Biochemistry Additional Techniques PDF

# Additional Techniques ## Introduction - In addition to electrophoresis, blotting, and chromatography, biochemistry employs a variety of other techniques to study the chemistry of biomolecules and living systems. - This lesson provides a broad overview of commonly used biochemical techniques, wit...

# Additional Techniques ## Introduction - In addition to electrophoresis, blotting, and chromatography, biochemistry employs a variety of other techniques to study the chemistry of biomolecules and living systems. - This lesson provides a broad overview of commonly used biochemical techniques, with an emphasis on methods that may be referenced on the exam. - A deep understanding of the technical aspects of each method is unlikely to be required, however, knowledge of the underlying principles of each method is still helpful in understanding passages and in interpreting data derived by a specific method. - Research methods in any natural science, but especially in biochemistry, often draw upon basic science principles covered in other courses (e.g., general and organic chemistry, physics, biology). - This lesson strives to be accessible to any student of biochemistry, referring to the relevant lessons of other subjects as needed. ## 14.4.01 Dialysis - Lesson 14.3 discusses chromatography as a means of protein purification. - Methods such as ion-exchange chromatography and affinity chromatography can separate a protein of interest from contaminating proteins; however, removal from the column yields a protein of interest collected in an elution buffer. - Depending on the method, the elution buffer may include contaminating ligands and other contaminants, or a pH or salt concentration that is incompatible with later experiments. - The purified protein must therefore be restored to conditions that are compatible. - One method of exchanging the elution buffer with a more compatible buffer is dialysis. - In dialysis, the sample of interest is loaded into a container with a porous membrane. - The pores must be large enough for salt, water, and small labels to pass through but small enough that the protein of interest cannot pass through. - The other side of the porous membrane is exposed to the desired final buffer *(i.e.*, the dialysate). - Undesired small molecules or ions diffuse out of the sample and into the dialysate fluid, and desired solutes diffuse in *(Figure 14.43)*. - Water also travels across the membrane by osmosis, equalizing osmotic pressure across the membrane. - Eventually, the small, permeable solutes reach a diffusive equilibrium wherein they are in equal concentrations across the membrane. ## When dialyzing a protein for purification purposes, several rounds of dialysis are usually employed. - In each round, sufficient time is given for the sample and dialysate buffers to equilibrate, which "cleans" the sample and "dirties" the dialysate. - Every new round replaces the dialysate fluid with fresh fluid, allowing the sample to become even cleaner, as shown in *(Figure 14.44)*. ## The principle of dialysis also has clinical relevance. - For example, hemodialysis is a treatment for patients whose kidneys cannot sufficiently filter their blood. - In this case, patient blood is diverted from the body and into a dialysis machine, where it is exposed to dialysate fluid across a porous membrane. - Excess waste products diffuse out of the blood and into the dialysate fluid, and nourishing electrolytes and other small molecules diffuse from the dialysate fluid and back to the blood *(Figure 14.45)*. - Cleaned blood exits the dialysis machine and is returned to the patient. ## 14.4.02 Biomolecule Quantitation - After a protein or any other biomolecule has been collected, most downstream applications require knowledge of sample concentration *(i.e.*, the amount of the specific biomolecule of interest in the sample needs to be quantified). - The absolute quantitation methods discussed in this lesson provide concentration values in terms of molarity *(e.g.*, mM) or in mass per volume *(e.g.*, mg/mL). - Importantly, absolute quantitation often must be used prior to electrophoresis and blotting to ensure that changes in band density are due to changes in expression level and not due to loading different amounts of total protein or nucleic acid into different lanes. ## Review of UV-Vis Spectroscopy Principles - Various methods exist to quantify biomolecules, many of which rely on ultraviolet-visual (UV/Vis) absorption spectroscopy. - The principles of UV/Vis spectroscopy are covered in detail in organic chemistry lesson 14.5. - In brief, different molecules absorb specific energies and wavelengths of ultraviolet and visible light. - The wavelength that a molecule absorbs most strongly, its lambda max *(λmax)*, is determined empirically. - The amount of absorbance A is related to concentration of the molecule *(c)* by the equation: $A = εcl$ - where ε represents the absorption coefficient *(i.e.*, the absorptivity) of the analyte and l is the pathlength *(i.e.*, the length of the sample through which the light passes). - This equation demonstrates that absorbance is directly proportional to sample concentration *(Figure 14.46)*. - Therefore, biomolecule concentration can be determined from sample absorbance. - For purified molecules that strongly absorb UV or visible light with known ε values, calculation of concentration is straightforward. - For impure samples, samples with poor absorptivity at their λmax, or samples with poorly defined ε values, however, additional measures must be taken. ## Absorbance and Quantitation of Purified Biopolymers - Nucleic acids *(i.e.,* DNA, RNA) and most proteins can be directly detected by ultraviolet spectroscopy. - Because tryptophan residues have a strong λmax at 280 nm, many proteins can be quantified by measuring absorbance at 280 nm *(i.e.,* A280). - In contrast, nucleotides absorb strongly at 260 nm, so DNA and RNA can be quantified by measuring absorbance at their λmax of 260 nm *(A260)*. - The values of the absorption coefficients of some proteins are known; however, when measuring the absorbance of a protein with an unknown absorption coefficient, it is common practice to use an average literature value. - Alternatively, the absorption coefficient of a protein of interest can be estimated based on the amount of tryptophan residues *(and, to a lesser extent, the amount of tyrosine and cysteine residues)* in that protein. - However, estimates of a protein's absorptivity based on these assumptions come with two caveats when interpreting data. ### The first caveat is that concentrations determined using average coefficient values may be inaccurate if the protein has a greater-than-average or less-than-average percentage of tryptophan residues. ### The second caveat is that the use of an average literature absorption coefficient value results in data that correlate more closely with the mass of protein in a sample than it does to the moles of protein in a sample. - A large protein with a typical percentage of tryptophan residues would absorb 280 nm light more strongly than a smaller protein that has the same tryptophan percentage, because the large protein contains more total tryptophan residues. - In other words, these measurements facilitate calculation of the protein concentration in mg/mL. - This concept is illustrated in *(Figure 14.47)*. ## When needed, the mass concentration *(e.g.*, mg/mL) of a sample of purified protein can be converted to molar concentration by dividing by the molar mass. - Recall that the molar mass of a protein in g/mol is numerically equal to its molecular mass in Da. - A protein with a mass of 50 kDa has a molar mass of 50 kg/mol. - Typical absorption spectra for purified protein and purified DNA are given in *(Figure 14.48)*. - Although both types of biopolymers have clearly different λmax values, the spectra do overlap *(e.g.,* tryptophan absorbs 260 nm light, just not as strongly as 280 nm light). - Consequently, protein contamination of DNA samples can affect DNA concentration measurements and vice versa. ## Consequently, although A280 and A260 measurements are quick and simple to set up, they are mainly used for relatively pure samples. - When quantifying impure samples *(e.g.*, crude lysates), different protocols are used that typically involve staining the biopolymer of interest. ## Absorbance and Quantitation of Impure Samples - Various methods of protein quantitation for impure samples exist *(e.g.*, Bradford assay, BCA assay, Lowry assay). - Each method differs slightly in their preparation, the resultant λmax, and their compatible contaminating reagents; however, the basic principle of each technique is similar. - Proteins within a sample react with reagents to produce an intense visible color upon interaction. - These proteins are said to be stained. - The intensity of many of these protein stains may vary with time or may be altered based on the environmental conditions of the experiment or the lab. - As such, literature values for the absorption coefficient are unreliable. - Instead, it is common practice to produce a new standard curve for each experiment. - Standard curves involve measuring the absorbance of samples with a known protein concentration. - For example, purified bovine serum albumin *(BSA)* can be obtained as a powder that can be accurately weighed out and diluted in the same buffer used for the protein of interest. - The absorbance measurements are plotted against the mass concentrations of the known standard, and a line of best fit *(i.e.*, the standard curve) is calculated. - From the standard curve, the measured absorbance of the sample containing the protein of interest can then be correlated to a mass concentration *(Figure 14.49)*. ## Instrument limitations often restrict the linear relationship between absorbance and concentration to a small range. - Therefore, it is common practice to choose concentrations of the protein standard that give absorbance measurements between 0.1 and 1 absorbance units *(AU)*. - If the sample of interest gives an absorbance measurement beyond that range, it can be diluted by a known factor. - The resulting absorbance and concentration value can be multiplied by the dilution factor to give the original concentration. - For example, a 10-fold dilution is prepared by mixing 1 mL of sample with 9 mL of dilution buffer *(10 mL total volume)*. - If the diluted sample yields an absorbance corresponding to 0.45 mg/mL of protein, then the undiluted sample has a concentration of 0.45 x 10 = 4.5 mg/mL. - In very concentrated samples, a diluted sample can be diluted again, multiple times if needed, in a process known as a serial dilution. - In this case the overall dilution factor is the product of the individual dilution factors. - For instance, mixing 1 mL of a protein with 9 mL of buffer, and then mixing 1 mL of that diluted solution with 9 more mL of buffer results in a 100-fold dilution. - *(Figure 14.50)* depicts an example of a concentration calculation using serial dilution. ## 14.4.03 Binding Assays - One of the most fundamental types of protein study is a binding assay, which analyzes the binding interaction between a protein and a ligand. - Binding interactions are also discussed in lesson 3.1. - In brief, the strength of a binding interaction between a protein and its ligand can be described by the Kd value *(i.e.*, the equilibrium dissociation constant) of the interaction. - The Kd value is defined as: $Kd = \frac{[P] [L]}{[PL]}$ - where [P] represents the concentration of free *(i.e.,* unbound) protein, [L] represents the concentration of free ligand, and [PL] represents the concentration of the protein-ligand complex. - From this relation, an equation describing the fraction of protein bound by ligand *(θ)* can be derived. $Fraction\ bound, \theta = \frac{[PL]}{[P] + [PL]} = \frac{[L]}{Kd + [L]}$ ## Binding Assays That Assume the Free Ligand Approximation - Importantly, the equations describing protein binding are defined using [L], which is the concentration of free, unbound ligand. - However, the experimenter usually only controls [Ltot], the total concentration of ligand, which is equal to [L] + [PL]. - The free ligand approximation can be used to assume [L] ≈ [Ltot], but this assumption is only valid if the ligand concentration is much higher than the total protein concentration. - When this approximation is valid, various methods can be used to measure protein binding. - For example, if the ligand-bound protein has a different absorption spectrum relative to unbound protein and ligand, then protein binding can be assessed using UV/Vis spectroscopy. - *(Figure 14.51)* shows how UV/Vis spectroscopy can be used to measure oxygen binding by hemoglobin. ## Isothermal Titration Calorimetry - Binding affinity between a protein and a ligand can also be determined by measuring the thermodynamics of the binding reaction. - Specifically, the enthalpy of the binding reaction (*ΔH*) can be measured by a technique called isothermal titration calorimetry *(ITC)*. - ITC measures the release of heat that occurs as aliquots of ligand are gradually injected *(or titrated)* into a sample of purified protein and bind. - In an isothermal titration calorimetry experiment, the small reaction vessel is maintained at a constant temperature *(i.e.*, isothermal conditions), and the power needed to maintain that temperature is monitored over time. - If the addition of ligand results in an exothermic binding reaction, the ITC machine records a decrease in machine power needed to maintain temperature because the heat released by the reaction helps maintain it. - Each time an injection occurs, the change in power needed is plotted as a deflection *(i.e.*, either a peak or an inverted peak) on a graph. - For any given injection, the area between the peak and the baseline represents the enthalpy change *ΔH* that results from the ligand injection *(Figure 14.52)*. ## As with other binding assays, this signal (the ∆H) eventually stops changing between injections, indicating that the protein is saturated and therefore no more binding can occur. - Because reaction thermodynamics are directly measured by this technique, rather than indirectly monitored through a reporter *(e.g.*, radioactivity, fluorescence), ITC determination of Kd does not require use of the free ligand approximation. ## 14.4.04 Melting Temperature Assays - Lesson 2.3 and Lesson 8.2 introduced the melting temperature *(Tm)* of proteins and nucleic acids as a descriptor of biopolymer stability. - Specifically, Tm is the temperature at which 50% of the biopolymer becomes denatured. - This concept describes assays that can be used to determine the Tm. ## In general chemistry and physics, "melting" is often used to describe the transition from a solid phase to a liquid phase. - As the temperature of a solid rises past its melting point, the average kinetic energy of the constituent molecules eventually surpasses the energy needed to break the intermolecular forces holding it together. - Therefore, the material changes from a rigidly packed solid to a fluid, freely diffusible liquid. - However, with biochemical melting temperatures, the polymers typically remain in an aqueous phase both before and after the transition. - Therefore, the use of the term "melting" in this context does not refer to a transition from one state of matter to another. - Instead, melting of biopolymers, like melting a solid to form a liquid, involves the breaking of noncovalent bonds *(e.g.*, intermolecular forces). ## As the temperature of a biopolymer sample increases, the bonds maintaining secondary, tertiary, and quaternary protein structure break apart, as do the hydrogen bonds maintaining nucleic acid base-pairing *(Table 14.1)*. - Due to the cooperative nature of folding, these bond-breaking events tend to occur near the same temperature *(i.e.,* as one bond breaks, it becomes easier for other bonds to break). - Therefore, the melting of biopolymers refers to the denaturing breakage of their noncovalent bonds, and the halfway point of this transition is the polymer's melting temperature, Tm. ## Like the melting of a solid to a liquid, the melting of a biopolymer involves the breaking of noncovalent bonds. ## Measuring Melting Temperature through Calorimetry - Like the solid-to-liquid melting transition *(in which heat added during a phase change does not increase the temperature)*, the melting transition of biopolymers is similarly marked by a rise in heat capacity. - These heat capacity changes can be measured using differential scanning calorimetry *(DSC)*. - DSC experiments increase the temperature at a constant rate while monitoring the power needed to maintain that rate. - As noncovalent bonds begin to break, the power needed increases, because some of the energy is used to break those bonds *(an endothermic process)* rather than increase temperature. - The point of highest power input, indicating the most energy per unit time, is taken as the Tm. - At temperatures above the Tm, when most polymers have already been denatured, the heat capacity begins to drop—few noncovalent bonds remain to absorb energy, and therefore more energy can be used to increase the kinetic energy *(and temperature)* of the sample *(Figure 14.53)*. ## Calorimetry can be used to measure the melting temperature *(Tm)* of biopolymers. - In calorimetry, Tm is the temperature that yields the largest heat capacity. ## Measuring Melting Temperature through Spectroscopy - Melting temperature can also be assessed with spectroscopic measurements that report on protein conformation *(i.e.,* native versus denatured). - Fluorescent techniques *(discussed in detail in concept 14.4.05)* can be used with both protein and nucleic acids. - Certain specialized fluorescent dyes bind specifically to double-stranded regions of DNA or RNA. - Therefore, intact nucleic acids provide a strong initial signal, but as temperatures increase and the nucleic acid denatures, the fluorescence signal decreases. - Protein folding, similarly, can be measured with fluorescence. - The fluorescence signal can come either from intrinsic tryptophan residues or from external fluorophores that have been covalently linked to the protein *(i.e.,* the protein is fluorescently labeled). - A fluorescence signal can be affected by the environment, which can include the conformational state of a protein that a fluorophore is attached to. - Consequently, fluorescence intensity and λmax changes can be monitored as temperature increases from low *(native conformation)* to high *(denatured conformation)*. - Circular dichroism is another spectroscopic technique that can be used to assess protein folding *(see Concept 14.4.06)*. - The spectroscopic data can be plotted as a function of temperature to produce a curve representing protein conformation. - Because folding is a positively cooperative process, the resulting graph has a sigmoidal shape, ranging from 0% denatured at low temperatures to 100% denatured at high temperatures. - The temperature that results in 50% denaturation is the Tm *(Figure 14.54)*. ## 14.4.05 Fluorescence - Fluorescence is a commonly utilized phenomenon in the natural sciences. - Fluorescence techniques are like UV/Vis spectroscopy in that they involve the absorption of photons in the UV/Vis spectrum. - Fluorescence differs in that the absorption of a photon by the fluorescent molecule *(also called excitation)* is followed by the release of a photon with lower energy *(and therefore a longer wavelength)*. - For example, a fluorescent molecule *(i.e.,* a fluorophore) might absorb a high-energy blue photon *(~488 nm)* and release a lower-energy green photon *(~510 nm)*. - In addition to chemical fluorophores, there are also genetically encoded fluorescent tags, such as green fluorescent protein *(GFP)*, that can be used to visualize localization and expression of the tagged proteins both in cell culture and in intact organisms *in vivo*. - Various applications of fluorescence are shown in *(Figure 14.55)*. - What causes the shift in wavelength between the excitation photon and the emission photon? - The absorbance of ultraviolet and visible light occurs when the photon's energy matches the energy needed to excite an electron to a higher energy state. - In fluorescent molecules some—though not all—of the absorbed energy is then released as vibrational energy, or heat, to the surroundings. - The remaining energy can then be emitted as light. - Because the remaining energy is less than the initial energy, it is emitted as longer-wavelength photon. - Additional heat energy is lost as heat after the photon is emitted due to vibrational relaxation *(Figure 14.56)*. ## 14.4.06 Circular Dichroism - Circular dichroism *(CD)*, introduced briefly in Concept 14.4.04, is a spectroscopic technique that reports the overall secondary structure of proteins. - Like other UV/Vis absorbance techniques, CD depends on the ability of the analyte *(in this case, regions of protein secondary structure)* to absorb electromagnetic radiation. - CD differs from other UV/Vis absorbance techniques, however, because CD relies on the absorbance of circularly polarized light. - Circularly polarized light differs from linearly polarized light *(also known as plane-polarized light)*. - Plane-polarized light oscillates in a two-dimensional plane, whereas circularly polarized light propagates in a spiraling, helical fashion. - Circularly polarized light can be conceptually understood as the combination *(i.e.*, interference) of two rays of equal-amplitude light, in which one ray's polarization axis is both rotated 90° *(e.g.*, vertical and horizontally polarized rays) and phase-shifted by 90° with respect to the other ray, as illustrated in *(Figure 14.57)*. ## For more on circular polarization of light, see Physics Concept 4.3.05. - CD spectra of proteins uses circularly polarized light in the far-UV range *(~180-250 nm)*. - Light in this region is absorbed by the peptide bonds of the polypeptide backbone. - Backbone interactions *(e.g.*, interactions that form secondary structure elements) alter the absorbance spectrum; therefore, different secondary structure elements *(e.g.*, a-helices, β-sheets) absorb right-handed and left-handed circularly polarized light to different extents. - The CD spectrum of a protein plots the difference in the individual right- and left-handed absorbance spectra. - This measurement describes the ellipticity of the protein sample. - Representative CD spectra for secondary structural elements are shown in *(Figure 14.58)*. ## Most proteins have a mixture of secondary structural elements, so their CD spectra will be a linear combination of the individual spectra shown in *(Figure 14.58)*. - For example, consider a protein with a structure that is 50% a-helix, 40% β-sheet, and 10% random coil. - Its CD spectrum will reflect those ratios, such that at any given wavelength the ellipticity value will be equal to 0.5 times the a-helix value plus 0.4 times the β-sheet value plus 0.1 times the random coil value. - Detailed analysis of a mixed-structure spectrum is not likely to be required on the exam. - Instead, the it is more likely to test identification of a structure with a single secondary structural feature given the reference spectra. - In addition, the exam may test for an understanding of how a spectrum changes in response to conformational changes *(e.g.*, ligand binding, denaturation). ## 14.4.07 Structure Determination - Determination of protein structure can have great scientific and biomedical significance. - For example, identification of a protein's structural features at the atomic level can allow scientists to discover the role each amino acid residue plays in protein function. - Not only can this help in understanding and identifying diseases, but structural information can be used in the design of therapeutics to target those diseases *(Figure 14.59)*. - Currently, most high-resolution structures are experimentally determined by cryogenic electron microscopy *(cryo-EM)*, x-ray crystallography, or nuclear magnetic resonance *(NMR)*. - Recently, computational techniques have advanced and allowed artificial intelligence models to use the data from experimentally determined structures to predict structures of novel proteins with relatively high accuracy. - Although it is unlikely that the exam will ask questions about the technical details of these methods, a brief overview of each may be instructive for synthesizing information throughout biochemistry and other subjects. ## Cryogenic Electron Microscopy (Cryo-EM) - The resolution of a microscope *(i.e.*, its ability to "see" small objects) is dependent on the wavelength of radiation used. - Because protein particles are much smaller than even the shortest wavelength of visible light *(~400 nm)*, visible light cannot be used to visualize proteins. - Instead, electron microscopy uses high-energy electrons to irradiate samples. - Because of the wave-particle duality of matter, these energetic electrons can have a wavelength that as short as 2 pm *(2 × 10-12 m)* or smaller, which is small enough to visualize proteins at atomic resolution. - Unlike other high-energy *(short wavelength)* radiation *(e.g.*, gamma waves), for which the development of optical lenses is difficult, electron beams can be precisely focused due to their charged nature. - In cryo-EM, samples are suspended in ice and examined at low temperatures. - Freezing must happen rapidly to prevent the water molecules from forming ordered, hydrogen-bonded crystal lattices, which would destroy proteins. - The low temperature slows the damaging effects of the high-energy electron beam. - Microscopy generally allows a researcher to take a picture of a sample; however, this picture is flat and two-dimensional. - Therefore, if a three-dimensional structure of a purified protein is needed, multiple pictures must be taken of a sample from different angles. - If multiple protein particles in solution exist in random orientations, then multiple angles can be visualized from a single picture; otherwise, a sample can be physically rotated in the microscope to provide the angles needed. - Once two-dimensional images or projections—have been obtained from various angles via many different particles, then a three-dimensional model can be reconstructed from those projections, as shown in *(Figure 14.60)*. ## X-ray Crystallography - Like high-energy electrons, x-rays have a very short wavelength and can be used to determine atomic resolution structures of proteins. - As the name suggests, x-ray crystallography *(XRC)* involves the formation of crystals. - To form the needed crystals, a protein-containing solution is purified and then slowly concentrated *(often with agents to prevent denaturation)* until crystals appear. - The solid crystal makes it difficult to take a direct "picture" of a molecule. - Instead, crystallography relies on the scattering of x-rays as they encounter atoms. - Most scattered rays interfere destructively; however, the regular spacing of biomolecules present in a crystal allows for certain scattered rays to experience constructive interference. - This is similar to the concept of multiple-slit diffraction *(see Physics Lesson 4.3)*, and the interference pattern seen is called a diffraction pattern *(Figure 14.61)*. ## From the observed diffraction pattern, the three-dimensional shape of the protein can be determined. - One complication of interpreting crystal structures is that the crystallization procedure traps molecules as a solid crystal, whereas most biomolecules exist *in vivo* as dissolved aqueous solutes. - This may introduce crystallization artifacts that are not relevant for aqueous molecules. ## Nuclear Magnetic Resonance (NMR) - As introduced in Organic Chemistry Lesson 14.7, nuclear magnetic resonance is a spectroscopic technique that uses very-low-energy radiofrequency waves to determine molecular structure. - Unlike cryo-EM and XRC, radiofrequency waves have a wavelength that is too long to directly report on the physical position of atoms. - Instead, NMR measures the absorption of the photons' energy by nuclei in a magnetic field. - The amount of energy an individual nucleus can absorb varies depending on the structural features of the molecule *(e.g.*, the degree of electron shielding), allowing NMR to report on chemical structure. - To summarize the NMR method, the electronic environment surrounding a nucleus causes a shift in the frequency at the λmax *(known as a chemical shift)* relative to some reference sample *(e.g.*, tetramethylsilane *(TMS)*). - This shift is measured in parts per million *(ppm)*. - In addition, the interaction of nuclear spins across three σ bonds or fewer *(called through-bond interactions)* can be seen as coupling of the spins through spin-spin splitting. - The information gained from one-dimensional proton *(1H)* and carbon-13 *(13C)* NMR data can be used to help in the determination of the primary structure of small organic molecules. - As molecules grow in complexity *(e.g.*, proteins), two-dimensional *(2D)* NMR and heteronuclear NMR techniques *(e.g.*, techniques that assess nuclei other than 1H) can be used. - Two-dimensional NMR allows for the explicit identification of which peaks interact by measuring the effects of each peak in a 1D spectrum on each peak on another 1D spectrum *(Figure 14.62)*. ## Heteronuclear NMR allows the probing of interactions between different elements. - In protein NMR, 15N is commonly used to probe the peptide bond *(Figure 14.63)*. ## Chemical shift data in 2D NMR can be used to monitor ligand interactions and conformational changes. - An example of this is shown in *(Figure 14.63)*. - The region shown captures interactions between the amide nitrogen and protons of the peptide bond. - A particular amide peak is tracked as more ligand is added to the protein sample. - At low ligand concentrations, the peak appears at the intersection of approximately 105 ppm on the 15N 1D spectrum and 8.75 ppm on the 1H spectrum. - The peak position shifts until, at high ligand concentrations, it appears at 125 ppm on the 15N spectrum and 7.25 ppm on the 1H spectrum. - This indicates changes in the chemical environments of these atoms, and therefore a change in the protein conformation, as the protein binds its ligand. ## To determine tertiary and quaternary structure, however, a specialized form of NMR must be used that can report on through-space interactions *(i.e.*, interactions between amino acid residues that are not close to each other in the primary structure but become close when the protein folds). - Details of this procedure are unlikely to be tested on the exam; however, the other fundamental principles of conventional NMR still apply to NMR for protein structure. ## Modeling and Artificial Intelligence - Based on the structural data obtained through cryo-EM, XRC, and NMR, a robust library of protein structures has been collected over several decades. - Many proteins without experimentally determined structures can be modeled after similar proteins, based on the biochemical properties of amino acids and on analysis of sequence similarity. - With the growing capabilities of artificial intelligence, predicted protein structures have increased in accuracy, as verified by subsequent experimental method