NESG Wiki - User contributions [en]

Setting up non-uniformly sampled spectra/NUS guide for Varian

2009-12-15T15:59:05Z

Aringh:

__NORICHEDITOR__
=== Setting up acquisition of non-uniformly sampled data on Varian spectrometers with BioPack ===
__NORICHEDITOR__

This section will give step-by step instructions on how to set up an NUS spectrum with BioPack for subsequent processing with Multidimensional Decomposition (MDD). In addition to this page, please consult the help files of BioPack itself. The recent BioPack versions (as early as May 2008), barring any bugs, have a few built-in options for Non-uniform sampling. See the section on Non-linear sampling according to V. Orekhov. If you have an older version of BioPack, please update to a more recent one.

#In VnmrJ, call a macro to load the desired experiment, such as a NOESY-HSQC. Set up the experiment in a regular fashion, except for the '''ni''' and '''ni2''' parameters.
#Go to the "Acquire" panel. Choose the "Sampling" tab (it used to be called "Digital Filter").
#In there, choose "Sparse (Orekhov) NLS Sampling" in the drop-down menu for the Indirect dimensions columns. See Figure 1 for details.[[Image:BPnusSnap2.jpg|thumb|center|1050px|Figure 1. Setting up Non-linear (non-uniform) sampling for BioPack experiments.]] This will automatically modify the pulse sequence by including the NUS portion. The new sequence will have "_S" at the end. The macro will also set up an additional dialogue for recording of NUS spectrum. The name of the "Sampling" tab will also change.
#*Alternatively (or if you have VNMR6.1C), you have to call the actual macro <tt>BP_NLSinit(3)</tt> from the command line. Here, 3 stands for 3D.
#*You may need to recompile the new pulse sequence by issuing <tt>seqgen(seqfil)</tt> command from the command line.
#You now have to modify the relevant parameters. Pressing "Show Parameters" button  or type <tt>dgnls</tt> and go to "Text Output" window to see them (see Figure 2). [[Image:BPnusSnap4.jpg|frame|Figure 2. Parameters for non-linear (non-uniform) sampling.]]
#*The most crucial ones '''nimax''' and '''nimax2''' define the desired resolution, while '''ni''' and '''ni2''' give the number of FIDs actually recorded. In fact, '''ni''' and '''ni2''' by themselves have no meaning here, rather only their product is relevant. The level of "sparsing" is thus given by the ratio <math>\frac{ni\cdot ni2}{nimax\cdot nimax2}</math>.
#*Two more imporatnt parameters may have to be adjusted: '''T2sp''' and '''T2sp2''', which govern the exponential bias of the corresponding dimension. Their values have to be only approximate. In the Arrowsmith lab we use 35ms for 1H, 20ms for 13C and 50ms for 15N, respectively. Other parameters are only important for R-MDD (recursive MDD), which is not yet implemented in the processing GUI, described [[Processing non-uniformly sampled spectra with Multidimensional Decomposition|elsewhere on this site]]. Please refer to the description in the BioPack help for further details.
#After the parameters are set, press the "Set Sampling Schedule" button or run the corresponding macro from the command line <tt>BP_NLSset</tt>. At this point two files should be created: "*.in" and "*.hdr_3". They contain the parameters and the actual schedule and '''MUST''' be saved together with the '''fid''' and '''propar''' files.
#*'''NB!''' if you change any of the '''nimax''', '''nimax2''', '''ni''', '''ni2''', '''T2sp''', '''T2sp2''' parameters, you have to run the schedule generator once again, otherwise you will not be able to process the data properly.
#Run the spectrum by typing <tt>go</tt>.
#In order to save all relevant files afterwards, use '''BPsvf''' macro, instead of <tt>svf</tt>.

[[Category:Multidimensional_Decomposition]] [[Category:Sparse]] [[Category:BioPack]] [[Category:Varian]] [[Category:Data_acquisition]] [[Category:Non-uniform_sampling]]

Conditioning procedure for cryogenic probes

2009-12-15T15:56:38Z

Aringh:

The information below is taken from '''Decoupling Noise''' in [http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199919600c.pdf HCN Cold Probe manual]

== '''Cryogenic Probe Conditioning''' ==

NMR experiments that require X-nucleus decoupling perform best when the rf coils are conditioned. Conditioning imparts just enough energy into the rf coils to disperse any extraneous condensed material into the vacuum space so that the material is carried away by the pump.

Use the following procedures to condition the rf coils if the Cold Probe has been left idle for a number of days or thermal cycled. Both procedures (run in the order presented) are required during the initial installation of the probe.

#Install the probe in the magnet.
#Start cryogenic operations, refer to the Cryogenic Systems Operation and Installation manual.
#Determine the approximate 90º pulse width and powers for all channels. Refer to the contract for pulse widths and use the procedures in Testing Probe NMR Performance, page 19.
#Take the sample out of the probe.
#Proton Channel
##Enter <tt>s2pul temp=25 tn='H1' pw=200 d1=0.1 at=0.1 nt=3000 dp='y'</tt>
##Set <tt>tpwr</tt> to the normal high-power level
##Run experiment with <tt>go</tt>
#X Nucleus Channels - Do this procedure for both X Nucleus Channels.
##Enter <tt>pwxcal</tt>, at the prompts select decoupler channel and nucleus.
##For channel 2 (13C) enter <tt>pw=0 pwx1=200 pwx2=0 temp=25 at=0.1 d1=0.1 nt=3000 dp='y'</tt> For channel 3 (15N) enter <tt>pw=0 pwx1=0 pwx2=200 temp=25 at=0.1 d1=0.1 nt=3000 dp='y'</tt>
##Set <tt>dpwr</tt> or <tt>dpwr2</tt> to the corresponding high-power level.
##Run experiment with <tt>go</tt>
#When completed, replace the sample in the magnet. Either monitor the real data as it comes in or set up the experiment but run the first increment a number of times using <tt>array('nt',200,1,0)</tt> to see if the conditioning has been successful.

== '''cryo_noisetest Macro Procedure (Lower Power, Longer Time)''' ==

This is an rf decoupling coil conditioning procedure that also quantifies the noise profile of a Cold Probe. The macro runs forever by cycling between periods of prolonged pulsing and testing.

#Remove the sample from the magnet.
#Ensure that the probe parameter is set to a valid probefile name.
#Enter <tt>cryo_noisetest</tt>. Enter the number of minutes that the decoupler coil conditioning will run before another quantitative test is done. Add five minutes (for the quantitative testing) to the number you entered to get a total recycle time. For example, 40 will run the conditioning for forty minutes followed by five minutes of tests and then start over. Results are printed out at the end of the tests. Reduce the number for more testing (per unit time), increase it for more.
#Enter desired number (use 40 during the initial installation of the probe). The procedure starts and a quantitative test is done straight away as a baseline measurement. Four mini-tests are done consecutively; two carbon and two nitrogen, if more than 2 channels are present. For each nucleus CW and WALTZ decoupling modes are selected. An array of <tt>dpwr</tt> and <tt>dpwr2</tt> is done for each nucleus and for each decoupling mode. The noise is estimated from each spectrum and saved in two date-stamped text files in the local users <tt>vnmrsys/data/testlib</tt> directory and also plotted out in graphical form.
#Enter <tt>aa</tt> or <tt>halt</tt> to stop the acquisition(s) and procedure.

-- Main.AlexEletski - 07 Mar 2008

Full probefile calibration

2009-12-15T15:53:42Z

Aringh:

== Full Probefile Calibration ==

This procedure is similar to [[NESG:AutomatedNMRParametersOptimization|Automated Pulse Width Calibration]], but the number of parameters to be optimized is much larger an the required time is much longer. Full calibration is usually needed only after hardware upgrades, e.g. probe or amplifiers.

#Insert a concentrated NC-labeled sample of a small protein, preferably in a Varian shigemi with proper filling height. Perform tuning/matching, lock and shim the sample.
#Load the <tt>ghn_co</tt> dataset and set the desired parameters - type <tt>man('BioPacklist')</tt> for a list.
#To start Autocalibration go to '''Setup''' -> '''Calibrations''' and select '''Full, using probefile values''' or '''Full, But Use Current 1H Offset and pw'''

 -- Main.AlexEletski - 14 Mar 2008

Installing and updating BioPack

2009-12-15T15:53:02Z

Aringh:

== '''Installing And Updating BioPack''' ==

You have the options of either performing a fresh install or an incremental update.

Incremental update is quicker, but it requires that you have the latest "base release" of BioPack installed

A fresh install is required if you have an earlier version of "base release" or you have just re-installed VNMRJ

=== '''Fresh Installation''' ===

#Go to the [http://www.varianinc.com/nav/products/nmr/apps/usergroup/toc/ul_image/psglib/BioPack&cid=KHPPJIIHFI&zsb=1206824864.usergroup BioPack download page]. Follow the installation instruction therein for a system-wide install. You will normally have to install the latest <tt>loadbiopack</tt> macro first.
#The new BioPack installation must be activated with a new probe file. The process is usually automatic if there is no user probefile (in <tt>~/vnmrsys/probes/</tt>). Otherwise you need to manually activate BioPack with the new probe file by calling <tt>BPbiopack1a('y')</tt>.
#Customize the new probefile according to the printed instructions. You can also view the instructions by typing <tt>man('BioPacklist')</tt>.
#Perform the full probefile calibration, if necessary.

=== '''Incremental Update''' ===

-- Main.AlexEletski - 31 Mar 2008

Simultaneous 13C,15N-resolved NOESY

2009-12-15T15:51:23Z

Aringh:

== Simultaneous 15N-, 13C(ali)-, 13C(aro)-resolved [1H,1H] NOESY ==

Modified from BioPack <tt>gnoesyCNhsqc.c</tt> by Main.AlexEletski, similar to the previous <tt>cnnoesy.c</tt> sequences by Youlin Xia and H. Atreya.

Requires pre-installed BioPack. BioPack power limits should be enabled on systems with cryogenic probes, otherwise wurst140 decoupling power is too high.

Make sure that 13C wurst decoupling power is 1 dB lower than the max rated decoupling power for the probe (wurst shape average is 80% of the max). 15N garp decoupling power must be 3 dB lower than the max rating.

*[[NESG:%ATTACHURL%/gnoesyCaliCaroNhsqc.tar|gnoesyCaliCaroNhsqc.tar]]: simNOESY package - tested with VnmrJ 2.1B

 Contents:

*vnmrsys/psglib/gnoesyCaliCaroNhsqc.c
*vnmrsys/parlib/gnoesyCaliCaroNhsqc.par/
*vnmrsys/manual/gnoesyCaliCaroNhsqc
*vnmrsys/templates/layout/gnoesyCaliCaroNhsqc/
*vnmrsys/maclib/gnoesyCaliCaroNhsqc

Changes as compared to <tt>gnoesyCNhsqc</tt> and <tt>cnnoesy</tt>:

*sw2N is the spectral width for 15N. Should be different from sw2 (13C)
*13C/15N t1 initial evolution is half-dwell only. The program will abort if <tt>f2180</tt> is not set to <tt>'y'</tt>. For testing purposes the initial evolution delay is set to a minimum if <tt>ni2=0</tt>.
*Only adiabatic 13C inversion pulses are used. You can choose either <tt>stC140</tt> or <tt>stC200</tt>. Composite inversion pulses are disabled.
*Simultaneous high power pulses on 13C and 15N are avoided, therefore, peak power can be used.
*Indirect 1H evolution modified to get correct initial points. Backward linear prediction is no longer required.
*15N inversion pulse changed to 90-180-90 composite. This should improve broadband performance at high fields, increasing the intensity of HE/NE strips of Arg, for example.
*<tt>jnh</tt> parameter is introduced for 1J_NH coupling constant. The INEPT delays are now determined by <tt>jnh</tt> instead of <tt>jch</tt>.
*<tt>jch</tt> is used to shift the 13C inversion pulses. The typical values are <tt>jnh=110 jch=155</tt>.
*Added a <tt>flipN</tt> flag. Setting <tt>flipN='y'</tt> will invert the sign of 15N peaks.
*Phase cycling is optimized for better axial peak and water suppression.
*SEDUCE decoupling on carbonyls is not used, since it yield only small sensitivity gains while leading to significant resonance shifts.

Installation:

*Save the <tt>gnoesyCaliCaroNhsqc.tar</tt> file into VnmrJ user home directory (e.g. <tt>/home/vnmr1</tt>).
*Extract with <tt>tar xvf gnoesyCaliCaroNhsqc.tar</tt> - existing files will be overwritten!
*<tt>cd</tt> to <tt>~/vnmrsys/psglib/</tt> directory and compile the pulse program by typing <tt>seqgen gnoesyCaliCaroNhsqc.c</tt>

Usage:

*In VnmrJ run <tt>gnoesyCaliCaroNhsqc</tt> script to setup experiment
*Record a 2D H-C/N plane first and optimize <tt>sw2</tt>, <tt>sw2N</tt>, <tt>dof</tt> and <tt>dof2</tt>, if necessary. Run <tt>calfa</tt> to find the optimal <tt>rof2</tt> and <tt>alfa</tt>, acheiving zero first-order phase correction.
*Set the desired mixing time, <tt>mix</tt>.
*Use at least <tt>nt=4</tt> for 3D acquisition, for best results use <tt>nt=8</tt>. The first two-step phase cycle suppresses axial peaks in the 13C/15N dimension, which is needed since 13C dimension is usually folded 3 times. The second two-step phase cycle suppresses axial peaks in the indirect 1H dimension. The third two-step phase cycle provides additional water suppression.

 

-- Main.AlexEletski - 26 Oct 2007

Estimation of measurement time

2009-12-15T15:47:39Z

Aringh:

The minimum measurement time of (4,3)D GFT experiments can be reliably predicted from the S/N distribution of a 2D [15N, 1H] HSQC and the rotational correlation time tau_c. First, you need to generate an integrated peaklist.

==== '''Peak Integration in 2D [15N, 1H] HSQC''' ====

Having processed 2D (15N, 1H) HSQC as described in data process, one can do the peak picking and integration of 2D (15N, 1H) HSQC manually or semi-automatically as following:

*Peak picking and integration by using program XEASY

##Use command <tt>ns</tt> to load the spectrum
##Use <tt>ls</tt> to load the corresponding sequence file (optional)
##Use <tt>in</tt> to automatic pick peaks with total peak number slightly more than expected by select adjust contour level; then manually remove side-chain amide peaks
##Use <tt>mn</tt> to measure noise level, and use <tt>tw</tt> to display the noise level value. Normally the value of 2.5 times of standard deviation ( ~250 if the noise level has been normalized ) is taken as noise level.
##Use <tt>ip</tt> to choose peak height (m) as integration mode, and use "ii" to integrate the whole spectrum. An [[NESG:XEASY|XEASY]] external program "PeakintI" can be used to obtain more accurate peak height values, which is described below.
##Use <tt>wp</tt> to save the peak list.

*More accurate peak height measurement by using program [[NESG:PeakintI|PeakintI]]. Type command: <tt>peakintI ../data/NHsqc6001 nhsqcsa_b.peaks 250 -i -t 2 2 0.1 </tt> where the <tt>nhsqcsa_b.peaks</tt> is the input peak list and the output file will be <tt>inhsqcsa_b.peaks</tt>.
*Combing atom name informaiton in the peak list for S/N distribution (optional). 
**by using [[NESG:UBNMR|UBNMR]], please check [[NESG:UBNMR|UBNMR]] macro
**by using script '''sim''', run macro '''comb_yang''' by typing <tt>sim comb_yang</tt>

==== '''S/N distribution analysis for 2D (15N, 1H) HSQC''' ====

The SN distribution of resonances in a NMR spectra can be fit to the Gaussian distribution:

[[Image:Sn nhsqc.jpg]]

 <tt>f= a*exp(-0.5*((ln(SN_i)-ln(SN_0))/b)^2) </tt> where <tt>SN_0</tt> is the most populated S/N observed, <tt>f is the expected population at a certain SN value SN_i</tt>, <tt>a and b</tt> are constants.

The SN of NHSQC <tt>SN_0</tt> can be obtained as following:

#Calculate <tt>ln(SN)</tt> for each peak from the peak height and noise level, e.g. by using EXCEL
#Obtain SN distribution (<tt>population v.s. ls(SN)</tt>) by using Sigma-Plot
#By using Sigma-Plot, fitting SN distribution to Gaussian distribution and obtain <tt>SN_0</tt>, constant <tt>a and b</tt>.

==== '''Calculation of Measurement Time''' ====

The SN distribution of resonances in other NMR spectra can also be fit to the Gaussian distribution as in 2D (15N, 1H) NHSQC: <tt>f= a*exp(-0.5*((ln(SN_i)-ln(SN_0))/b)^2)</tt> where <tt>SN_0</tt> is the most populated SN observed, <tt>f</tt> is the expected population at a certain SN value <tt>SN_i</tt>, <tt>a</tt> and <tt>b</tt> are constants. Based on this equation, one can calculate the expected SN_0 for a required peak detection yield.

Assuming that a peak shall have at least SN value of 2 in order to be observed or detected, and the average b for (4,3) GFT experiments is 0.8; if 95% peak detection yield is required, the expected SN_0 is: <tt>SN_0=exp(ln2+1.644*b)=7.4 </tt> 

[[Image:Sn2.jpg]]

NMR measurement time of (4,3) GFT HNNCABCA, (4,3)D GFT CABCAcoNHN and (4,3)D HABCABCONHN can be calculated from the following equation: <tt>T_43d= ((SN_43d* Tauc^2)/(SN_2d*A))^2 </tt> where <tt>T_43d</tt> is the required time for (4,3) GFT experiment, <tt>SN_43d</tt> is the expected SN value of (4,3)D GFT experiments, <tt>SN_2d</tt> is the SN per hour of 2D (15N, 1H) HSQC. =A= is constant, which has value of 0.8639 for (4,3) GFT HNNCABCA, 1.6019 for (4,3)D GFT CABCAcoNHN and 1.0153 for (4,3)D HABCABCONHN.

[[Image:Sn3.jpg]]

One can use [[NESG:UBNMR|UBNMR]] to run the measurement time prediction by the following command: <tt>predict T2d SN2d Tc</tt> where

*<tt>T2d</tt> is the acquisition time of 2D [15N,1H] HSQC in hours
*<tt>SN2d</tt> is the SN distribution average of 2D [15N,1H] HSQC
*<tt>Tc</tt> is the rotational correlation time of protein in nanoseconds.

 

-- AlexEletski - 03 Mar 2008

Measuring 15N T1 and T2 relaxation times (Varian)

2009-12-15T15:41:44Z

Aringh:

== '''Correlation Time and NMR Measurement Time Prediction''' ==

=== '''Experimental Setup''' ===

Run the <tt>HTP_t1t2_setup</tt> script to setup the following experiments:

*2D [15N, 1H] HSQC, ~1.5h for SN analysis
**<tt>NHonly='n' C13refoc='y' ni=256 f1180='n' phase=1,2 nt=8 ss=32</tt>
*1D 15N T1, ~30 min
**<tt>ni=1 phase=1 T1='y' f1180='n' NHonly='n' C13refoc='n' ss=256 nt=128</tt>
**<tt>relaxT=0.1, 0.2, 0.3, 0.4, 0.7, 1.0, 1.5, 2.0</tt>
*1D 15N T2, ~30 min
**<tt>ni=1 phase=1 T2='y' f1180='n' NHonly='n' C13refoc='n' ss=256 nt=128</tt>
**<tt>relaxT=0.01,0.03,0.05,0.07,0.09,0.11,0.13,0.15,0.17 maxrelaxT=0.17</tt>

 Key issues:

*For most proteins it is recommended to run T1 and T2 experiments for at least 30 min each to achieve adequate S/N on a room-temperature probe. Short measurement times may lead to underestimated tc values. With cryogenic probes or very concentrated (> 1 mM) samples the minimum measurement time may be smaller.
*Short <tt>d1</tt> delays (~1 s) may lead to incorrect integral for the first T1 point. However, long d1 values may lead to a large residual water line and non-uniform baseline, especially on a cryoprobe. See what works best for the particular sample and spectrometer.
*<tt>relaxT</tt> must be given as a multiple of 10 ms for T1 and '''odd''' multiple of 10 ms for T2.
*<tt>maxrelaxT</tt> is not used in T1 measurements.
*Avoid sampling T2 points beyond 250 ms - it may cause excessive sample heating.
*Intermediate tc values (between monomer and dimer) may indicate transient dimerization. Dilution studies are then required.
*The tc value is calculated under assumption of isotropic tumbling. For example, if a protein consists of long parallel a-helices the reported tc will indicate a larger molecular weight.

=== '''Correlation Time Measurement''' ===

Based on the Stokes's law, the isotropic rotational correlation time for approximately spherical globular proteins is a function of the effective hydrodynamic radius of the protein, which provides a simple way to check the oligomerizaiton status of protein in solution. Here is the basic law: the correlation time in nanoseconds is approximately half of the value of the protein's molecular weight in kilodaltons.

A protein's rotational correlation time (Tauc) can be quickly estimated from average 15N T1 and T2 relaxation times.

 To calculate Tauc:

*Process the data in Buffalo.VNMR with <tt>wft</tt> and adjust the phase for pure absorption.
*Invoke <tt>dc</tt> to correct baseline shift and slope. You can expand the spectrum for better results.
*Invoke <tt>dscale</tt> and <tt>setref</tt> to make sure the ppm scale is correct - required for the <tt>tc</tt> macro.
*Display all spectra with <tt>dssh</tt> and check that
**baseline is flat and uniform across points.
**Spectral intensity follows an exponential decay.
*Run <tt>tc([T1exp, T2exp, [ppm1, ppm2]])</tt>, where the first two arguments are T1 and T2 experiment numbers, and the last two optional arguments specify the integration range in ppm. By default, integration is performed over the range between 10.5 ppm and 8.5 ppm to exclude signals from side-chain CONH2 groups (6 - 7 ppm) and unfolded parts (8 ppm).

 

==== '''Tc macro''' ====

The [[NESG:%ATTACHURL%/tc|tc]] macro is invoked by typing <tt>tc([T1exp, T2exp, [ppm1, ppm2]])</tt> at the [[NESG:VnmR|VNMR]] prompt. <tt>T1exp</tt> and <tt>T2exp</tt> are the experiment numbers of T1 and T2 experiments, respectively. As an option, the integration range in ppm can be specified with the third and fourth arguments. The default integration range is chosen between 10.5 and 8.5 ppm to avoid the resonances from side chain -CONH2 groups and backbone amides from unfolded regions. If called with no arguments [[NESG:%ATTACHURL%/tc|tc]] will prompt for experiment numbers and use the default integration range.

Tc macro requires <tt>t1a</tt>, <tt>t2a</tt> and <tt>intav</tt> macros. The <tt>intav</tt> macro performs integration stores the integral values in the <tt>fp.out</tt> file. <tt>t1a</tt> and <tt>t2a</tt> macros extract T1 and T2 relaxation times from the exponential fitting to the data in <tt>fp.out</tt>. Manual phase and drift correction (<tt>dc</tt>) of 1D spectra should be performed first to get accurate results.

τ_c is then calculated as

τ_c = sqrt(6 * (T1 / T2) - 7) / (2 ω(N)),

where ω(N) is the 15N frequency. This formula is derived from equation (8) in [http://pubs.acs.org/cgi-bin/archive.cgi/bichaw/1989/28/i23/pdf/bi00449a003.pdf Kay, Torchia and Bax, Biochemistry '''1989''', 28, p8972-8979] by keeping only J(0) and J(ω(N)) terms and neglecting all higher frequencies. The complete equation (8) cannot be solved analytically for τ_c, and the simplified formula presented here should yield accurate results for systems with τ_c >> 0.5 ns.

The reported error range is from simple error propagation of the exponential fit error and is rather crude. Strictly speaking, such simple error propagation does not apply here, because the expected error distribution for τ_c is asymmetric.

To install tc macro:

#Download [[Media:Tc_macro.tar|tc_macro.tar]] into the user maclib directory (<tt>~/vnmrsys/maclib/</tt>)
#Unpack it with <tt>tar xvf tc_macro.tar</tt>

=== '''Measurement Time Prediction''' ===

The minimum measurement time of (4,3)D GFT experiments can be reliably predicted from the S/N distribution of a 2D [15N, 1H] HSQC and the rotational correlation time tau_c. First, you need to generate an integrated peaklist.

==== '''Peak Integration in 2D [15N, 1H] HSQC''' ====

Having processed 2D (15N, 1H) HSQC as described in data process, one can do the peak picking and integration of 2D (15N, 1H) HSQC manually or semi-automatically as following:

*Peak picking and integration by using program XEASY

##Use command <tt>ns</tt> to load the spectrum
##Use <tt>ls</tt> to load the corresponding sequence file (optional)
##Use <tt>in</tt> to automatic pick peaks with total peak number slightly more than expected by select adjust contour level; then manually remove side-chain amide peaks
##Use <tt>mn</tt> to measure noise level, and use <tt>tw</tt> to display the noise level value. Normally the value of 2.5 times of standard deviation ( ~250 if the noise level has been normalized ) is taken as noise level.
##Use <tt>ip</tt> to choose peak height (m) as integration mode, and use "ii" to integrate the whole spectrum. An [[NESG:XEASY|XEASY]] external program "PeakintI" can be used to obtain more accurate peak height values, which is described below.
##Use <tt>wp</tt> to save the peak list.

*More accurate peak height measurement by using program [[NESG:PeakintI|PeakintI]]. Type command: <tt>peakintI ../data/NHsqc6001 nhsqcsa_b.peaks 250 -i -t 2 2 0.1 </tt> where the <tt>nhsqcsa_b.peaks</tt> is the input peak list and the output file will be <tt>inhsqcsa_b.peaks</tt>.
*Combing atom name informaiton in the peak list for S/N distribution (optional). 
**by using [[NESG:UBNMR|UBNMR]], please check [[NESG:UBNMR|UBNMR]] macro
**by using script '''sim''', run macro '''comb_yang''' by typing <tt>sim comb_yang</tt>

==== '''S/N distribution analysis for 2D (15N, 1H) HSQC''' ====

The SN distribution of resonances in a NMR spectra can be fit to the Gaussian distribution:

[[Image:Sn nhsqc.jpg]]

 <tt>f= a*exp(-0.5*((ln(SN_i)-ln(SN_0))/b)^2) </tt> where <tt>SN_0</tt> is the most populated S/N observed, <tt>f is the expected population at a certern SN value SN_i</tt>, <tt>a and b</tt> are contants.

The SN of NHSQC <tt>SN_0</tt> can be obtained as following:

#Calculate <tt>ln(SN)</tt> for each peak from the peak height and noise level, e.g. by using EXCEL
#Obtain SN distribution (<tt>population v.s. ls(SN)</tt>) by using Sigma-Plot
#By using Sigma-Plot, fitting SN distribution to Gaussian distribution and obtain <tt>SN_0</tt>, constant <tt>a and b</tt>.

==== '''Calculation of Measurement Time''' ====

The SN distribution of resonances in other NMR spectra can also be fit to the Gaussian distribution as in 2D (15N, 1H) NHSQC: <tt>f= a*exp(-0.5*((ln(SN_i)-ln(SN_0))/b)^2)</tt> where <tt>SN_0</tt> is the most populated SN observed, <tt>f</tt> is the expected population at a certain SN value <tt>SN_i</tt>, <tt>a</tt> and <tt>b</tt> are constants. Based on this equation, one can calculate the expected SN_0 for a required peak detection yield.

Assuming that a peak shall have at least SN value of 2 in order to be observed or detected, and the average b for (4,3) GFT experiments is 0.8; if 95% peak detection yield is required, the exptected SN_0 is: <tt>SN_0=exp(ln2+1.644*b)=7.4 </tt> 

[[Image:Sn2.jpg]]

NMR measurement time of (4,3) GFT HNNCABCA, (4,3)D GFT CABCAcoNHN and (4,3)D HABCABCONHN can be calculated from the following equation: <tt>T_43d= ((SN_43d* Tauc^2)/(SN_2d*A))^2 </tt> where <tt>T_43d</tt> is the required time for (4,3) GFT experiment, <tt>SN_43d</tt> is the expected SN value of (4,3)D GFT experiments, <tt>SN_2d</tt> is the SN per hour of 2D (15N, 1H) HSQC. =A= is constant, which has value of 0.8639 for (4,3) GFT HNNCABCA, 1.6019 for (4,3)D GFT CABCAcoNHN and 1.0153 for (4,3)D HABCABCONHN.

[[Image:Sn3.jpg]]

One can use [[NESG:UBNMR|UBNMR]] to run the measurement time predition by the following command: <tt>predict T2d SN2d Tc</tt> where

*<tt>T2d</tt> is the acquisition time of 2D [15N,1H] HSQC in hours
*<tt>SN2d</tt> is the SN distribution average of 2D [15N,1H] HSQC
*<tt>Tc</tt> is the rotational correlation time of protein in nanoseconds.

 

-- AlexEletski - 03 Mar 2008

Long-range 15N-1H correlation experiments for histidine rings

2009-12-14T20:41:10Z

Aringh:

== 2D [15N, 1H] long-range HSQC for His rings ==

=== Varian ===

This pulse sequence correlates aromatic H-C protons of His with ring nitrogens.

Use the <tt>rna_WGgNhsqclr</tt> setup macro from BioPack. This macro loads the <tt>rna_WGgNhsqc.c</tt> pulsesequece with parameters for long-range correlations.

=== Varian (Old) ===

This section is left for historic reference.

Use the <tt>gNfhsqc_His.c</tt> pulse sequence. Modified from BioPack <tt>gNfhsqc.c</tt> by adding 90-180-90 composite 180 pulses on 15N and changing decoupling options.

*JNH=22.75 Hz, to refocus 1J_NH couplings
*15N carrier at 200 ppm
*15N spectral width is 200 ppm
*13C carrier at 125 ppm
*15N decoupling during acquisition is off
*13C 40 ppm WURST decoupling during acquisition

*[[NESG:%ATTACHURL%/gNfhsqc His.tar|gNfhsqc_His.tar]]: long range hsqc package - tested with VnmrJ 2.1B

Contents:

*vnmrsys/psglib/gNfhsqc_His.c
*vnmrsys/templates/layout/gNfhsqc_His/
*vnmrsys/maclib/gNfhsqc_His

Installation:

*Save the <tt>gNfhsqc_His.tar</tt> file into VnmrJ user home directory (e.g. <tt>/home/vnmr1</tt>).
*Extract with <tt>tar xvf gNfhsqc_His.tar</tt> - existing files will be overwritten!

Reference: [http://www.pubmedcentral.nih.gov/articlerender.fcgi?artid=2142369 J. G. Pelton, D. A. Torchia, N. D. Meadow and S. Roseman, ''Protein Sci'' 1993 2: 543-558]

-- Main.AlexEletski - 29 Oct 2007

Simultaneous 13C,15N-resolved NOESY

2009-12-14T20:39:32Z

Aringh:

== Simultaneous 15N-, 13C(ali)-, 13C(aro)-resolved [1H,1H] NOESY ==

Modified from BioPack <tt>gnoesyCNhsqc.c</tt> by Main.AlexEletski, similar to the previous <tt>cnnoesy.c</tt> sequences by Youlin Xia and H. Atreya.

Requires pre-installed BioPack. BioPack power limits should be enabled on systems with cryogenic probes, otherwise wurst140 decoupling power is too high.

Make sure that 13C wurst decoupling power is 1 dB lower than the max rated decoupling power for the probe (wurst shape average is 80% of the max). 15N garp decoupling power must be 3 dB lower than the max rating.

*[[NESG:%ATTACHURL%/gnoesyCaliCaroNhsqc.tar|gnoesyCaliCaroNhsqc.tar]]: simNOESY package - tested with VnmrJ 2.1B

 Contents:

*vnmrsys/psglib/gnoesyCaliCaroNhsqc.c
*vnmrsys/parlib/gnoesyCaliCaroNhsqc.par/
*vnmrsys/manual/gnoesyCaliCaroNhsqc
*vnmrsys/templates/layout/gnoesyCaliCaroNhsqc/
*vnmrsys/maclib/gnoesyCaliCaroNhsqc

Changes as compared to <tt>gnoesyCNhsqc</tt> and <tt>cnnoesy</tt>:

*sw2N is the spectral width for 15N. Should be different from sw2 (13C)
*13C/15N t1 initial evolution is half-dwell only. The program will abort if <tt>f2180</tt> is not set to <tt>'y'</tt>. For testing purposes the intial evolution delay is set to a minimum if <tt>ni2=0</tt>.
*Only adiabatic 13C inversion pulses are used. You can choose either <tt>stC140</tt> or <tt>stC200</tt>. Composite inversion pulses are disabled.
*Simultaneous high power pulses on 13C and 15N are avoided, therefore, peak power can be used.
*Indirect 1H evolution modified to get correct initial points. Backward linear prediction is no longer required.
*15N inversion pulse changed to 90-180-90 composite. This should improve broadband performance at high fields, increasing the intensity of HE/NE strips of Arg, for example.
*<tt>jnh</tt> parameter is introduced for 1J_NH coupling constant. The INEPT delays are now determined by <tt>jnh</tt> instead of <tt>jch</tt>.
*<tt>jch</tt> is used to shift the 13C inversion pulses. The typical values are <tt>jnh=110 jch=155</tt>.
*Added a <tt>flipN</tt> flag. Setting <tt>flipN='y'</tt> will invert the sign of 15N peaks.
*Phase cycling is optimized for better axial peak and water suppression.
*SEDUCE decoupling on carbonyls is not used, since it yield only small sensitivity gains while leading to significant resonance shifts.

Installation:

*Save the <tt>gnoesyCaliCaroNhsqc.tar</tt> file into VnmrJ user home directory (e.g. <tt>/home/vnmr1</tt>).
*Extract with <tt>tar xvf gnoesyCaliCaroNhsqc.tar</tt> - existing files will be overwritten!
*<tt>cd</tt> to <tt>~/vnmrsys/psglib/</tt> directory and compile the pulse program by typing <tt>seqgen gnoesyCaliCaroNhsqc.c</tt>

Usage:

*In VnmrJ run <tt>gnoesyCaliCaroNhsqc</tt> script to setup experiment
*Record a 2D H-C/N plane first and optimize <tt>sw2</tt>, <tt>sw2N</tt>, <tt>dof</tt> and <tt>dof2</tt>, if necessary. Run <tt>calfa</tt> to find the optimal <tt>rof2</tt> and <tt>alfa</tt>, acheiving zero first-order phase correction.
*Set the desired mixing time, <tt>mix</tt>.
*Use at least <tt>nt=4</tt> for 3D acquisition, for best results use <tt>nt=8</tt>. The first two-step phase cycle suppresses axial peaks in the 13C/15N dimension, which is needed since 13C dimension is usually folded 3 times. The second two-step phase cycle suppresses axial peaks in the indirect 1H dimension. The third two-step phase cycle provides additional water suppression.

 

-- Main.AlexEletski - 26 Oct 2007

Deuterium pulse width calibration and decoupling

2009-12-14T17:48:20Z

Aringh:

The most common application is deuterium decoupling in triple-resonance experiments with deuterated samples. Any time aliphatic 13C magnetization is transverse, you have to decouple it from the attached deuterons. Omitted or improperly calibrated decoupling would cause a sharp sensitivity drop due to scalar relaxation of the second kind (insert reference here).

Proper setup requires tuning the 2H channel and calibrating 2H 90-degree pulse.

== 2H 90° Automatic Calibration ==

See 'Chapter 9.0: '''2H AutoCalibration'''' in the BioPack manual.

The automatic calibration method of BioPack uses the same protein sample and calibrates the pulse directly on the HDO signal. It requires a K5022 relay to be present, which allows pulsing deuterium on the 4th channel, while detecting it. At SUNY Buffalo 600 MHz and 750 MHz instruments don't have this relay, therefore requiring manual calibration.

== 2H 90° Manual Calibration ==

The current protocol is similar to the '''Calibration via Indirect 2H Detection using 13C Observe''' section of Chapter 9.0 '''2H AutoCalibration''' in theBioPack manual. Though you can still use the <tt>ddec_pwxcal</tt> dataset as recommended in the BioPack manual, the method presented here is more accurate and easier to use.

Manual calibration of deuterium 90° pulse requires an ASTM standard test sample, which contains 60% benzene-d6 (C6D6) and 40% dioxane. The chemical shifts of benzene are 7.15 ppm for 2H and 128.0 ppm for 13C (http://www.sdsnmr.com/cs_table.html); 1J_CD = 24 Hz. Note that 2H and 1H shifts are essentially the same and that benzene-d6 contains 13C at natural abundance level.

#Insert the ASTM sample
#Tune the 13C and 2H channels as described on the [[NESG:RFCoilTuningAndMatching#13C_15N_and_2H_Channels_All_Prob|tuning page]].
#Lock the sample on the benzene-d6 signal; '''z0''' will be different from that when locking on D2O!
#Shim the sample as described on the [[NESG:SampleShimming|shimming page]]. You can use 1H gradient shimming on the dioxane resonance; set <tt>d1=10</tt> for best results.

=== 1D Deuterium Spectrum ===

Typical 2H 90-degree pulse width for decoupling is ~280 us. With a pulse that long, it is important to know the exact 2H offset of benzene-d6 to ensure the on-resonance condition. You can use a 1D deuterium spectrum of the test sample to find the resonance. This experiment will also test if you can pulse and detect deuterium on the first channel.

The current protocol is based on the "Directly Observing Deuterium" in the [http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199914100a.pdf Deuterium Decoupling Channel Installation] manual with small modifications. To record a 1D deuterium spectrum do the following:

#Turn lock off and connect the cable from '''J1 PROBE''' port on the 2H decoupler/lock diplexer to the '''J5311 PROBE''' probe port on the broadband preamp with the 2H RF filter in line.
#On 750 MHz instrument with a cryogenic probe you additionally have to:
##Disconnect the 13C cable from the BE188-20-7BB filter on the console side
##Connect this cable through the BE135-35-8BB filter to the '''XMTR J5313''' port on the broadband preamp
#Retrieve parameter set <tt>/vnmr/tests/C13sn</tt>. This will also set <tt>solvent='c6d6'</tt>
#Make the following changes: <tt>numrfch=4 rftype='dddd' tn='H2' tpwr=43 pw=250 dn=''dm='nnn' dn2='' dm2='nnn' dn3=''dm3='nnn' at=1 d1=1 nt=1 ss=2 in='n'''</tt</tt>
#<tt>Set <tt>tof=(7.15-5)*sfrq</tt> to set the carrier closer to the benzene-d6 line
#Acuire the spectrum with <tt>go</tt> and perform FT with a large <tt>fn</tt> value. You should see a signle line.
#Phase the spectrum and center the cursor on the line. Use <tt>movetof</tt> so set the carrier offset on the resonance. Write down the new <tt>tof</tt> value.
#Re-connect all cables to their original positions and activate lock

 

[[Image:C6D6 2H 1D spectrum.png|left|587x329px|1D deuterium spectrum of C6D6]]

 

 Even though you can now calibrate the deuterium 90-degree pulse directly with this experment, it will not be the one you need. In direct observe mode deuterium is on the first channel, while in a triple-resonance experiment it is on the fourth channel, and the RF path goes through different amplifiers.

=== Indirect Calibration - 1D 13C spectrum ===

Configure the spectometer for 13C direct detection:

On 600 MHz (SUNY Buffalo):

#Connect the cable from '''K5006''' port at the back of the box to the '''J5311 PROBE''' probe port on the broadband preamp with the 13C RF filter in line.

On 750 MHz (SUNY Buffalo):

#Locate the cable going from the 13C port on the probe to an RF filter, and disconnect it from the RF filter.
#Get a free cable and an identical filter, and connect it to the '''PROBE J5311''' port on the broadband preamp.
#Get a free cable and connect the '''first''' RF filter to the '''XMTR J5313''' port on the broadband preamp.

==== 13C In-Phase Triplet ====

At this step you will need the <tt>H2_pwxcal</tt> dataset. If you don't have this dataset, install it:

#Downloading [[NESG:%ATTACHURL%/H2 pwxcal.tar.gz|H2_pwxcal.tar.gz]] into your VNMRJ user home directory
#Ucompress and unpack it
#Go to <tt>~/vnmrsys/psglib/</tt> and compile the pulse sequence with <tt>seqgen H2_pwxcal.c</tt>

Before proceeding with deuterium 90 pulse calibration you need to verify that you can pulse and detect C13 as the 1st channel. You will also need do determine the 13C 90-degree pulse width.

#Type <tt>H2_pwxcal</tt> to load the custom dataset.
#Set <tt>anti='n' lkflg='y' dm3='nnn' pwx3=0 rftype='dddd' ampmode='dddp' tof=(128-92)*sfrq sw=10000 at=0.3 d1=5 ss=0 nt=1 fn=64k</tt>
#Set <tt>pw</tt> to the typical <tt>pwC</tt> value; keep the current <tt>tpwr</tt> or set <tt>tpwr</tt> to <tt>pwClvl</tt>, whichever is smaller.
#Run <tt>go</tt> to acquire spectrum
#Disable weighting functions and do FT. You should see an in-phase 1:1:1 triplet split by the 1J_CD coupling.
#Place the cursor on the middle line and type <tt>movetof</tt> to reset <tt>tof</tt>.
#Set <tt>sw=1000</tt> and acquire the spectrum again.
#Using the current dataset calibrate the 13C 90-degree pulse with the 360 method, along the lines described for 1H (1H cal). Adjust <tt>tpwr</tt> so that <tt>pw</tt> is equal or greater then the typical <tt>pwC</tt> pulse.

Keep in mind that 13C relaxation is so strong due to 2H quadrupolar mechanism that the effects on the accuracy of pulse width calibration are even stronger than for 1H.

[[Image:Inphase c6d6.png|frame|left]]

[[Image:C6d6 pw90 c13.png|frame|left]]

 

==== ====

----

==== 13C Anti-Phase Triplet - Indirect Calibration ====

Configure the pulse sequence to generate an anti-phase triplet.

#Set <tt>anti='y' pwx3=0</tt>
#Set <tt>jCD=24</tt> for benzene-d6 - you can measure is from the splitting.
#Set <tt>dof3</tt> to the <tt>tof</tt> value determined from deuterium 1D spectrum
#Run experiment with <tt>go</tt>
#Perform FT - you should see the outer lines in anti-phase and the middle line is missing

The pulse sequence now will applies 90x pulse on 13C followed by a delay tau=1/(4*1J_CD) and another 90x pulse on 13C. The outer lines will evolve during tau to antiphase with respect to 2H. Note that the J-coupling evolution here is "twice as fast" as with two spins 1/2! The second 13C pulse is used to suppress the middle line, which does not evolve under J-coupling.

Now you can calibrate the deuterium pulse width:

#Set <tt>dpwr3=40 ss=4 nt=1</tt>
#Array <tt>pwx3</tt> from 0 to 1000 in steps of 50
#Observe the antiphase triplet decrease linearly and then increase inverted. If you do not see any difference between scans you probably do not have pulsing on deuterium. Type <tt>rftype='dddd' BPsetampmode</tt> to fix it and repeat the experiment.
#The zero crossing point corresponds to the desired 90 pulse width. Adjust <tt>dpwr3</tt> so that <tt>pwx3</tt> is ~300 us and repeat the calibration.

<img alt="antiphase_c6d6.png" src="%ATTACHURL%/antiphase_c6d6.png" />

Fig. 4. Anti-phase triplet with the middle line suppressed.

<img width="834" alt="h2_pulse_calib.png" src="%ATTACHURL%/h2_pulse_calib.png" height="286" />

Fig. 5. Deuterium pulse width calibration. When deuterium pulse with is exactly 90 the outer lines are converted to unobservable MQ coherence.

Setting '''tof''' to the center line is important here, otherwise the triplet may not be phased properly.

Adjust '''pw''' to make the center line disappear.

==== Deuterium Decoupling Test and Probefile Update ====

To test 2H CPD decoupling:

#Set <tt>dmf3=1000000/pwx3</tt>, where pwx3 is the true 90 degree pulse width.
#Set <tt>rftype='dddd' BPsetampmode anti='n' dm3='nnn' dmm3='ccw' pwx3=0 at=0.06 sw=10000 nt=1 ss=0</tt>.
#Acquire and transform the spectrum. The result should be the familiar uncoupled in-phase triplet, though at a lower resolution.
#Set <tt>dm3='nny'</tt> to enable decoupling.
#Acquire and transform the spectrum. The result should be the decoupled single peak.

Here acquisition time is set to 60 ms, because deuterium decoupling should not be used with long duty cycles.

To update the probe file

#Set <tt>dof3=(3-5)*dfrq3</tt> to set default carrier at ~3 ppm
#In the '''Setup''' -> '''Globals&Calibration''' click on the '''H2''' button

Also see the instructions given in <tt>man('BioPacklist')</tt>.

For details on probefile update see [[NESG:BioPackProbefile#Manual_Probefile_Update|Manual Probefile Update]] page.

== Using 2H Decoupling With Deuterated Protein Samples ==

At this point you have calibrated deuterium 90-degree pulse width and updated the probefile.

For every new sample make sure to tune the 2H channel, including the next protein sample following the calibration with ASTM.

Deuterium decoupling parameters are read automatically when you load a <nop>BioPack dataset with a corresponding macro, provided that <tt>numrfch=4</tt> is set. Deuterium decoupling is off by default, however. After you load a dataset you should usually set <tt>dn3='H2' dm3='nyn'</tt> to turn it on. Refer to the pulse sequence manual for detailed instructions.

Make sure you are using the correct offset '''dof3''' for decoupling. The value used with C6D6 corresponds to 7.15 ppm. The default value in the probefile should correspond to ~3 ppm. If you are running an experiment like HNCA you may want to set it to 4.5 ppm.

To test 2H decoupling for a protein sample you may take a concentrated DCN sample and run the first increment of HNCA with constant-time Ca evolution, switching deuterium decoupling on and off. If decoupling is working properly, you should see a significant sensitivity difference.

-- Main.AlexEletski - 13 Jan 2007

Temperature calibration

2009-12-11T17:35:34Z

Aringh:

Temperature control is crucial for NMR experiments. There remains always a small discrepancy between the actual temperature of the sample and the temperature displayed in the monitor. The true sample temperature may depend on the airflow speed and the temperature of the air supply. In particular, discrepancies may be different for different spectrometers. We use temperature calibration from time to time and keep a record in order to calibrate the temperature accurately. The samples used for temperature calibration are methanol (for low temperature range) and ethylene glycol (for high temperature range).

'''IMPORTANT!''' In [http://www3.interscience.wiley.com/cgi-bin/fulltext/113510727/PDFSTART a recent MRC paper] it has been reported that radiation damping can significantly affect the accuracy of temperature calibration. Spectrometers equipped with cryogenic probes are affected the most. Systematic errors of up to 2-3 °C were observed. As a solution, the authors recommend to observe residual proton signals in a sample of highly deuterated methanol (typically 99.8% methanol-d4). At present (Sep 2007), NYSBC employs this method for calibration of its spectrometers. Thomas Szyperski's lab is also adopting this protocol.

== Temperature dependence of 1H HDO chemical shift in D2O ==

The general formula is

δ = 5.060 - 0.0122*T + (2.11 * 10^-5)T^2

In the range 0-50 °C it can be linearized to

δ = 5.051 - 0.0111*T

Here δ is the chemical shift relative to TMS, and T is the temperature in °C

For the reference see http://pubs.acs.org/doi/abs/10.1021/jo971176v

== Using highly deuterated methanol ==

Highly deuterated methanol is not yet provided as a standard sample, but you can order it from any supplier of isotope-labeled chemicals.

Consult you spectrometer's manual on how to prepare a sample. For Varian, a sample of not more that 3 cm in height in a regular tube is sufficient. You may want to avoid using very low sample height as it would affect the shimming.

You can tune/match the proton channel and lock the sample to the methanol signal. Because methanol spectrum has two lines, methyl and hydroxyl, the lock line will have a small modulation.

Gradient shimming on deuterium can always be used. It is possible to use gradient shimming on protons on spectrometers equipped with cryogenic probes. In this case you need to increase the gain and use a longer excitation pulse.

Because the sample is randomly deuterated to high degree, the hydroxyl signal mainly comes from the CD3OH species, while the methyl signal comes from CHD2OD. The methyl signal thus appears a 1:2:3:2:1 quintet due to 2J_HD scalar coupling of about 2 Hz. The chemical shifts of these two groups differ from those of pure CH3OH because of deuterium isotope effects. Therefore, macros for temperature calibration based on pure CH3OH (e.g. <tt>tempcal('m')</tt>) SHOULD NOT be used. The authors provide their own calibration function:

T = -16.7467 * (Δδ)^2 - 52.5130 * (Δδ) + 419.1381

where Δδ is the chemical shift difference between the two lines.

A macro <tt>tempcal_cd3od</tt> has been created to perform temperature calculation according to the above formula.

Since the lines are much more narrow than in pure CH3OH, simply placing the left and right cursors on the peaks is not accurate enough. A better procedure is the following:

#Zoom in on the -OH peak and position the left cursor exactly on the peak maximum.
#Type <tt>r1=cr</tt> to save the current cursor position.
#Zoom in on the CH3 peak and position the left cursor exactly on the peak maximum.
#Type <tt>r2=cr</tt> to save the current cursor position.
#Zoom out.
#type <tt>delta=r1-r2</tt> to calculate the frequency difference in Hz. This will also set the spacing between the left and right cursors.
#type <tt>tempcal_cd3od</tt> to launch the macro.

 

== Using standard samples ==

To verify the temperature do the following:

*Put the standard sample in the magnet and equilibrate it at the desired temperature.
*Record a basic 1D spectrum (use <tt>presat</tt>, <tt>s2pul</tt>, or <tt>water</tt> experiments).
*Select '''Box''' in the menu and set the cursors on 2 large peaks (one hydroxyl and one aliphatic)
*Key in <tt>tempcal('m')</tt> or <tt>tempcal('e')</tt> for methanol or ethylene glycol, respectively.

Important points to consider:

*The standard samples normally contain small volumes of pure methanol or ethylene glycol. There is no lock substance, so 1D is run without lock.
*Lineshapes will be poor, but shimming is not necessary either. Both peaks will have the same shape and it is the frequency difference between them that matters.
*Probe tuning and matching should be skipped. A detuned probe will diminish radiation damping. Use very small tip angle (for example, <tt>pw = 1</tt>) to avoid ADC overflow.
*When using experiments such as <tt>presat</tt> and <tt>water</tt>, turn presaturation off (<tt>satmode='n'</tt>), otherwise the hydroxyl signal may be distorted.
*The following keywords are acceptable arguments of <tt>tempcal</tt>: <tt>'methanol'</tt> or <tt>'m'</tt>; <tt>'glycol'</tt>, or <tt>'e'</tt>, or <tt>'g'</tt>.

 

[[Image:Tempcalb.jpg]]

 Fig. 1D spectrum collected for temperature calibration.

 -- Main.GaohuaLiu - 24 Jan 2007

Shimming

2009-12-09T17:24:02Z

Aringh:

Sample shimming is a process of optimizing magnetic field homogeneity for recording high resolution spectra. The shim system is essentially a set of coils with specific field profiles in 3D space. There are three shimming methods: shimming on the lock level, shimming on the FID (or spectrum) and gradient shimming.  The currents in the shim coils are adjusted to cancel any unwanted magnetic field gradients across the NMR sample as accurately as possible.

== Routine Shimming Protocol ==

#Retrieve the starting shims. This should be the closest set shims for your particular sample. e.g.:
#*Previous shims for the same sample.
#*Shims for a different sample of the same tube geometry and similar filling height.
#*Reference shims determined for a lineshape or doped water sample.
#Find and activate lock, while avoiding its saturation.
#Manually optimize <tt>Z1</tt> and <tt>Z2</tt> shims, then optimize <tt>X1</tt> and <tt>Y1</tt> shims.
#Start <tt>gmapsys</tt> and set up 1H gradient shimming. Optimize parameters and generate a new shimmap if necessary. Run gradient shimming with the appropriate <tt>gzsize</tt> setting. You can use a smaller <tt>gzsize</tt> value at this step.
#Manually optimize <tt>X1</tt> and <tt>Y1</tt> shims, then optimize <tt>XZ</tt> and <tt>YZ</tt> shims.
#Repeat step 4 with the optimal <tt>gzsize</tt> setting.

== Storing and Retrieving Shims ==

To save current shim parameters in a shim file named 'myshims', type in the command prompt

<tt>svs('myshims')</tt>

The shim file will be stored stored in <tt>~/vnmrsys/shims/</tt>.

To retrieve shim parameters type

<tt>rts('myshims') su </tt>

The <tt>rts</tt> command only loads the shim values into VNMR or VNMRJ. The <tt>su</tt> command is needed to send the new shim values to the shim unit. you are likely see a change in the lock level as the new shims are applied. See also information on the <tt>load</tt> parameter.

It may be a good idea to load old shim parameters before shimming. For example, if you want to acquire using a shigemi tube, but someone before you was using a regular NMR tube in the same spectrometer, loading old shim parameters should make the shimming easier. If you have shim parameters saved from an earlier measurement on the same sample, it is better to load those, since they are likely to be a much closer match.

== Manual shimming on the lock ==

Adjusting shim values to maximize the lock level is called shimming on the lock. As the name suggests, you need to have your sample locked for this procedure. Make sure the lock resonance is not saturated. The entire procedure is complex, and is described in '''Section 6.9. Using the Acquisition Window''' of the [http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199916000b.pdf VNMR 6.1C Getting Started] manual and '''Section 3.10. Shimming on the Lock Signal Manually''' of [http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199931300b.pdf VNMRJ Liquids NMR User Guide]

A routine procedure for any sample:

#Alternate between Z1 and Z2 to maximize lock
#Alternate between X and Y to maximize lock

This should improve convergence of gradient shimming.

Shimming of higher-order spinning shims is usually only done for standard lineshape samples.

== Gradient Shimming ==

In gradient shimming, like in MRI, gradients are used to probe the field profile within the sample. For that you need a single strong signal in your sample. For protein samples this is usually the H2O resonance. (In principle, gradient shimming can be done with any solvent, and even run on deuterium resonances). On our spectrometers gradient shimming only adjusts Z-shims (on some spectrometers with XYZ-gradient probes it is possible to use gradient shimming for all X-, Y- and Z-shims).

Even though it is not directly obvious, you MUST have the sample locked to run gradient shimming.

The procedure for gradient shimming is documented in the '''Section 11.6. Gradient Sample Shimming''' of [http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199916100c.pdf User's Guide: Liquids NMR]

=== Initial Setup and Parameter Optimization ===

#Make sure that your sample is locked and lock signal is not saturated.
#Turn spinning off.
#Make coarse adjustments of Z1, Z2, X, and Y shims as described above.
#Load <tt>gmapz</tt> dataset by typing <tt>gmapsys</tt>.
#Type <tt>dps</tt> or <tt>tn?</tt> to confirm that you have the dataset configured for 1H. Otherwise select 1H gradient shimming by clicking on the '''PFG H1''' button in the '''Acquire''' -> '''Gradient Shim''' tab in VNMRJ.
#Check parameters <tt>pw</tt>, <tt>tpwr</tt>, <tt>gain</tt>, <tt>d3</tt>, <tt>d1</tt>, <tt>nt</tt> and <tt>ss</tt>. For aqueous protein samples <tt>pw</tt> should be 1 or 2 microseconds, and <tt>gain</tt> should be low (< 6, otherwise ADC overflow may occur).
#Acquire test field profiles:
#*In VNMRJ Click on '''Acquire Trial Spectra''' in the '''Acquire''' -> '''Gradient Shim''' tab
#*In VNMR: Click on '''Set Params''' -> '''go, dssh'''. Click '''Return''' to get back to the main gmapz menu.

To further minimize radiation damping it may be useful not to tune the probe before shimming.

[[Image:Gmapsys.jpg|579x160px]]

You should see two field profiles, the second one smaller than the first. If the shims are already quite good the profiles would be symmetric. Room-temperature probes typically yield hat-like profiles, while cryogenic probes yield more rectangular or trapezoidal profiles. The profile width is determined by the filling height.

Make sure that the profiles are smooth and have a high SN before proceeding.

Some examples of bad field profiles:

*Noisy baseline and profile - insufficient SN, increase <tt>gain</tt> and/or <tt>pw</tt>.
*Jagged baseline and profile - ADC overflow, decrease <tt>gain</tt> and/or <tt>pw</tt>.
*Rough left edge of the profile - Likely sample precipitation.
*Rough right edge of the profile - Likely bubbles in a shigemi tube under the plunger, eject sample and remove bubbles.

For optimal performance the second profile should be about 50% lower than the first. This is controlled by the <tt>d3</tt> parameter, which sets the de-phasing time. This is an arrayed parameter: its first value is 0 and leads to the first profile, the second value is set to some delay and generates the second profile. If the starting shims are very good (this includes both axial and spinning shims!) and field homogeneity is high, then you need a longer delay to observe sufficient de-phasing. If, on the other hand, the starting shims are too far off, the de-phasing could be too large to be accurately quantified by the read-out gradient.

=== Creating Shim Maps ===

*Set <tt>gzsize = 5</tt> to select shims Z1 through Z5. Even though you may omit Z5 in subsequent gradient shimming, it is worth to have a map for it in the first place.
*Click '''Shim Maps''' -> '''Automake Shimmap''' to run automatic shim mapping. Give a name to your shim map (usually includes your sample name and the date).

=== Manual Calculation of gzwin and tof (Optional) ===

The <tt>gzwin</tt> parameter determines the size of the spectral region used to optimize shims, and <tt>tof</tt> in this particular case describes how it is centered. This region ideally corresponds to the active sample volume and is normally calculated automatically when you click on '''Set Params''' -> '''Find gzwin''', or '''Shim Maps''' -> '''Automake Shimmap'''.

For some samples, however, these parameters may be determined incorrectly due to imperfect field profiles. This is often observed for samples of suboptimal volume or samples with precipitation at the bottom.

In this case you can calculate <tt>gzwin</tt> and <tt>tof</tt> and map shims manually. The procedure is:

*Click on '''Set Params''' -> '''go, dssh'''.
*Type <tt>ds</tt> to display the first profile
*Use the box cursors to define the desired region
*Enter <tt>gmapsys</tt> and click '''Set Params''' -> '''Calculate gzwin'''.
*Click on '''Return''' -> '''Shim Maps''' -> '''Make Shimmap''' ([[NESG:ShimMapResult|ShimMapResult]])
*Click on '''Return''' -> '''Display''' -> '''Display Shimmap''' ([[NESG:DisplayShimMaps|Shim map example]])

=== Gradient Sample Shimming ===

First set <tt>gzsize</tt> to select the order of Z shims to be used:

*For large sample depths (e.g. in a regular tube) set <tt>gzsize = 5</tt>. If <tt>gmap_z1z4 = 'y'</tt> the first iterations will be carried out with Z1-Z4 shims and then with Z1-Z5 shims.
*For shigemi tubes with optimal sample volume (250-300 ul) set <tt>gzsize = 4</tt>.
*For shigemi tubes with lower than optimal sample depth use <tt>gzsize = 3</tt>.

Click on the '''Autoshim on Z''' button to start gradient shimming. Sample shimming will go on until rms error number is less than 1, or until maximum number of iterations (generally 5) are reached.

Using higher-order shims with a low sample depth may decrease convergence - you may even end up with worse shims than that you started with. Save the current shims with <tt>svs</tt> before testing to see if using higher-order shims has any benefits.

For best results, alternate between gradient shimming and adjusting X, Y, XZ and YZ shims by shimming on the lock.

For example, you have a sample of 220 ul in shigemi tube. Run gradient shimming with Z1-Z3 (<tt>gzsize = 3</tt>), then adjust X, Y, XZ and YZ, then do gradient shimming again with Z1-Z3 (<tt>gzsize = 3</tt>), save shims and run gradient shimming again with Z1-Z4 (<tt>gzsize = 4</tt>).

Click on '''Quit''' button to exit gradient shimming mode and go back to your experiment set up.

 

=== Re-cabling for Gradient Shimming on Deuterium ===

The 500 MHz instrument in Buffalo has a relay, which allows gradient shimming on 2H without switching cables.

The 600 MHz and 750 MHz instruments need to be wired for 2H observe in order to run gradient shimming on 2H. Assuming that you start with the instrument configured for 1H observe, do the following:

#Disconnect the cable from the "PROBE J1" port on the lock switch (bottom right on the picture below)
#Connect this cable to the "PROBE" port on the broadband preamp, as indicated by the red arrow (the left side of the magnet leg).

Also see the related topic on [[NESG:DeuteriumDecoupling#1D_Deuterium_Spectrum|directly observing deuterium]].

== Background Hardware Autoshim ==

Background hardware autoshim can only be active during acquisition. It starts with an acquisition start command (<tt>ga</tt>, <tt>go</tt> or <tt>au</tt>). This method is very useful for protein samples, whose B0 homogeneity is affected by continuous precipitation. It can also be useful for magnets with large drift rates, where large field corrections with Z0 coil introduce substantial Z1 perturbation.

Usage:

*<tt>hdwshim='y' su</tt> - Enable hardware autoshim during the first delay in the pulse sequence
*<tt>hdwshim='p' su</tt> - Enable hardware autoshim during the first presaturation delay in the pulse sequence
*<tt>hdwshim='n' su</tt> - Disable hardware autoshim

The shims used in hardware autoshim are determine by the global <tt>hdwshimlist</tt> parameter. If <tt>hdwshimlist</tt> does not exist, autoshim will only use <tt>z1</tt>.

To create the <tt>hdwshimlist</tt> type <tt>create('hdwshimlist', 'string', 'global')</tt>

You can set <tt>hdwshimlist</tt> to any combination of the following: <tt>z1</tt>, <tt>z1c</tt>, <tt>z2</tt>, <tt>z2c</tt>, <tt>x1</tt>, <tt>y1</tt>. The order is irrelevant. However, use of coarse shims (<tt>z1c</tt> and <tt>z2c</tt>) is not recommended. Examples:

*<tt>hdwshimlist='z1z2'</tt>
*<tt>hdwshimlist='z1z2x1y1'</tt>

In VNMRJ you can also control hardware autoshim from the '''Setup''' -> '''Shim''' tab.

Note that hardware autoshim is automatically switched off when other shim methods, like gradient shimming or software autoshim, are started. Check the <tt>hdwshim</tt> to see if hardware autoshim is still enabled.

 

== Sample Shimming on the FID ==

This is the option of last resort if you don't have a lock substance in your sample.

See '''Section 6.9. Using the Acquisition Window''' of the [http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199916000b.pdf Getting Started] manual of VNMR 6.1C for details

 

Shimming with the CHCl3 lineshape sample

 

-- Main.GaohuaLiu - 24 Jan 2007

Shimming

2009-12-09T17:17:01Z

Aringh:

Sample shimming is a process of optimizing magnetic field homogeneity for recording high resolution spectra. The shim system is essentially a set of coils with specific field profiles in 3D space. There are three shimming methods: shimming on the lock level, shimming on the FID (or spectrum) and gradient shimming.  The currents in the shim coils are adjusted to cancel any unwanted magnetic field gradients across the NMR sample as accurately as possible.

== Routine Shimming Protocol ==

#Retrieve the starting shims. This should be the closest set shims for your particular sample. e.g.:
#*Previous shims for the same sample.
#*Shims for a different sample of the same tube geometry and similar filling height.
#*Reference shims determined for a lineshape or doped water sample.
#Find and activate lock, while avoiding its saturation.
#Manually optimize <tt>Z1</tt> and <tt>Z2</tt> shims, then optimize <tt>X1</tt> and <tt>Y1</tt> shims.
#Start <tt>gmapsys</tt> and set up 1H gradient shimming. Optimize parameters and generate a new shimmap if necessary. Run gradient shimming with the appropriate <tt>gzsize</tt> setting. You can use a smaller <tt>gzsize</tt> value at this step.
#Manually optimize <tt>X1</tt> and <tt>Y1</tt> shims, then optimize <tt>XZ</tt> and <tt>YZ</tt> shims.
#Repeat step 4 with the optimial <tt>gzsize</tt> setting.

== Storing and Retrieving Shims ==

To save current shim parameters in a shim file named 'myshims', type in the command prompt

<tt>svs('myshims')</tt>

The shim file will be stored stored in <tt>~/vnmrsys/shims/</tt>.

To retrieve shim parameters type

<tt>rts('myshims') su </tt>

The <tt>rts</tt> command only loads the shim values into VNMR or VNMRJ. The <tt>su</tt> command is needed to send the new shim values to the shim unit. you are likely see a change in the lock level as the new shims are applied. See also information on the <tt>load</tt> parameter.

It may be a good idea to load old shim parameters before shimming. For example, if you want to acquire using a shigemi tube, but someone before you was using a regular NMR tube in the same spectrometer, loading old shim parameters should make the shimming easier. If you have shim parameters saved from an earlier measurement on the same sample, it is better to load those, since they are likely to be a much closer match.

== Manual shimming on the lock ==

Adjusting shim values to maximize the lock level is called shimming on the lock. As the name suggests, you need to have your sample locked for this procedure. Make sure the lock resonance is not saturated. The entire procedure is complex, and is described in '''Section 6.9. Using the Acquisition Window''' of the [http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199916000b.pdf VNMR 6.1C Getting Started] manual and '''Section 3.10. Shimming on the Lock Signal Manually''' of [http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199931300b.pdf VNMRJ Liquids NMR User Guide]

A routine procedure for any sample:

#Alternate between Z1 and Z2 to maximize lock
#Alternate between X and Y to maximize lock

This should improve convergence of gradient shimming.

Shimming of higher-order spinning shims is usually only done for standard lineshape samples.

== Gradient Shimming ==

In gradient shimming, like in MRI, gradients are used to probe the field profile within the sample. For that you need a single strong signal in your sample. For protein samples this is usually the H2O resonance. (In principle, gradient shimming can be done with any solvent, and even run on deuterium resonances). On our spectrometers gradient shimming only adjusts Z-shims (on some spectrometers with XYZ-gradient probes it is possible to use gradient shimming for all X-, Y- and Z-shims).

Even though it is not directly obvious, you MUST have the sample locked to run gradient shimming.

The procedure for gradient shimming is documented in the '''Section 11.6. Gradient Sample Shimming''' of [http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199916100c.pdf User's Guide: Liquids NMR]

=== Initial Setup and Parameter Optimization ===

#Make sure that your sample is locked and lock signal is not saturated.
#Turn spinning off.
#Make coarse adjustments of Z1, Z2, X, and Y shims as described above.
#Load <tt>gmapz</tt> dataset by typing <tt>gmapsys</tt>.
#Type <tt>dps</tt> or <tt>tn?</tt> to confirm that you have the dataset configured for 1H. Otherwise select 1H gradien shimming by clicking on the '''PFG H1''' button in the '''Acquire''' -> '''Gradient Shim''' tab in VNMRJ.
#Check parameters <tt>pw</tt>, <tt>tpwr</tt>, <tt>gain</tt>, <tt>d3</tt>, <tt>d1</tt>, <tt>nt</tt> and <tt>ss</tt>. For aqueous protein samples <tt>pw</tt> should be 1 or 2 microseconds, and <tt>gain</tt> should be low (< 6, otherwise ADC overflow may occur).
#Acquire test field profiles:
#*In VNMRJ Click on '''Acquire Trial Spectra''' in the '''Acquire''' -> '''Gradient Shim''' tab
#*In VNMR: Click on '''Set Params''' -> '''go, dssh'''. Click '''Return''' to get back to the main gmapz menu.

To further minimize radiation damping it may be useful not to tune the probe before shimming.

[[Image:Gmapsys.jpg|579x160px]]

You should see two field profiles, the second one smaller than the first. If the shims are already quite good the profiles would be symmetric. Room-temperature probes typically yield hat-like profiles, while cryogenic probes yield more rectangular or trapeziodal profiles. The profile width is determined by the filling height.

Make sure that the profiles are smooth and have a high SN before proceeding.

Some examples of bad field profiles:

*Noisy baseline and profile - insufficient SN, increase <tt>gain</tt> and/or <tt>pw</tt>.
*Jagged baseline and profile - ADC overflow, decrease <tt>gain</tt> and/or <tt>pw</tt>.
*Rough left edge of the profile - Likely sample precipitation.
*Rough right edge of the profile - Likely bubbles in a shigemi tube under the plunger, eject sample and remove bubbles.

For optimal performance the second profile should be about 50% lower than the first. This is controlled by the <tt>d3</tt> parameter, which sets the de-phasing time. This is an arrayed parameter: its first value is 0 and leads to the first profile, the second value is set to some delay and generates the second profile. If the starting shims are very good (this includes both axial and spinning shims!) and field homogeneity is high, then you need a longer delay to observe sufficient de-phasing. If, on the other hand, the starting shims are too far off, the de-phasing could be too large to be accurately quantified by the read-out gradient.

=== Creating Shim Maps ===

*Set <tt>gzsize = 5</tt> to select shims Z1 through Z5. Even though you may omit Z5 in subsequent gradient shimming, it is worth to have a map for it in the first place.
*Click '''Shim Maps''' -> '''Automake Shimmap''' to run automatic shim mapping. Give a name to your shim map (usually includes your sample name and the date).

=== Manual Calculation of gzwin and tof (Optional) ===

The <tt>gzwin</tt> parameter determines the size of the spectral region used to optimize shims, and <tt>tof</tt> in this particular case describes how it is centered. This region ideally corresponds to the active sample volume and is normally calculated automatically when you click on '''Set Params''' -> '''Find gzwin''', or '''Shim Maps''' -> '''Automake Shimmap'''.

For some samples, however, these parameters may be determined incorrectly due to imperfect field profiles. This is often observed for samples of suboptimal volume or samples with precipitation at the bottom.

In this case you can calculate <tt>gzwin</tt> and <tt>tof</tt> and map shims manually. The procedure is:

*Click on '''Set Params''' -> '''go, dssh'''.
*Type <tt>ds</tt> to display the first profile
*Use the box cursors to define the desired region
*Enter <tt>gmapsys</tt> and click '''Set Params''' -> '''Calculate gzwin'''.
*Click on '''Return''' -> '''Shim Maps''' -> '''Make Shimmap''' ([[NESG:ShimMapResult|ShimMapResult]])
*Click on '''Return''' -> '''Display''' -> '''Display Shimmap''' ([[NESG:DisplayShimMaps|Shim map example]])

=== Gradient Sample Shimming ===

First set <tt>gzsize</tt> to select the order of Z shims to be used:

*For large sample depths (e.g. in a regular tube) set <tt>gzsize = 5</tt>. If <tt>gmap_z1z4 = 'y'</tt> the first iterations will be carried out with Z1-Z4 shims and then with Z1-Z5 shims.
*For shigemi tubes with optimal sample volume (250-300 ul) set <tt>gzsize = 4</tt>.
*For shigemi tubes with lower than optimal sample depth use <tt>gzsize = 3</tt>.

Click on the '''Autoshim on Z''' button to start gradient shimming. Sample shimming will go on until rms error number is less than 1, or until maximum number of iterations (generally 5) are reached.

Using higher-order shims with a low sample depth may decrease convergence - you may even end up with worse shims than that you started with. Save the current shims with <tt>svs</tt> before testing to see if using higher-order shims has any benefits.

For best results, alternate between gradient shimming and adjusting X, Y, XZ and YZ shims by shimming on the lock.

For example, you have a sample of 220 ul in shigemi tube. Run gradient shimming with Z1-Z3 (<tt>gzsize = 3</tt>), then adjust X, Y, XZ and YZ, then do gradient shimming again with Z1-Z3 (<tt>gzsize = 3</tt>), save shims and run gradient shimming again with Z1-Z4 (<tt>gzsize = 4</tt>).

Click on '''Quit''' button to exit gradient shimming mode and go back to your experiment set up.

 

=== Re-cabling for Gradient Shimming on Deuterium ===

The 500 MHz instrument in Buffalo has a relay, which allows gradient shimming on 2H without switching cables.

The 600 MHz and 750 MHz instruments need to be wired for 2H observe in order to run gradient shimming on 2H. Assuming that you start with the instrument configured for 1H observe, do the following:

#Disconnect the cable from the "PROBE J1" port on the lock switch (bottom right on the picture below)
#Connect this cable to the "PROBE" port on the bradband preamp, as indicated by the red arrow (the left side of the magnet leg).

Also see the related topic on [[NESG:DeuteriumDecoupling#1D_Deuterium_Spectrum|directly observing deuterium]].

== Background Hardware Autoshim ==

Background hardware autoshim can only be active during acquisition. It starts with an acquisition start command (<tt>ga</tt>, <tt>go</tt> or <tt>au</tt>). This method is very useful for protein samples, whose B0 homogeneity is affected by continuous precipitation. It can also be useful for magnets with large drift rates, where large field corrections with Z0 coil introduce substantial Z1 perturbation.

Usage:

*<tt>hdwshim='y' su</tt> - Enable hardware autoshim during the first delay in the pulse sequence
*<tt>hdwshim='p' su</tt> - Enable hardware autoshim during the first presaturation delay in the pulse sequence
*<tt>hdwshim='n' su</tt> - Disable hardware autoshim

The shims used in hardware autoshim are determine by the global <tt>hdwshimlist</tt> parameter. If <tt>hdwshimlist</tt> does not exist, autoshim will only use <tt>z1</tt>.

To create the <tt>hdwshimlist</tt> type <tt>create('hdwshimlist', 'string', 'global')</tt>

You can set <tt>hdwshimlist</tt> to any combination of the following: <tt>z1</tt>, <tt>z1c</tt>, <tt>z2</tt>, <tt>z2c</tt>, <tt>x1</tt>, <tt>y1</tt>. The order is irrelevant. Using coarse shims (<tt>z1c</tt> and <tt>z2c</tt>) is not recommended. Examples:

*<tt>hdwshimlist='z1z2'</tt>
*<tt>hdwshimlist='z1z2x1y1'</tt>

In VNMRJ you can also control hardware autoshim from the '''Setup''' -> '''Shim''' tab.

Note that hardware autoshim is automatically switched off when other shim methods, like gradient shimming or software autoshim, are started. Check the <tt>hdwshim</tt> to see if hardware autoshim is still enabled.

 

== Sample Shimming on the FID ==

This is the option of last resort if you don't have a lock substance in your sample.

See '''Section 6.9. Using the Acquisition Window''' of the [http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199916000b.pdf Getting Started] manual of VNMR 6.1C for details

 

Shimming with the CHCl3 lineshape sample

 

-- Main.GaohuaLiu - 24 Jan 2007

Shimming

2009-12-09T17:14:24Z

Aringh:

Sample shimming is a process of optimizing magnetic field homogeneity for recording high resolution spectra. The shim system is essentially a set of coils with specific field profiles in 3D space. There are three shimming methods: shimming on the lock level, shimming on the FID (or spectrum) and gradient shimming.  The currents in the shim coils are adjusted to cancel any unwanted magnetic field gradients across the NMR sample as accurately as possible.

== Routine Shimming Protocol ==

#Retrieve the starting shims. This should be the closest set shims for your particular sample. e.g.:
#*Previous shims for the same sample.
#*Shims for a different sample of the same tube geometry and similar filling height.
#*Reference shims determined for a lineshape or doped water sample.
#Find and activate lock, while avoiding its saturation.
#Manually optimize <tt>Z1</tt> and <tt>Z2</tt> shims, then optimize <tt>X1</tt> and <tt>Y1</tt> shims.
#Start <tt>gmapsys</tt> and set up 1H gradient shimming. Optimize parameters and generate a new shimmap if necessary. Run gradient shimming with the appropriate <tt>gzsize</tt> setting. You can use a smaller <tt>gzsize</tt> value at this step.
#Manually optimize <tt>X1</tt> and <tt>Y1</tt> shims, then optimize <tt>XZ</tt> and <tt>YZ</tt> shims.
#Repeat step 4 with the optimial <tt>gzsize</tt> setting.

== Storing and Retrieving Shims ==

To save current shim parameters in a shim file named 'myshims', type in the command prompt

<tt>svs('myshims')</tt>

The shim file will be stored stored in <tt>~/vnmrsys/shims/</tt>.

To retrieve shim parameters type

<tt>rts('myshims') su </tt>

The <tt>rts</tt> command only loads the shim values into VNMR or VNMRJ. The <tt>su</tt> command is needed to send the new shim values to the shim unit. you are likely see a change in the lock level as the new shims are applied. See also information on the <tt>load</tt> parameter.

It may be a good idea to load old shim parameters before shimming. For example, if you want to acquire using a shigemi tube, but someone before you was using a regular NMR tube in the same spectrometer, loading old shim parameters should make the shimming easier. If you have shim parameters saved from an earlier measurement on the same sample, it is better to load those, since they are likely to be a much closer match.

== Manual shimming on the lock ==

Adjusting shim values to maximize the lock level is called shimming on the lock. As the name suggests, you need to have your sample locked for this procedure. Make sure the lock resonance is not saturated. The entire procedure is complex, and is described in '''Section 6.9. Using the Acquisition Window''' of the [http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199916000b.pdf VNMR 6.1C Getting Started] manual and '''Section 3.10. Shimming on the Lock Signal Manually''' of [http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199931300b.pdf VNMRJ Liquids NMR User Guide]

A routine procedure for any sample:

#Alternate between Z1 and Z2 to maximize lock
#Alternate between X and Y to maximize lock

This should improve convergence of gradient shimming.

Shimming of higher-order spinning shims is usually only done for standard lineshape samples.

== Gradient Shimming ==

In gradient shimming, like in MRI, gradients are used to probe the field profile within the sample. For that you need a single strong signal in your sample. For protein samples this is usually the H2O resonance. (In principle, gradient shimming can be done with any solvent, and even run on deuterium resonances). On our spectrometers gradient shimming only adjusts Z-shims (on some spectrometers with XYZ-gradient probes it is possible to use gradient shimming for all X-, Y- and Z-shims).

Even though it is not directly obvious, you MUST have the sample locked to run gradient shimming.

The procedure for gradient shimming is documented in the '''Section 11.6. Gradient Sample Shimming''' of [http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199916100c.pdf User's Guide: Liquids NMR]

=== Initial Setup and Parameter Optimization ===

#Make sure that your sample is locked and lock signal is not saturated.
#Turn spinning off.
#Make coarse adjustments of Z1, Z2, X, and Y shims as described above.
#Load <tt>gmapz</tt> dataset by typing <tt>gmapsys</tt>.
#Type <tt>dps</tt> or <tt>tn?</tt> to confirm that you have the dataset configured for 1H. Otherwise select 1H gradien shimming by clicking on the '''PFG H1''' button in the '''Acquire''' -> '''Gradient Shim''' tab in VNMRJ.
#Check parameters <tt>pw</tt>, <tt>tpwr</tt>, <tt>gain</tt>, <tt>d3</tt>, <tt>d1</tt>, <tt>nt</tt> and <tt>ss</tt>. For aqueous protein samples <tt>pw</tt> should be 1 or 2 microseconds, and <tt>gain</tt> should be low (< 6, otherwise ADC overflow may occur).
#Acquire test field profiles:
#*In VNMRJ Click on '''Acquire Trial Spectra''' in the '''Acquire''' -> '''Gradient Shim''' tab
#*In VNMR: Click on '''Set Params''' -> '''go, dssh'''. Click '''Return''' to get back to the main gmapz menu.

To further minimize radiation damping it may be useful not to tune the probe before shimming.

[[Image:Gmapsys.jpg|579x160px]]

You should see two field profiles, the second one smaller than the first. If the shims are already quite good the profiles would be symmetric. Room-temperature probes typically yield hat-like profiles, while cryogenic probes yield more rectangular or trapeziodal profiles. The profile width is determined by the filling height.

Make sure that the profiles are smooth and have a high SN before proceeding.

Some examples of bad field profiles:

*Noisy baseline and profile - insufficient SN, increase <tt>gain</tt> and/or <tt>pw</tt>.
*Jagged baseline and profile - ADC overflow, decrease <tt>gain</tt> and/or <tt>pw</tt>.
*Rough left edge of the profile - Likely sample precipitation.
*Rough right edge of the profile - Likely bubbles in a shigemi tube under the plunger, eject sample and remove bubbles.

For optimal performance the second profile should be about 50% lower than the first. This is controlled by the <tt>d3</tt> parameter, which sets the de-phasing time. This is an arrayed parameter: its first value is 0 and leads to the first profile, the second value is set to some delay and generates the second profile. If the starting shims are very good (this includes both axial and spinning shims!) and field homogeneity is high, then you need a longer delay to observe sufficient de-phasing. If, on the other hand, the starting shims are too far off, the de-phasing could be too large to be accurately quantified by the read-out gradient.

=== Creating Shim Maps ===

*Set <tt>gzsize = 5</tt> to select shims Z1 through Z5. Even though you may omit Z5 in subsequent gradient shimming, it is worth to have a map for it in the first place.
*Click '''Shim Maps''' -> '''Automake Shimmap''' to run automatic shim mapping. Give a name to your shim map (usually includes your sample name and the date).

=== Manual Calculation of gzwin and tof (Optional) ===

The <tt>gzwin</tt> parameter determines the size of the spectral region used to optimize shims, and <tt>tof</tt> in this particular case describes how it is centered. This region ideally corresponds to the active sample volume and is normally calculated automatically when you click on '''Set Params''' -> '''Find gzwin''', or '''Shim Maps''' -> '''Automake Shimmap'''.

For some samples, however, these parameters may be determined incorrectly due to imperfect field profiles. This is often observed for samples of suboptimal volume or samples with precipitation at the bottom.

In this case you can calculate <tt>gzwin</tt> and <tt>tof</tt> and map shims manually. The procedure is:

*Click on '''Set Params''' -> '''go, dssh'''.
*Type <tt>ds</tt> to display the first profile
*Use the box cursors to define the desired region
*Enter <tt>gmapsys</tt> and click '''Set Params''' -> '''Calculate gzwin'''.
*Click on '''Return''' -> '''Shim Maps''' -> '''Make Shimmap''' ([[NESG:ShimMapResult|ShimMapResult]])
*Click on '''Return''' -> '''Display''' -> '''Display Shimmap''' ([[NESG:DisplayShimMaps|Shim map example]])

=== Gradient Sample Shimming ===

First set <tt>gzsize</tt> to select the order of Z shims to be used:

*For large sample depths (e.g. in a regular tube) set <tt>gzsize = 5</tt>. If <tt>gmap_z1z4 = 'y'</tt> the first iterations will be carried out with Z1-Z4 shims and then with Z1-Z5 shims.
*For shigemi tubes with optimal sample volume (250-300 ul) set <tt>gzsize = 4</tt>.
*For shigemi tubes with lower than optimal sample depth use <tt>gzsize = 3</tt>.

Click on the '''Autoshim on Z''' button to start gradient shimming. Sample shimming will go on until rms error number is less than 1, or until maximum number of iterations (generally 5) are reached.

Using higher-order shims with a low sample depth may decrease convergence - you may even end up with worse shims than that you started with. Save the current shims with <tt>svs</tt> before testing to see if using higher-order shims has any benefits.

For best results, alternate between gradient shimming and adjusting X, Y, XZ and YZ shims by shimming on the lock.

For example, you have a sample of 220 ul in shigemi tube. Run gradient shimming with Z1-Z3 (<tt>gzsize = 3</tt>), then adjust X, Y, XZ and YZ, then do gradient shimming again with Z1-Z3 (<tt>gzsize = 3</tt>), save shims and run gradient shimming again with Z1-Z4 (<tt>gzsize = 4</tt>).

Click on '''Quit''' button to exit gradient shimming mode and go back to your experiment set up.

 

=== Re-cabling for Gradient Shimming on Deuterium ===

The 500 MHz instrument in Buffalo has a relay, which allows gradient shimming on 2H without switching cables.

The 600 MHz and 750 MHz instruments need to be wired for 2H observe in order to run gradient shimming on 2H. Assuming that you start with the instrument configured for 1H observe, do the following:

#Disconnect the cable from the "PROBE J1" port on the lock switch (bottom right on the picture below)
#Connect this cable to the "PROBE" port on the bradband preamp, as indicated by the red arrow (the left side of the magnet leg).

Also see the related topic on [[NESG:DeuteriumDecoupling#1D_Deuterium_Spectrum|directly observing deuterium]].

== Backgroung Hardware Autoshim ==

Background hardware autoshim can only be active during acquisition. It starts with an acquisition start command (<tt>ga</tt>, <tt>go</tt> or <tt>au</tt>). This method is very useful for protein samples, whose B0 homogeneity is affected by continuous precipitation. It can also be useful for magnets with large drift rates, where large field corrections with Z0 coil introduce substantial Z1 perturbation.

Usage:

*<tt>hdwshim='y' su</tt> - Enable hardware autoshim during the first delay in the pulse sequence
*<tt>hdwshim='p' su</tt> - Enable hardware autoshim during the first presaturation delay in the pulse sequence
*<tt>hdwshim='n' su</tt> - Disable hardware autoshim

The shims used in hardware autoshim are determine by the global <tt>hdwshimlist</tt> parameter. If <tt>hdwshimlist</tt> does not exist, autoshim will only use <tt>z1</tt>.

To create the <tt>hdwshimlist</tt> type <tt>create('hdwshimlist', 'string', 'global')</tt>

You can set <tt>hdwshimlist</tt> to any combination of the following: <tt>z1</tt>, <tt>z1c</tt>, <tt>z2</tt>, <tt>z2c</tt>, <tt>x1</tt>, <tt>y1</tt>. The order is irrelevant. Using coarse shims (<tt>z1c</tt> and <tt>z2c</tt>) is not recommended. Examples:

*<tt>hdwshimlist='z1z2'</tt>
*<tt>hdwshimlist='z1z2x1y1'</tt>

In VNMRJ you can also control hardware autoshim from the '''Setup''' -> '''Shim''' tab.

Note that hardware autoshim is automatically switched off when other shim methods, like gradient shimming or software autoshim, are started. Check the <tt>hdwshim</tt> to see if hardware autoshim is still enabled.

 

== Sample Shimming on the FID ==

This is the option of last resort if you don't have a lock substance in your sample.

See '''Section 6.9. Using the Acquisition Window''' of the [http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199916000b.pdf Getting Started] manual of VNMR 6.1C for details

 

Shimming with the CHCl3 lineshape sample

 

-- Main.GaohuaLiu - 24 Jan 2007

Deuterium Lock

2009-12-09T16:30:13Z

Aringh:

The lock system keeps the spectrometer operating at a constant net magnetic field. It compensates for the drift of the superconducting magnet's field and other instabilities. The current in the <tt>z0</tt> coil is adjusted so that the lock resonance matches a predefined frequency <tt>lockfreq</tt> . Before activating the lock system you have to find the lock resonance.

For detailed information on locking you sample see

*[http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199916000b.pdf VNMR 6.1C Getting Started Manual]
*[http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199931300b.pdf VNMRJ Liquids NMR Users Guide], section 3.7
*BioPack manual

Note that BioPack installation alters the interface described in the original manuals.

== Finding Lock Signal Manually ==

First you must start the lock signal display. The channel selector for probe tuning should be at 0.

In VNMR:

#Click on the '''Acqi''' button or type <tt>acqi</tt> to open the acquisition window.
#Click in the '''Lock''' button in the acquisition window

In VNMRJ:

#Go to the '''Setup''' tab; click on the '''Lock''' tab to see the adjustable parameters
#Click in the '''Lock Scan''' button in the middle of the screen

[[Image:Vnmrj lock.png|606x172px]]

Make sure the the lock system is not activated:

*In VNMR click on the '''LOCK:off''' button in the acquisition window;   LOCKED should change to  LOCK OFF 
*In VNMRJ uncheck the '''Activate Lock''' checkbox

Adjust <tt>z0</tt> to locate the lock resonance. You may increase lock power and gain to see the signal better. When manually adjusting <tt>z0</tt> the following rule holds: the closer you are to the resonance, the fewer sinusoid "beats" you see. When on-resonance you should see a straight line. The figures below show examples from a typical locking process.

[[Image:Lock a.jpg|205x273px|lock off-resonance]][[Image:Lock b.jpg|205x273px|lock closer to resonance]][[Image:Lock c.jpg|205x273px|lock on resonance]]''' '''

 

Activate the lock system:

*In VNMR click on the '''LOCK:on''' button in the acquisition window;  LOCK OFF   should change to  LOCKED  
*In VNMRJ check the '''Activate Lock''' checkbox

Adjust lock phase to maximize the lock signal. You may actually need to re-optimize lock phase after shimming.

Adjust lock power to maximize the signal while avoiding saturation. In the unsaturated state decreasing lock power by 6 dB should make the lock level drop by one half. Under saturating conditions the lock resonance is distorted and this may pose a problem for shimming on the lock.

Adjust lock gain to maximize the lock signal. Since it controls the amplification of the lock signal, increasing lock gain will increase both lock level and its noise.

Normally you want to have the highest non-saturating <tt>lockpower</tt> and the lowest <tt>lockgain</tt>, which yield an acceptable lock level. Lock level should be above ~20 for the sample to be locked. The lock level will usually increase after you shim the sample. It should be ~50 or higher before you run a real experiment, because if the level drops below 20 during acquisition (due to gradients, for example) lock may be lost.

Also, tuning the lock channel may improve sensitivity. Lock channel tuning is normally available on cryprobes.

The field produced by the <tt>z0</tt> coil has some slight z-dependence (mainly linear), which is usually compensated during shimming. On machines with very large field drift this may require use of hardware autoshim during long-term experiments.

== Finding Lock Signal Automatically ==

In VNMRJ:

#Go to the '''Setup''' -> '''Lock''' tab
#Click on the '''Find Lock''' button

This procedure usually takes longer, but may be useful if cannot find the lock resonance manually.

Note that you will still need to manually adjust lock power to avoid lock saturation.

== Temperature Dependence ==

The position of the water line in 1H or 2H spectra has a strong dependence on temperature. Since lock tries to match the deuterium resonance of water to a predefined frequency, <tt>z0</tt> will be different at different temperatures.

It also means the total <tt>B0</tt> field will be different even for the same spectrometer and sample locked at different temperatures. Thus, the DSS spectrum should be acquired at the same temperature as the protein sample being referenced. Alternatively, the DSS line position can be tabulated at various temperatures and it temperature dependence determined by a linear regression - this referncing method is employed by Agnus. This referencing method is also unaffected by <tt>lockfreq</tt> reset (see below).

== Solvent dependence ==

Deuterons in different solvents resonante at different ppm positions. The lock system, however, would adjust them to the same predefined frequency. Thus, a locked sample in D2O will experience a different B0 field than a locked sample in C6D6.

To make sure that the carrier offsets (<tt>tof</tt>, <tt>dof</tt>, <tt>dof2</tt>) remain unchanged, the lock solvent has to be specfied by setting the <tt>solvent</tt> variable. For example, for protein samples <tt>solvent='d2o'</tt>. For proper names of other lock solvents see the <tt>/vnmr/solventlist</tt> file in your VNMR or VNMRJ installation

== Resetting Lock Frequency ==

Locking requires setting <tt>z0</tt> so that the deuterium resonance of the solvent (D2O in protein NMR) matches the lock frequency <tt>lockfreq</tt> of the spectrometer. One day you may find that the lock resonance is beyond the operating range of <tt>z0</tt>. In this case it is necessary to reset <tt>lockfreq</tt> to a lower value. This parameter has to be changed on 500MHz, 600MHz and 750MHz instruments in Buffalo about once or twice a year depending on the field drift.

Run <tt>config</tt> to open a spectrometer configuration window and key in the new frequency there, then type <tt>su</tt> to enable the new setting. Record this event in the logbook and inform other users of the instrument. You must be running VNMR or VNMRJ as <tt>vnmr1</tt> users to do that. Regular users do not have permissions to modify configuration.

There is always an issue of how big the change in <tt>lockfreq</tt> should be. Obviously, it should not be so small that you will have to do it again next week, and should not be too large that your <tt>z0</tt> is out of range in on the opposite side.

For example, for both 600 MHz and 750 MHz spectrometers in Prof. Szyperski's lab one unit of <tt>z0</tt> is equivalent to approximately 0.05 Hz. Since <tt>z0</tt> has a range of -32767 to 32767, <tt>lockfreq</tt> can be safely changed by about 2700 Hz. Field drift rate of 600 MHz spectrometer was found to be ~315 <tt>z0</tt> units/day or ~16 Hz/day. The drift rate of 750 MHz is ~100 <tt>z0</tt> units/day or ~5 Hz/day. Therefore, <tt>lockfreq</tt> needs to be reset about twice a year on 600 MHz and once every year and a half on 750 MHz.

'''DO NOT''' type something like <tt>lockfreq = 99.999</tt> at the prompt. This would only be valid until you restart VNMR or VNMRJ and is grossly confusing.

'''IMPORTANT!''' Changing <tt>lockfreq</tt> significantly affects referencing with DSS, since you change the B0 field of the spectrometer. '''Always''' record a 1D DSS spectrum with a new <tt>lockfreq</tt>.

 

== Working with Bruker Instruments ==

The following description pertains to systems with Bruker TopSpin v. 2.0 and higher. 

After inserting the sample, the user opens the lock display using the command:
<pre>lockdisp
</pre>
This open the lock display (see below).  If unlocked, one observes the ringing signal (left); if locked, then the lock signal is a flat trace (right).

[[Image:Lock unlocked.jpg|400px]][[Image:Lock locked.jpg|400px]]

With the system unlocked:

*Manually adjust the "Field" and "Phase" settings on the BSMS panel, to make the unlocked signal symmetric and centered within the display.
*Then press the "lock" button on the BSMS panel to lock on deuterium.

With the system locked:

*Manually optimize the shims (BSMS panel) by optimizing the height of the lock signal prior to automated shimming routines.

 

 

== ==

 -- Main.AlexEletski - 24 Jan 2007

Inserting NMR Sample

2009-12-09T16:08:00Z

Aringh:

__TOC__

== Adjusting sample depth with a sample gauge ==

[[Image:Gauge sch.jpg|right|100px]]First, the NMR tube is needed to be inserted into a spinner turbine. For best results, NMR tubes should be positioned in the spinner turbine in such a way that the sample remains centered in the r.f. coil region inside the NMR probe. This can be done with the help of a sample gauge, which usually has a center line and rectangular markings to indicate the probe geometry. 

A sample gauge also has an adjustable bottom, which can be adjusted to allow quick positioning of a series of identical samples. Setting a sample tube too low may cause it to break, as well as contaminate and damage the probe!

It is also important to make sure that the tube sits firmly in the spinner. If your sample slides too easily inside a spinner turbine, it may indicate that the rubber o-rings of the spinner need replacement.

== Inserting and ejecting ==

To access the '''Insert''' and '''Eject''' buttons do the following: 

*VNMR - click on the '''Acqi''' button or type <tt>acqi</tt> 
*VNMRJ - click on the '''Setup''' tab

You can also type <tt>insert</tt> or <tt>i</tt> and <tt>eject</tt> or <tt>e</tt> at the command prompt in VNMR or VNMRJ.

Sample change procedure

#Eject the current sample first. Even if there is no sample in the magnet you need to use eject to create the upward thrust using the airflow.
#Remove the ejected sample. 
#Place the sample in the spinner in the bore. Verify that you have the airflow to support your sample while still holding the sample. Leave the sample.
#Click on Insert to lower your sample. 

Avoid leaving the spectrometer with an open bore for long periods of time, as this increases the risk of contamination of the bore with foreign particles. If you need to remove a sample and have nothing else to measure, either insert a standard NMR sample or cover the bore with a dedicated plug.

 

== External links ==

http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199916100c.pdf

http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199931300b.pdf

 

== Working with Bruker Instruments ==

As with Varians, the sample tube is loaded into a spinner and the depth is adjusted using a depth gauge.  The base of the depth gauge should be set to the minimum sample tube depth allowed by the probe in use.  DO NOT adjust this level.  The minimum volume depth for most Bruker probes is normally smaller than that for Varians, as a result it is important to use Shigemi tubes suitable for use with Bruker probes (they will have a shorter glass base).

 

Samples are manually ejected and inserted into the magnet using the sample eject/insert button on the BSMS panel; the button is located in the top left corner of the BSMS.  Samples can also be ejected using the command line in Topspin (command, ej). 

The procedure for inserting the NMR sample into the magnet is analogous to Varians:

#Eject the current sample.  Pressing the eject button on the BSMS (green light comes on) will initiate an upward airflow and the current sample will float to the top of the bore.
#Replace the current sample with the new sample.
#Pressing the eject button again on the BSMS (green light turns off) stops the airflow and the sample is slowly lowered into the magnet. 
#Look for the sample lock to ensure the sample is properly sitting in the probe.

Inserting NMR Sample

2009-12-08T22:35:04Z

Aringh:

__TOC__

== Adjusting sample depth with a sample gauge ==

[[Image:Gauge sch.jpg|right|100px]]First, the NMR tube is needed to be inserted into a spinner turbine. For best results, NMR tubes should be positioned in the spinner turbine so that the sample volume will be centered in the NMR probe. This can be done with the help of a sample gauge, which usually has a center line and rectangular markings to indicate the probe geometry. 

A sample gauge also has an adjustable bottom, which can be adjusted to allow quick posistioning of a series of identical samples. Setting a sample tube too low may cause it to break, as well as contaminate and damage the probe!

It is also important to make sure that the tube sits firmly in the spinner. If your sample slides too easily inside a spinner turbine, it may indicate that its rubber o-rings need replacement.

== Inserting and ejecting ==

To access the '''Insert''' and '''Eject''' buttons do the following: 

*VNMR - click on the '''Acqi''' button or type <tt>acqi</tt> 
*VNMRJ - click on the '''Setup''' tab

You can also type <tt>insert</tt> or <tt>i</tt> and <tt>eject</tt> or <tt>e</tt> at the command prompt in VNMR or VNMRJ.

Sample change procedure

#Eject the current sample first. Even if there is no sample in the magnet you need to use eject to create airflow.
#Remove the ejected sample. 
#Verify that you have the airflow to support your sample. Carefully insert the sample in the bore.
#Click on Insert to lower your sample. 

Avoid leaving the spectrometer with an open bore for long periods of time, as this increases the risk contaminating the bore with forieign particles. If you need to remove a sample and have nothing else to measure, either insert a standard NMR sample or cover the bore with a dedicated plug.

 

== External links ==

http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199916100c.pdf

http://www.varianinc.com/image/vimage/docs/products/nmr/apps/pubs/manuals/0199931300b.pdf

 

== Working with Bruker Instruments ==

As with Varians, the sample tube is loaded into a spinner and the depth is adjusted using a depth gauge.  The base of the depth gauge should be set to the minimum sample tube depth allowed by the probe in use.  DO NOT adjust this level.  The minimum volume depth for most Bruker probes is normally smaller than that for Varians, as a result it is important to use Shigemi tubes suitable for use with Bruker probes (they will have a shorter glass base).

 

Samples are manually ejected and inserted into the magent using the sample eject/insert button on the BSMS panel; the button is located in the top left corner of the BSMS.  Samples can also be ejected using the command line in Topspin (command, ej). 

The procedure for inserting the NMR sample into the magnet is analogous to Varians:

#Eject the current sample.  Pressing the eject button on the BSMS (green light comes on) will initiate an airflow and the current sample will float to the top of the bore.
#Replace the current sample with the new sample.
#Pressing the eject button again on the BSMS (green light turns off) stops the airflow and the sample is slowly lowered into the magnet. 
#Look for the sample lock to ensure the sample is properly sitting in the probe.

NMR Sample Preparation

2009-12-08T22:33:18Z

Aringh:

== '''Sample Transfer into NMR Tubes''' ==

Although our lab seldom produces NMR samples we frequently have to transfer the samples to an NMR tube. According to different requirements, three kinds of NMR tubes are generally used in the Szyperski Lab: regular 5 mm NMR tubes, Shigemi tubes and capillary tubes. Each kind of tube has its own special sample preparation procedure.

=== '''Regular and Shigemi Tubes''' ===

Regular NMR tubes:

*Pros
**Easy to transfer or titrate sample.
**Low price.
**Durable.
**Can be used with any spectrometer brand.
*Cons
**Requires large sample volumes - 0.4 to 0.6 ml.
**Suboptimal shimming, consequently, water suppression and sensitivity are poor (and smaller the volume, worse is the effect).
**Sample is subject to air oxidation.

Shigemi NMR tubes:

*Pros
**Ideal for small sample volumes: 0.2 to 0.3 ml
**Optimal shimming and water suppression
**Better sample stability due to reduced air oxidation
*Cons
**Complicated transfer and handling.
**High price.
**Very brittle - easy to break.
**Different tube types for different spectrometer brands.

In general, it is preferable to use Shigemi tubes for the [100% 13C, 100% 15N]-labeled samples. For [5% 13C, 100% 15N]-labeled samples, regular tubes are also fine, since these samples are used to record in general the 13C-HSQC spectrum of the methyl region, where water suppression is not an issue.

NOTE : It is advised to use long-end pipettes to transfer sample solution into and out of regular 5 mm NMR tubes and Shigemi tubes.

==== '''Protocol to transfer sample in a regular tube:''' ====

Fig.1 Sample in regular NMR tubes

[[Image:Sample1.jpg]]

#Place the sample at the bottom of the tube and cap the tube.
#Spin the tube in the hand centrifuge to collect residual sample from the walls and colapse the air pockets and bubbles, if any. Wrap the tube in a tissue for padding before inserting it into a holder of the centrifuge.
#If a part of the sample collected in the cap, transfer it into the tube and spin the sample again.
#Wrap a thin strip of Parafilm around the seam between the tube and its cap to prevent the sample from drying out.
#Put a label on the tube.

==== '''Protocol to transfer sample in a Shigemi tubes:''' ====

Fig.2 Sample in Shigemi tubes

[[Image:Sample2.jpg]]

 The Shigemi tube is composed of two parts: the outer tube and the insert (also called plunger). The insert is made of a special type of glass with magnetic susceptibility matching to that of the solvent (H2O for protein NMR). The matching magnetic susceptibility removes the edge effect at the interface of the sample and the glass, thereby improving the shimming. The whole set is expensive (~ $80 per set) and the glass type is more brittle as compared to the regular tubes. Extra care should therefore be taken while handling Shigemi tubes.

 

#Place the sample at the bottom of the tube and cap the tube.
#Spin it in the hand centrifuge to collect residual sample from the walls and collapse the air pockets and bubbles, if any. Wrap the tube in a tissue for padding before inserting it into a holder of the centrifuge.
#If a part of the sample collected in the cap, transfer it into the tube and spin the sample again.
#Slowly push the insert down the tube to drive the air out.
#At this point, it is common to get bubbles under the insert. To get rid of them, hold the tube at an angle (and maybe knock on it a few times) - the bubbles should then settle at the interface between the insert bottom and the tube wall. Push forward quickly, but gently, and simultaneously rotate the insert ('''I have never tried rotating - sounds a bit complicated. Does it work better this way?''' Main.AlexEletski).
#If you have a lot of bubbles (or foam) in the reservoir then let the sample stand still for some time. After a while the bubbles will rise on their own. Keep the inset in place by wrapping a thin strip of Parafilm around the tube rim.
#Pull the insert out as far as possible without letting any air back in.
#Repeat steps 5 - 7. if necessary.
#Wrap a thin strip of Parafilm around the tube rim. This should keep the insert in place as well as prevent the sample from drying out.
#Label the tube

'''Key notes:'''

*Rotate the insert when moving it in or out to reduce the resistance.
*Be careful not to drive the insert too far, especially when pushing air bubbles out. The neck of the insert forms a reservoir in the tube, but once it is full the sample will leak out.
*When immersed in liquid, the insert may easily slide under it own gravity. Therefore, always fix the insert with Parafilm when leaving the sample unattended
*It is important to push air bubbles out. Even the tiniest bubbles can lead to bad shimming and broad water lines - and that is exactly what Shigemi tubes are designed to avoid.
*Waiting for foamy solution to settle down before pulling up the insert can be crucial for maximizing the sample height. With certain viscous solutions, such as phage-based alignment media, the time required may be as long as several hours.

===== '''Maximum Volume''' =====

Varian tubes (BMS-005TV) have bottom length of 15-16 mm, leaving 16 mm as the maximul sample depth. This corresponds to 260 ul at 4.52 mm inner diameter. You still need to have some liquid filling part of the reservoir, meaning that you probably have to put 280-300 ul into the tube.

It is possible to use Bruker shigemi tubes with bottom length of 8 mm in Varian spectrometers. In this case max sample depth would be 24 mm, corresponding to 385 ul.

=== '''Capillary Tubes''' ===

We use 1.0mm and 1.7mm O.D. tubes from Wilmad. The capillary action (with a little help) is always sufficient to draw the solution into the tube. Normally, aliquot ~30uL of solution into a small Eppendorf tube, dip the capillary in, and tip or invert the whole setup if the solution does not completely go into the capillary. If needed, you can apply some suction from a syringe or pipette or even 'inject' the solution into the capillary with a syringe and very thin needle although these methods have a higher risk of squirting your solution onto the bench.

When sealing the tubes, first start with a good length of capillary tube (almost as long as the NMR tube) and put sample solution in it. Then tip the capillary until the solution moves to the middle of the tube. Now it is safe to seal the end of the tube in a Bunsen burner flame -- place only the very tip of the tube into the flame. To avoid the possibility of the protein heat-denaturing while sealing in case of insufficient space at the end of tube, wrapping the tube with some form of a 'heat-sink' (like a wet towel placed in a freezer) can be tried. After sealing one end of the tube, centrifuge the whole set-up (place sealed end into the centrifuge) by using a small hand-operated centrifuge for a few seconds to spin down the solution into the sealed end of the capillary tube. Seal the other end of the tube. As the second end begins to seal, essentially a closed container is heated. Normally, bubbles will generate on one end of the tube.

Some tips for handling capillary tubes and lowering the sample temperature:

#Don't use capillaries that have been broken on both ends (i.e., keep at least one machined end of glass to prevent particles of glass getting into your solution)
#Carefully clean and dry all of the capillaries
#Try to keep the tubes perfectly parallel to the magnetic field by using empty tubes or small pieces of paper wrapped around the top of the capillary tube to keep everything aligned and motionless in the NMR tube
#Try a few times with H2O/D2O and see how it goes
#Before you start, centrifuge the solution at ~14000g for ~30minutes
#Use a 1D (no water suppression) and array the temp from -5C to -20C in -0.1C steps to monitor the cooling. (if one out of ten capillaries freeze, then a decrease in the intensity of the water line could be observed)
#Don't spin the sample or bump the spectrometer during the cooling progress.

 

== '''Short-term and Long-term NMR Sample Storage''' ==

All NMR samples should be labeled and stored at 4C or -20C. The label should include the sample name and concentration, all buffer components, percentage of H2O:D2O, initials of the preparer and the date prepared. This should be neatly printed of prepared with a word processing program. You may also consider a code that is cross referenced to your lab journal giving more detailed information about the sample preparation and the sample conditions.

The optimal method for short-term sample storage, provided the sample is stable and sodium azide is present, is to simply keep them at 4C. There are plenty of 10 mm tube racks in the refrigerator. For valuable samples you should place the NMR tubes inside a second container, e.g., a 10mm capped NMR tube or a homemade container that will serve to catch any spilled sample in the event of glass breakage. This doubly protected NMR sample tube should then be placed in a 10 mm rack.

The optimal method for long-term sample storage, provided they are stable upon freezing, is to transfer the solution to a 1.5 mL plastic Eppendorf tube and then freeze at -20C. The sample can also be lyophilized at this stage. Lyophilized protein (r.t. or -20C) is probably the best condition to store biological sample for extended periods of time.

Precautions: DO NOT freeze samples directly in the glass NMR tubes, particularly in expensive Shigemi tubes. The risk of glass breakage and sample loss is high. If you need to freeze an NMR sample directly in the tube, then the following precautions must be taken: The liquid should be spread uniformly on the glass surface prior to freezing in order to prevent breaking of the thin glass walls; The NMR tube containing a frozen sample should be placed in a clean container, e.g., a 10 mm capped NMR tube or a homemade container that will serve to catch any spilled sample in the event of glass breakage; The sample must be stored in a 10 mm rack provided for this purpose.

 

 -- Main.Gaohua.Liu - 11 Feb 2007

[[Category:NMR_Acquisition]]

NMR Sample Preparation

2009-12-08T22:11:49Z

Aringh:

== '''Sample Transfer into NMR Tubes''' ==

Although our lab seldom produces NMR samples we frequently have to transfer the samples to an NMR tube. According to different requirements, three kinds of NMR tubes are generally used in the Szyperski Lab: regular 5 mm NMR tubes, Shigemi tubes and capillary tubes. Each kind of tube has its own special sample preparation procedure.

=== '''Regular and Shigemi Tubes''' ===

Regular NMR tubes:

*Pros
**Easy to transfer or titrate sample.
**Low price.
**Durable.
**Can be used with any spectrometer brand.
*Cons
**Requires large sample volumes - 0.4 to 0.6 ml.
**Suboptimal shimming, consequently, water suppression and sensitivity are poor (and smaller the volume, worse is the effect).
**Sample is subject to air oxidation.

Shigemi NMR tubes:

*Pros
**Ideal for small sample volumes: 0.2 to 0.3 ml
**Optimal shimming and water suppression
**Better sample stability due to reduced air oxidation
*Cons
**Complicated transfer and handling.
**High price.
**Very brittle - easy to break.
**Different tube types for different spectrometer brands.

In general, it is preferable to use Shigemi tubes for the [100% 13C, 100% 15N]-labeled samples. For [5% 13C, 100% 15N]-labeled samples, regular tubes are also fine, since these samples are used to record in general the 13C-HSQC spectrum of the methyl region, where water suppression is not an issue.

NOTE : It is advised to use long-end pipettes to transfer sample solution into and out of regular 5 mm NMR tubes and Shigemi tubes.

==== '''Protocol to transfer sample in a regular tube:''' ====

Fig.1 Sample in regular NMR tubes

[[Image:Sample1.jpg]]

#Place the sample at the bottom of the tube and cap the tube.
#Spin the tube in the hand centrifuge to collect residual sample from the walls and colapse the air pockets and bubbles, if any. Wrap the tube in a tissue for padding before inserting it into a holder of the centrifuge.
#If a part of the sample collected in the cap, transfer it into the tube and spin the sample again.
#Wrap a thin strip of Parafilm around the seam between the tube and its cap to prevent the sample from drying out.
#Put a label on the tube.

==== '''Protocol to transfer sample in a Shigemi tubes:''' ====

Fig.2 Sample in Shigemi tubes

[[Image:Sample2.jpg]]

 The Shigemi tube is composed of two parts: the outer tube and the insert (also called plunger). The insert is made of a special type of glass with magnetic susceptibility matching to that of the solvent (H2O for protein NMR). The matching magnetic susceptibility removes the edge effect at the interface of the sample and the glass, thereby improving the shimming. The whole set is expensive (~ $80 per set) and the glass type is more brittle as compared to the regular tubes. Extra care should therefore be taken while handling Shigemi tubes.

 

#Place the sample at the bottom of the tube and cap the tube.
#Spin it in the hand centrifuge to collect residual sample from the walls and collapse the air pockets and bubbles, if any. Wrap the tube in a tissue for padding before inserting it into a holder of the centrifuge.
#If a part of the sample collected in the cap, transfer it into the tube and spin the sample again.
#Slowly push the insert down the tube to drive the air out.
#At this point, it is common to get bubbles under the insert. To get rid of them, hold the tube at an angle (and maybe knock on it a few times) - the bubbles should then settle at the interface between the insert bottom and the tube wall. Push forward quickly, but gently, and simultaneously rotate the insert ('''I have never tried rotating - sounds a bit complicated. Does it work better this way?''' Main.AlexEletski).
#If you have a lot of bubbles (or foam) in the reservoir then let the sample stand still for some time. After a while the bubbles will rise on their own. Keep the inset in place by wrapping a thin strip of Parafilm around the tube rim.
#Draw the sample out as far as possible without letting any air back in.
#Repeat steps 5 - 7. if necessary.
#Wrap a thin strip of Parafilm around the tube rim. This should keep the insert in place as well as prevent the sample from drying out.
#Label the tube

Key notes:

*Rotate the insert when moving it in or out to reduce the resistance.
*Be careful not to drive the insert too far, especially when pushing air bubbles out. The neck of the insert forms a reservoir in the tube, but once it is full the sample will leak out.
*When immersed in liquid, the insert may easily slide under it own gravity. Therefore, always fix the insert with Parafilm when leaving the sample unattended
*It is important to push air bubbles out. Even the tiniest bubbles can lead to bad shimming and broad water lines - and that is exactly what Shigemi tubes were designed to avoid.
*Waiting for foamy solution to settle down before pulling up the insert can be crucial for maximizing the sample height. With certain viscous solutions, such as phage-based alignment media, the time required may be as long as several hours.

===== '''Maximum Volume''' =====

Varian tubes (BMS-005TV) have bottom length of 15-16 mm, leaving 16 mm as the maximul sample depth. This corresponds to 260 ul at 4.52 mm inner diameter. You still need to have some liquid filling part of the reservoir, meaning that you probably have to put 280-300 ul into the tube.

It is possible to use Bruker shigemi tubes with bottom length of 8 mm in Varian spectrometers. In this case max sample depth would be 24 mm, corresponding to 385 ul.

=== '''Capillary Tubes''' ===

We use 1.0mm and 1.7mm O.D. tubes from Wilmad. The capillary action (with a little help) is always sufficient to draw the solution into the tube. Normally, aliquot ~30uL of solution into a small Eppendorf tube, dip the capillary in, and tip or invert the whole setup if the solution does not completely go into the capillary. If needed, you can apply some suction from a syringe or pipette or even 'inject' the solution into the capillary with a syringe and very thin needle although these methods have a higher risk of squirting your solution onto the bench.

When sealing the tubes, first start with a good length of capillary tube (almost as long as the NMR tube) and put sample solution in it. Then tip the capillary until the solution moves to the middle of the tube. Now it is safe to seal the end of the tube in a Bunsen burner flame -- place only the very tip of the tube into the flame. To avoid the problems with the protein heat-denaturing if there is no enough space on the ends of tubes, try wrapping the tube in a 'heat-sink' (like a wet towel placed in a freezer). After sealing one end of the tube, centrifuge the whole set-up (place sealed end into the centrifuge) by using a small hand-operated centrifuge for a few seconds to spin down the solution into the sealed end of the capillary tube. Seal the other end of the tube. As the second end begins to seal, essentially a closed container is heated. Normally, bubbles will generate on one end of the tube.

Some tips for handling capillary tubes:

#Don't use capillaries that have been broken on both ends (i.e., keep at least one machined end of glass to prevent particles of glass getting into your solution)
#Carefully clean and dry all of the capillaries
#Try to keep the tubes perfectly parallel to the magnetic field by using empty tubes or small pieces of paper wrapped around the top of the capillary tube to keep everything aligned and motionless in the NMR tube
#Try a few times with H2O/D2O and see how it goes
#Before you start, centrifuge the solution at ~14000g for ~30minutes
#Use a 1D (no water suppression) and array the temp from -5C to -20C in -0.1C steps to monitor the cooling. (if one out of ten capillaries freeze, then a decrease in the intensity of the water line could be observed)
#Don't spin the sample or bump the spectrometer during the cooling progress.

 

== '''Short-term and Long-term NMR Sample Storage''' ==

All NMR samples should be labeled and stored at 4C or -20C. The label should include the sample name and concentration, all buffer components, percentage of H2O:D2O, initials of the preparer and the date prepared. This should be neatly printed of prepared with a word processing program. You may also consider a code that is cross referenced to your lab journal giving more detailed information about the sample preparation and the sample conditions.

The optimal method for short-term sample storage provided the sample is stable and sodium azide is present, is to simply place them at 4C. There are plenty of 10 mm tube racks in the refrigerator. For valuable samples you should place the NMR tubes inside a second container, e.g., a 10mm capped NMR tube or a homemade container that will serve to catch any spilled sample in the event of glass breakage. The latter, double-protected NMR samples should then be placed in a 10 mm rack.

The optimal method for long-term sample storage, provided they are stable to freezing, is to transfer the solution to a 1.5 mL plastic Eppendorf tube and then freeze at -20C. The sample can also be lyophilized at this stage. Lyophilized protein (r.t. or -20C) is probably the best condition to store biological sample for extended periods of time.

Precautions: DO NOT freeze samples directly in the glass NMR tubes, particularly in expensive Shigemi tubes. The risk of glass breakage and sample loss is high. If you need to freeze an NMR sample directly in the tube, then the following precautions must be taken: The liquid should be spread uniformly on the glass surface prior to freezing in order to prevent breaking of the thin glass walls; The NMR tube containing a frozen sample should be placed in a clean container, e.g., a 10 mm capped NMR tube or a homemade container that will serve to catch any spilled sample in the event of glass breakage; The sample must be stored in a 10 mm rack provided for this purpose.

 

 -- Main.Gaohua.Liu - 11 Feb 2007

[[Category:NMR_Acquisition]]

NMR Sample Preparation

2009-12-08T20:40:11Z

Aringh:

== '''Sample Transfer into NMR Tubes''' ==

Although our lab seldom produces NMR samples we frequently have to transfer the samples to an NMR tube. According to different requirements, three kinds of NMR tubes are generally used in the Szyperski Lab: regular 5 mm NMR tubes, Shigemi tubes and capillary tubes. Each kind of tube has its own special sample preparation procedure.

=== '''Regular and Shigemi Tubes''' ===

Regular NMR tubes:

*Pros
**Easy to transfer or titrate sample.
**Low price.
**Durable.
**Can be used with any spectrometer brand.
*Cons
**Requires large sample volumes - 0.4 to 0.6 ml.
**Suboptimal shimming, consequently, water suppression and sensitivity are poor (and smaller the volume, worse is the effect).
**Sample is subject to air oxidation.

Shigemi NMR tubes:

*Pros
**Ideal for small sample volumes: 0.2 to 0.3 ml
**Optimal shimming and water suppression
**Better sample stability due to reduced air oxidation
*Cons
**Complicated transfer and handling.
**High price.
**Very brittle - easy to break.
**Different tube types for different spectrometer brands.

In general, it is preferable to use Shigemi tubes for the [100% 13C, 100% 15N]-labeled samples. For [5% 13C, 100% 15N]-labeled samples regular tubes are also fine, since they are used to record in general the 13C-HSQC of the methyl region, where water suppression is not an issue.

Use long-end pipettes to transfer sample solution into and out of regular 5 mm NMR tubes and Shigemi tubes.

==== '''Regular Tubes''' ====

Fig.1 Sample in regular NMR tubes

[[Image:Sample1.jpg]]

#Transfer the sample to the bottom of the tube and cap the tube.
#Spin the tube in the hand centrifuge to collect residual sample from the walls and colapse air pocket and bubbles, if any. Wrap the tube in a tissue for padding before inserting it into a holder of the centrifuge.
#If a part of the sample collected in the cap, transfer it into the tube and spin the sample again.
#Wrap a thin strip of Parafilm around the seam between the tube and its cap to prevent the sample from drying out.
#Put a label on the tube.

==== '''Shigemi Tubes''' ====

Fig.2 Sample in Shigemi tubes

[[Image:Sample2.jpg]]

 The Shigemi tube is composed of two parts: the outer tube and the insert (also called plunger) made of a special type of glass whose magnetic susceptibility is matched to that of the solvent (H2O for protein NMR). The whole set is expensive (~ $80 per set) and the glass type is more brittle type than that of regular tubes. Therefore, extra care should be taken when handling Shigemi tubes.

 

#Transfer the sample to the bottom of the tube and cap the tube.
#Spin it in the hand centrifuge to collect residual sample from the walls and collapse air pocket and bubbles, if any. Wrap the tube in a tissue for padding before inserting it into a holder of the centrifuge.
#If a part of the sample collected in the cap, transfer it into the tube and spin the sample again.
#Slowly push the insert down the tube to drive the air out.
#At this point, you will likely get bubbles under the insert. To get rid of them, hold the tube at an angle (and maybe knock on it a few times) - the bubbles should then settle at the interface between the insert bottom and the tube wall. Push forward quickly, but gently, and simultaneously rotate the insert ('''I have never tried rotating - sounds a bit complicated. Does it work better this way?''' Main.AlexEletski).
#If you have a lot of bubbles (or foam) in the reservoir then let the sample stand still for some time. After a while the bubbles will rise on their own. Keep the inset in place by wrapping a thin strip of Parafilm around the tube rim.
#Draw the sample out as far as possible without letting any air back in.
#Repeat steps 5 - 7. if necessary.
#Wrap a thin strip of Parafilm around the tube rim. This should keep the insert in place as well as prevent the sample from drying out.
#Label the tube

Key notes:

*Rotate the insert when moving it in or out to reduce the resistance.
*Be careful not to drive the insert too far, especially when pushing air bubbles out. The neck of the insert forms a reservoir in the tube, but once it is full the sample will leak out.
*When immersed in liquid, the insert may easily slide under it own gravity. Therefore, always fix the insert with Parafilm when leaving the sample unattended
*It is important to push air bubbles out. Even the tiniest bubbles can lead to bad shimming and broad water lines - and that is exactly what Shigemi tubes were designed to avoid.
*Waiting for foamy solution to settle down before pulling up the insert can be crucial for maximizing the sample height. With certain viscous solutions, such as phage-based alignment media, the time required may be as long as several hours.

===== '''Maximum Volume''' =====

Varian tubes (BMS-005TV) have bottom length of 15-16 mm, leaving 16 mm as the maximul sample depth. This corresponds to 260 ul at 4.52 mm inner diameter. You still need to have some liquid filling part of the reservoir, meaning that you probably have to put 280-300 ul into the tube.

It is possible to use Bruker shigemi tubes with bottom length of 8 mm in Varian spectrometers. In this case max sample depth would be 24 mm, corresponding to 385 ul.

=== '''Capillary Tubes''' ===

We use 1.0mm and 1.7mm O.D. tubes from Wilmad. The capillary action (with a little help) is always sufficient to draw the solution into the tube. Normally, aliquot ~30uL of solution into a small Eppendorf tube, dip the capillary in, and tip or invert the whole setup if the solution does not completely go into the capillary. If needed, you can apply some suction from a syringe or pipette or even 'inject' the solution into the capillary with a syringe and very thin needle although these methods have a higher risk of squirting your solution onto the bench.

When sealing the tubes, first start with a good length of capillary tube (almost as long as the NMR tube) and put sample solution in it. Then tip the capillary until the solution moves to the middle of the tube. Now it is safe to seal the end of the tube in a Bunsen burner flame -- place only the very tip of the tube into the flame. To avoid the problems with the protein heat-denaturing if there is no enough space on the ends of tubes, try wrapping the tube in a 'heat-sink' (like a wet towel placed in a freezer). After sealing one end of the tube, centrifuge the whole set-up (place sealed end into the centrifuge) by using a small hand-operated centrifuge for a few seconds to spin down the solution into the sealed end of the capillary tube. Seal the other end of the tube. As the second end begins to seal, essentially a closed container is heated. Normally, bubbles will generate on one end of the tube.

Some tips for handling capillary tubes:

#Don't use capillaries that have been broken on both ends (i.e., keep at least one machined end of glass to prevent particles of glass getting into your solution)
#Carefully clean and dry all of the capillaries
#Try to keep the tubes perfectly parallel to the magnetic field by using empty tubes or small pieces of paper wrapped around the top of the capillary tube to keep everything aligned and motionless in the NMR tube
#Try a few times with H2O/D2O and see how it goes
#Before you start, centrifuge the solution at ~14000g for ~30minutes
#Use a 1D (no water suppression) and array the temp from -5C to -20C in -0.1C steps to monitor the cooling. (if one out of ten capillaries freeze, then a decrease in the intensity of the water line could be observed)
#Don't spin the sample or bump the spectrometer during the cooling progress.

 

== '''Short-term and Long-term NMR Sample Storage''' ==

All NMR samples should be labeled and stored at 4C or -20C. The label should include the sample name and concentration, all buffer components, percentage of H2O:D2O, initials of the preparer and the date prepared. This should be neatly printed of prepared with a word processing program. You may also consider a code that is cross referenced to your lab journal giving more detailed information about the sample preparation and the sample conditions.

The optimal method for short-term sample storage provided the sample is stable and sodium azide is present, is to simply place them at 4C. There are plenty of 10 mm tube racks in the refrigerator. For valuable samples you should place the NMR tubes inside a second container, e.g., a 10mm capped NMR tube or a homemade container that will serve to catch any spilled sample in the event of glass breakage. The latter, double-protected NMR samples should then be placed in a 10 mm rack.

The optimal method for long-term sample storage, provided they are stable to freezing, is to transfer the solution to a 1.5 mL plastic Eppendorf tube and then freeze at -20C. The sample can also be lyophilized at this stage. Lyophilized protein (r.t. or -20C) is probably the best condition to store biological sample for extended periods of time.

Precautions: DO NOT freeze samples directly in the glass NMR tubes, particularly in expensive Shigemi tubes. The risk of glass breakage and sample loss is high. If you need to freeze an NMR sample directly in the tube, then the following precautions must be taken: The liquid should be spread uniformly on the glass surface prior to freezing in order to prevent breaking of the thin glass walls; The NMR tube containing a frozen sample should be placed in a clean container, e.g., a 10 mm capped NMR tube or a homemade container that will serve to catch any spilled sample in the event of glass breakage; The sample must be stored in a 10 mm rack provided for this purpose.

 

 -- Main.Gaohua.Liu - 11 Feb 2007

[[Category:NMR_Acquisition]]