Protein glycoengineering enabled by the versatile synthesis of aminooxy glycans and the genetically encoded aldehyde tag

Article abstract of DOI:10.1021/ja206023e

Homogeneously glycosylated proteins are important targets for fundamental research and for biopharmaceutical development. The use of unnatural protein-glycan linkages bearing structural similarity to their native counterparts can accelerate the synthesis of glycoengineered proteins. Here we report an approach toward generating homogeneously glycosylated proteins that involves chemical attachment of aminooxy glycans to recombinantly produced proteins via oxime linkages. We employed the recently introduced aldehyde tag method to obtain a recombinant protein with the aldehyde-bearing formylglycine residue at a specific site. Complex aminooxy glycans were synthesized using a new route that features N-pentenoyl hydroxamates as key intermediates that can be readily elaborated chemically and enzymatically. We demonstrated the method by constructing site-specifically glycosylated variants of the human growth hormone.

Full text of DOI:10.1021/ja206023e

ARTICLE
pubs.acs.org/JACS
Protein Glycoengineering Enabled by the Versatile Synthesis of
Aminooxy Glycans and the Genetically Encoded Aldehyde Tag
Jason E. Hudak, Helen H. Yu, and Carolyn R. Bertozzi*
Departments of Chemistry and Molecular and Cell Biology and Howard Hughes Medical Institute, University of California, Berkeley,
California, 94720
S Supporting Information
b
ABSTRACT: Homogeneously glycosylated proteins are important
targets for fundamental research and for biopharmaceutical develop-
ment. The use of unnatural proteinꢀglycan linkages bearing struc-
tural similarity to their native counterparts can accelerate the synthesis
of glycoengineered proteins. Here we report an approach toward
generating homogeneously glycosylated proteins that involves chemi-
cal attachment of aminooxy glycans to recombinantly produced
proteins via oxime linkages. We employed the recently introduced
aldehyde tag method to obtain a recombinant protein with the
aldehyde-bearing formylglycine residue at a specific site. Complex
aminooxy glycans were synthesized using a new route that features
N-pentenoyl hydroxamates as key intermediates that can be readily elaborated chemically and enzymatically. We demonstrated the
method by constructing site-specifically glycosylated variants of the human growth hormone.
’ INTRODUCTION
We and others have explored the oxime-forming reaction of
aminooxy glycans with peptide-associated aldehydes or ketones
as a means of site-specific glycosylation.25,28,29 This chemistry
proceeds in mildly acidic aqueous buffers that are compatible
with most proteins and poses little risk of side reactivity with
endogenous protein or glycan functionalities.30,31 Despite pro-
mising initial work, however, the generality of the approach
remains limited by two challenges: (1) the need for a practical
means of introducing aldehydes or ketones into recombinant
proteins using standard expression systems, and (2) a reliable
route to synthesize higher-order aminooxy glycans.
The recently developed aldehyde tag methodology answers
the first challenge, as it enables site-specific introduction of
aldehyde-bearing formylglycine (fGly) residues into any recom-
binant protein of interest using conventional Escherichia coli32
or mammalian33 expression systems. The aldehyde tag is a five-
residue consensus sequence (CxPxR, where x is variable) that is
recognized by the formylglycine generating enzyme (FGE).34ꢀ36
Cotranslationally, FGE oxidizes the genetically encoded cysteine
residue to fGly, thereby providing a uniquely reactive site for
chemical modification. Although FGE is native to most cell types,
coexpression of additional FGE alongside a recombinant protein
substrate ensures high Cys-to-fGly conversion yields. Use of the
aldehyde tag method simply requires cloning the consensus
sequence into the protein’s gene at the desired site of modifica-
tion, followed by protein production in FGE-expressing cells.
In addition to its practicality, the aldehyde tag method bene-
fits from the resemblance of the fGly side chain to naturally
Precise molecular control of protein glycosylation would advance
the development of biopharmaceuticals as well as fundamental
studies of glycosylation-dependent biological processes.1ꢀ4 Bio-
logical methods on their own have failed to achieve this long
sought-after goal, as cellular biosynthetic pathways produce glyco-
proteins as a heterogeneous mixture of glycoforms. Glycoprotein
structures with optimal bioactivity are difficult to identify from
such complex mixtures and impossible to generate exclusively
using conventional cell hosts. The tools of synthetic biology have
been harnessed to construct heterologous glycan biosynthetic
pathways in simple host cells.5,6 While these systems have
produced glycoproteins that are globally modified with a single
glycan type, no recombinant expression system can yet pro-
duce proteins with discrete glycans at individual sites and with
complete homogeneity.
Chemical synthesis is currently the only plausible approach to
achieve homogeneously glycosylated proteins bearing tailored
glycans at specific sites.7ꢀ10 Several groups have successfully
combined frontier protein ligation techniques with chemical or
enzymatic glycan and glycopeptide synthesis methods to con-
struct glycoproteins that duplicate natural structures.11ꢀ13 The
valor of these efforts should not be underestimated since the
difficulties inherent to glycan and protein syntheses amplify each
other when both biopolymers coexist in the same synthetic target.14,15
A more practical, though less biologically authentic, approach
involves conjugating glycans to recombinant proteins via un-
natural but synthetically tractable linkages.16ꢀ18 Site-specificity
in the modification can be ensured by outfitting the protein with
a noncanonical amino acid side chain capable of chemoselective
ligation with an appropriately functionalized glycan.19ꢀ27
Received: June 28, 2011
Published: August 25, 2011
r
2011 American Chemical Society
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would require protection of the aminooxy group to avoid oxime
formation with the latent aldehydes or ketones associated with
free sugars. The lability of the NHS and NHPht groups under-
mined our efforts to deprotect late-stage glycan intermediates
without loss of the aminooxy protecting group.
Therefore, we sought to identify an alternative aminooxy
protecting group that possessed better chemical stability overall
but was readily removable under mild conditions that are compatible
with free sugars. Ideally, the nucleophilicity of the masked aminooxy
group would be superior to that of the N-hydroxyimides as well,
thus enhancing reactivity as a glycosyl acceptor. After screening a
variety of options, we found the N-pentenoyl protecting group to
be an excellent candidate. Originally introduced by Fraser-Reid
and co-workers,40ꢀ42 the pentenoyl amide is stable to many reac-
tion conditions and is removed by gentle treatment with aqueous
iodine. Despite its potential as a synthetic substrate, N-pentenoyl
hydroxamic acid (1, Table 1, also termed N-hydroxypenten-
amide, which we abbreviate NHPent) has seldom been utilized
in synthesis43 and, to our knowledge, never as a masked
aminooxy group.
To determine the scope of compound 1's reactivity as a
glycosyl acceptor, we subjected it to glycosylation reactions using
donor substrates that included glycosyl bromides and fluorides,
thioglycosides, and glycosyl acetimidates (Table 1). Most sub-
strates gave acceptable yields of NHPent glycoside product.
The highest yields were obtained using glycosyl N-phenyl
trifluoroacetimidates,44ꢀ46 as previously observed in the syn-
theses of glycosyl amides44 and hydroxamates.47,48 By contrast,
the less stable glycosyl trichloroacetimidates gave low yields in
glycosylation reactions with 1 under similar conditions (data not
shown).
We anticipated that the mild conditions reported for cleavage
of N-pentenoyl amides would be compatible with unprotected
glycans. To confirm that such conditions can cleave N-alkoxy-
pentenamides without harm to free sugars, we performed model
reactions on the test substrate N-pentenoyl aminooxy lactose
(15, prepared from the known disaccharide aminooxy lactose28
as shown in Scheme S1). Treatment of N-pentenoyl aminooxy
lactose with 3 equiv of I2 in 1:1 tetrahydrofuran (THF)/H2O as
reported41 resulted in iodocyclization of the NHPent group,
but hydrolysis of the iodoimidate intermediate to afford the
free aminooxy disaccharide was sluggish. Fortunately, the use of
nonaqueous solvent mixtures and acid catalysis, as shown
previously,49,50 alleviated this problem. We identified the optimal
deprotection conditions to be 3 equiv of I2 in wet MeCN/MeOH
with 0.1% formic acid (FA) at room temperature, which afforded
quantitative deprotection of N-pentenoyl aminooxy lactose in
under 1 h.
Chemical Synthesis of Aminooxy Lewis x. The utility of
NHPent glycosides as substrates for complex aminooxy glycan
synthesis was demonstrated with the construction of the tri-
saccharide Lewis x (Galβ1,4(Fucα1,3)GlcNAc), a leukocyte and
stem cell marker (Scheme 1).51,52 We began the synthesis with
the coupling of known trichloroacetimidate 16 to thioglycoside
17 to generate the lactosamine precursor 18 following related,
published routes.53,54 As seen with similarly protected glucosa-
mine derivatives,53,54 only glycosylation at the more reactive
4-hydroxyl of 17 was observed. The fucose residue was intro-
duced in the form of 19, which was outfitted with a p-methoxy-
benzyl (PMB) ether at the 2-position to enhance cis-glycosylation.
Similar to a previous report,55 TfOH-catalyzed coupling of 19
with the free C-3 hydroxyl of 18 proceeded smoothly to afford 20.
Figure 1. Natural glycan linkages resemble the linkage produced by
protein glycoengineering via the reaction of aminooxy sugars with the
aldehyde tag. (A) Linkages naturally found in the most abundant
mammalian N- and O-linked glycan structures. (B) The linkage formed
upon the reaction of an aminooxy glycan with the aldehyde of an fGly-
containing protein. The resulting oxime side chain bears high structural
similarity to the natural glycoprotein attachments.
glycosylated amino acid side chains. Most secreted and cell-
surface vertebrate glycoproteins are modified at Asn residues
with N-glycans or at Ser/Thr residues with O-glycans (Figure 1A).
We envisioned oxime-linked glycoproteins deriving from con-
jugation of the fGly aldehyde with synthetic aminooxy glycans as
illustrated in Figure 1B. Glycanꢀprotein linkages comprising
fGly-derived oximes are close structural mimics of N-glycan
linkages and just one atom longer than O-glycan linkages.
To capitalize on the aldehyde tag as a tool for site-specific
protein glycosylation, we sought to answer the secondchallenge—
a practical and general route to synthesize complex aminooxy
glycans. Here we report on the use of glycosyl N-pentenoyl
hydroxamates as intermediates in the synthesis of higher-order
aminooxy glycans. These analogues function well in both chemi-
cal and enzymatic glycosylation reactions, enabling the efficient
and high-yielding production of both simple and elaborated
aminooxy glycans as substrates for protein conjugation. The
new synthetic approach, together with the aldehyde tag technol-
ogy, enabled the generation of recombinant proteins modified
site-specifically with homogeneous glycans.
’ RESULTS AND DISCUSSION
Evaluation of N-Hydroxypentenamide as a Masked Amino-
oxy Group. In previous work, aminooxy glycosides were synthe-
sized by displacement of a glycosyl halide with N-hydroxysuccin-
imide (NHS) or N-hydroxyphthalimide (NHPht).29,37,38 Reaction
yields were acceptable using monosaccharide substrates but, in
our hands, fell precipitously with increasing glycan complexity.
Additionally, glycosylations of NHS and NHPht with other do-
nors, such as glycosyl acetimidates and thioglycosides, were
low yielding, even with monosaccharide substrates. In addition
to falling short as glycosyl acceptors, NHS and NHPht were
inadequate aminooxy protecting groups. They were labile to
basic and reductive conditions, which undermined their use
during complex glycan synthesis. Moreover, we envisioned using
one-pot multienzyme methods39 to convert simple aminooxy
glycans into more complex structures. However, such reactions
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Table 1. Reaction of Various Glycosyl Donors with
Scheme 1. Chemical Synthesis of Aminooxy Lewis x
N-Pentenoyl Hydroxamic Acid 1
a Reactions corresponding to entries 1ꢀ7 were conducted in CH2Cl2
(unless specified) at 0 °C, and those corresponding to entries 8ꢀ13 were
performed at ꢀ20 °C. b Isolated yields after column chromatography.
c Ratio determined by isolated yields or 1H NMR spectroscopy.
with the pentenoyl and thioglycoside groups, and the 2-hydroxyl
group of the fucose residue was acetylated to afford 21.
Motivated by the results in Table 1, we converted thioglycoside
21 to the corresponding glycosyl N-phenyl trifluoroacetimidate,
The PMB group was removed with 10% trifluoroacetic acid, thereby
avoiding reductive or oxidative conditions that are incompatible
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Scheme 2. Enzymatic Sialylation of N-Pentenoyl Aminooxy Lactose
Scheme 3. Deprotection of NHPent on Sialosides
22, for coupling with compound 1.46 The glycosylation reaction
was conducted with 1 equiv of TMSOTf in dichloromethane at
ꢀ20 °C to afford N-hydroxypentenoyl trisaccharide 23. The
Troc protecting group was removed with Zn in acidic MeCN,
and the amino group was acetylated. Notably, we observed no
reduction of the NꢀO bond during the Zn reduction step, in
sharp contrast to our previous experience with NHPth glyco-
sides. Removal of the TBDPS and acetyl groups of 24 proceeded
in high yield and without damage to the N-hydroxypentenamide
as well. Finally, aminooxy Lewis x was obtained by treatment of
25 with I2 in acidic MeCN/MeOH.
Chemoenzymatic Synthesis of Complex Aminooxy Glycans.
Enzymatic elaboration of synthetic core structures is an increas-
ingly popular strategy for complex glycan assembly.39,56 Enzymes
provide tight stereo- and regiochemical control and obviate the
need for intermediate protecting group manipulations and harsh
chemical conditions. The impact of enzymatic glycosylation
methods is most evident in cases where the chemical counterpart
is notoriously difficult, such as in the addition of terminal sialic
acid residues to a complex glycan.57 Given the biological importance
of sialosides,58 we sought to construct sialylated aminooxy glycans
as substrates for glycoprotein engineering.
Trisaccharides 29 and 30 (Scheme 3), termed aminooxy 2,6-
and 2,3-sialyllactose, respectively, were chosen as initial targets
with which to evaluate the performance of NHPent glycosides as
substrates for enzymatic sialylation. We used the one-pot, three-
enzyme system introduced by Chen and co-workers59,60 to append
sialic acid to N-pentenoyl aminooxy lactose (15, Scheme 2). This
methodology allows for the in situ generation of costly cytidine
50-monophosphate (CMP)-sialic acid from ManNAc, pyru-
vate, and cytidine 50-triphosphate (CTP) by the combined use
of E. coli aldolase and a CMP-sialic acid synthase, NmCSS.61
In the same pot, bacterial α-2,6-sialyltransferase (Pd2,6ST) or
α-2,3-sialyltransferase (PmST1) transferred sialic acid to 15,
affording NHPent glycosides 27 and 28, respectively, in excellent
yields. Notably, free aminooxy glycans would not be viable
substrates for this procedure, as the aminooxy group would undergo
side reactions with free reducing sugars as well as pyruvate. Finally,
we employed our previously optimized deprotection conditions to
obtain trisaccharides 29 and 30 (Scheme 3).
A higher level of complexity was embodied in the tetrasaccharide
sialyl Lewis x (Siaα2,3Galβ1,4(Fucα1,3)GlcNAc, abbreviated
SLex), a glycan known for its involvement in cancer progression
and leukocyte homing.62,63 We envisioned generating aminooxy-
functionalized SLex by chemical synthesis of an NHPent-modified
disaccharide core, followed by enzymatic installation of the terminal
sialic acid and fucose residues. Toward this end, compound 18
was transformed into NHPent lactosamine derivative 33 by
acetylation of its one free hydroxyl group, conversion of the
thioglycoside to the N-phenyl trifluoroacetimidate, and glyco-
sylation with compound 1 (Scheme 4). Deprotection and
N-acetylation gave N-acetyllactosamine NHPent glycoside 35
as a substrate for enzymatic glycosylation. The α-2,3-sialoside 36
was generated enzymatically as described above for compound
28. After purification by size exclusion chromatography, the
fucose residue was installed using the one-pot, two-enzyme
system described by Wu and co-workers.64 Briefly, guanosine
50-diphosphate (GDP)-fucose was generated in situ from adeno-
sine triphosphate (ATP), guanosine 50-triphosphate (GTP), and
fucose by the Bacteroides fragilis fucose kinase phosphorylase
(FKP). Subsequent glycosylation using Helicobacter pylori α-1,3-
fucosyltransferase (Hp1,3FT) generated tetrasaccharide 37. Fi-
nally, deprotection of 37 with I2 in acidic MeCN/MeOH gave
aminooxy SLex 38 in 65% isolated yield.
Conjugation of Aminooxy Glycans to Aldehyde-Tagged
Human Growth Hormone. With a route to complex aminooxy
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Scheme 4. Chemoenzymatic Synthesis of Aminooxy SLex
route to improving its pharmacokinetic properties.67,68 Indeed,
an artificially glycosylated hGH variant was recently shown to
have a prolonged serum half-life in mice.69
We produced aldehyde-tagged hGH with >95% Cys-to-fGly
conversion in E. coli with coexpression of the FGE from
Mycobacterium tuberculosis.32 The protein was purified by chro-
matography on Ni-NTA agarose and then reacted with aminooxy
lactose (S1) in 5% aqueous MeCN with 0.02% FA (pH ∼3.8).
The formation of the lactose glycoconjugate was confirmed by
Western blot probing with the galactose-binding lectin, soybean
agglutinin conjugated to fluorescein isothiocyanate (SBA-FITC)
(Figure 2B). Significant SBA reactivity was observed only for
products of aldehyde-tagged protein conjugation. A control
conjugation reaction using an hGH construct in which the Cys
residue required for the aldehyde installation was mutated to Ala
(CfA) produced no SBA-reactive product. This result con-
firmed the necessity of the Cys-derived fGly residue for site-
specific chemical glycosylation with aminooxy lactose. Further,
we analyzed the oxime conjugate by electrospray ionization mass
spectrometry (ESI-MS) (Figure 2C). Ions corresponding to
glycosylated (hGH+Lac) and unreacted (hGH) aldehyde-tagged
hGH were observed, the relative intensities of which indicated a
reaction yield of ∼81%. The protein glycoconjugate was found to
be highly stable when stored at ꢀ20 °C, and <10% hydrolysis of
the conjugate was observed after 8 days at 4 °C (Table S1).
We performed a similar reaction using the more complex
glycan aminooxy SLex (38). The formation of the glycosylated
hGH product was confirmed by Western blot probing with a
SLex-specific antibody as well as with the fucose-binding lectin,
Aleuria aurantia lectin (AAL), conjugated to biotin (Figure 2D).
Using the above conjugation conditions (5% MeCN, 0.02% FA),
the yield of glycosylated hGH as determined by ESI-MS was a
modest 20ꢀ25%. However, use of a sodium citrate (NaCit)
buffer at pH 3.5 improved the conjugation efficiency to ∼64%.
Despite the acidity of the reaction buffer, we did not detect any
loss of the potentially labile sialic acid and fucose moieties or any
damage to the aldehyde-tagged protein.
’ CONCLUSION
In summary, the facile generation of homogeneous glycopro-
teins was made possible by an improved route to aminooxy
glycans featuring glycosyl N-pentenoyl hydroxamate intermedi-
ates, combined with the genetically encoded aldehyde tag. The
versatility of the aldehyde tag with respect to expression system,
protein target, and site of aldehyde placement makes this
approach broadly useful for protein glycoengineering. In princi-
ple, multiple aldehyde tags can be introduced into a single
protein, allowing for the production of multivalent glycoforms,
a work currently in progress.
The method joins a growing toolkit for site-specific modifica-
tion of proteins with synthetic glycans, which combined could
produce myriad protein glycoforms with discrete glycans at
different positions. Other chemoselective ligation reactions, such
as azideꢀalkyne cycloadditions and disulfide exchange reactions,
have proven useful for modifying a single protein with multiple
moieties.19 Oxime formation is orthogonal to these and other
chemoselective ligation reactions (thiolꢀene additions, olefin
metathesis, Suzuki couplings, etc.). Exploring the tandem and
parallel use of these reactions for multiplexed protein modifica-
tion is a subject of future interest.
glycans established, we sought to combine the synthetic sugars
with recombinant aldehyde-tagged proteins as shown schemati-
cally in Figure 2A. As a protein substrate, we used a previously
reported construct of the human growth hormone (hGH) that
contains a C-terminal aldehyde tag as well as an N-terminal His
tag. The native protein is a well-established biotherapeutic
used in the treatment of human developmental diseases among
others.65,66 Site-specific chemical modification of hGH is a potential
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Figure 2. Generation of site-specifically glycosylated hGH. (A) Aldehyde-tagged hGH was reacted with synthetic aminooxy glycans to yield the oxime-
glycosylated conjugates. (B) Western blot of aldehyde-tagged hGH (lane 1) or the CfA mutant (lane 2) after reaction with aminooxy lactose S1 (5%
MeCN, 0.02% FA, 0.26 mg/mL aldehyde-tagged hGH, 1 mM S1, 16 h, 37 °C). The blot was probed with SBA-FITC (top), and total protein was
detected by Ponceau stain (bottom). (C) ESI-MS spectrum of the crude reaction mixture showing ions in the 20+ and 21+ charge states. Ions
corresponding to aldehyde-tagged hGH (hGH) and glycosylated hGH (hGH+Lac) are shown with arrows. (D) Western blot of aldehyde-tagged hGH
(lane 1) or the CfA mutant (lane 2) after reaction with aminooxy SLex 38 (50 mM NaCit, pH 3.5, 0.26 mg/mL aldehyde-tagged hGH, 1 mM 38, 16 h,
37 °C). The blot was probed with an anti-SLex antibody KM93 followed by a horseradish peroxidase (HRP)-conjugated secondary antibody (top), and
then stripped and reprobed with AAL-biotin followed by a FITC-conjugated antibiotin antibody (middle). Total protein loading was confirmed by
Ponceau staining (bottom). (E) ESI-MS spectrum of the SLex crude reaction mixture showing ions in the 20+ and 21+ charge states. Ions corresponding
to aldehyde-tagged hGH (hGH) and glycosylated hGH (hGH+SLex) are shown with arrows.
’ EXPERIMENTAL SECTION
Hex/EtOAc), and N-bromosuccinimide was added in 2 equiv portions
until the reaction was complete. Upon full hydrolysis, the resulting
hemiacetal was diluted in CH2Cl2 and washed sequentially with
Na2S2O3 solution, NaHCO3 solution, and brine. The organic layer
was dried over Na2SO4 and concentrated under reduced pressure. The
resulting residue was dissolved in CH2Cl2 (3 mL) and Cs2CO3 (150 mg,
0.44 mmol) was added, followed by N-phenyl trifluoroacetimidoyl
chloride (72 μL, 0.44 mmol) dropwise. The reaction mixture was stirred
for 12 h at room temperature under N2, after which it was diluted with
CH2Cl2, filtered, and concentrated under vacuum. The crude residue
was purified by silica chromatography to give the trifluoroacetimidate
Phenyl (2,3,4,6-Tetra-O-benzoyl-β-D-galactopyranosyl)-
(1f4)-6-O-(tert-butyldiphenylsilyl)-3-O-acetyl-2-deoxy-2-(2,
2,2-trichloroethoxy)carbonylamino-1-thio-β-D-glucopyrano-
side (31). A solution of disaccharide 18 (400 mg, 0.32 mmol) in
pyridine/Ac2O (2:1, v:v; 4.5 mL) was stirred for 3 h at room tempera-
ture under N2. Upon completion, the reaction was concentrated and
coevaporated with toluene to remove excess Ac2O and AcOH. The
crude residue was purified by silica flash chromatography (Hex/EtOAc
gradient) to obtain fully protected disaccharide 31 as a white powder
(400 mg, 97%): 1H NMR (500 MHz, CDCl3) δ 8.18 (d, J = 7.4 Hz, 2H),
8.07 (d, J = 7.4 Hz, 2H), 7.93ꢀ7.79 (m, 6H), 7.68ꢀ7.49 (m, 12H),
7.47ꢀ7.34 (m, 5H), 7.33ꢀ7.19 (m, 5H), 7.09 (t, J = 7.5 Hz, 2H), 7.03
(t, J = 7.3 Hz, 1H), 6.85 (t, J = 7.7 Hz, 2H), 6.69 (d, J = 9.0 Hz, 1H), 6.02
(d, J = 2.9 Hz, 1H), 5.81 (dd, J = 10.1, 8.2 Hz, 1H), 5.58 (dd, J = 10.4, 3.1
Hz, 1H), 5.50 (t, J = 9.8 Hz, 1H), 5.27 (d, J = 8.0 Hz, 1H), 5.11 (d, J =
10.4 Hz, 1H), 4.90 (q, J = 12.1 Hz, 2H), 4.74 (dd, J = 11.2, 4.6 Hz, 1H),
4.55ꢀ4.39 (m, 2H), 4.30ꢀ4.23 (m, 1H), 4.09 (q, J = 10.2 Hz, 1H), 3.95
(d, J = 11.2 Hz, 1H), 3.77 (d, J = 11.0 Hz, 1H), 3.40 (d, J = 9.7 Hz, 1H),
2.24 (s, 3H), 1.09 (s, 9H); 13C NMR (126 MHz, CDCl3) δ 171.54,
166.00, 165.56, 165.32, 164.45, 154.59, 136.05, 135.67, 135.29, 133.70,
133.52, 133.33, 133.17, 132.91, 131.69, 130.86, 130.25, 129.84, 129.77,
129.74, 129.51, 129.13, 128.77, 128.69, 128.41, 128.29, 128.22, 128.16,
127.78, 126.74, 100.10, 95.78, 85.76, 78.61, 77.41, 74.52, 74.18, 73.61,
72.02, 71.43, 69.90, 68.22, 61.95, 60.96, 54.87, 26.76, 21.24, 19.35;
HRMS (ESI) calcd for C67H65NO16Cl3SSi [M+H]+ m/z = 1304.2853,
found 1304.2883.
1
32 as a yellow oil (267 mg, 87%): H NMR (500 MHz, CDCl3) δ
8.16ꢀ8.03 (m, 4H), 7.87ꢀ7.68 (m, 7H), 7.68ꢀ7.31 (m, 14H), 7.19
(dddd, J = 28.0, 16.4, 10.1, 3.9 Hz, 7H), 6.98 (dt, J = 30.6, 15.6 Hz, 1H),
6.71 (dd, J = 23.5, 7.7 Hz, 1H), 6.00ꢀ5.92 (m, 1H), 5.83ꢀ5.73 (m, 1H),
5.60ꢀ5.40 (m, 1H), 5.33 (t, J = 9.9 Hz, 1H), 5.25 (dd, J = 22.4, 14.4 Hz,
1H), 5.18 (dt, J = 19.0, 9.4 Hz, 1H), 4.76 (dt, J = 28.6, 14.3 Hz, 1H),
4.63ꢀ4.51 (m, 2H), 4.47ꢀ4.35 (m, 1H), 4.29ꢀ4.17 (m, 1H), 4.13 (dt,
J = 13.7, 7.1 Hz, 1H), 4.05ꢀ3.69 (m, 2H), 3.58ꢀ3.26 (m, 1H),
2.20ꢀ1.92 (m, 3H), 1.16ꢀ1.02 (m, 9H); 13C NMR (126 MHz, CDCl3)
δ 173.12, 166.12, 165.51, 165.50, 164.72, 154.46, 136.03, 135.47, 133.88,
133.84, 133.62, 133.45, 133.35, 130.54, 130.02, 129.92, 129.88, 129.70,
129.58, 129.17, 129.03, 128.87, 128.73, 128.55, 128.44, 128.36, 127.96,
100.52, 95.48, 74.75, 73.37, 73.24, 72.20, 71.66, 69.90, 68.23, 62.14,
60.53, 54.23, 26.90, 20.97, 19.50. A mixture of trifluoroacetimidate 32
(257 mg, 0.186 mmol) and N-pentenoyl hydroxamic acid 1 (32 mg, 0.28
mmol) was dried by coevaporation with anhydrous toluene and left
under high vacuum. To the dried mixture was added 3 Å MS with stirring
in CH2Cl2 (4 mL) for 1 h at room temperature under N2. The solution
was cooled to ꢀ20 °C, and TMSOTf (34 μL, 0.19 mmol) was added
dropwise. The reaction was allowed to warm to room temperature over
1.5 h and was quenched with triethylamine, filtered, and concentrated
under vacuum. The resulting residue was purified by silica flash
chromatography (toluene/acetone gradient) to obtain N-hydroxypen-
tenoyl disaccharide 33 as a white solid (160 mg, 66%): 1H NMR (500 MHz,
N-Hydroxypent-4-enamide (2,3,4,6-Tetra-O-benzoyl-β-D-
galactopyranosyl)-(1f4)-6-O-(tert-butyldiphenylsilyl)-3-O-
acetyl-2-deoxy-2-(2,2,2-trichloroethoxy)carbonylamino-β-
D-glucopyranoside (33). Thioglycoside 31 (290 mg, 0.22 mmol) was
dissolved in acetone/CH2Cl2/H2O (15:1:1, v:v; 6 mL), and N-iodosuccin-
imide (NIS) (75 mg, 0.33 mmol) was added while stirring at room temperature.
The reaction was monitored by thin-layer chromatography (TLC) (2:1,
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CDCl3) δ 8.45 (s, 1H), 8.15ꢀ8.02 (m, 4H), 7.83ꢀ7.74 (m, 4H),
7.74ꢀ7.65 (m, 4H), 7.60 (dt, J = 21.6, 6.4 Hz, 2H), 7.54ꢀ7.36 (m,
12H), 7.25ꢀ7.20 (m, 2H), 7.20ꢀ7.16 (m, 2H), 5.95 (d, J = 3.2 Hz, 1H),
5.82ꢀ5.69 (m, 2H), 5.69ꢀ5.61 (m, 1H), 5.48 (dd, J = 10.3, 3.2 Hz, 1H),
5.20 (t, J = 9.3 Hz, 1H), 5.12 (d, J = 8.0 Hz, 1H), 5.01 (d, J = 16.5 Hz,
1H), 4.96 (d, J = 10.3 Hz, 1H), 4.86 (d, J = 12.0 Hz, 1H), 4.68 (t, J = 9.1
Hz, 2H), 4.53 (d, J = 6.4 Hz, 2H), 4.34 (t, J = 9.3 Hz, 1H), 4.17 (t, J = 6.0
Hz, 1H), 3.94ꢀ3.77 (m, 3H), 3.27 (d, J = 9.4 Hz, 1H), 2.35ꢀ2.15 (m,
4H), 2.12 (s, 3H), 1.08 (s, 9H); 13C NMR (126 MHz, CDCl3) δ 171.32,
170.85, 166.00, 165.51, 165.43, 165.21, 164.72, 136.58, 136.08, 136.06,
136.02, 135.95, 135.48, 135.44, 133.81, 133.51, 133.46, 133.43, 133.29,
130.54, 130.15, 130.09, 130.04, 129.98, 129.93, 129.85, 129.65, 129.49,
128.91, 128.83, 128.68, 128.64, 128.60, 128.45, 128.41, 128.32, 128.11,
127.85, 127.77, 115.81, 100.22, 95.41, 75.63, 74.87, 74.71, 73.14, 71.91,
71.50, 69.83, 68.08, 62.01, 60.64, 54.00, 53.53, 28.98, 26.86, 20.95, 19.42;
HRMS (ESI) calcd for C66H67N2O18Cl3Si [M+H]+ m/z = 1309.3296,
found 1309.3315.
N-Hydroxypent-4-enamide (2,3,4,6-Tetra-O-benzoyl-β-D-
galactopyranosyl)-(1f4)-6-O-(tert-butyldiphenylsilyl)-3-O-
acetyl-2-deoxy-2-acetamido-β-D-glucopyranoside (34). To a
solution of N-hydroxypentenoyl disaccharide 33 (140 mg, 0.11 mmol)
in 10% FA/MeCN (3 mL) was added activated Zn (500 mg), and the
mixture was stirred for 3 h. The suspension was filtered to remove
catalyst, concentrated under vacuum, and coevaporated with toluene to
remove excess FA. The crude free amine was dissolved in anhydrous
MeOH (3 mL), followed by dropwise addition of Ac2O (55 μL, 0.54
mmol) and N,N-diisopropylethylamine (DIPEA) (22 μL, 0.13 mmol) at
0 °C under N2. After 1.5 h, the reaction was allowed to warm to room
temperature, diluted with MeOH, and concentrated under vacuum. The
resulting residue was purified by silica flash chromatography (toluene/acetone
gradient) to give the N-acetylated disaccharide 34 as a white solid (97 mg,
77%): 1H NMR (500 MHz, CDCl3) δ 9.32 (s, 1H), 8.13ꢀ8.03 (m, 4H),
7.82ꢀ7.73 (m, 4H), 7.72ꢀ7.67 (m, 2H), 7.66ꢀ7.54 (m, 6H),
7.52ꢀ7.45 (m, 8H), 7.41ꢀ7.37 (m, 2H), 7.23 (t, J = 7.8 Hz, 3H),
7.17 (t, J = 7.8 Hz, 2H), 6.19 (d, J = 8.4 Hz, 1H), 5.96 (d, J = 3.1 Hz, 1H),
5.82ꢀ5.69 (m, 2H), 5.48 (dd, J = 10.4, 3.4 Hz, 1H), 5.11 (dd, J = 17.2,
7.9 Hz, 2H), 5.00 (dd, J = 17.2, 1.3 Hz, 1H), 4.94 (d, J = 10.2 Hz, 1H),
4.60 (d, J = 8.5 Hz, 1H), 4.58ꢀ4.47 (m, 2H), 4.36 (t, J = 9.1 Hz, 1H),
4.20ꢀ4.10 (m, 2H), 3.88 (q, J = 11.2 Hz, 2H), 3.26 (d, J = 9.4 Hz, 1H),
2.38ꢀ2.15 (m, 4H), 2.14 (s, 3H), 2.02 (s, 3H), 1.06 (s, 9H); 13C NMR
(126 MHz, CDCl3) δ172.53, 171.81, 170.88, 166.04, 165.46, 164.88, 137.93,
136.75, 136.06, 135.74, 135.43, 133.81, 133.56, 133.43, 133.40, 131.23,
130.58, 130.15, 129.94, 129.89, 129.85, 129.54, 129.11, 128.98, 128.82,
128.73, 128.65, 128.52, 128.41, 128.30, 128.26, 127.78, 125.38,
115.62, 102.03, 100.27, 75.89, 72.99, 72.35, 71.86, 71.59, 69.83, 68.17,
62.07, 60.56, 52.23, 32.87, 29.04, 26.82, 23.35, 21.53, 21.03, 19.41;
HRMS (ESI) calcd for C65H68N2O17Si [M+H]+ m/z = 1177.4360,
found 1177.4396.
4.75 (d, J = 8.8 Hz, 1H), 4.44 (d, J = 7.8 Hz, 1H), 3.98ꢀ3.92 (m, 1H),
3.92ꢀ3.80 (m, 3H), 3.77ꢀ3.66 (m, 5H), 3.63 (dd, J = 10.0, 3.4 Hz, 1H),
3.61ꢀ3.54 (m, 1H), 3.50 (dd, J = 9.8, 7.9 Hz, 1H), 2.32 (dd, J = 13.1, 6.5
Hz, 2H), 2.28ꢀ2.21 (m, 2H), 2.02 (s, 3H); 13C NMR (126 MHz,
CDCl3) δ 174.72, 172.85, 136.46, 115.89, 103.79, 102.82, 77.78, 75.33,
75.06, 72.45, 72.22, 70.91, 68.51, 61.00, 59.77, 52.90, 31.72, 28.81, 22.18;
HRMS (ESI) calcd for C19H32N2O12 [M+H]+ m/z = 481.2028, found
481.2036.
N-Hydroxypent-4-enamide (5-Acetamido-3,5-dideoxy-D-
glycero-α-D-galacto-2-nonulopyranosylonic acid)-(2f3)-β-
D-galactopyranosyl-(1f4)-2-deoxy-2-acetamido-β-D-gluco-
pyranoside (36). To N-pentenoyl aminooxy LacNAc 35 (21 mg,
0.044 mmol) were added N-acetylmannosamine (15 mg, 0.066 mmol),
sodium pyruvate (24 mg, 0.22 mmol), and CTP Na (37 mg, 0.066
3
mmol), and they were dissolved in H2O (3 mL). A concentrated stock of
Tris-HCl buffer pH 8.5 with MgCl2 was added to a final concentration of
100 mM Tris, 20 mM MgCl2. Recombinant E. coli K12 sialic acid
aldolase (4 U), Neusserua meningitidis CMP-sialic acid synthetase (4 U),
and Pasteurella multocida α-2,3-sialyltransferase (2 U) were added,
followed by H2O, to bring the volume to 4 mL. The reaction mixture
was incubated at 37 °C for 2 h, followed by shaking at room temperature
for 14 h. The reaction was monitored by TLC (4:2:1 EtOAc/MeOH/
H2O), and upon completion, calf alkaline phosphatase was added to
remove remaining nucleotide phosphate. After further incubation at
37 °C for 2 h, the reaction mixture was quenched with cold MeOH
(4 mL) and incubated on ice for 20 min. The mixture was centrifuged,
precipitates were removed, and the solution concentrated under
vacuum. The resulting residue was passed through a BioGel P-2 size
exclusion column and eluted with ddH2O to obtain 36 (28.9 mg, 86%)
as a white, fluffy powder after lyophilization: 1H NMR (600 MHz, D2O)
δ 5.80 (ddt, J = 16.9, 10.3, 6.6 Hz, 1H), 5.09ꢀ4.99 (m, 2H), 4.74 (d, J =
8.8 Hz, 1H), 4.52 (d, J = 7.9 Hz, 1H), 4.08 (dd, J = 9.9, 3.1 Hz, 1H), 3.96
(app d, J = 10.7 Hz, 1H), 3.92 (d, J = 3.0 Hz, 1H), 3.88ꢀ3.79 (m, 5H),
3.76ꢀ3.70 (m, 3H), 3.69ꢀ3.64 (m, 3H), 3.62ꢀ3.52 (m, 5H), 2.73 (dd,
J = 12.4, 4.6 Hz, 1H), 2.31 (dd, J = 13.5, 6.7 Hz, 2H), 2.24 (app t, J = 7.0
Hz, 2H), 2.02 (s, 3H), 2.00 (s, 3H), 1.77 (t, J = 12.1 Hz, 1H); 13C NMR
(151 MHz, D2O) δ 175.00, 174.73, 173.83, 172.51, 136.54, 115.87,
103.81, 102.54, 99.79, 77.73, 75.46, 75.16, 75.08, 72.87, 72.20, 71.74,
69.34, 68.31, 68.08, 67.45, 62.57, 61.01, 59.81, 59.42, 52.91, 51.67, 39.62,
31.73, 28.81, 22.19, 22.01; HRMS (ESI) calcd for C30H49N3O20
[MꢀH]ꢀ m/z = 770.2837, found 770.2812.
N-Hydroxypent-4-enamide (5-Acetamido-3,5-dideoxy-D-
glycero-α-D-galacto-2-nonulopyranosylonic acid)-(2f3)-β-
D-galactopyranosyl-(1f4)[(1f3)-α-L-fucopyranosyl]-2-deoxy-
2-acetamido-β-D-glucopyranoside (37). To a 5 mL solution of
100 mM Tris buffer (pH 7.5) containing 5 mM L-fucose (0.025 mmol),
5 mM MgSO4 (0.025 mmol), 10 mM GTP Na (0.05 mmol), and
3
10 mM ATP Na (0.05 mmol) were added Saccharomyces cerevisiae
3
N-Hydroxypent-4-enamide (β-D-Galactopyranosyl)-(1f4)-2-
deoxy-2-acetamido-β-D-glucopyranoside (35). A solution of
N-hydroxypentenoyl disaccharide 34 (94 mg, 0.080 mmol) in THF
(4 mL) was cooled to 0 °C, and tetrabutylammonium fluoride (1 M in
THF, 160 μL) was added dropwise under N2. The mixture was allowed
to warm to room temperature and stirred for 18 h, after which it was
diluted with EtOAc and concentrated under reduced pressure. The
resulting residue was dissolved in MeOH (4 mL), and NaOMe (25% in
MeOH, 40 μL) was added dropwise with stirring at room temperature.
After 8 h, the reaction was neutralized with Dowex 50W-X8 (H+) and
filtered to remove resin. The crude product was concentrated under
vacuum and purified by silica column chromatography (10ꢀ25%
MeOH/CH2Cl2). The purified residue was filtered to remove trace
silica and lyophilized to yield the deprotected N-hydroxypentenoyl
inorganic pyrophosphatase (40 U) and B. fragilis FKP (4 U). The
mixture was incubated at 37 °C for 2 h to preform GDP-fucose and
monitored by TLC (2:1:1 iPrOH/AcOH/H2O). To this was added
N-pentenoyl aminooxy sialoside 36 (5 mg, 0.07 mmol) and H. pylori
α-1,3 fucosyltransferase (1 U), followed by incubation at room tem-
perature for 16 h. The reaction was monitored by TLC (4:2:1 EtOAc/
MeOH/H2O), and upon completion, calf alkaline phosphatase was
added to remove remaining nucleotide phosphate. After further incuba-
tion at 37 °C for 2 h, the reaction mixture was quenched with cold
MeOH (5 mL) and incubated on ice for 20 min. The mixture was
centrifuged, precipitates were removed, and the solution was concen-
trated under vacuum. The resulting residue was passed through a BioGel
P-2 size exclusion column and eluted with ddH2O to obtain 37 (4.6 mg,
78%) as a white, fluffy powder after lyophilization: 1H NMR (600 MHz,
D2O) δ 5.88ꢀ5.77 (m, 1H), 5.06 (dd, J = 26.6, 14.1 Hz, 3H), 4.81 (app
s, 1H), 4.51 (d, J = 7.7 Hz, 1H), 4.12ꢀ4.01 (m, 2H), 3.97 (dd, J = 20.8,
1
LacNAc 35 as a white powder (32 mg, 84%): H NMR (500 MHz,
D2O) δ 5.80 (ddt, J = 12.8, 10.3, 6.5 Hz, 1H), 5.10ꢀ4.98 (m, 2H),
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Journal of the American Chemical Society
ARTICLE
10.7 Hz, 2H), 3.92ꢀ3.82 (m, 7H), 3.75 (d, J = 12.5 Hz, 1H), 3.70ꢀ3.55
(m, 10H), 3.51 (t, J = 8.7 Hz, 1H), 2.79ꢀ2.70 (m, 1H), 2.37ꢀ2.30 (m,
2H), 2.30ꢀ2.22 (m, 2H), 2.02 (d, J = 4.1 Hz, 6H), 1.78 (t, J = 12.2 Hz,
1H), 1.15 (d, J = 6.1 Hz, 3H); 13C NMR (151 MHz, D2O) δ 175.04,
174.51, 173.79, 172.53, 136.55, 115.88, 103.70, 101.62, 99.64, 98.61,
75.65, 75.57, 74.91, 74.58, 72.92, 71.89, 71.83, 69.24, 69.19, 68.26, 68.12,
67.70, 67.30, 66.70, 62.62, 61.44, 59.52, 59.33, 53.56, 51.70, 39.78, 31.73,
28.80, 22.29, 22.03, 15.25; HRMS (ESI) calcd for C36H59N3O24
[MꢀH]ꢀ m/z = 916.3416, found 916.3394.
’ ASSOCIATED CONTENT
S
Supporting Information. Experimental procedures for
b
1ꢀ30 and spectroscopic data for all new compounds; complete
ref 68. This material is available free of charge via the Internet at
http://pubs.acs.org.
’ AUTHOR INFORMATION
Aminooxy (5-Acetamido-3,5-dideoxy-D-glycero-α-D-galacto-
2-nonulopyranosylonic acid)-(2f3)-β-D-galactopyranosyl-
(1f4)[(1f3)-α-L-fucopyranosyl]-2-deoxy-2-acetamido-β-D-
glucopyranoside (38). A solution of N-hydroxypentenoyl SLex 37
(4.2 mg, 0.0046 mmol) in MeCN/MeOH/FA (3:1:0.001, v:v; 1.6 mL)
was stirred at room temperature with dropwise addition of I2 (0.014
mmol, 0.5 M solution in THF). The mixture was heated to 35 °C and
stirred for 2 h, after which the reaction was quenched with the addition
of aqueous NH4HCO3 (500 mM) and Na2S2O3 (50 mM) until the
disappearance of color. The solvent was removed under reduced
pressure, and the remaining residue was purified by silica flash chroma-
tography (4:2:1 EtOAc/MeOH/H2O). The desired fractions were
pooled and concentrated under vacuum. The purified product was
dissolved in ddH2O and lyophilized to give the free aminooxy SLex 38
(2.5 mg, 65%) as a white powder: 1H NMR (600 MHz, D2O) δ 5.11 (d,
J = 4.0 Hz, 1H), 4.72 (d, J = 8.7 Hz, 1H), 4.52 (d, J = 7.8 Hz, 1H), 4.09
(dd, J = 9.9, 3.0 Hz, 1H), 4.04 (dd, J = 14.8, 4.7 Hz, 1H), 3.98 (dd, J =
18.7, 9.6 Hz, 2H), 3.94ꢀ3.92 (m, 1H), 3.92ꢀ3.87 (m, 5H), 3.86 (dd, J =
13.6, 6.3 Hz, 2H), 3.78 (app d, J = 2.8 Hz, 1H), 3.70ꢀ3.63 (m, 8H),
3.61ꢀ3.56 (m, 3H), 3.55ꢀ3.51 (m, 1H), 2.76 (dd, J = 12.4, 4.6 Hz, 1H),
2.03 (s, 6H), 1.80 (t, J = 12.2 Hz, 1H), 1.17 (d, J = 6.6 Hz, 3H); 13C
NMR (151 MHz, D2O) δ 175.06, 174.51, 173.78, 103.04, 101.65, 99.95,
99.66, 98.61, 75.68, 75.37, 74.92, 74.71, 73.18, 72.95, 71.90, 71.84, 69.27,
69.20, 68.27, 68.15, 67.72, 67.32, 66.72, 62.65, 61.46, 59.69, 53.94, 51.72,
43.35, 39.79, 22.18, 22.05, 15.27; HRMS (ESI) calcd for C31H52N3O23
[MꢀH]ꢀ m/z = 834.2994, found 834.2997.
Corresponding Author
crb@berkeley.edu
’ ACKNOWLEDGMENT
We thank Dr. Xi Chen and Dr. Hai Yu for the gift of plasmids
encoding the NmCSS, Pd26ST, and PmST1 enzymes; Dr. Peng Wu
for supplying enzyme and plasmids for the FKP, Hp1,3FT, and
pyrophosphatase; Dr. David King for MS analysis and expertise;
and Brian Belardi, Karen Dehnert, David Rabuka, and Brian Carlson
for materials, helpful discussion, and manuscript critique. This
work was funded by NIH/NIGMS (R01GM059907). J.E.H. was
supported by a predoctoral fellowship from the U.S. National
Science Foundation.
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