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How to Run an SDS-PAGE gel

Okay now that you know how to cast an SDS-PAGE gel, how exactly do you use it? Here, Dan shows you how to set up your apparatus to run SDS-PAGE — everything in 7 easy steps.

WAIT!!
Don’t have a gel? Watch this video on How to Make an SDS-PAGE gel.

[see below for some important points to remember]

Some important points to remember:
- short glass plate faces inwards
- remove the comb carefully
- load samples inside the wells, and do not over fill (usually 10uL of sample is loaded)
- fill the inner chamber to the TOP with buffer
- fill the BOTTOM of the outer chamber with buffer
- attach power supply: RED to RED and BLACK to BLACK

DONE!

If you have any questions, feel free to comment below, or just email us: info@labtricks.com.

  1. confused
    i have a problem when i try loading my gel, my sample doesn't go all the way to the bottom of the well!! it like there's something stuck in my well. it's not a big problem cuz the gel runs fine, but it's annoying, so how do i fix/avoid this????
    • Alisha
      Another reason why this could be happening is that there might be some acrylamide precipitate on the wells. Before you load your sample, wash the wells. Try it, it might work.
  2. labtricks
    There are 2 possibilities for what is happening: 1) Your sample is really viscous -- having a high protein concentration, could cause this. The main problem would be pipeting the sample into the well. But in the end, it shouldn't affect your gel. If it's hard to pipette, try adding more loading buffer to your sample. And remember to boil and centrifuge your sample before loading. 2) Your using the wrong comb, or a bad comb -- make sure that if you have glass plates that say 1.0mm then use a 1.0mm comb. Using a smaller comb will cause a thin gel layer to form in the space between the comb and the glass plates. So you end up with wells that aren't as nicely formed, and your samples don't load well. An old comb can cause this problem too. But either way, the gel should run fine as the stacking gel should allow all the protein samples to catch up before entering the resolving gel....but yes it is annoying when this happens!
  3. Lily F.
    I just found you on YouTube. I'm a grad student and I wanted to show my undergrad assistant how to assemble the SDS-PAGE apparatus & how to run the gel. You even give away advice! You guys rock. Loving your site so far, hope to see more lab tricks!!
  4. labtricks
    Hi Lily, glad you like the site. Let us know if you have any questions :)
  5. trucskku
    My major is not in this field, but i did some related exp.. Your clips are so helpfully for me. Thank you so much. I hope I can learn more from your web.
    • labtricks
      Thanks! If you have any questions, just post them here or email us!
  6. Juls
    Excellent video tutorial! This is great for newbies as well as a teaching tool. I have bookmarked this site and can't wait to see more lab tricks.
  7. Squintz
    Excellent! Wish we could have saw the results.
  8. Sunny
    Hello, I've run gels before in one of my labs, but I'm just now trying to do it by myself. Your videos were very helpful in assembling the apparatus and preparing the gel, but when I connected the power supply, the entire inner chamber started bubbling and turned white/cloudy. If I remember correctly, the bubbles are expected, but I'm not sure about the cloudiness. The buffer was entirely clear when I poured it in, so I'm doubting it was contaminated, but I've never seen this happen before and was slightly confused. Any thoughts on this?
  9. linerss
    i would think it's normal regarding the cloudiness, as long as your buffer is correct.
  10. naresh
    Hi Which buffer you have used to fill the tank?
    • labtricks
      Hi Naresh, The buffer in the tank is the SDS Running Buffer which contains Tris, SDS, and Glycine.
  11. Hans Wespennest
    wow! nice!!!
  12. Naresh
    Hi can you please tell me how to prepare load sample.
    • labtricks
      Hi Naresh, Mix 10uL of your protein sample with 10uL of 2x loading buffer, then boil for about 5 minutes. After boiling, give your samples a short spin in the centrifuge to bring the sample to the bottom of the tube. Then you can load! (you can load 10uL of your sample, or all 20uL, it's up to you) The loading buffer contains Tris-HCl buffer, glycerol, SDS, bromophenol blue, and beta-mercaptoethanol.
  13. Rameez
    My protein is of a high molecular weight(~350kd).Its an huntingtons protein. Can u tell me which gel and buffer system should I use. I tried with SDS-PAGE and also native gel but its not working. Should I use a gradient gel?? Please guide me.
    • labtricks
      Hi Rameez, For SDS-PAGE, did you use a low percent acrylamide solution? That might work because it will help the protein migrate further on the gel.
  14. Ashwini
    hey..found you on youtube.. I have run the SDS PAGE before.. but I am facing problems with the well formation... the sample diffuses completely into the stacking gel.. Your video is very helpful ... keep updating.. Thank you..
    • labtricks
      Hi Ashwini, I'm confused about your question - do you see this happening when you load, or while running? Are you not seeing any bands in the resolving gel (after staining)? Let me know exactly what the problem is so I can be more helpful :) Also, always make sure your ammonium persulphate solution isn't too old or else your gels will not form properly. And double check your stacking gel buffer too.
  15. Zoey
    hi i have a little problem. after doing all, and applying 200volts my samples won't run down the gel, they just sit there in the wells. i also tried 120volts. what could the problem be?
    • labtricks
      Hi Zoey, What's in your running buffer? Make sure it contains Tris, Glycine, and SDS. Also, if you are using the standard Laemmli Tris-Glycine setup, make sure you have the following: -Running buffer with its natural pH of about 8.3 (no pHing step is usually required for this buffer). -Resolving/separating gel is a few tenths of a unit higher than running buffer (at pH 8.8). -Stacking gel is about 2 units below running buffer (at pH 6.8). -Sample buffer pH is usually the same as the stacking gel at pH 6.8.
  16. addy
    I work with protein expression in yeast and I'm facing the problem that apparently some proteins present in my samples are staying in the border of the stacking and resolving gels. When doing gel filtration of these samples there is any high molecular weight species. We suspected that some oligomers were forming when the sample got in contact with SDS gels that is an oxidizing environment so we tried reducing gels (using bis-tris in gel buffer and sodium bisulfite and mops in the running buffer). However, this partially resolved the problem. I have been suggested that some proteins don't like glycerol, so I was wondering if you could give me an alternative to glycerol for the sample buffer. Thank you for your help
    • labtricks
      Hi Addy, Quick question - does your loading buffer contain a reducing agent like b-mercaptoethanol (BME) or DTT? When you mix your sample with loading buffer containing BME or DTT, and then boil it for about 5 minutes before loading on the gel, that usually takes care of oligomers. So if your loading buffer doesn't have a reducing agent, try adding BME or DTT to it first, and hopefully that should solve the problem :)
  17. Fariha
    Hi, I am having a problem regarding SDS-PAGE. Usually I start running PAGE at 10mA for 30 min till the samples come out of stacking. Afterwards, current is increased to 20mA. Voltage stays the same throughout that is 250V. I see that some samples are running properly as can be seen by dye front (it should be uniform for proper running). Problem is that the dye does not stay uniform and makes strange smiley effect or sometimes even worst than that. In the end I do not get any result after staining and destaining :(. As I guess the problem is with running. Please help. Another problem is that sometimes while running two gels together, one gives the bands properly run but the second one has no band at all :(. Why is it so that one gel is properly run and the other one is not? while all the conditions are the same? I am using Laemillae system for sample buffer and running buffer. Kindly help
    • labtricks
      Hi Fariha, My first suggestions were going to be for you to lower your voltage, double check your buffer components, and make sure the inner chamber of your tank is full with buffer. However, it is interesting that you are able to get one gel to work when you run two of them. It seems like there may be a problem with your electrode assembly unit, for only allowing a gel on one side to work but not the other. Make sure to rinse the unit thoroughly and use fresh buffer for your next run. Also double check the buffer dam (plastic dummy plate) that you use when you only have one gel. If you notice one side of your electrophoresis unit to constantly cause problems while the other side works, then that would indicate that something is wrong with your unit. That's all I can suggest for now as I have never had this problem. Let me know how it goes. Good luck!
  18. Ann Li
    Hi, What is the difference between running a gel at constant current and running it a constant voltage?
    • labtricks
      Hi Ann, It depends on whether you are using a continuous or discontinuous system. For the procedure shown here (discontinuous, mini gel system) constant voltage is usually used. Some systems require constant current for better resolution. But note that high current will increase the amount of heat generated. Using a constant low current can help control the heat, but it may also take longer to run. There’s a nice table that summarizes this on page 351 of this manual: https://www.u-cursos.cl/faciqyf/2012/2/FBQI4204/1/material_docente/objeto/638204&ei=sRf3UIyELqGFiAKO1YBY&usg=AFQjCNGFreTr4YdXZhuBqzng1ZTJToG93Q -Suraaj
  19. jimy
    Hello, My name is jimy. I have a problem in SDS PAGE. My sample is protease, which I extracted and purified from mango. PH 6.11. Before running SDS PAGE, my protein concentration is 100 microgram/mL which I think enough for SDS PAGE. But unfortunately, the band doesn't appeared. I ad tried different protein concentration which is in tha range 100-500 micrgram/ mL, but still can get. I had used different % of resolving get (7.5-12.5%), 0.1% coammasie brilliant blue as staining solution and 10% acetic acid+ 25% methanol + 65% deionized water as destaining solution., but stiil can't get. My parameter is 200V, 30 min. Hopefully someone can help me.
    • labtricks
      Hi Jimy, If you've tried different concentrations, and different % resolving gels and your band still doesn't appear, I wonder if your sample is being degraded. Either due to self-cleavage (as it is a protease) or another contaminant which is cleaving it, and thus the fragments are running off the gel. This could be a possibility. Have you looked into the stability of your protein yet? Check different temperatures, pH, and double check your purification protocol - maybe you are losing the sample through that step. Good luck! -Suraaj
  20. Jey
    hey thanks a lot for the video! I wanna know can we reuse the running buffer and also pls tell me what i sthe running buffer level I'm using Medox dual Here is the link http://www.medoxbio.com/dual_vertical1.asp
    • labtricks
      Hi Jey, Yes you can use the running buffer a few times - I've reused buffer 2 or 3 times at the most. Your system looks similar to the Bio-Rad one shown in this video, so you can fill up the buffer to the same level as demonstrated here. However, double check with your system's manual. -Suraaj
  21. HuiShan
    Hi, i've recently carried out SDS-PAGE and noticed that airbubbles tend to form in the sample wells after about 10-15mins of inserting the sample well comb into the gel cast even though during insertion of the sample well comb there was no air bubbles, this was especially so during the cold season. could this be due to the weather? and this was more prominent in tricine-sds-page. thanks!
    • labtricks
      Hi HuiShan, This question has been asked many times and has also been answered on our YouTube channel. Here are a few things you can try to avoid this from happening: 1) Make sure your comb is clean and dry. Any water droplets can create bubbles in the gel and cause the wells to become deformed. 2) If you are using isopropanol/ethanol etc. to straighten your resolving gel, remember to pour it out before putting in the stacking gel. It can also help to wash with some water before adding stacking gel. 3) Increase your APS concentration. I like to use 20%, and that can help with polymerization. Good luck! -Suraaj
  22. Devina
    Hi.. I just did western blot, and I had problem that the protein didn't transfer to the membrane. Unfortunately, I didn't check whether the protein is still in the gel, but I stained the membrane and there was no protein. What I want to ask is, after I run the gel, I turned off the power, and left it for maybe 30 minutes, I just read in some guideline that we can not leave the gel because the protein will elute from the gel. Do you think it might be the reason? Thank you. Devina.
    • labtricks
      Hi Devina, Is your protein very small? If it was near the bottom of the gel, perhaps it could have leaked out of the gel. Or maybe you had low amounts of protein to start off with, so the transfer may not have been as great. Personally, I've had experience with leaving the gel in the tank after running SDS-PAGE and I can still see the protein on the gel after staining. But note, in those cases I had high concentration of protein in my lanes and didn't use the gel for Western blot. Are you sure there weren't any problems during transfer of the protein from gel to membrane? Maybe there's another reason for why you don't see protein on your membrane.
  23. Shruthi
    Hi, I am running gels for studying HeLa cell lysates. I use cruz marker and generally run 10% gels. Somehow my cruz marker shows dumb bell bands after the chemiluminiscent detection and I inferred referring that this may be due to the hot run. I use constant current conditions for the mini gel. So, I tried reducing the current conditions from 18mA to 16mA but no difference is seen. To what minimum value can I decrease the current value. Also, is this a good troubleshooting I am following or is there anything else I am supposed to rectify? Thank you!