John Forsberg

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 don't see why doing your PCRs off of a previous product will not work perfectly well.  I've done PCRs off of gel-extracted PCR product bands before (to avoid non-specific priming of my plasmid) in conventional restriction cloning and it worked beautifully.  In your case, your polymerase shouldn't have any problem adding the A overhangs.  You don't have to worry about your previous product being degraded either, as your PCR amplification will select for full-length templates.

I just used 1 µl of my gel extraction (which was probably massive overkill), but you could probably use even less than that to make sure you don't get partial ligations with your "template" that may only have an A overhang on one end.

There's a possibility that your gel will warp if it heats up significantly during transfer, which can distort your protein bands.  It really depends on how hot your transfer buffer gets.  I usually transfer at about 300 mA for 90 minutes, and I never use an ice block because it never gets that hot in the chamber.

Back when we ran transfers at 1 A, I always ran those transfers in the cold room with an ice block though.  Your markers coming out fine suggests to me that your gel didn't heat up excessively.

Based on that papers' calculations, the sizes of a base pair (accounting for loss of a water molecule on each strand in polymerization and a proton from each phosphate hydroxyl due to the pH) should be:

GC: 347.2207+307.1966-(4x1.0079+2x15.9994)-(2x1.0079)= 616.3711
AT: 331.2213+322.2079-(4x1.0079+2x15.9994)-(2x1.0079)= 615.383

average: 615.8771

Note that this reflects the loss of water and a proton from both strands.

Without accounting for the masses lost from the 2 waters or the 2 protons lost from the backbone phosphate, the masses end up being 654.4173 for GC and 653.4292, so an average of 653.9233.  So I could see your original estimate of 660 roughly matching the value obtained if you just counted DNA as if it were a chain of AMP, TMP, GMP and CMP.

So yeah, that paper looks right.


edit: I have found references to masses of DNA with salt bound giving an average mass for a base-pair at roughly 660, so perhaps that's where your number comes from.

In our lab we use Sequencher for aligning our sequences because our university has a site license for it, but you can align the sequences without purchasing software by putting your non-coding strand in a reverse-complement calculator (like http://www.bioinform...s/rev_comp.html).  Then you can use the output in any sequence alignment site you want (BLAST or ClustalW or whatever, although they may handle converting to reverse complement themselves; I didn't check), or you can just look at your forward sequence, find a series of 8 residues or so and search for them in the second sequence.  Once you know how they match up, you just have to copy and paste the extra sequence from your reverse primer reaction onto the end of your forward sequence.

Does that make sense?  I'm sure there are good sequence alignment sites out there that would do this for you, but I'm not familiar with them.

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).

I actually do the exact opposite with inserts and vectors.  If I'm only getting one good strong band from PCR, I usually just digest the insert, then PCR cleanup only to keep away from the contaminants from a gel extraction (I only gel extract if I'm getting multiple bands).  But I always gel extract the cut vector, since you want to avoid getting uncut, or supercoiled DNA in with your ligation at all costs (you don't want single-cut either, but that's usually hard to spot on a gel).  The contaminants I normally get from gel extraction are bad enough that I've re-done my PCR off my gel extracted insert, then run a PCR cleanup off that to keep the insert I use in the ligation as clean as possible.

As bob1 said, you almost surely just have uncut vector contaminating your ligation.  Without a control vector-alone ligation, you have no way of knowing how many colonies is reasonable to screen for insert.  If you don't see more than 2:1 colonies on your +insert plates versus your control plates, I'd consider it a failed ligation and just repeat the cloning from the "last known good" step (or even from the beginning if I'm feeling paranoid).  If you're only at 1.5:1, then you could probably try 10 colonies or something to try to salvage the work you've done, but I'd simultaneously restart the cloning so I can hit the ground running if none of them have insert.

If your cells have a plasmid with a Chloramphenicol resistance marker in it, you'll want to put your transformations on plates with both Ampicillin (I definitely prefer Carbenicillin, as I get fewer satellite colonies) and Chloramphenicol.  The RIPL plasmid in your BL21 cells is there to help with expressing proteins with codons that are rare in E. coli, so if you're expressing a eukaryotic protein, you'll definitely want to select for cells that keep that plasmid too.

We usually use 30 µg/ml Chloramphenicol and 100 µg/ml Carbenicillin for LB plates.

What detection method are you using for visualizing the antibody chains?

Using secondary antibodies on Western blots will vary wildly on how well they bind to the heavy and light chains, depending on whether the secondary antibody is a monoclonal or polyclonal and where its epitopes are.  I wouldn't expect quantitative measures of heavy:light using antibodies.

For Coomassie staining, assuming an equal amount of staining per gram of protein for the heavy and light chains, I'd guess offhand that the ratio should be about 2:1 heavy:light, as the heavy chain is usually around 50 kDa, and the light chain is usually around 25 kDa (forgive me if my numbers are a bit off, I don't remember exactly).  This is also assuming that the staining is linear in the concentration range of the proteins in the gel (I'm not sure offhand if this is the case, although I suspect it probably is).  Coomassie binding may be somewhat sequence-specific also, but I'm not sure how much it varies.

I've used the Miniblotter for probing sets of hybridoma supernatants for screening during monoclonal antibody production.

Generally speaking, it works best if you have one molecular weight standard lane to one side, then one big well running across the rest of the gel that you load with your protein sample/lysate.  That way you can cover as many of the Miniblotter wells as possible without worrying about whether a particular well is binding in the gap between lanes.

If you pour your own gels, you can simply take a piece of Scotch tape and tape around all of the wells but one, so when your gel solidifies, it leaves one long well instead of several.  You just load your sample in one huge volume instead of several smaller ones.  I usually ended up using about 50 µg per normal lane, or about 500 µg across a whole long lane.

Then when you line up your blot in the Miniblotter, you can line up your antibody wells with your large lane.

As we use commercial pre-cast gels most of the time, I just got the single-well pre-cast gels from Invitrogen, then cut a lane divider off another regular gel, and wedged it down between the 1-well gel plates to make a quick and dirty version.  You could probably also get away with cutting the lane dividers out from a regular gel (tear them out with a pipette tip?).

Alternatively, if you're set on using regular gels for your probing, you could probe with a removable stain like Ponceau S to reveal where your proteins are bound to the PVDF, then mounting it in the Miniblotter and aligning it using the stained areas.  Then you could wash the wells to remove the stain and probe with your antibodies.

One thing I found with the Miniblotter: make sure you have your antibody in block solution.  I always got nasty background on any antibody probings that were simply hybridoma supernatants without adding to 5% BSA (even after blocking the blot).  Not sure if you normally probe in block, but I figured I'd warn you anyway.

Hope that helps.

Ooh, for imaging, I'll usually have it in pretty deep water/liquid in my tray, then I'll either try to get both (gloved) hands under the edges to lift it out all at once or I'll tease up an edge with some filter forceps, then lift it enough with one hand to get the other hand under it.  That way the force of moving it is more distributed and less likely to tear.  I wet my gloves before handling gels to help prevent sticking (you should be able to easily slide your finger over the gel without it grabbing).

As for on the imager, I'll usually coat the surface with water first, then set the gel in that one edge first, then easing it down to prevent bubbles.

For getting your gel off the plates, you could also try holding the plate over your container, then teasing a little piece of edge away, then doing the water bottle trick.

One thing you could try is using a water squirt bottle to soak the gel while it's on one of the plates until an edge lifts up, then trying to use the water stream to kind of wedge the gel into your container for imaging.  Or just put the whole plate in with your gel, which might soften up how strongly the gel sticks to your plate, then gently teasing the gel off with a spatula/forceps after incubating/rocking for a bit.

I'm assuming you're using your own gels on glass plates?