Chemical Shift Referencing and Temperature Calibration.

Version11/27/00 – by Steve Hardies, from notes taken from Andy Hinck.

This document covers temperature and chemical shift calibration experiments that should be done in conjunction with protein spectra.

One is to take the spectrum of ethylene glycol, in which there are two peaks whose separation is highly temperature sensitive.  This measurement is used to compute the true temperature of the sample for comparison to the thermistor reading.  It reveals that there is a slow drift over time in the temperature control system reflected by a drifting difference between the reported temperature and the true temperature.  This is thought to be due to fluctuations in the air cooling system used to control sampe temperature.

One should take this reading about once every two weeks to be updated on the bias of the reported temperature.  You can then set the temperature for each protein determination so that the true temperature is constant and well determined across all of your experiments.  As of this writing, the thermistor is high by 1.6 degrees at 300K.  This much differential between two experiments may be sufficient to cause misidentification of specific resonnances.  As an example, the way I used this information was as follows.

I set the temperature at 301.6K for my protein determination (corresponding to a true temperature of 300K).  I then set the TE parameter to 300 to record the true temperature in the dataset parameter file.  I also used the settitle command to record that the temperature was 300.00 (true) and 301.6K (set) based on ethylene glycol calibration <date> in the title file.  This should alleviate any confusion about the temperature during subsequent examination of this dataset.  The thermistor bias may be different at other temperatures.  The thermistor can be physically recalibrated, but this is done sparingly because of the logistical problems of making sure every user understands the consequences of each adjustment to their experiments.

The second procedure specified below is to take the spectrum of 2,2-dimethylsilapentane-5-sulfonic acid (DSS) at a given temperature.  The purpose is to find the center position offsets (for hydrogen, nitrogen, and carbon) to incorporate during processing of protein spectra.  The hydrogen offset is measured directly.  The others are calculated indirectly from the hydrogen result.  This will cause your reported chemical shifts to be correctly scaled.

Note that DSS is the primary standard used in protein work, but a different but related standard (tetramethylsilane (TMS)) is the IUPAC standard for proton NMR.  To distinguish the difference, chemical shifts calibrated against DSS are denoted as dDSS or d(DSS) (that’s a lower case delta).  See John L. Markley et al., 1998. “Recommendations for the presentation of NMR structures of proteins and nucleic acids”. Pure  Appl. Chem., Vol 70 117-142.

Note that the cited recommendation is to include DSS at low concentration in the protein sample as an internal control, whereas the procedure below is to obtain the DSS offsets using an external standard.  The chemical shift of the water resonance relative to the DSS standard is temperature dependent by about -0.01 degrees C, and should always be the same at at given temperature.   An internal standard would also correct for salt and buffer effects, which are not expected to be very large under normal circumstances.

Temperature calibration with ethylene glycol.

A sample of 100% ethylene glycol in a  wilmad tube (round bottom NMR tube) is found in that flask full of standards at the operator’s table.

  • Use edte to set the temperature given best available information to achieve the true temperature at which you wish to operate.
  • Load the sample and leave some time for equilibration.
  • Turn SWEEP off.  There is no deuterium upon which to operate a lock, and the frequency shifts that are part of the lock operation will interfere with this spectrum.
  • Shim the sample by maximizing integrated fid. This methods does not require the deuterium lock signal.  It may also be a good method for improving higher order shims on proteins samples, sincethe integrated area of the fid may be more sensitive to improvements in these shims than is the lock signal.
    • Read (rsh) a suitable wilmad shim file, eg. aph_dss_wilmad.
    • If you haven’t already set up a specialized dataset for the dss calibration, do so now by selecting calib1h and using edc to save it by a different name, eg.
    • Make it issue a rectangular pulse of 2 dB and 7 usec.
    • Make TD (total data points) to 8K. This high value will increase the resolution by taking more data points.
      • The exact settings for the 90 degree pulse width and the shimming are not particularly critical.
    • Use gs and acqu to display a continuously acquired fid.
    • Noting the integrated fid value displayed in the info window, adjust the shims in the usual order to maximize this value.
    • Type stopto end the continuous acquisitions when your are through shimming.
  • Tune and match the proton probe as usual.
  • Measure the chemical shift between the two peaks in the ethylene glycol spectrum.
    • Take the spectrum (zg), fourier transform (ef), and phase.
    • Set the right peak to 0.
      • To zoom on the peak, left click to associate cursor with the spectrum (the cursor will seem to stick to the spectrum). Then middle click to the left and the right of the peak to zoom in on it.
      • Click on <calibrate> and put the cursor at the peak tip and middle click. This will open a box into which to enter a 0.
    • Zoom on the left peak in the same way, and position the cursor at the peak top.  Read the chemical shift from the information box.
  • For 100% ethylene glycol, y = 4.5677 – 0.0097723 x, where x is temp. in Kelvin. and y is the chemical shift.
    • In a separate shell type “bc -l” to get a calculator.
  • If necessary, reset the temperature to get the desired true temperature given the measured bias. Leave some time for equilibration.
  • Take several readings with some time in between to be sure that the sample is thermally equilibrated.  The thermistor comes to equilibrium before the sample does.

DSS standard.

A DSS sample in deuterated water is found in a wilmad tube in the flask of standards on the operator’s table.

  • Set the temperature and load the sample.
  • Starting from a suitable shim files, eg. aph_dss_wilmad, shim using the lock signal as usual.
  • Tune and match the proton probe as usual (remember to turn SWEEP back on).
  • If you haven’t already, make a specialized version of calib1h with edc (eg. and set it to issue a retangular pulse at 2 usec and 2 dB; set TD to 8K.
  • Take spectrum (zg), transform (ef), and phase.
    • The spectrum will have two tall peaks with 3 short peaks in between.  The one on the left is deuterated water (ie. [1]H in HDO); the one on the right is the relevant DSS resonance.  Ignore the short peaks.
  • Zoom on the rightmost DSS peak. Click <utilities> and <O1>.  Position the cursor on the peak top and read the absolute frequency from the information window.  For 300 K, it would be 500.1312532.  Be careful not to middle click, else you will change a carrier frequency setting in the active dataset.
  • Use <calibrate> to set this peak to zero, and then get the chemical shift to deuterated water as above.  For 300 K (true), it would be 4.7396.  This is the value to use for the center position shift of hydrogen.
  • To get center positions for nitrogen or carbon, multiply the absolute DSS frequency by the relative chemical shift constant for that nuclei to get the extrapolated reference frequency.  The relative chemical shift constants are given in the Markley et al. paper cited above and are based on choosing reference values (ppm = 0) for the other nuclei at a constant proportion to the hydrogen DSS standard under all conditions.  The nitrogen reference is 0.10132918 times the hydrogen reference.  The carbon standard is 0.251449530 time the hydrogen reference.
    • For nitrogen: Look in the pulse program and see on what channel nitrogen.  For hsqc_fb.ref, it is channel f3.  The carrier frequency will be found in parameter SFO3 = 50.683840.  Take the difference between the carrier and the nitrogen reference frequency and divide by the  reference frequency to get the center position for nitrogen. So for [15]N, the reference frequency is 500.1312532 * 0.101329118 =  50.677859.  The center position offset is (50.683840-50.677859)/50.677859 =118.05 ppm.
    •  [13]C is calculated similarly except that the carrier frequency may be a little more complicated to identify.
      • In some cases involving [13]C, The frequency is switched among several different frequencies.  If this is the case, there will be a list of frequencies listed specified by the parameter F2LIST.  The pulse program will advance through this list each time it exectues an “O2” command, cycling back to the beginning of the list as necessary.  Only one of these frequencies will be used as the [13]C carrier with respect to evolution of the [13]C  magnetization in te indirect dimension.  The other frequencies are used for decoupling.  It may not be obvious to the casual user which frequency on the frequency list is the correct one to use for the carrier position.  In this case, there should be a comment in the header of the pulse program clarifying which frequency to use.  As a rule of thumb, always read the comments in the pulse program first.
    • For example, suppose that you establish that the [13]C carrier was at = 125.764214.  500.1312532 * 0.251449530 = 125.757769.  (125.764214 – 125.757769)/125.757769 = 50.12 ppm
  • During processing on the NIS system, you can check your pulse program and/or use uxgrep to look in parameter files for information about the appropriate carrier frequencies.
  • You can expect center positions for all nuclei to move about – 0.01 ppm per increase of 1 degree centigrade.
  • Note: shifting towards a lower ppm is said to be shifting “upfield”. Shifting towards higher ppm is said to be shifting “downfield”.