Serianni_Ms_Edited_3_6_17_with_Figures_ASEdits2.docx Printed 3/6/17 1

RESERVE THIS SPACE

RESERVE THIS SPACE

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, IN 46556-5670 USA

*Email: aseriann@nd.edu

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 surprising 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. NMR studies of saccharide chemical transformations using stable isotopes reveal remarkable and stereospecific skeletal rearrangements such as C1–C2 transposition that defied prior detection, opening the opportunity to develop new catalysts and/or to better understand catalytic mechanisms of chemical and biochemical processes involving saccharides.

2

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 that is available for investigation with respect to carbon scaffold, configuration and substitution (1). This playground is particularly appealing because saccharides are biologically important molecules that are 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). 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 expedient 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. Sequentially 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 (3). The impactful 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 (4), 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 (5) that he had worked to develop. Researchers who have subsequently built on the solid foundation provided by Eliel’s seminal studies have benefitted from research tools and methods that

3

were unavailable, and 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 (6), multi-dimensional NMR data collection to resolve and assign complex 1H NMR spectra (7), polarization transfer methods to increase NMR sensitivity and selectivity (8), and routine access to highly enriched and pure stable isotopes such as 13C, 15N and 17,18O on large scales and at reasonable cost (9). A 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 -closing of aldoses and ketoses in solution is known as anomerization (10). This process involves the acyclic aldehydo and keto forms of reducing saccharides (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 (11). In addition to ring-opening and -closing, the acyclic carbonyl forms of aldoses and ketoses can also, in principle, react with solvent water to give acyclic hydrate forms (gem-diols) (Scheme 2). 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 make these determinations (12–14). This application is illustrated in Figure 1, which shows a 13C{1H} NMR spectrum (150 MHz) obtained on an aqueous 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

OHO

HOHO

OH

C1C3

C6

OHD-[1-13C]mannose (1)

! = 13C

!

4

ppm. The intense signals at ~95 ppm arise from the C1 carbons of the dominate aldopyranose forms, while the aldofuranose C1 signals appear slightly downfield of the aldopyranose C1 signals. The very weak signal observed at ~205 ppm arises from C1 of the acyclic aldehyde form, while that at ~91 ppm arises from C1 of the acyclic hydrate form (Tables 1 and 2). Integration of the six signals gives the following percentages of forms in solution at 30 oC: α-pyranose, 66.24 ± 0.05%; β-pyranose, 32.85 ± 0.06%; α-furanose, 0.64 ± 0.04%; β-furanose, 0.25 ± 0.04%; aldehyde, 0.0044 ± 0.0004%; hydrate, 0.022 ± 0.001% (12). 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. A source of error in the measurements shown in Figure 1 is the potential for signal mis-assignment, especially that for the acyclic hydrate form. This problem can be partly addressed by measuring the 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 chemical shift, can be used to make signal assignments. An example of this approach 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 and coupled to the 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. 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 (12,13). The results of studies of aldopentoses and aldohexoses are summarized in Tables 1–4, which list the C1 chemical shifts of the various forms, 1JC1,H1values, and percentages in solution, for the cyclic and acyclic forms of aldopentoses and aldohexoses in solution. A comparison of the percentages of acyclic forms in these aldoses is shown in Figure 3A. 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). In some cases, anomerization equilibria include other acyclic forms in addition to the carbonyl and hydrate forms. This behavior is displayed by the

5

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 oC is shown in Figure 4 (15). Labeled C2 signals arising from the pyranose forms of 2 appear 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 appear as doublets 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. Solution conditions affect anomerization equilibria, especially temperature and pH. For example, increasing the temperature of aqueous solutions of D-[1-13C]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) (16). 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 no acyclic hydrate can be detected by 13C NMR even when 4 is labeled with 13C at C2 (17). 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 oC). 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 (16,18). 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 (19) 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 (Table 1). For 5, only furanoses form upon ring closure of the acyclic aldehyde. If 5 is enriched with 13C at C1,

O

D-threose (3)

OHOH

HO

6

three 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 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, thus providing complete characterization of the anomerization kinetics under a specific set of solution conditions. This method is generally applicable 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) (19c,21). The effect of phosphate group ionization on the anomerization kinetics of pentose phosphates is shown in Figure 8 for D-[1-13C]ribose 5-phosphate (R5P) (6) (18). 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 molecule devoid of phosphate, suggesting a potential role for intramolecular catalysis in the anomerization of phosphorylated sugars in vivo (18). 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 (23).

O

D-ribose 5-phosphate (6)

OH

OHHO

H2O3PO

7

Relationships Between Saccharide Structure and Anomerization Kinetics

Ring opening is rate limiting for the interconversion of cyclic forms of aldoses in solution in most, if not all, cases. This behavior is demonstrated for D-talose (7) in Scheme 6, where kclose values are ~1000-fold larger than kopen values, and the smallest kclose (2.7 s-1) is ~60-fold larger than the largest kopen (0.046 s-1). However, exceptions to this behavior are likely to exist, especially in ketoses, as discussed below. In simple aldofuranoses such as those shown in Scheme 7, the anomer bearing hydroxyl groups in a cis arrangement at C1 and C2 opens more rapidly to the acyclic aldehyde than does the anomer bearing the same groups in a trans arrangement, although relative configuration at C2 and C3 affects the size of the difference (see below). This behavior (hereafter referred to as the “cis-1,2 effect”) is illustrated by the kinetics data for (5) in Figure 7, and in Table 5 where kopen values for D-[1-13C]erythrose (5), D-[1-13C]threose (3), and several C5-modified aldopentoses, measured under identical solution conditions, are compared. The cis-1,2 effect is most apparent in furanose rings having O2 and O3 trans, as found in the threo, arabino and xylo ring configurations. A model explaining this behavior invokes anchimeric assistance by O2 as a facilitator of proton extraction at O1, the latter stimulating ring-opening to the acyclic aldehyde (Scheme 8). In the erythro and ribo rings, the cis-1,2 effect is reduced considerably, however, and in some cases it is abolished (e.g., lyxo rings). Although the data are limited, deoxygenation at C5 appears to increase kopen slightly under the given solution conditions (H2O-catalyzed region) (10b). The effect of C4 substitution on kopen appears small; for example, kopen values for erythrose (5) and threose (3) range from 0.2 – 0.7 s-1, whereas those for the C5-modified aldopentoses range from 0.1 – 0.5 s-1, under the solution conditions given. The cis-1,2 mechanism shown in Scheme 8 may not be the only potential structural explanation for the observed differences in kopen between furanose anomers. Arguing from the principle of microscopic reversibility, furanose ring conformation may also influence kopen, with some conformations more prone to ring opening than others. The enhanced kopen values observed in the threo, arabino and xylo ring configurations might result from preferred ring conformations that also favor ring opening. This argument evolves from ring-closure arguments where trajectories of hydroxyl oxygen attack on the carbonyl carbon are highly constrained (25), thus leading to a small subset of ring conformations as the immediate products of closure. This

O

D-erythro-pentulose (8)(D-ribulose)

OH

OHHO

CH2OH

8

same subset of conformations would then be favored for ring opening, but pseudorotation may not favor them at equilibrium. When favored ring conformations in solution coincide with those favored for ring opening, kopen will be enhanced. Rates of ring pseudorotation relative to kopen will affect the potency of this factor. The effect of converting aldoses to ketoses on anomerization kinetics can be seen by comparing D-erythrose (5) and D-threose (3) with D-[2-13C]threo-pentulose (D-xylulose) (4) and D-[2-13C]erythro-pentulose (D-ribulose) (8) (Table 6). Ketopentoses 4 and 8 are essentially alkylated derivatives of aldotetroses 3 and 5, respectively, in which CH2OH groups replace the anomeric hydrogens in the aldoses. This type of ring alkylation affects kopen only to a small extent; for example, the average kopen value for 3 and 5 is 0.45 ± 0.17 s-1, whereas that for 4 and 8 is 0.23 ± 0.08 s-1 (Table 6). However, the effect of this alkylation on kclose is significant, with the aldoses ~30 times more reactive than the ketoses (average kclose of 0.34 ± 0.27 s-1 for the ketoses vs 10.8 ± 3.7 s-1 for the aldoses). The reduced rate of ring closure in the ketoses is likely due to the greater steric demands of the keto group relative to the aldehyde group and/or to the greater electrophilicity of the aldehyde. The former factor imposes significant constraints on the reaction trajectory required for productive ring closure, and thus the conformational dynamics of the acyclic keto form may affect kclose. The practical implications of the data in Table 6 are that, under identical solution conditions, ketoses anomerize more slowly than aldoses having related structures, largely because of reduced rates of ring closure. The cis-1,2 effect is also observed in ketose 4 in which O3 and O4 are trans, with ring-opening occurring more rapidly for the β-furanose than for the α-furanose (Table 6). As observed in the some aldoses (Table 5), this effect is essentially abolished in ketose 8 in which O3 and O4 are cis. Anomerization rates are affected by solution pH, with acid-catalyzed, H2O-catalyzed, and base-catalyzed regions (10b). The role of acid catalysis is apparent in kopen values for 5-deoxy-L-lyxose and 5-O-methyl-D-lyxose measured at pH 1.4 – 2.5 (24). Under acidic solution conditions, the rate constant for acid catalysis, kH3O+, is determined from eq. [1], kobs = kH2O + kH3O+ [H3O+] eq [1] where kobs is the observed rate constant and kH2O is the rate constant for the water-catalyzed reaction. A plot of kobs vs [H3O+] gives a line with slope equal to kH3O+. These plots are shown in Figure 9 for the four anomers, from which the following kH3O+ values for ring-opening were determined: for 5-deoxy-L-lyxose, 79 ± 3 s-1M-1 (α) and 184 ± 6 s-1M-1 (β); for 5-O-methyl-D-lyxose, 27 ±

9

2 s-1M-1 (α) and 69 ± 4 s-1M-1 (β). In both aldopentoses, kH3O+ values are ~2-fold larger for the β-anomer, that is, unlike behavior under water-catalyzed conditions (Table 5), the cis-1,2 effect is evident in acidic solution, attesting to the key role that solution conditions play in determining the relative reactivity of anomers. In addition, kH3O+ values are ~ 3-fold larger in the 5-deoxy derivative, consistent with a -CH3 substituent being less electron-withdrawing than -CH2OCH3, thereby promoting protonation at either O1 or O4 during catalysis.

The effects of furanose ring deoxygenation and alkylation on the percentages of acyclic aldehyde form in aqueous solutions of aldofuranoses are illustrated by the data shown in Table 7. Deoxygenation at C2 increases the percentage of

acyclic aldehyde in solution (3 vs 9; 14 vs 15), whereas deoxygenation at C3 exerts only a minor effect (3 vs 10). Ring alkylation at C3 and C5 reduces the percentage of acyclic aldehyde form appreciably (3 vs 11–14). The latter shift towards cyclic forms is a manifestation of the Thorpe-Ingold effect (27–29) (gem-dialkyl effect) that has been attributed to enthalpic and entropic factors. The ratio, [hydrate]/[aldehyde], varies between 0.4 – 10 within the group of compounds shown in Table 7; this ratio is influenced by steric factors in the acyclic hydrates, which vary with substitution pattern. For example, the very low percentage of hydrate form in solutions of 13 is caused, in part, by steric interactions between the two OH groups at C1 and the two CH3 groups at C3. The latter interactions are partly relieved in 11 and 12, and largely eliminated in 10, resulting in progressively higher percentages of hydrate in solution.

O

D-arabinose 5-phosphate (16)

OHOH

HO

H2O3PO O

D-lyxose 5-phosphate (17)

OHOHHOH2O3PO O

D-xylose 5-phosphate (18)

OH

OH

HOH2O3PO

O OH

OH

O OH

OHHO

O OH

HO

O OH

OHHO

H3COH2C

O OHOH

HO

H3CH3C

O OH

OHH3C

H3C O OH

HO

H3COH2C

2-deoxy-D-glycero-tetrose (9)

3-deoxy-DL-glycero-tetrose (10)

3-C-methyl-DL-erythrose (11)

3-C-methyl-DL-threose (12)

3-deoxy-3,3-di-C-methyl-DL-glycero-tetrose (13)

5-O-methyl-D-ribose (14) 2-deoxy-5-O-methyl-D-erythro-pentose (15)

10

The effects of furanose ring deoxygenation and alkylation on anomerization kinetics are illustrated by the data in Table 8. The cis-1,2 effect (Scheme 8) on kopen is maintained in 10–12, with 12 showing the greatest effect as expected, since O2 and O3 in 12 are trans. Interestingly, kopen for the α- and β-furanoses of 15 are essentially identical, whereas those of 9 differ, with the a-furanose opening more rapidly than the β-furanose. This behavior may be attributed to different preferred ring conformations of the α-furanoses of 9 and 15, with the former favoring 2E and the latter 3E based on 3JHH analysis (Scheme 9) (20). The former ring conformation orients both O1 and O3 quasi-axially, which presumably allows anchimeric assistance as shown in Scheme 8. In contrast, the preferred ring conformation of 15 orients O1 and O3 quasi-equatorially, thus disallowing anchimeric assistance and rendering similar kopen values for both anomers. The apiofuranoses behave like 11 and 12 with respect to relative values of kopen (30). Furanose ring alkylation enhances kclose values significantly; for example, kclose values range from 1.6 – 6.2 s-1 for 3 and 5, but values from 8.5 – 39.2 s-1 are observed for 11–13. This behavior is a further manifestation of the Thorpe-Ingold effect in saccharides (27–29). Ring-opening rates constants for the pentose 5-phosphates (Table 9) depend on ring configuration, with ribo (6) showing the greatest reactivity. The effect of pH on kopen is significant, with ~100-fold increases observed as pH is raised from 4.2 (mono-anion) to pH 7.5 (di-anion). A comparison of kopen values for the four 5-O-methyl aldopentoses in Table 5, obtained at pH 4.0 and 60 oC, to those found for the four pentose phosphates in Table 9, obtained at pH 4.2 and 40 oC, provides indirect evidence for catalysis by phosphate. The

average value of the eight kopen values in Table 5 is 0.21 ± 0.11 s-1 compared to the average of 0.44 ± 0.23 s-1 for the pentose phosphates, despite the 20o lower temperature used in the pentose phosphate measurements. Unlike neutral (uncharged) furanoses, the cis-1,2 effect (Scheme 8) is not observed in pentose phosphates; the α-anomer gives the larger kopen at pH 4.2 and 7.5 in all ring configurations except lyxo (17). In

17, kopen values for both anomers are essentially the same at either pH value. A possible mechanism that attempts to explain these observations invokes the phosphate group as a source of protonation of the ring oxygen in α-anomers, either directly (Scheme 10), or indirectly via H-bonding to a participating water

O

D-arabinuronicacid (19)

OHOH

HO

HOOC O

D-lyxuronicacid (20)

OHOHHOHOOC

O

D-xyluronicacid (22)

OH

OH

HOHOOCO

D-riburonicacid (21)

OH

OHHO

HOOC

11

molecule. Steric hindrance between the cis-oriented phosphate group and O1 in β-anomers presumably weakens this mode of catalysis. Studies of kclose in the pentose phosphates show that D-ribose 5P (6) is the most reactive, followed by 16 and 17/18 (18). Therefore, with respect to kopen and kclose, the ribo ring (6) is the most reactive pentose phosphate. This enhanced reactivity may have played a role in its evolutionary selection as a key sugar phosphate in biological metabolism (18). The different ring-opening behaviors of pentose phosphates compared to neutral furanoses stimulated interest in the anomerization kinetics of penturonic acids 19–22 (Table 10) (31). At pH 1.5, where the carboxyl group is mostly protonated, the cis-1,2 effect (Scheme 8) is observed, that is, the relative values of kopen mimic those found in neutral furanoses. A comparison of kopen values for the four 5-O-methyl aldopentoses in Table 5, obtained at pH 4.0 and 60 oC, to those found for the four penturonic acids in Table 10, obtained at pH 4.5 and 50 oC, provides indirect evidence for catalysis by the COOH group. The average value of the eight kopen values in Table 5 is 0.21 ± 0.11 s-1 compared to 1.38 ± 0.83 s-1 for the penturonic acids, despite the 10o lower temperature used in the penturonic acid measurements (the latter average could be enhanced somewhat by the slightly higher pH used for these measurements). At pH 1.5, intramolecular catalysis by the protonated carboxyl group and intermolecular catalysis by H+ enhance kopen values relative to those measured in neutral furanoses. A potential role for the COOH group in intramolecular catalysis involves protonation of O4 (Scheme 11, A). The effects of this catalysis may be offset by the electron withdrawing character of the COOH group. The latter factor would render O4 less prone to protonation and decrease kopen. In contrast, kopen values for 19–22 at pH 4.5, where the carboxyl group is largely ionized, favor β-anomers (Table 10), that is, anomers in which O1 and the COO- group are cis. While anomerization at pH 4.5 is largely water-catalyzed for neutral furanoses, kopen values may be enhanced in 19–22 by an intramolecular mechanism involving deprotonation of O1 (Scheme 11, B). The electron-donating property of the COO- group (relative to COOH) may also promote protonation of the ring oxygen.

12

N-Acetyl Side-Chain Cis-Trans Isomerization in Aminosugars and Conformational Properties of O-Acetyl Side-chains

Monosaccharides contain different types of substituents and side-chains that influence their chemical and biological properties (32). For example, the rotation of C–O bonds involving hydroxyl groups influences C–H and C–C bond lengths, which in turn influence the magnitudes and signs of NMR spin-spin (scalar; J-coupling) coupling constants such as JHH, JCH and JCC (33). These J-couplings, which are both abundant and redundant (i.e., multiple values report on the same conformational element) are valuable structural constraints to determine the conformational and dynamics properties of saccharides in solution (34–36). Examples of exocyclic groups such as hydroxymethyl (–CH2OH), O-acetyl (–O–CO–CH3), N-acetyl (–NH–CO–CH3), O-phosphate monoesters (–O– PO3-2), O-sulfate monoesters (–O–SO3-1), N-sulfamides (–NH–SO3-1), O-lactoyl (-OOC–CHOR–CH3), and C-glyceryl (–CHOH–CHOH–CH2OH) are shown in Scheme 12 (32). Here we limit discussion to N-acetyl and O-acetyl side-chains to illustrate recent work that aims to better understand their structural properties. A. N-Formyl and N-Acetyl Side-chains. The conformational behaviors of N-acyl groups in saccharides are characterized by two factors: (1) rotation about the Cx–NH bond θ1 that attaches the group to Cx of the saccharide, and (2) cis-trans isomerization of the amide bond θ2 in the acyl substituent (Scheme 13). Signals arising from the cis and trans forms of the N-formyl and N-acetyl groups in methyl 2-[13C]formamido-2-deoxy-D-glucopyranosides (23) (α) and (24) (β) and 2-[1-13C]acetamido-2-deoxy-D-[2-13C]glucopyranosides (25) (α) and (26) (β), respectively, can be observed by 1H and/or 13C NMR, with signal

assignments facilitated by selective 13C-enrichment at C2 of the saccharide and/or the carbonyl carbon of the side-chain (37). For example, the 1H NMR spectrum of methyl 2-

[13C]formamido-2-deoxy-β-D-glucopyranoside (24) at 600 MHz contains two sets of formyl hydrogen signals at ~8.15 ppm (trans) and ~7.93 ppm (cis), and two sets of anomeric hydrogen signals at 4.45 ppm (trans) and 4.42 ppm (cis)

(Figure 10). The former are split by the one-bond 1JCH of 195.6 Hz (cis) and 197.7 Hz (trans) that, along with their characteristic 1H chemical shifts, confirms their identity. In 24, ~74% of the amide bond exists in the trans

OOCH3

OH

HOHO

O

OCH3

OH

HOHO

NH

CCH3

ONH

CCH3

O

OOCH3

OH

HOHO

O

OCH3

OH

HOHO

NH

C

H

ONH

C

H

O23 24

25 26

! !

!

! !

!

"

"

#

#

! = 13C

13

configuration and ~26% in the cis configuration at 42 oC (Ktrans/cis = 2.84 ± 0.02) in aqueous (2H2O) solution (37). By comparison, ~81% trans and ~19% cis are observed in 23 (Ktrans/cis = 4.22 ± 0.03), showing that anomeric configuration affects the trans/cis ratio. At temperatures of 42 – 85 oC, aqueous solutions of 24 consistently contain more cis isomer than do those of 23 (Ktrans/cis ranges from 3.37 – 4.22 for 23, while Ktrans/cis ranges from 2.41 – 2.84 for 24), with increasing temperature increasing the percentage of cis form in solution in both anomers (Figure 11A). The detection of cis forms in aqueous solutions of 25 and 26 is challenging because significantly less of this form is present compared to aqueous solutions of 23 and 24. By incorporating selective 13C-enrichment (~99 atom-% 13C) at both C2 and the carbonyl carbon of the side-chain, not only is the detection of the labeled C2 and CO signals enhanced by ~100 fold, but J-coupling between the labeled carbons can be measured and used to confirm the assignments (assuming a non-zero value of the J-coupling). Note that 25 and 26 lack a characteristic 1H signal in the side-chain, unlike 23 and 24, thus favoring 13C NMR as the method of analysis. The carbonyl carbon region of the 13C{1H} NMR spectrum of 26 at 22 oC contains a strong (labeled) signal at ~177 ppm and a weak (labeled) signal at ~180 ppm (Figure 12A) (37). Both signals are split into doublets by 1.0 Hz and 0.8 Hz, respectively. The C2 region of the same spectrum contains a strong (labeled) signal at ~58 ppm and a weak (labeled) signal at ~63 ppm, and both are split into doublets by 1.0 Hz and 0.8 Hz, respectively (Figure 12B). The fact that identical signal splittings are observed for the paired weak CO and C2 signals, and the paired strong CO and C2 signals, confirms their assignments to the cis and trans forms of 25, respectively, with the splitting attributed to 2JC2,CO. Ktrans/cis values for 25 range from 45.3 – 72.5 over the temperature range 42 – 75 oC, and values of 31.1 – 45.3 were observed for 26 over the same temperature range (Figure 11B) (37). Thus, while significantly less cis form is observed in aqueous solutions of 25 and 26 compared to 23 and 24 (the latter have Ktrans/cis values of 2–4; see above), solutions of β-anomer 26 consistently contain more cis isomer than do those of α-anomer 25 at any given temperature, thus mimicking the behavior of the N-formyl compounds 23 and 24. van’t Hoff plots of the data shown in Figure 11 give slightly negative values for ΔHo (~ –1 – –3 kcal/mol) and ΔSo (–0.6 – –1.6 cal/K/mol) for the conversion, cis amide → trans amide, showing that the process is enthalpically favored but entropically disfavored. Conformation about θ1 in 23–26 is believed to favor structures in which H2 and the NH hydrogen are antiperiplanar (θ1 = 180o) based on the magnitudes of 3JH2,NH values (Scheme 14) (38,39). However, geometries in which H2 and NH are eclipsed (θ1 = 0o) give 3JH2,NH values similar in magnitude to those in

14

anti geometries (Figure 13) (40), so conformational conclusions on θ1 based solely on a single 3JH2,NH are not unequivocal. Efforts to provide more definitive assessments of θ1 await the application of redundant J-couplings such as 3JC1,NH, 3JC3,NH, 3JH2,CO, 3JC1,CO and 3JC3,CO that have been parameterized recently using density functional theory (DFT) (40). The observation of distinct signals arising from the cis and trans forms of 23–26 implies slow exchange on the NMR time-scale, rendering the system amenable to study by saturation-transfer (19) to measure the first-order rate constants, kcis→trans and ktrans→cis. As described above in studies of anomerization kinetics (Figure 7), selective 13C saturation of the carbonyl signal of the cis form gives ktrans→cis while saturation of the carbonyl signal of the trans form gives kcis→trans; saturation of the C2 signals could also be performed since they, like the carbonyl carbons, are reasonably well resolved at 150 MHz (Figure 12) (37). This application is shown in Figure 14 for 2-[1-13C]acetamido-2-deoxy-α-D-[2-13C]glucopyranoside (25), which shows the loss in carbonyl carbon signal intensity of the trans form with increasing saturation time of the carbonyl carbon in the cis form at different temperatures. Linearizing the data, as discussed in Figure 7, gives ktrans→cis values at each temperature. Side-chain cis-trans isomerization (CTI) rate constants in 23–26 determined by this method are summarized in Figure 15. Data show that CTI kinetics depends on anomeric configuration, with β-anomers more kinetically favored than α-anomers in both the N-formyl and N-acetyl compounds. Within the series 23–26, βGlcNAc structure 26 is most reactive, exhibiting the largest kcis→trans and ktrans→cis values at any given temperature. Energies of activation range from 16–19 and 19–20 kcal/mol for the cis→trans and trans→cis reactions, respectively (37). The biological implications of side-chain CTI remain to be established, but these data show that CTI equilibria and kinetics are both affected by side-chain structure (N-formyl vs N-acetyl) and anomeric configuration when the N-acyl side-chain is an equatorial orientation and when appended to C2 of an aldohexopyranosyl ring. It remains to be determined how these behaviors are influenced by other local environments and structural contexts (e.g., site of substitution; axial vs equatorial orientation; presence in larger oligosaccharides), and whether CTI influences the binding of N-acylated saccharide substrates to biological receptors. B. O-Acetyl Side-Chains. O-Acetyl side-chains resemble N-acetyl side-chains in that analogous θ1 and θ2 torsion angles characterize their conformations (Scheme 15). Recent work has shown (35) that the conformational behavior of θ1 can be investigated using redundant NMR J-couplings and circular statistics. For example, in the mono-O-acetylated compounds 27–29, selective 13C-labeling incorporated into the O-acetyl side-

15

chain allows the measurement of two or three JCC values (for example, in 28: 3JC2,CO, 3JC4,CO, 2JC3,CO) and simplifies the measurement of one or two JCH values (for example, in 28: 3JH3,CO). These J-couplings are sensitive to θ1, and DFT calculations (41) on model structures give parameterized equations that relate each to θ1. Pertinent equations derived for the 3-O-acetylated compounds 28α/β take the following forms (35): 3JH3,CO = 3.92 – 2.11 cos (θ) + 0.05 sin (θ) + 3.34 cos (2θ) + 0.07 sin (2θ) – 0.04 cos (3θ) + 0.05 sin (3θ) rms 0.22 Hz eq. [2] 3JC2,CO = 1.56 + 0.36 cos (θ) – 0.73 sin (θ) – 0.75 cos (2θ) – 1.22 sin (2θ) – 0.14 cos (3θ) + 0.02 sin (3θ) rms 0.13 Hz eq. [3] 3JC4,CO = 1.44 + 0.33 cos (θ) – 0.74 sin (θ) – 0.68 cos (2θ) – 1.21 sin (2θ) – 0.14 cos (3θ) + 0.16 sin (3θ) rms 0.04 Hz eq. [4] 2JC3,CO = –3.81 + 0.64 cos (θ) – 0.14 sin (θ) + 0.84 cos (2θ) + 0.05 sin (2θ) – 0.29 cos (3θ) – 0.03 sin (3θ) rms 0.10 Hz eq. [5] Similar equations were derived for compounds 27α/β and 29α/β (35). These equations and the experimental J-couplings measured from 1H and 13C{1H} NMR spectra were treated with a circular statistics package, MA’AT, in which different 2-parameter continuous circular probability distributions were used to model θ1 in 27–29. This modeling gave mean values of θ1 and circular standard deviations (CSD) for each compound. The mean identifies the most abundant θ1 torsion angle in aqueous solution, and the CSD provides a measure of the librational disorder about the mean, analogous to the order parameter S2 derived from NMR spin-relaxation measurements (42). This treatment is possible because four different J-couplings are available that display different functional dependencies on θ1 (redundancy). The results of this data analysis are shown in Figure 16. The position of the O-acetyl side-chain on the aldohexopyranosyl ring affects the mean value of θ1 and the CSD. The preferred conformation of the O-acetyl side-chain in 28α/β is shown in Scheme 16. In the trans configuration of the ester (θ2) shown, the C=O bond of the ester eclipses the C3–H3 bond in the preferred geometry about θ1. This situation contrasts with the behavior of θ1 in 29α/β, where the carbonyl carbon of the side-chain is roughly gauche to both H6R and H6S, and the C=O bond bisects the H6R–C6–H6S bond angle in the trans configuration of the ester (Scheme 17). The CSD is appreciably greater for 29α/β than for 28α/β, which indicates greater disorder about θ1 for O-acetyl groups appended to the primary alcoholic carbon. The

16

results of aqueous molecular dynamics simulations of 27–29 are in good agreement with these experimental models (35), which testifies to the reliability of the method used to fit the redundant J-couplings and provides important experimental validation of the MD results. Future applications of this type of J-coupling analysis are anticipated in structural studies of other types of saccharide side-chains, saccharide rings, and O-glycosidic linkages in oligosaccharides.

Stereospecific Carbon-Skeleton Rearrangements in Saccharides: C1–C2 Transposition Reactions

A. Molybdate-Catalyzed C2-Epimerization of Aldoses. In 1973, Bilik and coworkers reported that sodium molybdate catalyzes the C2-epimerization of aldoses in aqueous solution (43), and proposed a mechanism involving hydrogen shift to explain the reaction stereochemistry (44). However, unexpectedly, when the reaction was conducted with [1-13C]aldoses as reactants, it was found that C1–C2 transposition accompanies C2-epimerization (45). This remarkable

skeletal rearrangement is believed to involve initial complexation of bimolybdate with the hydrate (1,1-gem-diol) form of the aldose (Scheme 18). The reaction apparently occurs in two steps described by eqs. [6] and [7]. In the first step, monomeric molybdate dimerizes to form the bimolybdate species, Mo2O7H2. The latter subsequently binds with the aldose acyclic hydrate to give a negatively charged bimolybdate-aldose complex. This complex is catalytically active, yielding a putative transition state containing partial covalent bonds between C1–C2, C2–C3 and C1–C3. The ratio of starting aldose and its C2-epimeric product observed after equilibration is determined by their relative thermodynamic stabilities, since the reaction is freely reversible. In most cases, equilibration is reached in ~3 h at ~85 oC, and few if any by-products are observed. Studies have shown that hydroxyl groups at C1, C2 and C3 of the aldose reactant are required for molybdate-catalyzed C2-epimerization (MCE), while an OH group at C4 is not required but increases the reaction rate and reduces the formation of by-products (45). From a practical standpoint, MCE provides a powerful complement to cyanohydrin reduction (CR) reactions in the synthesis of stable isotopically labeled saccharides (Scheme 19) (46). Stable isotopes (13C, 2H) are introduced at C1 of aldoses by reacting K13CN (a relatively cheap, commercially available labeled precursor) with a starting aldose electrophile under solution conditions that stabilize the initially formed α-hydroxynitriles (cyanohydrins). The latter are hydrogenolyzed using a heterogeneous metal

2MoO3 Mo2O7H2 (bimolybdate)+ H2O

Mo2O7H2 + C6H14O7 [Mo2O5 . C6H10O7]-2 + 2H3O+

eq. [6]eq. [7]

17

catalyst (typically Pd/BaSO4) and H2 gas to give a pair of C2-epimeric [1-13C]aldoses in >80% yield, each containing one more carbon than the starting aldose (chain extension). The C2-epimeric products are purified by chromatography (47). If the hydrogenolysis is conducted with 2H2 gas in 2H2O solvent, the product [1-13C]aldoses will also contain 2H at C1 (Scheme 19) (46b,48). 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) (49). 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 (50–52). 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. 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 oC gives D-[2-13C]gluconate (31) and D-[2-13C]mannonate (32) in a 85/15 ratio (Scheme 22). Ozone 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 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 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-

18

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. 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 oC to give D-ribulose (33) (Scheme 24) (53). 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 (54). The reaction pathway presumably 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. 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]33 and H13COO- as the primary end-products. However, closer inspection of the C2 signals arising from D-[2-13C]33 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]33 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 (17) 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 33 and formate. However, approximately 10% of 30 undergoes C1–C1 transposition during degradation (Scheme 25). Potential mechanisms for this transposition involve inorganic phosphate as a catalyst in the initial formation of

19

a 1,3-dicarbonyl cyclic phosphate intermediate (Scheme 26) and subsequently as a tether during C1–C2 transposition (54). It is noteworthy that arsenate also appears to substitute for Pi in these reactions (54). 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 trace the fates of individual carbons during the reaction. In the original studies of molybdate-catalyzed C2-epimerization of aldoses (43,44), and of glucosone degradation (53), 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, 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 (55–57). 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. 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 Mo 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 (58), 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

20

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 to 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 the stereochemical control of chemical reactivity (4, 5, 59). 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.

Acknowledgements 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 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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2016, 59, 673–679. (b) Cyanohydrin reduction (CR) and molybdate-catalyzed epimerization (MCE) have found application in the commercial synthesis of isotopically labeled saccharides and their derivatives. See: www.omicronbio.com.

50. Hricovíniová-Bíliková, Z.; Hricovíni, M.; Petrusová, M.; Serianni, A. S.; Petrus, L. Carbohydr. Res. 1999, 319, 38–46.

51. Zhao, S.; Petrus, L.; Serianni, A. S. Org. Lett. 2001, 3, 3819–3822. 52. Wu, Q.; Pan, Q.; Zhao, S.; Imker, H.; Serianni, A. S. J. Org. Chem. 2007,

72, 3081–3084. 53. Wells-Knecht, K. J.; Zyzak, D. V.; Litchfield, J. E.; Thorpe, S. R.; Baynes,

J. W. Biochemistry 1995, 34, 3702–3709. 54. Zhang, W.; Serianni, A. S. J. Am. Chem. Soc. 2012, 134, 11511–11524. 55. Datta, A. G.; Racker, E. J. Biol. Chem. 1961, 236, 624–628. 56. Kochetov, G. A.; Solovjeva, O. N. Biochim. Biophys. Acta 2014, 1844,

1608–1618. 57. Kleijn, R. J.; van Winden, W. A.; van Gulik, W. M.; Heijnen, J. J. FEBS J.

2005, 272, 4970–4982. 58. Muller, O.; White, W. B.; Roy, R. J. Inorg. Nucl. Chem. 1964, 26, 2075–

2086. 59. Seeman, J. I. Chirality 2002, 14, 98–109.

24

Scheme 1. Hierarchies of molecular structure

atomic composition (molecular mass)

covalent bonding

configuration at chiral centers

conformational equilbria

conformational dynamics

intermolecular interactions(solvation, molecular recognition)

incr

easi

ng c

ompl

exity

O O

O O

acyclic aldehyde

!-pyranose

!-furanose "-furanose

acyclic hydrate

"-pyranose

CHO

OHOH

OH

OH

OH

CH(OH)2

OH

+H2O-H2O

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

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. 12.

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

a± 0.1 ppm, referenced (external) to the C1 chemical shift of !-D-[1-13C]mannopyranose (95.0 ppm); 1 M solutions of D-[1-13C]pentose in 2H2O at 28 oC. b±0.1 Hz. cPart of signal obscured; J-value could not be measured. Data were taken from ref. 13.

D-aldopentose NMR parameter "C1 (ppm)a 1JC1,H1 (Hz)b

D-arabinose !-furanose 102.6 171.8 #-furanose 96.7 174.2 !-pyranose 98.3 160.6 #-pyranose 94.1 169.0 hydrate 91.6 164.6 aldehyde 207.0 178.2 D-lyxose !-furanose 102.3 171.4 #-furanose 97.0 obscc !-pyranose 95.6 167.4 #-pyranose 95.8 161.8 hydrate 91.1 164.2 aldehyde 207.2 182.9 D-ribose !-furanose 97.8 172.6 #-furanose 102.5 173.0 !-pyranose 95.1 164.6 #-pyranose 95.4 165.4 hydrate 91.0 164.6 aldehyde 204.6 183.4 D-xylose !-furanose 97.1 obscc #-furanose 103.2 171.8 !-pyranose 93.8 169.4 #-pyranose 98.2 161.4 hydrate 91.3 163.8 aldehyde 206.0 179.0

25

Table 2. 13C Chemical Shiftsa of C1 in the Cyclic and Acyclic Forms of D-[1-13C]Aldohexoses in Aqueous Solutionb

a±0.1 ppm; referenced (external) to the C1 chemical shift of !-D-[1-13C]mannopyranose (95.0 ppm). b2 M solutions of D-[1-13C]aldohexose in 2H2O at 30 oC. c!f = !-furanose, "f = "-furanose; !p = !-pyranose; "p = "-pyranose. Data were taken from ref. 12.

hexose C1 chemical shift (ppm)c !f "f !p "p hydrate aldehyde

D-allose 96.8 101.6 93.7 94.3 90.4 207.1 D-altrose 102.2 96.2 94.7 92.6 90.8 206.2

D-galactose 95.8 101.8 93.2 97.3 91.1 206.4 D-glucose 97.6 103.2 92.9 96.7 90.3 205.9 D-gulose 96.0 101.4 93.7 94.7 90.3 206.3 D-idose 102.5 96.4 94.0 93.2 90.6 205.3

D-mannose 101.9 96.5 95.0 94.6 90.6 205.4 D-talose 101.8 97.3 95.5 95.0 91.0 204.3

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. 12 .

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

a1 M D-[1-13C]pentose in 2H2O at 28 oC. Two 13C{1H} NMR spectra, acquired on the same sample on separate occasions, were processed with 3 Hz and 5 Hz line-broadening functions, giving four spectra from which four percentages of each form were determined by signal integration. The average of the four percentages is shown for each form in the table. Standard deviations are given in parentheses. Data were taken from ref. 13.

pentose percent in solution !f "f !p "p hydrate aldehyde

D-arabinose 5.6 (0.0)

3.5 (0.1)

58.7 (0.2)

32.2 (0.2)

0.094 (0.003)

0.015 (0.002)

D-lyxose 1.7 (0.0)

0.60 (0.01)

70.8 (0.4)

26.9 (0.4)

0.086 (0.006)

0.011 (0.000)

D-ribose 7.4 (0.1)

13.2 (0.3)

20.2 (0.3)

59.1 (0.2)

0.088 (0.002)

0.042 (0.002)

D-xylose 0.86 (0.06)

0.69 (0.07)

36.5 (0.3)

62.0 (0.4)

0.062 (0.006)

0.009 (0.002)

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

a2 M D-[1-13C]hexose in 2H2O at 30 oC. Three 13C{1H} NMR spectra, acquired on the same sample on separate occasions, were processed, giving three spectra from which three percentages of each form were determined by signal integration. The average of the three percentages is shown for each form in the table. Standard deviations are given in parentheses. Data were taken from ref. 12.

hexose percent in solution !f "f !p "p hydrate aldehyde

D-allose 2.99 (0.04)

5.30 (0.04)

14.6 (0.1)

77.07 (0.04)

0.0063 (0.0001)

0.0032 (0.0009)

D-altrose 18.60 (0.04)

13.37 (0.03)

26.9 (0.2)

41.0 (0.1)

0.079 (0.002)

0.014 (0.001)

D-galactose 2.30 (0.05)

3.69 (0.05)

31.2 (0.3)

62.8 (0.2)

0.046 (0.006)

0.006 (0.003)

D-glucose 0.11 (0.01)

0.28 (0.03)

37.63 (0.05)

61.96 (0.03)

0.0059 (0.0004)

0.0040 (0.0006)

D-gulose 0.94 (0.05)

3.04 (0.03)

12.2 (0.1)

83.7 (0.2)

0.077 (0.005)

0.006 (0.002)

D-idose 12.14 (0.04)

16.12 (0.04)

33.7 (0.3)

37.4 (0.2)

0.7 (0.2)

0.094 (0.004)

D-mannose 0.64 (0.04)

0.25 (0.04)

66.24 (0.05)

32.85 (0.06)

0.022 (0.001)

0.0044 (0.0004)

D-talose 17.9 (0.3)

11.1 (0.2)

42.2 (0.4)

28.7 (0.2)

0.052 (0.002)

0.029 (0.003)

26

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. 12 and 13.

O

OH

COOHH3CCOHN

HO

O OH

COOH

H3CCOHNHO

!-pyranose (2!p) "-pyranose (2"p)

RR

OH

OH

OHOH

H HO

HOH3CCOHN

COOH

OH

OH

OHOH

H HOH

HOH3CCOHN

COOH

HO

OH

OH

OHOH

OH

HOH3CCOHN

COOH

keto (2k)

keto hydrate (2h)(gem-diol)enol (2e)

Scheme 3. Anomerization of Neu5Ac (2), and percentages of forms in aqueous solution at pH 2.0

R = -CHOH-CHOH-CH2OH

(5.8 %) (91.2 %)

(0.7 %)

(1.9 %)(0.5 %)

+H2O-H2O

H3ax

H3eq

H

# = 13C

# #

##

#

27

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 oC. (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. 15.

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. 16.

O

acyclic keto

(19.6 %)

!-D-xylulofuranose(18.1 %)

"-D-xylulofuranose(62.3 %)

acyclic hydrate

(0 %)

OH

+H2O-H2O

Scheme 4. Anomerization of D-[2-13C]xylulose (4), showing percentages of forms in solution determined by 13C NMR spectroscopy (0.3 M ketose, 85/15 v/v 1H2O/2H2O, 50 m M Na-acetate buffer, pH 4.0, 26 oC). Data were taken from ref. 17.

OH

HOOH

HOO

CH2OH

OHHO

OHCH2OH

CH2OH O OHOH

HO

CH2OH

HO

OH

OH

28

‘

O

acyclic aldehyde

(2.9 %)

!-D-erythrofuranose(28.2 %)

"-D-erythrofuranose(57.7 %)

acyclic D-erythrose

hydrate(11.1 %)

OH

+H2O-H2O

Scheme 5. Anomerization of D -erythrose ( 5), showing percentages of forms in 2H2O solution at 60o determined by 1H NMR spectroscopy (20)

OHHO

O OH

OHHOOHOHOH

CHO

OHOHOH

CH(OH)2

Figure 6. Percentages of cyclic and acyclic forms of D-[2-13C]threo-pentulose (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. 17.

29

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 oC. (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) (15 s) of saturation 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 <0.05 s-1 is obtainable for kdehydration. Data were taken from ref. 22.

Solution conditions: 50 mM Na acetate, pH 4.0,85/15 v/v 1H2O/2H2O, 28 °C. Data were taken from ref. 23.

!-furanose "-furanose

!-pyranose "-pyranose

aldehyde

0.0043 s-1

7.8 s-1

0.046 s-1

43 s-1

0.0019 s-1

2.7 s-1

0.037 s-1

22 s-1

(41%)

(18.5%)

(0.03%)

(29%)

(11.6%)

Scheme 6. Anomerization equilibria and kinetics for D-[1-13C]talose (7)

hydrate(0.03%)

30

Figure 8. pH Dependencies of kopen for the furanose forms of D-[1-13C]ribose 5P (6) (0.3 M in 85/15 v/v 1H2O/2H2O and 25 oC). Rate constants were measured using line-broadening methods except for values obtained at pH 2.3 and 4.0, which were measured by saturation transfer. Data were taken from ref. 18.

Table 5. Ring-opening Rate Constants for Several Aldotetroses and C5-Modified!Aldopentosesa

ak!o and k"o are ring opening rate constants for the !- and "-furanoses, respectively, of each compound. Solution conditions: 0.25 M aldose, 85/15 v/v 1H2O/2H2O, 50 mM Na acetate buffer, pH 4.0, 60 oC. Data were taken from ref. 24.

compound ring-opening rate constant

(s-1) kocis/kotrans ko! (±10%) ko" (±10%)

D-erythrose 0.69 0.53 1.3 D-threose 0.21 0.70 3.3

5-deoxy-L-arabinose 0.13 0.23 1.8 5-O-methyl-D-

arabinose 0.10 0.16 1.6

5-deoxy-L-lyxose 0.14 0.13 0.92 5-O-methyl-D-lyxose 0.13 0.14 1.1

5-deoxy-L-ribose 0.44 0.41 1.1 5-O-methyl-D-ribose 0.35 0.31 1.1

5-deoxy-L-xylose 0.53 0.20 2.7 5-O-methyl-D-xylose 0.37 0.15 2.5

OScheme 8. Anchimeric mechanism of furanose

ring opening (cis-1,2 effect). Note that O2 and O3 are trans; when cis,

the cis-1,2 effect is diminished or abolished.O-H

HO

HOHO-HO

OH

OHHO

R

R = H, CH3 or CH2OCH3

Scheme 7

31

Table 6. Unidirectional Rate Constants for Anomerization of D-[1-13C]Erythrose (5), D-[1-13C]Threose (3), D-[2-13C]Erythro-pentulose (4)and D-[2-13C]Threo-pentulose (8)

a0.3 M aldose or ketose, 85/15 v/v 1H2O/2H2O, 50 mM Na acetate buffer, pH 4.0, 55 oC. Date were taken from ref. 26.

compound rate constant (s-1) (± 5%)a kopen kclose

!-D-erythrofuranose 0.51 9.9 "-D-erythrofuranose 0.37 15 !-D-threofuranose 0.25 6.2 "-D-threofuranose 0.65 12

avg (± 1 SD) 0.45 (0.17) 10.8 (3.7) !-D-erythro-pentulofuranose 0.18 0.38 "-D-erythro-pentulofuranose 0.18 0.13 !-D-threo-pentulofuranose 0.20 0.13 "-D-threo-pentulofuranose 0.35 0.71

avg (± 1 SD) 0.23 (0.08) 0.34 (0.27)

Table 7. Thorpe-Ingold Effects on Anomerization Equilibria in Furanose Systems

aConditions: 0.1 M aldose, 2H2O, 50 mM Na acetate buffer, p2H 5.0, 25 oC. bConditions: 0.1 M aldose, 2H2O, 30 oC. cConditions: 0.25 M aldose, 2H2O, 25 oC. Data were taken from ref. 20.

compound percent in solution !f "f hydrate aldehyde

D-threose (3)a 51.8 37.6 9.6 0.96 2-deoxy-D-glycero-tetrose (9)b 25.3 59.2 2.4 3.1 3-deoxy-DL-glycero-tetrose (10)b 21.3 70.1 7.8 0.7 3-C-methyl-DL-erythrose (11)c 30.1 69.4 0.3 0.2 3-C-methyl-DL-threose (12)c 55.0 44.3 0.5 0.3 3-deoxy-3,3-di-C-methyl-DL-glycero-tetrose (13)c 29.0 71.0 0.04 0.1

5-O-methyl-D-ribose (14)c 34.5 64.6 0.8 0.1 2-deoxy-5-O-methyl-D-erythro-pentose (15)b 52.0 44.6 2.3 ~1.1

Figure 9. Plots of kopen against [H3O+] for the furanose anomers of 5-deoxy-L-lyxose (filled symbols) and 5-O-methyl-D-lyxose (open symbols). Squares = β-furanoses; circles = α-furanoses. Solution conditions: 0.25 M aldose, 50 mM KCl, 85/15 v/v 1H2O/2H2O, pH adjusted with HCl, 37 oC. Data were taken from ref. 24.

32

Table 8. Thorpe-Ingold Effects on Anomerization Rate Constants in Furanose Systems

aConditions: 0.1 or 0.25 M aldose, 2H2O, 50 mM Na acetate buffer, p2H 5.0, 60 oC. Data were taken from ref. 20.

compound rate constant (s-1) k!o k"o ko! ko"

D-erythrose (5) 0.49 0.31 4.8 6.2 D-threose (3) 0.090 0.45 1.6 6.0 2-deoxy-D-glycero-tetrose (9)b 0.43 0.30 1.4 1.9 3-deoxy-D-glycero-tetrose (10)b 0.23 0.15 3.0 7.0 3-C-methyl-DL-erythrose (11)c 0.27 0.12 12.1 9.4 3-C-methyl-DL-threose (12)c 0.18 0.74 12.6 39.2 3-deoxy-3,3-di-C-methyl- DL-glycero-tetrose (13)c 0.38 0.075 19.6 8.5

2-deoxy-5-O-methyl-D-erythro-pentose (15)b 0.15 0.16 5.2 4.6

O

OH

HOC1

2E confomerof 9; O1 and O1quasi-axial

O

C1

C3

3E confomerof 15; O1 and O3quasi-equatorial

OHHO

C3

Scheme 9. Preferred ring conformations of 9 and 15, and implications for kopen

O

OOHHO

OPO O-

O H

H

Scheme 10. Mechanism ofintramolecular catalysis ofring-opening of !-anomers

by phospate in pentosephosphate mono-anions

!

Table 9. Ring-Opening Rate Constantsa for Pentose Phosphates

aConditions: 0.15 M sugar phosphate. 85/15 v/v 1H2O/2H2O, 40 oC. Data were taken from ref. 18.

compound rate constant (s-1) pH 4.2 pH 7.5

k!o k"o k!o k"o arabinose 5-P (16) 0.64 0.49 50 33 lyxose 5-P (17) 0.24 0.22 13 13 ribose 5-P (6) 0.86 0.44 100 40 xylose 5-P (18) 0.42 0.17 33 9.6

33

Table 10. Ring-Opening Rate Constantsa for Penturonic Acids 19–22

aConditions: 0.3 M uronic acid, 70/30 v/v 1H2O/2H2O, 50 oC. Data were taken from ref. 31.

compound rate constant (s-1) pH 1.5 pH 4.5

k!o k"o k!o k"o D-arabinuronic acid (19) 0.46 1.52 0.67 3.22 D-lyxuronic acid (20) 0.40 0.55 0.59 1.12 D-riburonic acid (21) 1.65 0.76 1.03 1.27 D-xyluronic (22) 2.57 1.09 1.58 1.55

OO

OHHO

CH

Scheme 11. Potential modes of intramolecularcatalysis of ring-opening of penturonic acids (19)–(22).

(A) Protonated COOH group. (B) Ionized COOH group.

O O HO O

OHHO

C

HO O-

!

A B

O

OCH3

HOHOHO

trans

N CO

H CH3

O

OCH3

HOHO

HO

cis

N COH

CH3

Ktrans/ciskcis!trans

ktrans!cis

Scheme 13. Structural properties of N-formyl and N-acetyl side-chains (shown in blue) appended to methyl 2-amino-2-deoxy- D-glucopyranosylaminide. The freely rotatable "1 bond and the restricted bond "2 are identified. Signal integration of 1H and/or 13C{1H} NMR spectra give the equilibrium constants, Ktrans/cis. 1H and/or 13C saturation-transfer NMR experiments give the first-order rate constants, kcis!trans and ktrans!cis.37

O

OCH3

HOHOHO

trans

N CO

H H

O

OCH3

HOHOHO

cis

N COH

Ktrans/cis

H kcis!trans

ktrans!cis"1

"2

34

O

HO

HOHO

OHOCH3

methyl !-D-glucopyranoside

O

OH

CH3CCOHN

HO

O OH

HO

H8

H7 OO-

CH3C

O

O

OH

CH3CCOHN

HON-acetyl-neuraminic acid

HO OH

HO

H8

H7 OO-

O

OH

CHN

HO

N-acetyl-neuraminic acid

HO OH

HO

H8

H7 OO-

hydroxymethylO-acetyl

C-glyceryl

CH3C

O

O

-2O3P-O

HOHO

OH OHD-glucose 6-phosphate

O

HO

HOHO

NH-SO3-1OH

O

HO

HO

NHCCH3

OCOO-

O

H

H3C

O

-1O3S-O

HOHO OCH3

NHCCH3

O

N-acetyl

O-phosphomonoesterO-sulfomonoester

O-L-lactoyl

N-sulfamide

9-O-acetyl-N-acetyl-neuraminic acid

N-acetyl-muramic acid

!-D-glucopyranosylamineN-sulfamide

2-acetamido-2-deoxy-!-D-glucopyranose 6-sulfate

Scheme 12. Structures of different types of side-chains and substituents found inmonosaccharides at physiological pH

OH

35

Figure 10. 1H NMR spectrum (600 MHz; 2H2O) of 24 at 22 oC, showing signals arising from the formyl (HCO) and anomeric (H1) hydrogens in the cis and trans forms of the molecule (Scheme 13). The formyl hydrogen signals are split by 1JCH since 24 is selectively labeled with 13C (99 atom-% 13C) at the formyl carbon. Data were taken from ref. 37.

Figure 11. Equilibrium constants, Ktrans/cis, as a function of temperature for (A) 23 and 24, and (B) 25 and 26. Green = α-anomers; blue = β-anomers. Data were taken from ref. 37.

36

Figure 12. 13C{1H} NMR spectrum (150 MHz, 2H2O) of 26 at 22 oC. (A) Carbonyl signals assigned to the cis and trans forms, each split into doublets by 2JC2,CO. (B and C). C2 signals assigned to the cis and trans forms, respectively, each split into doublets by the same 2JC2,CO observed for the respective signals in (A). Data were taken from ref. 37.

C1C3N

Scheme 14. Newman projection (A) for the C2–N2 fragment (!1) in compounds 23–26. The anti arrangement between H2 and NH appears to be preferred in solution based on 3JH2,NH values. This geometry has the C2– H2 and C=O bonds eclipsed in the trans configuration of the amide (!2) (B).

H

COH2 H2

NC

O

R

C1

C3

R = H or CH3

H

eclipsedA B

!1 !2

37

Figure 13. Dependencies of 3JH2,NH on the H2–C2–N2–H torsion angle (θ1) in models of 23–26 determined by DFT. Green, amide in cis configuration; blue, amide in trans configuration (see Scheme 13). The difference between calculated couplings at θ1 = 0o and θ1 = 180o is < ~2 Hz in both plots. Data were taken from ref. 40.

Figure 14. (A) Plots of carbonyl carbon signal intensity in the trans form of 25 as a function of saturation time of the carbonyl carbon of the cis form at different temperatures. Black, 42.0 oC; blue, 52.7 oC; green, 64.4 oC; red, 75.1 oC; rose, 84.6 oC. (B) Linearizing the data shown in (A) from which ktrans→cis values at each temperaure were determined. Data were taken from ref. 37.

38

Figure 15. CTI rate constants measured at different temperatures by 13C saturation transfer. (A) 23 (circles) and 24 (squares). (B) 25 (circles) and 26 (squares). Black = ktrans→cis. Green = kcis→trans. Data were taken from ref. 37.

39

O

OHO

HO

COH3C

O

O

HO

CO

H3C

O

O

OH

HO

CO

H3C

HO

HOOH

OH

OH3-O-[1'-13C]acetyl-!,"-D-glucopyranoses (28!/")

2-O-[1'-13C]acetyl-!,"-D-glucopyranoses (27!/")

6-O-[1'-13C]acetyl-!,"-D-glucopyranoses (29!/")

#

#

#HO

HOScheme 15. 13C-Labeled mono-O-acetylated D-glucopyranoses27–29

# = 13C

$1

$1

$1

$2

$2$2

C4C2

O

Scheme 16. Newman projection (A) for the C3– O3 fragment ( !1) in compound 28"/#. The preferred geometry has the C3–H3 and C=O bonds eclipsed in the trans configuration of the ester (!2) (B).

COH3 H3

OC

O

CH3

C4

C2

eclipsedA B

!1 !2

~0o

40

Figure 16. Probability distributions across θ1 in 27–29 detemined from a fit of redundant experimental J-couplings using DFT parameterized equations similar to eqs. [2]–[5] and a von Mises model using the MAʼAT statistics package. Red, 27α; blue, 27β; green, 28α/β; black, 29α/β. Torsion angle θ in this plot is equivalent to θ1 in Scheme 15, where the torsion angle is defined as that between the O-acetyl carbonyl carbon and the hydrgogen attached to the carbon bearing the side-chain. For 29, the reference hydrogen is H6R. Data were taken from ref. 35.

C1'H6S

C5

H6RH6R

OC

O

CH3

H6S

C5

B

!1 !2

~-66o

Scheme 17. Newman projection (A) for the C6–O6 fragment ( !1) in compound 29"/#. The preferred geometry has the C=O bond bisecting the H6R–C6–H6S bond angle in the trans configuration of the ester (!2) (B).

A

41

OMo MoO

O

O

OO

CC

C

OO

R

H

HH

C

O

H

21

34

OH

OH-

OMo MoO

O

O

OO

CC

C

OO

R

H

HH

C

O

H

21

34

HO

Scheme 18. Interconversion of bimolybdate complexes involving the acyclic hydrate forms of D-glucose (complex A) and D-mannose (complex B). Arrows in complex A show S N2-like insertion and expulsion of OH - during its conversion to complex B. The attack of C3 on C1 in complex A is equivalent to attacking the re face of the C1 carbonyl equivalent. Likewise, in the reverse direction, attack of C3 occurs on the si face of the equivalent carbonyl at C2.

-2

-2

complex A

complex B

D-[1-13C]glucose hydrate (C6H14O7) + Mo2O7H2

!

!

D-[2-13C]mannose hydrate (C6H14O7) + Mo2O7H2

–2H++2H+

+2H+–2H+

42

D-[1,3-13C2]mannoseD-[1,3-13C2]glucoseD-[1-13C]ribose

+!

D-[2-13C]arabinose

! !HOHO

OHOHOH

OHOHOH

OHHO

OHOHOHOH MCE

OHOHOH

CHOHO

CHO CHOCHO

D-erythrose

D-[1-13C]arabinoseOHOHOH

HOCHO

+

!

!

MCE

"

"

D-[2-13C]riboseOHOHOHOH

CHO! [2'-13C]ribonucleosides

[2'-13C]2'-deoxyribonucleosides

D-[1,2-13C2]mannoseD-[1,2-13C2]glucose

+! !HO

HOOHOHOH

OHOHOH

OHHO

CHO CHO! !

! !

D-[2,3-13C2]glucose

!OHOHOH

OHHO

CHO!

MCE

Scheme 20. Synthetic routes showing the integration of cyanohydrin reduction (CR) and molybdate-catalyzed epimerization (MCE) in the synthesis of singly and doubly-13C-labeled aldopentoses and aldohexoses, and labeled nucleosides, from D-erythrose

MCE"

[1'-13C]ribonucleosides[1'-13C]2'-deoxyribonucleosides

K13CNCR

"

K13CNCR

CR K13CN! = 13C

Scheme 21. Two reactions (A) and (B) showing the application of MCE to interconvert 2-C-substituted D-erythroses with 2-ketoses

OOH

HO OH

H3C

OH

OHOH

CHOH3C

1-deoxy-D-xylulose

OHOH

OHO

CH3

pH 4.2, 70o C30 min

2-C-(methyl)-D-erythrose

Na2Mo2O7

OOH

HO OH

O

OHHO

HO

OO

HO

O

OHHO

HO

OHO

CH2

OHMo resin

67 oCH2O

OHOH

A

B

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

Pd/BaSO4pH 7.0-7.5+ H2 CHOHRR

CHOHKCNCN CHO

R

CHO xsH2O

R

CH(OH)2 xsH2O

R

CHO

43

D-[2-13C]gluconate (31)(~85%)

COO-

OHOHOH

HOOH

Na2Mo2O7

pH 4.5

D-[1-13C]glucosone (30)(dihydrate form)

CH(OH)2

OHOHOH

HOOH

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

D-[2-13C]mannonate (32)(~15%)

COO-

OHOHOH

HOHO

! = 13C!

! !

+ + minor 13C-labeledproducts

HO

Scheme 23. Hypothetical reaction of a 2,3-dicarbonyl sugar with bimolybdate to give branched-chain aldonates

OH

OH!

HO

R1OHHO

R2

HO

COO-

OH

OH!R1

R2

HO

COO-

OH

HO ! R1

R2

HO+pH 4.5

Na2Mo2O7?

! = 13C

CHOC

HOCH

OHOHOH

O

CHO

COH

HOC

OHOHOH

HCOO-

+

2,3-enediol 1,3-dicarbonyl

CHO

CHOH

COHOHOH

O

CH2OH

COHOHOH

O

D-[1-13C]ribulose (33)

OH-

HO-H

Scheme 24. Degradation of D-[2-13C]glucosone (30) to give D-[1-13C]ribulose (33) and unlabeled formate. The pathway presumably involves a 2,3-enediol and a 1,3-dicarbonyl sugar as intermediates.

D-[2-13C]glucosone (30)

! ! ! !

! = 13Cenolization

44

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 oC) after 20 days. (A) Full spectrum showing three signals “a” from D-[2-13C]33 (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]33 (signals “a”), the furanose C2 signals from D-[1,2-13C2]33 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]33 (three signals “a” each split by 1JC1,C2, [2-13C]glycolate (signal b) and an unidentified intermediate (signal “c”). Data were taken from ref. 54.

CHOC

HOCHOHOHOH

O

CH2OHC

OHOHOH

O

D-[2-13C]ribulose (33)

CH2OHC

OHOHOH

O

D-[1,2-13C2]ribulose (33)

~90%

~10%

+ H!COO-

+ HCOO-

Scheme 25. Reaction partitioning during thedegradation of D-[1,3-13C2]glucosone (30) inphosphate buffer

!

!

!

!

!

! = 13C

D-[1,3-13C2]glucosone (30)

45

HC

C

HOCH

OHOHOH

O

O

P

O

-O OHO-

H-OHHC

C

HOCHOHOHOH

O

OH

PO

-O

OH

O

H-OH

HC

C

COHOHOH

OP

O

O-O

OH

HO

HO

H

1,2-cyclic phosphate

HC

C

OH

COH

OH

OP

OHO

-O

OH

HO

HO

HC

C

COHOHOH

OP O-O

O

OH

HO

HO

1,3-cyclic phosphate

H

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

D-glucosone (30) 1-phosphate2,3-enediol

N S

CHO CHH

OH

N S

CHO CHHOH1

2 2

1

N S

CHO C

HH

OH2 1

N S

CHO C

H

OH12

H

Scheme 27. A potential mechanism for C1– C2 tranposition in 2-(1,2-dihydroxyethyl)-TPP during two-carbon (acylium anion) transfer catalyzed by transketolases. Formation of the symmetric cyclopropanediol intermediate may not be favorable.