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Description: Dialysis is a common technique used in biochemistry for separating molecules based on diffusion. In this procedure, a semipermeable membrane allows the ...

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Research

2D NMR: Heteronuclear Single-Quantum Correlation Spectroscopy (HSQC)

Heteronuclear single-quantum correlation spectroscopy (HSQC) is a 2D NMR technique that reveals one-bond correlations between hydrogen and a heteronucleus. The HSQC experiment is similar to the heteronuclear correlation experiment (HETCOR) but is more sensitive. In the HSQC spectrum, the proton chemical shift is plotted on the horizontal F2 axis, while the 13C chemical shift is plotted on the vertical F1 axis. The corresponding proton and 13C spectra are also shown. The HSQC contour plot does not have any diagonal peaks. All the cross-peaks indicate the protons directly attached to each carbon in a molecule. Similar to the COSY spectrum, imaginary lines are drawn in horizontal and vertical directions to identify the correlated peaks.
For example, the HSQC spectrum of 2-chlorobutane has a carbon signal at 11 ppm that can be assigned to the shielded methyl group and correlated to the proton multiplet at 1.01 ppm. The carbon signal at 24.8 ppm arises from the deshielded methyl group, which correlates to the proton multiplet at 1.52 ppm. Similarly, the methylene carbon signal at 33.3 ppm and methine carbon signal at 60.4 ppm correlate to the multiplets at 1.72 and 3.96 ppm, respectively.&#160; HSQC is a very useful technique for structurally elucidating complex molecules and is routinely used in protein NMR studies.

Heteronuclear single-quantum correlation spectroscopy (HSQC) is a 2D NMR technique that reveals one-bond correlations between hydrogen and a ...

2D NMR: Overview of Heteronuclear Correlation Techniques

Heteronuclear correlation spectroscopy is an analytical technique that investigates the coupling between different types of nuclei, often a proton and an X-nucleus, such as carbon-13 or nitrogen-15. This method is commonly used in nuclear magnetic resonance (NMR) spectroscopy to gain insights into complex chemical compounds&#39; structural and compositional aspects. A typical heteronuclear correlation spectrum displays X-nucleus chemical shifts on one axis and a proton spectrum on the other axis. Cross-peaks reveal the connection between specific protons and X-nuclei.
The main heteronuclear correlation experiments include HETCOR, HSQC, HMQC, and HMBC. HSQC, HMQC, and HMBC record the proton spectrum, whereas the X-nucleus spectrum is recorded by standard heteronuclear correlation spectroscopy or HETCOR. The HETCOR-HMQC-HSQC group of experiments cannot detect X-nuclei without an attached proton. HETCOR is used when ultrahigh peak resolution is required along the X-nucleus axis.
The heteronuclear single quantum correlation technique, or HSQC, investigates proton-to-X-nucleus single bond correlations. In an HSQC experiment, polarization is transferred from a proton nucleus to a neighboring X-nucleus and then back to the proton nucleus. The signal from the proton nuclei is then recorded. HSQC generates a spectrum like that of heteronuclear multiple-quantum correlation spectroscopy or HMQC but uses a different suppression method, providing better X-nucleus resolution along the axis.
Heteronuclear multiple-bond correlation, or HMBC, is a proton-detected experiment, like HMQC but with a longer initial delay time. This experiment is used to observe long-range proton to X-nucleus connectivity separated by 2-3 bonds, with some experiments going out to 4 or 5 bonds. In an HMBC experiment, direct one-bond correlations are suppressed as part of the sequence. The spectra of HMBC give rise to cross peaks that correlate which protons are coupled to what other specific X-nucleus is more than one bond away. The gradient HMBC (gHMBC) pulse sequence offers better suppression of interfering signals, making it particularly useful for analyzing complex molecules with overlapping signals.

Heteronuclear correlation spectroscopy is an analytical technique that investigates the coupling between different types of nuclei, often a proton and an ...

2D NMR: Homonuclear Correlation Spectroscopy (COSY)

Homonuclear correlation spectroscopy, or COSY, is a 2-dimensional NMR technique that provides information about coupled protons. Typically, the geminal and vicinal coupling are observed. For example, consider the COSY spectrum of ethyl acetate, where its 1D proton NMR spectrum is plotted along the vertical and horizontal axes with their corresponding chemical shift scale. Three spots on the diagonal corresponding to the three peaks in the 1D proton spectrum are called diagonal peaks. The COSY spectrum is symmetric about the diagonal, with signals above and below the diagonal called cross-peaks or off-diagonal peaks. An imaginary line is drawn from diagonal peaks parallel to the horizontal axis, leading to a cross peak. A vertical line is then drawn from the same cross peak towards the diagonal peak to correlate the peaks. The off-diagonal peaks correlating the multiplets at 1.25 and 4.2 ppm indicate that the methyl protons are coupled to the methylene protons. The signal at 2 ppm is not associated with cross-peaks because the protons are not involved in homonuclear coupling. A COSY spectrum is useful when overlapping multiplets complicate the corresponding 1D spectrum.

Homonuclear correlation spectroscopy, or COSY, is a 2-dimensional NMR technique that provides information about coupled protons. Typically, the geminal ...

Insensitive Nuclei Enhanced by Polarization Transfer (INEPT)

Insensitive Nuclei Enhanced by Polarization Transfer (INEPT) is an advanced Nuclear Magnetic Resonance (NMR) technique specifically designed to detect and enhance the signals of low-abundance nuclei, such as carbon-13 and nitrogen-15, in small molecules. The fundamental principle behind INEPT is the transfer of polarization from a more abundant and highly polarizable nucleus, typically hydrogen-1, to the low-abundance nucleus of interest. This process effectively boosts the NMR signal of the low-abundance nuclei, making them more easily detectable.
The INEPT technique involves a series of carefully timed radiofrequency (RF) pulses applied to the sample. These RF pulses excite the more abundant nuclei, causing them to align their spins in a specific manner. Through the process of spin coupling, the polarization of the more abundant nuclei is then transferred to the low-abundance nuclei. As a result, the NMR signal of these low-abundance nuclei is significantly enhanced, allowing for a more detailed analysis of the molecular structure.
Various pulse sequences have been developed to improve the observability of these low-abundance nuclei when coupled to a high-receptivity nucleus. The INEPT pulse sequence combines a spin echo and selective population inversion, which includes 90&#176; and 180&#176; pulses on the high-sensitivity (the I spins) and low-sensitivity nuclei (the S spins), respectively. Coupling-constant-based delays between pulses facilitate magnetization transfer through space or bond interactions. Additional RF pulses and delays invert the low-gamma magnetization, producing an observable NMR signal. The net result is an enhanced NMR signal from low-gamma nuclei that would otherwise be challenging to detect.
Various pulse sequences have been developed to improve the detection of low-abundance nuclei when coupled with high-receptivity nuclei such as protons. The INEPT sequence yields enhanced signals for insensitive nuclei, with half of the CH group producing negative peaks and the other half positive peaks. This pattern allows for discrimination between different types of carbon-hydrogen bonds within the molecule.
Decoupling methods can be employed to enhance sensitivity and resolution in distinguishing various carbon-hydrogen bonds, making INEPT an invaluable tool in the chemical and pharmaceutical industries.

Insensitive Nuclei Enhanced by Polarization Transfer (INEPT) is an advanced Nuclear Magnetic Resonance (NMR) technique specifically designed to detect and ...

Two-Dimensional (2D) NMR: Overview

The 1D NMR spectrum of large and complex molecules like natural products has complicated splitting patterns and overlapping signals, which can be easily interpreted using 2-dimensional (2D) NMR. Unlike 1D NMR, 2D NMR has two frequency axes that provide the coupling information between the nucleus A and nucleus B in a molecule. The process from which 2D spectra are obtained has four steps.
The first step is the preparation period, during which nucleus A is excited with a radiofrequency pulse. The second step is the evolution or wait period t1, which is incremented systematically, and no data is observed during this period. The next step is the mixing period, in which a second radiofrequency pulse is introduced, which transfers the magnetization to the spins of nucleus B. In the last step, the magnetization of nuclear spins from B is detected during the acquisition time t2 for each t1. Time domains t1 and t2 are Fourier transformed into frequency domains f1 and f2, and these correspond to two dimensions of 2D spectra, while the third dimension is the intensity. In 2D experiments, coupling between the same kind of nucleus is called homonuclear 2D experiments, whereas coupling between two types of nucleus is called heteronuclear 2D experiments. 2D spectra can be displayed as stacked or contour plots. When presented as stacked plots, 2D spectra are difficult to interpret. The contour plot is the horizontal cross-section of a stacked plot where the size of the circles shows the signal intensity. Its interpretation is straightforward and it is preferred for data analysis.

The 1D NMR spectrum of large and complex molecules like natural products has complicated splitting patterns and overlapping signals, which can be easily ...

2D NMR: Overview of Homonuclear Correlation Techniques

Homonuclear correlation spectroscopy (COSY) is a powerful technique used in Nuclear Magnetic Resonance (NMR) spectroscopy to study the correlations between nuclei of the same type within a molecule. It provides information about scalar couplings between adjacent nuclei, which helps determine connectivity and structural information. There are several COSY variants, each with its unique strengths and experimental parameters.
COSY90 is the standard two-dimensional (2D) COSY experiment that utilizes a second 90-degree pulse to achieve high sensitivity and resolution. It excels in analyzing small to medium-sized molecules and provides information on directly coupled protons. COSY45, similar to COSY90, uses 45-degree pulses instead of 90-degree pulses. This results in reduced sensitivity but improved resolution.
Long-range COSY (LR-COSY) extends the COSY experiment to detect scalar couplings between nuclei further apart in the molecule. It achieves this by combining multiple pulse sequences and magnetization transfers.
Relayed COSY is a variant of COSY that adds extra pulses to the standard pulse sequence to produce cross-peaks between cross-peaks, showing specific molecular connections hidden by overlapping signals.
TOCSY (Total Correlation Spectroscopy), also known as HOHAHA (Homonuclear Hartmann-Hahn Spectroscopy), reveals all through-bond connectivity between all coupled nuclei. TOCSY is especially useful for analyzing molecules with overlapping signals and complex coupling patterns.
NOESY (Nuclear Overhauser Effect Spectroscopy) measures the nuclear overhauser effect, which arises from dipole-dipole interactions between nuclei. It provides information about spatial proximity and can be used to determine the three-dimensional structure of a molecule.
ROESY (Rotating-frame Overhauser Enhancement Spectroscopy) is similar to NOESY but employs a different mechanism to obtain through-space correlations. It provides information about spatial proximity and is particularly useful for studying molecules with weak NOE signals.
In conclusion, choosing a COSY variant depends on the molecule or reaction under investigation and the desired information. The right COSY variant can be determined based on molecular size, coupling complexity, and the need for spatial or long-range connectivity data.

Homonuclear correlation spectroscopy (COSY) is a powerful technique used in Nuclear Magnetic Resonance (NMR) spectroscopy to study the correlations ...

Double Resonance Techniques: Overview

Double resonance techniques in Nuclear Magnetic Resonance (NMR) spectroscopy involve the simultaneous application of two different frequencies or radiofrequency pulses to manipulate and observe two distinct nuclear spins. One important application of double resonance is spin decoupling, which selectively suppresses coupling with one type of nucleus while observing the NMR signal from another nucleus, simplifying the spectrum and enhancing resolution.
Spin decoupling is usually achieved by irradiating the sample atoms with a suitable radio frequency (rf) pulse sequence, effectively eliminating all the coupling with one nuclide. This simplifies the observed spectrum, making it easier to analyze and understand the relationships between different nuclei.
Double resonance experiments can be classified as heteronuclear or homonuclear, depending on whether the two sets of nuclei are of different isotopes or the same isotope, respectively. They can also be selective or nonselective, depending on whether the irradiating frequency covers only a portion or all of the resonance frequencies.
In nonselective heteronuclear decoupling, samples are exposed to an appropriate radiofrequency range to remove all coupling with one nuclide. However, increased external field strengths necessitate more powerful irradiation across broader frequency ranges. In certain instruments, continuous irradiation creates enough heat to cause damage to thermally sensitive samples.
To overcome these problems, modern methods using a series of pulses and exact timing delays, like J-modulated spin echo pulse sequences, can be employed to eliminate or adjust coupling effects within the spectrum.
For example, the attached proton transfer (APT) experiment applies a J-modulated spin echo pulse sequence to focus on the phase of detected carbon signals. Carbon atoms attached to an even number of protons exhibit positive signals in the spectrum, while those bonded to an odd number of protons appear as negative signals.
It utilizes a combination of a 180&#176; proton pulse and broadband decoupling to simplify spectral interpretation and assign multiplicities. The 180&#176; pulse is applied to the protons during the APT experiment. Its role is to refocus the spin coupling evolution that naturally occurs due to interactions between protons and carbons.
Specifically, after the 90&#176; pulse creates transverse magnetization, the 180&#176; pulse ensures that the effects of coupling are frozen during the initial period where the decoupler is turned off. This means the chemical shifts evolve only during the time 1/J&#8203; (the coupling constant period) after the first 180&#176; pulse.
By carefully timing this pulse sequence, the experiment isolates carbon multiplicities by utilizing the J-coupling between carbons and their attached protons. After the coupling evolution period (1/J&#8203;), broadband decoupling is turned on. This removes the J-coupling signals, collapsing multiplets into single peaks for each carbon resonance.
The decoupling ensures that carbon signals are free of splitting, allowing for a clear differentiation of signals based on phase (positive or negative) rather than splitting patterns.
The APT technique selectively enhances carbon signals, providing valuable information about carbon-proton connectivity, the number of attached hydrogen atoms, and the overall molecular structure.

Double resonance techniques in Nuclear Magnetic Resonance (NMR) spectroscopy involve the simultaneous application of two different frequencies or ...

&sup1;H NMR of Labile Protons:  Deuterium (&sup2;H) Substitution

This lesson illustrates the role of deuterium substitution in simplifying the NMR spectrum of compounds comprising labile protons. One method employed is the use of deuterium. Amongst the three isotopes of hydrogen, deuterium (2H) has a nucleus composed of one proton and one neutron. When the D2O solvent is added to a pure dry ethanol solution, its labile proton is substituted with deuterium.
<img alt="Figure1" src="/files/ftp_upload/12280/12280-PT_Figure_1.svg" style="text-align: center; height: 700px;" />
Figure 1. The comparison of the proton NMR spectra of pure dry ethanol and the same in D2O solvent.
The signals from deuterium are not observed in a proton NMR spectrum, as deuterium&#8211;proton coupling is negligible, and no spin-spin splitting occurs between them. As shown in Figure 1, the NMR spectra of pure ethanol and ethanol in deuterium, the disappearance of the triplet in the latter, confirms that it corresponds to the labile &#8211;OH proton. Furthermore, the methylene protons that initially showed up as a multiplet transform to a quartet, proving that the multiplet is due to the coupling with the &#8211;OH proton and the methyl protons.

This lesson illustrates the role of deuterium substitution in simplifying the NMR spectrum of compounds comprising labile protons. One method employed is ...

Nuclear Overhauser Enhancement (NOE)

Irradiation of a spin-active nucleus causes an increase or decrease in the signal intensity of neighboring nuclei that are not necessarily chemically bonded or involved in J-coupling. &#160;This phenomenon, called the Nuclear Overhauser Enhancement (NOE), results from through-space interactions between the nuclear spins. The NOE effect decreases with increasing internuclear distance and is generally not observed beyond 4 angstroms. In NOE, dipole-dipole interactions between neighboring spin-active nuclei result in the transfer of nuclear spin polarization. These interactions can cause an increase or decrease in the signal intensity, which results in a positive or negative NOE, respectively.
NOE can be applied to improve the signal intensity of less sensitive nuclei such as carbon-13. Proton-decoupled carbon-13 spectra show a positive NOE effect, where the carbon signal intensities are substantially increased compared to those in proton-coupled spectra.
Targeted irradiation of specific nuclei can also reveal through-space proximity, which helps confirm stereochemistry and verify spectral peak assignments. The NOE effect is also applied in 2D NMR experiments, such as Nuclear Overhauser Effect Spectroscopy (NOESY), to determine the 3D structure of proteins.

Irradiation of a spin-active nucleus causes an increase or decrease in the signal intensity of neighboring nuclei that are not necessarily chemically ...

&sup1;H NMR of Labile Protons: Temporal Resolution

Protons bonded to heteroatoms such as nitrogen and oxygen exhibit a range of chemical shift values. This is due to the varying degree of hydrogen bonding between the proton and the heteroatom in other molecules. The extent of hydrogen bonding affects the electron density around the proton, thereby giving different chemical shift values for the protons in the proton NMR spectrum.
The &#8211;OH proton in alcohols typically appears in the range of &#948; 2 to 5 ppm but can vary depending on the specific alcohol and conditions. This &#8211;OH proton undergoes rapid proton exchange with other molecules and is referred to as a labile proton. So, the exact &#948; value of the &#8211;OH proton and the spin-spin coupling involving the &#8211;OH proton and the other protons in the alcohol depends on the rate of the &#8211;OH proton exchange. The proton exchange rate, in turn, depends on multiple factors, including the purity of the alcohol sample, temperature, and solvent.
<img alt="Figure1" src="/files/ftp_upload/12279/12279-PT_Figure_1.svg" style="text-align: center; height: 350px;" />
Figure 1. The triplet of a &#8211;OH proton in the proton NMR spectrum of neat ethanol.
For example, the &#8211;OH proton exchange in a pure dry ethanol sample is very slow on the NMR time scale, giving enough time for the spin-spin coupling. Consequently, as depicted in Figure 1, the proton NMR spectrum of pure dry ethanol shows a triplet for the &#8211;OH proton due to the spin-spin coupling with adjacent methylene protons. The methylene protons are seen as a multiplet due to the coupling with the &#8211;OH proton and the methyl protons.
The presence of trace amounts of an acid or base increases the proton exchange rate. The proton exchange becomes rapid, and the NMR spectrometer only records an average of all the possible environments for the &#8211;OH proton.
<img alt="Figure2" src="/files/ftp_upload/12279/12279-PT_Figure_2.svg" style="text-align: center; height: 325px;" />
Figure 2. The broad singlet of a &#8211;OH proton in the proton NMR spectrum of neat ethanol.
As a result, as recorded in Figure 2, the &#8211;OH proton exhibits a broad singlet. The protons from the adjacent methylene group present a quartet where the splitting is caused only by the adjoining methyl group.

Protons bonded to heteroatoms such as nitrogen and oxygen exhibit a range of chemical shift values. This is due to the varying degree of hydrogen bonding ...

&sup1;H NMR of Conformationally Flexible Molecules: Variable-Temperature NMR

The axial and equatorial protons in cyclohexane can be distinguished by performing a variable-temperature NMR experiment. In this process, except for one proton, the remaining eleven protons are replaced by deuterium. The deuterium substitution avoids the possible peak splitting caused by the spin-spin coupling between the adjacent protons. The remaining proton flips between the axial and equatorial positions.
<img alt="Figure1" src="/files/ftp_upload/13016/13016-PT_Figure_1.svg" style="text-align: center; height: 480px;" />
Figure 1. The temperature-dependent proton NMR spectra of cyclohexane.
Figure 1 depicts the proton NMR spectrum of the deuterium substituted sample, recorded at various temperatures. At room temperature, the rapid ring flipping of cyclohexane results in a single sharp peak. As the temperature is lowered to &#8722;60&#176;C, the rate of chair-chair interconversion reduces, resulting in the broadening of the peak.
Further lowering of the temperature broadens the peak, forming a saddle that splits into two peaks. On reaching a temperature lower than &#8722;89&#176;C, two sharp, well-resolved peaks corresponding to the equatorial and axial protons are formed. The peaks corresponding to the equatorial and axial protons occur at &#948; 1.62 and 1.14, respectively. At such low temperatures, the rate of chair-chair interconversion is reduced significantly, enabling the detection of the two distinct sets of protons on the NMR time scale.

The axial and equatorial protons in cyclohexane can be distinguished by performing a variable-temperature NMR experiment. In this process, except for one ...

&sup1;H NMR of Conformationally Flexible Molecules: Temporal Resolution

At room temperature, the chair conformer of cyclohexane undergoes rapid ring flipping between two equivalent chair conformers at a rate of approximately 105 times per second. These two chair conformers are in equilibrium. The rapid ring flipping results in the interconversion of the axial proton to an equatorial proton and an equatorial to the axial proton. Such interconversions are too rapid and cannot be detected on the NMR timescale. Hence, the NMR spectrometer cannot distinguish between the axial and equatorial protons, resulting in a single peak at &#948; 1.4.
Similarly, NMR averages all the conformations of any molecule with rapid conformational equilibria. For example, the CH3 protons in bromoethane have a single resonance and coupling constant for splitting by the CH2 protons due to the rapid internal rotation of the carbon-carbon bond. Lastly, besides conformational equilibria, the time-averaging effect of NMR spectroscopy is also observed in some chemical reactions.

At room temperature, the chair conformer of cyclohexane undergoes rapid ring flipping between two equivalent chair conformers at a rate of approximately ...

Mass Spectrometry: Carboxylic Acid, Ester, and Amide Fragmentation

The fragmentation patterns observed for compounds such as carboxylic acids, esters, and amides in the mass spectra include &#9082;-cleavage and McLafferty rearrangement. Fragmentation by &#9082;-cleavage preferentially occurs at the carbon-carbon bond at the &#9082;-position next to the carboxylic group to generate a neutral radical and a cation. Long chain compounds with hydrogen at their &#947;-carbon undergo McLafferty rearrangement to give a radical cation and a neutral alkene.
For example, the fragmentation of butyramide is demonstrated in Figure 1. The molecular ion peak of butyramide, which contains one nitrogen atom, shows an odd mass-to-charge ratio of 87. Fragmentation of butyramide molecular ion by &#9082;-cleavage produces a cation at a mass-to-charge ratio 44. In addition to &#9082;-cleavage, butyramide molecular ion also undergoes fragmentation via McLafferty rearrangement. The base peak at the mass-to-charge ratio 59 arises due to McLafferty rearrangement.
<img alt="Figure1" src="/files/ftp_upload/13051/13051_PT_Figure_1.svg" style="text-align: center; height: 250px;" />

The fragmentation patterns observed for compounds such as carboxylic acids, esters, and amides in the mass spectra include &#9082;-cleavage and McLafferty ...

Chemical Ionization (CI) Mass Spectrometry

The molecular ion peak of a molecule in the mass spectrum provides vital information for molecular identification. However, conventional electron impact ionization can lead to the rapid dissociation of some molecular ions before they reach the detector. A milder ionization method is required to increase the lifetime of such ionized analyte molecules. Chemical ionization (CI) is a gas-phase protonation reaction useful for mass-analyzing analyte molecules that are easily protonated to yield the corresponding conjugate acid. In this process, the sample is mixed with an excess reagent gas, which ensures the electron impact occurs primarily on the reagent gas. The charged species formed from the reagent protonates the analyte molecule, producing a relatively stable protonated analyte (conjugate acid) compared to the molecular ion. This results in an M+1 peak in the mass spectrum. The conjugate acid may then undergo fragmentation that generates additional signals.
For example, the CI of di-sec-butyl ether in methane gas illustrates this process. When the ether is mixed with excess methane gas as a reagent, electron impact occurs on methane rather than the ether. The resulting methane radical cation can react with another methane molecule to generate a methane radical and a methanium ion. The methanium ion is a source of gas-phase protons, which can protonate the ether to form its conjugate acid. The sequential reactions during the chemical ionization of di-sec-butyl ether are depicted in Figure 1.
<img alt="Figure1" src="/files/ftp_upload/12300/12300_PT_Figure_1.svg" style="text-align: center; height: 175px;" />
Figure 1: Chemical ionization of di-sec-butyl ether methane mixture.
This conjugate acid of the ether (m/z = 131) is relatively more stable than the molecular ion (m/z = 130) of ether formed via the conventional electron impact ionization. In the traditional route, the molecular ion undergoes fragmentation via &#945; cleavage, giving a signal at m/z = 101. Figure 2 illustrates the reactions that occur during electron impact ionization directly on di-sec-butyl ether.
<img alt="Figure2" src="/files/ftp_upload/12300/12300_PT_Figure_2.svg" style="text-align: center; height: 175px;" />
Figure 2: Electron impact ionization of di-sec-butyl ether and fragmentation of the molecular ion.
So, the mass spectra of di-sec-butyl ether ionized via CI feature an M+1 peak at m/z = 131. On the other hand, mass spectra of di-sec-butyl ether ionized via electron impact ionization do not show any peak at the m/z value of its molecular weight. Figures 3a and 3b show the mass spectra of di-sec-butyl ether ionized via the electron impact and chemical ionization methods, respectively.
<img alt="Figure3" src="/files/ftp_upload/12300/12300_PT_Figure_3.svg" style="text-align: center; height: 225px;" />
Figure 3:&#160;a) No m/z = 130 peak is visible in the mass spectrum obtained via electron impact ionization of di-sec-butyl ether. b) The m/z = 131 peak is clearly visible in the mass spectrum obtained via chemical ionization of di-sec-butyl ether.

The molecular ion peak of a molecule in the mass spectrum provides vital information for molecular identification. However, conventional electron impact ...

Mass Spectrometry: Alkyl Halide Fragmentation

Chlorine isotopes exist as 35Cl and 37Cl in a 3:1 ratio, while bromine isotopes exist as 79Br and 81Br in a 1:1 ratio. The mass spectrum of alkyl halides typically produces two distinct molecular ion peaks, the molecular ion peak, [M], and the molecular ion plus two, [M + 2] peak. The relative heights of these two peaks are proportional to the isotopic abundance ratios of the halide. For example, 2&#8208;chloropropane and 1&#8208;bromopropane display two peaks with relative peak heights in a 3:1 and 1:1 ratio, respectively.
Alkyl halides typically lose a non-bonding electron from the halogen to form molecular ions. The base peak results from the heterolytic cleavage of the carbon&#8211;halogen bond, where the bonding electrons go to the most electronegative atom, forming a cation and halogen atom. Interestingly, the &#9082;-cleavage is observed only in alkyl chlorides, not alkyl bromides. In alkyl chlorides, the&#160; C&#8211;Cl and C&#8211;C bonds exhibit similar strengths, while in alkyl bromides, the C&#8211;Br bond is weaker than the C&#8211;C bond. Consequently, the alkyl chlorides form a stable cation, where the positive charge is shared between the two atoms.
While less commonly discussed, fluoroalkanes and iodoalkanes exhibit unique fragmentation behavior. Fluoroalkanes, due to the strength of the C&#8211;F bond and the presence of only one isotope (19F), show limited fragmentation and lack isotopic peaks. Iodoalkanes, with their weak C&#8211;I bond and a single isotope (127I), predominantly fragment by losing iodine, resulting in simpler spectra with less isotopic splitting.

Chlorine isotopes exist as 35Cl and 37Cl in a 3:1 ratio, while bromine isotopes exist as 79Br and 81Br in a 1:1 ratio. The mass spectrum of alkyl halides ...

Electrospray Ionization (ESI) Mass Spectrometry

Higher molecular weight biomolecules are nonvolatile compounds that may decompose before ionizing or vaporizing during mass analysis with conventional electron impact ionization methods. Accordingly, electrospray ionization (ESI) is the favored method for vaporizing and ionizing biomolecules as it circumvents rapid fragmentation and enables the recording of mass signals for the entire biomolecule.
ESI utilizes electrical energy to transfer ions from the liquid phase of the sample into the gaseous phase. The analyte biomolecule is premixed with an ionic liquid&#8212;a nonmolecular compound serving as the ionization source. The solution is sprayed through a high-voltage capillary to generate a fine aerosol. The voltage difference applied between the capillary and the entrance of the analysis chamber causes the movement of charged droplets toward the analysis chamber. The ionic liquid from the aerosol evaporates during this movement, leaving behind a variably protonated or ionized analyte molecule. These charged species are detected in the mass spectrum at an m/z value equivalent to the sum of the analyte molecule&#39;s molecular mass and the ions&#39; molecular weight from the ionic liquid associated with the analyte molecule.
The position of mass signals varies depending on the type of cations present in the ionic liquid. For example, the presence of sodium ions in the ionic liquid leads to an (M+23) peak in the mass spectra, while the presence of a proton results in an (M+1) peak.

Higher molecular weight biomolecules are nonvolatile compounds that may decompose before ionizing or vaporizing during mass analysis with conventional ...

Matrix-Assisted Laser Desorption Ionization (MALDI)

Matrix-assisted laser desorption ionization (MALDI) is a powerful analytical technique used in mass spectrometry. It enables the identification and characterization of various biomolecules, including proteins, peptides, nucleic acids, and carbohydrates. MALDI spectrometry is widely employed in biological and medical research, as well as in fields like pharmacology and biochemistry.
The analyte of interest, a biomolecule or a mixture of biomolecules, is mixed with a suitable matrix material. The matrix is usually an organic compound, such as a small organic acid or a crystalline substance, that facilitates ionization. This analyte-matrix mixture is then deposited onto a target surface, inserted into the mass spectrometer, and dried. During drying, the matrix forms small crystals that incorporate the analyte molecules within their structure.
Once the sample is prepared, a pulsed laser is directed onto the target surface. The matrix molecules absorb the laser beam energy, causing them to heat up and vaporize rapidly. This process is known as laser desorption. As the matrix molecules vaporize, they carry the embedded analyte molecules. The vaporized analyte molecules become ionized through a process called proton transfer. The proton transfer occurs between the matrix and the analyte molecule. The matrix molecule donates a proton to the analyte molecule, resulting in the formation of protonated ions. The analyte ions are then accelerated by an electric field and further separated based on their mass-to-charge ratio in a time-of-flight mass analyzer.
The mass spectrum obtained from MALDI spectrometry provides valuable information about the mass and relative abundance of the analyzed biomolecules.

Matrix-assisted laser desorption ionization (MALDI) is a powerful analytical technique used in mass spectrometry. It enables the identification and ...

Mass Spectrometry: Aldehyde and Ketone Fragmentation

In mass spectrometry, the fragmentation of aliphatic aldehydes and ketones generally occurs through three key mechanisms: &#945;-cleavage, inductive cleavage, and the McLafferty rearrangement.
<ol>
	<li>&#945;-Cleavage 
	&#945;-Cleavage is a common fragmentation in carbonyl compounds where the bond adjacent to the carbonyl group is broken. This process produces an alkyl radical and an acylium cation, which is often detected in the mass spectrum due to its stability.</li>
	<li>Inductive Cleavage 
	In inductive cleavage, fragmentation yields an acyl radical and an alkyl cation. This occurs when electrons are drawn toward the carbonyl, weakening adjacent bonds and resulting in a distinctive fragmentation pattern. For example, in the mass spectrum of 5-methyl-2-hexanone, inductive cleavage produces an alkyl cation at a mass-to-charge (m/z) ratio of 71.</li>
	<li>McLafferty Rearrangement 
	The McLafferty rearrangement is a specific fragmentation pattern for carbonyl compounds with &#947;-hydrogen (hydrogen on the third carbon from the carbonyl group). The molecule undergoes a six-membered cyclic transition in this rearrangement, yielding a radical cation and a neutral alkene. For 5-methyl-2-hexanone, this rearrangement results in a peak at m/z 58.</li>
</ol>
Example: Fragmentation of 5-methyl-2-hexanone 
In the mass spectrum of 5-methyl-2-hexanone, the molecular ion undergoes these fragmentation patterns:
<ol style="list-style-type: circle;">
	<li>Inductive cleavage produces an alkyl cation at m/z 71.</li>
	<li>&#945;-Cleavage results in the base peak at m/z 43.</li>
	<li>McLafferty rearrangement creates a distinct peak at m/z 58.</li>
</ol>
An additional diagnostic feature in aldehydes is the M&#8722;1 peak, which arises due to the &#945;-cleavage of the aldehyde proton. In mass spectrometric analysis, this peak can help identify aldehydes, specifically among other carbonyl compounds.
Understanding these fragmentation pathways is crucial for interpreting mass spectra, particularly in identifying functional groups and distinguishing between similar compounds.

In mass spectrometry, the fragmentation of aliphatic aldehydes and ketones generally occurs through three key mechanisms: &#945;-cleavage, inductive ...

Mass Spectrometry: Alcohol Fragmentation

Alcohols (R-OH) ionize to lose one non-bonded electron from the oxygen atom, forming molecular ions. Due to their tendency to fragment rapidly, the intensity of the molecular ion peak in the mass spectrum is weak or sometimes absent. The fragmentation patterns for alcohols occur in two ways, i.e. &#9082;-cleavage and dehydration. During &#9082;-cleavage, the bond at the &#9082;-position adjacent to the hydroxyl group cleaves to give a resonance-stabilized cation and a radical. However, intramolecular dehydration involves losing a water molecule from an alcohol, forming an alkene cation, which shows a peak at M&#8722;18. Consider the fragmentation of butanol as shown below. &#9082;-cleavage produces a base peak at a mass-to-charge ratio of 31. In the case of dehydration, the fragmentation forms an alkene radical cation at a mass-to-charge ratio of 56.
<img alt="Figure1" src="/files/ftp_upload/13046/13046-PT_Figure_1.svg" style="text-align: center; height: 175px;" />

Alcohols (R-OH) ionize to lose one non-bonded electron from the oxygen atom, forming molecular ions. Due to their tendency to fragment rapidly, the ...

Mass Spectrometry: Amine Fragmentation

Amines can be identified using mass spectroscopy based on their characteristic fragmentation patterns. The molecular ions of amines undergo fragmentation via &#9082;-cleavage. The &#9082;-cleavage of the carbon-carbon bonds in amines generates an alkyl radical and resonance-stabilized nitrogen-containing cation.
In amines, the number of nitrogen atoms affects the mass of the molecular ion, which is described by the nitrogen rule of mass spectrometry. This rule states that a compound containing a single or an odd number of nitrogen atoms produces a molecular ion with an odd molecular weight, while a compound with no or an even number of nitrogen atoms generates a molecular ion with an even molecular weight.
For example, the mass spectrum of 3-methyl-1-butanamine, which contains one nitrogen atom, displays a molecular ion having an odd mass-to-charge ratio of 87. Fragmentation of this molecular ion via &#9082;-cleavage produces a base peak, which features a mass signal at a mass-to-charge ratio of 30, corresponding with the loss of the 2-methyl propyl radical.
Similarly, the fragmentation of triethylamine via &#9082;-cleavage generates a cation at a mass-to-charge ratio of 86. The molecular ion peak shows an odd mass-to-charge ratio of 101, as shown below.
<img alt="Figure1" src="/files/ftp_upload/13048/13048_PT_Figure_1.svg" style="text-align: center; height: 105px;" />

Amines can be identified using mass spectroscopy based on their characteristic fragmentation patterns. The molecular ions of amines undergo fragmentation ...

Mass Spectrometry: Cycloalkene Fragmentation

The molecular ions of cycloalkenes undergo fragmentation via a retro-Diels&#8211;Alder reaction.

The reaction proceeds via the cleavage of two carbon-carbon bonds in the cycloalkene to yield ethene. The remaining part of the cycloalkene structure is a dienyl radical cation with a molecular weight of 28 u lower than the molecular ion. This fragmentation pathway is similar to the fragmentation of cycloalkanes releasing ethene, differing only in the resultant radical cation. An alkyl species forms from the cycloalkane, whereas a dienyl species originates from the cycloalkene.

Retro-Diels&#8211;Alder reactions of cycloalkenes are also similar to the McLafferty-rearrangement-type fragmentation observed in acyclic alkenes with hydrogen on the &#947; carbon. In both cases, cleavage occurs at two positions, yielding an ethene molecule and an alkenyl species. In acyclic alkenes, the cleavage occurs at one carbon-carbon bond and a carbon-hydrogen bond, along with rearranging hydrogen from one carbon to another. In cyclic alkenes, cleavage occurs at two carbon&#8211;carbon bonds without any atomic rearrangement. Figure 1 depicts the fragmentation of cyclohexene via a retro-Diels&#8211;Alder reaction.
<img alt="Figure1" src="/files/ftp_upload/13044/13044-PT_Figure_1.svg" style="text-align: center; height: 90px;" />
Figure 1: Fragmentation of cyclohexene via a retro-Diels&#8211;Alder reaction.
In addition to the retro-Diels&#8211;Alder reaction, branched cycloalkenes exhibit fragmentation via cleavage of the side chain to form a cycloalkene cation. For example, the fragmentation of 1-methyl-1-cyclohexene via a retro-Diels&#8211;Alder reaction and side-chain cleavage are shown in Figure 2.
<img alt="Figure2" src="/files/ftp_upload/13044/13044-PT_Figure_2.svg" style="text-align: center; height: 225px;" />
Figure 2: Fragmentation of 1-methyl-1-cyclohexene via (top) a retro-Diels&#8211;Alder reaction; (bottom) side-chain cleavage.
The corresponding signals of the isoprenyl radical cation and the cyclohexenyl cation are evident in the mass spectrum of 1-methyl-1-cyclohexene shown in Figure 3.
<img alt="Figure3" src="/files/ftp_upload/13044/13044-PT_Figure_3.svg" style="text-align: center; height: 250px;" />
Figure 3: Mass spectrum of 1-methyl-1-cyclohexene.

The molecular ions of cycloalkenes undergo fragmentation via a retro-Diels&#8211;Alder reaction.

The reaction proceeds via the cleavage of two ...

Mass Spectrometry: Alkyne Fragmentation

The fragmentation of alkynes preferentially occurs at the carbon&#8211;carbon bond between the &#945; and &#946; carbon of the alkyne bond to generate a 3-propynyl cation (or propargyl cation). In terminal alkynes, there is the only type of fragmentation that yields the 3-propynyl cation. The unsubstituted 3-propynyl cation exhibits a peak at a mass-to-charge ratio of 39. In internal alkynes, the 3-propynyl cation is substituted. For example, 2-pentyne fragments into methyl-substituted 3-propynyl cation, which subsequently features as a mass signal at the mass-to-charge ratio of 56. The 3-propynyl cation is resonance-stabilized, and the corresponding mass signal is intense in the mass spectrum of alkynes. Figure 1 shows the fragmentation of 1-pentyne into an ethyl radical and a resonance stabilized 3-propynyl cation.
<img alt="Figure1" src="/files/ftp_upload/13045/13045-PT_Figure_1.svg" style="text-align: center; height: 115px;" />
Figure 1: Fragmentation of 1-pentyne molecular ion to yield the 3-propynyl cation.
Another fragmentation pathway observed in terminal alkynes is the loss of hydrogen from the terminal sp carbon. This results in a mass signal with a mass-to-charge ratio of one unit less than the molecular ion. This M&#8211;1 peak is very intense and becomes the base peak in the mass spectra of many alkynes, so it can be used to identify alkynes.
<img alt="Figure2" src="/files/ftp_upload/13045/13045-PT_Figure_2.svg" style="text-align: center; height: 275px;" />
Figure 2: The mass spectrum of 1-pentyne.

The fragmentation of alkynes preferentially occurs at the carbon&#8211;carbon bond between the &#945; and &#946; carbon of the alkyne bond to generate a ...

Mass Spectrometry: Aromatic Compound Fragmentation

Upon ionization, aromatic compounds generate a molecular ion that is observed as a prominent peak in their mass spectra. For example, the molecular ion peak for benzene appears at a mass-to-charge ratio of 78, while toluene is observed at a mass-to-charge ratio of 92. The molecular ion benzene is highly stable and does not readily undergo further fragmentation due to the significant amount of energy required to disrupt the aromatic stability of the benzene ring. In contrast, the molecular ion of alkyl&#8208;substituted benzenes, such as toluene, can undergo fragmentation by losing a hydrogen atom, forming a benzylic carbocation. This carbocation can rearrange to form the resonance-stabilized tropylium cation, which appears in the mass spectra as a strong peak. In the case of toluene, this strong peak is observed at a mass-to-charge ratio of 91.

Upon ionization, aromatic compounds generate a molecular ion that is observed as a prominent peak in their mass spectra. For example, the molecular ion ...

Mass Spectrometry: Long-Chain Alkane Fragmentation

The molecular ions of linear alkanes prefer to fragment at the carbon-carbon bond away from the end of the chain since the cleavage of an inner bond creates a stable carbocation and a stable radical. Consequently, the mass signals of linear alkanes feature intense peaks in the middle of the mass-to-charge ratio plot with weaker peaks on either end. The fragmentation of each carbon-carbon bond with the release of a methyl group in each splitting leads to prominent peaks in the mass spectra separated by 14 u. Smaller mass signals near each prominent peak result arise from secondary fragmentation at the carbon-hydrogen bond.
For example, consider the fragmentation pattern of n-hexane shown in Figure 1 and the corresponding mass spectra in Figure 2.
<img alt="Figure1" src="/files/ftp_upload/12299/12299-PT_Figure_1.svg" style="text-align: center; height: 500px;" />
Figure 1: Fragmentation of n-hexane molecular ion.
<img alt="Figure2" src="/files/ftp_upload/12299/12299-PT_Figure_2.svg" style="text-align: center; height: 275px;" />
Figure 2: Mass spectrum of n-hexane.
The signals at a mass-to-charge ratio of 71, 57, 43, 29, and 15 indicate that the cleavage can occur at all carbon-carbon bonds in the molecule. However, the relative abundance of these peaks varies. The difference between molecular ion signal and signal at 71 indicates the separation of methyl groups (M.W. = 15 u) group from molecular ions. Further release of a CH2 group from that fragment (M.W. = 14 u) results in a signal at 57. The signals at 43, 29, and 15 are due to the subsequent releases of CH2 groups. The abundance of signals at 15 and 71 is minimal due to the least probability for the formation of unstable methyl carbocation or a methyl radical, respectively. The base peak at 43 indicates the most preferred fragmentation, which yields a pair of stable carbocation and radical.

The molecular ions of linear alkanes prefer to fragment at the carbon-carbon bond away from the end of the chain since the cleavage of an inner bond ...

Dialysis: Diffusion Based Separation

Dialysis is a common technique used in biochemistry for separating molecules based on diffusion. In this procedure, a semipermeable membrane allows the movement of certain molecules based on size. This method can be applied to the removal of buffer, known as desalting, or exchanging buffer molecules or ions from a protein solution.
This video covers the principles of dialysis along with a general procedure. &#160;Several applications of dialysis are reviewed, including the removal of gradient reagents following ultracentrifugation, removing detergent after a membrane protein extraction, and the reconstitution of proteins by changing the solution environment.
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Biochemical samples typically have high buffer concentrations that can disrupt downstream processing and analysis. Dialysis is a common, inexpensive technique used to separate molecules based on diffusion. The method utilizes a semi-permeable membrane that allows the movement of certain components, based on size. This video will show the concepts of dialysis, a general procedure, and some of its uses in biochemistry.
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The most important aspect of dialysis is a semi-permeable membrane, which has pores that impose a molecular weight cut-off, allowing molecules below a certain size to pass through. For example, a 10k membrane will generally retain molecules larger than 10 kilodaltons. However, the molecular weight cutoff is not a discrete or precise boundary. The membrane typically contains a broad range of pore sizes, so a small fraction of molecules near the cutoff may be lost.
Since the molecular weight cutoff is defined using globular proteins, linear molecules of similar mass, like DNA or RNA, may slip through. Membranes are typically chosen one half to one third the molecular weight of the desired molecule.
To perform the procedure, a sample is placed into the membrane, which is in turn added to a large volume of solution, called the dialysate. Over time, smaller molecules will diffuse freely across the membrane between the sample and the dialysate, while the larger biomolecules are held within. Dialysis is a slow process. It is common to allow it to run overnight, or even across multiple days.
If the dialysate is pure water, the overall buffer concentration will decrease, a process known as desalting. If the solution contains other small particles, some will move into the sample, leading to buffer exchange. Because dialysis is an equilibrium process, the dialysate can be refreshed multiple times to further displace small molecules. Once the process is complete the sample is re-collected for further processing.
Now that you&#39;ve seen the basics of dialysis, let&#39;s take a look at a general procedure.
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Before beginning the procedure, the membrane is presoaked in dialysate. This makes it easier to use, and removes any preservatives. Once ready, the sample is collected, typically with a syringe and is then added to the dialysis container. This can be bare tubing, or contained within a cassette. Excess air is removed from the dialysis setup to maximize the sample&#39;s surface area with the membrane. The setup is then placed into the dialysate with stirring to maximize the diffusion. It should float to not inhibit stirring.
The dialysate is changed at relevant intervals as equilibrium between sample and dialysate is reached. After the last change, the reaction is typically left to run overnight. After a sufficient time period, the buffer-free or -exchanged sample is removed from the cassette. Once collected, the sample can be analyzed or further processed, depending on the nature of the experiment.
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Now that we&#39;ve looked at a general dialysis procedure, let&#39;s see some of the ways this technique is used in biochemistry.
Density gradients are a common way to separate complex biological samples. This concept relies on the distribution on small particles, typically sucrose or cesium chloride ions. Once complete, these reagents typically need to be removed before the collected sample can be processed. Dialysis makes it possible to utilize the purified sample for future analysis.
Certain proteins are found within a cell&#39;s lipid bilayer, and are usually studied by interspersing them into spherical lipid vesicles known as liposomes. The proteins and lipids are first extracted with a detergent. Dialysis can be used to slowly remove the detergent, forming proteoliposomes.
After purification, some proteins are misfolded, or denatured, leading to a loss in functionality. The compounds that cause these changes in structure can be removed with dialysis, leading to the reformation of functioning analytes.
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You&#39;ve just watched JoVE&#39;s video on dialysis. You should now understand this diffusion-based method, a simple experimental procedure, and the use of this technique.
Thanks for watching!
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Dialyse: Diffusion basiert Trennung

Dialysis is a common technique used in biochemistry for separating molecules based on diffusion. In this procedure, a semipermeable membrane allows the ...

Enzyme Assays and Kinetics

Enzyme kinetics describes the catalytic effects of enzymes, which are biomolecules that facilitate chemical reactions necessary for living organisms. Enzymes act on molecules, referred to as substrates, to form products. Enzyme kinetic parameters are determined via assays that directly or indirectly measure changes in substrate or product concentration over time.&#160;
This video will cover the basic principles of enzyme kinetics (including rate equations) and kinetic models. The concepts governing enzyme assays are also discussed, followed by a typical colorimetric assay. The applications section discusses an enzyme assay via F&#246;rster resonance energy transfer (FRET) analysis, characterizing extracellular enzyme activity in the environment, and investigating DNA repair kinetics using molecular probes.
<div class="chapter_text" id="script_0">
Enzymes are biochemical catalysts that are essential for life. Enzyme assays are used to study the kinetic properties of enzymatic reactions, elucidating the catalytic effects of enzymes. This video will cover enzyme kinetics and assays, go over a general procedure, and show some applications.
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<div class="chapter_text" id="script_1">
Enzymes are proteins, or less often RNAs, that act on a specific reactant, referred to as the substrate. An enzyme reduces the activation energy needed to initiate a biochemical reaction, causing the reaction to occur at a faster rate.
Enzymatic reactions can be broken up into three elementary components. The first is the formation of the enzyme-substrate complex, formed by the binding of the substrate to the enzyme active site. The complex can decompose into its original constituents. This is the second elementary reaction. Alternatively, the complex can form the product and recover the enzyme, the third elementary reaction.
The kinetics of an elementary reaction is given by the elementary rate law equation. Rate law equations give the rate in terms of the concentration of the reactants and a rate constant. Each of the elementary reactions has an individual rate law equation, with its own rate constant. These equations can be distilled down to a kinetic model known as the Michaelis-Menten equation. This gives the reaction rate in terms of the substrate concentration; which can be experimentally determined. Some general trends for enzyme reactions can be identified using the Michaelis-Menten equation. At high substrate concentration, a saturation point is reached, called Vmax. Here, the rate is limited by the total enzyme concentration, and the number of substrate molecules an enzyme converts into product per given time, also known as kcat. In Michaelis-Menten kinetics kcat is one of the two constants that govern reaction rate. The other constant, KM, is known as the affinity constant. KM is also equivalent to the concentration where the reaction rate is equivalent to one-half Vmax . An enzyme with a higher affinity will have a lower KM and reach Vmax faster, while an enzyme with lower affinity will have a higher KM and take longer to reach Vmax. Knowing kcat and KM allows for enzymes to be compared. To do this we use a ratio called enzyme efficiency. Higher kcat and lower KM result in higher efficiencies, while lower kcat and higher KM results in lower.
The factors used to elucidate enzyme kinetics must be determined experimentally. These assays are typically performed by mixing an enzyme and substrate solution in a controlled environment. Observations are made by measuring the changes in concentration of the substrate, product, or byproducts with respect to time.
The change in concentration over time is used to determine the reaction rate. In order to determine the kinetics, rate data must be obtained at multiple concentrations. If a plot of the inverse initial rate vs. inverse initial concentration, known as the Lineweaver-Burk plot, is linear, then the reaction follows Michaelis-Menten kinetics. The slope and intercept of the line allow for the determination of the kinetic parameters KM and Vmax, which can then be used to calculate kcat and the enzyme efficiency.
Now that the principles of enzyme kinetics have been discussed, let&#39;s look at how a typical enzyme assay is performed.
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In this procedure a colorimetric assay is demonstrated. The first step is to generate a standard curve, which will correlate absorbance with substrate concentration. Solutions of known concentration are prepared along with a control sample. A developer solution that reacts with the substrate is added to produce a colored compound. Absorbance is measured and plotted against concentration to generate the standard curve.
To perform the assay, a known concentration of substrate is prepared along with the appropriate amount of enzyme. The enzyme and substrate are mixed and allowed to incubate for a set time interval. pH and temperature are controlled with buffer solutions and heating blocks. A quenching agent is added to stop the reaction. Developer solution is then added to the reactions and mixed. The solutions are then placed in cuvettes and absorbance is measured. The amount of substrate consumed is determined by comparing the measured absorbance to the standard curve. Using the collected data, initial reaction rates are determined by plotting concentration over time. Finally, with the rate data and concentration, the Michaelis-Menten plot is made. This allows for the determination of kinetic properties for the enzyme such as turnover number and enzyme efficiency.
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Now that we&#39;ve reviewed an assay procedure, let&#39;s look at other ways assays are performed and their applications.
In this procedure FRET analysis is used to study the kinetics of a protease hydrolyzing a peptide bond of a protein. These emissions can be measured, allowing for a continuous and quantitative analysis of substrate consumption and production, aiding in the determination of the reaction kinetics.
Enzyme assays can be used in environmental science to determine the levels of extracellular enzyme activity in the environment. Waters, soils, and sediments can be collected from the environment and processed in the laboratory. Extracellular enzymatic activity of these materials can then be characterized using enzyme assays. This is a useful tool for understanding how the environment processes organic material.
A cell&#39;s DNA repair mechanism can be evaluated by studying the kinetics of enzymes found in the nucleus. The rate at which an enzyme removes DNA lesions, or damages, can be measured using fluorescent molecular beacons, which only fluoresce when bound to unique DNA sequences. The level of DNA repair can be measured in real time by detecting the fluorescently labeled cleavage products.
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<div class="chapter_text" id="script_4">
You&#39;ve just watched JoVE&#39;s video on enzyme kinetics and assays. This video explained enzyme kinetics, covered assay concepts, went over a general procedure, and described some applications.
Thanks for watching!
</div>

Enzyme Assays to Study Enzyme Activity and Kinetics

Enzym-Assays und Kinetik

Enzyme kinetics describes the catalytic effects of enzymes, which are biomolecules that facilitate chemical reactions necessary for living organisms. ...

MALDI-TOF Mass Spectrometry

Matrix-assisted laser desorption ionization (MALDI) is a mass spectrometry ion source ideal for the analysis of biomolecules. Instead of ionizing compounds in the gaseous state, samples are embedded in a matrix, which is struck by a laser. The matrix absorbs the majority of the energy; some of this energy is then transferred to the sample, which ionizes as a result. Sample ions can then be identified using a time-of-flight analyzer (TOF).
This video covers principles of MALDI-TOF, including matrix selection and how TOF is used to elucidate mass-to-charge ratios. This procedure shows the preparation of a MALDI plate, the loading of samples onto the plate, and the operation of the TOF-mass spectrometer. In the final section, applications and variations are shown, including whole-cell analysis, characterization of complex biological samples, and electron spray ionization.
<div class="chapter_text" id="script_0">
Matrix-assisted laser desorption ionization, or MALDI, is a mass spectrometry ion source ideal for the analysis of biomolecules. Most ion sources remove structural information from large, fragile biomolecules. MALDI maintains structural integrity, and therefore information, while accelerating the molecules into the mass analyzer, which separates the compounds based on size and charge. The most commonly coupled with MALDI is the time of flight, or TOF, mass analyzer. This video will show the concepts of MALDI ionization, a general procedure, and some of its uses in biochemistry.
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For mass spectrometry to function, molecules must be ionized into the gaseous state. In MALDI, the sample is embedded in a matrix, typically an organic compound containing aromatic and conjugated double bonds.
When a laser pulse strikes this mixture the matrix absorbs the majority of the energy, rapidly heats, and is desorbed, or released, from the surface. The energized matrix transfers some of its energy to the biomolecules, desorbing and then ionizing them.
MALDI is typically paired with a time of flight, or TOF, mass analyzer. An electric field applies kinetic energy to the ions, moving them into a field-free region called a drift tube. The velocity of the ions as they move through the tube is related to their mass-to-charge ratio, so heavier particles travel slower through instrument. A detector at the end of the tube measures each ion&#39;s flight time. With this knowledge, as well as the tube length and applied field strength, the mass-to-charge ratio of each ion can be elucidated.
This plot of signal intensity to mass-to-charge-ratio, known as a mass spectrum, can be compared to a library of collected spectra. If no matches are found, it can molecules can be identified by further techniques, such as tandem mass spectrometry. For more information, see this collection&#39;s video on the topic.
Now that the basics of MALDI-TOF have been discussed, let&#39;s look at the process in the laboratory.
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Before beginning an experiment, it&#39;s important to consider the choice of matrix from which samples will be desorbed. It must absorb the laser energy, be stable in a vacuum, not react with the target molecules, and be able to desorb. Depending on the sample, different matrices are preferred. For a large protein, a combination of CHCA and DHB has shown better separation of the peaks, called resolution, than the individual matrices.
There are a number of ways to prepare samples. We&#39;ll show what is known as the &#34;double-layer&#34;, or &#34;sandwich,&#34; method. To begin, clean the MALDI plate with ultra-pure reagents, as mass spectrometry is very sensitive to contamination. Dry the plate with a stream of inert gas.
Next, a saturated matrix solution is made, typically with an organic solvent . The solution is streaked onto the MALDI plate and dried. A second saturated solution of matrix containing trifluoroacetic acid, or TFA, is prepared. TFA helps ions into the gaseous phase.
Next, the sample solution is added on top of the dried matrix spot. Add the matrix solution containing TFA on top of the sample, thereby completing the matrix &#34;sandwich&#34;. Homogeneity of the spot can be verified under a low-powered microscope.
Plate a calibration standard, which is a mixture with a wide range of known masses and is used to correlate the time-of-flight to m/z. Finally, plate the matrix alone as a negative control.
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To analyze the spots, place the target plate into the instrument. Ensure there&#39;s no debris present, allowing for the formation of a tight vacuum. In the software, select the standard, negative control, and samples of interest. Label the spots with the correct identification.
The ion source and lens voltages can be manipulated to improve performance of the analysis. This will depend on the specifics of the instrument and sample. Focus on the standard spot and calibrate the instrument with the software.
Next, collect spectra from each of the sample spots. Try a few different locations on the spot to maximize the quality of the collected data. Once finished, the MALDI plate can be collected and reused after cleaning.
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Now that we&#39;ve reviewed a procedure, let&#39;s look at some of the ways MALDI is utilized, and a different ionization technique.
In addition to biomolecules, MALDI can be used to analyze living cells. Macrophages are immune cells that take on one of several different forms, based on their microenvironment. After exposing the cells to various signaling molecules, or cytokines, they can be added directly to the plate, and analyzed. The MALDI spectra can be used as unique &#34;fingerprints&#34;, depending on the cytokine used.
Complex biological samples like mammalian sebaceous secretions require a step of purification before MALDI analysis. Thin layer chromatography is one such technique that relies on the components&#39; polarity. The compounds are collected from the TLC pate, purified, and transferred to a MALDI matrix. The resulting spectra verify the identity and purity of the separated biomolecules from the mammalian sebaceous secretions.
Another common ion source for biomolecules is electrospray ionization, or ESI. In this method, the sample is injected into the instrument, where a high voltage is applied, creating an aerosol of charged droplets. As the solvent in the droplet evaporates, the charge is moved to the sample molecules, till they are completely gaseous. ESI doesn&#39;t require the spotting procedure, and the sample can be injected directly into the instrument. On the other hand, ESI is more sensitive to the presence of buffer components and other contaminants, meaning MALDI is more robust.
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You&#39;ve just watched JoVE&#39;s video on MALDI mass spectrometry. This video described the theory behind the instrument, went over a general procedure, and covered some of the uses of the technique. Thanks for watching!
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MALDI-TOF Mass Spectrometry-Principle, Operation and Uses

MALDI-TOF-Massenspektrometrie

Matrix-assisted laser desorption ionization (MALDI) is a mass spectrometry ion source ideal for the analysis of biomolecules. Instead of ionizing ...

Tandem Mass Spectrometry

In tandem mass spectrometry a biomolecule of interest is isolated from a biological sample, and then fragmented into multiple subunits in order to help elucidate its composition and sequence. This is accomplished by having mass spectrometers in series. The first spectrometer ionizes a sample and filter ions of a specific mass to charge ratio. Filtered ions are then fragmented and passed to a second mass spectrometer where the fragments are analyzed.
This video introduces the principles of tandem mass spectrometry, including mass-to-ratio selection and dissociation methods. Also shown is a general procedure for analyzing a biochemical compound using tandem mass spectrometry with collision-induced dissociation. The applications section covers selection reaction monitoring, determination of protein post-translation modifications, and detection of tacrolimus levels in blood.
<div class="chapter_text" id="script_0">
Tandem mass spectrometry links together multiple stages of mass spectrometry to first isolate a biomolecule, and then determine aspects of its chemical makeup. Biomolecules have large, complex structures, making it difficult to determine their molecular composition. Tandem mass spectrometry selects a molecule of interest that is later fragmented into multiple subunits, which can help elucidate its identification and sequence. This video will show the concepts of tandem mass spectrometry, a general procedure, and some of its uses in biochemistry.
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Tandem mass spectrometry begins as a typical mass spec instrument: with an ion source, which converts the sample into ions, and a mass analyzer, which separates the ions based on their mass-to-charge ratio. A common mass analyzer, the quadrupole, only allows ions with a specific ratio through, while the others crash into the rods of the apparatus. The species allowed through, called the precursor ion, is the biomolecule of interest. The ion moves into a collision cell, typically another quadrupole, where energy is applied to fragment the ion in a predictable pattern.
These fragments move into another mass analyzer, such as a time-of-flight, which separates these &#34;product ions&#34;. The product ions are then sent to the detector, as in a normal MS instrument. In the case of an unknown protein, the resulting spectrum contains numerous overlapping fragments, making a definitive complete sequence of the biomolecule difficult to generate. However, the spectral pattern is unique for a given protein. Analysis software compares the spectrum to a database of known peptide sequences, elucidating the unknown protein from the overlapping fragments.
Depending on the sample and desired degree of fragmentation, multiple fragmentation methods are possible. Fragmentation patterns depend on how the energy is transferred, its amount, and how it is distributed through the precursor ion. Energy can be transferred via neutral particles, radiation, or electrons. Using neutral atoms, a process called collision-induced dissociation or CID, primarily cleaves at the peptide bond between the amino acids, ideal for their identification.
Now that the basics of the technique have been covered, let&#39;s look at CID tandem mass spectrometry being used to study a component of bacterial cell envelopes.
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As with all mass spectrometric experiments, the first step is to ionize the sample. For biomolecules, this is typically done with matrix assisted laser desorption or electrospray ionization. The precursor ion signal is then optimized by tuning of the ion optics. Once done, the target is isolated and the fragmentation method is chosen, such as CID.
The strength of an applied voltage, which accelerates the precursor ion into the collision cell, affects the degree of fragmentation. This voltage is increased until the precursor is roughly 10% abundance compared to the highest product ion. Multiple spectra are acquired and averaged until a sufficient signal-to-noise ratio is achieved. The number of scans needed is dependent on the signal intensity of the original precursor ion and can range from 3 to 300.
The analyte in this example, lipid A from Escherichia coli K-12, had 19 major fragments after CID. Lipid A&#39;s general structure is well known, allowing software to reconstruct the specific composition from the sample.
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Now that we&#39;ve looked the procedure, let&#39;s look at some of the ways tandem mass spectrometry is used in biochemistry.
A common scanning mode in tandem mass spectrometry is selected reaction monitoring, or SRM. In SRM, both mass analyzers are fixed to a selected mass-to-charge ratio, focusing on specific precursor and product ions. Because of SRM&#39;s high degree of sensitivity, the spectra of peptide standards of known concentration can be utilized and compared to that of the unknown samples, allowing proteins of interest to be quantified.
Proteins are commonly modified after translation, typically by the addition of functional groups such as methyl groups, phosphate groups, or sugars, known as glycans. These are important in cell signaling processes, elucidating how cells communicate with one another. Because tandem mass spectrometry fragments the proteins into smaller components, it is possible to determine the location of the PTM to the specific fragment or even an amino acid. Some modifications, such as acetylation and trimethylation, are difficult to differentiate by mass alone, so chromatographic separation is performed before the mass spectrometry.
Many analytes in patient&#39;s blood are found at concentrations below the limit of detection for typical mass spectrometry. Another advantage of SRM is that it discards all but one product ion, increasing the sensitivity and enhancing the lower detection limit by up to 100 fold. In this example, the immunosuppressant drug, tacrolimus, could be detected at levels of 1 ng/mL.
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<div class="chapter_text" id="script_4">
You&#39;ve just watched JoVE&#39;s video on tandem mass spectrometry. This video described the theory of the instrument, went over a general procedure, and explained some of the ways the technique is currently being utilized. Thanks for watching!
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Tandem Mass Spectrometry: Principle, Instrumentation, Uses

Tandem-Massenspektrometrie

In tandem mass spectrometry a biomolecule of interest is isolated from a biological sample, and then fragmented into multiple subunits in order to help ...

Protein Crystallization

Protein crystallization, obtaining a solid lattice of biomolecules, elucidates protein structure and enables the study of protein function. Crystallization involves drying purified protein under a combination of many factors, including pH, temperature, ionic strength, and protein concentration. Once crystals are obtained, the protein structure can be elucidated by x-ray diffraction and computation of an electron density model.
This video introduces protein crystallization and shows a general procedure. Protein expression and purification, crystallization, and x-ray diffraction are covered in the procedure. Applications of protein crystallization include in silico drug design, binding site determination, and membrane protein structure analysis.
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Protein crystallization is the process of obtaining a latticed solid form of a protein. These crystals are especially valuable to structural biologists, assisting in the study of protein function. Other techniques, such as mass spec or SDS-PAGE, can only provide information on the one-dimensional structure of proteins. Protein crystallization is complemented by the techniques of recombinant protein expression and x-ray diffraction. This video will show the principles of protein crystallization, a general laboratory procedure, and several of its applications in the biochemical field.
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The first step required in the process is to obtain milligram quantities of very pure protein, typically using recombinant protein expression. The gene corresponding to the protein of interest is cloned into an expression vector, and the expressed protein is fused to an affinity tag, such as poly-histidine, to assist in the purification by affinity chromatography. To learn more, see this collection&#39;s video on affinity chromatography.
Formation of the purified protein into crystals is dependent on the proper combination of many factors, including pH, ionic strength, concentrations of precipitant and protein, temperature, and rate of equilibration. The most common method used is vapor diffusion, of which there are two categories: hanging drop and sitting drop. A droplet containing pure protein, buffer, and precipitant, which is an ionic solid that binds water molecules, reducing water availability for the protein and mimicking higher protein concentration, is in an enclosed microwell with a reservoir with a more highly concentrated mixture of the same buffer and precipitant. At the beginning, the concentrations of protein and precipitant are too low to cause crystallization. During the course of the experiment, water vaporizes from the droplet and collects in the reservoir; a decrease in the amount of water in the droplet causes the system to become supersaturated, and nucleation, followed by crystallization, can occur. The net transfer of water from the droplet is in equilibrium, and the system is maintained until the process is complete.
To visualize the 3D structure, x-ray diffraction is used. To obtain x-ray data from a crystal, it is placed in a monochromatic x-ray beam, where it is exposed to the beam at all angles. Each exposure provides an image, where each spot is a diffracted x-ray, which emerges from the crystal and is registered by a detector. The data are combined to produce a model of the arrangement of atoms within the crystal. The resulting crystal structure demonstrates the 3-dimensional placement of the atoms, with a typical resolution of 2 angstroms.

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Now that we have covered the principles of protein crystallization, let us look at a generalized protocol.
To begin the procedure an expression vector containing the gene of interest is transformed into cells. The cells are incubated and at mid-log phase, expression is initiated by adding an inducer, such as IPTG, which triggers transcription of the gene&#39;s mRNA. After protein expression, the crude material is suspended in lysis buffer, and then clarified by centrifugation.
The clarified lysate is then loaded onto a nickel column, and the polyhistidine-tagged protein binds to the column while all other biomolecules are washed away.
Once several milligrams of pure protein have been obtained, it is ready for crystallization by vapor diffusion. A 24-well hanging/sitting drop tray is filled with varying concentrations of sodium chloride and sodium acetate buffer solutions. For the sitting drop method, equal volumes of protein and reservoir solution are pipetted onto the shelf above each well, and then the tray is covered with transparent tape. The tray is then placed in an incubation chamber, and the wells are monitored for growth the following day, then every few days.
Once a proper crystal has been obtained it is ready for x-ray diffraction analysis. The crystal is mounted on a goniometer to position the crystal at selected orientations. The crystal is illuminated with a monochromatic beam of x-rays at all angles, producing a diffraction pattern. The software converts the two-dimensional images, taken at different orientations, to a three-dimensional model of the density of electrons within the crystal by determining the positions of the atoms in the crystal.
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<div class="chapter_text" id="script_3">
Now that we have reviewed a procedure, let&#39;s review some useful applications of protein crystallization, and another crystallization technique.
Protein crystallization may be used for in silico drug design. The three-dimensional structure of Influenza virus&#39;s polymerase basic protein 2, which has been linked to viral infection in mammals, was determined by crystallization and x-ray diffraction. Potential binding sites in the protein are visualized, and with the use of a docking program, a three-dimensional molecule was designed that would insert into a cleft in the protein.
Co-crystallization of protein-DNA complexes is also a useful technique. DNA-binding proteins modulate a wide variety of biological functions such as transcription and DNA polymerization and DNA repair; and crystal structures of these complexes can provide insight into protein function, mechanism, and the nature of the specific interaction. The E. coli protein SeqA, a negative regulator of DNA replication, was co-crystallized with hemimethylated DNA.
Integral membrane proteins such as G-protein coupled receptors, or GCPRs, are difficult to crystallize due to their limited amount of polar surface area available for forming crystal lattice contacts, which has led to the development of fusion-protein-assisted protein crystallization. Genes encoding &#946;2 adrenergic receptor, a GCPR, and a lysozyme were inserted into an expression vector. The crystallization of the &#946;2AR- lysozyme fusion protein was achieved due to the increased extracellular hydrophilic surface over the naturally hydrophobic &#946;2AR, provided by the lysozyme, necessary for forming packing interactions in the crystal lattice.
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<div class="chapter_text" id="script_4">
You&#39;ve just watched JoVE&#39;s video on protein crystallization. This video described its principles, a generalized protocol, and some its uses in the biomedical field. Thanks for watching!
</div>

Protein Expression, Crystallization, and X-Ray Diffraction

Protein-Kristallisation

Protein crystallization, obtaining a solid lattice of biomolecules, elucidates protein structure and enables the study of protein function. ...

Chromatography-based Biomolecule Purification Methods

In biochemistry, chromatography-based purification methods are employed to isolate compounds from a complex mixture. Two such methods used commonly by biochemists are size-exclusion chromatography and affinity chromatography. In size-exclusion chromatography, a column packed with porous beads separates components of a mixture based on size. On the other hand, affinity chromatography allows for a more specific separation of biomolecules by using a column that is composed of stationary phase, which contains target-specific ligands.
This video serves as an introduction to size-exclusion and affinity chromatography, as well as the concepts that govern them. A step-by-step procedure for the purification of a histidine-tagged protein by immobilized metal affinity chromatography is described. Applications for both of these chromatographic methods in biochemistry and biomedical research are also profiled.
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&#34;Chromatography&#34; refers to a wide range of methods used to isolate a component from a complex mixture, an essential step before a biomolecule&#39;s properties and activities can be determined. Each chromatographic technique has a different mechanism for separation, depending on the sample matrix and target compound. This video will focus on the principles and operation of two methods common to biochemistry: size-exclusion and affinity chromatography.
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<div class="chapter_text" id="script_1">
Size-exclusion chromatography or SEC is based on the size of the compounds in the sample. A mobile phase containing the sample is added to a column with a porous material: the stationary phase. The molecules in the sample fall into 1 of 3 categories.
Molecules too large to enter the pores travel the shortest distance through the column. Any species with a molecular weight above this &#34;exclusion limit&#34; will exit the column at the same time. Molecules small enough to freely enter the pores will be retained the longest, and will exit the column together. The molecular weight allowing complete pore entry is the &#34;permeation limit&#34;.
Only molecules between these limits will be separated from one another, as they spend varying amounts of time diffusing into and out of the pores. Smaller molecules are retained longer on the column because they spend more time in the stationary phase, whereas larger molecules within these limits exit earlier.
Molecular weights across 1 to 2 orders of magnitude fall within these limits. Columns are chosen with this in mind, or multiple columns can be used in series if there is a wide range of desired compounds.
Now that you&#39;ve seen the theory of SEC, let&#39;s look at how it is carried out.
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To begin the SEC procedure, the column must be equilibrated with deionized water and chromatography buffer. Once prepared, buffer containing the sample is injected onto the column. The buffer is then pushed through at a low flow rate. A detector monitors what exits the column to determine the presence of the desired analyte. Large molecules with molecular weights above the exclusion limit exit the column at the same time. Small fractions from the column are collected in tubes. Each fraction is tested for the quality of the target molecule by gel electrophoresis or other analytical techniques.
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<div class="chapter_text" id="script_3">
Now let&#39;s have a look at affinity chromatography or AC, one of the most efficient ways to purify proteins. Many biomolecules bind selectively to certain ligands-a property which AC utilizes by adhering a target-specific ligand to the stationary phase.
When the mixture flows through the column, the target molecules attach to the ligand, and the rest flow through. After the mixture has passed through the column, the target molecule can be collected through one of two elution methods based on specificity.
Biospecific elution can be said to a have a &#34;normal-&#34; or &#34;reverse-role&#34;. In normal-role biospecific elution, an agent is added that competes with the adhered ligand to bind with the target biomolecule.
In reverse-role biospecific elution, an agent competes with the target to bind to the adhered ligand. The second elution type, nonspecific elution, lowers the target-to-ligand binding by changing the solution&#39;s pH, ionic strength, or polarity. If a protein does not bind to a ligand that can be immobilized, the protein can be expressed containing a &#34;tag&#34;: short peptide sequences engineered to bind to the ligand.
One variety is immobilized metal ion affinity chromatography, &#34;IMAC&#34; for short, where an adhered metal ligand, like nickel or cobalt, binds to histidine residues on the modified protein. Through molecular biology techniques, target proteins are generated with repeating histidine residues, called a polyhistidine-tag, which binds to the metal via the imidazole side chain on histidine. Once bound, the protein can be collected with free imidazole via reverse-role specific elution and later used in a wide variety of downstream applications.&#160;Now that you&#39;ve seen the theory of affinity chromatography, let&#39;s look at an IMAC procedure in the laboratory.

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In IMAC, the stationary phase can be added directly to the mixture of mobile phase and sample, allowing the binding of the his-tagged protein. This slurry is then poured into the column, where the non-bound compounds drip into waste, while the slurry remains. The slurry container is rinsed to collect residual resin and sample, which is added to the column.
The resin is stirred to ensure the unbound components are free-flowing. Added wash buffer helps flush them away. Once all of the unbound components have been removed, the waste is replaced with a container to collect the target protein.
Buffer containing imidazole is added, stirred, and allowed to rest to unbind the target molecule. The imidazole binds to the metal, replacing and releasing the tagged protein. The freed protein is collected, and the imidazole step is repeated to ensure total collection. To further purify the sample, SEC can be run on the sample prior to analysis.
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Now that we&#39;ve seen the theory and procedure of these two techniques, let&#39;s look at some of the ways they&#39;re applied in the biochemical field.
A common reason to purify proteins is to study their role in disease. Cystic fibrosis is caused by defects in the cystic fibrosis transmembrane conductance regulator protein, or CFTR. After growing the protein with a his-tag in yeast, both affinity and size-exclusion chromatography allow the isolation of the protein, followed by the study of its function.
In some instances, the presence of a polyhistidine tag can change a protein&#39;s structure, thereby affecting its function. Another common tag is maltose-binding protein or MBP, which will bind to bound amylose in a column. Maltose is then used to release the complex. The MBP can then be cleaved and removed with SEC to produce the pure desired protein.
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<div class="chapter_text" id="script_6">
You&#39;ve just watched JoVE&#39;s video on size-exclusion and affinity chromatography. It covered the theory of the techniques, went over general procedures, and covered some of the uses of the techniques.
Thanks for watching!
</div>

Size Exclusion and Affinity Chromatography

Chromatographie-basierte Biomolekül Reinigungsverfahren

In biochemistry, chromatography-based purification methods are employed to isolate compounds from a complex mixture. Two such methods used commonly by ...

Two-Dimensional Gel Electrophoresis

Two-dimensional gel electrophoresis (2DGE) is a technique that can resolve thousands of biomolecules from a mixture. This technique involves two distinct separation methods that have been coupled together: isoelectric focusing (IEF) and sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE). This physically separates compounds across two axes of a gel by their isoelectric points (an electrochemical property) and their molecular weights.
The procedure in this video covers the main concepts of 2DGE and a general procedure for characterizing the composition of a complex protein solution. Three examples of this technique are shown in the applications section, including biomarker detection for disease initiation and progress, monitoring treatment in patients, and the study of proteins following posttranslational modification (PTM).
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Two-dimensional, or 2D, gel electrophoresis is a technique utilizing two distinct separation methods which can separate thousands of proteins from a single mixture. One of the techniques, SDS-PAGE or sodium dodecyl sulfate polyacrylamide gel electrophoresis, cannot fully separate complex mixtures alone. 2D gel electrophoresis couples the SDS-PAGE to a second method, isoelectric focusing or IEF, which separates based on isoelectric points, allowing for the resolution of potentially all proteins in a cell lysate. This video will show the principles of 2D gel electrophoresis, a general procedure, and some of its biomedical applications.
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2D gel electrophoresis begins with IEF as the first dimension. Every protein has a pH value, called the isoelectric point or pI, where the net charge is zero. When a protein is subjected to an electric field, it will move toward the electrode with opposite charge. Samples of interest are loaded onto immobilized pH gradient, or IPG, strips which have embedded ampholytes, molecules containing both acidic and basic groups. An electric field is then applied to the pH gradient strip, causing the proteins to migrate until they reach the pH value matching their pI, where they lose their net charge.
Prior to running the second-dimension, the embedded proteins are treated with SDS, denaturing them and providing a uniform negative charge. Once completed, the IPG strips are placed onto a polyacrylamide gel. An applied electric field draws the proteins toward the anode, with larger proteins moving more slowly through the gel.
Once the protein mixture has been separated according to pI and molecular weight, the proteome map is visualized using stains, and proteins of interest are identified.
Now that we have discussed the principles of 2D gel electrophoresis, let&#39;s go over a typical laboratory procedure.
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Before the experiment can be performed, the proteins must be solubilized into media. Solubilization of the sample is achieved by de-aggregating proteins with a combination of chaotropic agents for disrupting hydrogen-bonding interactions, nonionic detergents to prevent altering of the proteins&#39; charge, reducing agents to break disulfide bonds, and buffers. To remove interfering abundant proteins and other molecules, the material is sequentially extracted by centrifugation, and collection of the resulting pellet; followed by treatment with endonuclease, an enzyme used to consume any DNA that would interfere with the experiment.
Once the proteins have been solubilized, IPG strips are prepared by rinsing with a strip-cleaning solution and left upside-down to dry. Each strip is then assigned a strip holder number. Once ready, the cellular extract is loaded onto the strips in a slow, sliding motion from the negative to the positive end. For the purpose of rehydration, damp blotting paper is placed on top of the electrode and under the gel strips; the IPG strips are then lined onto the IEF instrument. A high electric current is applied, and the proteins begin to migrate.
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<div class="chapter_text" id="script_3">
Following completion of the first dimension, the gel for SDS-PAGE is prepared in a casting apparatus. The IPG strips are treated by placing them face down in SDS-containing equilibration buffer. The electrophoresis unit is readied with the addition of electrophoresis buffer. The treated IPG strips are collected using tweezers, placed on top of the gel plates, and sealed with agarose-sealing solution. A voltage source then applies an electric field, which is held until the fastest-moving proteins are 1 cm from the bottom of the gel.
After completion of the electrophoresis, the proteins must be visualized. Traditionally this is performed by staining with Coomassie blue or silver nitrate. Proteins of interest may be transferred from the gel, and analyzed by Western blot analysis.
A second identification approach involves excision of the proteins from the gel, digesting them, than analyzing them by mass spectrometry.
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<div class="chapter_text" id="script_4">
Now that we&#39;ve reviewed a procedure, let&#39;s look at some of the uses for 2D gel electrophoresis.
One of the most common uses for this technique is the identification of molecules involved in disease initiation and progression. 2D gel electrophoresis, coupled with mass spectrometry, can detect the up- or down-regulation of specific proteins in diseased areas in comparison to healthy ones.
Additionally, 2D gel electrophoresis is useful in following the progress of patients&#39; response to a potential therapeutic drug. Specimens may be taken from patients at various timepoints following administration of the treatment. In this way, 2D gel electrophoresis coupled with Western blot or mass spectrometry analysis, can detect proteins associated with negative responses such as inflammation; or the absence of proteins in an alleviated state.
Another use for 2D gel electrophoresis is in the study of protein structure and function following posttranslational modification, or PTM, which are additions to proteins following their translation from mRNA. PTM&#39;s can regulate a variety of functions, including protein signaling, gene expression, or cause oxidative damage. 2D gel electrophoresis is sensitive to modifications such as methylation or acetylation, which can cause a shift in pI as well as the molecular weight.
</div>
<div class="chapter_text" id="script_5">
You&#39;ve just watched JoVE&#39;s video on 2D gel electrophoresis. This video described the principles of the technique, a typical experimental procedure, and several of its applications in the field of biomedicine.
Thanks for watching!
</div>

Zweidimensionale Gelelektrophorese

Two-dimensional gel electrophoresis (2DGE) is a technique that can resolve thousands of biomolecules from a mixture. This technique involves two distinct ...

Metabolic Labeling

Metabolic labeling is used to probe the biochemical transformations and modifications that occur in a cell. This is accomplished by using chemical analogs that mimic the structure of natural biomolecules. Cells utilize analogs in their endogenous biochemical processes, producing compounds that are labeled. The label allows for the incorporation of detection and affinity tags, which can then be used to elucidate metabolic pathways using other biochemical analytical techniques, such as SDS-PAGE and NMR.
This video introduces the concepts of metabolic labeling and show two general procedures.&#160; The first uses isotopic-labeling, to characterize the phosphorylation of a protein. The second covers a photoreactive labeling to characterize protein-protein interaction within a Also three applications of metabolic labeling are presented: labeling plant material, labeling RNA to measure kinetics and labeling glycans in developing embryos.

<div class="chapter_text" id="script_0">
Metabolic labeling is used to investigate the machinery of a cell. This is accomplished using chemical analogs to probe the biochemical transformations and modifications that occur. This video will show the principles of metabolic labeling, typical isotopic and photoreactive labeling procedures, and some applications.
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<div class="chapter_text" id="script_1">
Metabolic labeling can be conducted using a number of strategies. Here we will describe isotopic, photoreactive, and bio-orthogonal labeling.
Isotopic labeling is performed using structural analogs that are chemically identical to their natural counterparts, but have uncommon isotopes incorporated into their structure. In this L-lysine analog the carbon and nitrogen atoms are replaced with carbon-13 and nitrogen-15. Cells cultured in the presence of isotopic analogs will incorporate them into their biochemical structures. Metabolites are collected from the cells and purified for analysis. Samples with stable isotopes are analyzed using techniques such as mass spectrometry or NMR spectroscopy. Samples with radioactive labels are analyzed using liquid scintillation counting and x-ray films, which will be demonstrated in the isotopic labeling protocol.
Photoreactive labels are functional groups incorporated into proteins, which are stable until exposed to ultraviolet light. The functional group forms a reactive radical, which binds to the nearest protein. A common example, L-photo-leucine, contains a diazirine ring, which is a photoreactive crosslinker. In contrast to isotopic labeling, there is some chemical dissimilarity between photo-reactive chemical analogs and their natural counterparts. Cells may preferentially incorporate natural compounds over analogs. Therefore, it is important to perform photo-reactive labeling in medium free of the compound being mimicked. Once exposed to ultraviolet radiation, the photo-reactive groups in a labeled protein become unstable and highly reactive, causing it to cross-link with interacting proteins, creating a protein complex. Cross-linked complexes, act as snapshots that can then be analyzed using SDS-PAGE and mass spectrometry methods. This provides insights into what reactions are occurring in the metabolic pathway by identifying reaction species and how they interact by determining binding sites.
Bioorthogonal labeling strategies utilize analogs with small functional groups that have little to no reactivity with natural biomolecules. For example, azides are small functional groups, whose reactivity is said to be orthogonal to biochemical reactions. In the Staudinger ligation, a phosphine group attacks the azido group. This yields a transition state that intramolecularly reacts with a nearby ester, resulting in an amine-bonded ligand. Bioorthogonal functional groups incorporated into biomolecules can be ligated with detection tags such as fluorescent functional groups, and affinity tags such as antigens.
Now that some concepts and strategies for metabolic labeling have been discussed, let&#39;s look at the process in the laboratory.
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The first step in a metabolic labeling experiment is to collect the protein of interest. To do this, cells are grown on a plate, and an expression method is used to promote synthesis of the desired protein. In this example, leucine-rich repeat kinases, or LRRK, are expressed. Disodium phosphate, containing radioactive phosphorus-32, is used as the analog. Proper measures must be taken to protect against ionizing radiation. This includes setting up a work area, wearing proper protective equipment, and checking for radioactive contamination. Once safety measures have been taken, the medium containing the isotopic analogs is prepared. The medium from the culture is removed and, replaced with one containing the isotopic chemical analogs and then incubated. Following incubation, the cells are lysed. The lysate is collected and purified.
After purification, proteins are resolved using SDS-PAGE and then transferred to a PVDF membrane. Autoradiography is performed by exposing the membrane to x-ray film and measured using a phosphor imager. Western blotting is used to measure relative protein levels in the PVDF membrane. In this example the phosphorylation levels of leucine-rich repeat kinases synthesized in 293T cells were measured. The autoradiogram shows how much phosphorous was incorporated into the protein. Western blotting elucidates the levels of the LRRK proteins. Image analysis software is used to obtain quantitative data of phosphorylation levels of the proteins.
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<div class="chapter_text" id="script_3">
In this next procedure, photoreactive labeling is demonstrated. First, the cells are prepared and cultured. The photoreactive analog is added to the cells at the mid-log phase and incubated. In this procedure p-benzoylphenylalanine is used. Samples are collected over intervals and put on ice. The samples are then exposed to obtain snapshots of the biochemical pathways over time. The proteins of interest are then purified and resolved using SDS-PAGE.
A photoreactive labeling strategy was used to identify compounds that interact with the protein of interest. Immunodetection with Western blotting shows protein bands that indicate higher molecular weight proteins are present in the irradiated samples. These are from cross-linking due to protein-protein interaction occurring during the UV irradiation.
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<div class="chapter_text" id="script_4">
Now that we&#39;ve reviewed metabolic labeling procedures, let&#39;s look at some of the ways the process is used.
Metabolic labeling concepts can be extended to multicellular organisms. Plants are grown in a sealed environment, rich in stable isotopes to produced labeled plant material. Carbon dioxide containing carbon-13 is added to the enclosure, while nitrogen-15 rich fertilizer is used. The resulting harvested plant material can help answer questions about carbon and nitrogen cycling from the ecosystem.
Labeling enables the separation of newly synthesized RNA from older RNA. By changing the initial concentration of analog, the kinetics of new RNA synthesis can be determined. The results show that the concentration of 4-thiouridine affects how much new RNA is transcribed. Additionally, incorporation rates of the label into RNA can be directly quantified with a spectrophotometer.
Using biorthogonal click chemistry, glycans in a zebra fish embryo can be labeled. The eggs are injected with a labeling compound that results in alkyne labels on the glycans. The glycans in the larvae are then ligated to a dye compound at the desired development stage. The glycans in the embryos are then imaged. Glycans produced at different time points can be identified by labeling using different colors at different stages of embryo development.
</div>
<div class="chapter_text" id="script_5">
You&#39;ve just watched JoVE&#39;s video on metabolic labeling. This video described the concepts behind metabolic labeling and their strategies, went over two general procedures, and covered some of the uses of the techniques.
Thanks for watching!
</div>

Isotopic, Photoreactive and Bioorthogonal Metabolic Labeling

Metabolische Kennzeichnung

Metabolic labeling is used to probe the biochemical transformations and modifications that occur in a cell. This is accomplished by using chemical analogs ...

Electrophoretic Mobility Shift Assay (EMSA)

The electrophoretic mobility shift assay (EMSA) is a biochemical procedure used to elucidate binding between proteins and nucleic acids. In this assay a radiolabeled nucleic acid and test protein are mixed. Binding is determined via gel electrophoresis which separates components based on mass, charge, and conformation.
This video shows the concepts of EMSA and a general procedure, including gel and protein preparation, binding, electrophoresis, and detection. Applications covered in this video include the analysis of chromatin-remodeling enzymes, a modified EMSA that incorporates biontinylation, and the study of binding sites of bacterial response regulators.
<div class="chapter_text" id="script_0">
EMSA, the electrophoretic mobility shift assay, also known as the gel shift assay, is a versatile and sensitive biochemical procedure. EMSA elucidates binding between proteins and nucleic acids by detecting a shift in bands in gel electrophoresis.
This video describes the principles of EMSA, provides a general procedure, and discusses some applications.
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<div class="chapter_text" id="script_1">
DNA replication, transcription, and repair, as well as RNA processing are all critical biochemical processes. They all involve binding between proteins and nucleic acids. Many serious diseases and disorders are associated with modifications in this binding. EMSA is a technique for qualitatively determining whether a specific protein binds to a specific nucleic acid. First, the nucleic acid is labeled, usually with radioactive phosphorus-32, to create a probe. Then the test protein and nucleic acid probe are mixed. When a protein binds to a nucleic acid probe, the resulting complex has greater mass and a different conformation than the nucleic acid alone.
Once bound, the complexes are analyzed with&#160;gel electrophoresis. In this technique, an electric field forces macromolecules to migrate through a gel matrix. The components separate based on mass, charge, and conformation. Electrophoresis can separate protein-DNA complexes from unbound probes. Since they have different masses and conformations, they will migrate through the gel at different rates and separate. The separation is easily detected, thanks to the presence of the radioactive phosphorus, and proves the protein successfully binds to the given nucleic acid. To verify the identification of the protein, a &#34;supershift assay&#34; uses an antibody with a known affinity to the protein. This has the added benefit of further shifting the complex, increasing resolution.
Now that we&#39;ve seen the principles, let&#39;s see it in the lab.
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<div class="chapter_text" id="script_2">
To begin the procedure, the protein must be isolated. To do this, molecular biology techniques are used to express the protein in cells, and then purify.
The nucleic acid is amplified and labeled to create a probe. Labeling is done through incubation for 10 min with dCTP containing radioactive phosphorus-32. A radiation-safe workbench and protective equipment are required.
The gel is then prepared. The gel needs to be non-denaturing, to prevent the protein from altering conformation and potentially unbinding from the probe during electrophoresis. Polyacrylamide gels have pore sizes of 5 to 20 nm and are useful for short probes up to 100 base pairs in length. Agarose gels have pore sizes of 70-700 nm and are useful for larger probes.
</div>
<div class="chapter_text" id="script_3">
With the protein and nucleic acid probe now prepared, we proceed to binding. A TRIS buffer solution is prepared and the protein and probe are added. The pH should be similar to physiological conditions, and salt concentration sufficient to prevent the protein from forming weak bonds with non-target nucleic acids. The reaction proceeds for 20-30 min at 4 &#176;C.
The next step is electrophoresis. A buffer of low ionic strength and a pH similar to that used in the binding reaction is utilized. It yields a &#34;caging effect&#34; that stabilizes complexes, increases mobility, and reduces heat generation. After electrophoresis, the components of the gel are transferred onto filter paper. In a dark room, the filter paper is then exposed to film. If the protein binds to the probe, two distinct labeled regions will be visible in the transfer. One representing the complex, and a separate one representing the unbound probe. The separation demonstrates that the protein successfully bound to the nucleic acid.
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<div class="chapter_text" id="script_4">
Now that we&#39;ve seen the basic procedure, let&#39;s look at some applications.
Chromatin is the tightly packed complex of DNA and proteins found eukaryotic cells. Chromatin-remodeling enzymes modify the structure to open the DNA to transcription. As this changes the mobility of the complex, EMSA can be used to explore the binding activity of the enzyme.
An alternate approach to labeling takes advantage of the interaction between DNA and the methyltransferase enzyme. A cofactor can be modified to bind permanently to DNA via methyltransferase. Rather than labeling with phosphorus-32, the cofactor is conjugated to biotin, which is advantageous because it&#39;s not radioactive.&#160;Because the cofactor is site-specific in its attachment, it&#8217;s relevant to genotyping, methylation detection, and gene delivery. The biotinilated nucleic acids are detected through ultraviolet fluorescence.
When environmental stimuli activate histidine kinases, a &#8220;response regulator&#8221; is phosphorylated. This in turn binds to DNA, affecting transcription, which can be studied by EMSA.&#160;For instance, a response regulator in Desulfovibrio vulgaris was demonstrated to bind to the gene of interest. EMSA was used to verify that the binding took place.
</div>
<div class="chapter_text" id="script_5">
You&#39;ve just watched JoVE&#39;s video on the electrophoretic mobility shift assay. You should now understand its principles of operation, the steps in its procedure, and its major operating parameters.
Thanks for watching!
</div>

Electrophoretic Mobility Shift Assay EMSA

Elektrophoretische Mobilität Shift Assay (EMSA)

The electrophoretic mobility shift assay (EMSA) is a biochemical procedure used to elucidate binding between proteins and nucleic acids. In this assay a ...

Photometric Protein Determination

Measuring the concentration is a fundamental step of many biochemical assays. Photometric protein determination takes advantage of the fact that the more a sample contains light-absorbing substances, the less the light will transmit through it. Since the relationship between concentration and absorption is linear, this phenomenon can be used to measure the concentration in samples where it is unknown.
This video describes the basics of photometric protein determination and introduces the Bradford Assay and the Lowry Method. The procedure in the video will cover a typical Bradford assay. Applications covered include direct measurement of very small volumes of nucleic acids to characterize concentration and purity, determination of coupling efficiency of a biomimetic material, and another variation of photometric protein determination using Remazol dye.
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	Determining the concentration of a protein in samples is a fundamental step in many biochemical assays. Photometric determination can be done with small sample sizes. The more a sample contains light-absorbing substances, the less the light will transmit through it. This provides a quantitative measurement of the absorbing substances. These concepts are so fundamental to science that the articles that introduced two of the techniques are in the three most cited papers of all time. This video will show the concepts behind some of the most common photometric protein determination techniques, how they are performed, and how the gathered data is analyzed.
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<div class="chapter_text" id="script_1">
	Photometric protein determination is based on the relationship between concentration and light absorbency. This is known as the Beer-Lambert Law, which states that the concentration of a light-absorbing species is proportional to its absorbance.
	This principle underlies all photometric protein determination methods. 
	For direct absorption analysis, the absorbance values of unaltered protein samples are measured. Because of their aromatic side chains, tryptophan and tyrosine residues give the highest absorbance readings at a wavelength of 280 nm.
	However, these amino acids-which are two of the least frequently found in proteins-are present in different amounts in every protein, so each determination is unique. To overcome this limitation, more complex assays-that are not dependent on these amino acids-were developed.
	One example is the Bradford Assay, where colored dye is added to the sample. The dye, known as Coomassie Blue, responds proportionally-the more protein present, the more binding events with the dye.
	Then, protein concentration is determined by measuring the absorbance of the bound Coomassie Blue dye, which absorbs light at 594 nm. However, the Bradford assay is linear over a short range of concentrations, so dilutions are often required before analysis.
	The Lowry Method combines the Biuret reagent, an alkaline solution of copper ions that react with peptide bonds, and the Folin-Cioc&#226;lteu reagent, which oxidizes aromatic protein residues. The resulting color change of the sample is proportional to the protein concentration.
	The absorbance of the reduced Folin reagent can be determined at 750 nm. Like direct absorption, each protein has a unique response, and must be calibrated for the protein of interest. Now that we've reviewed the basic principles behind some of the most common assays, let's look at how direct absorption and the Bradford assay are performed.
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	To begin a direct absorption analysis, the spectrophotometer is calibrated with a blank to determine zero absorbance. Standard solutions are prepared for use in creating the calibration curve. Then, an aliquot of the first standard is added to a cuvette, and placed into the spectrophotometer.
	The absorbance value at 280 nm is then recorded. This process is repeated for each standard, using a clean cuvette for each run. Once complete, a calibration curve is created by plotting the absorbance versus concentration. The slope of this line is the molar attenuation coefficient, which relates absorbance to concentration.
	Next, the unknown sample is added to a cuvette, and the absorbance value is recorded. As the data analysis for the different photometric determination methods is similar, we will cover that after we look at the Bradford assay.
	Here, the Bradford protein assay is performed with a BSA standard on a 96-well plate. To begin, BSA stock solutions are prepared.
	The unknown solutions are diluted with deionized water to ensure that the concentrations are within the assay's range. Depending on the kit, the Coomassie dye may also require dilution. Then, the calibration curve is set up by adding the BSA standards to the 96-well plate.
	Deionized water is added to reach the needed concentration to generate a standard curve. The unknown sample should be added to the plate in triplicates to ensure an accurate measurement is taken. Coomassie dye is next added to each well, mixing with the pipette.
	Deionized water is added to an empty well as a blank, to measure the absorbance. After waiting 5 min for the dye to bind, the absorbance is measured in a plate-reader at 590 nm.
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<div class="chapter_text" id="script_3">
	Now that we've performed a few assays, let's look at how to analyze the data. Each photometric protein determination method is based on the Beer-Lambert Law.
	The measured absorbance of the standards is used to create a calibration curve, which is then used to determine the concentration of unknown samples. This curve can be manually plotted, though newer spectrophotometric tools will create the calibration curve once all standards have been measured. These systems will also calculate protein concentration as unknown samples are analyzed.
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<div class="chapter_text" id="script_4">
	Now that we've reviewed how to analyze photometric protein determination data, let's look at some of the ways these procedures are utilized.
	The principles of photometric protein determination can also be used to directly measure nucleic acid concentration. The nanodrop spectrophotometer accepts samples of very small volume onto an optically active pedestal. The absorbance is then measured, and the system automatically determines the nucleic acid concentration. Because proteins and other sources can interfere with measurements, sample purity is determined by analyzing the 260 to 280 nm and 260 to 230 nm absorbance ratios. Pure nucleic acids typically yield ratios of approximately 1.8 and approximately 2.0 for DNA and RNA, respectively.
	Photometric protein determination can also be used in the production of biomimetic materials, which are inspired from nature to elicit specific cellular responses. Recombinant adhesins are bound to polystyrene beads to simulate bacterial attachment to host cells. The Bradford assay is used to determine the coupling efficiency of the recombinant adhesion to the beads in the production of the biomimetic material.
	Alternative photometric protein assays can be used in the detection and characterization of protein antimicrobials. Remazol brilliant blue R dye is covalently bonded to heat-killed bacteria. The protein antimicrobial is incubated in the dyed solution. Then, the sample is centrifuged, and the absorbance of the supernatant at 595 nm is measured using a microplate spectrophotometer. Increased absorbance, by the soluble dye released into the supernatant from the labeled bacteria, is a quantitative measurement of enzymatic activity.
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<div class="chapter_text" id="script_5">
	You've just watched JoVE's video on photometric protein determination. This video described the underlying principles of photometric determination, went over general procedures for some common assays, and covered some new advances in techniques. Thanks for watching!
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Photometric Protein Determination: Bradford Assay

Photometrische Proteinbestimmung

Measuring the concentration is a fundamental step of many biochemical assays. Photometric protein determination takes advantage of the fact that the more ...

Density Gradient Ultracentrifugation

Density gradient ultracentrifugation is a common technique used to isolate and purify biomolecules and cell structures. This technique exploits the fact that, in suspension, particles that are more dense than the solvent will sediment, while those that are less dense will float. A high-speed ultracentrifuge is used to accelerate this process in order to separate biomolecules within a density gradient, which can be established by layering liquids of decreasing density in a centrifuge tube.
The video will cover the principles of density gradient ultracentrifugation, including a procedure that demonstrates sample preparation, creation of a sucrose gradient, ultracentrifugation, and collection of fractionated analytes. The applications section discusses isolation of multi-protein complexes, isolation of nucleic acid complexes, and separation using cesium chloride density gradients.
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Density gradient ultracentrifugation is a common approach to isolate and purify cell structures for biochemical experiments. The technique uses a high-speed, or ultra, centrifuge to nondestructively separate cellular components in a density gradient. This video describes the principles of density gradient ultracentrifugation, provides a general procedure using a sucrose gradient, and discusses some applications.
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<div class="chapter_text" id="script_1">
Let&#39;s start by examining the principles of ultracentrifuges and density gradients. A suspension contains particles in a liquid solvent. Because of gravity, particles denser than the solvent sediment out while those less dense than the solvent float. The greater the difference in density between the particle and the solvent, the faster the separation.
An ultracentrifuge contains a unit called a rotor, which rotates at highly controlled speeds, simulating a strong gravitational field. Within this field, the differences in density between particles and the solvent are magnified.
The strength of the field depends on the speed of rotation. Even a small rotor at a relatively low rotational speed can create a force thousands of times stronger than the earth&#39;s gravitational field.
If a tube contains several liquids of different densities, centrifugation will keep them in separate layers in order of density, with the densest liquid closest to the base. Such a layering of multiple liquids is called a &#34;density gradient.&#34; There are two types. In step gradients, liquids of decreasing density are carefully layered on top one another. In continuous gradients, the liquids are mixed in varying proportions, so the density decreases smoothly from the base upwards.
Cellular organelles can be separated using a step gradient, through &#34;isopycnic density-gradient centrifugation.&#34; This is the simplest and most common centrifugation procedure.
This procedure is used to separate the cellular structures. The more dense the organelle, the further it descends-with mitochondria at the top and nucleic acids towards the bottom.
Now that you know the principles behind the technique, let&#39;s see it in the lab.
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<div class="chapter_text" id="script_2">
Before the procedure is started, the manufacturer&#39;s speed and density ratings should be noted, and the ultracentrifuge checked for corrosion. This procedure uses a swinging-bucket rotor.
First, the cellular material is prepared by homogenizing the cells, which nondestructively releases their organelles. The homogenate may be fractionated through preliminary low-speed centrifugation, to remove low-density components. Next, the sucrose solutions are prepared.
Sucrose is added in increasing amounts so each solution is more concentrated, and therefore denser, than the preceding one. The exact densities of the solutions will depend on the components to be separated, which vary between organisms. The solutions should have densities between those of the components to be separated, with the last solution denser than the densest component of the analyte. Techniques for separating components denser than sucrose, like nucleic acids, are described in the applications.
The sucrose gradient is now created in a clean centrifuge tube. A pipette is used to draw up the most concentrated sucrose solution. With the tube held upright, the pipette tip is placed high against the wall, and the liquid dispensed steadily down. It&#39;s important that the working area is kept free of vibrations and other disturbances.
After replacing the tip, the remaining solutions are added in order of decreasing density. They are dispensed carefully to form distinct layers and avoid mixing. Finally, about half a milliliter of the cellular sample is added atop the gradient, and the tube is weighed. This is used to balance the weight distribution, the next step of the process.
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<div class="chapter_text" id="script_3">
Centrifugation should begin as soon as possible. The tube is placed in the rotor, which is then balanced by placing blank solutions of equal weight in opposing slots. The rotor is placed in the ultracentrifuge and the system sealed. The temperature and rotation speed and time are set. Typical values are 4 &#176;C with a force of over 100,000 x g for 16 h.
After centrifugation, the tube is withdrawn from the rotor, taking care to keep it upright and undisturbed. The different cellular components have fractionated into discrete bands between the solution layers. The fractions can be collected with a syringe. Alternately, the bottom of the tube can be punctured with a fine, sterilized needle and the outflow collected in sterile tubes. The cellular components have now been isolated. They can be stored at -80 &#176;C.
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<div class="chapter_text" id="script_4">
Now that we&#39;ve seen the basic procedure, let&#39;s look at some applications.
A typical application is the isolation of multi-protein complexes in plant cells. In this example, complexes responsible for cyclic electron flow are being isolated from the thylakoid, the site of the light reaction in photosynthesis. This procedure uses discrete solutions of 14 to 45% sucrose. Centrifugation occurs over 100,000 x g for 14 h at 4 &#176;C.
Because nucleic acids are denser than sucrose, isopycnic centrifugation cannot separate them from organelles nondestructively.
A different technique, known as &#34;rate-zonal centrifugation&#34; is used. It separates organelles based on their sedimentation rates, which depend not only on their densities, but also on their conformations. A continuous gradient is used to separate the components based on this property.
The procedural steps are similar to those for isopycnic cases. In this example, RNA-ribosome complexes are isolated using a continuous gradient of 5% to 20%, centrifuged at 230,000 x g. Centrifugation is interrupted after a few hours to prevent co-precipitation.
Nucleic acid strands can be separated from each other on the basis of density.
This is because strands rich in guanine and cytosine are denser than those rich in adenine and thiamine. In this case, the gradient cannot be made of sucrose, because sucrose is less dense than nucleic acids. Instead, cesium chloride gradients, typically from 1.65 to 1.75 g/mL are used, as they have sufficient density and a low viscosity.
Here we see plankton DNA being purified using a continuous cesium chloride gradient. Centrifugation occurs at over 1,000,000 x g for 18 h under vacuum.
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<div class="chapter_text" id="script_5">
You&#39;ve just watched JoVE&#39;s video on ultracentrifugation with a sucrose density gradient. You should now understand how a density gradient works, how to construct a step gradient, and how to load and operate an ultracentrifuge. Thanks for watching!
</div>

Density Gradient Ultracentrifugation: Principle and Applications

Dichte Gradienten Ultrazentrifugation

Density gradient ultracentrifugation is a common technique used to isolate and purify biomolecules and cell structures. This technique exploits the fact ...

Co-Immunoprecipitation and Pull-Down Assays

Co-immunoprecipitation (CoIP) and pull-down assays are closely related methods to identify stable protein-protein interactions. These methods are related to immunoprecipitation, a method for separating a target protein bound to an antibody from unbound proteins. In CoIP, an antibody-bound protein is itself bound to another protein that does not bind with the antibody, this is followed by a separation process that preserves the protein-protein complex. The difference in pull-down assays is that affinity-tagged bait proteins replace antibodies, and affinity chromatography is used to isolate protein-protein complexes.
This video explains CoIP, pull-down assays, and their implementation in the laboratory. A step-by-step protocol for each technique is covered, including the reagents, apparatus, and instruments used to purify and analyze bound proteins. Additionally, the applications section of this video describes a procedure to study how myxovirus proteins inhibit influenza nucleoprotein, an investigation into the role of calcium ions in calmodulin via a pull-down assay, and a modified pull-down assay for characterizing transient protein interactions.
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Protein-protein interactions play a significant role in a wide variety of biological functions. The majority of protein-protein interactions and their biological effects have yet to be identified. Co-immunoprecipitation, or CoIP, and pull-down assays are two closely related methods for the identification of stable protein-protein interactions. This video will cover the principles of the two assays, their general laboratory procedures, and applications of these techniques.
</div>
<div class="chapter_text" id="script_1">
CoIP and pull-down assays are variants of immunoprecipitation, a method for selectively isolating a protein species from a complex solution. In an immunoprecipitation experiment, an antibody specific to a target protein is allowed to form an immune complex with that target in the sample. The complex is then captured on a solid support, typically protein A bound to a sepharose bead. Any proteins not captured are removed by centrifugation steps. The protein is then released from the antibody and solid support by boiling in reducing SDS-PAGE sample-loading buffer.
Co-immunoprecipitation is conducted in the same manner, except that intact protein complexes are captured onto the solid support. The antibody binds to the target protein, which, in turn, is bound to another protein that is not targeted by the antibody. As in immunoprecipitation, the protein complex is released from the antibody and solid support by boiling in reducing SDS-PAGE sample loading buffer.
Pull-down assays are similar to co-immunoprecipitation, differing only in the use of a &#34;bait&#34; protein, as opposed to an antibody. Through molecular biology techniques, this bait protein is engineered with an affinity tag, such as a series of histidine residues. These affinity tags are described in &#34;Chromatography-based Biomolecule Separations.&#34; The protein is then captured on an immobilized affinity ligand specific for the tag. The captured protein is then incubated with a sample that contains proteins that form complexes with the &#34;bait&#34;. The protein complex is released from the affinity support by washing with a solution containing a competitive analyte specific for the tag on the &#34;bait&#34; protein. Pull-down assays are useful for confirming protein interactions predicted by co-immunoprecipitation, and for discovering unknown protein interactions.
Now that the principles of co-immunoprecipitation and pull-down assays have been discussed, let&#39;s look at their laboratory procedures.
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<div class="chapter_text" id="script_2">
First let&#39;s discuss co-immunoprecipitation. To a series of microfuge tubes, the following is added: PBS buffer, and a 50% solution of protein A-sepharose, a resin-protein complex that binds to the antibody. The microfuge tubes are rotated to ensure proper distribution, and then the resin is washed with additional PBS buffer. Cell lysate, containing the desired protein, and 2 &#956;g of antibody are added to the microfuge tubes, and the mixture is rotated for 1 h at 4 &#176;C. The beads are pelleted by centrifugation, the supernatant is discarded, and the beads re-washed three times with buffer to remove non-bound protein. The beads containing the antibody-protein complex are in reducing SDS-PAGE sample-loading buffer, for removal of the complex from the antibody and for analysis by SDS-PAGE and immunoblotting.
</div>
<div class="chapter_text" id="script_3">
Now we will discuss the procedure for pull-down assays: The &#34;bait&#34; protein is expressed in a plasmid with the appropriate affinity tag. After reaching log-phase growth, the cells are lysed, and then centrifuged. Suspended streptavidin-sepharose beads, which capture biotin-tagged &#34;bait&#34; protein, are pipetted into a microfuge tube. The beads are then centrifuged and the supernatant carefully removed by aspiration. The beads are then washed with buffer, centrifuged, and the supernatant removed.
Cells containing the putative &#34;prey&#34; protein, which has an affinity for the &#34;bait&#34; protein, are harvested by centrifugation. The supernatant is then added to the microfuge tube containing the resin, and incubated at 4 &#176;C for 3 h on a shaker. The resin is then centrifuged, the supernatant removed, and the resin washed to remove non-bound protein. Elution buffer is added to the resin, and the mixture is incubated at room temperature for 30 min on a shaker. The resin is then centrifuged, and the supernatant containing the desired complex is analyzed by immunoblotting.
</div>
<div class="chapter_text" id="script_4">
Now that we&#39;ve reviewed the procedures, let&#39;s look at some of the useful applications of co-immunoprecipitation and pull-down assays.
Co-immunoprecipitation can be useful in better understanding of enzymes&#39; mechanism of action. Myxovirus-, or Mx-, resistance proteins inhibit a wide range of viruses, including influenza A, for which the mechanism is poorly understood. Co-immunoprecipitation was used to study the interaction between mouse Mx1 protein and influenza nucleoprotein.
Pull-down assays have proven useful in studying the effects of second messengers, which are proteins that communicate a signal from the cellular environment. They are a component of a signaling pathway where multiple proteins interact in response to environmental cues. Calcium ions act as secondary messengers by binding to calmodulin, which, in turn, binds to a wide variety of proteins, mediating many types of biological responses. Without the calcium, the protein can&#39;t bind to the calmodulin. A pull-down assay was conducted to test the ability of proteins to bind to calmodulin in the presence or absence of calcium ions.
Co-immunoprecipitation and pull-down assays are generally used for analyzing stable or strong protein interactions, but not transient ones. A recent development in pull-down assays, the HaloTag, has simplified the study of transient protein interactions; HaloTag is a genetically-encoded protein fusion tag, fused to the protein of interest, capable of chemically reacting with a haloalkane solid support. If functional analysis is desired, the protein complex, minus the tag, in its entirety could then be isolated by incubating with tobacco etch virus protease.
</div>
<div class="chapter_text" id="script_5">
You&#39;ve just watched JoVE&#39;s video on co-immunoprecipitation and pull-down assays. This video described the principles of the two methods, the general laboratory procedures, and some of their applications.
Thanks for watching!
</div>

Co-Immunopräzipitation und Pull-Down-Assays

Co-immunoprecipitation (CoIP) and pull-down assays are closely related methods to identify stable protein-protein interactions. These methods are related ...

Reconstitution of Membrane Proteins

Reconstitution is the process of returning an isolated biomolecule to its original form or function. This is particularly useful for studying membrane proteins, which enable important cellular functions and affect the behavior of nearby lipids. To study the function of purified membrane proteins in situ, they must be reconstituted by integrating them into an artificial lipid membrane.
This video introduces membrane protein reconstitution concepts and related procedures, such as protein isolation using detergent, formation of artificial vesicles using lipids, incorporation of the isolated protein into the artificial vesicle, and separation of the detergent from the solution. Finally, two applications are covered: reconstitution of membrane transport proteins and reconstitution of light-harvesting proteins.&#160;
<div class="chapter_text" id="script_0">
Reconstitution is the process of restoring an isolated biomolecule to its original form or functionality. This approach is often used when studying membrane proteins, which enable many important cellular processes and affect the behavior of neighboring lipids. However, the complexity of the cell environment makes membrane protein functions difficult to study in situ. The proteins can be extracted and purified, but their actual functions cannot be evaluated without a membrane. Therefore, isolated membrane proteins are reconstituted by integration into an artificial lipid membrane, such as a liposome. This video will introduce the principles of membrane protein reconstitution, a general reconstitution procedure, and a few applications in biochemistry.
</div>
<div class="chapter_text" id="script_1">
Cell membranes primarily consist of phospholipids and membrane proteins. The phospholipids form a bilayer in which the hydrophilic phosphate heads interact with the aqueous interior and exterior of the cell, while the hydrophobic fatty acid tails interact with each other in the bilayer.
Some membrane proteins only interact with the membrane by electrostatic or noncovalent interactions. Others, called &#39;integral proteins&#39;, are embedded in the lipid bilayer.
Like the bilayer, integral proteins have hydrophilic ends and a hydrophobic center, and are held in place by hydrophobic interactions. Integral proteins that span the entire membrane are known as &#39;transmembrane proteins&#39;.
The interactions between these proteins and the membrane are so strong that even lysing the cells will not separate them. A special surfactant called a detergent is used to extract the proteins. Similar to phospholipids, detergents have hydrophilic heads and lipophilic tails, and can enter the membrane freely.
Inside the membrane, the lipophilic tails of the detergent interact with the hydrophobic protein core. This surrounds the protein with a shell of the hydrophilic detergent heads, which disrupts the protein-lipid interactions.
The protein-detergent complex is now easily separated from the membrane. The detergent makes the complex soluble in aqueous solutions, and ready for reconstitution in an artificial membrane.
Proteins are often reconstituted in the membranes of liposomes, which are artificial vesicles. To prepare liposomes, dried lipids are hydrated and agitated to induce vesicle formation. When a detergent is added, it is incorporated into the liposome membranes.
To reconstitute the protein, the solubilized proteins and liposomes are combined, and then the detergent is removed from solution by dialysis or chemical adsorption. The proteins and liposomes rapidly assemble into proteoliposomes, so only the hydrophilic groups are exposed. The proteins then function as they would in a cell membrane, and can be investigated in isolation.
Now that we&#39;ve covered the basics of protein reconstitution, let&#39;s go over a protocol for reconstituting membrane proteins in liposomes.
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<div class="chapter_text" id="script_2">
To begin isolating the membrane proteins, the cells are lysed. Unbroken cells are removed with centrifugation.
The supernatant is centrifuged at a higher speed to pellet the membranes. The pellet is re-suspended and a detergent is added to extract the proteins.
The remaining cell debris is removed by additional centrifugation. The protein is purified from the supernatant with column chromatography and then concentrated or purified further as needed.
To begin preparing the liposomes, a suspension of phospholipids in organic solvent is dried under nitrogen or argon.
The phospholipids are hydrated with hydration buffer, and the mixture is sonicated to finish creating the liposomes.
Detergent is added to solubilize the liposomes, which is then combined with the proteins.
The detergent is then removed by adsorption onto polystyrene beads, dialysis, or a detergent-binding column. The resulting proteoliposomes are ready to be purified and used in subsequent experiments.
</div>
<div class="chapter_text" id="script_3">
Now that you are familiar with the basics of a membrane protein reconstitution procedure, let&#39;s look at a few applications of protein reconstitution in biochemistry.
A membrane-transport protein was reconstituted to gain a clearer understanding of its transport mechanism. Its function post-reconstitution was verified with an efflux of iodide ions. Then, the transport activity was studied in the presence of various small molecule ion channel inhibitors and potentiators. In this way, the direct interactions of these small molecules with the transport protein could be studied.
Chlorophyll and carotenoid-binding membrane proteins in plants harvest light, promote charge separation, and mitigate light damage. By reconstituting these light-harvesting proteins, their folding dynamics and interaction with pigments can be studied. The light-harvesting proteins reconstituted with this technique had very similar optical properties to the native proteins. Fluorescence emission spectroscopy can then be used to study energy transfer from pigments to the reconstituted light-harvesting proteins.
</div>
<div class="chapter_text" id="script_4">
You&#39;ve just watched JoVE&#39;s video on reconstitution of membrane proteins. Reconstitution is a way to transfer important proteins to a cell mimic for further investigation. This video covered the principles of protein reconstitution, a reconstitution protocol, and a few applications in biochemistry. Thanks for watching!
</div>

Extraction and Reconstitution of Membrane Proteins in Liposomes

Rekonstitution von Membranproteinen

Reconstitution is the process of returning an isolated biomolecule to its original form or function. This is particularly useful for studying membrane ...

F&#246;rster Resonance Energy Transfer (FRET)

F&#246;rster resonance energy transfer (FRET) is a phenomenon used to investigate close-range biochemical interactions. In FRET, a donor photoluminescent molecule can non-radiatively transfer energy to an acceptor molecule if their respective emission and absorbance spectra overlap. The amount of energy transferred&#8212;and consequently the overall emission of sample&#8212;depends on the proximity of an acceptor-donor pair of photoluminescent molecules. FRET analysis is combined with other biochemistry techniques to obtain detailed information of biomolecular structures and interactions from this &#8220;spectroscopic ruler.&#8221;
This video covers the principles and concepts of FRET analysis. The procedure focuses on preparing samples for FRET and ways to present and interpret data. Finally, the applications include monitoring conformational and cellular processes by labeling parts of a cell or protein, monitoring enzyme reactions that alter protein structures, and using FRET to monitor aggregation of monomers expressed by cells.
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F&#246;rster Resonance Energy Transfer, or FRET, is a non-radiative transfer of energy between light-emitting molecules, and is often used to investigate close-range biochemical interactions. FRET only occurs when fluorescent molecules are spaced within 10 nm of each other. FRET analysis can be combined with other techniques to obtain detailed structural information. This video will introduce the underlying principles of FRET, summarize a protocol and data presentation, and discuss some biochemical applications.
</div>
<div class="chapter_text" id="script_1">
A photoluminescent molecule such as a fluorophore is excited by absorbing electromagnetic radiation at a wavelength in its absorption spectrum. As it relaxes, it emits light at a wavelength within its emission spectrum. For more information about fluorescence, see JoVE&#39;s video on fluorescence microscopy. Different fluorophores absorb and emit light at different wavelengths, which frequently overlap. If the emission spectrum of a fluorophore overlaps significantly with the absorption spectrum of another fluorophore, the &#8220;donor&#8221; will release a virtual photon, which is absorbed by the &#8220;acceptor&#8221;. When an excited donor is within 10 nm of an acceptor, energy is transferred from donor to acceptor by dipole-dipole interactions. The release of energy by emission of light from the donor correspondingly decreases. Meanwhile, the excited acceptor emits light at its emission wavelength. The FRET response is evaluated in terms of efficiency, or the percentage of energy released from the donor by FRET rather than by fluorescence or other radiative processes. The efficiency depends strongly on the distance between the donor and acceptor, which allows FRET to act as a &#39;molecular&#39; or &#39;spectroscopic&#39; ruler.
In biochemistry, FRET is often used qualitatively to observe conformational changes in molecules by monitoring fluorophores as they move in and out of FRET range of each other. Similarly, cellular functions can be studied with molecules containing a FRET pair. If the labeled molecule is cleaved by enzyme activity, FRET stops and the observed fluorescence wavelength changes.
Now that you understand the principles behind FRET, let&#39;s look at an overview of a protocol and a few ways to present and interpret the data.
</div>
<div class="chapter_text" id="script_2">
Prior to the experiment, the biomolecules of interest, typically DNA or proteins, are engineered with fluorescent tags, using molecular biology techniques.&#160;Common ways to introduce the modified genetic material into the cells include transfection and electroporation.
Then, the cells are prepared for FRET visualization on a fluorescence microscope.&#160;For instance, the molecules may be immobilized on a slide for single-molecule FRET, or samples are loaded into wells for high-throughput screening.
Then, the excitation lasers, microscope, and associated equipment are prepared. (A) FRET experiments often involve powerful lasers; (B) so appropriate PPE and safety procedures should be used.&#160;The sample is then placed in the instrument and illuminated with the excitation laser.
</div>
<div class="chapter_text" id="script_3">
For experiments monitoring cell behavior, color images showing differences or changes in emission intensity are used. Donor and acceptor emission intensities are plotted together to track FRET response over time.
FRET data can also be fitted to various functions for more complex analyses. Depending on the experiment, data may be presented in multiple ways to best represent the results, making FRET a flexible experimental tool.
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Now that you&#39;re familiar with the basics of running and analyzing a FRET experiment, let&#39;s look at some applications of FRET in biochemistry research.
FRET can be used to study conformational changes or cellular processes by labeling parts of the protein or cell predicted to move within 10 nm of each other with a FRET pair. For example, protein sensors are prepared by labeling receptors with a pair of fluorophores. The FRET response is monitored live by confocal microscopy. Variation of emission wavelength and intensity indicate conformational changes.
FRET can also be used by preparing molecules with an active FRET pair and observing changes in the response. When the substrate is cleaved, FRET is disrupted, causing an increase in donor emission and a decrease in acceptor emission. The emissions are analyzed to determine contributions by donor, acceptor, and FRET. Once the direct emission factors are calculated for the cyan and yellow fluorescent proteins, the concentration and kinetic parameters of the substrate can be determined.
Cells designed to express monomers containing either of a FRET pair function as &#39;sensors&#39; for interactions between those monomers. If aggregation of those monomers is induced, a FRET response is observed. This can be used to investigate protein aggregation triggered by &#39;seeding&#39; of misfolded proteins. Here, cells were transduced with aggregates of the protein of interest, incubated, and analyzed with flow cytometry.
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You&#39;ve just watched JoVE&#39;s video on F&#246;rster Resonance Energy Transfer, or FRET. This video contained the underlying principles of FRET, preparation and analysis of a FRET experiment, and a few biochemical applications.
Thanks for watching!
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F&amp;#246;rster Resonance Energy Transfer (FRET)

Förster Resonance Energietransfer (FRET)

F&#246;rster resonance energy transfer (FRET) is a phenomenon used to investigate close-range biochemical interactions. In FRET, a donor photoluminescent ...

Surface Plasmon Resonance (SPR)

Surface plasmon resonance (SPR) is the underlying optical phenomenon behind label-free biosensors to evaluate the molecular affinity, kinetics, specificity, and concentration of biomolecules. In SPR, biomolecular interactions occur on a biosensor made of a thin layer of metal on a prism. Real-time interactions of biomolecules can be monitored by measuring the changes of light reflected off the underside of the metal.
This video describes the basic concepts of SPR and how it is used to analyze and visualize biomolecular interactions. This is followed by a sample preparation and experimental protocol for investigating binding rates using SPR. In the applications section, SPR imaging, localized SPR, and quantum dot enhanced SPR are explored.
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Surface plasmon resonance, or SPR, is the underlying phenomenon behind certain label-free biosensors for evaluating binding and adsorption interactions of biomolecules. Binding assays that require labeling, such as ELISA, can be a time-consuming process, and may alter the functionality of the analyte. In SPR, biomolecular interactions occur on a special sensor made of a thin layer of metal on one face of a prism. By monitoring the changes in light reflected off of the underside of the metal, SPR instruments visualize these interactions in real-time without the use of labels. This video will introduce the principles of SPR, a general procedure for SPR imaging, and some applications of in biochemistry.
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An SPR sensor is usually made of a thin layer of a noble metal atop the face of a prism. To take readings from the sensor, light is reflected off of the prism-metal interface into a photodetector. The reflected light will have a high intensity except at a certain angle, related to the electronic properties of the metal surface, known as the &#34;surface plasmon resonance angle&#34;.
As molecules bind to the surface, the electronic properties of the metal change, which in turn adjusts the angle. As new proteins attach, forming complexes, the angle will shift further. By measuring relative changes in the SPR angle, interactions like these can be monitored in real-time.
Another technique called localized, or &#34;L&#34;SPR, uses metal nanoparticles as the sensor surface. The properties that affect the SPR angle are highly localized to each nanoparticle, which improves sensitivity and signal resolution.
When investigating binding interactions with standard SPR, the sensor is generally mounted in a platform that becomes the floor of a flow cell in the instrument. The biomolecules of interest are carried through the flow cell by buffer solution. The sensor surface is often first coated with a substrate that has a high affinity for the metal. This ensures that a significant amount of ligand, which in turn binds to the analyte of interest, will be immobilized onto the sensor and reduces the likelihood that the ligand will dissociate during the procedure.
Once the ligand is immobilized on the sensor, the analyte is flowed over the sensor in buffer. By monitoring the change in the SPR angle over time as the analyte binds to the ligand, the binding rate and other kinetic information can be calculated.
The reflectance data can also be used for SPR imaging, or SPRi, by directing the reflected light to a CCD detector. This produces a high-contrast, high-resolution image of the entire sensor surface. Using SPR and the related techniques, questions can be answered about molecular affinity, kinetics, specificity, and concentration.
Now that you understand what is being measured in an SPR experiment, let&#39;s look at a procedure for investigating binding rates.
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Before beginning the procedure, the running and sample buffers must be prepared. The running buffer is used to deposit the ligand onto the sensor, and the sample buffer is used to deposit the analyte. The sensor chip is carefully cleaned and loaded into a sheath. Then, the device is placed into the instrument, where it becomes the bottom of the flow cell. The instrument software is set up for the experiment and subsequent analysis. If necessary, the sensor surface is primed with a substrate to capture the ligand. The ligand is flowed over the sensor surface in the running buffer, where it is captured by the substrate on the sensor surface.
Then, the analyte in the sample buffer is run through the flow cell, where it selectively binds to the immobilized ligand. The change in reflectance is plotted and compared against controls to determine rate constants and other reaction kinetics data for the investigated reaction.
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Now that you understand how an SPR experiment is performed, let&#39;s look at a few other applications of SPR in biochemistry.
Here, SPR imaging was used to evaluate proteins with an array of eleven receptors on a sensor. 3D graphs of reflectivity versus time and receptor concentration were prepared from the reflectivity data for each protein. These &#34;profiles&#34; are characteristic to each protein, and thus could subsequently be used for protein identification.
In this experiment, cell secretions were studied using a custom-made LSPR sensor. The sensor was also compatible with SPRi and fluorescence microscopy. Upon depositing the cell on the sensor, the interaction of cell secretions with the nanoparticle array could be measured with high spatial resolution.
Here, the use of quantum dots, nanoscale semiconductors, as an SPR signal enhancement agent mixed with the analyte was investigated. This enhanced &#34;nano-SPRi&#34; method was compared to assays by standard SPRi and the ELISA method. The nano-SPRi method significantly improved the sensitivity and limit of detection while still being less time-consuming than the ELISA method.
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<div class="chapter_text" id="script_4">
You&#39;ve just watched JoVE&#39;s video on surface plasmon resonance. This phenomenon is used to monitor and image biomolecular interactions without the use of labels. This video introduced the principles of SPR, a typical protocol for performing an SPR experiment, and a few applications of SPR in biochemistry.
Thanks for watching!
</div>

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