Rather than pipeting the gel in, I prefer to pour directly from a 15ml conical (or 50ml if pouring 4 gels from one tube). I find it more convenient and less bubbly - and it makes it faster if you're trying to pour 4 gels before your acrylamide polymerizes. Also if you have trouble with leaking, try turning the pads at the bottom upside down or lay a strip of parafilm (don't stretch it) along the bottom pad before seating the glass plate.
Most guides to hand-casting gels (as published by BioRad, etc) recommend that the distance from the bottom of the well to the top of the resolving gel should be at least double the depth of your sample once loaded into a well. As your sample runs down the gel, it gets backed up at the front between the stacking and resolving gels. This allows all of the proteins to start entering the resolving gel closer to the same time. So, the longer the stacking gel, the better the resolution of your bands.
Here are a few things you can try: 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!
That was incredibly useful. Performing a SDS-PAGE soon, I'm feeling content now that it's not going to be a desaster :D Thanks, good job for that video!
@TheLiona84 It can take from 15 to 45 minutes, depending on what percent gel you are making, and how much ammonium persulphate and TEMED is added. An indication that your resolving gel is ready is when you see a clean straight line form at the gel-isopropanol interface when the gel is ready.
Thank you so much for making this video!! We use butanol instead of isopropanol and rinse a buncha times with water and dry it before adding the stacking gel.
thatguy1w The width of the gel depends on the volume of the wells that you want. 1.5mm is usually ideal for 50ul wells, but 0.75mm is what we use in our lab. It's the suitable width for 30ul wells, which are the most common size for gel electrophoresis. @ dude in video, you should never allow the Acrylamide to spill over after you add the comb. That stuff is highly toxic (neurotoxic) and can even cause cancer. You should press a piece of 'Kimwipe' against the glass to suck up any spill overs.
In my lab, we just use spill trays so that the overflowing acrylamide is not an issue. It's all going to polymerise anyway, and the polyacrylamide is (comparatively) harmless. The danger is handling the monoacrylamide, for whic we have dispensers to avoid handling as much as possible.
If you have leakage, you can tape the bottom of the plates together after you place the two glass plates in the casting stands and before placing them in the casting frames. This helps ensure you wont have leakage when pouring the lower gel. It is really easy to remove the tape once the lower gel and stacking gel have polymerized.
Yes that can happen. Here are are few things you can try: 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!
1 cm is commonly used and taught, so that's what we suggest here, especially if you are making your first gel. You can leave a smaller space if it works, just make sure you have enough space so that the samples can enter the stacking gel, and have the opportunity to reach the resolving gel at the same time.
@alabonneheure1 It's probably a seal problem. Have you seen our "How to Avoid a Leaky SDS-PAGE gel" video? (there is a link in the description box above) Try the pen/microfuge tube trick. If you still have problems, let us know =)
@ishanrathore for conductivity. Distilled water acts as an insulator, therefore electrolytes are required to allow the current to flow. These are provided by buffer!
@TheLiona84 You can use water but be aware that water can slightly dilute the gels. So if you are working with a high molecular weight protein (the band appears near the top of the resolving gel), you don't want to use water because a gradient can form at the top of the resolving gel, and the protein will not separate properly. So you can use water, but if you have a high molecular weight protein, I would not recommend it.
@VegasPartier Yes, add the stacking layer until it almost overflows. Then when you slowly put in the comb, some of the mixture should overflow. And don't worry about the pipette tip - plastic is just fine! Good luck on your next gel =)
@ishanrathore well what is a buffer? Buffers provide us with an environment that tries to keep a consistent pH. So here we are using two different buffers: pH 6.8 for the stacking gel, along with pH 8.8 for the resolving gel. If we didn't use buffer, then the protein samples would migrate at the same rate in both the stacking and resolving gels.The wikipedia article on SDS-PAGE provides more detail on how this works (sorry can't post the direct link here, RUclips won't let me).
@tranceaddictallnight you want the pH to be about two pH units lower than your running buffer. In this case we are using Tris-glycine buffer for running the gel, which has a pH around 8. What is the pH of your resolving gel? And what are you using as your running buffer?
@zileburki Yes you can. Store the gel (inside the glass plates and with the comb) in a container, submerged in the Running Buffer that you use when running the gel. Cover the container with the lid. You can store it at room temp but I prefer at 4 degrees. Just remember to use the gel within a couple of days =)
I'm assuming you are doing a Western blot. I have never used a torn gel, but it should probably work as your samples are not running through the gel, but just transferring from gel to membrane. Just make sure you put the pieces back correctly! Let us know how it goes :)
@nana22124 Yes. As you see in the video at 3:10, add a layer of isopropanol to keep the gel straight, and wait until solidified (about 15 minutes). Then add the stacking gel.
You should be able to get these ingredients from any chemical company, so just ask the company from where you get your other lab supplies. Note: acrylamide can be purchased either in solid (powder) form or as a solution - I would suggest buying the liquid form to avoid handling the powder, as acrylamide is a neurotoxin. Regardless of whether you buy it in solution, or make your own solution from the solid form, be careful with acrylamide.
Hello, your video is so interesting and really helpfull to understand SDS PAGE's gel making. I have a question that I wanna ask. Why you should you tris HCl 8.8 in resolving gel and tris HCl 6.8 in stacking gel? What's difference between the two kind of tris HCl? Thank you
oops! sorry i accidentally deleted @admarshall617's question when responding: the question was: "how long does it take for the stacking gel and the resolving gel to solidify " our response is: "It can take from 15 to 45 minutes, depending on what percent gel you are making, and how much ammonium persulphate and TEMED is added. An indication that your resolving gel is ready is when you see a clean straight line form at the gel-isopropanol interface when the gel is ready."
We got the same apparatus for this. Did you remove the corners from the comb? Because we got some corners there and bubbles always come up over time and the outside bags are ruined then every time
Thank you for the demonstration. I have a question about the time limit that I can postpone loading the wells by my Lysate. Can I make the gel and load it the day after? How I need the keep it and prevent it from drying out? Thank you.
I generally make SDS gels in advance, wrap them up individually with paper towels, wet them with milli Q and then store it at 4 degrees. I’ve used 1-2 week old gels and they worked fine.
Using filter paper will not harm gel in any ways? What about if it leaves some of it's constituents in the liquid and it get solidified with the gel and can later effect your sample protein?
I do everything the same, but I usually get bubbles forming at one end of the combs, so I end up losing a well. Sometimes it's ok since there's enough gel wall for loading my ladder. Any recommendations?
Thanks for the nice video. Lately sometime I fail to make perfect well, and I realize it when I load my sample it won’t go through coz there are this sheet of the gel between the well. Might anyone have a suggestion? Many thanks
For some reason, when I prepare the gel, I notice small webs formed inside the well, which is why when I load the protein, the proteins don't run well. I prepared new apps and tried again, but somehow encountered the same problem, Could you suggest what should I do ?
Why do you put so much stacking gel solution on top? I know it condenses during/after polymerization, but is it dangerous to spill the solution when you put in the comb? Or is it the concentration & amount of Acrylamide that makes the difference?
@labtricks Can we use ethanol instead of isopropanol ? Does it affect the gel contents ? And if we can use ethanol is it absolute or diluted with water?
Question: Why 1 cm below the comb marked?? Why not, less?? Ive seen in my lab some people leave kind of .3cm between the comb and the solving gel... is it correct? thank you!! great videos!!
If the gel tears during transfer onto a cassette sandwich, can it still be salvaged if there is an attempt to put the pieces back together? Or should the gel be binned? I just transferred my samples from a torn gel onto a PVDF membrane and I hope my attempt to "reform" the gel is not in vain...
Hello, first of all thank you for this video. I am trying to get a vertical gel electrophoresis system from Bio-Rad but they have been so difficult to work with. It seems to be a mix of language barrier and laziness, or maybe I am too frustrated so I think they are being lazy and it's just the language barrier. Can anyone please tell me of their experience with this company and recommend other sources for the vertical gel electrophorsis system?
in our lab we have two casettes and a spacer. and we add water to create a cushioning effect or something..howz that different than what you have done here?
I always have a problem with stacking gel...i don't know it won't polymerize. could u pls let me know the recipe for preparing stacking gel. thanks i like ur videos
Hi, i have a question about Mini-PROTEAN TGX Stain-Free precast gels... there is number on the packet "4-15%"... What do the numbers mean separately? 4? 15?
+Shaig Shaggy The numbers correspond to the percentage of acrylamide in the resolving gel. A 4-15% gel is a gradient gel, with the lowest percentage of acrylamide just below the stacker being 4%, and the highest percentage is 15% near the bottom of the gel. The percentage increases from the top to the bottom of the gel. A single percentage gel would have one number, i.e. 10%.
When I add the acrylamide solution to the casette, I mark the level with a permanent marker then add propan-2-ol and let it polymerize. On polymerization, the actual gel gets lower than the mark so the size of the gel varies every time from one gel to another. Any suggestions?
Never had problems casting gels for westerns till this week where all my gels have this white opaque appearance instead of the normal clear appearance once harden. Do you have any idea what's going on?
I have a ques When i put plates in casting stand and check it with water either it put correctly or not...the water not leak..but when i starting puring gel it starts leaking from plates
I'm having problems with the stacking gel, it's not fully polymerizing. It gets a slimy consistency but never fully forms. We've tried to make new Acrylamide mix, new APS and new buffer but it still isn't working.
I hope you figured this out already! How old is your acrylamide stock? IT does go bad over time and may act as if you used a lower % of acrylamide. This might be more noticeable in the stacking gel as it never gets as solid as a resolving gel. Also do you have the right mix of bis/acrylamide?
For those who may read this later. Acrylamide will not polymerize in the presence of oxygen. Make sure it is fully sealed at the top by the comb (this is the true reason for the alcohol or water layer). If you need vigorous mixing, your recipe is bad. A quick swirl with the pipette tip after dispensing should be all that is needed. Double check the acrylamide/bis-acrylamide ratio, it is very possible you are mis-understanding the ratio and percentage of the stock. APS takes a while to go bad, I used to pour many very large gels, and I often used APS that was months old. Keep APS and Acrylamide in the fridge, and mix them right before you use them.
Thanks to Riasat202 and ***** for replying. Yes you having to put the stacking gel on top. That's where the samples are loaded. The purpose of the stacking gel is to let all samples flow through and reach the resolving gel at the same time. Then at the resolving gel, the proteins will separate out.
How did you prepare your solution? Maybe we can try to figure out what went wrong. Otherwise, here is an example recipe for the stacking gel (note that recipes can vary, so this is not the only way to make it): 1.5mL 30% acrylamide/bis-acrylamide solution 2.5mL 0.5M Tris-HCl, pH 6.8 0.1mL 10% SDS 5.9mL dH2O 50uL 20% ammonium persulphate 10uL TEMED
Jesus... The amount of uncertainty I have in the lab going off verbal descriptions sucks. It's so easy seeing it visually.
i feel that. all about visuals ;)
True
Amen
Ang
Same here! Reading it: Wait, what?. Seeing it: Oh. That's easy.
Rather than pipeting the gel in, I prefer to pour directly from a 15ml conical (or 50ml if pouring 4 gels from one tube). I find it more convenient and less bubbly - and it makes it faster if you're trying to pour 4 gels before your acrylamide polymerizes. Also if you have trouble with leaking, try turning the pads at the bottom upside down or lay a strip of parafilm (don't stretch it) along the bottom pad before seating the glass plate.
Most guides to hand-casting gels (as published by BioRad, etc) recommend that the distance from the bottom of the well to the top of the resolving gel should be at least double the depth of your sample once loaded into a well. As your sample runs down the gel, it gets backed up at the front between the stacking and resolving gels. This allows all of the proteins to start entering the resolving gel closer to the same time. So, the longer the stacking gel, the better the resolution of your bands.
Here are a few things you can try: 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!
At what% the isopropanol solution has to be added
Hi, I'm currently a biology student learning to be a lab technician with a professor who's bad at explaining things. This video is very helpful.
That was incredibly useful. Performing a SDS-PAGE soon, I'm feeling content now that it's not going to be a desaster :D Thanks, good job for that video!
My teacher used your video as an instruction video, be proud
@TheLiona84
It can take from 15 to 45 minutes, depending on what percent gel you are making, and how much ammonium persulphate and TEMED is added. An indication that your resolving gel is ready is when you see a clean straight line form at the gel-isopropanol interface when the gel is ready.
Thank you so much for making this video!! We use butanol instead of isopropanol and rinse a buncha times with water and dry it before adding the stacking gel.
thatguy1w The width of the gel depends on the volume of the wells that you want. 1.5mm is usually ideal for 50ul wells, but 0.75mm is what we use in our lab. It's the suitable width for 30ul wells, which are the most common size for gel electrophoresis.
@ dude in video, you should never allow the Acrylamide to spill over after you add the comb. That stuff is highly toxic (neurotoxic) and can even cause cancer. You should press a piece of 'Kimwipe' against the glass to suck up any spill overs.
1.وقدتملحصولI
In my lab, we just use spill trays so that the overflowing acrylamide is not an issue. It's all going to polymerise anyway, and the polyacrylamide is (comparatively) harmless. The danger is handling the monoacrylamide, for whic we have dispensers to avoid handling as much as possible.
If you have leakage, you can tape the bottom of the plates together after you place the two glass plates in the casting stands and before placing them in the casting frames. This helps ensure you wont have leakage when pouring the lower gel. It is really easy to remove the tape once the lower gel and stacking gel have polymerized.
Yes that can happen. Here are are few things you can try: 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!
1 cm is commonly used and taught, so that's what we suggest here, especially if you are making your first gel. You can leave a smaller space if it works, just make sure you have enough space so that the samples can enter the stacking gel, and have the opportunity to reach the resolving gel at the same time.
@alabonneheure1 It's probably a seal problem. Have you seen our "How to Avoid a Leaky SDS-PAGE gel" video? (there is a link in the description box above)
Try the pen/microfuge tube trick. If you still have problems, let us know =)
@ishanrathore for conductivity. Distilled water acts as an insulator, therefore electrolytes are required to allow the current to flow. These are provided by buffer!
Awesome video!! the plates and the casting frames looks exactly like the ones we have in UBC...
@TheLiona84 You can use water but be aware that water can slightly dilute the gels. So if you are working with a high molecular weight protein (the band appears near the top of the resolving gel), you don't want to use water because a gradient can form at the top of the resolving gel, and the protein will not separate properly.
So you can use water, but if you have a high molecular weight protein, I would not recommend it.
You just saved me from my practical today!!🙏🏼 thank you
Yes you can use ethanol instead of isopropanol to line the top of your resolving gel (as shown at 3:12 in the video)
@VegasPartier Yes, add the stacking layer until it almost overflows. Then when you slowly put in the comb, some of the mixture should overflow. And don't worry about the pipette tip - plastic is just fine! Good luck on your next gel =)
@TheLiona84 and yes, after it solidifies, pour off the isopropanol (or whatever you are using) and then add the stacking gel.
@ishanrathore well what is a buffer? Buffers provide us with an environment that tries to keep a consistent pH. So here we are using two different buffers: pH 6.8 for the stacking gel, along with pH 8.8 for the resolving gel. If we didn't use buffer, then the protein samples would migrate at the same rate in both the stacking and resolving gels.The wikipedia article on SDS-PAGE provides more detail on how this works (sorry can't post the direct link here, RUclips won't let me).
WOW thanks :) I'm doing my first gel tomorrow... it doesn't look so confusing now
@tranceaddictallnight you want the pH to be about two pH units lower than your running buffer. In this case we are using Tris-glycine buffer for running the gel, which has a pH around 8.
What is the pH of your resolving gel? And what are you using as your running buffer?
@zileburki Yes you can. Store the gel (inside the glass plates and with the comb) in a container, submerged in the Running Buffer that you use when running the gel. Cover the container with the lid. You can store it at room temp but I prefer at 4 degrees. Just remember to use the gel within a couple of days =)
I'm assuming you are doing a Western blot. I have never used a torn gel, but it should probably work as your samples are not running through the gel, but just transferring from gel to membrane. Just make sure you put the pieces back correctly! Let us know how it goes :)
@nana22124 Yes. As you see in the video at 3:10, add a layer of isopropanol to keep the gel straight, and wait until solidified (about 15 minutes). Then add the stacking gel.
You should be able to get these ingredients from any chemical company, so just ask the company from where you get your other lab supplies. Note: acrylamide can be purchased either in solid (powder) form or as a solution - I would suggest buying the liquid form to avoid handling the powder, as acrylamide is a neurotoxin. Regardless of whether you buy it in solution, or make your own solution from the solid form, be careful with acrylamide.
@pkgem we use neat (pure) isopropanol, straight out of the bottle
Hello, your video is so interesting and really helpfull to understand SDS PAGE's gel making.
I have a question that I wanna ask. Why you should you tris HCl 8.8 in resolving gel and tris HCl 6.8 in stacking gel? What's difference between the two kind of tris HCl? Thank you
The different pH-value might be because of the protein denaturated structure. But I am not sure what the exact reason is
oops! sorry i accidentally deleted @admarshall617's question when responding:
the question was: "how long does it take for the stacking gel and the resolving gel to solidify "
our response is: "It can take from 15 to 45 minutes, depending on what percent gel you are making, and how much ammonium persulphate and TEMED is added. An indication that your resolving gel is ready is when you see a clean straight line form at the gel-isopropanol interface when the gel is ready."
nigga you saved my life
Rodolfo Osuna Rodolfo shut up nigga. He saved mine too. And he's cute. Shiii.
LMAOO
We got the same apparatus for this. Did you remove the corners from the comb? Because we got some corners there and bubbles always come up over time and the outside bags are ruined then every time
@labtricks we use 8.8 pH Tris for both resolving and running gels. what is the purpose for the lower pH in the resolving gel?
Thank you for the demonstration. I have a question about the time limit that I can postpone loading the wells by my Lysate. Can I make the gel and load it the day after? How I need the keep it and prevent it from drying out? Thank you.
I generally make SDS gels in advance, wrap them up individually with paper towels, wet them with milli Q and then store it at 4 degrees. I’ve used 1-2 week old gels and they worked fine.
Using filter paper will not harm gel in any ways? What about if it leaves some of it's constituents in the liquid and it get solidified with the gel and can later effect your sample protein?
I do everything the same, but I usually get bubbles forming at one end of the combs, so I end up losing a well. Sometimes it's ok since there's enough gel wall for loading my ladder. Any recommendations?
Thanks for the nice video. Lately sometime I fail to make perfect well, and I realize it when I load my sample it won’t go through coz there are this sheet of the gel between the well. Might anyone have a suggestion? Many thanks
For some reason, when I prepare the gel, I notice small webs formed inside the well, which is why when I load the protein, the proteins don't run well. I prepared new apps and tried again, but somehow encountered the same problem, Could you suggest what should I do ?
Why do you put so much stacking gel solution on top? I know it condenses during/after polymerization, but is it dangerous to spill the solution when you put in the comb? Or is it the concentration & amount of Acrylamide that makes the difference?
Thank you for creating this video! very kind of u guys .
@labtricks
Can we use ethanol instead of isopropanol ? Does it affect the gel contents ? And if we can use ethanol is it absolute or diluted with water?
Those are automatic captions that youtube generates...so they are not accurate, but they are definitely entertaining to read sometimes :)
Hi. I hv 2 questions. Why add isopropanol to removes bubbles and why is there stacking and resolving gel ?
Also are the solutions purged with N2 beforehand? Cause the biorad manual says to do that
Why we are adding isopropanol ? Can we add water instead?
Please answer
What is the tris buffer for, you don’t need it for the polymerization and TEMED/APS work at RT?
thanks @labtricks
but even when we use single gel (native PAGE) or also agarose gel then also buffer is used instead of water
Used my split gel for a western and it worked fine :)
@poxyratarsed Did you remember to add isopropanol? That is what makes the resolving gel straight =)
You guys are awesome! Thank you!
Question: Why 1 cm below the comb marked?? Why not, less?? Ive seen in my lab some people leave kind of .3cm between the comb and the solving gel... is it correct? thank you!! great videos!!
Have u got any protocol for prepration of the reagents.
Put a blue tip round the back, forcing the spring to clamp down further.
If the gel tears during transfer onto a cassette sandwich, can it still be salvaged if there is an attempt to put the pieces back together? Or should the gel be binned? I just transferred my samples from a torn gel onto a PVDF membrane and I hope my attempt to "reform" the gel is not in vain...
how thick is the gel supposed to be? 1.5 mm? will all molds have the thickness?
Thank you so much for this video! it is incredibly helpful!
Hello, first of all thank you for this video. I am trying to get a vertical gel electrophoresis system from Bio-Rad but they have been so difficult to work with. It seems to be a mix of language barrier and laziness, or maybe I am too frustrated so I think they are being lazy and it's just the language barrier. Can anyone please tell me of their experience with this company and recommend other sources for the vertical gel electrophorsis system?
in our lab we have two casettes and a spacer. and we add water to create a cushioning effect or something..howz that different than what you have done here?
Yes, very helpful. This isn't very hard as compared to a lot of thins, but I am always worried that I will screw it up.
I always have a problem with stacking gel...i don't know it won't polymerize. could u pls let me know the recipe for preparing stacking gel.
thanks i like ur videos
Hi, i have a question about Mini-PROTEAN TGX Stain-Free precast gels...
there is number on the packet "4-15%"...
What do the numbers mean separately? 4? 15?
+Shaig Shaggy The numbers correspond to the percentage of acrylamide in the resolving gel. A 4-15% gel is a gradient gel, with the lowest percentage of acrylamide just below the stacker being 4%, and the highest percentage is 15% near the bottom of the gel. The percentage increases from the top to the bottom of the gel. A single percentage gel would have one number, i.e. 10%.
+Julie B Thank you for your answer...
when you cast the resolving gel do you wait for it to solidify before adding the stacking gel?
WHAT IS THE QUALIFICATION REQUIRED FOR THIS JOB , WHAT ARE THE PROSPECTS OF THIS AND WHAT ARE THE PRODUCTS THAT CAN BE MADE ?
Brilliant, I would like also to add that Bio-Rad company also sell Pr-prepared gels. good idea for assuring high quality results.
Again thank you,,
When I add the acrylamide solution to the casette, I mark the level with a permanent marker then add propan-2-ol and let it polymerize. On polymerization, the actual gel gets lower than the mark so the size of the gel varies every time from one gel to another. Any suggestions?
isopropanol absorbs the water from resolving gel.
how long do you leave the water in to check if the plates are leaky?
I've got a question..
How to prepare sample buffer and loading sample too for sds.??
Never had problems casting gels for westerns till this week where all my gels have this white opaque appearance instead of the normal clear appearance once harden. Do you have any idea what's going on?
Cool gut a molecular biology lab tomorrow and this was handy
We use 8.8 pH TRIS for our stacking gel in our lab. What is the reason for the lower pH in yours?
how I'm going to choose what percentage of polyacrylamide I need to use? I mean there ie 8% 12% and so on?
Every step of SDS PAGE is clearly demonstrated...!
I have a ques
When i put plates in casting stand and check it with water either it put correctly or not...the water not leak..but when i starting puring gel it starts leaking from plates
What is the reason
how long does it take for the gel to be solidify in 4C?
I use glicerol for the resolving gel. (0,4 mL for gel to 10 %)
can we use ethanol instead of isoproponal to remove bubbles?
plz can we use pure water in place of isopropanol?and what is the diference?
how m time we should wait for running gel solidification, and then we add thestaking gel?thx
@tranceaddictallnight we use 8.8 pH Tris for both resolving and running gels. what is the purpose for the lower pH in the resolving gel?
QUESTION: what do you do if you get leaks during the leak test?
Take your plates out and restart.
why are buffers used in gel preparation & not water ? plz with detailed reason
I'm at work so reading the subtitles. Starting from 0:29 it gets hilarious.
I'm having problems with the stacking gel, it's not fully polymerizing. It gets a slimy consistency but never fully forms. We've tried to make new Acrylamide mix, new APS and new buffer but it still isn't working.
I hope you figured this out already! How old is your acrylamide stock? IT does go bad over time and may act as if you used a lower % of acrylamide. This might be more noticeable in the stacking gel as it never gets as solid as a resolving gel. Also do you have the right mix of bis/acrylamide?
do you vortex your stacking gel mixture after adding aps and temed? I've had the same problem and now its gone
For those who may read this later.
Acrylamide will not polymerize in the presence of oxygen. Make sure it is fully sealed at the top by the comb (this is the true reason for the alcohol or water layer).
If you need vigorous mixing, your recipe is bad. A quick swirl with the pipette tip after dispensing should be all that is needed.
Double check the acrylamide/bis-acrylamide ratio, it is very possible you are mis-understanding the ratio and percentage of the stock.
APS takes a while to go bad, I used to pour many very large gels, and I often used APS that was months old.
Keep APS and Acrylamide in the fridge, and mix them right before you use them.
The amount of uncertainty I have with the music is greater than that for the procedure itself c:
Is the casting stand necessary?
"security accordion" dibs on my new band name
Why isopropanol is not added to stacking gel ?
Is it ok to reverse the order in which the resolving gel and the stacking gel are casted?
Stacking gel is where you stack your samples! So no.
Thanks to Riasat202 and ***** for replying. Yes you having to put the stacking gel on top. That's where the samples are loaded. The purpose of the stacking gel is to let all samples flow through and reach the resolving gel at the same time. Then at the resolving gel, the proteins will separate out.
3:04 "led the charge for the mars" lol these captions are hilarious.
Leakage at 3.14!...just when he added isopropanol😅😅😅
what is the function of APS and why we need to use it just 10%?
hello. I would like to know, where can I get those ingredients? thanks
THIS VIDEO CLEARLY EXPLAINS THE WHOLE PROCEDURE OF MAKING SDS PHAGE.....
im having problems with the wells leaking, when i wait for gel to harden and load my dye it leaks to another well.
Change your comb, take a wider spaced comb. Try removing gently after solidification.
Why do we add isopropanol?
Thank you! This helps a lot.
Thank you for the effort!
Hallo,
How to make a sds-page separating gel (5-14%)???
How did you prepare your solution? Maybe we can try to figure out what went wrong.
Otherwise, here is an example recipe for the stacking gel (note that recipes can vary, so this is not the only way to make it):
1.5mL 30% acrylamide/bis-acrylamide solution
2.5mL 0.5M Tris-HCl, pH 6.8
0.1mL 10% SDS
5.9mL dH2O
50uL 20% ammonium persulphate
10uL TEMED