Bromoketone C-glycosides, a new class of β-glucanase inactivators

Article abstract of DOI:10.1021/ja981580r

Although reliable methods have been developed for the labeling of the active site nucleophiles in glycosidases, no such reliable method has been developed for the identification of the acid/base catalyst. To address this problem two novel bromoketone affi

Full text of DOI:10.1021/ja981580r

10326
J. Am. Chem. Soc. 1998, 120, 10326-10331
Bromoketone C-Glycosides, a New Class of â-Glucanase Inactivators†
Steven Howard and Stephen G. Withers*
Contribution from the Protein Engineering Network of Centres of Excellence of Canada and Department
of Chemistry, UniVersity of British Columbia, VancouVer, BC V6T 1Z1, Canada
ReceiVed May 6, 1998
Abstract: Although reliable methods have been developed for the labeling of the active site nucleophiles in
glycosidases, no such reliable method has been developed for the identification of the acid/base catalyst. To
address this problem two novel bromoketone affinity labels based on a â-C-glucoside (6), and a â-C-cellobioside
(9), have been synthesized via a chemoenzymatic process and tested as inactivators of the â-glucosidase from
Agrobacterium sp. and the â-glucanases from Cellulomonas fimi. The â-glucosidase was inactivated by 6 in
a time-dependent manner according to kinetic parameters of ki ) 0.01 min-1 and KI ) 3.1 mM. Electrospray
ionization mass spectrometric analysis revealed that multiple labeling of the enzyme had occurred. The
â-endoglucanases, CenA and CenD, were inactivated stoicheometrically by 9 according to kinetic parameters
of ki ) 0.0155 min-1; KI ) 0.35 mM and ki ) 0.01 min-1; KI ) 6.0 mM, respectively. These should therefore
prove to be valuable reagents for the labeling of glycosidases.
Introduction
yields the product with retention of anomeric configuration.
Inverting glycosidases use a single-step mechanism in which
general base catalysis facilitates direct attack by water at the
anomeric center, with general acid-catalyzed departure of the
aglycon. All steps proceed via transition states with substantial
oxocarbenium ion character.
Structural information, especially with respect to the identities
of active site residues, is a key requirement for the understanding
of these important enzymes and also provides the basis for
engineering of new proteins with modified or improved function.
A variety of mechanism-based inhibitors and affinity labels has
been developed to label and identify catalytically important
residues in this class of enzymes.10,11 The catalytic nucleophiles
in many cellulases, including Cex, have been labeled using
2-deoxy-2-fluoro-â-D-glycosides and their identities subse-
quently determined by peptide mapping.12-16 In other cases
the catalytic nucleophile has been labeled and identified using
epoxyalkyl-â-D-xylosides and cellobiosides.17,18
There has been widespread interest in glycosidases in recent
years, largely due to their role in a multitude of biological and
industrial processes. In particular, enzymes such as cellulases
and amylases are important for the industrial degradation of
biomass. Microorganisms that degrade cellulose generally
secrete many different cellulases which act synergistically to
hydrolyze the substrate. One organism, Cellulomonas fimi,
produces an array of cellulases when grown on cellulose. Four
â-endoglucanases (CenA, CenB, CenC, and CenD),1-4 two
exoglucanases (CbhA and CbhB),5,6 and an exoglucanase/
xylanase (Cex)7 have been cloned and expressed in Escherichia
coli.
Glycosidases can be classified as “retaining” or “inverting”
depending upon the anomeric configuration of the initially
formed product with respect to the substrate. Both types possess
two catalytic carboxylic acid residues within the active site.
Retaining glycosidases employ a “double-displacement mech-
anism” in which, in the first step, one residue provides general
acid catalysis and the other provides nucleophilic catalysis to
form a covalent glycosyl-enzyme intermediate.8,9 In the second
step, general base-catalyzed hydrolysis of this intermediate
Affinity labels have also been used successfully in the labeling
and identification of the acid/base catalyst in some cellulases
and xylanases. Notable successes include the identification of
the acid/base catalytic residue in Cex with N-bromoacetyl-â-
D-cellobiosylamine19 and the determination of the crystal
* To whom correspondence should be addressed. Telephone: 604-822-
3402. Fax: (604) 822-2847. E-mail: withers@chem.ubc.ca.
† Abbreviations: C-glycoside, carbon glycoside; â-PNPG, 4-nitrophenyl-
â-D-glucopyranoside; â-DNPC, 2,4-dinitrophenyl-â-D-cellobioside; â-PNPC,
4-nitrophenyl-â-D-cellobioside; TLC, thin-layer chromatography; LC/MS,
liquid chromatographic/mass spectrometric; ESMS, electrospray mass
spectrometry; IPTG, isopropylthio-â-D-glucopyranoside.
(8) McCarter, J.; Withers, S. G. Curr. Opin. Struct. Biol. 1994, 4, 885-
897.
(9) Sinnott, M. L. Chem. ReV. 1990, 90, 1171-1202.
(10) Withers, S. G.; Aebersold, R. Protein Sci. 1995, 4, 361-372.
(11) Legler, G. AdV. Carbohydr. Chem. Biochem. 1990, 48, 319-385.
(12) Withers, S. G.; Warren, R. A. J.; Street, I. P.; Rupitz, K.; Kempton,
J. B.; Aebersold, R. J. Am. Chem. Soc. 1990, 112, 5887-5889.
(13) Tull, D.; Withers, S. G.; Gilkes, N. R.; Kilburn, D. G.; Warren, R.
A. J.; Aebersold, R. J. Biol. Chem. 1991, 266, 15621-15625.
(14) Miao, S.; Ziser, L.; Aebersold, R.; Withers, S. G. Biochemistry 1994,
33, 7027-7032.
(1) Damude, H. G.; Withers, S. G.; Kilburn, D. G.; Miller, J. R. C.;
Warren, R. A. J. Biochemistry 1995, 34, 2220-2224.
(2) Meinke, A.; Schmuck, M.; Gilkes, N. R.; Kilburn, D. G.; Miller, R.
C.; Warren, R. A. J. Glycobiology 1992, 21, 321-326.
(3) Tomme, P.; Kwan, E.; Gilkes, N. R.; Kilburn, D. G.; Warren, R. A.
J. J. Bacteriol. 1996, 178, 4216-4223.
(15) White, A.; Tull, D.; Johns, K.; Withers, S. G. Nat. Struct. Biol.
1996, 3, 149-154.
(4) Meinke, A.; Gilkes, N. R.; Kilburn, D. G.; Miller, J. R. C.; Warren,
R. A. J. Am. Soc. Microbiol. 1993, 175, 1910-1018.
(16) Mackenzie, L. F.; Brooke, G. B.; Cutfield, J. F.; Sullivan, P. A.;
Withers, S. G. J. Biol. Chem. 1997, 272, 3161-3167.
(17) Havukainen, R.; Torronen, A.; Laitinen, T.; Rouvinen, J. Biochem-
istry 1996, 35, 9617-9624.
(5) Meinke, A.; Gilkes, N. R.; Kwan, E.; Kilburn, D. G.; Warren, R. A.
J.; Miller, R. C. Mol. Microbiol. 1994, 12, 413-422.
(6) Shen, H.; Gilkes, N. R.; Kilburn, D. G.; Miller, R. C.; Warren, R. A.
J. Biochem. J. 1995, 311, 67-74.
(18) Sulzenbacher, G.; Schulen, M.; Davies, G. J. Biochemistry 1997,
36, 5902-5911.
(7) Tull, D.; Withers, S. G. Biochemistry 1994, 33, 6363-6370.
S0002-7863(98)01580-7 CCC: $15.00 © 1998 American Chemical Society
Published on Web 09/23/1998

Bromoketone C-Glycosides
J. Am. Chem. Soc., Vol. 120, No. 40, 1998 10327
structure of the endo-â(1,4)-xylanase II from Trichoderma reesei
in which the putative acid/base catalyst was covalently labeled
with an epoxyalkyl-â-D-xyloside.17 N-Bromoacetylglycosyl-
amines have the advantage that they are inert toward enzyme-
catalyzed hydrolysis, a potential problem in many other O-linked
affinity labels, and they have been used with varying success.
Keresztessy et al. have reported the successful labeling and
identification of the putative acid/base catalyst in the â-glu-
cosidase from Manihot esculenta Crantz using N-bromoacetyl-
â-D-glucosylamine.20 However, this compound was found to
cause unselective, multiple labeling in the â-glucosidase from
Agrobacterium sp.21 Furthermore, in the case of E. coli
â-galactosidase, N-bromoacetyl-â-D-galactosylamine was found
to label a single active site residue, Met-502, which was,
however, later shown not to participate in catalysis.22
Scheme 1. Synthesis of â-C-Glucoside (6) and
â-C-Cellobioside (9)a
Clearly, there is a need for new affinity labels with improved
selectivity. Here we report the synthesis of novel cellobioside
and glucoside affinity labels, using chemoenzymatic methods,
which are designed to label enzyme acid/base catalysts. These
new affinity labels 6 and 9 (Scheme 1) incorporate an R-bro-
moketone functionality into hydrolytically stable â-C-glycosides.
Of interest as targets for these novel â-C-glycosides are
â-glucosidases and endo and exo-glucanases. Agrobacterium
sp. â-glucosidase has undergone rigorous mechanistic investiga-
tions, but attempts to selectively label the acid/base catalyst have
so far been unsuccessful. In addition, â-(1,4)-glucanases from
C. fimi are attractive targets for these compounds since the acid/
base catalysts in most of these enzymes have not been identified
by labeling experiments. Although the acid/base catalyst in the
exoglucanase/xylanase, Cex, was labeled and identified using
N-bromoacetyl-â-D-cellobiosylamine, other glucanases, such as
CenD, are not inactivated by this compound. New compounds
are therefore required to probe the wide range of cellulases and
their different specificities.
a (a) mCPBA, CH2Cl2, 85%; (b) DIBAL/BuLi, THF, 92%; (c) CrO3,
H2SO4, 86%; (d) 5% Pd-C, H2, 10% AcOH-MeOH, 90%; (e) 5,
MeOH, Br2, 53%; (f) Agrobacterium sp. â-glucosidase-E358A mutant,
â-glucosyl fluoride, pH 7.0, 21%; (g) pyridine/Ac2O, 33%; (h) MeOH,
Br2, 58%.
Results and Discussion
The synthetic strategy was based on â-C-glycoside ketones
5 and 7 which were obtained from the allyl-â-C-glucoside (1)
via oxidation of the olefin function (Scheme 1). The final
bromoketones were obtained by acid-catalyzed bromination at
the R-positions with respect to the ketone functionalities.
Allyl-â-C-glucoside (1) was readily prepared from Grignard
addition to benzylated 1,5-gluconolactone.23 A one-step con-
version of this compound to the ketone 4 with a Wacker
oxidation was unsuccessful due to the low solubility of the
starting material under the reaction conditions. Epoxidation of
the olefin 1 with mCPBA followed by reduction with lithium
bis(diisopropylbutyl) aluminum hydride24 (prepared in situ) gave
the diastereomeric alcohols 3a,b. Both diastereomers were then
converted to the ketone 4 via the Jones’ oxidation. Deprotection
and bromination of 4 gave the desired material 6.
second glucose unit to the â-C-glycoside 5. This was achieved
by incubating the Glu358Ala mutant of Agrobacterium sp.
â-glucosidase26 with the glycosyl acceptor 5 and R-glucosyl
fluoride as the glucosyl donor, at concentrations of 100 and 50
mM, respectively. The desired cellobioside 7 was obtained in
21% yield (based on recovered starting material) with lesser
amounts of trisaccharide products. Repeating the reaction with
a higher concentration of 5 (150 mM), double the buffer
concentration (200 mM), and purification by flash chromatog-
raphy rather than HPLC provided a small improvement in the
yield (25%). The relatively low yields for this step are a
reflection of the strong preference of this enzyme for an aromatic
aglycon.27 Acetylation of the cellobioside 7 and analysis of
the 1H NMR-COSY spectrum revealed an unambiguous â(1,4)-
linkage. Bromination of 7 gave the desired product 9.
Best et al. have described the synthesis of allyl-â-C-
cellobioside in an analogous fashion to 1, but it was reported
that Grignard additions to â-D-cellobionolactone are low yield-
ing.25 As an alternative route to the target material 9, the â-C-
cellobioside 7 was synthesized via enzymatic addition of a
The â-C-glucoside 6 caused time-dependent inactivation of
Agrobacterium sp. â-glucosidase according to first-order kinet-
ics. Residual enzyme activity, at several different concentrations
of 6, was measured over time, and the observed pseudo-first-
order rate constants for inactivation were determined for each
(19) Tull, T.; Burgoyne, D. L.; Chow, D. T.; Withers, S. G.; Aebersold,
R. Analytical Biochem. 1996, 234, 119-125.
(20) Keresztessy, Z.; Kiss, L.; Hughes, M. A. Arch. Biochem. Biophys.
1994, 314, 142-152.
(21) Black, S.; Laszlo, K.; Tull, D.; Withers, S. G. Carbohydr. Res. 1993,
250, 195-202.
(25) Best, W. M.; Ferro, V.; Harle, J.; Stick, R. V.; Tilbrook, M. G.
Aust. J. Chem. 1997, 50, 463-472.
(22) Naider, F.; Bohak, Z.; Yariv, J. Biochemistry 1972, 11, 3202-3206.
(23) Lewis, M. D.; Cha, J. K.; Kishi, Y. J. Am. Chem. Soc. 1982, 104,
4976-4978.
(26) Mackenzie, L. F.; Wang, Q.; Warren, R. A. J.; Withers, S. G. J.
Am. Chem. Soc. 1998, 120, 5583-5584.
(24) Sunggak, K.; Ahn, K. H. J. Org. Chem. 1984, 49, 1717-1725.
(27) Kempton, J. B.; Withers, S. G. Biochemistry 1992, 31, 9961-9969.

10328 J. Am. Chem. Soc., Vol. 120, No. 40, 1998
Howard and Withers
the LC/MS mode (data not shown) revealed three major species
corresponding to protein molecules having 11, 12, and 13 labels
attached, a result very similar to that obtained with this enzyme
using N-bromoacetyl-â-D-glucosylamine.21
When the â-C-cellobioside analogue 9 was tested as an
inactivator of six different cellulases, its inactivation behavior
was found to be profoundly different from that with N-bro-
moacetyl-â-D-cellobiosylamine. Whereas N-bromoacetyl-â-D-
cellobiosylamine is an effective inactivator of the exoglucanase
(Cex),19 9 provided no inactivation. Further, both 6 and 9 were
found to inactivate the endoglucanase (CenA), yet only 9 was
able to inactivate CenD. Both CenA and CenD are readily
assayed with the chromogenic substrate â-DNPC. These were
therefore subjected to kinetic analysis in the presence of 9 using
methods analogous to those described above for Agrobacterium
sp. â-glucosidase.
The â-C-cellobioside 9 caused time-dependent inactivation
of CenD according to pseudo-first-order kinetics (Figure 2a).
Data were not recorded beyond 600 min due to indications from
TLC that 9 begins to decompose after this length of time.
Values of KI ) 6.0 ((0.9) mM and ki ) 0.01 ((0.001) min-1
were determined by fitting the data to Scheme 2 using a
nonlinear regression analysis (Figure 2b). Active-site-directed
protection was afforded by 50 mM cellobiose, the pseudo-first-
order rate constant for enzyme inactivation being reduced from
0.0053 to 0.0011 min-1 in the presence of 6.04 mM (9). ESMS
analysis of CenD using the spectrometer in the LC/MS mode
revealed one major species of 75 016 ((8) Da corresponding
to the native enzyme (Figure 3a). The same analysis of a
partially inactivated sample of CenD [9 (7 mM), 22 °C, 5 h]
showed two species, 75 021 ((8) Da and 75 394 ((9) Da, which
correspond to the unlabeled and labeled enzyme, respectively
(Figure 3b). The mass increase of 373 Da is consistent with
the expected mass increase of 381 Da as a result of the
stoichiometric labeling of CenD with one molecule of 9.
However, incubation of CenD with 9 for extended periods of
time (15 h) resulted in the addition of a second label. This is
not surprising given the presence of the inherently reactive
bromoketone functionality, and also given that the enzyme must
contain an extended oligosaccharide binding site into which the
label may bind.
Figure 1. Inactivation of Agrobacterium sp. â-glucosidase by 6. (a)
Plot of residual activity versus time at the following inactivator
concentrations: 3, 0 mM; O, 0.78 mM; b, 1.95 mM; 0, 3.9 mM; 9,
CenA was also inactivated by 9 in a time-dependent manner
(Figure 4a), values of KI ) 0.35 ((0.02) mM and ki ) 0.0155
((0.0004) min-1 being determined directly from a plot of kobs
versus inactivator concentration (Figure 4b). Again, cellobiose
(50 mM) was shown to afford protection from inactivation,
reducing the pseudo-first-order rate constant for enzyme inac-
tivation by 9 (0.51 mM) from 0.0093 min-1 to essentially zero
(Figure 4c). LC/MS analysis of CenA was conducted in an
analogous fashion to that described for CenD. Analysis of
unlabeled CenA (catalytic domain only) revealed a pair of peaks
of masses 43 829 ((5) and 44 102 ((5) Da (Figure 5a), the
different masses corresponding to slightly different positions
of cleavage when the cellulose binding domain was removed
proteolytically. Incubation with 9 (7 mM, 22 °C, 5 h) followed
by ESMS analysis revealed that both species were labeled to
an extent of approximately 70% as evidenced by the appearance
of two new species of masses 44 211 ((5) and 44 480 ((5)
Da (Figure 5b). The mass increase of 380 Da is again consistent
with stoichiometric labeling and, as with CenD, prolonged
incubation (15 h) resulted in some secondary labeling.
7.8 mM; 4, 11.7 mM; 2, 19.5 mM (activity
rate of â-PNPG
hydrolysis). (b) Plot of pseudo-first-order rate constants from panel a
versus inactivator concentration. (c) Plot of residual activity versus time
in the presence of (b) no inactivator, (0) 7.6 mM 6 + 15 mM IPTG,
and (9) 7.6 mM 6.
Scheme 2
concentration (Figure 1a). A plot of these rate constants versus
inactivator concentration is shown in Figure 1b. The saturation
kinetic behavior observed is compatible with the kinetic model
shown in Scheme 2.
Values of KI ) 3.1 ((0.53) mM and ki ) 0.01 ((0.0005)
min-1 were determined directly from this plot. Active-site-
directed protection against inactivation was demonstrated by
the use of 15 mM IPTG, the first-order rate constant for
inactivation by 6 (7.6 mM) being reduced from 0.0077 to 0.0048
min-1 (Figure 1c). ESMS analysis of the inactive enzyme in
The endocellulase, CenB, and the cellobiosyl hydrolases,
CbhA and CbhB, do not hydrolyze arylglycosides and so cannot
be assayed as readily. In addition to making assays more

Bromoketone C-Glycosides
J. Am. Chem. Soc., Vol. 120, No. 40, 1998 10329
Figure 2. Inactivation of CenD by 9. (a) Plot of residual activity versus
time at the following inactivator concentrations: 3, 0 mM; b, 0.3 mM;
O, 1.51 mM; 9, 1.68 mM; 0, 3.02 mM; 2, 6.04 mM; 4, 8.4 mM
(activity rate of â-DNPC hydrolysis). (b) Plot of pseudo-first-order
rate constants from panel a versus inactivator concentration. (c) Plot
of residual activity versus time in the presence of (9) no inactivator,
(O) 6.04 mM 9 + 50 mM cellobiose, and (b) 6.04 mM 9.
Figure 4. Inactivation of CenA by 9. (a) Plot of residual activity versus
time at the following inactivator concentrations: O, 0 mM; b, 0.05
mM; 0, 0.10 mM; 9, 0.20 mM; 4, 0.51 mM; 2, 2.3 mM (activity
rate of â-DNPC hydrolysis). (b) Plot of pseudo-first-order rate constants
from panel a versus inactivator concentration. (c) Plot of residual activity
versus time in the presence of (O) no inactivator, (9) 0.51 mM 9 + 50
mM cellobiose, and (0) 0.51 mM 9.
Figure 3. Electrospray mass spectrometry of CenD. (a) Reconstructed
mass spectrum of unlabeled enzyme. (b) Reconstructed mass spectrum
of enzyme partially inactivated by 9.
Figure 5. Electrospray mass spectrometry of CenA. (a) Reconstructed
mass spectrum of unlabeled enzyme. (b) Reconstructed mass spectrum
of enzyme partially inactivated by 9.
difficult, this also made it less likely that the affinity labels 6
or 9 would bind to and inactivate these enzymes. Nonetheless,
the possibility of labeling was monitored by ESMS. However,
after incubation with 9 for 15 h, no labeling was observed.
Clearly, there are profound differences between the reactivity
of these new affinity labels 6 and 9 and their N-(bromoacetyl)-
glycosylamine counterparts discussed earlier. What was not
clear a priori was whether these differences are the result of

10330 J. Am. Chem. Soc., Vol. 120, No. 40, 1998
Howard and Withers
mmol), and 2,6-di-tert-butyl-4-methylphenol (20 mg) were dissolved
in dichloromethane (60 mL), and the mixture was stirred at reflux for
4 h. After being cooled to room temperature, the mixture was diluted
with dichloromethane (60 mL) and washed successively with aqueous
10% sodium metabisulfite (50 mL), 10% aqueous NaHCO3 (3 × 50
mL), and saturated brine (50 mL). The organic layer was dried
(MgSO4) and filtered and the residue purified on silica gel (ethyl acetate:
petroleum ether (bp 35-60 °C), 1:6) to give the epoxide (2) (1.92,
3.40 mmol, 85%) as a colorless solid (mixture of diastereoisomers).
[R]D: +1.2 ° (c 0.92, CHCl3). 1H NMR (400 MHz, CDCl3): δ 7.13-
7.38 (m, Ar), 4.79-4.95 (m), 4.51-4.69 (m), 3.58-3.79 (m), 3.38-
3.40 (m), 3.26 (t, J 9.0 Hz), 3.07-3.16 (m), 2.74 (dt, J 15.3 and 4.5
Hz), 2.47-2.50 (m), 1.83-2.0 (m), 1.63 (ddd, J 14.1, 9.8 and 4.8 Hz,
H-3′).
2′-Hydroxy-3′-(2,3,4,6-tetra-O-benzyl-â-D-glucopyranosyl)pro-
pane (3a,b). Diisobutylaluminum hydride (1 M in hexane, 20 mL, 20
mmol) was dissolved in tetrahydrofuran (50 mL) and cooled to 0 °C,
and then n-butyllithium (1.6 M in hexane, 12.5 mL, 20 mmol) was
added dropwise. After being stirred for 30 min, the solution was added
to a stirred solution of the epoxide (2) (1.92 g, 3.40 mmol) in
tetrahydrofuran (50 mL) at 0 °C.
After 2 h, the reaction was quenched with ice-cold water (200 mL)
and then acidified with aqueous HCl (0.5 M). The aqueous mixture
was extracted with diethyl ether (3 × 200 mL), and the combined
organic fractions were washed successively with 10% aqueous NaHCO3
(200 mL) and saturated brine (200 mL). The organic layer was dried
(MgSO4) and filtered and the solvent removed in vacuo. Purification
on silica gel (ethyl acetate:petroleum ether (bp 35-60 °C), 1:3) gave
a 1:1 mixture of 3a and 3b (1.77 g, 3.05 mmol, 92%). Several of the
fractions contained pure samples of each diastereoisomer.
differences in inherent reactivity or a consequence of different
binding modes of these compounds in the enzyme active site
leading to different placement of the reactive functionalities with
respect to active site amino acids. In general R-halo esters and
amides tend to be less reactive than the corresponding R-ha-
loketones. However, the relative reactivities depend consider-
ably upon the nucleophile involved; thus, it is not easy to predict
which will react faster when the identity of the nucleophile is
unknown.28,29 In one case (Agrobacterium sp. â-glucosidase),
the bromoketone 6 and R-bromoamide exhibit very similar
inactivation behaviors. By contrast, only the R-bromoamide is
capable of inactivating Cex, the bromoketone having no effect.
It would therefore seem unlikely that the inherent reactivity of
the inhibitor is the dominant factor, but rather the different
binding modes likely mediated via hydrogen bonding to the
amide moiety.
It is also of interest to note that the second-order rate constants
for inactivation (ki/Ki ) 3.2, 1.7, and 44.3 M-1min-1 for CenD,
Abg, and CenA, respectively) are several orders of magnitude
greater than those measured previously for reaction of acetate
anion with bromoketones. For example, bromomethyl neopentyl
ketone, a similarly substituted derivative, reacts with acetate
anion at 54 °C with a second-order rate constant of 5.5 × 10-2
M-1 min-1 30
Reactions with acetic acid rather than acetate
.
are typically much slower still.31 The higher reaction rates at
the enzyme active site presumably reflect the location of the
carboxylic nucleophile in close proximity to the reactive center.
3a. Mp: 107-108 °C (ethyl acetate/petroleum ether, needles).
[R]D: -4.5° (c 1.5, CHCl3). 1H NMR (400 MHz, CDCl3): δ 7.18-
7.39 (20 H, m, OBn), 4.78-4.93 (4 H, m, CH2Ar), 4.47-4.64 (4 H,
m, CH2Ar), 4.02 (1 H, m, H-2′), 3.54-3.72 (5 H, m), 3.42 (1 H, dt, J
9.1 and 3.1 Hz), 3.37 (1 H, t, J 9.0 Hz), 2.09 (1 H, br, OH), 1.85 (1 H,
ddd, J3a′,3b′ 14.5 Hz, J 8.5 and 3.2 Hz, H-3a′), 1.71 (1 H, ddd, J 14.5,
7.3, and 2.4 Hz, H-3b′), 1.14 (3 H, d, J1′,2′ 7.0 Hz, H-1′). Anal. Calcd
for C37H42O6: C, 76.26; H, 7.26. Found: C, 76.47; H. 7.10. CIMS
(NH3): m/z 600 (M + NH4)+ (100%).
3b. Mp: 116-117 °C (ethyl acetate/petroleum ether, needles).
[R]D: +1.7° (c 1.2, CHCl3). 1H NMR (400 MHz, CDCl3): δ 7.15-
7.38 (20 H, m, Ar), 4.77-4.89 (4 H, m, CH2Ar), 4.48-4.63 (4 H, m,
CH2Ar), 3.97-4.03 (1 H, m, H-2′), 3.62-3.69 (2 H, m), 3.45-3.55 (4
H, m), 3.25 (1 H, t, J 9.1 Hz), 1.91 (1 H, d, J3a′3b′ 14.4 Hz, H-3a′), 1.57
(1 H, br, OH), 1.42 (1 H, dt, J3b′,2′ and J3b′,1′ 9.9 Hz, H-3b′), 1.14 (3 H,
d, J1′,2′ 6.2 Hz, H-1′). Anal. Calcd for C37H42O6: C, 76.26; H, 7.26.
Found: C, 76.42; H. 7.19.
Conclusion
These R-bromoketones therefore constitute a valuable new
class of affinity labels for glycosidases in general, and cellulases
in particular. The reactivity of this functionality is sufficiently
different from that of the known R-bromoamides and exocyclic
epoxides that they can be used in a complementary manner.
Further, the ability to synthesize R-bromoketones with the
R-anomeric configuration, an extremely difficult task for the
R-bromoamides, opens up the possibility of using this approach
with R-glycosidases. In addition to simply providing reagents
for the inactivation of the enzymes in question, they should be
valuable tools for the identification of the residues so tagged
through mass spectrometric analysis of proteolytic digests of
labeled enzymes.
3′-(2,3,4,6-Tetra-O-benzyl-â-D-glucopyranosyl)-2′-propanone (4).
A mixture of alcohols 3a,b (630 mg, 1.08 mmol) was dissolved in
acetone (10 mL), and then Jones reagent (15 g of CrO3 100 mL of
H2O, 24 g of concentrated sulfuric acid) (1.5 mL) was added while
being stirred at 0 °C. After 1 h, excess reagent was reduced using a
minimum amount of 5% sodium metabisulfite and then the mixture
diluted with water (50 mL). The aqueous mixture was extracted with
diethyl ether (3 × 50 mL), and the combined organic fractions were
washed successively with 10% aqueous NaHCO3 (50 mL) and saturated
brine (50 mL). The organic layer was dried (MgSO4) and filtered and
the solvent removed in vacuo. Purification on silica gel (diethyl ether:
petroleum ether (bp 35-60 °C), 2:3) gave the ketone (4) (540 mg,
0.93 mmol, 86%) as a crystalline solid. Mp: 72-73 °C (diethyl ether/
petroleum ether, needles). [R]D: -3° (c 1.4, CHCl3). 1H NMR (400
MHz, CDCl3): δ 7.16-7.40 (m, Ar), 4.80-4.95 (4 H, m, CH2Ar),
4.48-4.64 (4 H, m, CH2Ar), 3.78 (1 H, td, J1,2 9 Hz, J1,3b′ 9.0 Hz, J1,3a′
3.5 Hz, H-1), 3.63-3.72 (4 H, m), 3.44 (1 H, dt, J5,4 9.4 Hz, J5,6a and
J5,6b 2.9 Hz, H-5), 3.31 (1 H, t, J 9.1 Hz), 2.71 (1 H, dd, J3a′,3b′ 15.6
Hz, H-3a′), 2.56 (1 H, dd, H-3b′), 2.13 (3 H, s, Me). 13C NMR (50
MHz, CDCl3): δ 205.5 (C-2′), 138.3 (Ar), 128-130 (Ar), 86.9, 82.5,
78.5, 79.0, 75.5 (5 × CH), 75.0, 74.5, 73.5, 69.5 (5 × CH2, two signals
coincide), 45.5 (C-3′), 21.0 (C-1′). Anal. Calcd for C37H40O6: C,
76.53; H, 6.94. Found: C, 76.50; H, 6.86. CIMS (NH3): m/z 598 (M
+ NH4)+(40%), 581 (M + H)+ (20%).
Experimental Methods
Synthesis. All buffer chemicals and other reagents were obtained
from the Sigma/Aldrich Chemical Co. unless otherwise noted. Reac-
tions were monitored by TLC using Merck Kieselgel 60 F254 aluminum-
backed sheets. Compounds were detected by charring with 10%
ammonium molybdate in 2 M H2SO4 and heating. Flash chromatog-
raphy under a positive pressure was performed with Merck Kieselgel
60 (230-400 mesh) using the specified eluents. 1H NMR spectra were
recorded on a Bru¨ker WH-400 spectrometer at 400 MHz (chemical
shifts quoted relative to CDCl3 or DSS when taken in D2O). 13C NMR
spectra were recorded on a Varian XL-300 at 75 MHz or a Bru¨ker
AC-200 at 50 MHz and are proton-decoupled with CDCl3 or acetone
(30.5 ppm) as a reference.
1′,2′-Epoxy-3′-(2,3,4,6-tetra-O-benzyl-â-D-glucopyranosyl)pro-
pane (2). 3′-(2,3,4,6-Tetra-O-benzyl-â-D-glucopyranosyl)-1′-propene
(1) (2.39 g, 4.24 mmol), m-chloroperbenzoic acid (80%, 1.79 g, 8.32
(28) Lowry, T. H., Richardson, K. S. Mechanism and Theory in Organic
Chemistry; Harper and Row: New York, 1987; pp 381-382.
(29) Bordwell, F. G.; Brannen, W. T. J. Am. Chem. Soc. 1964, 86, 4645-
4654.
(30) Thorpe, J. W.; Warkentin, J. Can. J. Chem. 1973, 51, 927-935.
(31) Pusino, A.; Rosnati, V.; Saba, A. Tetrahedron 1984, 40, 1893-
1899.

Bromoketone C-Glycosides
J. Am. Chem. Soc., Vol. 120, No. 40, 1998 10331
3′-(â-D-Glucopyranosyl)-2′-propanone (5). Ketone 4 (540 mg, 0.93
mmol) was dissolved in 10% acetic acid-methanol (20 mL) and stirred
with 5% Pd-C (10 mg) under hydrogen at room temperature and
pressure. After 16 h, the mixture was filtered, the solvent removed in
vacuo, and the residue purified on silica gel (ethyl acetate:methanol:
water, 15:4:1) to give 5 (180 mg, 0.84 mmol, 90%) as a colorless syrup.
[R]D: -3° (c 1.3, MeOH). 1H NMR (400 MHz, D2O): δ 3.82 (1 H,
dd, J6a,6b 12.2 Hz, J6a,5 1.8 Hz, H-6a), 3.75 (1 H, td, J1,2 and J1,3b′ 9.5
Hz, J1,3a′ 3.1 Hz, H-1), 3.64 (1 H, dd, J6b,5 5.1 Hz, H-6b), 3.45 (1 H, t,
8.7 Hz), 3.31-3.41 (2 H, m), 3.19 (1 H, t, J 9.3 Hz), 2.98 (1 H, dd,
J3a′,3b′ 16.6 Hz, H-3a′), 2.68 (1 H, dd, H-3b′), 2.24 (3 H, Me). CIMS
(NH3): m/z 221 (M + H)+ (100%), 238 (M + NH4)+(30%). HRMS:
(M + H)+, Calcd 221.1020, found 221.1030.
1′-Bromo-3′-(â-D-glucopyranosyl)-2′-propanone (6). Ketone 5
(150 mg, 0.66 mmol) was dissolved in methanol (2 mL), to which was
added bromine (53 µL, 0.99 mmol) as a solution in methanol (0.5 mL).
After 3 h at 40 °C, the solvent was removed in vacuo and the residue
purified on silica gel (ethyl acetate:methanol:water, 17:3:1) to give the
R-bromoketone (6) (106 mg, 0.35 mmol, 53%) as a foamy solid. 1H
NMR (400 MHz, D2O) (assignments confirmed by COSY): δ 4.33 (2
H, s, H-1′), 3.81 (1 H, dd, J6a,6b 12.4 Hz, J6a.5 1.6 Hz, H-6a), 3.77 (1 H,
td, J1,2 and J1,3b′ 9.0 Hz, J1,3a′ 3.4 Hz, H-1), 3.64 (1 H, dd, J6b,5 5.1 Hz,
H-6b), 4.35 (1 H, t, J3,2 and J3,4 9.0 Hz, H-3), 3.33-3.48 (2 H, m, H-5
and H-4), 3.21 (1 H, t, H-2), 3.11 (1 H, dd, J3a′,3b′ 16.5 Hz, H-3a′),
2.87 (1 H, dd, H-3b′). 13C NMR (50 MHz, D2O): δ 204.5 (C-2), 70.2,
73.5, 75.8, 77.6, 80.0 (5 × CH), 61.2 (C-6), 36.9, 42.9 (2 × CH2).
CIMS (NH3): m/z 316 (M + NH4)+ (25%), 318 (M + NH4)+ (25%),
299 (M + H)+ (10%), 301 (M + H)+ (10%). Anal. Calcd for C9H15O6-
Br: C, 36.14; H, 5.05. Found: C, 36.07; H, 5.24.
3′-(â-D-Cellobiosyl)-2′-propanone (7). â-Glucosidase (E358A mu-
tant from Agrobacterium sp.) (0.96 mg) was incubated with 3′-(â-D-
glucopyranosyl)-2′-propanone (5) (150 mg, 0.682 mmol) and R-D-
glucosyl fluoride (62 mg, 0.34 mmol) in sodium phosphate buffer (150
mM, pH 7.0). After 24 h, the enzyme was reclaimed by dialysis using
a centrifugal ultrafilter with a molecular weight cutoff of 10 000
(Millipore). The filtrate was evaporated to dryness in vacuo and the
residue suspended in methanol (4 mL). The methanolic solution was
filtered through a silica plug which was further washed with methanol
(20 mL). The solvent was removed in vacuo, and the residue was
dissolved in methanol (4 mL) and then filtered (2.2 µm filter). The
solvent was removed in vacuo and the residue purified on a Dynamax
reverse phase column using a Waters HPLC system and eluting with
70% acetonitrile in water. Starting material 5 (84 mg, 0.38 mmol)
and â-C-cellobioside 7 (24 mg, 0.063 mmol, 21% based on recovered
staring material) were obtained as colorless syrups. Material with a
longer retention time, presumably trisaccharides, was also recovered
(15 mg). â-C-Cellobioside 7: [R]D: +1.9° (c 1.0, MeOH). 1H NMR
(400 MHz, D2O): δ 4.46 (1 H, d, J1′′,2′′ 8.0 Hz, H-1′′), 3.84-3.91 (2
H, m), 3,66-3.79 (3 H, m), 3.55-3.60 (2 H, m), 3.35-3.59 (4 H, m),
3.21-3.29 (2 H, m). 2.98 (1 H, dd, J3a′.3b′ 17.7 Hz, J3a′,1 2.5 Hz, H-3a′),
2.69 (1 H, dd, J3b′,1 9.2 Hz, H-3b′), 2.23 (3 H, s, Me). 13C NMR/APT
(75 MHz, D2O): δ 213.9 (C-2′), 103.3 (C-1′′), 70.2, 73.6, 73.9, 75.8,
76.2, 76.4, 76.7, 79.1, 79.5 (9 × CH), 60.8, 61.3 (C-6 and C-6′′), 46.3
(C-3′), 31.0 (C-1′). CIMS (NH3): m/z 400 (M + NH4)+ (100%).
HRMS: (M + NH4)+ calcd 400.1810, found: 400.1837.
3′-(2,3,6,2′′,3′′,4′′,6′′-Hepta-O-acetyl-â-D-cellobiosyl)-2′-
propanone (8). A crude sample of the â-C-cellobioside 7 (22 mg,
0.1 mmol) was dissolved in pyridine (1.5 mL) and acetic anhydride
(0.5 mL) and allowed to stand for 16 h. The solvent was removed in
vacuo and the residue purified on silica gel (ethyl acetate:petroleum
ether (bp 35-60 °C), 2:1) to give the acetylated â-C-cellobioside 8
(13 mg, 0.015 mmol, 33%) as a colorless syrup. 1H NMR (400 MHz,
CDCl3, assignments confirmed by COSY): δ 5.14 (1 H, t, J3,2 and J3,4
9.5 Hz, H-3), 5.11 (1 H, t, J3′′,2′′ and J3′′,4′′ 9.7 Hz, H-3′′), 5.04 (1 H, t,
J4′′,5′′ 9.6 Hz, H-4′′), 4.89 (1 H, dd, J2′′,1′′ 8.0 Hz, H-2′′), 4.79 (1 H, t,
J2,1 9.5 Hz, H-2), 4.48 (1 H, d, H-1′′), 4.42 (1 H, dd, J6a,6b 12.0 Hz,
(18 H, 6 × s, 6 × Me), 1.98 (6H, s). CIMS (NH3): m/z 694 (M +
NH4)+ (35%), 677 (M + H)+. HRMS: (M + H)+ calcd 677.2280,
found 677.2291.
1′-Bromo-3′-(â-D-cellobiosyl)-2′-propanone (9). The â-C-cello-
bioside 7 (20 mg, 0.052 mmol) and bromine (5 µL, 0.094 mmol) were
dissolved in methanol (0.5 mL) and allowed to stand at 40 °C for 1 h.
The solvent was removed in vacuo and the residue purified on silica
gel (ethyl acetate:methanol:water, 10:3:1) to give 1′-bromo-3′-(â-D-
cellobiosyl)-2′-propanone (9) (14 mg, 58%) as a colorless syrup.
[R]D: -10.9° (c 0.95, MeOH). 1H NMR (400 MHz, D2O): δ 4.46 (1
H, d, J1′′,2′′ 7.9 Hz, H-1′′), 4.31 (2 H, s, CH2Br), 3.83-3.91 (2 H, m),
3.66-3.80 (3 H, m), 3.55-3.61 (2 H, m), 3.43-3.53 (3 H, m), 3.34-
3.40 (1 H, m), 3.23-3.31 (2 H, m), 3.11 (1 H, dd, J3a′,3b′ 16.5 Hz, J3a′,1
3.1 Hz, H-3a′), 2.87 (1 H, dd, J3b′,1 8.9 Hz, H-3b′). 13C NMR/APT
(75 MHz, D2O): δ 204.2 (C-2), 102.7 (C-1′′), 69.7, 73.1, 73.4, 75.5,
75.7, 75.9, 76.2, 78.5, 78.9 (9 × CH), 60.2, 60.8 (C-6 and C-6′′), 42.6
(C-1′), 36.7 (C-3′). ESMS: m/z 483.5 (M + Na)+ (100%), 485 (M +
Na)+ (100%).
Enzyme Kinetics. Recombinant Agrobacterium sp. â-glucosidase
was purified as described previously.30 Recombinant CenA, CenD,
and Cex were generously donated by Dr. Neil Gilkes from the
Department of Microbiology, University of British Columbia, and were
purified as described previously.1-4,7,32 A continuous spectrophoto-
metric assay based on the hydrolysis of nitrophenyl glycosides was
used to monitor enzyme activity by measurement of the rate of
nitrophenolate release using a UNICAM 8700 UV-visible spectro-
photometer equipped with a circulating water bath.
Inactivation of enzymes by 6 or 9 was monitored by incubation of
the enzyme with various concentrations of the inactivator at 37 °C.
Residual enzyme activity was determined at the appropriate time
intervals by addition of a 10-µL aliquot of the inactivation mixture to
a solution of substrate and measurement of the corresponding nitro-
phenolate release at 37 °C to an extent no greater than 10% substrate
depletion. The same buffer conditions were used for both the
inactivation mixtures and assays. Individual conditions were as
follows: Agrobacterium sp. â-glucosidase, 50 mM sodium phosphate,
pH 6.8, 0.1% BSA, assay 800 µL × 1 mM â-PNPG (Km ) 0.1 mM);
CenA and CenD, 50 mM sodium phosphate, pH 7.0, 0.1% BSA, assay
200 µL × 2 mM â-DNPC (Km[CenA] ) 0.17 mM1, Km[CenD] ) 0.56 mM]);
Cex, same as for CenA/CenD except assayed using 800 µL × 6 mM
â-PNPC (Km ) 0.6 mM). Pseudo-first-order inactivation rate constants
at each inactivator concentration (kobs) were determined by fitting each
curve to a first-order equation using the program GraFit (Leatherbarrow,
R. J. GraFit Version 3.0; Erithacus Software Ltd.: Staines, U.K., 1990).
The values of ki and KI, assuming inactivation according to the kinetic
model shown in Scheme 2, were determined from a direct fit to the
plot of kobs against inactivator concentration.
Electrospray Mass Spectrometry. The analyses of the protein
samples were carried out using a Sciex API-300 mass spectrometer
interfaced with a Michrom UMA HPLC system (Michrom Bioresources,
Inc., Auburn, CA). Enzymes were incubated for 5 h (22 °C) under
the following conditions prior to LC/MS analysis: Agrobacterium sp.
â-glucosidase, 19.5 mM 6; CenA and CenD, 7 mM 9. The enzyme
(10-20 µg, labeled or unlabeled) was introduced into the mass
spectrometer through a microbore PLRP column (1 × 50 mm) and
eluted with a gradient of 20-100% solvent B at a flow rate of 50 µL/
min over 15 min (solvent A: 0.06% trifluoroacetic acid, 2% acetonitrile
in water; solvent B: 0.05% trifluoroacetic acid, 90% acetonitrile in
water). The MS was scanned over a range of 400-2300 Da with a
step size of 0.5 Da and a dwell time of 1 ms.
Acknowledgment. We thank Karen Rupitz for technical
assistance, Lloyd Mackenzie for advice, and Dr. Neil Gilkes
for providing enzyme samples and valuable discussion. We
thank the Protein Engineering Network of Centres of Excellence
of Canada and the Natural Sciences and Engineering Research
Council of Canada for financial support.
J6a,5 1.8 Hz, H-6a), 4.34 (1 H, dd, J6a′′,6b′′ 12.4 Hz, J6a′′,5′′ 4.4 Hz, H-6a′′),
4.06 (1 H, dd, J6b,5 5.3 Hz, H-6b), 4.03 (1 H, dd, J6b′′,5′′ 2.1 Hz, H-6b′′),
3.89 (1 H, td, J1,3a′ 9.5 Hz, J1,3b′ 3.1 Hz, H-1), 3.70 (1 H, t, J4,5 9.5 Hz,
H-4), 3.63 (1 H, ddd, H-5′′), 3.57 (1 H, ddd, H-5), 2.63 (1 H, dd, J3a′,3b′
16.3 Hz, H-3a′), 2.42 (1 H, dd, H-3b′), 2.13, 2.07, 2.06, 2.0, 1.99, 1.95
JA981580R
(32) Gilkes, N. R.; Langsford, M. L.; Kilburn, D. G.; Miller, R. C., Jr.;
Warren, R. A. J. J. Biol. Chem. 1984, 259, 10455-10459.