Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel

Chapter 7

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Saccharide Structure and Reactivity Interrogated with Stable Isotopes Wenhui Zhang, Reagan Meredith, Mi-Kyung Yoon, Ian Carmichael, and Anthony S. Serianni* Department of Chemistry and Biochemistry, University of Notre Dame, Notre Dame, Indiana 46556-5670, United States *E-mail: [email protected]

Several topics in saccharide chemistry and biochemistry, which were impacted by the work of Ernest Eliel and his contemporaries, are reviewed. We show how stable isotopic enrichment, NMR spectroscopy, and modern computational methods have been applied synergistically to reveal subtle and sometimes unexpected properties of saccharides in solution. Examples include the use of stable isotopes to detect and quantify the cyclic and acyclic forms of reducing sugars in solution, and to investigate relationships between saccharide structure, conformation and the kinetics of anomerization. Thermodynamic and kinetics studies of cis-trans isomerization of the N-acetyl side-chains of saccharides are enabled by selective 13C-enrichment and saturation-transfer NMR methods. Redundant NMR spin-couplings sensitive to the same molecular torsion angle can be interpreted collectively to derive conformational populations of flexible fragments such as O-acetyl side-chains and O-glycosidic linkages. NMR studies of saccharide chemical transformations using stable isotopes reveal stereospecific skeletal rearrangements such as C1–C2 transposition that defied prior detection, opening the opportunity to develop new catalysts and/or better understand catalytic mechanisms of chemical and biochemical processes involving saccharides.

© 2017 American Chemical Society Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


Introduction Carbohydrates provide a unique and expansive playground on which to investigate the intra- and intermolecular forces that dictate conformational equilibria and dynamics of molecules in solution. This opportunity evolves from the enormous structural diversity of saccharides with respect to carbon scaffold, configuration and substitution (1). This playground is particularly appealing because saccharides are biologically important molecules found in vivo in different degrees of polymerization (i.e., monosaccharides, oligosaccharides or polysaccharides) and in different modes of molecular conjugation (i.e., free in solution or appended to proteins, lipids and other biomolecules) (2, 3). This biological relevance provides a compelling argument to investigate saccharide structure, which plays key roles in determining many important biological functions and processes, including diseases such as diabetes and cancer. Unraveling the relationships between saccharide structure and their chemical and biological functions cannot be achieved, however, by restricting studies to only those saccharides found in biological systems. Such an approach, while efficient from a biological perspective, samples only a fraction of the total structural space, space that arguably must be sampled generously in order to derive reliable relationships between saccharide covalent structure and higher-order structural features such as conformational equilibria and dynamics. The term “structure” is hierarchical (Scheme 1). In its simplest definition, it describes the atoms comprising the saccharide and the covalent bonds between them. Higher-order definitions include the absolute configuration of their constituent chiral carbons, the available conformational options (conformational equilibria), and the kinetics of exchange between accessible conformational states (dynamics). These features are influenced by solvation, be it by simple solvent molecules like water or by functional groups present in the binding site of a biological receptor. If the saccharide contains ionizable functionality, solution pH may influence some or all of these properties (4, 5).

Scheme 1. Hierarchies of molecular structure 106 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


The enormous scientific achievements of Ernest Eliel in the field of organic stereochemistry benefitted from complementary studies of saccharides. Indeed, the book entitled Conformational Analysis by Eliel, Allinger, Angyal and Morrison, published in 1965 (6), testifies to this fact, wherein many of the stereochemical principles articulated by Eliel from his studies of general organic systems were applied, tested and refined with the use of saccharides. The inclusion of Stephen Angyal as a coauthor of this seminal book was no accident; Eliel realized the central role of saccharides in confirming and amplifying the principles of stereochemistry (7–9) that he had worked to develop. Researchers who have built upon the solid foundation provided by Eliel’s seminal studies have benefitted from research tools and methods that were unavailable, perhaps unimaginable, in the mid-20th century. These tools include, among others, very high field superconducting magnets (>14 Tesla) in NMR spectroscopy to improve spectral dispersion and sensitivity (10), multi-dimensional NMR data collection to resolve and assign complex 1H NMR spectra (11), polarization transfer methods to increase NMR sensitivity and selectivity (12, 13), and routine access to highly enriched and pure stable isotopes such as 13C, 15N and 17,18O on large scales and at reasonable cost (14). The timely convergence of these tools enabled modern NMR structural studies of complex molecules having molecular weights in excess of 50 kD, a remarkable development considering that 1H NMR spectrometers at 60–90 MHz (1.41 Tesla) using permanent magnets and operating in continuous wave modes were just coming of age in the 1960’s when Eliel was conducting his research. In this chapter, several topics pertinent to the field of saccharide chemistry and biochemistry are discussed which were impacted by early work of Eliel and his contemporaries. We show how the interplay of isotopic enrichment and modern NMR methods, coupled to modern computational methods, has been used to reveal subtle and surprising properties of these important biomolecules, including unusual skeletal rearrangements.

Saccharide Anomerization The spontaneous ring-opening and ring-closing of aldoses and ketoses in solution is known as anomerization (15, 16). This process involves the acyclic aldehydo and keto forms of reducing saccharides as central intermediates (Scheme 2). The types and distributions of cyclic forms produced depend on aldose and ketose structure; typically only five- (furanose) and six -membered (pyranose) rings form, since larger and smaller rings have unfavorable enthalpies and/or entropies of activation (17). In addition to ring-opening and ring-closing, the acyclic carbonyl forms of aldoses and ketoses can also react with solvent water to give acyclic hydrate forms (gem-diols) (Scheme 2).

107 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


Scheme 2. General scheme showing exchange between cyclic and acyclic forms of an aldose in solution during anomerization

Modern experimental measurements of anomerization equilibria are commonly made by NMR spectroscopy, and 13C NMR in conjunction with selective 13C-enrichment at anomeric carbons provides a superior approach to making these determinations (18–20). This application is illustrated in Figure 1, which shows a 13C{1H} NMR spectrum (150 MHz) obtained on an aqueous (2H2O) solution of D-[1-13C]mannose (1) in which six monomeric forms are detected in equilibrium.

Since 1 contains 99 atom-% 13C isotope at C1, the detection of the labeled carbons is ~100 times greater than for the remaining natural abundance carbons. Signals from the weak natural carbons can be observed between 60–80 ppm. The intense signals at ~95 ppm arise from the labeled C1 carbons of the dominant pyranose forms, while the furanose C1 signals appear slightly downfield of the pyranose C1 signals. The very weak signal observed at ~205 ppm arises from labeled C1 of the acyclic aldehyde form, while that at ~91 ppm arises from labeled C1 of the acyclic hydrate form (Tables 1 and 2). Integration of the C1 signals gives the following percentages of forms in solution at 30 °C: α-pyranose, 66.24 ± 0.05%; β-pyranose, 32.85 ± 0.06%; α-furanose, 0.64 ± 0.04%; β-furanose, 0.25 ± 0.04%; 108 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


aldehyde, 0.0044 ± 0.0004%; hydrate, 0.022 ± 0.001% (18). The large chemical shift dispersion of the C1 signals makes 13C NMR highly suitable for the detection of cyclic and acyclic forms of reducing saccharides in solution. When the reducing saccharide is a ketose, 13C NMR provides the only reliable means to determine anomeric equilibria, since these molecules lack anomeric hydrogens.

Figure 1. 13C{1H} NMR spectrum of D-[1-13C]mannose (1) in 2H2O, showing the assignment of the labeled C1 signals from the six monomeric forms in solution (α/β-pyranoses, α/β furanoses, and the acyclic aldehyde and hydrate forms). The weak signals between 60–80 ppm arise from the natural abundance C2–C6 carbons in the six forms. Data were taken from ref. (18). A source of error in the measurements shown in Figure 1 is the potential for signal mis-assignment, especially that of the acyclic hydrate form. This problem can be partly addressed by measuring the NMR J-coupling between C1 and its directly attached hydrogen (1JC1,H1) (Table 1). These 1JCH values are sensitive to structure near the C1 carbon, and their values, in addition to C1 chemical shift, can be used to make signal assignments. An example of this application is shown in Figure 2, which shows the C1 carbon signal of the hydrate form of 1 when its directly attached hydrogen (and other hydrogens two- and three-bonds removed from C1) are decoupled or coupled to the C1 carbon. The large splitting (164.2 Hz) is attributed to the one-bond 1JC1,H1. 1JC1,H1 values of 164 – 165 Hz are typically observed for hydrate forms, and 178 – 183 Hz for aldehyde forms (Table 1); significant deviations from these values constitute evidence that the assignment may be incorrect. 109 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


Table 1. C1 Chemical Shifts and 1JC1,H1 Values for the Cyclic and Acyclic Forms of D-[1-13C]Aldopentoses



Table 2.

13C

Chemical Shiftsa of C1 in the Cyclic and Acyclic Forms of D-[1-13C]Aldohexoses in Aqueous Solutionb

Figure 2. Appearance of the C1 signal of the hydrate form of 1 in 13C{1H} NMR spectra obtained with (A) and without (B) broadband 1H-decoupling. The signal in (B) is split by the large 1JC1,H1 (164.2 Hz) characteristic of hydrate forms. In this case, additional splittings are not observed, indicating that 2JC1,H2 and 3JC1,H3 values are probably small or zero. Data were taken from ref. (18).

13C{1H}

NMR spectra such as that shown in Figure 1 provide equilibrium constants for the component equilibria of aldose anomerization through signal integration, provided that the data were acquired under conditions that allow accurate quantitative analysis (18, 19). The results of studies of aldopentoses and aldohexoses are summarized in Tables 1–4, which list the C1 chemical 111 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


shifts of the various forms, 1JC1,H1 values, and percentages in solution, for the cyclic and acyclic forms of aldopentoses and aldohexoses in aqueous solution. A comparison of the percentages of acyclic forms of these aldoses is shown in Figure 3. Aldehydic content ranges from 0.0032 – 0.094% in solution, with solutions of allose and glucose containing the smallest percentages and those of ribose and idose the largest. Hydrate percentages range from 0.0059 – 0.7%, with solutions of allose, glucose and mannose containing very small percentages, and solutions of idose containing the largest (0.7 %; data not shown in Figure 3B).

Figure 3. Percentages of aldehyde (A) and hydrate (B) forms in aqueous solutions of aldopentoses and aldohexoses. Data were taken from Tables 3 and 4. The percentage of hydrate form for idose (0.7%) is not plotted in (B). Al = allose; Ar = arabinose; At = altrose; Ga = galactose; Gl = glucose; Gu = gulose; Id = idose; Ly = lyxose; Ma = mannose; Ri = ribose; Ta = talose; Xy = xylose. Data were taken from refs. (18) and (19).

In some cases, anomerization equilibria include other acyclic forms in addition to the carbonyl and hydrate forms. This behavior is displayed by the biologically important α-ketoacid, N-acetyl-neuraminic acid (2) (Scheme 3). The partial 13C{1H} NMR spectrum of [2-13C]2 at pH 2 and 25 °C is shown in Figure 4 (21). Labeled C2 signals arising from the pyranose forms of 2 appear 112 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


at ~96 ppm. The β-pyranose (2βp) is most preferred (91.2 %), followed by the α-pyranose (2αp) at 5.8% (Scheme 3). The weak signal at ~94 ppm arises from the acyclic hydrate form (2h) (1.9%; Scheme 3). The spectral region between 140 – 200 ppm contains signals arising from the acyclic keto form (2k) (198 ppm; 0.7%; Scheme 3) and, unexpectedly, the acyclic enol form (2e) (143 ppm; 0.5%; Scheme 3). Natural abundance C1 (COOH) signals from 2αp and 2βp also appear in this region (these signals are split by the one-bond 1JC1,C2), as do the signals arising from the amide carbons in both pyranoses.

Figure 4. Partial 13C{1H} NMR spectrum (150 MHz) of [2-13C]2 in 95/5 v/v 1H2O/2H2O at pH 2 and 25 °C. (A) Labeled C2 signals for the 2αp, 2βp and 2h forms (Scheme 3). (B) Carboxyl region showing signals arising from the labeled C2 carbons of 2k and 2e. Unlabeled carboxyl C1 carbons and N-acetyl carbonyl (COam) carbons in 2αp and 2βp appear at ~175 ppm. Data were taken from ref. (21). 113 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


Scheme 3. Anomerization of Neu5Ac (2), and percentages of forms in aqueous solution at pH 2.0. Data were taken from ref. (21).

Solution conditions affect anomerization equilibria, especially temperature and pH. For example, increasing the temperature of aqueous solutions of D-[113C]threose (3) exerts little, if any, effect on the percentages of furanose forms, but the percentages of the acyclic hydrate and aldehyde forms decrease and increase, respectively, with increasing solution temperature (Figure 5) (22).

In contrast, the ketopentose, D-threo-pentulose (D-xylulose) (4), anomerizes to potentially give solutions containing two cyclic ketofuranoses and two acyclic forms (Scheme 4), but the acyclic hydrate form cannot be detected by 13C NMR even when 4 is labeled with 13C at C2 (23). The percentages of the three forms depend on solution temperature as shown in Figure 6. As observed for 3, the percentage of acyclic carbonyl form increases appreciably with increasing temperature, at the expense of the β-ketofuranose. Compared to 3, solutions of 4 contain much more acyclic carbonyl form (2.4% for 3 vs 24% for 4 at 50 °C). 114 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


Figure 5. Percentages of cyclic (A) and acyclic (B) forms of D-[1-13C]threose (3) in aqueous solution (2H2O, 0.1 M tetrose, 50 mM Na-acetate, p2H 5.0) at different temperatures. (A) Filled circles, α-furanose; open circles, β-furanose. (B) Filled squares, aldehyde; open squares, hydrate. The sizes of the symbols provide estimates of the errors in each data point. Data were taken from ref. (22).

The kinetics for each component equilibrium in aldose and ketose anomerization is obtainable from NMR spectra of anomerizing systems at chemical equilibrium. Since the acyclic carbonyl forms of aldoses and ketoses are the presumed obligatory intermediates in the exchange of cyclic forms and the formation of hydrates, selective saturation of the well-resolved carbonyl carbon signals (or aldehydic hydrogens) results in the transfer of saturation to corresponding signals arising from the cyclic and acyclic hydrate forms due to chemical exchange (22, 24). The resulting rate of loss in signal intensity is determined by the ring-opening rate constants, kopen, and the spin-lattice relaxation times of the signals. This application of saturation-transfer NMR spectroscopy (25–27) is illustrated for the anomerization of D-[1-13C]erythrose (5), whose anomerization equilibrium is shown in Scheme 5. Note the significantly higher percentage of acyclic aldehyde and hydrate forms of 5 compared to systems in which pyranosyl rings can form (Tables 3 and 4). For 5, only furanoses form upon ring closure of the acyclic aldehyde. If 5 is enriched with 13C at C1, three 115 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


signals are observed in the anomeric carbon region (Figure 7A): α-furanose (αf), β-furanose (βf) and hydrate (h). The C1 signal of the acyclic aldehyde (not shown) is observed at ~205 ppm. Saturation of the aldehyde C1 signal for increasing amounts of time causes significant loss of signal intensity for C1 of the αf and βf forms (Figure 7, B and C). Linearizing the data (Figure 7D) allows kopen values for cyclic forms, and kdehydration for the hydrate (not shown), to be determined. Determinations of the individual Keq values for the component equilibria in Scheme 5 allow kclose and khydration values to be calculated, thereby providing complete characterization of the anomerization kinetics under a specific set of solution conditions. This method is generally appropriate to measure rate constants in the range 0.05 – 10 s-1; values >10 s-1 are obtained from quantitative treatments of line-broadening in the presence of chemical exchange (Gutowsky-Holm treatment) (27–31).

Scheme 4. Anomerization of D-[2-13C]xylulose (4), showing percentages of forms in solution determined by 13C NMR (0.3 M ketose, 85/15 v/v 1H2O/2H2O, 50 mM Na-acetate buffer, pH 4.0, 26 °C). Data were taken from ref. (23).



Figure 6. Percentages of cyclic and acyclic forms of D-[2-13C]threo-pentulose (D-[2-13C]xylulose) (4) in aqueous solution (see solution conditions in Scheme 4) at different temperatures. (A) β-furanose. (B) open squares, keto; filled circles, α-furanose. The sizes of the symbols provide estimates of the errors in each data point. Data were taken from ref. (23). The effect of phosphate group ionization on the anomerization kinetics of pentose 5-phosphates is shown in Figure 8 for D-[1-13C]ribose 5-phosphate (R5P) (6) (24).

This system is similar to that shown in Scheme 5 for 5 in that only two cyclic furanose and two acyclic forms of R5P are possible in solution. The effect of phosphate differs for both anomers, with the α-furanose more prone to ring-opening than the β-furanose at all solution pH values studied. Saturation-transfer experiments were conducted to measure kopen values at pH 2.3 and 4.0, and line-broadening experiments were conducted to make kopen measurements at the remaining pH values. In general, the presence of phosphate in the saccharide increases anomerization rate constants relative to the same 117 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


molecule devoid of phosphate, suggesting a potential role for intramolecular catalysis in the anomerization of phosphorylated sugars in vivo (24).

Scheme 5. Anomerization of D-erythrose (5), showing percentages of forms in 2H2O solution at 60o determined by 1H NMR. Data were taken from ref. (28).

Table 3. Percentages of Cyclic and Acyclic Forms of D-[1-13C]Aldopentoses in Aqueous Solutiona



Table 4. Percentages of Cyclic and Acyclic Forms of D-[1-13C]Aldohexoses in Aqueous Solutiona

Kinetic studies of anomerizing systems involving pyranosyl rings have also been reported, and data for the aldohexose, D-[1-13C]talose (7), are summarized in Scheme 6. From a thermodynamic perspective, this system is similar to that of D-mannose (1) (Figure 1), with pyranose forms dominating over furanose forms. Under the solution conditions indicated, kopen values range from 0.004 – 0.04 s-1, and kclose values range from 3 – 43 s-1. Interconversions of talopyranoses with the acyclic aldehyde occur more slowly compared to corresponding furanose interconversions. Thus, while talofuranoses are not favored thermodynamically, they are favored kinetically (32).



Figure 7. 13C Saturation transfer experiment conducted on D-[1-13C]erythrose (5) (0.1 M in 2H2O, 50 mM Na-acetate buffer at p2H 5.0) and 55 °C. (A and B) Partial 13C{1H} NMR spectra of 5 showing signals arising from C1 of the α- and β-furanoses (αf, βf) and hydrate (h) forms in the absence (B) and presence (B) of saturation (15 s) at C1 of the acyclic aldehyde form. (C) Plot of signal intensity vs saturation time, showing different rates of decay of the signals for αf (open circles) and βf (closed circles) forms. (D) Semilog plot of the data in (C) for αf, from which a kopen value of 0.40 s-1 is obtained; treatment of the data for βf gives a kopen of 0.19 s-1. Under these solution conditions, the effect of saturation on the C1 signal of the hydrate form is small, and only an upper limit of 80% yield, each containing one more carbon than the starting aldose (chain extension). The C2-epimeric products are purified by chromatography (60). If the hydrogenolysis is conducted with 2H2 gas in 2H2O solvent, the product [1-13C]aldoses will also contain 2H at C1 (Scheme 19) (59, 61). These [1-13C]-labeled aldoses can then be subjected to MCE to transfer the 13C and/or 2H to C2 of the C2-epimeric products. CR and MCE reactions have been effectively integrated into synthetic reaction pathways to provide access a wide range of selectively, multiply and/or uniformly labeled saccharides and their derivatives (e.g., nucleosides) (Scheme 20) (62, 63).

Scheme 19. Introduction of carbon (blue C), hydrogen (red H) and oxygen (green O) isotopes at C1 and/or C2 of aldoses through solvent exchange and cyanohydrin reduction (CR) (see color insert)

Scheme 20. Synthetic routes showing the integration of cyanohydrin reduction (CR) and molybdate-catalyzed epimerization (MCE) in the synthesis of singlyand doubly-13C-labeled aldopentoses and aldohexoses, and 13C-labeled nucleosides, from D-erythrose (see color insert) 144 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


Inspection of the bimolybdate complexes shown in Scheme 18 shows that the space enveloping H1 of the aldose reactant is largely unobstructed, such that replacement with a larger R-group (to give a 2-ketose reactant) should be possible without affecting reactivity. This expectation is realized in practice. Studies show that MCE interconverts 2-ketoses with 2-C-substituted aldoses with high stereospecificity, providing a convenient route to branched-chain aldoses (64–66). Two examples of this application are shown in Scheme 21. Reaction B demonstrates the high tolerance of the reaction to relatively bulky R-groups appended to C2 of the 2-ketose reactant.

Scheme 21. Two reactions (A) and (B) showing the application of molybdate-catalyzed rearrangement (MCE) to interconvert 2-C-substituted D-erythroses with 2-ketoses B. Molybdate-Catalyzed Conversion of Osones to Aldonates The C1–C2 transposition that accompanies MCE can be informally viewed as an internal redox process wherein the oxidation states of C1 and C2 are exchanged during the transformation. This mental construct for the reaction leads to the expectation that aldonates should be produced when 1,2-dicarbonyl sugars such as D-arabino-hexos-2-ulose (D-glucosone) (30) are used as reactants. Recent unpublished work from this laboratory indicates that the reaction of [1-13C]30 with molybdate at 90 °C gives D-[2-13C]gluconate (31) and D-[2-13C]mannonate (32) in a 85/15 ratio (Scheme 22). Osone 30 presumably binds bimolybdate in its dihydrate form to satisfy the hydroxyl group requirements discussed above. By analogy to the complexes that form with aldoses (Scheme 18), two different complexes with 30 are possible. One complex gives D-[2-13C]31, and the other D-[2-13C]32. Unlike the aldose reactions, however, the reaction with 30 is not reversible; the aldonates apparently cannot be converted to the osone, and thus an aldonate cannot be used to generate its C2-epimer. The negatively charged aldonates do not form bimolybdate complexes, presumably because of electrostatic repulsion (both partners are negatively charged). Since the reaction 145 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


is irreversible, the ratio of C2-epimeric aldonates is not determined by their relative stabilities, but rather by the relative stabilities of the two bimolybdate complexes (binding phase) and/or the relative catalytic efficiencies of the two complexes (catalytic phase). The rates of release of aldonate products from their complexes are assumed to be identical. It is interesting to note that, by analogy to the 2-ketose reactants shown in Scheme 21, 2,3-dicarbonyl sugars in their acyclic dihydrate forms should also form productive bimolydate complexes, leading to branched-chain aldonates (Scheme 23). This potential transformation, however, remains to be tested in the laboratory.

Scheme 22. 13C-Labeled products generated from the reaction of D-[1-13C]glucosone (30) with molybdate, observed by 13C NMR

Scheme 23. Hypothetical reaction of a 2,3-dicarbonyl sugar with bimolybdate to give branched-chain aldonates; this transformation remains to be tested in the laboratory.

C. Phosphate-Mediated Conversion of Osones to 2-Ketoses As discussed above, osones are reactive substrates in molybdate-catalyzed reactions where C1–C2 transposition occurs to give a pair of C2-epimeric aldonates. Recent work has shown, however, that this type of transposition in osones is not confined to molybdate-mediated reactions. Prior work has shown that D-glucosone (30) undergoes spontaneous degradation in dilute phosphate buffer at pH 7.4 and 37 °C to give D-ribulose (8) (Scheme 24) (67). Recent NMR studies conducted with D-[2-13C]glucosone (30) confirm this behavior, with unlabeled formate and D-[1-13C]ribulose observed as the major degradation products (68). The reaction pathway presumably 146 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


involves the formation of 2,3-enediol and 1,3-dicarbonyl intermediates, the latter undergoing attack at C1 by OH- with subsequent C1–C2 bond cleavage and protonation to give the 2-ketopentose and formate.

Scheme 24. Degradation of D-[2-13C]glucosone (30) to give D-[1-13C]ribulose (8) and unlabeled formate. The pathway presumably involves 2,3-enediol and 1,3-dicarbonyl sugars as intermediates.

Additional studies of this degradation pathway using other 13C-isotopomers of 30, however, indicated that the pathway shown in Scheme 24 is incomplete, and that, surprisingly, C1–C2 transposition also occurs during degradation. Initial indications of this transposition were found in the reaction shown in Scheme 24 in that a small amount of [13C]formate was observed by 13C NMR in the reaction mixture even though the mechanism shown does not explain its formation. A more definitive experiment was conducted in which D-[1,3-13C2]glucosone (30) was used as the substrate for degradation. Under these reaction conditions, the detection of D-[1,2-13C2]ribulose (33) in the reaction mixture would constitute clear evidence that C1–C2 transposition occurred during degradation. The 13C{1H} NMR spectrum of the products of this reaction is shown in Figure 17. These data show that most of the D-[1,3-13C2]30 degrades as shown in Scheme 24, giving D-[2-13C]8 and H13COO- as the primary end-products. However, closer inspection of the C2 signals arising from D-[2-13C]8 reveals weak satellites on each signal. The upfield region of the spectrum contains the C1 signals arising from each of the three forms of D-[1,2-13C2]8 present in solution (keto and two furanose forms). Each of these signals is split by one-bond 13C-13C J-couplings that are identical to those measured in authentic D-ribulose (23) and to the splittings measured from the C2 satellites (αf, 51.8 Hz; βf = 51.3 Hz; keto, 41.5 Hz). These and other lines of evidence indicate that during the degradation of 30, most of the carbon (~90%) flows down the pathway involving direct C1–C2 bond cleavage to give 8 and formate. However, approximately 10% of 30 undergoes C1–C2 transposition during degradation (Scheme 25). Potential mechanisms for this transposition involve inorganic phosphate as a catalyst in the initial formation of a 1,3-dicarbonyl cyclic phosphate intermediate (Scheme 26) and subsequently as a tether during C1–C2 transposition (68). It is noteworthy that arsenate appears to substitute for Pi in these reactions (68). 147 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


Figure 17. 13C{1H} NMR spectrum of a reaction mixture from the degradation of D-[1,3-13C2]30 (100 mM NaPi, pH 7.5, 37 °C) after 20 days. (A) Full spectrum showing three signals “a” from D-[2-13C]8 (keto and two furanose forms), and H13COO– (signal “b”). (B) The anomeric carbon region of (A) showing the furanose C2 signals of D-[2-13C]8 (signals “a”), the furanose C2 signals from D-[1,2-13C2]8 which appear as satellites on both “a” signals, and unreacted D-[1,3-13C2]30 (signals “b”). (C) Upfield region of (A) showing the C1 signals from D-[1,3-13C2]8 (three signals “a” each split by 1JC1,C2, [2-13C]glycolate (signal b) and an unidentified intermediate (signal “c”). Data were taken from ref. (68).

The preceding discussion serves to illustrate that C1–C2 transposition may be a more common skeletal rearrangement in saccharides than currently appreciated. These rearrangements are remarkable, but their detection requires the use of 13C-labeling in conjunction with NMR and other analytical methods to determine the fates of individual carbons during the reaction. In the original studies of molybdate-catalyzed C2-epimerization of aldoses (54–56), and of glucosone degradation (67), 13C-labeling was not employed, leading to erroneous or incomplete mechanisms for these reactions. It is interesting to note that the transfer of two-carbon fragments is a common occurrence in saccharide metabolism. For example, the coenzyme, thiamine pyrophosphate, promotes reactions catalyzed by the pentose phosphate pathway enzyme, transketolase, wherein the coenzyme functions as a carrier of a negatively charged acylium anion formed from the C1–C2 fragment of a 2-ketose, with the inherently unstable anion resonance-stabilized when covalently attached to the coenzyme (69–71). In principle, this carrier might also enable C1–C2 exchange during the two-carbon exchange as shown in Scheme 27, although, like the glucosone degradation pathway, only a small percentage of the catalytic cycles may follow this pathway. Studies with 13C-labeled substrates would be needed to test this possibility. 148 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.


Scheme 25. Reaction partitioning observed during the degradation of D-[1,3-13C2]glucosone (30) in phosphate buffer (see color insert)

Scheme 26. Proposed formation of phosphate complexes during the degradation of D-glucosone (30), showing its conversion to a phosphorylated 1,3-dicarbonyl intermediate (68) (see color insert)

Scheme 27. A potential (untested) mechanism for C1–C2 transposition in 2-(1,2-dihydroxyethyl)-TPP during two-carbon (acylium anion) transfer catalyzed by transketolases. Formation of the cyclopropanediol intermediate may not be favorable. 149 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.

Molybdenum-catalyzed skeletal rearrangements mimic enzyme-catalyzed reactions in their simplicity and high stereospecificity. Whether enzymes have evolved to exploit the inherent catalytic properties of molybdate in this fashion remains to be determined, as is the potential role of molybdate in chemical evolution. Other elements of the Periodic Table that lie in the vicinity of molybdenum have not shown an ability to catalyze C1–C2 transposition in aldoses. The one element that has not yet been tested is technetium, whose oxides have solution properties similar to those of molybdate (72), but whose rarity and radioactivity thus far have discouraged studies of its reactivity.


Concluding Remarks As discussed in the foregoing paragraphs, studies of the structures and reactivities of saccharides are enabled and/or strengthened when isotopically labeled substrates, especially 13C-labeled, are used to increase the information content of laboratory experiments. We have shown how these isotopes can be used to detect and quantify the cyclic and acyclic forms of reducing saccharides in solution and to investigate relationships between saccharide structure, conformation and the kinetics of tautomer exchange. With the use of 13C-labeled compounds, redundant NMR spin-couplings sensitive to the same molecular torsion angle can be interpreted collectively to derive conformational models of flexible fragments with minimal input from theory. The latter development provides needed experimental validation of conformational predictions derived from computational methods, especially MD simulations. Finally, studies of chemical reactivity using stable isotopes reveal remarkable skeletal rearrangements in saccharides that have defied detection, opening the opportunity to develop new catalysts and/or better understand catalytic mechanisms in chemical and biochemical systems. When Ernest Eliel and his contemporaries founded the field of stereochemistry, they established the fundamental principles of stereochemical analysis and of stereochemical control of chemical reactivity (6–9, 73). In the fifty or so years since Eliel’s pioneering work was conducted, enormous progress in isotope labeling and in analytical methods have provided new opportunities to test these fundamental principles and to extend their applications to increasingly more complex systems, including saccharides. It is safe to say that, fifty years hence, investigators looking back on work now being done will make the same claims, namely, fundamental principles remain so, but new tools and methodologies allow the discovery of new ways to exploit them.

Acknowledgments A.S. is indebted to many talented Notre Dame undergraduates, graduate students, postdocs and visiting scholars who conducted the studies discussed herein over a time period spanning more than thirty years. A.S. would also like to thank the National Institutes of Health and the National Science Foundation for their generous financial support over the same time period, with particular 150 Cheng et al.; Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel Volume 1 ACS Symposium Series; American Chemical Society: Washington, DC.

attribution given to current funding from NSF (CHE 1402744) and to continued material and intellectual support provided by Omicron Biochemicals, Inc. I.C. thanks the Department of Energy Office of Science, Office of Basic Energy Sciences, for financial support of the Notre Dame Radiation Laboratory (NDRL) under award number DE-FC02-04ER15533. This is document number NDRL 5169.

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Stereochemistry and Global Connectivity: The Legacy of Ernest L. Eliel

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