Recent Posts

I have tested Universal probes from Roche. Fortunately for one of the genes I had also regular TaqMan probe, synthesized on request. My result is totally diappointing. Most of the uniersal probes did not perform as I expected. Direct comparison of regular TaqMan with Universal probe showed much better result for TaqMan. Finally I had to order normal TaqMan probes for all genes I wanted to analyze.

Since 5 years I intensively use dual labelled TaqMan probes for my qPCR. However there are many other ways to detect PCR product (mol. beacons, scorpion tail probes, 2 oligos etc.). Did anybody make a comparison of any of these new approaches with classical TaqMan?

To generate a plasmid for expression of a recombinant protein can be challenging sometimes. In former times (prior to high fidelity polymerases) one had to find restriction enzymes that allow cloning of the coding sequence or its fragment into an expression vector. Expression levels were not always optimal due to the structure of the plasmid, not optimal Kozak sequence or some other reasons. Now generation of eukaryotic expression plasmids is really simple the strategy of this cloning can be divided into several steps.

1. Choose you favourite cloning vector. I usually use pcDNA3.1 it does relatively well in most of the cases.
2. Download the sequence of your gene from NCBI.
3. Locate coding sequence.
4. Identify restriction endonucleases that are present in pcDNA3.1 MCS and are absent from the coding sequence you want to clone.
5. Find start codon of your sequence (ATG). Check if there is a Kozak sequence around. In the best case it should look like accATGgnn, where ATG is your start codon.
6. If you have Kozak sequence, simply add a restriction site prior to ACC (for example GAATTCACCATGgnn for EcoRI) and order a primer that has 4 bases overhang, restriction site, gene sequence including Kozak, ATG and some coding sequence. The total length of 100% identical sequence should be at lease 15 at max 20.
7. If you do not have Kozak sequence, than simply add ACC in front of your ATG and GCG (Ala) immediately after the ATG. Then proceed as in the step 6 takin into account that your 100% identical part will start after GCG you added.

To be continued....

Strangly enough many researchers experience problems if they need to create deletions of mutations in plasmid DNA. It is really amazing how many help requests I got since I published PCR splicing method on my website.

This approach can be used to delete a fragment of any lenth or to introduce point mutations into a DNA seqence.

To delete a desired fragment from existing DNA fragment all you need is a pair of primers flanking the region where the deletion will be made (primers 1 and 4), 2 compltementary primers comprising a region of -15 bp to +15 bp related to the junction point (primers 2 and 3) and a high fidelity polymerase (a mix of Taq and Pfu for example). The procedure is shown on the picture. It is always helpful to create expected sequence using any sequence editing software first and then choose primers using this "virtual" construct. Following is important for carrying out the experiment:

    * If you use the plasmid as a template use about 500 ng. In this case you can make only 20 cycles to have a good product.
    * Primers 1 and 4 can be any, primers 2 and 3 should be about 30 bases (15 per flank), but you can anyway use annealing at 55°C.
    * Usually no purification of the first stage products is needed, just dilute them 1:100 to reduce the amount of the primers 2 and 3 which you don't need any more.
    * If the deletion is so short that you cant separate deleted and wt products on the gel, you have to gel-purify the products after the 1-st step.
    * Procedure for the point mutation is the same, but selecte primers 2 and 3 related to the mutation point and introduce the mutation in the primer sequence.



Original address of this article.

http://www.methods.info/Methods/Mutagenesis/PCR_splicing.html

This method allows precise analysis of methylation in a certain region by converting all nonmethylated cytosines into tymines, while methylated cytosines remain unchanged. This method requires small amount of genomic DNA and therefore seems to be very useful for the analysis of clinical samples, where the material amount is limited. However I suggest optimising the method using genomic DNA from a cell line and then apply it to valuable samples.

Before starting the experiment you have to develop primers for bisulphite converted DNA. You can generate a model of bisulphite treated DNA by substituting all cytosines which are not in CG context into tymines. And then design your primers in the way that they don not contain any CG. If this is impossible, you have to use C/T at the place of C in CG context. Usually primer selection is the most critical in bisulphite based methylation analysis, since the complexity of DNA is reduced. Therefore I would suggest to select 2-3 pairs of primers, check them on bisulphite modified DNA, and use the most specific ones.

   1.  Isolate genomic DNA with the quality sufficient for restriction enzyme digestion.
   2. Digest DNA (50 to 200 ng) with any enzyme which does not cut the region of interest, but resulting in as short fragments as possible in smallest possible volume. Note: increasing the amount of DNA will make denaturing not efficient enough and therefore make bisulphite reaction incomplete!
   3. Stop the reaction by boiling DNA for 5 min.
   4. Add 10N NaOH to 0.3N final concentration and denature DNA 37° C for 15 min.
   5. Prepare 4-5 eppendorf tubes with cold mineral oil.
   6. Add 2x volume of 2% low melting agarose to the DNA solution, mix by pipetting up and down.
   7. Form agarose beads by pipetting 10 µl aliquots of DNA/agarose mixture into cold mineral oil. Note: Don't pipett the second aliquot in the tube where you already have one bead!
   8. Transfer beads in the tube containing 1 ml of modifying solution (5M sodium bisulphite (2.5M sodium metabisulphite), 100mM Hydroquinon).
   9. Incubate the tubes 4 h at 50° C in the dark.
  10. Wash the beads 6 times for 15 min in TE pH 8.0.
  11. Complete the modification by incubating the beads 2 times for 15 min in 0.2 N NaOH.
  12. Wash the beads 3 times for 15 min in double distilled H2O.
  13. Use one ul of the obtained DNA for PCR with selected primers.
http://www.methods.info/Methods/DNA_methylation/Bisulphite_sequencing.html

This protocol for bisulphite treatment was kindly sent to me by Dr. Toshikazu Ushijima. I was intrigued, since in this protocol you can use relatively high amounts of DNA and there is no need of generating agarose beads – the most critical step in our old protocol. Unfortunately I was not able to test it myself yet, but my colleague, Dr. Natalia Kisseljova from the Russian Cancer Research Centre tested it. She was really impressed by the results, since this protocol leads to almost 100% conversion of nonmethylated cytosines. Even the region of DNA Dr. Kisseljova considers the most difficult, was converted the best when bisulphite treatment protocol Dr. Ushijima suggested. The usage of up to 2 mkg of DNA makes it much easier to amplify your target sequence.
Full protocol at methods.info
http://www.methods.info/Methods/DNA_methylation/Bisulphite_sequencing2.html

Dear methods.info community. Due to technical problems of the provider (or maybe some other reason) I had to re-install the forum. Unfortunately all the data got lost. So we have to start from scratch. Awaiting a lot of activity from old and new users.

Welcome to Simple Machines Forum!

We hope you enjoy using your forum.  If you have any problems, please feel free to ask us for assistance.

Thanks!
Simple Machines