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Why should I degas my gel solution for polyacrylamide gels?

Why should I degas my gel solution for polyacrylamide gels?



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In protocols for polyacrylamide gel electrophoresis (PAGE) I often see instructions to degas the gel solution by putting it under vacuum for 10-15 minutes before polymerizing the gel.

I usually don't do this, and when I tried it once I couldn't see any difference. So I'm wondering what exactly the degassing is meant to achieve and how big the effect should be.

  • What effect is degassing the gel solution supposed to have?
  • How important is degassing to achieve good gels?

Any literature that examines the effect of degassing would be appreciated.


The reason for degassing your gels is to remove oxygen. Oxygen in the gel interferes with polymerisation, slowing it down and making it less consistent, so degassing makes it faster and more uniform.

From the EncorBio SDS-PAGE protocol:

Polymerization is quicker and more uniform if you degas the first three solutions for a few minutes in an Ehrlenmeyer flask on a house vacuum prior to addition of the last three reagents. Molecular oxygen inhibits polymerisation by reacting with the free radical SO4- ions, which is actually the reason why PAGE gels are poured in tubes or between plates and not in open top horizontal apparatuses, as can be done with agarose. Also it's a good idea to layer some isopropanol on top of the gel as this prevents oxygen getting in and inhibiting polymerisation.

Oxygen can also lead to oxidation of protein products, which might be crucial if you then want to extract the products and use them for something else (e.g. Sun & Anderson, 2004).

Finally, having bubbles in your gel can distort the results and make them less reproducible, as the bubbles will not form consistently with each repetition and they disrupt the physical medium of the polyacrylamide. So another purpose of degassing is to ensure repeatability.

The Bio-Rad acrylamide polymerisation info sheet has the best info I could find:

The formation of polyacrylamide gels proceeds via free radical polymerization. The reaction is therefore inhibited by any element or compound that serves as a free radical trap (Chrambach 1985). Oxygen is such an inhibitor. Oxygen, present in the air, dissolved in gel solutions, or adsorbed to the surfaces of plastic, rubber, etc., will inhibit, and in extreme cases prevent, acrylamide polymerization. Proper degassing is critical for reproducibility. Therefore, one of the most important steps in the preparation of polyacrylamide gels is the evacuation, or “degassing” of gel solutions immediately prior to pouring the gel. This is done by placing the flask of gel solution in a vacuum chamber or under a strong aspirator. In some cases, a vacuum pump may be required.

Buffer stock solutions and monomer stock solutions are usually stored at 4°C. Cold solutions have a greater capacity for dissolved oxygen. The process of degassing is faster and more complete if the gel solution is brought to room temperature (23-25°C)' before degassing begins. Furthermore, if a cold gel solution is placed under vacuum, the process of evacuation tends to keep the solution cold. Pouring a gel with a cold solution will have a substantial negative effect on the rate of polymerization and on the quality of the resulting gel.

Polymerization in which riboflavin is used as one of the initiators calls for degassing. The conversion of riboflavin from the flavo to the leuco form (the species active in initiation) actually requires a small amount of oxygen (Gordon 1973).

This explains why polymerization initiated primarily by riboflavin can be completely blocked by exhaustive degassing. However, oxygen in excess of that needed to convert riboflavin to the active form will inhibit polymer chain elongation, as it does in reactions initiated only by ammonium persulfate and TEMED. Thus, while degassing is still important for limiting inhibition, it must not be so extensive that it prevents conversion of riboflavin to the active form. For polymerization initiated by riboflavin/TEMED, or riboflavin/TEMED/ammonium persulfate systems, degassing should not exceed 5 min.

A consequence of the interaction of riboflavin with oxygen is that riboflavin seems to act as an oxygen scavenger. This is supported by the observation that the addition of riboflavin (5 µg/ml) to stacking gel solutions containing ammonium persulfate/TEMED initiators results in cleaner, more uniform polymerization at gel surfaces exposed to oxygen (such as combs). The same effect could likely be achieved by more thorough degassing of solutions without riboflavin.

Whether using chemical polymerization (ammonium persulfate/TEMED) or photochemical polymerization (riboflavin/TEMED or riboflavin/TEMED/ammonium persulfate initiators), reproducible gel quality and separation characteristics require careful attention to gel solution temperature before degassing, and to degassing time, temperature, and vacuum. These parameters should be kept constant every time gels are prepared.

Sorry for the long quotes, but they are pasted here in case the original sources disappear.

References:


I have been running gels with different Acrylamide/Bisacrylamide ratios recently. People usually work with 1:37.5, 1:29 ratios which are commonly used for DNA and Protein gels. I have noticed that when you work with lower ratios 1:200 - 1:500, degassing becomes fundamental to guarantee reproducible resolution of my proteins. If I don't degass the mix in one of these gels some of the proteins I am resolving (which migrate really close to one another) won't separate well enough. Also, the time it takes to polymerize the gel can go from 20 to about 45 minutes if I don't degass the solution beforehand. Degassing solutions with normal 1:29, 1:37.5 crosslinker ratios doesn't, in my experience, seem to have much of an effect other than having quicker polymerization times. Perhaps it makes a difference with lower concentration gels (8-5%), but I honestly wouldn't worry too much about this if I was routinely running 10-15% gels and reproducible resolution wasn't a concern.


What could cause gel not to polymerize (solidify)? - gel making (Jun/23/2008 )

Am I missing anything? Do any of the above reagents require storage at a particular temperature prior to use? Why are my gels not solidifying like they used to?

Hi everyone. My running gels are not hardening. I've added

6 mL 30%acrylamide/0.8%bisacrylamide,
3.75 mL pH8.8 3M tris HCl,
100 uL 20% SDS
5 mL water

Am I missing anything? Do any of the above reagents require storage at a particular temperature prior to use? Why are my gels not solidifying like they used to?

I will try it and get back to you. u are potentially my day saver cellcounter!

Hi everyone. My running gels are not hardening. I've added

6 mL 30%acrylamide/0.8%bisacrylamide,
3.75 mL pH8.8 3M tris HCl,
100 uL 20% SDS
5 mL water

Am I missing anything? Do any of the above reagents require storage at a particular temperature prior to use? Why are my gels not solidifying like they used to?

Cellcounter is right. APS is the main culprit. It's very important that your store your dry APS in a dessicator. It will go bad over time if you don't (it happened to me!)
Clare

You guys were right on the money. As soon as I remade APS, I used fresh APS and what doyou know? My gel was rock hard(exaggerating) within 30 minutes.

You guys were right on the money. As soon as I remade APS, I used fresh APS and what doyou know? My gel was rock hard(exaggerating) within 30 minutes.

Hi everyone. My running gels are not hardening. I've added

6 mL 30%acrylamide/0.8%bisacrylamide,
3.75 mL pH8.8 3M tris HCl,
100 uL 20% SDS
5 mL water

Am I missing anything? Do any of the above reagents require storage at a particular temperature prior to use? Why are my gels not solidifying like they used to?

i've read somewhere that mixing the gel mixture around too much can cause oxygen saturation which then impedes gel polymerization. any truth in that?

Hi everyone. My running gels are not hardening. I've added

6 mL 30%acrylamide/0.8%bisacrylamide,
3.75 mL pH8.8 3M tris HCl,
100 uL 20% SDS
5 mL water

Am I missing anything? Do any of the above reagents require storage at a particular temperature prior to use? Why are my gels not solidifying like they used to?

i've read somewhere that mixing the gel mixture around too much can cause oxygen saturation which then impedes gel polymerization. any truth in that?

That is true. But unless you mix it like a nut, it should eventually polymerise.

Atmospheric oxygen is a free radical scavenger that scavanges SO4 - ions produced by the inherently unstable APS necessary for the polymerization.

In the good old days, people also used to deaerate acrylamide mix to make gels polymerize.

I was still deaerating the acrylamide mix to get the oxygen out from the solution last year (just in case). But now I found out it is not a necessity anymore. They can still polymerize even without deaeration!


Why won't my SDS-PAGE stacking gel polymerize? October 23, 2012 6:34 PM Subscribe

My lab manager has decided we will now be pouring our own SDS-PAGE gels. I pulled out the years-old equipment and tasked myself with figuring out how to do this. The resolving gel polymerizes nicely, but the stacking gel gets no farther than goo (and sometimes not even that far).

I'm using 12% resolving gel, 5% stacking gel. Recipes below:

Resolving gel (12%, 1ml solution) - does polymerize:
334ul DI H2O
250ul 1.5M Tris, pH 8.8
400ul 30% acrylamide/0.8% bis-acrylamide
5ul 20% SDS
10ul 10% APS
1ul TEMED

Stacking gel (5%, 1ml total) - Does not polymerize
549ul DI H2O
260ul 0.5M Tris, pH 6.8
170ul 30% acrylamide/0.8% bis-acrylamide
5ul 20% SDS
10ul 10% APS
1ul TEMED

Now, I tried doubling and tripling the amount of APS and TEMED (decreasing the water accordingly). I even set up a test series where I did 5x, 10x, and 20x APS and TEMED. Still no polymerization.

Solutions are at room temperature. APS is from a fresh bottle and made fresh with each attempt. TEMED is fresh. The only thing not fresh are the dry Tris, acrylamide, and bis-acrylamide reagents. The solutions themselves are fresh. Could the acrylamides have gone bad? But like I said, the 12% resolving gel is polymerizing.

Solutions are made in a yellow-light room, but when put out in white light polymerization is still unsuccessful.

Any experienced gel-makers have ideas?

Oxygen inhibits acrylamide polymerization, so maybe you need to degas your solutions. Or, if the volumes are impractically small, degas the water you use to make them.

How are you keeping air away from your stacker? Does the comb completely cover the top of the acrylamide solution? If any of it is exposed to air, you can carefully layer water-saturated butanol over the top with a pasteur pipet it will float and provide a barrier while the acrylamide polymerizes, then you can pour it off and rinse the gel with a little running buffer before loading your samples.
posted by Quietgal at 7:45 PM on October 23, 2012 [1 favorite]

Ask the lab manager to compare the cost savings vs. the amount of money wasted paying you to try to get this up and running, and the work lost by the delays. The savings are probably already more than offset.

I still make a lot of my own solutions and etc., but when it comes to this kind of thing my lab group buys precast gels. No safety worries about unpolymerized acrylamide, no wasted time, better consistency and reproducibility.
posted by caution live frogs at 6:52 AM on October 24, 2012 [1 favorite]

Response by poster: Purchasing is not an option, manager would prefer us spend the time getting it running to buying the gels.

I layered not-water saturated butanol over it but it didn't make a difference. Should I try water-saturating it first?
posted by schroedinger at 7:44 AM on October 24, 2012

A mix of comments and questions that may or may not be helpful:

-Are you mixing the gel solutions? I do invert my Falcon tubes to get the APS and TEMED mixed in a bit, but you don't want to aerate the gel mix with, say, vortexing.
-You say "APS bottle" - I assume you mean a container with dry APS, right, not a solution? (Because it totally does go bad when aqueous, and I make it fresh from powder every time because life's too short to spend re-pouring stupid gels.)
-How long are you waiting for the stacking gel to polymerize? A little longer doesn't necessarily hurt (heck, overnight polymerization is even preferable in some ways, though it can be inconvenient and isn't necessary for standard SDS-PAGE setups.) And a stacking gel will always be more gooey than a resolving gel - you can observe this when you take things apart after a run so that you can stain the gel.
-You say "solutions are at room temperature." Have you been storing your acrylamide solution at room temperature? The solution should be stored at 2-8°C. (In the dark, preferably, unless you go through it fast.)
-Have your acrylamide powders been stored in the dark (for plain acrylamide)? At 2-8°C (for the N,N′-Methylenebis(acrylamide))?
-Your volumes seem pretty small, and with a high stacking to resolving gel ratio. If you're using something other than more-or-less standardly sized mini gels (e.g. the ones you'd make for a Bio-Rad Mini-Protean rig), might that be relevant?

While yeah, you can degas your buffers, I have never needed to do so except for DNA sequencing gels, and I have never needed to layer the tops of my stacking gels with butanol or isopropanol. You shouldn't need to resort to extreme measures for standard protein SDS-PAGE. If I had to bet, I would bet on your acrylamide being iffy (either due to storage conditions/age of the powders or the solution), and that the problem is worse in the stacking gel because there is less acrylamide in it. Can you mooch a few mLs of premixed acrylamide solution off of a nearby lab that does gels regularly, or buy 100mL from Sigma or someone (since it looks like you are pouring pretty standard gels, not high C% Schägger gels or whatever)? That would at least confirm or refute the possibility that your acrylamide is at fault. Ditto for the Tris, if you're feeling paranoid.
posted by ubersturm at 9:36 AM on October 24, 2012

Best answer: I don't actually make only 1ml gels, those are just the recipes for 1ml to make it easier to compare the components. The stacking gel was left to polymerize overnight, and it was too gooey for any wells to form, period.

think I solved this one. The acrylamide powders were not stored in the dark and are about seven years old. I borrowed some fresh pre-made acrylamide/bis-acrylamide solution and everything polymerized just fine. All the info I looked at online addressed old APS and TEMED, but it looks like acrylamide can go bad too. Thanks all!
posted by schroedinger at 2:15 PM on October 24, 2012


How does it work?

There are several types of PAGE technique that are used, but the most common is called SDS-PAGE. In SDS-PAGE the detergent Sodium dodecyl sulfate is used to denature the proteins and normalise their mass-to-charge ratio. Without SDS, both the molecular weight and the charge of the protein would affect its separation in the gel. With SDS, only the molecular weight affects the migration speed and so samples separate according to this. PAGE without SDS is called native PAGE, as the proteins stay in their native conformation.

The Gel Matrix

As with agarose gel electrophoresis, the samples are separated using an electrical field, and pass through a gel matrix which influences the migration of the proteins. In PAGE, rather than agarose, we use a chemical called polyacrylamide. Varying the percentage of polyacrylamide in the gel lets us change the size of the pores in the gel, which means that we can separate different sizes of protein in different percentage gels. Typical gel percentages are shown in the table below.

Acrylamide PercentageSeparating Resolution
5 %60 – 220 Kd
7.5 %30 – 120 Kd
10 %20 – 75 Kd
12%17 – 65 Kd
15 %15 -45 Kd
17.5%12 – 30 Kd

Acrylamide is normally sold in a liquid form, as the powder form is neurotoxic an dangerous to handle. Polymerisation is achieved by mixing acrylamide with bis-acrylamide, which allows cross-links to form between the acrylamide molecules. Additional chemicals are added to initiate the polymerisation, usually ammonium persulphate as a source of free radicals and TEMED as a stabiliser. Once the polymerisation begins the gel is poured between 2 glass plates and allowed to completely polymerise.

The gel mixture is made up not in water but in electrophoresis buffer (Tris-HCl), that provides the ions for electrophoresis. Often, the gel is poured in 2 parts. The first parts is a resolving gel, with a pH around 8.8 which slows the migration of the proteins. Above the resolving gel, a stacking gel is poured with a pH of 6.8 and a larger pore size. This stacking gel works to compress the protein samples into a thin migration front, so that all the proteins in the sample arrive at the resolving gel at the same time, leading to an accurate relative migration.

Running the gel

Unlike in agarose gel electrophoresis, where the gels are cast in trays are run horizontally, SDS-PAGE gels are cast vertically using a casting apparatus. we cast the gels in this way so that the stacking and resolving gels form a continuous gel, which would be much more difficult in a horizontal gel. It also allows a much greater protein amount to be loaded onto the gel. The gel tank is also split into 2 sections. Depending on the manufacturer the tank will have an inner (or upper) buffer chamber, and an outer (or lower) buffer chamber. These two chambers are linked by the gel to create a continuous circuit. Each chamber contains and electrode, negative in the inner chamber and positive in the outer chamber. The inner chamber contacts the top of the gel, and when an electrical field is applied, the proteins will mode towards the positive electrode in the outer buffer chamber, due to the negative charges of the SDS molecules. Typical buffers for SDS-PAGE are Tris-Glycine for the buffer chambers, and Tris-HCl for the gel.

The speed of movement through the gel is then determined by the voltage gradient, i.e. the voltage between the electrodes. The required field strength is related to the size of the gel tank being used and the required voltage can be calculated using the simple equation E = V/d where E is the field strength, V the voltage and d the distance in cm between electrodes. Vertical gel tanks are generally run at 5 – 10 V / cm so if your tank has an electrode distance of 10 cm, you would run the gel at 50 – 100V. The exact value depends on your samples and should be determined empirically.

To apply this electrical field, we use a DC power supply. Most electrophoresis power supplies can be set to provide either a constant current or a constant voltage, with each having advantages and disadvantages. One potential issue is the production of heat due to the flow of current through the system which can be especially high with larger tanks that require higher voltage. For this reason, it is advisable to use some form of cooling, either passive in the form of a cooling block, or active such as a recirculating chiller, for larger electrophoresis systems.

Visualising the Protein

After migration, proteins must be visualised to determine their length and abundance. There are several methods commonly used to visualise proteins that are either specific or non specific. Non specific protein visualisation targets all proteins, using dyes that bind to common regions of the proteins such as the amino groups. Examples of non-specific protein stains include Coomassie Brilliant Blue and Ruby Pro. These stains are often non-reversible and can (but don’t always) interfere with downstream applications. Non-specific staining can be useful for quickly quantifying samples in a gel, or for ensuring a sample of interest is present.

To specifically visualise certain proteins, we need to use antibodies. Antibodies recognise unique 3 dimensional structures in the protein to distinguish them from others. By conjugating the antibody with a dye or enzyme, we can visualise just the protein that it recognises. The process of moving proteins from a gel to a membrane that can be probed with antibodies is called western blotting. To Learn more about western blotting, you can read our dedicated article soon. Whether you are western blotting or just non-specific staining, you will need to visualise the proteins using the gel documentation system. The type of system you use will vary based on whether you are using a visible stain like coomassie, or fluorescent or chemiluminescent dye attached to an antibody. As with agarose gel documentation systems, gel documentation systems for proteins can come in a variety of specifications depending on the requirements. For more advice on gel docs you can read our dedicated article.


Can someone tell me about extraction DNA from polyacrylamide gel? - (Nov/16/2006 )

Hey everybody, I'm very glad to join this forum. I'm a final year student and I have to do some experiments with SSR (simple sequence repeat) indicator. Now I'm preparing to cloning some genes and I have some troubles. It's the DNA extraction stage, but from 6% silver- staining polyacrylamide gel (normally, in my lab, they just extract from agarose gel). I need some advices about the protocol and the difference between the ethidium bromide - staining polyacrylamide gel and the silver - staining one which I have to noticed before I go to extract DNA.
The interested band is about 300 bp and I have to electrophoresis PCR products on polyacrylamide gel because there're many bands which have their size near the band I need (e.g 350bp, 270bp..). Can I use the QIAEX II Gel extraction Kit of QIAgen for this purpose? Other than this Kit, is there any procedure I can use? (this Kit is too expensive, so if there's any cheaper way, it's better)
Please help me, thank for your help!!!

if your DNA band were weak and contains less amount of DNA, whcih can only detected by silver stained band on PAGE, you might cut the band and put it into boiled water for elution, then reamplied for purification,
if your band is strong and can be detected by EB which is less sensitive than silver staining. you might followed instruction of QiaexII kit.

have you ever boiled polyacrylamide gel slice in water, I don't believe that the recovery DNA is enough to do the reamplification because in hot water, most of material in these slices will be denatured.
How can you stain gel with EB, or it's the same way with agarose gel?

yes
check how people recovery DD-RT band from PAGE gel , they even boil the gel slice for 15mins, I just put boiled wate into tube with cutted band.

stain PAGE with EB is the same as agarose gel, put gel in EB solution (it is better using the same buffer for electorporesis)
by the way SBRY green is more senstive for staining of DNA in PAGE gel.

, thank you so much for your answers! But my gel size is about 33 X 35 cm, so it's too difficult and unsafe when trying to stain it in EB solution. Do you know about another procedure which runs the gel slice in 2% agarose and obtain DNA on the DEAE membrane which is embeded below wells? i found it when I'm searching more information but I don't know how I can buy this membrane.

mini size PAGE should work for such size products (such as gel used on BioRad mini II)
alternatively low melting temperature agrose might be considered

I thought about it before and when i asked my teacher, she said that the mini size PAGE isn't appropriated because the length of it isn't enough to discriminate between this band and some another bands approach to it. I implied the electrophoresis apparatus which is used for protein (about 10 X 13 cm ?), it would be easier for me to work with it

10% non denature PAGE with better resolution for 300bp DNA around

You're a kind man! I will think more about all your advices. Hopefully I will successful with my experiments but I think that there will be more troubles I have to face, so I will post all of unclear things with me and glad to receive more help!


Acrylamide gel preparation - for SDS-PAGE (May/17/2007 )

Hello, I have some basic questions about Acrylamide gel preparation:

1. I would like to use a disposable 10ml/50ml tube for preparing the gel solution, but I'm worried that the surface area will be too small for degassing. Must I use an Erlenmeyer flask?

2. I'm buying acrylamide solution, but the solution, TEMED, and APS still seem like dangerous stuff. Must I prepare and cast the gel in a fume hood? When cleaning the plates etc., should I be concerned about monomers and take any precautions beyond wearing gloves?

3. Can I dispose of the polymerized gel in standard trash, or is it hazardous waste?

When dealing with TEMED or DTT, yes, do it in fume hood. And of course when cleaning the plates, use gloves. Well, they claimed that polymerised gel will be nothing at all. So you can basically throw in a bin.

Hello, I have some basic questions about Acrylamide gel preparation:

1. I would like to use a disposable 10ml/50ml tube for preparing the gel solution, but I'm worried that the surface area will be too small for degassing. Must I use an Erlenmeyer flask?

2. I'm buying acrylamide solution, but the solution, TEMED, and APS still seem like dangerous stuff. Must I prepare and cast the gel in a fume hood? When cleaning the plates etc., should I be concerned about monomers and take any precautions beyond wearing gloves?

3. Can I dispose of the polymerized gel in standard trash, or is it hazardous waste?

I usually do it in a 50 mL centrifugation tube. It's fine. you can also use the tube several times. try not to have foam on the surface of the liquid, unless you will aspirate a lot under vacuum. mix very gently. For most of purpose, I don't degas. I mix very very gently.

you can dispose the polymerized gel in standard trash, but only the polymerized gel. Sometime the gel left in the 50 mL tube doesn't polymerize so well, so I add some APS to the acrylamide left after pooling the gel, so it's perfeclty polymerized and I can throw it away.

Hello, I have some basic questions about Acrylamide gel preparation:

1. I would like to use a disposable 10ml/50ml tube for preparing the gel solution, but I'm worried that the surface area will be too small for degassing. Must I use an Erlenmeyer flask?

2. I'm buying acrylamide solution, but the solution, TEMED, and APS still seem like dangerous stuff. Must I prepare and cast the gel in a fume hood? When cleaning the plates etc., should I be concerned about monomers and take any precautions beyond wearing gloves?

3. Can I dispose of the polymerized gel in standard trash, or is it hazardous waste?

Here is some suggestions arised from my experience

I prefer to degass gel solution ( resolving ) to enhance band resolution ( keep in mind that many other factors influence resolution like sample preparation, and this is only one but important ). Only degass before adding SDS to prevent intensive foam formation. don,t worry about 10-50ml tubes - I usually degass in these tubes. For fast degasation put magnetic stirrer into solution because viscosity varied depends on %acrylamide in working solution and decrease rate of degasation. I only not recommend you to degass working solutions when you prepare gradient gels. Because one of mixing solutions usually have high % acrylamide ( near 20%) and if you remove air bubbles you remove so main polymerisation inhibitor oxygen and rate of polymerisation will increase and may clogg your mix chamber ( several years ago I had this problem).

Concerning purchasing acrylamide solution. I prefer to prepare fresh stock ( every month) from solid acrylamide. Because duringacrylamide solution storage acrylic acid concentration increase. It is also decrease resolution of your PAAG. But you can solve this problem by incubating your acrylamide solution with Amberlite MB 150 resin ( Supelco or analog in Merck or Fluka). Store your stock with this resin ( covering bottom of storage bottle) in dark bottle at 4C.

Concerning dangerous of APS . Don't worry about this (APS) and work in lab without hood. About Temed - it is only amine with strong smell and so only don't leave it open after using ( aliquote it into eppendorf tubes under hood if you buy 1L bottle and then keep in 4C and work easy).

GOOD LUCK! and don't big worry about this!

Thanks for the suggestions everyone, this is helpful. I'll try my first SDS-PAGE gel today.


What is the physical appearance of CHAPS detergent?

This biodetergent is supplied as a white crystalline powder.

What is the critical micelle concentration (CMC) of CHAPS detergent ?

The CMC value is 6-10 mM. Detergents with high CMC values are generally easy to remove by dilution detergents with low CMC values are advantageous for separations on the basis of molecular weight. As a general rule, detergents should be used at their CMC and at a detergent-to-protein weight ratio of approximately ten.

In 0-0.1 M Na+, what is the aggregation number?

The aggregation number at this level is 4-14.

What is the transition temperature?

The cloud point is greater than 100°C

What is the TLC of CHAPS (silica gel: methanol/NH4OH = 95/5)?

Is CHAPS detergent soluble in water?

Yes, this detergent becomes a clear and colorless to slightly-yellow solution in water. Solubility is 50 mg/mL at 20°C. It is preferable to avoid shaking and stirring mixtures when preparing CHAPS solutions.

How can I make a CHAPS 10% solution?

For a 10% solution (i.e., 1 g CHAPS solid into beaker followed by 9 g of water), cover beaker with watch glass and allow it to sit at room temperature for 30-60 minutes. Solutions of up to 1 M (60%) can be made in this way.

What is the stability of this detergent?

CHAPS biodetergent is moisture sensitive and hygroscopic.

Can CHAPS detergent be removed from samples by dialysis?

Yes, this detergent’s small micellar molecular weight and high critical micelle concentration (6-10 mM) allow it to be removed from samples by dialysis.

What is the difference between CHAPS and CHAPSO detergents?

A related detergent, called CHAPSO , has the same basic chemical structure with an additional hydroxyl functional group.

Both detergents have low light absorbance in the ultraviolet region of the electromagnetic spectrum, which is useful for laboratory workers monitoring ongoing chemical reactions or protein-protein binding with UV/Vis spectroscopy.

What concentration of CHAPS is used for preparing IEF gel?

Concentrations between 2-4 % are typically used in an IEF gel. CHAPS is commonly used for non-denaturing IEF (without urea) and has been shown to give excellent resolution of some subcellular preparations and plant proteins.

What is the difference between CHAPS and Triton X100?

Triton X100 is non-ionic. It has both hydrophilic and hydrophobic regions, but no net charges. CHAPS detergent is zwitterionic. It has hydrophobic regions but also a head group with a negative charge (in normal saline). In addition, Triton X100 forms large (greater than 90,000 MW) aggregates when concentration rises above 0.25 mM. CHAPS on the other hand forms smaller aggregates (6,000 MW) when concentration rises above 10 mM.


FAQ: Why are there extra bands visible on polyacrylamide gels?

For the 100bp DNA Ladder, the 500 and 517bp fragments, which run as a close doublet on agarose, separate very clearly on acrylamide, with the 517bp band running around midway between the 500 and 600bp fragments. This &ldquoanomalous&rdquo migration is an inherent characteristic of acrylamide gel electrophoresis and does not indicate any error in the stated size of the DNA fragments in our ladder.

For the 50bp DNA Ladder and the Low Molecular Weight Ladder, two or more of the bands comprising the 200bp reference band tend to run differently, resulting in one or more extra bands detectable around the 200bp range. As with the 100bp DNA Ladder, this is attributed more to the limitations of acrylamide gel technology than to a problem with the ladder composition. As defined in most molecular biology lab manuals (Maniatis&rsquo Cold Springs Harbor Molecular Cloning Manual, 2nd edition) and as acknowledged by the manufacturers of acrylamide gels, applications requiring precise sizing of DNA fragments should be performed using agarose gels whenever possible.


SDS-PAGE "Hall of Shame"

You may very well have prepared a nearly perfect gel, and would have a difficult time improving upon the product. If that is so, then by all means gloat about it! If you didn't do such a hot job don't despair. All good scientists learn from their mistakes, in fact, without making mistakes most of us wouldn't learn much at all!

The "Hall of Shame" presents examples of some of the worst gels students (and instructor) have run in past labs, with an example or two from a research lab. They represent many of the ways one can mess up a gel (but not all of them - we're still finding new ways!). See what features of your own gel(s) were unsatisfactory - or at least less than perfect - and use the illustrations to figure out what you might do to improve your technique.


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GelRed® is much more sensitive than EtBr, and can be used to stain dsDNA, ssDNA or RNA in agarose gels via either precast or post gel staining without destaining. GelRed® can also be used to stain polyacrylamide gels via post gel staining. GelRed® is also compatible with downstream DNA manipulations such as restriction digest, sequencing, and cloning. GelRed® and EtBr have virtually the same spectra, so you can directly replace EtBr with GelRed® without changing your existing imaging system. For detailed protocols for use, please download the GelRed® Product Information Sheet. Also see our GelRed® and GelGreen® Frequently Asked Questions (FAQs).

Non-Mutagenic and Safer for the Environment

A series of safety tests have confirmed that GelRed® is noncytotoxic, nonmutagenic and nonhazardous at concentrations well above the working concentrations used in gel staining. As a result, working strength GelRed® can be safely disposed of down the drain or in regular trash, providing convenience and reducing cost in waste disposal. For detailed test results, you may download the GelRed®/GelGreen® Safety Report.

How Safe is Your Gel Stain?

Many so-called “safe” DNA dyes like SYBR® Safe, Midori Green, GreenSafe, SafeView™, and RedSafe™ not only have low sensitivity, but also readily penetrate living cells to bind DNA, and some are cytotoxic. Unlike these dyes, GelRed® is cell membrane-impermeant, so it cannot enter living cells to interact with their DNA. See our Gel Stains Comparison Flyer or Gel Stains Comparison White Paper for details.

Choose the Right Stain for Your Application

GelRed® 3X in water (catalog no. 41001) is ready-to-use for post-electrophoresis gel staining, and is supplied in a 4L Cubitainer®. Biotium also offers GelRed® 10,000X in water (catalog no. 41003) and GelRed® 10,000X in DMSO (catalog no. 41002). GelRed® in water is a newer, safer formulation and our recommended format. We continue to offer GelRed® in DMSO for established users who do not wish to alter their protocols. We also offer GelRed® Agarose and GelRed® Prestain Plus 6X Loading Dye.

Also see GelGreen® Nucleic Acid Gel Stain, a safer replacement for SYBR® gel stains, which is compatible with visible light excitation.


Watch the video: Polyacrylamide Gel Electrophoresis- PAGE - Amrita University (August 2022).