Life of a Lab Rat

A UK-based reader writes

I'm a UK molecular biologist with DNA oriented experience who is about to try a western blot for the first time. Are you willing to share your protocol?

To be honest, it's pretty much the standard protocol. The antibody isn't fantastic (although that might be a function of my protein, of course) and I used to get a lot of background, but here's a quick guide to getting this particular antibody against this particular epitope in these particular samples to work (standard disclaimers, caveat emptor, this advice is worth exactly what you pay for it, etc. ad nauseam). I've emphasized the points that needed to be optimized:

1. Prepare protein from cells (it was a nuclear protein, so I used this method up to and including step 6)
2. Run gel (I used Invitrogen NuPAGE pre-cast Bis-Tris gels in MOPSMES running buffer)
3. Soak gel in Towbin buffer (Tris/Glycine/20 % MeOH) for 10 - 15 minutes
4. Blot onto PVDF at 200 mA for 2h at 4°C
5. Block in 3 - 6% milk powder in PBS/0.02% Tween20 overnight at 4°C
6. Wash membrane 3 – 6 x 5 minutes in PBS/0.1% Tween20 (PBS-T)
7. Incubate in primary antibody for 3 – 4 hours at room temperature (you'll have to work out the optimal concentration yourself)
8. Wash in PBS-T
9. Incubate in secondary antibody diluted 1/20,000, no more than 30 minutes
10. Wash 6 x 5 minutes in PBS-T
11. Develop signal with Pierce Femto substrate (1.5 ml each reagent per membrane) for 1 minute. Pat dry. Leave 5 minutes for signal to burn off a bit
12. Expose and develop film, whoop loudly.

The primary and secondary antibody diluent is PBS/0.05% BSA.

My friendly reader also said

I have to develop protocols for antibody purification- possibly involving formaldehyde crosslinking- can you give me any advice?

and I do have a protocol for that, here in fact, and am happy to answer questions on it.

As promised, my really rather simple miniprep method.

I should again point out that the GTE, NaOH/SDS and KOAc are functionally the same as the lysis reagents that you get in the commercial kits. So you can substitute in either direction.

As promised, this is the method I currently use for making maxiprep DNA.

I learned this method from an American in the lab where I did my DPhil. At the time, if I wanted to transfect eukaryotic cells I had to perform a double caesium chloride prep based on isopycnic ultracentrifugation.

That was as completely horrible as it sounds.

Marc introduced me to this method. It uses phenol, which is a bit scary (if you know what you're doing the risk is minimal), but produces superb quality plasmid DNA that can be transfected into cells with no further purification. And naturally you can use it for digests, transformation of bacteria, etc. It's cheaper than any commercial kit, probably faster, and certainly more robust. For the multi-tasking (or just plain lazy. . .) scientist, it is full of breakpoints where you can leave the prep and go and do something else, rather than being tied to the bench watching a column drip.

I couldn't do my job without it. Thanks, Marc.

I have been meaning to have a whinge about the scientific kit culture for more than a little while now.

Last year some time, I used a Qiagen miniprep kit (what can I say? I was in a hurry) and found that some noddy had added the blue dye to one of the reagents. Now, some of you will know what I'm talking about, and require no further explanation. The rest of you; seriously, you're better off not knowing.

The point is. . . well, no. Let's step back a bit.

Just over eight years ago I left a DNA extraction technologies company for reasons that do not concern us here. Eighteen months before that I had completed my first project at that company, which was to revamp a range of DNA extraction kits — minipreps, gel extraction, that sort of thing. I compared my efforts with those of the market leader, Qiagen (Promega and the rest, sorry, but even if your stuff is better it's not the benchmark). And even if I do say so myself, I did a pretty good job. My miniprep kit was faster, gave better yield, as good if not better quality, and was cheaper to produce.

It was, naturally (like the rest of the company down the line), torpedoed by a worse than useless sales and marketing department, and intellectually challenged management.

But I realized, last year as I used the Qiagen kit, that in more than eight years the market leader had failed to innovate at all. Minipreps are pretty critical to molecular biology research, and they still charge a 500% markup for something that, frankly, is crap.

Qiagen's idea of innovation is to add a blue dye, so that people can tell that something is mixed thoroughly. Which is bizarre to say the least; doing minipreps is one of the first things you will learn in a mol biol lab, and if you can't get that right first go you're going to have much more fundamental problems. Qiagen did nick one of my ideas, which was to add a collection tube to the kit. Now, I persuaded management that such a tube in the kit would be a good thing, and moreover, it should have a lid with an extended hinge so that it would actually fasten over the spin column while in the microfuge. Such a tube exists (and existed back then), so we just bought a job lot and packaged them with our stuff.

So Qiagen can't even get that right: Their collection tube is un-lidded, so you still have to fart around with different tubes at the end of the prep. To be fair, they did drop the five minutes on ice thing from the protocol, so I guess someone must be awake there.

Now, you might think I'm being hard on Qiagen, and you'd be right, but over the weekend I received the following (edited) email from my mate Tiddles:

Qiagen Maxi-Kit. first run = 1.3µg DNA/µl = ace. 4 times since ~0.3µg DNA/µl = a fornicating waste of my time. Fornicate. Fornicate. Fornicate. Fornicating fornicating kits. I fornicating hate fornicating kits. I have changed kit, changed solution, re-transformed my plasmid and still bollocks. Do you know what they think it is? I actually agree. . . It's growing too fornicating well and thus clogging their miracle fornicating filter system. What the fornication? Growing too well? What am I supposed to do? Put fornicating bleach in my media?

It's expensive, time-consuming, uses an imperial shedload of plastics, doesn't actually yield all that much DNA and, as m'friend Tiddles points out, can not cope with rich media. So to prep a lot of DNA for, say, multiple transfections, you need to do multiple preps. And if you try to use a decent amount of culture (like 101 ml instead of 100 ml) or rich media (2xTY) instead of that anæmic LB muck, the (bloody expensive) column throws up its beads in despair and fails miserably. Just like the Qiagen miniprep, in fact.

Y'see, back in '98, my miniprep kit could cope with buckets of rich media and still spew out loads of top-quality DNA, so I am very down on Qiagen for not coming up with something better.

But so that you know that I can do so much more than just whinge, my very next post will contain a link to the protocol that I currently use for maxiprep DNA. It makes loads, uses phenol, and gives you the feeling that you're doing some real chemistry. Great stuff.

I don't know if this is a good idea, whether people will like it, or even if I have the energy/inspiration to keep it up, but I thought a 'Weekly Protocol' feature might work.

Let's see how we go. I'll start with this one, for making splicing competent nuclear extracts. Cobbled together from a number of different sources, it's surprisingly easy and straightforward.

Printable: http://blogs.usyd.edu.au/labrats/pdfs/Nuclearextract.pdf