KCl can increase protein solubility. Volume: Each NMR sample should consist of a final volume of ? L, including all additives, for a standard tube e. Wilmad Sodium azide usually mM is added to prevent microbial contamination.
The internal standard 2,2-dimethylsilapentanesulfonic acid DSS can be added typically 50? M to reference the chemical shifts. New NMR tubes are not 'analytically clean' when delivered, but usually have organic or inorganic residues. Ensure tubes are clean a rinse with water or buffer is advisable and not chipped or warped by excessive heat. Shigemi tubes can be used for low volume samples, but require special care to eliminate bubbles. Tubes should be capped and the cap parafilmed to avoid evaporative loss.
Procedure — Operational Notes Use of concentrators: Proteins are often exchanged into their final solution conditions using concentrators with molecular weight cut off filters. The filters are stored with glycerol, which must be removed by at least three washes or spins of the concentrator with deionized water. Strong acids such as Nochromix now supplied under the name AlNochromix by Alconox, Inc are available to remove adhered materials and deposits by overnight soaking, followed by washes with water or buffer.
Long term storage of NMR tubes: We recommend flash freezing proteins in liquid nitrogen and storage at ? C to minimize risks of oxidation or degradation. A small scale freeze and thaw trial experiment is advised to assess the risk of protein precipitation during sample warm up.
Ensure that tubes are matched to the appropriate solvent. Protein solubility: AP Golovanov et al. The inserts are made from two materials, teflon white and Kel-F transparent. Choose the material that best matches the susceptibility of your material. If the sample is highly diamagnetic like water or chloroform then use teflon. The inserts are designed for use with liquid or soft samples. There are three types of caps fig.
The Boron nitride cap is very fragile and virtually unusable. The zircon cap is recommended for extreme temperatures because it is more durable that boron nitride. For solid state except CRAMPS , if there is not enough material, do not use restricted tubes but it is better to dilute the sample with something inorganic such as silica, alum or calcium sulphate. The restricted tubes are designed for soft samples and hard samples may damage them. For solid samples, grind them up with a mortar and pestle fig.
Place the rotor in its stand and put the powder in stages with a small spatula fig. Compress the powder after each stage using the long part of the special tool fig. Fill the rotor up to the cap's place. If the sample is correctly prepared it should spin up to a speed of 15 kHz. Putting the sample in its stand and putting the powder into the rotor.
Compressing the sample with the special tool. Use the long end. For liquids and gels the insert reduces the risk of bubbles that compromise resolution.
If the density of the liquid is known then the rotor, insert and screw can be weighed before and after filling to confirm that there are no bubbles. For soft samples such as biological tissue, the inserts hold the sample exactly in place.
If using an insert, put a drop of sample on the bottom of the insert and if not put a drop under the cap fig. Inserting a liquid sample with an automatic pipette. Place the pellet in the bottom of the rotor. The rotor can be placed in the bottom of an Eppendorf tube and spun in a swing-out centrifuge fig. A centrifuge should also be used to remove bubbles from gels, resins and liquid crystals. A rotor in an Eppedorf tube.
The sample in a centrifuge. Close the centrifuge when in use. For resins and other swelling samples, check the swelling ratio for the solvent typically 5.
Mix with a pin. For food and tissue samples, wash with D 2 O fig. Cut the sample to size with a scalpel figs. In general, we discourage the use of disposable NMR tubes , because the low tolerance of the outer diameter makes them slide around in the sample spinner and can result in more breakage inside the probe.
Use an internal standard. Residual 1 H in deuterated solvents can often be used for spectral calibration calibration table. However, in situations where an exact chemical shift is desired, or there is not solvent available for reference such as for 13 C conducted in D 2 O or 31 P , an additional internal standard must be used for chemical shift calibration. For nuclei other than 13 C or 1 H, additional standards can be used such as phosphoric acid for 31 P. Internal standards can be added directly to the sample if desired.
Add a drop of TMS to mL of deuterated solvent that can be used for several samples. Alternatively, if you are concerned about an internal standard reacting with the compound of interest, a capillary tube can be filled with an internal standard and placed in the NMR tube.
This situation is not appropriate if you are using the internal standard for quantitation purposes. For quantitation, the internal standard must be added directly to the sample in order to achieve the same filling factor in the coil. Indirect referencing can be used in lieu of an internal standard. Label your sample. However, you need to consider such issues as distortions arising from the solvent suppression and chemical shift calibration.
Dissolve your sample in a solvent selected by you. Mix it well and transfer it into an NMR tube. Filter the solution of your sample into the NMR tube if there are undissolved parts left. An appropriate volume is 0. Solvents are likely to contain a small amount of water when purchased and once the solvent container is opened, the water content will increase due to absorption from the atmosphere. You could store your solvents over molecular sieves, but be aware of the particles from the sieves.
The chemical shift of water and other trace impurities in various solvents can be found by searching for "NMR chemical shifts of solvents". Ensure that the polyethylene cap is pushed fully onto the tube avoid cuts and injuries! Seek advice if you do need to use parafilm. A thin strip of parafilm not more than two layers may be wrapped around the joint between the tube and the cap, not lower than 3 mm from the bottom of the cap.
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