Sample Preparation
Quantity
In general, more is better. Typical concentrations range from 100 µM to 100 mM, but working outside this range is also possible. Required volumes are 500 µL for 5 mm tubes and 35 µL for 1.7 mm tubes. Keep in mind that the signal-to-noise ratio increases with the square root of the number of scans, so halving your concentration will require an experiment four times as long to achieve the same sensitivity.
Purity
Generally, the purer the better, but this depends on your goals. Confirming the result of a reaction may simply require detecting a characteristic peak, and therefore might not need purification. If you’re trying to determine the structure of an unknown, your sample should be at least 85% pure and elute as a single peak from an LC. A biomolecular sample for 3D structure determination should exhibit a single band on a gel.
Organic solvents
Solvents should be deuterated to minimize the size of their signals and the likelihood of obscuring signals from your sample. If your sample has exchangeable groups that need detection (e.g., -NH, -OH), avoid using protic solvents like D2O and methanol-d4. Acetonitrile-d3 and DMSO-d6 are better options. If possible, steer clear of solvent mixtures, as they are more challenging to lock onto and shim.
Buffers
For biomolecular samples, phosphate buffers are preferred because they introduce no extra signals. However, Tris, HEPES, and similar buffers may be used with isotopically labeled samples since the signals from non-labeled compounds will be filtered out. The pH should be physiological or acidic to slow the exchange of amides with the solvent. Salts are often added, but be aware that high concentrations (greater than 150 mM) can make uniform excitation of the entire frequency range more difficult and may increase sample heating.
Remove insoluble material
Your solution should be clear. Cloudy solutions suggest that the sample hasn’t dissolved properly. Such solutions are more difficult to shim and produce broader peaks. If your sample doesn’t dissolve well, consider filtration or centrifugation to eliminate the insoluble material.
NMR tubes
Use high-quality tubes. Low-grade or disposable tubes will be harder to shim and will produce broader peaks than higher-quality tubes. Wilmad 535-PP-7 or an equivalent are acceptable. If using Shigemi tubes, ensure they are the Bruker versions with a small bottom length of 8 mm. The longer Varian Shigemi tubes can damage Bruker probes.
| Size | Supplier | Model |
|---|---|---|
| 5 mm | Wilmad | 535-PP-7 |
| 5 mm | Kontes | 897241-0000 |
| 5 mm | Norell | 509-UP-7 |
| 5 mm | New Era | NE-UP5-7 |
| 5 mm Shigemi | Sigma-Aldrich | BMS-005TB for water |
| Wilmad | CMS-005TB for chloroform | |
| Bruker | DMS-005TB for DMSO | |
| MMS-005TB for methanol |
Cleaning tubes
Avoid using a brush, as it may scratch the inner surface of the tube, leading to reduced magnetic homogeneity, poorer shimming, and broader lines. Generally, rinsing the tube with acetone, distilled water, and then methanol should suffice. A final rinse with deuterated water will replace protons adsorbed on the glass. Avoid drying tubes in an oven; heat can warp the tubes, making shimming difficult and potentially damaging the probe. It is best to dry tubes under a vacuum.
Volume
Sufficient volume to fill the active detection space must be used. Using less will make the samples difficult to shim and lead to broader peaks. Using too much will dilute your sample and waste solvent.
The table below lists the preferred volume and fill heights of different sized NMR tubes. Please note that the Duke NMR Facility does not have any probes capable of handling 10mm NMR tubes, that 3 mm tubes require a special adaptor to fit a 5mm spinner, and that while 1 mm tubes can be used with the 1.7 mm probe this arrangement is not optimal.
| Tube size | Volume | Fill height |
|---|---|---|
| 1 mm | 5 uL | |
| 1.7 mm | 35 uL | 30 mm |
| 3 mm | 160 uL | 30 mm |
| 5 mm Shigemi | 280 uL | |
| 5 mm | 600 uL | 45 mm |
| 10 mm | 4500 uL | 45 mm |
Filling 1.7 mm NMR tubes
The easiest way is to use a syringe or pipette. Long, fine pipettes can be drawn from a standard glass pipette. Disposable syringes are used by several Duke labs. These have the advantage of not requiring cleaning. Other Duke labs use Hamilton syringes. These can be purchased with an extended narrow gauge needle (22S,112 mm). If reusing a syringe it must be cleaned carefully before reuse. Its also posible to use a pippettor fitted with gel loading tips, but these don’t deliver the solution to the bottom of the tube. Also, if using organic solvents, the solvent may leach plasticizer and other small molecules from the tips into your solvent.
To reduce solvent evaporation from 1.7 mm NMR tubes polyoxymethylene balls can be used to plug the hole in the cap. They can be purchased from Bruker or other suppliers. Plugging the holes with parafilm is not a good idea, as the evaporating solvent extracts paraffin into your sample and onto the spinners.
