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There have been 5 items by vinylrooster (Search limited from 15-April 20)
10g NZ Amine (casein hydrolysate)
5 g yeast extract
5 g NaCL
add h20 up to 1 L, adjust pH to 7.5 with NaOH, and then autoclave.
Prior to use, I add filter sterilized in a hood:
12.5ml 1M MgCl2
12.5 ml of 1M MgSO4
10 ml of 2M glucose
I used it one time by opening quickly and taking what I needed and immediately resealing (all done in open air). A few days later I noticed that it was a little cloudy so I remade it exactly as described above, but this time when I aliquoted some I took it into the hood, sprayed everything with EtOH, had brand new gloves sprayed with EtOH and only had bottle open for 5 seconds. Again, about a day or two later I noticed it is cloudy again! Is this media very very very easy to contaminate, or is it some ingredient (maybe the glucose) that is making the liquid cloudy?? It's driving me a little crazy because I obviously cannot remake this every time I want to transform! Tonight, I will aliquot 5 mL into one falcon tube and shake 220RPM/37deg/overnight and a separate 5ml into another falcon tube which I will let sit on my lab bench. Tomorrow I will check to see if the one that I shook is more cloudy, which I would assume means that my stock is indeed contaminated?? Any help is appreciated, as always!
SalI is a known problem enzyme when cutting near the end of PCR products. If you can, switch to a different enzyme. There must be something wrong with your Qiagen column cleanup. You should never get 260/280 ratios that high. Does your wash buffer PE have EtOH? This could be where all of your DNA is going. You might want to wash columns first with buffer PB, then twice with PE followed by elution.
I would avoid AP digestion. The double digestion should be enough to inhibit religation. If it isn't, then I would change enzymes (see SalI above). Make sure you are doing your digestions in large enough volume, where the DNA added is a small fraction of the total digestion volume. This eliminates many problems from restriction enzyme inhibitors.
I originally wanted to avoid SalI as well because my post-doc has had bad experiences with it, but my PI insisted that we use it so I am kind of stuck with SalI whether I like it or not. Yes, my PE has EtOH and I do wash with PB, but only once with PE so I can try a double wash step with PE. I also tried doing without AP treatment and I got 50+ colonies on both my 'no ligase' and 'no insert' control plates, so I decided I must do AP (which confuses me a little bit, because I am obviously doing non-symmetric digestions and should not get self-ligation, but that is a story for a different day). The fact of the matter remains that I get plenty of supposedly positive colonies on my experimental plates and yet none of them appear to be what I want. The way I look at it is that my bacteria should only be able to grow on Amp if one of the following three things happens:
(1) uptake of empty vector
(2) uptake of vector with insert
(3) uptake of vector with something besides my insert
In any of the three cases my UNDIGESTED plasmid prep should run on a gel slower than my original vector, yet EVERY TIME my undigested are running quicker than my empty vector and thus are smaller than my vector? This doesn't appear to make sense if you follow my logic of only the above 3 possibilities. Thanks for your advice Phage and I will definitely use your hints and am welcome to any more you can think of!
I am a new graduate student who is currently having some troubles with my cloning. In particular, one discussion is very similar to my current problem. The discussion is here:
Persoanlly, I have a 6.9kB insert (generated from PCR) that I am trying to put into a 7kB vector (originally 5.6 PGK vector which I successfully put a 1.5kB insert into already and verified via sequecing thus 5.6+1.5=~7kB vector I am now trying to put my second insert--the 6.9kB one--into). Okay now for the details:
Insert is PCR generated, then gel purified with Qiagen kit, then digested with SalI O/N, then EtOH precipitated, then digested with NheI O/N, then gel purified via Qiagen kit which gives me a 260/280 ratio ~4.0 so I EtOH precipitate and lower the ratio to ~2.1 and concentation to ~13ng/uL in 30uL EB.. I start with ~9.4ng of DNA immediately after PCR/1st gel purification, but by the time I get to the 2nd gel purification, right before the ligation, I only have ~400ng! I've concluded that the columns seem to be bad so I have ordered low-melting Agarose and plan to do the 'Molecular Cloning' book Gel Extraction method next time to hopefully improve my yields. Also, during all of my gel extractions I use long band UV light and never expose for more than 20 seconds (normally 2-3 second burst, then turn off and cut, and repeat to get everything)
Vector is generated from maxiprep of previous clone that has my 7kb vector, then digest O/N with SalI (at same time as insert), then EtOH precipitated, and then digested O/N with NheI O/N (same time as insert), and then gel purified with Qiagen kit which gives me a 260/280 ratio of ~5.3 so I EtOH precipitate and lower the ratio to ~2.1 and concentation to ~13ng/uL in 30uL EB.
Also, throughout this whole process my insert and vector maintain 260/280 ratios between 1.8-2.1 except when I noted above.
Okay, after all of this I ligate using NEB T4 DNA ligase using a 1:1 ratio, 1:3 ratio, 'no insert' control, and 'no ligase' control, and PGK positive control for 30mins at room temp and then transform my cells. I originally tried DH5alhpa and they didn't work so I'll just explain the Xl-10 GOLD, which appeared to work. So, the XL-10 GOLD gave which gave me over 100 colonies on all plates, including the negative controls (I originally thought I didn't have to AP treat because my digestion is non-symetrical but obviously I had to). So, I took my vector, AP-treated for 45 mins, then heat inactivated, and then re-ligated for 30 mins at room temp. Then I transformed my XL-10 GOLD cells in the same way as before and got about 50-100 colonies on my the 1:1 and 1:3 plates, and absolutely zero colonies on both negative controls and plenty of colonies on the PGK positive control. So, I did miniprep and then test digestions (O/N) using single cutter enzymes on ~20 colonies and I get the following bands:
Original vector uncut - I get one band ~3kb and two bands greater than 12kB
Original vector NheI (single cutter)- I get one band ~7kb and two bands greater than 12kB
Original vector NotI (single cutter) - I get one band ~7kb and two bands greater than 12kB
Original vector SacII (double cutter which has an internal cut site in my insert as well as on my vector and for which I expect bands to come up around 3.6kB and 10.4kB if my insert is present and obviously will only cut vector if no insert is present so ~7kB band expect for empty vectors) -I get one band ~7kb and two bands greater than 12kB
ALL of those are correct and I get exactly what I expected....But, for all my minipreped clones I get weird results:
For some clones I get these bands:
Clone X uncut - I get multiple bands, some faint around 13 and 14 kB, one distinct band around 3kB
Clone X NheI (single cutter)- Get all the same bands as uncut but now distinct bands at 3kB AND 7 kB
Clone X NotI (single cutter) - Only one distinct band at ~5 kB
Clone X SacII (double cutter explained above) - Only one distinct band at ~4kB
these 'type' of clones confuse me, but I am pretty sure they are just empty vectors??
For other types of clones I get this pattern:
Clone Y uncut - Only one distinct band around 1.5kB
Clone Y NheI (single cutter)- Only one distinct band around 1.5kB
Clone Y NotI (single cutter) - Only one distinct band around 1.5kB
Clone Y SacII (double cutter explained above) - Only one distinct band at ~3.5kB
I've included a picture of my gel with a few representative colonies (I realize it is only 4 clones, but I have screen over 20 and get similar results, these are just representative of the what I get in general)
I selected these 4 colonies in particular because I did a colony PCR on 20 minipreped colonies using my primers from the original insert PCR and got faint bands at ~7kb (size of my insert) for the first three clones in the picture (two CLONE X and one 'similar to CLONE X') and one VERY distinct band at 7kB for CLONE Y. Obviously this was prior to me digesting and why I chose to digest these clones with so many enzymes at the same time.
Also, immediately after I transformed the cells with my AP-treated '30 minute room temperature' ligation cocktail, I took the ligation mixtures and put at 16deg cel O/N and then froze down in -20deg. I plan on doing a transformation with the now 'overnight 16deg' ligation mixtures tonight using XL-10 just to see what I get. Please please offer me any advice that you can give, I am quite desperate...I am a new grad student in a large lab and I fear my PI will begin to look down upon me if this does not start working soon! Also, I would like to note that we are planning to using this vector to build a new knock-in mouse that is based off of a knock-out mouse and I therefore must start with the orignal PGK vector which I inherited from the knock-out mouse. I am fairly certain that my insert is unique to humans and thus not present in bacterial genome, but I will ask some others in lab to find out for sure! This was quite the long post, but I wanted to make sure I got my whole story out there in the most detail so I could get the best help.... Thanks in advance for any advice you can offer!!