# Talk:Koch Lab:Protocols/Kinesin/Tubulin resuspension and polymerization

## 05-05-2009

Andy Maloney 00:49, 6 May 2009 (EDT): I have a few questions.

• The tubulin dimers are unstable? Are you talking about the $\displaystyle{ \alpha }$- $\displaystyle{ \beta }$-tubulin structure? What sort of magic occurs when you add GTP to tubulin that the tubulin heterodimer becomes stable?
• Steve Koch 01:28, 6 May 2009 (EDT): I know less than I used to, and don't think I ever knew a lot about it. All proteins have some degree of instability, and tubulin is perhaps more towards the unstable side. Basically, I think if you let the tubulin sit around at 4C it becomes less and less able to polymerize into MTs. Whether this is from aggregation or from partial unfolding or whatever, I don't know / don't remember. Here's a phrase from the kinesin home page: "Tubulin is labile and yields will be greatly improved by working rapidly in the initial steps of the preparation."
• Andy Maloney 12:48, 6 May 2009 (EDT): It seems weird that tubulin is okay in cells but not so in vitro. I guess it has to do with the fact that a cell regulates microtubules and tubulin while in vitro, we don't have the stuff in solution to regulate. Nor would we want that stuff in solution.
• In the Non-labeled tubulin resuspension section, why did you use $\displaystyle{ \lambda }$ as a unit? Does it mean something I don't know about yet?
• Steve Koch 01:28, 6 May 2009 (EDT): I use $\displaystyle{ \lambda }$ as shorthand for "microliters." All of my old protocols have it. I noticed today that Kelly does the same thing, so it's probably common enough that we don't need to quit doing it. It was pretty easy for me to get used to.
• Andy Maloney 12:48, 6 May 2009 (EDT): Yeah, I figured as much. But, we have to use $\displaystyle{ \lambda }$/$\displaystyle{ \mu }$ to mean µL/µg from now on.
• I am trying very hard to read some of your procedures and I can't quite understand some things. Is it alright if I put somethings in the language of chemistry, and not biology? In other words, stop naming every thing and say what it is instead of just a name I will forget.
• Steve Koch 01:40, 6 May 2009 (EDT): By "language of chemistry," do you mean saying "5,6-Dichloro-1-beta-D-ribofuranosylbenzimidazole" instead of "DRB"? (BTW: I initiated that wikipedia article, good for me.) Yeah, the acronyms can get crazy, especially sometimes when people use different ones. But, we should probably get used to them, since the people we'll be communicating with use them. I really like what you did with the hyperlinks to explanations / recipes. That's the way to go, I think. Learn what the acronyms mean, but use the acronyms. Or maybe I'm missing your point.
• Andy Maloney 12:48, 6 May 2009 (EDT): I think my understanding the acronyms is a good thing. Not only does it facilitate communication, but it will help me more effectively communicate what I want to say. What I do have a problem with is naming the same thing a new name. For instance, we have a buffer called BRB80. When we add GTP to it, it is transformed to GPEM. I'm not a fan of this and unfortunately it is rampant in biology to name the same thing several different names. Enough bitching...maybe we can stop the "legacy of naming" in our lab. What I wanted to say about the chemistry language is more for changing how you have written the protocols. You say "add x mL to y" without saying the final concentration in the sauce you want. I can understand using protocols for blindly making something but I'm not at the stage where I can do that yet. So, saying "add 1 mM of GTP to BRB80" makes more sense to me than saying "add 1 mL of GTP to 200 mL of BRB80". I can change this if it's okay.
• Steve Koch 16:11, 6 May 2009 (EDT): Yes, change it! It's very good to have final concentration mentioned in the protocol. I think it's good to have both kinds of information, such as "add 2 mL of 100 mM GTP to 198 mL of BRB80 for final concentration of 1 mM GTP." And then links on GTP and BRB80 to recipes for those stock solultions. This is out of place, but I want to point out that there is possibly an issue in the literature about whether to have potassium or sodium counter-ions in BRB80. I believe that potassium is more often used by the leading labs. The buffers cytoskeleton provides are sodium ions. It's easier to make with sodium, because you can buy sodium-PIPES but not potassium-PIPES. I think we should make potassium buffers. Another thing I remember is whether people use MgCl2 for their ATP/GTP or MgSO4. I seem to remember Block lab using MgSO4. Why would these little things matter? I don't know. But it's not unheard of for an enzyme to be picky about those things.
• Does it matter what ratios you use for rhodamine labeled tubulin? The more rhodamine tubulin you use just makes the tubulin brighter in the microscope. Shouldn't I try using different amounts to see what works best? I know from my experience with labeling liposomes, I usually only added 10% fluorescently labeled lipid when I wanted a strong signal and 1% when I wanted to take long exposures.
• Steve Koch 01:40, 6 May 2009 (EDT):Yes, the ratio matters. I read a paper recently, where they use the terms "brightly labeled" for 20% Alexa568-labeled tubulin and "dimly labeled" for 6.7%. Thus, I think your experience / intuition from fluorescent lipids work is a good foundation. I think we should start with 20% TRITC-labeled, based on that work or 10% based on your work. You can find my copy of the paper, with the supplemental methods added in my directory: \\controller\ ... My Documents\References\Kinesin, Microtubules\Plus-end directed\Nature 450_Bieling, Surrey et al_Reconstitution of a plus-end tracking system.pdf In general, we'd prefer to use un-labeled tubulin for studying the kinesin-microtubule interaction, because the fluorophores can disrupt that interaction. Thus, less % TRITC is better and cheaper, but more difficult to see.
• Andy Maloney 12:48, 6 May 2009 (EDT): Great! I just was wondering if there was problems associated with using fluorescently labeled tubulin with kinesin. You know Kiney doesn't like walking on crap down Microtubule Lane. It messes with his groove.
• Steve Koch 16:11, 6 May 2009 (EDT): I definitely worry about it messing with his groove!

## 05-06-2009

Andy Maloney 01:23, 7 May 2009 (EDT): More questions...

• Why do you store GTP in 100 mM MgCl2? Sigma sells the stuff in a Na-GTP form. Did you get it from somewhere that sold it as Mg-GTP? Did you just buy it from Cytoskeleton? Or, did you do ion exchange to make it Mg-GTP from Na-GTP?
• Steve Koch 01:35, 7 May 2009 (EDT): I'll have to think about this one some more...in near term, I think buying 100 mM liquid stocks is the way to go. We don't use a lot and it's better than weighing out yourself.
• We don't have to buy buffers from Cytoskeleton do we? Once we get the water purifier and pH meter, we shouldn't have to. Or, do you want to get the buffers from them anyway?
• Steve Koch 01:35, 7 May 2009 (EDT): I completely agree with you long-term. We'll make most of our own buffers, except in the case where a little can go a long way and it's more reliable / reproducible to use some commercial stuff (such as the GTP mentioned above). In the near-term, I want to buy their buffers so we can try to get things going ASAP.
• Why would you store 1 mL in a 100 mL tube for aliquots? Do they make PCR tubes that big? Is there a reason why they are "thin walled"?
• Steve Koch 01:35, 7 May 2009 (EDT): The should say 1 μL (aka λ) in 100 microliter tube. I've never used a tube smaller than 100 ul, and even those are sort of a bitch to open. The 100's fit well in the thermal cycler (which I like because it's easy to pick a temperature and it has a hot-lid (to prevent condensation)). I used thin-walled because the liquid will heat up more quickly reduce effects that may happen as the liquid heats from 4C to 37C. No idea whether it's really an issue. Thin-walled are tougher to open, and easier to break, but you can get used to it.
• Andy Maloney 02:04, 7 May 2009 (EDT): Argh. I'm already forgetting what that $\displaystyle{ \lambda }$ means... Just so you know (please don't berate me for not conforming) but I'm not going to use that notation.
• How much Taxol did you use? Did you get it from Cytoskeleton? If you did, that comes packed in DMSO. Sigma sells both Taxol and DMSO if we want to make the stuff ourselves. I'm thinking that making our own buffers is good because we won't have to wait to order new stock when we run out. Plus, it will be good for all of us to learn more chemistry in the lab and making our own buffers will help. By the way, DMSO is pretty cool.
• Steve Koch 01:35, 7 May 2009 (EDT): I don't recommend doing it, but Evan says if you get a drop of DMSO on your hand you will immediately taste garlic, just like wikipedia says. Who is validating whom? We definitely need to buy Taxol. Enough to make 100's of mLs of BRB80-T, which has 10 micromolar taxol. Since it's probably at least mildly toxic, I'd prefer buying it already in a suspension of known concentration, rather than having to deal with weighing out powders.

## 05-07-2009

Andy Maloney 23:24, 7 May 2009 (EDT): More follow up questions and statements.

• To follow up with the PIPES comment on 05-05-2009, Sigma sells PIPES in it's raw acid form. It's not soluble in pure water but it is soluble in 100 mM NaOH. I'm pretty sure that the solubility of PIPES is dependent on pH. So, if we wanted to use KOH to make PIPES soluble, we can.
• Steve Koch 00:58, 8 May 2009 (EDT):Yup, that's what I did at Sandia to make the K-PIPES. I have more detailed notes that I'll dig up.
• Steve Koch 01:00, 8 May 2009 (EDT): This is where I'm looking, BTW: (...My Documents\Archived Projects\,Project_Thermomyces_DARPA\Notebook\2003\08_August). (Hopefully can get all my notes mass uploaded to OWW soon, but for now, sorry they're on the server.) You can feel free to do keyword searches in those documents if you're ever looking for info an I'm not available. Of course, asking me is quicker because I have at least a faint memory of what I did back then.
• Steve Koch 01:04, 8 May 2009 (EDT): Actually, I can't really find any more notes other than the Excel recipe which I posted earlier, but which you may not have seen: Koch_Lab:Protocols/Kinesin/BRB80_Acid_Pipes
• Steve Koch 01:20, 8 May 2009 (EDT): I don't trust this data, but I was interested to find that when I tried kinesin heat-induced aggregation studies in K+ versus Na+ BRB80, the K+ aggregated much more quickly. However, it's a pretty tricky assay, so I'm not ready to say that K+ is worse than Na+ (or even that there is a difference). But interesting that I got the opposite result I was expecting: https://writer.zoho.com/public/skoch3/040204_DLS-on-kin797H%2C-various-buffers2
• Steve Koch 01:34, 8 May 2009 (EDT): https://writer.zoho.com/public/skoch3/040210_DLS-kin797h%2C-drosophila-temp-aggregation1 This is a subsequent file, indicating differences are unlikely to be due to the Na+ versus K+ counterion, but rather errors in amounts of EDTA or MgCl2. More interesting is the drosophila kinesin aggregation data, which was never published. Amanda Trent subsequently proved that it was not due to the 10x histidine tag, and thus likely the same motor domain aggregation seen with thermomyces kinesin. This is interesting to me, because it happened even at 40C, and probably lower, indicating that we need to be pretty careful with temperature of kinesin handling.
• Steve Koch 01:43, 8 May 2009 (EDT): OK, I should probably stop now, but I was energized finding this old stuff about kinesin aggregation. Here's a link to an experiment I did looking at the kinesin under DIC microscopy: https://writer.zoho.com/public/skoch3/040212_DIC-Microscopy-of-aggregated-kin797H1 In the heat-induced aggregate sample, I saw lots of clumps, which looked like 1-D polymers to me. But I also saw this in an unheated but old kinesin sample stored at 4C. To me, at the time, and now, this indicates that aggregation is a significant means of kinesin "denaturation" or "instability." It's why people store kinesin in ATP buffers, though I've never seen the aggregation mentioned. It's also pretty interesting to me that the images do look like "beads on a string" that are seen with protofibrils of amyloidosis (although probably on a bigger scale, I guess): http://www.pubmedcentral.nih.gov/articlerender.fcgi?artid=521943
• Taxol is soluble in ethanol as well as in DMSO.
• I know nothing about MgSO4 in buffers. It's soluble in water at least.
• Steve Koch 00:58, 8 May 2009 (EDT): We'll look up a Block Lab recipe.

Steve Koch 00:42, 3 May 2009 (EDT): I found the following note in one of my protocols from 2003: "Note: The cytoskeleton storage buffer has the correct amount of GTP, but does not have glycerol. So adding 2λ to 2 λ actually does not produce the “correct” final glycerol concentration. This seems to be OK, though." Whereas above, I say that the storage buffer does have cushion (glycerol). So, I don't know the answer, but this should be easily answerable by contacting cytoskeleton.

Andy Maloney 01:08, 7 May 2009 (EDT): Some notes about the Cytoskeleton buffers and Koch's above note.

• BST01 is what Cytoskeleton calls PEM but it is just BRB80. I note this because all their buffers for microtubules contain this buffer.
• Koch's note above is talking about Cytoskeleton's BST06 buffer. The one that has GTP but no glycerin. Cytoskeleton calls this GPEM, which is what we are calling it.
• Steve Koch 01:37, 7 May 2009 (EDT): Actually, what I think I meant by this comment is that the tubulin itself was not shipped w/ glycerol. I see now, though, that if we buy the liquid form, we can get it with glycerol. Talking with them on the phone today, though, they said they're moving away from providing the liquid form and trying to sell mostly lyophilized, since it's so much more stable.
• BST05 is Cytoskeleton's GPEM buffer (BRB80 + 1 mM GTP) plus 60% (v/v) glycerin. Koch uses this buffer plus GPEM to dilute the final concentration of glycerin to 6%. Somewhere around 5% glycerin promotes microtubule polymerization, quoted from Cytoskeleton's website.