Variable-Temperature NMR Spectroscopy, Conformational Analysis

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Variable-Temperature NMR Spectroscopy, Conformational Analysis, and Thermodynamic Parameters of cADPR Agonists and Antagonists Sarah-Marie Saatori, Tanner J. Perez, and Steven McGrath Graham J. Org. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.joc.7b02749 • Publication Date (Web): 24 Jan 2018 Downloaded from http://pubs.acs.org on January 25, 2018

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The Journal of Organic Chemistry

Variable-Temperature NMR Spectroscopy, Conformational Analysis, and Thermodynamic Parameters of cADPR Agonists and Antagonists

Sarah-Marie Saatori, Tanner J. Perez, and Steven M. Graham*

Department of Chemistry, St. John's University, 8000 Utopia Parkway, Queens, NY 11439, USA.

ABSTRACT: cyclic Adenosine 5'-diphosphate ribose (cADPR) is a ubiquitous Ca2+-releasing second messenger.

Knowledge of its

conformational landscape is an essential tool in unraveling the structureactivity relationship (SAR) in cADPR. Variable-temperature 1H-NMR spectroscopy, in conjunction with PSEUROT and population analyses allowed us to determine the conformations and thermodynamic parameters of the furanose rings, γ–bonds (C4'–C5'), and β–bonds (C5'–O5') in the cADPR analogs 2'-deoxy-cADPR, 7-deaza-cADPR, and 8-bromo-cADPR. A significant finding was that while the analogs are similar to each other and to cADPR itself in terms of overall conformation and population (∆G°), there were subtle, yet important differences in some of thermodynamic properties (∆H°, ∆S°) associated with each of the conformational equilibria. These differences prompted us to propose a model for cADPR in which the interactions between the A2'–N3, A5"–N3, and H2–R5' atoms serve to fine tune the N-glycosidic torsion angles (χ).

keywords: agonists; cADPR analogs; calcium release; nucleotides; signal messengers; synthesis.

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INTRODUCTION Beyond its role in redox metabolism, nicotinamide adenine dinucleotide (NAD+) has emerged of late as a regulator of numerous other biological pathways.1,2 NAD+ acts as an acyl acceptor in the decacylation of histone proteins in reactions catalyzed by silent information regulator-2 (Sir2) proteins (sirtuins), as the donor in the mono- and poly-adenosine diphosphoribosylation of proteins as catalyzed by monoADP-ribosyl transferases and poly-ADP-ribosyl polymerases (PARPs), as a 5'-cap to some prokaryotic RNAs,3 and, relevant here, as a substrate for cyclic adenosine 5'-diphosphate ribose (cADPR, Figure 1) synthases. At the phenotypic level, a recent mouse study suggested enhancing NAD+ levels can prolong lifespan.4 NAD+ levels can be depleted by the multifunctional transmembrane glycoprotein CD38 (and CD157), which is an NAD+ hydrolase, cADPR synthase, and a cADPR transporter.5,6 Whereas CD38 deficient mice have greatly enhanced NAD+ levels compared to wild type, overexpression of CD38 in human kidney cells led to the down-regulation of over 100 proteins involved in an array of cellular processes.7 Clearly NAD+ is a major player in the regulation of cellular metabolism and homeostasis, but we wish to focus on one aspect in particular: the role of its Ca2+-releasing metabolite cADPR. First discovered in 1987,8 cADPR quickly emerged as Ca2+-releasing second messenger signaling molecule.9 Changes in intracellular Ca2+ levels regulate a host of cellular processes that have physiological significance, notably heart muscle contraction, fertilization, and insulin secretion.10,11 The end result of the formation of cADPR is Ca2+-efflux from the ryanodine receptor (RyR), a large (~2 mDa total mass) tetrameric protein located in the sarco(endo)plasmic reticulum membrane. It is generally believed that cAPDR does not interact with the RyR directly, but rather through the agency of some other protein. RyR activity is modulated by multiple protein ligands, including kinases, phosphatases, calmodulin, and the FK-506 binding protein FKBP12.6 (calstabin).12,13 Which of these ligands – if any – is the cognate protein binding partner for cADPR remains to be established, although a recent photoaffinity labelling study utilizing a cADPR analogue identified glyceraldehyde 3phosphate dehydrogenase (GAPDH) as a low affinity (18 µM) cADPR binding protein.14 The SAR in cADPR. The structure-activity relationship (SAR) in cADPR continues to evolve. Over 100 cADPR analogs are known, and while the SAR is far from settled, certain patterns have emerged. The parent cAPDR (4a, Figure 1 and Figure 2) has an EC50 for Ca2+ release of 30–90 nM15,16 in the sea urchin homogenate (SUH) assay. Three early analogs – 2'-deoxy-cADPR (2'-dA cADPR, 4b)16,


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Figure 1. Synthesis of cADPR from NAD+ and breakdown of cADPR to ADPR. ADP ribosyl cylase (ADPRC) from the mollusk Aplysia californica is a soluble, commercially available enzyme. CD38 and CD 157 are transmembrane glycoproteins. cADPR can be hydrolyzed to ADPR in the continued presence of these enzymes; it can also undergoes spontaneous hydrolysis, with a half-life of 2.5 days at pH 7 at 37 oC

7-deaza-cADPR (4c)17, and 8-bromo-cADPR (8-Br-cADPR, 4d)15 (Figure 2) – are, respectively, a potent agonist (EC50 58 nM), a partial agonist (90 nM, with only 66% of the total Ca2+ released), and an antagonist (IC50 ~1000 nM). We wished to perform detailed conformational analyses on cADPR analogs, and we chose these three analogs for this variable temperature NMR study because (1) they span the range of pharmacological activity, (2) they are easily synthesized using the 'chemoenzymatic' approach (chemical synthesis of the appropriate NAD+ analog and enzymatic cyclization with the commercially available Aplysia californica ADP ribosyl cyclase, Figure 1 and Scheme 1), and (3) they are all based on a deoxyribofuranose or ribofuranoses scaffold and as such do not require substantial modification to the default parameters of PSEUROT18 (see Results and Discussion), the program used to convert 3-bond vicinal 1H-1H coupling constants to (deoxy)ribofuranose conformations. Compounds 4b, 4c, and 4d, along with 3-deaza-cADPR (9a, EC50 ~1 nM),19 and cyclic aristeromycin diphosphate ribose (cArisDPR, 10, EC50 80 nM)20 typify one class of cADPR analogs, the 'A-ring' modified analogs (see the legend to Figure 2).21 A common feature of deaza analogs 4c and 9a, carbocyclic 10, and cATPR (8) is an improved resistance to hydrolysis, which has implications for NMR studies (vide infra). A-ring and Adenine modified Analogs. Generally, substitution at the 8-position results in an antagonist. 8-amino-cADPR, like 8-Br-cADPR 4d, is an antagonist,15 in both the SUH assay and in Jurkat T cells (JTC).22 The 7-deaza alteration alters activity inconsistently; whereas 7-deaza-cADPR 4c is less effective at Ca2+ release than cADPR, 8-bromo-7-deaza-cADPR is a more effective antagonist than 8-bromo-cADPR 4d.23 Removal of the A-ring 2'-OH from the antagonist 8-NH2-cADPR 3 ACS Paragon Plus Environment


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Figure 2. Structures of cADPR analogs. Based on its name – cADPR, for 'cyclic adenosine diphosphate ribose', we prefer to call the ribofuranose ring derived from adenosine (the 'A' in cADPR) the 'A-ring' and the other ribofuranose ring (the 'R' in cADPR') the 'R-ring'. Others have called these rings the 'southern' and 'northern' ribose rings, respectively, but we fear this can lead to confusion in the subsequent discussion where the terms 'north' and 'south' refer to conformational descriptors of furanose ring geometries

led to a 30-fold drop in potency, whereas the same deletion in the antagonist 8-Br cADPR 4d had no effect on activity.24 In terms of modifications to the A-ring furanose, the Potter group found the A-ring 2'-OH could be deleted (2'-dA cADPR, 4b) with essentially no loss in activity, but deletion of the A-ring 3'-OH resulted in a ~100-fold drop in agonistic activity and replacing the 3'-OH with a methoxy group resulted in a weak antagonist.16 Additionally, analog potencies cannot be compared across different assays and organisms – agonists in for example a SUH assay may not show the same behavior in a JTC assay.22 cIDPR Analogs. A second class of cADPR analogs (Figure 2), the N1-cyclic inosine diphosphate ribose (N1-cIDPR, 5) analogs, has been studied extensively by the Potter group.25,26 The requisite dinucleotide precursor, nicotinamide hypoxanthine dinucleotide (NHD+), is a substrate for ADPRC but cyclizes at N7 rather than N1. Recognizing the cyclase's preference for a syn-oriented base, and knowing that bulky C8-substituents favor a syn orientation, they rationalized that 8-bromo-NHD+ might adopt a conformation more suitable for ADPRC-catalyzed cyclization at N1. The initially formed 8-bromo-N1-cIDPR was subsequently converted to N1-cIDPR (5), as well as to 8-phenyl-, 8-azido-, and 8-amino-cIDPR analogs. The cIDPR analogs show greatly enhanced hydrolysis resistance, are membrane permeable, and 5 behaved almost identically to cADPR in a JTC assay. cIDPR 5 is also an inhibitor (IC50 = 276 µM) of the cADPR hydrolase activity of soluble human CD38.27


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R-ring and Adenine modified Analogs. The analogs discussed above were all prepared via the chemoenzymatic approach: chemical synthesis of a nicotinamide dinucleotide precursor and subsequent cyclization using ADPRC. While some R-ring modified cADPR analogs have been prepared using this strategy28 (requiring the preparation of a nicotinamide mononucleotide (NMN) analog), generally total chemical synthesis is required. In the R-ring modified series (Figure 2), cyclic adenosine diphosphate carbocyclic ribose (cADPcR, 6, EC50 15 nM versus 50 nM for cADPR),29,30 and cyclic adenosine diphosphate thioribose (cADPtR, 7, EC50 39 nM versus 210 nM for cADPR),31 prepared by the Shuto group, are illustrative. The carbocyclic 6 series again showed hydrolysis resistance; both series showed interestingly that the 8-amino analogs were agonists. Pyrophosphate Modified Analogs. Comparatively speaking, pyrophosphate-modified cADPR analogs are rare (Figure 2). Phosphorothioate 13 (see Ring Deleted Analogs below and Figure 3) is one example; cyclic adenosine triphosphate ribose (cATPR, 8, 20 times more potent than cADPR),32 and two analogs33 where the pyrophosphate is replaced by methylenebisphosphonate, are others. The latter analogs, cADPR[CH2] (9b, EC50 111 nM versus 59 nM for cADPR) and 3-deaza-cADPR[CH2] (9c, EC50 302 nM versus ~1 nM for 3-deaza-cADPR 9a), as well as cATPR (8), were prepared via the chemoenzymatic approach. Ring Deleted Analogs. Some cADPR analogs (Figure 3) have been prepared that entirely lack and A-ring, an R-ring, or both. Members of this class of analogs are the cyclic inosine diphosphate ribose ethers (cIDPRE, 11, and their 8-substituted congeners 12),34 phosphorothioate 13 (one of the diastereomers is an agonist and the other an antagonist),35 cyclic triazole diphosphate ribose ether (cTDPRE, 14)36, and cyclic inosine diphosphate diethers (cIDPDE, 15). These 'R-ring deleted' analogs,

Figure 3. Structures of ring deleted cADPR analogs.



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prepared by the Zhang group, often retain agonistic activity. Especially remarkable is that the 'no ring' analog 15, is only 3-fold less potent an agonist than cADPR. The 'no ring' 15 notwithstanding, deletion of the A-ring is deleterious to activity. A-ring deleted N9-butyl-cIDPRE (16), like cIDPR 5 is an inhibitor of the cADPR hydrolase activity of soluble human CD3827. The cAcyloDPR analog 17, prepared by the Potter group, was found to be inactive in the SUH system. Rationale. Despite this wealth of pharmacological data, a key question in the SAR of the cADPR system remains mostly unanswered: in making cADPR analogs, do the alterations to the cADPR scaffold change the structure – specifically, its conformation – or is the structure largely preserved and consequently what changes is its interaction with the receptor? If a new conformation is adopted, has an analog become better or worse suited to the receptor binding site? Or is cADPR merely a scaffold upon which groups can be deleted from or appended to, allowing for better (or worse) contacts in the as yet unknown receptor binding site. In an extensive study by Moreau,22 a strong correlation was found between activity and the conformation of the 'southern' ribose ring (the 'A-ring' in our terminology),21 at least in the sea urchin system. Kudoh37 found cADPR (1a), cADPcR (6), and a 6-analog to be similar in A-ring structure in an NOE-restrained MD simulation. Less is known about the conformation of the R-ring, and other than our earlier38,39 and recent40 work on cADPR and its A-ring 2'-OMe and 3'-OMe analogs, even less is known regarding the conformation about the pyrophosphate backbone and the glycosidic bonds. Detailed structural information of the conformational landscape is lacking for the entire cADPR family, and missing completely is any thermodynamic data (∆Go, ∆Ho, ∆So) on the conformational equilibria – data could begin to explain why certain conformations are favored. Do conformational changes in one part of cADPR drive changes in another, that is, are the molecular motions coupled? This work, a variable temperature NMR spectroscopy investigation, aims to fill part of that gap by conducting a detailed conformational analysis and thermodynamic study on the known cADPR analogs 2'-dA-cADPR (4b), 7-deaza-cADPR (4c), and 8-Br-cADPR (4d).

RESULTS AND DISCUSSION Synthesis. The known cADPR analogs 4b, 4c, and 4d were synthesized using a chemoenzymatic approach consisting of 5'-phosphorylation of an adenosine analog (1) to give an AMP analog (2), formation of a pyrophosphate bond by activation of the 5'-phosphate of 2 to nucleophilic attack by NMN, thus providing the requisite NAD+ analog (3), and cyclization using the commercially available Aplysia ADPRC (Scheme 1) to yield a cADPR analog (4). Starting from 7-deazaadenosine (1c) or 6 ACS Paragon Plus Environment

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8-bromoadenosine (1d), 5'-selective phosphorylation was performed by the Yoshikawa method (TEP, 2 equiv. POCl3, 0 °C; water was omitted).41 Upon completion of the reaction (2–4 hours) the reaction mixture was slowly dripped via cannula into ice-cold, stirred, dry ether. After a brief centrifugation, the solid pellet was stirred in ice-cold water. Ion-exchange chromatography (formic acid gradient) afforded the AMP analogs as their free acids. In our hands the reaction was almost completely selective for 5'-phosphorylation and only trace amounts of inorganic phosphate could be detected by 31P NMR. The synthesis and purification of 7-deaza-5'-AMP (2c) proceeded uneventfully, but the phosphorylation of 8–bromoadenosine (1d) to the corresponding 5'-phosphate 2d proved problematic, as described below. Monitoring the conversion of 1d to 2d by reversed-phase (RP) HPLC analysis indicated the formation of two products with significantly different retention times (4.1 and 4.8 min with baseline separation) by RP HPLC but rather similar UV spectra (λmax 262 and 264 nm, respectively). Partial separation was achieved by ion-exchange chromatography (Sepharose Q resin eluted with formic acid). The early eluting fractions (early-2d) were sufficiently pure to be pooled, as were the later fractions (later-2d), as judged by RP HPLC analysis of the individual fractions. The 1H NMR spectra of early-2d and later-2d were remarkably similar, with H2 signals at 8.36 ppm and 8.34ppm, H1' signals at 6.14 (d, J = 5.6 Hz) and 6.13 ppm (d, J = 5.6 Hz), and H2' signals at 5.21 ppm (t, J = 5.5 Hz) and 5.25 ppm (t, J = 5.5 Hz). Ultimately we determined the acidic conditions of the Yoshikawa phosphorylation promoted an exchange of the 8-bromo substituent for chlorine (a phenomenon we were unaware of at the time) which had been previously noted in the literature.42 Comparison of the 13C NMR chemical shifts for C8,43 and, ultimately LCMS data (early-2d, 140 ppm, m/z MH+ 382; later-2d, 129 ppm, MH+ m/z 426), showed that early-2d was 8-Cl-AMP 2e and later-2d was the desired 8-Br-AMP 2d.44 With the desired monophosphates in hand, we then focused on the coupling step – joining the adenosine 5'-monophosphates 2b-2d to NMN via pyrophosphate bond formation to form NAD+ analogs 3b-3d. As in our previous synthesis of cADPR analogs, we chose,39 as had others,16 to use the Michelson45 procedure in which diphenyl phosphoryl chloride (DPPC, (PhO)2POCl) is used to activate the 5'-phosphate of a nucleotide towards nucleophilic attack via the formation of a P1-nucleoside-5'-



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Scheme 1. Synthesis of cADPR analogsa

a

Reagents and conditions: (a) (i) PO(OEt)3, POCl3, 0 °C, 2–4 h, (ii) H2O, 0 °C (iii) tri-n-octylamine, CH3OH; (b) (PhO)2POCl (diphenyl phosphoryl chloride, DPPC), tri-n-butylamine (TBA), dioxane, DMF; (c) (i) Ac2NMN, DMF, TBA, pyridine, (ii) NH3, CH3OH, 0 °C, 15 min; (d) ADPRC, HEPES buffer, pH 7.0, 2–3 h. adenosine 5'-monophosphate (AMP, 2a), 2'-deoxyadenosine 5'-monophosphate (dAMP, 2b), and NAD+ (3a), each as their free acids, were obtained from commercial sources. Ac2NMN was prepared as described.39

P2-diphenyl pyrophosphate (mixed anhydride, Scheme 1). Nucleophilic attack by the 5'-phosphate group of a second nucleotide displaces the weak base leaving group diphenyl phosphate from the mixed anhydride. In principle, either nucleotide could be the activated nucleotide and the other the attacking nucleophile; in practice we chose to activate the AMP analogs 2b-2d and use 2',3'-di-O-acetyl NMN (Ac2NMN; the acetyl groups enhance the solubility of NMN in organic solvents) as the attacking nucleophile, so as to minimize the exposure time of the somewhat fragile NMN glycosidic bond to the reaction conditions. The free acids of AMP analogs 2b-2d were thusly converted to their tri-noctylammonium salts to improve their solubility and reacted with DPPC. The mixed anhydride was precipitated into ether, the supernatant decanted, dried briefly under vacuum, and a solution of Ac2NMN then added. Ion-exchange chromatography as for 5'-AMP analogs 2c and 2d afforded NAD+ analogs 3b-3d in modest yield. Interestingly, the least challenging aspect of the synthesis was the enzymatic cyclization of NAD+ analogs 3b-3d to cADPR analogs 4b-4d. Cyclization reactions were typically run on a 50 µmol (30-40 mg) scale in ~ 6 mM HEPES buffer at room temperature with a substrate concentration of ~0.85 mM.46 All three NAD+ analogs were readily converted to the corresponding cADPR analogs over the course of several hours, in conversions exceeding 60% as monitored by RP HPLC. Ion-exchange chromatography on the Sepharose-Q column utilizing either TEAB or TEAA buffers gave the cADPR analogs 4b-4d, as their triethylammonium salts, in good yield. 8 ACS Paragon Plus Environment

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NMR Spectroscopy. 1H NMR (400 MHz) spectra for the triethylammonium salts of each of the cADPR analogs 4b-4d were recorded at 277, 298, 318, 338, and 353 K, though the lability of the R ring C1'-N1 bond in 4b and 4d limited data collection at the higher temperatures. The 1H NMR spectra at 298 K of cADPR analogs 4b-4d, as well as the 600 MHz spectrum (298 K) of cADPR (4a) for reference, are shown in Figure 4. (See Pages S23, S27, S28, and S34 for the spectra of each analog at each of the temperatures.) Samples were 10–20 mM in D2O containing TMSP as an internal standard. The spectra were all first-order (confirmed by spectral simulation, not shown), allowing the three-bond vicinal coupling constants (3J) needed for the conformational analysis (see Population Analysis) using PSEUROT to be extracted directly from the spectra. The information needed to perform conformational analysis on cADPR/analogs resides in eight signals (ten for 2'-dA analog 4b) : the A-ring H1' (henceforth A1') and R1' (d, J1'2), A3' and R3' (dd, J2'3', and J3'4'), A5' and R5' (dt, J5'5", J4'5', J5'P), A5" and R5" (ddd and dt, J5'5", J4'5", J5"P), plus A2" (J1'2" and J2"3') for deoxy-4b. The 1'2', 2'3', and 3'4' (and 1'2" and 2"3' for 4b) couplings are used to determine the furanose conformations via PSEUROT, and the 5'/5" couplings, via 'sum rules', to determine the γ– and β–bond conformations (see Population Analysis). The H4'-signals (broad multiplets due to coupling with H3', H5', H5", and sometimes phosphorus) and H2' signals (apparent triplet due to the near equivalence of J1'2 J2'3') are less than ideal. A combination of routine 1D 1H- and phosphorus-decoupled H1 (1H{31P}) NMR were used to extract the coupling constants, as generally the needed signals were well enough resolved. As we had done previously,38-40 additional information was obtained through the use of the 1D TOCSY47 experiment, useful especially at the higher temperatures as cADPR/analogs began to decompose. Two notable features of the 298 K spectra were apparent upon inspections. First, as has been noted,22 was the unusually high chemical shift for the A2' signal in the spectra of 4a, 4c, and 4d (5.4–5.5 ppm versus 4.6–4.8 for adenosine and AMP), characteristic of a syn-oriented base.48 The second feature was the consistency in the chemical shifts and coupling patterns across the analog series 4a-4d. The R1' and all 5'/5" signals, and to a slightly lesser extent the R2', R3', and R4' signals, were quite similar (See Tables 1 and 2 for coupling constants), suggesting comparable conformations and environments. (The differences in the A-ring chemical shifts can be attributed to the analog modifications, for example



D

A3 A1 R1

A2

R3

R2 R4

C

R4 R1

A1

A2

A3

B

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A4

A5”/R5’

R2

R5” A5’

8-Br cADPR (4d 4d) 4d

R5” A5’

7-deaza cADPR (4c 4c) 4c

R5' A5" R3

A4

R4

A1

R1

A3

R2

A2'

A5"/R3 R5' A4

R5” A5’

A2"

* ** 2'-dA cADPR (4b 4b) 4b

HOD

~

A

R5'/A4

A1 R1

A2

R2R4/A3

A5”/R3'

R5” A5’

cADPR (4 4a)

Figure 4. The 400 MHz 1D 1H NMR spectra of the cADPR analogs at 298 K. Panel A: cADPR (4a) reference spectrum (600 MHz).40 Panel B: 2'-dA-cADPR (4b). The peaks in the A2' signal marked with stars are from the 13 C satellite peak of the triethylammonium signal. Panel C: 7-deaza-cADPR (4c). Panel D: 8-Br-cADPR (4d). Primes in the 1', 2', 3', and 4' signals omitted for clarity. Note the generally overlapped A3'/R2'/R4' region in each analog, the overlapped A5"/R3' signals of 2'-dA cADPR (4b), and the overlapped A5"/R5'/A4' of 8-Br cADPR (4d). The 8-bromo sample was contaminated with a small amount of its hydrolysis product, 8-Br AppRib. Samples were 10–20 mM in D2O containing TMSP as an internal standard.

2'-deoxyribo versus ribo.) The R5' and R5" pair was consistently a doublet of triplets (dt), indicating JR4'R5', JR4'5", JR5'P, and J5"P, are similar in magnitude (~2 – 3 Hz). The A5' (dt) signal was similar in appearance to R5' and R5", but A5" is a ddd, indicating at a minimum a conformation about the A ring C4'-C5' bond different than that of the R ring C4'-C5' bond.49 These patterns remained largely intact throughout the variable temperature study, although the magnitude of the coupling constants changed. Accurate values for the J4'5' and J4'5" couplings were generally extracted from a 1H{31P} experiment (See pages S22, S23, S26, S31, S33); the J5'P and J5"P couplings were extracted by subtracting the appropriate J4'5' or J4'5" and J5'5" coupling from their corresponding signals.50 10 ACS Paragon Plus Environment

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As mentioned earlier, cADPR analogs 5–8, 9a, and, significantly here 7-deaza analog 4c, show increased resistance to hydrolysis of the R1'-N1 glycosidic bond (compare to cADPR; t1/2 = 2.5 d, pH 7, 37 °C). Pages S23, S27, S28, and S34 in the SI show the 1H NMR spectra of 4b, 4c, and 4d at the highest temperature from which coupling data could be extracted. The 2'-dA analog 4b and 8-Br analog 4d have completely broken down to their respective ADP-riboses (confirmed by HPLC; data not shown), whereas 7-deaza 4c is largely intact. Conformational Analysis Using PSEUROT. First proposed by Kilpatrick et al. in 1947,51 the pseudorotation concept is the basis for the conformational analysis of five-membered rings such as the furanose ring found in nucleosides and nucleotides. Five-membered rings avoid planarity by displacing one atom (envelope conformation, 2E, 3E, 2E, 3E) or two atoms (twist conformation, , ) from the ring plane defined by the remaining atoms, with atoms below ('exo') the ring plane indicated by subscripts and those above ('endo') by superscripts. (Figure 5a). Nucleos(t)ide furanose rings, lacking symmetry, allow for twenty possible 'ideal' or 'canonical' conformations – ten possible envelope forms

Figure 5. (a) Five-membered ring furanose geometries and their descriptors. In S-type conformers, note how H1' and H2' are anti while H3' and H4' are perpendicular, whereas in N-type conformers the opposite is true. (b) Endocyclic bond definitions. (c) The three canonical orientations of the γ– and β–bonds. (d) Atom, bond, and furanose definitions for cADPR. See ref. 53 for details.



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and ten possible twist forms – which alternate between E and T forms as a furanose traverses a full pseudorotational itinerary. The Altona-Sundaralingam (AS)52 formalism assigns a numerical value, the 'phase angle of pseudorotation' ('phase angle', P) to each of the twenty conformers. A second parameter – the maximum puckering amplitude, ψm, – describes by how much the ring deviates from planarity. (A planar ring would have ψm = 0o and idealized cyclohexane 60°; typical ψm's in nucleos(t)ides are 25o – 45o.) The allowed values of P are 0o – 360o; by analogy to a compass rose P = 0o (2T3; C2'-exo-C3'endo) is 'north' and P = 180o (2T3; C2'-endo-C3'-exo;) is 'south'. These relationships are shown in Figure 5a. It should be stressed that there is no high-energy barrier between adjacent 'conformers' on the pseudorotational wheel and that the notion of twenty 'stops' in pseudorotational space is merely a descriptive convenience. A nucleos(t)ide can adopt any value of P. The geometry53 of a furanose ring can also be based on the values of the endocyclic torsion angles ν0–ν4 (Figure 5b); indeed, this is the basis of AS formalism. The relationships are: tan =

° °

=

ν = cos # +

(1) (2)

%& '

( where . = 0, 1, 2, 3, 4

(3)

The elegance of the AS formalism resides in eq 1, as it reduces five endocyclic torsion angles to a single parameter, P, that simply describes which atom(s) are puckered out of the ring plane. Any individual endocyclic torsion angles can be re-created with eq 3. For reasons both steric and stereoelectronic54 not all of the 20 canonical forms are populated. The furanose ring of a nucleos(t)ides tend to localize to either a 'north-type' (N) or a 'south-type' (S) conformation. Under the generally true case of a two-state N S equilibrium, five parameters are needed to describe the system: the P of each conformer (PN and PS), the maximum puckering amplitudes of each (m N and m S ), and the mole fraction (of either conformer). For cADPR and its analogs, a complete description of the conformational landscape requires knowledge of the conformation of both furanose rings as well as the A-ring and R-ring γ– and β–bond conformations (Figure 5c and 5d). These six conformationally flexible subunits, plus the conformation about the glycosidic bonds χA and χR, (Figure 5d) drive the conformation. The aforementioned study22 of cADPR analogs examined only the A-ring conformation, and only to the extent of determining if the A-ring furanose was N-type or S-type. 12 ACS Paragon Plus Environment

Page 13 of 38 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


The equilibrium ratios and precise values for PN, PS, m N , m S and the γ– and β–bond conformations of the cADPR analogs 4b-d can be obtained through a more rigorous analysis of coupling constants, and to our knowledge this is the first such study to do so in significant detail. The furanose ring coupling constants for cADPR analogs 4b-d are summarized in Tables 1 and 2. Also included therein are the pseudorotation parameters for 4b-d as determined by PSEUROT, as well as the pseudorotation parameters for cADPR itself (4a), determined previously.40 The coupling constants from Tables 1 and 2 were the input for version 6.3 of the PSEUROT program.18 In addition to coupling constants related to the furanose conformation (A- and R-ring J1'2', J2'3', and J3'4', for analogs 4c and 4d, plus for analog 4b J1'2", and J2"3'), the user provides as input to PSEUROT substituent electronegativities, an initial guess as to the values of the five parameters that describe a two-state NS equilibrium (PN, PS, mN , PS, mS , and the S mole fraction), and a set of parameters to correlate the endocyclic ring atom torsions to the exocyclic 1H-1H torsions (the so-called 'A' and 'B' parameters). While the conformation of a furanose ring is described (Figure 5b and eq 1) by the endocyclic ring torsions, the observed 1H-1H coupling constants (Jobs) depend on the exocyclic 1H-1H torsion angles (Figure 5a). Combined with the initial guess of PN and PS, PSEUROT uses the A and B parameters to create exocyclic torsion angles (e.g., H1'-C1'-C2'-H2') by 'building in' hydrogen atoms onto the corresponding endocyclic torsions (e.g., ν4 = O4'-C1'-C2'-C3') specified by P. With the exocyclic torsion angles thus specified, a Karplus equation within PSEUROT calculates a predicted coupling constant, Jcalc, which it compares to Jobs. One of the reasons we chose these analogs to study first is that they contain only ribo- and deoxyribofuranose rings, for which the A and B values were specifically parametrized.55 (Furanoses different than ribo- and deoxyribo- generally require adjustment of the default A and B values or values specifically derived for the molecule under investigation.) PSEUROT then calculates the expected coupling constants for such a mixture and iteratively adjusts PN, PS, m N , m S , and the S mole fraction so as to produce the best fit between the calculated and observed coupling constants.



Page 14 of 38

Table 1. 400 MHz 1H-1H and 1H-31P Coupling Constants (J, Hz)a–c of the cADPR A-ring for 4b-d and PSEUROT 6.3 output (degrees) for cADPR analogs 4b-d

A-ring

4b 2'-dA

4c 7-deaza

4d 8-Br

PS 6.3h j

4a 4b 4c 4d

T (K) 277 298 318 338 353

J1’2’ 6.85 6.68 6.51 6.19 ndd

277 298 318 338 353

6.38 6.28 6.13 5.97 5.84

277 298 318 338 353

5.41 5.37 5.27 5.15 nd

PN

ψNm 35k 38 35 35

22.5 359.1 352.2 20.3

J1’2" 6.96 7.01 7.07 7.22 nd

J2’2" 14.20 14.19 14.22 14.32 nd

PS

177.1 170.7 159.1 173.7

J2’3’ 6.30 6.31 6.40 6.44 nd

J3’4’ 2.83 2.97 3.29 3.42 nd

J4’5’ 2.84 2.97 3.12 3.37 nd

J4’5” 7.45 7.21 7.00 6.72 nd

J5’5” 10.81 10.89 10.89 10.81 nd

J5’P 3.14 3.37 3.54 4.37 nd

J5” P 3.45 3.51 3.66 4.07 nd

5.06e 5.12 5.18 5.28 5.28

2.65e 2.81f 2.95 3.09 3.20

2.64 2.52 2.56 2.71 2.81

7.23 7.00 6.74 6.69 6.47

10.9 11.0 11.0 11.0 11.0

nd 3.57 3.74 3.78 3.98

nd 3.48 3.46 3.51 3.58

5.13 5.20 5.24 5.28 nd

3.44 3.50 3.53 3.57 nd

2.34 2.29 2.28 nd nd

6.17 6.10 5.92 nd nd

11.14 11.10 11.00 nd nd

2.93 3.31 3.61 nd nd

4.47g 4.34 4.58g nd nd

mS

RMSt

31.5 27.3 30.7 28.8

J2"3’ 3.58 3.78 3.96 4.22 nd

0.099 0.128 0.062 0.045

a

Values ± 0.04 Hz. bOnly J1'2', J2'3', and J3'4' are used in PSEUROT. cFuranose ring values are from the normal 1D H spectrum, J4’5’, J4’5”, and J5’5” values from the 1H{31P} spectrum. J5’P determined by subtraction of Σ(J4’5’ + J5’5”) from Σ(J4’5’ + J5’5” + J5’5P), the latter being the width of 5'-signal. J5"P determined analogously. d nd = not determined due to sample decomposition. eValue from 1D TOCSY with irradiation at A2'. fValue from A4'. gValue determined directly from A5". hErrors in PS6.3 output: cADPR (4a) PN ± 4.3o, PS ± 2.6o, m S ± 0.7o; 2'-dA cADPR (4b): PN ± 4.2o, PS ± 2.3o, m S ± 0.6o; 7-deaza cADPR (4c): PN ± 2.1o, PS ± 1.6o, m S ± 0.4o; 8-Br cADPR (4d): PN ± 2.4o, PS ± 1.6o, m S ± 0.4o. tRoot-mean square error of the observed vs. PSEUROT-calculated J values in Hz. jData for cADPR (4a) from ref. 40. kUnderlined values restrained in PSEUROT 6.3. Note the irregularity in the J5"P values for 4d (boxed). 1


Page 15 of 38 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Table 2. 400 MHz 1H-1H and 1H-31P Coupling Constants (J, Hz)a–c of the cADPR R-ring for 4b-d and PSEUROT 6.3b output (degrees) for 4b-d

R-ring

4b 2'-dA

4c 7-deaza

4d 8-Br

PS 6.3g 4ai

4b 4c 4d

T (K) 277 298 318 338 353

J1’2’ 4.00 4.04 4.06 4.02 nde

J2’3’ 5.07 5.05 5.08 5.13 nd

J3’4’ 2.11 2.17 2.25 2.36 nd

J4’5’ J4’5” 2.10 2.05 2.13 2.43 2.16 2.52 ~2.39d ~2.39d nd nd

J5’5” 11.90 11.96 12.02 nd nd

JP5’ 2.18 2.14 2.39 nd nd

JP5” 3.42 3.70 3.89 nd nd

277 298 318 338 353

4.00 4.04 4.06 4.07 4.06

5.00 5.02 5.03 5.05 5.04

2.45 2.54 2.61 2.67 2.74

nd 2.03 2.12 2.15 2.08

nd 2.49 2.59 2.70 2.74

nd 12.0 12.0 12.1 12.1

nd 2.39 2.53 2.60 2.73

nd 3.42 3.45 3.58 3.73

277 298 318 338 353

3.98 4.03 4.07 4.10 nd

5.06 5.08 5.10 5.09 nd

2.23 2.23 2.28 2.28 nd

2.12 2.05 2.16 nd nd

2.20 2.23 2.25 nd nd

12.0 12.0 12.1 nd nd

2.12 2.59 2.39 nd nd

3.46 3.65 4.00f nd nd

mS 36.6 36.6 34.5 36.2

RMSh 0.031 0.041 0.046 0.030

PN 340.0 337.4 343.2 337.2

mN 40j 40 40 40

PS 215.8 216.2 212.2 215.4

a

Values ± 0.04 Hz. bOnly J1'2', J2'3', and J3'4' are used in PSEUROT. cFuranose ring values are from the normal 1D H spectrum, J4’5’, J4’5”, and J5’5” values from the 1H{31P} spectrum. J5’P determined by subtraction of Σ(J4’5’ + J5’5”) from Σ(J4’5’ + J5’5” + J5’5P), the latter being the width of 5'-signal. J5"P determined analogously. d Value from R4'. end = not determined due to sample decomposition. fValue from 1D TOCSY with irradiation at R2'/R4'. gErrors in PS6.3 output: cADPR (4a) PN ± 0.8o, PS ± 0.3o, m S ± 0.3o; 2'-dA cADPR (4b) PN ± 1.2o, PS ± 0.5o, m S ± 0.5o; 7-deaza cADPR (4c): PN ± 1.1o, PS ± 0.5o, m S ± 0.4o; 8-Br cADPR (4d): PN ± 0.9o, PS ± 0.4o, m S ± 0.3o. hRoot-mean square error of the observed vs. calculated J values in Hz. iData for cADPR (4a) from ref. 40. jUnderlined values restrained in PSEUROT 6.3. Note the irregularity in the J4'5' and J5'P values for 4d (boxed). 1



Page 16 of 38

To determine the values of five parameters requires five observables (coupling constants); ribofuranoses, with only J1'2', J2'3', and J3'4' are thus said to be underdetermined, meaning their N S equilibria cannot be completely described. Generally being interested in the major conformer, one could restrain the P and m of the minor conformer. It has been shown56 that the coupling constants are much less sensitive to changes in m than to changes in P. If puckering amplitude information is available from other sources (e.g. a crystal or QM structure), such data can be used to restrain either or both m N , and m S . Regrettably, reliable crystal structure data is not available for either cADPR or analogs 4b–4d. The only crystal structure of cADPR in the Cambridge Structural Database (CSD Entry YINYID) is of the free acid and was deposited without coordinates. There was enough information in the text of the associated paper57a to estimate a PS ~175o for the A-ring and a PS ~208o for the R-ring, but the free acid raises concerns regarding the γ– and β–bond conformations in the crystal. Another study examined the crystal structure of cADPR bound to CD38, but retrieval of the cADPR ligand from the protein structure (Protein Data Bank ID: 2O3Q) returns an A-ring with the xylo-configuration rather than ribo.57b A third report,57c this time with cADPR bound to the Aplysia cyclase, has a cADPR structure with A-ring 2E (P ~162°)/γ+-bond conformations and R-ring 1E (P ~306°)/γt-bond conformations; the opposite γ-conformations were found here The option we ultimately chose was to collect NMR spectra over a range of temperatures sufficient to obtain significant changes in the coupling constants. Based on our earlier work38–40 and reluctant to use the crystal structure data, we restrained the m N of the A-ring and R-ring minor conformers to 35° (38° for 2b) and 40°, respectively. All other parameters were freely optimized. Assuming the identities of the two conformers involved in the NS equilibrium are temperature independent, a change in the coupling constants reflects a change in their equilibrium ratio (see Thermodynamic Parameters). Each temperature represents not only an additional PSEUROT observable but also the opportunity to perform van't Hoff analyses, from which ∆Ho and ∆So can be determined. Population Analysis. The PSEUROT output is summarized in Tables 1 and 2, and the changes in the north/south ratio as a function of temperature shown in Table 3. Also included in Tables 1 and 2 are couplings of each 5'- and 5"-proton to their respective 4'-protons (related to the γ–bond conformations)


Page 17 of 38 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Table 3. Equilibrium populationsa for the furanose rings, γ–bonds, and β–bonds in cADPR (4a) and analogs 4b-d

4a cADPRb

T (K) 277 298 318 338 353

NS 29:71 32:68 35:65 35:65 36:64 0.92

A-ring γt γ+ 58:42 57:43 56:44 54:46 53:47 0.97

277 298 318 338 353

28:72 30:70 33:67 36:64 nd 0.98

69:31 68:32 67:33 67:33 nd 0.98

8:92 9:91 11:89 17:83 nd 0.88

24:76 24:76 25:75 26:74 nd 0.85

6:94 10:90 11:89 12:88 nd 0.88

3:97 4:96 6:94 nd nd 0.99

277 298 318 338 353

25:75 26:74 28:72 30:70 32:68 0.96

65:35 61:39 59:41 60:40 59:41 0.80

nd 10:90 11:89 11:89 13:87 0.92

28:72 29:71 29:71 30:70 30:70 0.92

nd 10:90 11:89 13:87 13:87 0.88

nd 4:96 5:95 6:94 7:93 0.98

277 298 318 338 353

33:67 34:66 34:66 35:65 nd 0.90

51:49 49:51 47:53 nd nd 0.97

12:88 13:87 16:84 nd nd 0.95

26:74 25:75 26:74 25:75 nd 0.21

8:92 7:93 8:92 nd nd 0.29

3:97 6:94 7:93 nd nd 0.89

R2 c

4b 2'-dA R2

4c 7-deaza R2

4d 8-Br R2

–

t

β β 9:91 11:89 12:88 13:87 nd 0.99

NS 26:74 27:73 27:73 26:74 26:74 0.04

R-ring γt γ+ 4:96 5:95 6:94 7:93 nd 0.98

β+ βt 2:98 4:96 7:93 8:92 nd 0.94

a

N:S populations from PSEUROT 6.3. A-ring γ–bond population: fγ+ = [13.3 – (J4'5' + J4'5")]/9.7 (eq 4).58a R-ring γ–bond: fγ+ = [13.3 – (J4'5' + J4'5")]/9.4 (eq 5).58a A-ring β–bond: fβt = [26.4 – (J5'P + J5"P)]/21.4 (eq 6).58a R-ring β– bond: fβt = [25.5 – (J5'P + J5"P)]/20.5 (eq 7).58b Equations 4 and 6 were used as found in the associated references. Equations 5 and 7 were modified slightly (see the Supporting Information) to improve the fit between the observed and back-calculated coupling constants;58 the back-calculated coupling constants also allowed for identification of the minor conformer. b Data for cADPR from ref. 40. c Value not determined. d Correlation coefficient from the van't Hoff plots (Figure 6).



Page 18 of 38

and adjacent phosphorus atom (related to the β–bond conformations). The γ– and β–bond conformations and populations were elucidated using the 'sum rules'[58 shown in Table 3. In some cases these sum rules were slightly modified (see the Supporting Information) so as to provide a better fit between the back-calculated and observed coupling constants. We were concerned that the macrocyclic nature of the cADPR analogs would restrict their conformational freedom, but gratifyingly the majority of the coupling constants (Tables 1 and 2) showed a regular, albeit small, dependence on temperature. The exception was the R-ring furanose (Table 2), which showed either essentially no variation (J1'2' and J2'3' in 4b/4c/4d, and J3'4' in 4d) or very minor variation (J3'4' in 4b and 4c) in the couplings. All A-ring furanoses in 4b, 4c, and 4d favored south conformations, generally by a factor of 2:1, with PS = 166° ± 7°, and m S = 29° ± 2° (somewhat flatter than their free nucleoside counterparts). As we found for cADPR, the A-ring γ–bonds favored the trans conformation (percent γt at 298 K: 4a, 57%, 4b, 68%, 4c, 61%, but in 4d only 49%) and all β–bonds were heavily biased (83–92%) towards the βt conformer. For the R-ring furanoses, also similar to that of cADPR, the favored conformers were all south-type (N:S ~ 1:3), with PS = 214° ± 2°, m S = 35° ± 1°, a highly populated (~90% or more) βt state, but now an almost exclusively (>87–98%) populated the γ+ state. The favored conformers of the triethylammonium salts59s of 4a–d all show a remarkable degree of similarity in both identity and population, with an S-type furanose, γt bond, and βt bond for the A-ring and an S-type furanose, γ+ bond, and βt bond for the R-ring. Many of the solution conformations determined herein compare quite well to those of the published57a crystal structure of the free acid of cADPR (A-ring PS ~ 175°, β = –138°, R-ring PS ~ 208°, β = +160°), though both γ–bonds in the crystal structure were trans, and to our recent preliminary MD simulation of cADPR.40 Despite these surface similarities in cADPR 4a and cADPR analogs 4b-d, analysis of the thermodynamic parameters reveals some important distinctions relevant to the SAR of the cADPR system (next section).

Thermodynamic Parameters. Data for cADPR 4a is from reference 40. Inspired in part by the work of the Chattopadhyaya group, who have performed extensive conformational analysis and thermodynamic studies on monomeric nucleosides and nucleotides, as well as DNA and RNA,54a,60 we wished to establish if the same methodology could be applied to cADPR and its analogs. Having determined the conformer populations in 4b–4d we next sought to determine the thermodynamics (∆G°, ∆H°, and ∆S°) 18 ACS Paragon Plus Environment

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for each of the conformational equilibria. Qualitatively, the expression ∆G° = ∆H° – T∆S° predicts the conformer whose population increases with increasing temperature is the more disordered conformer. As was found for cADPR (1a) itself,40 it was always the minor conformer whose population increased with temperature, thus the minor conformers (N-type furanoses, β+ or β– bonds, A-ring γ+ bond, R-ring γt or γ–) are more disordered. It thus follows that, as was the case in cADPR,40 the major conformers are favored because of enthalpy. The exception to this trend was the R-ring furanoses in 4a, 4b, 4c, and 4d, whose N:S ratios were either essentially unchanged, or changed marginally, with increasing temperature. The van't Hoff equation (vide infra) predicts a temperature-insensitive population such as the R-ring furanoses in 4a–4d to have a zero or small ∆H° for their NS equilibrium and thus that the R-ring major conformer (here, S-type) is favored by entropy. The population data from Table 3 formed the basis of a more thorough analysis using van't Hoff plots (ln K as a function of 1000/T, Figure 6), revealing the precise values of ∆H°, and ∆S° for each equilibria; this data is shown in Table 4. The majority (15 of 18) of the plots in Figure 6 showed excellent linearity as reflected in correlation coefficients (R2) of approximately 0.90 or higher (Table 3). The moderately good fit (R2 = 0.80) for the A–ring γ–bond in 7–deaza analog 4c can likely be attributed the sum rule used to derive the populations, as the coupling constants changed with temperature in a consistent manner. The R–ring furanose and γ–bond in 8–bromo analog 4d displayed the smallest (and somewhat inconsistent) temperature-dependent variations in coupling constants, resulting in essentially unchanging populations and flat van't Hoff plot of poorer fit (R2 = 0.21 and 0.29, respectively). Thus, other than (1) the exceptional R-ring N S equilibria, which is indeed driven largely (in 4a and 4d) or significantly (in 4b and 4c) by an entropy term favoring the S-conformer and (2) the A-ring γt γ+ transition, where γt is only slightly favored over γ+ and ∆H° and ∆S° are roughly comparable, the remaining conformational equilibria are generally characterized by a negative ∆H° term that outweighs the T∆S° component by a factor of approximately two. This confirms that in the majority of cases the higher populated conformer is favored for reasons of enthalpy. Exceptions are noted in the subsequent discussion.



1.8 1.6

3.8

3.6

3.4

3.1

1.4

2.6

3.0

1.2

2.1

0.8 0.6

ln K

2.6

1.0

ln K

2.2

1.6 1.1 0.6

1.8

0.4

0.1 1.4

-0.4

1000/T (K)

B, β-bonds

1000/T (K)

C, γ-bonds

3.7

3.5

3.3

-0.9 3.1

3.7

3.5

3.3

3.1

2.9

3.7

3.5

3.3

3.1

2.9

2.7

A, furanoses

2.7

1.0

0.0

2.9

0.2

2.7

ln K

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 20 of 38

1000/T (K)

Figure 6. van't Hoff plots (ln χS/χN, χγ+/χγt, and χβt/χβ+/–, as a function of 1000/T) for 2'-dA cADPR (4b, •, ), 7-deaza cADPR (4c, , ), and 8-Br cADPR (4d, , ). Filled symbols represent the A-ring portion of each cADPR and open symbols the R-ring portion. The slope of each line is –∆Ho/R and the y-intercept is ∆So/R. For each conformation the slope, intercept, and correlation coefficient, R2, respectively, are: Panel A: furanose A-rings 4b (••) 0.57, –1.09, 0.98; 4c () 0.44, –0.48, 0.96; 4d () 0.12, 0.26, 0.90. Furanose R-rings: 4b () 0.17, 0.56, 0.85; 4c () –0.13, 0.49, 0.92; 4d () –0.05, 1.23, 0.21. Panel B: A-ring β-bond: 4b (••) 1.26, –1.99, 0.88; 4c () 0.44, 0.72, 0.92; 4d () 0.70, –0.48, 0.95. R-ring β-bond: 4b () 1.72, –2.68, 0.99; 4c () 1.17, –0.72, 0.98; 4d () 1.99, –3.75, 0.89. Panel C: A-ring γ-bond: 4b (••) –0.13, –0.32, 0.98; 4c () –0.30, 0.54, 0.80; 4d () –0.27, 0.95, 0.97. R-ring γ-bond: 4b () 1.25, –1.83, 0.88; 4c () 0.65, 0.02, 0.88; 4d () 0.24, 1.71, 0.29. Underlined values indicate a questionable fit.


Page 21 of 38 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


A-Ring Furanoses. The N S equilibria in the in the A-ring furanoses of cADPR (4a), 2'-deoxy 4b, and 7-deaza 4c are characterized by a larger negative ∆H° value (–0.97 ± 0.16 kcal/mol) and a smaller negative T∆S° value (–0.46 ± 0.18 kcal/mol), somewhat smaller than that of adenosine (Ado; ∆H°, T∆S° –1.05, –0.62 kcal/mol). The Chattopadhyaya60a group found that Ado, over the same temperature range as our study, experienced a broader change in equilibrium composition (10% decrease in S-type) with increasing temperature compared to A-ring of cADPR (7% decrease in S-type). One might expect that cADPR, being cyclic, has less conformational freedom than its acyclic adenosine counterpart. One stereoelectronic effect that drives the NS equilibrium is the anomeric effect (AE). The AE (donation of an O4' lone pair into the C1'-N9 antibonding σ* orbital) increases with increasing electron demand of the nucleobase, or with protonation of the nucleobase. The AE is generally more favorable in N-type conformations.60a–d In two studies60b,c of the effect of pH on the NS equilibrium, the same group found a dramatic decrease in the amount of S-type (or increase in N-type) conformer at pH values at or below the pKa of the adenine N1 atom. The NS ∆H° for protonated adenosine (Ado•H+) had dropped to a mere –0.05 kcal/mol, which they attributed to differing strengths of the AE in the neutral (weaker) and protonated (stronger) forms. The N6-amino group of cADPR/analogs is protonated at physiological pH,28,61 and yet their A-ring NS ∆H° values are hardly diminished relative to neutral Ado. It is curious that the A-ring furanoses in 4a, 4b, and 4c 'resist' the AE and seem thermodynamically to be more similar to neutral Ado. The ∆H° for the A-ring NS equilibria in 8-Br 4d, on the other hand, is significantly diminished (-0.24 kcal/mol) compared to 4a, 4b, and 4c, suggesting perhaps a steric interaction between the 8-bromo substituent and the A-ring furanose in the S-form. Interestingly, the 8-Br 4d NS equilibria is the only A-ring furanose with a positive T∆S° term, so while the – ∆H° contribution to ∆G° has been reduced, apparently a more disordered S-type conformer compensates. R-Ring Furanoses. The N S equilibria in the R-rings of 4a, 4c, and 4d are characterized by small values of ∆H° and positive T∆S° terms that favor S-type conformers, demonstrating that indeed the R-ring N S equilibria have a significant entropy component. These R-rings are inherently more disordered than their A-ring counterparts. This is somewhat surprising; in a comprehensive study of nine dideoxy-, deoxy-, and ribo nucleosides Plavec60a et al. found only one (deoxycytidine) whose



Page 22 of 38

Table 4. Thermodynamic parametersa at 298 K for the furanose rings, γ–bonds, and β–bonds in cADPR A Ring ∆S° T∆S° –1.20 –0.36 –2.17 –0.65 –0.95 –0.28 +0.52 +0.15 –0.62 +0.10

∆G° –0.45 –0.49 –0.60 –0.40

b

Ring T∆S° +0.64 +0.33 +0.29 +0.73

∆G° –0.61 –0.67 –0.54 –0.63

Pop.b 26:74 24:76 29:71 26:74

4a 4b 4c 4d

NS NS NS NS

∆H° –0.81 –1.13 –0.88 –0.24

Adoc Ado•H+c

NS NS

–1.05 –0.05

4a 4b 4c 4d

γt γ+ γt γ+ γt γ+ γt γ+

+0.52 +1.23 +0.37 +0.16 57:43 +0.25 –0.64 –0.19 +0.44 68:32 +0.60 +1.07 +0.32 +0.29 62:38 +0.54 +1.89 +0.56 –0.02 51:49

–1.81 –0.24 –0.07 –2.49 –3.64 –1.08 –1.30 +0.04 +0.01 –0.47 +3.39 +1.01d

-1.74 5:95 –1.41 8:92 –1.31 10:90 –1.48 8:92

4a 4b 4c 4d

β-/+ βt β-/+ βt β-/+ βt β-/+ βt

–1.03 +0.80 +0.24 –2.50 –3.96 –1.18 –0.88 +1.44 +0.43 –1.39 –0.96 –0.29

–4.51 –8.72 –2.60 –3.42 –5.32 –1.58 –2.33 +1.43 –0.43 –3.96 –7.46 –2.22

–1.91 –1.84 –1.91 –1.74

Pop. 32:68 31:69 27:73 34:66

∆H° +0.04 –0.33 –0.25 +0.10

R ∆S° +2.15 +1.12 +0.97 +2.45

–0.43 33:67 –0.15 44:56

–1.27 –1.32 –1.31 –1.10

10:90 10:90 10:90 13:87

4:96 4:96 4:96 5:95

∆Ho, T∆So, and ∆Go in kcal/mol, ∆S in cal/mol K. Underlined values denote the major contributor to ∆Go. Populations recalculated based on the calculated ∆Go; compare to Table 3. Values in bold are discussed in the A Model for cADPR section. cData from ref. 60b,c. Ado = adenosine, Ado•H+ = N1-protonated adenosine. d Poor fit in the van't Hoff plot; R2 = 0.29. a b

∆Ho value did not make a significant contribution to its ∆Go. But now the comparison to Ado•H+ is more appropriate. The positive charge in Ado•H+ led to a 0.72 kcal/mol diminishment of a T∆S° term that originally favored N-type neutral Ado (T∆S° = –0.62 kcal/mol in Ado and +0.10 kcal/mol in Ado•H+). The positive charge in cADPR/analogs is closer to the R-ring than to the A-ring and thus reasonably exerts more influence on the R-ring T∆S° term. Whether the ∆H° value for the N S equilibria in the 2'-deoxy-4b and 7-deaza-4c is truly negative, though, will require further study on additional analogs over greater temperature ranges. A van't Hoff plot lacking significant slope is extremely sensitive to additional data points, but it is safe to say, based on the general lack of effect of temperature on the R-ring N:S ratios that the ∆H° term is small and that the T∆S° term dominates the N S equilibria. While thermodynamically the R-rings resemble Ado•H+ (comparable ∆H° and T∆S° terms), neither the A-ring nor R-ring populations shift much from their approximate 1:3 N:S ratios, suggesting again some resistance to conformational change. 22 ACS Paragon Plus Environment

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γ–Bonds. In contrast to all the other conformationally mobile rotatable bonds, where a distinct preference one conformer is observed, the A-ring γ–bonds exists as a mixture of the γt and γ+ rotamers, with a slight preference for the trans-rotamer. The ∆H° and T∆S° terms are more evenly matched, with cADPR (1a), 7-deaza analog 4c, and 8-bromo analog 4d possessing positive ∆H° values that disfavor the γ+rotamer but positive T∆S° values that favor it. The T∆S° term, however, for 4b is negative, and the γ+ state is disfavored both enthalpically and entropically. Of all the analogs, 4b has the highest populated A-γt state, ~69%. The situation is different for the R-ring γ–bonds, most of which show a strong (>90%) ∆H°-driven preference for the γ+ rotamer. (While the R-ring γ–bond in cADPR analog 4d exists almost exclusively as the γ+ rotamer, the poor fit in the van't Hoff plot (R2 = 0.29) makes it impossible to conclude that it is too is favored for enthalpic reasons.) The crystal structure, of the free acid of cADPR, showed both γ–bonds as trans. Our recent MD simulation40 of cADPR found both γ– bonds to be γ+ and a north-favoring A-ring, though once the A–γ bond was restrained to trans a south A– ring was favored. We suspect this may be due to the bsc0 refinement of the AMBER force field which was designed to minimize the γt state.62 Thus three different experiments – this NMR study (R–γ+ and a mixture of A–γt and A–γ+), our recent MD simulation (R–γ+ and A–γ+ when unrestrained), and the crystal structure (of the free acid; R–γt and A–γt) – give three different outcomes for the γ–bonds. Clearly the A–γ bond, finely balanced between its two observed orientations, is sensitive to its environment. Analog 4b is also unusual in that its R-γ bond shows much larger favorable –∆H° and much larger unfavorable –T∆S° values for the γ+ state compared to the other three compounds, though the ∆G° values are comparable. The removal of the A-ring 2'-OH somehow greatly enthalpically favors the remote R-γ+ state while simultaneously penalizing it entropically. It is to be expected that there will be differences in the conformational preferences of the γ–bonds in cADPR compared to those of the free nucleosides. In the Chattopadhyaya60a group study they noted that without exception the population of the γ+ state decreased with increasing temperature, regardless of whether the γ+ state was the favored rotamer or not. The conclusion to be drawn is that in free nucleosides the γt state is the more disordered rotamer. Only in the R-ring γ–bonds of 4a–4d was this trend observed; the A-γ+ state became more populated with increasing temperature. Generally in 4a–4d we can say the more disordered rotamers are the A-γ+ and the R-γt. The adenine base is necessarily in the syn orientation in cADPR and the analogs. Given that in free nucleosides a syn conformation



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generally decreases the γ+ population48,58a it is remarkable the A-γ+ state in cADPR is populated to the extent that it is. β–Bonds. The βt is favored for enthalpy reasons and is highly populated in both rings of cADPR 4a and analogs 4b, 4c, and 4d. The T∆S° term is smaller than the ∆H° term by an approximate factor of two, and the favorable ∆H° for the R-ring β–bonds is greater than that of the A-ring β–bonds by the same factor. The T∆S° term for the A-ring β–bonds is somewhat variable in sign, but the R-ring β–bond is always negative. A Model for cADPR. Based on the above discussion, we would propose the following model (Figure 7) for cADPR that accommodates our data, potentially explains some of the SAR for cADPR, and provides direction for future efforts. What parts of the story should the model include? It is clear from the discussion above that the conformations of cADPR 4a and analogs 4b, 4c, and 4d are overall quite similar, meaning activity cannot be explained on basis of divergent structures. Changes to the preferred conformations of the A-ring β–bond, (A-β), R-β, and R-γ bonds will incur a substantial enthalpy penalty. In a sense, these subunits could be considered to be thermodynamically 'restrained'. Establishing that these conformers are kinetically restrained due to high barriers to rotation would require either lineshape analysis or substantial modeling efforts well beyond the scope of the present study. The furanose rings are certainly flexible, but there is no NMR-based evidence that the structural modifications (2'-deoxy-, 8-bromo, 7-deaza) had any impact on the preferred furanose conformations (A-ring PS ~ 168°; R-ring PS ~ 215°). If cADPR is indeed 'conformationally agnostic' as to the orientation of its furanose rings, this would seem to leave the A-ring γ–bond and the two N-glycosidic bonds, A-χ and R-χ (defined in the legend to Figure 7) as the only sites capable of variability. In our model (Figure 7), the nucleobase is positioned so that A-χ is ~60° and R-χ is ~10°, approximating the values from the crystal structure57a and our MD simulation40, with the adenine C2-N3-C4 edge pointing towards the area between A2' and A5" hydrogens. The phosphate backbone is behind the plane of the page. One attractive feature of this model is that it immediately provides a rationale for the 'unusually high' 1H chemical shift of A2', as it is edge on to the adenine and deshielded by the aromatic ring current of the nucleobase. Analogously, the higher chemical shifts of A5" and R5' relative to their geminal partners A5' and R5" are explained. Two models of cADPR (A-γt and A-γ+) were built in HyperChem and a simple vacuum geometry optimization (AMBER force field) was


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performed. The starting structures were built using the conformations listed in Tables 1 and 2: A-ring PS ~177°, R-ring PS ~216°, A-βt, R-βt, and R-γ+. Based on the very similar crystal structure/MD values, A-χ was set to +60° and R-χ to +10°. The remaining pyrophosphate backbone torsions and partial atomic charges were taken from their respective MD simulations. No attempt was made to determine if the structures were the global minimum structures, nor were alternate pyrophosphate backbone conformers considered. The resulting structures are shown in Figure 7. The close contacts and key torsion angles in the A-γt structure (Figure 7a,b) are: HA1'–H8 2.6-2.7 Å, HA2'–N3 2.5-2.8 Å, HA5"-N3 2.5-3.0 Å, HR5'–H2 2.5-3.0 Å, χA ~+60°, χR ~+5°, and HA1' and H8 eclipsed. Consider the case of an 8-bromo substituent (4d), which has the least populated (51%) A-ring γt-state of the series. The ∆H° for the A-ring NS equilibria in 4d is 0.57 kcal/mol less favorable than it was for 4a (∆∆H°A-ring = ∆H°4d – ∆H°4a = +0.57 kcal/mol), compensated for by a more favorable entropy change (T∆∆S° = +0.51 kcal/mol. What is the origin of the enthalpy penalty? To avoid an 8-Br/HA1' eclipsing interaction, the A-χ bond could rotate towards the A-ring C2', decreasing the distance between the A-ring C2' and the 8-bromo substituent, making A-χ less positive than +60°. Simultaneously this would decrease the HA5"–N3 distance, increasing the steric pressure and destabilizing the A-ring furanose S-state and γt states. The 7-deaza analog 4c retains its agonist properties perhaps because the exchange of N7 for a methine does not perturb χ. Why it should be that 2'-deoxy cADPR 4b, which has the most populated (68%) A-ring γt-state of the series, is more difficult to explain. We note that our earlier study of 3'-OMeA-cADPR39 clearly showed an altered A-γ bond, also difficult to rationalize. It would seem A-ring substituents in the cADPR system can have unpredictable effects on the conformation of other remote groups. This model – the notion that (1) agonistic/antagonistic potency is derived in part from an HA5"–N3 interaction that impacts the A-γt population and subtly alters the A-χ torsion and (2) that the R-ring S-type conformation is more disordered – can be tested. A hydrogen bond between the A-ring 2-OH and the amino group in the potent antagonist 8-amino cADPR would 'lock in' a reduced A-χ value, which should reduce the A-γt population more significantly than was the case for 8-bromo analog 4d. The 8-amino-2'-deoxy cADPR analog, a much less potent antagonist,24 lacks such an interaction, and would have a more relaxed A-χ torsion; it also accounts why a similar deletion in 4d does not lead to a reduction is antagonistic potency. Conversely, the 3-deaza analog 9a is the most potent analog known to date, and the increased steric bulk of the new methine group should make A-χ more positive than +60° 25 ACS Paragon Plus Environment


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as well as impact the A-γ conformation. To the second point, it could be that the R-ring is just a spacer between the adenine-N1 and the phosphate backbone, and in its entropically favored S-conformer allows local solvent to become more disordered. Certainly solvent is less organized about the cIDPRE analogs 11 and 12, where the R-ring has been replaced with an ether linkage. In contrast to the antagonistic activity displayed by most 8-substituted analogs, 8-NH2-cIDPRE and 8-N3-cIDPRE (12) are agonists, equipotent with cADPR, perhaps because the nucleobase, free of its R-ring, is not forced to adopt the χ angles that promote antagonism.

Figure 7. A model for cADPR. (a) Schematic representation of 4a–d based on our NMR data and the cADPR crystal structure/MD simulation values for the glycosidic bonds A-χ and R-χ. (b) and (c) HyperChem (HC) minimized structures (Amber99). The A-ring is in the γt (b) and γ+ (c) conformation. The A-χ bond was defined using standard accepted rules, but the R-χ bond defined as O4'R–C1'–N1–C2 so that it will have the same magnitude and sign as the A-ring.

Of course, the conformational behavior of a ligand in solution, even if correctly modeled, is never guaranteed to translate to the ligand in the bound state. The penalty cADPR or an analog would have to pay to bind in, for example, the (solution state) less favored A-γ+ conformation (< 0.4 kcal/mol) could easily be compensated for by a single favorable hydrogen bond contact to its binding protein. The point 26 ACS Paragon Plus Environment

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we would make is that this thermodynamic study and the model generated therein provide new insights and testable ideas for the study of the conformational behavior and SAR of cADPR. Additional detailed thermodynamic and structural studies of these and other cADPR analogs will be investigated in due course.

CONCLUSION A series of previously known cADPR analogs were synthesized and their solution state behavior studied utilizing variable temperature 1H NMR spectroscopy. Analysis of the coupling constants using PSEUROT and other methodologies allowed us to determine the conformations about majority of the rotatable bonds in this 20-membered macrocycle. Population and van't Hoff analyses provided us with the thermodynamics of the N S, γt γ+, and β+/– βt equilibria in both the A-ring and R-ring of cADPR. By combining the identities of the conformers with knowledge of ∆H°, ∆S°, and ∆G° for each of their conformational equilibria, we were able to propose a model that explains how small variations in analog structure can drive the associated equilibria in unexpected ways.

EXPERIMENTAL SECTION General Information. Moisture-sensitive reactions were performed under an atmosphere of dry nitrogen using syringe techniques. Phosphoryl chloride, triethyl phosphate (TEP), diphenyl phosphoryl chloride (DPPC), tri-n-octylamine (TOA), tri-n-butylamine (TBA), and anhydrous DMF, stored in a desiccator box, were used as obtained from commercial suppliers. Pyridine and triethylamine were distilled from KOH; 1,4-dioxane and diethyl ether were filtered through activated alumina prior to use. Deionized water was 18 MOhm/cm quality. Nucleoside/nucleotide starting materials were obtained from commercial sources and not dried prior to use. 2',3'-di-O-acetyl NMN was prepared as described (Graham, S.M; Macaya, D.J.; Sengupta, R.N.; Turner, K.B. Org. Lett. 2003, 6, 233–236). ADP ribosyl cyclase was from Sigma-Aldrich. Analytical HPLC utilized a 150 × 4.6 mm C18 column and UV-diode array detection. Eluant A was 20 mM pH 3 potassium phosphate buffer (KPB), eluant B a 90:10 mixture of eluant A:2-propanol, and eluant D 40:60 CH3CN:H2O. The flow rate was generally 1 mL/min. Preparative ion-exchange chromatography was done using a 10 × 1.6 cm Sepharose Q



Page 28 of 38

column (GE Healthcare/Pharmacia HiPrep Q HP 16/10, flow rate 5 mL/min) and water:0.25 M formic acid (FA), ammonium formate (AF), triethylammonium bicarbonate (TEAB), or triethylammonium acetate (TEAA) gradients. 'Lyophilized' signifies a frozen aqueous sample lyophilized three time from water; 'evaporation' signifies removal of water or other high-boiling solvents on a rotary evaporator connect to a mechanical oil pump. NMR Spectroscopy. NMR spectra (400 MHz) were acquired on a spectrometer outfitted with a variable temperature (VT) unit which had been recently calibrated. All 1H spectra (routine 1H, {31P}1H, 1D TOCSY) were collected with a spectral width of 8000 Hz, 64K time domain data points, and zerofilled to 64K points (spectral resolution of 0.12 Hz/pt). VT spectra were collected similarly but zerofilled to 256K points (spectral resolution of 0.03 Hz/pt). The 1D TOCSY mixing time was generally 100 ms. Resolution enhancement was routinely applied (lb ~–1 Hz and gb ~0.08-0.12) to all spectra prior to transform. The J values in Table 1 were obtained with more aggressive processing (lb -2, gb 0.2 for ring signals and up to lb -4 for 5'/5" signals. Samples were prepared by lyophilizing once from ~ 1 mL of 99.9% D2O and subsequent dissolution in 99.96% D2O. The internal standard used was 3-(trimethylsilyl)-2,2,3,3-tetradeuteropropionic acid (TMSP-d4), and spectra were generally shimmed so that the TMSP half-height linewidth was 0.7 Hz or less. All cADPR spectra were approximately 10-20 mM concentration. When given, assignments are based on either a COSY or a 1D TOCSY experiment. PSEUROT Calculations. Version 6.3 of the PSEUROT program was used. PSEUROT no longer runs on 64 bit PC-operating systems, but will run on a Windows XP 'virtual machine'. Version 6.3 utilizes the Donders form of the generalized Karplus equation (GKE). Substituent electronegativities were as follows: heterocyclic base (0.56), O4' (1.26), 2'-OH (1.26), C3' (0.62), C1' (0.62), C4' (0.62), C2' (0.62), 3'-OH (1.26), C5' (0.68). PSEUROT uses 'A' and 'B' parameters to convert the endocyclic ring torsion angles to the exocyclic H-C-C-H needed by the GKE to calculate expected J values. Standard values appropriate for ribofuranoses (4c, 4d, and 4b R-ring) were (reported as A,B): 1'-2' bond (1.102, 123.3), 2'-3' bond (1.090, 0.2), 3'-4' bond (1.095, –124.9). For the 4b A-ring deoxyribofuranose the values were: 1'-2' bond (1.030, 121.4), 1'-2: bond (1.020, 0.9), 2'-3' bond (1.060, 2.4), 2"-3' bond (1.060, 122.9), 3'-4' bond (1.090, –124.0). The puckering amplitude of the minor north conformers were restrained to 35° (A-ring) and 40° (R-ring). Full details and sample PSEUROT input files can be found in the Supporting Information. The program and an updated (2013) pdf version of the PSEUROT 6.3 manual is available from the corresponding author upon request.


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General Procedure for 5'-Phosphorylation (2). A suspension of adenosine analog 1c or 1d (~399 µmol) was suspended in 1400 µL of TEP, cooled to 0 °C in an ice-water bath, and POCl3 (77 µL, 824 µmol) added via syringe. The suspension clarified within approximately 15 minutes and was allowed to stir at 0°C. Reaction progress was monitored by HPLC (95:5 A:B, isocratic), following the disappearance of the starting material (tR ~11 min) and the appearance of product (tR ~4 min). After 1 h, the reaction was transferred via cannula to 50 mL of ice-cold rapidly stirred dry ether. The resulting precipitate was briefly centrifuged, supernatant decanted, and the process repeated with 2 × 2 mL portions of ether. After briefly drying under a stream of nitrogen, the precipitate was dissolved in water (40 mL) and stirred at room temperature for 1 h. The pH of the crude product solution was adjusted to 7.6 by the addition of KOH, and the reaction mixture purified on the Sepharose Q column (formate form) utilizing a gradient of 4% - 25% FA over 1000 mL. Product eluted at approximately 50 mM FA. Product fractions were evaporated to dryness, evaporated twice more from water, three times from methanol, and dried under high vacuum overnight. To improve solubility in the coupling with NMN, AMP analogs were converted their trioctylammonium salts. Thus, to a suspension of the free acids of AMP analogs 2b, 2c, or 2d (~200 µmol) in 5 mL of HPLC grade methanol was added one equivalent (~83 µL, 200 µmol) of TOA. After stirring at room temperature for 15 minutes, the resulting solution was evaporated to dryness and briefly dried under high vacuum. Final traces of water were removed by evaporation from dry dioxane. The resulting analog AMP•TOA salt was used without purification in the next step. 7-Deazaadenosine 5'-monophosphate tri-n-octyl ammonium salt (2c).17 From 106.3 mg (399 µmol) of 1c was obtained 63.3 mg (183 µmol, 46%) of the free acid of 2c as a clear glass, which was converted to its trioctylammonium salt. 1H NMR (400 MHz, CD3OD): δ 8.066 (s, 1H, H2), 7.576 (d, 1H, J = 3.80, H8), 6.629 (d, 1H, J = 3.76, H7), 6.255 (d, 1H, J = 5.88, H1'), 4.490 (t, 1H, J = 5.5, H2'), 4.351 (dd, 1H, J = 5.16, 3.40, H3'), 4.179 (m, 1H, H4'), 4.090 (m, 2H, H5', H5"), 3.045 (m. 6H, N–(CH2)3, 1.668 (m, 6H, N–(CH2CH2)3, 1.30 (m, 30H), 0.886 (t, 9H, J = 6.9 Hz, 3 × –CH3).

31

P NMR (162 MHz, D2O):

δ 0.956. 8-Bromo-adenosine 5'-monophosphate free acid (2d) and 8-chloroadenosine 5'-monophosphate free acid (2e). 8-Bromoadenosine (1d, 69.3 mg, 200.2 µmol) was reacted as described above. HPLC analysis after 15 min showed two product peaks with retention times of 4.1 min (UV λmax = 262 nm) and 4.8 minutes (UV λmax = 264 nm) in an approximate 26:74 ratio. After 30 min, the reaction was worked 29 ACS Paragon Plus Environment


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up and chromatographed (Sepharose Q column, FA gradient) to afford 54.1 mg of a mixture (58:42 by 1

H NMR) of 8-chloro-AMP 2e (79 µmol):8-bromo-AMP 2d (57 µmol), for a combined yield of 68%.

Repeated chromatographic purification gave sufficient quantities of the free acids of 8-chloro 2e and 8-bromo-AMP 2d for characterization. 8-Chloroadenosine 5'-monophosphate free acid (2e).42,43 1H NMR (400 MHz, D2O): δ 8.19 (s, 1H, H8), 6.10 (d, 1H, J = 5.6 Hz, H1'), 5.19 (t, J = 5.7 Hz, H2'), 4.64 (t, 1H, J = 5.1 Hz, H3'), 4.31 (app q, 1H), 4.24–4.14 (m, 2H, H5'/5").

13

C NMR (100 MHz, D2O): δ 152.1, 150.0, 149.3, 139.8 (C8), 117.2,

88.6, 83.6, 83.5 (d, J = 8.2 Hz, C5'), 71.2, 69.6, 64.4. LC MS (m/z): 382 (M+H+). UV (H2O, pH 3.0) λmax, nm: 262. HPLC (C18): tR ~ 4.1 min). 8-Bromoadenosine 5'-monophosphate free acid (2d).42,43 1H NMR (400 MHz, D2O): δ 8.18 (s, 1H, H8), 6.09 (d, 1H, J = 5.6 Hz, H1'), 5.24 (t, J = 5.8 Hz, H2'), 4.65 (dd, 1H, J = 6.0, 4.7 Hz, H3'), 4.31 (app q, 1H), 4.25–4.15 (m, 2H, H5'/5").

13

C NMR (100 MHz, D2O): δ 152.9, 159.8, 149.7, 129.0 (C8),

118.8, 89.7, 83.6, 835 (d, J = 8.4 Hz, C5'), 71.2, 69.6, 64.4. LC MS (m/z): 426 (M+H+). UV (H2O, pH 3.0) λmax, nm: 264. HPLC (C18): tR ~ 4.8 min). General Procedure for the synthesis of NAD+ analogs (3). Prior to coupling with NMN, AMP analogs were converted their trioctylammonium salts. Thus, to a suspension of AMP analog 2b, 2c, or 2d (~200 µmol) in 5 mL of HPLC grade methanol was added one equivalent (~83 µL, 200 µmol) of TOA. After stirring at room temperature for 15 minutes, the resulting solution was evaporated to dryness and briefly dried under high vacuum. Final traces of water were removed by evaporation from dry dioxane or DMF. The resulting analog AMP•TOA salt was used without purification in the next step. Thus, analog AMP•TOA salt (~200 µmol) was suspended in 200 µL of DMF and 1600 µL of dioxane. Next, diphenyl phosphoryl chloride (DPPC, 62 µL, 299 µmol, 1.5 equiv) and TBA (142 µL, 601 µmol, 3 equiv) were added via syringe. The stirred suspension clarified within 10 minutes. The reaction was followed by HPLC (100% A for 1 min, a gradient to 50% B over 10 min, then to 100% D over 10 min), monitoring the disappearance of starting AMP analog (tR ~4–6 min) and appearance of the activated mixed anhydride (tR ~21–25 min). After 2–3 h the reaction was evaporated and the crude activated mixed anhydride precipitated with 2 mL of dry ether. The ether was carefully removed with a glass pipet and the remaining solid dissolved in 1 mL of DMF, evaporated, and dried briefly under high vacuum. Next, 2',3'-di-O-acetyl NMN (Ac2-NMN,120.1 mg, 287 µmol, 1.4 equiv), dissolved in 1280 µL of dry DMF, was transferred via cannula to the activated AMP analog vessel, followed by pyridine 30 ACS Paragon Plus Environment

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(640 µL) and TBA (58 µL, 240 µmol, 1.2 equiv). The reaction was stirred rt and progress monitored by HPLC as above, this time monitoring the disappearance of starting activated mixed anhydride (tR ~21– 25 min) and appearance of NAD+ analog (tR ~13 min). Reaction times of 2–3 h were typical. Solvents were then evaporated and the residue dissolved in 2 mL of 1:1 CH3OH: conc. aq. ammonia, stirred for 15 min at rt, evaporated, and dried briefly under vacuum. 2’-Deoxy-A nicotinamide adenine dinucleotide ammonium salt (2'-deoxy-NAD+, 3b).16 The free acid of commercial 2'-deoxyadenosine 5'-monophosphate was converted to its TOA salt as described above. Thus, 136.7 mg (200 µmol) of 2'-deoxy-AMP•TOA 2b was activated with DPCC, coupled with Ac2-NMN, monitored by HPLC, deacylated with methanolic ammonia, and purified (4% 25% FA over 1000 mL) as described above. While the later-eluting fractions of were pure NAD+-3b, early-eluting 3b co-eluted with 2'-dAMP 2b. The early-eluting 3b/2b could not be separated after repeated Sepharose Q ion-exchange chromatographic steps using the FA gradient. Pure NAD+-3b ammonium salt (13.6 mg, 21 µmol, 10.5 %) was eventually obtained by Sepharose Q ion-exchange chromatography utilizing ammonium formate (0% - 25% AF over 1000 mL) as the eluant. 1H NMR

(D2O, 400 MHz): δ 9.258 (br s1H, Nic-H2), 9.056 (d, 1H, J = 6.3 Hz, Nic-H6), 8.737 (d, 1H, J = 8.1 Hz, Nic-H4), 8.329 (br s, 1H, Ade-H2), 8.093 (dd, 1H, J = 8.1 Hz, 6.3 Hz, Nic-H5), 8.038 (s, 1H, AdeH8), 6.337 (t, 1H, J = 6.9 Hz, A-H1'), 6.009 (d, 1H, J = 5.5 Hz, R-A1'), 4.663 (app qn, 1H, J = 3.1 Hz, A-H3'), 4.467 (br s, 1H, R-H4'), 4.410 (t, 1H, J = 5.3 Hz, R-H2') 4.350 (dd, 1H, J = 4.9, 2.7 Hz, R-H3'), 4.277 (br d, 1H, J = 11.9 Hz, R-H5'/5"), 4.215 (m, 1H, A-H4'), 4.17–4.05 (m, 3H, R-H5'/5", A-H5', AH5"), 2.755 (ddd, 1H, J = 14.00 Hz, 7.6 Hz, 6.1 Hz, A-H2'/H2"), 2.496 (ddd, 1H, J = 14.0 Hz, 6.3 Hz, 3.3 Hz, A-H2'/H2").

31

P NMR (162 MHz, D2O): δ -11.15 (d, J = 20.3 Hz), -11.53 (d, J = 20.3 Hz).

7-Deaza nicotinamide adenine dinucleotide ammonium salt (7-deaza-NAD+, 3c).17 The free acid of compound 2c (63.3 mg, 183 µmol) was converted to its TOA salt as described above, and a portion carried on to the coupling reaction. Thus, 122.2 mg (174.6 µmol) of 7-deaza-AMP•TOA 2c was activated with DPCC, coupled with Ac2-NMN, with both steps monitored by HPLC and as described above. After ammonia/CH3OH deprotection, Sepharose Q ion-exchange chromatography (ammonium formate gradient) afforded 31.3 mg (47.3 µmol, 27%) of the ammonium salt of NAD+ analog 3c containing a small amount of ammonium formate. 1

H NMR (D2O, 400 MHz): δ 9.297 (br s1H, Nic-H2), 9.093 (d, 1H, J = 6.3 Hz, Nic-H6), 8.715 (dt,

1H, J = 8.1, 1.5 Hz, Nic-H4), 8.104 (dd, 1H, J = 8.2 Hz, 6.2 Hz, Nic-H5), 7.950 (s, 1H, Ade-H2) 7.416 31 ACS Paragon Plus Environment


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(d, 1H, J = 3.9 Hz, Ade-H8), 6.475 (d, 1H, J = 3.8 Hz, Ade-H7), 6.129 and 6.114 (d, 1H, J = 6.5, A-H1' and d, 1H, J = 5.1 Hz, Nic-1'), 4.663 (dd, 1H, J = 6.3, 5.5 Hz, A-H2'), 4.587 (app qn, 1H, J = 2.4 Hz), 4.49–4.40 (m, 4H), 4.354 (app qn, 1H, J = 2.2 Hz), 4.29–4.17 (m, 3H).

31

P NMR (162 MHz, D2O): δ -

11.26 (br), -11.53 (br). 8-Bromo nicotinamide adenine dinucleotide (8-bromo-NAD+, 3d).22 The free acid of 8-bromo AMP (2d, 46.8 mg, 110 µmol) was converted to its TOA salt as described above. Thus, 8-bromo-AMP•TOA 2d (110 µmol) was activated with DPCC and subsequently coupled with Ac2-NMN (34 mg, 81.8 µmol, 670 µL DMF). The reaction was monitored by HPLC and as described above and after 2 h (~70% completion) a second portion of Ac2-NMN (34 mg, 81.8 µmol, dissolved in 340 µL DMF) was added, with continued stirring for an additional 2 h. The reaction was evaporated to dryness, deprotected with methanolic ammonia, and evaporated to dryness. Sepharose Q ion-exchange chromatography (AF gradient) afforded 47.6 mg (64 µmol, 58%) of the ammonium salt of NAD+ analog 3d containing a small amount of ammonium formate. 1H NMR (D2O, 400 MHz): δ 9.316 (s, 1H), 9.144 (d, 1H, J =

6.24 Hz), 8.850 (d, 1H, J = 8.12 Hz), 8.231 (dd, 1H, J = 7.98 Hz, 6.35 Hz), 8.079 (s, 1H), 6.024 (d, 1H, J = 5.16 Hz), 5.971 (d, 1H, J = 5.52 Hz), 5.222 (t, 1H, J = 5.86 Hz), 4.633 (t, 1H, J = 5.48 Hz), 4.47–4.40 (m, 3H), 4.38–4.20 (m, 5H).31P NMR (D2O, 162 MHz): δ -11.32 (d, J ~ 20 Hz), -11.76 (d, J ~ 19 Hz). General Procedure for the synthesis of cADPR analogs (4). An NAD+ free acid or ammonium salt analog (3b, 3c, or 3d, ~50 µmol) was dissolved in 15 mL of 25mM HEPES buffer, pH 7.0, 45 mL of deionized H2O, and the pH adjusted to 7.0 by addition of 0.1 M KOH. Next, 60 µL (approximately 15 Units; 1 Unit = 1µmol NAD+ converted in 5 minutes) of ADPRC stock solution was added. Reaction progress was monitored by HPLC (95:5 A:B, isocratic), following the disappearance of the NAD+ analog (tR ~5 min) and the appearance of cADPR analog (tR ~4 min). Once the amount of cADPR analog ceased to increase, the pH of the reaction was adjusted to 6.5 with acetic acid and the crude mixture loaded onto the Sepharose Q column. Elution was performed with a gradient of water and 0.25 M TEAB or TEAA, with cADPR analog eluting at ~75 mM salt. Product fractions were combined, lyophilized (3×), evaporated from CH3OH (3×), and dried under high vacuum. Yields were typically 60–75%.


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2’-Deoxy cyclic adenosine diphosphate ribose (2'-deoxy-cADPR, 4b).16 From 3b (21.4 mg, 33 µmol, 2 h reaction time) was obtained after chromatography (TEAB) 15.8 mg (24.7 µmol, 75%) of 2'-dA-cADPR 4b. 1H NMR (400 MHz, D2O, 277 K): δ 9.043 (s, 1H, H2), 8.387 (s, 1H, H8), 6.538 (t, 1H, J = 6.90 Hz, A1'), 6.165 (d, 1H, J = 4.00 Hz, R1'), 4. 946 (overlapping dt, 1H, J ~ 6.3, 3.3 Hz, A3'), 4.829 (dd, 1H, J = 5.04, 4.07, Hz, R1'), 4.791 (app qn, 1H, R4'), 4.568 (ddd, 1H, J = 10.87,

7.43 Hz, 3.45 Hz, A5"), 4.512 (dd, 1H, J = 5.06, 2.10 Hz, R3'), 4.401 (dt, 1H, J = 11.85, 2.03 Hz, R5'), 4.316 (dt, 1H, J = 7.36, 2.83 Hz, A4'), 4.147 (dt, 1H, J = 12.48, 2.81 Hz, R5"), 4.063 (dt, 1H, J = 10.88, 3.03 Hz, A5'), 3.353 (dt, 1H, J = 14.20, 6.30 Hz, A5'), 2.574 (ddd, 1H, 14.21, 6.96, 3.58 Hz). 1H{31P} NMR (D2O, 400 MHz) (Only the signals that changed upon 31P decoupling are listed): δ 4.791 (collapse to app q, 1H, J ~ 2 Hz), 4.567 (dd, 1H, J = 10.80 Hz, 7.44 Hz), 4.401 (dd, 1H, J = 11.86 Hz, 1.83 Hz), 4.147 (dd, 1H, J = 11.94 Hz, 2.06 Hz), 4.063 (dd, 1H, J = 10.82 Hz, 2.86 Hz).

31

P NMR (D2O, 162 MHz): δ -10.74 (d, J = 12.6 Hz), -11.5 (d,

J = 13.2 Hz)

7-Deaza cyclic adenosine diphosphate ribose, triethylammonium salt (7-deaza-cADPR, 4c).17 From 3c (31.3 mg, 47.3 µmol, 2 h reaction time) was obtained, after Sepharose Q ion-exchange chromatography (TEAA) and contaminated with a small amount of triethylammonium formate, 33.1 mg of cADPR analog 4b, in nearly quantitative yield. 1H NMR (D2O, 400 MHz, 298 K): δ 8.880 (s,

1H, H2), 7.429 (d, 1H, J = 3.72 Hz, H8), 6.843 (d, 1H J = 3.68 Hz, H7), 6.105 (d, 1H, J = 4.04 Hz, R1'), 5.884 (d, 1H, J = 6.28 Hz, A1'), 5.467 (dd, 1H, J = 6.18 Hz, 5.22 Hz, A2'), 4.75–4.67 (m, 3H, R4', A3', R2'), 4.560 (ddd, 1H, J = 10.89 Hz, 7.23 Hz, 3.55 Hz, A5"), 4.458 (dd, 1H, J = 5.00 Hz, 2.52 Hz, R3'), 4.414 (dt, 1H, J = 11.25 Hz, 2.22 Hz, R5'), 4.353 (dt, 1H, J = 7.04 Hz, 2.69 Hz, A4'), 4.153 (dt, 1H, J = 11.96 Hz, 2.89 Hz, R5"), 4.085 (dt, 1H, J = 11.52 Hz, 3.12 Hz, A5'), 3.21 (q, ~6H, J = 7.3 Hz, N–(CH2)3), 1.28 (t, ~9H, J = 7.3 Hz, 3 × N–CH2CH3). 1H{31P} NMR (400 MHz, D2O, 298 K) (Only the signals that changed upon 31P decoupling are listed): δ 4.559 (dd, 1H, J = 10.94 Hz, 6.97 Hz), 4.411 (dd, 1H, J = 11.98 Hz, 2.03 Hz), 4.153 (dd, 1H, J = 11.95 Hz, 2.49 Hz), 4.085 (dd, 1H, J = 10.98 Hz, 2.52 Hz).

31

P NMR (D2O, 162 MHz, 298 K): δ -

10.71, -11.36.



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8-Bromo cyclic adenosine diphosphate ribose (8-Bromo cADPR, 4d).15,22 Compound 3d (31.3 mg, 47.3 µmol) was incubated with 90 µL (~22 units) of ADPRC. After 90 min, HPLC analysis showed ~75% conversion of 3d (tR ~9 min) to 4d (tR ~6.5 min). After a total reaction time of 2 h, Sepharose Q ion-exchange chromatography (TEAA) and lyophilisation as usual afforded 31.4 mg (43.5 µmol, 68%) of 8-bromo-cADPR 4d containing a small amount of TEAA. 1H NMR (400 MHz, D2O, 277 K): δ

9.023 (s, 1H, H2), 6.192 (d, 1H, J = 5.44 Hz, A1'), 6.148 (d, 1H, J = 3.96 Hz, R1'), 5.493 (t, 1H, J = 5.26 Hz, A1'), 4.7972 (m, 3H, A3', R2', R4'), 4.505 (dd, 1H, J = 5.04 Hz, 2.24 Hz, R3'), 4.425 (m, 3H, A5"/ R5'/A4'), 4.160 (m, 1H, R5"), 4.053 (dt, 1H, J = 11.64 Hz, 2.71 Hz, A5'), 3.2 (q, ~6H, J ~7 Hz, N–(CH2)3), 1.3 (t, ~9H, J ~7 Hz, 3 × N–CH2CH3). 1H{31P} NMR (400 MHz, D2O, 298 K) (Only the signals that changed upon 31P decoupling are listed): δ 4.780 (collapse to app q, 1H, J ~ 2.1 Hz), 4.459 (dd, 1H, J = 11.20, 6.17 Hz, A5"), 4.416 (dd, 1H, J = 11.97, 2.12 Hz, R5'), 4.161 (dd, 1H, J = 11.98, 2.20 Hz, R5"), 4.053 (dd, 1H, J = 11.12, 2.34 Hz, A5').

ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge on the ACS Publications website at DOI:. Details of the PSEUROT calculations; details of the γ– and β–bond population calculations; all VT spectra (PDF).

AUTHOR INFORMATION Corresponding Author *E-mail: [email protected]. ORCID Steven M. Graham: 0000-0003-2058-2506

Notes The authors declare no competing financial interest.

ACKNOWLEDGMENTS


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Leonard Barasa and Sabesan Yoganathan for acquiring the LCMS spectra. Support from St. John's University, in the form of a sabbatical leave, is gratefully acknowledged. Uroš Javornik and Janez. Plavec (Slovenian NMR Centre) for hosting the corresponding author during the sabbatical leave.

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(21) Based on its name – cADPR, for 'cyclic adenosine diphosphate ribose', we prefer to call the ribofuranose ring derived from adenosine (the 'A' in 'cADPR') the 'A-ring' and the other ribofuranose ring (the 'R' in 'cADPR') the 'R-ring'. Most others call these rings the 'southern' and 'northern' ribose rings, respectively, but we fear this can lead to confusion in a conformational analysis study such as this the terms 'north' and 'south' are used extensively as conformational descriptors of the furanose ring geometries. (22) Moreau, C.; Ashamu, G.A.; Bailey, V.C.; Galione, A.; Guse, A.H.; Potter, B.V.L. Org. Biomol. Chem. 2011, 9, 278–290. (23) Zhang, B.; Bailey, V. C.; Potter, B. V. L. J. Org. Chem. 2008, 73, 1693–1703. (24) Zhang, B.; Wagner, G. K.; Weber, K.; Garnham, C.; Morgan, A. J.; Galione, A.; Guse, A. H.; Potter, B. V. L. J. Med. Chem. 2008, 51, 1623–1636. (25) Wagner, G. K.; Black, S.; Guse, A. H.; Potter, B. V. L. Chem. Commun. 2003, 1944. (26) Wagner, G. K.; Guse, A. H.; Potter, B. V. L. J. Org. Chem. 2005, 70, 4810–4819. (27) Swarbrick, J. M.; Graeff, R.; Zhang, H.; Thomas, M. P.; Hao, Q.; Potter, B. V. L. J. Med. Chem. 2014, 57, 8517–8529. (28) Guse, A. H.; Cakir-Kiefer, C.; Fukuoka, M.; Shuto, S.; Weber, K.; Bailey, V. C.; Matsuda, A.; Mayr, G. W.; Oppenheimer, N.; Schuber, F.; Potter, B. V. L. Biochemistry 2002, 41, 6744–6751. (29) Shuto, S.; Fukuoka, M.; Manikowsky, A.; Ueno, Y.; Nakano, T.; Kuroda, R.; Kuroda, H.; Matsuda, A. J. Am. Chem. Soc. 2001, 123, 8750–8759. (30) Shuto, S.; Fukuoka, M.; Kudoh, T.; Garnham, C.; Galione, A.; Potter, B. V. L.; Matsuda, A. J. Med. Chem. 2003, 46, 4741–4749. (31) Takano, S.; Tsuzuki, T.; Murayama, T.; Kameda, T.; Kumaki, Y.; Sakurai, T.; Fukuda, H.; Watanabe, M.; Arisawa, M.; Shuto, S. J. Med. Chem. 2017, 60, 5868−5875. (32) Zhang, F.-J.; Yamada, S.; Gu, Q.-M.; Sih, C. J. Bioorganic & Med. Chem. Lett. 1996, 6, 1203– 1208. (33) Xu, L.; Walseth, T. F.; Slama, J. T. J. Med. Chem. 2005, 48, 4177–4181. (34) Gu, X.; Yang, Z.; Zhang, L.; Kunerth, S.; Fliegert, R.; Weber, K.; Guse, A. H.; Zhang, L. J. Med. Chem. 2004, 47, 5674–5682. (35) Qi, N.; Jung, K.; Wang, M.; Na, L. X.; Yang, Z. J.; Zhang, L. R.; Guse, A. H.; Zhang, L. H. J. Chem. Soc. Chem. Commun. 2011, 47, 9462–9464. (36) Li, L.; Siebrands, C. C.; Yang, Z.; Zhang, L.; Guse, A. H.; Zhang, L. Org. Biomol. Chem. 2010, 8, 1843–1848. (37) Kudoh, T.; Fukuoka, M.; Ichikawa, S.; Murayama, T.; Ogawa, Y.; Hashii, M.; Higashida, H.; Kunerth, S.; Weber, K.; Guse, A. H.; Potter, B. V. L.; Matsuda, A.; Shuto, S. J. Am. Chem. Soc. 2005, 127, 8846–8855. (38) Graham, S.M.; Pope, S.C. Nucleosides, Nucleotides Nucleic Acids 2001, 20, 169–183. (39) Graham, S.M; Macaya, D.J.; Sengupta, R.N.; Turner, K.B. Org. Lett. 2003, 6, 233–236. (40) Javornik, U; Plavec, J; Wang, B; Graham, S.M. Carbohydr. Res. 2018, 455, 71-80. (41) Yoshikawa, M.; Kato, T.; Takenishi, T. Bull. Chem. Soc. Jpn. 1969, 42, 3505–3508. (42) Collier, A.; Wagner, G. Org. Biomol. Chem. 2006, 4, 4526–4532. (43) Uesugi, S.; Ikehara, M. J. Am. Chem. Soc. 1977, 99, 3250–3253. (44) Repeated ion-exchange chromatography allowed the isolation of sufficient 8-Br-AMP 1d to proceed. We briefly investigated Collier's procedure (ref. 42) and attempting the Yoshikawa procedure with POBr3 rather than POCl3, but ultimately decided to use commercially available for subsequent syntheses of 8-Br-NAD+ 2d. 36 ACS Paragon Plus Environment

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(45) Michelson, A. M. Biochim. et Biophys. Acta 1964, 91, 1–13. (46) (a) The other product of the cyclization, nicotinamide, is reported to inhibit the Aplysia californica cycle with an IC50 of ~ 1 mM. See Migaud, M. E.; Pederick, R. L.; Bailey, V. C.; Potter, B. V. L. Biochemistry 1999, 38, 9105–9114. (b) Another report, however, claims an IC50 of ~ 0.04 mM. See Sethi, J. K.; Empson, R. M.; Galione, A. Biochem. J. 1996, 319, 613–617. (c) The Aplysia kurodai cyclase is also reported to have an IC50 of ~ 1 mM. See Inageda, K.; Takahashi, K.; Tokita, K.; Nishina, H.; Kanaho, Y.; Kukimoto, I.; Kontani, K.; Hoshino, S.; Katadat, T. J. Biochem. 1995, 117, 125–131. (47) (a) Cavanagh, J.; Rance, M. J. Magn. Reson. 1992, 96, 670–678. (b) Thrippleton, M.J.; Keeler, J. Angew. Chem. Int. Ed. 2003, 42, 3938–3941. (48) Rosemeyer, H.; Toth, G.; Golankiewicz, B.; Kazimierczuk, Z.; Bourgeois, W.; Kretschmer, U.; Muth, H. P.; Seela, F. J. Org. Chem. 1990, 55, 5784–5790. (49) The stereospecific assignments of A5", R5', R5", and A5' are based on our previous work on cADPR (Ref. 39). (50) Since the difference between the first and last peaks in first-order multiplet is the sum of the individual coupling constants, we could extract, for example, JA5'P by subtracting JA4'5 and JA5'5", determine from the 1H{31P} experiment, from the A5' 'doublet of triplets' signal. (51) Kilpatrick, J. E.; Pitzer, K. S.; Spitzer, R. J. Am. Chem. Soc. 1947, 69, 2483–2488. (52) (a) Altona, C.; Sundaralingam, M. J. Am. Chem. Soc. 1972, 94, 8205–8212. (b) Altona, C.; Sundaralingam, M. J. Am. Chem. Soc. 1973, 95, 2333–2344. (53) There are several systems for describing pseudorotation parameters in the literature. The system used herein is what is found in PSEUROT, though it is not entirely consistent with the IUPAC-JCBN rules (wherein ν2 = C1'-C2'-C3'-C4', ν3 = C2'-C3'-C4'-O4', ν4 = C3'-C4'-O4'-C1', ν0 = C4'-O4'-C1'-C2', and ν1 = O4'-C1'-C2'-C3'). Variants for the puckering amplitude symbol besides ψ are Φ, Ψ, φ, and ν. The 1982 guidelines discouraged the use of 'C2'-endo' as a descriptor; no such prohibition appears in the 1998 guidelines. See "IUPAC Recommendations 1998", Markley, J. L.; Bax, A.; Arata, Y.; Hilbers, C. W.; Kaptein, R.; Sykes, B. D.; Wright, P. E.; Wüthrich, K. Pure Appl. Chem. 1998, 70, 117-142 and "IUPAC-IUB JCBN Recommendations 1982", Eur. J. Biochem. 1983, 131, 9–15. The 1998 guidelines use ψ for puckering amplitude, ν for an endocyclic torsion and φ for an exocyclic torsion. (54) (a) C. Thibaudeau, P. Acharya, J. Chattopadhyaya, Stereoelectronic Effects in Nucleosides and Nucleotides and their Structural Implications, Uppsala University Press, Uppsala, Sweden, 2nd edn, 2005. (b) M. Miljkovic, in Electrostatic and Stereoelectronic Effects in Carbohydrate Chemistry, ed. M. Miljkovic, Springer, Boston, 2014. (c) W. Saenger, Principles of Nucleic Acid Structure, SpringerVerlag, New York, 1984. (55) De Leeuw, H. P. M.; Haasnoot, C. A. G.; Altona, C. Isr. J. Chem. 1980, 20, 108–126. (56) Altona, C.; de Leeuw, F. A. A. M. J. Chem. Soc., Perkin Trans. 2 1982, 375–384. (57) (a) Lee, H.C.; Aarhus, R.; Levitt, D. Nat. Struct. Mol. Biol. 1994, 1, 143–144. (b) Liu, Q.; Kriksunov, I. A.; Graeff, R.; Lee, H. C.; Hao, Q. J. Biol. Chem. 2007, 282, 5853–5861. (c) Kotaka, M.; Graeff, R.; Chen, Z.; Zhang, L. H.; Lee, H. C.; Hao, Q. J. Mol. Biol. 2012, 415, 514–526. (58) (a) Altona, C. Recl. des Trav. Chim. des Pays-Bas. 1982, 101, 413–433. (b) Lankhorst, P. P.; Haasnoot, C. A. G.; Erkelens, C.; Altona, C. J. Biomol. Struct. Dyn. 1984, 1, 1387–1405. (c) Haasnoot, C. A. G.; de Leeuw, F. A. A. M.; de Leeuw, H. P. M.; Altona, C. Recl. des Trav. Chim. des Pays-Bas 1979, 98, 576–577. See the Supplementary data for details. (59) It should be noted that the solution structures of 4a-4d were all determined as their 37 ACS Paragon Plus Environment


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triethylammonium salts. No significant differences in coupling constants, at least in cADPR, were observed between the sodium salt (Ref. 39) and our recent variable-temperature study (Ref. 40) of the triethylammonium salt of cADPR. In response to a reviewer's comment as to presence of divalent cations and their possible effect on conformation, we would note that none of the syntheses nor purifications employed such cations. The possibility of contamination by divalent cations also seems unlikely, given that the final purifications of 4a-4d employed TEAB and TEAA buffers. (60) (a) Plavec, J.; Tong, W.; Chattopadhyaya, J. J. Am. Chem. Soc. 1993, 115, 9734–9746. (b) Thibaudeau, C.; Plavec, J.; Chattopadhyaya, J. Pure Appl. Chem. 1996, 68, 2137–2144. (c) Luyten, I.; Thibaudeau, C.; Chattopadhyaya, J. J. Org. Chem. 1997, 62, 8800–8808. (d) Thibaudeau, C.; Földesi, A.; Chattopadhyaya, J. Tetrahedron 1998, 54, 1867–1900. (e) Koole, L. H.; Buck, H. M.; Nyilas, A.; Chattopadhyaya, J. Can. J. Chem. 1987, 65, 2089–2094. (f) Thibaudeau, C.; Plavec, J.; Garg, N.; Papchikhin, A.; Chattopadhyaya, J. J. Am. Chem. Soc. 1994, 116, 4038–4043. (61) Kim, H.; Jacobson, E. L.; Jacobson, M. K. Biochem. Biophys. Res. Commun. 1993, 194, 1143– 1147. (62) Perez, A.; Marchan, I.; Svozil, D.; Sponer, J.; Cheatham, III, T.E.; Laughton, C.A.; Orozco, M. Biophys. J. 2007, 92, 3817-3829.


Variable-Temperature NMR Spectroscopy, Conformational Analysis

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