Peptides - Science topic

For making images I use the command:

ray 2400, 2400

png FileName, dpi=3000

This should save it without the background.

I try to minimize the extra space by focusing the GUI window on the protein/peptide. Alternatively, you can use an image editor. The simplest is to use PowerPoint, but you can also use SketchBook, Illustrator, Krita, Corel, Canva, etc., and crop out the background with their magic select tool.

I use SketchBook.

Note: You do need the PyMOL license to do this, especially if you want to publish the images.

Thank you Kais Khudhair al Hadrawi for the response. My initial inputs were not correct, I have resolved the issue.

Say you want to color just 2 objects in VMD different colors, it is relatively simple. You can do it in two main ways.

1) The easiest is to go to the VMD main window, to the "Graphics" tab, and then click on the "Representations..." tab.

A window will appear. In this window, you should select the coloring method ColorID from the drop-down menu. The drawing method and other options are down to you.

Now you need to make selections of the proteins. Go into the GRO or PDB file and write down the atom number with which each protein starts and ends. Into the "Graphical Representations" window that previously opened write those numbers into the "Selected Atoms" line like this: index (first atom) to (last atom) and then press enter.

For example: index 0 to 273

Next, you are going to click on the "Create Rep" button in the same window, I should create a new representation by copying the previous one. Change it by writing the atom numbers of the second protein (example: index 274 to 547) and press enter.

Afterward, you just need to change the color ID number by choosing from the drop-down menu the color you like for each protein.

In summary: Graphics -> Representations...-> select "index (first atom) to (last atom) -> ColorID coloring method -> repeat process after creating new representation just with the second proteins atom numbers

2) The more advanced method is to go to "Extensions" in VMD Main -> Tk Console and then paste this into the command prompt and press enter:

mol delrep 0 top

#First protein

mol selection {index 0 to 273}

mol color ColorID 0

mol representation Lines

mol addrep top

# Second protein

mol selection {index 274 to 547}

mol color ColorID 1

mol representation Lines

mol addrep top

Just change the code by adding your atom ID numbers, ColorID numbers, and drawing methods. You can copy-paste the section of code that has "mol selection" to "mol addrep top" for any number of proteins you want to color. I even used Excel to create code to color 200 peptides with different colors automatically.

NOTE: Be careful when selecting atom indices in different files. GRO files count the first atom from 0, PDB files from 1.

Possibly due to increased helicity and compact packing of the protein ligand complex

In general, one of the commonly used storage conditions for azide-labeled proteins or peptides is to keep them in buffer solution at a low temperature, usually at -20°C or even lower, such as -80°C. This helps slow down any potential degradation reactions and maintain the integrity of the labeled molecules. In addition, storing them in the dark can prevent any photochemical reactions that may occur... On the other hand, storing azide-labeled proteins or peptides at low temperatures in a suitable buffer solution and protecting them from light should help maintain their preservation. Its stability over time.

James M Fulcher Thanks for your answer! I agree it is probably not trivial, because you cannot assume that *all* ionisations are with sodium rather than a proton, so it sort of explodes combinatorically at the MS2 level.

Unfortunately, I cannot rerun the samples, because I am trying to apply a new analysis pipeline to existing data on PRIDE, and I have found this phenomenon in other researchers' published data.

I have tried setting up sodiation as a PTM on MaxQuant, and I think it could work, but it increases the analysis time from minutes per sample to days per sample, so might be useful for a proof-of-concept, but if anyone has suggestions for a better solution – or knows of a way to this in MaxQuant already – I would love to hear them!

I have also heard that the pH of phosphate buffer changes when freezing, so it is not preferred for a storage solution with sensitive materials.

If the solution is flash-frozen, the solutes do not have time to separate from the ice, so this method is preferable to slow-freezing.

To desalt peptide mixtures while preserving their hydrophilic and hydrophobic properties, you can use a reversed-phase solid-phase extraction (SPE) method. This method separates peptides based on their hydrophobicity using a hydrophobic stationary phase, while the mobile phase is a water-miscible organic solvent. The method allows for the elution of hydrophilic peptides first, followed by hydrophobic peptides using a stepwise increase in the organic solvent concentration. Alternatively, size exclusion chromatography (SEC) can also be used to separate peptides based on their size and hydrophilicity. SEC separates peptides by size, with larger peptides eluting first, and smaller peptides eluting later. This method can also be used to desalt peptide mixtures without affecting their hydrophilic and hydrophobic properties.

What do you plan to do with your 45mer peptide? Are you hoping it will have catalytic activity?

You don't really "design" an antibody. You grab your peptide or your mRNA and you vaccinate a critter. Its immune system then designs the antibody for you. Yes it really is possible. About $5k to hire a service to do it all for you, just off the sequences.

Designing antibodies involves several approaches aimed at generating antibodies with desired properties, such as specificity, affinity, and functionality. Here are some key methods and strategies used in antibody design:

By combining these methods and strategies, researchers can design and generate antibodies tailored to their specific research needs, facilitating a wide range of applications in basic research, diagnostics, and therapeutics.

I think you can use molecular docking tools like AutoDock or AutoDock Vina to calculate the binding affinity between a protein and a peptide. PyMOL can be used for visualization but not for calculating affinity. I hope this helps!

Okay, no problem.

There are a myriad examples of this sort of ORF structure in eukaryotic viral systems (particularly RNA ones), where it has evolved to diversify/expand coding capacity and/or regulate viral gene expression. At first pass you can predict relative expression levels by examining the Kozak consensus around each AUG codon. We usually consider the -3 and +4 positions to categorise them as strong, middling and weak. If the first AUG is strong, you typically won't get much of the second product. If the first AUG is medium or weak and the second is strong, you'll get lots of the second product. Confounders to look out for; 5'-UTR length and protein stability. AUGs very close to the 5'-cap are often not translated well. Shorter proteins are harder to detect and if they're too short (or the wrong sequence) to fold up into a coherent structure, they'll be turned over rapidly and can be hard to detect unless (or even if) the proteasome is turned off.

After that, it can get properly complicated with all sorts of ribosomal gymnastics depending on the exact sequence of the transcript.

Bugs - need to consider Shine-Dalgarno (sp?) sequences or lack of, I think. But my undergrad lectures on this were many years ago :)

Cheers

Paul

I might be a little concerned about the pH of the solution, since you dissolved the peptide aty high concentration in water with no buffer. Spot a few microliters of the stock onto a pH strip to see if it is strongly acidic, which might affect the stability of the peptide.

Somewhat less complicated than LC_MS I would suggest reversed-phase LC in combination with UV detection at 218 nm employing a proper (thus not to steep!) acetonitril gradient using 0,1% TFA.

Hi,

This is my go-to site for that purpose: https://prospector.ucsf.edu/prospector/cgi-bin/msform.cgi?form=msproduct

Why not look at monoisotopic mass?

The difference in masses obtained can be due to the way the elemental masses are set in the algorithm of the calculator: a difference in digits/rounding can result in large differences in peptide masses, let alone that of complete proteins..!

There is no 'correct' answer, I guess, it's more about the accuracy taken into consideration.

Hope this helps.

okay. Thanks for your reply.

Hi Talia,

I have looked into this particular question in the Methods paper I have published ; please see the link below. I did not use Proteome discoverer - I used MaxQuant and configured each of the TMT channel modification as a variable modification so that I could search my mass spec file to see how many peptides were not labeled.

Let me know if you can access the paper link below, if not I can also just email it to you.

All the best,

Hediye.

Thank you for your suggestions.

Dear Chen Lior

I believe you can try to use the holdup method adapted to cellular context. You can gie a look to Zembo et al. Sci. Adv. 2022 "Native holdup to measure binding affinities from cell extracts". The title alone already evokes all the requests you have.

Best

Hi Ibtissam,

did you defined the mentioned variable modifications in global parameters/protein quantification tab (peptides for quantification)?

Best,

Murat

Zahra Hajihashemi The lipid composition of eukaryotic and procaryotic cells is quite different. Generally, there is an higher proportion of anionic lipids in bacterial membranes than in mammalian membranes. Therefore, to target the bacterial membrane, you can add cationic amino acids on your peptide (lysine, arginine or histidine). What we have seen from our experiments, there must be a right balance between the positive net charge and the hydrophobic moment of the peptide. Increasing the positive net charge increases the performance (up to the a certain point). If the hydrophobic moment is too high, you lose in selectivity towards bacterial cell membranes. In order to improve the stability, you can incorporate non-natural amino acids.

Dear Fareed Sairi ,

Did you characterize the peptide, both with bioinformatics and experimental? I mean:

-Using for example Heliquest (https://heliquest.ipmc.cnrs.fr ) you get a proper idea about the peptide (its hydrophobicity, net charge, etc.). See for classification for example:

-Before checking in your assay, I think it is good to check the peptide in some way. If for example bioinformatics predicts membrane activity you can check quickly by using a monolayer technique or depending on what you have available in the lab check by circular dichroism (with and without lipids), see for example:

If this seems fine, then do realise that sometimes a pretty insoluble peptide (as described in the link above) that requires something like DMSO or TFE is all of a sudden reasonably soluble in aqua dest. as described here:

However, in your medium there will be a buffer containing salt (and a certain pH) and the ionic strength in the medium can cause aggregation (this can be checked with the peptide and the use of for example circular dichroism).

Obviously, something might have gone wrong with the synthesis, or the prediction is simply wrong, and the peptide has no antimicrobial activity but a lot can be checked before one have to draw this conclusion.

Best regards.

Assuming the peptide functions by binding. The target protein could be immobilized on a plate or transferred to a blot. Then the peptide (likely also with fluorescent / chemiluminescent tags) could be applied to bind to the protein for detection.

Try dissolving it in dimethylsulfoxide (DMSO). This is a water-miscible solvent that is generally very useful for this sort of thing, and it is compatible with subsequent biological experiments when diluted to a few % by volume.

Hi Jan Gebauer

I've also had some funny looking volcano plots from PD data. Discussion here: https://www.researchgate.net/post/Artefact_in_volcano_plot

I never got to the root cause, but the funny lines of points lining up were an artefact of normalisation - mind you, the underlying table of PSMs from PD was the original source.

Unfortunately I never got around to identifying why PD gave odd output.

Sorry I can't be more helpful!

Sam

Murat Eravci

Thank you for your kind reply.

I will try searching HLA peptide atlas.

Best regards

Jaekwan.

Dear Zeeshan Yaseen ,

Yes, this is a frequently asked matter. See for suggestions:

or for example https://www.softwaretestinghelp.com/increase-resolution-of-image/ (scroll down towards How to improve the resolution of a photo).

It is possible since I see papers with figures of PepDraw generated results, see for example

Article A short artificial antimicrobial peptide shows potential to ...

Best regards.

PNA have favorable binding conditions and -kinetics compared to oligonucleotides and are stable against nucleases and proteases.

Hello Mariano,

It is pretty common to have carry-over analyzing plasma/serum samples. Someone recommended me the use of Trifluoroethanol to wash out all retained peptides. Basically you just use it as a sample and inject 1 or 2ul direct onto the column with a relative short gradient. It does not damage the column in my experience, so maybe you can give it a try in the future.

Good luck!

Spectrophotometers have limits to how high an absorbance they can measure accurately. Depending on the instrument, this may be as low as 1 or as high as 3. If you are struggling to measure the absorbance of your peptide because of high background absorbance, one solution is to use a shorter pathlength. Remember that absorbance is directly proportional to the pathlength of light through the sample. If you are using a standard 1-cm-pathlength cuvette and get an absorbance of 2, if you switch to a 0.2-cm-pathlength cuvette, the absorbance will be 0.4, well within the useful range.

thank you for kind answers !

i will try paper chromatography, kaiser test, NMR/MS, all things in order.

Thank you very much! I will read this paper in detail, thanks for your help! Rob Keller

Thank you for your kind answer.

the article you recommend is great help to me.

have a nice day !!

Depends if they undergo glycation or not.

I am so glad to spend few time to enrich our knowledge about Covid vaccine

You should use:

-HPLC at 205 to 220 nm.

-LC-MS.

and other Colorimetric methodologies...

You may try the standalone version of HADDOCK then:

Also, you may try FlexPepDock from Rosseta. Maybe that one is easier for running a whole library

Bests,

Hi Eef Dirksen,

Thanks for your comment.

I have some serum and tissue samples from patients with a certain disease, and I want to identify signature proteins or peptides that can differentiate them from healthy controls. I am wondering what is the best method to extract and analyze (top down or bottom up) these biomarkers.We have a Q-Exactive mass spectrometer for this experiment. how to handle the data analysis and interpretation? Is there any specific software or tools to process and visualize the results? 

DCM is a less polar solvent than DMF. So, it could dissolve Fmoc-Osu.

To glycine, you could try to dissolve in DMF.

Remember that the solubility depends on the quantity of solute and solvent.

Note:

-Consult in the bibliography the solubility of Glycine, Fmoc-OSU, and Fmoc-Gly-OH.

-Do not use water to form Fmoc-Gly-OH.

-Usually, Fmoc-amino acids are soluble in DMF, DMF/DCM, and DCM (in some cases).

-You could use ACN as a reaction solvent.

-Do not use water to form Fmoc-Gly-O-. Later you could use HCl/H2O to protonate Fmoc-Gly-O- to Fmoc-Gly-OH.

Follow the reaction by TLC of HPLC.

I think you're saying you're getting an unacceptably high background reading from your negative control. It's good that you're concerned about this problem. It is common, especially for human blood samples.

If you are using human blood, be sure to take proper bloodborne pathogen precautions, including your vaccinations and PPE.

Consider using serum rather than plasma. And for human samples you should heat the sera to kill viruses.

You should be diluting your samples in blocker solution. For ELISA you probably can start with a 1:1000 dilution and work toward 1:1000000.

You can reduce non-specific binding by adding things to your blocker solution. Starting with a base of PBS, blockers usually contain a protein like casein or albumin at 0.1% (1 mg/mL) and a nonionic detergent like Tween at 0.01%. You can try using a different protein. You can safely increase the protein 10-fold. Depending on the ELISA format, you may want to include 1% control serum in your diluent and blocker as a blocker protein. Increasing the detergent (up to 10x) may decrease your overall signal, but it may increase signal/noise ratio. And sometimes it helps to add 0.1M glycine.

Dmitriy Berillo Thank you

Thank you @ Dr. John Carter for the response nd suggestions.

For large-volume applications, you may prefer batch-mode purification. Add the prequibilirated bulk resin into the sample and incubate for a while and shake continuously. AAs will adsorb on resin while Chlorides are not.

If the purpose is just analysis, you can derivate the AAs and LLE using organic solvents such as ıso-octane, chloroform, acetonitrile, MTBE, etc may give a chance of both chloride removal and enrichment of the AAs.

You could also consider mass spectrometry. Intact mass spectrometry can detect the mass of all proteins in a sample without fragmentation if the proteins ionize well. Most mass spec systems are setup to detect only positive charges and since DNA is heavily negatively charged, it will be invisible to the detection. Mass spec is very sensitive to low quantities of protein, but the more protein you have the better the detection as there is more to detect.

Bottom up Liquid chromatography with MS/MS and trypsinization can detect the mass of all peptide fragments after trypsinization of peptides and proteins, but deconvolution of what those peptides correspond to is kind of catch22 because you have to know the sequence of the proteins in your sample. Nevertheless, it will detect all peptide fragments.

Vladan Nedelkovski thanks for the reply. However, MDAnalysis takes in only PSF files and not PDF files. I have already computed the distance matrix and based on which I wanted to do the clustering. I have computed an RMSD-based distance matrix between every pair of structures. Still, I couldn't cluster it properly.

The reason I asked is that by docking peptides to protease binding sites this situation often occurs. Without knowing exactly why, I discard these results but if anyone knows better...

Since the compound is a peptide, terminal sterilization (autoclaving) at 121C for 15 mins resulted in gelling of the formulation although there is no significant change in purity. Himanshu Bhatt

Identifying a cell line that expresses VEGFR3 exclusively or with very low expression of VEGFR2 and VEGFR1 can be challenging, as many cell lines express multiple VEGF receptors. However, here are some approaches to help you identify suitable cell lines:

Remember that it is crucial to carefully evaluate and validate the expression profiles of VEGFR1, VEGFR2, and VEGFR3 in the selected cell lines to ensure the accuracy and reliability of your results.

Thank you very much for all the useful information, very much appreciated.

You could lyophilize the mixed solution TFA-ether that contains the peptide.

Hello! I think there is a biacore T-100 or (3000) at your university. This is good technology and if what are binding is large then it should work quite well. If both of the species are small then maybe not so much. Good luck. Here is the potential lab that has the instrument. https://espace.library.uq.edu.au/view/UQ:722458/UQ722458_OA.pdf

You can use a peptide solubility calculator to predict its solubility:

Since it is composed of several charged residues with a pI ~ 12. It should be soluble about 3 pH units from its pH since it is uncharged when pH = pI.

Should be soluble in water or Tris pH ~ 8. Use autoclaved sterile water and filter the peptide.

If the peptide was composed of a lot of nonpolar amino acids then it wouldn't be soluble in water.

Hello Alina

You can begin by looking at the change in structure over time and comparing g it with changes in your RMSD. Where ever there is a drastic jump or dip in the RMSD, you have see what are the changes especially where the mutation was introduced and compare with the wild type.

You can look at the bonding behavior of the mutant and the wild type also.

I hope this helps

Thanks @Ramirez

Tris buffer

Alain Mangé Ok thanks

Phase separation? You mean it goes from water to an organic phase solvent, or vice versa, and you want to stop it? A surfactant should do the trick. You might start with Tween.

EDANS (via its sulfo-group) can interact with a free amino group of the growing peptide chain (after its Fmoc deprotection) and mask and prevent it from the interaction with an added Fmoc amino acid. We are preparing a publication that discloses this event and suggests an approach for preventing it.

My name is Michael and I work for Biomatik. I highly recommend our custom peptide service for your research needs. Biomatik has been trusted by 10,000+ Scientists Since 2002.

Alternatively go back over your sample preparation protocol and determine the source of the PEG contamination.

No. You should have made a standard curve with several concentrations of a protein standard, such as BSA. The standards should also have been added as 10 µl samples. You can therefore compare the absorbance of your sample with the standard curve without calculating a dilution factor for the volume of Bradford reagent added..

It seems your suggestion is right. Laser exposure even at red wavelength (620-670 nm) could induce permanent damage to peptides or other biomolecules, and rate of possible signal degradation should depend on laser power in focal volume and total volume of the sample.

There are 3 of possible effects: 1) peptide fragmentation, 2) permanent peptide conformation switching 3) peptide precipitation/condensation out from solution and seeing FCS signal from impurities in the solution.

I also tried to dissolve the peptide in DMSO and it worked, however, cysteine containing peptides should be dissolved in DMF instead.

Hello Benjamin Justin Mildenberg

The number of cells must be adapted to the cell type as well as the frequency of secreting cells namely, the lower the expected frequency of responding cells, the higher the number of cells per well. For instance, if the expected frequency is 1 in 10,000 cells, then you will need approximately 200K cells per well. Conversely, when the expected frequency is high, say 1 in 100 cells, then it will be sufficient to seed 100K cells per well.

Another important factor I would like to mention is that antigen specific responses will require higher cell number (200K) while the use of mitogens may require lower cell number (100K). You also need to know that a minimum number of cells is required for proper cell-to-cell contact and thus for optimal stimulation. For example, if one is analyzing antigen-specific T cells in a PBMC sample, the cell number should be 200K cells per well.

So, cell number would play an important role in ELISpot assays in addition to the test peptide concentration.

Best.

Yes, your ratio is suboptimal on theoretical grounds. I don’t think there is much wrong with the pH at 7.4, but the reaction will still work at lower pH values. Whatever pH is used, you do need the peptide in molar excess over the maleimide functions to get the desired product in high yield. A better approach than manipulating pH to stop the N-terminal amine from triggering the side reaction is simply to block the amine.

Mode of action ?

Check :

The choice depends on available instrumentation and expertise, and some what less on the peptide structure. If your peptide is big enough, or if it includes some feature to limit structure, then you may observe characteristic alpha and/or beta absorption using CD. A short peptide, with only one beta strand or part of a helix, may not fold stably in the absence of a partner.

I recommend you minimize Cl- ion, as it exhibits high background absorption. I suggest you also predict secondary structure based on primary structure, using any of various computer tools.

Dear Muhammad Fikri Heikal,

I agree with Yun-Tzai. Both Raman spectroscopy and circular dichroism are powerful techniques used for analyzing protein folding and secondary structure, but they offer different types of information and have distinct advantages and limitations. I recommend you scientific articles that can illuminate for you the range of possibilities of these techniques:

Raman spectroscopy:

CD:

The choice between the two methods depends on the specific goals and requirements of your peptide folding analysis. Let's compare both techniques:

1. Raman Spectroscopy:

2. Circular Dichroism (CD):

However, CD might not provide as much detailed structural information as Raman spectroscopy. If your primary focus is on obtaining detailed information about the vibrational modes and tertiary structure of your peptide during folding, Raman spectroscopy might be a more suitable choice. On the other hand, if you want to primarily analyze the changes in secondary structure during folding, CD would be the preferred method. In some cases, combining both techniques can provide a more comprehensive view of the folding process.

Good luck

Comparing peptide solubility using different solvents the result was as follows:

Ammounia and DMF>DMSO>NH4HCO3

Ammonia 0.1 %

NH4HCO3 0.1 M

Edward Michelini It's a fluorophore, but I can't seem to find any HPLC machines available to me with an appropriate detector, or anybody with the right expertise in my school. I may have to have another think about doing preparative HPLC, I was getting quite good at the set up!!

Thank you :)

Well, of course you can do the usual syntheses also with having your half-built peptide scaffold swimming around instead of having it pinned to a substrate, but it is much harder to do it in a precise way, so probably your yield will decay while a larger fraction will end up as a side product and also you will probably have more in-process losses. In principle the usual coupling agents and protecting groups will still do their job.

Dear all, have you solved the problem? I have the same problem as Harry, could you please clarify how you solve this problem? Thanks much!

Are you 100% sure your sampling peak is homogeneous ?

Hi Bonnie,

We got you! Send us the peptide or protein sequences and we will make suggestions for your antigen candidate. Once the project is set, we will synthesize the peptide and then use it for immunization.

Our deliverable will include the purified antibody AND the peptide that we use in the antibody production!

Send us a request with your project details to initiate our collaboration: https://www.biomatik.com/services/custom-antibody-service/polyclonal-antibody-production.html

We look forward to hearing from you soon.

Best,

Biomatik Team