SDS-PAGE - Science method

Do you think that because the protein is in the soluble fraction the urea concentration can be less than 8M? Since 8M is generally used to remove from inclusion bodies.

Phosphorylated proteins often have different conformations or changes in their mobility, which could be more pronounced or less distinguishable depending on the gel system. Bis-Tris gels might alter protein conformation in a way that changes the separation of phosphorylated versus non-phosphorylated forms.

Sometimes the conditions for Western blotting, such as transfer efficiency or the blocking and antibody conditions, may not be fully optimized for the Bis-Tris gel system. It's possible that the proteins or phosphorylated forms are not transferred as efficiently, or there could be an issue with the antibody recognition in the context of the Bis-Tris gel.

Testing these possibilities one by one should help pinpoint the cause of the problem :

Check the pH and ionic strength of the buffers used in your Bis-Tris gels. You could try adjusting the running buffer or the gel composition to see if it makes a difference.

Bis-Tris gels require different transfer protocols.

Try Different Antibodies or Blocking Agents: If you suspect the phosphorylation-specific antibody may not be optimal for this new gel system, you could try another antibody or change the blocking conditions to see if this improves the results.

Yes removing the GdHCl part can be a good strategy

If you're using MG1655, it cannot make O-Antigen because it has a mutation in wbbL which is the first glycosyltransferase in the pathway for synthesizing the O-antigen repeating unit. Please see this paper - PMID: 37792901

I'm not shocked you can't see the ladder since wildtype MG1655 cannot make full LPS.

Tips: Avoid metal containers, protect silver nitrate from light, and stop development as soon as bands appear.

Dear Vasile Miha Sularea, have you detect MAVS aggregation successfully through western blot using a SDS-PAGE? In recent, I tried to detect MAVS aggregation through western blot post DSS treatement, but still in a trial sub stage. would you mind share your experience in MAVS aggregation detection?

The SDS will not dialyze in any reasonable amount of time from Laemmli sample buffer because its concentration is higher than its critical micellar concentration (CMC). This means that the detergent is in the form of micelles. These micelles are too large to pass through the pores of the dialysis membrane. Diluting the detergent to a concentration below the CMC (about 8 mM) should speed up the dialysis process. As for bromphenol blue, my guess is that the dye is incorporated into the SDS micelles, so it does not dialyze either.

Did you still have precipitate after boiling your samples in SDS sample buffer?

You could try native PAGE, with no SDS in the sample.

Since you first used 50nM MTX and then adapted your cells in 1uM MTX, to my understanding you made stable cell line and used MTX to create selection pressure to get high protein producers.

Loss of protein expression happens over time (although GOI has integrated into the cell genome and high copy no. of the gene would have been achieved due to MTX addition, MTX assures DHFR gene amplification but over time GOI amplified copies can be lost/kicked out of the cell due to metabolic burden). Also, in a mixed culture low producers can just outgrow the high producers ultimately leading to reduction in protein expression.

A way out is to develop single cell clones from this MTX adapted culture (preferably in absence of MTX) and to check the stability of cells/clones in maintaining the protein expression over few generations (atleast 30-40) depending upon the work that you do.

I faced a similar problem while working with Dictyostelium since there too stable clones were made and often expression is lost upon revival of the stocks.

Thanks Bo Pontoppidan for your suggestion. actually i have no other option available in lab right now. is there any recommendation to improve the resolution.

Sufiyanu Abubakar Jiga What antibiotic are you using and what is the half life of it? I have mainly used IPTG induction and PET plasmids. If you know your cassette, you can check it with the GCUA, here, https://gcua.schoedl.de/#google_vignette I think you are curing your plasmid overnight. If you have any fluorescent protein genes in constructs you could subclone that into your construct for a positive control. I hope you get it! good luck!!

There are no well-known endogenous proteins in mouse heart tissue around 70 kDa that have intrinsic fluorescence at Excitation 488 nm. However, there are some possibilities to consider:

1. Autofluorescent Molecules

2. Potential Proteins Near 70 kDa

3. Post-translational Modifications & Cofactors

Would you like me to refine this based on a specific application (e.g., microscopy, FACS, Western blot)?

DEar Amy E Griffiths

give a look to this review

best

Manuele

Dear Manuele Martinelli,

I appreciate your clarification. The way that I took the sample was wrong.

When taking both samples (induced Vs non-induced), I did not centrifuge to get the cells. I just took the sample (so it is probably just the media.

I think the way I took the sample was wrong, that is why there was no band of protein.

I try to get the sample, and do centrifugation to get the cells, and then resuspend it by DW. Finally, I got some bands from my protein.

Thank you.

Hello Lai Yen Fong I am facing the same issue. After protein purification, the flow through and supernatant fractions precipitate after I add the SDS loading dye to them. Could you suggest a solution?

Ammonium peroxydisulfate and ammonium persulfate are two names for the same thing. Use it for PAGE gel polymerization, not ammonium sulfate.

I remember that there is a distinct four-amino-acid insertion within the spike protein between S1 and S2, serving as a cleavage site for the protease furin. Published studies have discussed how this site is even proteolytically processed by various proteases (DOI: 10.1016/j.isci.2020.101212). If you included protease inhibitors in the lysis buffer, I doubt that cleavage occurs during cell lysis and instead suspect it happens during the cell culture phase. You may need to optimize your expression.

One possibility is that the protein was not expressed at all, which is not uncommon. You can't always judge by whether there is a minor band on SDS-PAGE at the expected position, without comparing to another sample in which the protein was not expressed (e.g., transformed with the empty vector).

Another possibility is that the expressed protein was degraded by proteases during storage for 2 weeks at 4^o. If you can't proceed directly from lysis to purification, you should store the sample frozen (preferably at -80 or in liquid nitrogen). You should also include protease inhibitor cocktail during the lysis step.

To find out whether the protein of interest was present in a small amount in the extract, a Western blot can be performed if an antibody is available.

If some samples are flowing out of the well, it indicates an issue with the composition of your loading buffer. Typically, this happens when the buffer is too diluted, likely due to adding too much dI water to the original solution. Adding glycerol to your loading buffer should help resolve the issue by improving sample density and keeping it in the well.

The protein's functional structure may have been disrupted during the process. SDS-PAGE is not a proper method for obtaining a pure and active protein.

SDS-PAGE denatures proteins by disrupting their native structure with SDS detergent, leading to a loss of native structure (bioactive). Afterward, it may not be possible to fully recover the protein's native structure and bioactivity.

Hope this helps! Let me know if you need more assistance.

Strong bases can cause structural and chemical changes in proteins, including:

Dear Ioannis Angelonidis

It was the only cisteine in your sequence or there are other cisteines?

if you are sure that the sequence is correct, the mean that the mutation do not introduce a stop codon due to a frameshit and there are other cisteines, you result may suggest that this Cystein is important for the structural stability of your protein (eg involved in S-S bonds) and with out it the protein is less folded and therefore more susceptible to proteases.

In this case, may not so simple to prevent the degradation.

You can try to reduce the induction temperature (use 17°c or 25°C) or identify the digestion site by performing mass spec and perform mutagensis in the digestion regions

best

Manuele

Hi.. I’m facing same issue right now. When I stored the gel in the fridge at +4C wrapping the gel plate in a wet issue and in a plastic pouch. when I checked after couple of days, the air pockets disappear.

But I dont know yet what’s causing this. Did you found out why?

You don't mention if your protein is present in the filtrate or not. If yes then it is a question of too large cutoff. If it is neither in the filtrate or concentrate then it is probably building a gel on the membrane because of too high local concentration. In that case UF is not the way to go. Try to concentrate it by chromatography - ion exchange can often work because the resins generally have very high capacity.

Ammonium sulphate precipitation is great as an initial purification step, but, as with your example, other proteins will frequently co-purify. You'll need to use a subsequent purification step such as ion exchange or hydrophobic interaction chromatography to remove the other proteins which you can see on your gel. Which method(s) to use will depend on the properties of your protein of interest.

Dear Liang Tang

first of all, if you can buy an acetic acid and methanol free staining solution, i suggest to you to use the microwave owen to accelletare the process. You really do not need to do overnight destaing.

regarding the voltage, it depends a little from the buffer that are you using (e.g Tris-glicine, MES, MOPS) and the strenght of your prower supply. However normally with standard SDS page costant voltage in the range 150-200V is ok. Lenght of run could be beetween 40 and 1h. I suggest you to use a pre-stained marker so you can monitor better the separation and decide when is it better to stop the gel.

best regards

Manuele

not vortex well,after add ACR and MQ,please vortex

Dear Liang Tang

as Junaid Nazir suggest, for a so small protein is preferable the use of an high % of acrilamide as 15% or 17%.

good luck

Manuele

To perform non-reducing SDS-PAGE for a native protein, prepare the sample without adding reducing agents like DTT or β-mercaptoethanol to the sample buffer, as these disrupt disulfide bonds. Use standard Laemmli buffer (with SDS to denature the protein) but omit the reducing agent. Load the protein samples mixed with this buffer onto a polyacrylamide gel (e.g., 10–12% depending on protein size). Run the gel under standard conditions (e.g., 80V for stacking, 120–130V for resolving) using SDS-PAGE running buffer. The absence of reducing agents preserves the protein's disulfide bonds, allowing you to analyze its native oligomeric or disulfide-linked state. After electrophoresis, stain the gel (e.g., Coomassie Blue) and proceed with visualization. Ensure fresh preparation of buffers to avoid contamination or oxidation artifacts.

1. Incomplete Polymerization of the Gel:

2. Residual Gel Components:

3. Staining and Destaining Artifacts:

4. SDS Precipitation:

To quantify ceruloplasmin in serum using SDS-PAGE, prepare serum samples, separate proteins on an SDS-PAGE gel, and stain the gel to visualize ceruloplasmin bands. Use densitometry to measure band intensity and compare it to a standard curve for quantification.

For the isolation and purification of LLO from Listeria monocytogenes first, grow the bacteria in an appropriate medium like Luria-Bertani (LB) broth and induce LLO expression if necessary. After harvesting the cells, lyse them using methods like sonication or chemical lysis, and solubilize the protein if it forms inclusion bodies. Purify LLO using techniques such as affinity chromatography (His-tag or GST-tag), ion exchange chromatography, and size exclusion chromatography. If the protein is expressed as inclusion bodies, refold it by gradually removing denaturing agents. Verify purity using SDS-PAGE and potentially a hemolysis assay, and then concentrate and store the protein at -80°C for further study.

Thank you, Beatrise Luīze Revina, for adding your response

Please run a cell lysate sample from cells without plasmid or just an empty vector (which does not have gene for TTR). Compare 3 samples on an SDS PAGE gel.

For induced and uninduced cell lysate, do you see similar level of expression for a 14 kDa protein? You could also verify whether that is your protein using western blot, if your protein has a His tag.

Dear, I hope you are doing well! Vertical streaks in 2DE electrophoresis are generally associated with excess salts in the samples. They can also result from issues during isoelectric focusing. In your specific case, I believe combining both factors is causing the vertical streaks. I suspect the first separation (isoelectric point) did not occur properly.

I recently encountered your problem. I would suggest you prepare 1X running buffer with this recipe: 14.4 grams of Glycine, 3 grams of tris base, and 1 gram of SDS in distilled water to 1L volume. You will get the result.

To prepare a collagen protein sample in powdered form for SDS-PAGE, dissolve the collagen in a suitable buffer that maintains its solubility and integrity. Start by weighing the required amount of collagen powder and dissolving it in a denaturing sample buffer, such as 1X Laemmli buffer (containing SDS and a reducing agent like β-mercaptoethanol or DTT), which breaks down secondary and tertiary structures. Heat the solution at 95°C for 5–10 minutes to ensure complete denaturation. If the collagen is difficult to dissolve, pre-dissolve it in a mild acid solution, such as 0.1 M acetic acid, before adding the sample buffer. Adjust the pH if necessary to avoid precipitation and centrifuge briefly to remove insoluble debris. Ensure the final sample concentration is appropriate for loading onto the gel for clear electrophoresis results.

Thank you Debajit Das, your words are helpful and I am going to check the parameters related to running buffer pH and ionic strength of the rehydration buffer.

I have always done extractions cold or near room temperature. With a 1% SDS (which is very high), you would have no trouble extracting, at least for mammalian cells in either whole tissues or culture. The big problem with using SDS or saponan (or other ionic detergent) greater the .5% is the complete dissolution of the nuclear membrane which then makes you crude extract very viscous (due to the released DNA) and difficult to work with (i.e. centrifuging the lysate to get a clear supernatant).

I'm going through the same too

Dear Abdulrazak Karabonde Umar, I do not think this gel will be used now. It is most likely that the gel has degraded to some extent, which could compromise the reliability of the results. I recommend running a fresh gel to ensure the accuracy and validity of your findings. This is especially important if you intend to use the results for publication purposes, as the integrity of the data is critical in such cases. However, this might be fine if the gel is only run for demonstration purposes to the BSc or MSc students.

Are these from two separate SDS-PAGE gels (each prepared using the same denatured samples) or from the same one (e.g using Biorad Stain Free gels to visualise the gel before transferring)? Was the protein reduced as well as denatured before loading on the gel? How was this protein that you are analysing expressed and purified?

In my opinion, looking at other literature on this it is most likely that the 180 kDa protein represents total spike protein, while the 120 kDa band is likely the glycosylated form of the S1 subunit protein (containing the RBD) and the 80 kDa protein the form of the S1 protein lacking glycosylation. You can easily check if this is the case by using PNGase F to remove the glycans from your protein and then comparing it on the same gel to a "mock" treated sample of the protein (i.e adding water instead of PNGase F). You should see the 120 kDa band disappearing in the PNGase F sample and appearing around 80 kDa.

Thank you Shefali Desai

Hi Can,

Howard Salis, 1st author of the manuscript I had linked in my prior response to the question 8 years ago, was at UCSF at the time his manuscript was published. The manuscript link to the prediction program is dead but he put it on a new server when he moved to Penn State.

I was using a prokaryotic GateWay Destination expression plasmid. The donor plasmid in which I had cloned the gene was primarily designed for recombining into eukaryotic expression vectors. Therefore, I added a prokaryotic Ribosome Binding Site (RBS or Shine-Dalgarno sequence) to the forward oligo to PCR and clone the gene I was expressing. Cloning was very straightforward (I actually had a high school student do it as a summer project). BUT it took almost 2 years to get expression (when I found Salis' program's new site). It correctly predicted that when transcribed, the RBS in the 5' leader sequence would form a very strong hairpin with an upstream NotI restriction site and flanking sequence (so lots of GCs in a row) in the original Donor plasmid. The hairpin sequestered the RBS, preventing the ribosome from assembling on it so translation was squelched. Eukaryotic ribosomes assemble at the mRNA cap and just plow their way to the Kozak site, secondary structure be damned. Prokaryotic ribosomes instead assemble on the mRNA on the RBS located directly before the initiating methionine, but only if the site is accessible.

Salis' program analyzed the inputted sequence and predicts expression levels but doesn't advise how to improve it. I used a separate program that maps RNA structure (I don't remember its name but there are certainly better programs now), and used intuition to manually tweak the sequence to eliminate secondary structure, and then tested it with Salis' program. I kept reiterating this process many times until I thought I had absolutely maxed predicted expression. I then repeated this for the initial 6xHis tag at the N-terminus because it too was predicted to form a very strong hairpin - that involved just changing some of the CAC codons to CAU, so pretty easy. I'm not sure if the His hairpin affected translation but as all the changes were incorporated into a single oligo, there was really no additional work.

The first time I tried the optimized vector, I got rid of all the desperation techniques I had tried - tight repression of toxic expressed proteins, medium formulations, temperature, inoculation protocols, fancy bugs. Also exotic purification techniques, protease inhibitors, and lysate formulations. Absolutely everything. Just 1 L of barebones LB on a shaker overnight and a simple lysate poured over a nickel column. Ended up with hundreds of milligrams of almost pure protein. I had *no* evidence of expression prior. So much protein that it overwhelmed the endogenous E. coli biotinylation system (I had included a C-tag for biotinylating the protein) so I had to add a 3rd plasmid to the system to express additional biotinylating enzyme (the 2nd plasmid expressed the 2 casein kinase subunits to phosphorylate the target expressed protein, so yeah, kinda complicated).

Quite long, but I hope this helps. I would certainly contact Howard Silas for advice - he's the expert. And I'm sure things have advanced significantly in the 8 years since this question was originally posted. https://www.researchgate.net/profile/Howard-Salis

Aakriti Singh you can use either L-Arg or L-Arg*HCl. Just look after pH of buffer

Satyendra Mondal yes, I was thinking that overheating could be an issue because I an running the PAGE at room temperature. However, my lab colleagues who run their gels at even higher voltages (120 V for instance) don't experience this problem so temperature does not seem to be an issue.

Hameer Khan Khaskheli Hi, thank you for your suggestions. i have tried using 15% gel also tried with 0.5mg/ml concentration as well as 5mg/ml concentration. But didn't work. Even though if the sample was overly concentrated, at least I should see smears if not defined bands. But I could observe nothing.

While the previous posts could explain your observation, in my experience many integral membrane proteins show anoumalous migration behaviour in SDS-PAGE, leading to smaller apparent MW and frequentyl also additional oligomeric bands of aggregated molecules with a regular size pattern of their monomeric size. One extreme example is bacteriorhodpsin (also 7 TM domains like GPCRs) with ~16-18 kDa on the gel instead of the correct ~26 kDa. The reason seems to be that the transmembrane helices are not bound to the average ~2 SDS per amino acid like "normal" proteins because of the highly hydrophobic properties which makes it difficult to fully unfold these regions. So you have an unfolded SDS-protein complex that appears more compact, i.e. smaller and migrates like a smaller protein.

Band stretching in SDS-PAGE can be caused by several factors related to gel preparation, sample loading, and running conditions. Here are some common causes and solutions:

Dear Elif Vera

MW of biotin is to small to detect the difference in MW

if you would like to detect it, you can try to add streptavidin and check the MW shift of biotinilated fraction. However it will allow you to disctiguish if a protein chain is biotinilated or not but not able to differentiate modo and multiple biotinilation.

good luck

Manuele

Gaurav sir and Sirvan sir, thank you.

yea sir i have duly given similar answers in my draft.

actually for cyclic protein with MW 95 coming above then lin non cyclic one.

when I tried the same method appr the protein size around 40 kDa comin lower then lin counterpart.

the cyclic protein confirmed with western blot as well.

so for manuscript i need very precise refrencce To justy the same statement which has been mentioned by both of you.

Thank you sir.

Best.

You can see from the marker protein ladder the size range of proteins that are separated in the gel. If the marker dye is still on the gel after electrophoresis, smaller proteins will most likely be located at the dye front. If the dye has run off the gel, the smaller proteins will have run off and will be lost.

Small proteins may be lost during blotting if the blotting time is too long.

Gaurav Chhetri Thanks for your suggestions

it is not usefull not to let the tracking dye exit the gel; there is no (resolved) material in front of the blue dye. you can run a 20% acrylamide gel . For small molecular weight protein you should run a tris-tricine gel to have a nice resolution (maybe the precast gel you use was a tricine gel) Also small protein are not stained as well as larger one so maybe you do not have enough of the protein you are looking for to see it by coomassy staining. Did you try western blot?

The two unexpected bands in your SDS-PAGE analysis of Bacillus subtilis LB supernatant could be linked to specific protein secretion systems active in the bacterium, such as the ESX (or Type VII) secretion system. This system, which is not exclusive to pathogenic bacteria, can secrete proteins like YukE from the yuk/yue operon. YukE is a member of the WXG100 protein family, known for being secreted even under nutrient-rich conditions like LB media. The yuk/yue operon in Bacillus subtilis encodes several proteins involved in this secretion pathway, and their expression and secretion could lead to distinct bands in the supernatant, which you are observing in your gel.

Other secreted proteins from Bacillus subtilis that might contribute to these bands include enzymes like subtilisin (encoded by the aprE gene) and alpha-amylase (encoded by the amyE gene). These proteins are commonly secreted into the culture medium and can often be seen in SDS-PAGE analyses as separate bands.

For precise identification of these bands, performing mass spectrometry would be the most effective approach. This method will allow you to identify the specific proteins present in the supernatant, confirming whether they are products of known secretion systems or other secreted enzymes, and provide a clear explanation for the unexpected results.

Hello Bhardwaj,

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

Hope you the best,

Nicolas

Thank you, everyone I am going to repeat it by next week, with all your kind suggestions, I hope I will get the strong band this time.

Hello Deepdarshan Urs

If I use the HRP-conjugated secondary antibody, can I develop the blots by using a TMB/ABTS substrate on the membrane? (not the chemiluminescent substrate)

Yes, you can. See Pg 2 (Fig 2) of the link attached below.

Best.

What is your gel percentage in native-PAGE and used gel chemistry? I suggest trying BN-PAGE to overcome the migration differentiation...Cover the proteins with negative charge using coomassie stain and hope to get a similar pattern with SDS-PAGE. Compare the proteins by only their shape and size, not by charge-induced mobility variation...

Hi. I have the same problem. If you find a solution, please write.

Hi! I am using 10% SDS-PAGE. But, It looks like there is a problem with my cell lysis protocol.

Hi Akash,

It is more convenient to use Biorad's ImageLab program (as noted above, it is free). There are tutorial videos on Biorad's YouTube channel on how to do this. The latest version of the program allows you to process .tif files (gray scale 8 bit) from other sources (but there may be problems with image settings, not all are simply imported). Or as an option - the free program ImageJ, there are a lot of video tutorials on it too

Have you heard of Valita Titer?

Plate-based assay that uses fluorescence polarization on a plate reader. Add, mix, & read for results in less than 15 mins.

The concentration of albumin in culture media containing serum is quite high. There are other proteins, too, but albumin is by far the most abundant. It can be difficult to get rid of all of it when purifying IgG from culture medium. There are resins available to help remove it, or you may be able to adapt the cells to grow in serum-free medium, which lacks albumin.

You mentioned a lysate, suggesting that the IgG was produced within the cells rather than excreted into the medium. If this is the method of production, thoroughly washing the cells to remove external albumin before preparing the lysate would probably help. Also, make sure there is no albumin in the lysis reagent, of course.

How do I wash the cells? And how to improve the lysis? thank you!

Hi Karthik! I also perform click reactions and detect my protein via SDS-PAGE. I first label the proteins in cells with AHA, lyse the cells in 1% SDS, boil and sonicate the sample, then use Click-iT™ Protein Reaction Buffer Kit from Thermo to click the protein to biotin-alkyne. Afterwards, I pull down the biotinylated proteins with Streptactin beads and release the protein by boiling it in 2% SDS for 10 min. As you can see, there are several boiling steps in the protocol. Unless your reporter tag is heat sensitive, I don't think boiling the samples before and after the click reaction would affect the quality or outcome of the reaction.

With 6xHis, your typical protein will elute at 150mM Imidazole rather quantitively, unless it has a high amount of histidines itself which strengthen the binding. With 250mM Imidazole, I got to elute pretty much any 6xHis Protein to completion this far. Some go to 500mM but seems unreasonable to me unless you got some stuff like 12xHis (9xHis for the most part retains at 75mM Imidazole, though this always depends on the length of these washing steps as your POI lets loose bit by bit).

When eluting with imidazole, don't vary the pH too much, there is no reason to. Depending on your pH, you might be too near to your protein isoelectric point, leading to precipitation. As IMAC resin doesn't work below pH7 (pH5 is typically used for pH-elution), make sure you are 7.5-10. If the pH is too high, your resin will reversibly turn to mud-green Ni(OH)2 while losing binding capacity.

In case you're not sure if the protein still clings to the resin, simply boil up a portion of the used resin in SDS-PAGE loading buffer.