These kind of "crystals" are very common with DDM, I guess they could be called
spherulites. Sometimes they develop one or a few straight edges after a while.
I don't think these are DDM crystals as DDM is very soluble, but
protein-detergent crystals that are dominated by weak detergent micelle
This has been discussed before, I guess more than once
I think most people (I'm sure i'll be corrected if wrong) would favor not
removing any atoms or setting occupancies to zero and let the invisible atoms
be accounted for by high B-factors (either set manually or just letting
refinement
To me this begs the question: why do this at all? A cost issue?
The Hampton coverslips work pretty well even with detergents that drastically
lower surface tension like C8E4. Those of a UK competitor that i shall not name
are not as good, in our humble experience.
Bert
Not quite sure what you mean but I suppose you refined with NCS restraints and
the red bar means that your chains in those regions are not identical. I would
turn NCS restraints off during refinement, with your resolution there is no
real good reason to include them. You probably have to do
I do think one has to consider whether there is a sufficient scientific advance
to justify publication. After all, peer reviewers are already overworked. From
the information given I don't think I would try to publish this but I'd
certainly consider depositing in the PDB and contaminant
I think it is very unlikely codon optimisation will improve solubility, so I'd
save my money and use it to try other things. Assuming you have tried (much)
lower temperatures for expression you could consider dialing down expression
via a different promoter or low-copy number plasmid. I assume
ert
You made the comment a few weeks ago not to boil helical membrane proteins for
SDS-PAGE. Could i please ask, does this also apply to type I membrane proteins
that only have a single a-helix, or is it just membrane proteins that are
predominantly helical?
Thanks
Rich
On 22 February 2017 at 19:
like others I'm not clear why you care where your protein runs on SDS-PAGE. I
think the band you're seeing is in fact the tetramer, suggesting your protein
(like KcsA) is very stable. Helical membrane proteins often migrate faster than
expected (by their Mw) on SDS-PAGE.
Also, never boil
It is entirely possible (fairly common) to have two different crystals in one
drop. If the habits are the same I'd expect the same cell, but that doesn't
have to be the case. Likewise, if the habits are different I'd expect different
cells, but again that is not necessarily the case.
bert
How long are you cryoprotecting for?
You don't really need more than a few seconds. If I have sensitive crystals i
tend to just swipe them slowly through the final CP solution and often that
works. If I would leave them in the CP solution for more than say 30 seconds
the crystals would behave
John, the lower-resolution datasets in your paper were generated by truncating
a high-res dataset, i.e. the lo-res datasets are of great quality. Would the
conclusions still be valid if the data are true low-res? (i.e. I/sigI 1.5-2
in last shell)?
Tx Bert
From:
Dear ,
If you want to get useful advice you have to give a bit more information. What
did you try?
Also, please do not attach large images to your message.
Thanks and good luck,
bert
From: CCP4 bulletin board [CCP4BB@JISCMAIL.AC.UK] on behalf of
Try running a regular SDS page gel w/o boiling your sample. Some P proteins run
differently on gel due to the conformational change that is induced. I agree
that you probably have a mixture in which the P form might be the minor
species. I'd probably try to get the IEX to work; given that the
This may be a useful paper to read, although it is a bit dated:
Acta Cryst. (1999). D55, 479-483
How many water molecules can be detected by protein crystallography?
In general, 3 A resolution is beyond where you can reliably add waters (I start
adding them only at 2.7-2.8 A resolution or
I think you'll find that this is not a naive question..
I doubt there is a consensus for this. Neither option is ideal, mainly because
of possible confusion generated for non-crystallographers. My preference is to
include the side chain but set the atoms i do not see to zero occupancy. The
Yes, this is tough. We mostly have used DMSO or DMF. You can try detergents,
but they tend not to be that effective in solubilisation and they might bind to
your protein rather than the compound you 'd like to bind. If you'd like to be
adventurous you could try using cyclodextrins as a
Hi,
I would stay away from the Mem-PER kit since the detergents in there are
possibly quite harsh (?). I doubt that detergents would cause lysis of yeast
cell walls in any case.
There is a budget-friendly alternative for mechanical breaking of yeast and
that is the bead beater, sold by
First, I wouldn't worry about the extent of SeMet incorporation. If you have
followed an established protocol (either using an auxotroph or regular strain)
the incorporation should be fine.
For your SeMet data, what is the redundancy and resolution? In my experience it
is often very hard to
In principle this is straightforward, but you'll need reasonably-sized
crystals. You'll have to wash them very well in mother liquor (protein buffer
will probably dissolve the crystals). I do at least 3 serial transfers in large
(5-10 ul) drops (with a loop), and for each transfer I move the
I wouldn't worry about oxidation of SeMet; at least I never have. A number of
years back there was a paper published actually claiming that oxidised SeMet
has a higher and sharper absorption edge than reduced SeMet. Of course mixed
forms could complicate things but I would just purify like you
Gerard's beautifully worded message underscores in my mind what a great
resource for all kinds of information this bulletin board is. I have many times
been impressed by the time people take to answer queries, even if they
sometimes seem perfectly googlable to me.
As a community we can be
Dear all,
I recently collected several datasets for a protein that needs experimental
phasing.
The crystals are hexagonal plates, and (automatic) data processing suggests
with high confidence that the space group is P622. This is where the fun begins.
For some datasets (processed in P622), the
You don't mention the condition, which is important (esp pH). Best pH values
are 6-8.
From personal experience: try K2PtCl4 and OsCl3.
Try ~5 mM for 30-60' (quick soak).
The advantage of osmium salts is that they give a nice color to your crystals
so you know something is binding.
HTH and GL,
Off the top of my head: Rhesus protein RhcG by the Stroud group (3HD6).
Steven White's website should be good to get this info relatively fast; in most
cases the expression system is listed.
bert
From: CCP4 bulletin board [CCP4BB@JISCMAIL.AC.UK] on behalf
depending on how extensively you have screened so far, the most efficient thing
to do may be to change the protein: different orthologs, truncations,
mutagenesis of entropy rich clusters, change of tag location or tag cleavage
etc.
From: CCP4 bulletin board
The problem you're having is a very common one, unfortunately. There are about
10,000 things you can try to improve diffractiongenerally going from low to
solvable resolution is very, very hard.
I would say the two most important ones are (i) to vary the detergent and (ii)
to go to another
The first thing is that you'll have to increase the salt concentration in your
buffer. 5 mM is way too low and may cause non-specific binding of the protein
to the resin. 100 mM is the minimum you should use. There is nothing in your
buffer that will precipitate, so you should't have to worry
Well, you'll never loose the protein in the flowthrough if you don't discard
the flowthrough.which i never do until I'm sure about what happened to the
protein.
But yes, try different Mw cutoff concentrators until you find the highest one
that works. Depending on the purification your
Hi Raji,
the amount of detergent used is more important than the concentration. The more
cells you have, the more detergent is required. To give you an idea, we tend to
use 100 ml of 1% detergent (=1 g) for 12 liters of cells (Ecoli) of OD600 1.
This is probably on the low side; if cost
I could add that even for membrane proteins that may not be stable in eg DM
after purification, DM may still be a viable option for extraction since there
will be large amounts of stabilising lipids present. Just make sure you change
to a milder detergent for the post-extraction steps.
Bert
since your protein aggregates even in a mild detergent you may have to find an
ortholog that is more stable.
however, there are a few things you can try before moving on (in arbitrary
order):
1. add glycerol during purification (5-20%)
2. get rid of the imidazole as fast as possible after Ni.
Disappearance of crystals is fairly common and not specific to DDM or indeed
any detergent as it also occurs for soluble proteins. Even after reaching
equilibrium conditions still change (evaporation, instability of components,
proteolysis, etc). The inverse also happens, that is formation of
. The successful candidate will have a
PhD or previous post-doctoral experience in X-ray crystallography and
biochemistry or biophysics and should have published at least one first author
paper in a leading journal.
Further information regarding this post can be obtained by contacting Prof Bert
van den
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