MASS SPECTRA - THE M+2 PEAK

MASS SPECTRA - THE M+2 PEAK This page explains how the M+2 peak in a mass spectrum arises from the presence of chlorine or bromine atoms in an organic compound. It also deals briefly with the origin of the M+4 peak in compounds containing two chlorine atoms.
	Note:  Before you start this page, it would be a good idea to have a reasonable understanding about how a mass spectrum is produced and the sort of information you can get from it. If you haven't already done so, explore the mass spectrometry menu before you go on.
The effect of chlorine or bromine atoms on the mass spectrum of an organic compound Compounds containing chlorine atoms One chlorine atom in a compound
	Note:  All the mass spectra on this page have been drawn using data from the Spectral Data Base System for Organic Compounds (SDBS) at the National Institute of Materials and Chemical Research in Japan. With one exception, they have been simplified by omitting all the minor lines with peak heights of 2% or less of the base peak (the tallest peak).
The molecular ion peaks (M+ and M+2) each contain one chlorine atom - but the chlorine can be either of the two chlorine isotopes, 35Cl and 37Cl. The molecular ion containing the 35Cl isotope has a relative formula mass of 78. The one containing 37Cl has a relative formula mass of 80 - hence the two lines at m/z = 78 and m/z = 80. Notice that the peak heights are in the ratio of 3 : 1. That reflects the fact that chlorine contains 3 times as much of the 35Cl isotope as the 37Cl one. That means that there will be 3 times more molecules containing the lighter isotope than the heavier one. So . . . if you look at the molecular ion region, and find two peaks separated by 2 m/z units and with a ratio of 3 : 1 in the peak heights, that tells you that the molecule contains 1 chlorine atom. You might also have noticed the same pattern at m/z = 63 and m/z = 65 in the mass spectrum above. That pattern is due to fragment ions also containing one chlorine atom - which could either be 35Cl or 37Cl. The fragmentation that produced those ions was:
	Note:  If you aren't sure about fragmentation you might like to have a look at this link.
Two chlorine atoms in a compound
	Note:  This spectrum has been simplified by omitting all the minor lines with peak heights of less than 1% of the base peak (the tallest peak). This contains more minor lines than other mass spectra in this section. It was necessary because otherwise an important line in the molecular ion region would have been missing.
The lines in the molecular ion region (at m/z values of 98, 100 ands 102) arise because of the various combinations of chlorine isotopes that are possible. The carbons and hydrogens add up to 28 - so the various possible molecular ions could be: 28 + 35 + 35 = 98 28 + 35 + 37 = 100 28 + 37 + 37 = 102 If you have the necessary maths, you could show that the chances of these arrangements occurring are in the ratio of 9:6:1 - and this is the ratio of the peak heights. If you don't know the right bit of maths, just learn this ratio! So . . . if you have 3 lines in the molecular ion region (M+, M+2 and M+4) with gaps of 2 m/z units between them, and with peak heights in the ratio of 9:6:1, the compound contains 2 chlorine atoms. Compounds containing bromine atoms Bromine has two isotopes, 79Br and 81Br in an approximately 1:1 ratio (50.5 : 49.5 if you want to be fussy!). That means that a compound containing 1 bromine atom will have two peaks in the molecular ion region, depending on which bromine isotope the molecular ion contains. Unlike compounds containing chlorine, though, the two peaks will be very similar in height. The carbons and hydrogens add up to 29. The M+ and M+2 peaks are therefore at m/z values given by: 29 + 79 = 108 29 + 81 = 110 So . . . if you have two lines in the molecular ion region with a gap of 2 m/z units between them and with almost equal heights, this shows the presence of a bromine atom in the molecule.

THE MASS SPECTRA OF ELEMENTS

THE MASS SPECTRA OF ELEMENTS This page looks at the information you can get from the mass spectrum of an element. It shows how you can find out the masses and relative abundances of the various isotopes of the element and use that information to calculate the relative atomic mass of the element. It also looks at the problems thrown up by elements with diatomic molecules - like chlorine, Cl2. The mass spectrum of monatomic elements Monatomic elements include all those except for things like chlorine, Cl2, with molecules containing more than one atom. The mass spectrum for boron
	Note:  If you need to know how this diagram is obtained, you should read the page describing how a mass spectrometer works.
The number of isotopes The two peaks in the mass spectrum shows that there are 2 isotopes of boron - with relative isotopic masses of 10 and 11 on the 12C scale.
	Notes:  Isotopes are atoms of the same element (and so with the same number of protons), but with different masses due to having different numbers of neutrons. We are assuming (and shall do all through this page) that all the ions recorded have a charge of 1+. That means that the mass/charge ratio (m/z) gives you the mass of the isotope directly. The carbon-12 scale is a scale on which the mass of the 12C isotope weighs exactly 12 units.
The abundance of the isotopes The relative sizes of the peaks gives you a direct measure of the relative abundances of the isotopes. The tallest peak is often given an arbitrary height of 100 - but you may find all sorts of other scales used. It doesn't matter in the least. You can find the relative abundances by measuring the lines on the stick diagram. In this case, the two isotopes (with their relative abundances) are: boron-10 23 boron-11 100 Working out the relative atomic mass The relative atomic mass (RAM) of an element is given the symbol Ar and is defined as: The relative atomic mass of an element is the weighted average of the masses of the isotopes on a scale on which a carbon-12 atom has a mass of exactly 12 units. A "weighted average" allows for the fact that there won't be equal amounts of the various isotopes. The example coming up should make that clear. Suppose you had 123 typical atoms of boron. 23 of these would be 10B and 100 would be 11B. The total mass of these would be (23 x 10) + (100 x 11) = 1330 The average mass of these 123 atoms would be 1330 / 123 = 10.8 (to 3 significant figures). 10.8 is the relative atomic mass of boron. Notice the effect of the "weighted" average. A simple average of 10 and 11 is, of course, 10.5. Our answer of 10.8 allows for the fact that there are a lot more of the heavier isotope of boron - and so the "weighted" average ought to be closer to that. The mass spectrum for zirconium The number of isotopes The 5 peaks in the mass spectrum shows that there are 5 isotopes of zirconium - with relative isotopic masses of 90, 91, 92, 94 and 96 on the 12C scale. The abundance of the isotopes This time, the relative abundances are given as percentages. Again you can find these relative abundances by measuring the lines on the stick diagram. In this case, the 5 isotopes (with their relative percentage abundances) are: zirconium-90 51.5 zirconium-91 11.2 zirconium-92 17.1 zirconium-94 17.4 zirconium-96 2.8
	Note:  You almost certainly wouldn't be able to measure these peaks to this degree of accuracy, but your examiners may well give you the data in number form anyway. We'll do the sum with the more accurate figures.
Working out the relative atomic mass Suppose you had 100 typical atoms of zirconium. 51.5 of these would be 90Zr, 11.2 would be 91Zr and so on.
	Note:  If you object to the idea of having 51.5 atoms or 11.2 atoms and so on, just assume you've got 1000 atoms instead of 100. That way you will have 515 atoms, 112 atoms, etc. Most people don't get in a sweat over this, and just use the numbers as they are!
The total mass of these 100 typical atoms would be (51.5 x 90) + (11.2 x 91) + (17.1 x 92) + (17.4 x 94) + (2.8 x 96) = 9131.8 The average mass of these 100 atoms would be 9131.8 / 100 = 91.3 (to 3 significant figures). 91.3 is the relative atomic mass of zirconium.
	Note:  If you want further examples of calculating relative atomic masses from mass spectra, you might like to refer to my book, Calculations in A level Chemistry.
The mass spectrum of chlorine Chlorine is taken as typical of elements with more than one atom per molecule. We'll look at its mass spectrum to show the sort of problems involved. Chlorine has two isotopes, 35Cl and 37Cl, in the approximate ratio of 3 atoms of 35Cl to 1 atom of 37Cl. You might suppose that the mass spectrum would look like this: You would be wrong! The problem is that chlorine consists of molecules, not individual atoms. When chlorine is passed into the ionisation chamber, an electron is knocked off the molecule to give a molecular ion, Cl2+. These ions won't be particularly stable, and some will fall apart to give a chlorine atom and a Cl+ ion. The term for this is fragmentation. If the Cl atom formed isn't then ionised in the ionisation chamber, it simply gets lost in the machine - neither accelerated nor deflected. The Cl+ ions will pass through the machine and will give lines at 35 and 37, depending on the isotope and you would get exactly the pattern in the last diagram. The problem is that you will also record lines for the unfragmented Cl2+ ions. Think about the possible combinations of chlorine-35 and chlorine-37 atoms in a Cl2+ ion. Both atoms could be 35Cl, both atoms could be 37Cl, or you could have one of each sort. That would give you total masses of the Cl2+ ion of: 35 + 35 = 70 35 + 37 = 72 37 + 37 = 74 That means that you would get a set of lines in the m/z = 70 region looking like this: These lines would be in addition to the lines at 35 and 37. The relative heights of the 70, 72 and 74 lines are in the ratio 9:6:1. If you know the right bit of maths, it's very easy to show this. If not, don't worry. Just remember that the ratio is 9:6:1. What you can't do is make any predictions about the relative heights of the lines at 35/37 compared with those at 70/72/74. That depends on what proportion of the molecular ions break up into fragments. That's why you've got the chlorine mass spectrum in two separate bits so far. You must realise that the vertical scale in the diagrams of the two parts of the spectrum isn't the same. The overall mass spectrum looks like this:

PAPER CHROMATOGRAPHY

PAPER CHROMATOGRAPHY This page is an introduction to paper chromatography - including two way chromatography. Carrying out paper chromatography Background Chromatography is used to separate mixtures of substances into their components. All forms of chromatography work on the same principle. They all have a stationary phase (a solid, or a liquid supported on a solid) and a mobile phase (a liquid or a gas). The mobile phase flows through the stationary phase and carries the components of the mixture with it. Different components travel at different rates. We'll look at the reasons for this further down the page. In paper chromatography, the stationary phase is a very uniform absorbent paper. The mobile phase is a suitable liquid solvent or mixture of solvents. Producing a paper chromatogram You probably used paper chromatography as one of the first things you ever did in chemistry to separate out mixtures of coloured dyes - for example, the dyes which make up a particular ink. That's an easy example to take, so let's start from there. Suppose you have three blue pens and you want to find out which one was used to write a message. Samples of each ink are spotted on to a pencil line drawn on a sheet of chromatography paper. Some of the ink from the message is dissolved in the minimum possible amount of a suitable solvent, and that is also spotted onto the same line. In the diagram, the pens are labelled 1, 2 and 3, and the message ink as M.
	Note:  The chromatography paper will in fact be pure white - not pale grey. I'm forced to show it as off-white because of the way I construct the diagrams. Anything I draw as pure white allows the background colour of the page to show through.
The paper is suspended in a container with a shallow layer of a suitable solvent or mixture of solvents in it. It is important that the solvent level is below the line with the spots on it. The next diagram doesn't show details of how the paper is suspended because there are too many possible ways of doing it and it clutters the diagram. Sometimes the paper is just coiled into a loose cylinder and fastened with paper clips top and bottom. The cylinder then just stands in the bottom of the container. The reason for covering the container is to make sure that the atmosphere in the beaker is saturated with solvent vapour. Saturating the atmosphere in the beaker with vapour stops the solvent from evaporating as it rises up the paper. As the solvent slowly travels up the paper, the different components of the ink mixtures travel at different rates and the mixtures are separated into different coloured spots. The diagram shows what the plate might look like after the solvent has moved almost to the top. It is fairly easy to see from the final chromatogram that the pen that wrote the message contained the same dyes as pen 2. You can also see that pen 1 contains a mixture of two different blue dyes - one of which might be the same as the single dye in pen 3. Rf values Some compounds in a mixture travel almost as far as the solvent does; some stay much closer to the base line. The distance travelled relative to the solvent is a constant for a particular compound as long as you keep everything else constant - the type of paper and the exact composition of the solvent, for example. The distance travelled relative to the solvent is called the Rf value. For each compound it can be worked out using the formula: For example, if one component of a mixture travelled 9.6 cm from the base line while the solvent had travelled 12.0 cm, then the Rf value for that component is: In the example we looked at with the various pens, it wasn't necessary to measure Rf values because you are making a direct comparison just by looking at the chromatogram. You are making the assumption that if you have two spots in the final chromatogram which are the same colour and have travelled the same distance up the paper, they are most likely the same compound. It isn't necessarily true of course - you could have two similarly coloured compounds with very similar Rf values. We'll look at how you can get around that problem further down the page. What if the substances you are interested in are colourless? In some cases, it may be possible to make the spots visible by reacting them with something which produces a coloured product. A good example of this is in chromatograms produced from amino acid mixtures. Suppose you had a mixture of amino acids and wanted to find out which particular amino acids the mixture contained. For simplicity we'll assume that you know the mixture can only possibly contain five of the common amino acids. A small drop of a solution of the mixture is placed on the base line of the paper, and similar small spots of the known amino acids are placed alongside it. The paper is then stood in a suitable solvent and left to develop as before. In the diagram, the mixture is M, and the known amino acids are labelled 1 to 5. The position of the solvent front is marked in pencil and the chromatogram is allowed to dry and is then sprayed with a solution of ninhydrin. Ninhydrin reacts with amino acids to give coloured compounds, mainly brown or purple. The left-hand diagram shows the paper after the solvent front has almost reached the top. The spots are still invisible. The second diagram shows what it might look like after spraying with ninhydrin. There is no need to measure the Rf values because you can easily compare the spots in the mixture with those of the known amino acids - both from their positions and their colours. In this example, the mixture contains the amino acids labelled as 1, 4 and 5. And what if the mixture contained amino acids other than the ones we have used for comparison? There would be spots in the mixture which didn't match those from the known amino acids. You would have to re-run the experiment using other amino acids for comparison. Two way paper chromatography Two way paper chromatography gets around the problem of separating out substances which have very similar Rf values. I'm going to go back to talking about coloured compounds because it is much easier to see what is happening. You can perfectly well do this with colourless compounds - but you have to use quite a lot of imagination in the explanation of what is going on! This time a chromatogram is made starting from a single spot of mixture placed towards one end of the base line. It is stood in a solvent as before and left until the solvent front gets close to the top of the paper. In the diagram, the position of the solvent front is marked in pencil before the paper dries out. This is labelled as SF1 - the solvent front for the first solvent. We shall be using two different solvents. If you look closely, you may be able to see that the large central spot in the chromatogram is partly blue and partly green. Two dyes in the mixture have almost the same Rf values. They could equally well, of course, both have been the same colour - in which case you couldn't tell whether there was one or more dye present in that spot. What you do now is to wait for the paper to dry out completely, and then rotate it through 90°, and develop the chromatogram again in a different solvent. It is very unlikely that the two confusing spots will have the same Rf values in the second solvent as well as the first, and so the spots will move by a different amount. The next diagram shows what might happen to the various spots on the original chromatogram. The position of the second solvent front is also marked. You wouldn't, of course, see these spots in both their original and final positions - they have moved! The final chromatogram would look like this: Two way chromatography has completely separated out the mixture into four distinct spots. If you want to identify the spots in the mixture, you obviously can't do it with comparison substances on the same chromatogram as we looked at earlier with the pens or amino acids examples. You would end up with a meaningless mess of spots. You can, though, work out the Rf values for each of the spots in both solvents, and then compare these with values that you have measured for known compounds under exactly the same conditions. How does paper chromatography work? Although paper chromatography is simple to do, it is quite difficult to explain compared with thin layer chromatography. The explanation depends to some extent on what sort of solvent you are using, and many sources gloss over the problem completely. If you haven't already done so, it would be helpful if you could read the explanation for how thin layer chromatography works (link below). That will save me a lot of repetition, and I can concentrate on the problems.
	Note:  You will find the explanation for how thin layer chromatography works by following this link. Use the BACK button on your browser to return quickly to this page when yhou have read it.
The essential structure of paper Paper is made of cellulose fibres, and cellulose is a polymer of the simple sugar, glucose. The key point about cellulose is that the polymer chains have -OH groups sticking out all around them. To that extent, it presents the same sort of surface as silica gel or alumina in thin layer chromatography. It would be tempting to try to explain paper chromatography in terms of the way that different compounds are adsorbed to different extents on to the paper surface. In other words, it would be nice to be able to use the same explanation for both thin layer and paper chromatography. Unfortunately, it is more complicated than that! The complication arises because the cellulose fibres attract water vapour from the atmosphere as well as any water that was present when the paper was made. You can therefore think of paper as being cellulose fibres with a very thin layer of water molecules bound to the surface. It is the interaction with this water which is the most important effect during paper chromatography. Paper chromatography using a non-polar solvent Suppose you use a non-polar solvent such as hexane to develop your chromatogram. Non-polar molecules in the mixture that you are trying to separate will have little attraction for the water molecules attached to the cellulose, and so will spend most of their time dissolved in the moving solvent. Molecules like this will therefore travel a long way up the paper carried by the solvent. They will have relatively high Rf values. On the other hand, polar molecules will have a high attraction for the water molecules and much less for the non-polar solvent. They will therefore tend to dissolve in the thin layer of water around the cellulose fibres much more than in the moving solvent. Because they spend more time dissolved in the stationary phase and less time in the mobile phase, they aren't going to travel very fast up the paper. The tendency for a compound to divide its time between two immiscible solvents (solvents such as hexane and water which won't mix) is known as partition. Paper chromatography using a non-polar solvent is therefore a type of partition chromatography. Paper chromatography using a water and other polar solvents A moment's thought will tell you that partition can't be the explanation if you are using water as the solvent for your mixture. If you have water as the mobile phase and the water bound on to the cellulose as the stationary phase, there can't be any meaningful difference between the amount of time a substance spends in solution in either of them. All substances should be equally soluble (or equally insoluble) in both. And yet the first chromatograms that you made were probably of inks using water as your solvent. If water works as the mobile phase as well being the stationary phase, there has to be some quite different mechanism at work - and that must be equally true for other polar solvents like the alcohols, for example. Partition only happens between solvents which don't mix with each other. Polar solvents like the small alcohols do mix with water. In researching this topic, I haven't found any easy explanation for what happens in these cases. Most sources ignore the problem altogether and just quote the partition explanation without making any allowance for the type of solvent you are using. Other sources quote mechanisms which have so many strands to them that they are far too complicated for this introductory level. I'm therefore not taking this any further - you shouldn't need to worry about this at UK A level, or its various equivalents.

GAS-LIQUID CHROMATOGRAPHY

GAS-LIQUID CHROMATOGRAPHY Gas-liquid chromatography (often just called gas chromatography) is a powerful tool in analysis. It has all sorts of variations in the way it is done - if you want full details, a Google search on gas chromatography will give you scary amounts of information if you need it! This page just looks in a simple introductory way at how it can be carried out. Carrying out gas-liquid chromatography Introduction All forms of chromatography involve a stationary phase and a mobile phase. In all the other forms of chromatography you will meet at this level, the mobile phase is a liquid. In gas-liquid chromatography, the mobile phase is a gas such as helium and the stationary phase is a high boiling point liquid absorbed onto a solid. How fast a particular compound travels through the machine will depend on how much of its time is spent moving with the gas as opposed to being attached to the liquid in some way. A flow scheme for gas-liquid chromatography
	Note:  You will have to imagine the coiled column in its oven. Drawing a convincing and tidy coil defeated me completely!
Injection of the sample Very small quantities of the sample that you are trying to analyse are injected into the machine using a small syringe. The syringe needle passes through a thick rubber disc (known as a septum) which reseals itself again when the syringe is pulled out. The injector is contained in an oven whose temperature can be controlled. It is hot enough so that all the sample boils and is carried into the column as a gas by the helium (or other carrier gas). How the column works The packing material There are two main types of column in gas-liquid chromatography. One of these is a long thin tube packed with the stationary phase; the other is even thinner and has the stationary phase bonded to its inner surface. To keep things simple, we are just going to look at the packed column. The column is typically made of stainless steel and is between 1 and 4 metres long with an internal diameter of up to 4 mm. It is coiled up so that it will fit into a thermostatically controlled oven. The column is packed with finely ground diatomaceous earth, which is a very porous rock. This is coated with a high boiling liquid - typically a waxy polymer. The column temperature The temperature of the column can be varied from about 50°C to 250°C. It is cooler than the injector oven, so that some components of the mixture may condense at the beginning of the column. In some cases, as you will see below, the column starts off at a low temperature and then is made steadily hotter under computer control as the analysis proceeds. How separation works on the column One of three things might happen to a particular molecule in the mixture injected into the column: It may condense on the stationary phase. It may dissolve in the liquid on the surface of the stationary phase. It may remain in the gas phase. None of these things is necessarily permanent. A compound with a boiling point higher than the temperature of the column will obviously tend to condense at the start of the column. However, some of it will evaporate again in the same way that water evaporates on a warm day - even though the temperature is well below 100°C. The chances are that it will then condense again a little further along the column. Similarly, some molecules may dissolve in the liquid stationary phase Some compounds will be more soluble in the liquid than others. The more soluble ones will spend more of their time absorbed into the stationary phase; the less soluble ones will spend more of their time in the gas. The process where a substance divides itself between two immiscible solvents because it is more soluble in one than the other is known as partition. Now, you might reasonably argue that a gas such as helium can't really be described as a "solvent". But the term partition is still used in gas-liquid chromatography. You can say that a substance partitions itself between the liquid stationary phase and the gas. Any molecule in the substance spends some of its time dissolved in the liquid and some of its time carried along with the gas. Retention time The time taken for a particular compound to travel through the column to the detector is known as its retention time. This time is measured from the time at which the sample is injected to the point at which the display shows a maximum peak height for that compound. Different compounds have different retention times. For a particular compound, the retention time will vary depending on: the boiling point of the compound. A compound which boils at a temperature higher than the column temperature is going to spend nearly all of its time condensed as a liquid at the beginning of the column. So high boiling point means a long retention time. the solubility in the liquid phase. The more soluble a compound is in the liquid phase, the less time it will spend being carried along by the gas. High solubility in the liquid phase means a high retention time. the temperature of the column. A higher temperature will tend to excite molecules into the gas phase - either because they evaporate more readily, or because they are so energetic that the attractions of the liquid no longer hold them. A high column temperature shortens retention times for everything in the column. For a given sample and column, there isn't much you can do about the boiling points of the compounds or their solubility in the liquid phase - but you do have control over the temperature. The lower the temperature of the column, the better the separation you will get - but it could take a very long time to get the compounds through which are condensing at the beginning of the column! On the other hand, using a high temperature, everything will pass through the column much more quickly - but less well separated out. If everything passed through in a very short time, there isn't going to be much space between their peaks on the chromatogram. The answer is to start with the column relatively cool, and then gradually and very regularly increase the temperature. At the beginning, compounds which spend most of their time in the gas phase will pass quickly through the column and be detected. Increasing the temperature a bit will encourage the slightly "stickier" compounds through. Increasing the temperature still more will force the very "sticky" molecules off the stationary phase and through the column. The detector There are several different types of detector in use. The flame ionisation detector described below is commonly used and is easier to describe and explain than the alternatives. A flame ionisation detector In terms of reaction mechanisms, the burning of an organic compound is very complicated. During the process, small amounts of ions and electrons are produced in the flame. The presence of these can be detected. The whole detector is enclosed in its own oven which is hotter than the column temperature. That stops anything condensing in the detector.
	Note:  This is simplified for clarity. There obviously has to be some way of lighting the flame. This is done with an electrically heated coil, but including it clutters the diagram.
If there is nothing organic coming through from the column, you just have a flame of hydrogen burning in air. Now suppose that one of the compounds in the mixture you are analysing starts to come through. As it burns, it will produce small amounts of ions and electrons in the flame. The positive ions will be attracted to the cylindrical cathode. Negative ions and electrons will be attracted towards the jet itself which is the anode. This is much the same as what happens during normal electrolysis. At the cathode, the positive ions will pick up electrons from the cathode and be neutralised. At the anode, any electrons in the flame will transfer to the positive electrode; and negative ions will give their electrons to the electrode and be neutralised. This loss of electrons from one electrode and gain at the other will result in a flow of electrons in the external circuit from the anode to the cathode. In other words, you get an electric current. The current won't be very big, but it can be amplified. The more of the organic compound there is in the flame, the more ions will be produced, and so the higher the current will be. As a reasonable approximation, especially if you are talking about similar compounds, the current you measure is proportional to the amount of compound in the flame. Disadvantages of the flame ionisation detector The main disadvantage is that it destroys everything coming out of the column as it detects it. If you wanted to send the product to a mass spectrometer, for example, for further analysis, you couldn't use a flame ionisation detector. Interpreting the output from the detector The output will be recorded as a series of peaks - each one representing a compound in the mixture passing through the detector. As long as you were careful to control the conditions on the column, you could use the retention times to help to identify the compounds present - provided, of course, that you (or somebody else) had already measured them for pure samples of the various compounds under those identical conditions. But you can also use the peaks as a way of measuring the relative quantities of the compounds present. This is only accurate if you are analysing mixtures of similar compounds - for example, of similar hydrocarbons. The areas under the peaks are proportional to the amount of each compound which has passed the detector, and these areas can be calculated automatically by the computer linked to the display. The areas it would measure are shown in green in the (very simplified) diagram. Note that it isn't the peak height that matters, but the total area under the peak. In this particular example, the left-hand peak is both tallest and has the greatest area. That isn't necessarily always so. There might be a lot of one compound present, but it might emerge from the column in relatively small amounts over quite a long time. Measuring the area rather than the peak height allows for this. Coupling a gas chromatogram to a mass spectrometer This can't be done with a flame ionisation detector which destroys everything passing through it. Assuming you are using a non-destructive detector . . . When the detector is showing a peak, some of what is passing through the detector at that time can be diverted to a mass spectrometer. There it will give a fragmentation pattern which can be compared against a computer database of known patterns. That means that the identity of a huge range of compounds can be found without having to know their retention times.

HIGH PERFORMANCE LIQUID CHROMATOGRAPHY - HPLC

HIGH PERFORMANCE LIQUID CHROMATOGRAPHY - HPLC High performance liquid chromatography is a powerful tool in analysis. This page looks at how it is carried out and shows how it uses the same principles as in thin layer chromatography and column chromatography.
	Note:  It is important to read the introductory page about thin layer chromatography before you continue with this one - particularly the part about how thin layer chromatography works. High performance liquid chromatography works on the same basic principle. HPLC is essentially an adaptation of column chromatography - so it might be a good idea to have a (very quick) look at that as well. Use the BACK button on your browser to return quickly to this page.
Carrying out HPLC Introduction High performance liquid chromatography is basically a highly improved form of column chromatography. Instead of a solvent being allowed to drip through a column under gravity, it is forced through under high pressures of up to 400 atmospheres. That makes it much faster. It also allows you to use a very much smaller particle size for the column packing material which gives a much greater surface area for interactions between the stationary phase and the molecules flowing past it. This allows a much better separation of the components of the mixture. The other major improvement over column chromatography concerns the detection methods which can be used. These methods are highly automated and extremely sensitive. The column and the solvent Confusingly, there are two variants in use in HPLC depending on the relative polarity of the solvent and the stationary phase. Normal phase HPLC This is essentially just the same as you will already have read about in thin layer chromatography or column chromatography. Although it is described as "normal", it isn't the most commonly used form of HPLC. The column is filled with tiny silica particles, and the solvent is non-polar - hexane, for example. A typical column has an internal diameter of 4.6 mm (and may be less than that), and a length of 150 to 250 mm. Polar compounds in the mixture being passed through the column will stick longer to the polar silica than non-polar compounds will. The non-polar ones will therefore pass more quickly through the column. Reversed phase HPLC In this case, the column size is the same, but the silica is modified to make it non-polar by attaching long hydrocarbon chains to its surface - typically with either 8 or 18 carbon atoms in them. A polar solvent is used - for example, a mixture of water and an alcohol such as methanol. In this case, there will be a strong attraction between the polar solvent and polar molecules in the mixture being passed through the column. There won't be as much attraction between the hydrocarbon chains attached to the silica (the stationary phase) and the polar molecules in the solution. Polar molecules in the mixture will therefore spend most of their time moving with the solvent. Non-polar compounds in the mixture will tend to form attractions with the hydrocarbon groups because of van der Waals dispersion forces. They will also be less soluble in the solvent because of the need to break hydrogen bonds as they squeeze in between the water or methanol molecules, for example. They therefore spend less time in solution in the solvent and this will slow them down on their way through the column. That means that now it is the polar molecules that will travel through the column more quickly. Reversed phase HPLC is the most commonly used form of HPLC.
	Note:  I have been a bit careful about how I have described the attractions of the non-polar molecules to the surface of the stationary phase. In particular, I have avoided the use of the word "adsorpion". Adsorption is when a molecule sticks to the surface of a solid. Especially if you had small molecules in your mixture, some could get in between the long C18 chains to give what is essentially a solution. You could therefore say that non-polar molecules were more soluble in the hydrocarbon on the surface of the silica than they are in the polar solvent - and so spend more time in this alternative "solvent". Where a solute divides itself between two different solvents because it is more soluble in one than the other, we call it partition. So is this adsorption or partition? You could argue it both ways! Be prepared to find it described as either.
Looking at the whole process A flow scheme for HPLC Injection of the sample Injection of the sample is entirely automated, and you wouldn't be expected to know how this is done at this introductory level. Because of the pressures involved, it is not the same as in gas chromatography (if you have already studied that). Retention time The time taken for a particular compound to travel through the column to the detector is known as its retention time. This time is measured from the time at which the sample is injected to the point at which the display shows a maximum peak height for that compound. Different compounds have different retention times. For a particular compound, the retention time will vary depending on: the pressure used (because that affects the flow rate of the solvent) the nature of the stationary phase (not only what material it is made of, but also particle size) the exact composition of the solvent the temperature of the column That means that conditions have to be carefully controlled if you are using retention times as a way of identifying compounds. The detector There are several ways of detecting when a substance has passed through the column. A common method which is easy to explain uses ultra-violet absorption. Many organic compounds absorb UV light of various wavelengths. If you have a beam of UV light shining through the stream of liquid coming out of the column, and a UV detector on the opposite side of the stream, you can get a direct reading of how much of the light is absorbed. The amount of light absorbed will depend on the amount of a particular compound that is passing through the beam at the time. You might wonder why the solvents used don't absorb UV light. They do! But different compounds absorb most strongly in different parts of the UV spectrum. Methanol, for example, absorbs at wavelengths below 205 nm, and water below 190 nm. If you were using a methanol-water mixture as the solvent, you would therefore have to use a wavelength greater than 205 nm to avoid false readings from the solvent.
	Note:  If you are interested, there is a whole section about UV-visible spectroscopy on the site. This explores the question of the absorption of UV and visible light by organic compounds in some detail.
Interpreting the output from the detector The output will be recorded as a series of peaks - each one representing a compound in the mixture passing through the detector and absorbing UV light. As long as you were careful to control the conditions on the column, you could use the retention times to help to identify the compounds present - provided, of course, that you (or somebody else) had already measured them for pure samples of the various compounds under those identical conditions. But you can also use the peaks as a way of measuring the quantities of the compounds present. Let's suppose that you are interested in a particular compound, X. If you injected a solution containing a known amount of pure X into the machine, not only could you record its retention time, but you could also relate the amount of X to the peak that was formed. The area under the peak is proportional to the amount of X which has passed the detector, and this area can be calculated automatically by the computer linked to the display. The area it would measure is shown in green in the (very simplified) diagram. If the solution of X was less concentrated, the area under the peak would be less - although the retention time will still be the same. For example: This means that it is possible to calibrate the machine so that it can be used to find how much of a substance is present - even in very small quantities. Be careful, though! If you had two different substances in the mixture (X and Y) could you say anything about their relative amounts? Not if you were using UV absorption as your detection method. In the diagram, the area under the peak for Y is less than that for X. That may be because there is less Y than X, but it could equally well be because Y absorbs UV light at the wavelength you are using less than X does. There might be large quantities of Y present, but if it only absorbed weakly, it would only give a small peak.

COLUMN CHROMATOGRAPHY

COLUMN CHROMATOGRAPHY This page shows how the same principles used in thin layer chromatography can be applied on a larger scale to separate mixtures in column chromatography. Column chromatography is often used to purify compounds made in the lab.
	Note:  It is important to read the introductory page about thin layer chromatography before you continue with this one - particularly the part about how thin layer chromatography works, although you will also need some idea about how to make a thin layer chromatogram. Use the BACK button on your browser to return quickly to this page.
Carrying out column chromatography The column In thin layer chromatography, the stationary phase is a thin layer of silica gel or alumina on a glass, metal or plastic plate. Column chromatography works on a much larger scale by packing the same materials into a vertical glass column. Various sizes of chromatography columns are used, and if you follow a link at the bottom of the page to the Organic Chemistry section of the Colorado University site, you will find photographs of various columns. In a school lab, it is often convenient to use an ordinary burette as a chromatography column. Using the column Suppose you wanted to separate a mixture of two coloured compounds - one yellow, one blue. The mixture looks green. You would make a concentrated solution of the mixture preferably in the solvent used in the column. First you open the tap to allow the solvent already in the column to drain so that it is level with the top of the packing material, and then add the solution carefully to the top of the column. Then you open the tap again so that the coloured mixture is all absorbed into the top of the packing material, so that it might look like this: Next you add fresh solvent to the top of the column, trying to disturb the packing material as little as possible. Then you open the tap so that the solvent can flow down through the column, collecting it in a beaker or flask at the bottom. As the solvent runs through, you keep adding fresh solvent to the top so that the column never dries out. The next set of diagrams shows what might happen over time.
	Note:  These diagrams are very simplified in order to make them easier to draw. In reality, the colours won't separate out into these neat blocks, but will probably be much more spread out - more so the further down the column they get.
Explaining what is happening This assumes that you have read the explanation for what happens during thin layer chromatography. If you haven't, follow the very first link at the top of the page and come back to this point afterwards. The blue compound is obviously more polar than the yellow one - it perhaps even has the ability to hydrogen bond. You can tell this because the blue compound doesn't travel through the column very quickly. That means that it must adsorb more strongly to the silica gel or alumina than the yellow one. The less polar yellow one spends more of its time in the solvent and therefore washes through the column much faster. The process of washing a compound through a column using a solvent is known as elution. The solvent is sometimes known as the eluent. What if you want to collect the blue compound as well? It is going to take ages to wash the blue compound through at the rate it is travelling at the moment! However, there is no reason why you can't change the solvent during elution. Suppose you replace the solvent you have been using by a more polar solvent once the yellow has all been collected. That will have two effects, both of which will speed the blue compound through the column. The polar solvent will compete for space on the silica gel or alumina with the blue compound. Any space temporarily occupied by solvent molecules on the surface of the stationary phase isn't available for blue molecules to stick to and this will tend to keep them moving along in the solvent. There will be a greater attraction between the polar solvent molecules and the polar blue molecules. This will tend to attract any blue molecules sticking to the stationary phase back into solution. The net effect is that with a more polar solvent, the blue compound spends more time in solution, and so moves faster. So why not use this alternative solvent in the first place? The answer is that if both of the compounds in the mixture travel quickly through the column right from the beginning, you probably won't get such a good separation. What if everything in your mixture is colourless? If you were going to use column chromatography to purify the product of an organic preparation, it is quite likely that the product that you want will be colourless even if one or more of the impurities is coloured. Let's assume the worst case that everything is colourless. How do you know when the substance you want has reached the bottom of the column? There is no quick and easy way of doing this! What you do is collect what comes out of the bottom of the column in a whole series of labelled tubes. How big each sample is will obviously depend on how big the column is - you might collect 1 cm3 samples or 5 cm3 samples or whatever is appropriate. You can then take a drop from each solution and make a thin layer chromatogram from it. You would place the drop on the base line alongside a drop from a pure sample of the compound that you are making. By doing this repeatedly, you can identify which of your samples collected at the bottom of the column contain the desired product, and only the desired product. Once you know this, you can combine all of the samples which contain your pure product, and then remove the solvent. (How you would separate the solvent from the product isn't directly relevant to this topic and would vary depending on their exact nature - so I'm not even going to attempt a generalisation.)

THIN LAYER CHROMATOGRAPHY

THIN LAYER CHROMATOGRAPHY This page is an introduction to chromatography using thin layer chromatography as an example. Although if you are a beginner you may be more familiar with paper chromatography, thin layer chromatography is equally easy to describe and more straightforward to explain.
	Note:  I'm taking a simple view of the way that thin layer chromatography works in terms of adsorption (see below) which should be adequate for students doing courses for 16 - 18 year olds. The reality is more complicated and the explanation will vary depending on what sort of solvent or solvent mixture you are using. Some similar problems are discussed on the page about paper chromatography, but I am unwilling to do the same thing on this page which is intended as a fairly gentle introduction to chromatography.
Carrying out thin layer chromatography Background Chromatography is used to separate mixtures of substances into their components. All forms of chromatography work on the same principle. They all have a stationary phase (a solid, or a liquid supported on a solid) and a mobile phase (a liquid or a gas). The mobile phase flows through the stationary phase and carries the components of the mixture with it. Different components travel at different rates. We'll look at the reasons for this further down the page. Thin layer chromatography is done exactly as it says - using a thin, uniform layer of silica gel or alumina coated onto a piece of glass, metal or rigid plastic. The silica gel (or the alumina) is the stationary phase. The stationary phase for thin layer chromatography also often contains a substance which fluoresces in UV light - for reasons you will see later. The mobile phase is a suitable liquid solvent or mixture of solvents. Producing the chromatogram We'll start with a very simple case - just trying to show that a particular dye is in fact a mixture of simpler dyes.
	Note:  The chromatography plate will in fact be pure white - not pale grey. I'm forced to show it as off-white because of the way I construct the diagrams. Anything I draw as pure white allows the background colour of the page to show through.
A pencil line is drawn near the bottom of the plate and a small drop of a solution of the dye mixture is placed on it. Any labelling on the plate to show the original position of the drop must also be in pencil. If any of this was done in ink, dyes from the ink would also move as the chromatogram developed. When the spot of mixture is dry, the plate is stood in a shallow layer of solvent in a covered beaker. It is important that the solvent level is below the line with the spot on it. The reason for covering the beaker is to make sure that the atmosphere in the beaker is saturated with solvent vapour. To help this, the beaker is often lined with some filter paper soaked in solvent. Saturating the atmosphere in the beaker with vapour stops the solvent from evaporating as it rises up the plate. As the solvent slowly travels up the plate, the different components of the dye mixture travel at different rates and the mixture is separated into different coloured spots. The diagram shows the plate after the solvent has moved about half way up it. The solvent is allowed to rise until it almost reaches the top of the plate. That will give the maximum separation of the dye components for this particular combination of solvent and stationary phase. Measuring Rf values If all you wanted to know is how many different dyes made up the mixture, you could just stop there. However, measurements are often taken from the plate in order to help identify the compounds present. These measurements are the distance travelled by the solvent, and the distance travelled by individual spots. When the solvent front gets close to the top of the plate, the plate is removed from the beaker and the position of the solvent is marked with another line before it has a chance to evaporate. These measurements are then taken: The Rf value for each dye is then worked out using the formula: For example, if the red component travelled 1.7 cm from the base line while the solvent had travelled 5.0 cm, then the Rf value for the red dye is: If you could repeat this experiment under exactly the same conditions, then the Rf values for each dye would always be the same. For example, the Rf value for the red dye would always be 0.34. However, if anything changes (the temperature, the exact composition of the solvent, and so on), that is no longer true. You have to bear this in mind if you want to use this technique to identify a particular dye. We'll look at how you can use thin layer chromatography for analysis further down the page. What if the substances you are interested in are colourless? There are two simple ways of getting around this problem. Using fluorescence You may remember that I mentioned that the stationary phase on a thin layer plate often has a substance added to it which will fluoresce when exposed to UV light. That means that if you shine UV light on it, it will glow. That glow is masked at the position where the spots are on the final chromatogram - even if those spots are invisible to the eye. That means that if you shine UV light on the plate, it will all glow apart from where the spots are. The spots show up as darker patches. While the UV is still shining on the plate, you obviously have to mark the positions of the spots by drawing a pencil circle around them. As soon as you switch off the UV source, the spots will disappear again. Showing the spots up chemically In some cases, it may be possible to make the spots visible by reacting them with something which produces a coloured product. A good example of this is in chromatograms produced from amino acid mixtures. The chromatogram is allowed to dry and is then sprayed with a solution of ninhydrin. Ninhydrin reacts with amino acids to give coloured compounds, mainly brown or purple. In another method, the chromatogram is again allowed to dry and then placed in an enclosed container (such as another beaker covered with a watch glass) along with a few iodine crystals. The iodine vapour in the container may either react with the spots on the chromatogram, or simply stick more to the spots than to the rest of the plate. Either way, the substances you are interested in may show up as brownish spots. Using thin layer chromatography to identify compounds Suppose you had a mixture of amino acids and wanted to find out which particular amino acids the mixture contained. For simplicity we'll assume that you know the mixture can only possibly contain five of the common amino acids. A small drop of the mixture is placed on the base line of the thin layer plate, and similar small spots of the known amino acids are placed alongside it. The plate is then stood in a suitable solvent and left to develop as before. In the diagram, the mixture is M, and the known amino acids are labelled 1 to 5. The left-hand diagram shows the plate after the solvent front has almost reached the top. The spots are still invisible. The second diagram shows what it might look like after spraying with ninhydrin. There is no need to measure the Rf values because you can easily compare the spots in the mixture with those of the known amino acids - both from their positions and their colours. In this example, the mixture contains the amino acids labelled as 1, 4 and 5. And what if the mixture contained amino acids other than the ones we have used for comparison? There would be spots in the mixture which didn't match those from the known amino acids. You would have to re-run the experiment using other amino acids for comparison. How does thin layer chromatography work? The stationary phase - silica gel Silica gel is a form of silicon dioxide (silica). The silicon atoms are joined via oxygen atoms in a giant covalent structure. However, at the surface of the silica gel, the silicon atoms are attached to -OH groups.
	Note:  If you aren't sure about it, you will find one possible structure of silicon dioxide towards the bottom of the page you will get to by following this link. Use the BACK button on your browser to return quickly to this page.
So, at the surface of the silica gel you have Si-O-H bonds instead of Si-O-Si bonds. The diagram shows a small part of the silica surface. The surface of the silica gel is very polar and, because of the -OH groups, can form hydrogen bonds with suitable compounds around it as well as van der Waals dispersion forces and dipole-dipole attractions.
	Note:  If you aren't sure about hydrogen bonds and van der Waals forces follow this link to the page about hydrogen bonding. You will find a further link to van der Waals forces at the bottom of that page. Use the BACK button on your browser to return quickly to this page.
The other commonly used stationary phase is alumina - aluminium oxide. The aluminium atoms on the surface of this also have -OH groups attached. Anything we say about silica gel therefore applies equally to alumina. What separates the compounds as a chromatogram develops? As the solvent begins to soak up the plate, it first dissolves the compounds in the spot that you have put on the base line. The compounds present will then tend to get carried up the chromatography plate as the solvent continues to move upwards. How fast the compounds get carried up the plate depends on two things: How soluble the compound is in the solvent. This will depend on how much attraction there is between the molecules of the compound and those of the solvent. How much the compound sticks to the stationary phase - the silica gel, for example. This will depend on how much attraction there is between the molecules of the compound and the silica gel. Suppose the original spot contained two compounds - one of which can form hydrogen bonds, and one of which can only take part in weaker van der Waals interactions. The one which can hydrogen bond will stick to the surface of the silica gel more firmly than the other one. We say that one is adsorbed more strongly than the other. Adsorption is the name given to one substance forming some sort of bonds to the surface of another one. Adsorption isn't permanent - there is a constant movement of a molecule between being adsorbed onto the silica gel surface and going back into solution in the solvent. Obviously the compound can only travel up the plate during the time that it is dissolved in the solvent. While it is adsorbed on the silica gel, it is temporarily stopped - the solvent is moving on without it. That means that the more strongly a compound is adsorbed, the less distance it can travel up the plate. In the example we started with, the compound which can hydrogen bond will adsorb more strongly than the one dependent on van der Waals interactions, and so won't travel so far up the plate. What if both components of the mixture can hydrogen bond? It is very unlikely that both will hydrogen bond to exactly the same extent, and be soluble in the solvent to exactly the same extent. It isn't just the attraction of the compound for the silica gel which matters. Attractions between the compound and the solvent are also important - they will affect how easily the compound is pulled back into solution away from the surface of the silica. However, it may be that the compounds don't separate out very well when you make the chromatogram. In that case, changing the solvent may well help - including perhaps changing the pH of the solvent. This is to some extent just a matter of trial and error - if one solvent or solvent mixture doesn't work very well, you try another one. (Or, more likely, given the level you are probably working at, someone else has already done all the hard work for you, and you just use the solvent mixture you are given and everything will work perfectly!)

What is the principle involved in chromatography

Chromatography (from Greek χρώμα: chroma, colour) is the collective term for a family of laboratory techniques for the separation of mixtures. It involves passing a mixture through a stationary phase, which separates the analyte to be measured from other molecules in the mixture and allows it to be isolated.

Chromatography involves a sample (or sample extract) being dissolved in a mobile phase (which may be a gas, a liquid or a supercritical fluid). The mobile phase is then forced through an immobile, immiscible stationary phase. The phases are chosen such that components of the sample have differing solubilities in each phase. A component which is quite soluble in the stationary phase will take longer to travel through it than a component which is not very soluble in the stationary phase but very soluble in the mobile phase. As a result of these differences in mobilities, sample components will become separated from each other as they travel through the stationary phase.

Techniques such as H.P.L.C. (High Performance Liquid Chromatography) and G.C. (Gas Chromatography) use columns - narrow tubes packed with stationary phase, through which the mobile phase is forced. The sample is transported through the column by continuous addition of mobile phase. This process is called elution. The average rate at which an analyte moves through the column is determined by the time it spends in the mobile phase.

Principles and Applications of Liquid Chromatography-Mass Spectrometry in Clinical Biochemistry

Liquid chromatography-mass spectrometry (LC-MS) is now a routine technique with the development of electrospray ionisation (ESI) providing a simple and robust interface. It can be applied to a wide range of biological molecules and the use of tandem MS and stable isotope internal standards allows highly sensitive and accurate assays to be developed although some method optimisation is required to minimise ion suppression effects. Fast scanning speeds allow a high degree of multiplexing and many compounds can be measured in a single analytical run. With the development of more affordable and reliable instruments, LC-MS is starting to play an important role in several areas of clinical biochemistry and compete with conventional liquid chromatography and other techniques such as immunoassay.

Introduction

Coupling of MS to chromatographic techniques has always been desirable due to the sensitive and highly specific nature of MS compared to other chromatographic detectors. The coupling of gas chromatography to MS (GC-MS) was achieved in the 1950s with commercial instruments available from the 1970s. Relatively cheap and reliable GC-MS systems are now a feature of many clinical biochemistry laboratories and are indispensable in several areas where the analysis of complex mixtures and unambiguous identification is required e.g. screening urine samples for inborn errors of metabolism or drugs. The coupling of MS with LC (LC-MS) was an obvious extension but progress in this area was limited for many years due to the relative incompatibility of existing MS ion sources with a continuous liquid stream. Several interfaces were developed but they were cumbersome to use and unreliable, so uptake by clinical laboratories was very limited. This situation changed with the development of the electrospray ion source by Fenn in the 1980s.

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Destroy user interface control1 Manufacturers rapidly developed instruments equipped with electrospray sources, which had a great impact on protein and peptide biochemistry. Fenn was awarded the Nobel Prize in 2002 with Koichi Tanaka who developed matrix assisted laser desorption ionisation, another extremely useful MS ionisation technique for the analysis of biological molecules.

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Destroy user interface control2

By the mid 1990s, the price and performance of LC-MS instruments had improved to the extent that clinical biochemistry laboratories were able to take advantage of the new technology. Biochemical genetics was one of the first areas to do so, and the analysis of neonatal dried blood spot samples for a range of inborn errors of metabolism was a major early application.

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Destroy user interface control3 There are a number of other clinical applications of LC-MS, and the technique is more generally applicable than GC-MS owing to the broader range of biological molecules that can be analysed and the greater use of LC separations in clinical laboratories. The reasons for choosing LC-MS over LC with conventional detectors are essentially the same as with GC-MS, namely high specificity and the ability to handle complex mixtures. Applications of electrospray MS were reviewed in The Clinical Biochemist Reviews in 2003.

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Destroy user interface control4 The current review focuses on the principles of LC-MS, practical considerations in setting up LC-MS assays and reviews some of the major applications in clinical biochemistry, concentrating on small molecule applications.

Mass Spectrometry Instrumentation

Mass spectrometers operate by converting the analyte molecules to a charged (ionised) state, with subsequent analysis of the ions and any fragment ions that are produced during the ionisation process, on the basis of their mass to charge ratio (m/z). Several different technologies are available for both ionisation and ion analysis, resulting in many different types of mass spectrometers with different combinations of these two processes. In practice, some configurations are far more versatile than others and the following descriptions focus on the major types of ion sources and mass analysers likely to be used in LC-MS systems within clinical laboratories.

Ion Sources

Electrospray Ionisation Source

Fenn developed ESI into a robust ion source capable of interfacing to LC and demonstrated its application to a number of important classes of biological molecules.

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Destroy user interface control1 ESI works well with moderately polar molecules and is thus well suited to the analysis of many metabolites, xenobiotics and peptides. Liquid samples are pumped through a metal capillary maintained at 3 to 5 kV and nebulised at the tip of the capillary to form a fine spray of charged droplets. The capillary is usually orthogonal to, or off-axis from, the entrance to the mass spectrometer in order to minimise contamination. The droplets are rapidly evaporated by the application of heat and dry nitrogen, and the residual electrical charge on the droplets is transferred to the analytes.

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Destroy user interface control5 The ionised analytes are then transferred into the high vacuum of the mass spectrometer via a series of small apertures and focusing voltages. The ion source and subsequent ion optics can be operated to detect positive or negative ions, and switching between these two modes within an analytical run can be performed.

Under normal conditions, ESI is considered a “soft” ionisation source, meaning that relatively little energy is imparted to the analyte, and hence little fragmentation occurs. This is in contrast to other MS ion sources, for example the electron impact source commonly used in GC-MS, which causes extensive fragmentation. It is possible to increase ESI “in-source” fragmentation by increasing voltages within the source to increase collisions with nitrogen molecules. This has been used in LC-MS analyses to identify components with common structural features e.g. the glycans in glycopeptides can be fragmented in-source to give 204 m/z reporter ions. This feature has been used to identify glycopeptides in tryptic digests of proteins in order to characterise the structure of the glycans.

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Destroy user interface control6 Although useful for some analytes, in-source fragmentation is limited for others, and more consistent fragmentation methods, such as collision induced dissociation (see below), are required to induce extensive fragmentation required for structural studies and tandem MS.

Small molecules (≈ <500 Da) with a single functional group capable of carrying electrical charge give predominantly singly charged ions. This can involve the addition of a proton to the analyte (M+H⁺) when the ion source is operated in positive ion mode or the loss of a proton (M-H⁻) when operated in negative ion mode. Adduction of cations (e.g. M+NH₄⁺, M+Na⁺, M+K⁺) and anions (e.g. M+formate⁻, M+acetate⁻) can occur when salts are present. Larger molecules and molecules with several charge-carrying functional groups such as proteins and peptides can exhibit multiple charging, resulting in ions such as M+2H²⁺, M+3H³⁺ etc. For proteins, this results in an envelope of ions with different charge states. This property can be used to accurately determine analytes with high molecular weights including proteins up to 100 kDa on mass spectrometers that scan up to only 4000 m/z. Indeed, it is unusual to detect ions with m/z values above this.

While ESI is the most widely used ion source for biological molecules, neutral and low polarity molecules such as lipids may not be efficiently ionised by this method. Two alternative ionisation methods developed for such analytes are described below.

Atmospheric Pressure Chemical Ionisation Source

In atmospheric pressure chemical ionisation (APCI), as with ESI, liquid is pumped through a capillary and nebulised at the tip. A corona discharge takes place near the tip of the capillary, initially ionising gas and solvent molecules present in the ion source. These ions then react with the analyte and ionise it via charge transfer. The technique is useful for small, thermally stable molecules that are not well ionised by ESI.

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