Proteomics analytical methods

2.1 Electrophoresis gels

Two-dimensional gel electrophoresis (2-DE) is currently one of the fundamental tools to display and evaluate the proteome complexity of any organism. It enables the separation of heterogeneous mixtures of proteins on a single polyacrylamide gel according to isoelectric point and molecular weight. The introduction of 2-DE in 1975 by O’Farrell, Klose and Scheele [4-6] allowed for separating between 2000 and 3000 proteins in a single gel. The isoelectric focalization (IEF) is the electrophoretic technique in which the proteins are separated according to their isoelectric point (pI) through a pH gradient. The pI of each protein corresponds to the pH at which the net charge of the protein is equal to zero. Under an electric field the proteins migrate until they reach their pI. Two methods are available for IEF.

In the first one, the pH gradient is created and maintained by passing an electric current through a solution containing amphoretic compounds. These compounds are molecules with particular pKs. By mixing them in the polyacrylamide gel, it is possible to form a mixture, which establishes a pH gradient. In the second IEF technique, an immobilized pH gradient (IPG) is used [7]. In this process, the pH gradient, which exists prior to the IEF, is co-polymerized within the fibers of the polyacrylamide matrix. This type of pH gradient is achieved through the use of a set of weak acids and bases named Immobilines. Different types of IPG strips can be used with different ranges of pH. Most of the laboratories running 2-DE gels are currently using strips with a pH range from 3.5 to 10 to display a wide range of proteins. However, IPG strips with narrow pH range can also be used to create a biochemical zoom on a part of the gradient range (like 4-7 IPG or 2-5 IPG). This provides higher protein concentration loading and the investigation of more proteins on a proteome “window”.

After the first dimension of separation (IEF), the IPG gel containing proteins needs to be equilibrated in a solution containing SDS (Sodium Dodecyl Sulfate). It is an anionic detergent that binds to the majority of proteins in a constant mass ratio (1.4g of SDS per gram of proteins). SDS allows the proteins to acquire a net charge per mass unit that is approximately constant. During this equilibration step, proteins are first reduced using DTT and then alkylated by incubation with Iodoacetamide. Once the equilibrium is achieved, the IPG strip is loaded on the top of a polyacrylamide gel. In this second dimension of the 2-DE technique,

the polypeptides enter into the polyacrylamide gel under an electric field and migrate through the porosity strictly according to their size (the molecular weight, M_r).

After the second dimension, the polypeptides must be detected either by staining, radiation (for radio-labeled proteins) or by immunologic assays. Silver staining is one of the most popular methods of detection due to its very high sensitivity (<1ng). The main drawback of this method is the incompatibility with mass spectrometry because of the presence of glutaraldehyde, which cross-links proteins. To overcome this problem, a silver staining method without glutaraldehyde has been introduced (about ten times less sensitive) [8].

Coomassie blue staining is also very popular. It is a very simple method, though 50 times less sensitive than standard silver staining. Other methods are also used such as fluorescence or negative staining with zinc. After staining, the 2-DE gels can be scanned and are ready to be analyzed through informatics software developed to perform quantitative protein analysis and automatic comparison of gels.

Even though 2-DE gels are considered the gold standard of proteomics, some issues remain uncertain. Due to analytical problems, in between-gel variations (up to 30% coefficient of variation) lead to the problematic detection and quantification of differences in protein expression, resulting in difficulties in distinguishing biological from experimental variations.

The two-dimensional difference gel electrophoresis (DIGE [9]) partially circumvents these problems. The DIGE method labels each sample with a spectrally resolvable fluorescent dye (Cy2, Cy3 or Cy5) prior to electrophoresis. The labeled samples are then mixed before IEF and resolved on the same 2-DE gel. Different images are acquired using different wavelengths and are compared with image analysis software. Variation in spot intensities due to gel-specific experimental factors is the same for each sample within a single DIGE gel.

Consequently, the relative amount of a protein in a gel in one sample compared to another is unaffected.

Yet, other issues remain unsolved in the 2-DE technique. This approach does not allow for identifying and characterizing all proteins present in a biological sample. Some proteins will not appear on the gel because of their extreme physico-chemical properties, such as exceedingly high or low molecular weight, exceedingly high pI, extreme hydrophobicity or insufficient abundance. Other techniques must therefore be applied to complete the picture, such as one-dimensional electrophoresis (1-DE, which separates samples only by molecular

weight), pre-treatment of the starting material, pre-fractionation, and early digestion of the protein mixture followed by liquid chromatography.

2.2 Liquid chromatography

The first chromatographic experiments were carried out in 1906 by the Russian botanist Mikhaïl Tswett, when he succeeded in separating various vegetal pigments. Chromatography is an analytical method for separating the different components of a mixture of molecules. It relies on differential affinities of these molecules for a mobile phase, the medium that carries the molecules, and for a stationary adsorbing medium, through which they pass. The stationary phase is either a solid or a liquid and the mobile phase is either a gas or a liquid.

The separation is achieved through the different rates of migration of each component of a complex mixture, depending on their affinity for the stationary phase. Three types of chromatography have been developed: thin layer chromatography (TLC); gas chromatography (GC) and liquid chromatography (LC) [10-13].

In LC the mobile phase is a liquid and the stationary phase can be either a liquid or a solid.

The role of the liquid mobile phase is first to solubilize the protein sample and second to carry the proteins along the column. Various types of LC have been developed, depending on the type of stationary phase. The separation can be based on partitioning, adsorption, ion exchange, size exclusion or affinity.

In partitioning LC, the stationary phase is composed of an organic liquid covalently fixed on an inert solid support (such as silica) or embedded in a porous inert support. The principle of separation is based on the partitioning coefficient of the proteins between the non-miscible stationary and mobile phases. The higher the partitioning coefficient, the higher the number of soluble proteins in the organic stationary phase and the higher the retention time for these proteins [14]

In adsorption LC, the stationary phase is composed of a solid adsorbent. The proteins are separated due to their varying degree of adsorption onto the solid surfaces. Different types of adsorbent can be used. In normal phase absorption, the stationary phase is composed of a polar compound (such as an amine or carboxyl) linked to silica beads. In contrast, in reverse phase LC the stationary phase is composed of a non-polar hydrophobic compound. In general,

the hydrophobic compounds are aliphatic chains such as C₈ or C₁₈ bound to silica beads.

When the proteins are loaded to the top of the column they bind to aliphatic compounds through hydrophobic interactions. The proteins are eluted with increasing concentration of a hydrophobic solvent such as acetonitrile. The low-hydrophobic proteins are eluted first by low concentration of solvent and the high-hydrophobic proteins are eluted only by a high concentration of solvent [15]

Ion exchange LC is used to separate proteins on the basis of their electric charge. In this method, ionic groups are covalently bound to the matrix column. These groups can be either positively or negatively charged and are compensated for by small concentrations of counter ions that are present in the buffer. When a sample is added to the column, an exchange with the weakly bound counter ions takes place. With a cationic exchange column the matrix is negatively charged. The higher the positive charges of proteins, the higher are their ionic interactions with the matrix. Increasing concentrations of salt solution are then used to elute the bound peptides. The principle is the same with anionic exchange, except that the matrix will be positively charged and the most negatively charged proteins have higher retention times [16]

Size exclusion LC is used to separate proteins by their size and shape. Its principle is to use a porous polymer composed of beads with very small holes. As the protein solution is poured on the column, small molecules enter the pores in the beads. The small molecules are eluted last because they have a longer path to go through, as they get stuck over and over again in the maze of pores running from bead to bead [17]

Finally, affinity chromatography relies on the biological ability of a protein to bind to a column. The most common type involves a ligand, a specific small biomolecule. This small molecule is immobilized and attached to a column matrix, such as cellulose or polyacrylamide. The target protein is then passed through the column and binds to it by its ligand, while other proteins elute out. This is a very efficient purification method since it relies on the biological specificity of the target protein, such as the affinity of an enzyme for its substrate [18]

Currently, a large number of proteomics projects couple LC with mass spectrometry, which is described in the following section.

2.3 Mass spectrometry

Mass Spectrometry is an analytical technique that is commonly used to identify unknown compounds, quantify known materials and elucidate the structural and physical properties of ions. It is a technique associated with very high levels of specificity and sensitivity. Analyses can often be accomplished with tiny quantities, sometimes requiring less than picogram amounts of material. Sir J. J. Thomson developed the first mass spectrometer in the first decade of the 20th Century, even before the determination of mass-to-charge ratios (m/z) of ions. The goal of all mass spectrometers is the exact determination and analysis of the m/z ratio of ions. A mass spectrometer is formed of three fundamental parts, namely the ionization source, the analyzer, and the detector. The first step in a mass spectrometric process is to generate ions from the analyte. Nowadays, the two ionization methods used most frequently for biochemical analyses are Electrospray Ionization (ESI) and Matrix Assisted Laser Desorption Ionization (MALDI). However, other methods also exist such as Chemical Ionization (CI), Electronic Ionization (EI) or Fast Atom Bombardment (FAB).

Electrospray Ionization (ESI), developed by Fenn and colleagues [19] is one of the Atmospheric Pressure Ionization (API) techniques and is well-suited for the analysis of polar molecules ranging from less than 100 Da to more than 1’000’000 Da in molecular weight. In ESI, the sample is introduced into a capillary and then submitted to a high voltage (~4000V).

The high voltage causes the formation of a cloud of charged droplets. Evaporation of the solvent from the initially formed droplet as it traverses a pressure gradient toward the analyzer leads to a reduction in diameter and an increase in surface field until the Rayleigh limit is reached. The latter corresponds to the point where the surface tension can no longer sustain the charge. At this point, an explosion occurs, and the droplet is ripped apart. This produces smaller droplets that can repeat the process until obtaining singly or multiply-charged ions.

The Nobel Laureate, Koichi Tanaka [20] has developed the Matrix Assisted Laser Desorption Ionization (MALDI) method in 1987. It is based on the bombardment of molecules with a laser light to provoke sample ionization. Most commercially available MALDI mass spectrometers now have a pulsed nitrogen laser of 337 nm wavelengths. In MALDI, the sample is first mixed with a highly absorbing matrix compound. The matrix co-crystallizes with the sample on the metal support. The matrix minimizes the degradation of the sample caused by the energy absorption of the incidental laser beam. The exact mechanism by which

the MALDI ionization occurs is not precisely understood. It is supposed that the matrix absorbs the energy transmitted by the laser, and this contribution of energy causes its expansion in gas phase by involving the molecules of the sample. The sample is mainly ionized by transfer of protons, either before desorption in the solid phase or by collision after desorption with the excited matrix to form singly-charged ions.

Once obtained, ions must be analyzed. The analyzer uses dispersion or filtering to sort out ions according to their mass-to-charge ratios. Several types of analyzers have been developed such as the Quadrupole, the Ion Trap or the Time of Flight (TOF). The Quadrupole mass filter consists in four parallel rods to which an electric field is applied. The resultant magnetic field allows for selecting ions according to their m/z ratio. Indeed, at one particular magnetic field strength, only ions with a particular m/z will pass trough the quadrupole filter and all other ions are thrown out of their original path. The Ion Trap mass analyzer consists of three hyperbolic electrodes: the ring electrode, the entrance end-cap electrode and the exit end-cap electrode. These electrodes form a cavity in which it is possible to trap and analyze ions. Both end-cap electrodes have a small hole in their center through which the ions can travel. Ions produced from the source enter the trap through the inlet focusing system and the entrance end-cap electrode. Various voltages can be applied to the electrodes to trap and eject ions according to their mass-to-charge ratios.

Time-of-Flight mass analyzers separate ions by virtue of their different flight times over a known distance. A brief burst of ions is emitted from a source. These ions are accelerated so that those with the same charge have an equal kinetic energy and then are directed into a flight tube. Since kinetic energy is equal to 1/2 mv² (where m is the mass of the ion and v is the ion velocity) the lower the mass ion, the greater is the velocity and the shorter is the flying time.

The travel time from the ion source through the flight tube to the detector, measured in microseconds, can be transformed onto an m/z value through the relationships described above. Because all ion masses are measured for each ion burst, TOF mass spectrometers offer high sensitivity as well as rapid scanning. They can also provide mass data for very high-mass bio-molecules [21]

Two analyzers can be combined to perform tandem mass spectrometry (MS/MS). The first analyzer selects a first ion, which is called the “parent ion”. After fragmentation of the parent ion by collision-induced dissociation (CID, a fragmentation obtained by the collision of a

molecule with neutral gas molecules), a second analyzer measures the m/z ratio of the ions resulting from this fragmentation. As fragmentation occurs mainly in the peptide bond of the amino-acid-chain backbone, a ladder of sequence ions is generated. The resulting peptide fragmentation masses differ by the mass of amino-acid residues, thus allowing stretches of the peptide sequence to be deduced, which can be very useful to identify compounds in complex mixtures [22]. Fragments will only be detected if they carry at least one charge. According to Roepstorff’s nomenclature [23] the product ions are denoted as a, b, and c, when the charge is retained on the N-terminal side of the fragmented peptide, and x, y and z when the charge is retained on the C-terminal side. As shown in Figure 2, ion types differ by the position of the fragmentation in respect with the peptide bond.

Figure 2: Fragmentation schema of a peptide with four amino acids. Fragments that carry the charge on the N-terminal side are denoted a, b and c, while those carry the charge on the C-terminal are denoted x, y and z.

Once the ion passes through the mass analyzer, the ion detector (the final element of the mass spectrometer) then detects it. The detector allows a mass spectrometer to generate an electric current signal from incident ions by generating secondary electrons, which are further amplified. Alternatively, some detectors operate by inducing an electric current generated by a moving charge. Among the described detectors, the electron multiplier and scintillation counter are the most commonly used and convert the kinetic energy of incident ions into a cascade of secondary electrons.

Ideally, the MS and MS/MS processes should generate a list of m/z values of all peptides (respectively fragmented ions) present in the analyzed sample. In reality, due to physico-chemical properties of some peptides or ions, not all m/z values are detected making the data interpretation much harder.

Finally, mass spectrometry can be used in combination with analytical separation methods such as liquid chromatography (LC-MS) or gas chromatography (GC-MS). In these cases mass spectrometry is used as a detector after chromatographic separation. Nowadays, this configuration is highly used in many laboratories because it allows for combining high separation and detection capacities [24]. Shotgun proteomics has evolved from the combination of some of these techniques, MS and multidimensional chromatography, and bioinformatics. In a typical shotgun proteomics approach, a complex protein mixture is digested with proteases to produce an even more complex peptide mixture. The peptides are loaded directly onto an LC/LC column placed in-line with an MS/MS spectrometer. The spectra are acquired “on the fly” as peptides are eluted from the column, ionized, and emitted into the mass spectrometer. Using elaborated algorithms, respective peptide sequences generated from MS/MS are automatically identified by comparison against protein databases, avoiding hence the manual interpretation. One such approach is the multidimensional protein identification technology (MudPIT [25]) which has been applied to the proteome study of various species such as yeast, E. coli [26], human extracts [27], Toxoplasma gondii [28]etc.

2.4 Protein chips

The necessity to miniaturize and to automate high-throughput analysis systems led to the development of microarray (biochips) technology. It is based on immobilizing small molecules, oligonucleotides or proteins onto surfaces for various high-throughput screening studies. Microarrays were initially developed for large-scale gene expression studies, for example to compare gene expressions between different tissue types, treatments, disease models, and human samples (see Chapters 25–28). Today, DNA microarrays are well established for studying the transcriptional state of a biological sample [29]. However, the level of expression of a transcribed mRNA does not always correlate with the protein expression level [30]. Protein microarrays were thus developed to analyze the expression level of a large number of proteins simultaneously and to study the interaction of proteins with a variety of molecules. It is already a successful method for the identification, quantitation and functional analysis of proteins in proteome research [31].

There are two main types of protein microarrays used: i) to measure the abundance of a protein (abundance-based microarray) by the use of a specific reagent, and ii) to study protein

function. Abundance-based microarrays are divided in two types: capture microarrays and reverse phase microarrays. In capture microarrays, a molecule (such as antibodies, peptide, RNA or DNA aptamers, or chemical molecules) is spotted on the surface of the microarray to catch and assay their target from a complex mixture. The relative amounts of the targeted proteins are then determined by comparison with a reference sample [32]. In this technique, the most popular capture molecules are antibodies because of their sensitivity and selectivity.

function. Abundance-based microarrays are divided in two types: capture microarrays and reverse phase microarrays. In capture microarrays, a molecule (such as antibodies, peptide, RNA or DNA aptamers, or chemical molecules) is spotted on the surface of the microarray to catch and assay their target from a complex mixture. The relative amounts of the targeted proteins are then determined by comparison with a reference sample [32]. In this technique, the most popular capture molecules are antibodies because of their sensitivity and selectivity.