SDS-PAGE - Science method

Hi everyone!

So I am trying to concentrate my recombinant proteins in order to make some assays with them, but I need a "considerable" concentration of each. One of my proteins is around 9kDa and the other one 15.8 kDa. They have been tagged with a 6 His and went already through a Nickel column. SDS PAGEs show considerable concentration of certain amount of protein of interest in many, fragments, hence, it was decided to concentrate them using a PES protein concentrator (3,000 MW concentrators) . However, when spinnning them as protocol suggests and collecting every flowthrough (FT) to localize any "lost" proteins, as well as the concentrated sample, rather than seeing a considerable larger band on the SDS , the bands are discrete, not much concentration is found and there is no protein whatsoever in any other FT to suggest protein is being lost through the filter. Am I facing degradation, or why is it that even concentrating it, is not concentrated.

Thank you all! Welcoming any suggestions or corrections!

Best wishes,

Jorge

I would like to know if anyone tried the western blot with the Instant Blue stained SDS-PAGE gels. Thanks.

Hi, all

I am doing the liposome flotation assay. In the end, I precipitated the protein and then ran the sds-page. But every time I could not see my protein in the gel, it was almost gone, maybe just like a shadow. I asked my colleagues, and they said something wrong happened during my precipitation. I want to find the reasons. Please provide some suggestions for me.

Here is my TCA precipitation protocol:

1. add 1 volume TCA to 10 volumes of my sample, and incubate 30min at 4C

2. centrifugate at 15000rpm, 20min, 4C

3. discard supernatant, and wash with acetone two times (then centrifugate at 15k, 5min, 4C)

4. remove acetone carefully; avoid touching the white precipitation

5. air dry overnight

6. dissolve in 2X loading buffer for SDS-page on the second day

Thank you!

April

Hi everybody!

I reads some papers or webs on protein sequencing using maldi top MS to sequence digested peptides. But I am wondering that all informations i collected is only about identification of protein bycomparing peptides sequence, % coverage sequence and matches.... No information on full sequence of tageted protein. Can I now exact protein sequence from excised protein band? Or only obtain via protein-coding gene sequencing??. Thank all.

Hello everyone,

unfortunately, Bio-Rad discontinued their Mini-PROTEAN 3 Multi-Casting Chamber, with which one could prepare up to 12 polyacrylamide gels for SDS-PAGE in parallel.

Does anyone know of a similar product with compatible dimensions?

Thanks for your input!

Best

Karina

I am running SDS-Page western blot using 10% acrylamide gels. However, my samples are not migrating more than 55 kDa. The bands are not defined. I am using 4x Laemmli buffer with LDS from Biorad. The cell lysates are human whole brain lysates. I am wondering if the LDS has something to do with this? I tried to boil the samples at 95 degrees for 5 min; heat at 70 degrees for 10 min, all did not work.

Write the composition of Resolving Gel

Hi !

I am trying to run some gels but everytime I do, my samples end up looking very wavy while the scale is ok.

For now my protocol is : sds page gel 12% acrylamide, run at 200V at room temperature for 40min. I prepare my sample by mixing them in a dye (commercial one) which I have to add beta-mercaptoethanol (BME) to it according to the notice. Then I put then 5 min at 95°C and I load them onto the gel.

I already tried to change few things in my protocol to improve my results but nothing worked I always have this huge blot in the end instead of thin strips. I changed :

- all the solutions and bought back every item so I am sure they are fresh.

- I ran the gel in the cold room at 4°C

- I ran the the gel at 120V

The only thing that comes to my mind now is to remove the BME because it is the only thing that I add compare to the scale.

Have someone had the same kind of issues ?

Thank you very much in advance ! :)

Jenny

I am facing an issue related to horizontal smearing on SDS gel. I have changed all the buffers and acrylamide with fresh ones, but I am still facing the issue. Below, the image is attached, where you can see the horizontal smear (thin line) appearing at the end of the gel. Other vertical smears in some wells are due to samples, but the horizontal one appears in every gel. Can you please provide a solution to solve this issue?

Cannot separate between 10~25kD, always have a line around 25kD...

(Resolving gel buffer: 30% Acrylamide/Bis, 1.5M Tris-Cl, pH 8.8, 10% SDS, 10% APS, TEMED)

I am working on expression and purification of one cytoplasmic protein with His tag, in E coli Bl21(DE3) host cell. Here is SDS picture. Line 2 and 9 are cell lysate, 3,4,7,8 wash steps and 5,6 are elutes using different purification procedure. The expected size is 48 kDa. For protein extraction I used a high pressure homogenizer, also I didn’t use any inhibitors. I was told to try to use Bugbuster protein extraction reagent supplemented with benzonase, do you think it might help?

I am running protein samples on SDS-PAGE that are tagged with fluorophores. If I place the gel in a destain solution of 60%water/30%methanol/10% acetic acid, the chemistry reverses and I lose the fluorophore. Is there an alternative storage solution I can use to get my gel to the imager to capture the fluorescence without chemical fixatives? After imaging, I then proceed to Coomassie Blue staining in acetic acid and methanol, at which point I am not worried about losing the fluorophore.

I have inserted crispr vector in BL21 competent cell with ampicillin resistance and after induction with IPTG for protein ...no cas protein band was obtained using SDS page.. not even near to its size both in the supernantant as well as in the pellet... Can anyone help me.. the vector I ordered contains 6x his tag... But no protein after NI Nta agarose...

Problem

After Transferring the membrane picture with Ponceau S Staining looks like the membrane is burning and has poor transfer. Can someone help with this?

I also noticed something weird on the gel after the transfer, it seems there are some blue spots with Coomassie Blue staining.

12% gel

I load 30 ug protein in each lane with six samples and two ladders separately.

Wet method: 19-hour/constant 30V at working fridge (4 degrees) with ice bucket

PVDF membrane active with the methanol > 1 mins

The transfer buffer is fresh with 25 mM Tris, 192 mM glycine, and 20%methaol but make a 10* stock solution of transfer buffer, which is 250mM Tris 1920mP Glycine, then add 700ml ddh20 and 200ml Methanol to make 1L transfer buffer.

I want to check the mTORC activity and for that I am using rat kidney. I used cell lytic buffer from Sigma Aldrich to extract the proteins from rat kidney (50mg sample homogenized in 200ul cell lytic buffer with Halt protease and phosphatase inhibitor cocktail followed by centrifugation at 13000 g for 15 minutes), took the protein concentration by BCA assay, and then ran SDS-PAGE at 80V for 1.5 hours. I used BioRad's 10X Tris/Glycine/SDS electrophoresis buffer. I then did the transfer to PVDF at 0.18A for 1.5 hours. I used 10X Tris/Glycine as a transfer buffer. I incubated the blot with S6 primary antibody at 4 degrees overnight, washed with TBST 3 times and incubated with secondary antibody for 2 hours at 4 degrees followed by TBST wash 3 times.

When I developed the membrane, I see the bands are not aligned. I see it smeared. Can anyone help me point out what could be the issue? Shall I lower the voltage while running the gel? or the problem could be something else?

Hi Everybody. I'm senior in a University in Vietnam. This is fisrt time I do SDS-PAGE, so I want to ask about the correction of my loading buffer recipe for SDS-PAGE as a following file.

In addition, If 0.5M Tris-HCl pH 6.8 isn't available, Can I replace it by 1M Tris-HCl pH 6.8 or any other concentrations of Tris-HCl pH 6.8.

Also, how should I mix these components? (In order or out of order)

(Samples I am using in my research is piglet intestines)

All my SDS Gels are having wave like thing and no prominent bands. I checked the pH of the buffers and they were optimum.

I have purified overexpressed protein from BL21 (DE3) cells using Ni-NTA column. When we run purified protein through native PAGE and SDS PAGE both which showed different result. In SDS page showed only one band of purified protein whereas two band in native PAGE. I have proceed whole experiment three times and found same results. What is the possibility to find two band in native page whereas it one in sds page. I have attached native PAGE image.

We have expressed our protein in chloroplast when we add extraction buffer it get precipitate soon after centrifugation as this is real quick even we don't have a time to prepare sample for SDS-PAGE and Bradford. We have changed the buffers and tried Tris, HEPES, Phosphate and got some how comparable results with HEPES then we check the concentration variables of HEPES from 50mM to 100mM, EDTA from 10mM to 20mM, PEFA-BLOC (protease inhibitor) from 1 to 5mM but it does't working. We have tried TCA/Aceton precipitation, 8M urea treatment in both extraction and sample buffer but still the problem is there

I have a protein extract that is not purified, I ran it through SDS-PAGE and observed bands of 150-200 kDa and other smaller ones of between 60 and 8 kDa. A colleague ran my samples through FPLC with a 6HR superose column, however in the FPLC protein profile I see peaks of less than 20 kDa, but not larger ones. What could have happened to the larger proteins?

In a Western Blot (WB) experiment, does the concentration of SDS-PAGE gel affect the position of protein bands? In the image, the same protein samples were used for all five lanes. The left side (lane 1, lane 2, lane 3) used a 10% gel, while the right side used a 12.5% gel, with all other conditions being consistent.

Thanks for your kindly help!

Hello,

For many phosphorylation studies of MAP kinases, cells are serum-starved overnight prior to stimulation. Serum-free media such as DMEM 1X is supplemented with 0.5% bovine serum albumin. Is there any drawback to including NEAA in the serum-free media? Is NEAA known to cause phosphorylation of MAP kinases?

Thanks!

In a Western Blot (WB) experiment, does the concentration of SDS-PAGE gel affect the position of protein bands? In the image, the same protein samples were used for all five lanes. The left side (lane 1, lane 2, lane 3) used a 10% gel, while the right side used a 12.5% gel, with all other conditions being consistent.

Thanks for your kindly help!

My current protein concentration is 13mg/ml. To load 30ug protein for SDS PAGE, only 2.15uL is needed, can i load 2.15uL sample + 2.15uL 2X Laemmli Buffer = 4.3uL in the well or do I dilute the sample first so that more volume is needed?

If so, how do I dilute the cell lysate, or can I just use PBS?

Help!

We are optimizing SDS-PAGE, i would like to know the best ratio and percentage for the Staking gel and separation gel of Acrylamide.

Its like 37.1:1 and 29.1 and 19.1 and also 30% and 40%,

So i am confused about it.

Your recommendation would help alot.

Hello everyone,

I am performing western blot (SDS-PAGE) for the phospho-mTOR protein, molecular weight 289 KD. I am using 6% resolving gel for it. I kept the transferring time around 4 to 6 hours at 60 volt (4 degree C). However I am not able to get the proper band. Please share any idea/knowledge.

I am trying to detect a secreted protein by Western blot in plan tissue. To this aim, we are using a simple protein extraction protocol based on TX-100 and DTT. Then, I am loading the samples in a SDS-PAGE and doing western blotting. However, I don´t detect my protein eventhough it should be overexpressed… We have read that this protein is secreted. Should I perform another protocol to identify it? Can this protocol somehow promote degradation of my protein ?

I am running SDS page gels using the mini-PROTEAN tetra vertical tank. Gels ran normally up until about 3/4 of the way down the gel. At this point, the dye changes from blue to yellow and became distort. I have inspected the module so this is not the problem. The electrophoresis buffer used has the correct pH. The loading dye was Laemmli Loading Buffer. All gels were run at 140V for 50-60 minutes.

Hey all

A few of my lab-mates store the protein after quantification (I used bradford assay) in cell lysis buffer containing protease inhibitor in -80. While others add loading dye and denature the protein in 95 degree and store it in -20.

I have quantified the protein in the cell lysate. However dye to time constraints and huge sample protein I couldn't add the loading dye after the quantification process. (2 hours had already passed While I was calculating the amount of loading dye required for each sample). I got panicked thinking the protein in the cell lysate would be degraded and hence upon an advice from a fellow senior I aliquoted 20uL of each sample into another 1.5 mL centrifuge tube and stored the the stock and the aliquot in -80. So that I need not freeze and thaw my stock again and again.

Following are my queries

1. At what stage is it recommended to store the protein?

2. Does the concentration differ after storage?

3. Do I need to do bradford assay once again after I thaw them from -80?

4. what is the incubation period for bradford assay? (after adding BSA to the bradford reagent how long should I wait to take the reading or should I take the reading straight away?

Thank you

Wishing you a happy christmas and a happy new year

Purpose of using stain and de-stain in SDS-PAGE gel

I am working on the brain tissue of mice (10% SDS-PAGE gel). The protein sample was extracted with lysis buffer (Tris-HCL, Triton, and B-mercaptoethanol). I think this leakage is due to the interaction between the protein sample and Triton. To be sure of this idea, I tried to do a control gel in which only the lysis buffer and the marker were loaded, where a well-separated marker refers to good electrophoresis, and after the staining step, the background was clear. what should I do?

I made two 10% sds-page gel (10% resolving and 4% stacking) and run them at 20mA. I am testing whether my newly edited recipe for gel casting is correct or not. Apparently, the gels took 3hr for the dye front to reach the bottom of the gels. The problem is somehow the protein ladders started to separate within the stacking gel, there are 4 to 5 bands above the interface and the remaining bands at the resolving gel.

May I ask anybody who knows how to resolve this issue or not?

I tried to express a Large fusion protein (about 223 KDa) in bacteria (BL21de3) in the pet28b vector, but it failed as the SDS PAGE shows just very few proteins successfully expressed. I wonder if may change the vector to pCold TF.

At a fixed voltage of 260V, electrophoresis of protein was faster in our previous batch of 1x SDS running buffer. However, the electrophoresis was much slower recently with much lower current (less than half of the previous one). The same issue occurs even with new dilution of freshly prepared 10x buffer to the 1x buffer. What would be the possible reasons of such issue?

I mixed EDTA mouse plasma with standard gel sample buffer and got very smeary image on Western with no distinct bands. Is there something in the plasma that needs to be removed before I run gels?

Hi everyone,

I purified Pfu and I treated samples in the heat block. however in the Pfu lane I can see multiple bands, can someone explains what could it be?

Hi all,

I was doing protein purification and after that I had to add 2 uL DTT (1mM), but instead, I added 1000 uL DTT (1mM) to my protein solution. I would like to ask you, if my mistake has any bad effect on my protein? I have to run SDS-Page, but I don't know if this amount of DTT would change the results?

best

Amir

Do you know of any of this software that is compatible with Apple computers? I tried GelAnalyzer4, PyElph, and the free online gel analyzer, but it is not working anymore.

What would be the minimum amount of protein needed to detect heme by TMBZ (3,3,5,5′-tetramethylbenzidine)? I am using cytochrome C from equine heart (positive control), chitinase (negative control), and Z-ISO (unknown). I want to do a quick sporting on filter paper of the presence of heme.

I would greatly appreciate any suggestions.

I am relatively new to conducting Western blots. I employed the alkaline lysis method to extract whole proteins from yeast. Subsequently, I introduced the Opti Protein Marker (ABM, CAT log NO: G252) and completed the gel electrophoresis. The proteins were then transferred to a nitrocellulose membrane using the wet transfer method. Following the transfer, I performed a Ponceau staining, during which the protein markers were clearly visible.

Moving forward, I proceeded to block the membrane for 1 hour in 5% BSA in TBST (Initially, I attempted blocking with 5% non-fat skimmed milk, but encountered high background signals). Subsequent to blocking, I washed the membrane with TBST (3 x 7 mins) and TBS (1 x 5 min). Following this, I incubated the membrane overnight at 4 degrees Celsius with a primary antibody (Beta-tubulin, Rabbit IgG Polyclonal), 1:1000 dilution in 5% BSA in TBST). Afterward, I repeated the washing steps with TBST (3 x 7 mins) and TBS (1 x 5 min).

For the next step, I incubated the membrane with a secondary antibody (Rabbit anti-Goat IgG (H+L), HRP, Polyclonal, 1:10,000 diluted with 5% non-fat skimmed milk in TBST) for 3 hours. Subsequently, I performed additional washes with TBST (3 x 7 mins) and TBS (1 x 5 min). Finally, I carried out chemiluminescence detection with an exposure time of 30 seconds.

However, my results were unsatisfactory as I observed multiple nonspecific bands, and the protein marker disappeared. I seek assistance from experienced researchers, especially those familiar with Western blotting of yeast proteins. I would appreciate any insights or suggestions to identify and resolve the issues.

Additionally, please refrain from suggesting the use of monoclonal antibodies, as it is currently beyond my budget constraints.

Thank you in advance for your help.

Gel electrophoresis, recombinant protein, expression in bacteria, molecular biology

I prepared GST-tagged protein. The molecular weight of my protein (50kda) with GST (26 kDa) tag is 76 kDa. After removal of GST tag the molecular weight should be 50 kDa but in SDS page I got 40 kDa. Can anyone please suggest me the cause?

I was doing transformation using plasmid with kanamycin resistance into BL21 DE3 E. coli cells with ampicillin resistance. I plated the culture on amp+kana plates and amp only plates. However, only colonies were found on the amp only plate. Since the transformation has fail several times before this, I decided to inoculate the colony from the amp only plate and continue the experiment (grow the culture to 0.55 OD600 and add IPTG). Afterwhich I ran a SDS-PAGE. I observed a difference between before IPTG and after, and the protein bands in the after IPTG lane corresponded to my protein of interest, which should not have been expressed... Does anyone have any knowledge about this? Please help.

Hi. Recently, as a mean to increase yield, I added 0.1% of Tx-100 into all of my IEX purification buffer including the sample injected. However, when analyzed on SDS -PAGE, I observed that there was no single band of my POI at the target size but instead as the picture shown; strong band in between 50 and 37kDa and a very faint band at 25kDa (is not visible in pic). FYI, my target protein is 49kDa and the GST tag is 26kDa. Is it possible for TX-100 to cleave my protein?

I need a detailed protocol of SDS-PAGE only for drosophila if anyone has done before

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.

Good afternoon,

I try to extract cell wall proteins from green macroalgae, I have tried different protocols and one of them included step of EGTA extraction, however this fraction is jelly like, so I expect polysaccharides from cell wall to be present. When loaded to SDS-PAGE, it was obvious that the gel pores were blocked and no bends were obtained, just light blue streak. According to Bradford, in this fraction I have quite a lot of proteins with which I would like to do Western Blot... I have already tried aceton precipitation, but still a lot of polysaccharides remained....So please do you have any recommendations how to get rid of these polysaccharides?

Thank you very much,

Tereza

In my procedure, I first label the protein with a probe and perform a Copper-catalyzed azide-alkyne cycloaddition (CuAAC) or CLICK reaction to append the protein-probe complex to a reporter tag. I have seen in several articles that the samples are not heated at 95°C in the loading buffer before running on an SDS-PAGE gel because of which I don't get crisp protein bands on gel. Is there any particular reason for not heating the sample? Can I do something to get better protein bands on the gel? [I have attached a gel image for this as well]

Thanks in advance.

Hi everyone,

I did SDS-PAGE for a series of proteins that I was sure about them. In my SDS-PAGE was added TCE (2,2,2-trichloro ethanol) as stain free for proteins. After running the gel I took it and put on UV transluminator for giving uv wavelenght (about 250-360 nm) excitation, AFTER exposure time about 1-5 min, I deteced for emition but no sign with proteins bands and gel was clear.

What should I do for visiting my proteins bands under UV?

I had proteins bands when I was staining gel by commassie blue.

Do I need a specific chemoDOC instrument? or I can do it by typical UV transluminator?

Best wishes

I am running some pulldown experiments using plasma samples(click biotin linker on protein, then blot against Strep-HRP). We started the experiment with a known protein (Lane 2-5) then plasma samples (Lane6-9), lane 5 and 9 both incubated with alkyne-biotin. Although the signal is very small and on WB I need to overexpose to see it, but the bands on lane 9 is giving a bizarre signal. Could anyone help me understand the reason for this and how to solve it?

I expressed an extracellular protein and I have obtained good readings on protein assay as it is GFP-tagged. However, I do not obtain any bands for the protein on the SDS PAGE. What could be the possible solution?

Hi all, I used pngase f to release glycan from transferrin. here is my protocol.

1. I prepared storage buffer for the pngase f without glycerol and aliquot them into different vial

2. I took 10 microgram of protein and denatured @ 100°C for 10 min

3. added Np40

4. then glycobuffer

5. pngase f added in the conc of 5mU, 50mU, 0.5U, 5U, 25U, 50U and incubated for 16 hrs.(1:10 v/v) total reaction vol 50 microlitre

6. I took 20 microlitre from that and prep for loading into sds page.

Here is my result. is my pngase f showing activity? or it looses activity because of no glycerol used in the buffer?

Lane 1 - standard

Lane 2 - without enzyme

Lane 3 - precipated after incubation

Lane 4 - 50U

Lane 5 - 25U

Lane 6 - 5U

Lane 7 - 0.5U

Lane 8 - 50mU

Lane 9 - 5U

Hello, we have been struggling with the lots of background in our wb membranes probed with an anti-Streptavidin-HRP from Thermofisher (Pierce 21134). Samples contained biotinilated proteins. Every time there is some blobs somewhere and so much background that it is hard so see our biotinilated proteins. I attached the same pic with different contrast. Did anyone face the same problem?

All stepts have been performed with PBS 1X and here the protocol:

Why is the reduced protein (with 2-ME) placed higher than the non-reduced one in SDS-PAGE?

I need to collect proteins after SDS-PAGE electrophoresis to quantify the proteins present in the target bright line using the bradford method. So how can I separate the protein and remove the polyacrylamide gel and SDS from the protein? Can you recommend some methods and ways to do that? Thanks for everything.

What would be the possible reason for this:

Following are our PAGE setup;

Protein Source:Rat Liver

Preparation : Ground in Lysis buffer and centrifuged and collected supernatant. Equalised all the samples with lysis buffer and loaded using 5x sample loading dye and beta mercaptoethanol.

Concentration : 30 microgram

SDS PAGE : 10%gel. Run 1.5 hr (120 V)

Sample preparation

Sample buffer (SDS reducing buffer)

3.55 ml deionized water

1.25 ml 0.5 M Tris-HCl, pH 6.8 2.5 ml glycerol

2.0 ml 10% (w/v) SDS

0.2 ml 0.5% (w/v) Bromophenol Blue

9.5 ml total volume

Store at room temperature.

Use: Add 50 µl ß-mercaptoethanol to 950 µl sample buffer prior to use. Dilute the sample at least 1:2 with sample buffer and heat at 95°C for 4 min.

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.

I want to identify a secretory protein that is secreted from min6 cell in the supernatant can anyone provide protocols?

Firs I want to get conditioned media and then precipitate after that I will do SDS-PAGE and CBB/silver staining. I am a newbie so I don't know how much medium, how much cells is needed for this assay, I couldn't find a good reference

Hello!

I have been doing some SDS-page analysis and before running my samples I added 5x sds sample loading buffer and then i boiled them at 95C for 5 minutes! The loading buffer fabricant told me it contains B-mercaptoetanol!

Can I say i runned my samples on a gel under denaturing and reducing conditions?

thank you

Dear all,

I have been trying to knockdown using siRNA (duplex) targeted against my protein of interest (# 2 lane shown in the image) and a random non-target siRNA ((# 1 lane shown in the image) treated cell as negative control and non-treated normal cell (# 3 lane shown in the image). I don't see silencing of the gene as could be seen in lane 2. Respective actin loading control is shown below the bands of interest. Currently, I am using 20 nM siRNA with interferring reagent protocol provided from thermofischer. If anyone can suggest any protocol that I can try, will be greatly helpful.

Thank you

best

Prem

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.

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!