General Solutions

Help for 3'-RACE PCR

PROBLEM: We have prolem to amplify the 3'-end of our interested cDNA by 3'-RACE PCR. The cDNA template was prepared from mRNA by using 3'-general primer and was proved good. But when a 5'-specific primer and this 3'-general primer were used to amplify a DNA fragment of about 0.7kb, the desired DNA product can not be detected on the gel. The nested PCR was also tried by changing 5'-specific primers, but no positive results were obtained due to the 3'-general primer which lower the specificity and was not changable.

If anybody has the successful experience to obtain 3'-end of cDNA by 3'-RACE PCR for cloning, please help us to overcome this problem. We will appreciate it very much.

Something that I've done that seems to help is the following:

Re: LB vs. TB?

I recently started using TB instead of 2xTY to grow 5 ml cultures for alkaline lysis minipreps and I'm quite happy with the results. I don't consider this to be a contradiction to what T.J. Murphy wrote since we could be using different host strains and the details of our alkaline lysis minipreps might vary. For the record, I use TG1 as host strain for most of my plasmid minipreps. My alkaline lysis miniprep is an amalgation of various procedures: alkaline lysis -> isopropanol ppt -> resuspend in RNase soln -> 1 x phenol/CIA extraction -> Sepharose CL-6B or Sephacryl 400 spun column. The spun column removes the digested RNA, is simpler and faster to do than PEG ppt and returns the DNA in TE in the original volume. Yields material of similar quality too. The spun columns are home-made and quite inexpensive. I'll be happy to post a more detailed account if people are interested (it's only yet-another-plasmid-miniprep). I've been using the miniprep DNA for restriction digests, sequencing, subcloning and bacterial transformation. I don't do mammalian cell transfections, so I can't comment on the quality of my miniprep DNA for that purpose.

One published modification that I have incorporated into the alkaline lysis part is the increased acetate concentration to precipitate the chromosomal DNA and RNA after SDS/NaOH treatment. Helge Weissig recently mentioned that reference (Feliciello and Chinali, Anal. Biochem., 212, 394-401, 1993) in response to a different question. I think this modification is even more useful for larger scale (100 ml, liter) preps since it reduces the amount of RNA in the isopropanol pellet and that makes resuspending the isopropanol pellet that much easier.

email: (Song Tan) tan@aeolus.vmsmail.ethz.ch

I've used the following protocol to isolate plasmid DNA when grown the bacteria in TB medium. The quality of DNA is great, I have directly used it for sequen- cing. Here it goes:

The yield of the DNA is appr. 30-40 ug/ 1.5 ml of TB culture, when using pBluescript

Suspension buffer:

25 mM Tris-HCl, pH 8
10 mM EDTA

Lysis solution:

20% SDS 125 ul/2.5 ml
2 M NaOH 250 ul/2.5 ml
dH2O 2.13ml/2.5 ml

Malaine@polaris.utu.fi

Re: differential display

>A rather painstaking analysis of the conditions used in the differential >display technique was recently published in Nucleic Acids Research by Bauer et >al. (vol 21, pp 4272-4280, 1993). These authors have analyzed virtually every >parameter of the technique and have apparently improved it considerably. The >results look impressive. We're fixin' to get into this game fairly soon, but >as yet have no practical experience so take this advice for what it is worth. > >Geoff Kitchingman >St. Jude Children's Research Hospital >Memphis, TN (aka, land of Elvis and bar-b-q) Has anyone tried this modification yet? There is one unclear point in that paper: Is the dT11VN primer from the RT rxn to be removed and added freshly to the PCR rxn or is it carried over?

Re: Cleaning sequencing gel plates

A convenient method for "siliconizing" sequencing plates is to use something called "RainX", a commercially available (Auto supply stores) product intended to be used on your car windshield to bead up water. It costs something like $2 per bottle and it goes a long way. This was featured in a BRL focus a few years ago. It lasts through several cycles of cleaning (up to 7 the focus article stated). It is less toxic than silane or other such chemicals, and doesn't have to be used in a hood. We routinely clan our plates with alconox detergent, rinse with water followed by 70% EtOH.

Best Mutagenesis Strategy --- SUMMARY

Thanks to the quick responses from: Jim Graham: earlier post on the subject by Julian Parkill, Gapped-duplex method by Stanssens et al [ Nuc. Acids. Res. (1989) Vol17 pp 4441-4454 ]. Luiz Oliveira: A simple and efficient method for the oligodeoxyribonu cleotide-directed mutagenesis of double-stranded plasmid DNA. R Jung, MP Scott, LO Oliveira and N Nielsen. Gene, 121 (1992) 17-24. hamel@cc.umanitoba.ca: Similar strategy + modification of Kunkel's. My pre-trial (of course biased) thoughts: 1) Gapped-duplex method: needs somewhat special vectors, still have to make ssDNA. Frequencies of mutagenesis not much improved over Kunkel's ( the only one I had experiences with). 2) Luiz's: needs to start with 1-5 ug plasmid to start with nitrocellulose binding steps may not be very consistent. Frequencies, fair. 3) After looking up couple PCR based strategies, I intend to try the following: a) Modified an inverse PCR protocol by Hemsley, A., et al, Nuc Acids Res, 1989, 17(16):6545-6551 on circular plasmid DNA. Briefly, 2 back-to-back primers near the intended target site running around the circle on dsDNA template to generated linear fragments, then remove the extra A at the 3'-ends with Klenow, then ligate. b) A modification of this is to use restriction sites for the recirculation of the PCR product (Stemmer and Morris, BioTechniques, 1992, 13(2):215.) c) My plan: *circular template (doesn't have to be whole plasmid, don't want to sequence the whole coding region anyway, so may use an insert fragment containing target site in the middle and circularize at the ends.) *Inverse PCR with 2 primers back-to-back (one containing the mutation). *Add some thermostable ligase in the PCR reaction, see if I can get away with ligation/circularization-w/o extra As attached(inserted). *If 1-step PCR/Ligation desen't work, try Klenow or RE type circular protocols. *Then use a fragment containing unique sites flanking the mutation to replace the counterpart in the WT already sitting in the proper plasmid for expression or whatever.

Re: Cleaning a contaminated primer stock

> > By dint of hard work and inexperience our primer stock > solutions have become contaminated probably with a 6kb DNA fragment, > rarther than giveing them a 1 minute silance and a Viking funeral > can anyone suggest a good way to cleanse them of the Foul Interloper > DNA? > > -- > Michael > ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ > NPC rights activist | Nameless Abominations are people too! > You could try 1) UV irradiating an aliquot of your contaminated primer stock, or 2) UV irradiating the PCR mix prior to addition of template, dNTPs, enzyme. I was able to eliminate false positives in nested RT-PCRs by irradiating the PCR mix (minus dNTPs, template & enzyme) in a clear 0.5ml tube for 2min in a UV Stratalinker (Stratgene). This eliminated all false positives. More than 2/3min will reduce product yield so the optimun irradiation time needs to be determined empirically. Also see "Removal of DNA contamination in PCR reagents by UV irradiation" Methods in Enzymology, 218, p381.

Re: Blocking nonspecifics on Southerns-your choice

Victor, When you say the salmon sperm DNA was degraded, what does that mean? I don't think you can effectively block membranes with nucleotides. Typically people use "sheared" or sonicated salmon sperm DNA. What I do to prepare sheared blocking DNA is to drop a known weight of the dry DNA (e.g. from Sigma) into a flask with a volume of TE (10, 1) to produce a 10 mg/ml concentration when dissolved. The flask with undissolved DNA is autoclaved for 20 min, which both dissolves and shears the DNA. This is then used both in prehybridization and hybridization at an approximate concentration of 250 ug/ml. I denature the salmon sperm DNA prior to addition to the buffer, but this is probably somewhat redundant as the autoclave step denatures the DNA. Regardless, blocking DNA ust be denatured. Ours ranges in size from 200-1000 bp. If you're seeing "spots" on your membrane, that's caused by unincorporated 32P. You need to do something (more?) to purify your probes from the unicorporated 32P (e.g. G50 sephadex columns). A general background (not spots; not restricted to lanes) IS generally a blocking problem, in my experience. The buffer I like to use, may not help you much if you're trying to save money, but it works great!! 0.25 M NaPhosphate 0.25 M NaCl 1 mM EDTA 10% PEG-8000 7% SDS 1% BSA The BSA presumably works as a blocking agent also, in this buffer. This buffer is somewhat tricky to make, but it's, by far, the best buffer I've evr used. If anyone wants details I could post them. BTW, I don't think SDS acts to "block" membranes.

Re: Home-brewed T7 RNA polymerase vs Commercial T7 RNA pol

>PS. I hope this doesn't violate any patent. Nonsense. "Owning" enzymes is only a recent proposal. We also routinely purifiy our own T7 polymerase. I will find out the details and get back to you. Lately I have been going with NEB T7 at 3ul/100ul rxn with 3mM NTP's and Hepes-KOH. Using the conditions as described in the Gurevich et al. (Analytical Biochem. 195, 207-213. 1991) and adding 3 G's to the intial 5' end of the transcript, I am getting about 3-10 ug of gel purified RNA from PCR generated 300 bp templates. I would expect a plasmid template to give higher levels if it were practical, from previous preps done by others here. Milligram quantities from a single reaction seems rather far off. However the Ambion procedure uses 7 mM NTPs. Perhaps a corresponding higher (>10 mM) Hepes would be appropriate.