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I have been a little concerned that the 7H9 media for growing mycobacteria becomes cloudy and crashes out of solution after autoclaving. It is fully dissolved before autoclaving and is clear, but once it has been autoclaved a sandy precipitate accumulates at the bottom of the bottle. Is this a standard problem? Is there anything we can do?
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Hey @sara I am experiencing the same issue although I am adding supplement and glycerol after autoclaving, Did you figure out the solution?
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Hey, I have been trying to design a peptide for a surface receptor using the RF-diffusion, But every time I am getting a peptide that has only glycine as the residue in results. I have looked at multiple literature but could not find relevant explanation.
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Hey! I know I'm late to reply but this is actually documented in the RFDiffusion code. The peptide is initially generated as a poly-glycine structure but then you can use the code at this repo:
to convert the poly-glycine into a valid protein structure. This code can also give you the PAE-interaction score which can help to qualify the "goodness" of the generated structure.
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I am currently learning about PyMol to utilize in my project. I used PyMol to visualize potential H-bond interactions in specific amino acid residues. However, I have discovered that Arg465 and Ser461 show a distinct interaction, as shown.
Please help identify this interaction.
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The broken yellow line with the distance indicator (6.2) looks like a simple distance monitor which you generate with a "measure" command, although I do not know how you generated the blue tubes around it. At 6.2Å, the Ca-Ca distance indicated by the broken line is far larger than the sum of the carbon Van der Waals radii (3.4Å). It is just about short enough that you might classify the contact as a solvent excluding contact (hydrophobic interaction)
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I am docking multiple ligands (new designed ligand) against my protein using Autodock Vina in Chimera. The results displayed in this program are somewhat strange, because in some cases the ligands with no hydrogen bond or fewer bonds has the most negative Score than the ligand with more hydrogen bonds!!? Please look at the picture to get my mean(The best model in ligand with 2 hydrogen bonds has a score -8.3, While the best model in ligand with one or zero hydrogen bond has score -8.7 and -10.1 respectively!).
I understand that checking with other software or tools like PyMOL or PDBSUM will better help to analyze the possible interactions, however since I have several ligands with almost similar score and interaction network or equal hydrogen bond numbers, I am curious to now how to pick the best one (based on the in silico analysis) among them. If any body has suggestion for this I will appreciated it.
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You should choose the pose with the highest score (-10.1) since it has a significant score difference with the second pose.
Docking algorithms like Autodock Vina evaluate multiple factors when calculating the docking score, including van der Waals interactions, electrostatic interactions, and hydrogen bonding. While the number of hydrogen bonds can be an important indicator of ligand binding, it's not the sole determinant of the docking score. Other factors such as the strength and geometry of the hydrogen bonds, as well as non-polar interactions, can also contribute to the overall score.
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What to do if ChimeraX software doesn't recognise the .chimerax file downloaded from SwissDock after docking?
Besides, the zip file of prediction done was empty.
Thank you.
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I had this issue - I used 7-zip to unpack teh folder instead and then it was fine
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I want to cluster multiple pdb files with respect to one reference pdb. I have to use Align > all this to *Ca in Pymol. However, how could i do this via scripting in Pymol? As I have multiple sets of pdb files with their own reference pdb files. Is it possible to do the same in Python via Jupyter Notebook in Pymol?
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Example script that demonstrates how to align and cluster multiple PDB files with respect to a reference PDB file using PyMOL and Python:
```python
import pymol
# Start PyMOL
pymol.finish_launching(['pymol', '-qc'])
# Load the reference PDB file
pymol.cmd.load('reference.pdb', 'reference')
# Iterate over the list of PDB files to align and cluster
pdb_files = ['file1.pdb', 'file2.pdb', 'file3.pdb']
for pdb_file in pdb_files:
# Load the current PDB file
pymol.cmd.load(pdb_file, 'current')
# Align all atoms to the reference
pymol.cmd.align('current', 'reference')
# Cluster the aligned structure based on CA atoms
pymol.cmd.cluster('current', cutoff=3.0, selection='name CA')
# Save the aligned and clustered structure
pymol.cmd.save('aligned_clustered_' + pdb_file, 'current')
# Delete the current structure from the PyMOL session
pymol.cmd.delete('current')
# Quit PyMOL
pymol.cmd.quit()
```
In this example, you would replace `'reference.pdb'` with the path to your reference PDB file. Similarly, update `'file1.pdb'`, `'file2.pdb'`, and `'file3.pdb'` with the paths to your own PDB files.
The script iterates over each PDB file, loading it into PyMOL as the `'current'` object. It then aligns the `'current'` structure to the `'reference'` structure using the `align()` function. The alignment is performed on all atoms.
After the alignment, the script clusters the aligned structure based on CA atoms using the `cluster()` function. The `cutoff` parameter specifies the clustering distance threshold.
Hope it helps
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I was using fragbuilder module in python to generate peptides of sizes 4, 6, and 10. However, the issue with fragbuilder module is that some of the bond angles are deviating from the standard values. For instance, C_alpha--C--N bond angle standard value is 121 degrees but fragbuilder assigns 111 degrees. This angle deviation causes a deviation in the distance between the nearest neighbor C_alpha---C_alpha and its value is 3.721 angstrom and the typical standard value is 3.8 A. Also another bond angle is a deviation from the standard value by 6 degrees which is the C_alpha---C---N whose value is 111.4 degrees and typical standard values are 117 degrees. My doubt is how much deviation is allowed for MD simulations of peptides (or proteins) while fixing the bond lengths and bonds angles ?
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Gary James Hunter Thanks for you reply.
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Recently I am stuggling to improve the kinetics of an artifical enzyme. I expressed this enzyme in E.coli and then test it's kinetics properties.
I noticed that if I pick single conies for testing, there will have a variation in both reaction rate and maxium reaction. indicating in fig.1 (the y axis represent the product that have been formed and the x axis represent the time in seconds.) 4 different conies have been picked and they all contain the same plasmid that transfection at the same time and same procedures. However there is huge differeces in the reaction rate and maxium reaction.
Then I wonder if it's due to the different conies would fold the protein differently, so I did another test by add multiple conies (actually all conies on one dish) into my culture medium. And then I test this mixed enzyme with different substrate concentration to test the affinity and kinetics at the same time. fig.2 (different color represent different concentration; the dash line represent a Imaginary limitation)
The problem that makes me wonder is that: what might be the reason for this reaction have a rate limitation?
I have few hypothesis about this phenomeon:
1. based on the Imaginary rate limitation; there might have steric effects preventing the binding of the substrate. (but I don't have see enough enzymatic reaction curve that have steric effects)
2. based on the varation between conies; this artifical enzyme might have many different ways of folding (I mean this enzyme would have many different prefered structures in different bacteria cells). maybe bactria from the same coniey would prefere similar stucture? and some stucture have better enzymatic performance, others do not.
I am really appreaciarte your reading and would be very happy to receive any response.
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Since you frequently refer to colonies ("conies"), it seems that you are not growing a culture of the cells and purifying the enzyme, just extracting the enzyme from the cells in the colony. In that case, there is not much point in studying the kinetics of the enzyme because it has not been purified. The signal obtained may be due to a mixture of multiple enzymes. The amount of the enzyme obtained from a colony may depend on factors such as how many cells are in the colony, the age of the colony, and how well the enzyme were extracted from each colony.
Generally speaking, an enzyme has a single overall conformation. It will not have different structures in different colonies. It might be expressed to different levels in different colonies. Some of it may be in an insoluble form due to failure to fold properly, and the proportion of insoluble, inactive protein may differ between colonies.
If you want to study this enzyme's kinetic properties, you really should purify it.
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I want to do DNA-Protein docking at specific active site. So please help me which tool and server can i use for this type of docking.
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There are several tools and servers available for DNA-protein site-specific docking, including:
  1. HADDOCK (High Ambiguity Driven biomolecular DOCKing): This is a widely used software for docking macromolecules, including proteins, nucleic acids, and their complexes. HADDOCK is capable of performing site-specific docking by specifying the active site residues of both the protein and the DNA molecule. HADDOCK can be downloaded and run locally or accessed through a web server.
  2. PatchDock: This is a user-friendly docking server that performs protein-DNA docking. It allows the user to specify the active site residues of both the protein and DNA molecule. The server generates several possible docking solutions and ranks them based on their predicted binding energies.
  3. HEX (High-speed EXhaustive docking): This is another widely used software for docking macromolecules, including proteins and nucleic acids. HEX is capable of performing site-specific docking by specifying the active site residues of both the protein and DNA molecule. It can be downloaded and run locally.
  4. ClusPro: This is a fully automated docking server that can perform docking between proteins and nucleic acids. ClusPro is capable of performing site-specific docking by specifying the active site residues of both the protein and DNA molecule. The server generates several possible docking solutions and ranks them based on their predicted binding energies.
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There are so many softwares for docking but which one is best? On which we have to rely?
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It's depended on the software algorithm.
The equipment listed below might be useful
AutoDock Vina, AutoDock
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I have solubilized my protein with 0.3% sarcosine and purified by using Ni-NTA,during purification most protein is going into flow through.
I have diluted my sonicated sample to 0.1% sarcosine but still I am unable to get binding of protein.
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Gaurav Chhetri Sir, I used this approach as well my protein is soluble in 1% sarcosine , but when i perform dialysis my protein get precipitated resulting in total loss of protein, what should I do
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Dear community.
I'm just studying for a master’s degree in the department of biochemical science & technology, at National Taiwan University.
When I try to refine the structure at Coot or search the paper about the protein structure or membrane protein.
I'm curious about what is the meaning of Fo and Fc, and the difference between the Fo-Fc map and the 2Fo-Fc map their meaning.
Why they are important when we try to fit the residues or molecules in electron density maps?
Also, I what to know does it has any program or application that can create the density map in this journal article.
The supplementary figure 6 in Astashkin, R., Kovalev, K., Bukhdruker, S. et al.Structural insights into light-driven anion pumping in cyanobacteria. Nat Commun 13, 6460 (2022). https://doi.org/10.1038/s41467-022-34019-9
It's my first time asking the question on this website and I’m not an English native speaker, sorry if I offended you.
Cordially,
Guan-Yi.
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2Fo-Fc is the best way to calculate an estimate of the true electron density from diffraction data and atomic model. (It is called 2Fo-Fc because the calculation involves combining the observed diffraction data, Fo, with the diffraction data calculated from the atomic model, Fc, in a way that gives the least-biased result).
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I had developed a homology modeled protein and identified the cavity of the protein through bioinformatic software. unfortunately, i couldn't calculate the cavity volume in my modeled protein. I want to get some information about the cavity volume and size in modeled protein for de novo drug design. kindly let me know the way to calculate the cavity volume of the homology modeled protein
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I was asked for my solution to this recently, so if someone else stumbles over this question, here's the R script to transcribe the CASTp json files into PQR files for import into ChimeraX, Pymol, etc.:
Let me know if you run into problems, I'll try my best to help!
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I want to visualise secondary structures of multiple proteins aligned (something similar to this figure). Any recommendations?
Thanks in advance
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try this one:
all the best
fred
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Why would AlphaFold give a protein model that contains homoserine instead of serine? Is there anything wrong with the predicted structural models? Thank you.
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Hi, how did you get the model? did you input the sequence or you downloaded the sequence from their website. It is unlikely that AFold should give homoserine rather than serine since its training material contains natural sequences.
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Can the differently coloured region around the ligand be saved so that I can directly see the coloured region the next time I open it? If so, what format would be the file? pdb format? Thank you.
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To preserve all the information of a ChimeraX session, you have to save it as a session file (extension .cxs). This enables you to continue the session from exactly the point where you saved it, colouring, orientation and all.
Saving as .pdb would only save the coordinates, no ChimeraX specific information.
You use the .pdb format (or .cif) to transfer the coordinates to other programs which cannot interpret the chimeraX specific information.
See https://www.cgl.ucsf.edu/chimerax/docs/user/commands/save.html to see a listing of all supported formats, and which part of the session they cover - the session file is the only one that covers all the information in your ChimeraX session - but it can only be opened by ChimeraX! All other formats only export the part of the information that can be interpreted by a particular class of other programs, e.g. images, that are 2D views that you can print or insert into a word processing file or a powerpoint presentation.
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I used AlphaFold to predict the structure of a protein that has not been well studied.
I have a pdb file of the predicted structure.
Now I would like to identify the domains of the protein using this pdb file.
Is there a suitable tool for my purpose?
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It depends on how you want to identify domains.
AlphaFold2 uses HMMER(http://hmmer.org/) to search seq DB(Pfam)s, and make alignment files, then based on these alignment files calculate the contact maps for folding. So...
1) aa seq-based domain info:
You can grab it directly from AlphaFold2 after HMMER calculation. However, simply you can put your aa seq into the Pfam website will give a similar result. This kinda method will work only when a similar seq is on the DB. AlphaFold2 does some stupid things when it is given multi-domain protein seq which is not on the Pfam DB.
2) Without having aa seq info, use only the PDB file to predict the domain:
well, this could be a harder question, a) either you write custom python code to calculate the center of mass and define the cutoff value to identify two domains,s or b) use at least two PDB files of the same protein and then align them then calculate the relative deviation of the other. We have a couple of unpublished multi-domain protein structures and compared them with AlphaFold2's performance. it predicted well on domains but not the relative position of each domain.
I think you need to provide more info about why and what is next step or what is the purpose of identifying the domain from AlphaFold2 prediction, if not Pfam domain info?
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Hii, Is there a way I can extract the alternative spliced protein isoform structures from PDB? Also can we mapped the structure to uniprot sequence So we can know which structure belong to which isoform sequence?
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Unfortunately, most of the databases contain 3D structures only for canonical isoforms. But, you would try Google Colab, a phyton-based online notebook running AlphaFold2, which can predict the structure of any custom sequence or noncanonical isoform. Check this out https://youtu.be/le7NatFo8vI
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Is AlphaFold accurate enough when the protein shares less than sequence similarity with the closest, structurally solved homologue?
Besides, how accurate is Alphafold in de novo structure prediction of protein families without solved structures?
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Several studies have found that AlphaFold predicts better structures than other approaches. The EBI alphafold database (alphafold.ebi.ac.uk/faq) displays the per-residue confidence score (pLDDT). It provides information about the model's accuracy. The majority of plant proteins in the EBI AlphaFold DB (alphafold.ebi.ac.uk/search/text/Oryza%20sativa%20subsp.%20japonica) have a pLDDT score of >50. Even proteins with less identity appear to be accurately predicted by the AF. For longer proteins and disordered proteins, AlphaFold has some limitations.
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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?
Thank you.
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Yes, it is mandatory to determine the proper molecular arrangement in space since the atomic coordinates of the protein structures are not energetically favorable. . The aim of energy Minimization is to find a set of coordinates representing the minimum energy conformation for the given coordinates b/w the amino acids. Lower the energy more stable they are.
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I want to perform targeted molecular docking of:
(a) a receptor (enzyme) whose structure is not availabe, and hence has been built by computational ab intio methods (and not homology, since the % identity is very low);
(b) a substrate whose structure is availabe.
Given that, the active sites of the enzyme are also not truly available, but has been obtained from (a) literature review; or
(b) inferred from cavities/clefts predicted by CASTp results, how exactly should I perform targeted molecular docking?
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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?
Thank you.
Sincerely yours,
Thai Leong
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A good decoy would be randomizing the sequence of your query peptide, an construct some of them. This would give you a set of same length peptides with the same aa composition. This is similar to the procedure in BLAST for estimating E-values for alignments
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Dear all, Hope you all doing well.
I was worried about the prediction of protein-protein interaction using online servers like ClusPRO. Actually recently we found that Protein A interact with Protein B using yeast two hybrid method. Protein B exist as a complex with Protein C, D, and E. Now, biochemically, Protein A didn't show interaction with Protein C, D and E. However, when I performed the protein-protein docking using ClusPRO, it shows interaction of Protein A with all Protein C, D and E. The question is how to rely on such online servers. Because it is giving interaction with whatever protein (as receptor and ligand) you feed to the Job server. It doesn't make any sense to me. How one can differentiate in-silico that Protein A and Protein B is the better interaction propensity than other member of the complex. I cannot do Molecular Dynamics simulation for such huge complexes with all permutation and combinations, because have no expertise in it and also it will be computationally very expensive. Please explain and give your valuable suggestions this in a simple manner.
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No personal experience but it seems to me that ClusPro as described in their paper https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5540229/ is one of the best performing programs available (and cited >1000 times). A general rule is to never rely on one (bioinformatics) method. So, I would:
-Use at least one other docking protein and realise that no method is 100% accurate (usually round 90% is the best you can get) and every method has their pros and cons
-Rely on the biochemical evidence (more). However, do realise that the method used might not be flawless or is unable to show subtle differences that might be relevant in your case
-Use another (biochemical/biophysical) experimental method to confirm your earlier experimental findings
Best regards.
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Let's say we docked Protein X with Ligand A, Ligand B and Ligand C respectively.
Meanwhile, we have three different desktops, i.e. Desktops 1-3 for molecular dynamic (MD) simulation. Can we make use of the desktops using the following way in speed up the process of getting the results from MD simulation?
First round of MD simulation:
Desktop 1 - Protein X-Ligand 1 complex
Desktop 2 - Protein X-Ligand 2 complex
Desktop 3 - Protein X-Ligand 3 complex
Second round of MD simulation:
Desktop 1 - Protein X-Ligand 3 complex
Desktop 2 - Protein X-Ligand 1 complex
Desktop 3 - Protein X-Ligand 2 complex
Third round of MD simulation:
Desktop 1 - Protein X-Ligand 2 complex
Desktop 2 - Protein X-Ligand 3 complex
Desktop 3 - Protein X-Ligand 1 complex
Thank you.
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It is okay to do that. The results are supposed to be machine independent, so that peers can also reproduce the results. Make sure that the software version is the same in all three devices.
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Hi,
I have been trying to minimize a protein showing invalid bond type in some residues. while minimizing it throw an error "restrained minimization has failed". please help
thanks in advance.
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You need to have Prime license to run the protein minimization.
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I generated the predicted structure of a dimer of proteins using AlphaFold2, but it has overlapped amino acids like the picture.
Because of this, SMOG ( https://smog-server.org/cgi-bin/GenTopGro.pl ) returns an error `FATAL ERROR: Contact between atoms 356 451 below threshold distance with value 0.192` when using this PDB as an input.
What I came up with was performing energy minimization on the entire protein by MD simulation software like Gromacs or resolving a part of the amino acid sequence using molecular viewers or modeling software like Pymol.
Which is a better way or is there another way to solve this issue?
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I checked your PDB file with both VMD and UCSF chimera. I could not find overlaps or missing .....
Then i checked it with gromacs 5.1.1 and generate a topology file for it ( OPLAS FF)...
Could you please discuss about your exact problem, becuase i think you can minimize your this PDB file with out any problem in Gromacs or VMD or ....
I also checked your PDB file in text format and i could not find any overlap residues,
Please read SMOG help or tutorial for specific instruction of pdb file, hope this help you ....
Please check the attached files
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If MD simulations converges to Boltzmann distributions ρ∼exp(−βϵ) after sufficiently long time why do we need MD simulations, as all the macroscopic quantities can be computed from the Boltzmann distribution itself. This question I am asking for short peptides of sequences of few amino acids.(tripeptide, tetrapeptide etc).
For instance in the given above (link) paper, they are using MD to generate Ramachandran distributions of conformations of pentapeptide at a constant temperature. So this should obey statistical mechanics. If it is so, then this should satisfy Boltzman distributions.So I should be able to write down the distributions using boltzmann weight as follows,
ρ({ϕi,ψi})∼exp(−βV({ϕi,ψi}))
.Here, all set of Ramachandran angle coordinates of the pentapeptides is given by {ϕi,ψi}{ϕi,ψi}.
Why should I run MD to get the same distributions?
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As
Behnam Farid
pointed out, you cannot know all the relationships (the functional form V({ϕi,ψi})) between the different amino acids (sterical clashes, interactions based on charges or hydrophobicity) to predict the energetically favorable combinations of phi and psi and hence need to sample them. The state distribution is affected by the amino acid sequence, may differ with the force fields and simulations methods, does depend on the solvent (ions etc.) and temperature.
Have a look here to see how complex the conformational space of small peptides (13-15) can already be:
Bests
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I am working on a ligand that is co-crystalized to parotein, but the ligand is missing some residues that makes it appear as if it was separated into two ligands!
what I need is to connect them into one to run molecular dynamics simulation, what is the best tool to do so, also what are the steps to make sure that it will be mostly accurate?
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Try Ligandscout or Maestro, Schrodinger.
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Hello all
Is there any reliable and free web server that runs molecular dynamics simulation of proteins?
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These days journals demand for 100 ns and Online tools are not reliable for MDS. The best solution for MDS is always gromacs or schrodinger.
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In the PubChem database I could find a 2D structure of the compound (CID 24763) but wants a 3D structure for docking, moreover the compound is too long so is there any technique to cut it short since the compound is starch like polysaccharide??
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Download the 2D and convert into 3D using Marvin sketch or chemsketch or chemdraw.
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I am trying to estimate the width of DNA from its crystal structure.
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Hi can you share the notebook ?
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I've used phyre2 to model the protein for the simulation using Gromacs, but later I found that 2/3 proportion of the structure ( except for IDR ) had been already determined by X-ray crystallography.
The known structure contains Zn2+ to stabilize the structure of the entire protein, so I doubt phyre2 can predict decent structure. How should I model the structure of proteins using a known structure as a part of it?
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Dear community,
As far as I understand, negative density features point at stuff that is in the model, but not supported by experimental data.
However, in many cases (e.g. attached image), when you visualize the densities, there are negative (red) density blobs not containing any atoms inside. Just empty space inside. There is nothing in the model in those regions of space. How it should be interpreted?
Could you tell me please what I am missing?
Would be grateful for any help,
Best wishes,
Aliaksei
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In layman's language, negative density is nothing but the regions where you build nothing, protein backbone or heteroatoms or anything because probability of finding atoms in these regions is zero so one should not build anything in red/negative density.
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I have used modeller to generate a pdb file. i have run it on PROCHECK. but the input format is not compatible. (brook haven format). How do i convert my file? or any other way? Please help. ASAP
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PROCHECK already accepts .pdb files. As far as I understand, Brookhaven might be an alternative way to call the PDB format (an old one, maybe?). I found some references to it around the internet, e.g. "Brookhaven Protein Databank format".
Anyway, just a remark: check to see if protein chains are specified in the MODELLER output file (sometimes MODELLER doesn't set chain IDs; this can cause problems with PROCHECK). If in doubt, don't use the chain parameter in the PROCHECK command, e.g. `procheck model.B99990004.pdb 2.0`. The resolution parameter is necessary, but any number is accepted (I've used 2.0 in the example above).
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While am running the EM, the minimizatin gets stopped before the forces get converged at step 14
Steepest Descents:
Tolerance (Fmax) = 1.00000e+03
Number of steps = 50000
Step= 14, Dmax= 1.2e-06 nm, Epot= 1.84916e+18 Fmax= inf, atom= 7809
Energy minimization has stopped, but the forces havenot converged to the
requested precision Fmax < 1000 (whichmay not be possible for your system).
It stoppedbecause the algorithm tried to make a new step whose sizewas too
small, or there was no change in the energy sincelast step. Either way, we
regard the minimization asconverged to within the available machine
precision,given your starting configuration and EM parameters.
Double precision normally gives you higher accuracy, butthis is often not
needed for preparing to run moleculardynamics.
You might need to increase your constraint accuracy, or turn
off constraints altogether (set constraints = none in mdp file)
writing lowest energy coordinates.
Back Off! I just backed up em.gro to ./#em.gro.9#
Steepest Descents converged to machine precision in 15 steps,
but did not reach the requested Fmax < 1000.
Potential Energy = 1.8491607e+18
Maximum force = inf on atom 7809
Norm of force = inf
What should I do?
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@Richard Mariadasse can you suggest me how do you solve the problem?
I'm having the same issue
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A number of post translational modifications can occur to a protein, including phosphorylation, methylation etc. I am aware that there are a number of servers and tools to predict positions for these modifications based on sequence and/or structural patterns or motifs. However, is there a tool (or server) available that can actually model them on a structure (or a model)?
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Hello,
If you want to add a post translational modification in an existing pdb file You can try Vienna 2.0
Best,
Christophe
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Given:
1. The nearest neighbor of 𝑝𝑖 then 𝑝𝑖-𝑝𝑗 is a Delaunay edge.
2. In a 3D set of points, if we know that consecutive points ie... 𝑝𝑖-𝑝i+1 are nearest neighbors.
3. The 3D points do not form a straight line
Assumption:
Each Delaunay tesselation (3D) has at least 2 nearest neighbor edges.
Is my assumption true? If not can you please explain to me the possible exceptions?
Thanks,
Pranav
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Are you trying to play chess in 3D?
Your idea is a good one, but your assumptions are not.
You need to give a clear definition of paths, so I suggest for you to start in one 3D box, it includes 8 points. I prefer to give each point the following notation
P(i, j, k), so the locations of the 8 points are at
(0,0,0) (1,0,0), (0,1,0), (0,0,1), (1,1,0), (1,0,1),(0,1,1) and (1,1,1).
Study this cube carefully, define each Delanoy edge (axioms of the path), and then add another box, which means 12 points, etc.
If you find the closed formula that allows you to calculate all possible paths from the starting point at the origin to the farthest point at the upper corner of the rectangular box, then you are on the right track.
I wish you good luck.
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I would like draw protein 3-D structure? Can anyone suggest me the best software that I can buy?
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I think for drawing of protein 3-D structure, softwares such as SPDBV and PyMOL are proper.
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Hi everyone,
I need to do MD simulation of wild type and ten variants at 50 ns. I am looking for a low-cost cloud service/ simulation environment. Would you please suggest me any?
Thanks in advance.
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you can check docker: https://www.docker.com/
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I had used SUMMA server for structure refinement.
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Hi,
Typically, the solvent accessibility of water-soluble protein is computed by using a water molecule with a size of 1.4 Å). For my research work, I need to compute the relative lipid (-CH2 as a probe molecule) accessibility for membrane protein structures through NACCESS program. In this regard, I don't know how to change the solvent parameters in NACCESS.
Is there anyone who can help me with this calculation?
Thank you in advance.
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Yuhong Mao I am not using Mac
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Hello everybody! Somebody knows an open source software for induced-fit docking? I have to dock ligands on receptor-models which have low sequence identity with the template (13-20%) and I was thinking that an induced fit docking can help me to improve the quality of my models, at least in the binding site.
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Nail Besli I used induced fit docking to sample different conformation of the binding site. Then I docked known active compounds and a set of decoys to all the models generated with the induced fit, selecting the model that ranks known active in the top ranked (according to docking score). ROC curve and enrichment factor were used to select the best model.
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I want to run protein-Ligand simulation where gold atoms are ligand. Gromacs is showing error for that. I tried to use Prodrug server also, but it is not generating coordinates for gold atoms. 
Can anyone help me in this?
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@Martiniano Bello , Thank you very much. I have download the golp.tgz and unzip it to the force field folder in gromacs or the input file folder . But I didn't find the relevant force field after runing the 'gmx pdb2gmx' command. Could you explain how to use the Golp for a greenhand? Thank you.
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I want to compare the active sites of some protein models in quantitative way (their volume or dimensions), Is this option available in the following softwares (MOE, YASARA, Pymol, Chimera) as I have access to them only. If any one had a similar experience with one of these softwares specifically.
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PyMOl is the batter solution for this for the countdown of the volume of the protein active site as I know. Furthermore, the total volume of the drug-protein complex or only protein volume with the unit is accurately counted by the YASARA dynamics suit.
Thanks
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Dear all,
I am a beginner and I have plotted dssp plot using cpptraj for the first time using following script;
trajin md-sim.ncsecstruct 1-263 out dssp.gnu sumout dssp.sum.agr
I wanted to know that what is None in dssp.gnu plot. I have read in amber manual that None is nothing just 0 integer. If it is correct so why cpptraj secstruct script plot None. Can I exclude this from dssp plot. Kindly help me.
[set cbtics {"None", "Para", "Anti", "3-10", "Alpha", "Pi", "Turn", "Bend"} ]
Thanks.
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Hi Saman,
"None" refers to no secondary structure predicted, sometimes referred to as random coil.
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Ignoring a proteins sequence, say I have a predicted contact map plotted from a useful tool like ConKit, it can be quite easy to determine general regions of secondary structural features and if is running paralell or anti-paralell etc etc..
However besides this, does anybody have any good tips for breaking the predicted contact map down further to see regions which could be contacts involving loops or helices-loop contacts? In essence I am seeking some tips for how someone would step-by-step analyse different factors from a predicted contact map.
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I have been analyzing 90-amino-acid fragment of a protein using HSQC and triple-resonance (1H-15N-13C) NMR. I know roughly where arginine, lysine, and tryptophan side-chain peaks lie on the HSQC, but how can I definitely distinguish what is a backbone and what is a side-chain? In addition to the HSQC, I have several 3D experiments (e.g. CBCANH and CACB(CO)NH).
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Arg Nε-Hε and at low pH Arg Nη-Hη and Lys Nζ-Hζ are visible in HSQC but these are aliased/folded peaks. Go for an open Sweep Width (in 15N dimension) HSQC where these peaks will appear at their exact 15N ppm values and you can identify them easily. If you go to the R/K side chains' 15N plane in CBCANH, you will observe CD and CG of R or CE and CD of K. The Trp side-chain usually shifted downfield and appear near the bottom left corner.
Start with the assignment, you will understand which one is the backbone NH because in CBCANH and CACB(CO)NH they will show only the CA and CB (these you have to identify based on their C ppm values). But the side chains will be different.
Best wishes.
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in many insect including Lepidoptera, there are two kind of sperm: Eupyrene and Apyrene. how can we distinguish two kind of sperm in pictures?
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Dear Fatemeh,
besides Lepidoptera the other group with well studied sperm dimorphism are the prosobranch gastropods. My studies of Serpulorbis strongly suggest, that the "atypical" spermatozoa supply (or are) nutrition to the typical spermatozoa (which are the regular, fertile spermatozoa). There might be some similarity in your objects of research.
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I need to find weather a region of a protein that I'm currently working on has a Calcium Binding Domain. The putative region has a similar amino acid profile as an EF domain.
I would like to run the sequence through a tool and get some proper perdition results 
I saw on many journal articles, showing the predicted EF hand domains with  EF hand test results. (Attached image)
Can anyone please suggest me a proper prediction server that I can use :)
Thank You
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Try IonCom (Zhang Lab) for any metal binding prediction.
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what is the best protein design program that can assess the effect of mutation in a globular protein structure? i am looking to change the specificity of a protein. but mostly i am interested to measure the outcomes of the mutation on the protein itself rather than the interaction. and if i want to look into the interaction as well should i use protein design or MD softwares?  
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Pymol software
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3d structure of a protein is made up several secondary structure elements like helices and sheets. How to find the number of these elements in a pdb file?
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You can open your pdb in Chimera, go to Tools - Sequence - Sequence and it will give you the answer
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Can't access CastP(an online tool for surface analysis of protein) from links i've found through google(http://cast.engr.uic.edu/castp/calculation.php, http://sts.bioengr.uic.edu/castp/)
I guess their server is down or i'm trying the wrong URL. Please suggest me any alternative.
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Thanks Jeff! Now we've upgraded CASTp into new 3.0 version, this link is correct and didn't change: http://sts.bioe.uic.edu/castp/
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I would like to understand the reason why proteins show frothing.
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Because proteins are amphiphilic and can act as surfactants in a similiar way detergents do. They can adsorb at air-water interface and lower the surface tension of the solution which allows formation of the foam.
Dawid
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I'm currently trying to get the 3D structure of a set of peptides (ranging from 12 to 20 aminoacids). Subsequently we want to make docking analysis against an enzyme.
Which software do you use for that? How do you refine the structure?
Thanks!
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Such short peptides usually do not assume a single defined structure in solution. The structure of the peptide in the complex is not necessarily the dominant conformation in the ensemble of structures in solution, but induced by the interaction with the binding partner. As a consequence, rigid docking of peptides usually is not possible.
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Hello structural biologists! I am interested in developing an inhibitor to target the assembly of a specific heteromultimeric complex. The complex exists in the cytoplasm of most eukaryotic cells. A crystal structure of the entire complex is available. I am interested in targeting the interaction surface between the two proteins.
The binding surface is comprised of two alpha helices, one from each protein. I am aware that PPIs are considered a particularly tough target for inhibitor development. However, I have an advantage here, as several residues in close proximity are known to be essential for complex formation. In a sense, these residues seem to form a binding pocket, albeit one shared between two proteins.
My goal is to develop a pharmacophore model of this 'binding site' so that I can apply virtual screening to look for compounds that can mask this interaction. I have not found much information on the generation of multi-protein pharmacophores. I would be grateful if someone could point me in the right direction.
Regards,
Patrick
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Go through this article:
Pharmacophore based virtual screening for identification of marine bioactive compounds as inhibitors against Mip protein of Chlamydia trachomatis. February 2016, RSC Advances 6(23):18946-18957
DOI: 10.1039/C5RA24999F
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I'm interested about a monoclonal antibody Rituximab, I've found pdb structure of Fc and Fab regions of this. Is there any too/method that I can use to append these and get the full pdb structure?
Fc fragment: 1L6X
Fab fragment: 2OSL
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Since you got both regions from PDB and the hinge region could be found at the files shown by Honegger, you may use any homology modeling tool (such as modeller) to create a full protein based on these tridimensional structures and the amino-acid sequence of your antibody.
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I need a software to draw chemical structures. I have been using the free trial version of ChemDraw, which has expired.  Now I am looking for an alternate software that can do chemical drawing. I am not looking for any special or advanced features. I just need a software that can  generate good quality images for use in publications? 
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There's a free online version of ChemDraw, which is kinda cool.
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Simulation has been extended till 70 ns, but still there is no sign for my protein to become stable. What could be the reason?
My protein is a monomer with two helices connected with a middle linker region of 3 amino acids.
Kindly throw some light.
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Few notes...
1. RMSD below one Angstrom is not big.
2. If you increase the y-axis upper limit from one to two Angstroms, the RMSD line will appear less noisy, and your conclusion about the (in)stability of the protein may change.
3. If you decrease the number of y-axis ticks from ten to five, the RMSD line will appear less noisy, and your conclusion about the (in)stability of the protein may change.
4. By plotting moving average, you can smooth out most of the noise, and your conclusion about the (in)stability of the protein may change.
5. Calculate RMSD after aligning different parts of the protein to identify which parts contribute most to the total RMSD value.
6. Some twenty years ago people were able to achieve excellent quantitative agreement with commensurable experimental data using structures and energies obtained from 500 ps (yes, picosecond) MD trajectories.
7. MD simulation is most useful if used for sampling of the configurational space near the initial structure---and for this purpose, in most cases, i would say, the shorter the trajectory, the better; rather than running one long MD simulation, it is generally better to run many short MD simulations, starting from different initial conditions---this is also very useful if one wants to assess the uncertainty of the quantitative results.
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input: amino acid sequence
output: potential protein binding partners to sequence
Thanks for taking the time to address my question.
Bre
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Dear BreAnne,
actually, it's highly difficult to predict an interaction between two proteins because, as said Steingrimur, it might be strongly related to the 3D structure.
However, there is case for which it becomes a bit less difficult: when the protein binds to a linear motif. A motif is a short amino acid sequence (of about 3 to 6 residues). In this case, the interaction is mostly driven by the sequence since the motif is unfolded most of the time.
You may find interesting details at the ELM website (http://elm.eu.org/), which stands for Eukaryotic Linear Motif. It's a manually curated database based on literature. You enter a protein sequence and you'll see all the putative motifs bound to it. Then it's also possible to search for a special motif if you already know it.
Best regards
Yves
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I have about 2000 of proteins ligands from Protein Data Bank and a lot of their characteristics like full name, molecular formula, SMILES, etc. I would like to classify them in several broad groups according to their chemical features. Is there any database with such data?
Thank you!
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Hi, Kirill! you can try fingerprints based clustering, e.g. RDkit for python can do this (http://www.rdkit.org/docs/Cookbook.html#clustering-molecules)
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How should I rename the ‘Zn’ as ‘M’ and provide energy coefficients for Zn in Autodock Tools? I have just started with autodock tools. My protein has Zn ion in it. In the tutorial, it says something about Grid macromolecules.
"ADT also determines the types of atoms in the
macromolecule. AutoDock can accommodate up to 7 atom
types in the macromolecule. It uses a standard set with two
customizable types, ‘X’ and ‘M’. If your macromolecule has a
non-standard atom type, ADT will prompt you to set up a
customizable type X or M for it by entering energy parameters.
For example, Zn is not in the standard set. If your
macromolecule has Zn, for AutoDock you have to rename the
‘Zn’ as ‘M’ and provide energy coefficients for Zn. ‘X’ can
be used as a second customizable type. It is not possible to
have more than 7 types in the macromolecule."
I do not know how to rename and give charge to Zn. Every time I try to save the file as .pdbqt, it gives 0.000 to Zn. [WARNING: These atoms have zero charge: Zn].
I tried using a PDB file editor, but I always do something wrong.
If someone could help in this aspect.
Thank you very much in advance.
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Few tricks be needed to carryout successfully completion.
1) Update the AD4_parameter.dat according to your necessary metal
2) Update the AD4.1_bound.dat also according to your necessary metal
3) then copy that two files into your installation folder
4) then during grid generation and docking dpf file generation steps, there at set map types option you should mention the required metal atom types and also there is a option named as other option; there u should mention the newly generated parameter file
5) finally the newly generated 2 files should be kept on your docking directory
Finally docking should successfully run.
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We are trying to join a dsRNA with a peptide using HyperChem. The moment we are trying to invoke the model builder, the double stranded structure just mashes up in a weird mesh like thing. Whenever we are trying to optimize the geometry using steepest descent or conjugate gradient or Newton-Raphson, a lot of Oxygen appears from nowhere (which do not show up in other visualization tools like PyMol or VMD). And when we are trying to join the peptide without doing the geometry optimization (the most unscientific thing I have ever heard of), the connecting bond just becomes a long stretch of line (which it should not be, because there's a permissible limit of bond length, isn't it???)
Can anybody shed any light on how to do this job in HyperChem??? We are open to other softwares also, if required.
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take a look at this video
best,
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I have a list of protein pairs that interact with each other, which was obtained using yeast-two-hybrid.No 3D-structure or other deeper structural information.
My tutor wants me to find out the protein-protein interaction site of these pairs using some bioinformatic tools.
I know nothing about bioinformatics or structures,so is this possible to do? can anyone recommend some good programs that can predict PPI sites?
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Thank you all for your kindly help!
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Dear researcher,
I want to simulate a selenium containing enzyme. But when i processed the selenium containing enzyme using pdb2gmx command so it is showing an error.
"Atom SE46 in residue CYS 597 was not found in rtp entry CYS with 11 atoms while sorting atoms"
please give me any suggestion how i can add the selenium during MDS.
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You should first check the literature to see if this selenocysteine is actually functional. Sometimes the heavy atom derivative of a side chain (e.g. selenomethionine, selenocysteine) is used to simply get better diffraction of crystals, and the actual enzyme has the normal sulfur-containing amino acids. If that's the case, just replace Se by S and proceed as normal.
If it really is a selenocysteine, you will have to parametrize it. To do this, you must follow the prescribed parametrization procedure of the parent force field. This can be quite laborious to do.
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Hi,
We are trying to draw a charged amine structure in HyperChem. We jotted down the structure as usual but it is showing without any charge.
Anybody has any idea about how it can be done???
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You should be careful with such a big system because the degrees of freedom is too high. Optimization process in Gaussian does not do a conformational search. It only does a single point optimization. Therefore, you should be very sure about the initial guess structure (input structure). Before going for QM, you may try conformational search or simulated annealing to generate a good starting structure.
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I have been reading that it is important for decoys (inactive molecules) to have similar physico-chemical properties to their matching ligands but different topology in virtual screening benchmark data sets
Why is it that? Does topology play a big role in discriminating ligands from decoys? 
What is the role for topolgy in protein-ligand interaction? 
Would you mind giving me a hint or recommend me some material that I could read? 
Thank you so much for all the help that can be provided :)
Hugs,
Stellamaris
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Hi Stellamaris, what I read from your question is that you are describing receptor antagonists, or competitors.
Matching the ligand binding site of a receptor relies on designing an antagonist that looks like a ligand (based on topology and computer docking programs), but  does not act as a natural ligand and thus inhibits the receptor.    
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I isolated my protein complex from mammalian cells. Purify it using affinity purification methods. but after running chromatography the concentration became very low to go further structural study. How can I I increase concentration without affecting the protein complex? 
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@Woojong Lee  I already tried Ultrafiltration using amicon filters according to the guideline. Thank you, I will try it again with more concern. 
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MODELLER website's  "difficult tutorial", available at link below, makes mention and use of "mGenThreader" software. Problem is: provided website by tutorial where mGenThreaded should be doesn't has the option to use mGenThreader, neither I've found any website for which I can download it. Instead of it, I've found "GenThreader" and "pGenThreader".
So I ask: Where Can I download mGenThreader? If this is not possible, may I use pGenThreader in place of mGenThreader?
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Dear Inacio, I think mGenThreader and GenThreader are the same thing: you need to create an alignment file as input and you can do it online: http://bioinf.cs.ucl.ac.uk/psipred/ 
if it doesn't work I suggest you to get in touch with the developer of GenThreader  Prof David Jones at UCL ( d.t.jones@ucl.ac.uk) I hope this help
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Dear researcher,
                           I want to dock a small protein in the interface of diamer protein. When i submit the diamer structure in to HADDDOCK server it terminates the jobs and do not produce the result. But when i delete the one chain of diamer and proceed the docking it accept and produce the result.
So question is:
is there any problem in diamer construction. If yes please suggest me how to create the diamer.
is there any problem in docking then how can i resolve it.
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HI Rohit,
There are other servers like Zdock , ,Hex, Cluspro, Gramm-X , as and alternative there is Megadock4.0 , (it does not have a server you would have to install it ). However, I think they still will have the same problem due to the large size of your dimer.
One of the options you may try is to extract the interface of the protein and use that as the protein receptor in the docking. Your next option is to establish a collaboration with one of the groups so they run their programs locally in their clusters, with the whole protein. 
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I am trying to calculate SASA scoring function using dock 6.7. However, I am confused about the interpretation of the score.
I have obtained:
SA_Descriptor Score: 14.15
How could I interpret it? What do highest  scores mean? What do slowest scores mean?
I would really appreciate all the help that can be provided!
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From the Dock 6 documentation:
"SASA_Score:
#percentage of ligand exposed times 100"
So your SASA_Descriptor Score of 14.15 is telling you that just over 14% of the molecule is exposed to ligand (or that just under 86% is buried in the receptor).
Higher scores mean that more of your molecule is sticking out into water.  Lower scores mean that more of the molecule is buried in the protein.  For drug applications, you'll probably want to go with the lowest score when choosing an ideal molecule to pursue.  Natural ligands lying on surface grooves might have very different priorities when deciding on an optimum pose.  So which score is 'best' will depend upon what question you're asking.
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I have a protein which has two chains each contains 422 residues, both chains are identical. I want to model this protein but the target sequence which I have contains more than 850 residues. When I combined the two chain residues as a single chain and aligned them with the target sequence it got aligned with the whole 850 residues target sequence and showed 56% identity. I want to model this two chains template protein into a single chain using my target sequence. How can i do it? Any suggestions?.
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This is embarrassing. I just realized that I never answered your follow-up question. Unfortunately, even if I gave the best possible answer now, it would most likely prove absolutely useless to you. I apologize.
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Is there a way to choose the site where we want to place the grid box using the command line?
When creating the grid parameter file in ADT, in the graphical interface, you go to Grid ---> Grid Box --> Center. Then you have to chose to center the grid box in either: 1) Pick an atom 2) Center on ligand 3) Center on macromolecule 4) On a named atom. Therefore, it is possible to chose to center the grid box in the ligand.
My question is how can you do it (chose the position of the grid box) with the command line? Is there a script that I could use? There is no option in the grid parameter file. I have more than 10000 compounds and it would take me a really big amount of time to do it graphically. Then, I would like to automatate the process. Is there a way that I can do it?
I am going to do rescoring which means that the grid will not be in the same place of the ligand automatically.
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Dear collegues,
I have received an answer from Proffessor Sanner (Michel Sanner), who works at the Scripps Research Institute. 
I hope it can  also be useful for you all:
"I have been  working in a version of AutoDock that supports making receptor side  chain flexible and as part of these software developments I wrote the  AGFR software program.
This command line program allows the compute grids (using AutoGrid4) and supports a range of options for automatically positioning of docking
boxes, including using: a known ligand, or a set of residues in the
receptor, a binding pocket identified by our pocket finding method
AutoSite, etc...
The software is not 100% ready for release yet but if you want to give
it a shot you can download the code from
please look at the Use cases on that web page for examples on how to use  the software. (Be aware though that currently the graphical user  interface AGFRgui is broken) but the command like version should be working.
The .trg file contains is a zip file that contains all the maps generated by AutoGrid4 and that can be used for AutoDock4.
So you have to unzip the .trg file and give AutoDock4 the the .map files located in the folder created by unzip.
> unzip myfile.trg
Archive: myfile.trg
inflating myfile/rigidReceptor.C.map
inflating .. "
Thank you all for your help! I really appreciate it!
Huge hugs 
Stellamaris
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I've isolated a new drug and would like to see the effect on the murine of S. aureus using Autodock
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I would suggest you to target, cell wall synthesizing protein. 
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For example, it is known that one protein is destroyed in ubiquitin-proteasome system, but the E3 ubiquitin-ligase that performes ubiqutilation of this protein is unknown. How can I predict which E3-ligase ubiqutilates this protein? I can find aminoacid sequence of the protein, I know sites for ubiqutilation in this sequence, I can find also information about secondary and tertiary structure of the protein.
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There several ubiquitin-related databases out there, and the most comprehensive one I know is https:/www.omicstools.com.  It has at least nine tools for use in querying about ubiquitin:
UbiNet:
Offers users an effective platform to efficiently study protein ubiquitylation networks among large-scale ubiquitylation data. The current version of UbiNet was designed specifically for humans to…
E3Net:
Provides a comprehensive collection of available E3-substrate specificities and a systematic framework for the analysis of E3-mediated regulatory networks of diverse cellular functions. Currently,…
mUbiSiDa:
mammalian Ubiquitination Site Database; Provides a scientific community with a comprehensive, freely and high-quality accessible resource of mammalian protein ubiquitination sites. The mUbiSiDa was designed to be a widely used tool for…
PlantsUBQ:
Describes the network of Arabidopsis proteins responsible for the covalent attachment of ubiquitin (Ub).  PlantsUBQ provides comprehensive information on the role of this post-translational…
hUbiquitome:
A public resource for the retrieval of experimentally verified human ubiquitination enzymes and substrates. hUbiquitome is the first comprehensive database of human ubiquitination cascades.…
plantsUPS:
A database of higher plants' ubiquitin 26S/proteasome system (UPS). Both automated search and manual curation were performed in identifying candidate genes. Extensive annotations referring to…
UUCD:
Ubiquitin and Ubiquitin-like Conjugation Database; A family-based database for ubiquitin and ubiquitin-like conjugation, which is one of the most important post-translational modifications responsible for regulating a variety of cellular processes,…
UbiProt:
A knowledge base of ubiquitylated proteins. UbiProt contains retrievable information about overall characteristics of a particular protein, ubiquitylation features, related ubiquitylation and…
SCUD:
Saccharomyces Cerevisiae Ubiquitination Database; A web-based database for the ubiquitination system in Saccharomyces cerevisiae (Baker's yeast). SCUD aims to represent a comprehensive yeast ubiquitination system, and is easily expandable with…
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Hi all,
I am learning a lot of things about the computational biology for the first time. I was curious, with the current technology, what is  the possible simulation time range for a research in computational biology using commercial force fields such as AMBER, GROMACS, CHARMM etc? Can we go more than ns without harming the results? 
Thanks!
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What is "possible" is limited only by the quality of the hardware and software (and how much patience you have for waiting on the results). Modern simulations often go well into the microsecond range pretty easily.
The better question is what is necessary to answer the question(s) at hand, and sampling is a general concern. A single, microsecond-length simulation may be less valuable than, say, 10 x 100-ns simulations, but of course that again depends on the purpose(s) of the simulation. Long simulations have exposed the fact that conformations often get trapped in local minima for long periods of time. This can be circumvented with enhanced sampling techniques or by simply running many more simulations from different starting conformations and/or initial (random) velocities.
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I am doing some researches about the 3D structure and binding site of DHFR, I wanted to have any 3D model for BH2 binding with DHFR. I could just find some models for TH2, but not for BH2.
Could some one provide me any model, or paper where I can find such modeling? it is not bad if there were any clear modeling for TH2 as well.
Thanks in advance!
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PDB codes 1DR1, 1DR3, 1DR4, 1DR6 in th eprotein data bank (rcsb.org) is what you are looking for. Relevant publication is:
Crystal structure of chicken liver dihydrofolate reductase complexed with NADP+ and biopterin.
McTigue, M.A., Davies 2nd., J.F., Kaufman, B.T., Kraut, J.
(1992) Biochemistry 31: 7264-7273
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Hi!
I was wondering if you can suggest program/script for linux or not, that can calculate the interactions between Ligand and protein from a pdb file. When I'm saying interactions i mean, h-bonds, aromatic interactions, hydrophobic bonds etc.
I found IChem on the internet, but it seems that it doesn't work properly. I couldn't even run it.
Thanks
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The Protein-Ligand-Interaction Profiler https://projects.biotec.tu-dresden.de/plip-web/plip/ adds some details that are not covered by HBplus. Besides the Eeb interface, a command-line driven version is available for download.
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Dear CHARMMers,
can anyone tell me what's the issue?
I try to run calculations with charmm for homology protein file, and when it comes to reading psf file (generated with par_app36_prot.prm file), the error occurs:
"Atom 1 0 NH3 problem in psf
***** LEVEL -3 WARNING FROM <psf_read_formatted> *****
***** atom type not found for atom"
Anyone might help to handle it?
Kind regards
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I noticed that you have 19 Atoms in topology file (psf) for the first residue, but in the PDB file you have 18 atoms. There should be the same number, are they both created in CHARMM?
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for protiens
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When you try to chose which software to use for running MD simulations, there are a few criteria that you may prioritize, such as:
1) possibility of modeling different biomolecular systems (e.g., protein, membranes, DNA, RNA and their complexes);
2) possibility on modeling the solvent environment explicitly and/or implicitly (water, ions, etc,);
3) flexibility on using different force fields for parametrization of the systems and efficient treatment of long-range electrostatic interactions;
4) computation efficiency (in terms of speed) for running long time dynamics (e.g., is the code parallelized, can you use GPU, etc,);
5) possibility on running the simulations in different statistical ensembles (e.g., NVT, NPT) and possibility of using different periodic boundary conditions based on the geometry of the simulation box;
6) implementation of stable numerical integrator methods for solving equations of motion;
7) being easy on building up equilibration protocols (e.g., through documentation and tutorials);
8) providing tools and/or modules for analyzing the trajectories;
9) being able to create outputs in a format which can be easy read from other data visualization and/or analyzing software;
10) being able to read and manipulate the inputs structures (e.g., PDB format files);
11) in some cases, being able to compute complex thermodynamic properties, such as free energies, using advanced molecular dynamics methods;
12) implementation of efficient conformation sampling algorithms;
13) and maybe others (depending on the problem).
Some molecular dynamics engines that satisfy the above mentioned criteria include CHARMM, GROMACS, NAMD, AMBER, DESMOND, TINKER, DL_POLY, and maybe others.
I hope that this helps this discussion!
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molecular docking
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thank you madhusudan
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Hi.. I have DNA solutions annealed. I wanna know the effect of pH on the stability of structures with PBS solution. Answers are appreciated. Thanks!!
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Measure the effect of temperature on the UV absorbance of the DNA at different pHs. As the DNA denatures with heating, the UV absorbance increases. This effect is used to measure the melting temperature of the DNA. The equipment required for this experiment is a spectrophotometer with a thermostatted cuvette chamber.
Another way to do the melting experiment is to use a differential scanning calorimeter, which measures the heat absorbed as the DNA melts during heating.
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What is the basic theory of peak shift (Blue or Red) in circular dichroism spectrum of proteins (208,222 nm of alfa-helix or 218nm of betta sheet). and what is the mean of this blue/red shifts? Could scattering lead to any shift in CD spectrum !?
As it is appear in attached figure, all spectrums belongs to a whole alpha-helix protein, but in different condition. in Uv-vis spectroscopy we could interpret the reason of any shift according to chemical environments and its effects on energy level of molecular orbitals n,pi,pi*. Now  I need to know the theory/reason of any shift in CD. Is it similar to UV-vis?  
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Please see
Decreasing intensity and broadening of both the positive and the negative CD bands leads to a shift in the x-axis intersection point towards higher wavelengths, and both bands also exhibit a slight red shift. These changes can be attributed to absorption flattening and differential scattering phenomena. Absorption flattening is a consequence of the non-random distribution of chromophores within the sample, i.e. to differential absorption from different portions of the sample. In peptide-membrane samples it occurs when the peptide chromophores are sequestered in discrete regions of the sample with high local density. The extent of absorption flattening is proportional to the concentration of absorbers in each particle [43]. Thus, with increasing binding, more and more peptides seem to form extended β-pleated aggregates on the bilayer and thereby enhance the absorption flattening effects in the CD spectrum. Differential scattering of the incident radiation becomes more and more important with increasing aggregate size [44]. Thus, the more extended the β-sheets, the more severe is the loss in spectral intensity.
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In my pdb file, there are 6 chain identifier. How can i select 2 identifier using vmd or pymol? then i will do md simulation. I want to take B and F identifier and will run for md simulaiton. I have attached my pdf file. can anyone help me about it ?                                                                                                                                     
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No need to bother with VMD or PyMOL when simple Linux commands will do it.
grep " B " 1brs.pdb > chain_b.pdb
grep " F " 1brs.pdb > chain_f.pdb
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Is it necessary to add missing residues in proteins before performing docking and MD
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First, you need to understand some points like.
  1. Proteins are dynamic in nature.
  2. The tertiary structure of protein is irregular and random and it is fashioned in such manner by many stabilizing forces due to the bonding interactions (hydrophobic and hydrophilic, covalent, salt bridge, ionic etc.) in between the side chain groups of amino acids. These interactions are held responsible for the stability of protein. So now imagine if any of the residue is missing  from its position, what will happen to your protein stability.
So, adding missing residue is essential in Molecular Dynamic Simulations as we are studying the change in structure with respect to time and by change in structure we mean change in interactions too.
But in case of Molecular Docking, story becomes bit different as in docking, we study the interaction in between the ligand and active site of protein,  if your missing residue is present in active site then you must need to add it but if the residue is missing from terminal position then it will not affect the results as in docking the protein is treated as rigid excluding the active site.
You can add missing residues by using spdbv (Swiss-PdbViewer). What you need is just to open your PDB file in spdbv and it will automatically add missing residues and save the file as current layer.  
You can also use Discovery studio by following the clean protein wizard
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I performed simulation of a protein and Protein-RNA complex for a 50ns. I have to interpret all my results RMSD, RMSF, Rg,SASA, binding energy with respect to  statistical error.
in other words,
statistical error in the fluctuations also need to take in to account for the MD analysis.
Could anyone please help me?
thanks in advance
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Did you perform multiple simulations of the same complex? That might be required before you venture into error estimations, because a single simulation can be stuck in one of the minima's. Error estimation can be done by calculating auto-correlation times of your data points and then using uncorrelated data points for bootstrap analysis.
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Hi,
I am trying to visualize my DNA bands run on agarose gels (stained with gel red during the gel preparation). Although I am using UV fluorescence to visualize the gel, I think some setting in my UV machine has been changed because I always obtain images like those attached. The bands of the marker are too thick and bright that I can't distinguish them. On what can this effect depend? What parameter could be involved? I am sure that it is not a problem with the amount of the marker loaded because all my gels are like this.Thanks
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I have found that decreasing the amount of Gel Red used, especially for the ladder, helps get clearer (and straighter) bands. If you are adding the Gel Red to the gel, try instead mixing it in the loading dye instead: 1 uL Gel red in 1 mL loading dye (mix well). 
Also, I think zooming in so that your gel fills up the whole screen will help, as will inverting the images so that you see black bands on a grey background instead of white bands (see attached image. Excuse the primer dimers, this was during optimization). That also uses less ink during printing. The manual for the imager will give tips on how to adjust the image, but play around with the brightness and shading functions until you get an image you are happy with.
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I am working with DNA-protein complex crystallization . I have tried fragments varying in length from 35 bp to 27 bp. DNA is specific in terms of affinity to protein. I am getting crystals with all the fragments but they are not diffracting . They form gelatinous like morphology when disturbed. How to improve for diffraction?
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HI,
Protein-DNA complexes can be tough to crystallize.  The fact that your crystals are becoming jelly-like reminds me of something that happened with my protein-DNA complex.  I worked with a nuclease, and found that cleavage was happening slowly in the crystal over time.  Crystals shot within a few days of formation diffracted well, but those harvested a week or two later were gelatinous (like a gummy candy) and didn't diffract.  This was because the DNA conformation was central to maintaining crystal contacts.  It would be helpful if you could describe the protein-DNA complex you are going after.  I think the first thing you should do is confirm that the protein-DNA complexes are forming in solution before crystallization: Appu's suggestion of using SEC_MALS is an excellent one.  I would also harvest your non-diffractive crystals, and run them on a gel to check if the crystals are your complex, and if your DNA is intact.  Playing with DNA length and sequence at the ends will be helpful, and you may also want to experiment with using sticky ends or backbone modifications that prevent hydrolysis, such as a phosphorothioate.  I added a phosphorothioate modification at each of the 3' and 5' ends: it disrupted a crystal contact for my P21212 crystals but that led to new interactions that ended up improving symmetry and resolution.  I have several colleagues who have found sticky ends or alternate DNA-structures to be helpful.  
Please follow-up to let us know if our suggestions help, or to provide some information we can use to help you.  Hang in there: I had a colleague who used more than 35 different Holliday junction substrates before he found the right one.
Happy Crystal Hunting!
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Hello! Anybody knows how to perform molecular replacement using all protein from the PDB as searching templates? which software can perform this function, or we write scripts ourselves?  In addition, i don't good at computer science.
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Dear Liu-Quing
We're actually working on a server to do exactly this, based on the experience reported in this paper
It will probably be called SIMBAD and will be hosted at CCP4 online.  Watch the CCP4 bulletin board for an announcement. If your case is urgent then you can contact us by e-mail.
Regards
Daniel Rigden
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I want to tag a protein molecule with a dummy hard sphere containing some charge or a point dipole instead of that sphere. I don't know if it can be done in GROMACS / NAMD packages. 
Is there any way to achieve this? 
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One way to do that would be to introduce a modified residue in the .rtp database in Gromacs files, prior to running pdb2gmx - after copying and renaming the residue you can add an atom, specify its charge and type and add a bond to any existing atom in that residue. (Note that you might need to edit several additional files, including .hdb and ffbonded.itp if you introduce a novel dummy type - this might be laborious, but offers the most flexibility.)
Another way is to use the pull code (see the Gromacs manual) to restrain the distance between an ion (or a dipole) and any desired group of atoms in your protein. It would be slightly harder, however, to restrain e.g. the orientation of the dipole (PLUMED might come in handy if the problem is relevant to your case).
The simplest approach would probably be to just edit the topology and add the charge to an existing atom in your protein. Here, though, a large charge might locally distort the geometry, especially if the modified atom has a small vdW radius.
It's hard to recommend any specific approach without knowing what your specific research question is, but at least one of these might do the job.
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I am trying to identify critical residues in my protein, that are responsible for its endonuclease activity. I have been having trouble purifying the protein so identifying the residues in vitro, by mutating each of the suspected residues, purifying each mutant protein and assaying it for its activity might not be possible at the moment. I was wondering if there is any way to predict the 3D structure of the protein, mutate all the shortlisted residues, and study DNA binding in silico, which might give some clues on the critical residues. 
Thank you!
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The best tools to use depends on whether the structures of related proteins are known. Do a homology search (blast/fastA) with your sequence on the PDB (http://www.rcsb.org/pdb/search/advSearch.do?search=new), if you find significant hits  over a fair length of your protein, you can use sites like SwissModel (https://swissmodel.expasy.org) to get a homology model of your protein. Also look at an alignment of your template structures, there may be structures of complexes that give you a hint what the active site may be. 
Also look carefully at a multiple sequence alignment of homologous proteins in other organisms - the sequence variability can give you hints of which positions are important for structure/function, and which ones are variable
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I wish to know in what terms can we define this sort of graph or diagram?
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  Before understandind Ramachandran plot It is very  important to  understand the structure of a peptide bond which is rigid and planar in nature .Simply   A  Ramachandran plot  is a  plot  to visualize energetically allowed regions for a polypeptide backbone torsion  angles  psi (ψ ) against (phi)  φ of amino acid residues  present in a  protein structure.It is used to analyze  the structure of  a protein , the conformation of the amino acid present in the protein  and close contacts between the atoms
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Hello,
I am solvating a short peptide with a dodecahedral box using the following GROMACS 4.6 commands:
pdb2gmx_mpi -ignh -f $1.pdb -o $1.gro -p $1.top -water spce -ff oplsaa
editconf_mpi -f $1.gro -o $1-BOX.gro -bt dodecahedron -d 2.5 -c
genbox_mpi -cp $1-BOX.gro -cs spc216.gro -p $1.top -o solv.gro
where $1 is the root name of my starting pdb
When I open solv.gro in Chimera or VMD I see a box that appears to be rectangular and the peptide is not in the center even though I used the -c flag in editconf.
I am wondering if this is normal or if I should do something differently with my setup.
Thanks!
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A dodecahedral box is a more efficient one because it adds fewer waters to maintain the same periodic distance. This is merely a visualization inconvenience and there is no need to change the approach taken.
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  • Online tools for secondary structure prediction only for soluble protein (not membrane protein)
  • Please provide some online servers/tools.
Thank you.
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Thank you everyone. 
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I am looking at  B-factors in two different PDB protein structures and I wonder if these B-factors are suitable for direct inter-PDBs comparisons, say to tell whether a particular loop in  protein A is more flexible than another loop  in protein B . Thanks in advance for your help.
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Hi Pedro,
Could you please specify which two structures are you comparing? I mean, are they two unrelated proteins (low structural homology), or are they different versions of the same protein (for example, an holoprotein, and its complexed counterpart)?
B-factors measure atom mobility inside a 3-D dynamic structure. They are related to atomic displacement, but also to resolution, so an atom's B-factor may be high not only because of high mobility, but also because of low structure resolution (if atom position was wrongly placed when inferring structure). Unless you have a high resolution structure, you usually don't know if your atom's B-factor is high due to high mobility, or due to poor resolution. That is why, if you have structures with different resolutions, I definitely unrecommend you comparing their B-factors. B-factor comparisons may be very tricky, so you should be very careful when interpreting the results.
But I could think of a situation in which B-factor comparison may be valuable: when comparing different states from the same protein (complexed-uncomplexed, for example). Even in this case, I would normalize the B-factors prior to comparison. In this case, you could identify protein regions whose mobility is reduced/increased because of structural changes, like when a holoprotein binds to a ligand.
I hope this helps.
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A protein pair (A1-B1) is known to strongly interact, and shows highly complementary interfaces (ie. one interface is mainly positive, the other mainly negatively-charged). On the other hand, a homologous protein pair (A2-B2) shows slightly less complementary interfaces (by visual inspection). What I want is to compare (A1-B1) and (A2-B2) in terms of this electrostatic/interfacial feature.
The APBS .log output file gives us atom-by-atom energy measures, as shown below:
Atom 0: 1.154954531700E+01 kJ/mol
Atom 1: -7.142811088081E-01 kJ/mol
Atom 2: 7.929991927597E+01 kJ/mol
Atom 3: 2.912266331817E+01 kJ/mol
Atom 4: 3.978532290650E+01 kJ/mol
Atom 5: 2.112787418063E+02 kJ/mol
(...)
Atom 5502: 6.670881588625E-01 kJ/mol
Is there any way to use such energy values to come out with a single measure to express how complementary the PPI interfaces are?
Best wishes,
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Hi,
Thank you very much for the information, Avinash.
Do you know any paper that has used the aforementioned strategy to analyse PPI electrostatic complementarity?
Could you please send us a link for such articles?
Best wishes,
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Speaking of radiation damage in the Cryo-EM, I dont understand the concept behind this dose rate.
For example in this line "The total exposure time for producing one image composed of 70 frames was 14s and required a dose rate of 2 e-/Å2/sec"
Can some one explain this one image of 70 frames and this dose rate.
For X-ray diffraction, a larger crystal can tolerate a high total dose and often diffracts to high resolution because more molecules contribute to the diffraction. For single-particle cryo-EM, the total electron dose used to image each molecule is set to a very low level to preserve structural information at the subnanometer-resolution level. The consequence of such low-dose imaging is that individual images have a very poor signal-to-noise ratio (SNR).
In the above lines very low level of electron dose here is defining the time limit or what?
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Dear Aamir,
as you mentioned one is trying to reduce radiation damage by imaging biological samples under low dose conditions.
The new DEDs as the Gatan K2 or the FEI Falcon II or III offer a very fast read out. 
That means that one can collect "movies" of a target composed of for example 70 frames / individual Images. These Images can now be summed up and corrected for beam induced motion. This summ of images is then treated as an individual images (one image , 70 frames). 5 frames were taken in 1 sec. The dose rate was set to  2 e-/Å2/sec, means in 1 sec 2e- are detected per Å2 of the detector . The summed Images suffered from a electrone dose of 28 e-/Å2.
If one later realizes that the beam damage is to high one can kick out the late frames and reduce the total dose.
Generally spoken a higher dose leads to better contrast but destroys the specimen. 
Best
Thiemo
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I am working with a membrane protein and crystallographic structure for the protein is not available yet. And am trying to predict the structure using in-silico methods and have predicted using Phyre2, I-TASSER, Robetta, etc. I want to develop a membrane model simulation, to proceed further with prediction. Could anyone help me in this and tell me some free tools or software for predicting the same?
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Materials Studio is a good software for model building, and lamps is a good software for running simulations. This software might be helpful. 
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I have some oligos (20-25 mer) with similarities to known G-quadruplex DNA forms. I want to build 3-D models of these oligos using the known ones as templates as in homology modeling. Is it possible? Can anybody guide me to relevant tools/references?
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Can you give me some parameter to adjust this structure on UNAFold web server? I tried many times but it's cannot for G-quadruplex.
Thank you
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I represent the protein structure in torsion-angle system(using torsion-angles phi+psi+w),during the simulation, I need the Cartesian coordinates of all heavy atoms in the backbone, how can I calculate?
thank you!
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Thanks a lot!
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Is there any method or tool to calculate total net charge and hydrophobicity of a given protein?I am trying to establish a link between protein disorder and net charge & hydrophobicity
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To calculate charge in a folded protein, you have to take into account the influence of neighboring charges on the pK values of titrable groups. The Adaptive Poisson-Boltzmann Solver (APBS) is a software package for modeling biomolecular solvation by solving the Poisson-Boltzmann equation (PBE). The PBE is a popular continuum model used to describe electrostatic interactions between solutes in salty, aqueous media.
http://www.poissonboltzmann.org. Work through the explanations, examples and tutorials on this site to understand what you are doing! Naccess ( http://www.bioinf.manchester.ac.uk/naccess/ ) can help you break down the solvent accessible surface of a folded protein by hydrophobic and hydrophilic contributions. Of course, protein folding and stability as well as protein aggregation are not directly governed by the average charge and hydrophobicity, the spacial distribution of charges and accessible hydrophobic patches play an important role!