E. Anoardo, G. Galli, G. Ferrante, "Fast Field Cycling NMR: Applications and Instrumentation." Appl. Magn. Reson.20:365-404 (2001)
F.A. Bovey, L. Jelinski, P.A. Mirau, Nuclear Magnetic Resonance Spectroscopy, Academic Press, NY, 1988.
Although the questions below are of a general nature, the instruments mentioned are the Varian spectrometers currently used in the NMR Centre of the Australian National University).
Please email your ideas for other questions to be included in this list to Chris Blake.
There is a much more comprehensive Basics of NMR document available at the Center for Imaging Science, Rochester Institute of Technology.
If you haven't used any NMR spectrometers at the NMR Centre before. Come down to room G48 or G42 in the Chemistry Faculties building, and ask Chris or Peta to arrange a training session for you. Even if you have used Varian spectrometers before, our setup here is likely to be slightly different, so in the long run you will save yourself time by having a short lesson. More information is available in the introduction to the NMR Centre. It is NMR Centre policy that you will not be given a license to operate a spectrometer until we assess your competence with Varian spectrometers.
How much solvent volume should I use? To get good resolution you need at least 0.7 ml of solvent in a 5 mm NMR tube. If you have a limited amount of sample you can increase its effective concentration by reducing the solvent volume to 0.4 - 0.5 ml however you will need to spend more time shimming. If you have a sample that will give a proton spectrum in 15 minutes when disolved in 0.7 ml, there is little point reducing sample volume if it means you need to spend an extra 10 minutes shimming! On the other hand if you then want to run a carbon spectrum of that sample it would certainly be worth reducing the sample volume. For 13C HMQC or HMBC runs with very small amounts of material, a 4mm tube and rotor is available.
What does the "ADC overflow" error message mean? The signal recieved from the NMR sample is first amplified by the reciever and then digitised by the analog to digital converter (ADC). If the signal is too strong for them to handle, either the receiver or ADC will "overflow", causing a RECEIVER OVERFLOW or ADC OVERFLOW message to be displayed. The acquired FID is likely to be clipped, resulting in a distorted spectrum. The solution is to use autogain (type gain='n' or on a Gemini GAIN=N) or to type in a lower value for the receiver gain. If overflow still occurs when the gain is set to zero, reduce the observe pulsewidth (PW) to half its present value. If overflow still occurs dilute your sample, or if the solvent signal is causing the ADC overflow use a solvent suppression technique.
How do I shim / tune the spectrometer? First of all, let's get our terminology straight. Shimming is adjusting the resolution of the signal by optimizing the homogeneity of the magnetic field. Tuning is adjusting the impedance of the probe. A poorly tuned probe reflects a lot of the power of the pulses, so that what should be a 90 degree pulse is in reality only (say) a 50 degree pulse. Probe tuning does not affect the resolution, however the signal to noise of a standard spectrum will be worse. Also, experiments such as DEPT or COSY that rely on accurate 90 degree pulses may produce artefacts or not work at all. (Note however that it is possible to adjust the pulse width to give a 90 degree pulse on a poorly tuned probe). Poor shimming on the other hand, results in broad NMR resonances. People often talk about "tuning the resolution" which is where some confusion between shimming and probe tuning arises. Shimming is adjusting the homogeneity of the magnetic field, so that every part of the sample in the NMR tube experiences exactly the same field strength.
OK, so how do I tune the probe? If you're using the broadband Gemini or Mercury spectrometers, you never need to tune the probe. The probes of these intruments are tuned at the factory, and further tuning is a specialised operation. For best results, you should tune all other spectrometers before acquiring a spectrum. Frequency, solvent and sample height all affect probe tuning. If you were running a set of similar samples in the same solvent, you might only bother to tune the probe before running the first spectrum. If however, half your samples were disolved in chloroform and half in D2O, you might run all of the chloroform samples and then quickly adjust the tuning after inserting the first D2O sample. Tuning involves setting up for the nucleus of interest and minimizing the reflected power shown on the meter on the magnet leg. Some recabling is required. Do not attempt to do this unless an NMR staff member has given you a lesson. This doesn't mean that Geminis have some great "automatic tuning" technology. It just means they are left in a state of tune that is good enough for the run-of-the-mill experiments they were designed for. On other spectrometers, tuning is necessary because
You can get the best possible tuning for your sample,
You may not know what nucleus the previous user left the probe tuned to, or whether he/she completely messed up the tuning,
More sophisticated experiments such as HMQC, HMBC etc. work best when the probe is tuned and short 90 degree pulses are required.
How do I tune for carbon or phosphorus? As mentioned above, if you're using a Gemini spectrometer, you don't need to tune the probe. First, check whether the probe you are using requires a tuning stick to be inserted. Tuning sticks are kept separate from the probe, and have a small capacitor on the end to change the tuning range of the probe. If a tuning stick is required, select the stick for the observe frequency and screw it gently all the way into the probe. You can find the observe frequency by setting up for the nucleus of interest and reading the value of sfrq from the dg display. Then make the cable connections for tuning, and adjust both the tuning and matching rods. These two tuning rods affect each other, so it is usually necessary to go back and forth between them to get a good minimum. There is a bit of a knack to it, so persevere! (Hint: make the tuning worse with one rod, then better with the other. Each dual operation should result in better tuning than before).
Also note that if you are decoupling protons while observing carbon or phosphorus, it is a good idea to check the proton tuning. If the probe is poorly tuned to protons, some decoupler power may be reflected, resulting in an improperly decoupled spectrum. On the Inova spectrometers you can tune the observe and decoupler channels at the same time. On older spectrometers you need to set up for and tune protons, then set up for and tune carbon or phosphorus. (Hint: always tune the highest frequency first and the lowest frequency last).
Which spectrometer should I use for carbon? When measuring carbon spectra, the main concern is usually signal to noise. You would expect higher field spectrometers to have a decisive advantage - for example a 500 Mhz spectrometer when compared to a 300 MHz spectrometer should have an advantage of (5/3) squared, or 2.8 times the signal to noise. However there are other considerations, including for example the type of probe. An indirect detection probe has the proton observe coil on the inside (that is, closer to the sample than the coil used for carbon). This improves the proton signal to noise, however if you use an indirect detection probe for directly observing carbon, the signal to noise will of course be worse than a standard probe which has the carbon coil on the inside. Regardless of the probe design, carbon and protons use different coils, and since the electronic circuit for the two nuclei is different it makes no sense to compare proton signal to noise on two instruments and extrapolate the results to carbon.
Also, signal to noise tests are usually performed by collecting a single scan on a concentrated sample, however this does not give the best indication of the results obtainable on "real" samples where the sample is scanned for several hours. When a sample is repeatedly pulsed, the relaxation times of the various carbons must be taken into consideration. Nuclei take longer to relax at higher fields, so the gain in signal to noise is less than expected. Also note that carbons that do not have directly bonded protons (i.e. carbonyls and quaternaries) have much longer relaxation times than protonated carbons.
In order to see how some of the spectrometers compare under "real life" conditions, a dilute sample was run for 256 scans on the Inova 500 (PFG indirect detection probe), broadband Gemini, and VXR300. A D1 delay of 1 second with a 45 degree pulse was used, and 16 dummy pulses were given to bring the system to a steady state before starting acquisition. The signal to noise ratios of three resonances were then measured.
It can be seen that there is not a large difference in the signal to noise you can expect to see on these instruments. Also remember that
if there is not much sample available, you should reduce the amount of solvent. (See How much solvent volume should I use?) A 4mm tube and rotor is available. This will allow you to use even less solvent than is necessary in a 5mm tube.
if you are interested in quarternary carbons, a longer D1 delay of 3 seconds or more is advisable.
If the signal to noise of your carbon spectrum is too low, try running a short and/or long range proton-carbon 2D correlation experiment. It has been known for a long time that this can give dramatic improvements in S/N. See J. Am. Chem. Soc. 101, 4481 - 4484 (1979).
Why are some of the peaks in my APT missing? The APT experiment relies as much on the size of the 1JCH coupling as the number of attached protons to generate the spectral pattern. This is because the delays in the experiment are matched to the inverse of the size of 1JCH. If 1JCH is much larger than the default 1JCH of the experiment (usually set to 140 Hz which is the average of 1JCH for sp3 and sp2 carbons) then peaks will either disappear or appear with incorrect phase. Carbons that may show this behaviour are terminal ethynyl groups (1JCH = 250 Hz approx.), epoxide carbons (1JCH = 175 Hz), furan, pyrone and isoflavone carbons (1JCH = 200 Hz), 2-unsubstituted pyridine and pyrolle carbons (1JCH = 180 Hz) and 2-unsubstituted imidazole and pyrimidine carbons (1JCH > 200 Hz).
I can't lock on.
You are using a deuterated solvent aren't you?
Can you see a lock signal? If not, make sure the lock is turned off, turn the lock power and lock gain to their maximum values, and look for a sine wave by adjusting Z0. If you find a sine wave, adjust Z0 until its frequency becomes zero. Then reduce the lock power (to avoid saturating the lock) and try to lock on.
If it loses lock as soon as you try to lock on, turn the lock off and adjust the lock phase as shown in the manual.
Is your tube spinning? It might not be spinning because you inserted the tube too quickly, causing it to break. Take the tube out and check that it is in one piece. While you have it out, use a depth gauge to check that the sample is centred in the probe.
It won't shim.
Check the linewidth of the narrowest line in your spectrum. If there are some broad lines and some narrow lines, the broad lines are probably broad because they are undergoing chemical exchange, not because the resolution is poor. Broad lines may also be caused by quadrupolar broadening if your compound has a transition metal.
If you have not already done so, load the standard shims. You don't know what sort of state the previous user left the shims in! All spectrometers in the NMR Centre have a macro rtss which loads the standard shims. This macro is equivalent to typing rts('stdshm') su. On the Inova 300 and 500, check that the probe parameter is set to the probe you are using, since the rtss macro uses this value to determine which shims to load.
If it still won't shim, take the tube out and inspect your sample. Is the tube scratched? Is there anything floating in the sample? Is the sample centred in the coil? If you are using a small amount of solvent to improve the concentration, you may need to add some more solvent to make it easier to shim.
Do you have paramagnetic ions in your sample?
Have other people been getting poor resolution? If so, report it to a member of the NMR staff. If not, change NMR tubes, filter your sample, and try again. If changing tubes solves the problem, throw the old tube away.
Have you placed your NMR tube in an oven to dry? If so, throw the tube away as it has distorted. (Remember, glass is a liquid. It flows at high temperature). The correct way to dry a tube is via a stream of dry nitrogen through a glass wool filter.
My tube broke when I inserted it into the magnet. During the day, phone an NMR staff member. Tell them the solvent and any hazards posed by your compound. After normal working hours tell the watchmen who will call in someone. Leave a note to warn others not to use the spectrometer. Remember - the more quickly you lower a tube into the magnet, the more likely it is to break! If necessary, use two hands on the sample eject button to make it easier to lower the tube slowly.
I can't phase correct my spectrum. The aph (automatic phase correction) command usually does a good job of correcting the phase, and should be the first thing you try. Sometimes (for example in noisy spectra) the aph command is unable to correct the phase, and in these situations it often leaves lp at a high value (say one or two thousand). In these situations you will have to correct the phase manually. First a couple of obvious things: if you ran a DEPT or APT experiment or something similar, there will be some positive and some negative peaks, so don't try and phase them all positive! Similarly in a 1:1 binomial solvent suppression sequence, half the spectrum will be positive and half negative.
Having established that you are not running an exotic pulse sequence that produces strange phases, the next thing to consider is foldback. Are you sure that you used a large enough spectral width when acquiring the spectrum? If one or more resonances occurred outside the observe region, the method used to digitise the signal results in these resonances appearing within the observed spectral width, but with a phase error. If in doubt, double or triple the spectral width, run the spectrum again, and see if the resonance that could not be phase corrected remains at the same chemical shift as before.
To perform manual phase correction, proceed as follows:
Type lp=0 rp=0. This sets the left phase and right phase to zero. On Varian spectrometers, "right phase" and "left phase" equate very roughly to the zero-order and first-order phase adjustments respectively. The zero-order phase affects the entire spectrum equally, while the first-order phase is frequency dependent. The zero-order phase should always be in the range -360° to +360° and the first order phase should also usually be in this range. If you have a first order phase correction of more than a thousand degrees, not only is it probably incorrect, but you will also probably be generating baseline roll. On the Gemini spectrometer, type QP to get into quick phase mode. On Sun based spectrometers or data stations, click the "phase" button with the mouse. Perform a zero-order phase correction on the largest peak as described in the manual for the spectrometer you are using. Now choose another peak some distance from the largest peak, and adjust the first-order phase. On Sun based systems you only get one shot at adjusting the zero-order phase - all subsequent corrections are made to the first-order phase, so there is no point clicking on the largest peak again. If you want to readjust the zero-order phase, get out of the phase-correction routine (by for example typing ds) then click on the phase button again. If for some reason a large first-order phase correction is required, it may be easier to choose a peak for the first-order adjustment that is close to the peak you used for the zero-order adjustment. On Sun based systems, set the phasing parameter to 100. This causes the effect of the phase values to be shown for the entire spectrum as you are making the adjustments, thus making it easier to see what you are doing.
I need to run my spectrum at a higher field to get better resolution. No you don't! The resolution of a high field spectrometer may even be worse than a low field spectrometer. What a high field instrument has more of is dispersion. This means that resonances with different chemical shifts are further apart. Multiplets due to coupling will not show any improvement unless the higher field instrument separates overlapping multiplets with different chemical shifts, or the multiplet showed strong coupling effects at lower field. Some nuclei such as 31P may have worse resolution because of a property called chemical shift anisotropy which increases with field strength.
There are no parameters for the solvent I want to use. If you're running a proton spectrum, set up for 1H / CDCl3, double the spectral width, run a quick spectrum, and put the two cursors around the spectrum. Then do a movesw and acquire the final spectrum. If you're running a carbon spectrum, set up for 13C / CDCl3, increase the spectral width by 20 percent, and run as normal. If your solvent has carbon nuclei which show up quickly, reference the solvent and check that the observed spectral range is correct.
If you are running a phosphorus spectrum, set up for 31P / CDCl3, increase the spectral width by 20 percent, and run as normal. Then supply an NMR tube containing the solvent to the NMR staff so that they can set up H3PO4 referencing parameters for you.
How can I suppress a strong solvent resonance in a proton spectrum? If the solvent signal is less than two or three times the size of the largest signal from your compound, it may not be worth bothering. On the Gemini, the usual method is to presaturate the solvent signal using the decoupler (instructions for doing this are in the folders near the spectrometer). Although it is simple, this method has the disadvantage that NH or OH protons that are exchanging with water also have their signals reduced or eliminated. Another method is the 1:1 binomial pulse sequence. The signals on one side of the solvent resonance are of opposite phase to the other side when this method is used. On the Inova spectrometers, the method of choice if a pulsed field gradient probe is in use, is watergate solvent suppression. Simply set the observe transmitter on the solvent position, type wgate and acquire a spectrum. The watergate sequence set up by the wgate macro uses hard pulses and therefore does not require pulse phases etc. to be optimised. The other watergate technique available uses shaped pulses. Simply type autowatergate, and wait while it automatically optimises the parameters and runs a final spectrum. Watergate is only available on the Inovas because it uses pulsed field gradients. If chemical exchange is very rapid, watergate may not be suitable, in which case a binomial pulse sequence is the best choice.