>> Hi. Welcome to the next in our OSTR series on Cutting-Edge Methodologies. Today we're
going to talk about structural mass spectrometry and its applications using cross-linking and
limited proteolysis. First, cross-linking mass spectrometry is an experimental workflow
used to identify constituents of large macromolecular complexes. It can be used to identify sites
of protein dimerization or disulfide formation, to compare protein confirmations in different
conditions, as well as to obtain distance constraints for modeling of protein structures
and confirmation of protein complexes. This is a methodology that can be applicable to
multiple different sample types including complexes with up to 100 different subunits
and is applicable with pull downs and in vivo applications both. The cross-linking reaction
can be done using multiple different types of cross-linkers with complementary chemistries
to target different types of resides. The methodology is also compatible with fractionation
and enrichment steps, which will increase the ability to identify the cross-linked peptides.
Then there's usually a mass spec acquisition step, some sort of data analysis, including
new software for differential quantitation, and visualization and modeling of protein
complexes. So this is just an example of a peptide, with lysine residues highlighted
in the light purple within the protein. A cross-linker is added which then allows, in
this case, cross-linking of proximal lysine residues together. When the protein is digested,
those residues stay locked together with a covalent modification and then they can be
analyzed to detect the sequence of the peptide as well as the site of the cross-link. It's
important to keep in mind that when a cross-linking agent is applied, there are multiple different
types of cross-links that can be formed. The most beneficial are intramolecular or intermolecular
cross-links in which two different peptide sequences are cross-linked. Because these
are the other ones that actually give conformational information. Less useful but more prevalent
are mono links in which one peptide is cross-linked but the other never cross-links to anything.
And loop links, in which two residues within the same peptide are cross-linked together.
Both of these types will unfortunately not give any useful information for conformational
changes or complex information. ^M00:02:26
There are multiple different types of cross-linkers that can be used. These start with reagents
such as EDC, which is a zero length cross-linker that will cross-link carboxylic to amine residues.
These are really nice because they're effectively cross-linking hydrogen bonds. Also common
are things such as DSS or BS3. These are traditional cross-linkers of size approximately 11 A,
that cross-link amine residues. And also there are photo cross-linkers, which can cross-link
an amine to any other amino acid after application of UV light. In addition, there are mass spectrometry
cleavable cross-linkers such as DSSO, which are commercially available. And these are
very convenient. When cross-linked into the protein and then analyzed in the mass spectrometer,
the fragmentation of the instrumentation will lead to cleavage of the cross-linker with
a very common and distinctive pattern of tagged forms of the protein. These can then be used
to set up the instrument to identify and target the peptides for further fragmentation and
lead to much less complicated mass spectra. In addition, there has recently been the use
of short distance cross-linkers for development of structure restraints for modeling of protein
complexes. And a few of those examples are given here. So there's lots of different types
of cross-linkers available and what you choose may depend on your experimental question.
^M00:03:57 After cross-linking, it's often a good idea
to do some sort of an enrichment step. This includes size exclusion or ion exchange chromatography,
which are used to enrich the cross-linked peptide over non-modified peptides. These
enrichment steps are very critical, especially for large protein complexes. Because the number
of cross-linked peptides as opposed to noncross-linked peptides is very small. And so without the
enrichment, it's very difficult to identify and detect the cross-linked peptides. Because
these peptides tend to be larger and have more charge, they can be enriched using either
the size exclusion or ion exchange chromatography. In my own recent set of data, just to show
you how important this enrichment step is, I found that the cross-link spectra tended
to be between 0.5 and approximately 8% of the total spectra identified. Whereas the
loop linked were about another 5%. The mono linked were 10 to 50%. And unmodified, somewhere
between 40 and 85%. So the peptides that we want to see with the cross-links that really
show us the confirmational information tend to be the low-end. And so an enrichment step
to bring them up is very useful. There are also multiple different software packages
that have been used to help identify and provide information about cross-linked peptides. Some
of these are commercially available and do a very nice job of analyzing cross-link data.
Part of the reason it's important to have nice cross-link data in software is because
the spectra, unless you're using a mass spec cleavable cross-linker, tend to be very complicated.
This is an example mass spectrum, and it's actually a very nice one. What you see are
the combination fragmentation spectra for two peptides that are cross-linked together.
Because they're attached, we see fragment ions coming from both parts of the cross-link,
both of the peptides that are cross-linked together. This is what we want because this
provides us with the sequence information to know which two peptides are being cross-linked
together and where the cross-linking occurs. However, this type of clean data with this
extent of cross-linking is very rare. And in fact, spectra often may look something
more like this, where we still have good distribution of ions across the cross-linker, but there's
a lot more fragmentation occurring, or they may even look like this. Again, where you
see a lot of other ions that aren't matching, not quite as good of coverage of the peptide,
but still enough to know what the sequence is and where the cross-link occurs.
^M00:06:30 So with a difficult spectra to interpret and
the low isolation rate of the cross-link peptides, that's why the addition of both enrichment
strategies help as well as mass spec compatible cross-linkers, which would cleave in the instrument
and give somewhat cleaner spectra, both of these help to give better data quality. So
as I said, cross-linking has been suggested to be used for structural analysis. Similar
to NMR experiments, cross-linking mass spec can be used to provide distance restraints
that can be used for ensemble of structural models to help with complexes that don't have
structures already. In these cases, short distance cross-links are often better for
restraining models than long-distance cross-linkings. Because you need that short distance to really
restrain the complex. The long distances don't really help because there are so many options
that could meet that restraint. In these cases, often one would need further experiments to
validate the model, which would include deuterium exchange mass spec, long-distance cross-linking,
and surface modification. These are ensembles of models, predicted structures, not true
structures themselves. So additional information to validate predictions are essential to have
to really know that you have a good model. ^M00:07:46
This is an example of using high density photo cross-linking for albumin, just to show you
again how it can work. In this case, this is the albumin protein. And you see all of
the red cross-links that occur. There's a very nice distribution of cross-links that
happen at the short distance. Whereas poorly matched or decoy falsely identified seem to
have long distances. And if one then looks at either the purified or the serum containing
albumin, you can see that they have very similar models that show similar distance restraints.
Whereas if there is no cross-linking, we just see this very odd conformation that shows
no relationship to the native protein. So this is just to show you that this type of
method can be used to get structural information about proteins.
^M00:08:32 Another method that can be used to try and
understand conformations and conformational changes is limited proteolysis coupled to
mass spectrometry. In this methodology, upon a conformational change or complex formation,
sites may become opened up or obscured for protease digestion. Limited proteolysis of
a protein under different conditions, followed by mass spectrometry analysis, can be used
to characterize these changes and look at where protein conformation changes are occurring.
Conformational parameters such as accessibility, segmental mobility, and solvent exposition
correlate quite well with limited proteolytic sites. So this would help you to see where
there are changes in protein conformation, where binding sites are taking place, and
what is exposed and what is not. In a standard limited proteolysis experiment, a protein
is incubated with a protease for a short amount of time. Because we don't want complete proteolysis
of the protein, just cleavage of accessible sites. After this, there's a mass spec analysis
performed, and this can be performed under -- the proteolysis can be performed under
different conditions for different times to look at different exposures of these residues.
Cleavage sites that are usually located in disordered regions or domains or in regions
with enhanced flexibility may see a lot more cleavage. In contrast to classic methods to
get structural information of a protein, limited proteolysis is quite nice because it doesn't
require as much of the protein or as much experimental effort to really get good data.
So this is sort of a nice initial approach to see if there are conformational changes
occurring and to get some initial data to help direct further downstream experiments.
It's important to keep in mind that the outcome of a limited proteolysis experiment is not
high-resolution information. Instead, partial digestion and other structure determination
methods are usually combined to allow monitoring of conformational transitions upon addition
of substrate, cofactor, or an inhibitor. It's a first step, it's not the last step.
^M00:10:45 This is an example looking at a specific protein
that's been incubated with trypsin to do limited proteolytic digestion. So as you can see,
as time passes, the input protein decreases in abundance and these lower fragments increase.
And then mass spectrometry analysis can be used to identify where the different cleavages
are occurring in each of these different fragments and what sites become exposed and affected
in limited proteolysis. For these experiments, there are several reaction conditions that
are critical. The pH of the sample is important because one wants to maintain the native conformation
of the substrate, maximize the stability of the substrate regardless of the optimum pH
for enzymatic activity. And one needs to keep in mind that a non-optimal pH may affect the
protease activity itself. So there's a lot of balancing to be done to keep native conformation
of the protein as well as some activity of the protease itself. The digestion time is
something that can also be optimized. As the incubation time increases, fragments become
released from the intact protein that are then subdigested by the protease, giving rise
to smaller peptides. These peptides originate from other proteolytic fragments. And so they
don't actually give any information about the native protein complex. They're really
only released once the protein starts being digested, and so they have little information.
So you don't want the digestion to go on too long, or these smaller, less interesting fragments
will start to dominate. Finally, the enzyme-to-protein ratio is very critical. In fact, some people
think it's the most important factor to modulate the selectivity of the protease. One can limit
the activity even when a broad specificity enzyme is used by changing just the ratio
of enzyme to protein. Usually the amount of enzyme you want to use will be different for
a protein alone versus a protein complex. And that's because the complex itself has
lower accessibility than the individual components because it has a tighter, more compact conformation.
So you may need to optimize the amount of enzyme across different experimental conditions.
^M00:12:57 Several different proteases are often used
and people will even use panels of proteases with different specificities to get different
and overlapping information. The selectivity of cleavage is not related to or limited by
the specificity of the enzyme. And instead grouping of the preferential cleavage sites
within the same region, if you use overlapping sites, can be used to help know that you're
really looking at the real accessible site and not just some sort of an off target information.
It's also important to quench the reaction. For both reproducibility and to avoid overhydrolysis,
complete quenching of proteolytic activity is critical. Because the quenching needs to
be both instant and complete, changing conditions, particularly pH, is the most common way to
do it. Often one would quench by addition of trifluoroacetic or acetonitrile. Denaturants
are also used, though some proteases will show residual activity in the presence of
denaturants. And so you need to know if you have good quenching conditions. These are
just some examples of different proteases and the specificity for cleavage as well as
common kinase-to-protein ratios that have been used for these types of experiments.
^M00:14:11 One very nice experimental design includes
the use of a reference sample to an experimental sample. And the reference sample, for example,
in this case, a denaturant kinase that's fully unfolded, digestion is done to completeness
to know what are the fragments that can be analyzed. The protein is also labeled with
isotopically labeled butyrate, so that the peptide masses will shift. Then there's an
experimental sample, which in this case would be a kinase with a small molecule. Which is
in the folded condition. Limited proteolysis is performed using a lower trypsin-to-protein
ratio for a short time. And then late labeling with a light form of the butyrate. The samples
are pooled and analyzed by mass spec. And in this case, the ratio of the limited proteolysis
peptides to the complete proteolysis peptides gives important information about conformational
differences. So in this case, you can see where they've applied this to looking at B-Raf
kinases induced by small molecule binding. You can see that for different peptides, the
different forms and different states show different cleavage propensities, which can
be used to identify binding sites on the protein and to understand how they affect conformational
change. In addition, people have used whole-cell limited proteolysis to analyze protein structural
changes directly in the biological context of the proteome on a whole-cell scale. This
is a very interesting methodology that includes some different limited proteolysis steps.
It does have some limitations. It gives a list of proteins that change, but it doesn't
tell you the nature of the change. You don't know if there's differences in accessibility
to ligand-binding protein aggregation or even altered interactors. Currently this is a method
that's only been used for medium to high abundance proteins and it tends to be biased against
membrane proteins because they tend to be less accessible. And it's best when used to
compare multiple treatment conditions. But it is something to keep in mind that may be
interesting in your own research. Just to show how they've used it, in this case there
are multiple different enzymes used to look at different proteins. And what you can see
are different sites of accessibility. And so in this case, the authors would first use
a generic proteolysis step to look at the proteolysis and then follow it up with tryptic
digestion. And in that case, where they have semi-tryptic peptides, those are the sites
of limited proteolytic activity. And so that's how they know where there is cleavage. And
then by comparing different conditions of the cell, they can look at how the protein
conformation is changed. But again, they don't know why the change has occurred. So for more
information about these types of methods and to think about how you could use them in your
own research, please feel free to contact your local mass spectrometrist. Within the
NCI, you can talk to me, Lisa Jenkins, or my colleague, Thorkell Andresson in Frederick.
Within the wider NIH community, I would direct you to visit the CREx for more information
about available resources. Thank you very much for your attention.
^M00:17:25 >> US Department of Health and Human Services,
National Institutes of Health, National Cancer Institute, cancer.gov. 1-800-4-cancer. Produced
August 2019.