=== '''Pulse Sequence (J H-N)''' ===

The sequence is modified from the fast HSQC sequence (gNfhsqc) in Varian BioPack).<ref>Tjandra N. et al, JACS (1996), 118, 6364-72</ref> 

A series of 2D experiments should be acquired with the J modulation delay (Delta) set to different values, preferably in an interleaved fashion. (Note that “Delta” is half the total modulation delay.)  For example, in Vnmr, this can be realized by:
<pre>Delta=(0.0005,0.002,0.0035,0.0062,0.0073,0.0084,0.0125,0.014)</pre>
which sets Delta from 0.0005 to 0.014, corresponding to J modulating delays from 1 to 28 msec. At least 3 modulation points are needed to extract the coupling constant, along with 15N T2 and a parameter related to pulse imperfection. It is recommended to take more than 3 points to get an estimation of experimental error. The maximum delay (28 msec in this case) should be bigger than 3/(2J) ( ~16 msec in case of JNH) to make sure the modulation curve crosses zero at least twice (Tjandra et al.). Our simulation shows that the modulation interval should not be close to 1/J (~10ms for JNH),  i.e. the stepsize for Delta should not be close to ~5 msec.  An example of the J-modulation experiment is shown below:

[[Image:Vnmrj-Jmod.png|Vnmrj-Jmod.png]]   

 

==== Considerations for best use of acquisition time ====

1. ni vs. nt

The acquisition time for the indirect dimension of this J-mod method can be shorter than that used for frequency separation methods such as IPAP. It just needs to be long enough to resolve the peaks of interest, and around 30ms is often reasonable. In our experience, linear prediction can be applied without causing noticeable errors.  The time saved from a smaller “ni” can be spent more productively on “nt” to increase S/N.

 2. number of modulation points vs. nt 

Our simulation shows that these two are equivalent, i.e., one can choose to spend more time averaging each spectrum, or one can use to the time to acquire more modulation points. The errors in both cases are comparable. 

==== Other general considerations ====

#Poor signal to noise (too few scans)
#Insufficient modulation points (at least 3, but >5 is suggested)
#Incorrect setting of Delta step size (should NOT be around 5ms)
#Incomplete modulation curve. (at least two zero crossings are required)

=== '''Data Processing''' ===

Data Processing Script: [[Media:NH-jmod4edit.txt|NH-jmod4edit.txt]] (change extension to .pl to run)

Detailed instructions for NH J-modulated data processing with NMRpipe and Sparky:[[Media:NHJ_README.txt|NHJ_README.txt]] 

 

1. Split FIDs

If the experiment is acquired in an interleaved fashion, the final fid file contains a series of 2D spectra and thus has to be split into n fids, where n is the number of modulation points. We can send the script for doing this upon request.

 

2. Process with NMRpipe 

The standard 2D processing for fast HSQC spectra will work. But make sure the same processing is applied to all spectra.

 

3.1 Convert to Sparky 

To convert an nmrpipe output file, say test.ft2, to a sparky file called test.ucsf, type: pipe2ucsf test.ft2 test.ucsf 

 

This should be applied to all 2D spectra from the J-modulation experiment. To load these ucsf files into a project, a convenient way is to go to the directory that contain these files, and issue: find `pwd`|grep ucsf|xargs sparky &

 

This way the full path is associated with each file and thus the project can be reloaded from any initial location after saving.

 

3.2 Convert to NMRViewJ (alternative)

For analysis in NMRViewJ, the experiment must be separated into distinct 2D files.  Each must be given a unique name; following the example above there will be 8 files, iso1.ft2 - iso8.ft2, in which iso1.ft2 was collected with a J-modulation delay of 4 ms and iso8.ft2 was collected with a 32 ms delay.  These files will be converted to NMRView format using the following command: nmrPipe -in iso1.ft2 | pipe2xyz -nv -out iso1.nv

 

=== Data Analysis ===

'''Sparky'''  

Create peaks in one of the spectra and center them by the “pc” command. Preferably the one with the shortest modulation delay which has the best S/N. Propagate the peaklist into other spectra. This can be done with the “pa” and “oc” commands with the source spectrum in the front, followed by “op” after bringing the destination spectrum to the front. Do not re-center the peaks in each spectrum. 

 

To save the peak lists containing the intensities, do an “lt”. In “Options”, check “Data Height” so that the output contains the intensities for each peak.

 

 

'''NMRViewJ '''

First, load the files into NMRViewJ.  Open and Draw iso1.nv (Datasets  Open and Draw Dataset).  Adjust the contour level to a reasonable level, and in the Attributes window (right click on spectrum  Attributes) go to the PeakPick tab, chose a name for the list (Protein1) and select the “Pick” button.  Using the cursor (right click, Cursors, select “PeakAdd” or “PeakDelete”) delete noise peaks or add low-intensity peaks. In a text editor, create a file that specifies the Jmod delays for each specific file.  This time file will present the unique portions of the 2D filenames and the delays (in s).  Remembering there are 8 files (iso1.ft2 - iso8.ft2) with delays from 1-28ms, the time file will look like this:

 

#1 0.001
#2 0.004
#3 0.007
#4 0.0124
#5 0.0146
#6 0.0168
#7 0.025
#8 0.028

 

The data analysis is performed using the Rate Analysis tool (Analysis  Rate Analysis).  In the “Prefix for matrix numbers” box, write the non-unique portion of the filenames (iso).  Select “.nv” for the suffix.  Select the peaklist (Protein1).  Select the button “Load Time File” and load the table created above.

 

To measure the intensity of each peak click the “Measure All” button.  The graph to the right will show points indicative of a decaying cosine function. You can click through the peaks to see each peak using the up and down arrows.  Unfortunately, the equation to fit this type of data has not yet been placed in NMRViewJ, but it will be available in a future version.  For now, the data should be saved using the "Save Table" option and will be analyzed separately.

 

=== '''Curve Fitting and RDC Calculation''' ===

Organize the data into a table format of Intensity vs. Delay for each peak: [[Media:Make_separate2.txt|make_separate2.txt]] (For Sparky format peaklists. Change extension to .pl to run)

With the data in a table format of Intensity vs Delay for each peak, the following equation may be fit to these data to determine the Jmod values: 

f(x) = c * (-a + cos(3.14159*2*j*x))*exp(-2*x*r) where ''a'' is a variable accounting for imperfection of pi pulses during the experiment (usually 0 < a < 0.05), ''c'' is the initial intensity of the peak, ''j'' is the one-bond H-N J-coupling value in Hz and ''r'' is the R2 relaxation rate. This equation may be fit individually or in a batch mode using any common curve-fitting algorithm.

Individual fitting: [[Media:FitR_1by1.txt|fitR_1by1.txt]] (Change extension to .pl to run)

Batch fitting: [[Media:FitR_all.txt|fitR_all.txt]] (Change extension to .pl to run)

 

To calculate RDCs, simply take the difference between the measured scalar couplings for the isotropic and aligned samples. The sign of the residual dipolar coupling is dependent on the gyromagentic ratios of the coupled nuclei. For 1H-15N couplings, a decrease in coupling upon alignment corresponds to a positive residual dipolar coupling.

Header to add to the isotropic peaklist: [[Media:Header-hsqc_ass.txt|header-hsqc_ass.txt]] (Change extension to .tab to use)

To calculate RDCs from isotropic and aligned coupling data: [[Media:I-jmod.txt|i-jmod.txt]] (Change extension to .tcl to run) 

=== References ===

<references />

 Updated by Hsiau-Wei Lee, 2011

Updated by Kari Pederson, 2013

2013-12-18T17:44:49Z

Kpederson: /* Data Analysis */

'''NMR Data Processing >  Jmod Measurement of RDCs'''

 

===== '''Pulse Sequence (J H-N)''' =====

The sequence is modified from the fast HSQC sequence (gNfhsqc) in Varian BioPack. For details see Tjandra N. et al, ''JACS ''(1996), 118, 6364-72.   

A series of 2D experiments should be acquired with the J modulation delay (Delta) set to different values, preferably in an interleaved fashion. (Note that “Delta” is half the total modulation delay.)  For example, in Vnmr, this can be realized by:
<pre>Delta=(0.0005,0.002,0.0035,0.0062,0.0073,0.0084,0.0125,0.014)</pre>
which sets Delta from 0.0005 to 0.014, corresponding to J modulating delays from 1 to 28 msec. At least 3 modulation points are needed to extract the coupling constant, along with 15N T2 and a parameter related to pulse imperfection. It is recommended to take more than 3 points to get an estimation of experimental error. The maximum delay (28 msec in this case) should be bigger than 3/(2J) ( ~16 msec in case of JNH) to make sure the modulation curve crosses zero at least twice (Tjandra et al.). Our simulation shows that the modulation interval should not be close to 1/J (~10ms for JNH),  i.e. the stepsize for Delta should not be close to ~5 msec.  An example of the J-modulation experiment is shown below:

[[Image:Vnmrj-Jmod.png|Vnmrj-Jmod.png]]   

 

===== '''General Comments''' =====

#Poor signal to noise (too few scans)
#Insufficient modulation points (at least 3, but >5 is suggested)
#Incorrect setting of Delta stepsize (should NOT be around 5ms)
#Incomplete modulation curve. (at least two zero crossings are required)

 

ni vs nt

 

 The acquisition time for the indirect dimension of this J-mod method can be shorter than that used for frequency separation methods such as IPAP. It just needs to be long enough to resolve the peaks of interest, and around 30ms is often reasonable. In our experience, linear prediction can be applied without causing noticeable errors.  The time saved from a smaller “ni” can be spent more productively on “nt” to increase S/N.

 

number of modulation points vs nt

 

 Our simulation shows that these two are equivalent, i.e., one can choose to spend more time averaging each spectrum, or one can use to the time to acquire more modulation points. The errors in both cases are comparable. 

 

===== '''Data Analysis''' =====

Data Processing Script: [[Media:NH-jmod4edit.txt|NH-jmod4edit.txt]] (change extension to .pl to run)

Detailed instructions for NH J-modulated data processing with NMRpipe and Sparky:  

Split FIDs

If the experiment is acquired in an interleaved fashion, the final fid file contains a series of 2D spectra and thus has to be split into n fids, where n is the number of modulation points. We can send the script for doing this upon request.

 

Process with nmrPipe

 

The standard 2D processing for fast HSQC spectra will work. But make sure the same processing is applied to all spectra.

 

Convert to Sparky

 

To convert an nmrpipe output file, say test.ft2, to a sparky file called test.ucsf, type:

 

pipe2ucsf test.ft2 test.ucsf 

 

This should be applied to all 2D spectra from the J-modulation experiment. To load these ucsf files into a project, a convenient way is to go to the directory that contain these files, and issue:

 

find `pwd`|grep ucsf|xargs sparky &

 

This way the full path is associated with each file and thus the project can be reloaded from any initial location after saving.

 

Convert to NMRViewJ 

 

For analysis in NMRViewJ, the experiment must be separated into distinct 2D files.  Each must be given a unique name; following the example above there will be 8 files, iso1.ft2 - iso8.ft2, in which iso1.ft2 was collected with a J-modulation delay of 4 ms and iso8.ft2 was collected with a 32 ms delay.  These files will be converted to NMRView format using the following command:

 

nmrPipe -in iso1.ft2 | pipe2xyz -nv -out iso1.nv

 

===== '''Spectral Measurments ''' =====

NMRViewJ 

 

 First, load the files into NMRViewJ.  Open and Draw iso1.nv (Datasets  Open and Draw Dataset).  Adjust the contour level to a reasonable level, and in the Attributes window (right click on spectrum  Attributes) go to the PeakPick tab, chose a name for the list (Protein1) and select the “Pick” button.  Using the cursor (right click, Cursors, select “PeakAdd” or “PeakDelete”) delete noise peaks or add low-intensity peaks.

 In a text editor, create a file that specifies the Jmod delays for each specific file.  This time file will present the unique portions of the 2D filenames and the delays (in s).  Remembering there are 8 files (iso1.ft2 - iso8.ft2) with delays from 1-28ms, the time file will look like this:

 

#1 0.001
#2 0.004
#3 0.007
#4 0.0124
#5 0.0146
#6 0.0168
#7 0.025
#8 0.028

 

 The data analysis is performed using the Rate Analysis tool (Analysis  Rate Analysis).  In the “Prefix for matrix numbers” box, write the non-unique portion of the filenames (iso).  Select “.nv” for the suffix.  Select the peaklist (Protein1).  Select the button “Load Time File” and load the table created above.

 To measure the intensity of each peak click the “Measure All” button.  The graph to the right will show points indicative of a decaying cosine function.   You can click through the peaks to see each peak using the up and down arrows.  Unfortunately, the equation to fit this type of data has not yet been placed in NMRViewJ, but it will be available in a future version.  For now, the data should be saved using the Save Table option and will be analyzed separately.

 

 

Sparky 

 

Create peaks in one of the spectra and center them by the “pc” command. Preferably the one with the shortest modulation delay which has the best S/N. Propagate the peaklist into other spectra. This can be done with the “pa” and “oc” commands with the source spectrum in the front, followed by “op” after bringing the destination spectrum to the front. It is not necessary to re-center the peaks in each spectrum. 

 

To save the peak lists containing the intensities, do an “lt”. In “Options”, check “Data Height” so that the output contains the intensities for each peak.

 

 

===== '''Curve Fitting''' =====

With the data in a table format of Intensity vs Delay for each peak, the following equation may be fit to these data to determine the Jmod values: 

f(x) = c * (-a + cos(3.14159*2*j*x))*exp(-2*x/t) where ''a'' is a variable accounting for imperfection of pi pulses during the experiment (usually 0 < a < 0.05), ''c'' is the initial intensity of the peak, ''j'' is the one-bond H-N J-coupling value in Hz and ''t'' is the T2 relaxation rate. This equation may be fit individually or in a batch mode using any common curve-fitting algorithm.

 Updated by Hsiau-Wei Lee, 2011

2013-06-27T17:29:36Z

Kpederson:

=== Brief Description ===

1H-15N residual dipolar couplings (RDCs) are easily acquired for the purpose of protein structure validation and refinement. The data can be collected on samples labeled only with 15N. Obtaining RDCs in two different alignment media greatly improve the quality of refinement and can aid in dimer structure determination; 500µl of a 0.5-1mM sample is usually sufficient for this purpose.

=== Data Acquisition and RDC Calculation ===

Refer to the linked pages for detailed descriptions of the [[Jmodulation Experiment RDC]] and [[HSQCTROSY RDC Measurement]] and RDC calculation methods. 

=== Alignment Media Preparation ===

==== Isotropic Sample ====

The first sample to be observed is an isotropic sample. You may want to dilute the isotropic sample by one third (to 75% of its original concentration) to match the concentration of most aligned samples. In the case of a weak dimer, this may be important. 

==== PEG Bicelle ====

Alignment of the protein sample in PEG(C12E5/hexanol) is used as a first alignment media because it produces primarily steric alignment (useful in dimer geometry predictions), the success rate is high, and it can be doped with other charged detergents to give a second alignment media .

Chemicals used for this preparation:

:'''Sigma Aldrich 76437''', Pentaethylene glycol monododecyl ether (C12E5 PEG)
:'''Sigma Aldrich H13303''', Hexanol
:'''Sigma Aldrich 855820''', Cetyltrimethylammonium bromide (CTAB)
:'''Sigma Aldrich O4003''', Sodium octyl sulfate (SOS)

The preparation procedure is as follows:

*Prepare the 16% PEG stock solution by first mixing 50µl of C12E5 (pentaethylene glycol monododecyl ether) with 200µl of buffer and 50µl of D2O by vortexing.<ref> Ruckert M and Otting G (2000), JACS, 122, 7793-7797</ref>
*Add approximately 16µl of hexanol to the stock solution, in aliquots of 2µl with vortexing after each addition. The solution will go from clear to milky, then to translucent and viscous with lots of bubbles. Continue to add hexanol until the solution goes clear again. If it becomes milky/turbid again, you have gone past the nematic phase.
*Sample Content (for 220 µL sample):

::145 µL of Protein stock solution 
::55 µL of 16% PEG stock solution
::20 µL of D2O
:::Final PEG concentration is 4.2%.

*Record the 2H splitting by running the s2pul expt with tn='lk' (for Varian instruments). The range of the splitting should be around +/-20Hz.

*PEG can be doped with either cetyltrimethyl ammonium bromide (CTAB) for positively charged proteins or sodium octyl sulphate (SOS) for negatively charged proteins. Charging the medium to be like the protein prevents association and gives higher resolution spectra. A suitable ratio of PEG:CTAB/SOS is ~30:1.

==== Pf1 Phage ====

Preparation of a Pf1 phage alignment sample is fairly straightforward.<ref>Hansen MR, Mueller L, Pardi A (1998), Nat Struct Biol, 5, 1065-1074</ref> The protein sample is diluted by the alignment medium.

Chemicals used for this preparation:

:'''ASLA Biotech P-50-P''', Pf1 phage 50 mg/mL

The preparation procedure is as follows:

*Start with a protein stock 0.5-1mM and a pf1 phage stock of 50 mg/mL. Prepare a sample of 12.5 mg/mL of phage.

*Sample Content (for 220 µL sample):

::145 µL of Protein stock solution 
::55 µL of Pf1 phage 50 mg/mL stock solution 
::20 µL of D2O
:::Final phage concentration is 12.5 mg/mL

*Record the 2H splitting by running the s2pul expt with tn='lk' (for Varian instruments). The range of the splitting should be around +/-8-10 Hz.

==== Polyacrylamide Gel (Compressed and Stretched) ====

The preparation of polyacrylamide gel samples is a two step process. First the gels must be polymerized, equilibrated to the correct pH and cut to the appropriate size before being dried. Second the gels are re-hydrated using the protein stock in the appropriate NMR tube. 

Chemicals used for this preparation:

:'''Bio-Rad 161-0144''', 40% Acrylamide/Bis solution 19:1
:'''Bio-Rad 161-0733''', 10X TBE
:'''Bio-Rad 161-0700''', APS
:'''Bio-Rad 161-0800''', TEMED
:'''Sigma Aldrich M7279-25G''', N,N'-methylenebisacrylamide (BIS)
:'''Sigma Aldrich 448281''', (3-Acrylamidopropyl)trimethylammonium chloride solution
:'''Sigma Aldrich 282731''', 2-Acrylamido-2-methyl-1-propanesulfonic acid

Other materials used for this preparation:

:'''Wilmad P-4.4965M-6.5135-0-0''', 4.4965mm +/-0.0065mm ID x 6.5135mm +/-0.0065mm. Ground polished and buffed OD. 40mm +/- 0.5mm long. Both ends wet saw cut only.

===== Preparation of Dried Acrylamide Gels =====

The preparation procedure is as follows:

*Prepare the positively and negatively charged 40% 19:1 bis:acrylamide solutions.
*Positively charged stock solution (2mL):

::31 mg of N,N'-methylenebisacrylamide (BIS)
::961 µL of (3-acrylamidopropyl)trimethylammonium chloride solution
::1039 µL of H2O

*Negatively charged stock solution (2mL):

::31 mg of N,N'-methylenebisacrylamide (BIS)
::802 mg of 2-acrylamido-2-methyl-1-propanesulfonic acid
::2000 µL of H2O

{| width="838" cellspacing="1" cellpadding="1" border="1"
|+ Standard formulas for preparing compressed and stretched polyacrylamide gels
|-
| Gel type
|  % acrylamide
|  % charged
| Vol. 40% charged stock solution (µL)
| Vol. 40% neutral stock solution (µL)
| Vol. 10x TBE (µL)
| Vol. 10% APS (µL)
| Vol. TEMED (µL)
| Vol. per casting tube (µL)
| Type of casting tube
|-
| charged compressed
| 7
| 50
| 43.75
| 43.75
| 400
| 7.5
| 5
| 140
| 2.8 mm ID  plastic
|-
| charged stretched
| 5
| 50
| 62.5
| 62.5
| 860
| 10
| 5
| 300
| 3.2 mm ID  glass
|-
| neutral stretched
| 5
| 0
| 0
| 125
| 860
| 10
| 5
| 490
| 4.5 mm ID  glass
|}

*Note: The APS solution should be prepared fresh and polymerization will begin as soon as TEMED is added. 

 

*Mix stock solutions of neutral and charged 40% acrylamide/bis 19:1 to achieve the desired overall charge ratio. 

*Dilute the mixtures 10x with TBE buffer (0.9 M TRIS, 0.9 M borate, 0.02 M EDTA, pH 8.2) to a final 7% or 5% concentration for compressed or stretched gels, respectively.

*Polymerization is initiated by the addition of ammoniumperoxide sulfate (APS) (0.15% or 0.1% for compressed or stretched, respectively) and tetramethylethylenediamine (TEMED) (0.1% or 0.05% for compressed or stretched, respectively). 

*Pipet the mixture into the casting tubes carefully, to avoid bubbles, and keep them 1-2 hrs, allowing polymerization to occur.
*Use a 200 µL pipet with a trimmed pipet tip (to avoid hitting the gel) and water to carefully push the compressed gels out of the casting tubes into a 1L Erlenmeyer flask filled with deionized water. For the stretched gels, use a 1000 µL pipet to carefully push the stretched gels out of the casting tubes into prepared racks (no more than 4-5 gels per rack) braced in 2L nalgene beakers filled with deionized water. 

*Wash the polymerized gels extensively in deionized water (two cycles over a period of 1 day). The gels will increase in size due to electro-osmotic swelling. Use cheesecloth to drain the water from the flask for the compressed gels. Be careful when draining and adding new deionized water not to damage the gels. The racks holding the stretched gels may be moved to a new 2L nalgene beaker with fresh deionized water. 
*Equilibrate the polymerized gels to the desired pH (to match the pH of the protein stock) by washing extensively in buffered solution (two cycles over a period of 1 day). The buffer should not contain only the major buffering species and no salts. 
*Wash the polymerized gels in deionized water overnight to allow them to swell to full size.
*Select the gels which have no cracks or imperfections and measure the diameter of the fully swollen gels and trim each gel to a length 5.7 times its diameter.

*Dry the gels over a 2 day period at room temperature on a teflon pan. 

===== Preparation of Aligned Compressed Gel Samples =====

*Sample content (compressed gel):
*:1 dried compressed gel equilibrated to the same pH as the protein stock
*:200 µL of Protein stock
*:20 µL of D2O 
*Measure a height of 12-13 mm for the sample volume in a 5 mm shigemi tube and mark with a sharpie. 
*Add one dried compressed gel to the tube, followed by the protein stock and D2O. 
*Insert the plunger to the marked height and hold in place with parafilm. 
*Allow the gel to swell in the fridge for 1 day. Inspect the final sample for cracks before collecting data. 

===== Preparation of Aligned Stretched Gel Samples =====

Chemicals used for this preparation:

:'''Sigma Aldrich 440272''', Dichlorodimethylsilane

Other materials required for this preparation:

:'''Wilmad 528-PP-2.8mm/Stem''', 528-PP NMR tube sealed to a 50mm long stem section (stem dimensions 2.8mm +/-0.025mm ID x wall 0.38mm +/-0.015mm). Overall length, 8 inches. Both ends open'''.'''

:'''Wilmad 528-PP-3.6mm/Stem''', 528-PP NMR tube sealed to a 50mm long stem section (stem dimensions 3.6mm +/-0.025mm ID x wall 0.38mm +/-0.015mm). Overall length, 8 inches. Both ends open.

:'''Wilmad 528-PP-3.9mm/Stem''', 528-PP NMR tube sealed to a 50mm long stem section (stem dimensions 3.9mm +/-0.025mm ID x wall 0.38mm +/-0.015mm). Overall length, 8 inches. Both ends open.

:'''Wilmad 5mm microprobe NMR tube style''', Upper tube 528-PP section 7 inches long sealed to a 50mm long stem section (427-PP). Both ends open.

*Sample content (stretched gel):
*:1 dried stretched gel equilibrated to the same pH as the protein stock
*:300 µL of Protein stock
*:20 µL of D2O
*Prepare an NMR tube by washing 3 times with dichlorodimethylsilane, then rinse with deionized water and dry.
*Place a dried stretched gel near the top of the 5mm end of the NMR tube and attach a syringe with tubing to the bottom of the tube.
*Carefully pipet in the protein stock and D2O allowing the solution's surface tension to hold it in place in the tube.
*Use the syringe to adjust the position of the solution, such that the gel is centered in the column of solution. Cap the NMR tube, and let swell at room temperature 1-2 days. Capping the tube may cause the solution to shift, it's position should be readjusted with the syringe, to keep the gel centered.

 

==== References ====

<references />

 

 

Updated by Hsiau-Wei Lee, 2011

Updated by Kari Pederson 2013