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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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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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When observing elastic modes of proteins, one of the results files shows deformation energy plot. What is the significance of deformation energy when studying protein dynamics?
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Deformation energy measures the rigidity of protein residues. Higher the Deformation energy, higher the rigidity residues have. Also, it gives idea about protein's local flexibility.
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I am currently working on a project on cryoEM single particle analysis of protein complexes. Though using HPLC size chromotography can separate the complexes and individual subunits, the structure is not homogeneous tested by negative staining. It might be due to the buffer used. However, how to choose the buffer? and how to access the stability of the protein complexes? Any suggestion is welcome.
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Unfortunately, things like glycerol or other additive that may stabilize a protein complex would add background to the images. A diagnosis for assessing the apparent heterogeneity stemming from the instability of the protein complex, not due to insufficient puification, is to check the distribultion from your 2D class-average images. However, since Relion has attractor effect---your complex vs complex minus a small subunit can be merged into the same class, we thus recommend ISAC or IMAGIC to do so. By the way, negative stain is tricky because uranyl acetate is acid (uranyl formate is neutral but the contrast is lower) and the supporting carbon would do something adverse. We will update you with other approaches that we are testing regarding this critical issues soon.
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Hello everyone,
I have a protein system (complex with a substrate) with a total charge of -9 (the charge of the ligand is -5 and the protein is -4). I want to do some QM/MM calculations using PM6 method in Gaussian.
Here are my questions and any help would be greatly appreciated:
1- I need to know whether I should neutralize the system before running the calculations or not.
2- And if so, can I add counterions manually without using any scripts?
or 3- Do I have to use a specific software for that?
P.S.: I am not interested in using AMBER to add those counterions, since I don't have the AMBER parameters for my ligands.
Thanks a lot in advance.
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Thanks so much Stanislav!
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Hi,
We were carrying out in vacuo energy minimization studies of a protein dimer (which is experimentally proven to be a dimer). Earlier, the same work has been done in our lab using an older version of GROMACS (4.5.5) and used Group cutoff schemes with coulomb type= cutoff and with no pbc.
When we reinitiated the work again and have to use the Gromacs 5.0.4, the default cutoff scheme is changed to Verlet. We are observing that using Verlet cutoff scheme, the monomers dissociate from each other which is not the case even in this version when using Group cutoff scheme.
I searched for literatures and found out the differences are probably in the pairlist generation. In my graduate courses, I have read about energy drift in molecular dynamics simulation and is aware (though not in details) that Verlet algorithm has something to do with it.
Can anyone elucidate on this problem? The minimization runs fine and the protein remains dimerized when using Group cutoff. This happens even after solvation. We have used an xyz pbc and grid neighbour searching type with default fourier spacing and rlist as we have not mentioned the last two parameters explicitly in the mdp file.
I want to know the theory in play behind this. Please help.
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I believe the answer to this question is covered here:
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I want to do normal mode analysis through ProDy i am not able to install on window platform please provide me step bu step method to install ProDy. I have tried website method it is not working in my case. i am not able to give python script path and prody path  
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Akhil Kumar, this is not surprising. ProDy often generates results for Calpha atoms and I imagine Bio3D would too. I guess you've found a way to deal with this my now, but here's some advice in case anyone else looks at this.
You can make the dots more visible by showing them as spheres. I assume the fluctuations are written into the B-factor column so should can use the spectrum command in PyMOL to colour by them e.g.:
spectrum b, rainbow
I'm not sure what the R-defor.pdb would give as I haven't used Bio3D, but I'd guess it gives a set of frames to visualise the deformation vector, which you can show by looping through states in PyMOL by pressing the next, previous and play buttons at the bottom right. You can also make this nicer with the mset command.
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How to generate dihedral constraints in gromacs for all protein?
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Hi, how did you freeze angles and dihedrals in gromacs using the freeze option?
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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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Is it valid to compare the wild type protein against the protein that has been truncated due to stop codon mutation using molecular dynamics simulations?
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As far as i think, if the truncation is towards the end terminals and if those terminals do not lie in the active site then possibly we can do te md. But i need more justification to this. Lets see if someone on RG answere this.
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Intermediates are very important and often found in protein unfolding or foldingbpath. Sometimes, the intermediates are characterized as molten globule states (native like secondary structure and partially perturbed tertiary structure), and pre molten globule states (probably intermediate state between molten globule and unfolded state). I have seen report to use the term highly ordered molten globule description for a-lactalbumin.
My question is if there is any attempt to categorise the intermediate states observed in the protein unfolding?
Secondly, I am unclear of the definitions of this molten globule or pre molten globule state.
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This is very extensive topic to answwe. There are many papers on molten globule state by Prof. Kunihiro kuwajima and others check on pubmed.
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hey, i am here with two basic questions:
I want to perform some analysis on monomers of pentameric protein. There is HSD residue in my protein but when i am performing hydrophobicity related analysis on it, i am getting error for the particular protonation state : like there is no hydrophobicity entry for HSD. is it possible to rename HSD residue to HIS in .XTC and .tpr files?
my second query is i have to select monomer and lets say run g_rmsf on it. How can i rewrite a trajectory with specific atom residues (monomer only) here? I didnt add chain identifiers to pdb before simulation ( A B C D).
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First question:
You cannot rename a residue in the trajectory as the topology is saved in the topology file your provided to the gmx tool, e.g. the .pdb file or the .top file. You just need to put these topology file in the commend line to let the gmx tool to read it.
Second one:
As the first answer writes, you can make an index file and use thr trjconv tool of gmx to extract the specific trajectory for the selected residues. The make_ndx command and trjconv command in gmx could do this.
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When I load repeat simulation of my mutated protein from 10ns-20ns-30ns, its RMSD graph is in picture. I watched my dcd, my protein goes out of the water box. I tried to put it inside of the box with "pbc wrap -centersel "protein" -center com -compound residue -all" code. The protein entered the box but the RMSD values doesn't change. How can I solve this problem?
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I asumed you use Gromacs package for rmsd calculation.
So first you should remove all pbc issue in your trajectory file. Try following steps:
1. If your system contains protein+ligand, make an Index file having group of protein+ligand
2. gmx trjconv -f xxx.xtc -s xxx.tpr -n index.ndx -o xxx_nowater.xtc -pbc cluster
select group number of protein+ligand.
3. now calculate rmsd using xxx_nowater.xtc
Hope it will work
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Hi All, I am trying to simulate one DMPG lipid bilayer in interaction with some fusion peptides. First I am preparing the membrane, and I am facing the problem: How to enable the NAMD to use NBFix parameters for SOD ions? Does anyone know how to set up this using NAMD and Charmm 36 FF?
Thanks in advance!
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Hi, now do you know how to add NB-fix or change it in the NAMD simulation? I am faceing the similar problem as you? Thanks a lot!
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I'm currently trying to label human p53 with small organic dyes (Alexa and/or Atto dyes). Does anyone have experience with labeling p53 with these dyes, and maybe would be able to suggest the best approach? I think one common way is to link these to Thiols, but I noticed a lot of cysteines are in the DNA binding domain and this might affect activity.
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Found this which might help:
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I am trying to do PCA from the MD trajectory of a multi-domain protein using GROMACS tools.
I am using gmx covar and gmx anaeig for PCA. My protein has many domains, but I would like to focus on only two particular domains. Thus, while doing gmx covar, for lsqft I am using protein-H as the group and for covariance analysis I am using index for one particular domain. Similarly for gmx anaeig, I am focusing only on that particular domain and writing as output (the filter.pdb) for only that domain.
I am not sure it this is physical and correct. In short my query is can I calculate PCA (covariance matrix) only for a particular domain of a protein? or should I always do PCA for the entire protein?
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This is perfectly physical and sound to analyze only a part of a protein. Keep in mind though that your extracted data should preserve variance that you are looking to characterize. For instance, if you intend to study how a part of a protein (e.g., a helix or a loop) moves with respect to the rest of the protein, then by extracting only coordinates related to this particular part would result in a complete loss of the information you are interested in.
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Is there any software avail to do nanoparticles with protein molecules...
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this article will help you so much i was search about the same question and this contain an informative information
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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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I have trajectories from Amber in netCDF format (.nc file). I wonder if there is a tool for general rotamer transitions? it will be a lot of labor to make code for each amino acid torsional angles one by one and then annotate it by rotamer library.
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Wow, I had to solve that one myself. So here it is:
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Hi Folks,
I am doing an MD of a protein-ligand complex. But during the simulation, the ligand passed the PBC wall and entered another PBC box. Anyone know how to process the trajectory to restore the ligand to original position?
I know Gromacs has such function:
trjconv -s ... -f ... -pbc whole -center
But how to do it in NAMD?
Thanks!
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Hello everyone,
Does anyone know why we need to fix the pbc in our analysis?
Do I also need to fix the pbc for my contactmap analysis?
Best wishes,
Feng
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So i have my two proteins and a protein linker all in PDB format. I need to combine them in as precise as possible way. What software would you suggest me to use? The orientation is not the most important part, i can trial and error until my orientation satisfies my parameters as i am doing corse grain simulations.
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Suggest me LAMMPS tutorial with commands exclusively for the protein dynamics. i have found out some document but that is not exclusively having the commands for the protein dynamics as like in GROMACS. Please suggest me some link or document.
Thank you....
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If you download the LAMMPS source code, the "examples" directory within the LAMMPS folder has one called "peptide" which might be helpful to you. Among the files in this directory you should find "data.peptide" and "in.peptide". "data.peptide" is a LAMMPS data file which contains all the information about the atom positions, bond parameters, etc... In GROMACS terminology this file is kind of like the combination of a .gro and a .top file. The "in.peptide" file contains all of the parameters directing how to perform the MD run. In GROMACS terminology it is kind of like the .mdp file. If you play around with this example and read up about it in the manual (http://lammps.sandia.gov/doc/Manual.html), this should help you get started.
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hello
kindly tell me what is the significance of B-factor and mode shape in protein dynamic study. And what are the parameters to know which is the best score for both of these .
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I have simulated a DNA-Protein Complex structure with 150mM NACL salt concentration for 100 nanoseconds. The simulation is completed with not much problem, but while visual analysis in VMD of the trajectory, we found the DNA is wavering a lot, even more than protein. Especially 3 nucleotides from one end, which come apart and distort the B-DNA structure. Usually, terminal residues of DNA wavers but in my case, the DNA structure is distorting.  What could be the possible reason for such behavior of DNA?
I used AVOGADRO to generate the B-type 12 nucleotide double stranded DNA. The sequence is ATATATATATAT.
I used HADDOCK to dock the protein and DNA, based on some experimental data for the interface.
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parmbsc1 is an improved DNA parameters but not for protein. You can either use amber14sb or amber99sb. But I cannot see a good reason why these versions of amber will perform bad/differently to charmm36. Terminal base pairs are always flexible. I would suggest extending the length of the DNA by 3-5 base pairs to mitigate there effect on core DNA behavior. Can you post a vmd snapshot of the final structure?
before starting with trjconv, first make sure that your reference structure (must be tpr as Justin has suggested, tpr is required for -pbc mol) is also not broken (pbc fixed). I recommend using tpr used for energy minimization. Dump the gro/pdb to ensure that the protein-dna complex is whole. A possible work flow...
gmx trjcat prun.xtc -n index.ndx -o protein-dna.xtc (removing water first assuming water is not of interest)
gmx trjconv -s protein-dna.xtc -pbc mol -o nojump-protein-dna.xtc -s em-protein-dna.tpr
or
gmx trjconv -s protein-dna.xtc -pbc nojump -o nojump-protein-dna.xtc -s em-protein-dna.tpr
---that can fix most issues, if not then go with this
step1: gmx trjconv -s protein-dna.xtc -pbc whole -o whole-protein-dna.xtc -s em-protein-dna.tpr
step2: gmx trjconv -s whole-protein-dna.xtc -pbc cluster -o cluster-protein-dna.xtc -s em-protein-dna.tpr
output of step2 should fix even the most crazy pbc issues....
##### RMSD calculation for dna-peptide
better calculate the rmsd for protein and dna separately. Dump an index with protein backbone and dna-backbone and then use gmx rms
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Can we observe the effect of the ligand on the protein through RMSD, RMSF and radius of gyration graphs? Does it induce any change in the protein?
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gmx rms, gmx rmsf, gmx gyrate. GROMACS also proposes the group modules which can be helpful in grouping atoms of the system for specific analysis for example when it comes to measuring distances.
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Hi All, I started simulation of reverse micelles in gromacs. After simulation I tried to visualise my system in VMD (it look like fig min.RM.png), to visualise properly I tried trjconv tool in gromacs but unable to relocate with in box. So I check with starting system, my initial system are not in the  center to box (see fig: pack.png)
Is there any way to visualise my system
Thanks
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Trajectory files need some process before using to extract information.
Trajconv is used as an editing tool for the trajectory file to correct periodicity or modify the trajectory file manually like time unites,frame frequance, etc). Through the MD , the protein diffused in the unit cell so it sometimes jump from one side of the unite cell to the other side. So the following commands used to correct this:
gmx trjconv -f md_0_50_raw.xtc -s md_0_50.gro -o md_nojump.xtc -center -pbc nojump -n index.ndx
then type 0 | 0 ,
gmx trjconv -f md_nojump.xtc -s md_0_50.tpr -o md_0_50_processed.xtc -center -pbc mol -ur compact -n index.ndx
22 | 0
22(represent the protein option)
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Visualizing protein folding in gromacs
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Yes, DSSP tool is in gromacs but don't know is it gonna work in your PC or not. In my case, it is installed in super computer facility, so I use it without any problem. When I try to use it in my PC it shows some kinda error related to executable file.
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I am trying to simulate do a Umbrella sampling simulation in GROMACS. I have the starting structure from XRD. The ligand sits almost at the middle of the lipid bilayer. I want to generate the initial  configuration for performing US. I am stuck with what I should give as the reference so that the ligand traverses from the extracellular side to the intracellular side. 
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You have to define 2 groups f.ex center of mass of the bilayer and ligand. If you perform, pulling simulation, then ligands starts moving. It is easy to extract windows by means of a script for the umbrella sampling simulation. https://www.researchgate.net/publication/315788220_Phosphatidylserine_flip-flop_induced_by_oxidation_of_the_plasma_membrane_A_better_insight_by_atomic_scale_modeling
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Hello all,
I would like to know how replica exchange umbrella sampling methods works , and how can i start by using plumed plugin to gromacs.
Thanks in advance,
Sincerely,
ANJI BABU
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thank you
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I am simulating Aβ peptide (2BEG) with inhibitor, how to add molecules at random position prior to solvation, which was done by lemkul in his study.
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 The easiest way to do that is to use packmol.
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Removed Question as account inactive and questions cannot be deleted
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Is xray data available? I assume you want to align the MD density to the crystal data. To do that you need to superpose your model onto the Xray model, then calculate Fmod using the superposed MD model in the unit cell of the xray data. With two isomorphous data sets you can inspect the density easily using Coot. You can create a difference Fourier using CNS tools or Phenix.
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I am working on a protein whose structure is determined from NMR and its in pentameric state.
We believe that It should exist as tetramer in real life.
Is it possible to model the tetramer from  Pentameric NMR structure?
If so, please let me know how to proceed?
Thanks in advance
P. Saravanan
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Could you try the Rosetta symmetry module (https://www.rosettacommons.org/manuals/rosetta3.1_user_guide/symmetry.html)? If you know what sort of symmetry the tetramer has, you could define that symmetry in the module and then include intramolecular constraints taken from pentameric structure. That way a symmetric tetramer is built to be as much like the pentameric structure as possible. Rosetta also apparently has symmetric docking (but I have not used this feature) so you could try docking the pentamer monomers into tetramer symmetry.
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I am trying to do protein folding studies for approx 70aa length protein using Gromacs. However, when I perform simulations, my protein goes out of the box and interacts with its images when simulated at 1.00nm box.
Can anyone please guide me as to how to calculate (if there is a formula or so) how big the PBC box should be such that no interactions are allowed between the periodic images?
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You should determine the water box sizes based on the protein unfolded structure, using for example one of the unfolded configurations. Based on this unfolded structure, you can determine (x_max, x_min), (y_max, y_min) and (z_max,z_min) of protein. Then, construct a water layer around the protein with thickness at least the cutoff of non-bonded interactions size: let's say if the cutoff=11-12 Angstrom, then the thickness of the layer is 15 Angstrom (assuming that the water box may shrink a little if the water box is not initially equilibrated at normal conditions, i.e., density = 1 gr/cm^3). Then, the big water box sizes are defined as:
big_sizeX = x_max-x_min + 2*thickness
big_sizeY = y_max-y_min + 2*thickness
big_sizeZ = z_max-z_min + 2*thickness
Then, after building a water box with these dimensions (big_sizeX, big_sizeY, big_sizeZ), you can solvate the folded protein structure in this 'big' water box. With this, then, you are sure that even if the protein gets unfolded, the images will not see each other.
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Hi everybody,
I would like to pull a ligand in a channel from my protein and calculate the PMF with Umbrella sampling. This channel is probably not linear and thus have to be refined. Too much colisions appears along a simple one axis pulling and causing a catastrophic PMF calculation.
Currently I'm working with Gromacs and I wondered if it is possible to define statically or dynamically the coordinates for the center of the channel and use it during the simulation (Pulling).
One solution could be to define in the md_pull.mdp configuration file all the transition points of my pathway through which my ligand must pass from my APO structure determined with CAVER.
Instantiation of Gromacs parameters of md_pull.mdp is not enough clear for me and PLUMED is may be a more direct solution. Currently I have no experience with PLUMED but It seems on paper to be able.
An other choice would be to define dynamically the channel center, on-the-fly. PLUMED seems also capable to manage this sort of challenge but once again I have no idea of the feasibility of a such work.
What do you think about this problem and of the tracks evoked above ? 
Cheers, FR.
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Hi Francois
None of the  magic recipes proposed with a lot of hype for calculating free energies of
such complex systems from  canned MD like Gromacs,Charmm and the like do not
survive a true statistical  physics examination  because concepts like entropies,chemical
and potentials are not quantities derived from Newton's laws which deliver only mechanical
quantities and time averages of them .In simple systems one ASSUMES that these averages are ergodic (strictly proved only for a system of hard spheres by Ya Sinai)  but
even this is not true for a protein-solvent-ligand system  with multiple time scales ,strong
long range electrostatic interactions and  a solvent like water.The major problem is not the
inaccuracy of the   force fields but the very nature of the quantity you want to estimate.It
would much better do  Monte Carlo Simulation but this is much more difficult for your system so nobody suggests or develops such and hides the garbage under the carpet in the name of quick success.A more modest approach would be to search for clever
approximate and  softer modelling combining physics geometry and experimental data.
Of cource it is always possible to believe in MacMD and run everything,  this is usefull
for many things and I have done it for decades myself but I  don't believe the free energy
results for systems like yours.
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I have seen many papers on folding and unfolding of peptides using Replica exchange with solute tempering (REST). How do we implement this method in GROMACS > 5.1?
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You might want to refer to the two papers linked below.
The first one tells you how to scale your force field parameters by specific factors; basically, you'll have to make local copies of your force field files and write a script to scale the parameters for protein atom types.
The second presents HREX, a patched version of Gromacs (AFAIK limited to 4.6, so it might not be possible to run REST in gmx > 5.1) that allows to perform replica exchange with arbitrary energy functions; from what I recall, the problem is that standard Gromacs has replex limited to the lambda code, so this workaround was necessary.
I assume you're fluent with Gromacs and scripting - setting up REST might not be as straightforward as, say, running standard REMD.
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Calculating Protein-lingand interation using MM/GBSA and MM/PBSA
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Dear Shadrack
All the best
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Hello, everyone!
    Now I'm studying on the MD simulation of interface diffusion. After I got the trajectory of the atoms, I didn't find the solution to obtain the atoms' RMSD along the z direction(vertical to the interface). The software I used is VMD. I want to know if there're some skill to deal with the problem.
Thanks a lot
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When you use the formula for RMSD, instead of using the vector position r_i, use its projection along the direction you are interested (e.g., any direction n), so that in formula of rmsd, you replace r_i with
z_i = dot_product(r_i, n)
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Hi,
I want to perform molecular dynamics simulation of protein-RNA complex at deifferent pH condition in gromacs. 
When I am using Amber force field, It is not detecting protonated residues while OPLSaa force field is not detecting nucleotides. 
Is there any other software available to perform this simulation??
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Hi, Karolina.
I want to perform the simulation of protein-RNA complexes at different-2 pH conditions. 
I used OPLSaa force field to do that for protein alone, but when I am doing that for protein-RNA complexes, it is showing errors.
Amber94 is asking me to delete all extra hydrogen atoms, which I have added during protonation of protein wherever OPLSaa is not detecting nucleotides. 
 So, How can I  do that without modifying force field? 
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 I am using this command in cpptraj
rms ToFirst :1-81&!@CA,C,N,O*= first out rmsd1.agr mass
Plot is generate using  number of frames.How to plot rmsd of amber trajectories in time unit  ns using cpptraj ?
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Hi,
you could try the "time" keyword in rms command.
I don't know how many frames there are in your trajectory, but assuming you are saving your coordinates after each 1 ps, you could try typing:
rms ToFirst :1-81&!@CA,C,N,O*= first out rmsd1.agr time 0.001 mass
Hope that helped.
Karolina
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Dear all, 
which method would you prefer to determine changes in protein tertiary structure (no alteration in secondary structure)?
I don't have the possibility for NMR, X-ray or SAXS.
Thanks =)
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The large change in shape should be detectable by gel filtration chromatography and analytical ultracentrifugation, as well.
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I need to study the effect of PTMs on the biophysical properties of microtubules. I have tried PyTMs, but it has limited features. Even if we just modify the PDB file and later equilibrate the system by simulating it using GROMACS, it would be great.
Looking forward to reply soon, please!
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I agree with Tanuj Sharma. There are some software, such as Discovery Studio, Chimera, and others, in which you can add modifications at protein terminals. However, if you want to see the modification happening, it is needed to use a quantum chemistry method.
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A total open question. How would one go about linking a function to a protein given only its structure? 
I understand this is a million-dollar question, so vagueness is appreciated. 
I have a predicted structure of a protein from de-novo/threading (I-TASSER) followed by extensive simulated annealing MD simulations. There is only DNA work (sequencing), no actual protein experimental work - nothing at all, this is something knew we found.  There are also no template structures, i.e., homology modelling efforts have been awful. 
Some thoughts I've had:
Are there di-cysteine bonds? Typically extra-cellular proteins have these over intra-cellular proteins. 
What is the secondary structure content? i.e., helical vs beta sheet with regards to transmembrane proteins
What is the tertiary structure? Globular, fibrillar, etc.
Are their stretches of exposed hydrophobic residues? i.e., integral membrane proteins.
Are there are metal binding sites e.g., 3x HIS 1 ASP which is very typical for zinc?
Stretches or grooves for protein-protein interaction and/or potential hydrogen bonding?
Given that I have a structure, is there any way of comparing this to 3D structures in the PDB databank?  Having said that, if there was, and a match was found, you could safely assume the homology modelling would have worked to begin with.
I appreciate any thoughts you have.
Thanks 
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I am running a MDS using Gromacs ver. 5.0.2 on a 42 residue peptide with a computer of 24 CPUs and one GPU. The GPU performance is always around 70%. Is there any way to improve this situation?
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you can also utilize both cpu and gpu while performing simulations using command
gmx mdrun -v -s md.tpr [tpr file] -c md.pdb [output pdb] -nb cpu gpu
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I am trying to perform a PCA analysis using side-chain heavy atoms, instead of the conventional CA-based PCA. The question is in that case, do I need to fit the trajectory on a side chain reference or I need to do the fitting again on the backbone. My protein is rigid but interface residues adopt different rotameric states upon binding with different partners and my goal is to quantify these states through PCA to remove the noise associated with the MD trajectory. It will be great if someone can suggest some tools as the tools I am using (bio3d, Wordom) somehow gave different results...Thanks
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Yes, we have implemented in CHARMM program (www.charmm.org).
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Hi everyone, I am using gromacs 5.1 for COM pulling of the ligand from a lignad-protein complex. I am getting an abrupt fluctuation in the position of the ligand near 110ns and 270 ns (as you can see in the attached figure) which should not be there. So my question is, What is the possible reason that leads to fluctuation and how it can be fixed?
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Why shouldn't those increases be there? Nothing seems abnormal to me. Watch the trajectory and see what is happening on those frames. Probably a side chain is flipping out of the way or something, allowing for a quicker movement of the ligand.
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I want to make h-bond analysis to show a formation and movement of an interface between two molecules (protein&RNA).  
The common h-bond analysis gives me only a duration of it's life. But I need time frames, a plot of their durations on the time axis with points of formation and breaking of every bond.
 How can I do this? Help me, please! 
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1. Make a list of all possible hydrogen bonds.
2. Calculate distances and angles for all possible hydrogen bonds for each frame.
3. Determine the presence/absence of each hydrogen bond in every frame using cutoff values for the distances and angles.
4. Statistics, plots . . . 
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Hi,
   I was trying to run heating of a protein keeping all protein heavy atom fixed in its' crystal structure position using GROMACS.I found that gmx genrestr helps to make a .itp file which contain list of atoms to be restrained with certain force constant values but even if I apply high force constant value (e.g 1000 kcal/mol/A^2) to the heavy protein atoms rmsd calculation on obtained trajectory shows that protein havy atoms fluctuate.How can I completely cease protein heavy atom motion in constrained MD simulation in GROMACS ?
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1. Create a group of atoms in index.ndx file that you want to keep frozen
2. use freezegrps in your mdp file.
That should help. Positions of the atoms from this group will be frozen during simulation.
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I want to simulate a RNA. It has GTP as it's 5' capping.
I want to simulate it gromacs in gromacs using AMBER99ff.
Can suggest me how to add GTP in force field ?
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Hi Sunil,
You should try antechamber module in AmberTools, and ACPYPE.
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How long does it take, using a single processor, or compute cluster, to calculate the change in gibbs free energy on binding (ΔΔG) for an average sized dimer protein (already pre-docked), with a single snapshot using the MM/PBSA and/or MM/GBSA methods?
Additionally, how accurate is a single snapshot for MM/PBSA and MM/GBSA, compared to non-MM complex energy scoring functions?
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Hi Aaron, 
There is no direct answer to your question. I will try explaining.
If you want to use 1 single snapshot to calculate -- it would be something like energy of complex minus energy of protein minus the energy of the ligand. 
The answer to your first part - about time - it should not take long at all - but it really depends on the size of your structure. The smaller they are the quicker - I would estimate a couple of minutes. If you use GPUs it would be much much faster. 
The answer to you accuracy question is not simple. 1) Both of these methods are implicit solvent methods so they will not be good at all. 2) GB is an approximation of PB which is in itself a approximation so your results should be interpreted with care. 3) Single snapshot is not a good idea. You should at least do some form of averaging. e.g. you should simulate your complex, protein and ligand in explicit water seperately. Then desolvate the trajectories and calculate the average energy for each of the 3 simulations and then do Complex energy - Protein energy - Ligand energy to get the free energy of binding ... PB is preferred to GB. What this process will do is allow for your structures to sample a few conformations in explicit solvent - as a result the average you will get for each structure would be a better reflection than a single snapshot. 
But when you ask about non-MM based scoring function - the comparison would be hard - without looking at the specifics of the scoring function. Better methods exist e.g thermodynamic integration etc for calculation of free energies. I know this maybe not the exact answer you were expecting - but it may give you a start. 
Hope this helps.
Best,
/A
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I have performed MD simulations for a protein at different conditions. I would like to identify time-averaged conformational differences among the multiple trajectories I got. I would like to do it in a quantified way rather than simply trust in visual analysis. An idea I had is to align the time-averaged structures (e.g., using PyMOL), measure the distance between equivalent atoms, average the distances according to each residue, and then generate a plot of average residue distance versus sequence. Does anyone know a tool that does something similar?
Thank you in advance.
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Normalized covariance matrix of atomic positions, root mean square fluctuations of atomic positions, and configuration entropy could also help quantitatively to determine structure differences between trajectories.
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In the tutorials, it's mostly the trajectories of the same structure whose MSM are being built. I was wondering if MSMBuilder could be used to cluster trajectories of similar proteins?
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Thanks! I'm also thinking along similar lines to what you suggested i.e. getting pre-processed/intermediate files (as numpy arrays) of trajectories via MSMBuilder and then using those in ML algorithm implementations in sci-kit.
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I used mindist of gromacs to find the distance between the proteins (native and mutants) and the metal (metalloprotein) over the simulation period. I have the output in form of graphs. 
Is there a way to get the average distance moved?
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First of all, you have to create an index file for protein and metal. On the next step you can use the following command: g_dist -f traj.xtc -s topol.tpr -n index.ndx
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Hi Dear all;
Is there any server to find the Kd (or binding affinity) between a protein and a RNA (in the pdb file of a complex that is reach after docking with haddock)? 
The Best.
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Hi Farzaneh, sorry, but Kd measurements have to be done in solution with actual reagents if you want to publish the results.
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i want to know the interface residue of protein 1ns5. thank you
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There is a molecular visualization software called 'PyMol'. Open your whichever protein in that and follow the below link for the further work.
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Dear all,
I'm using Desmond to simulate membrane proteins and I'm interesting in changing simulation conditions in order to see how the system will act.
To equilibrate my system, before lunching MD, I'm using the lipid-protein equilibration/relaxation protocol that was developed by Dmitry Lupyan in collaboration with Schrodinger Inc researchers. Unpublished, 2009" - Desmond membrane relaxation protocol.
I would like to change the initial velocities, in the equilibration protocol, in order to see if my system will explore other conformational spaces. What parameter should I change in this protocol in order to realise this?
Thank you in advance for your help.
Kind regards.
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Change the value of "seed" in "randomize_velocities" plugin.
---
Not a Desmond user.
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I have extended the simulation from 10ns to 20ns using .cpt file during molecular dynamics run,when i am trying to analyze the result i am getting the graph only from 10ns to 20ns instead of 0-20ns. How can i get the graph from 0-20ns??
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You might not have used "-append" option, that means you have two trajectories (0-10ns and 10-20ns). So, before analysis concatenate them.
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I have a protein and want to know which one of the Fe, Zn or As ions bind to the receptor faster. I am wondering if I can use molecular dynamic tools such as GROMACS to find which one of these ions go to the binding site faster and make interaction with the protein?
I thought I can define an environment with similar concentration of Zn, Fe and As and run the simulation and find which one of these ions make an interaction with receptor faster. Do you have any suggestion to solve this problem using MD?
Thanks
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Dear Mohamad
You can do this study using Gromacs. You have to maintain your ions concentration in your water box and run the simulation. Prepare separate topology file manually for your ions and add in to your main topology file. You can prepare ion topology manually from previous literatures. Recently I'm working on such problems. If further clarification needed you can contact me.
Thanks
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The protein is a GPCR and MD simulation was carried out under vacuum.
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Because in vacuum, protein structures are often unstable. Watch the trajectory and you'll probably see structural changes, or at minimum large oscillations in the positions of those atoms (which is what RMSF is directly calculating).
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I was trying to understand and would be needing help to understand if cation -pi and pi-pi interactions can be as strong as electrostatic interactions between a phosphate group and its pocket. For example in the case of Stat-3 or Grb-2.
The pocket has 2 Arg, 3 Ser, 1 Glu, 1 Lys.
Thank you
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There is no such thing as a simple pi-pi interaction (see more below). Cation-pi interactions are somewhat simpler, so let's begin with those. The atomic nuclei in aromatic rings are enclosed in a molecular orbital system where the outermost orbitals are these pi-orbitals. The next level above these pi-orbitals are the empty so-called anti-bonding orbitals (or pi*-orbitals). They are not really anti-bonding at ll because they can be filled by overlapping orbitals from a metal ion forming what is (or used to be) called a dative bond. In like manner, a filled pi-orbital from an aromatic ring can overlap an empty bonding orbital on the metal atom. Since the atomic nucleus of the metal ion can be regarded as a single point, this kind of interaction (the "cation-pi interaction) is relatively "simple". The principles of pi-pi interaction are the same, mutual overlap of a filled pi-orbital with an empty pi*-"antibonding"-orbital on the other aromatic ring. The complication arises from the fact that there are many ways to do this, depending on the mutual orientation of the two aromatic rings. They can be parallel to each other as in graphite, or perpendicular as in many crystals, or anything in between, as in proteins.
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I have calculated solution NMR structure of 45 residue membrane protein using cyana-3.97, but the backbone RMSD of 20 lowest target function conformers is about 3.5. Can someone suggest me what do I have to do so that I get backbone RMSD below 1.00?
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Dr. Hiqmet,
During the Structure calculation using cyana I set macro to get me 20 conformers with lowest target functions. I get the PDB file for the 20 conformers having least target functions.
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Hi,
I want to perform the last step of Molecular Dynamic Simulation (MDS) with different.mdp parameters by gromacs 5.1.4. Do I have to change all post-steps parameters, including EM and Equilibration as well and restart them?
Best Regards
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Energy minimization is not that important. However, if you change cut-offs it is better to perform a short equilibration simulation. But if you change thermostat, you do not need to do so. It all depend what you going to change in mdp file. 
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I did a MD simulation in gromacs for a protein and I would like to calculate the RMSD just for a part of the protein instead that for the whole protein. Any idea how to do that?
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All you need to do is run gmx make_ndx on any coordinate or .tpr file and select the region of interest (type "help" for examples, the syntax is quite simple). Then you pass the index file to gmx rms -n and choose the group for output. You don't need to run the simulation again, you just have to analyze it properly (note that much of this is discussed in the GROMACS manual, either the PDF or online at manual.gromacs.org)
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In PyMol how can I determine:
1) the diameter of a membrane protein at the height of the membrane and/or the area in that plane (so that I can estimate how much space the protein takes in the membrane and how many lipids it displays)
and
2) the distance of a residue (or protein bound metal) to the membrane surface?
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Hi Josy,
Apologies, I should've just included some programs in the initial answer. One of the better programs (and has a web interface, but requires you register with them) is Crysol.
Hopefully those help!
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Hello everyone,
Is it possible to find the points of interaction between the DNA and protein in DNA linked proteins using VMD or we have to use the pdb file of those DNA linked proteins to find such points of interaction. Thanks
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If you have PDB file of the protein DNA complex, then you can use PDBSum
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Hello All
I wonder if there is docking programs that take into account the hydrophobic environment of transmembrane domain... Shouldn't docking simulations to the surface of TM domains have different desolvation penalty term than that of docking simulations for soluble proteins?
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You may use a hydrophobic implicit  solvation model by applying  a reasonable  dielectric constant...
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I am trying to find the inhibition constant Ki using time dependent inhibition equation given in the paper (Yu-Ting Lee, Hang Thu Ta, Ronald G. Duggleby, Cyclopropane-1,1-dicarboxylate is a slow-, tight-binding inhibitor of rice ketol-acid reductoisomerase, Plant Science, Volume 168, Issue 4, April 2005, Pages 1035-1040). In order to find Ki i have to find Koff and that is where the prblem starts. the Kon value is fine but the error for koff value is higher than actual koff value. I am using MATLAB to produce graphs and calculate the values.
Is there any other way to find the koff value where the error is not higher than actual value. I  am using only one assay to find all the values.
I am new to enzyme kinetics so might need detailed information for an answer :)
Thank you !
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For a truly irreversible inhibitor, koff is zero. However, some covalent inhibitors are not irreversible.
One method for measuring koff that has been applied to beta-lactamase inhibitors, for example, is the "jump-dilution" experiment. The inhibitor and the enzyme are co-incubated. The inhibitor concentration should be in excess over the enzyme, and the incubation time should be long enough, so that 100% inhibition is achieved during the incubation. The complex is then diluted into the substrate to start the reaction. The dilution factor should preferably be high enough that the inhibitor is essentially fully dissociated at equilibrium so that the enzyme activity could eventually recover fully, although this is not essential. The progress of the reaction is followed over time. The reaction rate will be seen to increase with time as the inhibitor dissociates. The resulting reaction progress curve is fit to an equation containing an exponential function (see below). The exponential coefficient is koff.
[P]=Vst + (Vi -Vs)[1-exp(-kofft)]/koff
[P] is product concentration, Vi is initial rate, Vs is steady-state rate, t is time
The mathematical treatment is the same as for onset of slow-binding inhibition, except the reaction rate speeds up instead of slowing down. The treatment of slow-binding inhibition can be found on p.141 of Copeland's Evaluation of Enzyme Inhibitors in Drug Discovery. I have just replaced kobs with koff.
The method to measure Ki from the onset of inhibition progress curves is given in section 6.3 of the same book. It can be difficult to implement, however, if the affinity of the inhibitor's equilibrium binding (Ki) is low relative to its covalent reactivity (kon). Under those conditions, the kobs vs [I] plot is linear, so that no estimate of Ki is obtained.
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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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I have the AMBER 10ns MD trajectory, I want to calculate the average RMSD over all snapshots for each residue. Would you please tell me where I can get the cpptraj script to plot RMSD vs Residue Number. I have attached sample plot below.
Thanks
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AMBER tools have an utility called cpptraj. This utility can calculate "Per residue RMSD". You can go through the Manual of AmberTools15 or any of its version but not less than 12, in the chapter "cpptraj" you will get the "rmsd" option and details of how to calculate "Per res RMSD".
To get an average for each residue, you can use R. Getting average is pretty simple.
Hope this help.
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I have been working on BCL2l9 protein structure prediction, since its sequence similarity was less than 30% with the template on pdb database, I decided to go by hhpred and get the best alignment there and then use that best alignment in modeller. How can I do this?
Thanks in advance!
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TM-align:
Zhang & Skolnick (2005) Nucleic Acids Res 33: 2302-2309.
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I have a very basic understanding on this matter. I am currently using online servers for APBS calculations. I tried several ways but PDB2PQR server always exclude metal ions from calculations. The only working way I found is to edit .pqr file but I am not confident on results.
Kindly Help
regards
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In fact, PDB2PQR exclude heteroatoms because it takes a library of known atomic charges and radii to generate .pqr files and this only covers the usual amino acids and nucleotide bases. But, as you correctly guessed, editing the .pqr file do the job: just  keep the proper format in the added lines to include your metal ions.The key issue, of course, is selecting the proper charge and atomic radii for these ions but a search in the web will provide you acceptable results.
APBS solve numerically the Poisson-Boltzmann equation approximating any molecular system by a set of spheres with atomic charges and radii given in the .pqr file. Therefore, you can feel confident on APBS results provided you include proper values for these parameters in the .pqr file.
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Hi all,
I am little confused about detergent's off rates. What does that mean when we say one detergent has lower or higher off rates than the other?. And is there any correlation between different off rates and membrane protein dynamics particularly conformational flexibility? 
Thanks 
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After reading this paper now I understood this better. I think detergents binds to the protein more or less like a ligand and when micelles are formed monomer detergent species can exchange with free detergents in the solution (the one not being part of micelle). The fact that DDM has faster exchange rates may also mean that it is more flexible and therefore GPCR conformational switches are facilitated. On the other hand LMNG with very very low CMC and slow off rates is more like lipid and therefore more stabilising as reported in many papers. Thanks 
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I understand that in this case the simulations will be biased, however, I would like to check how these known Kd's will affect the conformation of the peptide.
Usually I work in Gromacs but I don't know the straight forward way to do it there. Amber? I am checking their mail lists as well.
Thank you.
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To the best of my knowledge, there is no straight way to produce or recalculate force field parameters with the usage of standard tools of  GROMACS. In general, I think it is not a simple task to optimize force field parameters based on Kd, especially in case you want to check its several values.
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WB showed that protein A alters protein B expression but we found no colocalization at all between them in immunofluorescence analysis (CoIP wasn't performed yet).
Any suggestions, please?
Does anyone has a good literature/paper they could share about "Intermittent protein-protein interaction"?
Help is much appreciated.
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I agree with Sebastian, yo may use BiFC and, in parallel any of the resonance transfer techniques using fusion proteins: A with a donor and B with an acceptor (to then perform BRET or FRET).
But I wonder: You are assuming that A affects B expression because they interact but, in the absence of further information, this is more a wish than a solid hypothesis. 
If you indeed have some more information that makes likely this hypothesis I would go first to a natural source or a cell coexpressing the two proteins and perform colocalization studies.
On the other hand transient interactions do occur but usually due to a triggering factor, for instance when you activate a GPCR then beta arrestin may be recruited. In the absence of "activators"  that allow conversion of non ineracting A and B to AB (or viceversa), it would be quite difficult to identify the transient event.
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Which experiment would be the most appropriate?
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Dear all,
First, let me thank you for your helpful answers and suggestions.
What I am seeking is to determine whether the endogenous protein is physiologically forming homodimers. Thus I prefer avoiding manipulating the system in any way. I will thus go for a Western blot after cross-linking. 
Again, many thanks for your help. 
Best luck in your research
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Dear Sir, I want to find the intramoleculcular interaction energy of a protein in vacuum in namd. I would like to include both the bonded and non-bonded terms in the intramolecular interactions. Is there a way to find the interaction directly from the simulations and if so, i would like to know which ensemble shall i use.
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You can use vmd energy plugin to calculate the energy of a snapshot extracted from a trajectory ( http://www.ks.uiuc.edu/Research/vmd/plugins/namdenergy/) . The average over trajectory after equilibration should be equivalent to the ensemble average. The ensemble will be the same as the one used to generate the trajectory.
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I downloaded a crystal structure from PDB but it seems a bit odd. The residues are doubled:
Does anyone knows what is the meaning?
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If you open the .pdb in a text editor and inspect the REMARK lines you will find the explanation, here the relevant section from 5BS8.pdb header :
THE DOUBLE-STRANDED DNAS ARE SUPERPOSED COPIES OF EACH
OTHER, OVERLAID IN OPPOSITE ORIENTATIONS (BECAUSE DIRECTIONALITY
 COULD NOT BE RESOLVED). THEREFORE IN REALITY, RATHER THAN IN THE
 MODEL, THE BIOLOGICAL ASSEMBLY WOULD IN FACT BE HEXAMERIC - FOUR
 PROTEINS AND ONE DOUBLE-STRANDED DNA.
 COORDINATES FOR A COMPLETE MULTIMER REPRESENTING THE KNOWN
 BIOLOGICALLY SIGNIFICANT OLIGOMERIZATION STATE OF THE
 MOLECULE CAN BE GENERATED BY APPLYING BIOMT TRANSFORMATIONS
GIVEN BELOW. BOTH NON-CRYSTALLOGRAPHIC AND
 CRYSTALLOGRAPHIC OPERATIONS ARE GIVEN.
REMARK 350
REMARK 350 BIOMOLECULE: 1
REMARK 350 AUTHOR DETERMINED BIOLOGICAL UNIT: OCTAMERIC
REMARK 350 APPLY THE FOLLOWING TO CHAINS: A, B, C, D, E, F, G, H
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Does it affect TEER or expression of transport proteins?
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You may use them, but you should check for TEER and expression of ABC-transporters, e,g, by Western blotting.
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