SDS-PAGE - Science method

I have been purifying proteins which have an N-terminal His-tag using Ni-NTA affinity chromatography. I carried out size exclusion chromatography using Superdex HiLoad, Superdex, and Superose columns. I also carried out dialysis of the Ni-NTA purified proteins.

Both the above experiments were performed for buffer exchange and to remove non-specific proteins.

When I ran SDS-PAGE gels for the Ni-NTA purified and dialysed proteins, I could see a single band corresponding to the protein of my interest. Multiple bands beneath the protein of interest could be seen on SDS-PAGE gel for the SEC fractions.

I used appropriate controls to rule out the possibility of degradation due to prolonged exposure at room temperature and the effect of varying salt concentrations.

Please help... to all the protein purification experts out there!!

Hello,

I am wondering if anyone here who performs SDS-PAGE has seen this before on their gels post-staining? We make our 12% tricine gels in house, fix them in 25% isopropanol 10% acetic acid, then stained overnight in Coomassie G250 35mM HCl. The gels are then destained in distilled water. We are noticing what seems like "halos" or zones of white around our proteins in the gels. I have images and notes attached regarding the issue. When the peptide is in its neat form, it is in a 1M imidazole, 500mM NaCl, 20mM Tris buffer at pH 8. The peptide has a final concentration of 200mM imidazole when it is in its 1/5 diluted form. We have seen this effect many times before, but are not sure what is it causing it. Is it perhaps due to the presence of imidazole; can the imidazole, or maybe just an overall high salt concentration, cause this effect? We use fresh running buffer, fresh fixative and fresh gel reagents (e.g. new aliquots of APS) for each run. Coomassie is reused and made fresh every month and a half; the Coomassie used here is less than a month old.

Any input or words of wisdom would be greatly appreciated! Many thanks in advance.

Leisha

I am looking for an alternative to kerosene (CMR) and mineral oil (too viscous) used as cooling fluid for IEF and SDS-PAGE electrophoresis performed on multiphor apparatus (flatbed).

Does anyone have an idea, solution and experience ?

Thanks !!

I recently performed protein extraction with RIPA buffer of some cells I collected. The intended purpose was to use it to load in an SDS PAGE gel for subsequent Western Blot. I made the mistake of lysing the cells in too much RIPA buffer and although there is protein, the sample is too diluted. I want to load 20 micrograms of total protein per lane, and I have roughly 400 microlitres of a protein solution at 0.13 microgram/microlitre. Since there is no gel that supports having 150 microlitres of sample loaded into a lane, I am looking for a way to concentrate what I have rather than just discarding it.

The purified protein is exhibiting a trimeric state based on my NATIVE PAGE gel and SDS PAGE shows monomeric configuration. Can you recommend detergents to destabilize the H-bonds mediated by sulfate ions (based on literature)? I have already tried BME and DTT and they don't work.

My gel running is going well but as the maker running down towards the end of the run, it got spread out towards the outside of its own lane, not so sure how could it can happen, even I tried to load the empty lane with 1x loading buffer

SDS-PAGE details are available for collagen type I and I could get the bands of subunits of the same. I need the protocol for native-PAGE too.

how many bands does BSA give in sds page?

Hello,

I am using the costar gel loading tips round 4853 and an eppendorf pipette (100 µl) to load my gels. The volume of my samples is 25 µl. Before loading, I usually set my eppendorf pipette to 25 µl and use the gel loading tips as mentioned above. But everytime I get air bubbles in my tips that result non-optimal western blots.

Do you have any tips/ideas how to avoid air bubbles in the tips?

What I've already tried:

I resuspend the tip to remove the air bubbles but in most cases the resuspending process results in more bubbles.

I pull the pipette slowly to avoid capture any air, but still isn't working.

I readjusted the volume of the pipette to 23,5 µl to set a lower volume to avoid sucking excess air, but then I lose sample since a small amount is still in the tube. So that doesn't work either.

I am thankful for any help and tips!

This is an SDS-PAGE silver stained gel. The sample is from an FPLC fractions mixed with sample buffer (has SDS and DTT). I'm not sure why the lane has darker edges all the way through (vertically). Would anyone know what is causing this? Could this be from overloading the wells?

I have cloned a gene in pET28a vector, and induced it for 5 hrs with 1mM IPTG. The cells were centrifuged from 1 ml culture (Uninduced and induced both), and the pellet was suspended in 200 microlitres 2X SDS loading dye and kept at -20 overnight for further use.

The next day, run on 12% SDS PAGE, on staining induced band was visible.

To have a better image, the same protein was run again on 12% SDA PAGE, the induced band disappeared.

pls suggest, how to stabilize induced protein.

Hi everyone

Looking for some suggestions on bench top protein analyzers for titer determination. Currently using SDS-Page which is time consuming and allows for a large amount of operator error. I have previously used CEDEX Bio/Bio HT but and looking for something with a similar IgG titer determination functioning. Looking for something that has a small footprint and is relatively 'quick' to run titer analysis on centrifuged cell culture samples (Hybridoma, CHO). HPLC and other LC analyzers also not an option.

If anyone has any suggestions I would greatly appreciate it.

Thanks

I would like to measure the protein concentration using the Bradford assay. To do this I have to resuspend the isolated protein pellet in the sample buffer. However, at this stage, I do not have ampholyte reagent to make the rehydration buffer (I do have Urea, DTT, CHAPS and Bromophenol Blue). After this, I want to rehydrate the IEF gel strips as the first dimension gel and then run 2nd dimension gel. I am wondering if missing ampholyte in the rehydration buffer will considerably affect the result. How important is the role of ampholyte?

Any suggestions and comments would be greatly appreciated.

Dear all,

I was trying to see what is going on with my commercial protein sample so I ran SDS PAGE to check. However, I have no idea what is going on with lane 5&6 (they're almost the same sample). I'm looking forward to seeing all the insights from you!

I ran out of Bromophenol Blue that is given in the standard recepie of Laemmli Buffer by CSH, now what concentration should I take Commassie Brilliant Blue G-250 as an alternative, should I take it the same concentration as Bromophenol Blue?

Hi, currently I'm working with E. coli Bl21 (DE3) pET-26b(+)-N20-aiiA-6xHis. I'm trying to improve the secretion rate of the aiiA to the extracellular fraction.

"pET-26b(+)-N20" already provide the signal for the extracellular secretion. so i thinks its normal to have thick band in the SDS. but how about the native protein? why it also looks thick? especially the one in the sample are more thicker, than the one in the control.

Both sample and control are undergo the same process. The difference only in the plasmid (control doesn't have plasmid).

thank you in advance!

This is the used protocol:

•Spin down an aliquot of Amyloid 42 HFIP** (~ 5 mg/ml), then 1 µL of Amyloid 42 was diluted in 20 µL PBS.

•5 µL of 4x Laemmli protein sample buffer for SDS-PAGE was added. The sample was boiled to 95 degrees for 5 minutes. Then, short spin down.

•Then, the sample was run along the ladder (BLUeye PrestainedProtein Ladder) in 4–20% precast polyacrylamide gel, 8.6 × 6.7 cm (W × L), for use with Mini-PROTEAN Electrophoresis Cells.

•The gel was stained with Quick Coomassie Stain for 1 hr.

•Lastly, the gel was scanned.

I use gel 16.5%, 120V, 10 µL sample. Thank you

I have been observing double bands for protein molecular marker(especially 15 kDa band) every time I am doing SDS-PAGE. Sometimes even the samples appear to have a double band. These are 12.5% handmade gels and sample running voltage is 160 V. Can anyone suggest how to resolve this issue.

Thank you for your answers.

I am trying to dimerize a synthetic peptide (22 amino acids) with a N-terminal cysteine, that was added for this purpose. I use the BM-(PEG)₃ crosslinker from Thermo Fisher, which is based on maleimide-thiol chemistry. I reduce the sulfhydryl-bonds using TCEP, add the linker and stop the reaction with DTT. All according to the instructions provided by Thermo Fisher. I check the results with an SDS PAGE, but so far the protein bands stay on the same height before and after the reaction. I tried to get a positive control with insulin, lysozyme and murine SAA, but only the SAA shows a very faint band that could be a dimer.

Has anyone used this linker successfully or has any tips on how to get the reaction working?

stacking gel 5%,running gel 8%

first band includes catalase(400KDa） and glucose oxidase（160KDa) , small molecular crosslinker(MW<1000) is used ,but no aggregates showed in this band(should have been above than most other bands),why the band below showed like dumbbell-shaped?

for other bands, is the concentration of protein too high because the color of bands is deep and bands are long? why it showed a funnelform?

could any beautiful people help me out of this, I would be so appreciated of your kind answers. your advice is of great help for me as a beginner. Thanks for help!

Hi it's my second time running sds page. I use 90 volt, 10 ul sample and 5 ul protein ladder

Thank you

MAP Tau, htau40, 2N4R, has the actual weight of 45 kDa but runs as 67 kDa on SDS-PAGE. What can explain this much weight difference?

Is it specifically about Tau's unique structure effecting charge, or possible post translational modifications?

Dear researchers,

When I run SDS-PAGE, sometimes I found that the high molecular weight bands (including bands of the molecular weight marker) disappear from the gel, but low molecular weight bands are visible (see figure below). I've made new running buffer and new component solutions used to make the gel, but this still happen occasionaly.

Has anyone run into the same problem? I would be very appreciated if anyone knows the cause and possible solution. Thanks in advance.

Hello everyone,

I'm having some conceptual misunderstandings regarding non-reducing SDS-PAGE. In this situation, we omit reducing agents such DTT or BME from the loading buffer to preserve disulphide bonds in the proteins' structure. However, in every protocol i've seen, SDS is present and sample heating is still performed. Wouldn't this result in disrupting the disulphide bridges, since we are still denaturing the samples? I know that disulphide bonds are more heat resistant than hydrogen bonds (since they are covalent bonds) and that heating in the presence of reducing agents is only done to facilitate the disruption of those bonds. But I couldn't understand if high temperature alone is sufficient or not to break these linkages.

Thank you kindly for your attention.

Best regards,

Miguel

Hi! I'ts one of the first times i run SDS page for proteins and i got this result. what could be wrong? i attach a picture.

thank you in advance!

I have my construcsee a band at the correct molecular weight but nott in pGT7 plasmid, transformed into BL21AI when I do a test expression and check on SDS-PAGE I don't see a thick expression band. I do see a band at the correct molecular weight but I don't see the protein after purification on gel. am also getting micromolar concentrations in the end. am starting to think the problem could be in expression. my conditions are 16degrees overnight after induction with 0.04% and have also tried 0.1% and 0.01% with similar results. KIndly help.

I did this SDS PAGE GEL and my results are not precise or improving.

can someone guide me through the trouble?

Hi

I am currently studying the expression of the TEV protease recombinant protein, and unfortunately, I have encountered a somewhat illogical problem during my work. I would greatly appreciate it if you could help me based on your experience and knowledge.

One of my colleague’s previously used the soluble fusion tag “GST” for expressing the TEV protein. In their design, they were able to express TEV/GST protein in SHuffle strain by using the pBAD A series vector under araBAD promoter and ori: p15A.

However, the protein was totally expressed into inclusion body. In order to optimize that project, we decided to use the previous backbone, however with an alternative tag based on an article by Dr. Yutaka Kuroda entitled " A SEP tag enhances the expression, solubility and yield of recombinant TEV protease without altering its activity " Consequently, GST fusion tag was replaced with SEP tag, incorporated in C-terminal. This article claimed that this tag significantly enhances the solubility of the TEV protein.

It should be noted that Dr. Kuroda used the pET15b vector under T7 promoter in their design.

After changing the solubility fusion tag, the integrity of the target fragment was confirmed by Sanger sequencing. In spite of the confirmation of critical elements within the expression vector, no protein was expressed in Shuffle (induced by Arabinose at 30 and 16 C for 4 and 18 h, respectively), even into inclusion body forms. I have included the gel images of my colleague's vector and my own below for your reference. (The expected size of TEV/SEP is ~ 29 kDa, while GST/TEV is approximately ~58 kDa.)

Furthermore, since we don't need to purify the TEV protein in my project, this protein is not fused to His- tag.

the important question for us at this moment is the lack of protein expression by the vector.

Can you please help me why we have not any band in our SDS-page gel of our recombinant protein?

Hello everybody,

I purified my protein and when I run it on SDS-PAGE, a small molecular weight sized band (10 kDa) appears. Do you have any idea what that could be?

an image of the SDS-PAGE is attached.

RIPA buffer was used for protein extraction from rat liver samples. After calculating the protein content of the homogenate through Bradford assay (BSA), 5-30 ug of protein was loaded to 12% SDS-PAGE. Then the protein bands were transferred through the wet-transfer method followed by the blocking buffer [1 % BSA (Himedia) in 1X PBS for 1 hrs], followed by washing thrice for a period of 10 min, later adding primary antibody (overnight) and again same washing step followed by secondary antibody (1.5 hrs). The antibody we ran for was β-actin with anti-mice specificity. Since the protein ladder transferred quite well on the blot, we arent speculating on the transfer issue and also ponceau proves the same (visible after stain). In addition, we did a dot blot, checking the affinity of the antibody. Bands are also visible with CBB stain on the gel. Still we arent getting it on the blot. Please suggest a solution to this. Thank you!

After immunoprecipitation with specific antibody and Protein G, samples were eluted with elution buffer, SDS sample buffer, and reducing agent at 70 °C for 5 minutes. Also, samples were incubated at 95 °C for 10 minutes before loading on SDS-PAGE.

The bubbles did not exist in the gel, and the replicate experiment shows the same result.

This is confusing because the left lane is a negative control, and the right lane is a positive control that should show immunoprecipitated protein.

Is it an protein aggregation?

I am trying to check a purified recombinant protein on SDS PAGE coomasie Blue and silver staining. Before SDS PAGE, I have checked the total protein concentration using lowry and it shows good amount of concentration. But when I run it on SDS PAGE the coomasie blue stain does not show any bands, while the silver staining one shows only 2 bands. I also check the cell lysate before purification but it shows the same result as the purified lane, only 2 bands and not on the desired MW band. How can I resolve the problem? Thank you in advance for your responses.

When I'm running SDS-PAGE 12%, my sample moves to the other well (it's formed as a small curve with the following well) , even if I put it slowly,carefully, 15ul per well and I'm using a 1mm glass. I think it may be the sample buffer i use, it is dense. I look forward to your recommendations.

Sample buffer recipe (5x):

For 1ml:

- Tris (1M, pH 6.8) 0.25ml

- SDS 0.1 g

-Bromophenol blue 0.005 g

-Glycerol 99.5% 0.502 ml

- H2OMiliQ 0.25 ml

I use sample Buffer 1X

I have performed SDS-PAGE using precast Bis-Tris gels and MOPS running buffer. The bands of my reduced samples are always very weak compared to the non-reduced samples. I loaded 2ug for each sample. Does anyone have any idea why?

I am currently trying to express Turritopsis dhornii TET enzyme with a 6x HIS tag in a BL21 E.coli host. Every time I purify my protein it shows double bands on SDS-PAGE, when testing its activity on ELISA there seems to be no activity. My lysis buffer contains 50mM HEPES ph 7.5, 30 mM imidazole, 500mM NaCl, and 1mM DTT. I purify with 800uL of Nickel Sepharose beads. Are there any adjustments I could make to stop this double band from showing?

Hello everyone!

In a SDS-PAGE, what is the more effective way to separate high molecular weight protein st (>300 kda) and get a good resolution at the end? I am using Tris-acetate 4-8% gel and MOPS Tis acetate running buffer to run the gel. Could anyone suggest what voltage should I use? Is the voltage should be separate for stacking and separating gel and for how long? I am using semi-Dry Bio-rad transfer system, Is 10 min transfer would be enough with high mol. wt. settings?

Thanks :)

hi everyone... i am working on bovine collagen protein but i am not getting proper bands in SDS PAGE. please give me some suggestion . thanks.

SDS-PAGE is used to separate the particles in the mixture, under the influence of the electric field applied to it.

When I load a protein sample, the sample drifts up the sides of the sample well.My Running Buffer is Tris-Tricine-Hepes system.And the sample strip will become U-shaped after starting to run.

Buffer component：

100mM Hepes

100mM Tris

100mM Tricine

0.1%SDS

I expressed and purified a recombinant human enamelin protein in bacteria and I wrote a protocol/recipe on how I did it. I also have an SDS-PAGE showing the final purified product. I was wondering to which journal could I submit the manuscript ?

I am currently working on a thermostable polymerase which I am overexpressing in the BL21(DE3) strain. The polymerase has a histag at the N-terminus. However, during purification, I am not obtaining a single band but rather several bands of different molecular weights ranging from 5-85 kDa. The expected molecular weight of my polymerase is 92 kDa. The additional bands observed on the SDS-PAGE gel after purification on TALON or AKTA systems, some appear to have the histag (confirmed by westernblott). I have attempted to optimize the purification conditions by adjusting buffers, using protease cocktails, DMSO, Betaine, low induction temperatures with longer time, shorter time of induction, and optimised times and amplitude of sonication, but none of these measures have yielded the desired results. Do you have any suggestion what should I try?

I have prepared 12% of SDS gel and a sample concentration that needs to load 24 ul in each lane to fulfill the requirement of 4uM protein concentration and the maximum volume of the lane is 20 ul. I have also prepared 2X sample loading dye. How much loading dye should I add to the sample so that it will work?

Molecular weight of my protein of interest is 7KDa. I am not able to get this on 18% SDS-PAGE electrophoresis. I want to do western blotting of my samples having this. Kindly suggest the method to separate the low molecular wt protein and western blotting of it.

Thank you

A huge bubble forms in the gel during running as you can see in the photo. This was occured after 1 hour it started to run. The bubbles looks like a second line in the gel. Why does that occur and how can i fix it?

In SDS - PAGE is common to have a stacking and a resolving gel, each with different pH. Is there such thing in a gradient polyacrylamide gel? Since it is only one gel... Is there a stacking portion of the gel and a resolving portion of the gel? If so... How do they work?

For example... How does this gel work?

https://www.thermofisher.com/us/en/home/life-science/protein-biology/protein-gel-electrophoresis/protein-gels/nupage-bis-tris-gels.html?gclid=CjwKCAjwsvujBhAXEiwA_UXnAAik72PNIWnDwcIpa4D2POpuMDEPL1UIe1c_JqwcE14fpMowT-vdtRoCcE0QAvD_BwE&ef_id=CjwKCAjwsvujBhAXEiwA_UXnAAik72PNIWnDwcIpa4D2POpuMDEPL1UIe1c_JqwcE14fpMowT-vdtRoCcE0QAvD_BwE:G:s&s_kwcid=AL!3652!3!481383713744!e!!g!!nupage%204%2012%20bis%20tris%20gel!11496899015!113567084778&cid=bid_pca_gel_r01_co_cp1359_pjt0000_bid00000_0se_gaw_bt_pur_con

Thanks

We were doing western with cell lysate and used 5% BSA in TBST for blocking. Also, primary and secondary dilution of antibodies has been prepared in BSA in TBST. With this, we are getting a prominent non-specific band at 42 kDa while developing with any primary antibody.

The attached picture is a blot of PTEN (50-55 kDa), but the band is at 42 kDa. How can I get rid of this non-specific band?

Any kind of help will be appreciated.

I am working on a recombinant protein using SDS-PAGE. I used Coomassie brilliant blue for visualization of the respected protein bands. The voltage was set at 150V and the gel was run for 1 hour. Acetic acid was used for decolorization of the gel.

All SDS-PAGE buffers were appropriately prepared and used fresh. However, there is a weird curved line at the center of the gel. How can I get rid of it?

I’m working on recombinant protein expression.

My bacteria lysis protocol:

The grown cells (Bl21) were harvested by centrifugation at 5000g for 20 min at 4 ◦C, resuspended in 5mL of PBS buffer, pH 7/4. Then cells were incubated with lysozyme (0.3 mg/mL). After that, bacteria were disrupted by 16 cycles of sonication of 30 s each(30s pulse,30s stop total time: 8min). Then, the lysed cells were centrifuged at 12,000 rpm for 15 min. at 4 ◦C.

The supernatant and Precipitation dissolved in ureatasted by SDS-PAGE 7/5% But I have seen NO BAND for the supernatant.

Protein molecular weight: 116kDa

If you have any experience can you help me?

How do I analyse the gel result I have obtained from an SDS-PAGE electrophoresis?

I followed the exact protocol for kidney, liver samples and it worked. But it is not working for brain samples. I extracted the histone proteins using pre-lysis, lysis, and balanced buffer from Bio-Rad. Then checked the protein concentration followed by SDS-PAGE (80V for 1.5 hours). Then I did the transfer (0.18A for 1.5 hours) on PVDF. I used H3K9BHB primary antibody (1:1000)dilution and kept in in cooler for 12-16 hours followed by secondary antibody (Anti-goat Rabbit 1:10000 dilution) for 2 hours. I developed the blots using LICOR's fluorescence-based detection.

When I put the sds gel in all the wells, even the empty wells, it shows a continuous band from the beginning of the gel to the end.

What is the problem?

Hello all, first post on ResearchGate!

I have been performing immunoprecipitations with FLAG beads (abcam, ab270704), eluting using SDS loading buffer (with DTT) and boiling. The eluted products are then used for SDS-PAGE and downstream Western Blots. We're concerned our proteins of interest are not being fully eluted from the beads using our current method. Abcam suggests a competitive elution using the DYKDDDDK peptide, however I'm having trouble finding it for purchase from them. Has anyone used that peptide product from another company and had success?

Thanks!

Hey, i've been doing a western blot like from 3 weeks ago, and i can't get good results, i'm getting this type of runs, and i don't know what can it be, maybe degraded proteins?.

I'll be very grateful if someone can help. Thanks

Hi all,

Just wondering if anyone has experienced similiar issues when running SDS-PAGE gels? The ladder stops running straight toward to bottom of the gel? Ponceau staining of the nitrocellulose membrane showed some lanes had expanded while others had shrunk (don't have a picture of this unfortunately). I could see my target protein however the bands wern't uniform. Anyone have any advice?

Some helpful info -

I pour my own gels using standard recipe (16%)

Buffers are always made fresh (although not PHd)

Ladder is from ThermoFisher, catalogue #26616

Just let me know if any other information would be helpful!

Thankyou!

After successful purification of my protein of interest by affinity chromatography (first picture, right side), yielding 50ml of 0.4 mg/ml (estimated by nanodrop), I went on to perform concentration on an Amicon Ultra 15 device. The MWCO is 3kDa, while my protein is 13KDa. We take extra care to centrifuge at low speed (2500 rpm instead of the 4000g that the supplier recommands), and to "wash" the filter every 10 minutes by up-and-downs to reduce adsorption and protein loss in the filter. Every hour, the concentration in the retentate is measured against the filtrate as a blank, by Nanodrop.

The problem is, it looks like our protein concentration is not increasing at all, even though nothing is found in the filtrate as well. An SDS-PAGE was performed to confirm that the protein is not lost in the filtrate. However, in the retentate (after a 30x concentration factor), the protein band's intensity is almost the same as before the concentration (2nd picture).

Is anyone familiar with this problem ? How can we prevent it ?

Hello,

My Western blot turned out smeary the other day following the semi-dry transfer using a Trans-Blot® SD Semi-Dry Electrophoretic Transfer Cell. The transfer was performed at 15V for 50 mins.

I don't know how to describe it, but the bands in the marker are clearly bleeding, when they are supposed to be (kind of) straight lines. The gel was actually fine after I ran the samples, the marker bands were all straight, hence I am trying to figure out why this might've happened (this is the 2nd time it has happened).

Also, when filming the membrane, it seems like the protein (105 kDa) did not even transfer off the gel, as nothing came up besides the loading control which is a lower molecular weight (37 kDa).

I attached 2 images - one image is how my Westerns always look, and the other is the problematic Western.

Has anyone else ever encountered this problem?

Hi! I have trouble with the SDS-PAGE. I do not know why my protein band and size marker are not visible in the high molecular weight above 50 kDa.

At this moment, I have tried to change the SDS-Running buffer, but as you can see in Figure I attached the result is the same. Furthermore, i am following the online protocol of Sigma (https://www.sigmaaldrich.com/VN/en/technical-documents/protocol/protein-biology/gel-electrophoresis/sds-page).

Could you show me the way to improve it?

Hi all, with MBP tag at N-terminus and 6xHis at C-terminus, I have been trying 5 protein expressions in NEB Exp cells, optimized at different temperatures and IPTG conc. Upon Ni-NTA purification, I'm seeing several truncated bands with strong MBP signal and very weak band at right size bands with several intermediary sizes on SDS-PAGE. I avoided to use amylose resin because mpb is at N, and only using Ni-NTA (to avoid pulling down non-full length molecules.

I used pMAL-c5x several times with several proteins (from 10 kD to 350 kD proteins) in the past with very good success, but never saw this happen. If anyone faced this problem and solved it, your insights would be highly appreciated. Thanks!

As a researcher, I am writing to request your help in obtaining a protein ladder for my research experiments (SDS-PAGE). Unfortunately, my current budget does not allow for the purchase of a protein ladder, and I am hoping that someone may be able to assist me in finding a low-cost or free option.

My westerns haven't worked for the past three to four months, which is an issue. I attempted using two different antibodies from two different companies, but I got no blots. I load 50 to 100 ug of protein onto gel using SDS-PAGE concentrations ranging from 12 to 17%. I started with the Poncaeu stain until I was confident before moving on the westerns. What might be the cause of that then?

My protein of interest is fused with MBP tag, and it does not contain tryptophan but MBP does. After factor Xa digestion, my protein was separated from MBP, and I eluted my protein while MBP is still in the MBP column.I ran SDS-PAGE with TGX stain-free gel.

Theoretically, my protein should not be visualized in Bio-Rad ChemiDoc imaging system machine because it doesn't have tryptophan. Stain-Free gels (Bio-Rad) contain trihalo compound and they interact with tryptophan residues of your protein and show fluorescent signal by UV detection.

There is no MBP contamination as it stays in the MBP column and it will only be eluted by maltose. What could be the reason?

Thank you in advance!

We are working on the purification of recombinant hemagglutinin by lentil lectin resin. After purification, a double bond HA is seen on the SDS-PAGE gel. Does anyone know what the second bond is? and how we can get rid of it?

Iam unable to get any bands.. So it will be helpful if i know about alternatives..

Tried with 7.5%, 10%, 12% and 15% gel percentages.. Protein concentration was checked through biuret and it is about 1.7mg/ml.. Used coomassie blue for staining (1hr) followed by destaining for overnight.. Any suggestion will be very helpful..

I have been running gel zymography (7.5% polyacrylamide co-polymerized with 1% gelatin) loaded with 250 µg of protein / well from colonic lysate with the hopes of visualizing MMP-2 and -9. I had a few good gels, but recently I have been getting these white spots on my gels (see attached) post-Coomassie staining and de-staining, following a protocol by Frankowski et al., 2012 (doi: 10.1007/978-1-61779-452-0_15).

Can someone explain where these big spots are coming from? It makes it impossible to quantify my images. I thought it was the Coomassie itself, perhaps due to chunks of undissolved Coomassie; however, I filtered the stain and still have the same problems. My most recent gel looks the exact same as this image, with the same two white circles.

Additionally, is using 1% gelatin appropriate? I have seen researchers use 0.1-0.2% gelatin co-polymerized with the polyacrylamide, but the source I listed above used 1%.

Thank you!

Hello. I wanted to prepare whole cell extract for western blotting. Currently, I

Step 1. Lyse cells in detergent (e.g., TritonX-100),

Step 2. Take some lysate for determining total protein amounts (e.g., by Bradford assay),

Step 3. Add SDS sample buffer to the rest of the sample, boil.

However, this method has caused viscosity and made sample loading quite difficult, and the WB membrane showed band streaks upon Ponceau staining. And I think the genomic DNA content that remains in the sample is the reason.

Does anyone know how to effectively remove DNA during preparation of whole cell extract? Thanks very much.

As part of my western blot procedure, I ran my SDS-PAGE electrophoresis (16% polyacrilamide for resolving gel, and 6% stacking gel) at 124V.