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Buffers and Buffering Theory How to Prepare a Buffer Common Buffers Common Buffer Preparations Buffer Theory Most biological systems will function only within a quite narrow range of conditions, and their activity can vary widely within that range. The acidity, or free proton concentration, of the environment is an important parameter. To prevent the proton concentration of a solution from changing, compounds can be added to a solution that "buffer" or minimize such changes. A compound will act as a proton concentration buffer if it limits changes in proton concentration by binding protons when the proton concentration of the solution increases and releasing bound protons when the proton concentration decreases (Eqn 1). + H+ + Buffer (Eqn 1) Unfortunately, any one compound will be effective as a buffer only for a limited range of proton concentrations, and so the first step in preparing a buffer is deciding which buffer + to use. ) and half in the base form (Buffer), i.e., when half of its proton binding sites are filled, a buffer should be chosen that will be about half filled at the proton concentration desired. The affinity of a compounds for protons is often expressed as its acid dissociation constant (Ka), defined in Eqn 2. This is convenient because, as can be seen from Eqn 2, the value of the acid dissociation constant is equivalent to the proton concentration of a solution at which the compound will have half of its proton binding sites filled, (Eqn 2) If the proton concentration and the acid dissociation constant are both expressed as their negative log, Eqn 2 becomes; (Eqn 3) and if, for reasons largely historical, the negative log operator is called "p", the expression becomes; (Eqn 4) rearranging Eqn 4 and inserting the values for the pH and pKa; (Eqn 5) Further rearranging of Eqn 5 gives Eqn 6; (Eqn 6) After choosing a buffer, the next step is to decide its concentration. The buffer concentration must be sufficient to maintain the pH within acceptable limits with the changes in proton concentration expected to occur. For biological systems, this generally means that the total buffer concentration ([Buffer]total) is within a range from 1 mM to 200 mM. Knowing that the total buffer concentration is equal to the sum of the concentrations of its forms; (Eqn 7) by substituting for the [Buffer] term in Eqn 7 with its equivalent from Eqn 6, we see; (Eqn 8) and by rearranging, (Eqn 9) So for any pH we choose, by finding a buffer whose pKa is within around one pH unit of that pH, Common buffers Molecular Weights of Different Forms Buffer pKa Neutral form HCl salt Na+ salt H3PO4 / NaH2PO4 (pKa1) 2.12 98.0 - 120.0 Glycine (pKa1) 2.34 75.07 111.5 - Citric acid (pKa1) 3.13 Acetic acid 4.75 60.05 - 82.0 Citric acid (pKa2) 4.76 192.1 - 294.1 MES 6.15 195.2 - 217.2 Cacodylic acid 6.27 H2CO3 / NaHCO3 (pKa1) 6.37 62.01 - 84.01 Citric acid (pKa3) 6.40 Bis-Tris 6.50 209.2 245.7 - ADA 6.60 190.2 - 212.1 Bis-Tris Propane (pKa1) 6.80 282.4 318.9 - PIPES 6.80 302.4 - 325.3 ACES 6.90 182.2 218.7 - Imidazole 7.00 68.1 104.5 - BES 7.15 213.2 - 235.2 MOPS 7.20 209.3 - 231.2 NaH2PO4 / Na2HPO4 (pKa2) 7.21 120.0* - 142.0* TES 7.50 229.3 - 251.2 HEPES 7.55 238.3 - 260.3 HEPPSO 7.80 268.3 - 290.3 Triethanolamine 7.80 149.2 185.7 - Tricine 8.10 179.2 215.6 - Tris 8.10 121.1 157.6 - Glycine amide 8.20 74.1 110.6 - Bicine 8.35 163.2 199.7 - Glycylglycine (pKa2) 8.40 132.1 168.6 - TAPS 8.40 243.2 - 265.3 Bis-Tris Propane (pKa2) 9.00 ? 355.3 - Boric acid (H3BO3 / Na2B4O7) 9.24 61.8 - 201.24 CHES 9.50 207.3 - 229.3 Glycine (pKa2) 9.60 75.07 - 96.1 NaHCO3 / Na2CO3 (pKa2) 10.25 84.01 - 105.99 CAPS 10.40 221.3 - 243.3 Piperidine 11.12 Na2HPO4 / Na3PO4 (pKa3) 12.67 142.0* - 164.0* *The anhydrous molecular weight is reported in the table. Actual molecular weight will depend on the degree of hydration. PROTEIN PURIFICATION AND ANALYTICAL TECHNIQUES -Studies on pure proteins are essential for understanding structural and functional properties of proteins. -Method for each protein worked out by trial and error on small samples goal: separate the protein you want from other proteins and small molecules mild conditions to avoid denaturation (usually low temperature, 0–4° C, and avoiding extremes of pH) need detection method (e.g. biological activity, or spectroscopy) usually use several purification methods, one after another start with (mixture of) proteins in buffered solution, e.g. extract of proteins from cells that have been lysed (broken open) Source of Protein In order to purify a protein you need a source: could be blood or some other biological fluid, but most often whole cells, usually a specific type (liver, muscle, yeast, bacteria, etc.) Cells must be broken open (lysed, e.g., by osmotic shock or by mechanical disruption such as with a "French press" or a tissue homogenizer) to disrupt cell membranes to release proteins in soluble form without damaging the protein. Membrane-bound proteins can also be purified, but different approaches are required. Initial fractionation of homogenate usually by differential centrifugation --> several fractions (successive pellets, supernatants) of decreasing density, each with lots of proteins assay each fraction to find which fraction contains most of the protein of interest, and fractionate that further by more discriminating methods. CHROMATOGRAPHIC TECHNIQUES Column Chromatography o Invention of column chromatography a critical event in biochemistry, because it was the basis for development of procedures for obtaining pure proteins. o Different kinds of chromatographic separations based on one of the following: size of protein (molecular sieve chromatography = gel filtration = size exclusion chromatography), or net charge of protein (ion exchange chromatography), or specific ligand binding properties of protein (affinity chromatography) o o o o o In column chromatography a solid phase ("matrix", "resin", generally some kind of polymer, often a polysaccharide)(see below) is placed in a glass tube, the column. terminology: adsorbent: solid material/matrix, a "stationary phase" that some molecules bind to (adsorb to) elution: the process of washing something off an adsorbent (with an eluting buffer; the solution coming off the column is the eluate.) Protein mixture is passed into the column. Either due to molecular size differences or different binding affinities for column matrix, some proteins are retained longer on the column (e.g., some bind more tightly than others) The properties of some different types of column packing materials (for separations based on molecular size, charge or specific ligand binding) are described below. Gel Filtration Chromatography (also called "Molecular Sieve" chromatography, or "Size Exclusion" chromatography) o stationary phase (column matrix) = "beads" of a polysaccharide material that separates proteins based on size and shape. Different column packing materials (hydrated, porous beads of carbohydrate polymer (e.g. dextran or agarose) or polyacrylamide) available, with wide range of molecular exclusion limits, for separating proteins of all sizes. Solution of mixture of proteins, small molecules, etc. "filters" through the beads: Large molecules can’t get into the smaller pores in the beads and move more rapidly through the column, emerging (eluting) sooner. Smaller molecules and ions can enter all the pores in the beads with the buffer, and thus have more space to "explore" on their way down the column, and elute later. o For any particular column dimensions and material, volume of buffer required to elute a specific protein depends mostly on molecular weight of the protein (but shape plays an important role also -- separation is really based on differences in hydrodynamic volume). Thus, one can separate proteins by size. o Size exclusion chromatography o This animation illustrates how size exclusion chromatography works. Note how the small red spheres p into the channels in the beads, whereas the large blue spheres do not. Thus, the small spheres have longer "distance" to transverse than large spheres to get to bottom of column, which means that a larger volume of solvent must pass throu the column before the red spheres eluted. o The following plot of relative amo of the large solute (blue) and of th smaller solute (red) goes with the animation. Larger solutes elute EARLIER, smaller solutes LATER, from a si exclusion column. o o o calibrate the column: determine elution volumes of proteins with known molecular weights construct a calibration curve relating(known) molecular weight to (measured) elution volume specifically for that column. o o Such a calibration curve can then be used to estimate the molecular weight of an unknown protein. Ion Exchange Chromatography o o o o o o o Ion exchange resins have charged groups covalently attached to the stationary phase (adsorbent, matrix), either positive or negative. Obviously, if ionizable groups are weak acids or bases, the pH of the buffer determines the charge state of the matrix. Proteins bind to the matrix by electrostatic interactions. Strength of these interactions depends on net charge on the protein (a function of buffer pH and the nature of the ionizable groups on that protein, reflected in the pI of the protein), and salt concentration of the buffer (high salt concentrations reduce the interaction and can be used to elute the proteins by competing with the protein groups for binding to the charged groups on the matrix). The higher the net charge on the protein at the pH of the environment on the column, the more tightly it sticks to an oppositely charged matrix, and the higher the salt concentration required to elute it from the column. The further the "working pH" is from the isoelectric point (pI) of a protein, the greater the net charge on the protein, and the more tightly it will stick to an ion exchanger of opposite charge. By proper choice of eluting buffer (often a gradient with increasing salt concentation, or changing the pH), specific proteins can be eluted from the column and separated from other proteins in the mixture. Ion Exchange Chromatography Example in figure is cation exchange chromatography -- column packing beads have covalently attached negatively charged groups Negatively charged solutes move down the column more or less without sticking, so they elute first. Positively charged solutes bind, and the higher the positive charge on a molecule, the tighter it binds, so the later it elutes. Example: Suppose you have 5 different proteins, with relative isoelectric points as indicated on the pH scale below. pH SCALE (working pH = 6.5 for these examples): 0 -----------pI#5----------pI#4------- 6.5 --------pI#1----------pI#2----------pI#3------------- 14 Suppose that your column is equilibrated and being eluted at pH 6.5 (the working pH is 6.5), by washing the column with a gradient of buffer of increasing salt concentration. Protein What's the RELATIVE net charge at pH 6.5? 1 2 3 4 5 o o ANION EXCHANGE Anion exchange matrix has + charged groups (e.g., DEAE (diethylaminoethyl) groups). A molecule with a net + charge won't stick, so will wash on through and elute before anything else (proteins 1, 2 and 3 in the current example). Molecules with net - charge will elute in the order of their pI values, because of differences in net charge: the most charged one (the one whose pI is furthest from the working pH) sticks the most tightly (elutes last). See elution profile below. CATION EXCHANGE Cation exchange matrix has – charged groups (e.g., carboxymethyl (CM) groups). A molecule with a net - charge won't stick, so will wash on through and elute before anything else (proteins 4 and 5 in the current example). Molecules with net + charge will elute in the order of their pI values, because of differences in net charge: the most + charged one (the one whose pI is furthest from the working pH) sticks the most tightly (elutes last). See elution profile below. Label the peaks below with #1, #2, #3, #4, and/or #5, based on the expected order of elution of proteins #1-5 from a cation exchange column, or from an anion exchange column, at pH 6.5. Affinity Chromatography o a more specific adsorbent in which a ligand specifically recognized by the protein of interest is covalently attached to the column material o When a mixture of proteins is passed through the column, only those few that bind strongly to the ligand stick, while the others pass through the column. o Protein of interest is eluted with a buffer containing the free ligand, which competes with the column ligand to bind to the protein, and protein washes off (with bound ligand). o Affinity Chromatography o some variations: immunoaffinity chromatography: an antibody specific for a protein is immobilized on the column and used to affinity purify the specific protein. "polyHis tags" on recombinant proteins: a sequence of His residues is placed (by genetic engineering of a cloned gene) at the Cterminus of a specific recombinant protein to be produced in vivo, and that protein can be purified on a column with Ni2+ ions (or Cu2+ or Co2+ or Zn2+) held in chelated form on an affinity column; the His imidazole groups on the end of the recombinant protein bind with high affinity, but other proteins don't stick. The recombinant protein can then be eluted with an imidazole buffer. ELECTROPHORESIS Electrophoresis In an electric field, a protein or other charged macromolecule will move with a velocity that depends directly on the charge on the macromolecule and inversely on its size and shape. pH obviously important in determining net charge Gel electrophoresis is carried out in some supporting media, usually polyacrylamide or agarose, with pores of big enough to allow passage of the macromolecule. Electric field is applied, and molecules move toward electrode opposite to their net charge, but they’re slowed down ("friction") by the gel o larger or more elongated shaped molecules move the most slowly o smaller, most compact molecules move faster. The proteins in the gel are easily stained for detection purposes. Because the net charge on a protein and its molecular weight are characteristic properties of a protein, electrophoresis is a powerful method for characterizing degree of purity of a protein preparation, but can also be used for purification of small amounts of proteins. Discontinuous Gel Electrophoresis ("disc gel electrophoresis") 3 experimental variations to ordinary gel electrophoresis: 1) 2 gel layers, a lower or resolving gel and an upper or stacking gel 2) The buffers used to prepare the 2 gel layers are of different ionic strengths and pH 3) The stacking gel (upper gel) has a lower acrylamide concentration, so its pore sizes are larger. These variations cause formation of highly concentrated bands of sample in stacking gel and greater resolution of sample components in lower (resolving) gel. o Stacking and separation in a discontinuous gel: o Buffer compositions control stacking and separation: o Glycine equilibria: o Formation of an ion front: o The voltage gradient sharpens the ion boundary: o What happens to the proteins? Proteins have mobilities between those of Gly and Cl-. o In separating gel, o Glycine mobility increases, becomes greater than protein mobility, but still slower than Cl-. o Protein sample, now in a narrow band, encounters both the increase in pH and decrease in pore size. Increase in pH would tend to increase electrophoretic mobility, but smaller pores decrease mobility. Relative rate of movement of ions in lower gel is chloride > glycinate > protein. Proteins separate based on charge/mass ratio and on size and shape parameters. SDS-PAGE (Sodium Dodecyl Sulfate-PolyAcrylamide Gel Electrophoresis) a variant of electrophoresis in which the buffers contain SDS, a detergent that binds to proteins. Sodium dodecyl sulfate, SDS CH3(CH2)10CH2-SO4-, Na+ Most proteins bind SDS at a constant ratio of about 1.4 g SDS/g protein, i.e., about 1 SDS for every 2 amino acid residues, unfolding the proteins Sample treatment before running gel included bmercaptoethanol reduction (so no disulfide bonds left) and heating to ensure complete unfolding and complete separation of different polypeptide chains large negative charge resulting from the bound SDS masks the native charge on the protein, so that all proteins have essentially the same charge to mass ratio (very negative), and same shape ("random coil") so rate of movement in the electric field (toward the + pole because of – charge on sulfates) depends only on the molecular weight of individual polypeptide chains (which travel separately) Protein mobility INVERSELY proportional to the log of the MASS of individual polypeptide chains, and net charge of protein itself hardly makes any difference at all. SDS-PAGE often used to o o ESTIMATE PURITY (number of stained or radioactive or fluorescent bands on the gel) and to o o DETERMINE MOLECULAR WEIGHT of INDIVIDUAL POLYPEPTIDE SUBUNITS of proteins (using standards of known polypeptide chain mass) o o Purification of small amounts of polypeptide for sequence analysis Estimating protein molecular weight from SDS gel electrophoresis a) Diagram of a stained SDS gel: standards of known molecular weight (lane 1) and pure protein of unknown M.W. in lane 2 b) "standard curve" (calibration) to relate M.W. to mobility on THIS GEL o o o o Elution profile from an anion exchange resin (binds negatively charged proteins) Proteins were eluted by increasing NaCl concentration in the eluting buffer. Total protein was measured by determining the absorbance at 280 nm. In order to "assay" (identify) the fatty acid-binding proteins, they were o o o labeled by binding radioactive fatty acids (CPM=counts per minute - gray shading). Purity of each peak was assessed using SDS-PAGE (insert/overlay). There are two nearly pure proteins that bind fatty acids. The two proteins were obtained in pure form following one additional step (not shown). Western blotting is an immunological technique for detecting a specific protein in a mixture separated by gel electrophoresis, using antibodies specific for that protein to detect it on the gel. Isoelectric Focusing separation based on differences in ISOELECTRIC POINT (pI) (so based on CHARGE DIFFERENCES) Isoelectric Focusing pH gradient set up first (using purchased mixture of ampholytes, different molecules designed to have range of pIs, which are first electrophoresed on the gel to form the pH gradient) Mixture of molecules (proteins) is then applied, electric field is turned on, and each protein moves to the position (pH) at which its net charge is zero, i.e., its pI. Two-dimensional Electrophoresis isoelectric focusing in first dimension, followed by SDSPAGE at 90o to that (2nd dimension) Ultracentrifugation Molecular Weight and Shape = fundamental physical properties of a protein. Estimates of molecular weight can be obtained using SDS-PAGE or gel filtration, as described above. One very useful technique for measuring molecular weight and shape is centrifugation. A particle that's subjected to a centrifugal field by being spun in a centrifuge is subjected to a force, where m is the mass of the particle, r is the distance of the particle from the center of rotation, and w is the angular velocity. = buoyancy factor, which accounts for the fact that particle is buoyed up by the surrounding solvent of density r (g/ml). is the specific volume of the particle (ml/g) (= 1/density of the particle). If = r then the particle will not move. Movement of particle through the solvent is resisted by a frictional coefficient, f, that depends on the shape of the particle. Frictional coefficient is an important factor in any transport process, such as centrifugation or gel filtration. A spherical particle has f = 1.0, whereas a cigar-shaped or cylindrically-shaped particle will have f > 1.0. Movement of any particle under the influence of a centrifugal field is characterized by its sedimentation coefficient, S, which is directly proportional to its molecular mass, M, and inversely proportional to f. , where N is Avogadro's number. Ultracentrifugation is used in two ways to characterize proteins: In sedimentation equilibrium experiments, the centrifuge is operated at a relative low speed so that the forces of sedimentation and diffusion balance and the protein distributes in the centrifuge cell in a manner proportional to its molecular weight. In sedimentation velocity experiments, the centrifuge is operated at maximal speed, which causes the protein to sediment to the bottom of the tube. The rate at which the boundary moves gives S, which when combined with M gives f, a measure of the shape of the protein. Spectroscopic Methods Ultraviolet-visible spectroscopy (uv-vis frequency range) o The principles of absorption spectroscopy. (a) Electronic and vibrational transitions in a diatomic molecule. (b) The electromagnetic spectrum. Introduction Many compounds absorb ultraviolet (UV) or visible (Vis.) light. The diagram below shows a beam of monochromatic radiation of radiant power P0, directed at a sample solution. Absorption takes place and the beam of radiation leaving the sample has radiant power P. The amount of radiation absorbed may be measured in a number of ways: Transmittance, T = P / P0 % Transmittance, %T = 100 T Absorbance, A = log10 P0 /P A = log10 1 / T A = log10 100 / %T A = 2 - log10 %T The last equation, A = 2 - log10 %T , is worth remembering because it allows you to easily calculate absorbance from percentage transmittance data. The relationship between absorbance and transmittance is illustrated in the following diagram: So, if all the light passes through a solution without any absorption, then absorbance is zero, and percent transmittance is 100%. If all the light is absorbed, then percent transmittance is zero, and absorption is infinite. The Beer-Lambert Law Now let us look at the Beer-Lambert law and explore it's significance. This is important because people who use the law often don't understand it - even though the equation representing the law is so straightforward: A=αlc Where A is absorbance (no units, since A = log10 P0 / P ) α is the molar absorbtivity with units of L mol-1 cm-1 l is the path length of the sample - that is, the path length of the cuvette in which the sample is contained. We will express this measurement in centimetres. c is the concentration of the compound in solution, expressed in mol L-1 The reason why we prefer to express the law with this equation is because absorbance is directly proportional to the other parameters, as long as the law is obeyed. We are not going to deal with deviations from the law. Let's have a look at a few questions... Question : Why do we prefer to express the Beer-Lambert law using absorbance as a measure of the absorption rather than %T ? Answer : To begin, let's think about the equations... A=αlc %T = 100 P/P0 Now, suppose we have a solution of copper sulphate (which appears blue because it has an absorption maximum at 600 nm). We look at the way in which the intensity of the light (radiant power) changes as it passes through the solution in a 1 cm cuvette. We will look at the reduction every 0.2 cm as shown in the diagram below. The Law says that the fraction of the light absorbed by each layer of solution is the same. For our illustration, we will suppose that this fraction is 0.5 for each 0.2 cm "layer" and calculate the following data: Path length / cm 0 0.2 0.4 0.6 0.8 1.0 %T 100 50 25 12.5 6.25 3.125 Absorbance 0 0.3 0.6 0.9 1.2 1.5 A = tells us that absorbance depends on the total quantity of the absorbing compound in the light path through the cuvette. If we plot absorbance against concentration, we get a straight line passing through the origin (0,0). Note that the Law is not obeyed at high concentrations. This deviation from the Law is not dealt with here. The linear relationship between concentration and absorbance is both simple and straightforward, which is why we prefer to express the Beer-Lambert law using absorbance as a measure of the absorption rather than %T. Theoretical principles Introduction Many molecules absorb ultraviolet or visible light. The absorbance of a solution increases as attenuation of the beam increases. Absorbance is directly proportional to the path length, b, and the concentration, c, of the absorbing species. Beer's Law states that A = αlc where α is a constant of proportionality, called the absorbtivity. Different molecules absorb radiation of different wavelengths. An absorption spectrum will show a number of absorption bands corresponding to structural groups within the molecule. For example, the absorption that is observed in the UV region for the carbonyl group in acetone is of the same wavelength as the absorption from the carbonyl group in diethyl ketone. Electronic transitions 1. The absorption of UV or visible radiation corresponds to the excitation of outer electrons. When an atom or molecule absorbs energy, electrons are promoted from their ground state to an excited state. In a molecule, the atoms can rotate and vibrate with respect to each other. These vibrations and rotations also have discrete energy levels, which can be considered as being packed on top of each electronic level. Absorbing species Absorption of ultraviolet and visible radiation in organic molecules is restricted to certain functional groups (chromophores) that contain valence electrons of low excitation energy. The spectrum of a molecule containing these chromophores is complex. This is because the superposition of rotational and vibrational transitions on the electronic transitions gives a combination of overlapping lines. This appears as a continuous absorption band. Transitions FLUORESCENCE SPECTROSCOPY o : Fluorescence. (a) The principle of fluorescence. (b) Absorption and fluorescence emission spectra of tyrosine. In most cases, molecules raised to an excited electronic state by absorption of radiant energy return to ground state by radiationless transfer of the excitation energy to the surrounding molecules in the form of heat. o o o o o o o Sometimes an excited-state molecule will lose only part of its energy by transfer (yellow arrow below), and will re-radiate the larger part as light (green arrow below). That emitted light is fluorescence. o Since energy of emitted light is always lower than energy of absorbed light, fluorescence emission is always at a longer wavelength than wavelength of the exciting (absorbed) light (Fig. 6A.4(b) below). o terminology Fluorophore = a molecule that absorbs light but then returns to the ground state by emitting some of the light as a photon rather than losing all the energy as heat o wavelength and intensity of emitted light both very sensitive to the environment of the fluorophore (e.g., hydrophobic vs. aqueous environment can shift emission spectrum) o measurements very sensitive so can detect small amounts of protein or other fluorophore o Fluorophores in proteins Trp (maximum wavelength of fluorescence emission (lmax ~340 nm) is the strongest source of intrinsic fluorescence in proteins without fluorescent prosthetic groups, but tyrosine also contributes to intrinsic fluorescence (see Fig. 6a.4(b) above. o o Some ligands and prosthetic groups are fluorescent, e.g. the chromophore in green fluorescent protein USES of fluorescence spectroscopy -- examples: detect conformational changes e.g. during protein folding (environment of chromophore affects lmax and intensity of Trp fluorescence; the more hydrophobic the environment, e.g. as Trp residues get buried in the interior of the protein during folding, the shorter the wavelength of maximum fluorescence emission) detect and quantitate ligand binding NUCLEAR MAGNETIC RESONANCE (NMR) SPECTROSCOPY (microwave, i.e. radio, frequency range) o Basis: A spinning charged particle (in this case, a nucleus) behaves as a magnet, and can interact with an externally imposed magnetic field such that absorbance of electromagnetic radiation of appropriate energy (in the microwave, i.e. radio, frequency range) can flip the spin. o Nuclei used in biochemical studies include 1H (proton NMR), 2H, 13C, 14 N, 17O, 31P, and 19F (in 19F-Tyr). o To get an NMR spectrum (in ppm), you "sit" on a magnetic field strength and change the radio frequency to get resonance. The type of nucleus you're observing, but also the molecular structural environment of the nucleus (including its solvent and surroundings in 3 dimensional space) affect the width and position (position = "chemical shift") of the NMR signal (peak) for that nucleus. Interaction with a nearby nucleus within the molecule can cause spin coupling, which is seen as splitting of the NMR signal (double peak). Altering the spin on one nucleus can affect the spin on a nearby nucleus (< ~5Å away), and for small proteins it is possible by NMR to do enough distance measurements between nuclei within the tertiary structure to determine the entire 3-dimensional structure. o USES of NMR: complete 3-D structure of small proteins in solution (< 25,000 daltons) conformational changes (e.g., during folding) determination of pKa of an ionizable group, e.g. His follow ligand binding dynamics (motion in solution), e.g. Tyr and Phe ring flips o ABSORPTION SPECTROSCOPY terminology: Absorption = transfer of energy from a photon (light) to a molecule Chromophore = a molecule or a group on a molecule that absorbs light Chromophores in proteins include the peptide bond (maximum wavelength of absorbed light, lmax, ~220 nm, "far" uv) aromatic a.a. residues (lmax ~280 nm for Trp, "near" uv; aromatics also absorb ~220 nm) some prosthetic groups (tightly bound non-amino acid components in proteins, e.g., the heme in hemoglobin and myoglobin is red -- it absorbs visible light.) USES of absorbance spectroscopy: determine concentration (Beer's Law) conformational changes (environment of chromophore affects lmax and absorbance) detect and quantitate ligand binding (e.g., O2 binding to hemoglobin changes absorbance of the heme Theoretical Principles Introduction The term "infra red" covers the range of the electromagnetic spectrum between 0.78 and "wavenumbers", which have the units cm-1. wavenumber = 1 / wavelength in centimeters It is useful to divide the infra red region into three sections; near, mid and far infra red; Region Wavelength range) Wavenumber range (cm-1) Near 0.78 - 2.5 12800 - 4000 Middle 2.5 - 50 4000 - 200 Far 50 -1000 200 - 10 The most useful I.R. region lies between 4000 - 670cm-1. Theory of infra red absorption IR radiation does not have enough energy to induce electronic transitions as seen with UV. Absorption of IR is restricted to compounds with small energy differences in the possible vibrational and rotational states. For a molecule to absorb IR, the vibrations or rotations within a molecule must cause a net change in the dipole moment of the molecule. The alternating electrical field of the radiation (remember that electromagnetic radation consists of an oscillating electrical field and an oscillating magnetic field, perpendicular to each other) interacts with fluctuations in the dipole moment of the molecule. If the frequency of the radiation matches the vibrational frequency of the molecule then radiation will be absorbed, causing a change in the amplitude of molecular vibration. Molecular rotations Rotational transitions are of little use to the spectroscopist. Rotational levels are quantized, and absorption of IR by gases yields line spectra. However, in liquids or solids, these lines broaden into a continuum due to molecular collisions and other interactions. Molecular vibrations The positions of atoms in a molecules are not fixed; they are subject to a number of different vibrations. Vibrations fall into the two main catagories of stretching and bending. Stretching: Change in inter-atomic distance along bond axis Bending: Change in angle between two bonds. There are four types of bend: Rocking Scissoring Wagging Twisting Vibrational coupling In addition to the vibrations mentioned above, interaction between vibrations can occur (coupling) if the vibrating bonds are joined to a single, central atom. Vibrational coupling is influenced by a number of factors; Strong coupling of stretching vibrations occurs when there is a common atom between the two vibrating bonds Coupling of bending vibrations occurs when there is a common bond between vibrating groups Coupling between a stretching vibration and a bending vibration occurs if the stretching bond is one side of an angle varied by bending vibration Coupling is greatest when the coupled groups have approximately equal energies No coupling is seen between groups separated by two or more bonds Infrared Spectrophotometry In the emission spectra of hydrogen, helium, and mercury, the colored lines are the light energy emitted when an electron falls from a higherenergy state down to a lower energy state Electronic absorption spectroscopy in the visible area of the EM spectrum measures the energy of the reverse process, i.e., the wavelength of light required to promote an electron to a higher energy orbital. This light is in the visible and UV regions of the electromagnetic spectrum. If molecules are excited by photons of infrared light (IR), 600-3500 cm^-1, the electrons in the covalent bonds of those molecules will vibrate. Different types of bonds (C-C, CO, C-H, C=O, etc.) vibrate at distinct IR frequencies. The absorption of infrared light results in bending, stretching, scissoring, rocking, and other vibrations of the bonds. The energy of the vibration depends on the type of vibrational mode (bending, rocking, etc.), the mass of the atoms across the bond, and the strength of the bond.An instrument which measures the absorption of infrared light by molecules is called an Infrared Spectrophotometer. For example, C-O stretching vibrations appear within the same region of the spectrum, , regardless of what else are bonded to those atoms . C=O stretches occur at higher frequencies, around 1600-1700 cm^-1, because the bond is stronger. Due to this characteristic behavior, one can usually get a rough idea of the types of atoms and bonds present in a molecule by looking at the wavelengths (frequencies) of the IR absorption bands. The IR spectrum is divided roughly into two sections. The area from 3500-1500 cm^-1 is called the functional group area; it identifies the presence of alkanes, alkenes, alkynes, aldehydes, alcohols, etc. The rest of the spectrum is known as the fingerprint region. The peaks in this part are fairly unique to the substance being analyzed and a peak-by-peak correspondence to a known spectrum confirms identification. Group -CH3 -CH2-O-H Principal Transmittance Bands (cm^-1) 2962 2872 1460 1375 2926 2863 1455 3350+/- -C-Oaromatic ring 150 1050-1150 3050+/-50 1601 1500 730 690 This bond can be found in a variety of functional groups shown below: ketone 1715 aldehyde 1727 ester 1190-1245 Recycling Structure Polymer Name Symbol -C=O polyethylene terephthalate Uses soda bottles milk , detergent, high density bleach, polyethylene water, and vinegar bottles plumbing fixtures, some water bottles, polyvinylchloride glass cleaner bottles (usually clear) grocery and other shopping low density bags, polyethylene bread bags, food wrap indooroutdoor carpeting, polypropylene some yogurt and margarine nonrecyclable at this time containers, shampoo and syrup bottles toys, insulated cups and polystyrene containers (as styrofoam) drink resins, complex boxes and composites, squeezable laminates catsup bottles Introductory theory Introduction Few methods of chemical analysis are truly specific to a particular analyte. It is often found that the analyte of interest must be separated from the myriad of individual compounds that may be present in a sample. As well as providing the analytical scientist with methods of separation, chromatographic techniques can also provide methods of analysis. Chromatography involves a sample (or sample extract) being dissolved in a mobile phase (which may be a gas, a liquid or a supercritical fluid). The mobile phase is then forced through an immobile, immiscible stationary phase. The phases are chosen such that components of the sample have differing solubilities in each phase. A component which is quite soluble in the stationary phase will take longer to travel through it than a component which is not very soluble in the stationary phase but very soluble in the mobile phase. As a result of these differences in mobilities, sample components will become separated from each other as they travel through the stationary phase. Techniques such as H.P.L.C. (High Performance Liquid Chromatography) and G.C. (Gas Chromatography) use columns - narrow tubes packed with stationary phase, through which the mobile phase is forced. The sample is transported through the column by continuous addition of mobile phase. This process is called elution. The average rate at which an analyte moves through the column is determined by the time it spends in the mobile phase. Distribution of analytes between phases The distribution of analytes between phases can often be described quite simply. An analyte is in equilibrium between the two phases; Amobile Astationary The equilibrium constant, K, is termed the partition coefficient; defined as the molar concentration of analyte in the stationary phase divided by the molar concentration of the analyte in the mobile phase. The time between sample injection and an analyte peak reaching a detector at the end of the column is termed the retention time (tR ). Each analyte in a sample will have a different retention time. The time taken for the mobile phase to pass through the column is called tM. A term called the retention factor, k', is often used to describe the migration rate of an analyte on a column. You may also find it called the capacity factor. The retention factor for analyte A is defined as; k'A = t R - tM / tM t R and tM are easily obtained from a chromatogram. When an analytes retention factor is less than one, elution is so fast that accurate determination of the retention time is very difficult. High retention factors (greater than 20) mean that elution takes a very long time. Ideally, the retention factor for an analyte is between one and five. We define a quantity called the selectivity factor, α , which describes the separation of two species (A and B) on the column; α = k 'B / k 'A When calculating the selectivity factor, species A elutes faster than species B. The selectivity factor is always greater than one. Band broadening and column efficiency To obtain optimal separations, sharp, symmetrical chromatographic peaks must be obtained. This means that band broadening must be limited. It is also beneficial to measure the efficiency of the column. The Theoretical Plate Model of Chromatography The plate model supposes that the chromatographic column is contains a large number of separate layers, called theoretical plates. Separate equilibrations of the sample between the stationary and mobile phase occur in these "plates". The analyte moves down the column by transfer of equilibrated mobile phase from one plate to the next. It is important to remember that the plates do not really exist; they are a figment of the imagination that helps us understand the processes at work in the column.They also serve as a way of measuring column efficiency, either by stating the number of theoretical plates in a column, N (the more plates the better), or by stating the plate height; the Height Equivalent to a Theoretical Plate (the smaller the better). If the length of the column is L, then the HETP is HETP = L / N The number of theoretical plates that a real column possesses can be found by examining a chromatographic peak after elution; where w1/2 is the peak width at half-height. As can be seen from this equation, columns behave as if they have different numbers of plates for different solutes in a mixture. The Rate Theory of Chromatography A more realistic description of the processes at work inside a column takes account of the time taken for the solute to equilibrate between the stationary and mobile phase (unlike the plate model, which assumes that equilibration is infinitely fast). The resulting band shape of a chromatographic peak is therefore affected by the rate of elution. It is also affected by the different paths available to solute molecules as they travel between particles of stationary phase. If we consider the various mechanisms which contribute to band broadening, we arrive at the Van Deemter equation for plate height; HETP = A + B / u + C u where u is the average velocity of the mobile phase. A, B, and C are factors which contribute to band broadening. A - Eddy diffusion The mobile phase moves through the column which is packed with stationary phase. Solute molecules will take different paths through the stationary phase at random. This will cause broadening of the solute band, because different paths are of different lengths. B - Longitudinal diffusion The concentration of analyte is less at the edges of the band than at the center. Analyte diffuses out from the center to the edges. This causes band broadening. If the velocity of the mobile phase is high then the analyte spends less time on the column, which decreases the effects of longitudinal diffusion. C - Resistance to mass transfer The analyte takes a certain amount of time to equilibrate between the stationary and mobile phase. If the velocity of the mobile phase is high, and the analyte has a strong affinity for the stationary phase, then the analyte in the mobile phase will move ahead of the analyte in the stationary phase. The band of analyte is broadened. The higher the velocity of mobile phase, the worse the broadening becomes. Van Deemter plots A plot of plate height vs. average linear velocity of mobile phase. Such plots are of considerable use in determining the optimum mobile phase flow rate. Resolution Although the selectivity factor, α, describes the separation of band centres, it does not take into account peak widths. Another measure of how well species have been separated is provided by measurement of the resolution. The resolution of two species, A and B, is defined as Baseline resolution is achieved when R = 1.5 It is useful to relate the resolution to the number of plates in the column, the selectivity factor and the retention factors of the two solutes; To obtain high resolution, the three terms must be maximised. An increase in N, the number of theoretical plates, by lengthening the column leads to an increase in retention time and increased band broadening - which may not be desirable. Instead, to increase the number of plates, the height equivalent to a theoretical plate can be reduced by reducing the size of the stationary phase particles. It is often found that by controlling the capacity factor, k', separations can be greatly improved. This can be achieved by changing the temperature (in Gas Chromatography) or the composition of t α can also be manipulated to improve separations. When α is close to unity, optimising k' and increasing N is not sufficient to give good separation in a reasonable time. In these cases, k' is optimised first, and then α is increased by one of the following procedures: 1. Changing mobile phase composition 2. Changing column temperature 3. Changing composition of stationary phase 4. Using special chemical effects (such as incorporating a species which complexes with one of the solutes into the stationary phase) Introduction Gas chromatography - specifically gas-liquid chromatography - involves a sample being vapourised and injected onto the head of the chromatographic column. The sample is transported through the column by the flow of inert, gaseous mobile phase. The column itself contains a liquid stationary phase which is adsorbed onto the surface of an inert solid. Have a look at this schematic diagram of a gas chromatograph: Instrumental components Carrier gas The carrier gas must be chemically inert. Commonly used gases include nitrogen, helium, argon, and carbon dioxide. The choice of carrier gas is often dependant upon the type of detector which is used. The carrier gas system also contains a molecular sieve to remove water and other impurities. Sample injection port For optimum column efficiency, the sample should not be too large, and should be introduced onto the column as a "plug" of vapour - slow injection of large samples causes band broadening and loss of resolution. The most common injection method is where a microsyringe is used to inject sample through a rubber septum into a flash vapouriser port at the head of the column. The temperature of the sample port is usually about 50°C higher than the boiling point of the least volatile component of the sample. For packed columns, sample size ranges from tenths of a microliter up to 20 microliters. Capillary columns, on the other hand, need much less sample. For capillary GC, split/splitless injection is used. Have a look at this diagram of a split/splitless injector; The injector can be used in one of two modes; split or splitless. The injector contains a heated chamber containing a glass liner into which the sample is injected through the septum. The carrier gas enters the chamber and can leave by three routes (when the injector is in split mode). The sample vapourises to form a mixture of carrier gas, vapourised solvent and vapourised solutes. A proportion of this mixture passes onto the column, but most exits through the split outlet. The septum purge outlet prevents septum bleed components from entering the column. Columns There are two general types of column, packed and capillary (also known as open tubular). Packed columns contain a finely divided, inert, solid support material (commonly based on diatomaceous earth) coated with liquid stationary phase. Most packed columns are 1.5 - 10m in length and have an internal diameter of 2 - 4mm. Capillary columns have an internal diameter of a few tenths of a millimeter. They can be one of two types; wall-coated open tubular (WCOT) or support-coated open tubular (SCOT). Wall-coated columns consist of a capillary tube whose walls are coated with liquid stationary phase. In support-coated columns, the inner wall of the capillary is lined with a thin layer of support material such as diatomaceous earth, onto which the stationary phase has been adsorbed. SCOT columns are generally less efficient than WCOT columns. Both types of capillary column are more efficient than packed columns. In 1979, a new type of WCOT column was devised - the Fused Silica Open Tubular (FSOT) column; These have much thinner walls than the glass capillary columns, and are given strength by the polyimide coating. These columns are flexible and can be wound into coils. They have the advantages of physical strength, flexibility and low reactivity. Column temperature For precise work, column temperature must be controlled to within tenths of a degree. The optimum column temperature is dependant upon the boiling point of the sample. As a rule of thumb, a temperature slightly above the average boiling point of the sample results in an elution time of 2 - 30 minutes. Minimal temperatures give good resolution, but increase elution times. If a sample has a wide boiling range, then temperature programming can be useful. The column temperature is increased (either continuously or in steps) as separation proceeds. Detectors There are many detectors which can be used in gas chromatography. Different detectors will give different types of selectivity. A non-selective detector responds to all compounds except the carrier gas, a selective detector responds to a range of compounds with a common physical or chemical property and a specific detector responds to a single chemical compound. Detectors can also be grouped into concentration dependant detectors and mass flow dependant detectors. The signal from a concentration dependant detector is related to the concentration of solute in the detector, and does not usually destroy the sample Dilution of with make-up gas will lower the detectors response. Mass flow dependant detectors usually destroy the sample, and the signal is related to the rate at which solute molecules enter the detector. The response of a mass flow dependant detector is unaffected by make-up gas. Have a look at this tabular summary of common GC detectors: Support Dynamic Detector Type Selectivity Detectability gases range Flame ionization (FID) Hydrogen and air Most organic cpds. 100 pg 107 Thermal conductivity Concentration Reference (TCD) Universal 1 ng 107 Electron capture (ECD) Halides, nitrates, nitriles, peroxides, anhydrides, 50 fg 105 Mass flow Concentration Make-up organometallics Nitrogenphosphorus Mass flow Hydrogen and air Nitrogen, phosphorus 10 pg 106 Flame photometric Mass flow (FPD) Hydrogen and air possibly oxygen Sulphur, phosphorus, tin, boron, arsenic, 100 pg germanium, selenium, chromium 103 Aliphatics, aromatics, ketones, esters, aldehydes, amines, 2 pg heterocyclics, organosulphurs, some organometallics 107 Photoionization (PID) Concentration Make-up Hall electrolytic Mass flow conductivity Hydrogen, Halide, nitrogen, oxygen nitrosamine, sulphur The effluent from the column is mixed with hydrogen and air, and ignited. Organic compounds burning in the flame produce ions and electrons which can conduct electricity through the flame. A large electrical potential is applied at the burner tip, and a collector electrode is located above the flame. The current resulting from the pyrolysis of any organic compounds is measured. FIDs are mass sensitive rather than concentration sensitive; this gives the advantage that changes in mobile phase flow rate do not affect the detector's response. The FID is a useful general detector for the analysis of organic compounds; it has high sensitivity, a large linear response range, and low noise. It is also robust and easy to use, but unfortunately, it destroys the sample. DNA technology Determining the molecular sequence of DNA that makes up the genome of different organisms is an international scientific goal, several laboratories are participating worldwide in this task Recombinant DNA Technology Techniques for - Isolation - Digestion - Fractionation - Purification of the TARGET fragment - Cloning into vectors - Transformation of host cell and selection - Replication - Analysis - Expression of DNA How do we obtain DNA and how do we manipulate DNA? Quite straightforward to isolate DNA For instance, to isolate genomic DNA 1. Remove tissue from organism 2. Homogenise in lysis buffer containing guanidine thiocyanate (denatures proteins) 3. Mix with phenol/chloroform - removes proteins 4. Keep aqueous phase (contains DNA) 5. Add alcohol (ethanol or isopropanol) to precipitate DNA from solution 6. Collect DNA pellet by centrifugation 7. Dry DNA pellet and resuspend in buffer 8. Store at 4°C Enzymes that can cut (hydrolyse) DNA duplex at specific sites. Current DNA technology is totally dependent on restriction enzymes. Restriction enzymes are endonucleases Restriction enzymes recognise a specific short nucleotide sequence This is known as a Restriction Site The phosphodiester bond is cleaved between specific bases, one on each DNA strand Examples of restriction enzymes and the sequences they cleave Source microorganism Enzyme Arthrobacter luteus Alu I Bacillus amyloiquefaciens H Recognition Site Ends produced AG CT Blunt Bam HI G GATCC Sticky Escherichia coli Eco RI G AATTC Sticky Haemophilus gallinarum Hga I GACGC(N)5 Sticky Haemophilus infulenzae Hind III A AGCTT Sticky Providencia stuartii 164 Pst I CTGCA G Sticky Nocardia otitiscaviaruns Not I GC GGCCGC Sticky DNA fractionation Separation of DNA fragments in order to isolate and analyse DNA cut by restriction enzymes Electrophoresis Linear DNA fragments of different sizes are resolved according to their size through gels made of polymeric materials such as polyacrylamide and agarose. For instance, agarose is a polysaccharide derived from seaweed - and gels formed from between 0.5% to 2% (mass/volume i.e. 0.5 to 2.0g agarose/100 ml of aqueous buffer) can be used to separate (resolve) most sizes of DNA DNA is electrophoresed through the agarose gel from the cathode (negative) to the anode (positive) when a voltage is applied, due to the net negative charge carried on DNA When the DNA has been electrophoresed, the gel is stained in a solution containing the chemical ethidium bromide. This compound binds tightly to DNA (DNA chelator) and fluoresces strongly under UV light - allowing the visualisation and detection of the DNA. Analysing complex nucleic acid mixtures (DNA or RNA) The total cellular DNA of an organism (genome) or the cellular content of RNA are complex mixtures of different nucleic acid sequences. Restriction digest of a complex genome can generate millions of specific restriction fragments and there can be several fragments of exactly the same size which will not be separated from each other by electrophoresis. Techniques have been devised to identify specific nucleic acids in these complex mixtures Southern blotting - DNA Northern blotting - RNA These techniques are not to be confused with Western blotting, which is used to analyse PROTEINS which have been immobilised on nitrocellulose/nylon filters. Proteins which have been separated by polyacrylamide gel electrophoresis (PAGE) are transferred to nitrocellulose/nylon filters and the filter is probed with antibodies to detect the specific protein - similar to the method used for expression library screening. Southern blotting This technique, devised by Ed Southern in 1975, is a commonly used method for the identification of DNA fragments that are complementary to a know DNA sequence. Southern hybridisation, also called Southern blotting, allows a comparison between the genome of a particular organism and that of an available gene or gene fragment (the probe). It can tell us whether an organism contains a particular gene, and provide information about the organisation and restriction map of that gene. In Southern blotting, chromosomal DNA is isolated from the organism of interest, and digested to completion with a restriction endonuclease enzyme. The restriction fragments are then subjected to electrophoresis on an agarose gel, which separates the fragments on the basis of size. DNA fragments in the gel are denatured (i.e. separated into single strands) using an alkaline solution. The next step is to transfer fragments from the gel onto nitrocellulose filter or nylon membrane. This can be performed by electrotransfer (electrophoresing the DNA out of the gel and onto a nitrocellulose filter), but is more typically performed by simple capillary action. In this system, the denatured gel is placed onto sheet(s) of moist filter paper and immersed in a buffer reservoir. A nitrocellulose membrane is laid over the gel, and a number of dry filter papers are placed on top of the membrane. By capillary action, buffer moves up through the gel, drawn by the dry filter paper. It carries the single-stranded DNA with it, and when the DNA reaches the nitrocellulose it binds to it and is immobilised in the same position relative to where it had migrated in the gel. The DNA is bound irreversibly to the filter/membrane by baking at high temperature (nitrocellulose) or cross-linking through exposure to UV light (nylon). The final step is to immerse the membrane in a solution containing the probe either a DNA (cDNA clone, genomic fragment, oligonucleotide) or RNA probe can be used. This is DNA hybridisation - in other words the target DNA and the probe DNA/RNA form a 'hybrid' because they are complementary sequences and so can bind to each other. The probe is usually radioactively labelled with 32P, often by removal of the 5' phosphate of the probe with alkaline phosphatase, and replacement with a radiolabelled phosphate using α -[32P]ATP and polynucleotide kinase. The membrane is washed to remove non-specifically bound probe (see washing & stringency conditions), and is then exposed to X-ray film - a process called autoradiography. At positions where the probe is bound, α -emissions from the probe cause the X-ray film to blacken. This allows the identification of the sizes and the number of fragments of chromosomal genes with strong similarity to the gene or gene fragment used as a probe. The principle of Southern blotting What Southern blotting can tell us 1. Whether a particular gene is present and how many copies are present in the genome of an organism 2. The degree of similarity between the chromosomal gene and the probe sequence 3. Whether recognition sites for particular restriction endonucleases are present in the gene. By performing the digestion with different endonucleases, or with combinations of endonucleases, it is possible to obtain a restriction map of the gene i.e. an idea of the restriction enzyme sites in and around the gene- which will assist in attempts to clone the gene. 4. Whether re-arrangements have occurred during the cloning process Northern blotting Northern blotting is a simple extension of Southern blotting - and derives its name from the earlier technique. It is used to detect cellular RNA rather than DNA. Initially, it was thought that RNA would not bind efficiently to nitrocellulose, and other modified materials were synthesised for use as a membrane. However, it was then shown that when RNA was denatured, that it would also bind efficiently to nitrocellulose. This means that the RNA has to be unfolded into a linear strand before it will bind efficiently to nitrocellulose. Chemicals such as formaldehyde and methylmercuric hydroxide can be used to denature the RNA - breaking down hydrogen bonding structure in the molecule. Alkali is not used to denature the RNA - since RNA is degraded under alkaline conditions. Isolating RNA RNA is extracted from the cells of interest - but precautions must be taken to avoid degradation of the single-stranded RNA by ribonuclease (RNase), which is found on the skin and on glassware. Wear gloves, use specially treated plastics and glassware to avoid accidently introducing ribonuclease to extraction prep. Addition of diethylpyrocabonate (DEPC) inhibits ribonuclease activity and baking at high temperature destroys ribonuclease activity (only useful for treating heat resistant equipment, such as glassware). DNA sequencing: Maxam & Gilbert sequencing For this method, need to use DNA fragments ~500 nucleotides - for instance by isolating restriction fragments of the DNA that is to be sequenced. The method is reliable for sequencing up to ~250-300 nucleotides at a time. The technique requires that the target DNA is end-labeled (usually radioactively). Either at the 5' end: add alkaline phosphatase to remove 5' phosphate, and polynucleotide kinase to add back a radiolabeled phosphate to the 5' OH group using [α 32P]ATP Or at the 3' end: add a homopolymeric tail using terminal tranferase and [α 32 P]dATP (or another 32P-labeled deoxyribonucleotide triphosphate) Both single-stranded (ss) and double-stranded DNA (ds) can be sequenced. If ds DNA is used, then the label must be removed from one end, so that fragment sizes can be gauged by the distance to the end of the molecule from one unique label at the other end. The M&G method involves the chemical degradation of DNA The process requires addition of chemicals that bring about cleavage of DNA at specific positions. (The Sanger method involves DNA synthesis). Either 4 or 5 separate chemical reactions are performed. The reactions are carried out in two stages: Stage 1: Specific chemical modification of bases in the DNA Stage 2: Chemical cleavage of sugar-phosphate backbone at modification site Stage 1: Specific chemical modification of bases in the DNA Base Specific modification modified G Methylation of base with dimethyl sulfate at pH 8.0. Makes base susceptible to cleavage by alkali A+G Treatment with piperidine formate at pH 2.0. Results in removal of purine bases C+T Hydrazine opens pyrimidine rings and causes their removal from DNA C In high ionic strength (1.5 M NaCl) only cytosine reacts with hydrazine Treatment with 1.2 M NaOH at high temperature (90°C) gives strong cleavage at A, less at C Stage 2 When bases are modified and destroyed by the treatments in stage 1, piperidine at 90°C is used to cleave the sugar-phosphate backbone at the site. A>C For example: The 'trick' in the reaction is to limit incubation times with base-modifying reagents and/or the concentrations of reagents used, so that a ladder of progressively longer molecules is generated in the M&G sequencing reactions. The differently sized/labelled fragments are separated by polyacrylamide gel electorphoresis (PAGE) Limitations: Resolving power of the gel. A typical gel is composed of 6% acrylamide (6g/100ml). By altering the % of the acrylamide, shorter fragments (use higher %) or longer fragments (use lower %) can be resolved. Samples can also be electrophoresed for different lengths of time to resolve fragments of different sizes. Dangers: The chemicals used destroy DNA - they are mutagens Sanger dideoxy sequencing A DNA synthesis reaction. Needs ss DNA as the template. If only ds DNA is available, can be treated with alkali to separate it into single strands (denature it). Rather more bases can usually be read by the Sanger technique: ~500 vs ~300 by the M&G technique. The Sanger method The reaction requires an oligonucleotide primer (typically 16-17 nucleotides long). The primer is annealed to the ssDNA template. Four separate synthesis reactions are set up for each DNA template/oligo primer. Method uses 4 different dideoxyribonucleotide triphosphates (ddNTPs), one for each reaction mix Remember: for DNA polymerase to continue synthesising the DNA strand, it requires a free -OH group at the 3' position of the sugar. Look carefully - this is missing from the dideoxyribonucleotide. This is the chemistry behind the technique: the phosphodiester bond that forms between successive nucleotides in a DNA chain requires a 3' hydroxyl group. The oligonucleotide primer provides the first free -OH group and each successive nucleotide that is added provides the next. ddNTPs differ from ordinary dNTPs in that they have a hydrogen (rather than a hydroxy) on the 3' position of the sugar. Once a ddNTP is incorporated, the nucleotide chain can't be extended from that point. In each of 4 reaction mixtures the following reagents are mixed 1. Template ssDNA 2. Primer 3. Each of the 4 dNTPs (dATP, dCTP, dTTP, dGTP) 4. DNA polymerase (Klenow fragment or T7 DNA polymerase) 5. One of the 4 possible ddNTPs: 'G mix': add ddGTP 'A mix': add ddATP 'T mix': add ddTTP 'C mix': add ddCTP Sanger chain termination technique The newly synthesised strand is labeled with a radionucleotide. Either the primer is 5' end-labelled with 32P or a radiolabeled deoxyribonucleotide is incorporated in the growing chain ([α 35S]dATP is popularly used). In each of the 4 mixes, the ratio of dNTP:ddNTP is critical. The ratio is usually ~100:1. This results in a reasonably random termination of chains of DNA at different positions. Each of the 4 synthesis mixes will contain a series of differently sized radiolabeled DNA fragments, in each mixture all will terminate at the same type of nucleotide. The length is dependent on how far the nucleotide chain is extended before a ddNTP is incorporated. Chain termination at different G residues: the 'G mix' reaction Like M&G sequencing, the reactions are loaded into 4 adjacent lanes on a long gel for resolving the different sized fragments by PAGE. After electrophoresis the gel is dried and exposed to X-ray film and the sequence read from the bottom to the top of the gel