Sorry for taking me ages to reply. I have been on vacation.
beads experiment: take the pellet from 50 mL of culture, resuspend in 5 mL of buffer, sonicate, spin at highest centrifugation speed your table centrifuge allows (at 4oC) and then incubate 2 mL of supernatant for ~2h with 20 uL of affinity beads (equilibrated in the proper buffer). Then wash the beads 3-5 times with the buffer while centrifugating at low speed for 2 min in between washes. Load this on the SDS-PAGE.
Interaction partners: well, it might not work as your professor thinks. You need a very stable interaction and if you have a transient interaction as in regulatory pathways, these are not stable complexes (trust me, I also have the nightmare of transient complexes in my PhD thesis). Moreover, even if you have a stable interaction, if you have more than 150 mM salt (NaCl or KCl) in the binding buffer, this complex might disassemble. Furthermore, if you have to elute the protein from the column, you need to add imidazole, which is a salt and will increase the total amount of salt in the buffer so your interaction partner will dissociate before your his-tagged protein.
Your frozen pellets are good forever...ok not forever, but until the end of your project as long as it does not last 10 years. So, yes you can use the ones you have in the freezer.
As I've never used those markers before, I'm not sure I'm confident of the band I'm pointing out, but if your markers are what I think they are I'd say your protein is the band in the induced total and soluble lanes that is just above the third obvious marker from the bottom.
I've attached the band I'm thinking is correct, and what I think the markers are. I can't be positive, as I usually use multicolored markers, and I usually get a feel for how the markers run on a given gel type over time, so I can tell at a glance which markers are which.
I'll also note that with many protein expressions that I do, I do not see my protein in the total extract at all, but only see it after enrichment after binding to affinity resin and eluting (for many His-tagged proteins in particular). So you may want to try a trial run of your first purification step to see if you get a more prominent band at the right size.
I am also confused myself. Is that a 15% gel? I guess it must be if you want to see your protein of 16 kDa Anyhow, what I can tell you is that sometimes in the uninduced fractions you could also see your protein when you have leaky expression which can happen as T7 from pET system is a very strong promoter and it is very difficult to inhibit before expression is induced. I would do this experiment with the beads that I have suggested above, like this you can observe whether the enriched fraction contains a protein around your size. I would guess that the pink band did not separate enough and that you have from bottom to top: 6, 15 and so on and the top marker bands that run all together contain the pink band as well. I would not continue until I see the enriched band on the gel from the beads experiment. However, you could also try direct large scale expression. Most people around me are lucky enough for it to work from the first trial
For the first part: I usually measure undiluted, but it works like you too; actually, if you are trying to get OD600 = 1 then you should dilute. I induce myself at 0.6
For the second part: you can keep your pellets also at -20 (in case you need space in your -80; I surely always have this problem)
I always resuspend the pellet in 5 mL of buffer and take 100 uL of this + 50 uL 3X loading buffer (load only 3 uL on a gel because it is very concentrated); sonicate these 5 mL and spin down for 30 min at the max speed of the centrifuge I have at 4oC; take 100 from supernatant and treat as above; from the pellet, resuspend it in 2 mL of water and take 100 uL and treat as above; the rest of the 5 mL of the soluble fraction I incubate with 20 uL affinity beads for my tag on rotation in the cold room for 1-2 h and then wash 3-5 times and then boil the beads in sample buffer and load 10 uL of this on a gel.
When you're transforming bacteria from purified plasmid DNA, you should get tons of colonies (unless your DNA or cells are poor quality).
There shouldn't be too much colony-to-colony variability in expression, so you can pick any colony and it should be okay. I usually inoculate 2 colonies in 2 tubes whenever doing a new growth in case one of them doesn't grow (for whatever reason). I would avoid any colonies that look significantly different from the others (may be a sign of contamination).
Loading buffer I prepare and aliquot in 1 mL Eppis at -20; the rest I keep at RT. I never prepare gels ahead of time and then store them. People do that and keep cast gels in the fridge but over time the H+ gradient between the stacking and the resolving (which are usually at different pHs) will diffuse and the resolution of the gels will be sub-optimal.
PMSF is a serine protease inhibitor only, there are quite a number of other proteases (cysteine, metallo, aspartic) out there that won't be inhibited by PMSF at all. I recommend using a cocktail tablet such as Complete from Roche or any suitable one from Sigma-Aldrich. Dirt on some wells would affect your reading by not allowing all the light into the well thereby getting a reduced reading at the end. You could just put an empty plate in the reader and see how it comes out at your wavelengths.
Freeze/thaw of any protein is a good way to degrade activity typically. If you need to do these assays, it is best to make sure that all the lysates have been treated in the same manner, this includes the number of times you have freeze/thawed the lysate. Storage at -80 is also recommended.
Green things in the lysate could well be chlorophyll or chloroplasts (or as you say, bits of leaf), these could absorb in the red spectrum (660 nm is red) but shouldn't absorb in the green (505 nm), though discrete particles could scatter the light which might account for a lower reading than in the blank.
Ok. That is a bit of a problem. I was assuming that you weren't using an extraction buffer control (and I did mean extraction buffer when I talked about lysis buffer). In the case that my assumption was true, the negative absorbances would be due to the buffer being less absorbent than water at the read wavelength.
Would it be possible for there to be something in the plants that might inhibit the reaction (proteases perhaps)?
Thanks, that was very clear - what you actually need to do is very simple - you need to compare your samples to a standard composed of pure forms or known activities of lysate containing the relevant enzyme (or a similar one) that you can set as the relative 100% strength. Using water as a 100% is not ideal for this sort of thing as it should actually have an activity of 0 (presumably). The water should be treated as a blank to set no activity, it would be better to use the lysis buffer as the blank rather than water, as this will affect the absorbance to some extent.
The negative values you got are probably from the lysis buffer.
You got it all right with the culture, as I explained.
Do not worry about how different people do the same thing in many ways. Choose whatever works for you, and stick to it.
I do 50-100 mL of preculture. Depends on what flask I have available I usually make 1/10 of the volume of the flask i.e. for 500 mL flask I do 50 mL preculture.
Usually, I measure OD600 in a cuvette not with the microplate reader (too complicated and takes too long); I just take 1 mL culture, put in a cuvette and stick it in the spectrophotomer. Moreover, measure the culture every 1 hour or even 30 min. Keep in mind that the duplication time of BL21 derived bacteria is 20 min (around, depends on what protein you are expressing) so if you are measuring every 2 hours you get 0.1 and next point is >6.
I usually do 6 L of main induced culture but you need the flasks and shaker capacity for this kind of expression and BTW, I do X-ray crystallography so I need tons of my protein Another important aspect: I use 1/5 of the volume of the flask for expression i.e. in a 5 L flask, I put 1 L of media. This is needed for good aeration.
And another thing: before I waste my time with 6 L of culture, I make an expression test in 100-200 mL of culture from which I take several time points 50 mL and harvest the cells, resuspend in 5 mL, brake them with a sonicator, and run a gel with pellet, soluble protein (supernatant of the disrupted cells) and I also incubate for 1-2 hours on rotation 2 mL of supernatant = soluble protein solution with 20 uL of the affinity beads i.e. Ni-sepharose for His-tagged proteins to get a sample of enriched protein (for the SDS-PAGE, I wash these beads 3 times at low centrifugation speed and then boil them in loading buffer)