comes with glue on it so all I did was stick it to the objective. There are 2 alligator clips attached to the heating element that supply current to it from the control box. The thermistor on top is epoxied into place with Arctic Silver Thermal Epoxy.
The power supply is an OEM power supply and so it has exposed terminals on it. I enclosed the power supply in a box and chose to have a main switch to power the power supply (green indicator LED) and another DPST switch on the front to turn on the +12VDC output to the temperature controller. This allows the power supply to warm up before switching power to the temperature controller. To the right is a picture of the circuit in the box.
If I could have found a bigger switch on the front, then I would have used it. Why? Because everybody loves switches.

Experiments

Once I have all my stock solutions prepared, I am now in a position to start putting them all together. There is still some preparatory work that needs to be done before an experiment can be observed in the microscope however.

Assay checklist

Before starting an experiment, I always ensure that I have the following solutions and stocks prepared ahead of time.

1. 10x PEM
2. H₂O - just a convenient vial of water is necessary to dilute the 10x PEM.
3. PEM
4. 29% labeled tubulin
5. Antifade
6. 10 mM Taxol in DMSO
7. PEM-Glu
8. α-PEM
9. PEM-A
10. PEM-T
11. PEM-α
12. PEM-αA
13. Kinesin

I have not spoken about PEM-T, PEM-α, PEM-αA, or kinesin yet and before I outline my recipe to make the motility assay, I will talk about these solutions.

PEM-T is what I use to fix microtubules that have been recently polymerized. The recipe for this is:

• 10 μM Taxol in PEM.

This Taxol solution is in an aqueous environment. This means that I never keep a stock solution of it lying around and I will make a fresh batch of it every time I polymerize microtubules. I will make up a 199 μL solution of PEM-T by mixing up the following.

• 198.8 μL PEM
• 0.2 μL of the 10 mM Taxol in DMSO.

I make this solution about 5 minutes before the microtubules are supposed to come out of the thermal cycler. I do this to help ensure that there will not be a large number of Taxol crystals in solution and that most of the Taxol will be used to stabilize my fresh microtubules.

PEM-α is just a dilution of α-PEM. Instead of 1.0 mg/mL of alpha casein in PEM, it is 0.5 mg/mL alpha casein in PEM. PEM-α is used to dilute the kinesin in and I make up

• 500 μL α-PEM
• 500 μL PEM

1 mL typically.

PEM-αA is PEM-α plus 1 mM PEM-A. PEM-αA is what is used to store the kinesin that will be used in an assay in. I will make up the following:

• 99 μL PEM-α
• 1 μL PEM-A.

Kinesin was supplied by Dr. Haiqing Liu at a concentration of 275 μg/mL should be stored in the -80°C freezer at all times. When I'm ready to conduct experiments, I will take an aliquot of kinesin out of the freezer and place it in the electronic ice bucket. Once the kinesin has been removed from the freezer, I never put it back as in doing so will cause the kinesin to deteriorate faster than if it stays in the electronic ice bucket at 4°C.

Once all the solutions I need are ready, I'll start an experiment.

Making an assay and microtubule polymerization

1. The first step I take is to turn on the mercury lamp and setup the microscope for Kohler illumination. I will then make sure that the camera software is ready to take data. The mercury lamp should be on for at least 30 minutes before taking measurements to ensure that it is warmed up and ready for experiments.
2. Once the microscope is setup, I'll make sure that the temperature controller is on and the software associated to it is working properly.
3. While I'm setting up the microscope, I'll ensure that the thermalcycler is on and warming up for polymerization. Once it is, I will take an aliquot of tubulin from the freezer and place it in the thermalcylcer. The tubulin should be in the thermalcycler for a total of 30 minutes.
4. Around 20-25 minutes I will prepare the solution of PEM-T.
5. Once the 30 minutes are up, I will add the PEM-T to the microtubules while in the thermalcycler. I will then remove the tube and protect it from ambient light and store it at room temperature.

Once I have microtubules, I am now ready to start the preparation of a slide for an assay.

1. I add 10 μL of the α-PEM to the flow cell and allow it to sit for 10 minutes.
2. Before the 10 minutes are up, I will dilute 1 μL of kinesin in 9 μL of PEM-αA and store it in the electronic ice bucket. This diluted solution of kinesin is at a concentration of 27.5 μg/mL.
3. Still during the 10 minutes, I will prepare a motility solution. Motility solutions consist of
   • 90.5 μL PEM
   • 1 μL PEM-Glu
   • 1 μL PEM-A
   • 2.5 μL Antifade
   • 0.1 μL Taxol
   • 5 μL of microtubules
4. Once the 10 minutes are up, I will add the diluted kinesin in PEM-αA to the flow cell by fluid exchange. This then is allowed to sit for another 5 minutes.
5. After the 5 minutes are up, I add 10 μL of the motility solution to the flow cell by fluid exchange.
6. I seal the flow cell with nail polish and put it on the microscope to observe the system.

Conclusion

The gliding motility assay is fickle to put it politely. It is full of possible issues that can cause the completed flow cell to not exhibit motility. This is the worst case scenario as it takes anywhere from 1 to 2 hours to produce the first flow cell of the day. Thankfully making another one only take any where from 15-30 minutes. The above recipe works well for me as it has allowed me to create checks with chemical aliquots that help prevent failure of an experiment.

References

1. Ace of Cakes
2. The Adventures of Baron Munchausen paraphrase
3. Ray, S., Meyhöfer, E., Milligan, R., & Howard, J. (1993). Kinesin follows the microtubule's protofilament axis. The Journal of Cell Biology, 121(5), 1083-1093. doi: 10.1083/jcb.121.5.1083.
4. Woehlke, G., Ruby, a K., Hart, C. L., Ly, B., Hom-Booher, N., & Vale, R. D. (1997). Microtubule interaction site of the kinesin motor. Cell, 90(2), 207-16.
5. Böhm, K. J., Steinmetzer, P., Daniel, A., Baum, M., Vater, W., & Unger, E. (1997). Kinesin-driven microtubule motility in the presence of alkaline-earth metal ions: indication for a calcium ion-dependent motility. Cell motility and the cytoskeleton, 37(3), 226-31. doi: 10.1002/(SICI)1097-0169(1997)37:3<226::AID-CM4>3.0.CO;2-4.
6. Fox, P., & McSweeney, P. (1998). Chapter 4: Milk Proteins. Dairy chemistry and biochemistry (1st ed., pp. 146-238). London: Blackie Academic & Professional.
7. Holt, C., Ormrod, I. H., & Thomas, P. C. (1994). Inorganic constituents of milk. V. Ion activity product for calcium phosphate in diffusates prepared from goats’ milk. The Journal of dairy research, 61(3), 423-6. doi: 10.1017/S0022029900030855.
8. Olmsted, J. B., & Borisy, G. G. (1975). Ionic and nucleotide requirements for microtubule polymerization in vitro. Biochemistry, 14(13), 2996-3005. doi: 10.1021/bi00684a032.
9. Böhm, K. J., Stracke, R., & Unger, E. (2000). Speeding up kinesin-driven microtubule gliding in vitro by variation of cofactor composition and physicochemical parameters. Cell biology international, 24(6), 335-41. doi: 10.1006/cbir.1999.0515.
10. Aitken, C. E., Marshall, R. A., & Puglisi, J. D. (2008). An oxygen scavenging system for improvement of dye stability in single-molecule fluorescence experiments. Biophysical journal, 94(5), 1826-35. doi: 10.1529/biophysj.107.117689.
11. Rasnik, I., McKinney, S. a, & Ha, T. (2006). Nonblinking and long-lasting single-molecule fluorescence imaging. Nature methods, 3(11), 891-3. doi: 10.1038/nmeth934.
12. Shelanski, M. L., Gaskin, F., & Cantor, C. R. (1973). Microtubule assembly in the absence of added nucleotides. Proceedings of the National Academy of Sciences of the United States of America, 70(3), 765-8.
13. Keates, R. A. B. (1980). Effects of Glycerol on Microtubule Polymerization Kinetics. Biochemical and Biophysical Research Communications, 97(3), 1163-1169. doi: 10.1016/0006-291X(80)91497-7.
14. Arnal, I., & Wade, R. H. (1995). How does taxol stabilize microtubules?. Current biology, 5(8), 900-8. doi: 10.1016/S0960-9822(95)00180-1.
15. Yvon, a M., Wadsworth, P., & Jordan, M. a. (1999). Taxol suppresses dynamics of individual microtubules in living human tumor cells. Molecular biology of the cell, 10(4), 947-59.
16. Foss, M., Wilcox, B. W. L., Alsop, G. B., & Zhang, D. (2008). Taxol crystals can masquerade as stabilized microtubules. PloS one, 3(1), e1476. doi: 10.1371/journal.pone.0001476.
17. Castro, J. S., Trzaskowski, B., Deymier, P. a, Bucay, J., Adamowicz, L., & Hoying, J. B. (2009). Binding affinity of fluorochromes and fluorescent proteins to Taxol™ crystals. Materials Science and Engineering: C, 29(5), 1609-1615. Elsevier B.V. doi: 10.1016/j.msec.2008.12.026.
18. Castro, J. S., Deymier, P. a, Trzaskowski, B., & Bucay, J. (2010). Heterogeneous and homogeneous nucleation of Taxol crystals in aqueous solutions and gels: effect of tubulin proteins. Colloids and surfaces. B, Biointerfaces, 76(1), 199-206. doi: 10.1016/j.colsurfb.2009.10.033.
19. Passivation paper place holder.
20. Mahamdeh, M., & Schäffer, E. (2009). Optical tweezers with millikelvin precision of temperature-controlled objectives and base-pair resolution. Optics Express, 17(19), 17190. doi: 10.1364/OE.17.017190.
21. Sambrook, J., Russell, D.W. (2001). Molecular cloning : a laboratory manual (3rd ed.). Cold Spring Harbor, N.Y. Cold Spring Harbor Laboratory Press.
22. Alberty, R. A. (1968). Effect of pH and Metal Ion Concentration on the Equilibrium Hydrolysis of Adenosine Triphosphate to Adenosine Diphosphate. The Journal of Biological Chemistry, 243(7), 1337-1343.

---

1. Böhm, KJ, Steinmetzer, P, Daniel, A, Baum, M, Vater, W, & Unger, E (1997). Kinesin-driven microtubule motility in the presence of alkaline-earth metal ions: indication for a calcium ion dependent motility. Cell motility and the cytoskeleton, 37(3), 226-31. doi: 10.1002/(SICI)1097-0169(1997)37:3<226::AID-CM4>3.0.CO;2-4.
2. Böhm, KJ, Stracke, R, Baum, M, Zieren, M, & Unger, E (2000). Effect of temperature on kinesin driven microtubule gliding and kinesin ATPase activity. FEBS letters, 466(1), 59-62. doi: 10.1016/S0014-5793(99)01757-3.
3. Böhm, KJ, Stracke, R, & Unger, E (2000). Speeding up kinesin-driven microtubule gliding in vitro by variation of cofactor composition and physicochemical parameters. Cell biology international, 24(6), 335-41. doi: 10.1006/cbir.1999.0515.
4. Ozeki, T, Verma, V, Uppalapati, M, Suzuki, Y, Nakamura, M, Catchmark, JM, et al. (2009). Surface-bound casein modulates the adsorption and activity of kinesin on SiO2 surfaces. Biophysical journal, 96(8), 3305-18. doi: 10.1016/j.bpj.2008.12.3960.
5. Woehlke, G, Ruby, AK, Hart, CL, Ly, B, Hom-Booher, N, & Vale, RD (1997). Microtubule interaction site of the kinesin motor. Cell, 90(2), 207-16. doi: 10.1016/S0092-8674(00)80329-3.
6. Moorjani, SG, Jia, L, Jackson, TN, & Hancock, WO (2003). Lithographically Patterned Channels Spatially Segregate Kinesin Motor Activity and Effectively Guide Microtubule Movements. Nano Letters, 3(5), 633-637. doi: 10.1021/nl034001b.
7. Hess, H, Clemmens, J, Qin, D, Howard, J, & Vogel, V (2001). Light Controlled Molecular Shuttles Made from Motor Proteins Carrying Cargo on Engineered Surfaces. Nano Letters, 1(5), 235-239. doi: 10.1021/nl015521e.
8. Ray, S, Meyhöfer, E, Milligan, RA, & Howard, J (1993). Kinesin follows the microtubule’s protofilament axis. The Journal of Cell Biology, 121(5), 1083-1093. doi: 10.1083/jcb.121.5.1083.
9. Verma, V, Hancock, WO, & Catchmark, JM (2008). The role of casein in supporting the operation of surface bound kinesin. Journal of biological engineering, 2, 14. doi: 10.1186/1754-1611-2-14.
10. Howard, J., Hudspeth, A. J., & Vale, R. D. (1989). Movement of microtubules by single kinesin molecules. Nature, 342(6246), 154-158. Nature Publishing Group. doi: 10.1038/342154a0. doi: 10.1007/BF00421079.

Quick reference materials list

Below is a quick reference list of the items used in this assay.

Chemicals

• PIPES
• EGTA
• MgCl₂
• NaOH
• Catylase
• Glucose Oxidase
• β-mercaptoethanol
• D-glucose
• Un-labelled tubulin
• Rhodamine labelled tubulin
• GTP
• Glycerol
• Taxol
• DMSO
• α-casein

Equipment

• Microscope
• -80°C freezer
• Hot plate/Stirrer
• Fridge
• -20°C freezer
• pH meter
• Electronic Ice Bucket
• Analytical balance
• Thermal cycler
• Autoclave
• Minicentrifuge
• Barnstead Easypure RoDI

Supplies & Tools

• Pipette tips
• Pipettors
• Microcentrifuge tubes
• Microscope slides
• Microscope slips
• Double stick tape
• Tissues
• Weigh boats
• Weigh paper
• Razor blades
• Box cutter
• PP jars
• Screw top vials
• Centrifuge tubes
• Parafilm
• Beakers
• Gloves
• Chemistry clamps
• Clamp holder
• Support stand
• Condenser
• Flask
• Stir bars
• Syringes
• Syringe filters
• Small and Large freezer storage boxes.
• Desiccator
• Slide box

Need to fix

• W-PEM
  • 1.0 mg/mL of whole casein in PEM.
• [math]\displaystyle{ \alpha }[/math]-PEM
  • 1.0 mg/mL of α casein in PEM.
• [math]\displaystyle{ \beta }[/math]-PEM
  • 1.0 mg/mL of β casein in PEM.
• [math]\displaystyle{ \kappa }[/math]-PEM
• 1.0 mg/mL of κ casein in PEM.
• M-PEM
  • This solution has the appropriate mixture of α-PEM, β-PEM, and κ-PEM in it such that the amount of proteins equal roughly those in the average cow's milk⁽⁶⁾.