- Maximilian Peters added an answer:How can one solve the problem of getting expression bands lower than expected size?
I am working with a protease gene which is his-tagged at c-terminal and is ~50Kda size. I transformed the recombinant construct (pET23b+Gene) in Rosetta DE3 pLysS. When I am trying to check induction I am getting results like that n given SDS PAGE.
The conditions are:
Induction with 0.6mM IPTG when od reaches 0.6
Lane-2 marker (97.4kda, 66kda 43 kda 29kda 20.1 kda and 14 kda)
lane-3 uninduced cells 200ul
lane-4 induction of 2 hours
lane-5 induction for 4 hours
lane-6 induction for 6 hours
lane-8 20 hour induction.
As you can see the gel image expression is going on but its giving band of much lower size than expected. I have done sequencing/ primer walking of my gene, they are intact and 100% correct.
In your 20h induction sample you have a 2nd band with a higher molecular weight. This band is absent or very faint in your other samples. Is it possible that this is your protease?Following
- Amelia Trimarco added an answer:Why has PCR amplifying 4kb DNA using cDNA as a template failed?
I used the SMARTer RACE kit from clontech to get the 3' and 5 ends of my target gene. Then I tried to PCR amplify the whole gene. I tried both the platinum Taq High fidelity DNA polymerase and SeqAmp DNA polymerse system but didn't work. The cDNA I am using is the 5'RACE ready cDNA. Primers were designed based on the 5' and 3' UTR, 18bp long and with Tm around 50 degree. I know my target gene has low abundance. So I always add about 400ng cDNA to each reaction. I tried annealing temperature ranging from 42 to 55 degree. Extension at 68 degree for 4-5min. Initial denature at 94 degree for 30sec to 2min. And Ma2+ concentration is 2mM for platinum Taq DNA poymerase and 1mM for SeqAmp DNA polymerase.
Can you give me some suggestions on PCR a 4kb product? Thanks.
Do you have no product or some not-specfic products?Following
- Stephanie Misquitta added an answer:Is it possible that a very low amount of protein is expressed in uninduced culture too ?
I induced a culture (Rosetta gami pLysS) with IPTG ( 0.5 mM and 1 mM) at 37 degrees. Together with it I also kept a control ( uninduced). After lysis, when I run SDS-PAGE gel, I get a band at appropriate position in induced culture pellets, but I also get a faded band at the same position in control as well. Is it possible that my protein gets expressed in control without any induction ?
Yes, It is common to get expression in uninduced cells. You need to determine if this is "leaky" expression or the endogenous protein. Using an empty plasmid would be a good control.Following
- Sobia Ejaz added an answer:Why is there no protease activity in my protein sample?
I ran gelatin zymography of my plant protein sample but there was no zone of hydrolysis. Moreover, the bands appeared similar as that of sds page bands. I ran sds as well as native zymography and both showed the same result. Is there is no protein activity in my crude sample?
Thanku Amaj .Inshallah I will follow your suggestion......Following
- Gary Laco added an answer:What is the reason of getting three bands in the 12-14 kDa position instead of one in SDS gel ?
I expressed 11 of my mutated proteins at 16C for 3days in 2YT media with 1mM IPTG; purified them following His tag purification method. Always I maintained cold environment, did not do freeze-thaw cycle. My protein size is 13.87 kDa. After running SDS gel I am getting three bands instead of one. I am wondering whether my proteins are degraded any reason. I have attached the gel picture here.
I would run the WT protein on the same gel with the mutants using the same SDS/PAGE buffer to make sure you are not getting artifacts from the SDS/PAGE loading buffer. It looks like the mutant proteins are being cleaved, but I have gotten doublets that were due to the SDS/PAGE loading buffer (need to make with Tris acid and NaOH to adjust pH, do not use Trizma).Following
- Lipsy Chopra added an answer:I am getting vertical streaking in my SDS-PAGE. Is there any method to remove them?
Thanks in advance for your replies.Following
- Ying Zhang added an answer:How can I change an insoluble recombinant protein to a soluble protein?
How can I change insoluble recombinant proteins to soluble proteins? My expression vector and host are pet21a and Bl21 respectively. Can I resolve this problem by host change?
You may try autoinduction at low temperature which doesn't need IPTG, so that the induction will be very mild and protein expression will not be a great quantity within a short time. For Autoinduction you need a special media.
You may also fuse your protein to a larger well soluble protein, which may increase solubility of your target protein. After purification you can cut your protein by a protease cut sit between the two protein-sequences.Following
- Mangesh Bawankar added an answer:I require clarification for induced and uninduced cells concerning my SDS-PAGE of a protein. What do you think?
I run the gel of my protein. My protein is cloned in a pET-16b vector and transform in BL21 (DE3) strain. I induced with 1mM IPTG at 0.54 OD and incubate further at 37 degree for 16 hrs in LB broth with ampicillin. The protein has a His-tag and was not purified by Ni-column yet. As you can see in the picture, there are two bands separated by a few kD which are present in the induced culture but not in the uninduced culture. One of the two bands were of my protein. The molecular weight of my protein is of 17kD approximately. I did not understand why the two bands are coming when only one protein is there and how I can get rid of it.
Thanks all..I got the single band after Ni column purification from the 3 hrs induced culture.Following
- Nick S Berrow added an answer:How can we increase the solubility of E. coli overexpressed protein?
We are trying to purify an artificial protein. Very good expressed, but mostly in the spun down pellet. If anyone has experience to make it a bit more soluble, with a kind of mild buffer, additives, will be helpful. Thank you.
First-check the Q&A forums there are hundreds of similar threads on ResearchGate that that have already been answered very well!
Can you be a bit more specific about your tag, fusion protein, expression system etc?
As you already have a construct that expresses then there are a few things that you can try (I am assuming you are using T7, LB and IPTG at 37°C so far with a globular protein and not an IDP).
Firstly if you want to continue to use IPTG for induction use a richer, buffered medium , terrific broth is simple to make but can improve soluble yields dramatically over LB.
Try reducing temperatures post-induction to 16-20°
Try reducing IPTG concentration-although in most e.coli expression strains induction is not really titratable.
Try auto-induction media instead of IPTG
If none of these work then think of changing your fusion protein or tag, I particularly like SUMO, thioredoxin and Z-tag (and much prefer them to GST and MBP) and/or changing the boundaries of your construct to avoid e.g. unstructured domains. There is no sure way of getting soluble protein that stays soluble after cleavage from the fusion but these should help.
Of course the effects of all of these are dependent on your protein and it may be that you ultimately have to accept re-folding from inclusion bodies.
P.S this answer is a copy of my answer to a similar older threadFollowing
- Ravi Gupta added an answer:How do I concentrate protein from lamelli buffer?
I already extracted my protein directly in SDS buffer containing Bromophenol blue and also B-mercaptoethanol. After running SDS page, I am having problem with protein concentration. Is there any way by which i can concentrate my proteins in the buffer like we use vaccum centrifuge for DNA? I have more than 100ul of sample volume...please suggest
Protein conc can be determined using 2D quant kit (GE Healthcare). We regularly use it for quantifying proteins in SDS loading or lysis bufferFollowing
- Alice Moscovici added an answer:Does anyone know how to run SDS PAGE for glycoroteins?
I have been trying to purify glycoroteins but they are fail to appear on SDS PAGE because they are highly coated with sugars. How can i Stain them through manual method?Following
- Paul E Harris added an answer:If panning to change the thickness of Western Blot gels from 1.0 mm to 1.5 mm, how much more time should the SDS-PAGE and WB be operated?
Recently, I have been planning to switch the thickness of SDS-PAGE gels for Western Blot from 1.0 mm to 1.5 mm to accommodate for larger samples. I use Life Technologies' NuPage Tris-Bis 4-12% gels. If I had been running for 100 min at 70 V previously with 1.0 mm, how much longer (and perhaps with higher voltage) should the gels be run if the thickness is bumped up now to 1.5 mm? As for WB, I had been running for 60 min at 30 V.
The company's protocol does not say much about any modification regarding a change in the gel thickness.
I am wondering if anyone had to make such a change as well and has already figured out how much more time relatively these two steps would have to be taken.
Regardless of any advice given here. You really should check the transfer of protein from gel to membrane using the prestained molecular weight standards. We too us the NuPage 4-12 gels and get good transfer of 20-30 ug on a 1.0 mm Gel to nitrocellulose in 20-30 minutes and 45 minutes for the 1.5 mm gels..but it all depends on your blotting system.Following
- Daying Wen asked a question:What is your program to clean the HPLC RP column between blood sample injections?
Mobile phase acetic acid-MeOH-acetonitrile 70:15:15
How many CV for high concentrated solvent cleaning and re-equilibrating between sample runs?Following
- Craig Skinner added an answer:How can I identify and characterize a monoclonal antibody further?
I immunized BalB/C mice with extra-cellular part AXL protein[BL21(DE3)-pET32a-AXL, ~63kDa)] which is a recombinant fusion protein. After fusion of spleen cells and SP2/0, I screened positive clones with indirect ELISA using the same recombinant fusion protein and un-purified induced tag proteins[BL21(DE3)-pET32a] as control. Finally I got 20 clones and they are positive in AXL coated ELISA wells and negative in control wells.
I know the routine way to identify an antibody is Co-Immunoprecipitation, Western Blot and then Mass Spectrometry. But same antibodies are not suitable for Western Blot, so how can I identify these antibodies? THANKS!
You could try a native PAGE western, though I'm assuming AXL is a membrane-bound protein so this might be difficult. Also, you can get the dissociation constant by various methods, or use it for immunofluorescence. IMF is really fun and makes beautiful figures too!Following
- Weber Lin added an answer:Can a protein degrade only at the C-terminal?
I want to produce a recombinant protein about 65 kDa by BL21 E.coli and purify by Ni-sepharose beads (GE healthcare). (I fused myc and his tag at C-terminal end of my protein)
After elute my sample with elute buffer contains 250 mM imidazole, I dialysis my protein to PBS (pH 7), enhance my protein concentration by 50 kDa condensor, run 10% SDS-PAGE to confirm the purity of recombinant protein and storage at -20℃.
I detected a major band between 55-70 kDa in molecular weight ladder (I think that is my protein)
But I cannot detect the protein signal by Western blot no matter using anti-myc or anti-his tag antibody.
I think there is something wrong in my recombinant protein, but I don't know what's happening.
I have checked the plasmid contruct by restriction enzyme digestion, primary and secondary antibody's function by positive control.
Is there any possible protein degrade only at C-terminal of protein?
Thanks for everyone.
I will add protease inhibitor to inhibit the protease activity during purification and try it again.
I'll also use 30 kDa condensor or S-200 gel filtration column to enhance the purity of my sample.Following
- John C Schmitz added an answer:What is the best method to normalize between gels?
Hi, I'm running Western blots, looking at various proteins involved in the mTOR pathway and I'm looking at phosphorylated versus total protein ratio.
I have 8 replications (animals) and 4 treatments and am using gels with 15 wells so I can only load 3 reps (each with their 4 treatments) per gel. I'm using the same procedures but sometimes exposure times will differ somewhat on each occasion.
What would be a good way to normalize between different gels? How can I account for differences between gels.
I was also wondering if anyone ever uses a positive control and normalizes to that. Right now my data is organized as phosphorylated protein as a ratio of the positive control (which is a pooled random muscle sample), total expressed as a ratio of the positive control and then the final value is a ratio of those ratios as in Phosphorylated versus Total.
Another option is to use both antibodies (phospho and total) on the same blot. The antibodies need to be derived from different species (a rabbit antibody and a mouse antibody). The secondary antibodies contain infrared dyes that are detected at different wavelengths. We use the Infrared scanner from LICOR to detect both signals simultaneously (many core facilities or big labs have one of these scanners). The primary antibodies must also recognize different protein epitopes (to avoid interference). Admittedly, this takes some trouble shooting to determine the optimal conditions, but once the conditions are set, the technique eliminates normalization between two gels. We typically test for GAPDH just to make sure our protein concentrations were similar but we don't normalize to GAPDH. Just the ratio of phospho/total protein.
- Gaurav Kumar Singh added an answer:What should be the ideal dialysis buffer composition for a His-tagged purified protein to remove imidazole salts?
My His-tagged purified protein contains 10mM trisCl, 300mM NaCl and 200mM imidazole at which the protein was eluted at pH 8.0. What should be ideal dialysis buffer composition to remove imidazole?
I am using 10mM tris and 300mM NaCl at pH 8.0.
Does NaCl affect protein function? Do I need to remove it also?
Thanks Antonio. I really appreciate your answer. Yes, I did centrifugation and I have lost some of my protein in the pellet. I wish I could put them back in solution!Following
- Scheherazade Sadegh-Nasseri added an answer:How do I couple proteins to cnbr-sepharose for making a protein specific resin?
Firstly, I wanna purify one kind of antibody from serum which is specific to one kind of protein. Ex) protein XX. So, I coupled this protein XX to CNBr sepharose resin to make protein XX affinity column. I coupled 1 mg of this protein to 0.75 g CNBr resin, but when I checked the protein XX column's affinity, I found the capacity of this affinity column is so weak to bind with anti-protein XX antibody in serum. Then I wanna know what's the problem? If I coupled so small amount of protein to CNBr resin?
One thing to warn you is that the recommended amount of protein by GE is on the high side and can cause cross reactivity. So, do not go too high on protein amount to couple-stay within 2-5 mg/ml wet volume, no more.Following
- Miranda Ween added an answer:Does somebody have any suggestions to overcome outer membrane protein purification problem?
We are trying to purify a membrane protein expressed in BL21. We tried both N and C terminal his tag, but the protein doesn't bind well to the Ni column. Along with the protein of interest, many bands are appearing in the elution. Strangely, the protein doesn't have (or very poor) UV absorbance. Also, I don't see any color change when I add SDS sample buffer to this protein, but still see a band in SDS-PAGE. I tried Co instead of Ni, and got a band smaller than expected, but it didn't show activity. I think something is happening during the purification because I see enough protein in the membrane suspension. I use DDM to solubilize it from membrane because we want to crystallize the protein. Proteins pI is 9. 2, buffer pH is 8 in all cases (from cell lysis to purification). I am new in membrane protein field. Any advice would be appreciated. Thank you.
Membrane proteins are tricky. they are at such a low ratio compared with other host proteins even when overexpressed. My suggestions would be these.
1. increase your binding buffers level of imidazole. If you have at least 6xHis as your tag it should elute at 250mM Imidazole or above. so you could try 80 or 100mM Imidazole in binding buffer. This prevents those host proteins from ever binding to your column and clogging it up, preventing your protein from binding in the next mls of sample.
2. Increase your sample load. I have been known to solubilise 200ml of membranes before and only get a very small, but fairly pure UV peak of protein. if you are only loading a few ml of solubilised membranes, it is possible the amount of your protein you elute is so weak it can't be seen on coomassie but will be able to be seen by His antibody
- Leo J.L.D. Van Griensven added an answer:Does anyone know about the Sevag method for protein removal?I am extracting Lentinan from Shiitake and the method says that I need to remove protein usin the Sevag method. I have searched and the only thing that I have found is the usage of chloroform and octanol, but I haven´t found the exact method
Interphase is the junk layer between the water layer and the chloroform layer. Denatured protein is for large part in the interphase (and might stick to the wall of the tube; take care)Following
- Lin Yuan added an answer:Why is it that my biotinylated protein cannot be purified?
I am woking on IP recently, but one of my target protein which has a biotin and HA tag never can be found after being purfied using streptavidin dynabeads (purified using Streptavidin beads, and tested using HA antibody by WB), even though the protein expression level is high enough before purification.
however, other proteins are happy with the same lysis buffer (20 mM Tris, 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1% Triton X-100, 2.5 mM sodium pyrophosphate, 1 mM β-Glycerolphosphate, 1 mM Na3VO4, 1 μg/ml Leupetin, 1 mM phenylmethylsulfonyl fluoride (PMSF), pH7.5), and similar megenic beads.
I am really confused about that, and need your help! Thank you in advance!
Hi everyone, I tried evey suggestion you give. Finally this problem has been solved, it ia because the lysis buffer I used is not fit for this Streptavidin beads (Dynabeass M-280). now i find the right buffer and everything goes well.
Thank all of you very much!Following
- Adam B Shapiro added an answer:Why is the zero'th time deacetylation reaction showing higher florescence intensity than 1hour reaction?
I am trying to standardize de-acetylation assay for my enzyme and I have taken no enzyme reaction as blank. My substrate is labelled with AMC and using NAD as cofactor.
I am not getting saturation curve for both varied substrate and varied enzyme concentrated catalytic reaction.
I am not sure whether my enzyme is working efficiently because only peptide and test reaction with enzyme giving same intensity. One more problem is '0'th time reaction is showing higher florescence intensity instead of the 1hr reaction?
I am not able to understand the problem please help
If you see a linear dependence of the reaction rate on the substrate concentration, rather than saturation, a probable reason is that the Km for the substrate is much higher than the range of substrate concentrations you are using.
You should see that the reaction rate is directly proportional to the enzyme concentration. That is expected.
If you are seeing no reaction, your enzyme may be inactive due to denaturation, misfoldiing or aggregation. Or the buffer conditions you have chosen are far from optimal for the enzyme. Or you have omitted an essential ingredient that the enzyme requires for activity, such as Mg2+.
As for the observation that the fluorescence intensity is higher at t=0 than at t=1 hour, is this a large enough difference to be meaningful, or just random variation? Is it possible that you have allowed the fluorophore to become photobleached during the hour through excessive exposure to light? Has the sample evaporated during that time, reducing the volume of it that the light passes through? Has the peptide adsorbed to the surface of its container, reducing the concentration left in solution?Following
- Mona Naeema Rahman added an answer:Nickel column takes too long to runI am trying to purify my protein using Nickel column after expressing it using a Baculovirus expression system in Insect cells. I use detergent to lyse the cells. Spin it down to remove DNA and other cell debris and then apply the sample to a Nickel column. The problem is, the sample runs through the column very slowly and the whole purification process (gravity flow) takes me several hours. I use 5mM and 20mM of Imidazole to wash and elute using 500mM Imidazole. The flowthrough and wash steps take several hours. Since I am not seeing my protein on the SDS gel, I believe it might be getting denatured due to the length of the procedure. What am I doing wrong?
One thing you may think about trying in order to minimize the time of each step, is to do your purification in a "batch-wise" fashion rather than on the column. I do this for my protein which I have to keep in detergent as well. Basically, after rocking the clarified lysate with the resin for an hour in the cold room, I spin down the resin and carefully pipette off the "flowthrough" i.e. unbound protein. Then, I add the appropriate wash buffer, let that incubate with rocking for 10-15 minutes and then collect the resin by centrifugation again, pipetting off the solution above i.e. the "wash". I do this for each step. I do this more because of limited time (kids need to be picked up etc) and one has to be very careful to ensure all the buffer is removed before adding the buffer for the next step. You may want to increase the number of washes if you are not confident in your ability to remove all the unbound portion. For 5 mL of resin, I generally use 25 mL buffer for each step so the "trace stuff' really is trace. Also, when you are done, be sure to clean your resin of detergent.Following
- Martiniano M Ricardi added an answer:Can anyone share his/her experience purifying GFP (or GFP variants) in e-coli?
I will soon start expressing GFP-quimeric proteins in E. coli (BL21) and then purifying them using a 6xHIS tag. Does anybody have experience working with recombinant GFP proteins it complicated? Do they tend to form inclusion bodies? Do I have to work always on Dark?. Any comments and advices will be greatly appreciated.
Thank you very much for all the answers! Let´s hope my fusion proteins behave!Following
- Alessandro Chiadò added an answer:How do I minimize protein sample loss during filtration?
I am filtering my protein sample through 0.2um filter as preparation for injection onto size exclusion column. The column specification suggests that the sample shall be no more than 500uL.
The filter I used was Thermo Scientific's Nalgene™ Syringe Filters - 25mm Diameter, which is recommended for 10~50mL solution. No wonder when I filtered my 500uL sample through the filter, a great amount was lost on the filter.
Do any one have suggestions on what filters would be suitable for this purpose?
Thanks a lot!
As Gautam Krishnan you can try to find the proper filter on Merk Millipore website. They sell you filter with low volume retention (such as the very small 4mm filters) and with membrane made with different materials so you can reduce protein adsorption into the filter.
- Mohammed Hakim Jafferali added an answer:When do I add protease inhibitors to my lysate?
I'm trying to 'optimise' the way I prep brain tissue lysate for Western blots. I plan to use MMP-9 Ab- so don't want to use EDTA/EGTA in my cocktail. I'm happy to concoct my own mix (in the first instance- rather than buy commercial EDTA-free tabs).
I just want to know when I should add the protease inhibitor cocktail... once I've homogenised and aliquoted but prior to frozen storage, OR add once I've thawed aliquots and run on gels?
As others mentioned, its best to add protease inhibitor (better use protease inhibitor cocktail, that inhibits different proteases activity) before homogenization. I would also recommend you to add protease inhibitors before and after each step once you ruputre the plasma membrane i.e after homogenisation, after lysis and solubilisation.Following
- Scheherazade Sadegh-Nasseri added an answer:Can you suggest a low-cost way to measure the activity of highly-concentrated proteins?
As part of a quality control endeavor, I’m looking for a low-cost way to measure the activity of highly-concentrated proteins. (IE: purified proteins, sitting in vials in a lab). So far, the best idea I've had is to dilute samples of the proteins (sometimes by as much as 1:1,000,000) and then use off-the-shelf assays kits to detect activity. But the serial-dilutions are a hassle, and the assay kits are not cheap. Are there easier ways? Most of the stuff I've seen seems to be designed for high sensitivity and multiplexed assays – for detecting lots of stuff in serum, for example. But I don't need high sensitivity, and I don’t need multiplexed detection.
Depending on what activity you are aiming for different assays can be used. More information is needed here.Following
- Lauren S Sherman added an answer:How can I isolate a protein band which is not visible by staining methods but shows immunoreactivity?
I need to isolate a particular kDa protein from a crude antigen (whole cyst). I'm not able to locate that particular kDa protein in SDS gel (10% separating gel of 10X10 size mini gel) stained by coomassie stain or silver staining. But if I blot the gel in NCM and perform ELISA using the corresponding patient serum as primary antibody, I am getting a prominent band in positive samples.
I tried increasing the volume of antigen load, since its a crude protein they get clogged. Also, I tried increasing the concentration of separating gel and the length of the gel (mini to maxi gels) yet I m not able to pick the particular band by staining. I get a smeared region. I need to excise the particular band and carry out downstream processing but I face difficulty in viewing the band in gel.
Have you tried a negative stain? As Javier mentioned, these stains are easy to use, and are not restricted by the chemistry of your protein of interest. BioRad makes a couple of negative stains that have worked well for me: http://www.bio-rad.com/en-us/product/negative-stainsFollowing
- Gernot Kaber added an answer:How to strengthen/activate signal peptide cleavage during recombinant protein expression in CHOK1 cells?
I’m expressing human recombinant proteins in CHOK1 cells. The expression level is high and the proteins are secreted efficiently to medium. But the molecular weight and N-terminal protein sequencing of one of the proteins suggests that the protein which is present in medium still contains the signal peptide.
Can anyone tell me if there is a method to persuade cells to cleave the signal peptide? I have no idea why they don’t do it naturally. The signal peptide which is used is a native peptide for those proteins.
Thanks in advance,
All the best,
As i understand the current model of co-translational processing and secretion through the ER-Golgi it seems unlikely that secretion would be efficient without cleavage. Could the additional band be a post translational modification? And really only a minor percentage is ucleaved but shows up in the N-terminal sequencing?
Maybe the protein can be secreted through one of the describe non-classical, ER independent pathways. You can try blocking export through the ER-golgi with Brefeldin A and see what happens to the second band.Following