Fundamentals of Biochemistry - A. A. Anisimov 1986

Proteins
Physicochemical Properties of Proteins

2.3.1. Protein Extraction and Purification. The initial step in protein Isolation and Purification is their extraction from Cells. To achieve this, cells are disrupted, i.e., converted into a homogenate (breaking Cell walls, outer membranes, and cytoplasmic structures). Various Methods of cell and tissue homogenization exist, and the choice of a specific method depends on the properties, composition, and structural strength of the starting material. Most animal cells are disrupted relatively easily, whereas working with plant and bacterial cells presents significant challenges due to the presence of cell walls.

A fairly simple and accessible disruption method is homogenization via grinding Cells and Tissues with a solid abrasive material (e.g., quartz sand) in a mortar and pestle in the presence of a suspension medium. The drawback of this method is that it can disrupt The Structure of larger Organelles, such as METABOLISM/14.html">Chloroplasts.

To disrupt animal cells, homogenizers with rotating blades or a pestle—typically made of an inert material such as Teflon—are frequently employed. For the disintegration of microbial cells, mechanical shaking of The Cell suspension with small Glass beads (ranging from 50 to 500 µm in diameter) is used. Shaking at a frequency of 300–3000 oscillations/min destroys the cells; however, the resulting intense vibration can sometimes also cause the destruction of cellular organelles. Homogenization of microbial cells can likewise be achieved through high pressure using specialized presses. In this process, the cell suspension is placed in a chamber, excessive pressure is applied (up to 1⋅10⁸ Pa), and it is then rapidly released. The abrupt pressure drop causes the cells to rupture. Cells can also be homogenized by forcing them through microscopic orifices.

Specialized devices known as ultrasonic disintegrators are widely utilized for Cell Disruption. However, because ultrasound is a protein-denaturing agent, the operating parameters of the device must be selected with great care to prevent protein inactivation. Additional methods for cell lysis include osmotic Shock, alternate freezing and thawing, Treatment with organic Solvents (chilled acetone, toluene), and autolysis. For the removal of plant and bacterial cell walls, enzymatic preparations such as Lysozyme, cellulolytic enzyme complexes, and chitinase can be applied.

Throughout all Stages of Protein isolation and purification, one must account for their high instability, lability, and propensity to lose their natural, native properties—in other words, to undergo Denaturation. Proteins are exceptionally sensitive to many chemical Reagents (including acids, alkalis, and organic solvents) and denature upon heating. They adsorb readily onto various surfaces (such as the porcelain of mortars), leading to protein loss. Protein autolysis (self-Digestion) and degradation by microorganisms introduced into the solution from air or Water are also potential risks.

In most cases, the cell disruption process is accompanied by the release of heat; therefore, all Procedures involving cell disintegration and the isolation of cellular organelles should be performed at reduced temperatures (around +4°C) in thermostated cold rooms to prevent thermal denaturation. Maintaining a constant pH within a specific range is equally crucial, and to this end, suspension media are prepared using Buffer solutions. To eliminate the effects of heavy Metal Ions that may be introduced via water and reagents during homogenization, chelating agents are employed, such as EDTA (ethylenediaminetetraacetic acid) and its sodium salt, Trilon B, among others.

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To prevent The oxidation of sulfhydryl groups within proteins, reducing agents such as Cysteine, ß-mercaptoethanol, dithiothreitol, and others are added to the extraction medium.

Proteins can lose their biological activity As a result of surface denaturation, which occurs during vigorous foaming of the reaction mixture. Foam formation is prevented by adding neutral Surfactants.

It is well established that many proteins are localized within cellular organelles. To isolate them, the homogenate obtained after cell disruption is subjected to centrifugation. Depending on the centrifugal acceleration applied, specific organelles pellet out and can subsequently be used for protein extraction. Centrifugation also serves to separate solid debris and fragments of cellular structures. The supernatant (extract) represents an aqueous solution of diverse organic and Inorganic Compounds—including cell components and solubilization medium ingredients.

In the cell, many proteins exist in an associated state with other substances or cellular structures; mild detergent solutions (surface-active agents) or nonpolar solvents (such as ether or butanol) are applied to achieve their optimal solubilization.

Nonionic detergents, such as sodium deoxycholate and Triton X-100, which are widely used for the solubilization of Membrane Proteins, weaken hydrophobic protein-lipid and Protein-Protein Interactions, thereby releasing proteins from membranes. Anionic and cationic detergents also destabilize ionic bonds between the charged groups of membrane components. An example of an anionic detergent utilized for membrane Protein Isolation is sodium dodecyl sulfate (SDS).

Following material homogenization, protein extraction is performed. Depending on The properties of the target protein and the research objectives, water, saline solutions, various buffer mixtures, organic reagents, aqueous-alcoholic solutions, or weak acids and alkalis are employed as solvents. Protein extraction is very frequently carried out concurrently with material homogenization.

The extraction yields a complex mixture of proteins and other substances. Various purification methods are applied to isolate individual proteins. The Selection and sequence of these methods are determined empirically in each specific case, depending primarily on the PHYSICOCHEMICAL PROPERTIES OF the protein being isolated. Fractionation can be achieved by exploiting differences in Protein solubility at varying concentrations of neutral salts (salting out), organic solvents (ethanol, diethyl ether, acetone, dioxane, and others), different pH levels, and temperatures (see Section 2.3.4).

Salted-out proteins contain high amounts of salts; to remove these, the protein pellet obtained via centrifugation is subjected to dialysis against a buffer solution or to Gel filtration on molecular sieve columns. Desalting using molecular sieves proceeds many times faster.

An effective approach for Protein Purification is fractionation via chromatographic techniques. Chromatography is defined as a physicochemical method of substance Separation based on the distribution of mixture components between two phases: one stationary and the other a Mobile phase filtering through the stationary layer. The mobile phase is most commonly a liquid (liquid chromatography) or a gas (gas chromatography). Depending on the Properties of the stationary phase, several types of chromatography are distinguished. The characteristics of these chromatographic methods are outlined below.

The last Three types of chromatography are widely applied in Protein Chemistry. In ion-exchange chromatography (IEC), protein separation occurs due to their interaction with oppositely charged particles of ion-exchange resins (ionites)—solid, insoluble substances used to pack columns after appropriate preparation.

Ionites contain active (ionogenic) groups with mobile ions capable of exchanging with the ions of the analyzed mixture under specific conditions. Ionites containing active acidic groups and capable of exchanging mobile cations are termed cation exchangers (cationites). Conversely, ionites possessing basic active groups and mobile anions are called anion exchangers (anionites).

where R'—⁺NH₃ and R'—СОО^- represent the protein, — SO^-₃ —СН₂—N⁺≡ (СН₃)₃ denote the functional (sulfo- and trimethylaminomethyl) groups, H⁺ and OH^- are the mobile ions, and R stands for the base (matrix).

The application of size-exclusion chromatography (gel filtration) enables the Separation of proteins according to their molecular sizes. For this purpose, various porous insoluble Materials are used, such as synthetic resins, porous glass (Bio-Glas), porous silica (Porasil), polysaccharide derivatives based on dextran (Sephadex, Molselect) or agarose (Sepharose), and synthetic polyacrylamide polymers (Bio-Gel P), among others.

Porous materials are produced in the form of granules of a specific size; when suspended in a solvent, two phases are formed: one inside the granules and the other outside. The Column is packed with the suspended granules, a protein mixture is applied, and the solvent is passed through. Depending on the size and shape of the protein molecules, they are distributed between the two phases in different ways: larger and asymmetric molecules penetrate less frequently or do not penetrate at all (depending on the pore size of the support) into the internal solvent phase. Molecules for which the internal solvent is inaccessible remain in the external phase and move down the column at a much higher speed than smaller molecules. Proteins with relatively small molecular sizes (less than the pore diameter) or low-molecular-weight compounds—such as salts, sugars, Amino Acids, etc.—inevitably enter the pores of the granules and elute from the column more slowly, as they are in a state of diffusion equilibrium with the internal solvent (Fig. 2.3). Porous materials act as a molecular sieve, or more precisely, as an antisieve, since large molecules pass through them faster.

Sephadex, a derivative of the synthetic polysaccharide dextran, is most frequently used as a molecular sieve. As a result of chemical treatment, cross-links are formed in dextran, rendering it insoluble in water while allowing it to swell readily to form a gel. There are several grades of Sephadex, which differ in the pore diameter within the granules (Table 2.2).

Fig. 2.3. Diagram of protein solution desalting using Sephadex:

I, II, III — filtration stages; 1 — Sephadex granules, 2 — protein molecules, 3 — inorganic salts

Table 2.2. Types of Sephadexe

Affinity chromatography is based on the biospecific interaction between the target biopolymer (or group of Biopolymers) and a specific substance. This type of chromatography is applicable to Enzymes, IMMUNOGLOBULINS, Lectins, and receptor proteins that are capable of selectively binding to substrates, inhibitors, Cofactors, Antigens, and receptors. THE PRINCIPLE OF this chromatographic method is that a substance capable of specifically binding to the target protein is immobilized on an insoluble support. This substance is termed a Ligand. The support treated in this manner (the adsorbent) is placed in a column, and a protein mixture is passed through it. Only the protein that exhibits an affinity for the specific ligand binds to the adsorbent (Fig. 2.4). Subsequently, the protein is eluted from the column using a solution that induces dissociation of the formed protein-ligand complex.

Electrophoretic methods are widely used for protein separation; they are based on the migration of charged protein molecules in an electric field at a velocity that depends on the magnitude of their charges at a given pH and Ionic strength of the solution. The migration rate is also influenced by the shape and mass of the protein molecules, though to a lesser extent than by their net charge.

Electrophoretic separation can be performed in solution or on solid wet Supports, such as starch gel, polyacrylamide gel, silica gel, or chromatographic paper. Polyacrylamide gel Electrophoresis (PAGE) is characterized by high resolution and sensitivity; it features a standard pore size determined by the concentration of the initial monomers used for gel polymerization. Therefore, in this method, substance separation occurs based on both charge magnitude and molecular size.

Electrophoretic methods also include isotachophoresis and isoelectric focusing (IEF). In isotachophoresis (from the Greek isos meaning equal, and tachos meaning speed), charged ions are first separated according to their charge and mobility, and then migrate in an electric field at equal and constant velocities for a given experiment.

The isoelectric focusing method allows proteins to be separated simultaneously in both a voltage gradient and a pH gradient. The pH gradient is maintained by adding a mixture of carrier ampholytes (brand names Ampholine and Servalyt)—synthetic substances with amphoteric properties that differ from one another in their pI values—to the column. Specifically, ampholytes are polyaminopolycarboxylic acids with molecular weights ranging from 300 to 1000, synthesized from polyethylenepolyamines. Under The Influence of an electric field, the carrier ampholytes distribute themselves along the height of the column in a strictly defined sequence, establishing a linear pH gradient. The gradient range depends on the COMPOSITION OF THE ampholine or servalyt mixture and their pI values. A steep pH gradient is created by a mixture of ampholytes with a wide pI interval, whereas a shallow gradient is produced by a mixture with a narrower spectrum of ampholyte isoelectric points (Fig. 2.5). In the latter case, a higher resolution of the method is achieved, enabling the separation of proteins that differ in pI by 0.01 pH units or less.

Fig. 2.4. Diagram of the affinity chromatography method:

1 — support (matrix), 2 — "spacer arm", 3 — ligand, 4 — protein

The test proteins loaded onto the column migrate through it under the Influence of the applied electric field

(Fig. 2.6). Protein molecules in the region where the pH is higher than their pI are negatively charged and migrate downward (toward the anode). Conversely, protein molecules in a region with a pH lower than their pI migrate upward. As a result, each type of protein becomes concentrated in a zone where the pH equals its pI. Here, the protein focuses, as the electric field ceases to exert a force on molecules that have lost their net charge.

The application of the aforementioned extraction and purification methods makes it possible to isolate individual proteins with a high degree of purity. In certain cases, well-purified Proteins can be obtained in crystalline form. Proteins are crystallized from salt solutions or organic solvents. Protein recrystallization further increases their purity.

Isolated protein preparations must be tested for homogeneity. Direct Methods for Assessing protein purity do not exist; sample homogeneity is evaluated based on the absence of impurities. A substance is considered pure if none of the methods used to determine homogeneity reveal the presence of impurities. For this purpose, it is mandatory to use several Methods based on different principles. Sample homogeneity with respect to molecular weight is established by ultracentrifugation, and with respect to charge by PAGE and IEF. A pure substance yields only a single symmetrical peak upon ultracentrifugation. Peak Asymmetry, as well as the presence of a shoulder on the peak or multiple peaks, indicates heterogeneity of the studied protein solution. Electrophoretic methods for assessing homogeneity should also yield only a single protein zone following separation. In addition, the criterion of functional homogeneity—the determination of specific activity of proteins—is utilized. There is also a biological method for establishing the purity of protein preparations, which is based on the fact that most proteins possess antigenic properties, i.e., The ability to induce the appearance of specific Antibodies in an animal after administration. Injection of a homogeneous substance causes The formation of a single type of antibody, whereas a heterogeneous substance causes the formation of several types.

Fig. 2.5. pH limits of carrier ampholyte mixtures

Fig. 2.6. Migration process of protein molecules during isoelectric focusing. A — beginning of the process; B — end

The isolated and purified protein must be stored at a low Temperature. If the protein is crystallized from an ammonium sulfate solution, it can be stored in this solution as a suspension. Most commonly, proteins are stored in a dried state; drying is performed under high vacuum from the frozen state (lyophilization).

2.3.2. Molecular Weight of proteins and methods for its determination.

The molecular weight of proteins can be determined by various physical methods, among which the gravitational, gel-filtration, and electrophoretic methods are the most commonly used. Modern techniques allow determining molecular weights with an error not exceeding 5%, which amounts to several thousand and occasionally hundreds of daltons. Elucidating the Introduction/19.html">Primary Structure of many individual proteins has made it possible to calculate their molecular weight with high accuracy.

In the gravitational method, analytical ultracentrifuges are used to determine the molecular weight, allowing researchers to observe The process of particle sedimentation (settling). Modern ultracentrifuges develop centrifugal accelerations exceeding the acceleration of gravity by up to 500,000 times. Protein molecular weight determination by ultracentrifugation is carried out based on the sedimentation velocity of protein molecules and Sedimentation Equilibrium.

When determining molecular weight by sedimentation velocity, the protein solutions under study are centrifuged at high speeds. This causes protein particles uniformly distributed throughout the volume at the beginning of centrifugation to gradually move toward the bottom of the cell. A boundary layer forms between the Regions of the solvent free of protein molecules and those containing them. By recording the movement of this boundary during centrifugation, the sedimentation velocity of the molecules is determined, on The basis of which the molecular weight of the protein is calculated using the Svedberg equation.

In the Sedimentation equilibrium method, centrifugation is performed at relatively low rotational speeds of ~7–8 thousand rpm, which corresponds to a centrifugal acceleration of 40–50 thousand g. At such acceleration, even particles with a large molecular weight do not settle to the bottom. Centrifugation is continued until an equilibrium distribution of the studied protein along the entire length of the cell is achieved, where the movement of the protein toward the bottom of the cell under centrifugal force is balanced by the upward movement caused by diffusion. Based on the resulting protein concentration gradient in the centrifuge cell, the molecular weight of the protein is calculated using a special formula.

The Determination of protein molecular weight by gel filtration is carried out by packing a column with a porous gel (such as Sephadex, Sepharose, etc.) or applying it in a thin layer onto a solid support (Thin-Layer Chromatography). In the column method, the elution volume (V_e) of the target protein and proteins with known molecular weights is recorded; then, a calibration plot of $\lg M$ versus $V_e/V_o$ is constructed to determine the molecular weight of the protein under study. $V_o$ (void volume) is the volume of the solvent between the beads, defined as the volume required to elute a compound that does not penetrate the gel pores. The molecular weight values of most proteins fit onto the calibration line, with the exception of proteins having a highly asymmetric molecular shape. This method is also used to determine the molecular weights of subunits in proteins with a quaternary structure (see Section 2.4.7). In this case, 6 M guanidine hydrochloride is used, which causes protein dissociation into subunits and the loss of native conformation, thereby avoiding the influence of molecular shape.

In thin-layer chromatography on porous gels, the distance traveled by the protein in a given time is measured, and the molecular weight is calculated using a calibration curve plotted in coordinates of $\lg M$ versus $x$ (the distance traveled by the protein) using standard proteins.

In the electrophoretic METHOD FOR DETERMINING protein molecular weight in PAGE, the mobility of the target protein is also compared with the mobility of marker proteins. The graphical dependence of protein mobility on the logarithm of its molecular weight is expressed as a straight line. To eliminate the influence of molecular shape and charge on the electrophoretic separation of proteins, as well as to determine the molecular weights of individual subunits in a protein with a quaternary structure, proteins are pre-denatured using sodium dodecyl sulfate (SDS). Approximately 1.4 g of SDS binds to 1 g of protein, with the hydrophobic groups of SDS located inside the resulting complex and the sulfonic acid groups On the surface. The latter provide a net negative charge to the complex, which depends solely on the protein mass. During electrophoresis in gels with uniform porosity, these complexes migrate at a rate inversely proportional to the molecular weight of the protein. SDS-polyacrylamide gel electrophoresis is widely used to study complex mixtures of membrane proteins and to determine their molecular weights, as many membrane proteins are poorly soluble in the absence of SDS.

Each of the aforementioned Methods for determining molecular weight has its drawbacks; therefore, accurate results can be obtained by combining different methods.

The application of these methods has established that the molecular weights of various proteins vary over a wide range: from 6 thousand to several million daltons. The molecular weights of some proteins are listed below.

2.3.3. Amphoteric Properties of Proteins. Proteins act as amphoteric electrolytes because their molecules contain both acidic and basic groups. The acid-base properties of proteins are primarily determined by the ionizable side chains of amino acids. THE CONTRIBUTION OF terminal NH₂ and COOH groups is extremely negligible. The $\text{p}K$ values of amino acid side chains within proteins differ slightly from those of free amino acids, as the degree of group ionization in proteins depends on The Nature of neighboring side chains, i.e., on the electrostatic environment.

The presence of dissociating groups in proteins imparts a specific net charge to the molecule, which depends on the ambient pH. Most natural proteins are classified as acidic proteins due to a significant content of dicarboxylic acids and therefore, at pH values close to neutral, carry a net negative charge, similar to the Cytoplasm.

Amino acids with dissociating side chains are components of all proteins, where they are predominantly located on the globule surface, thereby determining the overall hydrophilicity of the molecule as well as several other physicochemical properties of protein molecules. The charge distribution on the molecular surface is non-uniform, and its various regions may carry opposite charges stabilized by water dipoles. A shift in environmental pH leads to Changes in the dissociation pattern of radicals and the redistribution of charges on the molecular surface, resulting in alterations to the Spatial Structure of Proteins and the degree of their biological activity.

The presence of numerous dissociation sites also determines the ability of protein molecules to interact with small ions (particularly metal ions) and other charged macromolecules, which is crucial for protein functioning. The ability of proteins to interact with both cations and apathogens (anions) is of great biological importance. For instance, the ability of proteins to bind calcium cations plays a vital role in many physiological processes and is directly related to the Regulation of Metabolic processes.

For each protein, There is a specific value of active medium reaction at which the positive and negative charges within the molecule are balanced. The pH value at which a protein carries no net charge and does not move in an electric field is called the isoelectric point (IEP) and is denoted as pI. The IEP value can be determined from curves obtained during acid-base titration of a protein solution, as well as by isoelectric focusing of the protein (see Section 2.3.1). The isoelectric points of some proteins are given below.

The pH value corresponding to a protein's IEP is determined by the number of its ionogenic groups and their dissociation constants ($\text{p}K$). The IEP is above 7 if the protein contains A large number of basic amino acid residues, and below 7 if acidic amino acids predominate. For most Globular proteins, the IEP lies in the acidic range (4.5–6.5). However, there are exceptions. For example, the enzyme pepsin, which Functions in the strongly acidic environment of The Stomach, has an IEP of about 1, whereas protamines have an IEP of about 12.

The net charge of a protein molecule is positive at $\text{pH} < \text{pI}$ and negative if $\text{pH} > \text{pI}$. Because a protein molecule carries no net charge at its IEP and electrostatic repulsion between neighboring molecules is absent, proteins are easily precipitated from solutions at a pH equal to their IEP.

The isoelectric point of a protein should be distinguished from the isoinic point, as these values do not always coincide. The isoinic point of a protein is defined as the pH value at which the number of protons bound to basic groups equals the number of protons released by dissociated acidic groups in the protein molecule. Thus, the isoinic point corresponds to the pH at which the net charge of the protein molecule is zero in the complete absence of electrolytes in the solution. Consequently, the isoelectric and isoinic points coincide only when the protein solution contains no ions other than the ionized residues of the protein molecule's Amino Acids and the ions resulting from water dissociation. Direct measurement of the isoinic point is difficult; therefore, it is determined by measuring the IEP at various salt concentrations and extrapolating to zero concentration. In the absence of salts and when the isoinic point is near pH 7, it will nearly coincide with the isoelectric point. However, in the presence of salts, the difference between these points can be quite significant.

Being amphoteric electrolytes, proteins exhibit buffering properties.

2.3.4. Solubility of proteins. The vast majority of proteins are hydrophilic substances that dissolve well In aqueous solutions. Their solubility, like that of other macromolecular substances, is determined by the Nature of the groups exposed on The surface of the molecule during its spatial folding into the native conformation. Most of the protein molecular surface is formed by groups capable of Hydration.

Hydration refers to the binding of water dipoles to ionic and polar groups. The charged ionic groups in proteins are dissociable radicals. In the dissociated state, they attract water molecules via ion-dipole interactions. Non-ionic polar amino acid side chains (asparagine, glutamine, Serine, Threonine) form Hydrogen Bonds with water. Oxygen and hydrogen atoms belonging to peptide groups are also capable of forming hydrogen bonds with water because these atoms carry excess negative ($ ext{O}$) and positive ($ ext{H}$) charges. However, water access to peptide groups is hindered by steric hindrance created by amino acid side chains, meaning they do not make a significant contribution to protein hydrophilicity.

Protein solubility in water increases upon The addition of low concentrations of neutral salts [$( ext{NH}_4)_2 ext{SO}_4$, $ ext{Na}_2 ext{SO}_4$, $ ext{MgSO}_4$, etc.]; this effect is known as salting-in. The dissolution of proteins, like other substances, is promoted by factors that decrease interactions between solute molecules. Neutral salts at low concentrations increase the degree of dissociation of ionized protein groups, shield the charged groups of protein molecules, and thereby reduce protein-protein interactions. It is known that the degree of dissociation of electrolytes (including proteins) is directly proportional to the Dielectric Constant of the solvent, which, in turn, is proportional to the degree of polarization of solvent molecules and their dipole moment. For instance, strongly polarized water molecules have a dielectric constant of 80 at room temperature, whereas acetone and ethanol have a dielectric constant of 20–30. Neutral salts at low concentrations further increase the dielectric constant of water. As a result, water enhances the dissociation of the solute, particularly the protein. By interposing between charged groups and orienting around them, water dipoles prevent their interaction.

High concentrations of neutral salts, conversely, precipitate (salt out) proteins from aqueous solutions; this process occurs most actively at the protein's IEP. The relative salting-out efficiency of various ions depends on their size, charge magnitude, and hydration capacity. At high ion concentrations in solution, they draw polarized water molecules away from the charged groups of the protein, thereby partially stripping the protein of its hydration shell, which normally prevents its precipitation from solution. At high salt concentrations, this factor plays a decisive role. According to their ability to salt proteins out of aqueous solutions, anions and cations can be arranged into specific series—the Hofmeister series, or lyotropic series. For anions in an alkaline environment compared to the protein's IEP, this series is as follows: $ ext{SO}_4^{2-} > ext{F}^- > [ ext{citrate}]^{2-} > [ ext{tartrate}]^{2-} > [ ext{acetate}]^- > ext{Cl}^- > ext{NO}_3^- > ext{Br}^- > ext{I}^- > ext{CNS}^-$. The lyotropic series for cations under the same conditions is as follows: $ ext{Ba}^{2+} > ext{Sr}^{2+} > ext{Ca}^{2+} > ext{Mg}^{2+} > ext{Cs}^+ > ext{Rb}^+ > ext{K}^+ > ext{Na}^+ > ext{Li}^+$.

The solubility of proteins also depends on the pH of the solvent, its composition, and temperature. As noted above, proteins exhibit minimum solubility at their pI, which is explained by the absence of electrostatic repulsion between protein molecules.

The addition of water-miscible organic solvents (such as ethanol or acetone) to a protein solution decreases protein solubility, and at high solvent concentrations, protein precipitation is observed. This phenomenon is attributed to a decrease in the dielectric constant of the medium and, moreover, a reduction in the degree of protein hydration, which leads to enhanced attraction between oppositely charged groups within the protein molecule and subsequent protein aggregation. Proteins with varying solubility properties precipitate at different intervals of salt or organic solvent concentrations, and at distinct values of the active reaction of the medium. This property of proteins is exploited in their isolation and purification via fractionation (see Section 2.3.1).

In solution, proteins exhibit colloidal properties: they diffuse slowly, do not pass through semipermeable membranes, scatter light, and are characterized by high viscosity. However, it should be noted that protein solutions are not typical colloidal solutions, as proteins are dispersed down to single molecules and form a homogeneous solution. In contrast, typical colloidal solutions are heterogeneous and biphasic (consisting of a solute and a solvent), and the colloidal particles (micelles) of the solute are composed of multiple molecules. The similarity between protein solutions and true colloidal solutions stems from the fact that protein molecules have dimensions approaching the size of colloidal micelles (10^-4—10^-7 cm).

The formation of colloidal solutions by proteins and other biological macromolecules determines many physicochemical phenomena observed in biological fluids and organisms as a whole. Protein solutions, like all colloidal solutions, can lose their fluidity under certain conditions and form gels, or jellies. Gels arise as a result of the association of molecules into a network whose internal space is filled with a large amount of solvent, while phase separation into liquid and solid phases—such as occurs during coagulation—does not take place. It is believed that in A number of PLANT AND ANIMAL tissues, proteins exist not only as solutions but also as gels (in cell protoplasm, the lens of the eye, Connective Tissue, etc.). Protein solutions, such as milk, can transition into a gel state under the influence of microbial enzymes, resulting in the formation of curdled milk or kefir. During the preparation of plants for the winter season, in the process of so-called autumn "hardening," a portion of proteins transitions from a dissolved state to a gel-like state.

Gels age over time, synerese water, and divide into two phases: a compacted gel and a diluted sol. This process is known as syneresis and occurs, for example, when kefir is left standing. One of the properties of gels is their ability to swell—to increase in volume due to the binding of a large amount of water. Such a process occurs, for example, during seed germination. The connective tissue proteins of animals absorb water, thereby enabling its storage by the Organism. Swelling plays a major role in leather manufacturing technology and baking.

In solutions of proteins and other high-molecular-weight compounds, The phenomenon of coacervation may be observed—the fusion of the aqueous envelopes of several particles without the aggregation of the particles themselves. Coacervates arise when the solubility of solution components is limited, which causes the formation of coacervate droplets. Coacervates are formed, for example, from proteins with opposite charges. They are ascribed great importance in The Theory of THE ORIGIN OF life; however, unlike biological nonequilibrium structures, coacervates are thermodynamically equilibrium systems.

Owing to their hydrophilic and hydrophobic groups, proteins can influence the solubility of other substances, acting as emulsifiers—substances that stabilize emulsions formed by mutually insoluble liquids (water and oil). In The Human Body, fats in the Blood AND Lymph are in an emulsified state. The protein forms a thin film on the surface of the fat droplets, which attracts water and prevents the fat particles from clumping together. One of the causes of urinary and gallstones can be a deficiency in the body of mucins—acidic Glycoproteins that coat hydrophobic microparticles and thereby facilitate their elimination from the organism. Milk can also be viewed as an emulsion, representing fat droplets emulsified by caseinogen in water.

2.3.5. Protein Denaturation. Protein denaturation is defined as the disruption of the native spatial structure of a protein molecule, leading to a decrease or complete loss of its solubility, alteration of other physicochemical properties of the protein, and loss of its specific biological activity. Denaturation is accompanied by the Cleavage of covalent bonds in the polypeptide chain backbone. A breakdown of disulfide bridges, as well as hydrophobic, ionic, and hydrogen bonds, occurs. As a result, the native tertiary structure and, to a significant extent, the Secondary structure are disrupted. Protein denaturation is caused by both certain chemical compounds and physical factors. The Mechanism of the denaturing action of chemical agents is determined by their structure. For example, urea, guanidine hydrochloride, and formamide, owing to the presence of an amide group, compete with the peptide groups of the protein for hydrogen bonds, diverting them to themselves. These reagents also disrupt hydrophobic interactions. Urea and other amides exhibit their maximum denaturing effect at high concentrations (6M—10M).

The denaturing action of moderately polar organic solvents (lower alcohols, Ethylene glycol, dioxane, dimethyl sulfoxide, etc.) is associated with a sharp decrease in the dielectric constant of aqueous protein solutions, which increases the electrostatic interaction forces between charged groups. The formation of stable intra- and intermolecular ionic bonds leads to protein denaturation. Organic solvents disrupt hydrophobic interactions within protein molecules. Nonpolar aliphatic and aromatic Hydrocarbons destroy the intramolecular Hydrophobic bonds of proteins through direct interaction with their hydrophobic groups.

Ionic detergents, such as sodium dodecyl sulfate, bind to oppositely charged groups of the protein. The electrostatic repulsion of like-charged groups remaining in the polypeptide chain leads to the cleavage of Hydrogen bonds and other weak interactions that stabilize the native conformation of the protein.

Denaturing Agents also include heavy metal cations and iodide and thiocyanate anions. These substances apparently form fairly stable compounds with the polar groups of proteins, distorting The system of ionic and hydrogen bonds.

Trichloroacetic acid and tannin are characterized by a complex denaturation mechanism that includes both a direct effect on hydrogen bonds and the blocking of polar groups.

Many proteins undergo denaturation upon strong acidification (pH < 2—3) or alkalinization (pH > 10—11). Under these conditions, virtually all dissociating groups of the protein carry a like charge (predominantly H₃N⁺ groups at low pH values and COO^- groups at high pH values). The mutual repulsion of like charges causes the cleavage of some weak bonds, thereby disrupting the native structure. However, it must be taken into account that certain proteins are quite stable at extreme pH values. For example, Histones, protamines, and lysozyme are relatively stable at pH 2, whereas pepsin exhibits maximum activity at pH 1.5—2.2. Lysozyme, histones, and protamines are also stable at pH 10, while the pH optimum for the catalytic activity of alkaline Phosphatases and arginase lies in the range of 9.5—9.7.

Among the Physical factors of denaturation, heating is the most universal. The intensification of the thermal motion of polypeptide chains leads to the cleavage of hydrogen bonds and the disruption of hydrophobic interactions. The rate of thermal denaturation depends significantly on the active reaction of the medium and the presence and concentration of salts. Thermal denaturation is accompanied by protein aggregation and precipitation, which represent a secondary phenomenon.

There is extensive data indicating that the majority of proteins in thermophilic microorganisms exhibit enhanced thermostability. They do not denature upon heating up to 60°C, undergoing only minor conformational changes, whereas in mesophiles (organisms inhabiting normal, moderate temperatures), significant denaturation is observed at +60°C. Only rare proteins in mesophiles possess enhanced thermostability.

Proteins also denature under many mechanical stresses: high pressure (5000—10 000 atm), grinding of dry preparations, vigorous shaking of solutions, and irradiation with high-frequency sound waves. All of this must be kept in mind during biochemical studies of proteins. During lyophilization, most proteins do not denature, which allows this drying method to be used to obtain long-term storable protein preparations.

Other physical factors of protein denaturation include ultraviolet light (especially within the range of 260—310 nm) and ionizing radiation. Protein denaturation also occurs when it is distributed at a two-phase interface; such surface denaturation is observed during the foaming of protein solutions during protein isolation and upon foam formation on the surface of water bodies.

During protein denaturation, many physicochemical properties of the protein change: solubility, sedimentation constant, viscosity, optical properties, etc. In the process of denaturation of proteins with quaternary structure, their dissociation into subunits may occur. The protein exhibits a substantial decrease in the number of regions with regular secondary structure types, a reduction in the number of intramolecular hydrogen bonds, and an increase in the number of these bonds between the protein and water. Because denaturation involves the disruption of hydrophobic interactions during the unfolding of the protein molecule, leaving hydrophobic groups exposed on the molecular surface, this causes a loss of Protein solubility and swelling capacity in aqueous solutions.

Protein denaturation exposes reactive groups that are not fully accessible to detection by conventional methods in the native protein (sulfhydryl, phenolic, imidazolyl, etc.). The change in the number of reactive groups upon denaturation is also manifested in a shift of the protein's pI. As a rule, the isoelectric point shifts toward alkaline pH values. Protein denaturation is accompanied by an increase in negative optical activity.

The Transformation of a compact molecule into a random coil, which occurs during denaturation, makes most peptide bonds accessible to the action of Proteolytic Enzymes. Consequently, the proteolysis of denatured proteins proceeds at a faster rate than that of native ones.

The loss of biological activity by proteins is expressed as the inactivation of enzymes, Hormones, and Viruses. This serves as an important criterion of denaturation, although some exceptions exist. Enzymes such as Papain and pepsin retain their activity upon denaturation by urea, while ribonuclease and lysozyme retain their activity upon heating in dilute acid. Denaturation of a number of proteins results in a decrease in antigenicity while retaining immunological Specificity.

Slow protein denaturation occurs during prolonged seed storage, resulting in a decrease in their swelling capacity and, consequently, a reduction in germination intensity.

Complete protein denaturation is irreversible in most cases; however, under appropriate conditions, proteins can sometimes be renatured. The ability to undergo renaturation after heating has been demonstrated, for example, for the enzyme Trypsin. A prerequisite for its renaturation is very slow cooling of the protein to room temperature ("annealing"), which restores the native conformation and specific biological function of trypsin.

2.3.8 Optical Properties. All proteins generally absorb ultraviolet (UV) light in three regions. Absorption at wavelengths (λ) greater than 250 nm with a maximum around 280 nm is due exclusively to the presence of aromatic amino acids—Tryptophan, Tyrosine, and phenylalanine. The major contribution is made by tryptophan (λ¹_max = 278 nm, ε² = 5600) and tyrosine (λ_max = 275 nm, ε = 1300), which is present in large amounts in the composition of almost all proteins. Absorption at 210—250 nm is of a complex nature and is determined by the presence of aromatic and several Other Amino Acids, as well as the presence of hydrogen bonds, α-helical regions, etc., in proteins. The absorption band with a maximum located near 190 nm is due mainly to the presence of peptide bonds in the protein. The absorption intensity in this region varies primarily depending on The amount of α-Helical structures in the protein.

The spectrophotometric method for Quantitative Protein Determination (based on the absorption intensity at 280 nm) is founded on the property of proteins to absorb light in the UV region of the spectrum. It is not very precise because the amounts of tryptophan and tyrosine vary quite widely among different proteins. When working with proteins, it is conventionally assumed that 1 optical density unit of a solution at 280 nm corresponds to a concentration of approximately 1 mg/ml (at a cuvette path length of 1 cm). Despite its insufficient precision, this method is widely used in modern biochemistry because it is simple and rapid to perform, and allows for the further use of the protein solution after optical density determination. The method is used particularly often for monitoring changes in protein concentration during column separation.

Spectrophotometry also makes it possible to determine how the optical density of solutions depends on the wavelength of absorbed light—in other words, to obtain absorption spectra—and is used to ascertain the spatial arrangement of aromatic amino acids within the protein globule (whether interior or exterior). This latter application relies on the fact that the absorption spectra of aromatic amino acids (their λ_mах and ε) undergo noticeable changes depending on their environment, specifically the solvent polarity and the pH of the medium. Tryptophan, tyrosine, and phenylalanine located in a less polar environment (for example, inside the protein globule) exhibit higher λ_mах and ε values. If a protein's spectrum is sensitive to changes in solvent polarity, it can be concluded that the amino acids showing shifts in λ_mах and ε are predominantly located on the surface of the protein globule.

¹ λ_mах is the wavelength at which maximum absorption is observed.

² ε is the molar absorption coefficient.

Analyses of this type are usually performed by recording differential spectra—that is, by directly measuring the difference in UV Light absorption by a protein in various solvents. Differential spectrophotometry makes it possible to determine the degree of polarity of the microenvironment surrounding an aromatic amino acid situated inside the globule. For instance, if THE SPECTRUM OF an amino acid incorporated into a protein in a polar solvent shows higher λ_mах and ε values than those of the free amino acid in the same solvent, this amino acid resides in the interior of the protein and is surrounded by nonpolar amino acids.

In the visible spectrum (380–760 nm), light absorption is a property exclusive to colored proteins, known as Chromoproteins—such as Hemoglobins, Cytochromes, flavoproteins, the "blue protein" from Pseudomonas, which intensely absorbs light at around 600 nm, and several others.

In the infrared (IR) region of the spectrum (760–10,000 nm), all proteins absorb light. IR spectroscopy is widely used to determine the relative content of α-helices, β-structures, and amorphous regions within a protein molecule. This can be estimated from the intensity of certain spectral bands, notably the amide I band (~1680 cm^-1). When representing IR spectra, frequencies (v) or wave numbers (1/λ) are used instead of wavelengths, which is why band positions are specified in reciprocal centimeters.

The Use of polarized IR light makes it possible to determine the orientation of hydrogen bonds relative to the polypeptide chain: they are parallel in the α-Helix and perpendicular to the chain in the β-structure. Infrared dichroism registers the absorption spectra of a protein for two mutually perpendicular directions of polarization of the incident light. In one case, the electric field vector is parallel to the peptide chains, and in the other, it is perpendicular to them. This method employs so-called stretched protein films, in which all protein molecules are oriented in the same direction, to measure the dichroic ratio—that is, The ratio of band areas obtained when the electric vector of the light wave is parallel and perpendicular to the molecular axis. The —C—O (1660 cm^-1) and —N—H (3300 cm^-1) groups absorb most strongly when they are parallel to the vector. The magnitude of the dichroic ratio serves to evaluate the probability of an α-helical structure within the protein.

Proteins are optically active substances: they rotate plane-polarized light passing through them and absorb left- and right-circularly polarized light to different extents. Plane-polarized light is light in which the electric field vector for all photons in the light beam lies in a single plane, whereas circularly polarized light is light in which this vector traces out a helix. This property of proteins is attributed to the presence of chiral carbon atoms in their molecules. The interaction of polarized light with proteins is studied using optical rotatory dispersion (ORD) and circular dichroism (CD). These methods measure the capacity of an optically active substance to rotate plane-polarized light (ORD) and to differentially absorb right- and left-circularly polarized light (CD) as a function of wavelength. ORD and CD methods are applied to broadly characterize the helical content of proteins and to investigate conformational changes.

Proteins and many protein structures are characterized by optical anisotropy, meaning a variation in optical properties depending on the direction. In some cases, optical anisotropy stems from the internal architecture of proteins, while in others it results from their shape or artificially induced orientation.

Protein solutions also exhibit fluorescence—the emission of a light quantum upon transition from an electronically excited state to the ground state. This protein property is the basis of fluorescence spectroscopy, which measures the intensity of emitted light upon sample excitation (the absorption of a light quantum through UV irradiation). The fluorescence spectrum is always shifted toward longer wavelengths relative to the absorption spectrum because the molecule loses energy during the transition from the excited state to the ground state.

The fluorescent amino acid residues in proteins are tryptophan, tyrosine, and phenylalanine. The fluorescence of each can be distinguished by the wavelength at which it is observed. In addition, conjugated (complex) proteins may contain other fluorescent components. Fluorescence measurements provide valuable insights into protein conformational transitions, ligand-binding sites, solvent interactions, molecular flexibility, intermolecular distances, and more.