Your band is residual RNA with strong secondary structures which are resistant to RNAse A. To get rid off it:

1.As much as it possible remove all culture media above cell pellet before DNA extraction.

2.Do not exceed cell biomass proposed for the kit. Excess of cells gives excess of RNA that are not digested by RNAse and lowers quantity of purified DNA.

3.Be sure to follow the protocol of the kit, i.e. use the indicated volumes of reagents and incubate with them during indicated times.

4.Increase concentration of RNAse A or prolong time incubation with RNAse A containing solution.

I had similar problems with purification of yeast genome DNA and prolongation of time of incubation gives better results.

What is your RNAse A digestion protocol?

Dear Maisano,

You need to give more information first to get a proper answer. Did you use Alkaline Lysis method for your miniprep? Here are a few possibilities

1) Most likely, what you see is still RNA. RNAseA does not completely degrade RNA but depending on the amount of RNAse used and incubation time you will get short RNAs upto 20nt long. If you want to get rid of these very short RNAs you will need to incubate your DNA with RNAseI (NEB sells this enzyme).

2) You might have made mistake in the amount of SDS you are using in your protocol and this band might be SDS. SDS being negatively charged binds to EtBr during the electrophoresis.

3) If you used silica based separation of your DNA from the first miniprep lane you have shown in your gel, it is possible that you have a lot of silica contamination and this also gives such signals.

Please let me know which one of these are actually turns out to be the reason.

What are you using to extract your plasmid DNA?

I would have thought RNA, does the band not diminish at all when you add RNase? What kit did you use and does it include an RNase clean up step?

Likely it is tRNA, double strand RNAs are difficult to digest using only RNAase A, you can add RNAase H, in your digestion mix.

Another possibility is you are using an old loading buffer, which produces ghost bands in agarose gels, try changing LB with a fresh mix.

Have you changed ALL the reactives of your protocol?

That´s the first step when you get a contamination in your process; first you have to prove that your reactives are OK and if that doesn´t solve your problem, you should analyze the procedure

it is RNA!! try open new kit..perhaps, the RNAse that you use is damaged already. I have this kind of experience before.

I've seen a similar type of "contamination" before that actually came from the loading dye. I'm not sure what the band was caused by, but simply changing the loading dye fixed my issue. I'm not sure from your picture, if in your case the kit DNA and the mini-prep DNA were loaded in the same loading dye; but if not, it would be worth your time to run a lane with just loading dye and see if you get the same "contaminating" band.

Make sure ur RNase is working properly. And subsequently do Phenol:chloroform: isoamyl step followed by Cholroform:Isoamyl and precipitate..You will surely get rid of this..

May be I miss something

could be denatured plasmid? I mean ssDNA?

try to do a short incubation in sol 2 (NaOH-SDS)

or use a single strand DNA nuclease such as S1

The regular minipreps that we do using the alkaline lysis method gives us a band exactly like this of RNA. Was this sample run very shortly after the prep/RNase treatment? In which case the RNase will not have had time to act (I need to incubate the sample for some hours in Tris-EDTA buffer pH7.5 with RNase (Roche) at a final concentration of 2 mg/mL to see a reduction in this band). What is your downstream processing? Restriction digestions are not affected by this RNA, neither is subsequent transformation etc.

If you do want to get rid of it, you can try PEG precipitation, exploiting the correlation between the concentration of PEG , duration of precipitation and temperature, with the size of molecules which are precipitated. We use a 1:1 mixture of 20% PEG8000 in 2.5M NaCl and DNA solution, incubated at 4 C for 20 min and spun and washed with EtOH. This process mostly precipitates supercoiled plasmid DNA (4-9 kb) and the small molecular weight RNA remains in solution.

All the best!

Your band is residual RNA with strong secondary structures which are resistant to RNAse A. To get rid off it:

1.As much as it possible remove all culture media above cell pellet before DNA extraction.

2.Do not exceed cell biomass proposed for the kit. Excess of cells gives excess of RNA that are not digested by RNAse and lowers quantity of purified DNA.

3.Be sure to follow the protocol of the kit, i.e. use the indicated volumes of reagents and incubate with them during indicated times.

4.Increase concentration of RNAse A or prolong time incubation with RNAse A containing solution.

I had similar problems with purification of yeast genome DNA and prolongation of time of incubation gives better results.

I agree with et al., could be the reactants. Run controls using your reactants and also check the pippettes you are using. Once all your reactants are clear THEN move to check your source of DNA.

Thanks everyone for the insightful suggestions! Yes I did run with Rnase A or without Rnase A comparisons and I definitely saw a difference between them. However, this band still exist with 4min Rnase A treatment. It is not due to loading buffer because my midiprep DNA (followed promega kit) ran great in the gel. It must be some type of RNAs or small DNAs that's resistent to Rnase A.

Also, I made the solution 1 to 3 myself. In the past it worked like a charm, since I made new solutions it started acting like this. Could it be PH related? Since the protocol says use solution 3 (NaoAC and acetic acid) to neutralize in order to precipitate DNA with SDS, I wonder if the PH is not optimal after adding solution 3 so that something else stayed in the solution with plasmid DNA... adjusting the volume of solution 3 to bring the PH to ~7 is beyond my math ability...

Make a new batch of RNaseA. Unless you have problem with the subsequent steps of your experiment, I would not worry too much about it.

The subsequent sequencing was not affected at all. But I am curious to find out what it is and how to get purer miniprep DNA. Somehow the midi prep was not successful with either qiagen kit or promega kit, I suspect that the plasmid DNA binds to the column. I have seen this problem multiple time in which plasmid DNA do not elute from the colume. So I decided to do multiple minipreps and then pool them and concentrate so that I can get enough high quality DNA for transfection. But I have this problem of impurity for miniprep...

Is your plasmid big? low copy number? they are the tricky one to purify.

I am in full agreement with the answers of Najla Arshad and Dmitry Karpov.

Kindly do what suggested by Dmitry

If this band correlates with low pH it might be product of acid hydrolysis of DNA (in case you dissolve/elute with water, water can be as low as pH 5.0 when prepared by reverse osmosis). Try time course incubation to see is the band expanding in time. In case of RNA of complex secondary structures try incubation temperatures of 56 to 60 degrees Celsius - RNase works well at that temps.

I qoute Dmitry Karpov for Ram Nagina:

Your band is residual RNA with strong secondary structures which are resistant to RNAse A. To get rid off it:

1.As much as it possible remove all culture media above cell pellet before DNA extraction.

2.Do not exceed cell biomass proposed for the kit. Excess of cells gives excess of RNA that are not digested by RNAse and lowers quantity of purified DNA.

3.Be sure to follow the protocol of the kit, i.e. use the indicated volumes of reagents and incubate with them during indicated times.

4.Increase concentration of RNAse A or prolong time incubation with RNAse A containing solution.

I had similar problems with purification of yeast genome DNA and prolongation of time of incubation gives better results

The best way to Mini-Prep...Is by BOILING...

-Prepare lysozyme (10mg/ml in TE) and a boiling water bath...

-Place 1.5ml of bacterial culture into a 1.5ml tube, spin 30Sec at RT... Aspirate off medium

-Resuspend in 0.35ml of STET buffer (up/down)

STET buffer cold (4C) : for 100ml total, add 8% Sucrose, 0.5% Triton X-100, 50mM EDTA (pH=8), and 10mM Tris (pH=8)

- Add 25ul of freshly prepared lysozyme... Mix by Vortex...

-Place tube in boiling water for 40sec (only)

-Centrifuge immediately for 4min at RT....

-Remove pellet with toothpick, and add 0.5-0.7ml of ethanol... Centrifuge for 10min in cold.

-Remove supernatant liquide. and resuspend the pellet in 50-100ul of TE (or water)...

All The best

CE0702

The boiling method is good but be careful of your sample being contaminated with genomic DNA (ie. chromosomal). Can be problematic if your subsequent step is PCR, sequencing.

Xu Maisano.If your plasmid is high copy no then double the volume of medium using for low plasmid bacterial culturing.But all these are mentioned in every kit.

Second..Elution of your plasmid if your plasmid is high molecular weight pre-heated the elution buffer for 3 minutes at 75c.Then incubate it for 3-5 minutes at room temperature. Another thing your plasmid elution will be decreased 30% if you are using water.So keep in mind all these thing when isolating the plasmid. Hope you will get very good result.

Dmitry Karpov..Informative point you mentioned during Plasmid mini prep..so helpful to me.thank,s.

Could it be the ghost band:-)

Tibs

Volume 21, Issue 11, November 1996, Pages 441–442

Methods and reagents: Eliminating ghost bands from plasmid preps

Paul N. Hengen

National Cancer Institute, Frederick CancerResearch and Development Center, Frederick, MD 21702-1201, USA

http://dx.doi.org/10.1016/S0968-0004(96)30041-

Methods and reagents is a unique monthly column that highlights current discussions in the newsgroup bionet.molbio.methds-reagnts, available on the Internet. This month's column updates the discussion of a ghost DNA band that continues to haunt netters. For details on how to partake in the newsgroup, see the accompanying box.