Electrophoresis - Science method

If you are running 4ng of dna per well then this is spread over much of the length of the gel (assuming some degradation) and there is just not enough dna to see a signal. You will need to load a lot more dna to se it on a gel.

Hello Seda Homurlu

Standard tank will suffice. It's not worth investing in a specialized electrophoresis tank. You just have to incorporate a few points in your protocol.

During the electrophoresis step of comet assay, the high current can cause several problems.

High current can generate heat which can cause an increase in temperature that can denature DNA or cause smearing of DNA on the gel. It can also cause uneven migration of DNA fragments. To overcome this problem, specialized electrophoresis tanks for the Comet Assay are equipped with connections for a recirculating water chiller to keep the tank at a constant temperature during electrophoresis and eliminate joule heating effects. These tanks can be used with ethylene glycol, allowing cooling to 4°C in the electrophoresis buffer. Also, they are constructed from opaque acrylic, so that no light can penetrate the tank during electrophoresis.

Without using these specialized tanks, you too can get rid of these problems. Do the following.

1. After the lysis step, carefully remove the slide carrier from the dish. Gently place the slide carrier in a washing dish pre-loaded with ice-cold double distilled H2O and leave it for 30 min ensuring that slides are completely covered with double distilled H2O.

2. Insert a frozen cooling pack inside the sliding drawer under the electrophoresis tank to maintain optimal buffer temperature.

3. Carefully add ice-cold electrophoresis working solution to the electrophoresis tank and transfer the slide carrier into the electrophoresis tank.

4. Allow the slides to sit in the electrophoresis tank for 20 min so that the DNA relaxes and unwinds. Keep the power supply switched off during this step.

5. If needed, insert a new frozen cooling pack to maximize chilling.

6. Maintaining the current will be influenced by the buffer volume in the electrophoresis tank. Instead of filling the tank as you would if running a gel, the buffer level should start by just barely covering the slide. Adjustments to the level can then be made until a current can be maintained.

7. Perform electrophoresis for 20 min at 1.19 V/cm, or whatever conditions you may have optimized for the run.

Another point which I would like to mention is that covering samples to minimize UV light exposure is necessary, which can induce additional DNA damage and result in high background. This would be important when preparing your samples, but it is probably fine to run the electrophoresis uncovered.

Best.

I would suggest to try blocking with BSA for detecting phosphorylated proteins specifically because of the presence of phosphatases in milk.

If the amplimer is small it will diffuse and spreadout and the signal becomes weaker and may disappear. Also if you leave the gel on the UV light for too long then it bleaches the fluorescence and no signal will be seen

Paul Rutland Thank you so much for all your suggestions! I will try using denaturing gels and SYBR Gold.

Hello Bhardwaj,

I couldn't give a definitive answer but I could give 1 potential answer for this. It could be due to incomplete dissolving of agarose. Would recommend looking at a similar post that was made on research gate on Dec 14, 2016. Posted by Lorenz Kempeneers.

Hope you the best,

Nicolas

It is supplied in DMSO which is quite volatile so if it is very old you may have lost some dmso to evaporation and reached the point where it cannot dissolve the dye cold but the dye is more soluble in warm solvent. If you have not used much and if the supplier material sheet gives you the total volume in the vial then you could measure the volume in the vial to see if it is what it is expected to be

I run a lot of SDS-PAGE gels every day and hope to prepare as many SDS-PAGE gels as possible in one go. However, when I prepare a large number of gels, they start solidifying before I can pour a few, leading to waste. Does anyone have any tips to prevent the gels from solidifying too quickly? I know there are companies that specialize in large-scale production of precast SDS-PAGE gels, and there must be a solutions to address my issue.

Chinmay Hazare thanks alot

DH5alpha does not carry any plasmid, but of course it is a host for recombinant plasmids. So only you would know what size the plasmid was depending upon what plasmid you put in it (or was in the strain that was given to you). But DH5alpha alone does not carry a plasmid.

But from the figure you show, the very bright fluorescence in the middle of the gel is probably RNA and the faint bands at the top might be plasmid but most likely are just some genomic DNA.

The band in your water is contamination. You need to throw away ALL (and I do mean ALL) of your reagents (enzyme, buffer, water, primers, dNTPs, etc.). Make a new batch of TE buffer & make a fresh aliquot of your primers.

Get new boxes of pipet tips & other plasticware. Wipe down your bench area & clean your micropipettes. Open a new box of gloves.

Common sources of contamination:

Good luck!

I am unclear from your question whether there is any added primer in the ntc as there should be but it may also be possible if you set the baseline too low and there is a small amount of signal released from the mastermix then it could look like amplification

"You can't rush quality".

Seriously. It just takes time to separate out the bands. If you turn up the voltage too high, then you will get smeared bands and poor resolution.

I faced something wierd my RNA qubit concentration increased after DNase I treatment. Can anyone help why is it so?

I would guess that the transfer is slower from the higher percentage gel, potentially leading to a lower signal due to incomplete transfer. This would reduce the signal-to-background ratio, making the background seem higher. If this is correct, a longer transfer time may improve the result.

Hello,

After many attempts I managed to solve some of the problems. Right now, Lambda ladder is quite good (although not fully separated probably due to to short swith time), but the sample bands are smeary, tick and distorted. Current electrophoresis conditions were:

Switch time 6-36s

Voltage 6V

Angle 120

Buffer temperature 14C

Run time 22 h

I have tried adding slimmer slices but it didnt improove anything. I would be realy grateful for any suggestions, because im running out of ideas.

We currently suspecting that the longer run PCR (confirmation) might be incompatible with our product. We are trying different protocols in order to (hopefully) achieve confirmation.

A low 260/230 ratio may indicate the presence of contaminants that absorb at 230 nm or less, such as EDTA, carbohydrates, phenol, or guanidine HCL. A low 260/280 ratio may indicate the presence of contaminants that absorb at 280 nm or less, such as proteins or phenol.

First, I would quantify the DNA sample before starting the PCR, reduce the total volume of the reaction, and perform a temperature gradient.

you do appear tp have weakly visible bands in this image so I would check that your sample loading dye is dense enough to hold the sample in the well. If so then run 3 or 4 more cycles of pct to get more product. You mention that the gel is not covered with running buffer. It is usual to cover the gel completely with running buffer to get good results. You could just load more sample and the gel would look better in this case but more cycles would be better

Jeremy Mullesa I think your plasmid is ok. PDR111 is overloaded, therefore, it appears to run faster and looks smaller/different from your clone. Load equal amounts. To ensure you have the correct fragment cloned, sequence your construct.

Kais Khudhair al Hadrawi you answer is off-topic and simply generated by ChatGPT. The RG community for sure knows how to use this tool.

Hello,

I agree with Dr. Paul above, this happens due to formation of heteroduplexes. Your original band contains more than one product, with no noticeable difference in mobility, but the slow moving band is a heteroduplex. On cutting the expected band and Re-PCR, all the three combinations are generated again. We observed this and resolved in our study on such observed heterogeneity during ribosomal DNA ITS region amplification in Asiatic Vigna species (see Saini et al., Genet. Res., Camb. (2008), 90, pp. 299–316.). We also proved the differences (indel lengths, 2 bp and above) among clones by doing heteroduplex analysis by mixing different clones, in that study.

If you are interested in getting the amplicons, instead of band purification go for cloning and sequence.

all the best

In Native PAGE (Polyacrylamide Gel Electrophoresis), the pH value of each component plays a crucial role in ensuring the proper separation and migration of proteins based on their native charge and size. maintaining appropriate pH values for each component in Native PAGE is essential for preserving the native structure and charge of proteins, ensuring accurate separation and analysis. Any deviation from the optimal pH range can lead to protein denaturation, aggregation, or altered migration patterns, affecting the reliability and reproducibility of the results.

To Adron Ung

"Just switch the leads of the electrodes: black to black and red to red going to the electrophoresis gel assembly with ... red to black and black to red going towards the battery. Now the cathode(-) is at the bottom of the gel and the anode is at the top. Science!"

What positive charged dye can be used for such case?

what exactly is the problem You are running the gel too high a voltage causing some smearing and you have primer dimer that would probably benefit from using a hot start enzyme so the only problem that I can see is that you look to have over amplification of your samples. Where is the no template control on this picture because it may be that you have pcr contamination in your pcr and any amplification of the negative control would indicate this.

How many cycles of pcr are you running and how much dna (ng) are you amplifying in each pcr reaction?

If you use unstained markers, you can stain the blot temporarily with Ponceau S to locate and mark the positions of the markers before developing the blot.

Thank you very much for the suggestions, I will try again. Best wishes. Malcolm Nobre Stefanie Meyer

Make sure your cDNA synthesis process is correct. Also the loading dye.

If you're performing asymmetric PCR with only one primer (in your case, the FAM-labeled reverse primer), and you're not seeing any product bands on a denaturing polyacrylamide gel electrophoresis (PAGE), there could be several potential reasons for this:

By carefully reviewing these factors and optimizing your experimental conditions, you should be able to troubleshoot the issue and visualize your PCR products on the denaturing gel.

I'd denature them again, to be on the safe side, as long as that does not interfere with the interpretation of your experiment.

I don't see the point in using multiple size markers. Size markers are used to estimate the size of your fragment. If the bands of your marker are too close in the region of your fragment, adjust the concentration of your gel. For instance, for an agarose gel, if the fragment size is in several kilobase pairs, use a low-concentration gel (between 0.4% and 0.8%). If the fragment size is in a few hundred base pairs, use a more concentrated gel (between 1.5% and 2%). I think it's more useful to adjust the gel concentration rather than using multiple different markers.

Paul Rutland Thanks, dear sir.

Given that the ladder on the lhs of the gel is trying to work well and the single sample on the rhs of the gel is spreading in a similar shape to the mess in the middle it looks like some salt or other contaminant in your samples is causing the samples to spread and the central samples being close to each other the effect is much greater I would not worry about the gel and pay particular attention to what is remaining or being added to your samples prior to running them but in answer to your question,,no I have never seen this effect before

Hi Constanza,

Throughout the electrophoresis steps, please use powder free gloves to prevent contamination from skin keratins. It is also important to dedicate apparatus for electrophoresis and not to use any surfaces used for Western blots. Otherwise, you will encounter eg., casein and BSA contamination arising from blocking of membranes during Western blots.

The best way to clean any glass apparatus to remove background proteins is the following procedure: this is the same procedure our proteomics laboratory at MSKCC used, for high sensitivity sequencing of peptides:

1: RInse glassware with 80% acetic acid (v/v)

2: Follow 1 above with MQ filtered water

3. Follow 2 with 100% methanol

4: Follow 3 with MQ filtered water.

This protocol of stringent washes can completely remove any protein/peptide contamination from your elelctrophoresis apparatus. Please use a fume hood for this cleaning experiment. Also, make sure your electrophoresis containers are glass eg;, pyrex glass containers that can be used for stainng and washing of the gels.

The plastic containers used for electrophoresis can be rinsed with dilute methanol and water. You can omit the acetic acid for these plastic containers.

Good luck!

Hediye.

Dcrease the cycle of pcr less than 30 cycle not more make it 29 cycle

Can you share a picture of what you are observing?

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