Download Protein Isolation and Analysis

Protein Isolation and Analysis
Cell Lysis
• To study a protein, you need to obtain
large amounts of it, purified away from all
other proteins.
• First, you need to lyse the cells: break
them open.
• Grinding (mortar and pestle)
• Sonication (high frequency sound)
• Force through a small orifice at high
pressure: French press and plunger
homogenizers
• Detergent to disrupt membranes:
especially for membrane proteins
• The point is to break open the cells while
leaving membrane-bound organelles
intact.
• After this process, you have a cell extract
or homogenate.
Centrifugation
• Most proteins have a particular location within the cell:
are associated with a some organelle, or bound to the
cell membrane, or just loose in the cytoplasm.
• The cell homogenate can be fractionated into these
different components using the centrifuge.
• Centrifuges spin the samples at high speeds, subjecting
the homogenate to forces up to 600,000 x g (where 1 g is
the normal gravitational force at the Earth’s surface).
• But usually we use 100 – 10,000 x g.
• The pellet is what goes to the bottom of the centrifuge;
the supernatant is what is left suspended in the liquid.
• Differential centrifugation: centrifuge the homogenate
several times, using successively stronger forces.
Different organelles pellet at different gravitational
levels, depending on their size, density, and shape.
Velocity Centrifugation
• Velocity centrifugation, often called rate-zonal centrifugation,
separates cell components based on how fast they sediment.
• The Svedberg (S) is a measure of sedimentation rate, and not
additive. For example, the eukaryotic ribosome is 80S, and is
composed of a 40S small subunit and a 60S large subunit.
• The sample is carefully layered on the top of the centrifuge
tube. The tube is filled with a sucrose gradient, from about 5%
to 30% sucrose. The sucrose forms a density gradient, which
helps keep the cell components in tight bands and not diffusing.
• Sometimes compounds other than sucrose are used: Ficoll and Percoll are
common ones. Unlike sucrose they can’t diffuse into membrane-bound
organelles.
• The samples are then centrifuged until different bands of cell
components have separated due to their different
sedimentation coefficients.
• Note that if you keep centrifuging, eventually everything will end up in the
pellet. This is NOT an equilibrium method.
• You can then puncture the bottom of the tube and remove the
bands drop by drop.
Equilibrium Centrifugation
• Equilibrium centrifugation (isopycnic) separates
on the basis of buoyant density: the density
where the cell components float.
• Buoyant density is independent of size and shape. It is
measured in g/ml, with water having a buoyant
density of 1.0.
• The samples are mixed with a high concentration
of sucrose or cesium chloride (CsCl), then
centrifuged until everything in the mixture has
floated to its equilibrium density position.
• The centrifugal forces generate a density gradient
from the CsCl solution, because there is a large
difference in the gravitational force at the top of the
tube than at the bottom.
• This is an equilibrium technique: once the
components have floated to their positions, they stay
there.
• This is a very common technique for purifying
DNA.
Protein Separation
• Each cell contains a complex mix of proteins,
and it usually takes several steps to isolate one
particular protein.
• Separation is based on differences in size, charge,
hydrophobicity, and affinity for other molecules.
Proteins differ widely in these properties; this is not
true of nucleic acids.
• Analytical separation methods are used to
detect and identify a protein, while preparative
methods are used to isolate large amounts of a
specific protein for further study.
• Column chromatography is an important preparative
method. A mixture of proteins is applied to the top
of a column containing a permeable matrix that binds
some proteins more tightly than others.
• Electrophoresis is a common analytical method.
Proteins are separated by an electrical field, using a
gel matrix.
Column Chromatography
• Proteins are separated because they flow through the
column at different speeds, based on how they interact
with the column material.
• Three basic types: ion exchange (as well as the very
similar hydrophobic interaction); gel filtration; and affinity.
• Ion exchange chromatography uses small beads that are
covered with charged groups, either positive or negative.
• These groups form ionic bonds with charged groups on the
surface of the proteins.
• Proteins are eluted from the column by increasing the salt
concentration of the solvent. At some salt concentration,
the charged groups on the protein bind to Na+ or Cl- ions
more tightly than to the column material, and they get
washed out of the column into a fraction collector.
• Hydrophobic interaction chromatography is very similar:
the gel beads are covered with hydrophobic groups.
Proteins bind in high salt, and are eluted by decreasing the
salt concentration. The high salt solution is very hydrophilic,
and forces hydrophobic regions to stick together.
Gel Filtration
Chromatography
• Sometimes called size exclusion
chromatography.
• The gel matrix consists of beads with many
pores of different sizes.
• Principle: Large proteins are too big for the
pores, so they just go around the beads.
Smaller proteins can spend time inside the
pores of the beads. This means that smaller
proteins have to travel through a larger
volume than larger proteins. Thus, large
proteins elute first and small proteins last.
• Sephadex is the main brand name for the
beads used in gel filtration chromatography.
Affinity Chromatography
• Many proteins bind to specific small molecules,
referred to as ligands. If the ligand your protein
interacts with is covalently bound to the column
material, it will bind to (adsorb to) the column while all
other proteins wash through without binding. Your
protein can be eluted by raising the salt concentration
or pH, or by simply adding a lot of ligand molecules to
the elution buffer.
• Most affinity methods are batch methods: the protein is
adsorbed and washed under one set of buffer conditions,
then eluted with another, different set of buffer conditions.
• Antibodies raised against your specific protein work
very well for affinity columns: they are very specific for
a single protein. It is often possible to go from a crude
mixture to a pure protein in a single step of immuneaffinity chromatography.
Antibodies
• This is a very simplified treatment of the subject!
• Antibodies bind very tightly and specifically to antigens. Each
antibody binds to a single antigen.
• Antibodies are proteins, composed of two identical light chains
and two identical heavy chains.
• They form a characteristic Y shape, with the antigen binding
regions at the ends of the two arms.
• Thus, each antibody molecule can bind to 2 antigen molecules. This allows
large complexes of antibodies and antigens to form. These large complexes
are insoluble, and cells of the immune system find them and destroy them.
• Each antibody binds to a specific region of its target antigen, the
epitope. A given antigen might have several different epitopes,
each binding to a different antibody.
• Specificity of the antibodies comes from the variable regions of
the heavy and light chains. There are billions of possible antigen
binding sites created by these regions.
More Antibodies
• Antibodies are made in the B
lymphocytes (white blood cells).
• Each B cell makes only a single kind of
antibody.
• The B cells display their antibody on the
cell surface. When an antigen binds to
that antibody, the B cell is stimulated to
divide and to produce large amounts of
its antibody, which get secreted into
the blood.
• Each individual can make millions of
different antibodies, but most of them
just present on the surface of their B
cell, waiting for attack by foreign
antigens.
Making Antibodies for Research
• It is a simple process: you inject your protein (the
antigen) into a rabbit (or mouse, goat, sheep,
whatever). The rabbit’s immune system detects
the foreign protein and produces several different
antibodies that react with it.
• Most proteins have several different epitopes on their
surfaces. An epitope is simply a region that happens to
interact with an antibody. Thus, the immune serum
raised in a rabbit will contain many different antibodies
to your protein. This is called polyclonal antibodies.
• After a few weeks, the rabbit has lots of antibody against your
protein in its blood. Usually all you need to do is remove the
red blood cells from the blood to have a usable antibody
preparation (immune serum).
• You can increase the immune response by giving the rabbit
booster shots of your protein every few weeks.
Monoclonal Antibodies
• Monoclonal antibodies are derived from a
single B cell, and thus they are all identical and
react to only one epitope on your protein.
• Monoclonal antibodies are made by injecting
your protein into a mouse, waiting until an
immune response appears, then taking the B
cells from the mouse’s spleen and fusing them
with an immortal myeloma (tumor) cell.
• This produces a hybrid cell, a hybridoma, that
produces a single type of antibody (from the B cell)
and is immortal (from the myeloma cell).
• You can then grow as much of this hybridoma as
you like, giving you as much of this specific antibody
as you like.
Using Antibodies
• Because they are so specific, antibodies have many uses
in studying proteins.
• Affinity chromatography
• Labelled antibodies to detect specific proteins.
• Antibody molecules can be labelled with fluorescent
dyes; or with enzymes like horseradish peroxidase that
can be used with an artificial substrate to produce an
insoluble and easily visible dye, or with gold particles,
which are easy to see in the electron microscope, or
with a radioactive tag.
• It is common to use a conjugated secondary antibody: a
labelled antibody from goats or sheep that reacts with
the common region of rabbit antibody molecules. This
antibody is conjugated (covalently attached to) the label
molecule.
• The idea is to use your painfully hand-made rabbit
antibody to bind to your protein, then buy some labelled
goat anti-rabbit antibody, so you don’t have to do the
labelling process over and over: you can use the goat
antibody with all of your rabbit antibodies.
Electrophoresis
• Electrophoresis is a protein separation technique based on movement
of the charged protein molecule in an electric field.
• Electrophoresis is done using a gel matrix, which prevents the proteins
from diffusing due to convection (temperature-driven movement of the
electrophoresis buffer).
• The most common gel matrix is polyacrylamide, giving rise to the acronym
PAGE; polyacrylamide gel electrophoresis
• Most proteins have a net negative charge, so they move to the positive
pole.
• The simplest form of electrophoresis is native gel electrophoresis.
Here, the proteins are suspended in a buffer and pulled through the gel
by the electric field.
• The speed each protein moves is determined by its size and charge.
• Individual proteins can be detected by antibodies, or by their own
enzymatic activity, or by a general protein stain like Coomassie Blue.
• This technique is useful for detecting different alleles of the same gene.
SDS Gel Electrophoresis
• SDS-PAGE is a method for separating proteins
according to their molecular weight.
• As opposed to native gel electrophoresis, which
separates on the basis of both molecular weight
and charge.
• SDS = sodium dodecyl sulfate (a.k.a. sodium
lauryl sulfate), a detergent that unfolds proteins
and coats them in charged molecules so that
the normal charge on the protein is completely
overwhelmed by the charges on the SDS
molecules.
• This means that the charge to mass ratio of all
proteins is essentially identical.
• SDS also unfolds the proteins, so their
shape does not affect migration speed.
• Also, a reducing agent like betamercaptoethanol (BME) is included, to break
any disulfide bridges between cysteines.
• SDS denaturation isn’t perfect: some proteins
behave anomalously,
Isoelectric Focusing
• Isoelectric focusing (IEF) separates proteins by their
isoelectric point, the pH at which the net surface charge is
zero.
• Recall that whether an acidic or basic amino acid is charged
depends on its pKa and the pH of the solution.
• IEF uses a mixture of ampholytes, chemical compounds that
contain both acidic and basic groups.
• When an electric field is applied, the ampholytes move to a
position in the gel where their net charge is zero, and in the
process they set up a pH gradient.
• Proteins also move down the pH gradient until they reach a
pH where they have no net charge, their isoelectric point.
• Isoelectric focusing is thus an equilibrium process: proteins
move to a stable position and stay there. (But in practical
terms, the gradient breaks down over time).
Two Dimensional Gel Electrophoresis
• Two dimensional gels are a way of separating proteins into individual
spots that can be individually analyzed.
• Proteins are first separated by their isoelectric point and then by their
molecular weight. These characteristics are relatively independent of each
other.
• First, isoelectric focusing is performed on a protein sample, running the
proteins through a narrow tube or strip of acrylamide.
• Then the IEF gel is placed on top of an SDS gel, allowing the proteins to
be separated by their molecular weight at right angles to the isoelectric
point separation.
• Then the gel is stained with a general protein stain such as Coomassie
Blue.
• Some issues:
• While a cell might contain up to 100,000 proteins, at best only 3000 spots
can be resolved.
• Proteins expressed at a low level (such as regulatory proteins) don’t show
up well: spot size is proportional to the amount present in the cell
• Special techniques are needed for membrane proteins, which aren’t easily
solubilized by the usual techniques.
• Comparing spots between 2D gels require image analysis software (and
well-standardized running conditions).
Western Blots
• The Western blot is an extension of electrophoresis: a
way of using antibodies to detect a specific protein.
• First, the proteins are separated by electrophoresis.
Usually SDS gels, but sometimes 2-D gels.
• Then the proteins are transferred to a membrane that
binds them tightly. This puts all the proteins in a single
layer, so the antibodies don’t have to diffuse through
the gel to find them.
• Usually done by electroblotting: using electricity to pull
the proteins out of the gel and onto the membrane
surface. Or, just by capillary action.
• The membrane is then incubated with the antibody,
followed by a labelled secondary antibody to detect
the protein of interest.
Mass Spectrometry
• How can you identify all those protein spots on a 2D gel?
• The general principle of mass spectrometry is that if you ionize a group of atoms or molecules, you can
separate them on the basis of charge to mass ratio, by accelerating them in an electric field in a vacuum.
• The original mass spectrometers were used to separate isotopes, based on slightly different masses.
• During the ionization process, proteins tend to break up in characteristic ways, producing molecular ions
whose molecular weights can be measured very precisely.
• Assuming you are working with an already sequenced genome, you can predict the size of fragments that
will be generated by any gene. Thus you can identify the gene product by matching the actual fragments
with list of predicted fragments.
More Mass Spectrometry
• For most protein work, the proteins are first digested into
small fragments (say , 5-10 amino acids), separated by HPLC
(high performance liquid chromatography), and then run
individually through the mass spec.
• Protein sequencing and older protein identification methods
also start with proteolytic digestion
• Endopeptidases that digest proteins at known sites are used,
such as trypsin (cleaves after Lys or Arg) and chymotrypsin
(cleaves after Phe, Trp, or Tyr).
• Ionizing the peptide needs to be done rather gently. One
common technique is MALDI (matrix-assisted laser
desorption/ionization). The proteins are mixed with the
matrix molecules, which efficiently absorb the UV laser
energy and encourage ionization of the proteins. When
irradiated with the laser, they vaporize along with the protein,
but their small size makes them easy to detect and ignore.
• Time-of-flight mass spectrometry is generally used (so the
whole thing is MALDI-TOF). The molecular ions are
accelerated in an electric field, and the time it takes them to
cross a chamber of known length is proportional to their mass
(actually, charge to mass ratio). This technique works well for
the wide range of sizes seen with peptides.
Probing Protein Structure
• The three dimensional structure of a protein determines its function.
• But, we don’t necessarily know how to determine the exact function even with very good data on the
structure.
• Three dimensional structure can be determined by two methods: X-ray crystallography and
nuclear magnetic resonance (NMR).
• Both methods require knowledge of the amino acid sequence of the protein.
• If I haven’t said this before, almost all protein sequences are derived from sequencing the DNA, then
finding genes, then translating them into amino acid sequences. Direct sequencing of proteins is
difficult, and you can only get 15-20 amino acids at the N-terminus.
X-ray Crystallography
• You can’t see any details of protein
structure with light or electron
microscopes: the atoms are too small
compared to the wavelengths used.
• X-rays have wavelengths in the proper
range: 0.1 nm, which is about the size of a
hydrogen atom.
• However, X-rays can’t be focused.
• X-rays are scattered in a very precisely
known manner by electrons: the higher
the density of electrons, the more
scattering occurs.
• By shining an x-ray beam through a crystal,
the scattering from each atom in the
repeating array reinforce or cancel each
other to form a diffraction pattern, which
looks like a series of spots.
More X-ray Crystallography
• Mathematical techniques (Fourier analysis) can
convert the spot pattern into a three dimensional
electron density map.
• Using the known protein sequence, you can fit the
amino acids to the electron density map, producing a
structure for the protein.
• But, you need crystals of the protein, and it can be
very difficult to crystallize some of them
• Some regions of protein are not held in rigid positions—they
flop around and so don’t form regular crystals.
• Nevertheless, there are maybe 100,000 known
protein structures derived from this technique.
• You can also crystallize a protein along with its ligand
molecules, to see where and how the ligand binds.
Nuclear Magnetic Resonance
• Done in aqueous solution, not on crystals
• Atomic nuclei with odd mass numbers, such as 1H, 13C, and 15N, have the
property of spin, which means that they act as tiny magnets, with a
north and south pole.
•
1H
is normal hydrogen, but 13C and 15N are isotopes that require labelling of the protein
(done by adding isotopically-labeled nutrients to the growth medium used by the
organism that makes the protein).
• When put into a strong magnetic field, these magnetic nuclei can align
either with the magnetic field or directly opposite. One orientation has a
higher energy than the other.
• Adding additional energy in pulses (radio frequency) will cause the
orientation to flip, but only if the pulse frequency matches the resonant
frequency of the nuclei
• The flipping can be detected as a absorption of energy.
• The position of a given nucleus relative to other nuclei affects the
resonant frequency: electrons in different orbitals partially shield the
nucleus from the magnetic field.
• For example, it is possible to distinguish an H in a C-H bond from a H in a O-H bond.
• The difference in resonant frequency between the atom in isolation and the atom
bound in a compound is called the chemical shift. It is measured in parts per million
(ppm)
More NMR
• Using multi-dimensional NMR on several
different types of atom, it is possible to
work out a three-dimensional structure
of a molecule, up to about 25,000
molecular weight. (with the help of a lot
of mathematics)
• Once the structure has been solved, the
result is a set of slightly different
structures. The differences come from
regions of the protein that are not rigidly
fixed: areas where the polypeptide takes
on many different conformations in
solution.
• This is a big contrast to X-ray crystallography,
which assumes proteins are very rigid
structures.