3.5

The three most widely used characteristics for separating proteins are [blank_start]size[blank_end], defined as either length or mass; [blank_start]net electrical charge[blank_end]; and [blank_start]affinity[blank_end] for specific ligands.

The first step in a typical protein purification scheme is [blank_start]centrifugation[blank_end]

Proteins vary greatly in mass, but not in [blank_start]density[blank_end].

The [blank_start]sedimentation constant[blank_end], s, of a protein is a measure of its sedimentation rate. The sedimentation constant is commonly expressed in [blank_start]Svedberg[blank_end] units (S), where a typical large protein complex is about 3–5S, a proteasome is 26S, and a eukaryotic ribosome is 80S.

The most common initial step in protein purification from cells or tissues is the separation of water-soluble proteins from insoluble cellular material by [blank_start]differential centrifugation[blank_end]. A starting mixture, commonly a cell homogenate (mechanically broken cells), is poured into a tube and spun at a rotor speed, and for a period of time, that forces cell organelles such as nuclei as well as large unbroken cells or large cell fragments to collect as a pellet at the bottom; the soluble proteins remain in the supernatant

On the basis of differences in their masses, water-soluble proteins can be separated by centrifugation through a solution of increasing density, called a [blank_start]density gradient[blank_end].

All the proteins start from the thin layer of the sample that was placed at the top of the tube and separate into bands (actually, disks) of proteins of different masses as they travel at different rates through the density gradient. In this separation technique, called [blank_start]rate-zonal centrifugation[blank_end], samples are centrifuged just long enough to separate the molecules of interest into discrete bands, also called zones.

rate-zonal centrifugation is effective in determining precise molecular weights

In [blank_start]two[blank_end]-[blank_start]dimensional[blank_end] gel electrophoresis, proteins are separated sequentially, first by their [blank_start]charges[blank_end] and then by their [blank_start]masses[blank_end]

In two-dimensional gel electrophoresis, proteins are separated sequentially, first by their charges and then by their masses (Figure 3-39a). In the first step, a cell or tissue extract is fully denatured by high concentrations (8 M) of urea (and sometimes SDS) and then layered on a strip of gel that contains urea, which removes any bound SDS, and a continuous pH gradient. The pH gradient is formed by ampholytes, polyanionic and polycationic small molecules that are cast into the gel. When an electric field is applied to the gel, the ampholytes will migrate. Ampholytes with an excess of negative charges will migrate toward the anode, where they establish an acidic pH (many protons), while ampholytes with an excess of positive charges will migrate toward the cathode, where they establish an alkaline pH. The careful choice of the mixture of ampholytes and careful preparation of the gel allows the construction of stable pH gradients ranging from pH 3 to pH 10. A charged protein placed at one end of such a gel will migrate through the gradient under the influence of the electric field until it reaches its [blank_start]isoelectricpoint[blank_end] (pI), the pH at which the net charge of the protein is [blank_start]zero[blank_end]. With no net charge, the protein will migrate no further. This technique, called [blank_start]isoelectric focusing[blank_end] (IEF), can resolve proteins that differ by only one charge unit. This method is sensitive enough to separate phosphorylated and nonphosphorylated versions of the same protein.

Proteins that have been separated on an IEF gel can then be separated in a second dimension on the basis of their molecular weights. To accomplish this separation, the IEF gel strip is placed lengthwise on one outside edge of a square or rectangular slab of polyacrylamide gel, this time saturated with SDS to confer on each separated protein a more or less constant [blank_start]charge[blank_end]:[blank_start]mass[blank_end] ratio.

In this technique, called [blank_start]liquid chromatography[blank_end] (LC), the sample is placed on top of a tightly packed column of spherical beads held within a glass, metal, or plastic cylinder ( Figure 3-40). The sample then flows down the column, driven by gravitational or hydrostatic forces alone or sometimes with the assistance of a pump.

Proteins that differ in mass can be separated on a column of porous beads made from polyacrylamide, dextran (a bacterial polysaccharide), or agarose (a seaweed derivative)—a technique called [blank_start]gel filtrationchromatography[blank_end]

Although proteins flow around the beads, they spend some time within the large depressions that cover a bead’s surface. Because smaller proteins can penetrate these depressions more readily than larger proteins can, they travel through a gel filtration column more [blank_start]slowly[blank_end] than larger proteins do (Figure 3-40a). (In contrast, proteins migrate through the pores in an [blank_start]electrophoretic[blank_end] gel; thus smaller proteins move [blank_start]faster[blank_end] than larger ones.)

In ion-exchange chromatography, proteins are separated on the basis of differences in their pH.

The ability of proteins to bind specifically to other molecules is the basis of affinity chromatography. In this technique, ligands or other molecules that bind to the protein of interest are covalently attached to the beads used to form the column.

The purification of a protein, or any other molecule, requires a specific [blank_start]assay[blank_end] that can detect the presence of that molecule as it is separated from other molecules (e.g., in column or density-gradient fractions or gel bands or spots).

Many assays are tailored to detect some functional aspect of a protein. For example, assays of enzymatic activity are based on the ability to detect the loss of substrate or the formation of product, these are called [blank_start]Chromogenic Enzyme Reactions[blank_end]

The classic method for determining the amino acid sequence of a protein is [blank_start]Edman degradation[blank_end]

In this procedure (Edman degredation), the free [blank_start]amino group[blank_end] of the N-terminal amino acid of a polypeptide is labeled, and the labeled amino acid is then cleaved from the polypeptide and identified by [blank_start]high-pressure liquidchromatography[blank_end]. The polypeptide is left one residue shorter, with a new amino acid at the N-terminus. The cycle is repeated on the ever-shortening polypeptide until all the residues have been identified.

A [blank_start]peptide mass fingerprint[blank_end] is the list of the molecular weights of peptides that are generated from the protein by digestion with a specific protease, such as trypsin.

In regards to X-Rar Crystallography: Elaborate calculations and modifications of the protein (such as the binding of heavy metals) must be made to interpret the diffraction pattern and calculate the distribution of electrons (called the [blank_start]electron density map[blank_end]).

Protein crystals are relatively easy to crystallize making X-Ray Crystallography an almost universal solution for determining the 3D structure of proteins.

In this technique, a dilute protein sample in an aqueous solution is applied in a thin layer to an electron microscope sample holder (a “grid”) and rapidly frozen in liquid helium to preserve its structure. It is then examined in the frozen, hydrated state in a [blank_start]cryoelectron[blank_end] microscope.

An important distinction between x-ray crystallography and NMR spectroscopy is that the former method directly determines the [blank_start]locations[blank_end] of the atoms, while the latter directly determines the [blank_start]distances[blank_end] between the atoms, from which the structure is deduced.

Centrifugation separates proteins on the basis of their rates of [blank_start]sedimentation[blank_end], which are influenced by their masses and shapes

[blank_start]Electrophoresis[blank_end] separates proteins on the basis of their rates of movement in an applied electric field. SDSpolyacrylamide gel electrophoresis (SDS-PAGE) can resolve polypeptide chains differing in molecular weight by 10 percent or less (see Figure 3-38). Two-dimensional gel electrophoresis provides additional resolution by separating proteins first by [blank_start]charge[blank_end] (first dimension) and then by [blank_start]mass[blank_end] (second dimension).

[blank_start]Liquid chromatography[blank_end] separates proteins on the basis of their rates of movement through a column packed with spherical beads. Proteins differing in mass are resolved on gel filtration columns; those differing in charge, on ionexchange columns; and those differing in ligand-binding properties, on affinity columns (see Figure 3-40).

Antibodies are powerful reagents used to detect, quantify, and isolate proteins.

[blank_start]Immunoprecipitation[blank_end], often abbreviated as IP, permits the separation of a protein of interest from other proteins in a complex mixture using antibodies specific for the protein of interest. The antibodies are used to precipitate their target protein out of solution for subsequent analysis. Molecules tightly bound to the target protein can precipitate with it

[blank_start]Pulse[blank_end]-[blank_start]chase[blank_end] experiments can determine the intracellular fate of proteins and other metabolites

[blank_start]Autoradiography[blank_end] is a technique for detecting radioactively labeled molecules in cells, tissues, or electrophoretic gels using two-dimensional detectors (photographic emulsion or electronic detectors).

Isotopes, both radioactive and nonradioactive, play a key role in the study of proteins and other biomolecules. They can be incorporated into molecules without changing the chemical composition of the molecule or as add-on tags. They can be used to help detect the synthesis, location, processing, and stability of proteins

[blank_start]X-ray crystallography[blank_end] provides the most detailed structures but requires protein [blank_start]crystallization[blank_end]. [blank_start]Cryoelectron[blank_end] microscopy is most useful for large protein complexes, which are difficult to crystallize. Only relatively small proteins are amenable to [blank_start]NMR[blank_end] three-dimensional structural analysis.