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FluffyMember Since 18 May 2011
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Posted phage434 on 10 August 2012 - 05:13 AM
Posted Papaver on 19 July 2012 - 10:03 PM
I believe the buffer has ATP as one of its ingredients and it is as old as the enzyme. I am thinking of getting a new one Friday ( an enzyme with a new buffer to be extra careful, I might get the 10X so that way I will stick to the 1.5uL)
Also for my transformation I do incubate it on ice for 5 min before heat shock. After heat shock I incubate again on ice for 2 min. Also according to my protocol after that I add 80uL SOC medium to ensure better spreading, and then I put on 37 at 250rpm speed for 1 hour. Thought I should be more clear
So if I add SOC medium do you think I should still add LB medium ( I am guessing it has to have ampicillin antibiotic)?
Thanks again dear!
SOC Medium is even better than LB, and for both: no antibiotics before plating the cells on LB/Amp-plates.
You can always play a bit around with your transformation mix. Incubation longer on ice before heat shock would give you cells more time to "get in contact" with the DNA. You can also add more of your ligation mix...1 µl is a small amount.
You can also plate more...
Posted Papaver on 19 July 2012 - 06:46 AM
so first...if you buffer is 5 x , you would use 2 µl ; final concentrations should always be 1 x. You should also check if the buffer contains ATP (ligation reaction depends on it), otherwise you have to add it to the reaction. If the buffer is as old (and often reused) as the enzyme it might be better to add ATP.
Do you incubate your transformation mix prior to heat shock? If not, do this for around 30 min. After heat shock, add ~ 1 ml LB medium and incubate for at least one hour. I often incubate two. Try different amounts for plating...say 50 µl, 100 µl, 200 µl. Then pellet the remaining cells, remove a bit of LB and resupend the pellet in the remaining ~ 150 µl of LB. So you would plate all your cells. But I would do these "plating series" only for your ligation reaction
Considering your colonies you got with the DD-plasmid...maybe some plasmids weren't double digested in your reaction and so they self ligated...if there are only 5 I would not worry...it can always happen that you get colonies with empty plasmid even if you do a double-digestion cloning.
Posted Papaver on 12 July 2012 - 10:12 AM
there is no need to worry. The concentrations after gel extractions are often that low but it's absolutely enough for ligation.
Use around 40 ng of your double digested vector, that's a good concentration to work with...less would be also ok. The molar ratio of vector vs. insert should be 1:3 to 1:5. pET14b is ~ 4.6 kb, your fragment ~0.5 kb. So I would use 2.5 µl of your vector, 5.5 µl of your insert, 1.5 µl of ligase buffer (10x) and 0.5 µl ligase (I usually set up this reaction without using additional water and fill up final concentration with vector and/or insert). For transformation I would use 5 µl.
I'm only wondering why you digested your vector sequentially. If both enzymes are compatible with the buffer you can do this in one reaction as you have done with the Topo vector. I have also never dephosphorylated my vector before...even not when I digested with only one enzyme (but that's my way
But anyways...just keep doing you stuff....that's fine
Posted Papaver on 05 July 2012 - 10:25 PM
I don't think it's weird..
Your EcoRI hasn't digested the Plasmids completely...except No. 9 of the digested sample. There you can see you fragment very nice and the plasmid part show only one rich band. The other digestions also show your insert although the bands are more or less weak. Furthermore you can see that all undigested plasmids move to the same size which implies that all contain the insert...
So everything is ok. I would take the plasmid/clone of the digestion No 9 and go ahead with the work. If possible sequence your insert to make sure that there are no mutations in.
Posted phage434 on 02 July 2012 - 11:28 AM
Posted Papaver on 02 July 2012 - 11:01 AM
Yes! Or, you look at the reverse complement...your complement sequence is from 3' to 5' and the EcoRI site is there if you read from 3' to 5' which is not correct and leads to misunderstandings
Altogether...your insert has no EcoRI site...that's why your fragment in the digestion is ~ 500 kb and not smaller...it would be if there was a EcoRI site.
Good luck for your further experiments.
Posted phage434 on 02 July 2012 - 05:26 AM
Posted Papaver on 01 July 2012 - 10:19 PM
Posted Papaver on 01 July 2012 - 07:42 AM
Posted Papaver on 30 June 2012 - 04:39 AM
Every digested Plasmid shows a fragment of ~ 500 kb which fits to your information of the first post.
Considering multiple bands: In general I would say that all except No. 3 (with two bands) are not completely digested since I don not exactly know if control three and digested three are the same. On the other side the fragment of No. 3 is a bit smaller than the others.
So I would take No 3 and one of the others and sequence them to make sure everything is right.
Now some last words to your restriction protocol...
400 ng are sometimes not enough...the smaller the fragment the more difficult it is to see the band in the gel. I once had to deal with that...I couldn't see my insert (it was ~ 300 kb) and then I ran a PCR and I got it. Since then I always try to use at least 500 ng, better 600. Usually the companies supply a 10x buffer so 2 µl of buffer would be too much in a 10 µl reaction and 1 µl enzyme is also too much. It can cause unspecific cuts, especially with EcoRI.
A good set up would be (in my opinion) for your DNA:
6 µl DNA (à 100 ng/µl)
2 µl EcoRI-buffer (10x)
0.5 µl EcoRI
11.5 µl water
--> 37°C for 1-2 h (or less if it is a fast digest enzyme)
Then load as much as possible onto the gel.
Posted Papaver on 27 June 2012 - 10:31 PM
The 1 kb ladder of which company? They always differ a little bit...
Posted Papaver on 27 June 2012 - 10:17 AM
the marker is not described so I cannot say anything about the size of your fragments. Also the Topo-Fragment is not well detected...I can, at least guess, where the band could be.
By the way, it is also helpful to load the undigested Plasmid to compare the sizes.
I would say the digestion worked but only because the first then lanes look similar and the bands are located in the 400 bp area of a general agarose gel
Posted allynspear on 07 October 2011 - 08:55 AM
As far as similarity and identity goes, for nucleotide alignment, there is no difference unless your sequence has ambiguous characters like Y, H, W, N, etc. For protein alignments, identity means exactly the same amino acid in a position, but similarity can mean amino acids with similar properties:
Serine and Threonine: Both hydrophillic, hydroxyl
Isoleucine and Leucine: Both hydrophobic, aliphatic
Aspartic acid and Glutamic acid: Both hydrophillic, acidic
When you get a protein alignment, repost and people can take a look.
Best of Luck.
Posted Adrian K on 27 August 2011 - 12:53 AM
There are six lanes, For band number 2 &3, there are 2 bands.
Maybe I am wrong, but I do not find that your band sizes were corresponded with the size you wanted. it is like 1.2kb and 1.8kb. Which brand of 1kb ladder you use, can provide the catalog number as well as a reference?
Let's say the band you wanted is the bottom band, I suggest you directly just pool all your samples, run it in agarose gel, do a gel purification and elution and finally do a cloning into your desired vector. There is no need to waste so much time on PCR optimization.
Is hard to say in this stage that whether your cDNA is degraded or whether you should increase it. I suggest you try to use your genomic DNA as control and see if there is any differences, provided your primer was not designed on the splicing site. Also, addition of Betaine or DMSO might help to eliminate the unspecificity binding and perhaps reduce smear. This is not the issue of primer dimmer.
Thanks for your reply. Much appreciated. I am sorry for my questions, I noticed you mentioned you work long hours in the lab. I am really sorry if I am being a bother.
The ladder I am using is a 1kb ladder from Invitrogen and its Cat # is 15615-016.
Also I do notice two bands in each lane, you are right, but I thought the second band on the bottom would be my right size, since the in the ladder the bright band is 500bp and right above it would be close to 585bp?
OK so you think gel purification is a better idea than PCR clean up using the PCR sample itself, rather than gel?
And when you say genomic DNA do you mean I order a DNA extract of my gene only and use that as a positive control with my primers?
I was thinking of using the DMSO and or formamide today but I thought I should receive some feedback from the forum
But really thanks alot, you are great help
Haha, don't don't apologize, is not your fault. When I plan my work, I forget that almost the next whole week is my country's holiday, and I can't use most of the instruments. Most of the staff will be absent, and thus the office closed.. I got no choice but to stay back for "lab marathon".
Yes, gel-excised purification is always better as it only cut out and isolate the band you wanted. Sorry for make you confuse in my previous post. The shortcut (a.k.a cheat) I was trying to say is you can use a PCR clean up spin column, load all of the remaining PCR products from your gradient PCR (would be roughly 15ul from each tube times 6 equals to around 90ul total volume), do a clean up and elute the product (using around 40ul elution buffer). After the elution, load all the eluted product onto agarose gel, run it, and gel-excised-purified the band you required (your case, cut only the 585bp band).
Alternatively, you just load all your ~90ul product onto the agarose gel, run the electrophoresis and proceed with gel-excised-purification for the 585bp band. During your elution step, try elute with less than 40ul elution buffer (30ul would be even better as it 3x concentrated your initial volume of ~90ul). Use the product for your cloning purposes.
My suggestions is mean for you to speed up your work rather than just focus on PCR optimization. Those were the few tricks I use for my work, yet to fail me for the almost 30 different clones I did. Just imagine the time you required for optimizing 30 independent PCR, unless you are using for endpoint detection.
However for your cloning purposes, as phage434 mentioned earlier, you might not get it done due to those RE required extra 5-6 bases. I suggest you do a TA / Blunt end cloning first, send for sequencing and after your new primers arrived, you PCR it again using the cloned plasmid, so that you do not need to use your cDNA again.
For the PCR addictives, I never heard of using "formamide". The addictives commonly used is betaine, DMSO and BSA.
Hope this helps.