Molecular Biology of the Cell - Volume 1 - Alberts B., Bray D., Lewis J., Raff M., Roberts K., Watson J. 1994

Introduction to Cell Biology
How Cells Are Studied
Fractionation of Cell Contents

Biochemical analysis often involves the disruption of the delicate Structure of Cells. However, gentle Methods for fractionating Cell contents have now been developed, aiming to preserve the function of various cellular components. Just as tissue can be separated into its constituent cell types, cells can be separated into their functional Organelles and macromolecules. In this section, we will focus on methods that allow for the purification of organelles and Proteins. Related techniques for labeling macromolecules with radioisotopes and Antibodies, as well as highly efficient methods for analyzing DNA and Gene function, are discussed in subsequent sections.

4.4.1. Ultracentrifugation can be used to separate organelles and macromolecules [24]

There are several ways to disrupt cells: they can be subjected to osmotic Shock, ultrasonic vibration, forced through a small orifice, or ground up. During this process, cell membranes (including The Plasma Membrane and The Endoplasmic reticulum membrane) break into fragments that immediately reseal to form tiny vesicles. When disruption methods are applied gently, some organelles remain intact (nuclei, Mitochondria, the Golgi apparatus, Lysosomes, and Peroxisomes). Thus, The Cell suspension is converted into a soluble extract containing a rather crude suspension of membrane-bound particles with characteristic sizes, charges, and densities. It has been shown that with the proper choice of homogenization medium (which requires careful trial-and-error analysis for each organelle), the particles in the extract retain most of the biochemical properties characteristic of intact organelles in the cell.

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Fig. 4-42. Schematic representation of the fractionation of subcellular components from cell extracts by repeated centrifugation at progressively higher speeds. In general, the smaller the subcellular component, the higher the centrifugal force required to sediment it. Typically, the following conditions are required at different stages of centrifugation: low speed - 1000 g - 10 min, medium speed - 20,000 g - 20 min, high speed - 80,000 g - 1 h, very high speed - 150,000 g - 3 h.

After preparative centrifuges came into widespread use in the early 1940s, the Separation of various Components of the homogenate became highly feasible. Extracts of disrupted cells are fractionated by subjecting them to high-speed centrifugation (Fig. 4-41). This Treatment separates cellular components by size: larger particles move faster during centrifugation. Large components of the extract, including nuclei or unbroken cells, sediment rapidly at relatively low speeds, forming a pellet at the bottom of the centrifuge tube. At higher speeds, mitochondria sediment, and at even higher speeds and longer centrifugation periods, small closed vesicles (microsomes) and then Ribosomes are pelleted (Fig. 4-42). All of these fractions are contaminated, but if the Procedure of resuspending the pellet and centrifuging is repeated several times, many impurities will be removed.

Centrifugation is usually the first step in fractionation, separating only components that differ significantly in size. To achieve a higher degree of resolution, the homogenate must be layered in a thin band on top of a salt solution. During centrifugation, different fractions sediment at different rates, forming distinct bands that can be isolated (Fig. 4-43). To prevent mixing of the sedimenting components, the salt solution must contain an inert and highly soluble material (such as sucrose) whose density increases progressively from top to bottom, forming a density gradient.

When sedimenting through such sucrose gradients, different cell components collect into separate bands that can be isolated (see Fig. 4-43). The Sedimentation Rate of each component is determined by its size and shape and is typically expressed as a sedimentation coefficient, denoted by S (see Table 4-7). The rotor in modern centrifuges spins at speeds up to 80,000 rpm, subjecting the separating particles to forces greater than 500,000 times gravity. Under such immense forces, even relatively small macromolecules, such as tRNA or simple Enzymes, separate and distribute strictly according to their size. Measuring the sedimentation coefficient of macromolecular complexes is commonly used to determine their total mass and the number of subunits they contain.

The ultracentrifuge separates cell components not only by mass but also by buoyant density. In this case, the sample sediments in a steep gradient formed by a highly concentrated solution of sucrose or cesium chloride. Cell components move down the gradient until they reach a region where the density of the solution equals their own density. No further sedimentation of the components occurs, and they "band" at this level. Thus, a series of distinct bands is formed in the centrifuge tube, with the bands closest to the bottom of the tube containing components of maximum buoyant density. This method is so sensitive that it can separate unlabeled macromolecules from those containing heavy isotopes (¹³C or ¹⁵N). The cesium chloride density gradient centrifugation method was developed in 1957 to separate labeled and unlabeled DNA synthesized by Bacteria in the presence of NUCLEOTIDES labeled with ¹⁵N. This now-classic experiment demonstrated that METABOLISM/36.html">DNA Replication occurs in a semiconservative manner (see Section 3.2.3).

Fig. 4-43. A sample of subcellular components is layered on top of a diluted sucrose solution and sedimented at different rates depending on size. A continuous sucrose density gradient is established in the tube, with the concentration increasing toward the bottom of the tube (typically, sucrose concentrations in the range of 5-20% are used). The sucrose concentration gradient is necessary to stabilize the solution and the sedimenting bands against convection. After centrifugation, the different components can usually be collected individually. To do this, the plastic centrifuge tube is punctured and drops are collected from the bottom, as shown in the figure.

4.4.2. The details of complex intracellular processes can be deciphered at THE MOLECULAR LEVEL in cell-free systems [25]

Studying organelles or other large subcellular components isolated by ultracentrifugation is extremely important for understanding their Functions in the cell. Obtaining purified fractions of Mitochondria and Chloroplasts made it possible to establish the key role of these organelles in energy conversion processes. The separation of closed vesicles formed by fragments of smooth and rough endoplasmic reticulum allowed these vesicles to be used as functional models of intact organelles. Fractionated cell extracts, also called cell-free systems, are widely used to study intracellular processes. Only by working with cell-free extracts can the Molecular Mechanism of biological processes be established, since only in this case can the mechanism under study be investigated in its pure form—without Interference from Side Reactions occurring in the cell. The Use of cell-free systems brought the first triumphant success in studying the Mechanisms of Protein Biosynthesis. The starting point in this case was a crude cell extract capable of translating RNA molecules into protein. After repeated fractionation of this extract, ribosomes, RNA, and various enzymes were obtained, which collectively constitute the Protein Biosynthesis machinery. Once individual components were obtained in pure form, they could be added to or omitted from the system, thereby clarifying The Role of each component in protein biosynthesis. This same "in vitro Translation system" proved useful for deciphering The Genetic Code, using artificial polynucleotides of known composition as Messenger RNA (mRNA). Today, various in vitro translation systems are also used to determine the mechanisms of protein targeting to different intracellular compartments (see Section 8.6.6), as well as to identify proteins encoded by purified mRNA preparations (mRNA purification is an important step in Gene cloning Procedures (see Section 5.6.7)). Table 4-8 lists some Milestones in the history of The Development of cell extract Fractionation Methods.

Much of what we know about the molecular biology of the cell has been discovered through The Study of cell-free systems. It was in this way that the mechanisms of DNA replication, Introduction/24.html">DNA Transcription, RNA splicing, Muscle contraction, and particle transport along microtubules were elucidated. Analysis in cell-free systems implies the complete separation of all its individual macromolecular components, and in particular, all the proteins that make up the system. Methods for protein separation are discussed in subsequent sections.

Table 4-7. Some Typical Sedimentation Coefficients

Sedimentation coefficients are measured in seconds and are given by the equation (dx/dt)/w²∙ x, where x is the distance from the center of rotation in cm, (dx/dt) is the sedimentation rate (cm/s), and w is the angular velocity of the centrifuge rotor in radians per second (rad/s). Since these coefficients are represented by very small numbers, they are usually expressed in Svedberg units (S), where 1S equals 1 x 10^-13 s.

4.4.3. Chromatography can be used for protein fractionation [26]

Chromatography is currently one of the most widely used methods for protein fractionation. Initially, this method was developed for fractionating low-molecular-weight compounds, such as sugars and Amino Acids. Partition chromatography became the most widespread method, finding broad application for the separation of small molecules. In general terms, the method is as follows. A drop of the sample is applied to a special paper (paper chromatography) or a Glass or plastic plate coated with a thin layer of an inert sorbent, such as Cellulose or silica gel (Thin-Layer Chromatography). One end of the plate is then placed in a mixture of Solvents (e.g., Water and alcohol). As the solvents migrate up the plate, they carry along those sample molecules that dissolve in them. The solvents are chosen so that they are bound differently by the sorbent. As a result, sample molecules that are more soluble in the bound solvent move more slowly, while others, more soluble in the weakly sorbed solvent, move faster. After a few hours, the plate is dried, stained, and the positions of the various molecules are determined (Fig. 4-44).

Proteins are most commonly separated by Column chromatography. In this method, a mixture of molecules in solution is passed through a column containing a solid, porous matrix. Due to interactions with the matrix, different proteins pass through the column at different rates. Once the various proteins reach the bottom of the column in a specific sequence, they are collected as individual fractions (Fig. 4-45). Currently, many Different types of matrices have been developed and are in use, allowing proteins to be separated according to their charge (Ion-exchange chromatography), Hydrophobicity (hydrophobic chromatography), size (gel-filtration chromatography), or ability to bind to specific chemical groups (Affinity Chromatography).

Table 4-8. Major Milestones in the Development of Ultracentrifugation and Cell-Free Extract Preparation

Fig. 4-44. Separation of low-molecular-weight compounds by paper chromatography. The sample is applied to the origin and dried, and then, utilizing capillary action, a mixture of two solvents is passed through the paper. Different components of the sample migrate along the paper at different rates, depending on the relative solubility of the components in the solvent that is more strongly adsorbed by the paper. The introduction of this method revolutionized biochemical analysis in the 1940s.

A wide Selection of different types of matrices is commercially available (Fig. 4-46). Ion-exchange columns are packed with small, positively or negatively charged beads. When using such columns, protein fractionation occurs According to the distribution of charges On the surface of the protein molecules. Hydrophobic columns are packed with beads from which hydrophobic chains project; these columns retain proteins with exposed hydrophobic regions. Columns designed for Gel filtration are filled with tiny porous beads; when using these columns, proteins are separated by size. Small molecules penetrate inside the beads as they pass through the column, while larger molecules remain in the spaces between the beads. As a result, they pass through the column faster and emerge first. Gel filtration is commonly used both for separating molecules and for determining their size.

Figure 4-45. Separation of molecules by column chromatography. The sample is applied to the top of a cylindrical glass or plastic column filled with a permeable matrix (such as cellulose) immersed in a solvent. A large volume of solvent is then slowly pumped through the column and collected in individual test tubes from the bottom of the column. Different components of the sample migrate through the column at different rates, which forms The basis of their fractionation.

Figure 4-46. Three types of matrices used for chromatography. In ion-exchange chromatography (A), the insoluble matrix contains ions that retard molecules of the opposite charge. The following matrices are used for molecular separation: diethylaminoethyl cellulose (DEAE-cellulose) - positively charged; carboxymethyl cellulose (CM-cellulose) and phosphocellulose - negatively charged. The interaction forces between the molecules in solution and the ion exchanger are determined by the Ionic strength and pH of the eluting solution, which can be varied in a specific manner to achieve effective separation (as shown in Figure 4-47). In gel-filtration chromatography (B), the matrix is inert but porous. Low-molecular-weight compounds penetrate into the matrix particles. Finding themselves in a relatively larger volume, they pass through the column more slowly. Beads of a cross-linked polysaccharide (dextran or agarose) can be used as the matrix. Since commercial Polysaccharides are available with A wide variety of pore sizes, they can be used to fractionate molecules with molecular weights ranging from 500 to 5 x 10⁶ daltons. In affinity chromatography (C), an insoluble matrix is used that is covalently coupled to specific ligands (antibodies or enzyme substrates) that bind a specific protein. Enzyme molecules bound by the immobilized substrate can be eluted with concentrated solutions of the free substrate, while molecules bound to Immobilized Antibodies can be eluted by dissociating the antibody-antigen complex with concentrated salt solutions or solutions of low or high pH. A single run through such a column often yields a very high degree of purification.

At each step of column chromatography, the purity of a protein in the mixture increases by no more than 20-fold, making it virtually impossible to isolate an individual protein from a complex mixture in a single run. Since any single protein typically constitutes less than 1/1000 of the total cellular protein, its purification requires the successive use of several different types of columns (Figure 4-47). The method of affinity chromatography is far more efficient. This method is based on biologically important interactions occurring at The surface of protein molecules. For example, when an enzyme substrate is covalently linked to a matrix, such as polysaccharide beads, the enzyme is specifically retained by the matrix and can be eluted (washed off) in a virtually pure state. Similarly, short DNA oligonucleotides of a specific sequence can be immobilized (see Section 4.6.8) and used to purify DNA-binding proteins that recognize this nucleotide sequence on Chromosomes (see Section 9.1.8). Specific antibodies can also be coupled to the matrix; such a carrier is highly convenient for purifying proteins recognized by these antibodies. Affinity columns offer a high degree of Specificity, allowing for a very high level of purification (1000- to 10,000-fold) in a single chromatographic step.

The resolution of conventional column chromatography is limited by the heterogeneity of the matrices (such as cellulose), which causes uneven solvent flow through the column. Recently developed chromatographic resins (usually silica-based) consist of tiny spheres 3 to 10 µm in diameter, packed into a special casing to form a homogeneous column. Such columns for High-Performance Liquid Chromatography (HPLC) provide an exceptionally high level of resolution.

Because the support particles in HPLC columns are packed very tightly, the flow rate through them is negligible without high pressure. For this reason, these columns are typically housed in steel cylinders connected to a sophisticated system of pumps and tubing that provide the pressure required for a high flow rate. In traditional column chromatography, the flow rate through the column can be quite low (approximately one column volume per hour), giving the separating solutions sufficient time to equilibrate with the interior of the large matrix particles. Under HPLC conditions, rapid equilibration occurs between the solutions and the interior of the tiny spheres, so that solutes with different affinities for the matrix are efficiently separated even at high flow rates. Thus, while achieving even poor separation with conventional column chromatography used to take hours, high-quality fractionation via HPLC now takes only minutes. This is why this method is currently extremely popular for separating both proteins and small molecules.

Figure 4-47. Typical results obtained during Protein Purification by various chromatographic methods. In this case, the cell extract to be fractionated was first passed through a column packed with an ion-exchange resin (A). The column was then washed, and the bound proteins were eluted with a solution containing a gradually increasing salt concentration. Proteins with the lowest affinity for the ion-exchange resin pass through the column without being retained and are collected from the bottom of the column in the first fractions of the eluate. The remaining proteins elute according to their affinity for the ion-exchange resin. The highest salt concentration is required to elute the proteins that bind most strongly to the resin. The protein of interest eluted as a narrow peak, identified by its enzymatic activity. Fractions containing this activity were pooled and applied to a second column for gel filtration (B). The fraction of the still partially purified protein was detected by enzymatic activity; the active fractions were collected and purified to homogeneity on a column (C) containing an immobilized enzyme substrate.

Figure 4-48. The detergent sodium dodecyl sulfate (SDS) in its ionized form and the reducing agent ß-mercaptoethanol. These two Reagents are used to solubilize proteins during SDS-Polyacrylamide gel Electrophoresis.

4.4.4. Sodium dodecyl sulfate polyacrylamide-gel electrophoresis (SDS-PAGE) can be used to determine the size and subunit composition of proteins [27]

Proteins usually carry a net positive or negative charge, determined by the presence of positively or negatively charged amino acid side chains on their surface. If protein molecules are placed in an electric field, they migrate at a rate determined by their net charge, as well as by their shape and size. This phenomenon forms the basis of electrophoresis—a method for separating mixtures of proteins in free aqueous solutions or in a solid porous matrix, such as starch.

In the mid-1960s, a modified method of electrophoresis was developed: sodium dodecyl sulfate polyacrylamide-gel electrophoresis (SDS-PAGE). This method was a major step forward compared to the conventional protein analysis methods known at the time. In this technique, proteins migrate through an inert matrix—a highly cross-linked polyacrylamide gel. The gel is typically prepared by polymerizing monomers immediately before use. The pore size of the gel can be adjusted to retard the migration of specific molecules. In this process, the proteins are dissolved in a solution containing a powerful, negatively charged detergent, sodium dodecyl sulfate, or SDS (Figure 4-48).

By binding to hydrophobic Regions of the protein molecule, this detergent causes the protein molecules to unfold into long, extended chains. Upon unfolding, individual protein molecules are released from complexes with other proteins or lipid molecules and are solubilized in the detergent solution. A reducing agent, usually mercaptoethanol (Figure 4-48), is added to disrupt S-S Disulfide Bonds in the proteins. This allows for the analysis of individual Polypeptides that make up multi-subunit molecules.

What happens when a mixture of SDS-solubilized proteins is subjected to electrophoresis in a polyacrylamide gel slab? Each protein molecule binds A large number of negatively charged detergent molecules, whose total charge overwhelms the protein's intrinsic charge. Consequently, when an electric field is applied, the proteins migrate toward the positive electrode. Proteins of the same size behave similarly because, first, their native structure is completely disrupted by SDS so that their shapes are identical, and second, they bind the same amount of SDS and acquire the same negative charge. Large proteins, which carry a greater charge, experience larger driving forces but also face much greater resistance. In free solution, these effects usually cancel each other out, but in the pores of a polyacrylamide gel, which acts as a molecular sieve, larger proteins are retarded much more than smaller ones. As a result, a complex mixture of proteins is resolved into a series of bands arranged according to their molecular weight. Staining the gel with Coomassie blue dye reveals the major polypeptide fractions. Minor Proteins can be identified by silver staining, which can detect as little as 10 ng of protein in a band. Specific proteins can be identified on these gels by labeling them with antibodies coupled to radioactive isotopes, enzymes, or fluorescent Dyes. This identification is often performed after transferring the proteins from the gel onto a sheet of nitrocellulose (via 'blotting'). This method is described in more detail below in relation to nucleic acid studies (see Section 4.6.8). This protein detection method is called Western blotting.

Figure 4-49. SDS-polyacrylamide gel electrophoresis. Individual proteins form complexes with negatively charged sodium dodecyl sulfate molecules and migrate through the porous polyacrylamide gel as negatively charged SDS-Protein Complexes. Because The rate of migration under these conditions is higher for smaller polypeptides, this method can be used to estimate the molecular weight of a polypeptide chain and to study the subunit composition of a protein.

Figure 4-50. Analysis of Protein samples by SDS-polyacrylamide gel electrophoresis. The photograph shows a gel used to detect proteins present at successive stages of Enzyme Purification. The leftmost lane (lane 1) contains the complex mixture of proteins in the starting cell extract; each subsequent lane contains proteins obtained after chromatographic fractionation of the protein samples analyzed in the preceding lane (see Figure 4-47). An equal amount of protein (10 µg) was loaded into the well of each lane. Individual proteins normally appear as sharp, stained bands; the bands broaden if too much protein is present. (Courtesy of Tim Formosa.)

The method of SDS-polyacrylamide gel electrophoresis of proteins is far more powerful than any previously known protein fractionation technique, if only because it can be used to detect any protein regardless of its water solubility. This method allows for the separation of Membrane Proteins, cytoskeletal protein components, and proteins that are part of large macromolecular aggregates. Because polypeptides are separated strictly by size in this method, it can provide information about the subunit composition of any complex and the molecular weight of the proteins that form it (Figure 4-49). A photograph of a gel used to analyze the successive steps of protein purification is shown in Figure 4-50.

4.4.5. Two-dimensional gel electrophoresis can resolve more than 1000 proteins on a single gel [28]

It is well known that closely spaced bands in a gel can overlap. This effect limits the number of proteins that can be resolved by one-dimensional Separation Methods (usually no more than 50). Two-dimensional gel electrophoresis, which combines two different separation procedures, allows for the identification of more than 1000 proteins. The results are obtained in the form of a 'two-dimensional' protein map.

In this method, the first step separates proteins according to their charge. To do this, the sample is dissolved in a small volume of a solution containing a nonionic (uncharged) detergent, mercaptoethanol, and urea as a denaturing agent. In this solution, all polypeptide chains without exception are solubilized, denatured, and dissociated, without altering their intrinsic charge. The dissociated polypeptide chains are then separated by isoelectric focusing, a technique based on The change in a protein's charge as the pH of the environment changes. Each protein is characterized by an isoelectric point—the pH value at which the net charge of the protein molecule is zero, and consequently, the protein ceases to migrate in an electric field. In isoelectric focusing, proteins are subjected to electrophoresis in a narrow tube filled with a polyacrylamide gel in which a pH gradient is established using specialized buffers. Under the Influence of the electric field, each protein migrates to the zone of the gradient that corresponds to its isoelectric point and remains focused there (Figure 4-51). This constitutes the Separation of proteins in the first dimension of two-dimensional gel electrophoresis.

Figure 4-51. Separation of protein molecules by isoelectric focusing. At low pH values (high H⁺ ion concentration), the carboxyl groups of proteins tend to remain uncharged (—COOH), while the basic, nitrogen-containing groups of proteins are fully charged (e.g., -NH₃⁺), giving the proteins a net positive charge. At high pH values, the carboxyl groups are negatively charged (—COO^-), and the basic groups tend to remain uncharged, e.g., (NH₂). As a result, proteins acquire a net negative charge (see Fig. 2-8). At its isoelectric point, a protein is uncharged because its positive and negative charges are balanced. Consequently, if a tube containing a solution with a fixed pH gradient is subjected to a strong electric field, each type of protein will migrate until it forms a sharp band in the pH zone corresponding to its isoelectric point, as shown in the figure.

In the second step, the gel tube containing the separated proteins is subjected to electrophoresis again, this time in a direction perpendicular to that of the first step. In this case, electrophoresis is carried out in the presence of SDS, and the proteins are separated according to their molecular weight, as in one-dimensional SDS-PAGE. The original gel is equilibrated with sodium dodecyl sulfate and, after placing it on top of an SDS-PAGE gel slab, electrophoresis is performed, during which each polypeptide chain migrates through the gel slab to form a distinct band. This achieves separation in the second dimension of two-dimensional gel electrophoresis. Consequently, only those proteins that are indistinguishable in both their isoelectric point and molecular weight remain unresolved; such a combination is extremely rare.

Figure 4-52. Fractionation of E. coli cell proteins by two-dimensional polyacrylamide gel electrophoresis. Each spot corresponds to an individual polypeptide chain. First, proteins were separated according to their isoelectric points by isoelectric focusing from left to right. Then, in the presence of SDS, they were separated by electrophoresis from top to bottom according to the molecular weight of their subunits. Note that the Abundance of different proteins in the cell varies. (Courtesy of Patrick O'Farrell.)

Table 4-9. Major milestones in the development of chromatography and electrophoresis methods and their application to the separation of biological macromolecules

Using various Protein Staining Methods, or autoradiography in the case of radioactively labeled proteins (see Section 4.5.2), trace amounts of virtually all polypeptide chains can be detected. Up to 2000 individual polypeptide chains can be resolved at one time by two-dimensional gel electrophoresis; this is sufficient to detect most bacterial proteins (Fig. 4-52). The resolution of this method is so high that it can separate two virtually identical proteins differing by a single charged amino acid. Table 4-9 introduces the major milestones in the development of chromatography and electrophoresis methods.

Figure 4-53. Peptide mapping ("fingerprinting") of proteins. In this case, the protein was cleaved with Trypsin to yield a mixture of small polypeptide fragments. This mixture was fractionated in two dimensions: by electrophoresis and by partition chromatography. The resulting spot pattern is characteristic of the protein.

4.4.6. Selective Protein Cleavage yields a characteristic set of peptide fragments [29]

Molecular weight and isoelectric point are characteristic parameters of a protein. However, precise identification of a protein molecule relies on determining its amino acid sequence. Even at the first step of this process, which involves cleaving the protein into small fragments, significant information about the protein can be obtained. Currently, Proteolytic Enzymes and chemical reagents that cleave proteins at specific amino acid residues are commercially available (Table 4-10). For example, the enzyme trypsin cleaves Lysine and Arginine residues on the carboxyl side; the chemical reagent Cyanogen bromide cleaves peptide bonds following Methionine residues. Because these specific enzymes and reagents cleave a limited number of bonds in a protein molecule, their action yields a mixture of large Peptides. By separating this mixture using electrophoresis or chromatography, a peptide map characteristic of the protein under study can be obtained. Such peptide maps are sometimes called protein "fingerprints" (Fig. 4-53).

Table 4-10. Some reagents used for the cleavage of peptide bonds in proteins

Specificity is shown for the amino acids on each side of the cleaved bond. After cleavage, the carboxyl group of amino acid 1 is released; this amino acid is located to the left of the peptide bond in standard notation (see Scheme 2-5).

This method was developed in 1956 to compare normal hemoglobin with the mutant form of the same protein found in the Blood of sickle-cell anemia patients. The mutant protein was found to differ from the normal one by a single amino acid. This provided the first proof that a mutation can result in the substitution of just one amino acid in a protein.

4.4.7. Automated instruments can analyze short Amino acid sequences [30]

Once the protein has been cleaved into small fragments, the next step is to determine The amino acid sequence of each isolated peptide fragment. This is done through a series of Chemical Reactions first proposed in 1967. First, the peptide is treated with a reagent that reacts only with the free amino group at its N-terminus. Next, this reagent is activated by exposure to a weak acid. It then specifically cleaves the peptide bond joining the N-terminal amino acid to the peptide chain; the released amino acid is subsequently identified by chromatography. The remaining peptide is thus shortened by one amino acid. It is subjected to the same sequence of reactions repeatedly until every amino acid in the peptide has been determined.

The cyclic nature of these reactions allows the entire process to be automated. Today, instruments (amino acid sequenators) are commercially available that automatically sequence amino acids in peptide fragments. In the final step of the analysis, the amino acid sequences obtained for the peptide fragments are aligned in the order they occurred in the intact chain. This is achieved by comparing the sequences of overlapping sets of fragments generated by cleaving the same protein with different proteolytic enzymes.

Improvements in protein sequencing technology have significantly increased its speed and sensitivity, allowing the analysis of minute sample amounts. For example, a sequence of several dozen Amino acids can now be determined using only a few micrograms of protein—the amount that can be recovered from a single band on an SDS-polyacrylamide gel. This has proven extremely important for studying many low-abundance cell proteins, such as steroid or polypeptide Hormone Receptors. Today, determining just 20 amino acids of a protein is sufficient to design a DNA probe for cloning the corresponding gene (see Section 5.6.5). Once the gene is isolated, the remaining undetermined portion of the protein's amino acid sequence can be deduced from The nucleotide sequence according to the genetic code. This is a major achievement, because even with full automation, determining the complete primary sequence of a protein remains a highly challenging task. For example, if a protein consists of 100 amino acids, its sequence can be determined in about a month of hard work. However, as the amino acid chain length increases, the complexity rises rapidly, preventing amino acid sequencing from becoming a routine procedure. Given that DNA Sequencing is easier and less time-consuming (see below), the amino acid sequences of most proteins are now typically deduced from The nucleotide sequences of their corresponding genes.

Summary

Cell populations can be analyzed biochemically by disrupting the cells and fractionating their contents by ultracentrifugation. Further fractionation allows the Reconstitution of functional cell-free systems; such systems are essential for defining the molecular details of complex cellular processes. For example, protein synthesis, DNA replication, RNA splicing, and various types of Intracellular Transport have recently been analyzed using this approach.

Major proteins in soluble cell extracts can be purified by column chromatography; depending on the type of column matrix, biologically active proteins can be separated based on their molecular weight, hydrophobicity, net charge, or affinity for other molecules. During purification, the sample is typically passed through several columns in succession, with the enriched fractions from one column applied to the next. Once a protein is purified to homogeneity, its biological activity is thoroughly characterized. A small fragment of the protein's amino acid sequence can also be determined to clone its gene; the remainder of the amino acid sequence is then deduced from the nucleotide sequence of the gene.

Table 4-11. Use of some radioactive isotopes in biological research

¹⁾ Isotopes are listed in order of decreasing energy of their emitted electrons. ¹³¹I also emits γ-rays. Half-life is the time required for 50% of the atoms of a given isotope to decay.

Even if The amount of protein is extremely small, its molecular weight and subunit composition can be determined using SDS-polyacrylamide gel electrophoresis. In two-dimensional electrophoresis, proteins are separated into individual fractions by isoelectric focusing in the first dimension, followed by SDS-electrophoresis in the second dimension. This method can be used to separate proteins that are normally considered insoluble.