Questions related to Peptides
I am computing Van der Waal interactions in python for a peptide of size 10 residues for various conformations. The total conformations (or the number of PDB files is 300,000). Is it possible to compute only the 1-4 atom distances to compute Van der Waals interactions as the bonded and 1-3 atom distances are irrelevant when it comes to Van der Waal interactions using some python module?
Hi . I was wondering if anyone could help me determine N terminal and C terminal end of a new peptide sequence that I'm looking to synthesise.
1. if DIEFRVLH is the peptide sequence I'm interested in, how can I determine the ends and directionality of these peptides?
2. What programs/softwares are available to study more characterstics of peptide and to determine its drug potential? I have already used ProtParam and Swiss ADME program.
when I docked peptide and protein in the patch dock, it gives me ten best structures. should I work with all these ten structures?
I want to perform a molecular dynamics simulation of a peptide inhibitor in complex with a protein from Leishmania donovani (pdb id: 3t4p). The only protocol I have is a protein-ligand complex simulation in GROMACS with a CHARMM force field package. I want to know if this same package can be used to run the peptide-protein dynamics.
I cannot purify a peptide synthesized by solid-phase peptide synthesis using HPLC or Biotage. I tried both ways, and still, the trace is not pure. What can I do differently to get a pure peptide fraction?
I'm running a nanoLC-MS/MS system (from Thermo Fisher), while for all samples (peptides), we only observe high TIC at 95% buffer B (acetonitrile), nothing was eluted at 2-50% buffer B in 60 min. the TIC chromatogram of samples were similar to the blank one (pure water), although we find different MS spectrum. Whet we have done:
(1) Clean the whole system with 70% buffer B, actually NC pump and loading pump pressures kept quite normal.
(2) Sample injection model is micro-pickup, we use buffer A (2% ACE/water) as the transport liquid.
(3) Previously, during elution, the NC pump flow is from valve 3 (in)-2-5-4 (out) then to the analytic column. The trap column was set between 5 and 2 (flow direction 5-2, sample loading was from 6 (in)-5-2-1 (out, discharge liquid). Therefore, in this way, the sample concentrated in the trap column was reversely washed out during the elution step. Actually, I feel very confused about this, can the trap column be operated at a reverse flow direction?? Therefore,
(4) I changed the flow route during the elution, i.e., 4 (in)-5-2-3(out).
(5) None of above step works, so, I thought the trap column might be damaged, and then replace it with a new one, exactly the same P/N.
Then, again, it does not work.
Regarding the samples, we detected the protein concentration, which were around 2 mg/mL, 2 or 4 ul of injection volume. All the samples were checked using HPLC, most peptides were eluted before 30% buffer B.
What I'm thinking:
(1) The analytical column is damaged? while, as I know, if nothing blocked, such a column should work for a long time, right? If yes, what is the reason for the damage of analytical column?
(2) The trap column should work at a reversed flow direction during the elution??? ( I did not try this for the new trap column). If so, what is the reason behind??
Any tips, comments, helps are welcome, MANY THANKS in advance.
Hi everyone, I am currently in the process of designing a stapled peptide. The sequence itself is very hydrophobic, thats why I am currently measuring an comparing the helicity in TFE/water (1
:1). The unstapled peptide is showing good helicity since the solvent mixture is already inducing/stabilizing the alpha helix. Measuring my stapled peptide now (stapled in i,i+7 position with reported suitable linker) is giving me a lower helicity. Are there cases where stapling a peptide is not increasing the helicity of the peptide compared to the unstapled version? Could this be an effect of the solvent mixture and the hydrophobicity of the peptide?
Hi everyone! I want to do a chemical reaction that involves the sidechains of arginine and lysine.
The desired chemical reaction is a molecule with COOH groups reacting with Amino groups of the amino acid sidechains of lysine and arginine.
Unfortunately, Lysine and arginine have the alpha-amino-groups, which normally are forming the peptide bond. Those will also react with the COOH groups of the molecule...
So my plan is
1) block alpha-amino-groups in lysine and arginine
2) now add the other molecule and do the desired chemical reaction
I am a geneticist, not a biochemist. I would highly appreciate ideas how to chemically react the alpha-amino-groups away (block them). Possibly with acetic acid that reacts with the alpha-amino-group?
I was thinking of going to a specific pH near the isoelectric point, where the alpha-amino-group is not protonated. Thus the alpha-amino-group might react with the molecule, while the amino groups in the sidechain do not??
I am looking to get a peptide synthesized and have gotten many quotes from various companies. One company, WatsonBio, provided a very appealing quote, so it seems too good to be true. I am trying to do some recon on the company, but I thought I'd reach out to this community to see if anyone has worked with them before.
Thank you in advance!
Could you please advise me on what would be a typical composition of neopeptone?
I understand that it provides vitamins, nucleotides, minerals, and peptides, but is it possible to get more detailed information?
Thank you very much in advance,
Im willing to do peptide sequencing of my sample using edmand degradation method for which i need to characterize and purify the peptide sample,I have access only to 2D gel electrophoresis and HPLC ,will there be diffference in the results or purity of the sample using these techniques when comparing with mass spectrometry analysis
I'm looking for the best/most suitable code of column /resin to use in chromatography steps for separation of freeze-dried peptides in the ÄKTA pure. The manufacturers brochure has about 5 to 6 options, but I could not find what is the best for may case. Someone has experience int this case and this device? The fractions were ultafiltraded in MWCO raging from 10 to 3 kDa. and kept freeze dried in -80 °C
I have encountered some problem when doing md for protein-peptide. Gromacs cannot recognized the peptide as the peptide also composed with benzoyl group. Any suggestions for my problem?
I'm performing my Master degree in biochemistry and I had to to find a way to map the interaction between 2 proteins (so that it will be then possible to design a specific peptide inhibitor to disrupt this bond), but I do not know how to proceed.
Does somebody know an experimental approach to map a binding site between two proteins precisely ?
Thank you very much in advance for your help!
I am looking for a LC-MS spike-in dataset with :
- two or more classes.
- at least a hundred of samples.
- a list of the spiked peptides (mz and RT).
I found a dataset in Leepika Tuli et al, 2012 (https://www.researchgate.net/publication/221865421_Using_a_spike-in_experiment_to_evaluate_analysis_of_LC-MS_data) which corresponds perfectly to what I am looking for, but it contains only 10 samples.
in my project i want to investigate the proteolytic cleavage of a specific peptid through skin bacteria. For that i firstly want to collect the proteases of the bacteria and measure concetration with the Bradford-Assay before i incubate them with the peptid of interest.
My problem is that i cant really get the bacteria (mainly Staphylococcus) to secrete sufficient amount of the protease.
I tried to incubate the baterial pellet for various duration after suspending it in PBS (pH = 7,6). The amounts which i got from where all pretty low ranging from 0,23 mg/ml - 0,028mg/ml.
Does anyone have any idea how to get larger amounts.
Thank you for your time and help.
We take a na-caseinat sample and digest it for 15 min with trypsin. On SDS-gel (stained with Coomassie) we can nicely see the fading of the main band.
When using comparable samples for bradford assay, we can also see the digestion of the protein. My question is now, why do we see this, as the content of aminoacids (in the form of large protein or smaller peptides) stays the same and bradford stains does not peptide bonds?! Thanks.
I am trying to complex small peptides (~7000-9000 MW) with plasmids (roughly 6000 bp) to condense the later down to form particles. Recently, I've run into an issue when trying to concentrate my materials. If I use a higher concentration of plasmid and scale the peptide to it (using N/P ratios) a precipitate will form in solution after mixing. The solutions do contain some salt in the form of PBS and the precipitate does lessen when the salt is removed. Does anyone have any tricks for getting these materials to play nicely with each other and stay in solution? Thanks!
The Dyna-bead (M-280) protocol calls for a PBS with 0.05% Tween-20 solution to be used to wash and elute mAb labeled beads. Ultimately these beads will be used to enrich peptides out of samples for detection by mass spectrometry. Will the Tween contaminate my peptide samples and inhibit ionization? Is it necessary to use the detergent or can a different one be used?
Does anyone know if oxytocin can be measured in body fluids (specifically saliva) using Fourier-transform infrared spectroscopy?
Triplicate peptide samples were dissolved in human serum at a concentration of 100 microM and incubated at temperature of 37 C. Then, the aliquots of each peptide were taken out at 0, 12, 24, 36, and 48 h. Each aliquot was quenched with 6 M urea and incubated for 10 min at 4 C. Then, 20% TFA was used to precipitate serum proteins for an extra 10 min at 4 C. All aliquots were centrifuged at 14,000 g for 15 min, and the supernatant was analyzed by RP-UPLC using a linear gradient of 5–30% buffer B for 5 min.
I am attempting to express a Plasmodium gene which is extremely AT rich with several repetitions of the same. The peptide I am attempting to express is 506 amino acids long.It has 2 distinct overlapping domains(which i am trying to express together). However I have tried Ecoli BL21(24*C for 12hrs) and Arctic strains(12*C for 24hrs) but I have not been able to generate it. It has a C terminal 6x his tag and not being picked up by western blot. I am using pet24b vector.
I am new at these procedures and any help or suggestion from the community to shed light into this would be of immense help!
We performed a peptide-Doxorubicin conjugation reaction. The mechanism of the reaction's conjugation is based on solid-phase peptide synthesis.
In a nutshell, we combined DOX, HATU, DIPEA, and FRRG [the peptide] in DMF to produce DOX-FRRG and a few other small molecules were also produced as by-products . Preparative RP-HPLC is the conventional method used for purifying frrg-dox. I'm curious whether there are any other methods for purifying the solution, particularly cheaper ones.
I'm planning on running my peptide samples on a high-resolution LC-MS instrument. I'm going to use a ZipTip C18 tip for the extraction of peptides and desalting of the sample. However, I'll be sending these samples overseas and it might take 2-3 days for them to reach their destination. I can potentially keep them in dry ice throughout their delivery, but it is very costly and we had few issues before where the dry ice evaporated until it reached the destination.
If I free-dry my peptide samples, do you think they are going to be stable for couple of days? Considering there won't be any humidity where the enzymes can work on the peptides, but I just wanted to get the opinion of people who has lots of proteomics experience.
I am synthesising the tetrapeptide, GPOG, following Fmoc strategy in the Solid Phase Peptide Synthesis (SPPS). After the final cleavage from the resin, I evaporate the cleavage mixture (TFA, m-cresol, m-thiocresol, water) in a rotary evaporator. Then I add cold diethyl ether to the evaporated peptide solution to precipitate the white peptide powder.
I then centrifuge the mixture (white peptide powder dispersed in diethyl ether) at 8000 rpm for 10 min at 20 C. In this process, the peptide powder changed to a light brown sticky mass. The yield of this mass is very less in comparison to the amount of white powder that I had first got by adding cold diethyl ether.
Can anyone please provide any explanation and solution for this?
I'm about to construct some peptide sequences ending in 2-azidoacetic acid. In the literature there are several papers with more or less standard procedures with this compound. For example:
Yu, M. et al. Inorg. Chem. 50, 12823–12835 (2011)
Lundquist IV, J. T. et al. Org. Lett. 3, 781–783 (2001)
However, none of those papers utilise 2-Cl-chlorotrityl resin. All use Wang. My question is WHY? Is there some reason that you cannot couple an azido-AA on a peptide using 2-Cl-Trt resin???
Thanks ahead for comments.
I have a neutral peptides that is insoluble in pH 7.4 solvent. I need solution to be pH 7.4 for the subsequent reaction. I have tried DMSO to help dissolve the peptide, but it would be precipitate if I regulated the pH to 7.4. What can I do to try solving this problem?
I'm trying to build outer membrane of e. coli with small peptides above it, with CHARMM-gui. Peptides should be positioned above the membrane, but charmm-gui creates space for them between LPS (lipopolisacharides) pushing LPS-s to the edges of simulation box.
If I use only one peptide, it is not problem, empty spaces is not big, and charmm-gui sucessfully finishes job. But when using more peptides, empty space created underneath them is large, so LPS-s are pushed to the edges, tightly packed, which obaviously causes problem for charmm-gui and gives error:
"Charmm terminates abnormally. Please check the output or report this failure to the CHARMM-GUI developers. The bilayer generation is stopped to prevent an infinite loop. Please refresh the browser to restart the bilayer generation with different random seed."
I tried several times, with the same outcome. It looks like membrane builder thinks that peptides are "inserted" in LPS, so it leaves empty space for them in between LPS-s.
I report error to charmm-gui developers, and got short answer: "View “step3_packing.pdb”. "
I looked, and see nothing strange, nothing to lead me to solve the problem.
Any idea where to look, or how to solve problem?
Third picture is outer membrane without peptides
I am carrying His-Pull down Assays with purified peptides to investigate binding. I repeat the same protocol meticulously, prepare fresh imidazole every time I carry out a GST-PD, have BSA (0.5mg/ml) in the binding buffer, etc. I do have positive results so I know the assay works and that there is binding but repeating the assay gives me some or the other error such as non-specific binding with the negative controls. So, I really want to know if there's anything I can do to rescue the irreproducibility?
Also, I would like to know the usual method of normalizing the GST-PD data for binding.
We are currently working on the characterization of a complex protein mixture by MS/MS and when searching in the sequence database (transcriptome) with PEAKS X+ software. He does not seem to be able to assign peptides for sequences containing unknown amino acids (noted "X" in the sequence) neither with "*" were the software simply remove those sequence from the database.
How can we solve this problem, which annotation for unknown amino acid is required by PEAKS software ?
Thank in advance if someone can help us.
I have performed docking of a peptide epitope with MHC complex using Galaxypepdock software and have obtained 9 different models along with protein structure similarity (TM score) ranging from 0.970 to 0.995, Interaction similarity score (220-250). How to go about the analysis as I am not able to see my peptide docked with the protein structure?
I am very new to the FTIR technique and recently stumble upon an observation that I cannot explain.
So when I analyzed the native proteins, the spectra showed no transmittance in the Amide A range. But when the protein was hydrolyzed first prior to measurement, it showed a peak in Amide A range.
From reading several books and publications, I understand that Amide A corresponds to the bond between N and H in peptides. So why does it require hydrolysis of a protein into peptides in order for this peak to show?
Even though I have used very little concentration of p-NPA, saturation has not been attained for 5000 seconds. But researchers have done a kinetics study of peptide catalyst with the same concentration of substrate i.e., p-NPA (that I am using for my work), and monitored catalysis for 400 seconds. But, I am unable to attain saturation even within 5000 seconds for 0.2 mM, 0.4 mM, 0.6 mM, 0.8 mM, 1.0 mM and so on. So, if saturation is not attained, then how to calculate the value of Vmax, Km, Kcat? Can anyone kindly suggest to me, if I am doing anything wrong at any step? It would be really helpful.
I am looking for a tool (online, R, Python, or otherwise) which I can use to highlight peptide sequences on the full protein sequence in a visually nice way for publications and presentations.
Extended description: In several of my bottom-up proteomics research projects, I have identified proteins of interest for a given condition/disease. Often, these proteins are activated/deactivated by cleavage (e.g. the complement system, coagulation system, angiotensinogen, etc.). Therefore, I commonly perform a peptide-centric analysis after the protein centric analysis, to identify changing peptides and then I manually map these to the protein sequence. I am looking for a tool to help me with this; where I can submit the list of peptide sequences and have these visually mapped to the full protein sequence of origin. Ideally, the tool should include known cleavage products (e.g. from UniProt KB).
Any advice is most welcome and thank you for your time.
Tue Bjerg Bennike
I am wondering if my proteins are digested into small peptides, will the protein concentration remain the same if I use BCA or Nano-drop to detect the concentration.
To provide some context:
Once an antigen is processed endogenously or exogenously by the APC's (ex: dendritic cells), it is then presented on MHCI or MHCII as a peptide.
specific naive Tcells have a matching Tcell receptor that is complimentary to the presented peptide. considering the incredible diversity of peptide antigens that can be presented, what governs the production of Tcells having the exact match to the presented peptide?
What is the mechanism of Tcell receptor diversity considering that somatic hypermutation is designated to Bcells?
(picture adapted from: Molecular biology of the cell - Garland science 2008)
I am studying protein corona formation and composition on coated gold nanoparticles. I coat them with polymers and incubate in FCS or plasma. I now want to compare between differently coated particles what relative amounts of proteins are bound. I don't need nanograms, it's enough to say "Particle A binds twice as much BSA/protein X as particle B".
I use MaxQuant to process my Orbitrap data and I can see what proteins bind in general. But how can I quantify and compare? I read about the Top 3 approach. Did I understand it correctly? I should add a known amount of a reference protein (they use BSA mostly, but I would need to use something else, because BSA is a protein of interest. Any suggestions?) and then they use three peptides of a protein that show the highest intensities, add the intensities up and compare it to the reference?
How do I know which peptides belong to which protein? Do I need to do that all manually in an Excel list of like 300 proteins? What kind of reference protein should I use? Does every protein have at least 3 peptides? Would I need to look at unique peptides only? It's quite hard to find a detailed explaination on this.
Thank you very much for your support!
I am doing an MD simulation of a peptide with 3 unnatural amino acids and a linker using Gromacs. I have already generated the itp files for unnatural amino acids and the linker using ATB web server.
Can anyone suggest to me how to generate topology(.itp and .top) for the whole peptide chain ?
I am trying to purify a hydrophobic peptide (A-beta 42) that is 4kDa in size. So, I am using reverse-phase chromatography equipped with a C18 column (1.7 um). I am expressing the peptide in BL21 cells and I have confirmed that the induction was successful by running the cell lysate on a tricine system SDS gel. However, I don't see any peaks on my chromatogram when I run the total cell lysate. I tried to run a commercially available sample of the same peptide dissolved in water and I can clearly see the peak on the chromatogram. I am not sure why is that... so I reran the same sample on a gel, and I can still see my cell lysate present in the sample. Is there any suggestion to troubleshoot this... I am thinking to add His tag and use the affinity column but I see that most papers use C18 columns...
I am attaching the chromatograms of the sample and the standard for reference.
I want to fuse the P2A DNA sequence between 2 genes of interest. Can the P2A peptide have the ability to self excise when the polpeptide chain is synthesized in prokaryotes?
Most examples for this procedure are typically done in eukaryotic system. I am not sure if this ability of self excision would happen in prokaryotes? Maybe someone has/have experience(d) this phenomenon.
Any ideas are most weclome, if any :)
I use 3 peptides, I dilute them in Dh20.
When I put in the media after 3 hours my peptides precipitate in the cell culture media.
When I remove the serum is possible to avoid it. But I need to put serum in my cells.
Hi. Is it always invalid to get the docking result obtained using structures that have not undergone energy minimisation? What would be the factors considered in this decision?
YASARA Structure allows the users to build the missing residues in protein crystal structures using BuildLoop and OptimiseLoop command. If the missing residues are modelled using BuildLoop and OptimiseLoop command only without energy minimisation, is the subsequent docking result valuable and valid. In other word, should the data generated this way should be discarded as invalid?
Alternatively, is it meaningful to retain the docking results generated using the same modelled structures with and without energy minimisation, and hypothesise that the best-ranked peptides obtained in both methods can be one of the best peptide inhibitors, which can only be confirmed experimentally?
I have been running a förster resonans emission transfer (FRET) assay, using a dnp/mca conjugated peptide probe for my protease of interest. However I have problems with the assay; sometimes the assay works fine and I get really nice and clear signals, but sometimes the signal from both the FRET-peptide control (w/o protease) and buffer only control (i.e. only the buffer) declines dramatically during the timecourse of the experiment, and the signal from the protease is much weaker than normal (after correction for the declining background...).
Peptide: dnp-NH-GTQVKLIGHR-CO-mca. Detected fluorometrically at 392 nm using an excitation wavelength of 325 nm according to litterature. 20ug/ml final concentration
Buffer: 50mM Hepes (pH 7.4), 200 mM NaCl, and 2 mM DTT
Enzyme: Ctss, 0.1 ug
Plate and buffer pre-heated to 37 degrees.
Read in Spectramax M2e using fluorescence kinetik read with 325 exitation, 392 emission and 325 auto-cut off filter.
Any idea of what might be going wrong?
Im using a known peptide sequence of gene of interest from phytozomw to tblastn with SRA data in NCBI. My sequence results doesnt cover the query and arent highly similar.
After downloading the fasta sequence of all the aligned hits I blastn it but my plant of interest is not in the results.
What can I do to find the desired sequence for my plant?
I have a peptide with non standard amino acids and their parameters are in CHARMM36 format (.prm and .rtf). moreover, the parameter file contains a PATCH to add a thioether bond.
Is there a way (other than CHARMM-GUI, because it cant apply a custom PATCH in the uploaded files) to prepare the files manually for GROMACS while applying the PATCH?
Hi fellow RGers, in my experiments, I'm trying to express a peptide and I'm curious about the general stability of the expressed peptides. Is there a rule of thumb concerning peptide stability in bacteria? I have heard from other colleagues that peptides are generally more stable when it's secreted vs in the cytosol. However, I cannot find any papers confirming this. Have anyone came across any data in literature that supports it? Thank you and have a nice day.
Hello, I am interested in working with dyphenilalanine but I am having a hard time identifying the correct item on Merck or other site to place an order. Specifically:
- Small diphenylalanine, NH2−Phe−Phe−COOH (FF), peptide monomers from the work Strong Piezoelectricity in Bioinspired Peptide Nanotubes, Kholkin et al 2010, I have not been able to identify this one anywhere
- For L-Phenilalanyne from Simultaneous Synthesis and Self-assembly of Cyclic Diphenylalanine at Hydrothermal Condition, Togashi et al 2006, it this it https://www.sigmaaldrich.com/PT/en/product/aldrich/s452777 ?
I would appreciate it if anyone could guide me towards the correct peptide as well as help me understand how to correctly identify similar things in the future.
Thank you in advance.
my peptide molecule having free acid and gunidine group ,How can we crystallize and what are the best solvents for crystallization ?
Hi. I am currently designing peptide ligands.
I noticed that DUD-E (Directory of Useful Decoys, Enhanced) can be used to generate decoys for small molecules. I wonder if DUD-E is equally useful in generating decoys for the peptide ligand.
Is there other way of decoy generation for peptides?
Hello, does anybody use the OPA reagent to quantify peptides by fluorescence?
I'm struggling with the conditions, specially with the ratio between OPA/peptide to do the calibration curve. I follow the Thermo Fisher protocol but it's not working. Thanks in advance
Our primary hippocampal cells are simply too happy in the presence of Abeta1-42. While the neurites are definitely destroyed in the presence of micromolar quantities of oligomers (prepared with the standard method, ie resuspension of denatured film into DMSO to 5 mM, sonication, then dilution/incubation to 100 uM in cold Ham's/DMEM overnight at 4C, and spinning before application), 4 days after receiving micromolar concentrations of oligomers, the cells still give the same viability response using WST-1 or CCK8. We've tried two peptide sources. Has anyone solved this problem?
I am currently working on expressing and purifying recombinant strep-tagged AMPs peptides in plants. My usual strategy is after whole protein purification, I do strep-resin based strep-tagged peptide purification, then perform on-resin digestion with the protease followed by His-Trap purification to get rid of my His-tagged protease, then size-exclusion chromatography with the column for resolving peptides lesser than 3 KDa. After that I finally perform RP-HPLC using C18 columns to finally purify my peptide and concentrate it.
I have heard about flash chromatography for peptides and I was wondering for purification, I can do Strep-Resin based peptide purification followed by on resin digestion and then go straight for RP-HPLC bypassing the His-trap and Size-exclusion chromatography step? Is it a feasible option to do this as I am going to purify 1 gm of peptide and I am trying to find a good and less time consuming strategies but to get good quantity and quality peptides.
Any suggestions on this would be most welcome.
Are structure-based peptide drug design and ligand-based peptide drug design mutually exclusive? If we have the crystal structure of the protease and design the peptide inhibitor based on the sequence of its ligand (protease cleavage site), is it a structure-based peptide drug design and ligand-based peptide drug design or a hybrid of both? Thank you.
I should label peptides with cyanine dyes, which are very unstable molecules. Do you have any advice to handle and purify the product?
I'm trying to measure the concentration of my pure peptides with Thermo Scientific NanoDrop One. The peptide sequence is ELAGIGILTV, which does not have the key amino acids required for measuring proteins at A280, so I measured the concentration at A205nm instead. However, based on the readout from NanoDrop, the peptide absorbs higher at A280, and the concentration measured at A205nm is also unreasonably low.
Does anyone know the general rule of choosing the proper program to measure peptide concentration?
i have a peptide mixture ( bought from a company) and i would like to purify it
The sequence is Acetyle-KCYCSIGGGTRQCYATLAECR-NH2
i expected to have 2 disulphide bonds in this peptide
i would like to purify it using C8 column(9.4*250mm).
I dissolve the sample in 90% Water and 10% ACN in 1 millilitre and filter it .
Mobile phase ACN with 0.1 TFA
Water with 0.1 TFA
Flow rate :2 ml/min
Injected volume : 20 microliter
detected at 3 wavelength : 210nm -220nm -280 nm
Time Water ACN
0 87% 13%
5 87% 13%
35 40% 60%
45 40% 60%
50 87% 13%
i collected all the fractions and diluted it 10 times for ESI MS .
but i do not see the peptide with all peaks i collect in ESI MS (find attached of chromatogram i got ) . Any ideas please
Hello, Research Community (RA),
I am looking to use a Pepsite2 bioinformatic tool for computing the binding residues of receptors (ACE) when used with food peptides. Please let me know if the enzyme model is required to prepare for that. I mean to say this tool asks for the PDB code and chain id of an enzyme that can be easily found from the protein data bank. But is there any treatment needed for this PDB structure like removal of a lisinopril inhibitor or water molecules?
As you probably know the "Tryptone" used as component of LB media and alike are the product of digestion of casein. Similarly Cas-Amino acids are the product of acidic hydrolysis of casein.
Given the above, and knowing that casein is highly phosphorylated I am trying to understand if the peptides in Tryptone and the amino acids in "Cas-Amino acids" are also phosphorylated.