NMR Sample Preparation

In nuclear magnetic resonance (NMR), unlike other types of spectroscopy, the quality of the sample has a profound effect on the quality of the resulting spectrum. Follow these few simple rules to ensure that the sample you prepare gives a spectrum in which useful information is not lost or obscured.

Use proper tubes and caps

Please do not use chipped, scratched or cracked NMR tubes. Please use good quality tubes from Wilmad, Norell, or New Era. NMR tubes are available from the chemistry research stockroom. Minimum length is 7". Tubes must be capped. Please use a non-glass secondary container when transporting NMR samples between your lab and the NMR lab.

There are, generally, three tiers of NMR tube quality:

High-throughput. These are the least costly tubes.
Economy tubes. These are more costly than High-throughput tubes, but they have better quality control.
Precision tubes. These are more costly than Economy tubes, but they have the best quality control of all the tiers.

I have been asked a few times lately about economy or high throughput tubes. In my opinion, they are fine for routine work; however, they have some drawbacks:

They typically take longer to shim than a higher quality tube. Also, they may not shim as well as a higher quality tube.
Their outer diameter varies more than that of high quality tubes, sometimes causing spinner fitment problems.
Their thick walls decrease S/N when compared to a better tube.
They must not be used for variable temperature (VT) experiments.
Sample Tube Grade Affects the Resolution has helpful and important information.

High throughput or economy tubes are allowed in the NMR lab. If you use high throughput or economy tubes for routine spectroscopy (no VT!), please be sure that the tube is neither too loose nor too tight in the spinner. If it does not fit well in the spinner, please try a different spinner. If you cannot find a spinner that fits that tube well, discard that tube.

Use the correct quantity of material

For ¹H spectra of organic compounds, the quantity of material required is typically 5 to 25mg. It is possible to obtain spectra from smaller quantities; but, at very low concentrations, more spectrometer time will be required (halving the sample concentration will require 4x the time to acquire data with the same S/N) and peaks from common contaminants such as water and grease tend to dominate the spectrum.

Remove all solid particles

Solid particles distort the magnetic field homogeneity because the magnetic susceptibility of a particle is different from that of the solution. A sample containing suspended particles has a field homogeneity distortion around every single particle. This causes broad lines and indistinct spectra that cannot be corrected. So that there are no solid particles in your samples, you might consider filtering all samples into the NMR tube. This can be done with a small plug of glass wool tightly packed into a Pasteur pipette. If the plug is not tight enough, the filtration will be ineffective. If it is too big, some of your sample will remain trapped in it. Do not use cotton wool: most NMR solvents dissolve material from it, which can easily be seen in ¹H spectra.

Make samples to the correct depth

In the magnet, the main field direction is vertical, along the length of the sample. Each end of the sample causes a major distortion of the field homogeneity, which is corrected using the spectrometer's shim controls. A partial correction is done for every sample (shimming), and takes a few minutes (can take longer--quite sample dependent). For our Bruker spectrometers, the optimum sample filling height is 4 cm, or ~0.55 mL. Shorter samples may be very difficult or impossible to shim, and cause considerable delay in recording the spectrum. Samples that are too long can also be difficult to shim (thermal gradeints can induce convection) and are a waste of costly solvent. You should consider checking your sample depth using a ruler. After preparation, you should ensure that the cap is pushed fully onto the tube to minimize solvent loss through evaporation.

Use deuterated solvents

Samples are typically prepared using solvents that contain deuterium in place of hydrogen. The NMR signal from the deuterium nuclei is used by the spectrometer for stabilization ("Lock"). Many deuterated solvents are available from the stockroom. The NMR lab does not supply you with solvents.

It is possible to acquire NMR data without a deuterated solvent--we call this "No-D" NMR. This can be done reasonbly well in automation on the AX-400--to do so, select "none" as the solvent. When the sample is run, instead of the typical shimming routine, an au program will run that acquires a 1-scan proton, picks the largest signal in the spectrum, shims on that signal, then the spectrometer runs whichever experiment you've selected. This gives reasonable results; however, better No-D results may be achieved manually on the hands-on spectrometers (AM-400 or AV-500).

Specialty NMR tubes

For small sample quantities, you may consider a Shigemi tube or a 3 mm NMR tube. I have Shigemi tubes you can borrow. Tricks for dealing with small sample sizes: "Solvents, NMR Tubes, and Susceptibility Matched Plugs Sets" courtesy of Charlie Fry at the University of Wisconsin-Madison.

For light sensitive samples, amberized NMR tubes may be helpful. I have a small number of these in the lab.

For air/moisture sensitive samples, a valved tube may be necessary. Valved tubes are only allowed on the AM-400 and AV-500. If you want to use a valved tube on the AV-500, it must be short enough to fit on the autosampler, part number: Wilmad 528-LPV-200M. The autosampler can be disabled if longer tubes must be used on the AV-500, but it needs to be coordinated with NMR personnel.

Washing NMR tubes

After use, NMR tubes should be washed by rinsing with acetone or some other suitable solvent. NMR tubes are ideally dried while lying flat in a vacuum oven at a low temperature. If a vacuum oven isn't available, please try drying your tubes with a blast of dry air or nitrogen. If you must use a hot oven, please lay the tubes flat and place them in the oven for as short a time as possible. Long periods of high heat can distort NMR tubes.

Label your samples

Labels on your samples are best done with a permanent marker directly on the top of the tube, or on the cap. If you use a sticker or a piece of tape, your label must stick smoothly on the tube. Do not leave a flap.

Use an internal reference (optional)

Usually, a small amount of reference is added to the solvent by the supplier. However, the amount of tetramethylsilane (TMS) or any other reference material that is required for a ¹H spectrum is far less than can be added after the sample has been prepared. One drop of TMS in a sample causes serious problems due to distorted baseline and exceeded dynamic range. Even the standard amount of TMS added to a bottle of CDCl₃ is too much. Think about adding 2-3 mLs of CDCl₃ containing TMS to a bottle that does not contain TMS and then use that bottle for sample preparation. This provides a small TMS signal; you never want your reference signal to be taller than your solvent signal. Alternatively, the residual protons in the deuterated solvent may be used as a secondary reference. For samples in D₂O, DSS or TSP is used as an internal reference. Remember that the chemical shift of water is highly temperature dependent.

Degassing samples

Some samples need to be degassed or have oxygen removed. The most effective way of doing this is by using the freeze-pump-thaw technique, at least three cycles. It is sometimes sufficient to flush the space above the sample surface with nitrogen. This should be done with great care to avoid blowing the solution out of the tube. Do not bubble nitrogen through the solution in an NMR tube. This wastes costly solvent through evaporation, and is not an effective method of removing oxygen.

Modified from Alan S.F. Boyd's document, 4 October, 1995, NMR Services at Heriot-Watt University Chemistry Dept., Edinburgh, Scotland