User:Andy Maloney/Notebook/Lab Notebook of Andy Maloney/2009/09/21/The chemistry of kinesin and microtubule buffers
Understanding how to store and handle the chemicals used for kinesin and microtubule experiments is crucial to being able to track down problems that may occur. As everyone that works with kinesin and microtubules will say, things can and will go wrong with the experiments if the experimenter is not careful with their assays. In order to alleviate issues with the experiment that I have been having, I have taught myself some important chemical realities that need to be addressed with what we have done and ways that we can alleviate variables in the experiments.
I will structure this page first with how I clean the items used to prepare samples and buffers. I will then go over and link to as many references as possible the chemicals used in the experiments. I will list what I find as the most valuable information for handling these chemicals and their proper storage and possibly their disposal. Finally, I will go over the buffers used for the experiments and what I think is the best way to prepare and store them.
Any and all questions or comments should be directed to the talk page. Please do comment on this page as I would like to ensure that no one has to go through the same mistakes that I have gone through. Also, if anyone has a better, more efficient methodology for preparing any of the buffers we use, please comment on the talk page. This is open science after all and you shouldn't feel afraid to voice your opinion!
Preparation of items used
To ensure that the items I make are free from unwanted contaminants, I clean everything excessively. Almost to the point of ludicrousness. All steps are repeated for each item used and at any point during the process of making something you need to reuse an item, such as a spoonula used to scoop a different chemical, that item is cleaned before it touches anything else. My process is as follows:
- Clean the item with a sponge and a 1% solution of Alconox.
- Rinse in the faucet for about 30 seconds to ensure that all soap is gone.
- Repeat steps 1 and 2. Excessive but it must be done.
- After rinsing in faucet water, proceed to the 18.2 MΩ water dispenser and rinse with this water. Thankfully the water that comes from our dispenser is UV treated and filtered with a 0.2 µm filter. So, we can feel confident that there are no bugs in the water. I typically rinse an item 10 times. This means that if you are rinsing a beaker, you need to rinse said beaker with 10 volumes of water. Yep, crazy but I do it.
- After rinsing in pure water, proceed to dry the item with nitrogen.
I shouldn't have to say this but,
Always wear gloves!
I don't care if you are just cleaning something in the sink, you need to wear gloves to prevent any thing from your hands getting into or on the item you are cleaning. What I typically do between the sink and ultra pure water station is dry my gloved hands and then rinse my hands in the ultra pure water. I then rinse the item I am interested in. Something else I shouldn't have to say is
Never touch what you have cleaned with your hands or your gloved hands.
Try to handle the item in a way so that you don't destroy all your hard work of cleaning. Koch mentioned to me that he has had lots of discussions with previous experimenters about cleaning items. One thing that he did say is to make sure that what ever you clean doesn't produce bubbles when rinsed. If it does, then you have not rinsed away all the soap. In my procedure I rinse my items 10 times which ensures that no soap remains. However, making sure there are none is a quick way to determine if what you cleaned is soap free.
As a final step, you can autoclave the items you just cleaned. Sometimes it makes sense to just autoclave items, such as microcentrifuge tubes or pipet tips, without having to go through the cleaning process.
Weigh boats and weigh paper
This is slightly tricky because lots of people in the lab handle the weigh boats and weigh paper. I'm almost convinced that you cannot trust that every lab member will handle the boats and paper with the same care that an individual would. So, I'm going to go on a limb and say that each person that uses these items, needs to have their own stock. That way you can keep people from using yours and you can handle them in the manner you feel fit to.
Some things to note about these items is that you should never ever pick one up without...guess...clean gloves/or a fresh pair of gloves on. Don't do it because you risk contaminating the stock. One thing to note about the weigh boats is that there will always be plastic splinters along the sides. To remove these splinters, go around the boat with your gloved hand to remove them. This way you won't have any in your solutions.
I have not go so far as to clean the boats and paper in the same way I do glassware but, if things don't work this time around, you better believe that I'm going to start.
This is tough to consider as well. Koch says that the best way to get chemicals from the stock is to use a weigh boat and dump out some in it and take from that. Once you have the amount you need, you toss the rest that might be left in the boat. I'm beginning to believe that this is the best way as well. The reasoning behind this is that if for some reason you didn't clean the spoonula well enough, you will not contaminate the stock.
There are problems with this. Specifically for really expensive chemicals. If you dump out half your stock inadvertently, then that's it, you have to get rid of it. More thought needs to be considered when dealing with expensive chemicals.
Below is a list of the chemicals we use and a link to their product page.
- Rhodamine tubulin
- Glucose oxidase
Somethings to note about PIPES is that it is an acid. Don't get this stuff on your skin because it will irritate it. We use acid PIPES because there was a discussion with Koch about what type of ions are needed in solution for motility to work. Some say that you need potassium and others, sodium. Since we have a free acid form of PIPES, we can add either NaOH or KOH to pH the buffer to PIPES's pKa of 6.89 thus adding the appropriate counter ion we want.
PIPES should be stored in the desiccator at room temperature in its original container.
EGTA is also an acid so be careful. The reason we use the acid form is for the same reason we use the acid form of PIPES. It should be stored in its original container in the desiccator.
MgCl2 is very hygroscopic. This means it really likes to be in water and it will pull moisture from the atmosphere so it can be in it. For this reason, we use a 1 M solution of MgCl2 in water. This way we do not have to fuss around with trying to weigh out an appropriate amount of MgCl2, since what we weigh would also include water.
MgCl2 should be stored in its original container at room temperature. You do not have to desiccate it since the MgCl2 is in solution already.
NaOH is again hygroscopic and thus likes to pull moisture from the atmosphere. I store it in its original container and in the desiccator. When ever I open the jar and close it, I blow in nitrogen so it is stored in a nitrogen rich environment (i.e. no moisture). It is a strong base so be careful when handling it. I like to make up 1 Normal solutions of this and use it in this form. Normal is in bold because chemists like to change molar to normal when dealing with strong acids and bases.
When making a 1 N solution, I typically store it in a glass jar. I sometimes will also make up a 12 N solution and dilute it from that but, this is not necessary.
Glycerol is a form of sugar. It should be stored in its original container and I would think it should be flushed with nitrogen as well.
We were previously using Cytoskeleton's tubulin storage buffer that has glycerol in it. Unfortunately I do not trust what we got from Cytoskeleton because Cytoskeleton is just littered with typos which make me nervous. From now on any solutions that need glycerol in it will come from the EMD stock.
Some things to note about glycerol is that it is very viscous. Almost to the point where trying to use a pipet tip to get some out is pointless. Since our stock is greater than 99% pure, I think weighing it out is a better bet than trying to pipet it out.
This is a sugar and has the consistency of powdered sugar. Nothing special here except that it should be stored in its original container at room temperature. It is hygroscopic and the stuff we have is anhydrous (without water) but I think that it has been opened for long enough for the stuff in the bottle to be full of water now. Oh well. This is where proper storage in a desiccated environment would be best.
ATP & GTP
Originally we purchased ATP and GTP from Cytoskeleton. Unfortunately Cytoskeleton doesn't tell us what exactly it is we purchased and they have mixed protocols about how to put ATP and GTP into solution. So, building on my belief that I should know everything about all the things I use so that it makes my life easier to hunt down problems, I purchased more from Sigma. We got the magnesium salt kind of ATP because for our assays to work, we need magnesium in solution. We got the sodium salt version of GTP.
Since we have salt forms for both ATP and GTP, they should go into solution very easily. Right now I am not sure if they should go into our PEM buffer, or if they should be reconstituted in pure water. I do know that the GTP needs to be in at least a 2 mM solution of MgCl2. At any rate, I have both stored in a secondary container that is filled with desiccant and under a nitrogen environment in the -20˚C freezer. I also wrapped Parafilm around the cap to ensure that no moisture gets in.
This we purchased from Cytoskeleton and it says to store at -20˚C. Right now I have them in the -80˚C freezer and I am not sure if this has ruined it or not. Sigma says to store it from 2-8˚C so there is absolutely no consensus here on the storage temperature. Sigma does say that stored in DMSO, Taxol is stable for "several" months. Oddly enough it was my belief that a solution with Taxol in it caused all my woes. I wonder if it is because of me putting it in the -80˚C freezer. I can't see why but it may be the case.
What's nice about getting this from Cytoskeleton is that it is already aliquoted in convenient sizes. I'm pretty sure I do not want to mess with weighing out Taxol at all.
Some things to note about Taxol is that it is very toxic. You definitely do not want to handle anything with this stuff in it without gloves.
Since Taxol is soluble in DMSO, we have to have this stuff. DMSO is hygroscopic and reacts with just about everything. Originally we got this from Cytoskeleton however, I found out that the way we were storing it may have caused me problems. The stuff we got from Cytoskeleton is in the -80˚C freezer and the stuff we got from Sigma is in the desiccator.
Sigma packs DMSO in ampules. Since DMSO likes just about everything, figuring out how to store it in something other than the ampules has been an interesting road. It turns out that DMSO can be stored in HDPE, LDPE, and PP without any reaction. Thankfully, the cryo vials from Nalgene are made of HDPE and PP. I have the DMSO from one ampule in 3 cryo vials. These cryo vials are then in a secondary HDPE container filled with desiccant in a nitrogen environment. The jar then has Parafilm around it as well. This is then going to be stored in the desiccator.
When handling this stuff, you need to be crazy careful. Apparently if you get it on your skin you will taste garlic almost immediately. But that won't happen because you are wearing gloves.
This comes as a lyophilized powder. It should be stored in its original container in the -20˚C freezer. Other than that, there isn't very much to talk about.
We really have 2 types of casein. A technical grade and vitamin free grade. I'm not clear on the differences of these caseins. They should be stored at room temperature in their original containers.
One thing to note about the whole caseins is that they are very difficult to get into solution. Sigma says to dissolve them in a 1 molar NaOH solution. Since we need the casein to be in PEM at pH 6.89, this is not a great idea. So, I have devised another method. I have successfully gotten casein to go into our PEM buffer by stirring it and heating it to 60˚C for about 30 minutes. Everything I have read about casein is that it does not have a tertiary structure and thus will not denature. Why it does this is beyond my ability to understand at this moment but, it does work and we don't have to use a strong base to dissolve it.
As for the shelf life of this stuff, I can't find one.
This is an anti-caner drug. It should be handled with extreme care and never ever handled without gloves. It is insoluble in water and thus needs to be reconstituted in DMSO. One of the good things that comes from Cytoskeleton is that it they aliquot Taxol in vials that are stored in our -80˚C freezer. Just add DMSO and your Taxol solution is ready to go.
I cannot find anything that states that our Taxol will go bad in a -80˚C freezer. It is lyophilized from Cytoskeleton so I cannot see it going bad in there. I do think that the DMSO is bad.
We have two types of tubulin both from Cytoskeleton. Both are made from bovine brain and one is labeled with rhodamine. Tubulin is very unstable and should never leave the freezer unless you are aliquoting or polymerizing.
For a compilation of what other people do for their buffers, please visit here.
This is the base buffer used for the kinesin and microtubule experiments. Please see this page for a description behind the naming conventions.
Since we use acid PIPES and acid EGTA, we need either NaOH or KOH to get both PIPES and EGTA into solution. If you prepare a stock solution of 1 N NaOH, then all you have to do is add the 1 N NaOH to get the chemicals into solution. The procedure to make this buffer is as follows.
- Weigh out the appropriate amount of PIPES and EGTA.
- Add them to a fraction of the total volume of the buffer. Remember acids go into water.
- Add approximately 125 mM NaOH from the 1 N stock solution. I say approximate because it is. There really is no way of saying definitively how much to add because of weighing errors of the PIPES and EGTA. You need to add enough to get both the PIPES and EGTA into solution but at the same time, not too much so that you make the pH too basic. I would start with an amount just below 125 mM and adjust from there.
- Add ultra pure water to almost the volume you need.
- pH to 6.89.
- Add the remaining water.
There you have it. The base buffer used in all other buffers for the experiments.
This is PEM with 10 µM Taxol in solution. To make this buffer,
- Follow the recipe for PEM.
- Dissolve the Taxol in DMSO. Following Cytoskeleton's procedure, you will make a 2 mM stock.
- Add Taxol to make a 10 µM solution of Taxol in PEM. You don't have to make a special solution of PEM to add Taxol to to prevent dilution of the PEM. This is because adding the amount of Taxol needed to make a 1.5 mL stock of PEM-T only dilutes the PEM by a factor of 0.5% which is okay.
Taxol can be dissolved in DMSO to a concentration of almost 60 mM. I think I might try doing this with the Taxol we have just so that I don't have to add as much to the PEM buffer when making PEM-T.
Actually, I am going to do this. Instead of adding 100 µL of DMSO as Cytoskeleton suggests, I am going to add 20 µL of DMSO. This will give me a total concentration of Taxol of 10 mM. That means that I will be diluting PEM by 0.1% with this stuff which is 5 times better than before.