So, here goes...
Week 4 (seems a very long time ago now!):
This week I did more of some of the stuff I’ve already talked about – I set up some more transfections in tissue culture, transformed and plated out some bacterial cells and started a new scheme of PCR mutagenesis.
The main new things were immunoprecipitation and western blotting, which I’ve learnt about but have never actually done before. We took a batch of cells I had transfected, spun them down and resuspended in lysis buffer, sonicated to aid the lysis, and then spun down again. We then measured the protein concentration with a Bradford assay, and added the correct volume of lysate to FLAG resin (which had been thoroughly washed). This was then left to incubate, the idea being that the anti-FLAG resin will bind to the FLAG-tag on the HDAC binding partner. If the binding partner is able to interact with the HDAC, it will form a complex and the HDAC will also be ‘pulled out’ when we isolate the beads. So, we’re hoping that the mutants don’t interact, or show impaired interaction, with the binding partner, proving the importance of the key mutated residues.
After incubation, the beads are washed several times and they are added to SDS sample buffer for loading onto a protein gel. I discovered that loading samples onto a polyacrylamide gel is not really a skill of mine...I was missing the wells completely at first!
Having run the gel, we then did a Western blot to transfer the proteins to a nitrocellulose membrane for antibody probing. This was another fiddly technique – an air-bubble- free sandwich needs to be made with the gel adjacent to the membrane between blotting paper and foam, and then all clipped into a cassette which can be submerged in the blotting apparatus. The gels we use are pre-made, and very thin and flimsy – I’m a bit scared of touching them as they have a tendency to fall apart!
The transfer was successful, so we then moved on to the antibody probing – we split the membrane to probe different areas with different antibodies. The blots are incubated with primary and then secondary (HRP-conjugated) antibody, with many important rounds of washing in blocking buffer in-between (our blocking buffer is made from milk powder (!) and PBS). It’s a bit strange seeing powdered milk sitting on a shelf in a lab!
Finally, the blot can be visualised with the HRP substrates, and then captured on photographic film and developed.
The blot was a mixed success really – some of the results were what we expected, others not quite.
Week 5:
This week I finished the new round of PCR mutagenesis I had started – this involved setting up the PCR reactions with appropriate primers and templates, running a sample of the product on the gel to check it had worked, and then using the product as a template for the next round. It was relatively problem-free, with only one round needing to be repeated – I’ve sent the final product off for sequencing now, so we’ll soon see if it worked!
I also helped to set up some HDAC assays to look at the effect of some of the mutations we had made on the enzymatic activity of the HDAC. The assay uses a substrate with an acetylated lysine, and the principle is that the more functional HDAC your sample contains, the more of this substrate will be deacetylated by it. The deacetylated substrate can then react with a developer solution to give a fluorescent product. So, more HDAC activity leads to a higher fluorescence reading. The set-up of the assay involved a bit of maths to work out how much of each component needed to be made up, and then a 96-well plate was used to create the separate drops with different mutant samples. The results we got were quite good – as expected, the mutants showed much lower deacetylase activity than the wild type (indicated by a much lower fluorescence reading obtained by a plate reader).
Week 6 - protein prep:
This week my supervisor was away, and so I worked with one of the PhD students, purifying out some proteins from bacterial cells which we had previously transformed and induced protein expression in. We had 3 different bacterial pellets in the freezer, each of which contained a different expressed protein, and the process of getting the final purified protein samples from these pellets took the whole week.
The proteins were all His tagged, so we followed a basic protocol for purifying out His-tagged proteins. I had to make up lots of buffers first (lysis buffer, wash buffer, elution buffer, dialysis buffer) and then followed the protocol to try and purify out the proteins.
First the cells were lysed by resuspending in lysis buffer and emulsifying, and then the samples were centrifuged to remove debris and the lysate was incubated with a nickel slurry (nickel binds the His-tag on the desired protein). The incubated protein-nickel mixture is then passed down a column and washed with wash buffer several times (lots of watching things drip...) and finally the protein is eluted by adding a buffer containing a high concentration of imidazole (which competes with the tagged protein for binding to the nickel).
A protease was then added to cleave off the His tag, and the sample was then dialysed overnight to remove small molecules such as salts. Setting this up was a bit fiddly – the sample is pipetted into tubing, which needs to be double-clipped at each end to make sure no protein can leak out - I saw the importance of this when one morning I found one of the clips had come off (luckily the second was still in place!). Then the dialysis bag is left spinning gently in the buffer overnight.
Finally, the protein is removed from the dialysis bag and further purified by gel filtration. The principle of this is that proteins travel down a column containing lots of small beads, and small proteins can pass through these beads whereas larger proteins can’t. So, smaller proteins take longer to pass down the column due to taking a longer path through all of the beads, and different sizes of protein therefore come off the column at different times (larger ones first). Glass tubes collect these different fractions, and a trace is obtained which shows you which fractions contain protein. You can then run a gel using samples from the tubes which give peaks on the spectrum to confirm that a particular peak indicates presence of your desired protein.
The whole purification process took about a day and a half per protein, and there were a lot of new techniques for me. So, the PhD student supervised me closely for the first one, but as I kept repeating the techniques I got the hang of it and by the end was doing a lot of it myself. I’ve also got much better at loading the protein gels! (Although there was one slight mishap, where I didn’t add enough buffer and so the current didn’t run properly, with the result of a very interesting looking smiling gel with very distorted lanes...). At the end, we concentrated down all of the proteins by spinning through a concentrator tube containing a membrane which retains proteins above a particular size, and then finally we measured the protein concentration. We found that we had obtained a very large amount of one of the proteins (which we knew we would, as we got massive chunky bands on the gels we’d run) and much smaller amounts of the other 2. From the gels we ran of the products, it looks like this is because only one of the proteins actually expressed properly – unfortunately the other 2 didn’t really express and were full of contaminants. Never mind, we can always try again....
So, that's about it - I'd be surprised if anyone's actually read this far what with the length of this post! Just shows how busy the last few weeks have been....only 2 more to go now!
