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Why was one part of insert missing? - (Aug/11/2008 )

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Dear all,

I am trying to ligate a 2.2kb insert with 10kb vector. The insert is from another plasmid after double PCR. I used BamHI and NheI double digestion for the vector and the insert. And then I ligated them, I got the colonies, the vector control was empty. But I checked about 20 colonies, there was only one I got the right band with another enzyme digestion. And then I checked all the enzymes that I could find restriction sites in the insert. most of them were right but some of them were wrong. So I was confused and then sent it to sequence. The result showed the same as my digestion results. About 600bp of the insert lost.

I have some problem with gel extraction. I could not get any pure product from gel extraction. So I used purification kit to purify the vector and the insert instead of running on the gel and then extraction. I know that would be some problem. I digested the vector with NheI, and add CIAP, and then purify. And then digested with BamHI, add CIAP, and then purify. The part between NheI and BamHI is only 20bp. I think it would be cleaned out during purification. The problem I was thinking is that CIAP might destroy the ends. ??? Because I didn't inactivate it after digestion. And also because I used double PCR, so I got a 500bp band all the time in my PCR product. But I didn't run the gel and also only purify it with the kit. I wonder if it would disturb the ligation either.

Do you think I need to use CIAP? (I don't know how to calculate and I am sure that I used too much of it.)

Because the BamHI has very strong star activity I can find if I use them together, so I have to digest sequently. And after the first digestion, I would lose some of the product. And after the second digestion, I would lose more. And if I run the gel, I probably could not get very pure product. The ration of 260/280 was only 1.40 in my previous ones and I gave up this method. Do you think the low purity would affect ligation a lot? or, I can still use the purification kit??

I don't know why some part of the insert would disappear. And I also checked the insert after double digestion, it was intact. That means something happened during ligation and transformation.

I have done it over one month. I don't know what I should do now.

Thank you very much.


there a two reason why a large segment of an PCR amplified insert can go missing

1- a restriction site was not taken into account.
When you conducted the restriction digest on the PCR amplified product, there was an extract restriction site within the PCR product. Consequently your PCR product was cut short.

2- error in PCR amplification
PCR always favours the shortest PCR product. Consequently there will always be short PCR products products. Occasionally this problem can be quite severe. This is why you also gel purify the inserts before cloning.


To perneseblue: I would disagree with your first point. uterus says at the end that he/she checked the insert after digestion and it was intact.

To uterus:
1 - A quick fix to get your clone: In the first paragraph you say that you got the colonies, checked 20, one of them had an insert. If you have any more colonies left, you could quickly screen them by a crack gel and identify more of those with inserts. I could dig out a good cracking protocol for you.

2. - Here is where I agree with perneseblue: After the PCR you need to run a gel and cut out the right band. Otherwise your PCR byproduct will compete for the vector with your insert.

3. - Your restriction schemes: Double digestion with BamHI NheI is OK. As long as it is not longer than 30 - 60 min. By running so many restriction-purification rounds you are loosing DNA. Also, why CAIP? Phosphatase treatment is only necessary for a vector that could lock onto itself.

Finally, torturing yourself for a month is not a good thing. Go have an ice cream break, try one of the suggestions and ask more questions.


QUOTE (Andriy @ Aug 11 2008, 07:13 PM)
To perneseblue: I would disagree with your first point. uterus says at the end that he/she checked the insert after digestion and it was intact.

Ah, my mistake. sleep.gif I missed that last sentence. I though that there was no description of the treatment of the insert.

And back to question, aside from losing vector DNA from multiple rounds of DNA purification as mentioned by Andriy, over exposure to CIP will damage your DNA ends. CIP treatment can sometimes be more trouble than it is worth. Which in this case, isn't necessary.

I find it odd that BamHI is giving trouble. You might want to increase the volume of your digest or decrease the amount of BamHI enzyme you are using. The total volume of restriction enzyme should not exceed 5% of the total volume of the restriction digest. The glycerol that the enzymes come in are inhibitory to enzymatic action and does encourage star activity.

I am also curious about the insert. I am unclear about the nature of the insert. Is it a PCR product? How was it digested? Sequentially?


I am really a newbie in molecular biology. I have many problems in every step. Thank you very much.

Today, I will try double digestion again and do gel extraction. My supervisor said he didn't use CIAP as regular way. Well, let me try his way. And also, I should definitely get out of the byproduct from the gel.

Andriy, I have no idea with 'cracking protocol'. Could you tell me in detail? Thank you.

perneseblue, the major part of the insert is in plasmid. And then I need to add another two parts to it. So I run two PCR.


QUOTE (uterus @ Aug 12 2008, 07:50 AM)
perneseblue, the major part of the insert is in plasmid. And then I need to add another two parts to it. So I run two PCR.

So you add your restriction sites with PCR to the fragment?
Did you keep in mind to add some additional base pairs to your primers (additionally to the restriction sites)? Otherwise you won't be able to amplify the whole restriction site and your enzymes won't cut.

I would run the PCR, put it onto a gele, double digest it, purify it with a column and then ligate it.

@Andriy: Can you use your cracking protocol to distinguish 10 and 12 kb? Maybe you have a better protocol, mine I can only use for smaller plasmids. Can you share it with us, please?



Gentlemen, thanks for your interest in the cracking protocol! biggrin.gif

Usage: Tough cloning situations when every colony matters.

Sensitivity: The size difference between parental and expected clone has to be in the range of 4 kb vs 4.3 kb. Not 4 kb vs 4.1 kb.

To Vista: Yes, it will distinguish between 10 and 11 kb. Run 0.7% agarose gel.

---------------Colony Cracking:----------------------
A. Cell suspension
1. Put 20 ul of water into an eppendorf tube (one per colony)
2. Touch a colony with a 01-10 ul pipette tip. Put a tip with the tube with water. Repeat for each colony
2a. For control, use a colony from control plate or 1 ul of parental plasmid.
3. Vortex at low speed to swirl the cells off into water.
4. Take the pipette tip out, touch a fresh LB plate. Tips go into garbage, plate goes into 37C incubator so the next day I have replicas of all good colonies.

B. Add 20 ul of crack buffer to each of the tubes.
Crack buffer: 2 ml 1M NaOH, 2 ml 10% SDS, 400 ul 0.5M EDTA, water to 40 ml.
No vortexing or pipette mixing at this step. Just drop it in there and proceed immediately to C.

C. Add 20 ul of loading dye (60% glycerol in TAE with bromophenol blue).

D. Run a gel.
Note: If you added to many cells in the step A, the final mix might try to float out of the wells. Use wide combs.

Picture: Here is how the gel eventually looks like. Note genomic DNA band at the top and RNA at the bottom.
Plasmid size: 4.5 vs 5.2 kb


Dear people:

We have a trouble with sequence analysis that intrigue us:

A 585 bp fragment was successfully TA cloned from mouse mRNA into a pcDNA3.1/NT-GFP-TOPO vector, as the PCR analysis shows. The 6762 bp supercoiled construction was sent to sequence with a primer which hybridizes exactly at the 3' end of the cloned fragment. All the sequence seems perfect from 3' to the 5' end of the cloned insert, and shows the correct nucleotides of the vector following the 5' end of the insert (we get the expected sequence some tens of bp forward of the primer site, as usually). The problem is that a 22 bp sequence including the 16 bp of the 5' primer used for the original cloning is missing. We repeated the process two more times with the same results. In the other hand, the PCR analysis of the clone amplifies the expected 585 bp by using the same primers for the original cloning, including the 5' primer whose sequence disappeared from the sequencing. The restriction of the sequenced clones releases an expected fragment for a correct construction (it doesn’t analyses the 22 bp missing region; it releases a fragment that includes the whole insert).

The cloning was repeated onto a pENTR/D-TOPO, and then recombined with a pLenti4/TO/V5-DEST vector. A 669 bp fragment was PCR amplified using a different 5' primer than the one used in the pcDNA3.1 cloning (hybridizes plasmid backbone), but the same 3' primer. The 5' primer that hybridizes to the 5' end of this fragment was used to sequence at the same facility as the pcDNA3.1 was. All the sequence of the insert and surrounding vector's nucleotides are perfect, but almost the same 22 bp missing in the pcDNA3.1 sequencing disappeared again: the pLenti construction lacks of only 20 bp of the 22 missing nucleotides of the pcDNA3.1. The next two expected cytosines (bases 21-22 of the pcDNA3.1 22 bp missing fragment) are substituted in the analysis by two thymines. The expected CGG immediately before the start of the 5' oligo (where the sequence begins missing) are changed by GGC. However, the PCR (that uses the 5' primer which included into the 22 bp absent sequence) was perfectly amplified.

The missing sequence (in the pLenti vector) is (5'-3'): ATGGATACAGCACCTGCATC. It has not very much CG, but forms some loop (as shown by the OLIGO software).

Is it possibly to have some secondary structure that avoids the sequencing polymerase to pass through, but allows ours?

Our major interest is to know the region of the fusion of the insert and the vector, and know the correct ORF of our insert and the GFP one (GFP is not properly part of the pLenti vector, but we’re talking here as if it were).

Thanks in advance. Any comment would be greatly appreciated



If I am reading that correctly you need to use a sequencing primer about 50 -100 bases upstream of where your region of interest starts not at the 5' end of the region of interest as sequencing results will not show this.


Hello, all!

Yes, stevo, I forgot some detail: the 5' primer we used to sequence the last time, was also used to amplify a 669 bp PCR fragment enclosing the complete insert (585 bp) into the pLenti vector; this 669 bp product was the one sent to sequence. The closer tail of this primer is 70 nucleotides upstream of the begin of the 5' missing end of the insert. The sequence analysis, as usually, and as you tell, shows only 58 of the last bases between the sequencing primer, and the insert; the 58 bp include the end of the GFP sequence, as well as a 8 bp connector fragment. The 58 bp are shown almost in a perfect way: just the immediate CGG before the missing 20 bp 5’ end of the insert (that start with the 16 bp of the primer used to amplify the former 585 bp insert) are changed by GGC. Then, the sequence shows our insert at position 21, and the first CC are substituted by TT, but all the next nucleotides are in perfect correspondence with the insert. So, the sequence just modifies those two small fragments, but keeps the correct number of nucleotides, except for the 20 bp missing 5’ end of the insert.

Well, we’re guessing about the nature of the construction since the sequence analysis shows the lack of 20 bp of our insert, including the 16 bp of the 5’ primer used to amplify the 585 bp originally cloned, while later, the PCRs still perfectly amplify the 585 bp fragment, using the 16 bp primer which the sequencing demonstrate is not there.

Comments, suggestions?

Thanks everybody. Thanks stevo!



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