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The concept of shotgun proteomics is shown above. Instead of separating the
proteins, the entire cell extract is digested with proteases and then the complex
mixture is separated. The peptides are eluted from the final separation method,
usually reversed-phase chromatography directly into the mass spectrometer
where they are automatically subjected to MS/MS analysis. The peptides are
identified in a similar way to how proteins are identified. Maybe 10 peptides
are entering the mass spectrometer. The MS picks automatically the most
intense, isolates it (throwing away the other 9 peptides) and then smashes it
into pieces. The mass of the peptide is used to search the database to find all
peptides with the same mass. The fragmentation spectra of all these peptides
are then calculated and compared to the experimental fragments observed. The
best matching peptide sequence is then selected.
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AC; affinity chromatography. SEC; Size exclusion chromatography. IEC;
Ion exchange chromatography. HIC; Hydrophobic interaction
chromatography. . HILIC; Hydrophilic interaction chromatography.
RPC; Reversed phase chromatography.
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IEX separates molecules on the basis of differences in their net surface
charge. Molecules vary considerably in their charge properties and will
exhibit different degrees of interaction with charged chromatography
media according to differences in their overall charge, charge density
and surface charge distribution. The charged groups within a molecule
that contribute to the net surface charge possess different pKa values
depending on their structure and chemical microenvironment. Since all
molecules with ionizable groups can be titrated, their net surface charge
is highly pH dependent. In the case of proteins, which are built up of
many different amino acids containing weak acidic and basic groups,
their net surface charge will change gradually as the pH of the
environment changes i.e. proteins are amphoteric. Each protein has its
own unique net charge versus pH relationship which can be visualized
as a titration curve. This curve reflects how the overall net charge of the
protein changes according to the pH of the surroundings. IEX
chromatography takes advantage of the fact that the relationship
between net surface charge and pH is unique for a specific protein. In
an IEX separation reversible interactions between charged molecules
and oppositely charged IEX media are controlled in order to favor
binding or elution of specific molecules and achieve separation. A
protein that has no net charge at a pH equivalent to its isoelectric point
(pI) will not interact with a charged medium. However, at a pH above its
isoelectric point, a protein will bind to a positively charged medium or
anion exchanger and, at a pH below its pI, a protein will behind to a
negatively charged medium or cation exchanger..
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When all the sample has been loaded and the column washed so that
all non-binding proteins have passed through the column (i.e. the UV
signal has returned to baseline), conditions are altered in order to elute
the bound proteins. Most frequently, proteins are eluted by increasing
the ionic strength (salt concentration) of the buffer or, occasionally, by
changing the pH. As ionic strength increases, the salt ions (typically Na
+ or Cl-) compete with the bound components for charges on the
surface of the medium and one or more of the bound species begin to
elute and move down the column. The proteins with the lowest net
charge at the selected pH will be the first ones eluted from the column
as ionic strength increases. Similarly, the proteins with the highest
charge at a certain pH will be most strongly retained and will be eluted
last. The higher the net charge of the protein, the higher the ionic
strength that is needed for elution. By controlling changes in ionic
strength using different forms of gradient, proteins are eluted
differentially in a purified, concentrated form. A wash step in very high
ionic strength buffer removes most tightly bound proteins at the end of
an elution. The column is then re-equilibrated in start buffer before
applying more sample in the next run.
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For molecules such as nucleic acids which carry only negativelycharged groups, an anion exchanger is the obvious choice. However,
since the net charge of molecules such as proteins (carrying positively
and negatively charged groups) depends on pH, the choice is based on
which type of exchanger and pH give the desired resolution within the
constraints of sample stability. For example, the slide shows a
theoretical protein which has a net positive charge below its isoelectric
point and can bind to a cation exchanger. Above its isoelectric point the
protein has a net negative charge and can bind to an anion exchanger.
However, the protein is only stable in the range pH 5–8 and so an anion
exchanger has to be used.
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The term “reversed phase” derives from “normal phase”
chromatography, a technique utilizing a hydrophilic stationary phase
together with mobile phases consisting of organic solvents such as
hexane or methylene chloride. In RPC the stationary phase is
hydrophobic so that a water/organic solvent mobile phase is used, that
is, the stationary phase is more hydrophobic than the mobile phase.
RPC media may be referred to as adsorbents while eluent solutions
may be referred to as mobile phases. The separation of biomolecules
by RPC depends on a reversible hydrophobic interaction between
sample molecules in the eluent and the medium. Initial conditions are
primarily aqueous, favoring a high degree of organized water structure
surrounding the sample molecule. Frequently, a small percentage of
organic modifier, typically from 3–5% acetonitrile, is present in order to
achieve a “wetted” surface. As sample binds to the medium, the
hydrophobic area exposed to the eluent is minimized. Separation relies
on sample molecules existing in an equilibrium between the eluent and
the surface of the medium. The distribution of the sample depends on
the properties of the medium, the hydrophobicity of the sample and the
composition of the eluent (mobile phase), as illustrated in the slide.
Initially, conditions favor an extreme equilibrium state where essentially
100% of the sample is bound. Since proteins and peptides carry a mix
of accessible hydrophilic and hydrophobic amino acids and are rather
large, the interaction with the medium has the nature of a multi-point
attachment. To bring about elution the amount of organic solvent is
increased so that conditions become more hydrophobic. Binding and
elution occur continuously as sample moves through the column. The
process of moving through the column is slower for those samples that
are more hydrophobic. Consequently, samples are eluted in order of
increasing hydrophobicity.
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Sample is applied under conditions that favor binding, typically using an
aqueous solution containing an ion-pairing agent, such as trifluoroacetic
acid (TFA), to enhance the hydrophobic interaction and a low
concentration of organic modifier such as 5% acetonitrile. After
application, and when all non-bound molecules have passed through
(i.e., the UV signal has returned to baseline), conditions are altered in
order to elute the bound sample. Elution begins by increasing the
concentration of organic modifier, such as acetonitrile. Molecules with
the lowest hydrophobicity will elute first. By controlling the increase in
organic modifier, molecules are eluted differentially. Those molecules
with the highest degree of hydrophobicity will be most strongly retained
and eluted last. A wash step removes most of the tightly bound
molecules at the end of elution.
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Ion-pairing agents A common way to increase the hydrophobicity of
charged components, enhance binding to the medium, and so alter
retention time, is to add ion-pairing agents to the eluent. These agents
bind via ionic interactions with charged groups and thereby suppress
their influence on overall hydrophobicity. Since most proteins and
peptides are slightly basic, ion-pairing agents are often acids such as
trifluoroacetic acid (TFA) whereas a base such as triethylamine is used
for negatively charged molecules.
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Ion-pairing agents A common way to increase the hydrophobicity of
charged components, enhance binding to the medium, and so alter
retention time, is to add ion-pairing agents to the eluent. These agents
bind via ionic interactions with charged groups and thereby suppress
their influence on overall hydrophobicity. Since most proteins and
peptides are slightly basic, ion-pairing agents are often acids such as
trifluoroacetic acid (TFA) whereas a base such as triethylamine is used
for negatively charged molecules.
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To prevent the OH groups from binding to peptides in a hydrophilic
mode, the OH groups are often blocked or end capped to give acetyl
groups.
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Water is a good solvent for polar substances, but a poor solvent for
non-polar substances. In liquid water a majority of the water molecules
occur in clusters due to hydrogen bonding between them- selves
(Figure 3). Although the half-life of water clusters is very short, the net
effect is a very strong cohesion between the water molecules, reflected,
for example, by a high boiling point. At an air-water interface, water
molecules arrange themselves into a strong shell of highly ordered
structure. Here, the possibility to form hydrogen bonds is no longer in
balance, but is dominated by the liquid side of the interface. This gives
rise to an ordered structure that manifests itself as a strong surface
tension. Anything that influences the stability of the water shell also
affects the surface tension. When a hydrophobic substance such as a
protein or hydrophobic ligand is immersed in water some- thing
analogous to the surface tension phenomenon happens. The water
molecules cannot “wet” the surface of the hydrophobic substance.
Instead they form a highly ordered shell around the substance, due to
their inability to form hydrogen bonds in all directions. Minimizing the
extent of this shell leads to a decrease in the number of ordered water
molecules, that is, a thermodynamically more favorable situation in
which entropy increases. In order to gain entropy, hydrophobic
substances are forced to merge to minimize the total area of such
shells. Thus hydrophobic interaction depends on the behavior of the
water molecules rather than on direct attraction between the
hydrophobic molecules. The hydrophobic ligands on HIC media can
interact with the hydrophobic surfaces of proteins. In pure water any
hydrophobic effect is too weak to cause interaction between ligand and
proteins or between the proteins themselves. However, certain salts
enhance hydrophobic interactions, and adding such salts brings about
binding (adsorption) to HIC media. For selective elution (desorption),
the salt concentration is lowered gradually and the sample components
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The hydrophobic ligands on HIC media can interact with the
hydrophobic surfaces of proteins. The ligands immobilised onthe beads
are, in order of increasing hydrophobicity: hydroxypropyl < methyl <
benzyl = propyl < iso-propyl < phenyl < pentyl!
These ligands are much more less hydrophobic than C8 and C18
ligands. In pure water any hydrophobic effect is too weak to cause
interaction between ligand and proteins or between the proteins
themselves. However, certain salts enhance hydrophobic interactions,
and adding such salts brings about binding (adsorption) to HIC media.
For selective elution (desorption), the salt concentration is lowered
gradually and the sample components elute in order of hydrophobicity.
HIC media are composed of ligands containing alkyl or aryl groups
coupled to an inert matrix of spherical particles. The matrix is porous, in
order to provide a high internal surface area, while the ligand plays a
significant role in the final hydrophobicity of the medium. Interaction
between the protein and the medium is promoted by moderately high
salt concentrations, typically 1–2 M ammonium sulfate or 3 M NaCl. The
type of salt and the concentration required in the start buffer are
selected to ensure that the proteins of interest bind to the medium and
that other less hydrophobic proteins and impurities pass directly
through the column.
Antichaotropic salts are employed, eg. (NH4)2SO4. Chaotropic salts
(NaCl, NH4SCN) do not retain solutes
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Interaction between the protein and the medium is promoted by
moderately high salt concentrations, typically 1–2 M ammonium sulfate
or 3 M NaCl. The type of salt and the concentration required in the start
buffer are selected to ensure that the proteins of interest bind to the
medium and that other less hydrophobic proteins and impurities pass
directly through the column.
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Binding: buffer conditions are optimized to ensure that the target
molecules interact effectively with the ligand and are retained by the
affinity medium as all other molecules wash through the column.
Elution: buffer conditions are changed to reverse (weaken) the
interaction between the target molecules and the ligand so that the
target molecules can be eluted from the column. Wash: buffer
conditions that wash unbound substances from the column without
eluting the target molecules or that re-equilibrate the column back to the
starting conditions (in most cases the binding buffer is used as a wash
buffer). Ligand coupling: covalent attachment of a ligand to a suitable
pre-activated matrix to create an affinity medium. Pre-activated
matrices: matrices which have been chemically modified to facilitate the
coupling of specific types of ligand.
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The basis for purification of IgG, IgG fragments and subclasses is the
high affinity of protein A and protein G for the Fc region of polyclonal
and monoclonal IgG-type antibodies.
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