Electrophoresis - Science method
An electrochemical process in which macromolecules or colloidal particles with a net electric charge migrate in a solution under the influence of an electric current.
Questions related to Electrophoresis
I can see marker bands on my 1% agarose gel but no sample bands. What's weird is I can see fluorescent in loading wells although no sample bands are visible, seems like my sample was clogged into loading wells. Does anyone meet the same situation and know how to deal with it? My sample is just ~1,500 bp and I'm pretty sure the direction of the current of electrophoresis is correct.
I only used 2 wells of 12-well gel for my first electrophoresis. Can I reuse the remaining wells at the end of this electrophoresis? Could the gelred dye in the gel have been affected by the previous electrophoresis? In my view, the dye molecule cannot move without binding with DNA, so I can use the free well one more time. But someone told me I should turn around the gel if I wanna use it one more time to avoid the influence on gelred dye after the first electrophoresis.
I have run multiple agarose gels in which I have at least two combs.
I continually see the lower part of the gel visualising much fainter than the top portion.
The last band of my ladder (250 bp) often disappears. It is quite a problem as I often don't see faint bands in this area if I have loaded my PCR products in the lower portion of the gel.
See images attached - look especially at the ladders in the top vs bottom
In the one image you can see the gel itself after a run in the tank, where the loading dye is significantly lighter in the bottom portion than the top - so it's not a problem with the visualising equipment.
I have run the gel for different times (30-60 min) as well as at different voltages (100V vs 120V) and see the same.
The TAE buffer has been changed
I have observed the same with another gel tank.
The intercalating dye, SBYR Safe, has been replaced with a new aliquot.
Other individuals have also experienced the same issue
I cut a vector that already contain the promoter using BstBI enzyme. The electrophoresis gel result showed that no different between control and vector+plasmid cut BstBI enzyme. There are more than 1 band found in the gel. Does anyone have any idea why it is this way?
I am in the process of conducting an electrophoresis and I have a Safer dye at my disposal. However, I am uncertain whether it would be more suitable to combine the Safer dye with the 6X DNA LOADING Buffer. Would it be advisable to include the Safer dye in the loading DNA buffer mixture before incorporating it into the DNA sample? Or is such a procedure unnecessary?
Strain: Sphingomonas, Gram negative
The organisms were harvested during the logarithmic growth period and centrifuged at low temperature to remove the medium.Extraction of RNA revealed bands of abnormal size, no band at 23s, but two very close bands near 16s, and electrophoresis results showed no degradation.
I would like to ask if anyone has encountered this?
Thanks for looking and replying!
DNA amplification and electrophoresis, the target band was not as expected, the fragment was recovered and re-amplified with the same primers, and the correct size band appeared, but the recovered fragment was ligated into the vector and picked for cloning for Bacteria Liquid PCR, and fragment larger than the expected size was amplified (?). . I am no longer able to answer such a situation with my current knowledge, does anyone know what went wrong?
Usually, I use a time of 120 minutes and 120 V, but I am testing and standardizing new primers and I realized that perhaps the sample might have run off the gel. I would like to know if you use any protocol considering base pairs to decide on the time and voltage.
I prepared TAE for running electrophoresis and would like more information about how long of an expiration period I can consider.
Is it possible that the amplification failure products can be visualized in electrophoresis? Due to the failed amplification results it shows bands in my electrophoresis with bands that are quite clear. My amplification curve clearly shows amplification failure, but when I look back at it with electrophoresis there are some obvious bands, how is that possible?
During the electrophoresis run on a 1% agarose gel at 100 volts for 25 minutes, I was unable to clearly visualize the expected 28S and 18S bands representing ribosomal RNA. Instead, a smear-like pattern appeared on the gel. The ladder was run correctly, indicating that the gel and electrophoresis setup were functioning properly.
To provide context, I measured the concentration of my total RNA using a nanodrop spectrophotometer, which yielded a reading of 2.07 in A260/A280. The concentration was determined to be 1352 ng/μl, and for the gel analysis, I loaded 3 μl of RNA with 1 μl of loading buffer and only 3 ul Ladder.
Considering these results, I am unsure about the integrity of my total RNA and whether the observed smear is indicative of degradation or other factors affecting the sample quality. I am reaching out to this esteemed community in the hopes that someone with experience in RNA electrophoresis can offer insights into potential causes for the observed smear and absence of distinct 28S and 18S bands.
If nanodrop facility is not available, can we predict the dna conc. from the electrophoresis picture?
Molecular weight of my protein of interest is 7KDa. I am not able to get this on 18% SDS-PAGE electrophoresis. I want to do western blotting of my samples having this. Kindly suggest the method to separate the low molecular wt protein and western blotting of it.
I want to do MALDI-TOF mass for my protein. For its procedure I need to do bis tris electrophoresis of immunoprecipitated protein, then excise protein band. For running bis tris electrophoresis, I need to solve my protein in LDS sample buffer, but I think that would i replace SDS with LDS in this buffer? Is it too different for doing mass spect?
When purifying my DNA amplicons, I need to excise my DNA of interest from my gel (1.2% agarose).
I've observed that the DNA is distributed top and bottom of the thickness. Out of curiosity, I've done some research into why, but I can't find it. Does anyone have the answer?
I did several times gel agarose electrophoresis to find the best N/P ratio of chitosan/tRNA polyplex. I don't know why I could not observe the tRNA in the well of gel agarose electrophoresis and the whole gel matrix? Even there is no smearing.
Would you please help me to interpret my result?
Thanks for your time and consideration.
Hey everyone. I need to mark LC3 protein, which has 16-14 kDa. When I do a 14% gel for the electrophoresis after the 20 kDa, my gel bands stay wavy (as shown in the photo) so it is not possible to get the protein.
Has anyone experienced this and could give some advice?
Good afternoon! We are trying to do bisulfite conversion in a new way and we want to evaluate DNA integrity after bisulfite conversion by agarose gel electrophoresis. For tests, we use 5 µg of mouse liver DNA. At the output, we get a concentration of 50 ng/µl. For electrophoresis, we take 4 µl of converted DNA bisulfite. We use 1% agarose gel, 1x TAE buffer and ethidium bromide okara. The duration of electrophoresis is 1 hour at a voltage of 120. After electrophoresis, a five-minute incubation on an ice bath enabled partial hybridization and visualization of the DNA. But we don't see any bands. Perhaps someone can tell what we are doing wrong?
When the PCR product was electrophoresed, such a band was visible and smear was seen under the band.
Cycle conditions were (1) 94.0°C for 3 min, (2) 94.0°C for 10 sec, (3) 55.0°C for 10 sec, (4) 72.0°C for 7 sec, (5) 72.0°C for 1 min, and (6) 20.0°C for 10 min.
The number of cycles is 40 cycles from (2) to (4).
The swimming time was 15 min.
Gels were 3% agarose gels.
I would like to know what causes such smears and how to improve them.
We did comet assay based on joves protocol. Under fluorescence microscope, cells were visible but without tail. The drugs showed toxicity when grown in 6-well plate. Lysis was done for overnight at 4 degree celsius. Electrophoresis was run at 25V, 300mA for 30mins.
I'm doing a SOE PCR. As soon as the PCR is done, I run electrophoresis of the samples. The gel shows the marker but no sample, as if I didn't load the sample with the loading buffer. But then, I run again the same samples in the same electrophoresis chamber with the same conditions and it shows results. What can it be? This happened to me 4 times, and I'm losing my mind. I couldn't find any answers.
Looking in to alternatives for Etbr for our DNA electrophoresis. I've done some research on GelRed and Sybersafe but I've been hearing conflicting reviews. I'd like some insight in to other labs and their results. Is green dye better than red? Are you seeing bleeding? All insight is welcomed.
I am trying to use the Quiaxcel advanced capillary electrophoresis system for fragment analysis of PCR products that have been cut with a restriction enzyme.
I dissolve my ladder (size marker) in PCR buffer, Restriction enzyme buffer (containing BSA) and EDTA in similar concentrations to the ones in which my samples are dissolved. However, doing this the resolution of the ladder is not clear and I can't see the expected peaks in my samples.
Does anyone knows if any of the buffers I am using influence the electrophoretic run, or does anyone that has used the machine before has any suggestions?
Thanks a lot!
I´m haveing trouble with some samples I´m trying to determine Serpina 5 content by ELISA assay. I´m using a comercial strip kit.
With my director we are starting to think the protein is forming some kind of agregate that is blocking the recognition site for the antibodye or something like that, so we were thinking of doing a cracking step, like in denaturing electroforesis, to the samples before diluting for the assay.
Is this posible? has someone done it and can share your experience?
Thanks a lot for any help you can give!
I was working on vertical electrophoresis using polyacrylamide gel to separate DNA fragments, but I have encountered different problems, so I need your technical advice to resolve my problems.
The first problem is that the polyacrylamide solution takes a very long time to polymerize or solidify in the plate. I used 50ml polyacrylamide solution per plate by mixing 13.3ml (30% acrylamide bisacrylamide), 10ml (10x TBE), 0.350ml (10% ASP), 26.35ml (water) and 5µl (TEMED).
The second problem is that migrating DNA form a parabola shape after moving halfway from the well. So it ultimately gives the band of different sizes (those expected to be the same size).
The third problem is that the obtained band was not bold enough for scoring. I used ethidium bromide for staining the gel after electrophoresis.
I think about tool can automatically interpretate my hemoglobin electrophoresis diagram
Is this exist or applicable ?
I am doing PCR-RFLP to detect SNP, after the restriction enzyme digestion I check with 15% polyacrylamide gel, if the restriction occurs I should see two bands (126 and 20bp) but I only see the 126bp band, what could I be doing wrong or how do I solve it?
We want to know the charge of a dipeptide (Mw < 1kD). Dipeptide is based on aspartic acid and its negatively charged. So, to prove that we have carried out 3% agarose gel experiment, tracking dye we used was bromophenol. We applied 150 V during electrophoresis. The idea is since the peptide is negatively charged, so it can move towards positive electrode during electrophoresis. Thus we can experimentally prove that the peptide is negatively charged.
We made the wells in the middle of agarose and loaded the dipeptide along with tracking dye which is bromophenol in 1:1 ratio. So upon electrophoresis,we observed the band was dragged towards the positive end. After that we stained gel in Coomasive Brilliant Blue (CBB) for 3 to 4 h and followed by destaining using a reported protocol. The problem we observed that after destaining for longer time also we can not see the peptide band where it actually moved, still after 18 h, the gel fully looked blue and no bright spot for peptide.
So, I think staining may not be proper as I donot know what type of interaction can be between CBB and my peptide (-vely charged).
So, please suggest a dye which can stain a -vely charged dipeptide to see its band in gel electrophoresis. Is agarose or SDS PAGE better to visualize a small charged peptide during electrophoresis. Also suggest, if any major concern with this method.
I was wondering if the routinely used NativePAGE sample buffer with a pH of 6.8 causes protein precipitation if my protein has a pI of 7? Could I increase the buffer pH without affecting the electrophoresis? Thanks a lot for your help.
I have done a blue-white screening with E.coli and pUC19 with lambda DNA. Then, I selected some white colonies (and blue colonies as a control) and gel electrophoresed them next to pure pUC19. All lanes have a band similar to pure pUC19. I am stuck on why this is the case. Any help is appreciated!
I have a small but old electrophoresis device that is full of a thick mass of salts from the TBE buffer, I have tried to remove it just with water but it doesn't come off and it is starting to make the device to leak. Maybe someone had a similar issue?
We have performed southern blot analysis for several years but this problem appeared first time. The 1-1.5% agarose gel running for 18-20 hours in cold room(4C) under the 55V melted.
One day I and my colleague switched on two separate gels on one Power Supply and this happened (picture).
50xTAE ( Tris 242g/L, 0.5M EDTA pH 8.0, 57.2 ml/l glacial acetic acid).
After that we started to set up only one electrophoresis gel but the picture was the same. We thought that something happened with the buffer ( by the way we used 1xTAE buffer), so we change the buffer to a fresh one and saw the same picture. The same picture was when we change the Power Supply, the place where we switched on the electrophoresis and also when we changed the gel box(tank) to a new one. So we have changed everything except the agarose itself but we used it before and nothing bad happened.
Did something similar happen to anyone?
Thank you so much in advance for any response.
I induced hairy root by A. rhizogenes ATCC15834, then I did PCR test in hairy roots to detect rolB, rolC and virG. However, there was only rolC and virG shown in electrophoresis result. I have repeated 3 times, however there were nothing changes. How can I explain for this result ?
There is a non-specific band with the extracted plasmid going to run electrophoresis. About 500bp. non-specific bands for both enzyme cut identification and control. Extraction reagent is fine.15000bp marker.
I have done electrophoresis of my PCR product 70 V for 40 min and the bands were moving away from the ladder range what is the correct way to do electrophoresis?
I am currently extracting a specific protein (hordein) from whole barley grain and would like to determine the molecular weight using SDS-gel electrophoresis. Before running the gel, I need to quantify the protein but need to dilute it beforehand. I decided to use 75% EtOH for the dilution since that was the percentage of EtOH that I used for the extraction. I know most dilutions use DI water but since it is a prolamin protein, it is not water-soluble. Any thoughts on whether I am proceeding this correctly? Many thanks!
I' m facing some problems with my western blot. The bands of markers seems normal in the gel after electrophoresis, but they drift after transferred to the membrane (showing in the picture)? Does anyone know what could be the reason? I've met this a few times.
Thank you so much!
Dear colleagues. I had a problem with gel electrophoresis, namely with a dna ladder (100 bp and 1 kb). Usually I put electrophoresis on 115 V and for 30 minutes. You can see the results in the attached picture. What I also tried to do: 1) change the voltage to 80V and increase the time to 40 minutes; 2) replaced the TAE buffer with a new one (I use 1X TAE); 3) added different volumes (0.5; 1; 1.5; 2 µl) of simple rulers or diluted in loading buffer in 1/8. But the results are still bad. In the attached photo, I used 1.2% agarose, 1X TAE buffer, 115 V and 30 minutes. The volume of the ruler varied from 0.5 to 2 µl (the first 4 strips are 1 kb ruler, and the second 4 strips are 100 bp).
I have extracted the gDNA of E. coli 25922 using Qiagen DNeasy Blood & Tissue kit. In order to confirm successful extraction, gel electrophoresis was used to visualise the length of extracted genomic material. Theoretically, according to gen bank gDNA of E. coli 25922 is 5,152,857 bp. However, the largest DNA ladder available to me is the Lamdba Hind iii ladder, possessing a range from 125 - 23,130 bp. When the extracted gDNA and DNA ladder undergo electrophoresis, the bands produced by the gDNA samples align with the 23,000 bp band.
Gel electrophoresis conditions were 0.7% agarose, 80 V, 1.5 hours
I would like to perform, among other things, electrophoresis with preserved proteins (not denatured).
To do so, I need to first extract proteins with a "soft" buffer.
The best would have been M-TER, but I was wondering if B-TER would do the job (because that's what I have in the lab).
(In order to increase the yield of proteins extraction, I planned to perform griding through Precellys and sonication with a probe.)
Thank you for your help.
I am running western gel. I have experienced some issues, some happened repeatedly.
1. The samples are not running at the same speed. All the samples are prepared the same time. The voltage I used for running the gel is 100V. However, the samples close to one of the electrodes run lower than the samples close to the other one. The trails of all the samples are not horizontal but tilted to one side, could someone give me some suggestions?
2.The current has been really low. When I start running gel at 100V, the current would show around 12, that's not high. Current would lower to 2/3 when the samples reached the half way of the gel. The protein would barely move down afterwards. Does someone know how it happens?
The wire/cable in my chamber (for protein electrophoresis) was cut in half. I tried to re-join it by knotting the two ends. But the current still run unevenly among running wells.
I want to ask "what is the material makes up the wire/cable?" so that I can try to weld it. The brand of the chamber is Biorad.
I would appreciate any other suggestions to repair the wire/cable.
Thank you so much!
Anyone suggest electrophoresis for interaction of Protein with DNA give detail procedure and material?????
I have performed experiment on optimisation of PCR for one of the SNPs and while performing electrophoresis, the issues occurred that I'm trying to figure out right now, and need some help with that.
The methods followed while preparation of the gel:
In order to prepare agarose gel, first preparation of 100ml of 10X Tris-Borate-EDTA (TBE) was needed. To obtain that, 1.080g of Tris Base, 0.550 of Boric Acid and 0.095g of EDTA Na2·2H2O was added to the beaker. It was then filled with reverse osmosis (RO) water, after which HCl has been added to adjust the pH of the substance to 8. Substance was stirred well while HCl and RO water has been added to achieve final volume of 100ml of the TBE. Four new, clean beakers were taken, to prepare two 2% agarose gels with 12 wells, which had the best porosity for the size of amplicon that was studied in this experiment. 5ml of TBE has been pipetted carefully into each one of them, and then diluted with 45ml of RO water to achieve 1X TBE buffer. Then carefully measured 1g of agarose has been added to two of the beakers, and 2ul of GelRed, before microwaving them for 2 minutes to dissolve all particles, after which they were poured into the gel tanks, and left to set for around 20 minutes, before removing end plates and combs. TBE buffer from the other beakers has been poured all over set gels.
4ul of loading dye has been added to each PCR tube and briefly vortexed. 5 ul of DNA ladder has been loaded into first well of each gel. Then one gel was filled in order with 12ul of PCR samples of set A, the other one with duplicate set B. Loaded gels then had been run at highest current for approximately an hour, until the samples has travelled at least halfway through the gel, after which photos of the gel image has been taken.
The problem already was obvious once the run was over, visible smiley blue line on top of the gel, which I assumed was either the issue with the pH of the TBE (however as I had TBE that I used and checked it for confirmation, it was still at perfectly 8). Another thing is that the blue line has decolorized in few spots, into greenish/grayish colour which I cannot find any explanation for in troubleshooting pages.
Then once trying to obtain gel images, it turned out gels were melted, even though the tanks were not warm at all when I took them to do the images, which was straight after turning the power pack off.
The images don't show any good results, they are having the burn line, DNA ladder didn't separated itself, there are some faint bands visible in one gel, there are visible DNA bands in the wells, so some DNA didn't even left the well.
Did something similar happen to anyone and what is the main issue starting with for this experiment?
Thank you so much in advance for any response.
The electrophoresis is OK, the gel seems OK and also the running buffer. But my samples and the marker are stucked in the well.
Any idea? many thanks.
From my understanding, proteins in the blood have distinct charges as they move through the bloodstream. If a protein had a sufficiently, say positive charge, and a negative charge was applied outside the bloodstream on the skin, would it cause that positively charged protein to accumulate near that location on the body? I am unaware of any current research on this topic and would be very grateful if anyone could point me to some.
One potential issue I see with this is if charged proteins are balanced by counterions in vivo and if this may prevent the process from working.
I´m having problems using ExoSAP-IT™ PCR Product Cleanup Reagent for PCR purification for the first time.
I´m using 5ul of PCR product with 2ul ExoSAP-IT and then two steps of incubation 50 min 37C and 15 min 80C.
The problem is I did an electrophoresis with the PCR products (1 to 5-PCR, as seen on gel image below) vs the cleaned with ExoSAP-IT (1 to 5-EXO) to physically see the cleaning but got this results:
- An inespecific band apears in the "cleaned" product, just below the interest fragment at ~400 bp.
- It does clean the primer excess though.
Has anyone had this problem?
It can be some type of contamination even though my ExoSAP-IT is brand new?
Hello everyone. I would like to clarify and get advice on single-stranded DNA electrophoresis. A 2% solution of agarose gel was used, which contained about 12 µl of ethidium bromide. The DNA marker (or ladder) was from 10-300 bp. DNA samples were used with a 4X buffer for loading in the amount of 1 µl of buffer and 4 µl of DNA sample. 1 sample from the ladder (5.5 OE), 2 (10.0) and the extreme (5.5 OE). 1X TBE was used as a buffer solution. The strength of the intense field was 90 V. And so on for 1.5 hours. But the ladder, alas, did not work. The lines with the samples are blurred at the end. There is an assumption that the concentration of agar must be increased and the voltage reduced. But honestly I don't know. Do you have any suggestions
Hello everyone, I want to make the RT-PCR Master Mix with SYBR Green l powder, so I have questions: What is the difference between the SYBR green dye used in RT-PCR Master Mix and the dye used in the gel stain for the electrophoresis??? What grade of SYBR Green l powder I must use?? Can I use a grade of Chemical raw materials, pharmaceutical to make Master Mix SYBR Green powder???
I´m in the process of troubleshooting for my cloning experiment.
One problem I stumbled across is that the size of my fragments is different before and after gel extraction. I cut a 2271bp and a 1590bp fragment each from a plasmid (I use BamHI+SalI or HindIII+SalI for this, the sizes are as expected). I run the digestion on a 1% agarose gel. After gel extraction (NEB kit) I run 1µl of the purified fragment again on a 1% gel (I use loading buffer containing Gel red). Now both fragments run slower, at about 3500bp and 2500bp respectively.
What could be the problem, maybe the buffer? Did anyone of you face a similar problem in the past?
I am having a lot of problems when running the gel . It looks like the proteins are kind of precipitating, and I don’t know why. Can you please help me to understand what is going on?
Thanks in advance!
Proteins are extracted to perform electrophoresis, so I want to save the supernatant for the other day to do Bradford. Proteases can damage it?
I performed PCR yesterday afternoon and let it run over night on setting at 6 C in the thermocycler. This morning I prepared my 2% agarose gel and after it set, I placed it in the electrophoresis chamber and loaded 10ul of the samples into the wells. The volts used were 104-105v for about an hour and a half. When I took it to the imager, I got these faint bands showing. I just would like to know where in the process did I go wrong.
I performed site direct mutagenesis for single base substitution and after DpnI digestion (2hours at 37 0C), control and mutant samples were electrophoresed in 1% agarose gel. As a control I used pWhitescript 4.5kb control plasmid. The expected band size for the control is 4.5kb, but instead the band was at 9kb. The band for the mutant plasmid was as I expected. Could you please tell why this could happened?
I am having some issues with transfer efficiency to nitrocellulose membranes. To get all the technical info out of the way:
Transfer method: Wet, tank transfer overnight at 30V with normal tris-glycine transfer buffer + 20% methanol. I do this at RT with stirring. Of note: I have not recently been equilibrating my gel in transfer buffer, as this was reported to decrease transfer of basic proteins due to the stripping of SDS (I can furnish this reference if anyone is interested).
Sample: In all cases these are in vitro reconstituted nucleosomes with or without certain enzymes--the main transfer issues are the core histones, which are highly basic (pI 10-11). There should be no more than about 100 ng of each histone being transferred per lane. The salt in the sample is <25 mM. They are boiled prior to loading. There is no noticeable precipitate in the samples before or after boiling, however the volume is quite small (20 ul) so I would not necessarily detect precipitation.
Nitrocellulose is 0.2 um and is pre-equilibrated in transfer buffer prior to transfer.
The gel: I use a 15% gel, 1.5 mm thick. The acrylamide I use is 37.5:1 acrylamide:bis. Some issues could arise here--the acrylamide solution is about 1 year old and has been stored at RT that entire time (which is what is recommended on the label). In addition, we typically keep a stock of 10% APS at 4C, so I cannot guarantee my APS has not gone bad.
I have been quite careful generally to ensure there are no bubbles between any components of the sandwich and filter paper/foam pads are soaked in transfer buffer.
Now to the issue. Attached are images of the membrane after transfer (and also after processing, thus the relatively high background from the blocking buffer) as well as the gel after transfer. As you can see from lane to lane I seem to have differing transfer efficiencies. This has occurred with my last three blots. My first thought was sample precipitation, but I have not had this issue before despite running essentially the same blot many many times over the past year.
Any thoughts? Quite frustrating, as it has appeared suddenly with identical protocols/reagents as before.