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Research

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: 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 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 ...

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: 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 ...

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 ...

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:  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 ...

&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 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: 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 ...

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: 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 ...

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: 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: 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: 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: Branched Alkane Fragmentation

This lesson delves into the mass spectrometry of branched alkane fragmentation. Branched alkanes possess secondary or tertiary carbon atoms, which generate relatively stable carbocations if the cleavage occurs at the branching point. The high stability of carbocations drives the instant fragmentation of branched alkanes. Accordingly, the branched alkane&#39;s molecular ion peak is very weak or invisible in the mass spectra, especially in comparison to a linear alkane.
<img alt="Figure1" src="/files/ftp_upload/13041/13041-PT_Figure_1.svg" style="text-align: center; height: 275px;" />
Figure 1. Fragmentation pathway of 2-methylbutane (top), neopentane (middle), and n-pentane (bottom) molecular ions.
Figure 1 shows the most feasible fragmentation pathway observed in 2-methyl butane, neopentane, and n-pentane molecular ions. 2-methyl butane and neopentane fragment to yield secondary and tertiary carbocations, respectively. The stability of these carbocations drives the fragmentation reaction, even though the co-produced methyl radical is relatively unstable. In contrast, the cleavage of n-pentane leading to the methyl radical is difficult since the stability of a primary carbocation is low.
<img alt="Figure2" src="/files/ftp_upload/13041/13041-PT_Figure_2.svg" style="text-align: center; height: 105px;" />
Figure 2. Fragmentation of 2,2-dimethylpentane.
As shown in Figure 2, the fragmentation of 2,2-dimethyl pentane involves the loss of either methyl or propyl radical, resulting in a tertiary carbocation. Here, the stability of the co-produced radical determines the cleaving bond. So, the cleavage that produces a propyl radical is favored, and the signal from the 2-methyl propyl carbocation becomes the base peak.

This lesson delves into the mass spectrometry of branched alkane fragmentation. Branched alkanes possess secondary or tertiary carbon atoms, which ...

Flow Cytometry and Fluorescence-Activated Cell Sorting (FACS): Isolation of Splenic B Lymphocytes

Source: Perchet Thibaut1,2,3, Meunier Sylvain1,2,3, Sophie Novault4, Rachel Golub1,2,3 
1 Unit for Lymphopoiesis, Department of Immunology, Pasteur Institute, Paris, France 
2 INSERM U1223, Paris, France 
3 Universit&#233; Paris Diderot, Sorbonne Paris Cit&#233;, Cellule Pasteur, Paris, France 
4 Flow Cytometry Platfrom, Cytometry and Biomarkers UtechS, Center for Translational Science, Pasteur Institute, Paris, France
The overall function of the immune system is to defend the body against infectious organisms and other invaders. White blood cells, or leukocytes, are the key players of the immune system. Upon infection, they are activated and initiate an immune response. Leukocytes can be divided into various sub-populations (e.g., myeloid cells, lymphocytes, dendritic cells) based on different parameters that can be biological, physical, and/or functional (e.g., size, granularity, and secretion). One way to characterize leukocytes is through their surface proteins, which are mainly receptors. Each leukocyte population expresses a specific combination of receptors (e.g., cytotoxic, activating, migration receptors) that can define subsets among populations. As the immune system encompasses a wide range of cell populations, it is essential to characterize them to decipher their participation in the immune response.
Flow cytometry (FC or FCM) is a widely used method for analyzing the expression of cell surface and intracellular molecules, characterizing and defining different cell types in a heterogenous cell mixture. Flow cytometers are composed of three main sub-systems: fluidics, optics, and electronics. The fluidics system transports the cells in a stream such that they pass in front of a laser one by one. The optics system consists of light sources (lasers) to illuminate the particles, optical filters to direct the resulting light, and fluorescent signals to appropriate detectors. Finally, the electronics system converts the detected light signals into electronic signals that can be processed by the computer. As an individual cell passes in front of the laser beam, it scatters light. A detector in front of the beam measures forward scatter (FS) and several detectors to the side measure side scatter (SC). The FS correlates with cell size and SC is proportional to the granularity of the cells. In this manner, cell populations can often be distinguished based on differences in their size and granularity alone.
In addition to analyzing a cell&#39;s size, shape, and complexity, flow cytometry is widely used for detecting the expression of cell surface receptors (1). This is accomplished by using fluorochrome-labelled monoclonal antibodies which bind to known cell-specific receptors. Upon excitation, these bound fluorochromes emit a light of specific wavelength, called emission wavelength, which can be detected and scored. Fluorescence measurements provide quantitative and qualitative data about fluorochrome-labeled cell surface receptors. Hematologists were first to use FC for the therapeutic follow up of immune cell populations (2). Now, it is used for a wide range of applications such as immunophenotyping, cell viability, gene expression, cell counting, and GFP analysis.
FACS (Fluorescent Activated Cell Sorter) is a specialized type of flow cytometry, which sorts a population of cells into subpopulation using fluorescent labelling. Just like conventional flow cytometry, first FS, SC, and fluorescent data are collected. Then, the machine applies a charge (negative or positive) and an electrostatic deflection system (electromagnets) facilitates the collection of charged droplets containing cells into appropriate tubes.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10494/10494fig1.jpg" /> 
Figure 1: Schematic representation of FACS. Sample (1) is aspirated in the FACS (2) and passed in front of the laser (3). Cell fluorescence is sensed by fluorescence detectors (4). Finally, cells are incorporated in droplets and the cells of interest are deflected by deflection plates (5) and collected in a collection tube (6). The remaining cells go into the trash (7). <a href="https://www.jove.com/files/ftp_upload/10494/10494fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The sorting aspect of the FACS presents many advantages. Many tests can help understand the role of specific cells in immune system, such as analyses of gene expression like RT-qPCR, cell cycle, or cytokine secretion. However, cells should be purified upstream to obtain clear and specific results. Here, FACS comes in useful and the desired cells can be sorted with great purity, yielding highly reliable and reproducible results. FACS can also be used to sort cells based on nuclear or other intracellular staining and according to the presence, absence, and density of surface receptors. FACS is now a standard technique for the purification of subpopulations of cells and has the ability sort up to four populations simultaneously.
This lab exercise demonstrates how to isolate splenic leukocytes and then how to specifically sort B lymphoid cells from the splenic leukocyte cell mixture using FACS.

Flow Cytometry and FACS: Isolation of Splenic B Lymphocytes

Cytométrie en flux et tri cellulaire activé par fluorescence (FACS) : isolement des lymphocytes B spléniques

Source: Perchet Thibaut1,2,3, Meunier Sylvain1,2,3, Sophie Novault4, Rachel Golub1,2,3
1 Unit for Lymphopoiesis, Department of Immunology, Pasteur ...

Magnetic Activated Cell Sorting (MACS): Isolation of Thymic T Lymphocytes

Source: Meunier Sylvain1,2,3, Perchet Thibaut1,2,3, Sophie Novault4, Rachel Golub1,2,3 
1 Unit for Lymphopoiesis, Department of Immunology, Pasteur Institute, Paris, France 
2 INSERM U1223, Paris, France 
3 Universit&#233; Paris Diderot, Sorbonne Paris Cit&#233;, Cellule Pasteur, Paris, France 
4 Flow Cytometry Platfrom, Cytometry and Biomarkers UtechS, Center for Translational Science, Pasteur Institute, Paris, France
Defence against pathogens depends on surveillance by the immune system. This system is complex and comprises many cell types, each one with specific functions. This complex composition enables immune responses to a large diversity of pathogens and injuries. Adaptive immunity allows specific responses against specific pathogens. The majority of cells responsible for this type of immunity are the lymphocytes (B cells and T cells). Usually, B cells respond to extracellular infections (such as bacterial infections), and T cells respond to intracellular infections (such as viral infections). The different types of cells in lymphocyte populations can be characterized by the combination of cell surface proteins they express and/or by a panel of secreted cytokines.
Magnetic sorting allows enrichment of targeted cell populations using magnetic properties and expression of one or several cell surface proteins (1, 2). This technique consists of three steps. First, the cells are incubated with magnetic beads that are coupled with one or several monoclonal-specific antibodies. Cells that express surface proteins that bind to these antibodies attach to the magnetic beads. Then, the targeted cell populations are captured with a magnet. To finish, the targeted cells are eluted from the magnet. At the end, two sorting products are obtained, one containing unlabeled cells and the second containing the target cells coupled with the magnetic beads. Columns can be used to improve the efficiency of magnetic sorting. In the column, a non-magnetic element lengthens the path of cell through the column. Hence, cell flow is slowed down, facilitating cell capture by the magnet.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10495/10495fig1.jpg" /> 
Figure 1: Schematic representation of magnetic separation. Thymic leukocytes are stained with anti-CD3 biotinylated antibodies. After washing, streptavidin (SAV) coupled beads specifically fix the biotin on anti-CD3 antibodies. (1) Cells are transferred in a column. (2) The magnet does not retain unlabeled cells, while CD3-positive cells remain in the column. Finally, the column is separated from the magnet and (3) CD3-positive cells are eluted in medium. <a href="https://www.jove.com/files/ftp_upload/10495/10495fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
There are two types of magnetic sorting (3). In positive sorting, cells of interest are captured with the magnetic beads. In negative sorting, unwanted cells are removed by capturing with the magnetic beads carrying the appropriate antibodies. This MACS technique permits good enrichment of targeted cells and improves the percentage of recovered cells from 1-20% to 60-98% in an organ. After sorting, it is necessary to verify cell purity and sorting by different methods (e.g. flow cytometry). The MACS technique is ideal to enrich a target population for other experiments such as cell culture or cell cycle analysis.
In this lab exercise, we demonstrate how to isolate thymic leukocytes and thereafter enrich thymic CD3-positive cells from the mix using magnetic cell sorting technique.

MACS: Isolation of Thymic T Lymphocytes

Tri cellulaire magnétique (MACS) : isolement des lymphocytes T thymiques

Source: Meunier Sylvain1,2,3, Perchet Thibaut1,2,3, Sophie Novault4, Rachel Golub1,2,3
1 Unit for Lymphopoiesis, Department of Immunology, Pasteur ...

ELISA Assays: Indirect, Sandwich, and Competitive

Source: Whitney Swanson1,2, Frances V. Sjaastad2,3, and Thomas S. Griffith1,2,3,4 
1 Department of Urology, University of Minnesota, Minneapolis, MN 55455 
2 Center for Immunology, University of Minnesota, Minneapolis, MN 55455 
3 Microbiology, Immunology, and Cancer Biology Graduate Program, University of Minnesota, Minneapolis, MN 55455 
4 Masonic Cancer Center, University of Minnesota, Minneapolis, MN 55455
Enzyme-linked immunosorbent assay (ELISA) is frequently used to measure the presence and/or concentration of an antigen, antibody, peptide, protein, hormone, or other biomolecule in a biological sample. It is extremely sensitive, capable of detecting low antigen concentrations. The sensitivity of ELISA is attributed to its ability to detect the interactions between a single antigen-antibody complex (1). Moreover, the inclusion of an enzyme-conjugated antigen-specific antibody permits the conversion of a colorless substrate into a chromogenic or fluorescent product that can be detected and easily quantitated by a plate reader. When compared to the values generated by titrated amounts of a known antigen of interest, the concentration of the same antigen in the experimental samples can be determined. Different ELISA protocols have been adapted to measure antigen concentrations in a variety of experimental samples, but they all have the same basic concept (2). Choosing the type of ELISA to perform, indirect, sandwich, or competitive, depends on a number of factors, including the complexity of the samples to be tested and the antigen-specific antibodies available to use. The indirect ELISA is frequently used to determine the outcome of an immunological response, such as measuring the concentration of an antibody in a sample. The sandwich ELISA is best suited for analyzing complex samples, such as tissue culture supernatants or tissue lysates, where the analyte, or antigen of interest, is part of a mixed sample. Finally, the competitive ELISA is most often used when there is only one antibody available to detect the antigen of interest. Competitive ELISAs are also useful for detecting a small antigen with only a single antibody epitope that cannot accommodate two different antibodies due to steric hinderance. The protocol will describe the basic procedures for the indirect, sandwich, and competitive ELISA assays.
The indirect ELISA assay is commonly used to measure the amount of antibodies in serum or in the supernatant of a hybridoma culture. The general procedure for the indirect ELISA assay is:
<ol>
	<li>Coat wells with antigens</li>
	<li>Add serum or hybridoma culture supernatant containing antibody (primary or 1&#176; antibody)</li>
	<li>Incubate and wash</li>
	<li>Add secondary (or 2&#176;) enzyme-conjugated antibody</li>
	<li>Incubate and wash</li>
	<li>Add substrate</li>
</ol>
The sandwich ELISA assay differs from the indirect ELISA assay in that the method does not involve coating the plates with a purified antigen. Instead, a &#34;capture&#34; antibody is used to coat the wells of the plate. The antigen is &#34;sandwiched&#34; between the capture antibody and a second &#34;detection&#34; enzyme-conjugated antibody - where both antibodies are specific for the same antigen, but at different epitopes (3). By binding to the capture antibody/antigen complex, the detection antibody remains in the plate. Either monoclonal antibodies or polyclonal antisera can be used as the capture and detection antibodies. The main advantage of the sandwich ELISA is that the sample does not have to be purified before analysis. Moreover, the assay can be quite sensitive (4). Many commercially available ELISA kits are of the sandwich variety and use tested, matched pairs of antibodies. The general procedure for the sandwich ELISA assay is:
<ol>
	<li>Coat wells with capture antibody</li>
	<li>Add test samples containing antigen</li>
	<li>Incubate and wash</li>
	<li>Add enzyme-conjugated detection antibody.</li>
	<li>Incubate and wash</li>
	<li>Add substrate</li>
</ol>
Most commercially available sandwich ELISA kits come with enzyme-conjugated detection antibodies. In cases where an enzyme-conjugated detection antibody is not available, a secondary enzyme-conjugated antibody specific for the detection antibody can be used. The enzyme on the secondary antibody performs the same role, which is to convert the colorless substrate to a chromogenic or fluorescent product. The above-mentioned secondary enzyme-conjugated antibody would more like to be used in a &#34;homemade&#34; sandwich ELISA developed by an investigator who has generated their own monoclonal antibodies, for example. One drawback to using a secondary enzyme-conjugated antibody is to be sure it only binds to the detection antibody, and not the capture antibody bound to the plate. This would result in a measurable product in all wells, regardless of the presence or absence of antigen or detection antibody.
Finally, the competitive ELISA assay is used to detect soluble antigens. It is simple to perform, but it is only suitable when the purified antigen is available in a relatively large amount. The general procedure for the competitive ELISA assay is:
<ol>
	<li>Coat wells with antigen</li>
	<li>Incubate and wash</li>
	<li>Preincubate test sample with enzyme-conjugated primary antibodies</li>
	<li>Add mixture to well</li>
	<li>Incubate and wash away any unbound enzyme-conjugated primary antibody</li>
	<li>Add substrate</li>
</ol>
The &#34;competition&#34; in this assay comes from the fact that more antigen in the test sample used in step 3 will result in less antibody available to bind to the antigen coating the well. Thus, the intensity of the chromogenic/fluorogenic product in the well at the end of the assay is inversely related to the amount of antigen present in the test sample.
A key component in any type of ELISA is the titrated standards of known concentrations that will allow the user to determine the antigen concentration present in the test samples. Typically, a series of wells are designated for creating a standard curve, where known amounts of a purified recombinant protein are added to the wells in decreasing amounts. When these wells are processed at the same time as the test samples, the user can then have a reference set of absorbance values obtained from a microplate reader for known protein concentrations to go along with the absorbance values for the test samples. The user can then calculate a standard curve to which the test samples can be compared for determining the amount of protein of interest present. The standard curve can also determine the degree of precision of the user&#39;s dilution making.
Finally, the last step in each of the ELISA types listed above calls for the addition of a substrate. The degree of conversion of the substrate to product is directly related to the amount of enzyme present in the well. Horseradish peroxidase (HRP) and alkaline phosphatase (AP) are the most common enzymes found conjugated to antibodies. As expected, there are a number of substrates available specific for either enzyme that produce a chromogenic or fluorescent product. Moreover, substrates are available in a range of sensitivities that can increase the overall sensitivity of the assay. The user must also take into consideration the type of instrumentation available for reading the plate at the end of the experiment when picking the type of substrate to use, along with its corresponding enzyme-conjugated antibody.
Commonly used chromogenic substrates for HRP include 2,2&#39;-Azinobis [3-ethylbenzothiazoline-6-sulfonic acid]-diammonium salt (ABTS) and 3,3&#39;,5,5&#39;-tetramethylbenzidine (TMB), whereas p-Nitrophenyl Phosphate (PNPP) is used for AP. ABTS and TMB produce water-soluble green and blue colored reaction products, respectively. The green ABTS product has two major absorbance peaks, 410 and 650 nm, while the blue TMB product is best detected at 370 and 652 nm. The colors of ABTS and TMB change to yellow upon the addition of an acidic stop solution, which is best read at 450 nm. Color development for ABTS is slow, while it is fast for TMB. TMB is more sensitive than ABTS, and may produce a higher background signal if the enzymatic reaction proceeds too long. PNPP produces a yellow water-soluble product after AP conversion that absorbs light at 405 nm.

Tests ELISA : Indirect, en sandwich et par compétition

Source: Whitney Swanson1,2, Frances V. Sjaastad2,3, and Thomas S. Griffith1,2,3,4
1 Department of Urology, University of Minnesota, Minneapolis, MN 55455
...

ELISPOT Assay: Detection of IFN-&#947; Secreting Splenocytes

Source: Tonya J. Webb1 
1 Department of Microbiology and Immunology, University of Maryland School of Medicine and the Marlene and Stewart Greenebaum Comprehensive Cancer Center, Baltimore, Maryland 21201
ELISPOT is a standardized, reproducible assay used to detect cellular immune responses. The assay utilizes an enzyme-linked immunosorbent assay (ELISA)- based method to detect single-cell immune responses which can be visualized by spots, hence the name ELISPOT. ELISPOT was first described in 1983, by Czerkinsky, as a method of enumerating the number of B cell hybridomas producing antigen-specific immunoglobulins (1). The same group further developed the assay to measure the frequency of cytokine producing T lymphocytes. Now ELISPOT has become a gold standard for measuring antigen-specific T cell immunity in clinical trials and vaccine candidates. For example, after vaccination or during an infection, plasma cells and memory B cells secrete antibodies that provide protection. Typically, these B cell responses are assessed by measuring serum titers of antigen-specific antibodies. However, this type of analysis, typically measured by ELISA, may not include memory B cells, which can be present even in the absence of detectable serum antibody levels. Furthermore, it has been well-established that circulating memory B cells are important for the rapid and protective antibody response observed following pathogen re-exposure, thus it is critical to be able to detect these cells. Therefore, to clearly assess antigen-specific memory B-cell responses, both ELISA and ELISPOT should be used (2).
ELISPOT assay uses a plate containing membrane-lined wells that are coated with antibodies in order to capture secreted proteins of interest. Then, the plate is loaded with cells and stimuli to induce protein production. The secreted proteins are captured by the antibodies coated on the surface. After appropriate incubation time, cells are removed and the secreted molecule is detected by using a biotinylated antibody that is specific for a different epitope, as compared to the capture antibody. Next, streptavidin peroxidase is added, followed by the addition of a substrate that allows the detection of the spots (Figure 1). The strength of this assay is that it allows one to quantitate the number of cells producing the protein of interest. Importantly, one can assess if there are changes in the total number of cells producing a specific protein or if individual cells within a population are producing more protein. Moreover, it can provide information regarding kinetics and can be used to assess overall immune activation (mitogen stimulation) relative to antigen-specific responses (antigen simulation). The ELISPOT assay will permit the detection one activated cell amongst 300,000 cells following mitogenic or antigen-specific activation.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10497/10497fig1.jpg" /> 
Figure 1: ELISPOT protocol overview.
The major advantages of this assay are its- a. Simplicity- the protocol is relatively simple and straightforward. It does not require technical expertise, b. Sensitivity- it permits the detection of immune cells at the single cell level and requires very few cells compared to other methods such as flow cytometry, c. Functionality- it provides quantitative data regarding immune function.
This lab exercise demonstrates the ELISPOT protocol for detection of IFN-&#947; secreting-splenocytes, but as mentioned above this assay can also be used to assess antibody secretion by B cells (3).

ELISPOT Assay: Detection of IFN-&amp;#947; Secreting Splenocytes

Test ELISPOT : Détection des splénocytes sécrétants l'IFNgamma

Source: Tonya J. Webb1
1 Department of Microbiology and Immunology, University of Maryland School of Medicine and the Marlene and Stewart Greenebaum ...

Immunohistochemistry and Immunocytochemistry: Tissue Imaging via Light Microscopy

Source: Michael S. Lee1 and Tonya J. Webb1 
1 Department of Microbiology and Immunology, University of Maryland School of Medicine and the Marlene and Stewart Greenebaum Comprehensive Cancer Center, Baltimore, Maryland 21201
Immunohistochemistry (IHC) and immunocytochemistry (ICC) are techniques used to visualize the expression and localization of specific antigens using antibodies. The first published use of IHC was in 1941 when Albert Coons used the technique to visualize the presence of pneumococcal antigen in tissue sections from mice infected with Pneumococcus (1). The name, immunohistochemistry, is derived from the roots &#34;immuno-,&#34; in reference to antibodies, and &#34;histo-,&#34; in reference to the tissue sections used in IHC. The root &#34;cyto-&#34; in immunocytochemistry highlights the key difference between ICC and IHC. Whereas IHC uses sections of whole tissue, ICC uses cells that have been isolated from tissue or grown in culture. The difference in samples used means sample preparation technically differs between IHC and ICC, but otherwise the protocols for ICC and IHC are identical and one will find the terms are frequently used interchangeably.
In both IHC and ICC, antibodies with either chemical or fluorescent tags, such as peroxidase or rhodamine, respectively, are used to visualize the distribution of any antigen of interest through specific binding of the tagged antibody to the antigen. In the case of IHC, thin slices of tissue are immobilized on a slide to maintain the structure of the tissue before being stained, allowing the visualization of antigens in the context of whole tissues (Figure 1). In the case of ICC, cells are distributed evenly on a slide before being stained, allowing the visualization of antigen distribution within individual cells but not within the structure of any specific tissue. Due to the similarities between the two protocols, this protocol will focus on IHC to address the additional complexities of sample preparation involved in IHC.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10498/10498fig1.jpg" /> 
Figure 1: Outline of IHC Protocol. Visual outline of an IHC protocol for paraffin-embedded tissue dissected from a mouse. This protocol uses a biotinylated secondary antibody and strepavidin-HRP to visualize the location of antibody binding. Other options, such as fluorescently tagged antibodies, are also possible. <a href="https://www.jove.com/files/ftp_upload/10498/10498fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The first major decision when performing IHC is how to prepare the tissue sections in order to maintain the structure of the tissue throughout the staining process. The two main choices are formalin-fixed sections of paraffin-embedded tissue or fresh sections of frozen tissue. There is no simple answer as to which method to use as it depends on what downstream analysis will be conducted. Formalin-fixation of paraffin embedded tissues is generally thought to better preserve tissue morphology for optimal imaging while freezing fresh tissue can preserve protein function for subsequent assays outside of IHC. In addition, fresh frozen tissue sections have been shown to be more suitable for gene expression analysis (2). A third consideration is whether or not the antibodies for your antigen of interest are suitable for either fixed or frozen tissue sections, as some antibodies have only been optimized for a specific type of section and may not work for others. Finally, one also needs to determine how long they need to store the tissue sections, as fresh frozen samples must be kept at -80&#176;C and may not last beyond one year while fixed sections can be stored for much longer at room temperature. These are a few of the major considerations for determining whether to use formalin-fixed sections of paraffin-embedded tissue or fresh sections of frozen tissue. Ultimately, if one has enough tissue, it may be best just to have some of both.
In this experiment, we set out to determine if cyclin D1 expression was increased in enlarged spleens from a spontaneous mouse model of lymphoma development. Splenic tissue samples were first isolated from either wild-type mice, transgenic mice that do not have lymphoma, or transgenic mice that have spontaneously developed lymphoma. The spleen tissue samples were fixed in paraformaldehyde, embedded in paraffin, sectioned, stained using a mouse anti-cyclin D1 primary antibody followed by a horse anti-mouse secondary antibody, and developed using 3,3-diaminobenzidine (DAB). The sections were then counterstained in Harris Hematoxylin Solution and then the sections were imaged at 20X magnification.
Reagents
Paraffin-Embedded Sections
<ol type="a">
	<li>4% Paraformaldehyde (PFA)</li>
	<li>Ethanol (anhydrous denatured, histological grade 100%, 95%, 80%, 75%, and 50%). Can be diluted from 100% stock using double distilled water (ddH2O)</li>
	<li>Xylene</li>
	<li>IHC compatible glass slide to ensure tissue section remains attached throughout the entire procedure. IHC compatible glass slides have a specialized coating and are readily available from multiple retailers. If performing ICC, use a chambered slide. Chambered slides allow for cells to be seeded in the chambers and placed in the incubator until cells attach to the slide and reach proper confluency, at which point the chambers can be removed and staining can proceed in the same manner as IHC.</li>
	<li>Paraffin</li>
	<li>0.3% Hydrogen peroxide (H2O2)/methanol: To prepare, add 1 mL 30% H2O2 to 99 mL methanol. Store at -20&#176;C</li>
	<li>Antigen retrieval buffer: IHC citrate buffer pH 6.0</li>
</ol>
Fresh Frozen Sections
<ol type="a">
	<li>Optimal cutting temperature (OCT) embedding compound</li>
	<li>Optimal fixative: 4% PFA or acetone that has been cooled to -20&#176;C</li>
</ol>
Staining
<ol type="a">
	<li>Blocking buffer: Should be determined by user. One example is horse serum diluted in 1X PBS</li>
	<li>Diluted primary antibody: see manufacturer specifications</li>
	<li>Diluted biotinylated secondary antibody: see manufacturer specifications</li>
	<li>Diluted avidin-Horseradish Peroxidase (HRP): Only for peroxidase visualization. See manufacturer specifications.</li>
	<li>DAB or another compatible substrate</li>
	<li>Counterstain (optional)</li>
	<li>Ethanol (anhydrous denatured, histological grade 100% and 95%)</li>
	<li>Xylene</li>
	<li>Organo/Limonene Mount</li>
</ol>

Immunohistochemistry &amp; Immunocytochemistry: Tissue Imaging

Immunohistochimie et immunocytochimie : Imagerie tissulaire par microscopie optique

Source: Michael S. Lee1 and Tonya J. Webb1
1 Department of Microbiology and Immunology, University of Maryland School of Medicine and the Marlene and ...

Antibody Generation: Producing Monoclonal Antibodies Using Hybridomas

Source: Frances V. Sjaastad1,2, Whitney Swanson2,3, and Thomas S. Griffith1,2,3,4 
1 Microbiology, Immunology, and Cancer Biology Graduate Program, University of Minnesota, Minneapolis, MN 55455 
2 Center for Immunology, University of Minnesota, Minneapolis, MN 55455 
3 Department of Urology, University of Minnesota, Minneapolis, MN 55455 
4 Masonic Cancer Center, University of Minnesota, Minneapolis, MN 55455
Polyclonal antibodies are defined as a collection of antibodies directed against different antigenic determinants of an antigen or several antigens (1). While polyclonal antibodies are powerful tools for identifying biological molecules, there is one important limitation - they are unable to distinguish between antigens that share antigenic determinants. For example, when bovine serum albumin is used to immunize an animal, B cells with different surface Ig will respond to different antigenic determinants on bovine serum albumin. The result is a mixture of antibodies in the antiserum. Because bovine serum albumin shares some epitopes with human serum albumin in evolutionarily conserved regions of the protein, this anti-bovine serum albumin antiserum will also react with human serum albumin. Therefore, this antiserum will not be useful for distinguishing between bovine and human serum albumins.
Several approaches have been taken to overcome the specificity issue of polyclonal antisera. One is by absorbing the unwanted antibodies by passing the antiserum through a chromatography column of immobilized antigens (2). This method is tedious and frequently unable to completely remove the unwanted antibodies. Another approach is to isolate individual antibody-producing B cells and expand them in culture. However, like most normal untransformed cells, B cells do not survive in long-term culture.
To overcome the inability of B cells to survive in culture, one approach is to prepare a myeloma-B cell hybridoma. In 1847, Henry Bence-Jones discovered that patients with multiple myeloma, a lymphoid tumor, produced a large quantity of antibodies (3). B cells in these patients have become malignant and grow uncontrollably. Since the malignant B cells are derived from a single clone, they are identical and produce only a single type of antibody (i.e., a monoclonal antibody, or mAb). However, most of these myeloma cells produce antibodies of unknown specificities. In 1975, by fusing a myeloma cell to a B cell, Cesar Milstein and Georges Kohler succeeded in producing a hybridoma that can be cultured indefinitely in vitro and produces an unlimited number of monoclonal antibodies of known antigenic specificity (4). The rationale behind their approach is to combine the immortal properties of the myeloma cell and the antibody producing properties of the B cell. Their technique revolutionized antibody production and provides a powerful means for identification and purification of biological molecules using monoclonal antibodies.
Generally, preparing a monoclonal antibody requires several months. The general procedure includes the following steps:
<ol>
	<li>Immunization and screening of antibody titer</li>
	<li>Fusion of antibody-producing B cells and myeloma cells</li>
	<li>Selective growth of the hybridoma</li>
	<li>Screening the hybridomas for producing the desired monoclonal antibody</li>
	<li>Cloning by limiting dilution - a process whereby cells are diluted to a concentration to statistically allow for less than 1 cell to be added to the wells of a 96-well plate. Some wells will end up with 0 cells and some will have 1 cell. The wells with seeded with 1 cell will eventually grow into a monoclonal population of cells.</li>
	<li>Growth of the hybridoma and preparation of monoclonal antibody</li>
</ol>
This protocol focuses on the last step - growth of the hybridoma and preparation of the monoclonal antibody. The antibody is purified from the culture supernatant by ammonium sulfate precipitation (often referred to as salting out) - a commonly used method of removing proteins from a solution. Proteins in solution form hydrogen bonds, along with other hydrophilic interactions, with water through their exposed polar and ionic groups. When concentrations of small, highly charged ions (such as ammonium or sulfate) are added, these groups compete with the proteins for binding to water. This removes the water molecules from the protein and decreases its solubility, resulting in precipitation of the protein.

Producing Monoclonal Antibodies Using Hybridomas

Génération d'anticorps monoclonaux à l'aide d'hybridomes

Source: Frances V. Sjaastad1,2, Whitney Swanson2,3, and Thomas S. Griffith1,2,3,4
1 Microbiology, Immunology, and Cancer Biology Graduate Program, ...

Immunofluorescence Microscopy: Immunofluorescence Staining of Paraffin-Embedded Tissue Sections

Source: Thomas Chaffee1, Thomas S. Griffith2,3,4, and Kathryn L. Schwertfeger1,3,4 
1 Department of Lab Medicine and Pathology, University of Minnesota, Minneapolis, MN 55455 
2 Department of Urology, University of Minnesota, Minneapolis, MN 55455 
3 Masonic Cancer Center, University of Minnesota, Minneapolis, MN 55455 
4 Center for Immunology, University of Minnesota, Minneapolis, MN 55455
Pathologic analyses of tissue sections can be used to obtain a better understanding of normal tissue structure and contribute to our understanding of mechanisms of disease. Tissue biopsies, either from patients or from experimental in vivo models, are often preserved by fixing in formalin or paraformaldehyde and embedding in paraffin wax. This allows for long-term storage and for the tissues to be sectioned. Tissues are cut into thin (5 &#181;m) sections using a microtome and the sections are adhered to glass slides. The tissues sections can be stained with antibodies, which allow for the detection of specific proteins within the tissue sections. Staining with antibodies conjugated to fluorophores (also known as fluorochromes) - compounds that emit light at specific wavelengths when excited by a laser - is known as immunofluorescence. The ability to detect proteins within a section can provide information such as cell type heterogeneity within the tissue, activation of specific signaling pathways, and expression of biomarkers. Depending on the fluorophores used and type of microscope available for analysis, multiple colors can be used, which allows for multiplexed analysis of targets.
The following protocol outlines the basic steps involved in immunofluorescence staining of paraffin embedded tissue sections. It is important to note that this protocol will not include any details on the fixation of tissue, process of paraffin embedding, or sectioning of the tissues. Once tissues have been sectioned and placed on glass slides, they are rehydrated through a series of graded ethanol (EtOH) incubations. The sections are incubated with a blocking reagent to reduce non-specific binding of antibody to the tissue section. The sections are then incubated with a primary antibody that may or may not be directly labeled with a fluorophore. If the primary antibody is not directly labeled, the sections are then incubated with a secondary antibody labeled with a fluorophore. Different antibodies may require different staining conditions, thus suggestions for optimization of antibodies are included. Following washing to remove all unbound antibody, the slides are mounted with media containing DAPI to fluorescently label the nucleus. Once the mounting media has dried, the slides can be imaged using a microscope with lasers that can detect the different fluorophores.

Immunofluorescence Staining and Microscopy

Microscopie à fluorescence : coloration par immunofluorescence des sections de tissus inclus en paraffine

Source: Thomas Chaffee1, Thomas S. Griffith2,3,4, and Kathryn L. Schwertfeger1,3,4
1 Department of Lab Medicine and Pathology, University of Minnesota, ...

Confocal Fluorescence Microscopy: A Technique to Determine the Localization of Proteins in Mouse Fibroblasts

Source: Dominique R. Bollino1, Eric A. Legenzov2, Tonya J. Webb1 
1 Department of Microbiology and Immunology, University of Maryland School of Medicine and the Marlene and Stewart Greenebaum Comprehensive Cancer Center, Baltimore, Maryland 21201 
2 Center for Biomedical Engineering and Technology, University of Maryland School of Medicine, Baltimore, Maryland 21201
Confocal fluorescence microscopy is an imaging technique that enables increased optical resolution as compared to conventional &#34;wide-field&#34; epifluorescence microscopy. Confocal microscopes are able to achieve improved x-y optical resolution through &#34;laser scanning&#34;- typically a set of voltage-controlled mirrors (galvanometer or &#34;galvo&#34; mirrors) that direct laser-illumination to each pixel of the specimen at a time. More importantly, confocal microscopes achieve superior z-axial resolution by using a pinhole to remove out of focus light originating from locations that are not in the z-plane being scanned, thus enabling the detector to collect data from a specified z-plane. Because of the high z-resolution achievable in confocal microscopy, it is possible to collect images from a series of z-planes (also-called z-stack) and construct a 3D image through software.
Before discussing the mechanism of a confocal microscope, it is important to consider how a sample interacts with light. Light is composed of photons, packets of electromagnetic energy. A photon impinging on a biological sample can interact with the molecules comprising the sample in one of four ways: 1) the photon does not interact and passes through the sample; 2) the photon is reflected/scattered; 3) the photon is absorbed by a molecule and the absorbed energy is released as heat through processes collectively known as nonradiative decay; and 4) the photon is absorbed and the energy is then rapidly reemitted as a secondary photon through the process known as fluorescence. A molecule whose structure permits fluorescence emission is called a fluorophore. Most biological samples contain negligible endogenous fluorophores; therefore exogenous fluorophores must be used to highlight features of interest in the sample. During fluorescence microscopy, the sample is illuminated with light of the appropriate wavelength for absorption by the fluorophore. Upon absorbing a photon, a fluorophore is said to be &#34;excited&#34; and the process of absorption is referred to as &#34;excitation&#34;. When a fluorophore gives up energy in the form of a photon, the process is known as &#34;emission&#34;, and the emitted photon is called fluorescence.
The light beam used to excite a fluorophore is focused by the objective lens of a microscope and converges at a &#34;focal point&#34; where it is maximally focused. Beyond the focal point the light again diverges. The entering and exiting beams may be visualized as a pair of cones touching at the focal point (see Figure 1, left panel). The phenomenon of diffraction imposes a limit on how tightly a beam of light can be focused - the beam actually focuses to a spot of finite size. Two factors determine the size of the focal spot: 1) the wavelength of the light, and 2) the light-gathering ability of the objective lens, which is characterized by its numerical aperture (NA). The focal &#34;spot&#34; extends not only in the x-y plane, but also in the z direction, and is in reality a focal volume. The dimensions of this focal volume define the maximum resolution achievable by optical imaging. Although the number of photons is greatest within the focal volume, the conical light paths above and below the focus also contain a lower density of photons. Any fluorophore in the light path can thus be excited. In conventional (wide-field) epifluorescence microscopy, emission from fluorophores above and below the focal plane contribute out-of-focus fluorescence (a &#34;hazy background&#34;), which reduces image resolution and contrast, as demonstrated in Figure 1, with the red cube representing fluorophore emission above the focal plane (red star) that results in out-of-focus fluorescence (top right). This problem is ameliorated in confocal microscopy, due to the utilization of a pinhole. (Figure 2, bottom right). As depicted in Figure 3, the pinhole allows emissions originating from the focal place to reach the detector (left), while blocking the out-of-focus fluorescence (right) from reaching the detector, thus improving both resolution and contrast.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10501/10501fig1.jpg" /> 
Figure 1. Optical resolution of epifluorescence versus confocal microscopy. <a href="https://www.jove.com/files/ftp_upload/10501/10501fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The light beam used to excite a fluorophore is focused by the objective lens of a microscope and converges at a focal volume and then diverges (left). The red star represents the focal plane of a sample that is being imaged while the red square represents fluorophore emission above the focal plane. When capturing an image of this sample using an epifluorescent microscope, the emission from the out-of-focus red square will visible and contribute to a &#34;hazy background&#34; (top right). Confocal microscopes have a pinhole which prevents the detection of light emitted outside of the focal plane, eliminating the &#34;hazy background&#34; (bottom right).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10501/10501fig2.jpg" /> 
Figure 2. Pinhole effect in confocal microscopy. <a href="https://www.jove.com/files/ftp_upload/10501/10501fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Although the highest intensity of the excitation light is at the focal point of the lens (left, red oval), other parts of the sample not in the focal point (right, red star) will get light and fluoresce. To prevent light emitted from these out-of-focus regions to reach the detector, a screen with a pinhole is present in front of the detector. Only the in-focus light (left) emitting from the focal plane is able to travel through the pinhole and reach the detector. The out-of-focus light (right) is blocked with the pinhole and fails to reach the detector.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10501/10501fig3.jpg" /> 
Figure 3. Principal components of a confocal laser scanning microscope. <a href="https://www.jove.com/files/ftp_upload/10501/10501fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
For the sake of simplicity, the mechanistic description of a confocal microscope will be limited to that of the Nikon Eclipse Ti A1R. Although there may be minor technical differences between different confocal microscopes, the A1R serves well as a good model for describing confocal microscope function. The excitation light beam, produced by an array of diode lasers, is reflected by the primary dichroic mirror into the objective, which focuses the light onto the specimen being imaged. The primary dichroic mirror selectively reflects the excitation light while allowing light at other wavelengths to pass through. The light then encounters the scanning mirrors that sweep the light beam across the specimen in an x-y manner, illuminating a single (x,y) pixel at a time. Fluorescence emitted by fluorophores at the illuminated pixel is collected by the objective lens and passes through the primary dichroic mirror to reach an array of photomultiplier tubes (PMTs). Secondary dichroic mirrors direct the emission light to the appropriate PMT. Excitation light scattered by the sample back into the objective is reflected by the primary dichroic mirror back towards the specimen, and thus prevented from entering the detection light path and reaching the PMTs (see Figure 3). This allows the relatively feeble fluorescence to be quantified without contamination by light scattered from the excitation light beam, which is typically orders of magnitude more intense than the fluorescence. Because the pinhole blocks light from outside of the focal volume, the light arriving at the detector comes from a narrow, selected z-plane. Therefore, images can be collected from a series of adjacent z-planes; this series of images is often referred to as a &#39;z-stack&#39;. By using the appropriate software, a z-stack can be processed to generate a 3D image of the specimen. A particular advantage of confocal microscopy is the ability to distinguish the subcellular localization of staining. For example, the differentiation between membrane staining from intracellular staining, which is very challenging with conventional epifluorescence microscopy (1, 2, 3).
Sample preparation is an important facet of confocal imaging. A strength of optical microscopy techniques is the flexibility to image live or fixed cells. When attempting to produce 3D images, because of the number of images that must be acquired for a z-stack, the difficulty of maintaining cell health, and the movement of live cells and their organelles, the use of fixed cells is typical. The procedure for fixing and staining cells for confocal fluorescence is similar to that conventionally used in immunofluorescence. After culture in chamber slides or on coverslips, cells are fixed using paraformaldehyde to preserve cellular morphology. Non-specific antibody binding is blocked using bovine serum albumin, milk, or normal serum. In order to maintain the specificity of the secondary antibodies, the solution used should not originate from the same species in which the primary antibodies were generated. The cells are incubated with primary antibodies that bind the antigen of interest. When labeling several cellular targets, the primary antibodies must each be derived from a different species. Antibodies tagging an antigen are then bound by fluorophore-conjugated secondary antibodies. Fluorophore-conjugated secondary antibodies should be selected so that they are compatible with the wavelengths of laser excitation available in the confocal microscope. When visualizing multiple antigens, the excitation/emission spectra of the fluorophores should differ enough so that their signals can be discriminated by microscopic analysis. The stained specimen is then mounted on a slide for imaging. A mounting medium is used to prevent photobleaching and specimen dehydration. If desired, a mounting medium containing a nuclear counterstain (e.g. DAPI or Hoechst) can be used (4).
In the following protocol, mouse fibroblasts transfected to express CD1d (LCD1) were stained with antibodies recognizing CD1d and CD107a (LAMP-1). CD1d is a major histocompatibility complex 1 (MHC 1)-like receptor present on the surface of antigen presenting cells that presents lipid antigens. LAMP-1 (lysosomal associated membrane protein-1) is a transmembrane protein primarily present in lysosomal membranes. For proper antigen presentation, CD1d is trafficked through the low pH lysosomal compartment, so LAMP-1 is being used as a marker of the lysosomal compartment for this protocol. By probing the LCD1 cells with anti-CD1d and anti- LAMP-1 that were produced in different species, secondary antibodies with unique fluorophores can be used to determine the localization of each protein in the cell and whether CD1d is present in the LAMP-1 positive lysosomal compartments.

Confocal Fluorescence Microscopy and Localization of Proteins

Microscopie confocale à fluorescence : une technique pour localiser les protéines dans les fibroblastes de souris

Source: Dominique R. Bollino1, Eric A. Legenzov2, Tonya J. Webb1
1 Department of Microbiology and Immunology, University of Maryland School of Medicine ...

Immunoprecipitation-Based Techniques: Purification of Endogenous Proteins Using Agarose Beads

Source: Susannah C. Shissler1, Tonya J. Webb1 
1 Department of Microbiology and Immunology, University of Maryland, Baltimore, MD 21201
Immunoprecipitation (IP, also known as a &#39;pull-down&#39; assay) is a widely used technique that has applications in a variety of fields. First conceived in 1984, it was refined in 1988 (1, 2). The fundamental goal of IP is purification and isolation of a specific protein using an antibody against that protein. The word &#34;immuno&#34; refers to the use of an antibody while the word &#34;precipitation&#34; refers to pulling down a specific substance from a solution. The target protein might be endogenous or recombinant. Most recombinant proteins have an epitope tag (i.e. myc or flag) attached to them to simplify subsequent purification. Typically, it is easier to optimize recombinant protein IP because the antibodies against recombinant epitope tags are very strong and effective. Antibodies against endogenous proteins have extremely variable efficacy - making it much more difficult to optimize these IPs. A necessary step after immunoprecipitation is verification of purification. The isolated protein is resolved using SDS-PAGE and subsequently probed for purity by western blots (Figure 1). An important control is the use of a different antibody during the Western blot to verify pull down of the correct protein. The combination of IP with subsequent techniques is a powerful analysis tool. The goal after purification may be characterization of the protein itself by NMR, mass spectrometry, and in vitro assays, or analysis of the protein&#39;s interacting partners (i.e. protein, DNA, RNA) (3, 4, 5).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10502/10502fig1.jpg" /> 
Figure 1: Overview of Immunoprecipitation Procedure. Immunoprecipitation is the isolation of a specific protein using an antibody. After production of lysate from cells, there are two major steps- pre-clearing and pull down. During pre-clearing step, the cell-lysates are pre-cleared of proteins that bind to antibodies non-specifically using an isotype control antibody. In pull down step, the target protein is pulled down using a protein-specific antibody. The isolated protein is then analyzed by Western blot. Isotype antibodies and protein specific antibodies have the same constant domain, but different antigen binding domains. A key component of this protocol is Protein A/G agarose beads that bind the constant domain of antibodies- allowing immunoprecipitation of the target protein. <a href="https://www.jove.com/files/ftp_upload/10502/10502fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Antibodies are the key component of an immunoprecipitation that differentiate it from other forms of protein purification (i.e. nickel affinity column purification). Antibodies are molecules made by B cells that can recognize specific protein epitopes. Antibodies have two domains: constant (Fc) and antigen binding (Fab) (Figure 1). The constant domain identifies the type of antibody and dictates function in vivo. Usually, the constant domains of antibodies used for IP are mouse, rat, or rabbit IgG. The antigen binding portion of the antibody recognizes a specific epitope of a specific protein. Antibodies can recognize epitopes on folded proteins that may not exist when the protein is denatured and vice versa. Therefore, the availability of the epitope depends on protein folding - identifying an important factor to consider when choosing antibodies and conditions for IP.
Both prokaryotic and eukaryotic systems have antibody-binding proteins. In eukaryotic systems, the purpose is immune protection from bacteria while in prokaryotic systems, the purpose is protection from the immune system. Antibody-binding proteins affect IP methodology in two ways. First, there is a necessary pre-clearing step (Figure 1) to rid the lysate of proteins that bind antibodies - thereby reducing non-specific binding in the final product. This step uses an isotype antibody that has the same constant domain as but a different antibody binding domain than your protein-specific antibody. Bacterial antibody-binding proteins are the second key component of this method. After the protein-specific antibody binds the target protein, the antibody: protein complex must be pulled down (Figure 1). Proteins A, G, and L are bacterial proteins that bind the constant domain of antibodies. While bacteria use this to subvert the immune system, researchers have co-opted this system for easy antibody purification, and it is used during both the pre-clearing and pull-down steps. These proteins have different binding affinities for different species and different constant domain subtypes - another factor to consider when choosing conditions for IP. Many companies sell Protein A/G labeled agarose beads (Figure 1), pre-made spin columns, or resins to make columns. In general, beads and spin columns are used for smaller sample sizes while resins are used for bulk purification.
In this lab exercise, we demonstrate how to purify the endogenous protein c-myc, from primary murine thymocytes, using Protein A/G Plus agarose beads based basic immunoprecipitation technique. The protocol starts from cell lysate preparation and ends with the with verification of successful protein pull down using Western blot analysis.

Immunoprecipitation Techniques: Purification of Proteins

Techniques basées sur l'immunoprécipitation : purification des protéines endogènes à l'aide de billes d'agarose

Source: Susannah C. Shissler1, Tonya J. Webb1
1 Department of Microbiology and Immunology, University of Maryland, Baltimore, MD 21201
Immunoprecipitation ...

Cell Cycle Analysis: Assessing CD4 and CD8 T Cell Proliferation After Stimulation Using CFSE Staining and Flow Cytometry

Source: Perchet Thibaut1,2,3, Meunier Sylvain1,2,3, Sophie Novault4, Rachel Golub1,2,3 
1 Unit for Lymphpoiesis, Department of Immunology, Pasteur Institute, Paris, France 
2 INSERM U1223, Paris, France 
3 Universit&#233; Paris Diderot, Sorbonne Paris Cit&#233;, Cellule Pasteur, Paris, France 
4 Flow Cytometry Platfrom, Cytometry and Biomarkers UtechS, Center for Translational Science, Pasteur Institute, Paris, France
The cell cycle is a universal process of life. During the cell cycle, a cell undergoes several modifications to divide into two daughter cells. This mechanism occurs throughout an organism&#39;s life in response to its needs. Cell divisions and embryonic development produce a full organism from a single-celled zygote. During adulthood, the cell cycle is central to many critical biological processes, such as tissue repairs.
Mechanisms of cell division are tightly controlled events where the cell undergoes stepwise modifications before final division. Cells that are not yet in the cycle are described as being in the Gap 0 (G0) phase. During this stage the cell is considered quiescent. When the cell starts to cycle, four distinct phases are recognized: Gap 1 (G1), Synthesis (S), Gap 2 (G2) and Mitosis (M). G1 phase is a checkpoint for resources needed by the cell for DNA synthesis. Then, S phase occurs, and DNA replication starts, followed by the G2 interphase, another checkpoint that controls all elements necessary to for the cell to divide. Finally, the cell enters mitosis and divides into two daughter cells.
Cell division is a highly informative parameter in many different biological systems. In the field of immunology, analysis of leukocyte proliferation can indicate the mechanism of the immune response. Other domains of investigation also rely on cell cycle analysis. For instance, analysis of the cell cycle during tumor development has improved our understanding of cancer.
Many fluorescent dyes are now available for tracking cell proliferation. These dyes differ in their chemical and spectral properties. Two different classes of dyes exist: protein dyes permanently combine with protein by forming a covalent bond, and membrane dyes stably intercalate within cell membranes via strong hydrophobic associations. In vitro and in vivo studies of immune cell proliferation by flow cytometry are among the most common applications of both classes of cell tracking dyes (1, 2).
CFSE (Carboxyfluorescein succinimidyl ester) is a fluorescent dye that marks dividing cells. Initially, all the cells receive the same amount of dye; dividing cells evenly split the dye they received between their two daughter cells. Consequently, the cell cycle can be followed by the progressive decrease of dye intensity in the cells. CFSE staining is followed by conventional multi-parametric flow cytometry, a high-throughput, fluorescence-based technology that allows phenotypic and functional characterization of cells based on their degree of CFSE staining (3).
In the following experiment, we assess the proliferation of CD4+ and CD8+ T cells in-vitro, following CD3 stimulation, using CFSE staining and flow cytometry.

Cell Cycle Analysis: Assessing CD4 and CD8 T Cell Proliferation

Analyse du cycle cellulaire : utilisation de la coloration CFSE et de la cytométrie de flux pour évaluer la prolifération des lymphocytes T CD4 et CD8 après stimulation

Source: Perchet Thibaut1,2,3, Meunier Sylvain1,2,3, Sophie Novault4, Rachel Golub1,2,3
1 Unit for Lymphpoiesis, Department of Immunology, Pasteur ...

Adoptive Cell Transfer: Introducing Donor Mouse Splenocytes to a Host Mouse and Assessing Success via FACS

Source: Meunier Sylvain1,2,3, Perchet Thibaut1,2,3, Sophie Novault4, Rachel Golub1,2,3 
1 Unit for Lymphopoiesis, Department of Immunology, Pasteur Institute, Paris, France 
2 INSERM U1223, Paris, France 
3 Universit&#233; Paris Diderot, Sorbonne Paris Cit&#233;, Cellule Pasteur, Paris, France 
4 Flow Cytometry Platfrom, Cytometry and Biomarkers UtechS, Center for Translational Science, Pasteur Institute, Paris, France
Adoptive cell transfer is a method for introducing cells into a patient or study organism in order to treat a disease or study a biological process, such as haematopoiesis. Aims of adoptive transfer are various; it can be used in fundamental biology as well as in medical sciences (1, 2). In mouse models, migration and distribution of transferred cells can be studied and followed by a tracking system (cell surface marker, staining by CFSE, etc.). In cancer studies on mouse models, transfer of specific cell populations can be used as experimental treatment against tumors. Another example for this technique is creation of chimeric mice by transfer of bone marrow cells to irradiated mice or mice with a severe immunodeficiency phenotype. This mouse model can be used to assess the impact of gene deletion on a specific cell population for instance. Transfer of bone borrow cells is also used in human medical treatment. When patients are irradiated in case of cancer therapy, adoptive transfer of bone marrow allows immune system reconstitution.
The first step in this technique is to obtain the cell population of interest. The technique chosen to isolate this population depends on the level of specificity of the targeted population. The largest level of selection is the whole organ, in which all cell populations present in the organ are taken. A more precise method is selection of a target cell population, often selected by one cell surface marker. The ideal method to sort cells in this case is by magnetic sorting. Finally, the most stringent level is the selection of cells by several cell surface markers to sort very specific cell populations. Flow cytometry sorting is the most popular method for this level of selection. Once population of interest is obtained, it can be transferred to the host. Before adoptive transfer it is essential to ensure compatibility between host and donor. Indeed, regardless of transfer goal, compatibility is crucial to assure cells adoption by the host without cells rejection.
In this lab exercise, we demonstrate the adoptive cell transfer technique by transferring splenocytes from a CD45.2 mouse into a CD45.1 Rag&#947;c mouse (lacking lymphocytes) and four days later confirm the splenocytes transfer using flow cytometry (see Figure 1).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10504/10504fig1.jpg" /> 
Figure 1: Schematic representation of adoptive transfer. (1) Splenocytes are isolated from CD45.2 mice and (2) transferred in CD45.1 Rag&#947;c mouse, control mouse is injected with PBS only. (3) 4 days after adoptive transfer, splenocytes are recovered from mice and (4) analyzed by flow cytometry. <a href="https://www.jove.com/files/ftp_upload/10504/10504fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Adoptive Cell Transfer Technique

Transfert adoptif de cellules : introduction de splénocytes d'une souris donneuse vers une souris hôte et évaluation du taux de succès au FACS

Assay for Cell Death: Chromium Release Assay of Cytotoxic Ability

Source: Frances V. Sjaastad1,2, Whitney Swanson2,3, and Thomas S. Griffith1,2,3,4 
1 Microbiology, Immunology, and Cancer Biology Graduate Program, University of Minnesota, Minneapolis, MN 55455 
2 Center for Immunology, University of Minnesota, Minneapolis, MN 55455 
3 Department of Urology, University of Minnesota, Minneapolis, MN 55455 
4 Masonic Cancer Center, University of Minnesota, Minneapolis, MN 55455
One of the main functions of the cells of the immune system is to remove target cells that have been infected with viruses or have undergone transformation into a tumor cell. In vitro assays for measuring the cytotoxic capacity of immune cells have been a staple in laboratories for many years. These assays have been used to determine the ability of T cells, NK cells, or any other immune cell to kill target cells in an antigen-specific or -nonspecific manner. Death ligands (e.g., Fas ligand or TRAIL), cytokines (e.g., IFNg or TNF), or cytotoxic granules (i.e., perforin/granzyme B) expressed by effector cells are some ways in which target cell death can be induced. With the explosion in tumor immunotherapy research in recent years, there is growing interest in finding agents to increase the cytotoxic activity of immune cells to improve patient outcomes. Conversely, some diseases are hallmarked by the overexuberant activity of immune cell cytotoxic activity, resulting in efforts to identify agents to temper these responses. Thus, having an assay in which the user can easily integrate any number of different effector cells, target cells, and/or response modifiers into the experimental design can serve as a valuable means of quickly assessing the cytotoxic capacity of effector cells and/or the responsiveness of the target cell.
These in vitro assays involve the mixing of different cell populations, as well as using a relatively low number of both effector and target cells. Thus, one necessity of the assay is to label the target cells in a manner that can easily be detected and quantitated, allowing the user to then determine the &#39;percent specific lysis&#39; mediated by the effector cells. Radioactivity - especially, chromium 51 (51Cr) in the form of Na251CrO4- is an inexpensive way to rapidly and nonspecifically label cellular proteins within the target cells (1). The short labeling and total assay times reduces the potential for significant changes in the number and/or phenotype of the target cells, which could influence the outcome of the assay. Upon the loss of membrane integrity of the target cells as a result of the cytotoxic activity of the effector cells, the 51Cr-labelled cellular proteins within the target cells are released into the culture supernatant, becoming available for quantitation. As with any assay examining the function of immune cells in vitro, there are a number of important considerations to consider improving the performance of the experiment. One of the most critical features is to use healthy effector (for maximal cytotoxic activity) and target (for maximal responsiveness and minimal spontaneous death/51Cr release) cells. Effector and target cell contact is required (leading to the common use of round-bottom 96-well plates to encourage cell-cell contact) (2). Finally, data analysis is dependent on the inclusion of positive and negative control target cell populations.
The following protocol will outline the steps for performing a standard 51Cr release assay to measure the cytotoxic ability of a population of effector cells, though a nonradioactive version using Europium has recently been developed. 51Cr is a powerful &#947;-radiation emitter. Consequently, the use of this assay requires proper radiation safety training, dedicated laboratory space, a gamma counter, and disposal of radioactive samples.
The general sequence of events in this assay are: 1) prepare 51Cr-labeled targets; 2) prepare effector cells and add to plate while target cells are labeling; 3) add labeled targets to plate; 4) incubate plate; 5) harvest supernatants; and 6) analyze data after running samples on counter. Samples are commonly prepared in triplicate, and then averaged to account for any subtle pipetting differences.
Proper PPE is important for this assay. Specifically, the user should wear a lab coat and gloves. Safety glasses may be required based on the laboratory or institution. There should be ample lead shielding for safe storage and use of the 51Cr during all steps. Finally, there should be dedicated lab space and equipment set aside for using 51Cr, including all the proper signage to indicate where samples with 51Cr are being kept and a Geiger counter equipped with gamma probe to survey the space for possible contamination.
In this lab exercise, we will determine the ability human peripheral blood mononuclear cells (PBMCs), (CpG stimulated vs. unstimulated) to kill melanoma cells, using human melanoma cell line WM793 as model and the chromium release assay.

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