Electrophoresis - Science method

20 cycles of pcr can be as much as 1000000 times more pcr product so it is really easy to over amplify your template to the point where product molecules anneal to each other and form longer concatamers which will look like a smear of pcr product. Set up some pcr second round amplification tubes and remove tubes from the pcr machine at 18, 18. 20......26 cycles and see where you get a clean product. As template you could use 1:100 dilution of the first round product

Firstly, I think you may need to dilute your DNA before doing the PCR if it is luminescent. Second, make sure you haven't forgotten to add anything to the PCR mix. Thirdly, you need to use a positive control for accurate detection. I think if you take all these things into account, you'll be fine.

Us ethe gel again. Small dyes will move in the electric field but there should be enough in the gel to intercalate with the dna and if not you can post stain if the signal is a bit weak

I use ethidium bromure (BET) to visualize PCR products. This dye migrate in opposite direction of DNA. If my gel run for a too long time the dye risk to be over of small sizes. I supose it is your case.

What you can try: increase agarose concentration at 2% or 3%.

Reduce the time for the migation : 15 min should be enough. You can adjust the voltage.

You can increase your SBYR amount.

Another tip : don't use detergent to clean your gel tank

Thank you very much Prof. Liger for your answer

I assume you mean it's a stain that is a safer alternative to Ethidium bromide, like SYBR Safe?

You do not need to mix it into your DNA loading buffer or the DNA samples. Just add the SYBR to the melted agarose, swirl well to mix, and pour the gel.

Thou Chen Loh :)Thank you very much for your reply. I ran the agarose gel test directly and I didn't heat it to 70 degrees before electrophoresis. Do I need to mix the RNA and lodding buffer and heat it to 70 degrees before testing?

For the bacterial liquid PCR, did you use primers that are on the ends of your fragment or is one in the vector? That would add some length.

There is no fixed time or voltage as conditions are set depending on the size of the band and the sizes of any other amplimers that need to be separated from the band of interest. The percentage of the agarose gel is important depending on the band sizes to be separated. I use 5v/cm where the length is the distance between the electrodes in the gel tand but this can be reduced when separating closely sized amplimers or increased if just checking for amplification. Small fragments are better run in cold gels to minimise thermal diffusion, A rough guide as to waht percentage gel should be used to run pcr products/restriction fragments is

%gel Range kb

0.3 5 - 60

0.5 1 – 20

0.7 0.8 – 10

0.9 0.5 – 7

1.2 0.4 – 6

1.5 0.2 – 3

2.0 0.1 – 2

50X TAE is very stable, we make it 1-4 times a year (it depends on how many gels we are running). It hasn't gone moldy/bad yet.

The 1X TAE can eventually grow microbes, if it starts to smell odd it likely has mold. If you use a clean container and keep the lid shut it will keep for weeks.

The 1X TAE in the gel running rigs tends to go bad faster than in a storage bottle. I would suggest changing it daily (or if you run infrequent gels just store the rigs dry and empty)

I think this band is because of the primer bind with itself we call it dimer

Trizol contains phenol and an acidity detecting dye. At the right pH this dye is pink Storage or light affects phenol. Its OH group activates the ring structure to allow addition of electrophilic ions such as OH- , This forms quinone which then reacts with 2 molecules of phenol to form phenoquinone which changes both the nature of the phenol and its pH leading to poor rna purification. I think that you will do better with a new bottle of Trizol

https://socratic.org/answers/607619 will show you images of the transformation

Yes, it can be calculated based on the ladder

Namita Chutani https://info.gbiosciences.com/blog/tris-tricine-sds-page-what-is-it-and-how-to-perfrom-it#:~:text=Tricine%20gel%20should%20be%20run,room%20or%20using%20precooled%20buffers.

You may replace LDS and SDS for PAGE. It is not applicable for mass spec whether lithium or sodium salt. Li-cor uses LDS in PAGE products and applications they recommend LDS instead of SDS due to enhanced solubility but also more applicable for plant proteomic research. For MS getting rid of all types of dodecyl salts and in-gel digestion by applying wash replicates after excising the band would be an efficient protocol.

I think this is due to contact zones of the gel and a buffer which may create conditions slightly different than internal layer, such as ions..

To analyze the results of an SDS-PAGE gel, you can follow these general steps:

These video playlists might be helpful to you:

There could be several reasons why you are unable to observe tRNA in the well of agarose gel electrophoresis:

It is worth noting that tRNA is relatively small in size, and depending on the experimental conditions, it may migrate differently compared to larger DNA fragments. Ensure that your gel electrophoresis system and parameters are suitable for the size range of tRNA.

Overall, it is important to troubleshoot and optimize various factors including sample concentration, staining method, gel conditions, and loading techniques to enhance the visualization of tRNA on the agarose gel.

These video playlists might be helpful to you:

I think that this due to the use of high volt during running, and the repeated use of the sameTBE or TAE buffer

These video playlists might be helpful to you:

Here are some potential reasons why you may not be seeing any bands after electrophoresis:

I hope these suggestions help you identify the issue and improve your results.

These video playlists might be helpful to you:

A smear under the band in electrophoresis can be caused by multiple factors, including incomplete amplification, excess template DNA, too many cycles, inappropriate annealing temperature, and degradation of the DNA during the PCR reaction.

In your case, the PCR reaction has a short extension time of only 7 seconds, which could contribute to incomplete extension and result in smearing. Additionally, the number of cycles may be too high for the specific PCR reaction, leading to over-amplification and the production of nonspecific products. The annealing temperature could also be contributing to nonspecific amplification, as it is only 55.0°C, which may be too low for the specific primers being used.

To improve your results, you may consider optimizing the extension time and annealing temperature for your PCR reaction, reducing the number of cycles, and adjusting the amount of template DNA used. Additionally, you could try using a different DNA polymerase or optimizing the PCR conditions to reduce degradation of the DNA during the reaction.

These video playlists might be helpful to you:

If cells are visible but without tail in the comet assay, it usually means that DNA is not fragmented or the level of fragmentation is very low. There are several reasons why comet assay tails may not appear:

To troubleshoot the issue, you may want to adjust the electrophoresis conditions, such as lowering the voltage or extending the electrophoresis time, to encourage the formation of tails. It may also be helpful to optimize the lysis conditions to ensure complete DNA release. Additionally, you can try reducing the number of cells loaded on the slide to ensure proper migration of DNA fragments. Finally, it may be helpful to confirm the presence of DNA damage by performing other assays, such as the alkaline or neutral comet assay.

These video playlists might be helpful to you:

Hello Bahaa Kargule

Protein Electrophoresis is done to identify some unique kind of protein in the body to detect specific diseases. Body fluids, such as blood, urine, and cerebrospinal fluid (CSF) contain many different proteins that have various roles, such as transporting nutrients, removing toxins, and controlling body functions.

The various proteins in body fluids are subjected to a controlled electric current (electrophoresis) fractionating them into a typical pattern of bands or peaks that then can be measured. The proteins are divided into six groups, called prealbumin (rarely detected on serum or urine protein electrophoresis), albumin, alpha 1 globulins, alpha 2 globulins, beta globulins, and gamma globulins. The beta fraction may be further divided into beta 1 and beta 2 subgroups. Each of these protein groups (electrophoresis fractions) is distinct and at specific concentrations. The patterns typically seen in certain conditions and diseases can help with diagnosis.

Various conditions and diseases can affect protein production and/or protein loss, thus changing the pattern of bands seen on protein electrophoresis. For instance, any problems with the kidney or liver, or if one is having trouble with the uncontrolled growth and division of a malignant plasma cell leading to the production of large amounts of a single type of immunoglobulin (multiple myeloma). The doctor may use the results of protein electrophoresis to make a diagnosis or decide on the course of treatment. But further investigation will usually be needed to make a definitive diagnosis, for instance, use of immunofixation electrophoresis or immunosubtraction electrophoresis to identify abnormal bands seen on protein electrophoresis.

Best.

Hello Adriana, it would also be helpful to know what the expacted band size is. Accordingly, did you use a 0.7x, 0.8x, or 1.2x agarose buffer? The lower the concentration of the agarose and higher the voltage used during electrophoresis, the faster is the movement of DNA across the gel, so that could be the reson why there is no bands seen.

Mixing the agarose well in the buffer until the mixture appears transparent, adding the DNA staining dye directly into this mixture, and then cooling the gel mixture on the gel tray until all of the gel turns completely solid (waiting 30-40 minutes at least) could help in the uniform creation of the agarose gel for your purposes.

Also, a well-designed gel run usually has only one band of DNA sample per lane, as agarose gel electrophoresis is only a qualitative confirmation of the presence of your DNA of interest, with the DNA ladder indicative of the approximate size of the DNA concerned. Having multiple bands in the same lane is not considered a good PCR result (as seen in your last lane), so I personally would also check whether the primers used are unique for your amplicon of interest with no danger of forming 3' loops, hairpin structures, etc.

There are a few aspects that you may want to consider. There is no easy answer to what dye to use because it depends on your risk aversion, financial status and the sensitivity of the gels that you are running. So sybr gold is 9 times more expensive than EtBr while the sensitivity of EtBr is 1ng while sybr gold is 25pg so cost/sensitivity may be important. Gelstar,sybr gold and sybr safe can all be excited using blue visible light avoiding UV risk to skin and eyes and also avoiding expensive UV transilluminators. Sybr green can detect 60pg of dna but is best used post staining after the run which may be inconvenient if you are watching a gel in order to excise one band Methylene blue is very cheap but can only be used post electrophoresis while nile blue can be used during electrophoresis but both these ( and fast blast) can be used with visible light to detect the dna. I think that the evidence against EtBr is poor and that some of the other dyes ( not the SAFE dyes) do not look perfectly safe and that it can be used with gloves quite safely although when I went round our electrophoresis area with a hand help UV lamp it was astonishing where splashes of etbr had ended up ( even the ceiling)

The problem with electrokinetic sample loading is that the salts are small and highly charged so load first and the dna only loads later in the injection process and you get a weak signal due to low loading. Often this means that running the same sample twice may desalt the sample and it loads a lot better the second time. The other problem is that the ladder has a different salt concentration so loads early so sizing is an issue. If you cannot dilute the sample then you can precipitate and wash the dna and redissolve in water before preparing the sample for running or run the pcr mix through a disposable gel filtration column as used for cleaning up sequencing reactions

Thanks everyone!

We´ve tryed changeing the buffer by diafiltration, but not tested yet EDTA on that condition. I will try that next.

The SDS/b-mercaptoetanol is a little to harsh I think, but was sugested by someone at the lab and it gives me no good results so far.

Hi Muhammad Fozan

1. You can DEPC-treated water to prepare your gel, assuming your RNA is getting degraded in the gel.

2. You can use a different buffer system like MESA + formaldehyde agarose gel, which as per my experience gives best results for RNA.

3. Also take note of your visualization reagent's sensitivity. For Gel Red it is ~50ng DNA/RNA.

Hope it helps!

To answer your questions:

To increase the polymerization time of your gel:

1. make sure your APS stock (crystals) is dry and make fresh APS. If the crystals are wet then throw the stock away and get new. This is usually the problem when gels do not polymerize in a reasonable amount of time.

2. Add more TEMED.

3. Put a flame (Bunsen burner) near the gel. Heat will increase the rate of polymerization.

Migrating DNA:

1. Be sure to load all lanes of the gel with at least loading dye such that the current/resistance is the same across the gel.

Gel staining:

1. Acrylamide gels stain very quickly with EtBr since they are so thin. If the gel is bright pink after staining, you may need to destain the gel so you can visualize the DNA. You should be able to see a few nanograms of DNA.

Since HPLC can answer questions of both small and large molecules even if they are ionized or neutral, it may be mentioned as more universal. Already ionized or ionizable compounds are needed to perform electrophoresis which may be counted as a limitation. But, indeed, in some cases, depending on the application, improved resolution can be achieved at electrophoresis than LC. In my opinion, they must be used orthogonal and complementary (e.g. antibody characterization, QbD workflows)

A short 20 mer traps very little intercalating dye so is diffuse due to its short size and weak. You might see it better by running the gel for a shorter time so maximising the signal and using silver staining to detect the bands but why do you need to see the 20mer. If you run an uncut 146 size amplimer then any sample generating a 126 base product must have been cut by the enzyme and 146 and 126 will separate well on a wide range of PAGE gels

Sabnam Kar You don't need 2D-DIGE equipment. This was just an example for the use of these Cy-dyes and intended to help you to identify them. After labeling your peptide, you should be able to visualize it in your experiment as you have described it, just by using a transilluminator with a suitable spectrum (e.g., blue or UV for dyes with a fluorescein like spectrum).

The smiling bands in positions 3 and 4 look like there is too much salt in the sample. Try diluting the sample in water and running it shorter and slower ( less time lower voltage) to see if a proper shaped band is seen

I used pH 8 sample buffer, pH 8 running buffer, and pH 8 Tris-Glycine gel for for Native PAGE and it worked well for my protein.

Dear Emilie,

The solution may be very simple: if you clone in Lambda DNA digested with PstI into PstI-digested pU19, and then digest all of the plasmid DNA samples produced from cultures inoculated with blue or white colonies and selected with ampicillin, then all of your samples will include a band at 2.7 kb corresponding to ... pUC19 ... which is the source of the ampicillin resistance and will turn the colonies blue if there is no insert of DNA at the PstI site.

Your gel images suggests that a number of your samples contain Lambda DNA inserts, but since the band intensities vary a lot, I'd be a little careful in interpreting some lanes with bands above the pUC19 band, as this may represent partially-digested plasmid DNA. Lanes 3, 6, 7, 9, 10, 13, 15 and 16 look interesting. You could look at a Lambda restriction map to see the range of PstI fragments it would produce, and map them onto the fragments you can see in your gel.

Regards, Andrew

Thanks to all of your answers, I was able to get rid of the build-up of salts with a 0.1 M solution of HCl. The tank keeps leaking but I have been able to find the crack. Hopefully, I can solve it with an adhesive as Katie A S Burnette suggests.

Hellow B R Siddharth

I am using 0.25% (w/v) Bromophenol blue, 0.25% (w/v) xylene cyanol FF, and 30% (w/v) glycerol in H20. Preparing according to "Molecular Cloning: A laboratory manual" by Joseph Sambrook. Refer to the appendices in this book.

1. - Prepare sufficient 1 x TBE electrophoresis buffer (1:10 dilution of TBE:DEPC H2O) - Clean all tools using DEPC H2O. - Position the comb 0.5-1 mm above the plate so that a complete well is formed when the agarose is added.

2. - Prepare agarose gel for a 1.2% agarose gel: 1.2 g agarose / 100 ml 1 x TBE buffer in Erlenmeyer flask - Cover the flask with kimwipes/ parafilm and heat with microwave until the agarose dissolves. Measure it again and complete the evaporated liquid with distilled water. - Leave it to cool down to about 60 °C on the bench for several minutes but do not leave it too long so the agarose should not start to solidify. - Stain the agarose solution: 5 µl ECO Safe Nucleic Acid Staining Solution / 100 ml gel - Mix the agarose solution well by swirling the flask. Pour the agarose into the mold. (3-5 mm thickness)

3. - After 30 minutes at room temperature carefully remove the comb. - Position the gel into the gel electrophoresis tank. Avoid bubbles!

Add enough TBE buffer to cover the gel to a depth of about 5 mm.@

We changed the buffer again and the problem was solved. Looks like someone made the buffer wrong :))

Actually I do not sure. I used strain provied from my university’s LAB

Your plasmid bands are overexposed, what my advisor would call "screaming bright". Whatever the tiny bands are, they are honestly too diluted to be a concern.

It seems that too much PCR product has been uploaded. Have you run the controls along? Please check with the PCR product conc. and load only 1ul if you dont want to dilute. You can lower the cycle number in PCR reaction in future as well. Good luck

Can we see a picture of the gel. A good pcr band should work at many times and voltages and I wonder if you just have primer dimer and no amplification

Phenol red works, basically any dye with a negative net charge should work. Look for sulfonated azo-, triphenylmethane- or xanthene- dyes.

I'm not sure how much EtOH can be tolerated in SDS-PAGE, but I don't think you should be concerned about solubility. Just dilute the protein directly in sample buffer and boil, which will completely solubilize the protein anyway. This approach also minimizes the concentration of EtOH, which is worth doing just in case EtOH negatively affects SDS gels.

It looks like something is wrong with your transfer setup. All the marker dyes are being pulled to the same point in the top middle of your gel. Something there acts as a stronger cathode than the rest of the cathode during your transfer. Perhaps the insulation on one of the wires has degraded.

Dear Maxim, your bands are still not look linear, this may cause the wrong results in determining the size. I thing your gel's problem is originated from electric field. Are you degasing your buffers? If your electric cable is working properly, you can count this as a possibility.

Bests.

There will always be some degradation of genomic dna in the purification process but the main reason is that agarose cannot distinguish very high molecular weight dna so although there will be ahuge range of dna sizes they will all appear as a band at the limit of the gels ability to separate large dna. Even the mixing and shaking of the dna causes dna breakage which is why making dna for PFGE means suspending the cells in agarose plugs and doung all of the purification steps on immobilised dna

Hi Noel,

I don't think that you will have the same extraction efficiency for your stratum corneum samples with B-PER buffer since this buffer is optimized to extract proteins from bacteria. I would recommend to use M-PER buffer since it is optimized to mildly extract proteins from mammalian samples.

Best,

Murat

I wouldn't risk it. Either toss the gel and start over, or if it's multiple gels and you are willing to go through the trouble you could remelt the gels in the microwave in a larger flask and add H2O to get back to 1x TAE and add more agarose to reach 1% and pour the gels that you need now. So you'll end up making 10x as much as you had planned but you can store the remaining agarose as a stock in the fridge and melt and pour gels as you need them. Just be careful to not let too much evaporate out when remelting or you'll end up drifting towards higher %gels as you continue using the stock.

for the uneven migration, your tank may be slowly leaking (the buffer between your gels) for the current may be your power supply is not on constant voltage but on constant current (and as the resistance increase the voltage decrease) ...

Suggestion.

Platinum wire can be very expensive and difficult to find or replaced by the manufacturer of your gel electrophoresis tank. Moreover, the wire sold by Sigma (Merck-EP1422 Sigma-Aldrich® VS20 replacement platinum wire) is expensive, very thin (0.2 mm) and very brittle.

I suggest you visit your local dentist or search online for a company selling Nickel-Titanium Orthodontic wire in your country, order a spool (bobbin) and use it in your gel tank. Wire measuring a diameter of .16", .18" or .20" will be suitable for the purpose. The Nickel-Titanium alloy from which the wire is made is highly acid/ corrosion resistant and not expensive at all.

Good Luck

Hello Laxmi Kashyup

Electrophoretic Mobility Shift Assay (EMSA) is used to analyze nucleic acid–protein interactions. EMSA is based on the principle that DNA–protein complexes are larger and move slowly when subjected to nondenaturing polyacrylamide gel electrophoresis (PAGE), compared to the unbound (free) DNA probe. Since the rate of DNA migration is shifted or retarded when bound to protein, the assay is also referred to as a gel shift or gel retardation assay.

Please refer to a representative protocol provided in the article below. Commonly used variants and expected outcomes are also discussed in the paper.

Best.

You do not need to titrate TBE with HCl to adjust the pH. By doing that, you introduce fast moving Cl- ions, which might be responsible for heating the gel and distorting the bands. Just mix the specified amounts of Tris, Boric acid, and EDTA, and add water to them to the final volume. Without any adjustment, the pH value should be within 8.3-8.6, which is perfectly fine for nucleic acid electrophoresis.

The presence of protein in the gel and not being transfered on to the PVDF membrane. Now you have to work out in order to ensure that the protein is efficiently transferred to PVDF membrane.

I would suggest the followings issue to be considered:

I) Check the membrane pore size.

II) Double check that the transfer apparatus is on and functioning normally.

II) Check once again the transfer buffer, just to make sure that it has sufficient ions.

III) According to the size of protein of your interest and gel/membrane size adjust the voltage and transfer time

IV) Check that the heat is not being generated that normally increase the resistance and destabilize the transfer buffer.

Best

I would expect that the high speed of blood flow (mainly) and also turbulence and the random thermal movement of the molecules would overcome any small movement due to movement in an electric field

Hello Alejandro,

I have not seen this problem before and the only things that I can think of are that the amplimer has a polymorphism somewhere near the middle. In the reannealing step of the pcr when the primer anneals then at the later cycles the product reannneals with itself and mismatched strands re-anneal to form a pcr product that has a single stranded bubble in it. For example if we call the 2 strands N and M for the 2 base mismatches then we get NN and MM and twice as much NM plus MN strands. Then either there is some nuclease contamination or unexpectedly the EXO1 itself cuts the single stranded bubble giving a shortened double stranded minor product. Hopefully someone will come up with a better answer.

best wishes,

Paul

Maxim Kutyrev Please ignore my previous about degradation and RNA. I misunderstood what you are doing. For samples that small or unusually charged due to solvation effects and possibly by secondary structure if you want to use agarose then I would use 3% agarose (3% low gelling or at least 2% low gelling and 1% ordinary grade agarose,) run at low voltage to minimise thermal diffusion and run the gel less far ( shorter time) to maximise the band intensity. Is there a good reason for not running PAGE gels? ....they separate oligos better than agarose and if using denaturing page secondary structure is minimised. Are these gels diagnostic/analytical or preparative ? .If preparative and you are trying to separate small moieties and salts perhaps size exclusion mini columns would be applicable using something like sephadex g25

A 1% (w/v) solution of bromophenol blue is easier to prepare (10 mg/mL) and stable for years in the fridge. As the concentration is not too critical, it suffices to weigh near 10 mg of the dye and then add 100 x the volume of water (example: if your dye weighs 9.2 mg, add 920 µL of water).

May be there was insufficient quantity or concentration of DNA loaded on the gel. Increase the amount of DNA, but don't exceed 50 ng/band. The DNA may be degraded. Avoid nuclease contamination Sonu Singhal

Thank you very much for your time and answers to my questions

It may be that your original fragment before purification has significant secondary structure and is supercoiled and knotted to a small spherical shape. In DNA purification kits the sample to be purified is added to chaotrophic salts which linearise the molecule to facilitate the binding of the dna to the membrane.If the eluted dna remains linear on elution from the membrane then it will run slower through the agarose and will appear as a larger size than the natural coiled version

Membrane proteins are mainly water insoluble, even If you run a native or SDS-PAGE, they will smear on the gel.

I would give you some suggestion depending on your next step of analysis with the isolated mitochondrial membrane proteins

If you would like to isolate these membrane proteins in a native-like state, you should maybe consider to use detergents like triton x114 which is principally able to isolate/seperate membrane proteins from hydrophilic ones.

If no need for the native-like state you should maybe try BAC-PAGE, which has totally contrary to SDS-PAGE regarding the conditions,

here you can also try 2D Gelelectrophoresis in which 1.Dimension: IF and 2.Dimension: BAC-PAGE

No, We are using bradford methods as well as other methods.

Run an assay with 2 samples which have no dna in them and one normal dna sample.If the negative controls amplify bands of the right size then you have contamination

pWhitescript plasmid was supplied with the kit of agilent and I searched it, but I could not find something helpful. Probably, I will try it with a different control. Thank you very much for your answer. It was very helpful!!

Hello Ezinne Deborah

For your study you can refer to the methodology section of the research article given below. It may be helpful.

The link below may also be helpful.

Good Luck.

-For WB 10% ethanol in western blot transfer buffer works well and 100% ethanol for wetting the membrane.

Also dry or seminary transfer work well reducing waste. Or alcohol free transfer Dunn (doi:10.1016/0003-2697(86)90207-1, 10 mM NaHCO3, 3 mM Na2CO3, with 50 µM SDS for large transmembrane proteins). Alcohol is more important for small proteins.-Or 336 Tris, 260mM Glycine, 140mM Tricine and 2.4mM EDTA is a suitable rapid protein transfer buffer. Not necessary to pH. One 60 cm2 gel ~ 1.3A for 12-15 min