Questions related to Proteins
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,
what is the best method to extract proteins from serum and tissue sample?
what is the best method to identify the signature protein/peptide between serum and tissue samples by using mass spectrometry?
I want to stain muscle cells for presence of glycoproteins. For this I am trying to do lectin staining. However, right now I do not have a working protocol and I have come across 2 lectins in literature that can bind N-glycans (glycoproteins i am interested in). I want to know which of these two could be a better choice? And if anyone has done lectin staining before and can suggest me a protocol or a product? I am ok to use a fluorescence conjugate lectin or a regular one with the use of secondary antibody biding.
I'm purifying some mutants of the protein I study. The wild type protein exists as a monomer and is 28kDa.
I have two mutants (same protein, same number of amino acids but with 8 amino acid substitutions at defined positions), one of the mutants (mutant 1) analysed using size exclusion chromatography with multi-angle static light scattering (SEC-MALS) and its MW was shown to be 33kDa and has an oligomerization state of 1.2. The other mutant, mutant 2, also measured by SEC-MALS was 59kDa with an oligomerization state of 2.2.
For the wild type to measure the concentration I've just been using the MW (28kDa) and extinction coefficient (calculated by entering the sequence into online software ProtParam) and using a NanoDrop measuring absorbance at 280. This gives the concentration in mg/ml which I then convert to molar concentration.
For the mutants I want to measure their concentration the same way - measuring A280 on the NanoDrop using the mutants MW and extinction coefficient and calculating molarity from mg/ml. I'm not sure if this is an obvious/stupid question but what MW weight and extinction coefficient would you use for the mutants on the NanoDrop? E.g. For example mutant 2 molecular weight (MW) of the protein based on its amino acids (AA) composition is predicted to be 28kDa, but SEC-MALS shows it is 59kDa as the protein forms an oligomer.
My instant is to use 59kDa and the computed extinction coefficient predicted from the AA composition - is this correct?
Thanks in advance!
Hello, I hope analytical chemistry people or biochemistry major fellows could help me. Kindly advise on how to prepare the following denaturation buffer: [2% SDS, 1 M β-mercaptoethanol (β-ME)]?
The buffer is expected to be used with Endoglycosidase H enzyme, extracted from Streptomyces plicatus, CAS 52769-51-4 | 324717.
Earlier I had used Serva IPG Strips in which it was easy to decipher the gel side of the strips but recently we shifted to Bio Rad and here we are encountering this problem. Your valuable suggestions are required.
I have extensively searched google scholar but I am struggling to find any groups who have previously used Rosetta to conduct ab-initio structure modelling of single-pass or membrane anchored proteins and I'm specifically not talking about homology modelling just ab-initio.
Please let me know if you have read any papers or know anyone who has done this,
2nd year PhD student at University of Liverpool.
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!
My stock protein concentration is 2.13mg/ml. I have to have a final concentration of 0.83 nanomoles and my final volume for the protein would be 50 microlitres. The molecular weight of the protein is 55kDa. I did a few calculations based on this and I got a volume of protein to add to my experiment, but it is impossible to measure it using pipettes. Do I have to dilute the protein more to get to a measurable volume for my final concentration?
Does anyone know an accurate technique to quantify intracellular protein concentration besides ELISA? I need a reliable and highly quantitative method to measure protein concentration to gather data from cell culture experiments and incorporate them into my model of protein-protein interactions.
Hi, I need to check microbial growth on wastes rich in proteins that tends to ferment pretty quickly. To get an idea of what and how much it's growing, I thought about using Plate Count agar, YS or YM for screening for yeasts, maybe Violet Red Bile for gram negative. What can I use to check protease activity? Since it's a mixed colture, I cannot use super selective media.
Could this course of action work for a totally uknown mixed colture? The goal is to identify the microrganisms responsible of spoilage and fermentation.
Thanks in advance for the help.
I need to prove that my protein is localized on the cell-surface but not inside the cell in soluble form.
How can I make it?
There is an opinion to wash cells, to label all cell-surface proteins with NHS-biotin, and to collect labelled fraction of cell-surface proteins with streptavidin magnetic beads. After that I can make western blot with specific Ab.
Maybe there is another way?
Protein and cell lines are human.
Does anyone have experience with Casein kinase 1, I am working with its homolog in yeast (Yck). Is it regulated or just a constitutively active protein?
I checked its expression levels, they don't change in response to different carbon source. But I am trying to determine if its activity is modulated by phosphorylation or not.
I am trying to detect the secretion of certain tagged soluble protein following transfection in 293T cells. These proteins are epitope tagged and can be visualized via Western Blotting. Therefore, I plan on using the supernatant to run a western blot to determine whether these proteins are secreted.
I am torn between TCA precipitation, ultracentrifugation, or just taking the supernatant and running it on a protein gel.
For context, these transfections are done in 6 well plates and in Optimem.
Does anyone have any suggestions as what the best way to detect these secreted proteins is?
Recently I run a stain-free gel (Bio-Rad) to separate my protein of interest and do total protein normalization. After separation, I imaged the gel on ChemiDoc. Unfortunately, I overexposed the gel. Please could you advise on how to do total protein normalization in the setting of overexposed gel? I have attached the image of the gel for your reference.
The ones we have tried are either against the signalling peptide or do not recognise small levels of expression. Or show only one band in Western analyses.
I've conjugated a PEG4-maleimide (MW 613.66) onto an antibody fragment (52 kDa), but I'm trying to figure out how this affects A260/280 measurements on a Nanodrop afterwards. After conjugation and removal of unconjugated material on a PD-10 column, I measure the concentration of the eluted aliquots but the measurements seem really off, and the A260/A280 ratio is nowhere close to 0.5. I blanked the Nanodrop with the PD-10 elution buffer. Seems strange that such a small molecule could affect the readings that much. Does anyone have experience with PEG and if/how it could affect concentration measurements after conjugation?
So I have a peptide with ~14 kDa molecular weight.
It contains one Cysteine in the middle of the sequence.
I tried to desulfurize this Cysteine.
But how do I know if it worked or not? I use UPLC with ESI-MS, H2O/ ACN.
Since the mass diference is very small, I suspect that mass spectra of product will not be much different from the starting material.
Retention time should also be very similar, at least I can't see the second peak. Or perhaps conversion is 100% and there won't be a second peak, I don't know :)
I have a system with two proteins. I want to compute the energies of these proteins separately. So, I create two separate groups, say, Prot-1 and Prot-2, and add these to my rerun mdp file as energygrps. However, when I perform gmx energy on the rerun.edr file, any energy term associated with Prot-2 returns 0. Prot-1 returns energy values for the total protein system (i.e. for both proteins). I am not sure what is going wrong here. Any suggestion is highly appreciated.
I have done SMD of protein at applying constant velocity using NAMD software and CHARMM ff. Since, this is my first time in performing Steered MD, I am not sure as to should I do umbrella sampling alongwith SMD? Are the results of SMD without any umbrella sampling significant ? It would be helpful if I could get some references as well.
Andree Hubber (2004) described effector-coding genes with signal aa for both T3SS and T4SS.
Kindly discuss your ideas and viewpoints on the origin of life and the RNA world hypothesis.
What are the contradictory views on why researchers are still unsure about the origin of life through RNA or such analogous molecular intermediate pre-cursors preceding its existence?
"The general notion of an “RNA World” is that, in the early development of life on the Earth, genetic continuity was assured by the replication of RNA and genetically encoded proteins were not involved as catalysts. There is now strong evidence indicating that an RNA World did indeed exist before DNA- and protein-based life. However, arguments regarding whether life on Earth began with RNA are more tenuous. It might be imagined that all of the components of RNA were available in some prebiotic pool and that these components assembled into replicating, evolving polynucleotides without the prior existence of any evolved macromolecules. A thorough consideration of this “RNA-first” view of the origin of life must reconcile concerns regarding the intractable mixtures that are obtained in experiments designed to simulate the chemistry of the primitive Earth. Perhaps these concerns will eventually be resolved, and recent experimental findings provide some reason for optimism. However, the problem of the origin of the RNA World is far from being solved, and it is fruitful to consider the alternative possibility that RNA was preceded by some other replicating, evolving molecule, just as DNA and proteins were preceded by RNA." - Robertson and Joyce
[This is as per the explanation by Michael P Robertson and Gerald F Joyce in the article: "The origins of the RNA world." published in the Cold Spring Harb. Perspect. Biol. 4, a003608 (2012).]
The scientific community must resolve this contradicting conjecture through rational discussion and debate backed by strong experimental evidence on what must be the pre-cursor molecule to the Origin of Life if it is not RNA!
I am just a new student doing research about carbonic anhydrase, I found so many papers classify their CA protein into alpha, beta, and gamma classes, but they didn't say the exact method they used to classify them. I guess when they once found a new CA, they did sequence alignment with different classes CA sequence. But I am not sure which CA sequence they aligned with, and what the signature of the alignment that makes the new CA just this class type. Could you give me some hints or papers that illustrate the classification method?
Long story short, I need to degrade 30ug of RNA and i need to do it at 4C. I want to use only as much RNAse A as necessary.
So if performing the reaction at 4C, how long of incubation time and how much RNAse A would i need to degrade 30ug of RNA?
For example, would 1ug of RNAse A(20ug/ml conc.) for 15min at 4C be enough?
I would like to perform, among other things, electrophoresis with preserved proteins (not denatured).
To do so, I need to first extract proteins with a "soft" buffer.
The best would have been M-TER, but I was wondering if B-TER would do the job (because that's what I have in the lab).
(In order to increase the yield of proteins extraction, I planned to perform griding through Precellys and sonication with a probe.)
Thank you for your help.
I'm using Proteome Discoverer 2.1 to analyze my peptide spectrum from Q-Exactive. But after the process complete, I get two error warnings in consensus workflow. In Peptide Validator node, it says "no decoy search was performed for the following search nodes:-SequestHT(A2) in workflow Workflow.FDR+fixed threshold validation is used instead". Meanwhile in Protein FDR Validator node, the error warning says "Cannot validate proteins for group SequestHT, because at least one search node has no decoy PSMs."
Do you have any idea what is the problem?
Thank you :)
I was wondering if anyone knows-
which statistical test I should use in order to find whether a sample is an outlier in my proteomic data? It's obvious when looking at the PCA, but how should one calculate this?
Many thanks for your help!
I am looking for a specific method I read years ago,, but cant find it right now,,
it was basically incubating the protein with some concentration of non-polar molecules,, have them attach to the protein hydrophobic patches, and i think they got a crystal structure or something,,
Any chance anyone recalls something similar???
I am studying proteins bound to large (~500 nm) nanoparticles. I just want to know if there are any special recommendations for sample prep, to get a good CD signal, and whether near UV-CD or far UV-CD is better suited for this, considering the scattering effects I have to overcome.
Thanks in advance!
Our old but not to old DynaPro Plate reader I does not work anymore and Wyatt does not want to investigate the problem as the instrument is 10 years old. We have the money to pay them but they really do not want to loose time on it...
We would like to know if some of you know DLS instruments that are compatible with the measurement of several conditions (at least 30 conditions) in parallel. Of course the goal is to find a company that is able to do a maintenance.
The bacterium that I am working with is a motile bacteria. I constructed over-expression plasmid constructs into the bacterium. I want to check is there any effect of overexpression of the proteins on bacterial motility. Can anyone suggest some reference papers?
the assay was done using TMBZ solution which was made by dissolving 0.01 g of TMBZ in 5 ml of methanol and 15 ml of 0.25 M sodium acetate buffer (mixing sodium acetate and glacial acetic acid) (pH 5.0) and finally adding PBS and 3% Hydrogen peroxide.
I am about to define an experiment where we want to investigate 10 - 20 de novo small proteins. We are mainly interested in affinity but also want to show that proteins are folding properly. For that we are thinking about using circular dichroism. I am having seconds thoughts though if this is the right method in the long run. When it comes to publishing, I have the gut feeling that reviewers might ask for a crystal structure of the protein or even the complex. I am working on getting an impression myself by reading nature and science papers but I would like to get to know your advice and experience concerning the matter. What methods are best suited to give our research credibility that might be expected in high impact journals?
I'm looking for a commercial partner who is able to determine log P (partition coefficient) of the protein with the use of bioinformatics tools. The protein mass is 2,4 kDa, it includes 22 amino acids and three S-S bridges.
Thank you in advance.
I have been working with TNBS(2,4,6-Trinitrobenzenesulfonic acid, proteins detection) for three years and I'm wondering if TNBS has any expiration date?
I couldn't find anything on the box or Gbioscience's website which I purchased from.
I have doubts my TNBS doesn't work well anymore.
All extant protein alignment algorithms that I am aware of perform poorly for complex proteins that have many possible isoforms. There are better ways of making these assignments. Do you know of a new approach?
I have the problem in endogenous nuclease contamination in my protein preparation purified from E. coli BL21(DE3). I would like to ask your experts in avoiding or removing such contamination. Can you suggest me the protocols or an alternative E. coli host strain with tagged-nucleases?
Thank you in advance.
I would like to find whether two proteins have a common interaction path in the graph using the STRING database. There is software that is doing it?
Demographers estimate that by 2050, the number of people on Earth will reach 10 billion. With such a number of people, the agricultural economy, logistics of food supplies and people's eating habits will have to change. It is likely that economics will force these processes, which will result in the transition of the majority of humanity to nutrition mainly based on vegetable and vegetarian diets. Meat production is many times more expensive than the production of cereals, fruits and vegetables. In addition, according to scientific research and the theory of futurologists, the production of traditional meat, e.g. pork and beef, may be replaced by the production of protein from insect breeding. Research shows that there are more proteins in the bodies of insects than in traditional meat dishes. In addition, the logistics of food supplies, agri-food products will have to improve. Systems for matching agricultural and reptile production to the current needs of the industry and the nutritional needs of people will be improved so as to reduce the scale of food wastage. The biggest threat to the implementation of this plan may be unexpected atmospheric phenomena, natural disasters, droughts, hurricanes, tropical heat in the areas in which agricultural crops have been cultivated so far. In addition, industrial exploitation of arable land and climate change causes soil depletion and the disintegration of areas suitable for agricultural production. Therefore, it will be necessary to continue the technological progress in the production of crops, in biotechnology, in the creation of new plant varieties resistant to pests and adverse climatic changes.
Please, answer, comments. I invite you to the discussion.
I have 3 mutant proteins I have designed, cloned, sequenced, expressed and purified. Now I sent the proteins off for mass spec analysis just a tryptic digest as this was predicted to generate fragments adequate to confirm mutations. I got my results back today and have found that 2 of the 3 have wild-type residues instead of mutant. I am confused as the DNA sequencing following cloning was correct how can my protein have reverted back? I use the term native loosely as the native in this case was just the mutant I started my project with. Any ideas are greatly appreciated!
I want to preface this that I am a PhD student working on a bacterial genetics project and am in no way an expert in protein structures - I'm just messing around with this to see if it can provide any interesting insights that I may have missed just looking at sequence and homologs! Anyway, I am working with a protein that has no defined crystal structure, but we know which ligand it primarily binds. I was able to generate some predicted structures using I-TASSER, but since the ligand I'm interested in is not in the ligand-protein binding database it didn't predict any binding sites for it (though it did find how related ligands would bind which is great!). Does anyone know of (free) software in which you can provide a protein sequence/structure and a ligand name/structure and it would predict how they would likely bind? It seems like something that should exist, but I can't find it! Thanks :)
Right now, I am working with protein, which we obtained by lysating cells in a protease inhibitor mix with RIPA-Buffer. In the beginning we lysated 10^6 cells in a 0.5ml vessel, but because of slow growth we reduced the amount to 5*10^5 cells without changing any of the protocol. I determined the amount of protein with the Pierce 660nm Assay. Afterwards I performed Western-Blots with 30 µg protein per lane.
Here you see my SDS-Page and in Lanes 3, 7, 8 and 9 I used protein obtained out of 5*10^5 cells, while in Lanes 2, 4, 5, 6 and 10 I used protein obtained out of 10^6 cells. The dye runs at a different speed and even in Total-Protein-Stain you see, that the lanes with 5*10^5 cell-protein contain significantly less protein than the other lanes. The same goes for staining with a antibody.
My question is: Why is there less total protein in the lanes with 5*10^5 cells-protein than in the others, even though there should be the same amount everywhere? Is the Pierce-Assay at fault? Could there be problems with too much protease inhibitor for 5*10^5 cells? And most importantly, can you think of a way to still use the protein samples and quantify it with the 10^6 samples? Because we have 40 reaction vessels full of protein obtained by this method.
Your help would be greatly appreciated!
I may have a luxury problem, but i can see my protein bands before i add the developer. Why is tha?. I follow a protocol that i got from my PI which in essential is the protocol published by Wray et al 1981. I followed it because i wanted to try it and see if i can use it to visualize my DNA protein crosslink.
I followed the protocol to the letter though when i add the pre-stain solution ( mix to the gel i see my protein bands within 5 min. 21ml of 0.36% sodium hydroxide to 2.5 ml of 30% ammonium hydroxide and dropwise added 4 ml of 20% silver nitrate solution to the mix while vortexing and raised volume to 100 ml using distilled water).
Can anyone explain this to me? i have tried searching for answers but the usual problem is that bands are not showing up or background his high.
Thank you so much for helping me.
I know that there are libraries for immune antibody libraries in scFv, Fab, or VHH format, but was looking for a way to buy comercial libraries for affibodies. Please let me know if you know a way.
I have a protein system where a single water molecule can play a role in a ligand stabilization. To check whether this single water molecule is important in binding, I wanted to perform TI (thermodynamics integration) calculations, starting from system with water and annihilate it. My plan was to use AMBER software (PMEMD) for this purpose. I would replace water molecule with dummy atoms during TI. After many trials I finally got my tleap output (parmtop & prmcrd), but while running the script I am getting an error.
My question is: am I doing it right or there is a simpler way than TI to calculate an impact on energy after cutting our water molecule from the system?
I will be grateful for any tips!
I came across papers using only transblot turbo transfer for TGX stain free. We dont have transblot turbo transfer, so am trying to optimize TGX stain free using wet transfer. I activated stain free gels for 45 sec, 2.5 min, 5 min (exposure for Intense) in Chemi Doc imager and transfered in Biorad LF PVDF with Towbin buffer at different transfer condition: 30 volt overnight, 75 volt for 3 hrs. Though transfer looks complete in both the transfer condition as I didnt see total protein bands in gel post transfer but I m getting a very faint total protein bands in membrane after transfer, which I think I cant use for normalization. I am concerned if the wet transfer conditions reduces total protein fluorescence on the membrane. Chemiluminescence image for specific protein in the same blot, however, is very good. So it would be very helpful if anyone could suggest wet transfer conditions for TGX stain free. My proteins of interest are within the range of 15 to 150kD.
If the trans blot turbo is much better than wet transfer (with regard to transfer efficiency and post transfer total protein intensity on blot), we might have to install one in our lab.
I did ITC experiments with BSA and Its difficult to get same results with same concentration of protein as well as binding ligand.
For an experiment i need to stop the initiation of the protein synthesis without stopping the already ongoing synthesis (so cycloheximid is not an option).
I am looking for a physical explanation on the fact that if prokaryotes had some kind of endomembranes it would not have been feasible for them to perform their functions. Any suggestions on this fact.
I would like to do an EMSA on a known promoter binding site (biotinylated oligo) and we suspect that we need 3 proteins to be present in the complex to bind DNA. We will use transfected HEK-293 extracts overxpressing our proteins of interest. What strategy is the best :
1) Transfect HEK cells with one gene each time and have separate nuclear extracts with one of the 3 proteins and then combine them with the oligos before loading on the gel ?
2) Transfect HEK cells with all 3 genes combined and have just one nuclear extract to add to oligos ?
My main concerns are the stability of the complex during nuclear extraction and the capacity of the proteins to interact with each other and with DNA in the binding buffer before gel loading (we will use the LightShift® Chemiluminescent EMSA Kit from ThermoFisher. Thanks for your advice !
Hello, so i am basically doing a time course and seeing the optimal time to solubilise my protein in DDM + CHS. Not really thinking at the time I solubilised my proteins for x hours then when they were done put them in a freezer until all samples were ready to be examined. Would they continue to solubilise at -20C, thus meaning I should restart the experiment?
These concerns MD simulations using Gromacs. Suggestions involving Amber/LAMMPS/etc., aren't going to be of any help.
I'm looking for strategies that might save me on lines of code.
As I am sure you are aware there are several strategies for relaxing a protein in a bilayer. For example, a population approach is placing position restraints on the backbone which are decreased between simulations whilst preserving velocities. Finally, all position restraints are removed and the simulation is left to run unrestrained. This is useful for a sequential simulation->simulation->....->simulation, where the velocities are preserved.
My concern is when I want to run e.g., 4 identical simulations, each with different starting velocities. That way I can cover more phase space and I can calculate a metric of uncertainty when I'm measuring a property. This is very easy to do by hand. Each of the four production runs will use preserved velocities for a separate set of equilibration runs i.e.,
Sim1: eq_run -> eq_run -> eq_run -> production_run
Sim2: eq_run -> eq_run -> eq_run -> production_run
Sim3: eq_run -> eq_run -> eq_run -> production_run
Sim4: eq_run -> eq_run -> eq_run -> production_run
But this becomes very difficult when I have e.g., 100 models to build and run. That's 4*100 + 4*100 + 4*100 + 4*100 simulations with all corresponding files. Ultimately, I am looking for a strategy where I can introduce a bias or short external influence so that each model shares the same equilibration runs before spawning 4 unique production runs but without destroying the velocities preserved from the equilibration simulations i.e.,
Sim1,2,3,4: eq_run -> eq_run -> eq_run -> production_run1/2/3/4
I have a feeling, something like replica-exchange might work? Introducing 4 different temperature that gradually relaxes back to what they should be over a very short period of time just so that the preserved velocities diverge.
Thoughts are most welcome.
I have tried a few times to couple proteins to my MagPlex beads. Each time I reach at the coupling step (in MES pH 5), I loose most of my beads (70-90%), since they do not stick to the magnet anymore, only diping a tip in the supernatant is enough to remove the beads from the magnet.
Does anyone encountered this problem?
Thanks for your answers
I want to test whether one protein has pro-tumorgenesis effect in pancreatic ductal adenocarcinomas (PDACs). I want to over express the protein in PDAC cell line first but I don't know which cell line should I do.
I checked ATCC website, there are many different cell lines. Anyone can share your experience in choosing cell line?
I'm currently using Bradford method for a protein assay. And I am measuring the proteolytic activity in roots of certain plants, as many studies found plants have in situ proteases to aid the nitrogen uptake. To determine the proteolytic activity, I will incubate the roots in a known ovalbumin standard solution. Then, after a period of time, the Bradford Assay will be conducted on the incubated solution to look for a decrease in the concentration. I am stilldetermining what concentration to use. In case you don't know, the dye itself without proteins has a maximum absorbance at 495nm, with proteins, it will have a maximum absorbance at 595nm. The dye also has a protein detecting range of 200 - 1500 μg/mL, which is the case for albumins (often BSA is used, but I was restricted to use ovalbumin). When I tested the ovalbumin solution with a concentration of 1500 μg/mL, no color change could be seen, the peak from spectrophotometric analysis also remained at 495nm. I then tested with a range of ovalbumin solutions, then realising with a concentration at or above 50mg/mL, a color change can be seen. The expected results are, there should be a color change & a peak at 495nm when the ovalbumin concentration is at 1500 μg/mL, but there wasn't, and it also requires a high concentration to detect protein concentration, which is odd for a sensitive assay like the Bradford method. Do you have any insights or ideas of what is causing this problem? My current idea is, there is something abnormal about the dye (this is the product I'm using from Bio-rad), or the ovalbumin powder that I used to make the solution has impurities. The impurities idea might be incorrect though, as the powder I'm using is lab grade, and the impurities shouldn't be too significant to this extent.
I am performing western blots on some old brain tissue and I did a BCA and I basically would need to add 200ul of each protein sample to achieve the right concentration which obviously too much for gel electrophoresis. I think the person who isolated this tissue a few years ago added too much lysis buffer because the sample is clear and usually it is at least a little cloudy. I am trying to figure out the best way to concentrate these protein samples and I am not sure if the lysis buffer will play a role. It looks like the simplest method is TCA or acetone precipitation... Does anyone have any input?
In my sample preparation of protein sample I am using simultaneous TCEP and CAA for a reduction and alkylation in one step.
Using DTT and CAA a simultaneous reduction and alkylation is not possible - the DTT and the CAA will react with each other (right?) and the disulfide bridges are still there.
Now my question - Why DTT and CAA react with each other, but TCEP and CAA not? Why is this simultaneous reduction and alkylation possible? What are the differences?
- TCEP reacts with the free electron pair of the phosphorus
- thiol of DTT gets rid of a proton and the sulfur anion reacts
- both reactions are nucleophillic attacks
I am trying to find kinetic information (such as rate of enzyme production etc.) on the expression of taq polymerase in E-Coli after inserting a recombinant plasmid.
Any help would be appreciated
I'm trying to measure the concentration of a protein I've purified and then labeled with Atto 647 flourescent dye.
The protein has no tryptophans or tyrosines (although does have phenylalanines) so the absorbance is very low at 280nm. I can get around this using the Bradford assay to work out protein concentration when the protein is unlabeled. However I want to label the protein with an Atto 647 flourescent dye, but can not think of a way to calculate the dye to protein ratio as there is no absorbance at 280nm as mentioned. I'm pretty sure Bradford or BCA assays will not be useful either as the solution is blue in colour because of the 647 dye.
Anybody got any tips?
I am trying to perform homology modeling using the swiss-model but I have found that the swiss-model gives the same number of residues as that of the template. Whereas my sequence has more residues than that of the model predicted by the swiss-model. What should I do? I am confused as it does not give me an exact number of residues in the 3D structure.
I am trying to characterize the microstructure (localization of carbohydrates, fats, and proteins) of coconut milk powder using confocal microscopy. From the review of the available literature, I have found that fats and proteins can be non covalently labeled using fluorescent dyes. However, I am having trouble deciding the labeling protocol for the carbohydrate. Two common methods that I have seen in the papers are as follows:
1. Use antibodies or lectins conjugated with fluorophores to label carbohydrates
2. Covalent labeling of carbohydrates with FITC
If I use the first method, the signal from antibodies or lectins may interfere with the signal from protein present in the sample.
In the second method, the FITC (probe used for covalent labeling) also shows an affinity for proteins. So, I am afraid that the use of such a probe may also interfere with the protein signal (from the food sample). Is there any other probe available that I can use?
What would be the best way to label carbohydrates such that I can simultaneously image three components in the food mixture?
Can I dissolve freeze-dried catalase or lysozyme directly into de-ionised water or do I need to use a buffer (separate solutions, not both proteins in one)? I will not be storing the solutions and will be using them straight away.
If I need a buffer what should I use?
What would be the max concentration solution I can prepare?
It is well known that heating can denature proteins. However, what does happen to proteins in the case of short and ultrashort (microsecond or nanosecond scale) heating to extreme temperatures (100-1000 degrees of Celsius) ? Such heating occurs for example when applying ultra/short laser pulses or pulsed intense electric fields.
I am using DESMOND for molecular dynamics on protein LRP6 and its ligand. At stage 4 I am having a problem, "job failed due to backend error". The job has failed multiple times. We took the original pdb id 3s8v that is complex of dkk1 and lrp6, and from that we removed dkk1 and extracted the lrp6 and docked to get the appropriate ligand. But while using DESMOND it shows a backend error. Why is this happening?