Chemistry

Protein purification

Protein purification


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The centrifugation

Centrifuges have a rotor made of metal or plastic, which contains several bores for centrifuge tubes or devices for hanging swing-out buckets. Most centrifuges can be temperature controlled during centrifugation, since the temperature can increase considerably, especially with long centrifugation times or at high speeds. At very high speeds the air resistance becomes very high, which is why centrifuges are used here, the rotor chamber of which can be evacuated (ultracentrifugation).

The rotor types

In most cases, so-called fixed-angle rotors are used, in which the sample tubes are at a fixed angle (approx. 30 °) to the rotor axis. The sediment collects on one side of the tube. Disadvantage of these types of rotors is that the liquid is pressed against the tube lid during centrifugation, especially when the tubes are very full, and may leak out. Fixed-angle rotors are used almost exclusively for the pelleting of materials (cells, membrane residues after cell lysis, precipitation of DNA, proteins, etc.).

With the swing-bucket rotor, the tubes are hooked into a device in which they align themselves with gravity during centrifugation, i.e. the gravity field is always at the bottom of the tube. No liquid can leak out during this centrifugation.

In the vertical rotor, the tubes are parallel to the rotor axis during centrifugation. After centrifugation, the liquid has to reorient itself from vertical to horizontal, and for this to happen exactly, a density gradient (e.g. a cesium chloride gradient when isolating plasmid DNA) is required in the tube. Vertical rotors are therefore used for density gradient centrifugation in an ultracentrifuge.


Non-mechanical digestion process

Non-mechanical disruption processes are used for cells that cannot be easily broken (for example yeasts). They are usually gentler than the mechanical processes.

  • In non-plant cells, variants of phenol-chloroform extraction such as Trizol digestion can be carried out by dissolving the membrane lipids out of the cell membrane, but in the presence of guanidinium thiocyanate it is denaturing. & # 911 & # 93
  • In yeast cells, autolysis is induced with toluene, which holes in the cell membrane and enzymatic lysis with zymolyase destroys the glucan cell wall, while Triton X-100 destroys the cell membrane.
  • In the case of Gram-positive bacteria, treatment with lysozyme destroys the peptidoglycan envelope, then the cell membrane can be loosened with Triton X-100.
  • In Gram-negative bacteria, the lipopolysaccharide of the outer cell membrane is dissolved by treatment with EDTA, then the peptidoglycan envelope is destroyed by treatment with lysozyme, then the cell membrane can be destroyed with Triton X-100.
  • In the case of Gram-negative bacteria, the membrane lipids can be saponified using alkaline lysis; due to the high pH value, this method is also denaturing.

Green fluorescent protein (GFP) has become an indispensable part of modern cell biology. In 2008, the Nobel Prize for Chemistry was awarded for its discovery and further development. The inestimable value of GFP lies in its ability to bind to any other protein in a gene-specific manner. Coupled with a virus, for example, it can explain the route of infection and the spread of a disease. In addition, GFP and its color variants serve as a tool for marking various cellular components and proteins or for visualizing transport processes in living organisms. Dr. Jürgen Braun and the students from the Johanna Wittum School in Pforzheim show with their experiment how GFP can be purified from a protein mixture and even observed.

Information on the experiment, the loanable experiment set from the Johanna Wittum School and the Biotechnology Working Group is available at www.modernebiotechnologie.de

Detailed descriptions of the experiments and a safety data sheet are available for download.


Protein purification. 2.1 Properties of proteins

1 Protein purification 2 Science has been studying the structure and function of proteins for more than two hundred years, when the French chemist Pierre J. Macquer used the term albumins to summarize all substances that exhibit the peculiar phenomenon of changing from liquid to solid when heated. These substances included chicken egg white, casein and the blood component globulin. As early as 1787, around the time of the French Revolution, the purification of coagulable, protein-like substances from plants was reported. In the early nineteenth century, many proteins such as albumin, fibrin or casein were purified and analyzed, and it soon became apparent that these compounds were structured in a considerably more complex manner than the other organic molecules known at the time. The word protein was probably coined by the Swedish chemist J & oumlns J. von Berzelius around 1838 and then published by the Dutch Gerardus J. Mulder together with a chemical formula that Mulder considered to be generally valid for proteinaceous substances at the time. Of course, the homogeneity and purity of these proteins, which were then purified, did not meet today's requirements, but they showed that individual proteins can be distinguished from one another. At that time, cleaning could only be successful because you could use simple steps: extraction for enrichment, acidification for precipitation and crystallization when a solution is simply left to stand. Hofmeister received the chicken album in crystalline form as early as 1889. Although Sumner was able to crystallize enzymatically active urease in 1926, the structure and structure of proteins remained in the dark until the middle of the twentieth century. It was only the development of powerful purification methods with which individual proteins can be isolated from complex mixtures, accompanied by a revolution in techniques for the analysis of the separated proteins, that we have made it possible to understand protein structures today. In this chapter these cleaning methods are described and it should be clear how they are used systematically and strategically. It is extremely difficult to look at the topic from a general perspective, as the physical and chemical properties of different proteins can differ immensely. However, this diversity is biologically necessary because proteins, the actual tools and building materials of a cell, have to perform a wide variety of functions. 2.1 Properties of proteins Size of proteins The size of proteins can vary greatly, from small polypeptides, such as insulin, which consists of 51 amino acids, to very large multifunctional proteins, for example apolipoprotein B, a cholesterol-transporting protein that consists of a Chain consists of over amino acids, with a molecular mass greater than daltons (500 kda). Many proteins consist of oligomers of the same or different protein chains and have molecular weights of up to several million Daltons. In general, it can be expected that the larger a protein, the more difficult it will be to isolate and purify. This is due to the analytical methods, which show very low efficiencies with large molecules. In Figure 2.1, the separation capacity of individual separation processes (the maximum number of analytes that can be separated from one another under optimal conditions) is plotted against the molecular weight. It can be seen that for small molecules such as amino acids or peptides, some chromium molar mass = molar mass (M) is incorrectly often referred to as molecular weight, not mass, but the quotient of the mass of a substance divided by the amount of substance of the substance. Unit: g / mol. Absolute molecular mass (m M) is the molar mass (M) of a molecule divided by the number of particles in one mole (Avogadro constant N A): m M = M / N A. Unit: g. Relative molecular mass (M r) is the molecular mass standardized to 1/12 of the mass of 12 C (dimensionless).

2 14 Part I: Protein Analysis 1D Electrophoresis 2.1 Separation Methods for Biomolecules. The separation capacity of individual separation methods (the maximum number of substances separated from one another in an analysis) is significantly different for different molecular weights of the substances. SEC exclusion chromatography HIC hydrophobic interaction chromatography IEC ion exchange chromatography RPC reversed phase chromatography CE capillary electrophoresis. Dalton (Da) is a non-SI-compliant unit of mass named after the English naturalist John Dalton (). A dalton is equal to the atomic mass unit (u = 1/12 of the mass of 12 C) and roughly corresponds to the mass of a hydrogen atom (1, g). The most common use in biochemistry is kda (kilodalton = Da). Proteome Analysis Chapter 41 Affinity Chromatography Section matographic methods are quite able to separate more than 50 analytes in a sample. In the area of ​​proteins it can be seen that of the chromatographic techniques only ion exchange chromatography is able to separate more complex mixtures reasonably efficiently and that in this molecular mass range the electrophoretic techniques are far more powerful. For this reason, proteome analysis (the analysis of all proteins in a cell), in which several thousand proteins have to be separated, is now practically exclusively based on electrophoretic processes (one and two-dimensional gel electrophoresis). The figure also shows that there are no efficient separation processes for large molecules, for example for protein complexes with molecular weights of more than 150 kda, or for organelles. However, the separation efficiency of a method is not always the relevant parameter that plays a role in cleaning. If selective cleaning steps are available, the importance of the separation capacity takes a back seat and the selectivity becomes the decisive factor. For example, an affinity purification based on the specific interaction of a certain substance with an affinity matrix, for example an immunoprecipitation or an antibody peraffinity chromatography, has a very poor separation capacity of 1, but an extremely high selectivity in a very complex mixture with which a protein can be extracted from a isolate in a single step. Since the most important purification techniques, electrophoresis and chromatography, require the analytes to be present in dissolved form, the solubility that the protein possesses in aqueous buffer media is another important parameter when planning a protein purification. Many intracellular proteins located in the cytosol (e.g. enzymes) are easily soluble, while proteins that have structure-forming functions, such as the proteins of the cytoskeleton or membrane proteins, are usually much more difficult to dissolve. The very hydrophobic, integral membrane proteins whose natural environment is lipid membranes and which aggregate and precipitate without solubilizers such as detergents are particularly difficult to handle in aqueous media. Available amount The amount available in the starting material plays a decisive role for the effort that has to be made for protein purification. A protein intended for purification may only be present in a few copies per cell (e.g. transcription factors) or in a few thousand copies (e.g. many receptors). Common protein -

3 2 Protein purification (e.g. enzymes) can make up percentages of the total protein in a cell. Over-expressed proteins are often present in significantly higher quantities (> 50%), as are some proteins in body fluids (e.g. albumin in plasma> 60%). Since the purification normally becomes much easier with increasing amounts of a protein, various sources of starting material should be examined for the content of the protein of interest, especially when isolating rare proteins. Acid / base properties Proteins have certain acidic or basic properties due to their amino acid composition, which is used for separation via ion exchange chromatography and electrophoresis. The net charge of a protein depends on the ph value of the surrounding solution and is positive at a low ph value, negative at a high ph value and zero at the isoelectric point at this ph value the positive and negative charges compensate each other. Biological activity The purification of a protein is often made more difficult by the fact that a certain protein can only be recognized and localized in the variety of other proteins on the basis of its biological activity. Therefore, in every phase of protein isolation, care must be taken to preserve this biological activity. It is usually based on a specific molecular and spatial structure. If it is destroyed, one speaks of denaturation, this is often irreversible. In practice, in order to avoid denaturation, the use of some cleaning processes must be ruled out from the outset. The biological activity is often differently stable under different environmental conditions. Too high or too low buffer concentrations, temperature extremes, contact with non-physiological surfaces such as glass or missing cofactors can change the biological characteristics of proteins. Some of these changes are reversible: even after denaturation and loss of activity, small proteins in particular are often able to renature under certain conditions, that is, to regain their biologically active form. In the case of larger proteins, this rarely works and often only with poor yield. The measurement of the biological, such as the enzymatic, activity gives the possibility of tracking the purification of a protein: with increasing purification steps, a higher specific activity is measured. In addition, the biological activity itself can be used to purify the protein. It is often associated with binding properties to other molecules, for example enzyme substrate or cofactor, receptor ligand, antibody antigen, etc. These very specific bindings are used to design affinity purifications (affinity chromatography) and are characterized by high levels under optimal conditions Enrichment factors and thus through a great efficiency that is hardly achievable in any other way. Enzymatic Activity Tests Chapter 4 Affinity Chromatography Section Stability If proteins are extracted from their biological environment, their stability is often noticeably impaired, as they are degraded by proteases (proteolytic enzymes) or almost always lead to an irreversible loss of biological aggregates, which leads to an irreversible loss of biological activity leads. For these reasons, protease inhibitors are often added in the first steps of protein isolation and cleaning is generally carried out quickly and at low temperatures. If you consider this variety of properties, it quickly becomes clear that protein purification cannot follow a schematic rule. For a successful isolation strategy, in addition to an understanding of the behavior of proteins in the various separation processes and a minimal knowledge of the solubility and charge properties of the protein to be purified, a clear idea of ​​the purpose for which the protein is to be purified is necessary. Purification Purpose The first steps in a purification process, the purity to be striven for and the analytics to be used are largely dependent on the intention with which a certain protein is to be purified. When isolating a protein for therapeutic purposes (e.g. insulin, growth hormones or anticoagulants), the requirements for purity are far higher than for a protein that is used in the laboratory for structural investigations. In many cases one only wants a protein for unambiguous identification or for the clarification of a few

4 16 Part I: Protein Analysis Proteome Analysis Chapter 41 isolate fewer amino acid sequence segments. A very small amount of protein is sufficient (usually in the microgram range). The sequence information can be used to identify the protein in protein databases or to produce oligonucleotide probes and isolate the protein's gene. This can then be expressed in a host organism in a much larger amount (up to a gram amount) than was present in the original source (heterologous expression). Many of the further investigations are then not carried out with the material from the natural source, but with the recombinant protein. Strategically new approaches to the analysis of biological issues, such as proteome analysis and subtractive approaches, require completely new types of sample preparation and protein isolation, since the quantitative relationships of the individual proteins must not be changed here.A great advantage of these new strategies is that it is no longer necessary to ensure that biological activity is maintained. Even if each protein purification is to be regarded as an individual case, some general rules and procedures can be found, especially for the first purification steps, which have already been used frequently in successful isolations and which should be discussed in detail below. 2.2 Protein localization and purification strategy The first step in any protein purification is to bring the desired protein into solution and to separate out all particulate and insoluble material. Figure 2.2 shows a scheme for different proteins. For the purification of a soluble extracellular protein, cells and other insoluble components have to be separated in order to obtain a homogeneous solution, which can then be subjected to the purification or analysis processes discussed in the following sections (precipitation, centrifugation, chromatography, etc.). Sources for extracellular proteins are, for example, culture debris from microorganisms, plant and animal cell culture media or also body fluids such as milk, blood, urine or cerebrospinal fluid. Usually extracellular proteins are present in solution in relatively low concentrations and require an efficient concentration as the next step. In order to isolate an intracellular protein, the cells must be disrupted in a manner that releases the soluble contents of the cell and leaves the protein of interest intact. The methods of cell disruption (cell disruption) differ primarily depending on the type of cell and the number of cells to be disrupted. Membrane proteins and other insoluble proteins Membrane-associated proteins are usually purified from the relevant membrane fraction after it has been isolated. In addition 2.2 purification scheme for various proteins. Depending on the localization and solubility of the proteins to be purified, various pre-purification steps have to be carried out before selective and highly efficient steps can follow.

5 2 Protein purification 17 peripheral membrane proteins that are only loosely bound to membranes are separated from the membrane by relatively mild conditions, e.g. high pH value, addition of EDTA or low concentrations of a non-ionic detergent, and can then often continue like Soluble proteins are treated. Integral membrane proteins, which aggregate outside their membrane via hydrophobic amino acid sequence regions and become insoluble, can only be isolated from the membrane with the help of high detergent concentrations; Structural proteins (e.g. elastin), which are sometimes also cross-linked via post-translational functional groups (post-translational modifications). The first and very efficient cleaning step here is the removal of all soluble proteins. Further steps are usually only possible under conditions that destroy the native structure of the protein. Further processing often takes place after the cross-linking on the denatured proteins has been broken up and using chaotropic reagents (e.g. urea) or detergents. Recombinant proteins A special situation arises in the production of recombinant proteins. A very simple purification results after the expression of recombinant proteins in inclusion bodies. These are dense aggregates of the recombinant product that are in a non-native state and are insoluble, be it because the protein concentration is too high, because the expressed protein cannot be correctly folded in the bacterial environment or because the formation of the (correct) Disulfide bridges in the reducing environment inside the bacterium is not possible. After a simple purification by differential centrifugation (Section 2.5.1), in which the other insoluble cell components are separated off, the recombinant protein is obtained practically pure, but it still has to be converted into the biologically active state by renaturation. If the expression of recombinant proteins does not lead to inclusion bodies, the protein is in a soluble state inside or outside the cell, depending on the vector used. Here the cleaning is very similar to the cleaning of natural proteins, only with the advantage that the protein to be isolated is already present in relatively large quantities. Recombinant proteins can be purified very easily by using specific marker structures (tags). Typical examples are the fusion proteins, in which at the DNA level the coding regions for a tag structure and for the desired protein are ligated and expressed as a protein. Such fusion proteins can be purified in a single step using specific affinity chromatography with antibodies against the tag structure. Examples of this are GST fusion proteins with antibodies against GST or biotinylated proteins via avidins. Another frequently used tag structure are polyhistidine residues, which are attached to the N- or C-terminal end of the protein chain and which can be easily isolated via immobilized metal affinity chromatography. Metal Affinity Chromatography Section Homogenization and Cell Disruption In order to be able to purify biological components from intact tissues, these complex cell assemblies have to be destroyed in a first step by homogenization. This creates a mixture of intact and broken cells, cell organelles, membrane fragments and also small chemical compounds that come from the cytoplasm and damaged subcellular compartments. Since the cellular components are transferred into an unphysiological environment, the homogenization medium should meet various basic requirements: protection of cells from osmotic bursting, protection from proteases, protection of biological activity (function), prevention of aggregation, as little interference with organs as possible biological analyzes and functional tests.

6 18 Part I: Protein analysis Table 2.1 Protease inhibitors Substance concentration Inhibitor of phenylmethylsulfonyl fluoride (PMSF) 0.1 1 mm aprotinin 0.01 0.3 μm serine proteases ε-amino-n-caprons ε 5 mm antipain leupeptin 70 μm 1 μm cysteine ​​proteases Pepstatin A 1 & mum aspartate proteases Ethylenediaminetetraacetic acid (EDTA) 0.5 1.5 mm Metalloproteases Usually this is done with isotonic buffers at a neutral pH value, to which a cocktail of protease inhibitors is often added (Tab. 2.1). If one wants to isolate intracellular organelles such as mitochondria, nuclei, microsomes etc. or intracellular proteins, the (still) intact cells have to be broken open. This is achieved by mechanically destroying the cell wall, during which frictional heat can arise and which should therefore be carried out with cooling as far as possible. The technical implementation of the digestion varies depending on the starting material and the localization of the desired target structure (Table 2.2). In the case of very sensitive cells (e.g. leukocytes, ciliates) it is often sufficient to repeatedly pipette the cell suspension or to press it through a sieve to allow digestion by means of black table 2.2 Biological starting materials and digestion methods Material Digestion method Comments Bacteria gram-positive enzymatic with lysozyme peptidoglycan cell wall EDTA / Tris Makes the cell wall permeable French press gram negative Cell grinder with glass beads Mechanical destruction of the cell wall Freeze-thawing Ultrasound for large quantities unsuitable due to local overheating Yeast autolysis with toluene French press several times, as inefficiently enzymatic inhibition of plants with glass beads Knife homogenizer high protease content in plants + dithiothreitol + phenol oxidase inactivators polyvinylpyrrolidone + protease inhibitors fibrous tissue grind in liquid nitrogen cold homogenization buffer do not phase Large tissue higher eukaryotes Grind, possibly after drying, cells that grow in suspension culture Osmolysis with hypotonic buffer Very sensitive cells Press through sieve Repeated pipetting of the suspension Add protease inhibitors Fibrous cells Chop up Dounce homogenizer Muscle tissue Chopping, meat grinder difficult to break down Enzymology, Vol. 182 Guide to Protein Purification, Academic Press 1990.)

7 2 Protein Purification 19 to achieve high shear forces. For the somewhat more stable animal cells, the shear forces are generated with a glass pestle in a glass tube (Dounce homogenizer). These methods are not suitable for plant and bacterial cells. Cells that do not have a cell wall and that are not associated in cell clusters (e.g. isolated blood cells) can be osmolytically disrupted by placing them in a hypotonic environment (e.g. in distilled water). The water penetrates the cells and makes them burst. In the case of cells with cell walls (bacteria, yeasts), the cell walls must be broken down enzymatically (e.g. with lysozyme) before osmolytic digestion is successful. This type of digestion is very gentle and is therefore particularly suitable for isolating cell nuclei and other organelles. For bacteria, repeated freezing and thawing is often used as a digestion method, whereby the change in the state of aggregation deforms the cell membranes in such a way that they break open and the intracellular content is released. Microorganisms and yeasts can be dried in a thin layer for two to three days at C, whereby the cell membrane is destroyed. The dried cells are ground in a mortar and can be stored at 4 C for a longer period of time. Soluble proteins can be brought back into solution in a few hours from the dry powder with an aqueous buffer. With cold, water-miscible organic solvents (acetone, 15 C, 10-fold volume) cells can be dehydrated quickly, whereby the lipids are extracted into the organic phase and the cell walls are destroyed. After centrifugation, the proteins remain in the precipitate, from which they can be recovered by extraction with aqueous solvents. In the case of stable cells such as plant cells, bacteria and yeasts, grinding with a mortar and pestle can be used to break down the cells, although larger organelles (chloroplasts) can also be damaged. The digestion is facilitated by adding an abrasive (sea sand, glass beads). For larger quantities, a knife homogenizer is suitable, in which the cell tissue is cut up by a rapidly rotating knife. This creates considerable heat, so that there should be an opportunity for cooling. For small objects such as bacteria and yeasts, the efficiency of the digestion is significantly improved by adding fine glass beads. Vibrating cell grinders are used for a relatively rough digestion of bacteria. These are lockable steel vessels in which the cells are vigorously shaken together with glass beads (diameter 0.1 0.5 mm). Here, too, the heat generated must be dissipated. Cell organelles can be damaged by this digestion method. Rapid changes in pressure break up cells and organelles very efficiently. For example, ultrasonic waves in the frequency range of khz are used to generate strong pressure changes in the suspension of a cell material via a metal rod. Since a lot of heat is released with this method too, only relatively small volumes and short sonication pulses of no more than ten seconds should be used. DNA is fragmented under these conditions. In another digestion method, which is particularly suitable for microorganisms, up to 50 ml of a cell suspension are pressed under pressure through a narrow opening (<1 mm), the cells being destroyed by the shear forces that occur (French press). Depending on the objective, the desired proteins are subjected to further purification steps in soluble form. For this purpose, the homogenate is usually roughly separated into different fractions using differential centrifugation methods (Section 2.5.1).

8 20 Part I: Protein Analysis 2.4 Precipitation The precipitation (precipitation) of proteins is one of the first techniques that was used for the purification of proteins (the salting out of proteins happened for the first time over 130 years ago!). The method is based on the interaction of precipitating agents with the proteins in solution. These agents can be relatively unspecific and precipitate practically all proteins from a solution, which is used in the first steps of a cleaning process to obtain the total proteins from a cell lysate. However, the precipitation can also be carried out in such a way that a fractionation of the constituents of a solution is possible. An example of this is the Kohn fractionation of plasma, which was worked out as early as 1946 and is still used today for the production of plasma protein on a large scale. Increasing amounts of cold ethanol are added to blood plasma and the respective precipitated protein is centrifuged off in fractions. With the exception of the precipitation of antigens with antibodies, the precipitation is not protein-specific and is therefore only used for a rough pre-purification of protein mixtures. Depending on the question and the starting material, the filling can be carried out under different conditions. In doing so, not only the efficiency of the filling itself, but also other aspects should always be taken into account: Is the biological activity impaired by the filling agent and the filling conditions? Under what conditions can the filler be removed? Determining the Concentration of Proteins Chapter 3 Salting Out The property of a salt to precipitate proteins is described in the so-called Hofmeister series (Fig. 2.3). The salts further to the left (so-called antichaotropic or kosmotropic) salts are particularly good and gentle filling agents. They increase hydrophobic effects in the solution and promote protein aggregations through hydrophobic interactions. The (chaotropic) salts on the right reduce hydrophobic effects and keep proteins in solution. The most common method of precipitating proteins is to salt them out by adding ammonium sulfate. Before the precipitation, the proteins should be present in a concentration of about 0.01-2%. Ammonium sulfate is particularly well suited, since in concentrations above 0.5 M it protects the biological activity of even sensitive proteins. It is easy to remove from the proteins again (dialysis, ion exchange) and, moreover, it is inexpensive, which is why it can also be used for fillings from larger volumes and thus already in the first cleaning steps. Ammonium sulfate is usually added in portions to the protein solution under controlled conditions (temperature, pH value), which enables fractional precipitation and thus an enrichment of the protein of interest. It should be noted that a complete filling can take a few hours! Ammonium sulfate precipitates should normally be centrifuged off tightly and well (100 g, see section 2.5). The only major disadvantage of ammonium sulfate concerns the precipitation of proteins, which require calcium for their activity / structure, since calcium sulfate is practically insoluble and is thus removed from the proteins. These proteins must therefore be filled with other salts (e.g. acetates). Filling with organic solvents It has been known for over a hundred years that proteins can be filled with cold acetone or short-chain alcohols (mainly ethanol). Long-chain alcohols (larger than C 5) are not soluble enough in water and cannot be used for filling. No general rules can be given for the choice of the organic filler or the optimal temperature. Ethanol has antichaotropic chaotropic cations: NH 4 + K + Na + C (NH 2) 3 + (guanidine) 2.3 The Hofmeister series. Anions: PO 3 4 SO 2 CH 3 COO 4 Cl Br NO 2 ClO 3 I SCN

9 2 Protein purification 21 has proven particularly useful for the precipitation of plasma proteins. For protein solutions that still contain lipids, acetone is often used, since in addition to the precipitation of the proteins, the lipids are extracted at the same time. In order to avoid too high local concentrations of the organic solvent, which can lead to the denaturation of the proteins, the solvent should be added slowly. Good cooling and slow addition are also useful, as adding the organic solvent (e.g. ethanol to water) can generate heat that leads to undesired denaturation. The precipitate is pelleted by centrifugation (see below) and taken up again in aqueous buffers. A frequently used protocol for acetone precipitation adds a 5-fold volume excess of 20 C cold acetone to the protein solution and incubates overnight at 20 C. It is then centrifuged for 30 min at g. This precipitation usually gives excellent results even for very small amounts of protein.The yield of the precipitation must be checked with analytical methods (SDS gel electrophoresis, activity tests, etc.). SDS gel electrophoresis Section Enzymatic activity tests Chapter 4 Filling with trichloroacetic acid A frequently used method to precipitate proteins from solutions is filling with ten percent trichloroacetic acid, whereby a final concentration of 3 4% should be achieved. After centrifugation, the precipitate is resuspended in the desired buffer and used further, whereby the pH of the solution should be checked. This method denatures the proteins and is therefore mainly used for concentration for gel electrophoresis or before enzymatic cleavage. The minimum sample concentration should be 5 & mug ml 1. Filling of nucleic acids Protein solutions from cell disruptions, especially from bacteria and yeasts, contain a large proportion of nucleic acids (DNA and RNA), which can interfere with protein purification and therefore usually have to be separated. Since nucleic acids are highly negatively charged polyanions, they can be filled with strongly basic substances (e.g. polyamines, polyethyleneimines or anion exchange resins) or very basic proteins (protamines). By optimizing the precipitation and washing conditions, it must be avoided that proteins of interest are also filled by the precipitation reagent or in complex with the nucleic acids (e.g. histones, ribosomes). 2.5 Centrifugation Centrifugation is not only one of the most common techniques for separating insoluble components, but also for cell fractionation and the isolation of cell organelles. It is based on the movement of particles in a liquid medium by centrifugal forces. The central component of a centrifuge is the rotor, which is used to hold the sample beakers and is driven by a motor at high speed. There are different designs of the rotors, such as fixed-angle rotors, vertical or vibrating bucket rotors (Fig. 2.4), which are available in different sizes and materials. They allow separations from a few microliters to a few liters and can be operated with different, adjustable rotational speeds depending on the task at hand. Coolable centrifuges are mostly used for working with biological materials. High-speed centrifuges, the ultracentrifuges, are always operated connected to a vacuum system in order to avoid the frictional heat that occurs at high speeds as a result of air resistance. When operating centrifuges, certain safety measures must be observed, above all the opposing sample vessels must be well balanced in order to avoid any imbalance that could destroy the centrifuge. Basics The physical principle of centrifugation is a separation according to size and density. On a particle that is moved around an axis of rotation with constant angular velocity ω,

10 22 Part I: Protein analysis Fixed-angle rotor A B C D E Vertical rotor Swing-out rotor 2.4 Rotors for centrifugation. Fixed-angle rotor, vertical rotor and swing-bucket rotor when loaded (A) under centrifugation conditions at the beginning of the separation (B) during the separation (C) when braking (D) and after the end of the centrifugation (E). Fractions containing protein are shown in red. a centrifugal force acts, which accelerates the particle outward. The acceleration B depends on the angular velocity ω and the distance r from the axis of rotation: B = ω 2 r (2.1) The acceleration is related to the acceleration due to gravity g (981 cm s 2) and is expressed as a relative centrifugal acceleration RZB in multiples of the acceleration due to gravity (g ) specified: 2r RZB = 981 (2.2) The relationship between the angular speed and the rotational speed in rotations per min (rpm) is given by:

11 2 Protein purification 23 = rpm 30 (2.3) which results from substitution: RCF = 1, r (rpm) 2 (2.4) It must be taken into account that normally during centrifugation the distance of the particles from the axis of rotation and thus also the RCF change. The mean value is therefore often used for conversions. The sedimentation speed of spherical particles in a viscous liquid is described by the adjacent Stokes equation. Here v is the sedimentation velocity, g is the relative centrifugal acceleration, d is the diameter of the particle, & rho p and & rho m are the density of the particle or the liquid and η is the viscosity of the medium. The sedimentation speed increases with the square of the particle diameter and the difference in density between particles and medium and decreases with the viscosity of the liquid. If the sedimentation takes place in a medium such as 0.25 M sucrose, which is less dense than all particles and also has a low viscosity, the diameter of the particles is the dominant factor for the sedimentation rate. The sedimentation coefficient s is the sedimentation speed under geometrically given conditions of the centrifugal field. It is given in Svedberg units (S). Stokes equation: d = 2 () g p 18 m (2.5) v s = r 2 (2.6) 1 S corresponds to seconds. Various biological molecules lie in this order of magnitude. The Svedberg unit of a biomolecule is sometimes included in its name (e.g. 18S rRNA), which then allows a conclusion about the size of the particle. Table 2.3 shows the size and centrifugation conditions for cleaning cells and some cellular compartments. A good overview is also given by the representation of the particles in a density / sedimentation coefficient diagram (Fig. 2.5) or in a density / g-value diagram. The various techniques of centrifugation can easily be understood from Stokes' equation. Density (in g / ml) Sedimentation coefficient (in S) 2.5 Density and sedimentation coefficient of some cell compartments. The figure shows the distribution of different cell components with regard to their density and their sedimentation coefficient.


Protein Purification Basics

1 Basics of protein purification I. Why do you clean enzymes at all? Detailed studies on the mode of action of an enzyme only lead to success if the protein can be isolated and separated from the hundreds of other proteins of a liver cell, a yeast or a bacterium. For example, in order to clearly characterize the conversion of compound A to compound B catalyzed by enzyme x, it is necessary to remove all activities that also have an influence on A and B (dashed arrows). Only conversions of compound A with homogeneous enzyme x enable an unequivocal qualitative and quantitative detection of product B and its rate of formation. The determination of the amino acid composition and the N-terminal sequence, attempts to crystallize the enzyme x A B X-ray structure, mass spectrometric investigations and numerous other techniques of protein characterization also require homogeneous protein. Otto Warburg laid the foundation for the enzyme purification discipline with numerous inspiring publications (1930) and techniques. Since then, the repertoire of methods has developed by leaps and bounds, with the focus of the last decade primarily on miniaturization, automation and optimization of known principles. The standard of every enzyme purification is the achievement of the maximum possible specific activity, the ratio of activity to protein amount. This ratio should increase with each successful cleaning step until it finally reaches a maximum value with the homogeneous enzyme preparation. In practice, however, a loss of activity of the enzyme that is often associated with the purification, clouds the theoretically expected increase in the enrichment factor. The loss of catalytic activity is the most common undesirable side effect. It is the reason why enzyme purifications usually have to be carried out quickly. Once released from the protective physiological state of the cell interior, enzymes often come into contact with high ion concentrations, absorbing glass surfaces, elevated temperatures, metals, oxygen and other potentially harmful influences during purification. All of these factors often lead to irreversible denaturation of the fragile structure of enzymes. Enzymes should not be viewed as stable chemical compounds but rather as labile biomolecules, which require special attention when handling. A dirty, inadequately cleaned glass container to which traces of a surfactant or a metal salt still adhere can irreversibly destroy the activity and thus destroy days, weeks or months of work in one fell swoop. II. Cultivation, Harvesting and Production of Extracts of Prokaryotic Cells a) Cultivation Cell disruption is preceded by the cultivation and harvesting of the corresponding organisms. When selecting the substrate and the growth conditions, it must first be ensured that the desired enzyme is induced and expressed. 1

2 Although Pseudomonas putida grows excellently on NB complex medium, it does not express any enzymes for aromatic degradation on this nutrient medium. Enzymes such as pyrocatechol-1,2-dioxygenase or muconate cycloisomerase are only induced when growing on benzoate as the sole energy and carbon source or at least in its presence. b) Harvesting It is also crucial that the cells are harvested at a point in time at which the expression and thus the concentration of the desired protein is at its maximum. In the case of metabolic enzymes, this is usually the case during the exponential growth phase. In order to achieve a maximum cell yield at the same time, harvesting is normally carried out in the late exponential growth phase (Fig. 1). Fig. 1: The phases of growth of microorganisms in batch culture and the most favorable harvest phase The cells are harvested by centrifugation. This and all following steps of protein purification should, if possible, be cooled (+4 C) in order to minimize loss of activity. Repeated washing (= resuspension and renewed centrifugation) of the cell pellet with the buffer suitable for later cell disruption prevents residues of the culture medium from being dragged along which negatively affect the subsequent purification or stability of the protein. The choice of the digestion buffer and any stabilizing additives are of great importance. The activity and stability of the protein to be purified should not be negatively influenced by the buffer (e.g. by complexing metals or cofactors). High buffer concentrations should be avoided as they usually interfere with later chromatographic steps. In addition, various additives should be tested for their potential protective effect in preliminary tests. Examples are phenylmethylsulfonyl fluoride and the reducing dithiothreitol (DTE) as protease inhibitors or antioxidants. The addition of certain metal ions can also have a stabilizing effect on the activity (e.g. Mn 2+ in the case of muconate cycloisomerase). If harvested cells cannot be further processed directly or if the required amount of cells can only be obtained over a longer period of time, it can be stored at -20 to -70 C. However, storage mostly means loss of activity and should be avoided if possible. 2

3 c) Cell disruption The cell disruption for the production of cell-free lysates is the introductory step in the protein enrichment process. Since it has an impact on the total amount of protein to be enriched, its biological activity, its integrity against proteolytic digestion, its association with other cellular components and the presence of additional contamination, the method used is of great importance. A general distinction is made between methods of enzymatic lysis and those of mechanical lysis. Methods of enzymatic cell lysis are based on the digestion of cell wall components (peptidoglycan framework) of the cell wall, for example by lysozyme. The exposed, highly sensitive protoplasts are then opened by detergents, osmotic shock or mechanical treatment. Enzymatic methods of cell lysis minimize protein denaturation, work regardless of the size of the batch and, in certain cases, allow a certain selectivity in the release of cellular components. Disadvantages are a large number of factors that can influence the success of the lysis and, on the other hand, the addition of substances that can interfere with later cleaning steps. Methods of mechanical cell lysis include methods of shaking with simultaneous shaking and methods of shearing. As part of the shaking method, a cell suspension is shaken in a closed chamber together with fine glass beads at high frequency (vibrating mill). The force of the impact of the pearls as well as the simultaneous shear forces fragment the cells. Care must be taken to ensure adequate cooling. A common device for opening cells by shearing in the liquid phase is the so-called French press. In it, the cell suspension is passed under high pressure through a tiny opening, behind which there is sudden relaxation and thus the occurrence of strong shear forces. Variables are the height of the selected pressure (7,000 to psi) and the number of passes. Ultrasound is another common method of cell opening. An ultrasound probe is immersed directly into the cell suspension to be treated and pulses lasting 30 to 45 seconds are generated. The disadvantage of this method is that it generates a lot of heat. Intensive cooling and working with pulse intervals are necessary. Fig. 2: French press with a small digestion cell The advantages of the purely mechanical digestion methods lie in addition to the absence of possibly disturbing additives, in particular in the large quantities of cells that can be processed. Cells also differ comparatively little with regard to the parameters necessary for opening. Depending on the decomposition method selected, the lysate obtained contains more or less chromosomal DNA. Due to the high viscosity, their removal is slow and easy

4 necessary due to interactions with chromatographic purification steps and is generally carried out by adding DNAse. The removal of insoluble membrane and cell wall components is achieved by ultracentrifugation. The clear supernatant obtained in this way is referred to as cell-free crude extract. III. Determination of protein content and activities of cell-free enzymes a) Activity determination The specific detection of the protein to be enriched in the crude extract as well as in fractions of chromatic separations is done with the help of activity determinations. Depending on the procedure, these can be divided into continuous, coupled and discontinuous verifications. Continuous evidence is the simplest in terms of the technical effort involved. With them, the enzymatic reaction can be followed photometrically online by measuring either the decrease in the substrate, the increase in the product or the decrease / increase in a required cosubstrate. Continuous verifications are also possible with the help of other suitable measuring techniques (fluorescence, ph value, viscosity, heat development). Coupled evidence is required if none of the components involved in the reaction can be detected by UV / VIS spectroscopy. By coupling the reaction with a second (or third) reaction, which in turn has a chromophoric reactant, the speed of the original reaction can be deduced under certain framework conditions (Fig. 3). Fig. 3: Principle of a coupled enzyme test for phosphofructokinase with lactate dehydrogenase as indicator enzyme (NADH decrease is measured) Discontinuous detection requires the isolation of the product due to its special detection (e.g. detection of radionuclides, HPLC and GC detection). In order to determine conversion rates, several parallel cycles have to be stopped (quenched) at different times and the concentration of substrate or product has to be determined in each case. 4th

5 The enzyme activity is influenced by the concentration of the substrates, activators, inhibitors, the salt concentration, the pH value, the ionic strength and the temperature. Although the enzyme test runs under optimized conditions, it is usually not necessary to take all factors into account. Fig. 4: Effect of the substrate consumption on the enzyme activity: (a) Dependence of the conversion rate on the substrate concentration (b) Measured initial conversion rates depending on the substrate concentration Formally, the activity of an enzyme is at its highest at an infinitely high substrate concentration (Fig. 4a), a fact that is not is feasible. If the enzyme obeys Michaelis-Menten kinetics, the substrate concentration used experimentally should be at least 10 times the k m value of the substrate. At [substrate] = 10 x km, the speed should theoretically be 91% of the maximum possible at [substrate] =.The use of high substrate concentrations has several advantages, on the one hand, greater inaccuracies in the measurement of the substrate addition only have a relatively minor effect on the measured conversion rate, on the other hand, the decrease in substrate at the beginning of the reaction does not cause any noticeable reduction in the conversion rate (Fig. 4b). The inhibitory effects of products are also lower in the presence of high substrate concentrations. The measured initial speed of the reaction, which occurs within the first minute (s) after all components have been mixed, serves as the basis for determining the activity. The enzyme sample may have to be diluted beforehand to such an extent that the initial speed can still be conveniently measured, i.e. the course of the reaction is neither too fast nor too slow. Any blind reaction that may be observed must be subtracted from the measured activity. A blank reaction results from an unspecific decrease in the substrate / cosubstrate, for example through disintegration or through reaction with other enzymes in the protein sample. Enzyme activities are usually given in enzyme units (units, U). An enzyme unit U is defined as the activity which catalyzes the conversion of 1 µmole substrate or the formation of 1 µmole product in one minute at 25 ° C. The specific activity is given in units per mg protein (U / mg protein). b) Protein determination In addition to the specific quantification of the desired enzyme through activity measurement, the determination of the total protein content is necessary for the determination of the specific 5

6 specific activity required. The latter provides information about the success and efficiency of a cleaning step. The most common methods for protein determination are: - Biuret reaction (relatively low sensitivity 0.05-5 mg / ml) - Lowry method (sensitive 0.05-0.5 mg / ml, susceptible to interference) - UV absorption (simplest measuring principle , no sample consumption, 0.05-2 mg / ml (280 nm), 0.01-0.05 (205 nm)) - dye binding (very sensitive 0.01-0.05 mg / ml, very popular) - BCA reagent (very sensitive 0.005-0.05 mg / ml, similar to the Lowry method) The most frequently used dye method uses the reagent Coomassie Blue G-250. The reagent is dissolved in acid and has a red-brown color in the protonated state. When it binds to positively charged protein residues, the color changes to blue and can be detected at 595 nm. Linearity is only given in a certain concentration range (approx. & Microg protein / ml). The deposition of blue dye residues on the surface of the cuvette can be problematic, as it suggests that the measured values ​​are too high in subsequent measurements. The use of disposable plastic cuvettes is therefore advantageous. c) Summary of the measurement values ​​The course of enrichments is usually summarized in the form of an enrichment table (Tab. 1). Tab. 1: Typical enrichment table of a protein purification (here: maleyl acetate reductase from Pseudomonas sp. B13 purification step total volume [ml] total protein [mg] total activity [U] specific activity [U / mgprot.] Yield enrichment factor [%] raw extract, ion exchange, 3 4.2 83 Chromatography Hydrophobic Interaction, 2 45.2 64 Chromatography Affinity Chromatography 75 7,, 3 145.5 36 Gel Filtration 12 4,, 9 149.5 23 IV. Precipitation Methods In the early days of protein purification, precipitation methods of proteins were , based on changing the solvent properties, the only available strategy of enrichment. This has been reflected in some of the original protein names (e.g. globulins, serum globulins, albumins, prolamines) ) and hydrophobic (water-repellent nder) structural features are determined on its surface. Although hydrophobic structural elements are preferably present in the interior of the enzyme, a not insignificant amount of them (depending on the type of protein) is on the surface and comes into contact with the solvent (Fig. 5A). The solvent used in precipitation methods is in most cases water. The dissolving properties of proteins can be changed by changing the ionic strength, the pH value, the temperature, by adding water-soluble organic solvents, organic polymers or a combination of different factors. The most common is the use of neutral salts. 6th

7 ABC hydrophobic surface Fig. 5: A) Distribution of charges and hydrophobic structures on the surface of a typical enzyme B) Electrostatic interactions between protein molecules and small aggregates C) Solubility of a protein of the globulin type near its isoelectric point a) The solubility of proteins at low salt concentrations Most proteins are in dissolved form inside the cell. The cytoplasm contains a high concentration of various proteins (40%) under physiological conditions (ionic strength 0.15 0.2 M, neutral ph value). If one extracts the cell contents one can change these conditions. The solubility of a protein is the result of several effects: - polar interactions of charged structural elements with the solvent - ionic interactions with dissolved salt - repulsive electrostatic forces between similarly charged molecules or small aggregates (Fig. 5B) Some enzymes (e.g. globulins) already form the ionic strength of the medium is between 0 and the physiological value. This can have two causes: - large hydrophobic surface fractions reduce polar and ionic interactions with solvent or salt - a net charge of zero (isoelectric point) minimizes electrostatic repulsion and promotes hydrophobic interactions with one another.Therefore, proteins generally have their lowest point of solubility in the isoelectric point and can (provided that the solubility is sufficiently low) precipitate by changing the pH accordingly (Fig. 5C). b) Protein precipitation by salting out at high salt concentrations This technique is the most frequently used. In contrast to the salting-in method, which depends largely on the charge distribution on the protein surface and the resulting polar interactions with the solvent, the salting-out effect is primarily based on the presence of hydrophobic surface structures. A suitable model for illustration is the following: water molecules orient themselves in the direct vicinity of a hydrophobic surface and form ordered hydration envelopes (Fig. 6). This prevents the approach and association between two such hydrophobic surfaces. To the extent that salt is added to such a system, the water molecules are required to hydrate the much more polar ions and are increasingly removed from the protein shell. The free hydrophobic areas can now aggregate, the protein 7

8 fails. Proteins with a higher proportion of hydrophobic surface structures aggregate earlier with increasing salt concentration. Water molecules Hydrophobic structural element Fig. 6: Anomaly of water molecules around hydrophobic surface structures The type of ions plays a major role in salting out. Salts from polyvalent anions (sulfate, phosphate) and harmless cations (ammonium, potassium, sodium) are beneficial. Ammonium sulfate is the most commonly used ionic filler for several reasons. It is sufficiently soluble in water (up to 4 M), shows no significant heat of dissolution, has a favorable density in a watery solution (important for the centrifugation of precipitated protein), stabilizes enzyme activity in microbes and prevents concentrated enzyme growth. Various methods exist for salting out proteins. If the volume of the protein solution is large, we recommend filling with solid salt. It is added in finely powdered form while stirring the cooled precipitate in portions until the desired concentration is reached. After sufficient equilibration, precipitate and solution are separated by centrifugation and examined for activity. In the case of a fractionated procedure, the supernatant is subjected to another salt addition, centrifuged again, etc. If the sample volume is small and does not matter or if the salt concentration required for the filling is low (50% saturation), it can alternatively be filled with concentrated salt solution. The advantage lies in faster equilibration and the avoidance of punctual overconcentrations, which inevitably arise when solid salt is added. A great advantage of ammonium sulfate precipitation is the frequently observed stabilization of enzymes. An enzyme precipitate containing 2 3 mol / l (NH 4) 2 SO 4 often retains its activity unchanged for years (typical commercial form of enzymes). If an enrichment procedure is to be interrupted for a longer period of time, storage as ammonium sulfate precipitate is therefore preferable. Due to the high concentrations of ammonium sulphate necessary for an accident, contamination with heavy metals can have an unfavorable effect on the enzyme activity, even in trace amounts. Elimination is achieved by adding small amounts of EDTA. c) Protein filling with organic solvents The filling of proteins with water-soluble organic solvents (e.g. ethanol, acetone, methanol, n-propanol) is used comparatively little. Since it is based on different phenomena than precipitation by salting out, it represents an additional possibility for enrichment. An advantage of water is 8

Mixing water / solvent means that you can work at temperatures below 0 C, but this is also necessary because denaturation becomes significant at temperatures above 10 C. d) Protein precipitation through denaturation Heat precipitation If the enzyme to be purified has above-average heat stability at (50 C), heat precipitation is an extremely simple option for enrichment. Proteins that cannot withstand this temperature and coagulate can be centrifuged off. V. Separation by Adsorption Chromatographic Techniques Proteins adsorb on a multitude of solid phases, mostly in a specific way. The use of adsorptive techniques, especially in the context of column chromatography, is popular and usually provides the highest levels of enrichment. Adsorptive batch processes offer a comparatively poorer resolution, but are characterized by their simpler and faster implementation and are particularly justified in the case of large initial quantities. The following are used as specific adsorbents for proteins: - ion exchangers - hydrophobic materials - inorganic compounds (calcium phosphate, hydroxyapatite) - fixed ligands - affinity adsorbents (immobilized substances, inhibitors, cofactors, antibodies), these compounds are mostly immobilized on a gel matrix (agular rose). The latter should prevent the adsorbent from being washed out, guarantee a high eluent flow due to its spherical shape and at the same time offer a high surface area and thus capacity. If the particles also have pores, the resulting molecular sieve effect can bring about an additional selection. a) Ion exchange chromatography Proteins bind to ion exchangers due to electrostatic interactions between their charged surface and the charged residues of the ion exchanger, whereby a corresponding amount of counterions is displaced (Fig. 7). To give a feeling for the order of magnitude of the number of these interactions: the average distance between charged groups of a diethylaminoethyl cellulose ion exchanger (DEAE cellulose) is approx. 1.5 nm, the diameter of a globular protein is approx With a potentially high protein binding capacity, ion exchange chromatography is a suitable introductory step in enzyme enrichment. However, the capacity of a certain protein depends essentially on its size. The larger a protein, the lower its binding capacity. Particularly large proteins (& gt 10 6 Da) only bind to the surface, but not in the pores of the exchanger and are excluded (Trisacryl). 9

10 Fig. 7: Binding of a negatively charged protein to an anion exchanger. Seven positively charged ions (e.g. HTris +) associated with the protein are exchanged together with seven negative counterions (Cl-) of the ion exchanger. The two decision criteria for the choice of adsorbent are: - the charge, + or and the nature of the charged group - the type of matrix (particle shape and size), flow rate, capacity, costs The majority of proteins have a pH value between 7 and 8 have a negative overall charge and can be adsorbed under these conditions with an anion exchange material. Tab. 2: Choice of ion exchanger for purifying a protein with a known isoelectric point isoelectric ion exchanger buffer ph point 8.5 cationic 7.0 7.0 cationic 6.0 anionic 8.0 5.5 anionic 6.5 The pH value within an ion exchanger is not identical to that of the elution buffer. Responsible for this is the Donnan effect, which describes the adsorption and release of protons from the matrix. In general, the ph value is 1 unit higher within anion exchangers and 1 unit lower within cation exchangers. The difference is greater, the lower the ionic strength of the buffer used. A buffer should therefore generally be selected which has the highest buffer effect for a given ionic strength. The Donnan effect must be taken into account with regard to the optimum pH stability of an enzyme. A bound protein can generally be eluted using two methods: - Changing the pH value of the eluent (lowering the pH for anion exchangers, increasing the pH for cation exchangers) - Increasing the ionic strength of the eluent (reducing the electrostatic interactions between protein and adsorbent) The ph method is associated with numerous difficulties and restrictions, apart from a few exceptions, the high salt gradient method is used, mostly using sodium and potassium chloride. 10

11 The desorptive effect of the salt can on the one hand be explained by the ion exchange effect (the ions of the salt displace the charged protein residues), on the other hand the increasing ionic strength weakens the electrostatic interactions necessary for binding (see also salting out). b) Hydrophobic interaction chromatography Hydrophobic interactions are of great biochemical importance. They are essentially involved in the stabilization of three-dimensional tertiary structures of proteins, in antibody-antigen reactions and in enzyme-substrate binding reactions. By hydrophobic interaction one understands the phenomenon that 2 hydrophobic molecules aggregate spontaneously in a polar environment (e.g. water). The driving force for this is an increase in entropy, which is always thermodynamically favored. It follows from this that hydrophobic interaction is not a force per se, but is only enforced by a polar environment. By dissolving a salt and increasing the ionic strength of the medium, the hydrophobic interaction of two non-polar molecules also increases. Proteins have more or less high proportions of hydrophobic surface structures. With a correspondingly high ionic strength, they are therefore able to adhere to hydrophobic adsorbents. In addition to the salt content, the strength of the interaction can be controlled by the choice of adsorbent. Materials with low hydrophobicity (linked butyl residues) are preferably used for strongly hydrophobic proteins, materials with high hydrophobicity (e.g. octyl residues) correspondingly for less hydrophobic proteins. Gels with phenyl residues can be classified roughly in the middle in terms of their hydrophobicity and are suitable for most proteins. The adsorption takes place in the presence of high salt concentrations, the elution accordingly with a descending salt gradient. The same criteria apply to the choice of ions as to the protein filling, ammonium sulfate is the most common salt. c) Hydroxyapatite chromatography, affinity chromatography, dye ligand chromatography, immunoadsorption Reference is made to general literature on protein purification (see point VII.). VI. Separation in solution gel filtration (size exclusion chromatography), electrophoretic methods a) Gel filtration (size exclusion chromatography) The fractionation of proteins on the basis of size exclusion techniques gives gel filtration a special position among the techniques for protein filtration. The term gel filtration is unfortunate because, in contrast to conventional filtration, no components are retained. The absence of adsorptive phenomena has the advantages and disadvantages of gel filtration. On the one hand, sensitive proteins are not impaired by binding, on the other hand, the lack of a specific binding worsens the chromatographic resolution of this technology.Gel filtration requires a porous gel matrix of as precisely defined pore sizes as possible (Fig. 8A). A column packed with such spherical gel particles has 2 different measurable liquid volumes. The exclusion volume, which corresponds to the volume of the liquid outside and between the gel particles 11

12 corresponds, and the inclusion volume, which corresponds essentially to the liquid within the gel particles. A mixture of proteins is applied in the smallest possible volume to the surface of a gel filtration column and allowed to flow through it. Large molecules cannot diffuse into the pores and are the first to migrate through the column within the exclusion volume. Small proteins diffuse into the porous gel particles and elute accordingly later (Fig. 8B). Both the pore size and the diameter (Stokes radius) of the protein are decisive for the separation. The more appropriate term size-exclusion chromatography is increasingly being used. Fig. 8: Course of the size-exclusion chromatography A B Two-dimensional representation of the pores of a gel matrix and their accessibility for molecules of various sizes (no realistic size ratios). Simple illustration of the time course of a size-exclusion chromatography. Large molecules are excluded from the largest part of the existing bed volume and migrate almost unhindered with the solvent front. If all proteins in a mixture have a similar spherical structure, the sequence of ripening corresponds to the elution of their reciprocal molecular weights. The elution profile of a gel filtration is sketched in Fig. 9A. The empty or excluded volume V 0 of the column is determined by eluting a compound with a very high molecular weight (e.g. dextran blue 2000 MW & gt 2 million). The elution volume V t for the smallest standard protein corresponds to the sum of the empty volume V 0 and the inclusion volume of the gel matrix. Elution volumes of proteins between these border areas are denoted by V e. The mobility of a protein can be given in the form of its distribution coefficient K av: K av = V e - V 0 / V t - V 0 The semi-logarithmic plot of K av against the corresponding molecular weight (Fig. 9B) results in a sigmoid curve. The separation of proteins is greatest in the linear range (K av: 0.2 0.8); this range is therefore also indicated as the separation or fractionation range of a gel matrix. The steeper the sigmoid area within the fractionation area, the higher the dissolving power of the column material. For the separation of proteins with low 12

For differences in molecular weight, material with the smallest possible fractionation range is used. Fig. 9: Chromatographic performance of a size exclusion matrix A B describes a simple elution profile. Component 1 is completely excluded from the matrix (V 0), component 2 partially (V e) and component 3 not (V t). describes the sigmoid dependence of the distribution coefficient K av on the logarithm of the molecular weight. The separation process of a gel filtration column is limited to a maximum of 10 proteins and therefore only allows moderate enrichment factors. Gel filtration is therefore mostly used at a relatively late point in time within an enrichment, when the number of remaining contaminations is already low. b) Electrophoretic methods Electrophoretic methods enable the high-resolution separation of proteins primarily in the analytical area. In particular, SDS polyacrylamide gel electrophoresis is predestined for the purity control of preparations in the course of protein purification and is used routinely for this purpose. The electrophoretic principle: A protein molecule has a certain net charge in aqueous solution at every ph value that does not correspond to the isoelectric point. This causes the protein molecule to move in the electric field. The specific mobility & upsilon is proportional to the number of net charges per molecule z and inversely proportional to the viscosity of the medium & eta and the particle radius r (Stokes radius): & upsilon = z / 6 & pi & etar The separation of proteins during electrophoresis in free solution occurs due to their different solutions Charge and its different sizes and is deteriorated by diffusion and intermolecular electrostatic interactions

14 tert. The diffusion is minimized by the lowest possible working temperatures, the electrostatic interactions by high ionic strength or an anionic detergent. Since an increase in the ion strength also causes an increase in the current strength during electrophoresis, care must be taken to ensure adequate heat dissipation. In addition, buffers are used as electrolytes, in which the ions have a high molecular size and thus a low mobility and conductivity (e.g. HTris +). Electrophoreses are usually carried out at a neutral or slightly alkaline pH value, during which most proteins migrate to the anode. Gel electrophoresis is a technique that is far superior to electrophoresis in free solution in terms of separation performance. Here the electrophoresis takes place within a network-like matrix, which has pores of different diameters. Depending on the size of the molecules, these pores lead to different effective viscosities of the medium. Gel electrophoresis therefore separates on the basis of both the charge and the size. The gel can be optimized for a specific separation problem (a size range to be separated) via the degree of crosslinking. A main advantage of gel systems is the minimization of convection and diffusion of proteins, which is evident in the high band sharpness during the entire electrophoresis process. ABC Cathode buffer tank Tubes with gel cylinders Protein-containing gel zones after coloring Sample in sample buffer Buffer supernatant Cathode gel between glass plates Anode buffer tank Fig. 10: (A) Experimental setup for disk gel electrophoresis (B) Localization of separated proteins in disk gels (C) Storage of a slab Early starch gels gave way in the 1960s (1964) to synthetic polyacrylamide gels, the properties of which were more controllable and reproducible. The most favorable gel form developed in the thin-slab form (Fig. 10C), which, in contrast to the long-used disk form (Fig. 10A, B), enables the application of several samples, and at the same time enables better heat dissipation due to the high surface area. Simple (native) gel electrophoresis: The proteins are separated here in their active, unchanged (native) form. Sample and running buffers contain neither SDS nor urea. The separation takes place according to size and load. Most proteins are negatively charged in the weakly alkaline running buffer (ph 8-9) used and migrate to the anode. All proteins that are positively positioned under these conditions do not get into the gel but rather diffuse into the cathode buffer. Native gel electrophoresis provides the basis for detecting the activity of proteins on the gel. 14th

15 SDS gel electrophoresis (SDS-PAGE): In this, the most popular of all gel electrophoretic methods, the enzymes are denatured with the aid of the detergent SDS (sodium dodecyl sulfate) before separation. The addition of 2-mercaptoethanol serves to cleave existing disulfide bridge bonds. Compared to native gel electrophoresis, it offers two advantages: - The separation is only dependent on the size of the protein, very high resolution (1% of MW) - Aggregates and insoluble particles are then separated and the protein can be separated into the individual proteins + SDS denatured polypeptide in the form of a strongly negatively charged chain Mobility [mm] Fig. 11: The denaturing effect of SDS log (MG) Fig. 12: Plot of mobility versus molecular weight for a number of proteins (size standards) SDS binds strongly Proteins. The resulting polypeptide chains contain 1 molecule of SDS on 2 amino residues each. Oligomers in which the polypeptides are not covalently linked to one another are cleaved into the individual subunits. Each SDS molecule has a negative charge, so for a typical polypeptide of this mass there are 180 negative charges. This far exceeds the number of net charges on the protein at neutral pH. Accordingly, the charge-to-size ratio is practically the same for all proteins and the separation is solely a result of the molecular sieve effect of the gel pores. Despite the fact that SDS-PAGE does not allow separation of proteins of identical sizes, it offers the highest resolution as a simple standard method. The entire gel can be calibrated by applying size standards at the same time. The graphical single-logarithmic plot of the mobility of proteins against their molecular weight provides a linear dependence within a certain range. The degree of crosslinking of the polyacrylamide gel and thus its pore size can be adapted to the size of the proteins to be separated over a wide range. Two variations are possible: - Change in the acrylamide content (3-20%) - Change in the crosslinker content bisacrylamide (0.1-1%) Very large proteins (1000 kda) are mixed with low concentrations (3-4% acrylamide, 0.1% bisacrylamide) gels separated, very small proteins (10 kda) require strongly cross-linked gels (20% acrylamide, 1% bisacrylamide). 12% gels are used as standard. 15th

16 Isoelectric focusing: With IEF there is a watery solution of various ampholytes between the cathode and anode. When an electric field is applied, these form a linear ph gradient. Ampholytes are highly complex mixtures (several hundred to thousands) of very small proteins, each of which has its own individual pi and builds up when an electric field is applied. The linear ph gradient is actually based on the formation of a large number of discrete ph values ​​that cover the specified range relatively evenly. Admitted proteins migrate in this field up to the point at which the prevailing pH value corresponds to their isoelectric point. The neutral overall charge prevents further migration, and a focus effect is also achieved through particularly high fields. The gel used here only has the task of stabilizing the ph gradient. Proteins should not be impaired in their movement by molecular sieve effects, the gel concentrations used are therefore as low as possible. The IEF usually uses columnar gels. One advantage of the IEF is its high resolution, which can already be seen in the term focus. Most other protein separation methods are associated with diffusion and mixing phenomena, which show negative effects over time. Diffusion no longer plays a role in the IEF. If a protein molecule moves out of the range of its isoelectric point, it is charged and immediately moves back. Once the IEF is over, the system is in equilibrium and theoretically no more movement takes place (with the field applied!). Due to the absence of conductive ions, almost no current flows and the resulting high voltages cause additional band sharpness. Certain phenomena can limit the use of IEF: - the protein is not stable at its isoelectric point (e.g. disintegration into subunits) - the protein is not sufficiently soluble at its isoelectric point - the protein forms complexes with the ampholytes (which occurs during formation several bands noticeable) The IEF is the 1st stage in the highest resolution 2D gel electrophoresis, in which the separation of even the most complex protein mixtures is possible through combination with SDS-PAGE as the 2nd stage. Coloring and detection of proteins after gel electrophoresis: After gel electrophoresis, protein-containing zones must be made visible, which is usually the case. happens with the help of an organic dye, which binds firmly to proteins. The choice of the dye depends on: - its detection sensitivity - its ability to color all types of proteins evenly. Known dyes are Coomassie Blue R-250, Coomassie Blue G250, Amido Black, Nigrosine. The coloring with org. Dyes include the following steps: - Immediately after electrophoretic separation, the proteins are fixed by denaturation (otherwise the bands would diffuse). Usual: methanol / acetic acid / water in a ratio of 3: 1: 6. - The gel is incubated in a dye solution until it is completely saturated. - Excess, unbound dye is removed by swirling in a decolorizing solution. A dyeing method that is one factor more sensitive than Coomassie blue is the silver stain method. Treating the gel with silver nitrate causes 16

17 Non-stoichiometric binding of silver to proteins. These complexes become visible as black to brown bands after reduction. Due to the high sensitivity, the necessary sample amount is minimized and overload effects are avoided. The disadvantage is that proteins can behave very differently in terms of their coloring behavior towards silver stain. The specific detection of a protein on the gel succeeds with the help of a suitable activity test (only with native gels) or with the help of appropriate monoclonal antibodies. VII. In-depth literature: Protein Purification Robert K. Scopes. 3rd edition. Springer-Verlag, New York Methods in Enzymology. Vol Guide to protein purification ed: Murray P. Deutscher. Academic Press San Diego 17


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5 Current Topics Typical projects are based on a biological question or a medical problem, identify the molecules involved or a specific receptor as a target and pursue the solution with model systems, synthetic ligands and intelligent assays.

6 Modern interdisciplinary content You learn from nature, e.g. About molecular evolution, enzymatic specificity and catalysis, medical mechanisms of action, molecular communication, biosynthetic processes and uses methods of macromolecular biochemistry, biomimetic synthesis, biophysics. Structure elucidation of combinatorial assays

7 Structure of the new Bachelor's degree program (6 semesters) Professional objective Master (4 semesters) Professional objective PhD (6 semesters) Professional objective Habilitation / junior professorship

8 Bachelor's degree in Basics of Chemistry: Further Basics: Anorg. and analytical chemistry Physics Organic chemistry Mathematics Physical chemistry Computer applications Biochemistry Basics of biology: i Soft skills: Physiology Presentation, rhetoric Molecular and cell biology Seminars on specialist literature Genetics Communication (English) Microbiology Teaching skills Bachelor thesis in the 6th semester Academic degree: Bachelor of Science

9 Master's degree in advanced study e.g. in: Biochemistry Organic Chemistry Modern Biology Physical Chemistry Master's thesis in the 10th semester Academic degree: Master of Science

10 Please note Limited number of participants (approx. 30 Bachelor students) Modularization, final examination for each module Introduction of the credit point system: t 180 credit points Bachelor 120 credit points Master Horizontal change to pure biology or chemistry Change to / from other universities or abroad

11 Example: Biochemistry module Main lecture: Internship: Seminar: Elective lecture: Elective internship: Biochemistry I & amp II (2 SWS each) 2 2 weeks Biochemical topics and soft skills e.g. Structural biology e.g. NMR driving license Contents of the main lecture: Proteins, enzymes, lipids, biomembranes, carbohydrates, metabolism, nucleic acids, genetic engineering

12 Contents of the internship Gene Protein Structure Function Cloning & Expression Protein Purification & Enzyme Kinetics Spectroscopy & MD Simulation Antibacterial or hemolytic

13 Career prospects Bachelor: Basic knowledge Master: Extended knowledge Dr. rer. nat: specialization chemical industry biotech industry pharmaceutical industry i agro-industry & ecology, environment academic career

14 Topicality New specialist sections Chemical Biology in the GBM, GDCh, DECHEMA, DPhG in Dortmund new course in Chemical Biology, in Erlangen and W & uumlrzburg Molecular Medicine, in Heidelberg and L & uumlbeck Molecular Biotechnology Virtual Faculties Chemical Biology in the Internet (University of Bonn, Leipzig, Germany) Marburg, M & uumlnchen) AVENTIS initiates Lab Initiative Chemical Biology, large number of new textbooks, h trade journals, h if conferences

15 Textbooks Chemical Biology: Learning through Case Studies Herbert Waldmann, Petra Janning (2009) Essentials of Chemical Biology: Structure t and Dynamics of Biological il Macromolecules Andrew Miller, Julian Tanner (2008) Chemical Biology: From Small Molecules to Systems Biology and Drug Design Stuart L. Schreiber, Tarun M. Kapoor, G & uumlnther Wess (2007) Chemical Biology: Applications and Techniques Banafshe Larijani, Colin A. Rosser, Rudiger Woscholski (2006) Chemical Biology: A practical course Herbert Waldmann, Petra Janning (2004)

16 contact persons pa Prof. Marcus Elstner, chairman of the Dr. Birgid Langer, Chair for Biochemistry edu Prof. Anne S. Ulrich, Chair for Biochemistry Teaching / Chemical _ Biology / ChemBio00.html

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Non-mechanical digestion process

Non-mechanical disruption processes are used for cells that cannot be easily broken (for example yeasts). They are usually gentler than the mechanical processes.

  • In non-plant cells, variants of phenol-chloroform extraction such as Trizol digestion can be carried out by dissolving the membrane lipids out of the cell membrane, but in the presence of guanidinium thiocyanate it is denaturing. & # 911 & # 93
  • In yeast cells, autolysis is induced with toluene, which holes in the cell membrane and enzymatic lysis with zymolyase destroys the glucan cell wall, while Triton X-100 destroys the cell membrane.
  • In the case of Gram-positive bacteria, treatment with lysozyme destroys the peptidoglycan envelope, then the cell membrane can be loosened with Triton X-100.
  • In Gram-negative bacteria, the lipopolysaccharide of the outer cell membrane is dissolved by treatment with EDTA, then the peptidoglycan envelope is destroyed by treatment with lysozyme, then the cell membrane can be destroyed with Triton X-100.
  • In the case of Gram-negative bacteria, the membrane lipids can be saponified using alkaline lysis; due to the high pH value, this method is also denaturing.

Use

The protein foaming agents are considered to be the foaming agents with the best adhesive effect, but they can only be used as low-expansion foam. However, they are sensitive to high temperatures, which denature the proteins and thereby disintegrate. However, the finished foam has very good heat resistance. If only good flowability and film formation are required, FFFP foaming agents can also be applied unfoamed if necessary.

Protein foaming agent is hardly used by the public fire brigades these days, nor do they usually have it in stock. Protein foams are often used by plant fire departments.

The proportioning rate is usually between 3 & # 160% and 6 & # 160%.

Sub Surface

Fluorinated protein foams (FP and FFFP) are suitable for the "base injection" method (also called "sub surface") for fighting fires in mineral oil tanks: the finished foam is fed into the bottom of the tank and rises either freely through the liquid or through one Hose up to the surface. Due to the lipophobic effect of the fluorosurfactants, the foam is hardly loaded with fuel, as would be the case with non-fluorinated protein and multigrade foams. This enables a good extinguishing success to be achieved - otherwise adhering fire material would continue to burn on the foam surface and destroy the foam.


Protein purification. 2.1 Properties of proteins

1 Protein purification 2 Science has been studying the structure and function of proteins for more than two hundred years, when the French chemist Pierre J. Macquer used the term albumins to summarize all substances that exhibit the peculiar phenomenon of changing from liquid to solid when heated. These substances included chicken egg white, casein and the blood component globulin. As early as 1787, around the time of the French Revolution, the purification of coagulable, protein-like substances from plants was reported. In the early nineteenth century, many proteins such as albumin, fibrin or casein were purified and analyzed, and it soon became apparent that these compounds were structured in a considerably more complex manner than the other organic molecules known at the time. The word protein was probably coined by the Swedish chemist J & oumlns J. von Berzelius around 1838 and then published by the Dutch Gerardus J. Mulder together with a chemical formula that Mulder considered to be generally valid for proteinaceous substances at the time. Of course, the homogeneity and purity of these proteins, which were then purified, did not meet today's requirements, but they showed that individual proteins can be distinguished from one another. At that time, cleaning could only be successful because you could use simple steps: extraction for enrichment, acidification for precipitation and crystallization when a solution is simply left to stand. Hofmeister received the chicken album in crystalline form as early as 1889. Although Sumner was able to crystallize enzymatically active urease in 1926, the structure and structure of proteins remained in the dark until the middle of the twentieth century. It was only the development of powerful purification methods with which individual proteins can be isolated from complex mixtures, accompanied by a revolution in techniques for the analysis of the separated proteins, that we have made it possible to understand protein structures today. In this chapter these cleaning methods are described and it should be clear how they are used systematically and strategically. It is extremely difficult to look at the topic from a general perspective, as the physical and chemical properties of different proteins can differ immensely. However, this diversity is biologically necessary because proteins, the actual tools and building materials of a cell, have to perform a wide variety of functions. 2.1 Properties of proteins Size of proteins The size of proteins can vary greatly, from small polypeptides, such as insulin, which consists of 51 amino acids, to very large multifunctional proteins, for example apolipoprotein B, a cholesterol-transporting protein that consists of a Chain consists of over amino acids, with a molecular mass greater than daltons (500 kda). Many proteins consist of oligomers of the same or different protein chains and have molecular weights of up to several million Daltons. In general, it can be expected that the larger a protein, the more difficult it will be to isolate and purify. This is due to the analytical methods, which show very low efficiencies with large molecules. In Figure 2.1, the separation capacity of individual separation processes (the maximum number of analytes that can be separated from one another under optimal conditions) is plotted against the molecular weight. It can be seen that for small molecules such as amino acids or peptides, some chromium molar mass = molar mass (M) is incorrectly often referred to as molecular weight, not mass, but the quotient of the mass of a substance divided by the amount of substance of the substance. Unit: g / mol. Absolute molecular mass (m M) is the molar mass (M) of a molecule divided by the number of particles in one mole (Avogadro constant N A): m M = M / N A. Unit: g. Relative molecular mass (M r) is the molecular mass standardized to 1/12 of the mass of 12 C (dimensionless).

2 14 Part I: Protein Analysis 1D Electrophoresis 2.1 Separation Methods for Biomolecules. The separation capacity of individual separation methods (the maximum number of substances separated from one another in an analysis) is significantly different for different molecular weights of the substances. SEC exclusion chromatography HIC hydrophobic interaction chromatography IEC ion exchange chromatography RPC reversed phase chromatography CE capillary electrophoresis.Dalton (Da) is a non-SI-compliant unit of mass named after the English naturalist John Dalton (). A dalton is equal to the atomic mass unit (u = 1/12 of the mass of 12 C) and roughly corresponds to the mass of a hydrogen atom (1, g). The most common use in biochemistry is kda (kilodalton = Da). Proteome Analysis Chapter 41 Affinity Chromatography Section matographic methods are quite able to separate more than 50 analytes in a sample. In the area of ​​proteins it can be seen that of the chromatographic techniques only ion exchange chromatography is able to separate more complex mixtures reasonably efficiently and that in this molecular mass range the electrophoretic techniques are far more powerful. For this reason, proteome analysis (the analysis of all proteins in a cell), in which several thousand proteins have to be separated, is now practically exclusively based on electrophoretic processes (one and two-dimensional gel electrophoresis). The figure also shows that there are no efficient separation processes for large molecules, for example for protein complexes with molecular weights of more than 150 kda, or for organelles. However, the separation efficiency of a method is not always the relevant parameter that plays a role in cleaning. If selective cleaning steps are available, the importance of the separation capacity takes a back seat and the selectivity becomes the decisive factor. For example, an affinity purification based on the specific interaction of a certain substance with an affinity matrix, for example an immunoprecipitation or an antibody peraffinity chromatography, has a very poor separation capacity of 1, but an extremely high selectivity in a very complex mixture with which a protein can be extracted from a isolate in a single step. Since the most important purification techniques, electrophoresis and chromatography, require the analytes to be present in dissolved form, the solubility that the protein possesses in aqueous buffer media is another important parameter when planning a protein purification. Many intracellular proteins located in the cytosol (e.g. enzymes) are easily soluble, while proteins that have structure-forming functions, such as the proteins of the cytoskeleton or membrane proteins, are usually much more difficult to dissolve. The very hydrophobic, integral membrane proteins whose natural environment is lipid membranes and which aggregate and precipitate without solubilizers such as detergents are particularly difficult to handle in aqueous media. Available amount The amount available in the starting material plays a decisive role for the effort that has to be made for protein purification. A protein intended for purification may only be present in a few copies per cell (e.g. transcription factors) or in a few thousand copies (e.g. many receptors). Common protein -

3 2 Protein purification (e.g. enzymes) can make up percentages of the total protein in a cell. Over-expressed proteins are often present in significantly higher quantities (> 50%), as are some proteins in body fluids (e.g. albumin in plasma> 60%). Since the purification normally becomes much easier with increasing amounts of a protein, various sources of starting material should be examined for the content of the protein of interest, especially when isolating rare proteins. Acid / base properties Proteins have certain acidic or basic properties due to their amino acid composition, which is used for separation via ion exchange chromatography and electrophoresis. The net charge of a protein depends on the ph value of the surrounding solution and is positive at a low ph value, negative at a high ph value and zero at the isoelectric point at this ph value the positive and negative charges compensate each other. Biological activity The purification of a protein is often made more difficult by the fact that a certain protein can only be recognized and localized in the variety of other proteins on the basis of its biological activity. Therefore, in every phase of protein isolation, care must be taken to preserve this biological activity. It is usually based on a specific molecular and spatial structure. If it is destroyed, one speaks of denaturation, this is often irreversible. In practice, in order to avoid denaturation, the use of some cleaning processes must be ruled out from the outset. The biological activity is often differently stable under different environmental conditions. Too high or too low buffer concentrations, temperature extremes, contact with non-physiological surfaces such as glass or missing cofactors can change the biological characteristics of proteins. Some of these changes are reversible: even after denaturation and loss of activity, small proteins in particular are often able to renature under certain conditions, that is, to regain their biologically active form. In the case of larger proteins, this rarely works and often only with poor yield. The measurement of the biological, such as the enzymatic, activity gives the possibility of tracking the purification of a protein: with increasing purification steps, a higher specific activity is measured. In addition, the biological activity itself can be used to purify the protein. It is often associated with binding properties to other molecules, for example enzyme substrate or cofactor, receptor ligand, antibody antigen, etc. These very specific bindings are used to design affinity purifications (affinity chromatography) and are characterized by high levels under optimal conditions Enrichment factors and thus through a great efficiency that is hardly achievable in any other way. Enzymatic Activity Tests Chapter 4 Affinity Chromatography Section Stability If proteins are extracted from their biological environment, their stability is often noticeably impaired, as they are degraded by proteases (proteolytic enzymes) or almost always lead to an irreversible loss of biological aggregates, which leads to an irreversible loss of biological activity leads. For these reasons, protease inhibitors are often added in the first steps of protein isolation and cleaning is generally carried out quickly and at low temperatures. If you consider this variety of properties, it quickly becomes clear that protein purification cannot follow a schematic rule. For a successful isolation strategy, in addition to an understanding of the behavior of proteins in the various separation processes and a minimal knowledge of the solubility and charge properties of the protein to be purified, a clear idea of ​​the purpose for which the protein is to be purified is necessary. Purification Purpose The first steps in a purification process, the purity to be striven for and the analytics to be used are largely dependent on the intention with which a certain protein is to be purified. When isolating a protein for therapeutic purposes (e.g. insulin, growth hormones or anticoagulants), the requirements for purity are far higher than for a protein that is used in the laboratory for structural investigations. In many cases one only wants a protein for unambiguous identification or for the clarification of a few

4 16 Part I: Protein Analysis Proteome Analysis Chapter 41 isolate fewer amino acid sequence segments. A very small amount of protein is sufficient (usually in the microgram range). The sequence information can be used to identify the protein in protein databases or to produce oligonucleotide probes and isolate the protein's gene. This can then be expressed in a host organism in a much larger amount (up to a gram amount) than was present in the original source (heterologous expression). Many of the further investigations are then not carried out with the material from the natural source, but with the recombinant protein. Strategically new approaches to the analysis of biological issues, such as proteome analysis and subtractive approaches, require completely new types of sample preparation and protein isolation, since the quantitative relationships of the individual proteins must not be changed here. A great advantage of these new strategies is that it is no longer necessary to ensure that biological activity is maintained. Even if each protein purification is to be regarded as an individual case, some general rules and procedures can be found, especially for the first purification steps, which have already been used frequently in successful isolations and which should be discussed in detail below. 2.2 Protein localization and purification strategy The first step in any protein purification is to bring the desired protein into solution and to separate out all particulate and insoluble material. Figure 2.2 shows a scheme for different proteins. For the purification of a soluble extracellular protein, cells and other insoluble components have to be separated in order to obtain a homogeneous solution, which can then be subjected to the purification or analysis processes discussed in the following sections (precipitation, centrifugation, chromatography, etc.). Sources for extracellular proteins are, for example, culture debris from microorganisms, plant and animal cell culture media or also body fluids such as milk, blood, urine or cerebrospinal fluid. Usually extracellular proteins are present in solution in relatively low concentrations and require an efficient concentration as the next step. In order to isolate an intracellular protein, the cells must be disrupted in a manner that releases the soluble contents of the cell and leaves the protein of interest intact. The methods of cell disruption (cell disruption) differ primarily depending on the type of cell and the number of cells to be disrupted. Membrane proteins and other insoluble proteins Membrane-associated proteins are usually purified from the relevant membrane fraction after it has been isolated. In addition 2.2 purification scheme for various proteins. Depending on the localization and solubility of the proteins to be purified, various pre-purification steps have to be carried out before selective and highly efficient steps can follow.

5 2 Protein purification 17 peripheral membrane proteins that are only loosely bound to membranes are separated from the membrane by relatively mild conditions, e.g. high pH value, addition of EDTA or low concentrations of a non-ionic detergent, and can then often continue like Soluble proteins are treated. Integral membrane proteins, which aggregate outside their membrane via hydrophobic amino acid sequence regions and become insoluble, can only be isolated from the membrane with the help of high detergent concentrations; Structural proteins (e.g. elastin), which are sometimes also cross-linked via post-translational functional groups (post-translational modifications). The first and very efficient cleaning step here is the removal of all soluble proteins. Further steps are usually only possible under conditions that destroy the native structure of the protein. Further processing often takes place after the cross-linking on the denatured proteins has been broken up and using chaotropic reagents (e.g. urea) or detergents. Recombinant proteins A special situation arises in the production of recombinant proteins. A very simple purification results after the expression of recombinant proteins in inclusion bodies. These are dense aggregates of the recombinant product that are in a non-native state and are insoluble, be it because the protein concentration is too high, because the expressed protein cannot be correctly folded in the bacterial environment or because the formation of the (correct) Disulfide bridges in the reducing environment inside the bacterium is not possible. After a simple purification by differential centrifugation (Section 2.5.1), in which the other insoluble cell components are separated off, the recombinant protein is obtained practically pure, but it still has to be converted into the biologically active state by renaturation. If the expression of recombinant proteins does not lead to inclusion bodies, the protein is in a soluble state inside or outside the cell, depending on the vector used. Here the cleaning is very similar to the cleaning of natural proteins, only with the advantage that the protein to be isolated is already present in relatively large quantities. Recombinant proteins can be purified very easily by using specific marker structures (tags). Typical examples are the fusion proteins, in which at the DNA level the coding regions for a tag structure and for the desired protein are ligated and expressed as a protein. Such fusion proteins can be purified in a single step using specific affinity chromatography with antibodies against the tag structure. Examples of this are GST fusion proteins with antibodies against GST or biotinylated proteins via avidins. Another frequently used tag structure are polyhistidine residues, which are attached to the N- or C-terminal end of the protein chain and which can be easily isolated via immobilized metal affinity chromatography. Metal Affinity Chromatography Section Homogenization and Cell Disruption In order to be able to purify biological components from intact tissues, these complex cell assemblies have to be destroyed in a first step by homogenization. This creates a mixture of intact and broken cells, cell organelles, membrane fragments and also small chemical compounds that come from the cytoplasm and damaged subcellular compartments. Since the cellular components are transferred into an unphysiological environment, the homogenization medium should meet various basic requirements: protection of cells from osmotic bursting, protection from proteases, protection of biological activity (function), prevention of aggregation, as little interference with organs as possible biological analyzes and functional tests.

6 18 Part I: Protein analysis Table 2.1 Protease inhibitors Substance concentration Inhibitor of phenylmethylsulfonyl fluoride (PMSF) 0.1 1 mm aprotinin 0.01 0.3 μm serine proteases ε-amino-n-caprons ε 5 mm antipain leupeptin 70 μm 1 μm cysteine ​​proteases Pepstatin A 1 & mum aspartate proteases Ethylenediaminetetraacetic acid (EDTA) 0.5 1.5 mm Metalloproteases Usually this is done with isotonic buffers at a neutral pH value, to which a cocktail of protease inhibitors is often added (Tab. 2.1). If one wants to isolate intracellular organelles such as mitochondria, nuclei, microsomes etc. or intracellular proteins, the (still) intact cells have to be broken open. This is achieved by mechanically destroying the cell wall, during which frictional heat can arise and which should therefore be carried out with cooling as far as possible. The technical implementation of the digestion varies depending on the starting material and the localization of the desired target structure (Table 2.2). In the case of very sensitive cells (e.g. leukocytes, ciliates) it is often sufficient to repeatedly pipette the cell suspension or to press it through a sieve to allow digestion by means of black table 2.2 Biological starting materials and digestion methods Material Digestion method Comments Bacteria gram-positive enzymatic with lysozyme peptidoglycan cell wall EDTA / Tris Makes the cell wall permeable French press gram negative Cell grinder with glass beads Mechanical destruction of the cell wall Freeze-thawing Ultrasound for large quantities unsuitable due to local overheating Yeast autolysis with toluene French press several times, as inefficiently enzymatic inhibition of plants with glass beads Knife homogenizer high protease content in plants + dithiothreitol + phenol oxidase inactivators polyvinylpyrrolidone + protease inhibitors fibrous tissue grind in liquid nitrogen cold homogenization buffer do not phase Large tissue higher eukaryotes Grind, possibly after drying, cells that grow in suspension culture Osmolysis with hypotonic buffer Very sensitive cells Press through sieve Repeated pipetting of the suspension Add protease inhibitors Fibrous cells Chop up Dounce homogenizer Muscle tissue Chopping, meat grinder difficult to break down Enzymology, Vol. 182 Guide to Protein Purification, Academic Press 1990.)

7 2 Protein Purification 19 to achieve high shear forces. For the somewhat more stable animal cells, the shear forces are generated with a glass pestle in a glass tube (Dounce homogenizer). These methods are not suitable for plant and bacterial cells. Cells that do not have a cell wall and that are not associated in cell clusters (e.g. isolated blood cells) can be osmolytically disrupted by placing them in a hypotonic environment (e.g. in distilled water). The water penetrates the cells and makes them burst.In the case of cells with cell walls (bacteria, yeasts), the cell walls must be broken down enzymatically (e.g. with lysozyme) before osmolytic digestion is successful. This type of digestion is very gentle and is therefore particularly suitable for isolating cell nuclei and other organelles. For bacteria, repeated freezing and thawing is often used as a digestion method, whereby the change in the state of aggregation deforms the cell membranes in such a way that they break open and the intracellular content is released. Microorganisms and yeasts can be dried in a thin layer for two to three days at C, whereby the cell membrane is destroyed. The dried cells are ground in a mortar and can be stored at 4 C for a longer period of time. Soluble proteins can be brought back into solution in a few hours from the dry powder with an aqueous buffer. With cold, water-miscible organic solvents (acetone, 15 C, 10-fold volume) cells can be dehydrated quickly, whereby the lipids are extracted into the organic phase and the cell walls are destroyed. After centrifugation, the proteins remain in the precipitate, from which they can be recovered by extraction with aqueous solvents. In the case of stable cells such as plant cells, bacteria and yeasts, grinding with a mortar and pestle can be used to break down the cells, although larger organelles (chloroplasts) can also be damaged. The digestion is facilitated by adding an abrasive (sea sand, glass beads). For larger quantities, a knife homogenizer is suitable, in which the cell tissue is cut up by a rapidly rotating knife. This creates considerable heat, so that there should be an opportunity for cooling. For small objects such as bacteria and yeasts, the efficiency of the digestion is significantly improved by adding fine glass beads. Vibrating cell grinders are used for a relatively rough digestion of bacteria. These are lockable steel vessels in which the cells are vigorously shaken together with glass beads (diameter 0.1 0.5 mm). Here, too, the heat generated must be dissipated. Cell organelles can be damaged by this digestion method. Rapid changes in pressure break up cells and organelles very efficiently. For example, ultrasonic waves in the frequency range of khz are used to generate strong pressure changes in the suspension of a cell material via a metal rod. Since a lot of heat is released with this method too, only relatively small volumes and short sonication pulses of no more than ten seconds should be used. DNA is fragmented under these conditions. In another digestion method, which is particularly suitable for microorganisms, up to 50 ml of a cell suspension are pressed under pressure through a narrow opening (<1 mm), the cells being destroyed by the shear forces that occur (French press). Depending on the objective, the desired proteins are subjected to further purification steps in soluble form. For this purpose, the homogenate is usually roughly separated into different fractions using differential centrifugation methods (Section 2.5.1).

8 20 Part I: Protein Analysis 2.4 Precipitation The precipitation (precipitation) of proteins is one of the first techniques that was used for the purification of proteins (the salting out of proteins happened for the first time over 130 years ago!). The method is based on the interaction of precipitating agents with the proteins in solution. These agents can be relatively unspecific and precipitate practically all proteins from a solution, which is used in the first steps of a cleaning process to obtain the total proteins from a cell lysate. However, the precipitation can also be carried out in such a way that a fractionation of the constituents of a solution is possible. An example of this is the Kohn fractionation of plasma, which was worked out as early as 1946 and is still used today for the production of plasma protein on a large scale. Increasing amounts of cold ethanol are added to blood plasma and the respective precipitated protein is centrifuged off in fractions. With the exception of the precipitation of antigens with antibodies, the precipitation is not protein-specific and is therefore only used for a rough pre-purification of protein mixtures. Depending on the question and the starting material, the filling can be carried out under different conditions. In doing so, not only the efficiency of the filling itself, but also other aspects should always be taken into account: Is the biological activity impaired by the filling agent and the filling conditions? Under what conditions can the filler be removed? Determining the Concentration of Proteins Chapter 3 Salting Out The property of a salt to precipitate proteins is described in the so-called Hofmeister series (Fig. 2.3). The salts further to the left (so-called antichaotropic or kosmotropic) salts are particularly good and gentle filling agents. They increase hydrophobic effects in the solution and promote protein aggregations through hydrophobic interactions. The (chaotropic) salts on the right reduce hydrophobic effects and keep proteins in solution. The most common method of precipitating proteins is to salt them out by adding ammonium sulfate. Before the precipitation, the proteins should be present in a concentration of about 0.01-2%. Ammonium sulfate is particularly well suited, since in concentrations above 0.5 M it protects the biological activity of even sensitive proteins. It is easy to remove from the proteins again (dialysis, ion exchange) and, moreover, it is inexpensive, which is why it can also be used for fillings from larger volumes and thus already in the first cleaning steps. Ammonium sulfate is usually added in portions to the protein solution under controlled conditions (temperature, pH value), which enables fractional precipitation and thus an enrichment of the protein of interest. It should be noted that a complete filling can take a few hours! Ammonium sulfate precipitates should normally be centrifuged off tightly and well (100 g, see section 2.5). The only major disadvantage of ammonium sulfate concerns the precipitation of proteins, which require calcium for their activity / structure, since calcium sulfate is practically insoluble and is thus removed from the proteins. These proteins must therefore be filled with other salts (e.g. acetates). Filling with organic solvents It has been known for over a hundred years that proteins can be filled with cold acetone or short-chain alcohols (mainly ethanol). Long-chain alcohols (larger than C 5) are not soluble enough in water and cannot be used for filling. No general rules can be given for the choice of the organic filler or the optimal temperature. Ethanol has antichaotropic chaotropic cations: NH 4 + K + Na + C (NH 2) 3 + (guanidine) 2.3 The Hofmeister series. Anions: PO 3 4 SO 2 CH 3 COO 4 Cl Br NO 2 ClO 3 I SCN

9 2 Protein purification 21 has proven particularly useful for the precipitation of plasma proteins. For protein solutions that still contain lipids, acetone is often used, since in addition to the precipitation of the proteins, the lipids are extracted at the same time. In order to avoid too high local concentrations of the organic solvent, which can lead to the denaturation of the proteins, the solvent should be added slowly. Good cooling and slow addition are also useful, as adding the organic solvent (e.g. ethanol to water) can generate heat that leads to undesired denaturation. The precipitate is pelleted by centrifugation (see below) and taken up again in aqueous buffers. A frequently used protocol for acetone precipitation adds a 5-fold volume excess of 20 C cold acetone to the protein solution and incubates overnight at 20 C. It is then centrifuged for 30 min at g. This precipitation usually gives excellent results even for very small amounts of protein. The yield of the precipitation must be checked with analytical methods (SDS gel electrophoresis, activity tests, etc.). SDS gel electrophoresis Section Enzymatic activity tests Chapter 4 Filling with trichloroacetic acid A frequently used method to precipitate proteins from solutions is filling with ten percent trichloroacetic acid, whereby a final concentration of 3 4% should be achieved. After centrifugation, the precipitate is resuspended in the desired buffer and used further, whereby the pH of the solution should be checked. This method denatures the proteins and is therefore mainly used for concentration for gel electrophoresis or before enzymatic cleavage. The minimum sample concentration should be 5 & mug ml 1. Filling of nucleic acids Protein solutions from cell disruptions, especially from bacteria and yeasts, contain a large proportion of nucleic acids (DNA and RNA), which can interfere with protein purification and therefore usually have to be separated. Since nucleic acids are highly negatively charged polyanions, they can be filled with strongly basic substances (e.g. polyamines, polyethyleneimines or anion exchange resins) or very basic proteins (protamines). By optimizing the precipitation and washing conditions, it must be avoided that proteins of interest are also filled by the precipitation reagent or in complex with the nucleic acids (e.g. histones, ribosomes). 2.5 Centrifugation Centrifugation is not only one of the most common techniques for separating insoluble components, but also for cell fractionation and the isolation of cell organelles. It is based on the movement of particles in a liquid medium by centrifugal forces. The central component of a centrifuge is the rotor, which is used to hold the sample beakers and is driven by a motor at high speed. There are different designs of the rotors, such as fixed-angle rotors, vertical or vibrating bucket rotors (Fig. 2.4), which are available in different sizes and materials. They allow separations from a few microliters to a few liters and can be operated with different, adjustable rotational speeds depending on the task at hand. Coolable centrifuges are mostly used for working with biological materials. High-speed centrifuges, the ultracentrifuges, are always operated connected to a vacuum system in order to avoid the frictional heat that occurs at high speeds as a result of air resistance. When operating centrifuges, certain safety measures must be observed, above all the opposing sample vessels must be well balanced in order to avoid any imbalance that could destroy the centrifuge. Basics The physical principle of centrifugation is a separation according to size and density. On a particle that is moved around an axis of rotation with constant angular velocity ω,

10 22 Part I: Protein analysis Fixed-angle rotor A B C D E Vertical rotor Swing-out rotor 2.4 Rotors for centrifugation. Fixed-angle rotor, vertical rotor and swing-bucket rotor when loaded (A) under centrifugation conditions at the beginning of the separation (B) during the separation (C) when braking (D) and after the end of the centrifugation (E). Fractions containing protein are shown in red. a centrifugal force acts, which accelerates the particle outward. The acceleration B depends on the angular velocity ω and the distance r from the axis of rotation: B = ω 2 r (2.1) The acceleration is related to the acceleration due to gravity g (981 cm s 2) and is expressed as a relative centrifugal acceleration RZB in multiples of the acceleration due to gravity (g ) specified: 2r RZB = 981 (2.2) The relationship between the angular speed and the rotational speed in rotations per min (rpm) is given by:

11 2 Protein purification 23 = rpm 30 (2.3) which results from substitution: RCF = 1, r (rpm) 2 (2.4) It must be taken into account that normally during centrifugation the distance of the particles from the axis of rotation and thus also the RCF change. The mean value is therefore often used for conversions. The sedimentation speed of spherical particles in a viscous liquid is described by the adjacent Stokes equation. Here v is the sedimentation velocity, g is the relative centrifugal acceleration, d is the diameter of the particle, & rho p and & rho m are the density of the particle or the liquid and η is the viscosity of the medium. The sedimentation speed increases with the square of the particle diameter and the difference in density between particles and medium and decreases with the viscosity of the liquid. If the sedimentation takes place in a medium such as 0.25 M sucrose, which is less dense than all particles and also has a low viscosity, the diameter of the particles is the dominant factor for the sedimentation rate. The sedimentation coefficient s is the sedimentation speed under geometrically given conditions of the centrifugal field. It is given in Svedberg units (S). Stokes equation: d = 2 () g p 18 m (2.5) v s = r 2 (2.6) 1 S corresponds to seconds. Various biological molecules lie in this order of magnitude. The Svedberg unit of a biomolecule is sometimes included in its name (e.g. 18S rRNA), which then allows a conclusion about the size of the particle. Table 2.3 shows the size and centrifugation conditions for cleaning cells and some cellular compartments. A good overview is also given by the representation of the particles in a density / sedimentation coefficient diagram (Fig. 2.5) or in a density / g-value diagram. The various techniques of centrifugation can easily be understood from Stokes' equation. Density (in g / ml) Sedimentation coefficient (in S) 2.5 Density and sedimentation coefficient of some cell compartments. The figure shows the distribution of different cell components with regard to their density and their sedimentation coefficient.



Comments:

  1. Kisho

    Other variant is possible also

  2. Odhran

    Why are there so few comments on such a good posting? :)



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