An Introduction to Mass Spectrometry

1. What is mass spectrometry (MS)? What Information does mass spectrometry provide?
2. Where are mass spectrometers used?
3. How can mass spectrometry help biochemists?
4. How does a mass spectrometer work?
   1. Introduction
   2. Sample introduction
   3. Methods of sample ionisation
   4. Analysis and separation of sample ions
   5. Detection and recording of sample ions
5. Electrospray ionisation
   1. Electrospray ionisation
   2. Nanospray ionisation
   3. Data processing
6. Matrix assisted laser desorption ionisation
7. Positive or negative ionisation?
8. Tandem mass spectrometry (MS-MS): Structural and sequence information from mass spectrometry
   1. Tandem mass spectrometry
   2. Tandem mass spectrometry analyses
   3. Peptide sequencing by tandem mass spectrometry
   4. Oligonucleotide sequencing by tandem mass spectrometry
9. Background reading

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1. What is mass spectrometry (MS)? What information does mass spectrometry provide? Mass spectrometry is an analytical tool used for measuring the molecular mass of a sample.
For large samples such as biomolecules, molecular masses can be measured to within an accuracy of 0.01% of the total molecular mass of the sample i.e. within a 4 Daltons (Da) or atomic mass units (amu) error for a sample of 40,000 Da. This is sufficient to allow minor mass changes to be detected, e.g. the substitution of one amino acid for another, or a post-translational modification.
For small organic molecules the molecular mass can be measured to within an accuracy of 5 ppm or less, which is often sufficient to confirm the molecular formula of a compound, and is also a standard requirement for publication in a chemical journal.
Structural information can be generated using certain types of mass spectrometers, usually those with multiple analysers which are known as tandem mass spectrometers. This is achieved by fragmenting the sample inside the instrument and analysing the products generated. This procedure is useful for the structural elucidation of organic compounds and for peptide or oligonucleotide sequencing.
2. Where are mass spectrometers used?
Mass spectrometers are used in industry and academia for both routine and research purposes. The following list is just a brief summary of the major mass spectrometric applications:

• Biotechnology: the analysis of proteins, peptides, oligonucleotides
• Pharmaceutical: drug discovery, combinatorial chemistry, pharmacokinetics, drug metabolism
• Clinical: neonatal screening, haemoglobin analysis, drug testing
• Environmental: PAHs, PCBs, water quality, food contamination
• Geological: oil composition

3. How can mass spectrometry help biochemists?

• Accurate molecular weight measurements:
  sample confirmation, to determine the purity of a sample, to verify amino acid substitutions, to detect post-translational modifications, to calculate the number of disulphide bridges
• Reaction monitoring:
  to monitor enzyme reactions, chemical modification, protein digestion
• Amino acid sequencing:
  sequence confirmation, de novo characterisation of peptides, identification of proteins by database searching with a sequence "tag" from a proteolytic fragment
• Oligonucleotide sequencing:
  the characterisation or quality control of oligonucleotides
• Protein structure:
  protein folding monitored by H/D exchange, protein-ligand complex formation under physiological conditions, macromolecular structure determination

4. How does a mass spectrometer work?
4.1 Introduction
Mass spectrometers can be divided into three fundamental parts, namely the ionisation source , the analyser , and the detector.
The sample has to be introduced into the ionisation source of the instrument. Once inside the ionisation source, the sample molecules are ionised, because ions are easier to manipulate than neutral molecules. These ions are extracted into the analyser region of the mass spectrometer where they are separated according to their mass (m) -to-charge (z) ratios (m/z) . The separated ions are detected and this signal sent to a data system where the m/z ratios are stored together with their relative abundance for presentation in the format of a m/z spectrum .
The analyser and detector of the mass spectrometer, and often the ionisation source too, are maintained under high vacuum to give the ions a reasonable chance of travelling from one end of the instrument to the other without any hindrance from air molecules. The entire operation of the mass spectrometer, and often the sample introduction process also, is under complete data system control on modern mass spectrometers.

Simplified schematic of a mass spectrometer

4.2 Sample introduction
The method of sample introduction to the ionisation source often depends on the ionisation method being used, as well as the type and complexity of the sample.
The sample can be inserted directly into the ionisation source, or can undergo some type of chromatography en route to the ionisation source. This latter method of sample introduction usually involves the mass spectrometer being coupled directly to a high pressure liquid chromatography (HPLC), gas chromatography (GC) or capillary electrophoresis (CE) separation column, and hence the sample is separated into a series of components which then enter the mass spectrometer sequentially for individual analysis.
4.3 Methods of sample ionisation

Many ionisation methods are available and each has its own advantages and disadvantages ("Ionization Methods in Organic Mass Spectrometry", Alison E. Ashcroft, The Royal Society of Chemistry, UK, 1997; and references cited therein).
The ionisation method to be used should depend on the type of sample under investigation and the mass spectrometer available.
Ionisation methods include the following:
Atmospheric Pressure Chemical Ionisation (APCI)
Chemical Ionisation (CI)
Electron Impact (EI)
Electrospray Ionisation (ESI)
Fast Atom Bombardment (FAB)
Field Desorption / Field Ionisation (FD/FI)
Matrix Assisted Laser Desorption Ionisation (MALDI)
Thermospray Ionisation (TSP)
The ionisation methods used for the majority of biochemical analyses are Electrospray Ionisation (ESI) and Matrix Assisted Laser Desorption Ionisation (MALDI) , and these are described in more detail in Sections 5 and 6 respectively.
With most ionisation methods there is the possibility of creating both positively and negatively charged sample ions, depending on the proton affinity of the sample. Before embarking on an analysis, the user must decide whether to detect the positively or negatively charged ions (see section 7).
4.4 Analysis and Separation of Sample Ions
The main function of the mass analyser is to separate , or resolve , the ions formed in the ionisation source of the mass spectrometer according to their mass-to-charge (m/z) ratios. There are a number of mass analysers currently available, the better known of which include quadrupoles , time-of-flight (TOF) analysers, magnetic sectors , and both Fourier transform and quadrupole ion traps .
These mass analysers have different features, including the m/z range that can be covered, the mass accuracy, and the achievable resolution. The compatibility of different analysers with different ionisation methods varies. For example, all of the analysers listed above can be used in conjunction with electrospray ionisation, whereas MALDI is not usually coupled to a quadrupole analyser.
Tandem (MS-MS) mass spectrometers are instruments that have more than one analyser and so can be used for structural and sequencing studies. Two, three and four analysers have all been incorporated into commercially available tandem instruments, and the analysers do not necessarily have to be of the same type, in which case the instrument is a hybrid one. More popular tandem mass spectrometers include those of the quadrupole-quadrupole, magnetic sector-quadrupole , and more recently, the quadrupole-time-of-flight geometries.
4.5 Detection and recording of sample ions.
The detector monitors the ion current, amplifies it and the signal is then transmitted to the data system where it is recorded in the form of mass spectra . The m/z values of the ions are plotted against their intensities to show the number of components in the sample, the molecular mass of each component, and the relative abundance of the various components in the sample.
The type of detector is supplied to suit the type of analyser; the more common ones are the photomultiplier , the electron multiplier and the micro-channel plate detectors.

5. Electrospray ionisation

5.1 Electrospray ionisation
Electrospray Ionisation (ESI) is one of the Atmospheric Pressure Ionisation (API) techniques and is well-suited to the analysis of polar molecules ranging from less than 100 Da to more than 1,000,000 Da in molecular mass.

Standard electrospray ionisation source (Platform II)

During standard electrospray ionisation (J. Fenn, J. Phys. Chem., 1984, 88, 4451), the sample is dissolved in a polar, volatile solvent and pumped through a narrow, stainless steel capillary (75 - 150 micrometers i.d.) at a flow rate of between 1 �L/min and 1 mL/min. A high voltage of 3 or 4 kV is applied to the tip of the capillary, which is situated within the ionisation source of the mass spectrometer, and as a consequence of this strong electric field, the sample emerging from the tip is dispersed into an aerosol of highly charged droplets, a process that is aided by a co-axially introduced nebulising gas flowing around the outside of the capillary. This gas, usually nitrogen, helps to direct the spray emerging from the capillary tip towards the mass spectrometer. The charged droplets diminish in size by solvent evaporation, assisted by a warm flow of nitrogen known as the drying gas which passes across the front of the ionisation source. Eventually charged sample ions, free from solvent, are released from the droplets, some of which pass through a sampling cone or orifice into an intermediate vacuum region, and from there through a small aperture into the analyser of the mass spectrometer, which is held under high vacuum. The lens voltages are optimised individually for each sample.

The electrospray ionisation process

5.2 Nanospray ionisation
Nanospray ionisation (M. Wilm, M. Mann, Anal. Chem., 1996, 68, 1) is a low flow rate version of electrospray ionisation. A small volume (1-4 microL) of the sample dissolved in a suitable volatile solvent, at a concentration of ca. 1 - 10 pmol/microL, is transferred into a miniature sample vial. A reasonably high voltage (ca. 700 - 2000 V) is applied to the specially manufactured gold-plated vial resulting in sample ionisation and spraying. The flow rate of solute and solvent using this procedure is very low, 30 - 1000 nL/min, and so not only is far less sample consumed than with the standard electrospray ionisation technique, but also a small volume of sample lasts for several minutes, thus enabling multiple experiments to be performed. A common application of this technique is for a protein digest mixture to be analysed to generate a list of molecular masses for the components present, and then each component to be analysed further by tandem mass spectrometric (MS-MS) amino acid sequencing techniques (see Section 8).

ESI and nanospray ionisation are very sensitive analytical techniques but the sensitivity deteriorates with the presence of non-volatile buffers and other additives, which should be avoided as far as possible.

In positive ionisation mode, a trace of formic acid is often added to aid protonation of the sample molecules; in negative ionisation mode a trace of ammonia solution or a volatile amine is added to aid deprotonation of the sample molecules. Proteins and peptides are usually analysed under positive ionisation conditions and saccharides and oligonucleotides under negative ionisation conditions. In all cases, the m/z scale must be calibrated by analysing a standard sample of a similar type to the sample being analysed (e.g. a protein calibrant for a protein sample), and then applying a mass correction.
5.3 Data processing
ESI and nanospray ionisation generate the same type of spectral data for samples, and so the data processing procedures are identical.
In ESI, samples (M) with molecular masses up to ca. 1200 Da give rise to singly charged molecular-related ions, usually protonated molecular ions of the formula (M+H)⁺ in positive ionisation mode, and deprotonated molecular ions of the formula (M-H)^- in negative ionisation mode.
An example of this type of sample analysis is shown in the m/z spectrum of the pentapeptide leucine enkephalin, YGGFL. The molecular formula for this compound is C₂₈H₃₇N₅O₇ and the calculated monoisotopic molecular weight is 555.2692 Da.
The m/z spectrum shows dominant ions at m/z 556.1, which are consistent with the expected protonated molecular ions, (M+H⁺). Protonated molecular ions are expected because the sample was analysed under positive ionisation conditions. These m/z ions are singly charged, and so the m/z value is consistent with the molecular mass, as the value of z (number of charges) equals 1. Hence the measured molecular weight is deduced to be 555.1 Da, in good agreement with the theoretical value.

Positive ESI-MS m/z spectrum of leucine enkaphalin, YGGFL.

The m/z spectrum also shows other ions of lower intensity (ca. 25 % of the m/z 556.1 ions) at m/z 557.2. These represent the molecule in which one ¹²C atom has been replaced by a ¹³C atom, because carbon has a naturally occurring isotope one atomic mass unit (Da) higher. The intensity of these isotopic ions relates to the relative abundance of the naturally occurring isotope multiplied by the total number of carbon atoms in the molecule. Additionally the fact that the ¹³C ions are one Da higher on the m/z scale than the ¹²C ions is an indication that z = 1, and hence the sample ions are singly charged. If the sample ions had been doubly charged, then the m/z values would only differ by 0.5 Da as z, the number of charges, would then be equal to 2.
The m/z spectrum also contains ions at m/z 578.1, some 23 Da higher than the expected molecular mass. These can be identified as the sodium adduct ions, (M+Na)⁺, and are quite common in electrospray ionisation. Instead of the sample molecules being ionised by the addition of a proton H⁺, some molecules have been ionised by the addition of a sodium cation Na⁺. Other common adduct ions include K⁺ (+39) and NH₄⁺ (+18) in positive ionisation mode and Cl^- (+35) in negative ionisation mode.
Electrospray ionisation is known as a "soft" ionisation method as the sample is ionised by the addition or removal of a proton, with very little extra energy remaining to cause fragmentation of the sample ions.
Samples (M) with molecular weights greater than ca. 1200 Da give rise to multiply charged molecular-related ions such as (M+nH)ⁿ⁺ in positive ionisation mode and (M-nH)^n- in negative ionisation mode. Proteins have many suitable sites for protonation as all of the backbone amide nitrogen atoms could be protonated theoretically, as well as certain amino acid side chains such as lysine and arginine which contain primary amine functionalities.
An example of multiple charging, which is practically unique to electrospray ionisation, is presented in the positive ionisation m/z spectrum of the protein hen egg white lysozyme.

Positive ESI-MS m/z spectrum of the protien hen egg white lysozyme.

The sample was analysed in a solution of 1:1 (v/v) acetonitrile : 0.1% aqueous formic acid and the m/z spectrum shows a Gaussian-type distribution of multiply charged ions ranging from m/z 1101.5 to 2044.6. Each peak represents the intact protein molecule carrying a different number of charges (protons). The peak width is greater than that of the singly charged ions seen in the leucine enkephalin spectrum, as the isotopes associated with these multiply charged ions are not clearly resolved as they were in the case of the singly charged ions. The individual peaks in the multiply charged series become closer together at lower m/z values and, because the molecular weight is the same for all of the peaks, those with more charges appear at lower m/z values than do those with fewer charges (M. Mann, C. K. Meng, J. B. Fenn, Anal. Chem., 1989, 61, 1702).
The m/z values can be expressed as follows:
m/z = (MW + nH⁺)/n
where m/z = the mass-to-charge ratio marked on the abscissa of the spectrum;
MW = the molecular mass of the sample
n = the integer number of charges on the ions
H = the mass of a proton = 1.008 Da.
If the number of charges on an ion is known, then it is simply a matter of reading the m/z value from the spectrum and solving the above equation to determine the molecular weight of the sample. Usually the number of charges is not known, but can be calculated if the assumption is made that any two adjacent members in the series of multiply charged ions differ by one charge.
For example, if the ions appearing at m/z 1431.6 in the lysozyme spectrum have "n" charges, then the ions at m/z 1301.4 will have "n+1" charges, and the above equation can be written again for these two ions:

1431.6 = (MW + nH⁺)/n and 1301.4 = [MW + (n+1)H⁺] /(n+1)

These simultaneous equations can be rearranged to exclude the MW term:

n(1431.6) - nH⁺ = (n+1)1301.4 - (n+1)H⁺
and so:
n(1431.6) = n(1301.4) +1301.4 - H⁺
therefore:
n(1431.6 - 1301.4) = 1301.4 - H⁺
and so:
n = (1301.4 - H⁺) / (1431.6 - 1301.4)

hence the number of charges on the ions at m/z 1431.6 = 1300.4/130.2 = 10.
Putting the value of n back into the equation:

1431.6 = (MW + nH⁺) n
gives 1431.6 x 10 = MW + (10 x 1.008)
and so MW = 14,316 - 10.08
therefore MW = 14,305.9 Da

The observed molecular mass is in good agreement with the theoretical molecular mass of hen egg lysozyme (based on average atomic masses) of 14305.14 Da. The individual isotopes cannot be resolved when the ions have a large number of charges, and so for proteins the average mass is measured.
This may seem long-winded but fortunately the molecular mass of the sample can be calculated automatically, or at least semi-automatically, by the processing software associated with the mass spectrometer. This is of great help for multi-component mixture analysis where the m/z spectrum may well contain several overlapping series of multiply charged ions, with each component exhibiting completely different charge states.
Using electrospray or nanospray ionisation, a mass accuracy of within 0.01% of the molecular mass should be achievable, which in this case represents +/- 1.4 Da.
In order to clarify electrospray/nanospray data, molecular mass profiles can be generated from the m/z spectra of high molecular mass, multiply charged samples. To achieve this, all the components are transposed onto a true molecular mass (or zero charge state) profile from which molecular masses can be read directly without any amendments or calculations.
The m/z spectrum of lysozyme has been converted to a molecular mass profile using Maximum Entropy processing and the data are shown. The mass profile is dominated by a component of molecular mass 14,305.7 Da, with a series of minor peaks at higher mass, which is usually indicative of salt adducting e.g. Na (M+23), K (M+39), H₂SO₄ or H₃PO₄ (M+98). The molecular masses can be read easily and unambiguously, and a good idea of the purity of the protein is obtained on inspection of the molecular mass profile.

Molecular mass profile of lysozyme obtained by maximum entropy processing of the m/z spectrum

Proteins in their native state, or at least containing a significant amount of folding, tend to produce multiply charged ions covering a smaller range of charge states (say two or three). These charge states tend to have fewer charges than an unfolded protein would have, due to the inaccessibility of many of the protonation sites. In such cases, increasing the sampling cone voltage may provide sufficient energy for the protein to begin to unfold and create a wider charge state distribution centering on more highly charged ions in the lower m/z region of the spectrum.
The differences in m/z spectra due to the folded state of the protein are illustrated with the m/z spectra of the protein apo-pseudoazurin acquired under different solvent conditions.
Analysis of the protein in 1:1 acetonitrile : 0.1% aqueous formic acid at pH2 gave a Gaussian-type distribution with multiply charged states ranging from n = 9 at m/z 1487.8 to n = 19 at m/z 705.3, centering on n = 15 (lower trace). The molecular mass for this protein was 13,381 Da. Analysis of the protein in water gave fewer charge states, from n = 7 at m/z 1921.7 to n = 11 at m/z 1223.7, centering at n = 9 (upper trace). Not only has the charge state distribution changed, the molecular weight is now 13,444 Da which represents an increase of 63 Da and indicates that copper is remaining bound to the protein. Many types of protein complexes can be observed in this way, including protein-ligand, protein-peptide, protein-metal and protein-RNA macromolecules.

Positive ESI-MS m/z spectra of the protein apo-pseudoazurin analysed in water at pH7 (upper trace) and in 1:1 acetonitrile:0.1% aq. formic acid at pH2 (lower trace).

6. Matrix assisted laser desorption ionisation

Matrix Assisted Laser Desorption Ionisation (MALDI) (F. Hillenkamp, M. Karas, R. C. Beavis, B. T. Chait, Anal. Chem., 1991, 63, 1193) deals well with thermolabile, non-volatile organic compounds especially those of high molecular mass and is used successfully in biochemical areas for the analysis of proteins, peptides, glycoproteins, oligosaccharides, and oligonucleotides. It is relatively straightforward to use and reasonably tolerant to buffers and other additives. The mass accuracy depends on the type and performance of the analyser of the mass spectrometer, but most modern instruments should be capable of measuring masses to within 0.01% of the molecular mass of the sample, at least up to ca. 40,000 Da.

MALDI is based on the bombardment of sample molecules with a laser light to bring about sample ionisation. The sample is pre-mixed with a highly absorbing matrix compound for the most consistent and reliable results, and a low concentration of sample to matrix works best. The matrix transforms the laser energy into excitation energy for the sample, which leads to sputtering of analyte and matrix ions from the surface of the mixture. In this way energy transfer is efficient and also the analyte molecules are spared excessive direct energy that may otherwise cause decomposition. Most commercially available MALDI mass spectrometers now have a pulsed nitrogen laser of wavelength 337 nm.

Matrix assisted laser desorption ionisation (MALDI)

The sample to be analysed is dissolved in an appropriate volatile solvent, usually with a trace of trifluoroacetic acid if positive ionisation is being used, at a concentration of ca. 10 pmol/�L and an aliquot (1-2 �L) of this removed and mixed with an equal volume of a solution containing a vast excess of a matrix. A range of compounds is suitable for use as matrices: sinapinic acid is a common one for protein analysis while alpha-cyano-4-hydroxycinnamic acid is often used for peptide analysis. An aliquot (1-2 �L) of the final solution is applied to the sample target which is allowed to dry prior to insertion into the high vacuum of the mass spectrometer. The laser is fired, the energy arriving at the sample/matrix surface optimised, and data accumulated until a m/z spectrum of reasonable intensity has been amassed. The time-of-flight analyser separates ions according to their mass(m)-to-charge(z) (m/z) ratios by measuring the time it takes for ions to travel through a field free region known as the flight, or drift, tube. The heavier ions are slower than the lighter ones.

The m/z scale of the mass spectrometer is calibrated with a known sample that can either be analysed independently (external calibration) or pre-mixed with the sample and matrix (internal calibration).

Simplified schematic of MALDI-TOF mass spectrometry (linear mode)

MALDI is also a "soft" ionisation method and so results predominantly in the generation of singly charged molecular-related ions regardless of the molecular mass, hence the spectra are relatively easy to interpret. Fragmentation of the sample ions does not usually occur.

In positive ionisation mode the protonated molecular ions (M+H⁺) are usually the dominant species, although they can be accompanied by salt adducts, a trace of the doubly charged molecular ion at approximately half the m/z value, and/or a trace of a dimeric species at approximately twice the m/z value. Positive ionisation is used in general for protein and peptide analyses.

In negative ionisation mode the deprotonated molecular ions (M-H^-) are usually the most abundant species, accompanied by some salt adducts and possibly traces of dimeric or doubly charged materials. Negative ionisation can be used for the analysis of oligonucleotides and oligosaccharides.

Positive ionisation MALDI m/z spectrum of a peptide mixture using alpha-cyano-4-hydroxycinnamic acid as matrix

7. Positive or negative ionisation?

If the sample has functional groups that readily accept a proton (H⁺) then positive ion detection is used
e.g. amines R-NH₂ + H⁺ = R-NH₃⁺ as in proteins or peptides.

If the sample has functional groups that readily lose a proton then negative ion detection is used
e.g. carboxylic acids R-CO₂H = R-CO₂^- and alcohols R-OH = R-O- as in saccharides or oligonucleotides

8. Tandem mass spectrometry (MS-MS): Structural and sequence information from mass spectrometry.

8.1 Tandem mass spectrometry
Tandem mass spectrometry (MS-MS) is used to produce structural information about a compound by fragmenting specific sample ions inside the mass spectrometer and identifying the resulting fragment ions. This information can then be pieced together to generate structural information regarding the intact molecule. Tandem mass spectrometry also enables specific compounds to be detected in complex mixtures on account of their specific and characteristic fragmentation patterns.

A tandem mass spectrometer is a mass spectrometer that has more than one analyser, in practice usually two. The two analysers are separated by a collision cell into which an inert gas (e.g. argon, xenon) is admitted to collide with the selected sample ions and bring about their fragmentation. The analysers can be of the same or of different types, the most common combinations being:

• quadrupole - quadrupole
• magnetic sector - quadrupole
• magnetic sector - magnetic sector
• quadrupole - time-of-flight.

Fragmentation experiments can also be performed on certain single analyser mass spectrometers such as ion trap and time-of-flight instruments, the latter type using a post-source decay experiment to effect the fragmentation of sample ions.

8.2 Tandem mass spectrometry analyses.
The basic modes of data acquisition for tandem mass spectrometry experiments are as follows:

Product or daughter ion scanning:
the first analyser is used to select user-specified sample ions arising from a particular component; usually the molecular-related (i.e. (M+H)⁺ or (M-H)^-) ions. These chosen ions pass into the collision cell, are bombarded by the gas molecules which cause fragment ions to be formed, and these fragment ions are analysed i.e. separated according to their mass to charge ratios, by the second analyser. All the fragment ions arise directly from the precursor ions specified in the experiment, and thus produce a fingerprint pattern specific to the compound under investigation.

This type of experiment is particularly useful for providing structural information concerning small organic molecules and for generating peptide sequence information.

Precursor or parent ion scanning:
the first analyser allows the transmission of all sample ions, whilst the second analyser is set to monitor specific fragment ions, which are generated by bombardment of the sample ions with the collision gas in the collision cell. This type of experiment is particularly useful for monitoring groups of compounds contained within a mixture which fragment to produce common fragment ions, e.g. glycosylated peptides in a tryptic digest mixture, aliphatic hydrocarbons in an oil sample, or glucuronide conjugates in urine.

Constant neutral loss scanning:
this involves both analysers scanning, or collecting data, across the whole m/z range, but the two are off-set so that the second analyser allows only those ions which differ by a certain number of mass units (equivalent to a neutral fragment) from the ions transmitted through the first analyser. e.g. This type of experiment could be used to monitor all of the carboxylic acids in a mixture. Carboxylic acids tend to fragment by losing a (neutral) molecule of carbon dioxide, CO₂, which is equivalent to a loss of 44 Da or atomic mass units. All ions pass through the first analyser into the collision cell. The ions detected from the collision cell are those from which 44 Da have been lost.

Selected/multiple reaction monitoring:
both of the analysers are static in this case as user-selected specific ions are transmitted through the first analyser and user-selected specific fragments arising from these ions are measured by the second analyser. The compound under scrutiny must be known and have been well-characterised previously before this type of experiment is undertaken. This methodology is used to confirm unambiguously the presence of a compound in a matrix e.g. drug testing with blood or urine samples. It is not only a highly specific method but also has very high sensitivity.

8.3 Peptide Sequencing by Tandem Mass Spectrometry.

The most common usage of MS-MS in biochemical areas is the product or daughter ion scanning experiment which is particularly successful for peptide and nucleotide sequencing.

Peptide sequencing: H₂N-CH(R')-CO-NH-CH(R")-CO₂H

Peptides fragment in a reasonably well-documented manner (P. Roepstorrf, J. Fohlmann, Biomed. Mass Spectrom., 1984, 11, 601; R. S. Johnson, K. Biemann, Biomed. Environ. Mass Spectrom., 1989, 18, 945). The protonated molecules fragment along the peptide backbone and also show some side-chain fragmentation with certain instruments (Four-Sector Tandem Mass Spectrometry of Peptides, A. E. Ashcroft, P. J. Derrick in "Mass Spectrometry of Peptides" ed. D. M. Desiderio, CRC Press, Florida, 1990).

There are three different types of bonds that can fragment along the amino acid backbone: the NH-CH, CH-CO, and CO-NH bonds. Each bond breakage gives rise to two species, one neutral and the other one charged, and only the charged species is monitored by the mass spectrometer. The charge can stay on either of the two fragments depending on the chemistry and relative proton affinity of the two species. Hence there are six possible fragment ions for each amino acid residue and these are labelled as in the diagram, with the a, b, and c" ions having the charge retained on the N-terminal fragment, and the x, y", and z ions having the charge retained on the C-terminal fragment. The most common cleavage sites are at the CO-NH bonds which give rise to the b and/or the y" ions. The mass difference between two adjacent b ions, or y"; ions, is indicative of a particular amino acid residue (see Table of amino acid residues at the end of this document).

Peptide sequencing by tandem mass spectrometry - backbone cleavages

The extent of side-chain fragmentation detected depends on the type of analysers used in the mass spectrometer. A magnetic sector - magnetic sector instrument will give rise to high energy collisions resulting in many different types of side-chain cleavages. Quadrupole - quadrupole and quadrupole - time-of-flight mass spectrometers generate low energy fragmentations with fewer types of side-chain fragmentations.

Immonium ions (labelled "i") appear in the very low m/z range of the MS-MS spectrum. Each amino acid residue leads to a diagnostic immonium ion, with the exception of the two pairs leucine (L) and iso-leucine (I), and lysine (K) and glutamine (Q), which produce immonium ions with the same m/z ratio, i.e. m/z 86 for I and L, m/z 101 for K and Q. The immonium ions are useful for detecting and confirming many of the amino acid residues in a peptide, although no information regarding the position of these amino acid residues in the peptide sequence can be ascertained from the immonium ions.

An example of an MS/MS daughter or product ion spectrum is illustrated below. The molecular mass of the peptide was measured using standard mass spectrometric techniques and found to be 680.4 Da, the dominant ions in the MS spectrum being the protonated molecular ions (M+H⁺) at m/z 681.4. These ions were selected for transmission through the first analyser, then fragmented in the collision cell and their fragments analysed by the second analyser to produce the following MS/MS spectrum. The sequence (amino acid backbone) ions have been identified, and in this example the peptide fragmented predominantly at the CO-NH bonds and gave both b and y" ions. (Often either the b series or the y" series predominates, sometimes to the exclusion of the other). The b series ions have been labelled with blue vertical lines and the y" series ions have been labelled with red vertical lines. The mass difference between adjacent members of a series can be calculated e.g. b3-b2 = 391.21 - 262.16 = 129.05 Da which is equivalent to a glutamine (E) amino acid residue; and similarly y4 - y3 = 567.37 - 420.27 = 147.10 Da which is equivalent to a phenylalanine (F) residue. In this way, using either the b series or the y" series, the amino acid sequence of the peptide can be determined and was found to be NFESGK (n.b. the y" series reads from right to left!). The immonium ions at m/z 102 merely confirm the presence of the glutamine (E) residue in the peptide.

Peptide sequencing by tandem mass spectrometry - an MS-MS daughter or product ion spectrum.

A protein identification study would proceed as follows:

• a. The protein under investigation would be analysed by mass spectrometry to generate a molecular mass to within an accuracy of 0.01%.
• b. The protein would then be digested with a suitable enzyme. Trypsin is useful for mass spectrometric studies because each proteolytic fragment contains a basic arginine (R) or lysine (K) amino acid residue, and thus is eminently suitable for positive ionisation mass spectrometric analysis. The digest mixture is analysed - without prior separation or clean-up - by mass spectrometry to produce a rather complex spectrum from which the molecular weights of all of the proteolytic fragments can be read. This spectrum, with its molecular weight information, is called a peptide map. (If the protein already exists on a database, then the peptide map is often sufficient to confirm the protein.)
  For these experiments the mass spectrometer would be operated in the "MS" mode, whereby the sample is sprayed and ionised from the nanospray needle and the ions pass through the sampling cone, skimmer lenses, Rf hexapole focusing system, and the first (quadrupole) analyser. The quadrupole in this instance is not used as an analyser, merely as a lens to focus the ion beam into the second (time-of-flight) analyser which separates the ions according to their mass-to-charge ratio.
  
  Q-TOF mass spectrometer operating in MS (upper) and MS/MS mode (lower) modes.
• c. With the digest mixture still spraying into the mass spectrometer, the Q-Tof mass spectrometer is switched into "MS/MS" mode. The protonated molecular ions of each of the digest fragments can be independently selected and transmitted through the quadrupole analyser, which is now used as an analyser to transmit solely the ions of interest into the collision cell which lies inbetween the first and second analysers. An inert gas such as argon is introduced into the collision cell and the sample ions are bombarded by the collision gas molecules which cause them to fragment. The optimum collision cell conditions vary from peptide to peptide and must be optimised for each one. The fragment (or daughter or product) ions are then analysed by the second (time-of-flight) analyser. In this way an MS/MS spectrum is produced showing all the fragment ions that arise directly from the chosen parent or precursor ions for a given peptide component.
  
  An MS/MS daughter (or fragment, or product) ion spectrum is produced for each of the components identified in the proteolytic digest. Varying amounts of sequence information can be gleaned from each fragmentation spectrum, and the spectra need to be interpreted carefully. Some of the processing can be automated, but in general the processing and interpretation of spectra will take longer than the data acquisition if accurate and reliable data are to be generated.

The amount of sequence information generated will vary from one peptide to another, Some peptide sequences will be confirmed totally, other may produce a partial sequence of, say, 4 or 5 amino acid residues. Often sequence "tag" of 4 or 5 residues is sufficient to search a protein database and confirm the identity of the protein.

Peptide sequencing in summary:

Peptides fragment along the amino acid backbone to give sequence information.

Peptides ca. 2500 Da or less produce the most useful data.

The amount of sequence information varies from one peptide to another. Some peptides can generate sufficient information for a full sequence to be determined; others may generate a partial sequence of 4 or 5 amino acids.

A protein digest can be analysed as an entire reaction mix, without any separation of the products, from which individual peptides are selected and analysed by the mass spectrometer to generate sequence information.

About 4 �L of solution is required for the analysis of the digest mixture, with a concentration based on the original protein of ca. 1-10 pmol/�L. MS/MS sequencing is a sensitive technique consuming little sample.

Sometimes the full protein sequence can be verified; some proteins generate sufficient information to cover only part of the sequence. 70 - 80% coverage is reasonable.

Often a sequence "tag" of 4/5 amino acids from a single proteolytic peptide is sufficient to identify the protein from a database.

The final point in this summary means that mass spectrometers have been found to be extremely useful for proteomic studies, as illustrated below.

The proteomics procedure usually involves excising individual spots from a 2-D gel and independently enzymatically digesting the protein(s) contained within each spot, before analysing the digest mixture by mass spectrometer in the manner outlined above. Electrospray ionisation or MALDI could be used at this step.

The initial MS spectrum determining the molecular masses of all of the components in the digest mixture can often provide sufficient information to search a database using just several of the molecular weights from this peptide map.

If the database search is not fruitful, either because the protein has not been catalogued, is previously uncharacterised, or the data are not accurate or comprehensive enough to distinguish between several entries in the database, then further information is required.

This can be achieved by sample clean-up and then MS/MS studies to determine the amino acid sequences of the individual proteolytic peptides contained in the digest mixture, with which further database searching can be carried out.

8.4 Oligonucleotide sequencing by Tandem Mass Spectrometry.

Oligonucleotide sequencing: P-S(B)-P-S(B)-P-S(B)

Oligonucleotide sequencing can also be achieved by tandem mass spectrometry although it is not so well documented. However fragmentation patterns have been established and reported (S. Pomerantz, J. A. Kowalak, J. A. McClosky, J. Amer. Soc. Mass Spectrom., 1993, 4, 204). The experimental principle is similar to that of peptide sequencing, in that individual species are mass measured in MS mode of instrument operation, and then their molecular-related ions selected by the first (quadrupole) analyser to be transmitted into the collision cell where they undergo fragmentation after bombardment with a collision gas. The fragments are analysed by the second (time-of-flight) analyser to produce an MS/MS product, or daughter, ion spectrum showing all the fragment ions that arise directly from the chosen parent or precursor ions.

Negative electrospray ionisation is often the preferred ionisation method. The optimisation of the fragmentation conditions varies from component to component and diligence must be taken to ensure the best conditions are employed.

Data processing and interpretation is again of paramount importance for accurate, reliable results and hence sequence information.

9. General reading

"Mass Spectrometry: A Foundation Course", K. Downard, Royal Society of Chemistry, UK, 2004.

"An Introduction to Biological Mass Spectrometry", C. Dass, Wiley, USA, 2002.

"The Expanding Role of Mass Spectrometry in Biotechnology", G. Siuzdak, MCC Press, San Diego, 2004.

"Ionization Methods in Organic Mass Spectrometry", A.E. Ashcroft, Analytical Monograph, Royal Society of Chemistry, UK, 1997.

http://www.astbury.leeds.ac.uk (A.E. Ashcroft's MS web pages and tutorial)

Table of amino acid residues .

Symbol	Structure	Mass (Da)
Ala A	-NH.CH.(CH3).CO-	71.0
Arg R	-NH.CH.[(CH2)3.NH.C(NH).NH2].CO-	156.1
Asn N	-NH.CH.(CH2CONH2).CO-	114.0
Asp D	-NH.CH.(CH2COOH).CO-	115.0
Cys C	-NH.CH.(CH2SH).CO-	103.0
Gln Q	-NH.CH.(CH2CH2CONH2).CO-	128.1
Glu E	-NH.CH.(CH2CH2COOH).CO-	129.0
Gly G	-NH.CH2.CO-	57.0
His H	-NH.CH.(CH2C3H3N2).CO-	137.1
Ile I	-NH.CH.[CH.(CH3)CH2.CH3].CO-	113.1
Leu	-NH.CH.[CH2CH(CH3)2].CO-	113.1
Lys K	-NH.CH.[(CH2)4NH2].CO-	128.1
Met M	-NH.CH.[(CH2)2.SCH3].CO-	131.0
Phe F	-NH.CH.(CH2Ph).CO-	147.1
Pro P	-NH.(CH2)3.CH.CO-	97.1
Ser S	-NH.CH.(CH2OH).CO-	87.0
Thr T	-NH.CH.[CH(OH)CH3).CO-	101.0
Trp W	-NH.CH.[CH2.C8H6N].CO-	186.1
Tyr Y	-NH.CH.[(CH2).C6H4.OH].CO-	163.1
Val V	-NH.CH.[CH(CH3)2].CO-	99.1

Mass Spectra Interpretation: ALDEHYDES

Mass Spectra Interpretation: ALDEHYDES

No. of Questions= 8

ELECTRON IMPACT MASS SPECTROMETRY PRODUCED THE SPECTRA BELOW. MANY PEAKS ARE LABELLED TO AID INTERPRETATION: M/Z (RELATIVE ABUNDANCE). LOOK AT THE HINTS AND FOLLOW THE WEB REFERENCE LINK FOR HELP. YOU WILL NEED A PERIODIC TABLE, A TABLE OF COMMON ISOTOPES (WHICH YOU CAN FIND IN THE WEB REFERENCE), A CALCULATOR, AND SCRATCH PAPER TO WORK THIS QUIZ.

1.	How many oxygens are in the molecular ion peak in this spectrum?
		Web Reference
		Click on image to enlarge
	a)	0
	b)	1
	c)	2
2.	For this same spectrum, choose the compound that the spectrum represents.
		Web Reference
		Click on image to enlarge
	a)	butanal
	b)	propanal
	c)	2-propenal
	d)	methoxy-ethene
	e)	ethanediol
3.	Find the alkyl ion series in the spectra below. (Check the hint!)
		Web Reference
		Click on image to enlarge
	a)	15, 29, 43
	b)	29, 43, 57
	c)	none
4.	Find the alkyl LOSS ion series in the same spectra shown below. (Check the hint!)
		Web Reference
		Click on image to enlarge
	a)	15, 29, 43
	b)	29, 43, 57
	c)	none
5.	For this same spectrum, choose the compound that the spectrum represents.
		Web Reference
		Click on image to enlarge
	a)	2-propenal
	b)	2-methyl propanal
	c)	pentanal
	d)	2-oxo-propanal
	e)	butanal
6.	Choose the compound that this spectrum represents.
		Web Reference
		Click on image to enlarge
	a)	hexanal
	b)	3-methyl butanal
	c)	pentanal
	d)	2-methyl propanal
	e)	2,2-dimethyl propanal
7.	Choose the compound that this spectrum represents.
		Web Reference
		Click on image to enlarge
	a)	2-methyl propanal
	b)	butanal
	c)	2-oxo-propanal
	d)	2-propenal
	e)	pentanal
8.	Choose the compound that this spectrum represents.
		Web Reference
		Click on image to enlarge
	a)	2,2-dimethyl propanal
	b)	hexanal
	c)	2-oxo-propanal
	d)	3-methyl-butanal
	e)	pentanal

The Basics Nuclear Magnetic Resonance Spectroscopy

Nuclei possessing angular moment (also called spin) have an associated magnetic moment. A few examples of magnetic isotopes are ¹³C, ¹H, ¹⁹F,¹⁴N, ¹⁷O, ³¹P, and ³³S. Please note that not every isotope is magnetic. In particular, you should note that ¹²C is not magnetic. If a nucleus is not magnetic, it can't be studied by nuclear magnetic resonance spectroscopy. For the purposes of this course, we will be most interested in ¹H and ¹³C. I will limit my discussions to ¹H in this short treatment. Generally speaking, you should think of these special nuclei as tiny, atomic, bar magnets.
Nuclear Magnetic Spectroscopy is based on the fact that when a population of magnetic nuclei is placed in an external magnetic field, the nuclei become aligned in a predictable and finite number of orientations. For ¹H there are two orientations. In one orientation the protons are aligned with the external magnetic field (north pole of the nucleus aligned with the south pole of the magnet and south pole of the nucleus with the north pole of the magnet) and in the other where the nuclei are aligned against the field (north with north, south with south). The alignment with the field is also called the "alpha" orientation and the alignment against the field is called the "beta" orientation. From my description of the poles, which orientation do you think is the preferred or lower in energy? If you guessed the "alpha", you are correct. It might be worth noting at this point that before the nuclei are placed in the magnetic field they have random orientation

random orientation outside of field	alpha and beta orientation in field

Since the alpha orientation is preferred, more of the population of nuclei are aligned with the field than against the field. You might wonder why any spins would align against the field. Realize that we are talking about atomic magnets. These are very, very weak magnets. The energy difference between the alpha and beta orientations is not large. There is enough energy for nuclei to exchange between the two orientations at room temperature, though a slight excess on average is in the lower energy, alpha state.
The nuclear magnetic resonance (NMR) spectroscopy experiment involves using energy in the form of electromagnetic radiation to pump the excess alpha oriented nuclei into the beta state. When the energy is removed, the energized nuclei relax back to the alpha state. The fluctuation of the magnetic field associated with this relaxation process is called resonance and this resonance can be detected and converted into the peaks we see in an NMR spectrum.
What sort of electromagnetic radiation is appropriate for the low energy transition involved in NMR? Well believe it or not, radio waves do the trick. Radio waves are at the very low energy end of the electromagnetic spectrum and are sufficient to induce the desired transition. It is for this reason that NMR is considered to be a safe method of analysis. The same technology is now used in hospitals in MRI (Magnetic Resonance Imagining - people are afraid of the word nuclear). If you have ever had an MRI done, realize that you were placed in a magnetic field and all your magnetic nuclei lined up in the manner described above. Excess nuclei were pumped to higher energy states as you were exposed to radio waves.
The following are two very, very important points to accept and learn if you are going to understand the rest of the discussion.
1.    Electric currents have associated magnetic fields.
2.    Magnetic fields can generate electric currents.
If you haven't had physics yet, try to accept these two points. Certainly most people have at least heard of electromagnets and if so, you probably have some idea about the first statement.
The following is a very important NMR relationship. This expression relates the external field to the frequency of resonance.

n=	mHo
	2p

In this equation, n is frequency, m is the magnetogyric ratio (not needed for this discussion - a constant for each nucleus). The big thing to glean from this equation is that the external field and the frequency are directly proportional. If the external field is larger , the frequency needed to induce the alpha to beta transition is larger. It follows then that in a larger field, higher frequency radio waves would be needed to induce the transition.
In this context, it is relevant to note that different nuclear magnetic resonance spectrometers have different magnetic field strengths. For example, the NMR on the first floor of Park Hall has a relatively high field, superconducting magnet. Because the field is high (high enough to erase bank cards and interfere with pacemakers and watches), the frequency range needed to excite protons is relatively high. It is called a 300 MHz (MHz = megahertz, a hertz is a cycle per second - a frequency unit) spectrometer, referring to the excitation frequency. The NMR on the second floor of Park Hall has a much weaker electromagnet associated with it. It is a 60 MHz instrument. Since different NMRs have different operating frequencies, spectra cannot be compared from different machines if they are reported in frequency units. For this reason, the universal ppm (parts per million) units are used in NMR. Please note the following relationship between ppm and frequency. The fact that frequency and ppm are directly proportional is all you need to retain for the future discussion and the course in general.

Chemical shift in ppm =	peak position in Hz (relative to TMS)
	spectrometer frequency in MHz

Now let us use these basic ideas to better understand and interpret NMR spectra.
1. Why do we see peaks? When the excited nuclei in the beta orientation start to relax back down to the alpha orientation, a fluctuating magnetic field is created. This fluctuating field generates a current in a receiver coil that is around the sample. The current is electronically converted into a peak. It is the relaxation that actually gives the peak not the excitation.
2. Why do we see peaks at different positions? Realize that in principle, a peak will be observed for every magnetically distinct nucleus in a molecule. This happens because nuclei that are not in identical structural situations do not experience the external magnetic field to the same extent. The nuclei are shielded or deshielded due to small local fields generated by circulating sigma, pi and lone pair electrons.
To understand this concept better, consider a "run of the mill" hydrogen like that in ethane or methane. When this sort of hydrogen is placed in a magnetic field, the sigma electrons start to circulate. Remember : Magnetic fields generate currents. When the electrons circulate, they generate a small magnetic field that happens to point in the opposite direction to the external field. Remember: Currents have associated magnetic fields. Since magnetism is a vector quantity (vector quantities have direction and magnitude), this local field reduces the overall field somewhat. Therefore, the described hydrogen experiences a reduced magnetic field. If we reconsider the important NMR equation given on page two of this document, we can only conclude that if the external field is lower then the frequency of the electromagnetic radiation needed to induce the alpha to beta transition must be lower. Remember that frequency and ppm are directly proportional. Therefore, if a hydrogen requires a lower frequency, then it will show up as a peak at a lower ppm value. Hydrogens like those in methane are at around 1.0 ppm in the NMR spectrum.

methane	bromoethane

Now consider a hydrogen near a halogen as in bromoethane. This type of hydrogen is in a magnetically altered situation as compared to the hydrogen in methane. Due to its inherent electronegativity, the halogen atom has the effect of pulling sigma electron density away from the hydrogens in the molecule. The effect is largest for the hydrogens closest to the halogen atom. Though the little local opposing sigma field is still generated next to the hydrogens, it is partially pulled away by the electronegative bromine . Therefore, the hydrogens experience less of the local field and more of the external field. In other words, the vector in the vicinity of the hydrogen has been reduced as compared to methane. After you do the vector addition you end up with a larger overall field (again, as compared to methane). So going back to the fact that field and frequency are directly proportional, hydrogens near an electronegative atom should require a higher frequency to flip from the alpha to beta orientation. Therefore, they should appear at a higher ppm in the spectrum. Hydrogens like those in bromoethane should appear from ca. 2.5-4.0 ppm in the NMR spectrum.

1H NMR Spectrum of Bromoethane

Now as a last example, let us consider the NMR spectrum of benzene. Benzene and aromatics in general are very interesting because their hydrogens appear around 7 ppm even though they have no electronegative atoms. Why is this so? It has to do with the pi electrons. Because benzene and its relatives are aromatic, the p orbitals at each carbon in the ring overlap forming one continuous pi system. When the benzene ring is placed in a magnetic field, the external field induces a current in the pi system and that current generates a secondary magnetic field (or induced magnetic field). Once again, remember that electric currents have associated magnetic fields and that magnetic fields generate currents. The secondary magnetic field is such that it adds to the external field in the vicinity of the aromatic hydrogens as diagrammed below.

Benzene

If the local field is in the same direction as the external field, the resulting field is larger than the external field. This means that the frequency needed to flip those hydrogens experiencing that field is larger. Larger frequency translates into higher ppm position.
It is really interesting to consider 18-annulene diagrammed below. 18-Annulene is a large enough ring to have both cis and trans double bonds. This means that some of the hydrogens are pointing in toward the center of the aromatic ring. Reconsider the diagram of benzene above. If you look at it carefully, you will see that the magnetic field opposes the external field on the inside of the ring!!! If 18-annulene is aromatic like benzene, the inner hydrogens should absorb at lower frequency (ppm) and guess what? They do - they appear at -1.7 ppm!! Isn't that neat!!

[18]-Annulene

So summing up, the different hydrogens of a molecule appear at different positions because small local magnetic fields are generated when local electrons begin to circulate due to the effect of the external magnetic field. These small fields either add to or subtract from the external field altering the frequency needed for excitation. Some of the effects are due to the circulation of sigma electrons while others are due to the circulation of pi electrons. The pi effects can be the most dramatic as was demonstrated in the preceding examples.
3. What causes splitting?
Many peaks in NMR spectra appear as symmetric patterns called doublets, triplets, quartets, quintets, etc. When you see these patterns it tells you about the number of adjacent (usually on the carbon next door to that bearing the absorbing hydrogen(s)), but different hydrogens. In simple spectra such as those we will be studying in organic chemistry lab, the number of peaks you see is one more than the number of adjacent, but different hydrogens. This is the so called n+1 rule. Different means that the adjacent hydrogens have a unique magnetic environment and absorb at a distinct frequency compared with the hydrogens in question. For example, consider bromoethane (structure given below).

Bromoethane has two different types of hydrogens so we expect two absorptions in the NMR spectrum. One absorption corresponds to the two hydrogens that are closest to the halogen atom. The other to the hydrogens comprising the methyl group that is farther away. Based on what I described above with regard to chemical shift (the ppm value), the hydrogens nearer the bromine should be at a higher ppm position. The hydrogens further from the bromine should be at lower ppm position.
Anyway, getting back the splitting, the hydrogens closer to the bromine will appear as a quartet because they are near three different hydrogens (the hydrogens on the methyl group). Those adjacent hydrogens are communicating their presence to the hydrogens being flipped. They are saying, "We are your neighbors and there are three of us." The reason they are able to communicate their presence is that they are little magnets and as such, they either add to or subtract from the external magnetic field depending on their orientation. Since there are many protons in a sample, the following are the possibilities for the neighboring hydrogens during excitation:

Please note that in the above diagram the "a" hydrogens are the ones near the bromine being flipped from the alpha to beta orientation. The "b" hydrogens are the three neighbors. As shown above, it is possible that a given "a" hydrogen will have three "b" hydrogens nearby that are aligned with the applied field during excitation. It is also possible that the three neighbors could all be aligned against the applied field. More probable is that either two protons will be aligned against the field or two with the field. These situations are more probable because there are more combinations of the three nuclei that give rise to these two possibilities. Since there are three combinations of each of these two, they are each three times more probable than having all three adjacent nuclei aligned with or against the field.
Now let us think about what these neighboring, local magnets do to the overall field. The "a" hydrogens that have all three neighbors aligned against the field have a lower overall magnetic field. Going back to the fundamental nuclear magnetic resonance equation (see page 2 ), you would conclude that these "a" hydrogens would have a lower frequency requirement for the alpha to beta transition and therefore appear at lower ppm. For the "a" hydrogens having three neighbors with all three "b" hydrogens aligned with the external field, the cumulative local field adds to the external field. This resultant field is larger than the external field so higher frequency electromagnetic radiation is needed to induce the alpha to beta transition. For the "b" hydrogens near two nuclei aligned with the field and one nucleus aligned against the field there is a slight increase in overall field leading to slightly higher frequency requirements. Similarly, two spins aligned against and one aligned with the field leads to slightly lower frequency requirements. So in the end, the "a" population is divided into four groups appearing at slightly different frequencies. The intermediate frequency peaks are taller than the higher and lower frequency peaks because they reflect more probable situations for local hydrogens. Hence, a quartet is observed.
Now if you understand why the "a" hydrogens give a quartet can you figure out why the "b" hydrogens give a triplet? Try to work it out using vectors as done in the above diagram.
For simple systems like bromoethane, n + 1 peaks will be observed for a given absorption, where n = the number of neighboring, but different hydrogens. This formula can be very useful when interpreting simple spectra.

The Interpretation of Simple NMR Spectra

This year, we will abstract the following information from NMR spectra to determine structures of products from organic reactions and isolations.
1. The number of peaks. The number of peaks is directly related to symmetry. If a compound has three significantly different types of hydrogens, it should have three different NMR absorptions.
2. The area under each absorption (the integral). The relative areas (or integrals) of the various absorptions in an NMR spectrum equals the relative number of hydrogens absorbing. If we know the molecular formula of a compound, we can use this ratio to figure out the actual number of each type of hydrogen. From the numbers of each type, we can infer the carbon structure. For example, with bromoethane, the relative areas under the NMR peaks are 2:3. This tells us that there is a group of two hydrogens that are the same and another group of three hydrogens that are the same.With your current knowledge of organic chemistry, it seems most likely that the compound has a methyl (-CH₃) and a methylene (-CH₂-) group. In other words, the most probable way to have three identical hydrogens is on a methyl group. The most probable way to have two identical hydrogens is in a methylene group.
Suppose you have a compound with the formula C₅H₁₂O and you are told that there are two NMR peaks, having the relative areas of 1:3. Can you come up with the structure of the compound?
3. The splitting pattern. For this semester, we will be using the n+1 rule as it applies to the simple structures we will be determining. You will see one more peak than the number of adjacent, but different hydrogens. Therefore, you can look at any peak and automatically know how many neighbors there are. This is crucial information because it allows you to start to hook atoms together in your structure. The problem is that people often confuse integral with splitting. So you must always remember this saying "Integral tells you what is here and splitting tells you what is near" This means that the integral tells you about the absorbing hydrogens and the splitting tells you about the neighbors. So what does it mean if you see a quintet with an area of two in a spectrum?
4. The position of the peak or the chemical shift ( ). This tells you about the electronic environment (the electronic environment directly relates to the magnetic environment) of the absorbing hydrogens. It will tell you if there are pi bonds or electronegative atoms nearby, etc. There are nice tables available that organize how different groups effect the frequency of absorptions and in lab you will always have these tables available to you. Yes, you will even have them on exams. A good rule of thumb when you are solving spectra is that the closer a hydrogen is to an electronegative atom the higher the ppm position. This little rule only works if the hydrogen is two or more bonds away from the atom. You will soon see the utility of this when you begin your problems in the workshop. It is also useful to keep in your head that aromatic hydrogens absorb at around 7 ppm.
A few tricks of the trade that are generally useful for spectral problem solving......
1. Always calculate the index of hydrogen deficiency or unsaturation number at the beginning of a problem ( you will normally be given the formula of the compound). Determining the unsaturation number is very helpful in regard to knowing which structural elements need to be present in your final solution. The unsaturation number is where you compare the actual formula with the theoretical saturated formula and compute the number of pairs of hydrogens that are missing. This topic should have been covered in class by now.
2. It is a good idea to interpret your IR spectrum before you do the NMR spectrum so that you have an idea about which functional groups are present in your molecule.
3. Organize your ideas about the structure of the unknown as you go along. For some people it is helpful to set up the following table for the NMR data and conclusions. The important part of the table is the conclusion column in which you are drawing a structural conclusion about the absorbing hydrogens and their neighbors. You should write a structural fragment down as has been done below for bromoethane.

ppm	integral	splitting	conclusion
1.6	3	triplet	CH3CH2-
3.4	2	quartet	-CH2CH3 near electronegative atom

4. You will notice as we do problems in class that we tend to emphasize and draw the most information from the integral and splitting. Chemical shift (ppm position) in many cases is the last point of interest. There are a few relevant chemical shifts that should be interpreted at the outset of a problem.. One is the aromatic chemical shift. Aromatic hydrogens absorb at ca. 7 ppm. This is a very distinct and characteristic shift and should be interpreted immediately. If you observe a peak at seven chances are you have an aromatic ring. The most common aromatic ring is benzene. Another very distinctive shift is that of the aldehyde functional group. Aldehydic hydrogens appear at ca. 9 ppm in the spectrum. if you see a shift of nine ppm assume that you have an aldehyde functional group.
5. Solving spectra rapidly involves making good educated guesses. If you get an integral of three there is really only one probable way to have three identical hydrogens - a methyl group. If you get an integral of nine it is most likely three methyl groups that are the same by symmetry. If you get aromatic absorptions, you probably have one or more benzene rings. Always start with the simplest ideas and work your way toward more exotic solutions.

Spectroscopy Types

Absorption

Absorption spectroscopy is a technique in which the power of a beam of light measured before and after interaction with a sample is compared. Specific absorption techniques tend to be referred to by the wavelength of radiation measured such as ultraviolet, infrared or microwave absorption spectroscopy. Absorption occurs when the energy of the photons matches the energy difference between two states of the material.

Fluorescence

Fluorescence spectroscopy uses higher energy photons to excite a sample, which will then emit lower energy photons. This technique has become popular for its biochemical and medical applications, and can be used for confocal microscopy, fluorescence resonance energy transfer, and fluorescence lifetime imaging.

X-ray

When X-rays of sufficient frequency (energy) interact with a substance, inner shell electrons in the atom are excited to outer empty orbitals, or they may be removed completely, ionizing the atom. The inner shell "hole" will then be filled by electrons from outer orbitals. The energy available in this de-excitation process is emitted as radiation (fluorescence) or will remove other less-bound electrons from the atom (Auger effect). The absorption or emission frequencies (energies) are characteristic of the specific atom. In addition, for a specific atom, small frequency (energy) variations that are characteristic of the chemical bonding occur. With a suitable apparatus, these characteristic X-ray frequencies or Auger electron energies can be measured. X-ray absorption and emission spectroscopy is used in chemistry and material sciences to determine elemental composition and chemical bonding.
X-ray crystallography is a scattering process; crystalline materials scatter X-rays at well-defined angles. If the wavelength of the incident X-rays is known, this allows calculation of the distances between planes of atoms within the crystal. The intensities of the scattered X-rays give information about the atomic positions and allow the arrangement of the atoms within the crystal structure to be calculated. However, the X-ray light is then not dispersed according to its wavelength, which is set at a given value, and X-ray diffraction is thus not a spectroscopy.

Flame

Liquid solution samples are aspirated into a burner or nebulizer/burner combination, desolvated, atomized, and sometimes excited to a higher energy electronic state. The use of a flame during analysis requires fuel and oxidant, typically in the form of gases. Common fuel gases used are acetylene (ethyne) or hydrogen. Common oxidant gases used are oxygen, air, or nitrous oxide. These methods are often capable of analyzing metallic element analytes in the part per million, billion, or possibly lower concentration ranges. Light detectors are needed to detect light with the analysis information coming from the flame.

• Atomic Emission Spectroscopy - This method uses flame excitation; atoms are excited from the heat of the flame to emit light. This method commonly uses a total consumption burner with a round burning outlet. A higher temperature flame than atomic absorption spectroscopy (AA) is typically used to produce excitation of analyte atoms. Since analyte atoms are excited by the heat of the flame, no special elemental lamps to shine into the flame are needed. A high resolution polychromator can be used to produce an emission intensity vs. wavelength spectrum over a range of wavelengths showing multiple element excitation lines, meaning multiple elements can be detected in one run. Alternatively, a monochromator can be set at one wavelength to concentrate on analysis of a single element at a certain emission line. Plasma emission spectroscopy is a more modern version of this method. See Flame emission spectroscopy for more details.
• Atomic absorption spectroscopy (often called AA) - This method commonly uses a pre-burner nebulizer (or nebulizing chamber) to create a sample mist and a slot-shaped burner that gives a longer pathlength flame. The temperature of the flame is low enough that the flame itself does not excite sample atoms from their ground state. The nebulizer and flame are used to desolvate and atomize the sample, but the excitation of the analyte atoms is done by the use of lamps shining through the flame at various wavelengths for each type of analyte. In AA, the amount of light absorbed after going through the flame determines the amount of analyte in the sample. A graphite furnace for heating the sample to desolvate and atomize is commonly used for greater sensitivity. The graphite furnace method can also analyze some solid or slurry samples. Because of its good sensitivity and selectivity, it is still a commonly used method of analysis for certain trace elements in aqueous (and other liquid) samples.
• Atomic Fluorescence Spectroscopy - This method commonly uses a burner with a round burning outlet. The flame is used to solvate and atomize the sample, but a lamp shines light at a specific wavelength into the flame to excite the analyte atoms in the flame. The atoms of certain elements can then fluoresce emitting light in a different direction. The intensity of this fluorescing light is used for quantifying the amount of analyte element in the sample. A graphite furnace can also be used for atomic fluorescence spectroscopy. This method is not as commonly used as atomic absorption or plasma emission spectroscopy.

Plasma Emission Spectroscopy
In some ways similar to flame atomic emission spectroscopy, it has largely replaced it.

• Direct-current plasma (DCP)

A direct-current plasma (DCP) is created by an electrical discharge between two electrodes.
A plasma support gas is necessary, and Ar is common. Samples can be deposited on one of the electrodes, or if conducting can make up one electrode.

• Glow discharge-optical emission spectrometry (GD-OES)

• Inductively coupled plasma-atomic emission spectrometry (ICP-AES)

• Laser Induced Breakdown Spectroscopy (LIBS), also called Laser-induced plasma spectrometry (LIPS)

• Microwave-induced plasma (MIP)

Spark or arc (emission) spectroscopy - is used for the analysis of metallic elements in solid samples. For non-conductive materials, a sample is ground with graphite powder to make it conductive. In traditional arc spectroscopy methods, a sample of the solid was commonly ground up and destroyed during analysis. An electric arc or spark is passed through the sample, heating the sample to a high temperature to excite the atoms in it. The excited analyte atoms glow, emitting light at various wavelengths that could be detected by common spectroscopic methods. Since the conditions producing the arc emission typically are not controlled quantitatively, the analysis for the elements is qualitative.
Nowadays, the spark sources with controlled discharges under an argon atmosphere allow that this method can be considered eminently quantitative, and its use is widely expanded worldwide through production control laboratories of foundries and steel mills.

Visible

Many atoms emit or absorb visible light. In order to obtain a fine line spectrum, the atoms must be in a gas phase. This means that the substance has to be vaporised. The spectrum is studied in absorption or emission. Visible absorption spectroscopy is often combined with UV absorption spectroscopy in UV/Vis spectroscopy. Although this form may be uncommon as the human eye is a similar indicator, it still proves useful when distinguishing colours.

Ultraviolet

All atoms absorb in the Ultraviolet (UV) region because these photons are energetic enough to excite outer electrons. If the frequency is high enough, photoionization takes place. UV spectroscopy is also used in quantifying protein and DNA concentration as well as the ratio of protein to DNA concentration in a solution. Several amino acids usually found in protein, such as tryptophan, absorb light in the 280 nm range and DNA absorbs light in the 260 nm range. For this reason, the ratio of 260/280 nm absorbance is a good general indicator of the relative purity of a solution in terms of these two macromolecules. Reasonable estimates of protein or DNA concentration can also be made this way using Beer's law.

Infrared

Infrared spectroscopy offers the possibility to measure different types of inter atomic bond vibrations at different frequencies. Especially in organic chemistry the analysis of IR absorption spectra shows what type of bonds are present in the sample. It is also an important method for analysing polymers and constituents like fillers, pigments and plasticizers.

Near Infrared (NIR)

The near infrared NIR range, immediately beyond the visible wavelength range, is especially important for practical applications because of the much greater penetration depth of NIR radiation into the sample than in the case of mid IR spectroscopy range. This allows also large samples to be measured in each scan by NIR spectroscopy, and is currently employed for many practical applications such as: rapid grain analysis, medical diagnosis pharmaceuticals/medicines, biotechnology, genomics analysis, proteomic analysis, interactomics research, inline textile monitoring, food analysis and chemical imaging/hyperspectral imaging of intact organisms, plastics, textiles, insect detection, forensic lab application, crime detection and various military applications.

Raman

Raman spectroscopy uses the inelastic scattering of light to analyse vibrational and rotational modes of molecules. The resulting 'fingerprints' are an aid to analysis.

Coherent anti-Stokes Raman spectroscopy (CARS)

CARS is a recent technique that has high sensitivity and powerful applications for ''in vivo'' spectroscopy and imaging.

Nuclear magnetic resonance

Nuclear magnetic resonance spectroscopy analyzes the magnetic properties of certain atomic nuclei to determine different electronic local environments of hydrogen, carbon, or other atoms in an organic compound or other compound. This is used to help determine the structure of the compound.

Photoemission

Mössbauer

Transmission or conversion-electron (CEMS) modes of Mössbauer spectroscopy probe the properties of specific isotope nuclei in different atomic environments by analyzing the resonant absorption of characteristic energy gamma-rays known as the Mössbauer effect.