J.1747 0285.2009.00940.x

NMR-Based Screening of Membrane ProteinLigands

NaveenaYanamala1,†,ArpanaDutta1,†,BarbaraBeck2,BartvanFleet2,KellyHay2,AhmadYazbak3,Rieko Ishima1, AlexanderDoemling2,* andJudithKlein-Seetharaman1,*

1Departments of Structural Biology, 2Pharmaceutical Sciences andChemistry, University of Pittsburgh, Pittsburgh, PA15260, USA3Synthatex, Shefa-Amr Industrial Park, PO Box 437 Shefa Amr20200, Israel*Corresponding author: Judith Klein-Seetharaman, [email protected];Alexander Doemling, [email protected]†These authors contributed equally to this manuscript.

Membrane proteins pose problems for the appli-cation of NMR-based ligand-screening methodsbecause of the need to maintain the proteins in amembrane mimetic environment such as detergentmicelles: they add to the molecular weight of theprotein, increase the viscosity of the solution, inter-act with ligands non-specifically, overlap with pro-tein signals, modulate protein dynamics andconformational exchange and compromise sensitiv-ity by adding highly intense background signals. Inthis article, we discuss the special considerationsarising from these problems when conducting NMR-based ligand-binding studies with membrane pro-teins. While the use of 13C and 15N isotopes isbecoming increasingly feasible, 19F and 1H NMR-based approaches are currently the most widelyexplored. By using suitable NMR parameter selec-tion schemes independent of or exploiting the pres-ence of detergent, 1H-based approaches requireleast effort in sample preparation because of thehigh sensitivity and natural abundance of 1H inboth, ligand and protein. On the other hand, the 19Fnucleus provides an ideal NMR probe because of itssimilarly high sensitivity to that of 1H and the lackof natural 19F background in biologic systems.Despite its potential, the use of NMR spectroscopyis highly underdeveloped in the area of drug discov-ery for membrane proteins.

Received 29 July 2009, revised 30 November 2009 and accepted forpublication 30 November 2009

Membrane proteins are encoded by up to 30% of typical genomesand constitute the most important class of drug targets: morethan 60% of current drugs are targeting membrane receptors,

channels or transporters. Among these, the G-protein-coupledreceptors (GPCRs) are the largest group of drug targets becauseof their important role in mediating communication between theinside and outside of the cell in response to an enormous varietyof different ligands, ranging from small proteins and peptides tosmall organic molecules, ions and even light. These ligands canbe hormones, odorants, neurotransmitters or other functional clas-ses of biologically active compounds. Despite the importance ofmembrane proteins as drug targets, they have not been very ame-nable to structure-based drug design. This is because the hydro-phobic nature of their transmembrane regions hamperscrystallization as well as NMR-spectroscopic analysis. Progress inmembrane protein structure determination by NMR is steadilybeing made, with some recent spectacular breakthrough achieve-ments in the sizes of protein structures obtained for both b-barrelmembrane proteins (1,2) and a-helical proteins (3). Because thestructure determination of membrane proteins involves extensivedetergent screening and the selection of suitable buffer condi-tions, it is not a routine application. Thus, NMR structure-baseddrug design involving membrane protein targets still remains afuture goal. However, this does not preclude the application ofNMR techniques to membrane protein drug discovery. In particular,NMR spectroscopy can yield high-quality ligand-binding informationeven in the absence of the structures of the targets. This articlewill explore the applicability of different NMR-spectroscopicapproaches to the study of ligand–membrane protein interactionsfrom a fundamental perspective keeping in mind their potentialuse in drug discovery.

This review is organized as follows. First, in 'NMR-basedapproaches to drug screening', we will briefly review differentNMR-based approaches to the study of ligand binding to solubleproteins. In 'Challenges in membrane protein NMR spectroscopy',we will highlight the applicability and special considerations forNMR-based approaches in the context of membrane protein stud-ies. '1H NMR-based approaches for membrane proteins' describes1H NMR-spectroscopic approaches, and '19F NMR-basedapproaches' provides an overview of 19F NMR-spectroscopicapproaches. 'Comparison of 1H and 19F-NMR-based versus conven-tional screening of membrane proteins' will discuss the advanta-ges and disadvantages of 1H and 19F NMR-based screeningmethods when compared to other high-throughput screening (HTS)approaches. 'Synthesis of 19F containing small molecule com-pounds' will describe the practical aspects of obtaining 19F con-taining small molecule compound libraries. Finally, we willconclude with 'Summary and outlook'.

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Review Article

ª 2010 John Wiley & Sons A/S

doi: 10.1111/j.1747-0285.2009.00940.x

NMR-based approaches to drugscreening

Although NMR-based screening is only one of many screeningtools in drug discovery, its simplicity, wide range of application(including protein–protein and protein–nucleic acid interactions)and superior ability to detect weakly bound molecules haveattracted much attention. Nuclear magnetic resonance allows themeasurement of multiple parameters at different levels of com-plexity and information content. Thus, NMR-based methods differsignificantly from one another as a result of the particularapproach used. Excellent reviews of different NMR-screeningmethods are provided for example in (4–7). Here, we brieflyreview the different methods that have been mainly employedwith soluble proteins to provide an idea of the scope ofapproaches with potential or realized applicability to membraneprotein ligand screening. Fundamentally, two types of experimentscan be distinguished in NMR-based screening approaches: one todetect protein signals (Screening of ligands by detecting target-protein signals) and the other to detect ligand signals (Screeningof ligands by detecting ligand signals). There are also specializedimprovements in technology to increase throughput or to studyparticular types of ligands such as those that disrupt protein–pro-tein interactions (Other NMR-based screenings). Because sensitivityof the observed NMR signals in the ligand–protein interacting sys-tems depends on binding affinity, the estimation of the ligand dis-sociation constant (or binding constant) is also described(Determination of ligand-binding constants by NMR), before weend with a Summary.

Screening of ligands by detecting target-protein signalsIn protein-detection based screening, the identification of ligandbinding is based on changes in NMR signals arising from proteins,typically in one-dimensional 1H spectra or two-dimensional 1H,15N-heteronuclear single quantum correlation (HSQC) spectra. Becauseof the large number of peaks in proteins, two-dimensional experi-ments will afford better resolution of signals but require that theprotein is labeled. The longer data acquisition times for higherdimensional spectra are also a drawback, especially when screeninglarger numbers of ligands. Recent efforts are therefore aimed atdecreasing the acquisition time, including 'SOFAST- HMQC' or'Ultra-fast experiments' (8,9).

Binding information can be obtained for one- or two-dimensionalspectra regardless of whether the signals are assigned or not by sim-ply recording if signals show altered chemical shifts or line broad-ness, and many screening programs are based on this approach (6).Broadening of the NMR signals is observed when the exchange rate(defined by the population weighed on ⁄ off-rate of the ligand) is simi-lar to the difference in chemical shifts between the free and boundforms (10). Changes in signal positions are only observed when theexchange rate is slow, i.e., the ligand binds tightly. Broadening andchanges in chemical shifts of signals upon ligand binding aredetermined by the differences in chemical shifts, the relativeprotein ⁄ ligand molar ratio and the on ⁄ off-rate of the ligand. For anin-depth discussion of the different regimes, see (7,11).

Chemical shift changes and line broadening are parameters thatcan be used for screening even if resonance assignment is not fea-sible. More information, however, can be obtained when signalsare assigned. In that case, the changes in chemical shift or broad-ness of lines can be used to generate testable hypotheses on whatare the residues in contact with the ligand, or which are allosteri-cally modulated by ligand binding.

Even more information can be extracted, if the protein structure isknown. In particular, the pioneering work of Fesik at Abbott Laborato-ries (Illinois, USA) opened a new field in the area of fast andefficient drug discovery, a technique coined structure–activity rela-tionships by nuclear magnetic resonance ('SAR by NMR') (12). TheAbbott group uses 2-dimensional 1H,15N-HSQC spectra to screensmall molecular weight compounds for binding to 15N-labeled pro-teins of determined structures. Structure–activity relationships byNMR locates the binding site for the ligand on a protein's surfacebecause the resonances have been assigned prior to ligand screen-ing, and the structure of the protein is known. Comparing the struc-tures of compounds that bind to the same site on a protein providesinformation about the functional groups involved in ligand bindingand can guide the synthesis of lead compounds by medicinal chemis-try. This technique is restricted, however, to protein sizes of less than30 000 D because of limitation by the molecular rotationalcorrelation times leading to broad NMR lines for larger proteins.Many compounds have been discovered by this technique (13), andseveral compounds emerged in human clinical trials (14).

In cases where protein signals have not been or cannot be identi-fied, other lead optimization methods such as Inter-Ligand NOE(ILOE) and ILOE for Pharmacophore Mapping (INPHARMA) can beused to detect protein-mediated ligand–ligand interactions bydetecting ligand signals (15,16). The principle of these methods isbased on two ligands binding to the same protein. ILOE is used toidentify pairs of small molecules that bind to adjacent sites on thesurface of the target protein (15). In contrast to the ILOE's detectionof simultaneous ligand binding at two different but proximal sitesin the protein (15), the INPHARMA technique is specialized to iden-tify ligands that compete for the same ligand-binding site (16). Theidea in the ILOE approach is similar to the SAR by NMR approachin that the occupation of proximal but initially independent ligand-binding pockets can be combined with a single ligand targetingboth pockets to obtain higher affinity ligands. In the INPHARMAapproach, the two ligands are never close in space or bound to theprotein simultaneously, but rather the observed NOEs are mediatedby spin diffusion via the protons on the protein. The advantage ofthe ILOE and INPHARMA methods is that assignment and structureof the protein do not need to be known for lead optimization.

Screening of ligands by detecting ligand signalsProtein-detection-based methods suffer from the general drawbackthat NMR lines become broader with the increased size of the mol-ecule under study. This makes it desirable to measure the ligandinstead of the protein, because ligands are typically small mole-cules and will give rise to much sharper and more intense signals.Thus, NMR-based screening has often made use of detecting

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238 Chem Biol Drug Des 2010; 75: 237–256

signals of the ligands that interact with target proteins. There aremultiple ways by which ligand signals can carry information on pro-tein binding, and these can be detected by classical NMR parame-ters such as chemical shift and relaxation. Excellent overviews areprovided for example in (6,7).

A popular approach for ligand screening is based on the transferredNOE (trNOE) mechanism. Proton-proton cross-relaxation exhibitspositive NOE peaks for small molecules alone (MW < 2000 D) thatundergo fast molecular tumbling, whereas negative NOE peaks areobserved when the molecular tumbling becomes slow by forming acomplex with the target protein. Because ligands are at equilibriumbetween the free form and bound to the target protein, the NOEintensity that is encoded during the bound state is transferred bythe exchange and observed at the free ligand signal position. Othermethods that are based on the cross-relaxation mechanism includesaturation transfer-difference (STD) experiments, Water-LOGSY,cross-saturation, transient trNOE and NOE pumping (7).

Saturation transfer-difference (STD) experiments detect inter-molec-ular magnetization transfer by taking the difference of two NMRspectra recorded with and without saturation of protein signals(17). The mechanism of this approach is based on rapid proton spindiffusion in proteins: in large proteins, once a part of the proteinsignal is irradiated, the saturation is transferred to the entire pro-tein within 0.1 seconds (18). The application of STD to membraneproteins is discussed in 1H NMR-based approaches for membraneproteins.

Another mechanism for communication between ligand and proteinis via water molecules (19,20). This approach is based on the obser-vation that ligands are often hydrated when bound to protein, orspecifically mediate the interactions between ligand and protein viahydrogen bonds. Thus, by excitation of water, ligand and proteinsense their proximity. This mechanism is the basis for theWater-Ligand Observation with Gradient Spectroscopy (Water-LOGSY) technique that detects water-ligand NOE transfer. For

water-ligand molecules that are located on the target-protein sur-face, the NOE is negative (19,20).

Other NMR-based screeningsTo improve the HTS capabilities of NMR-based approaches, target-immobilized NMR screening (TINS) has been proposed (21). Here,the protein target is immobilized on a gel-based solid support. Thisis associated with several potential advantages: the target does notneed to be soluble or even be a protein; the quantity of requiredtarget is reduced, as a single sample of the target is sufficient fora flow-through screen. With TINS, compound libraries can bescreened much faster than using a traditional NMR sample in solu-tion.

In addition to screening, the binding of ligands to single proteinssuch as enzymes or receptors, it is becoming increasingly importantto investigate ligands interfering with protein–protein interactions,as the importance of protein–protein interactions as targetsincreases. A fast and information-rich NMR-based technique toscreen antagonists of protein–protein interactions has recently beendescribed by Holak et al. (22). This experiment has been coinedNMR-based Antagonist Induced Dissociation Assay (AIDA) for thevalidation of inhibitors acting on protein–protein interactions(Figure 1). Antagonist Induced Dissociation Assay detects signalsappearing upon the dissociation of the target-protein complexes.The approach requires a large protein fragment (larger than 30 kDa)to bind to a small reporter protein (less than 20 kDa). This method-ology has been successfully used to discover novel p53 ⁄ mdm2antagonists (23). A cost of goods saving 1D AIDA technique hasbeen described recently as well, in which tryptophan resonancesare used as reporters for ligand-binding events because of theirseparation from most other signals in proton NMR spectra ofproteins (24).

In contrast to the earlier mentioned in vitro assays, there are alsoefforts to conduct screening in vivo. The approach is called small

A

B

C

D

Figure 1: Schematic outlining the principle of the AIDA technique to screen for ligands. Here, AIDA was used to discover antagonists ofthe protein–protein interaction between p53 and mdm2. Left: structure (pdb identifier 1YCR) of the complex between p53 (blue helix) andmdm2 (yellow surface). Nuclear magnetic resonance screening of chemical compounds schematically drawn in the middle yields the 1D AIDAproton NMR spectra of the p53 ⁄ mdm2 complex on the right. Spectra labeled A–D exhibit signals from p53 in the presence of augmentingconcentrations of an antagonist. (A) no antagonist added (W23 is buried and does not give a signal). (B–D) increasing concentrations of anantagonist are added and more and more complex dissociates. This can be seen by the increase in the intensity of the W23 peak.

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Chem Biol Drug Des 2010; 75: 237–256 239

molecule interactor library (SMILI)-NMR (25). This method recordsNMR signals of a protein that is over-expressed in Escherichia coliand elucidates changes in signal positions and broadening uponligand interactions (26). The in-cell NMR approach has also beenapplied to observe and disrupt protein–protein interactions, coinedStructural Interactions (STINT) NMR (27). The advantages of the invivo studies are the detection of signals of unpurified proteins andinformation for more biologically relevant in vivo protein structuresand interactions. Expansion of in vivo ligand-binding studies tomammalian cells has recently enhanced the relevance and informa-tion content of the technique (28).

Determination of ligand-binding constants byNMRTypically, ligand-protein titration is conducted by observing proteinsignals to determine ligand association ⁄ dissociation constants. First,based on the equation of the dissociation constant, when the disso-ciation constant of the ligand is around 1 lM (tentatively definingmoderate binding), approximately 99% of the protein binds theligand at 0.1 mM protein concentration with almost equal amountof the ligand. Upon varying the ligand concentration, the populationof the bound form is consequently changed. Therefore, the titrationcurve is generated by plotting changes in the peak positions or sig-nal intensities to determine the dissociation constant. Next, whenthe dissociation is above 1 lM (tentatively defining as weak bind-ing), larger amounts of the ligand is required to saturate the proteinsignals to the bound form. Because of limitations in ligand solubilityor appearing of non-specific interactions at high ligand concentra-tions, it is possible that the dissociation constant is not well deter-mined by NMR for very weakly interacting systems. Finally, whenthe dissociation constant is significantly lower than lM, such as nM

(strong binding), the titration curve becomes so sharp that anaccurate dissociation constant is not obtained.

Determining ligand affinity using ligand signals is not straightfor-ward. When the binding is strong, the ligand-saturated point is dif-ficult to detect because ligand signals become broadened uponbinding to the protein. When the binding is weak, interaction isbetter detected using the STD technique and other experimentsdescribed earlier. However, it is difficult to determine the dissocia-tion constant accurately because other rate constants, such ascross-relaxation rates, are involved in such experiments.

These issues are illustrated by the case studies of different ligandsbinding to the model protein bovine serum albumin (BSA). Bovineserum albumin binds a variety of different ligands including moder-ate-affinity (lM), high-affinity (nM) and low and ⁄ or varying affinitymultisite binding ligands. For example, L-tryptophan is a moderate-affinity ligand, while naproxen is a high-affinity ligand, and salicy-late has been proposed to bind to 76 binding sites in total (29). Asystematic review of 1H NMR spectroscopy of these different typesof ligands and combinations thereof (30) has yielded the followingconclusions: when measuring 1H NMR chemical shifts and linewidths, titrations of different ligand ⁄ protein ratios are needed toobtain an accurate binding constant. Particularly, careful measure-ments and analyses have to be carried out for multisite ligands: awrong 1:1 binding model can provide a visually acceptable fit to

the experimental salicylate binding data, while in-depth studiesreveal the multiple site binding modes of this ligand (30).

For sub-micromolar affinity ligands where the free ligand peak isunaffected by the bound state, reporter ligands can be used forscreening (31). In this approach, the known ligand is prebound andthe new ligands in the screen are tested for their ability to displacethe bound ligand. For example, in the case of BSA, this approachhas been taken to study tryptophan binding: complementary to the1H NMR studies of BSA described earlier, 19F NMR-based studiesof L-5-tryptophan (32) and L-6-tryptophan (33) binding to BSA havebeen carried out. The extreme sensitivity of the 19F chemical shiftresulted in the observation of two distinct peaks, indicating thepresence of multiple tryptophan binding sites, a low-affinity and ahigh-affinity binding site. Competition with non-fluorinated trypto-phan can be used to establish relative affinities of these ligandswith respect to tryptophan at both sites. Thus, while the 19Fapproach – unlike the 1H approach – is restricted to ligands thatbind at the same sites as 19F-containing ligands do, the 19F NMRstudies proved useful in revealing an additional tryptophan-bindingsite that went undetected with 1H NMR, showing the complemen-tary nature of the approaches.

SummaryIn summary, NMR techniques for drug discovery are high-contentmethods: they potentially provide binding information, the locationof the binding site and the conformation of the bound ligand.Nuclear magnetic resonance can also supply structural informationthat enables the docking of the ligand to the protein's bindingpocket. In addition, NMR provides very valuable information aboutthe general behavior of the ligands that other HTS methods do notreveal, including solubility, binding behavior (promiscuous ligands),precipitation potential and aggregation. Because NMR-basedscreening is sensitive toward finding medium-affinity to low-affinityligands, the approach can also serve as an effective prescreeningtool for subsequent assay-based HTS. Thus, NMR-based screeningfor small molecular weight drugs is now well established in indus-try and can be used complementary to HTS methods and computa-tional screening methods.

Challenges in membrane protein NMRspectroscopy

While 1H NMR-based methods to study ligand binding can be car-ried out with unlabeled protein, more sophisticated applications ofNMR-spectroscopic techniques such as SAR by NMR require label-ing, typically the biosynthetic introduction of 13C and 15N nuclei.However, many proteins cannot be successfully expressed in E. colior Pichia pastoris that make uniform 13C, 15N labeling affordable.When proteins need to be expressed in mammalian or insect celllines to obtain them in functional form, uniform labeling becomesprohibitively expensive when the protein expression levels are notunusually high. In such cases, specific 15N-labeled and ⁄ or13C-labeled amino acids are introduced (34–36). Such proteins arenot amenable to structure determination by NMR spectroscopy.Mammalian membrane proteins often belong to this group, e.g.,

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when they are glycosylated or otherwise post-translationally modi-fied in their native form and require the mammalian or insect cellmachinery for proper folding.

NMR signal assignment requires well-resolved mono-disperse spec-tra as a prerequisite, in which a large number of the NMR-activenuclei in the sample are visible and resolved from each other, andthe signal intensity for different peaks is as uniform as possible.This in part is the reason for the limit in size of biomolecules thatcan be studied, but poor quality spectra can also arise from sys-tems that are dynamic and ⁄ or prone to aggregation even when thesize of the monomeric unit is relatively small, depending on thepropensity of the proteins and choice of detergents. Thus, it is criti-cal to choose suitable detergents for each membrane protein. Afteror complementary to light-scattering experiments, 1H,15N-HSQCspectra are typically recorded to screen for detergents and otherconditions, such as salt concentration and pH, under which reason-able NMR spectra can be obtained. Recent developments in micro-coil NMR technology have the potential to make the screening of alarge number of different detergents for their suitability to supportNMR studies more feasible (37).

We will demonstrate these issues using the GPCR rhodopsin as anexample. Rhodopsin is a glycosylated and palmitoylated 43 kDa pro-tein containing 348 amino acids. 1H,15N-HSQC spectra of either15N-lysine-labeled or 15N-tryptophan-labeled rhodopsin are shown inFigure 2A and B, respectively. The protein was dissolved in 20 mM

sodium phosphate (pH 6.0) and 10% D2O containing octyl glucosideor dodecyl maltoside detergent micelles. The quality of both NMRspectra is quite poor as evidenced by the heterogeneity in numberand intensity of signals (Figure 2). Site-directed mutagenesis andscreening of solvent conditions has led to the improvement in spec-tral quality for some membrane proteins, e.g., diacylglycerol kinase,where the E. coli origin and expression system made such studiespossible (38). When an optimal condition for NMR study is notfound for the membrane protein of interest, fragments of the pro-teins may be studied instead (39). Although such fragments studieswill gain some limited insight into the structure of the membrane

proteins, they typically do not bind ligands in functional formexcluding such systems from NMR-based ligand screeningapproaches.

The reason for the difficulties in obtaining membrane protein struc-tures by NMR is largely based on the fact that NMR signalsbecome broader as the molecular mass increases, leading to thereduction in sensitivity of NMR experiments. Because membraneproteins are studied under conditions surrounded by micellesformed by the detergents, the apparent molecular mass becomeslarger than the protein molecular weight. Also, when membraneproteins form biologically functional or non-functional oligomers, theapparent molecular mass, including the surrounding deter-gent ⁄ micelles, results in further broadening of NMR signals. Thus,several efforts are underway to detect protein NMR signals of largeproteins, which are useful for drug screening and ⁄ or signal assign-ment purposes: fast experiments, TROSY methods and various iso-tope labeling techniques. TROSY in particular has been crucial in allof the recent determinations of membrane protein structures butrequires deuteration. Efforts to detect NMR signals in shorter time,such as 'SOFAST- HMQC' or 'Ultra-fast experiments' may prove use-ful for drug-screening or drug validation purposes (8,9). Because theline widths of methyl signals in these experiments are relativelynarrow as a result of methyl three-site jump and the TROSY selec-tion can further increase sensitivity (40–42), observing the methylsignals becomes advantageous for large macromolecular systems,including membrane protein systems. Several excellent review arti-cles describe these techniques (43–46).

Despite such difficulties in protein expression and sample prepara-tion, there is increasing success in the determination of membraneprotein structures by NMR spectroscopy. To illustrate this progress,we downloaded a list of membrane protein structures determinedwith the help of NMR spectroscopya and analyzed the structureswith respect to their transmembrane organization (Figure 3). Of 44structures, 28 structures were determined using solution NMR (theothers utilized solid-state NMR). While these numbers are encour-aging, it is important to realize that the majority of these structures

A B

Figure 2: 1H,15N-heteronuclear single quantum correlation (HSQC) spectrum of rhodopsin labeled with (A) a-15N-lysine and (B) a,e-15N-tryp-tophan. Rhodopsin contains 11 lysine residues but only one of these, Lys339, gives rise to a high intensity peak (labeled in the figure) (35).There are a total of five tryptophan residues in rhodopsin, the signals corresponding to backbone and side-chain signals are represented by'a' and 'e', respectively (36). Reprinted with permission from the Proceedings of the National Academy of the United States of America (Copy-right ª 2002, The National Academy of Sciences, Copyright ª 2004, The National Academy of Sciences).

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still represents either b-barrel proteins (Figure 3, '0¢ bin) or singletransmembrane helices (Figure 3, '1' bin). A recent success was thestructure determination of diacylglycerol kinase (Figure 3, '3' bin),which although only consisting of three transmembrane helicesforms a trimer. The trimeric organization is significant because it isformed via domain-swapping of helices. Thus, the structure actuallyrepresents with 9 (!) transmembrane helices the largest membraneprotein whose structure has been determined by NMR spectroscopyto date (3). These results are highly encouraging: a decade ago,only the structures of small membrane proteins with molecularweights less than 10 kDa could be determined by NMR because ofthe decrease in the molecular tumbling by the addition of deter-gents (47). However, recent developments of NMR methodology andefforts of protein expression and sample preparation enabled theearlier mentioned structure determinations for membrane proteinswith molecular weight >20 kDa.

Finally, it should be noted that in the application of NMR-screeningmethods to membrane proteins by looking at ligand signals, it isimportant to distinguish whether signal changes are because ofligand–detergent interaction or ligand–protein interaction. It is thuscritical to record a suitable reference spectrum in each case.

1H NMR-based approaches for membraneproteins

Solution NMR spectroscopy has dramatically advanced in the scopeof its applicability to proteins, especially when studying proteins ofincreasingly larger size or membrane proteins, by way of usingNMR-active isotopes of hydrogen, carbon and nitrogen. While 1H is100% abundant, 15N and 13C isotopes are used to replace the moreabundant 14N and 12C isotopes in proteins, respectively. The abilityto introduce these isotopes is therefore one constraint on the

applicability of NMR spectroscopy to the study of proteins in gen-eral, including protein-ligand interactions. The natural abundance ofthese isotopes in the detergents and solvents used can significantlyadd to the background, in particular for 1H NMR spectroscopy,where the 1H isotope is 100% abundant. Additional problems arethe low signal-to-noise ratio because of slow molecular tumbling ofthe protein–detergent complex discussed earlier. In the followingparagraphs, we summarize current efforts in overcoming these con-straints, with major emphasis on recording 1H NMR spectra. Similarconsiderations however would also apply to the direct detection ofother isotopes such as 13C.

Suppression of background signals in NMRexperiments for membrane proteinsAs described in 'Challenges in membrane protein NMR spectros-copy', in the case of membrane proteins, a membrane mimetic isrequired, provided by detergent micelles when they are studied withsolution NMR methods. The detergent concentrations are typically100 times higher than the protein concentrations to ensure that onlyone functional protein or protein complex is present per micelle foruniformity purposes. The high signal intensity originating from thedetergent leads to the suppression of signal intensities from the pro-tein (dynamic range problem) and also results in overlapping withthat of protein peaks. Over-sampling is a feature available in mostrecent commercial NMR instruments, but if it is not available, largedetergent signals also cause other artifacts such as baseline rollingand insufficient digitization of the signal (48). An example is shownfor a 0.7 mM solution of rhodopsin in 1% octyl glucoside (Figure 4).At the scale used, the protein signals are not even visible in thisFigure, and the spectrum is dominated by the detergent signals. Avalue of 1% for the detergent concentration is in fact relatively low;in many cases, much higher detergent concentrations are used,making the dynamic range problem even more severe.

A biochemical solution to the detergent background problem is theuse of deuterated detergents. However, their synthesis is typically

Figure 3: Analysis of integral membrane protein structuresdetermined by NMR spectroscopy deposited in the protein data-bankb. The y-axis represents the number of protein structures witha particular transmembrane segment organization plotted on the x-axis. The x-axis represents the total number of transmembrane heli-ces in each structure. The '0' category corresponds to b-barreltransmembrane proteins. The PDB identifiers that represent eachcategory are '0' (1G90, 2JMM, 2K0L, 1MM4, 1MM5, 1Q9F, 2JQY,2K4T, 2JK4), '1' (1AFO, 2RLF, 1ZLL, 2HAC, 2J5D, 2JO1, 1JP3, 2JWA,2KIK, 2K1L, 2K21, 2K9J), '2' (1WAZ, 2A9H, 2JX4, 2K9P), '3' (2KDC)and '4' (2K73, 2K74). The data for the plot were downloaded onNovember 26, 2009 from Dror Warschawski's websitea.

Figure 4: One-dimensional 1H NMR spectrum of bovine rhodop-sin acquired in 20 mM sodium phosphate buffer (pH 6.0) and 1%octyl glucoside. The spectrum was acquired using a 800 MHz Bru-ker spectrometer, at 20�C. At the scale shown, only the detergentsignals are visible, demonstrating the large difference between theintensity of detergent and protein signals.

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very expensive. Unless the protein can be studied in commonly useddetergents for which deuterated forms can be purchased off theshelf, custom-synthesis is often required. In addition, use of deuter-ated detergent for screening large numbers of samples mayincrease the screening cost significantly. The type of detergent thatwill give rise to optimal NMR spectra while maintaining the func-tion of the protein is largely empirical, requiring extensive screeningof different detergents and detergent ⁄ lipid mixtures and may settleon non-standard detergents (37,49,50). Membrane proteins have tobe continuously maintained in the presence of membrane mimeticsduring cell extraction (or after refolding from inclusion bodies).Further, all purification and concentration steps require largevolumes of buffers. Because of these reasons, typically the proteinwill be purified in a non-deuterated detergent, followed byexchange with the deuterated detergent. This adds an additionalstep of complexity to the NMR sample preparation to ensure effi-cient, homogenous and complete replacement of detergent withminimal protein loss. Thus, use of deuterated detergent may notalways be practical based on cost and preparative effort, especiallyat the relatively large quantities needed for NMR-based screening.

When deuterated detergent is not available, too expensive or notpractical, application of multiple solvent suppression experiments,such as WET (51), selective pulse experiments including sculpting(52,53) or coherence selection (54–56), is required. If possible, satu-ration by radio-frequency is not applied to suppress the water, sol-vent or detergent signals in protein samples because proteinsignals underneath the solvents are also saturated and the signalreduction is propagated to the entire protein by the spin-diffusionmechanism (57). Among the water suppression techniques, pulsetechniques that use relatively long durations are not efficient to beincorporated into various 3D NMR experiments and coherenceselection in combination with pulsed-field gradient is commonlyapplied.

Because one-dimensional NMR-spectroscopic approaches currentlyhave (and in the foreseeable future will continue to have) broader

applicability to membrane proteins, solvent suppression schemessometimes with loss of information in some regions of the spec-trum are particularly important. The earlier described AIDA method(53) also makes use of focusing on a particular spectral region (seeFigure 1). Here, we demonstrate the utility of such an approachusing selective excitation sculpting studies of full-length rhodopsinin octyl glucoside micelles as a model system. Rhodopsin is themost extensively studied G-protein-coupled receptor, and knowledgeabout its structure serves as a template for other related receptors.Because of the large numbers of members of the GPCR family andtheir importance as drug targets (see Introduction text of this articleunder Abstract), these studies are highly relevant for drug discoveryefforts involving these receptors.

One-dimensional 1H NMR spectra recorded by selectively excitingthe protein NH region by applying a selective excitation pulse cen-tered around 10–12 ppm show 1H chemical shifts from both back-bone and side-chain regions of rhodopsin in octyl glucoside micelles(Figure 5A). Further, excitation of the same region using the hyper-bolic secant shaped pulse to remove detergent and water signalssignificantly increased the intensities of the NH peaks in the rangefrom 6.0–8.5 ppm (Figure 5B) (58,59). Note, however, that the num-ber of peaks observed in the 1D 1H NMR spectrum is significantlyreduced. We tentatively propose that the observed signals arisemostly from the backbone C-terminus residues and flexible loopregions. This hypothesis is based on the previous observation (35)that sharp, highly intense and thus slowly relaxing signals arefound only for Lys339 in a uniformly 15N-lysine labeled rhodopsinsample (Figure 2). Furthermore, comparison between the observedsignals and those obtained with a peptide corresponding to thesequence of the C-terminal residues reveals extensive similaritiesbetween the rhodopsin C-terminus and the free peptide in solution(60).

One-dimensional 1H NMR spectra of bovine rhodopsin recorded atdifferent concentrations of octyl glucoside indicated chemical shiftdependence of the C-terminus backbone peaks (data not shown),

A B

Figure 5: One-dimensional NMR spectra of unlabeled bovine rhodopsin in octyl glucoside micelles. (A) Selective excitation of the NHregion by employing a selective excitation pulse. (B) Selective excitation of the NH proton peaks with sculpting using hyperbolic secantshaped pulse (58,59). A total of 0.5 mM (7 mg in 350 lL) concentration of bovine rhodopsin was used to acquire the spectra. The NMR exper-imental parameters pulse width, excitation bandwidth and acquisition time are as provided in the legend to Figure 7 and detailed in (61).

NMR-Based Screening


highlighting the need to control the detergent environment quantita-tively to obtain reproducible NMR results. To investigate possibledetergent–protein interactions, we recorded one-dimensional andtwo-dimensional 1H-1H selective excitation NOE spectra. Weobserved differential interactions of the rhodopsin backbone signalswith those of the detergent micelles (Figure 6). In particular, a setof strong NOE peaks was observed from rhodopsin protons(Figure 6B, represented by arrows) to a detergent peak at �1.85ppm (Figure 6A, indicated by arrow). The identity of this detergentsignal is shown as an inset in Figure 6A, a -CH2- group near thesugar head group. We did not observe intramolecular rhodopsinprotein NOE peaks. A potential solution to detect such NOEs couldbe provided by detergent deuteration.

Using the sculpting experiments, we have successfully identifiednovel ligands binding to rhodopsin and interacting with cytoplasmicloop and C-terminal residues by measuring chemical shift and line-broadening effects in selectively excited 1H spectra as a function ofadded ligand, the anthocyanin cyanidin-3-glucoside (61). In thisstudy, we were able to identify chemical shift and intensity changesin receptor and ligand. In dark-adapted rhodopsin an upfield shift ofthe chemical signals (Figure 7, peaks at position 3, 4, 7, 8, 9 and10) of the protein was observed. In the case of ligand, some of thepeaks corresponding to ligand (compare signals at position 2, 11,14, 18 and 19 in Figure 7A with 7D) experienced decrease in inten-sity and some of them disappeared (peaks marked as 'x' and atpositions 22, and 24 in Figure 7) in the presence of rhodopsin, indi-cating restriction in mobility upon binding. Further, the comparisonof the 1H NMR spectra of rhodopsin upon light activation both inthe absence and presence of ligand indicated decrease in peakintensities at peak positions represented as '+' in Figure 7C. Usingthe selective excitation sculpting method, this study suggested thatthe binding of anthocyanin ligand, cyanidin-3-glucoside, modulates

both the structure and the dynamics of rhodopsin in two differentstates, the inactive dark state and the light-activated Metarhodop-sin II state. The approach is extendable to other conformations,such as G-protein-bound or opsin structures.

The results obtained with rhodopsin show high promise for theextension of the approach to other GPCRs. We have already demon-strated with rhodopsin that multiple conformations can be studied,because the life-time of these conformations under the NMR condi-tions studied are known. For other GPCRs, it also needs to beestablished what the stability of resting, activated or G-protein-bound states are, to ensure that the time it takes to acquire anNMR spectrum is meaningful for the particular conformation ofinterest. Furthermore, while the cytoplasmic loops and the C-termi-nus of rhodopsin are functionally important regions in the protein(critical for receptor activation and G-protein binding), it remains tobe shown whether the same approach is also suitable to studyligands such as retinal that bind in the transmembrane domain ofrhodopsin.

Saturation transfer-difference (STD) NMRapplication to membrane proteinsOf the many techniques developed for screening by NMR, summa-rized in 'NMR-based approaches to drug screening', a particularlypromising technique for application to membrane proteins is STD.The technique requires very small amounts of protein (in the nM–lM range) because the ligand is present in 100-fold excess overthe protein (7). Protein signals are saturated by irradiation around)1 ppm, which is transferred within �0.1 seconds to the rest ofthe protein and the ligand. When the ligand off-rate is fast, theinformation is quickly transferred to the ligand in solution where itdecays slowly (within �1 seconds), so that during saturation, the

A

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Figure 6: (A). One-dimensionalsolution selective NOE 1H NMRspectrum of bovine rhodopsin in0.15% octyl glucoside recorded at600 MHz, 25�C. (B). Two-dimen-sional solution 1H – 1H NOE spec-trum of bovine rhodopsin in 1%octyl glucoside. The NOEs from oneof the detergent peaks (marked withan arrow in Figure 6A) to the 1Hpeaks from rhodopsin (representedin box in Figure 6A) are indicated byarrows in 6B.

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proportion of saturated ligands in solution increases, amplifying thedifference signal, up until the ligand excess concentration isreached. Thus, the intensity of the STD spectrum will be higher forligands with fast off-rates, but even tight binding can still be mea-sured, giving the technique a wide dynamic range. This approachhas already been used for study of ligands targeting membrane pro-teins by NMR (18,62). In one study, integrins were embedded inDMPC ⁄ DMPG liposomes and binding of cyclic peptides was tested(18). An affinity of 30–60 lM was obtained, typical for this class ofmembrane receptors and demonstrating the particular utility ofNMR-based approaches to reliably detect relatively low affinities.From differences in STD responses of individual protons in the cyc-lic peptide, it was even possible to map the epitope that is in

direct contact with the receptor to a phenyl ring in the peptide.Only 0.25 nmol of the integrin was sufficient per assay. Anotherspectacular application of STD to membrane proteins is the recentstudy of the interaction of the sweet brazzein protein with thehuman sweet receptor (62). This receptor is a Class C GPCR, con-taining a large extracellular ligand-binding domain, coupled to theseven-transmembrane helical bundle typical for GPCRs. These recep-tors are challenging and interesting because they contain multiplebinding sites in both transmembrane and extracellular domains andhave very low affinity for their ligands, ranging from lM to mM.The ligands can bind simultaneously and affect each other's affinity,thus it is imperative that the full-length native receptor is studied.One-dimensional 1H,15N HSQC STD experiments demonstrated the

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Figure 7: 1H NMR spectra acq-uired using selective excitation sc-heme with sculpting. (A) Rhodopsinbefore (black solid line) and afterthe addition of ligand, cyanidin-3-glucoside (red dotted line). (B)Rhodopsin in the presence ofcyanidin-3-glucoside before (blacksolid line) and after light activation(red dotted line). (C) Illuminatedrhodopsin in the absence (blacksolid line) and presence of cyani-din-3-glucoside (red dotted line).(D) Cyanidin-3-glucoside alone inphosphate buffer and 0.6% dodecylmaltoside. Each spectrum wasobtained after applying two 180�hyperbolic secant pulses, followinga 90� rectangular pulse, withcarrier frequency at 11.5 ppm. Thefirst and second 180� pulses wereemployed to invert 6000 and 8000Hz spectral ranges, respectively.The last rectangular pulse wasapplied for 9.9 ls. Echo delay forthe first and the second 180�pulses were set to be 0.2 and1 ms, respectively. A total of 2048scans were acquired with0.5 seconds repetition delay usinga 800 MHz proton resonance fre-quency. Reprinted with permissionfrom the Blackwell Publishing.

NMR-Based Screening


binding of brazzein to the sweet receptor (�100 lg) in membranesuspensions with high intensity, while a non-sweet mutant brazzeinprotein did not give rise to strong STD signals. This level of proteinamounts without purification requirement (because membrane prep-arations were used) is in our experience relatively straight-forwardto obtain for many GPCRs. Thus, the approach is likely to havebroad applicability to other membrane receptors. Given that theSTD technique is highly sensitive and neither limited by protein sizenor requires the assignment of the protein, this technique shouldfind wide applicability to screening of ligands for membrane pro-teins that have lipid or detergent environments surrounding them.

19F NMR-based approaches

19F NMR spectroscopy can be a viable alternative for one-dimen-sional NMR-spectroscopic measurements, providing complementaryresults. Because there is no background from 19F nuclei in neitherbiomolecules such as proteins nor detergents used to dissolvemembrane proteins, the applicability range of 19F NMR to studyligand binding in soluble and in membrane proteins is identical. Inthe following paragraphs, we therefore review the extensive litera-ture on 19F NMR-based approaches to study ligand binding to pro-teins, regardless of the proteins under investigation being solubleor membrane proteins. First, we will review 19F ligand-observe stud-ies using fluorinated ligands, including fluorinated phospholipids.We will then cover studies of structure and dynamics of proteins by19F NMR. These studies will involve not only ligand-inducedchanges in structure and ⁄ or dynamics but also those involving otherconformational changes, such as during protein function or proteinfolding, because the principles are the same.

19F NMR studies of protein structure, dynamics and ligand bindingoffer several advantages over other NMR-spectroscopic approachesas a result of the unique chemistry of the 19F atom. 19F has 100%natural abundance, and its sensitivity to NMR detection is 83% thatof 1H. The presence of nine electrons surrounding the 19F nucleusmakes it very sensitive to minor changes in its environment, includ-ing both Van-der-Waals and electrostatic interactions, which isreflected in its wide range of chemical shifts. This characteristicincreases the probability of obtaining well-resolved peaks of fluo-rine atoms in different environments. Another major advantage of19F NMR over other conventional NMR techniques is the appear-ance of its NMR signals in the absence of any background signals,including membrane mimetic environments and even entire cells.The information content of 19F NMR ligand-based screening, whilenot as high as SAR by NMR, is higher than that of HTS methods,in particular those employing cell-based approaches. These uniqueproperties of the 19F nucleus suggest that 19F NMR spectroscopycould provide a highly desirable alternative to HTS by conventionalNMR-spectroscopic techniques, in cases where the latter methodsmay not be applicable, such as for many membrane proteins or forin-cell studies. From a practical perspective, 19F labeled compoundsare easily accessible by different chemical methods (see 'Synthesisof 19F containing small molecule compounds').

Ligand–protein interaction studies include (i) evaluating binding ofligands, (ii) characterizing binding kinetics of the ligands and (iii)

determining the structural changes of a protein on ligand binding.These are probed by changes in line shape and ⁄ or chemical shiftof a free fluorinated ligand on binding to a protein (19F ligand-observe studies) or that of a fluorinated residue in a protein onligand binding (19F protein observe studies). Both approaches canbe employed in the context of drug screening (19F NMR-basedligand screening).

19F ligand-observe studiesSpectral changes of a free fluorinated ligand on binding to a pro-tein – like in the case of 1H NMR – can be either broadening ofits line width or changes in its chemical shift depending on thebinding affinity of the ligand. Fluorine signals of the ligand boundto the protein are expected to show restricted motion compared toits free state and hence give a broader line shape. It may alsoundergo chemical shift changes upon binding that may be eitherupfield or downfield depending on the nature of the change ofinteractions of the fluorine atom with its environment. A downfieldshift indicates a more hydrophobic environment or a greater extentof Van-der-Waals interaction of the fluorine atom. Changes in elec-trostatic interactions of the fluorine atom with its environment caninfluence either a downfield or an upfield shift (63). Note however,structural information of the binding site can only be procured byobserving changes in fluorinated protein on ligand addition.

Ligands with a low binding affinity rapidly exchange between boundand free forms that may lead to broadening of its resonances. Theadvantage of characterizing ligand–protein interactions of suchweak binding ligands by studying changes in fluorinated ligandsrather than protein observed changes is the requirement of lessamount of protein. Binding constants can be determined by T2 mea-surements that contain a weighted average of relaxation rates ofthe free and bound forms of a ligand at different concentrations(64). The utility of T2 measurements has for example been demon-strated for BSA in binding studies of isoflurane, a volatile anes-thetic (64). A Kd of 1.4 mM was obtained from T2 measurements ofthe free ligand and that bound to the protein (64). Another interest-ing case is the influenza virus M2 membrane protein, which formsproton channels that lead to the disruption of the matrix proteinand the release of the viral genome (65). Amantidine is an inhibitorof this process. 1H NMR of amantidine or the protein could not pro-vide information on ligand binding because very broad signals wereobtained (66,67). 19F T2 relaxation measurements were used in thiscase to reveal interactions between the fluorinated amantidineligand, and the M2 protein as well as interactions between theligand and the dodecylphosphocholine micelles the protein wasdissolved in (67).

Inhibitors of enzymatic reactions may be detected by a methodcalled fluorine-based biochemical screening (FABS) (68,69). In thismethod, a substrate is tagged with a fluorinated moiety, andchanges in distinct 19F signals for the substrate and product are fol-lowed with the progress of an enzymatic reaction in presence oftest inhibitors. This method is particularly suited for screening inhib-itors with low-binding affinity that remain undetected by regularNMR ligand screening methods. The sensitivity of the method isenhanced in the case of weak affinity ligands by having moietieswith three fluorine atoms attached to the ligand and the method is

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named 3-FABS (69). IC50 value of the inhibitors is obtained by tak-ing the ratio of the integrals of the 19F peaks of the substrate andthe product as a function of inhibitor concentration. In addition toscreening mixtures of inhibitors, it is also possible to screen mix-tures of closely related enzymes to determine selectivity of aninhibitor provided the substrate is specific for the different enzymes.This method has been applied in several cases such as screeninginhibitors for kinase AKT1 and protease trypsin (69), caspases (70)and thymidine phosphorylase (71).

Information on binding constants and stoichiometry of binding canbe obtained by titrating fluorinated ligand and monitoring thechanges in the protein-bound peaks and free peaks of the ligand by19F NMR. In the slow exchange regime, we will observe two peaks,which may be sufficiently resolved in their chemical shift values tobe useful for quantitation. Binding constants are determined fromthe ratios of bound and free ligand concentrations quantified byintegrating 19F NMR signals (72).

19F protein observe studiesStudying ligand binding by monitoring changes in 19F signals report-ing on protein conformation can be useful under conditions whereaccurate affinities and binding modes cannot be unambiguouslydetermined from ligand-observe methods, or where it is desirableto increase the information content that can be obtained from 19FNMR studies. If 19F labels are placed on the protein, one can studywhere the ligand binds, and whether the ligand induces conforma-tional changes, oligomerization or folding transitions. There are twoapproaches to introduce 19F labels into proteins. In the firstapproach, a 19F label is introduced biosynthetically as a fluorinatedamino acid. As for incorporation of other isotope-labeled aminoacids (see above), this method may not be very cost effective formammalian membrane proteins (including GPCRs) because insect ormammalian cell expression required for such systems in fluorinatedamino acid-rich medium can be very expensive. In the secondapproach, a 19F label is introduced through chemical reaction with

activated cysteines. This approach has been shown to work wellwith GPCRs (73). However, this method is limited to labeling onlysurface exposed amino acids or those amino acids in membraneproteins for which side chains are exposed to the membrane forease of entry of labeling reagents. The principle is shown inFigure 8. A receptor will have endogenous cysteines, shown in ahomology model of the corticotropin-releasing factor receptor inFigure 8A. The cysteines can be derivatized with a 19F containingligand directly, but a less invasive approach is to first activate theaccessible cysteines and then introduce a trifluoroethylthiol groupthrough disulfide exchange (Figure 8B). This procedure contains min-imal perturbation from added chemical groups and retains maximalflexibility from the ethyl side chain.

Using 19F NMR to observe the protein can be useful, for example,if it is of interest to determine whether a receptor is in an activeor inactive conformation upon ligand binding. If the specific chemi-cal shifts associated with each state are known, then the appear-ance of the respective peaks can be used as an indicator whethera ligand is, for example, an agonist or antagonist or an inhibitor orinducer of oligomerization. This idea is illustrated with bovine rho-dopsin: 19F NMR spectroscopy was used to study the conforma-tional changes in rhodopsin upon light activation to which the 19Fchemical shifts were very sensitive (73). In this case, the 19F labelwas introduced through chemical reaction of trifluoroethyltiol withactivated cysteines (Figure 8B), here on rhodopsin. Distinct chemicalshifts are found for the dark, inactive and the light-active states atnumerous sites on the rhodopsin surface (Figure 9).

Determining structural changes in specific regions of a protein onligand binding requires the introduction of a 19F label into the pro-tein. More common than the chemical cysteine-labeling approach,is to substitute amino acids in the protein with fluorinated analogsand track the chemical shifts and line widths in 19F NMR spectra.The small size of the fluorine atom has enabled the substitution ofresidues such as Trp, Tyr, Phe with their fluorinated analogs withoutperturbations of the native structures of proteins. The observed

A B

Figure 8: Selective cysteineCF3-derivatization of G-protein-coupled receptors (GPCRs). (A) Asan example, the five endogenouscysteine residues in the corticotro-pin-releasing factor receptor (CRFR)are shown in yellow. (B) Chemicalprocedure of selective cysteinederivatization via activation andthiol exchange (73,74). A sulfhydrylgroup on the protein (here GPCR)is activated by reaction withdithiodipyridine. The thiopyridinylderivative undergoes disulfideexchange with a fluorine-contain-ing sulfhydryl reagent (73).

NMR-Based Screening


chemical shift range, expression of the labeled protein in sufficientamount and integrity of the fluorine labeled protein are some ofthe factors that should be considered when choosing an isomer ofa fluorinated amino acid. For example, of the 4-fluoro, 5-fluoro and6-fluoro isomers available for fluoro-tryptophan, the 6-fluoro isomerhas a very narrow chemical shift range and also shows broadunresolved spectra compared to the other two fluoro-tryptophans inlactate dehydrogenase enzyme (75). Moreover, the 6-fluoro isomer-labeled protein shows perturbations in its secondary structure asdetected by circular dichroism spectroscopy, and a broad peak isobtained in the 19F NMR spectrum (75). On the other hand, the

4-fluoro isomer labeled protein can be produced in much largerquantity and shows no perturbations of the native structure (75).Assignments of the 19F peaks can be performed by either mutatingthe fluorinated residue or by nudge mutations, whereby a mutationin an adjacent position changes the chemical shift of thefluorinated residue as a result of change in its environment, or bycomplexation of a solvent accessible fluorinated residue withparamagnetic ions such as Gd3+ leading to line broadening of thatresidue (76).

19F NMR has been used to track both allosteric and non-allostericchanges on ligand binding to a protein. For example, in studies ofthe binding of D-glucose and D-galactose to the fluoro-tryptophan-labeled aqueous chemosensory receptor of E. coli (77), it was seenthat sugar binding resulted in changes in chemical shifts of not onlythose fluoro-tryptophan residues that are adjacent to the bindingsite but also those tryptophan residues that are distant from thebound sugar by as much as 15 � (77). These results indicate thatsugar binding leads to a global change in the structure of the pro-tein that is translated from the binding site to distant regions onthe surface, and this global change can be tracked by 19F NMR(77). A different way of probing conformational change is toobserve line broadening by the addition of Gd(III)-EDTA that indi-cates solvent accessibility of the fluorine-labeled residue (78). Infor-mation on binding constants and stoichiometry can be obtained bytitrating the ligand and monitoring the shifts in the peaks of fluori-nated amino acids (78). 19F NMR has also proved to be suitable forstudying protein dynamics by monitoring relaxation rates of fluori-nated residues, as illustrated by the study of ligand binding in iono-tropic glutamate receptor (GluR2) (76).

Structure and function of membrane proteins in particular are lar-gely influenced by their interactions with lipid bilayers, and 19FNMR can be used to study the detailed mechanisms of theseeffects. For example, line widths of lactate dehydrogenase becomesharper on adding increasing concentrations of lysolecithin in anon-linear fashion (75). Because there was no change in the chemi-cal shifts of the tryptophan residues, it was concluded that lysoleci-thin is only solvating the protein and not causing a conformationalchange (75). The number of lipid molecules bound to a protein canbe calculated from the variation in line width with lipid concentra-tion. In the case of lysolecithin binding to lactate dehydrogenase,this number was found to be lower than the aggregation number oflysolecithin, suggesting that lactate dehydrogenase is not insertedin the micelles but binds individual lipid molecules that shieldexposed hydrophobic surface patches from initiating aggregationand inactivation of the enzyme (75).

19F NMR is a suitable technique in mapping the sites of the inter-action of proteins with membranes. The use of solvent induced iso-tope shifts can provide information on solvent exposure of aresidue. However, residues that are not solvent exposed could beeither buried in a protein core or face the membrane or be mem-brane bound. This ambiguity can be overcome by the use of fattyacids in which a paramagnetic spin label is incorporated into themembrane under study, and its interaction with a fluorine probe inthe protein is detected by the broadening of the corresponding fluo-rine peaks in a 19F NMR spectrum (79). The paramagnetic electron

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Figure 9: 19F one-dimensional NMR spectra of trifluoroethylthi-ol-labeled bovine rhodopsin, and its various cysteine mutants indark (red lines) and after illumination (blue lines) (73). The spectrumwas referenced with respect to trifluoroacetic acid (TFA). Reprintedwith permission from the Proceedings of the National Academy ofthe United States of America (Copyright ª 1999, The NationalAcademy of Sciences).

Yanamala et al.


of the labeled fatty acid 7 � from either end of the lipid phase willcause broadening of a fluorine nucleus that is within 15 � from thelabel i.e., either in or near the lipid phase (79,80). By labeling spe-cific amino acids with 19F and by their mutagenesis analysis, inter-actions with lipids can be followed, thus helping in mapping sitesof protein–lipid interaction. The amount of broadening observed isinversely proportional to the distance between the label and fluori-nated residue raised to the power of six (78). 8-doxylpalmitic acidincorporated in lysophosphatidylcholine is used as the nitroxidespin-labeled fatty acid to map the site of interaction of lactatedehydrogenase with lysophosphatidylcholine (80). Another use ofsuch spin-labeled fatty acids, in the case of lactate dehydrogenase,is to determine whether substrate binding has any effect on theresidues in the lipid binding region. Lactate dehydrogenase oxidizesD-lactate, and the electrons produced reduce the nitroxide labeledfatty acid, disrupting its interactions with the fluorine nucleus andrecovery of the peak that was lost ⁄ broadened because of its inter-action with the label (80,81).

19F NMR-based ligand screeningThe ease of obtaining information from ligand-binding studies by19F NMR, as mentioned earlier, has extended its applicability toHTS of chemical libraries that is a routine procedure in the field ofdrug discovery. The broad chemical shift dispersion of the fluorinenucleus allows for identifying 'hits' in a screen with less chances ofencountering the problem of spectral overlap from different chemi-cal compounds. The simplicity of the 19F spectra, unlike 1H spectra,decreases the time for deconvoluting the spectra when a large mix-ture of chemicals is being screened. Changes in chemical shift val-ues and ⁄ or line widths of the free fluorinated ligand upon theaddition of a protein will indicate whether a compound is bindingto the protein or not. Thus, monitoring free ligand peaks allows theuse of very low protein concentrations, in tens of lM range. Infor-mation on binding constants and stoichiometry of binding fromligand titration experiments can be further used to rank orderligands in a screen. Such information was obtained while screeninga library of compounds for chaperones PapD and FimC, involved inthe assembly of pili on E. coli, and are essential proteins thatrepresent targets for the development of antibacterial agents (82).19F NMR studies can also be used to provide further information onbinding sites to optimize the lead compound by characterizing thestructural changes induced by their binding. This is performed byusing proteins substituted at different positions by fluorinatedamino acids and monitoring their chemical shift changes on ligandbinding. This is much less expensive and easier compared to1H NMR where the spectra are complicated and furtherdeconvolution requires expensive isotope labeled samples of highconcentration.

There is a concern regarding availability of a library of fluorinatedcompounds. However, about 12% of the compounds in AvailableChemical Directory of Screeningd compounds contain fluorine. Asdescribed earlier, there are a few drawbacks of the ligand-basedscreening methods if the ligand (i) has very high affinity because ofthe insensitivity of NMR to detect ligand peaks in sub-lM concen-tration ranges (ii) has slow kinetics and (iii) binds to the protein viaa covalent bond. However, these problems are overcome by ligand-

based competition-binding experiments in which 19F NMR signals ofa spy molecule, which has medium to weak affinity for the proteinof interest, is monitored as it is displaced by higher affinity ligandsduring a screen (83,84). This places a constraint on the types ofligands that can be identified with this method, as the ligands haveto exhibit sufficient affinity to compete with the spy molecule,thereby limiting the affinity range of binders. Another limitation isthat as in other competition binding experiments, this method canonly study ligand binding to previously known binding site. Controlmolecules, which do not interact with the protein, can also be usedalong with the spy molecule in this method. Therefore, the screensare performed by monitoring the relative signal intensities of thespy and the control molecule (83,84). The protein is then added tothe mix of spy and control molecule and the NMR signal of the spymolecule disappears as a result of binding to the protein (83,84). Ahit in the screening process is indicated by the reappearance of thespy molecule signal at the same place as before the protein wasadded indicating displacement of the spy molecule with a com-pound of higher affinity from the library (83,84). The extent of dis-placement can be measured from the ratio of the control to spymolecule signal intensity that will in turn provide the binding con-stant of the hit (83,84). The choice of the spy and control moleculescan be decided by their solubility in aqueous solution so that non-specific binding to proteins can be ruled out. A major advantage ofthis method is the requirement of only the spy molecule to be fluo-rinated and not the ligands being screened. This approach is knownas fluorine chemical shift anisotropy and exchange for screening(FAXS) (83). The FAXS method has been successfully used to screenlibraries for human serum albumin where the binding constant of ahit was found to be in good agreement with that obtained fromother techniques such as fluorescence spectroscopy and isothermaltitration calorimetry (85). Human serum albumin concentrations aslow as 600 nM were used (85), showing that the use of very lowprotein concentrations is a major advantage of FAXS over otherNMR-screening methods. This is especially beneficial for findingpotential ligands for membrane proteins that are important drug tar-gets but are difficult to be purified in large amounts. This methodwas also used to screen ligands for the kinase domain of p21-acti-vated kinase (84). Apart from its use in HTS, FAXS has been verysuitable for fragment-based screening of potent ligands. The usehas been illustrated in screening fragments against v-Src SH2domain that has a high affinity for phosphotyrosine (86).

For HTS of ligands, ligand titrations to obtain binding affinities arenot always feasible because of (i) time-consuming titration proce-dure and performing relaxation experiments for each titration point(ii) aggregation arising from addition of excess ligand during titra-tions for ligands with medium affinities and (iii) loss of native char-acteristics of a protein by the addition of the increasingconcentrations of ligands dissolved in organic solvents. A different'titrationless' method has been developed based on gxy and R2

measurements (87). gxy is transverse cross-correlation rate constantof a fluorine attached to an aromatic ring and its ortho-proton andR2 is the transverse relaxation rate constant. The ratio gxy ⁄ R2 givesa more accurate estimation of the exchange rate constant than thatobtained from the more conventional R1q (rotating frame relaxationrate) measurement. This in turn gives a more accurate dissociationconstant of the ligand (87).

NMR-Based Screening


As a proof of concept for extending these approaches to membraneproteins, we screened binding of 19F-labeled small molecules torhodopsin by mixing the ligands with the receptor. Ligands were ina mixture of 10 compounds at 50 lM concentration each. Thereceptor concentration was 0.2 mM in detergent solution (fivefoldexcess). For a ligand with micromolar affinity, these conditionsensure that the majority of the ligand will be bound, and thereforea maximal peak shift is expected for a hit. Excellent signal-to-noiseratio can be achieved with 7 min acquisition time (Figure 10). Bothline-width and chemical shift changes were observed.

A fragment-based library can be considered complimentary to alibrary of compounds for HTS purposes. Such a library is a collec-tion of fluorinated fragments based on Local Environment of Fluo-rine (LEF) (88). The collection of chemical fragments covers awider chemical space than HTS libraries, and the 'hits' obtainedin a fragment library screen would lead to faster 'lead' optimiza-tion. Many parameters are kept in mind during the building ofsuch a fluorinated fragment library. For example, local substituentsaround the fluorine atom influence its chemical shift dispersionand solubility. Usually, a single chemically equivalent fluorine ispreferable, because more than one non-equivalent fluorine atomwould lead to complex 19F NMR spectra. The fragments are clus-tered according to their global structural features and local envi-ronmental fingerprints into different global and local clusters sothat the library has a good coverage of different environmentsaround the fluorine atom. These fragments are then mixed intotwo batches: one for CF3 containing molecules and the other forCF-containing molecules. The fragments are screened by collecting19F NMR spectra in the absence and presence of a protein andconsidering those signals as 'hits' that are perturbed on proteinaddition. The screening can be further confirmed by recording thesame spectra in the presence of a known ligand. The advantageof this method is that it uses fewer concentrations of the frag-ments, thus enabling the testing of a large compound mixture andalso lowering the protein concentration to be used. The low frag-ment concentration is also helpful in not limiting the use ofligands that have low water solubility.

Comparison of 1H and 19F-NMR-basedversus conventional screening ofmembrane proteins

The main advantage of drug discovery by NMR spectroscopy whencompared to traditional HTS methods using other spectroscopic orcell-based assays is its high-information content: in addition toligand binding itself, the location of binding, affinities and confor-mational changes induced in the protein can be observed. Further-more, as a result of the high sensitivity of NMR spectroscopy tomolecular size, artifacts arising from low solubility of the ligand orability of the ligand to precipitate the protein virtually never goundetected, unlike in traditional HTS approaches. However, thestringent requirements are also the main disadvantage, limiting theapplicability of traditional NMR-based approaches to small solubleproteins. However, these difficulties can be overcome by using spe-cialized 1H-based approaches and 19F-NMR-based approaches,which open the door to study of proteins that are otherwise out ofreach for NMR, including large and ⁄ or multimeric soluble proteincomplexes and full-length membrane proteins in detergent micelles.

An example demonstrating the limitations in traditional HTS meth-ods is the most common membrane protein drug discovery targetfamily the GPCRs. Because GPCRs are not enzymes and have tradi-tionally in the past been difficult to obtain in soluble form, all cur-rent HTS assays are cell-based. Several different approaches aretypically employed. Changes in intracellular calcium concentrationare measured for Gq coupled receptors, the cAMP assay is used forGi or Gs coupled receptors. More recently, reporter genes havebeen employed, beta-arrestin redistribution has been measured, andreceptor internalization has also been used as a reporter for GPCRligand binding and activity (89). The most sensitive and widelyemployed assay is the cAMP assay, but it is restricted to Gs and Gicoupled receptors. The calcium-based assay employed for Gq cou-pled receptors has the problem of not distinguishing constitutiveactivity from basal levels of intracellular calcium concentration, it isbeing difficult to quantitate pharmacological effects. The reportergene assay requires long incubation with ligands, and there are

Figure 10: Example of a screening of a 19F-labeled compound library (eight compounds are visible in the particular range shown). The 19FNMR spectra were acquired both in the absence (colored blue) and presence of bovine rhodopsin (colored red). The buffer used to acquireboth the spectrum contained 50 mM phosphate buffer (pH 6.0) and 0.5% dodecyl maltoside micelles.

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many problems associated with this, including many false positives,issues with stability, redistribution of ligands and receptors, etc.Arrestin redistribution is a protein interactions-based assay: arrestinbinding to the GPCR is initiated by the phosphorylation of the C-ter-minus of the GPCR. It has been demonstrated in many instancesthat binding of proteins at the cellular side of the receptor, includ-ing arrestin binding to the C-terminus, but also other proteins, e.g.,those involving PDZ domains, alters the ligand-binding propertiesand pharmacology of receptors. Finally, receptor internalization is acomplicated process, and efficient and fast internalization is notalways given. In addition to these assay-specific disadvantages, allof these assays are necessarily indirect and are therefore error-prone. Moreover, compound libraries have limited solubility, andhigh concentrations of DMSO are needed to solubilize them. Thesehigh DMSO concentrations alter the cell surface properties. Finally,while an HTS will almost always yield a hit, especially whenscreening large libraries, the quality of the compounds identifiedmay be low and time-extensive and cost-extensive procedures arerequired to transform the hit to a lead.

NMR-based screening has found increasing application to solubleproteins not only because of the enormous amount of informationthat can be obtained from such a screen (12,90), but most impor-tantly, NMR-based assays are not prone to artifacts brought about bydenaturation, aggregation or precipitation of proteins induced by theligands. There are estimates that 20% of all hits in HTS are based onunspecific ligand effects. Such effects are immediately recognized inNMR-based screens because of the direct measurement of proteinsignals. Furthermore, solubility of the compounds is directly visiblefrom the NMR samples. Another advantage is the fact that weakligands can be identified easily. A weak but selective ligand canbecome the starting point for successful screening, such as isexploited in the fragment-based screening approach. Thus, eventhough an NMR-based screen may seem more expensive because ofthe large protein requirements, in the long run, successful compoundsmay be found easier and cheaper when viewed from the end-productperspective. Typically, HTS is evaluated based on the number of com-pounds screened versus number of hits, but one really has to criticallyevaluate how many of the hits have actually led to a lead or drug. Infact, there are many cases where HTS in pharmaceutical industry hasnot yielded drugs against a desirable target.

Synthesis of 19F containing smallmolecule compounds

The access to diverse and drug-like screening libraries labeled with19F is the prerequisite for 19F NMR-based screening technology. Arecent database search revealed that more than million fluori-nated small molecular weight compounds are commercially avail-ablec. However, many of those compounds do not satisfy drug-likecriteria and are rather unlikely to yield expandable hits duringscreening. A notable exception is the trifluoromethyl group contain-ing compound nitisinone. This compound was originally developedand is still used as an herbicide. It was recently found to be usefulto treat the hereditary orphan disease tyrosinemia type 1 (HT-1)(91). Since its first use for this indication in 1991, it has replacedliver transplantation as the first-line treatment for this rare and

mostly deadly condition. This is an interesting example because thecompound is by no means 'drug-like', containing three strong elec-trophiles in addition to a nitro group. Nevertheless the compound iswell tolerated, no severe side effects are reported and the drugcomprises a major therapeutic advancement by increasing the for-mer 4-year survival rate of 29% of newborns with HT-1 to 88%.Because nitisinone is probably an exception rather than the ruleand many of the fluorinated compounds that are commercially avail-able will not have the desired properties to make a drug or evenscreen for biologically relevant compounds, the development of newlibraries containing 19F is highly desirable. Introduction of fluorinein organic compounds is an established area of organic chemistryand can be accomplished by a plethora of techniques (92). Manyuseful reactions exist to selectively introduce fluorine in organiccompounds (Figure 11). To this end specific reagents have beenintroduced, e.g., the recently described Togni's reagent for the elec-trophilic introduction of trifluormethyl groups (93) or Buchwald'snucleophilic aromatic substitution of triflates (94) (Table 1).

Because of the exceptional physico-chemical nature of fluorine,however, organic chemistry of fluorine often takes different reactionpathways (Table 1). Thus, fluorine introduction is commonly used inmedicinal chemistry to alter the drug compound's profile, includingits solubility, metabolism, pKa and logD (lipophilicity). In addition, itis well known that there are distinct stereochemical effects in fluo-rine compounds as opposed to their non-fluorine counterparts, e.g.,the trifluormethyl group in phenols has an energetic preference foran out-of-plane geometry as opposed to the methyl group (Table 1,

Figure 11: Overview of some current fluorine chemistries. A lar-gely underdeveloped way to access fluorine-containing organic com-pounds is by using multicomponent reaction chemistry (MCR) andemploying fluorine building blocks (95). Many fluorinated buildingblocks are commercially available in large diversity, e.g., aldehydes,carboxylic acids, amines, alcohols, cyanates, phenols and heterocy-cles. Based on the scaffold diversity amenable by MCR chemistryone can easily imagine the accessible fluorine chemical space(Figure 12).

NMR-Based Screening


entry 9) (96). Fluorine introduction into organic molecules is verypopular to protect metabolically labile positions, e.g., in benzenerings. Finally, positron emission tomography has to be mentionedas a special application of fluorine in drug discovery because ofits excellent properties to follow the fate of drugs in the humanbody in a time and site resolved manner. Positron emissiontomography has found its clinical application as a modern diag-nostic form in several indication areas and will gain increasinglymore importance with the rise of molecular markers in clinicaltrials (97).

Summary and outlook

The prospects of NMR-based screening of small molecule ligandbinding to membrane proteins are very good: 1H and 19F NMR-spec-troscopic approaches have been developed to overcome many ofthe challenges associated with solution NMR studies of membraneproteins in detergent micelles. Solvent suppression schemes andSTD spectroscopy are powerful 1H-NMR-based approaches to studyligand binding to membrane proteins that are not accessible to

Table 1: Compilation of some extreme physico-chemical proper-ties of fluorine, and fluorine moieties that make them so attractivein medicinal chemistry.

1 C–F strongest bond in organic chemistry,485 kJ ⁄ mol cf. CH 416 kJ ⁄ mol

2 C–F (1.41 �); C–O (1.43 �); C–H (1.09 �)4 van-der-Waals volume of the trifluoromethyl (CF3) group is

similar (42.6 �2) to that of the ethyl group (CH3CH2;axially anisotropic), but the shapes of the two groupsare very different

5 CF is bio-isosteric to COH, COMe6 Single H ⁄ F exchange raises the logD value by approximately 0.257 18F labeled compounds for PET investigations (t 18F �110 min)8 19F (spin 1 ⁄ 2) 0.83 of 1H9

Stereochemical effect of trifluoromethylphenolesand methylphenoles

Figure 12: 19F-tagged variations of the Groebcke reaction. All derivatives amenable from the differentially 19F-substituted starting materi-als isocyanide, benzaldehyde and amino pyridine can be synthesized ('fluorine dance').

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structure determination of NMR spectroscopy. At the same time,the prospects for structure determination of membrane proteins byNMR spectroscopy are steadily increasing. Complementary to 1H-NMR spectroscopy, the future of 19F-NMR-based drug discovery isparticularly bright. Fluorine is an extremely versatile element withmany advantages in the drug discovery pipeline. At the chemicalsynthesis stage, 19F derivatives are easily obtained. The fluorinesubstitution modulates ligand properties that can lead to betterdrugs. At the screening stage, the 19F nucleus provides a sensitiveligand binding as well as conformational probe without backgroundsignals. At the more biologic level, the cellular uptake and fate of19F-tagged compounds can be detected in a time-resolved andspace-resolved as well as otherwise in a label-free manner. Thiscan be extended to in vivo 19F imaging. In this review, we focusedmostly on the first three stages and the particular challenges andopportunities for 19F NMR in the context of screening membraneproteins. In the future, such approaches are likely to gain furtherpopularity as the instrumentation capabilities further improve.Nuclear magnetic resonance instrumentation companies are devel-oping increasingly more sensitive and versatile 19F NMR cryoprobes.Screening of 19F libraries will become fast and cost efficient, andthe discovery of novel small molecular weight ligands by NMR willbecome possible even for the difficult membrane bound targetsGPCRs and ion channels. The broad and general availability of 19F-labeled compound libraries is currently an issue and has to besolved in the future. An inexpensive and efficient method usingmulticomponent reaction chemistry involving labeled building blocksis proposed.

Acknowledgments

Kelly Hay is grateful to the Fluorine division of the American Chemi-cal Society for a Moissant Summer Internship. This work was inpart supported by the National Science Foundation CAREER grantCC044917, National Institutes of Health Grants NLM108730 and1R21GM087617-01 and the Pennsylvania Department of Health.

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Notes

ahttp://www.drorlist.com/nmr/MPNMR.html.bhttp://www.pdb.org/pdb/home/home.do.chttp://www.emolecules.com.dhttp://www.mdli.com/products/acdsc.html.

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