Frequently Asked Questions

  1. Do I have to use a specific type of NMR tube?
  2. How should I prepare my NMR sample?
  3. My sample has a high ionic strength (> 200 mM). What do I need to know?
  4. I want to quantitate my 1D proton NMR spectrum. What do I need to know?
  5. I want to quantitate my 1D 13C NMR (or other nucleus) spectrum. What do I need to know?
  6. How should I clean my NMR tube?
  7. How do I view and process my data on my lab or personal computer?
  8. How did you get the 800 into the lab?

  1. Do I have to use a specific type of NMR tube?

    Yes, you must use a Wilmad "Precision Grade" tube (e.g. 506-PP) that is in perfect condition: chipped, cracked, or warped tubes are not allowed. Tubes are readily available from Chemistry Stores and the NMR Centre. Users violating this rule will have NMR access suspended, and damage resulting from ignoring this rule may be billed to the users' supervisor.

    Other tubes (e.g. Shigemi, quartz, etc.) may only be used with the explicit permission of the NMR Centre manager.

  2. How should I prepare my NMR sample?

    • Tube: Use only an allowed NMR tube (e.g. Wilmad 506-PP) that is not chipped or cracked.

    • Concentration: 1-2 mg/mL for 1H NMR, and 20-50 mg/mL for 13C NMR. Don't make samples too concentrated, especially when using the 600 MHz cryoprobe. Overconcentrated samples may suffer from "radiation damping" which distorts peakshapes and baselines.

    • Preparation: Dissolve (or mix) your sample in the solvent of choice in a vial, add any desired internal standard, and mix very thoroughly before transferring to the NMR tube. If necessary, filter the sample to remove insoluble particles. (Dissolving/mixing directly in the NMR tube will lead to terrible lineshapes, while insoluble particles can also degrade lineshapes.)

    • Volume: Too little and the lineshape/shims will be poor, but too much and there can be issues/artifacts from sample convection, especially with cryoprobes.
      • When using the cryoprobe, the volume in the tube should be 560 uL.
      • For other probes (e.g. 300 MHz, 600 MHz in MacN), between 600-650 uL is fine.
      • If you are using a volatile solvent (e.g. CDCl3, DCM) and are running long experiments, be mindful of sample evaporation during your experiment.
      • For 3 mm NMR tubes, use 200 uL.

    • Aqueous (H2O) samples: Ensure that there is at least 5-10% D2O in the sample for spectrometer lock.

    • Organic solvent samples: Some solvents (e.g. DMSO and D2O) are more likely to take on H2O from the atmosphere, leading to an intense water peak in the NMR spectrum. To reduce this, prepare a sample using a fresh vial of deuterated solvent and then wrap the sample tube cap with Parafilm.

  3. My sample has a high ionic strength (> 200 mM). What do I need to know?

    High ionic strength samples are problematic, particularly when using the 600 MHz cryoprobe or the 400 MHz Prodigy probe. This is because:
    1. More RF power is required to perform certain NMR experiments, and in some cases the power required exceeds what is safe for the NMR.
    2. Signal intensity is reduced. As an estimate, 100 mM NaCl reduces signal by 50%, while 200 mM reduces signal by 66%.
    3. 13C NMR and any other experiments involving decoupling cannot be performed because the decoupling will actually cause the sample to heat up!

    OK, but I still need to analyze this sample. What are my options?
    1. Use a 3 mm NMR tube to reduce the overall amount of salt in the NMR probe. They are fragile and tough to clean, but they are ideal for mass-limited samples.
    2. Use the 600 MHz in MacN 305, which does not have a cryoprobe.
    3. Reduce the ionic strength of your sample, if possible.

  4. I want to quantitate my 1D proton NMR spectrum. What do I need to know?

    First, you will need to add an internal standard of known concentration to the sample. The ideal standard is one that does not chemically interact with the sample (including interactions due to sample pH), is non-volatile, and has a simple NMR spectrum with at least one quantifiable peak that does not overlap with analyze peaks. For aqueous solutions a common choice is DSS, while organic solvents may require more careful selection.

    Second, you will need to alter the acquisition parameters to ensure the relaxation delay between scans (parameter D1) is long enough to allow the sample to return to equilibrium before collecting each scan. According to NMR theory, D1 should be at least five times the longest T1 relaxation time constant of the peaks that you wish to quantify.

    Our standard parameters/experiments use a short D1 and do not produce quantitative spectra.

    In determining what value of D1 to use, be aware that T1 values depend on the chemical environment, solvent, magnetic field, and other parameters, and so you may not be able to use literature values. Two commonly employed methods are: measure the T1 values for your sample (e.g. using an inversion recovery experiment) or assume T1 values are under 5 seconds (a reasonable assumption for most common 1H chemical environments). In the latter case, you would set D1 to 25 seconds.

  5. I want to quantitate my 1D 13C NMR (or other nucleus) spectrum. What do I need to know?

    First, everything in the above FAQ on quantitation of 1D proton NMR also applies here. For 13C nuclei, T1 values are considerably longer; for example, aromatic carbons can have T1 of > 30 sec, meaning D1 must be a few minutes!

    Second, you will need to change the actual pulse sequence. 13C NMR is low sensitivity, so our standard parameters/experiments boost the signal-to-noise by transferring magnetization from nearby 1H atoms. This boost is non-quantitative as not every 13C chemical environment enjoys the same magnetization transfer. For quantitative 13C NMR one must use a pulse sequence without the boost, which means more NMR time is required (prior to attempting this, please contact NMR Centre staff for the details).

    Given the harsh realities of quantitative 13C NMR, spectra can take a VERY long time to collect. You may want to consider an alternative method of quantification unless your sample is extremely concentrated or it is 13C-labelled.

  6. How should I clean my NMR tube?

    Use an NMR tube washer to rinse the tube with at least three volumes of the same (non-deuterated) solvent. Then, rinse one final time with a low boiling point solvent (e.g. acetone). Dry the tubes by laying them flat in a 70 C oven. Avoid higher temperatures which can warp the tubes (warpage of even a few micrometers can cause breakage inside the NMR probe).

    For biomolecular aqueous samples, soak the tubes overnight in 5% HNO3, then rinse with multiple volumes of water until the pH of the rinsate is netural. If a film remains in the NMR tube, let the tubes soak in 1 M NaOH for one hour, then repeat the HNO3 and rinsing steps. Do not use stronger NaOH or longer times or else you will etch the tubes.. Dry the tubes as described above.

    If you would like to minimize the HOD peak in a biomolecular NMR sample (e.g. polysaccharide NMR) it may help to fill the NMR tube with D2O and let it sit for a short while prior to removing the D2O and drying the tube in a 70 C oven.

  7. How do I view and process my data on my lab or personal computer?

    The New Users' Guide describes how you can download your file from the NMR servers and also how to install and use a free, academic version of Topspin 4.0 from Bruker.

  8. How did you get the 800 MHz NMR into the lab?

    A large opening was made in the side of the Science Complex, directly into the nearby AAC staff lounge. Large double-doors were installed in the lounge and the NMR lab, allowing the magnet to be brought in.