NMR Thermometry Part 2

Yesterday I wrote a part 1 of a blog post regarding a Nature paper from October 2013.  You might want to check out that post before reading this one.

http://sitspinnmr.blogspot.com/2014/01/nmr-thermometry.html

These two posts review/critique the paper "Thermal maps of gases in heterogeneous reactions" by Jarenwattananon et al.

http://www.ncbi.nlm.nih.gov/pubmed/24153305

http://www.nature.com/nature/journal/v502/n7472/full/nature12568.html

My post yesterday was an attempt to review the publication.  I tried to minimize my comments and let the authors speak for themselves.  Today I plan to air my grievances and concerns about this paper.  

Before I begin I want to note that this paper got tons of press.  For example ..

http://www.nature.com/news/magnetic-map-1.13992

http://cen.acs.org/articles/91/i43/Taking-Reactor-Temperatures.html

http://www.rsc.org/chemistryworld/2013/10/nmr-thermometer-takes-reactor-temperature

http://www.spectroscopynow.com/details/ezine/141fed63ae8/MR-thermometry-Catalytic-insight.html?tzcheck=1&tzcheck=1

http://scitation.aip.org/content/aip/magazine/physicstoday/article/67/1/10.1063/PT.3.2231?dm_i=1Y69,229E6,E1MX8G,7GOMF,1

With so much attention, I'm sure my puny critiques will not be too rough for Jarenwattananon and co-workers.  I will divide them into three categories: accuracy, precision and sources of error.

#1) Accuracy.

In the body of the paper the authors state "The disagreement between NMR-derived temperatures and fibre-optic sensor measurements was at most 4%."  In the abstract they say "measurement error [is] less than four per cent of the absolute temperature."  I feel that it a bit misleading use agreement with external temperature sensors as a metric of accuracy.

Drill down into the paper and really ask yourself "what is the limit of accuracy in the temperature measurement?"  It has to be how accurately you can measure line widths of the analyte.  For example, Fig 1c shows data and Lorentzian fit for propylene in temperature calibration experiment.

Look carefully and tell me, what is the smallest line width difference you think that you can assess on such a spectrum?  Could you distinguish two spectra with line widths that differ by 1 Hz, 5 Hz or 10 Hz?    By inverting the linear equation (I'll use calibration curve for PtNP) , it is possible to calculate how much the temperature changes if line width changes by 1, 5 or 10 Hz.


T =  1 Hz / 0.16 Hz K^-1 = 6.25 K
T =  5 Hz / 0.16 Hz K^-1 = 31.25 K
T = 10 Hz / 0.16 Hz K^-1 = 62.5 K

4% of 400 K is 16 K, so the authors seem to acknowledge an accuracy of line width measurement of about 3 Hz.  I question if they can really measure the line width so accurately.  If the error is closer to +/- 25 K, then their false color thermal maps, particularly on the microreactor in Fig. 4b, could be quite misleading.

#2) Precision

In the caption for Fig. 1 and 3, the authors state that "temperature calibrations were performed for over 30 different systems." and "thermal maps were generated for more than 15 systems."  How much do the results vary from system-to-system?  Are these variations an indication of the variation in each reactor or the precision of the measurement.  If you repeat the calibration or mapping experiment several times how much variation would you see?

#3) Sources of error

I think of this experiment as almost an anti-DOSY.  In this experiment, the NMR signal in the presence of a gradient is sharpened not attenuated by molecular motion.  I surmise that this experiment works because gas molecules at elevated temperatures diffuse fast enough average gradients.  The problem as I see it is that this experiment only works if everyone agrees that the only factor that impacts the line widths is molecular motion.  You could fool yourself and think you see a cool spot if regions of the sample have any reason other than the gradient to have broad lines.  Some examples that spring to mind with heterogeneous catalysts is local concentration of paramagnetic substances. 

A related issue that troubles me is that the slope of the calibration curve (deltaF vs. T) is different for the two catalysts (PtNP and Pd-MOF).  I don't understand why.  It seems like if gradient-induced line width is narrowed by motional averaging (which is related to T) the specific catalysts should not matter.  Why is deltaF equal to -87 Hz at 400 K and G equals 0.05 G cm^-1 for PtNP and -79 Hz for Pd-MOF.  Either the material impacts the line width (which is clearly what the authors think) or these values represent errors in line width measurement.

***

In the end, I really love this paper.  It really made me think about stuff I don't get to think about too much line mean free path and the Maxwell distribution.  I get persnickety about a few issues, but you shouldn't let me distract you from the unprecedented view into reactor energetics this new NMR thermometry technique offers.    

NMR thermometry

The paper for this week was published back in late October in Nature.  I'm just getting to it now though,  because a) I'm always a bit behind and b) the title "Thermal maps of gases in heterogeneous reactions" does not alert the reader that the content of this paper is highly relevant to anyone who wishes to be fluent in contemporary NMR research.  The full citation is

"Thermal maps of gases in heterogeneous reactions"

by

Jarenwattananon, Gloggler, Otto, Melkonian, Morris, Burt, Yaghi and Bouchard

Nature 2013 502, 537

http://www.ncbi.nlm.nih.gov/pubmed/24153305

http://www.nature.com/nature/journal/v502/n7472/full/nature12568.html

This paper describes a novel NMR thermometry technique to measure gas temperatures at gas-solid interfaces in heterogeneous catalysts.  For engineers who optimize reactor design, temperature gradients on catalyst beds contain vital information on reaction energetics (so I am told).  The key insight of the authors is that in the presence of a weak magnetic field gradient (<< 1 G cm^-1) motional averaging will result in an inverse relationship between line width of a given peak and temperature of the system.  Figure 1a provides an excellent pictorial description of this technique.

To demonstrate the feasibility of NMR thermometry, the authors design a simple calibration experiment using a 10 mm NMR tube with catalyst-loaded glass wool (more on their catalysts later) and an analyte of propylene gas flowing at 15 cc min^-1, 40 PSI.  A 1D 1H spin echo spectrum is recorded with gradient applied during acquisition on a 400 MHz NMR (with microimaging capabilities) and the probe temperature set from 303 to 413 K in 10 K steps.  A sum of Lorentzians is fit to the olefin region of the propylene spectrum to get line widths.  Figure 1b shows linear fit of a plot of the change in linewidth relative to 303 K (deltaF) versus temperature (T) for three different gradient strength.

In the absence of a gradient, the slope of deltaF versus T is negative but small.  For a gradient strength of 0.1 G cm^-1, the slope is -0.13 Hz K^-1.  A stronger gradient results in a larger slope.

The authors settle on 0.005 G cm^-1 for gradient strength for their NMR thermometry setup.  They measure deltaF vs. T for two catalysts: Pt nanoparticles (PtNP) and Pd metal-organic frameworks (Pd-MOF).  Least squares fit results in a slope and intercept of -0.16 Hz K^-1 and 151 Hz and -0.10 Hz K^-1 and 119 Hz for PtNP and Pd-MOF, respectively.  Now the authors have the tools in place to measure T by NMR.  Simply measure the line width (of propylene olefin peaks) in 1D 1H spectrum with a gradient of 0.05 G cm^-1 during acquisition and use the appropriate linear equation to calculate temperature.

Having convincing demonstrated feasibility, the authors move to an application: thermal maps of lab-scale demo reactors with each catalysts.  In this experiment, propylene and hydrogen gas (actually para-hydrogen) are flowed through the reactor at 15 cc min^-1, 40 PSI and catalysis (hydrogenation) takes place.  The microimaging experiment (which takes ~30 min) divides the reactor into voxels and in each voxel the spin echo/gradient experiment is performed.  Line widths and, subsequently, temperature is determined for each 0.73 x 0.73 mm pixel in the reactor.  Figure 3 shows false color image of axial, coronal and sagittal views of the reactors.

Significant variations in the temperature of the catalyst bed is observed.  There are spots with T > 600 K and other spots with T < 400 K!  To validate these results, the authors carefully place a fiber optic temperature sensor at 3 spots in the demo reactor (you can see where in the coronal view of PtNP and sagittal view of Pd-MOF reactors).  The sensor agrees well with NMR thermometry-derived T.  The error is at most 4%.

Finally, the authors demonstrate NMR thermometry on a 1 mm microreactor with catalyst supported on silica gel.  Para-hydrogen preheat to 418 K is flowed into the reactor.  I assume that there is also propylene.  I have no clue what the flow and pressure is because the authors do not go out of their way to bore us with any details.  The temperature calibration experiment results in a slope of -0.2 Hz K^-1.  No mention of the intercept.  Figure 4 show a false color image thermal map from the microimaging experiment.

Once again, NMR thermometry reveals significant temperature variations in the reactor from 30 K higher than gas T at the inlet to 30 K lower at the outlet.

This is an amazing paper!  It is easy to understand why it was published in Nature.  The thermal maps in Fig. 2 and 4 are incredible.  I am no expert, but  don't see any other way to measure these types of temperature gradients on this scale other than NMR thermometry.  In Fig. 4 for instance, there is a gradient of ~60 degrees over a space of ~2.5 mm (which is less than 1/10 of 1 inch).

***

I'm going to make this a two-part blog and return tomorrow and sharpen my critique up a little bit.  See you then.

  

Wrapping up 2013

There were a lot of papers that I read last year and really liked.  I would like to write a full critique, but I just don't have time.  So here is my twitter critique (<150 words - I know that twitter uses characters, but give me a break.)

The quiet renaissance of protein nuclear magnetic resonance

by

Paul Barrett and a cast of thousands

http://www.ncbi.nlm.nih.gov/pubmed/23368985

http://dx.doi.org/10.1021/bi4000436

Super well-written review.  A call to arms to NMR spectroscopists to embrace NMR as a broad problem-solving tool rather than machine that generates PDB files.



Mycalol: A natural lipid with promising cytotoxic properties against human anaplastic thyroid carcinoma cells

By

Adele Cutignano and co-workers

http://www.ncbi.nlm.nih.gov/pubmed/23857783

http://onlinelibrary.wiley.com/doi/10.1002/anie.201303039/abstract;jsessionid=E7C1B2F402EF5A4BA96E06EFFEF70D1F.f04t03

A great natural products structure elucidation paper.  This paper is what really inspired me to start this blog.  The authors isolated a lipid from a sponge they found in Antarctica.  The molecule has five stereocenters and it would have been a pain in the ass to assess stereochemistry using NOE and J coupling.  The authors use CD and Mosher's method and a lot of clever logic.  A+


Characterization of Cyclopentyllithium and Cyclopentyllithium Tetrahydrofuran Complex

By

Su, Hopson and Williard

http://www.ncbi.nlm.nih.gov/pubmed/23875807

http://pubs.acs.org/doi/abs/10.1021/ja4059102

The first of three JACs papers published by these authors this fall.  Good grief.  I won't get three JACS papers in a lifetime.  These guys got three during the football season!  This one is probably my favorite for the cool use of DOSY to explore the equilibrium between a hexameric and tetrametric molecule.   


The influence of electronic modifications on rotational barriers of bis-NHC-complexes as observed by dynamic NMR spectroscopy

by

Kolmer, Kaltschnee, Schmidts, Peeck, Plenio and Thiele (so many Germans and no umlauts!)

http://www.ncbi.nlm.nih.gov/pubmed/24000182

http://onlinelibrary.wiley.com/doi/10.1002/mrc.4002/abstract

A great physical organic chemistry paper regarding Grubbs catalysts.  Quantitative dynamics to measure rotational barriers and a clear relationship between redox properties and rotational barrier.  Nice work.

Direct Measurements of the Mn(II) hydration state in metal complexes and metalloproteins through 17O NMR linewidths

by

Gale, Zhu and Caravan

http://www.ncbi.nlm.nih.gov/pubmed/24088013

http://pubs.acs.org/doi/abs/10.1021/ja4094132

NMR tricks to assess the hydration state of metal complexes.  Relaxivity measurements.  Sign me up. 

Xenon-based Molecular Sensors

This weeks paper is a hot-off-the-press JACs article by Garimella et al.  The full title is

Hyperpolarized Xenon-Based Molecular Sensors for Label-Free Detection of analytes.

by

Garimella PD, Meldrum T, Witus LS, Smith M, Bajaj VS, Wemmer DE, Francis MB, Pines A.

The full citation is

http://www.ncbi.nlm.nih.gov/pubmed/24313335

http://pubs.acs.org/doi/abs/10.1021/ja406760r

A few months ago I reviewed a very nice paper by Perrone et al. regarding NMR-based sensors (see http://sitspinnmr.blogspot.com/2013/10/nmr-chemosensing.html).  This weeks paper describes an alternative approach to designing an NMR-based sensor.  Garimella et al. design a sensor with two binding sites.  One site will bind a specific analyte, in this case the dye Rhodamine 6G.  The other site will bind xenon gas molecules dissolved in solution.  It turns out that xenon NMR is so sensitive that subtle changes on the other end of the sensor, such as the binding of a ligand, alter the chemical shift.  Below it the TOC graphic to explain the concept visually.

  

What do the author's actually do?

They begin by creating a peptide receptor library with 1.6e5 (=20^4) molecules based on the sequence KXXPGXXGWKKG.  Their hope is that some of these molecules (~2% or so - which is 3,200) will bind with a dye.   These peptides are bound to polystyrene beads at the C-terminus and pyrene at the N-terminus.  Since the peptides are attached to beads, the authors incubate the bead library with 10 uM dye for 1 hour, wash away the dye and manually sort the colored beads.  (I am glad I don't have that job.)  They screen the bead library multiple times and sequence the hits by mass spectrometry.  I am sure that this part is pretty clever, but the details are a bit lost on me.  In the end, their consensus sequence to bind Rhodamine 6G is H2N-KDDPGDEDWKKG-CO2H, which they call the "D-peptide."  The make another peptide as a control with the sequence H2N-KNNPGNQGWKKG-CO2H.  They call this peptide the "N-peptide."  The figure below shows visual confirmation that the D-peptides binds with Rhodamine 6G using the bead incubation assay, whereas the N-peptides does not show any color change.  So are we the audience to assume that the N-peptide does not bind the dye?  At this stage, I think the answer is yes. 

'

Garimella et al. can now move away from the beads and focus on the peptides.  They do some smart controls like replacing the pyrene with the Xenon binding cryptophane cage at the N-terminus to confirm that the dye binds to the peptide, not the pyrene.  They also confirm specificity by trying many other dyes, none of which bind with the D-peptide (no word about the N-peptide, though).  

Now we can get into some spectroscopy.  The authors use NOESY to confirm the interaction between the dye and D-peptide.  Their sample is 250 uM dye and 200 uM D-peptide-cage.  They also record a NOESY of 250 uM dye and 200 uM N-peptide-cage.  Here is what the authors report:

"The spectra of both the D- and N-peptides in the presence of dye revealed cross peaks in the region corresponding the dye-peptide interactions.  However, there were many more cross peaks in the D-peptide-cage sample, and they were more intense than those in the N-peptide-cage sample."

and then a bit later

"The NOE data .. verifie(s) that there was a difference in interaction of the dye with the D- and N-peptides, confirming our visual evidence that the library produced a receptor for the Rhodamine 6G analyte."  

 
So much for a negative control!  Still you have to give the authors a lot of credit for turning a bug into a feature.  (I am stealing that phrase from my friend Todd, who used it to describe a research projects around here.  It seems to me that the ability to turn a bug into a feature is a key trait in good science.)  So the N-peptide binds Rhodamine 6G, albeit with less affinity than the D-peptide.  As a final note on this topic, it seems to me that there is something funny going on with the interactions between the dye and peptide.  I spent some time staring at the NOESY spectra in the SI and I can't make heads or tails of this interaction. 

On to the Xenon-based detection - The Pines lab has developed a really cool system to deliver optically hyperpolarized xenon gas into their NMR samples.  The net result is a HUGE signal.  In the introduction to this paper the authors say that the NMR signal has the "strength compatible to those of water in conventional experiments."  I am assuming that they only need one scan!

At any rate, the authors make ten NMR samples, five for the D-peptide-cage, five for the N-peptide-cage each with increasing dye concentration from 0 to 1000 uM in steps of 250 uM.  The peptide concentration equals 200 uM.  Let's first consider the Xe NMR spectrum for the peptide-cage samples with no dye.  There are three peaks.  One for hyperpolarized Xe bound to the cage at ~60 ppm.  There is a sample for aqueous Xe (some the gas pumped into the solution dissolves in the solvent like CO2 in soda pop) at ~200 ppm.  There is also so undissolved Xe gas signal, which is set to 0 ppm.  (Since these signals are in slow exchange, I guess the dissolution process is slow on the NMR time scale.)  The gaseous signal is used as a reference and set to zero ppm in all spectra.  What I find strange is that the exact chemical shift of the aqueous Xe is at ~190.75 for the D-peptide and 191.25 for the N-peptide.  This is not a good quality for a sensor to possess.  I think that you would want any extra unbound sensor to be impervious to other conditions in the solution.  Is Xe sensing the pH of the solution (presumably the pKa of the D- and N- peptide is different)?  The authors did warn us that Xe NMR was sensitive.  The exact chemical shift of the Xe bound to cage equals ~60.5 ppm for the D-peptide and 62.25 ppm for the N-peptide.  I'll say it again - The authors did warn us that Xe NMR was sensitive.

Let's first consider the D-peptide.  When dye is added to the solution both peaks shift.  The aqueous Xe moves upfield and the Xe-cage moves downfield.  I think it is strange that the aqueous Xe shifts, but it is what it is.  The Xe-cage moves quite a bit more than the aqueous, Xe by the way.  Now let's consider the N-peptide.  As the dye concentration increases, the Xe-cage peak move upon binding with the N-peptide, albeit less than with the D-peptide.  The authors advocate using the difference between the peaks as a readout of the assay.  Using this metric, the Xe peaks move 0.78 and 0.33 ppm for the D-peptide-cage and N-peptide-cage upon binding 1 mM dye.  See figure below ...  

I am going to criticize this paper a bit, so I'll issue my usual caveat: I am a nobody doing LN2 fills and teaching 1st year graduate students how to acquire 1D 1H spectra, whereas the authors of this paper are real scientists doing the hard, creative work of producing scientific knowledge.  My purpose in starting this blog is not to trash people's work so that I feel better about my life.  I wanted to force myself (on a very public forum) to critique great science in hope of improving my science.  So here is my critique in one sentence: the negative control is not so negative.  Traditionally, in a paper like this one, you would have one positive and one negative control, proving that you get a signal when the analyte is present and no signal when the wrong analyte present.  In this paper the authors see a delta delta of 0.78 ppm for a carefully chosen strong binder and a delta delta of 0.33 ppm for a weaker binder.  If they could make a non-binder would the delta delta equal 0 ppm?  What would they see if they tried another dye molecule? 

Overall, I give the authors a lot credit for using a SELEX type procedure to hunt out a binder.  I kind of glossed over this point earlier to get to the spectroscopy, but it was a substantial effort to find a needle in a stack with 20^4 pieces of hay.  Clearly, this paper is a starting point.  The authors set out to show that their sensor-cage-Xe trick would is feasible.  I think they succeeded.  I wonder if their problem isn't going to be that Xe NMR is just too sensitive, though.     

Carbon CEST and low Spin-lock Field R1rho

Great paper this week by Zhao, Hansen and Zhang from UNC Chapel Hill (& the U of T).  Here is the citation:

The full title is

Characterizing Slow Chemical Exchange in Nucleic Acids by Carbon CEST and Low Spin-lock Field R1rho NMR Spectroscopy.

by Bo Zhao, Alexander Hansen and Qi Zhang

http://www.ncbi.nlm.nih.gov/pubmed/24299272

http://pubs.acs.org/doi/abs/10.1021/ja409835y

First of all, I've got to give the Zhao et al. credit for a descriptive title.  This JACs communication introduces a new pulse sequence (2D 13C CEST) useful for characterizing slow exchange in nucleic acids.  Then this experiment is compared and contrasted to other tools for characterizing slow exchange, namely R1rho RD and ZZ-exchange.  So as you can see, the title really sums up the paper.

I'll admit that I really like this paper, so let's jump in and see what it is all about.

This paper describes "a nucleic-acid-optimized 2D 13C CEST experiment."  I discussed the concepts behind the CEST experiment a couple of weeks ago, albeit in a slightly different context (see http://sitspinnmr.blogspot.com/2013/11/catalycest.html).  The experiment described in this week's paper is essentially a selective 2D HSQCs with a long exchange time (Tex) before t1 during which weak B1 is applied to the 13C channel.  This experiment is repeated and the offset of this B1 is arrayed.  The worked-up data (CEST profile) is a plot of the intensity of a given peak vs. offset.  Note that there are as many CEST profiles as peaks in the HSQC spectrum, in principle.  The authors also repeat at different B1 powers.  If the system does not undergo slow chemical exchange the CEST profiles is just a selective saturation experiment (like presat).  When the offset equals the resonance frequency, the peak disappears.  If the system does undergo slow chemical exchange, the saturation of either peak will impact the intensity of the other.  As a result, there are two dips in the CEST profile.

The authors use their new pulse sequence to characterize the ligand-free state of the fluoride riboswitch.  The structure of the ligand-bound form has been determined using crystallography - it is a pseudoknot.  The ligand-free form is hypothesized to have two stem loops based on the 1D 1H NMR.  There are a few extra peaks in the 1D 1H spectrum of the ligand-free RNA, presumably due to slow exchange with another species.  The unstated hypothesis is that this 2nd species is a pseudoknot identical to the ligand-bound form.
 

 
One cool trick employed by the authors is to selectively 13C/15N label guanosines in their RNA.  This selective labeling along with a clever use of shaped pulses "to selectively invert and refocus carbon magnetization of interest and to refocus carbon-carbon couplings from neighboring carbons" isolates two G resonances: H8-C8 (base) and H1'-C1' (sugar).  I have played this trick myself, so I know how cool it is to reduce a complicated RNA to a pretty simple and easy to look at spectrum.  If you study the primary sequence of the fluoride riboswitch  you will find 12 G residues.  If you classify these twelve based on the hypothesized secondary structure of the ligand-free RNA and the measured structure of the ligand-bound RNA you would find that - a) there are 4 G residues (G7, G8, G10 and G39) with different secondary structure in free and bound form; b) there are 8 G residues (G1, G2, G4, G14, G23, G30, G31, G33) with the same secondary structure in free and bound form.  These are probes we can use to monitor the slow exchange between these two states using the CEST experiment.

The authors record 13C CEST profiles at 30 C with Tex = 300 ms. (I wonder how they chose these values!).  There is only one dip in the CEST profile of one of the control probes (G33) with the same secondary structure (and presumably chemical shift) in either the ligand-free or bound form.  There are two distinct dips in the CEST profile of two probes (G8 and G10) that transition from a tetraloop secondary structure to a stem-loop upon ligand binding.

The interpretation of this data is that there is an "invisible" excited state (ES) in slow exchange with the ground state (GS) for G8 and G10 (seen in both base and sugar resonances).  For G33 on the other hand, there is no chemical exchange and, thus, no excited state.

The authors make a throw away comment that made me pause: "except for residues from P2, we observed either asymmetrically broadened intensity dips or more than one intensity dip for all other guanosine residues."  I guess this observation makes sense.  After all P1 is a stem-loop in the ligand-free structure and right in the business end of the pseudoloop in the ligand-bound state.  So although it is helical in both structures, it might have different chemical shifts.  So what I am saying is that my division of the 12 guanosines into two easy categories is an oversimplification, at best, and just plain wrong, at worst.

The authors focus their attention on G8 and G10.  Two-state Bloch-McConnell equations are fit to the CEST profiles to extract the ES chemical shift, the exchange rate constant and equilibrium constant.  For the base resonances, the ES chemical shift equals 134.3 and 133.8 ppm for G8 and G10, respectively, which is ~4 ppm upfield from the GS chemical shift.  The average C8 chemical shift for a G residue in a helix equals 133.47.  The exchange rate constant (k1 + k-1) equals 112 +/- 10 Hz.  The population of ES equals 9.8%.  (By the way, with a population that large, why don't you see ES cross-peaks in the HSQC?)  It is trivial to calculate K = 0.112 and deltaG = 5.5 kJ/mol or 1.3 kcal/mol.  Also it is trivial to calculate k1 = 11.2 Hz and k-1 = 100.8 Hz.  (Remember that these values are at 30 C.)

I think it is important to consider these numbers in context.  Arguably, the most prominent NMR dynamics measurement was published WAY BACK in 2005 by Eisenmesser et al. (Nature 2005 438, 117 - http://www.ncbi.nlm.nih.gov/pubmed/16267559).  This publication discusses the protein cyclophilin A (CypA).  CypA is in equilibrium between a major and minor form and the authors use relaxation dispersion NMR to measure K = 0.055, k1 = 60 Hz and k-1 = 1080 Hz.  Man, I thought this work was the bee's knees back in the day!  At any rate, this protein refolds with rate constants ~ one-fold larger than the fluorine riboswitch.     

For another example Wenter et al. (Angew. Chem. Int. Ed. 2005 44, 2600 - http://www.ncbi.nlm.nih.gov/pubmed/15782371) followed refolding of a bistable 20-mer using real time NMR.  (This paper is one of all-time favorites).  They determine K = 0.236 , k1 = 0.031 +/- 0.006 Hz and k-1 = 0.131 +/- 0.024 Hz at 25 C.  By the way, to ease comparison with the present paper I've flipped the reaction from the Wenter paper so that the more stable fold is the reactant and the less stable fold is the product.  However you look at it, this bistable hairpin refolds with rate constants THREE ORDERS OF MAGNITUDE LESS than the fluoride riboswitch. 

It kind of stuns me that the Wenter 20-mer has such smaller rate constants!  To unfold, the 20-mer does have to break 6 Watson-Crick base pairs and form four new ones.  (I don't know if these two events happen sequentially or simultaneously).  Returning to the Zhao paper, only three or four Watson-Crick base pairs are broken, but nine new base pairs are formed (if we assume the "invisible" species is a pseudoknot identical to the ligand-bound form).  Additionally, some tertiary elements need to form.  So how does the fluoride riboswitch do it so fast?  (This point seems like a good place to admit that I am reading this paper in ASAP form and do not have access to the SI, so I have no idea how much Mg2+ Zhao et al. uses vis-a-vis Wenter et al.)  I guess I should also turn the question around and ask why does the Wenter 20-mer refold so slowly.  Obviously, there is something with the transition state!

To finish up, Zhao and co-workers end their communication by comparing their novel pulse sequence to two more established techniques, low spin-lock field R1rho RD and ZZ exchange.  The results are essentially identical, though ZZ exchange is a bit dicey.  They conclude "The currently presented 2D 13C CEST experiments ... provide powerful tools to investigate slow chemical exchange.  The robustness of these methods promises a unique opportunity to facilitate atomic understanding of slow conformational interconversion that is essential to many vital nucleic acid functions."

I agree.  I like this communication an awful lot and I want to try CEST next time I deal with slow chemical exchange.

CatalyCEST

This week we'll be discussing the following recent JACs communication:

A CatalyCEST MRI Contrast Agent That Detects the Enzyme-Catalyzed Creation of a Covalent Bond

by

Dina Hingorani, Edward Randtke and Mark Pagel

http://dx.doi.org/10.1021/ja400254e

http://www.ncbi.nlm.nih.gov/pubmed/23601132

I am a sucker for NMR-based bioanalytical assays.  These experiments appeal to my desire to move NMR beyond its role in structural biology and into other arenas of biology.  Additionally, I have watched these CEST methods with a passing interest, but I have never had a chance to sink my teeth into this kind of work.

In this JACs communication Hingorani et al. describe a MRI contrast agent designed to detect catalysis by the enzyme transglutaminase (TGase).  The authors call this technique CataylCEST.  Here is a rough sketch of how it works.  The authors design a paramagnetic tag with a moiety that looks like the substrate to an enzyme.  (In this case the substrate is lysine and the enzyme is TGase).  Then the enzyme covalently attached the paramagnetic tag to a second substrate (in this case a protein or peptide via glutamine residues).  Once the tag is attached, protons (and 13C, 15N, etc) in the neighborhood to it broaden and resonate at dramatically different frequencies due to hyperfine contact shift.  Let's assume that some of these protons are in slow exchange with the water.  If you saturate these spins using a presat-type experiment, then during the saturation pulse, some of this saturation is transferred to the water by chemical exchange and the integral of the water signal will decrease relative to a control.  Figure 1A of the paper (below) describes this experiment:

 

The specific enzyme the author's query, TGase, catalyzes covalent bond formation between side chains of glutamine (Q) and lysine (K).  TGase forms cross-links in the extracellular matrix in tissues and in cancer.  The authors do not discuss their motivation for monitoring TGase in the introduction and only touch on it in the conclusion.  I can only surmise that their long-term goal is to develop in vivo CEST MRI with exogenous CEST agents that do not suffer from rapid pharmacokinetic washout.  

The results of this study - 

The author's synthesize TM-DO3A-cadaverine paramagnetic CEST agent and couple this molecule to 5 different substrates using inexpensive microbial TGase.  The substrates are: Boc-Gln-OH, CBZ-protected QG peptide, QR peptide, GQR peptide and bovine serum albumin (BSA).  They acquire their NMR data on a 600 MHz NMR at 37 C using (essentially) the presat experiment with a saturation time of 4 s and a field of 20 uT (or (20 uT/14.1 T)*600 MHz = 850 Hz field).  The saturation offset is arrayed in 1 ppm steps from -30 to 30 ppm, relative to water, which is set at 0 ppm.  The data is presented as a saturation profile showing the intensity of the water signal as a function of the saturation offset.  Finally, the authors fit Lorentzian line shapes to assess the chemical shift of the species in slow exchange with water.  The authors do not report error bars, which I find troubling, because the peaks are quite broad.  As we'll see later, the authors will argue that a difference of 1.8 ppm is significant, but a difference of 1 ppm is not.  I am not sure what to make of this difference, myself.

The most convincing result is with BSA.  The authors make an NMR sample with 25 mM CEST agent, 0.75 mM BSA and 10 mM glutathione (to maintain the reducing environment required by the eznyme) in pH 7 Tris buffer.  Then they react with 0.327 uM TGase for 24 h under aerobic conditions.  The authors repeat this experiment in triplicate.  Before catalysis, the saturation profile shows CEST at +4.6 ppm upfield from water.  After cataysis, the saturation profiles shows a CEST at +4.6 ppm and -9.2 ppm.

 

  

The interpretation is that the signal at +4.6 ppm is diamagnetic amines and amides in BSA.  TGase catalyzes the formation of a covalent bond between the CEST agent and Q side chains on BSA.  Hyperfine contact shift of Tm(III) induces an upfield shift in the chemical shift of the amides to -9.2 ppm.  (By the way - do you notice how much easier it is to look at the Lorentzian fits on the right than the saturation profiles on the left!)

If the publication ended right here, I'd be sold.  This paper confuses me, though, when the authors report their controls.  Their results reinforce a long held suspicion about CEST: how do you assign the peaks in your saturation profiles?

Let's discuss their controls.  The most logical control is each of the components individually.  The CEST agent alone shows a CEST at -5.2 ppm, BSA alone shows a CEST at +5.6 ppm and glutathione alone shows a CEST at +5.4 ppm.  By the way, I'll note that the text says that BSA and glutathione have a CEST at +4.6 ppm, but the figure caption says +5.6 and +5.4 ppm, respectively.  You see what I mean about error bars!  I guess 1 ppm is not significant.  So why isn't the CEST profile of the reaction mixture before catalysis equal to a superposition of the reactants?  (I think it probably is, if you correct for concentration, but the authors do not do this for the readers and we are left wondering).  Next the authors try control peptides.  The authors mix 25 mM CEST agent, 25 mM peptide, 10 mM glutathione in pH 7 Tris buffer with and without TGase.  For GQR and QR, there is a CEST at -9.0 ppm before catalysis and CESTs at +5.8 ppm and -10.8 ppm after.  (In the discussion, the authors assign the signals at -9 ppm and -10.8 ppm to a supramolecular adduct and paramagnetic amides, respectively, implying that they can tell a difference between signals the differ by 1.8 ppm.  See what I mean about error bars!)  For CBZ-protected QG, there is no CEST signal before catalysis and CESTs at 4.6 ppm, 9.8 ppm and 22.5 ppm after.  Finally for Boc-protected Q there is a CEST at +7.2 ppm before and a broad CEST between -10 ppm and -20 ppm after catalysis.  Why do none of the controls show the same signal before catalysis as BSA?  (Once again, I presume the issue is concentration, but the authors leave it up to their reader to discern).  Also, after covalent attachment of the CEST agent why does the CEST differ depending so dramatically for different substrates?    

The author's interpretation of their results falls into two categories (they don't make this explicit, I am interpreting).  1) Aggregation/Heterogeneity - There is a noncovalent supramolecular adduct or some type of conformational heterogeneity that dramatically alters the chemical shift of the water exchangeable protons.   This effect is responsible for CESTs at -5.2 for the CEST agent alone, at -9 ppm for GQR and QR peptides prior to TGase catalysis, at +4.6 ppm, +9.8 ppm and +22.5 ppm for the CEST agent-linked ZQG peptide and at -10 to -20 ppm for the CEST agent-linked Boc-Gln-OH.  2) Change in chemical exchange rate contants -  The authors assert that chemically modifying the substrate alters the rate constant and rates of chemical exchange.  This effect is responsible for the CEST at +5.8 ppm for GQR and QR peptides after TGase reaction.

As you can tell, I am skeptical of these explanation.  I am not saying the authors are incorrect.  They know a lot more about this subject than I do.  I only mean to say that I believe more justification is needed.  For example, there are a few additional controls the authors could do to validate their assignments.  If they are concerned about "noncovalent supramolecular adducts", why not reduce concentration or increase ionic strength to break up these interactions?  If they are concerned about hydrophobicity causing conformational heterogeneity or heterogeneous ligand conformations, then perhaps the peptides they are using are not good controls?  If they are concerned about rate constants, why not play with temperature or field?  

In the end, this paper DOES convince me of its main objective: the authors have a catalyCEST MRI contrast agent that can be used to detect the formation of a covalent bond in BSA mediated by TGase.  After studying this paper, though, I am concerned that the CEST varies from substrate to substrate - from -9.2 ppm for BSA to 22.5 for ZQG.  Let me end my critique with a question: If you wanted to check for TGase activity using a new protein and you were not sure if it was a substrate or if you wanted to test TGase activity in vivo, then what results do you expect from the CEST assay?  The fact that you do not know shows how far we have to go. 

Dissecting the stereocontrol ...

My blog post this week will critique the paper:

Dissecting the Stereocontrol Elements of a Catalytic Asymmetric Chloroactonization: Syn Addition Obviates Bridging Chloronium.

By

Yousefi, Ashtekar, Whitehead, Jackson and Borhan

http://pubs.acs.org/doi/abs/10.1021/ja4072145

http://www.ncbi.nlm.nih.gov/pubmed/24025085

Funny story about this JACs communication (JACs 2013 135, 14524): I was reading this paper while riding a stationary bike at my campus gym, when a certain faculty member approached (name withheld to protect the guilty).  He asked "What are you reading?".  I showed him the paper and he said "That looks brutal!"  I laughed.  This science in this paper is excellent, but it was a poor choice by me for a blog that focuses on NMR.  By the time I realized that the NMR methodology was not novel, I was too deep into the paper to pull back.  Nevertheless, I think this publication is worth discussing, because it presents an interesting solution to a difficult diastereospecific assignment problem.

As an exercise for myself, I'll see if I can briefly describe the problem and why this paper is relevant to people interested in NMR.  Then I'll show the author's data and interpretations.

Yousefi et al. are interested in the reaction below: an enantioselective halocyclizations of alkenes.

This type of chemistry is important as "a robust, versatile route a wide range of heterocycles."  The specific question that the authors address is the following:  Why is this reaction enantioselective?  To address this question, the author use NMR spectroscopy of isotopically labeled precursors to assess the mechanism.  Figure 1 describes the two proposed intermediates: path a - three-membered chlorenium ion intermediate; path b - carbocation intermediate.  SPOILER ALERT (this part is in the title): It is path b.

These two mechanism offer some predicable consequence.  If the reaction proceeds by path a, the "enantioselectivity would be controlled in parallel with the initial asymmetric chlorenium delivery, yielding anti addition".  If the reaction proceeds by path b, "the reaction's enantioselectivity would be determined at the (presumably catalyst controlled) ring-closing step."  If the author's can show syn addition, then path a can be ruled out.  The problem demonstrating syn addition is that "the chlorine resides on a nonstereogenic carbon with no record of its attach path on the alkene."  So it is not easy to determine the addition, unless you can somehow label the product!    

The authors synthesize a E-deuterated analog of the alkene.

Then they perform the halocyclization reaction and analyze the products by NMR.

At this point I'll note that the publication itself has ZERO NMR spectra in any figure.  So you'll have to dig through the SI to see any of their data.  Here is the spectrum of their products.

No integrals or peaks picked, but I see 2 methylenes from the lactone at 2.5 and 2.8 ppm and the 5 phenyl protons at 7.4 ppm.  At 3.75 ppm is the CH adjacent to the Cl.  To remind you, below is the expected molecule for the protonated starting material:

  
One relevant factor to consider is that their deuterated analog is not perfect.  They report 94:6  E:Z and 88% D-incorporation.  So they have a bit of protonated product in with their deuterated product.  Looking at this data I see a "roofed" non-first order CH2 from the protonated product (peaks at 3.815, 3.805 and 3.772 ppm with the final leg hidden under the big peak at 3.74 ppm) overlapping with two CHD diastereomers at 3.805 and 3.74 ppm. 

The author's interpretation is explained in the following figure from the SI:

   
You may be asking how they know that c is the R,R or S,S diastereomer and f is the R,S or S,R diastereomer?  This gets to the heart of the difficult problem in diastereospecific resonance assignment.  Let me quote directly from the paper: "The absolute stereochemistry of the CHDCl group in the major isolate was straightforwardly established via NOE analysis of epoxide 3-D obtained from the chemically transformed chloroactone product 2-D"

To translate, the deuteratd epoxide (3-D) has a peak at 2.98 ppm in 1D 1H NMR spectrum.  The protonated epoxide (3) has two peaks at 2.98 and 2.73 ppm.  Using NOE, the peak at 2.98 is established as trans the phenyl group.  (This data is nowhere to be found in the body of the paper or SI, by the way.)  Hence the deuteron in 3-D must be cis.  Retrosynthetic analysis can be used to establish the stereochemistry of the major product 2-D.

I'll leave it to the organic chemists to fight through their mechanistic arguments.  Their NMR argument is interesting to me, though.  Basically, the authors cannot complete diastereospecific assignment of 2-D using conventional approaches, so they modify the molecule in a stereochemically predictable manner to simplify the problem.  I find this approach to be clever.

Like I said at the beginning, this article is not for the faint of heart.  I don't know that I am really all that interested in mechanistic details of this reaction.  My interest is in the NMR.  Frankly, the authors do not wow me with their NMR, but I am intrigued by their solution to the diastereospecific assignment problem.