Strain-Promoted Alkyne-Azide Cycloadditions (SPAAC) Reveal New Features of Glycoconjugate Biosynthesis

Article abstract of DOI:10.1002/cbic.201100117

We have shown that 4-dibenzocyclooctynol (DIBO), which can easily be obtained by a streamlined synthesis approach, reacts exceptionally fast in the absence of a Cu^I catalyst with azido-containing compounds to give stable triazoles. Chemical mod

Full text of DOI:10.1002/cbic.201100117

DOI: 10.1002/cbic.201100117
Strain-Promoted Alkyne–Azide Cycloadditions (SPAAC)
Reveal New Features of Glycoconjugate Biosynthesis
Ngalle Eric Mbua, Jun Guo, Margreet A. Wolfert, Richard Steet,* and Geert-Jan Boons*[a]
We have shown that 4-dibenzocyclooctynol (DIBO), which can
easily be obtained by a streamlined synthesis approach, reacts
exceptionally fast in the absence of a CuI catalyst with azido-
containing compounds to give stable triazoles. Chemical modi-
fications of DIBO, such as oxidation of the alcohol to a ketone,
increased the rate of strain promoted azide–alkyne cycloaddi-
tions (SPAAC). Installment of a ketone or oxime in the cyclo-
octyne ring resulted in fluorescent active compounds whereas
this property was absent in the corresponding cycloaddition
adducts; this provides the first example of a metal-free alkyne–
azide fluoro-switch click reaction. The alcohol or ketone func-
tions of the cyclooctynes offer a chemical handle to install a
variety of different tags, and thereby facilitate biological stud-
ies. It was found that DIBO modified with biotin combined
with metabolic labeling with an azido-containing monosac-
charide can determine relative quantities of sialic acid of living
cells that have defects in glycosylation (Lec CHO cells). A com-
bined use of metabolic labeling/SPAAC and lectin staining of
cells that have defects in the conserved oligomeric Golgi
(COG) complex revealed that such defects have a greater
impact on O-glycan sialylation than galactosylation, whereas
sialylation and galactosylation of N-glycans was similarly
impacted. These results highlight the fact that the fidelity of
Golgi trafficking is a critical parameter for the types of oligo-
saccharides being biosynthesized by a cell. Furthermore, by
modulating the quantity of biosynthesized sugar nucleotide,
cells might have a means to selectively alter specific glycan
structures of glycoproteins.
Introduction
Metal-free cycloadditions between cyclooctynes and azides
that give stable 1,2,3-triazoles, have found wide utility in label-
ing glycans in proteins and lipids of living cells, glycoprotein
enrichment for proteomics, protein and oligonucleotide modi-
fication, and tissue reengineering.[1] These reactions, which
have been coined “strain-promoted alkyne–azide cycloaddi-
tions” (SPAAC) have also made an entry in material sciences
and have, for example, been employed for the assembly, cross-
linking and surface modification of dendrimers,[2] derivatization
of polymeric nanostructures,[3] and patterning of surfaces.[4]
The attraction of SPAAC is that it does not require a toxic
metal, is highly efficient even in a very complex milieu and
proceeds efficiently at ambient temperature. Density functional
theory (B3LYP) calculations of the transition states of cycloaddi-
tions of phenyl azide with acetylene and cyclooctyne indicate
that the fast rate of the strain promoted cycloaddition is due
to a lower energy required for distorting the 1,3-dipole and
alkyne into the transition-state geometry.[5]
these reagents is rather limited (Scheme 1). It has, however,
been found that electron-withdrawing fluorine groups at the
propargylic position of a cyclooctyne (DIFO, 2) dramatically in-
crease the rate of strain-promoted cycloaddition with azides.[6]
The attractiveness of this methodology has, for example, been
demonstrated by visualization of glycans in vivo at subcellular
resolution during the development of zebrafish embryos.[7]
We have found that derivatives of 4-dibenzocyclooctynol
(DIBO, 3) react fast with azido-containing saccharides and
amino acids, and can be employed for visualizing metabolically
labeled glycans of living cells.[8] While the fluorine atoms of
DIFO (2) influence rate enhancement by increasing interaction
energies, the aromatic rings of 3 accomplish a similar increase
in reaction rate through conformational effects that result in
decreasing the distortion energy. Attractive features of DIBO
(3) include easy access to the compounds by a simple synthe-
sis approach, nontoxicity and straightforward attachment of a
variety of probes. Furthermore, dibenzocyclooctynes can be
generated photochemically by short irradiation by UV light of
corresponding cyclopropenones, and thereby provide opportu-
nities for the spatially and temporally controlled labeling of
The first generation of cyclooctynes (1) suffered from rela-
tively slow reaction rates and as a consequence the scope of
[a] N. E. Mbua, Dr. J. Guo, Dr. M. A. Wolfert, Prof. Dr. R. Steet,
Prof. Dr. G.-J. Boons
Complex Carbohydrate Research Center, University of Georgia
315 Riverbend Road, Athens, GA 30602 (USA)
E-mail: rsteet@ccrc.uga.edu
gjboons@ccrc.uga.edu
Supporting information for this article is available on the WWW under
http://dx.doi.org/10.1002/cbic.201100117.
Scheme 1. Reagents for labeling azido-containing biomolecules.
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Labeling Cellular Glycoconjugates by SPAAC
the target substrates.[9] We have
also shown that by employing
nitrones and nitrile oxides as
1,3-dipoles, the rate of cycload-
dition can be further enhanced
and this technology has, for ex-
ample, made it possible to se-
lectively tag proteins at the
N terminus or perform sequen-
tial modifications of complex
compounds.[10]
Furthermore,
several analogues of DIBO have
been reported that exhibit even
higher rates of cycloaddition
with azides.[11]
We report here a streamlined
approach for the preparation
and modification of DIBO, and
show that modification of the
eight-membered ring by, for ex-
ample, installment of a ketone,
affects the rate of cycloaddition.
The presence of a ketone or
oxime resulted in compounds
that are fluorescent. Interesting-
ly, the corresponding cycloaddi-
tion products are nonfluores-
cent, and hence compounds
such as 11 and 12, provide a
novel metal-free alkyne–azide
fluoro-switch click reaction,
which, for example, can be ex-
ploited in monitoring reactions
in real time. Metabolic labeling
combined with SPAAC of wild-
type cells and those that have
known defects in their glycosy-
lation machinery showed that
relative quantities of sialylation
of glycoconjugates can easily
and reliably be established. Fur-
thermore, a combined use of
metabolic labeling/SPAAC and
lectin staining revealed that a
defect in the conserved oligo-
meric Golgi (COG) complex af-
fects terminal processing of N-
glycans to a greater extent than
modification of O-glycans.
Scheme 2. Reagents and conditions: a) TMSCHN2, BF3·OEt2, DCM, ꢀ108C, 3 h, 71%; b) NaBH4, EtOH/THF, 7 h,
100%; c) Br2, CHCl3, 0.5 h, 58%; d) LDA, THF, 0.5 h, 57%; e) 4-nitrophenyl chloroformate, pyridine, DCM, 18 h, 92%;
f) propanol amine, Et3N, DCM, 3 h, 89%; g) tris(ethylene glycol)-1,8-diamine, Et3N, DCM, 3 h, 80%; h) Dess–Martin,
DCM, 0.5 h, 92%; i) (carboxymethyl)hydroxyamine, DCM/MeOH, 61%; j) N-{2-[2-(2-amino-ethoxy)ethoxy]ethyl}-2-
aminooxy-acetamide, AcOH, DCM/MeOH, 63%; k) biotin-OSu, Et3N, DCM/MeOH, 89% for 14, 92% for 15.
this resulted in ring expansion by carbene insertion to give tri-
eneone 5.[12] The latter compound was reduced with NaBH4
(!6), brominated with bromine in chloroform (!7), and then
treated with LDA in THF[13] to give target compound 3 in a
yield of 57%. In an alternative approach, intermediate 6 was
obtained by base-mediated ring opening of Kagan’s ether,
which was prepared by a double Friedel–Crafts alkylation phe-
nylacetaldehyde.[14] The former synthesis route provides a
Results and Discussion
Chemical synthesis and physical properties of DIBO
4-Dibenzocyclooctynol (3) was easily prepared starting from
commercially available dibenzosuberenone (4, Scheme 2),
which was treated with TMSCHN2 in the presence of BF3·Et2O;
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R. Steet, G.-J. Boons et al.
higher overall yield (71% vs. 42%), requires fewer chemical
steps, and is more scalable.
to a lower energy required for distorting the 1,3-dipole and
alkyne into the transition-state geometry.[5] This finding has
been exploited in the design of more reactive cyclooctynes
and, for example, it has been found that installment of an
amide or cyclopropanation in the eight membered ring leads
to higher rates of cycloaddition.[16] Surprisingly, oxime 12 did
not exhibit a faster rate of reaction than alcohol 3. Computa-
tional studies will be required to provide a rationale for these
observations.
The hydroxyl of 3 provides an opportunity for further func-
tionalization and, for example, can be activated with 4-nitro-
phenyl chloroformate to give activated carbonate 8, which can
be treated with amines, such as propanol amine and tris(ethyl-
ene glycol)-1,8-diamine, to give carbamates 9 and 10, respec-
tively. Alternatively, the alcohol of 3 can be oxidized to ketone
11 by using Dess–Martin periodate, which can then be modi-
fied by aminooxy derivatives. This procedure was employed
for the preparation of compounds 12 and 13. It was observed
that oxime formation was rather sluggish and the rate of reac-
tion could not significantly be increased by the addition of ani-
line.[15] Finally, acylation of the amino groups of compounds 10
and 13 with N-(biotinyloxy)succinimide (biotin-OSu) and Et3N
gave biotin-labeled derivatives 14 and 15, respectively.
The acid labile oxime linkage of compound 15 provides op-
portunities for catch and release strategies, which, for example,
are required for glycoproteomic applications. In this respect,
treatment of triazole 21, which was formed by treatment of 15
with benzyl azide, at pH 2 for 10 h resulted in hydrolysis of the
oxime linkage. On the other hand, compound 20, which was
formed by treatment of derivative 14 with benzyl azide, was
stable under these conditions.
It was observed that compound 3 has an excellent shelf life
and remained intact after treatment with nucleophiles, such as
thiols and amines. However, upon exposure to azides a fast
reaction took place to give the corresponding triazoles in high
yield. Rate measurements of cycloaddition of compounds 3, 9,
11 and 12 were conducted by UV spectroscopy at 25ꢁ0.18C.
A calculated amount of 0.25m solution of benzyl azide, which
was required to achieve a desired azide concentration (6ꢁ
10ꢀ4–1.5ꢁ10ꢀ2 m), was added to a thermally equilibrated 6ꢁ
10ꢀ5 m solution of cyclooctyne in methanol. The progress of
the reactions was monitored by the decay of the characteristic
absorbance of acetylenes at approximately 317 nm. Consump-
tion of starting material followed a first-order equation and
pseudo-first-order rate constants were obtained by least-
square fitting of the data to a single exponential equation. The
rate dependence as a function of the concentration of azide
was linear and least-square fitting of the data to a linear equa-
tion produced the bimolecular rate constants summarized in
Table 1.
It was observed that compound 11 exhibits fluorescent
properties and in methanol emits light with a maximum fluo-
rescent intensity of 436 nm and a quantum yield of 36% when
excited at 375 nm. Oxime 12 showed a maximum fluorescent
intensity of 380 nm and a quantum yield of 34% when excited
at 313 nm (Figure 1). Interestingly, triazoles 18 and 19, which
were formed by treatment of 11 and 12, respectively, with
Table 1. Bimolecular rate constants for the reactions of acetylenes with
benzyl azide in methanol at 25ꢁ0.18C.
Figure 1. Fluorescence spectra of compounds 11 and 18 (lex =375 nm) and
12 and 19 (lex =313 nm) in methanol (10ꢀ5 m); AU: arbitrary fluorescence
units.
Cyclo-
octyne
Product Rate
[mꢀ1 sꢀ1
Cyclo- Product Rate
octyne
]
[mꢀ1 sꢀ1
]
3
11
16
18
0.0567ꢁ0.0027
0.2590ꢁ0.0067
9
12
17
19
0.0696ꢁ0.0019
0.0611ꢁ0.0035
benzyl azide, showed only very weak fluorescence. Previously,
a number of CuI-mediated fluorogenic alkyne–azide click
(CuAAC) reactions were described, which were based on the
formation of a fluorescent 1,3-triazole from a nonfluorescent
precursor, such as azide or alkyne substituted coumarins, 1,8-
naphthalimides or anthracenes.[17] Treatment of 11 and 12 with
benzyl azide is the first example of a metal-free alkyne–azide
fluoro-switch click reaction. A unique feature of this reaction is
that the triazole moiety is responsible for fluorescent quench-
ing whereas this effect is not observed for the parent cyclooc-
tyne. It is to be expected that fluorogenic SPAAC will provide
opportunities to monitor the progress of reactions in real time.
Such an approach will in particular be advantageous for appli-
cations in which a fluorescent cyclooctyne is attached to a sur-
face or solid support.[4,18]
The rate constant of cycloaddition of DIBO (3) with benzyl
azide is two orders of magnitude higher than that of cyclooc-
tyne 1, and similar to that of DIFO. Conversion of the alcohol
of DIBO into a carbamate, as in compound 9, did not influence
the rate of cycloaddition. However, ketone 11 reacted three-
times faster than the parent alcohol 3. Probably, oxidation of 3
to 11 induces a small change in the conformation of the cyclo-
octyne ring, which might be responsible for the observed en-
hancement of reaction rate. In this respect, density functional
theory (B3LYP) calculations of the transition states of cycloaddi-
tions of phenyl azide with acetylene and cyclooctyne indicate
that the fast rate of the strain promoted cycloaddition is due
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Labeling Cellular Glycoconjugates by SPAAC
Evaluation of DIBO for labeling glycoconjugates of living
cells
The bioorthogonal chemical reporter strategy is emerging as a
versatile method for labeling biomolecules, such as nucleic
acids, lipids, proteins, and carbohydrates.[1a,c,d] In this approach,
an abiotic chemical functionality (reporter) is incorporated into
a target biomolecule, which can then be reacted with a com-
plementary bioorthogonal reagent linked to a probe. Azide is
commonly employed as a reporter and can be installed into
biomolecules by using azido-containing biosynthetic precur-
sors that can be accepted by the cell’s native or engineered
biosynthetic machinery. For example, azido-containing glyco-
conjugates can be biosynthesized by metabolic labeling with
peracetylated N-a-azidoacetylmannosamine (Ac4ManNAz),
which is an appropriate substrate for the cell’s glycosylation
machinery.[19] A subsequently bioorthogonal reaction can then
covalently attach a probe to the azido function, which in turn
makes it possible to conduct a multitude of functional studies.
A number of bioorthogonal reactions have been described for
reactions with azides, however, SPAAC is emerging as a partic-
ularly attractive approach as it can be performed under physio-
logical conditions and does not require a toxic metal catalyst.
To establish biotin-modified DIBO derivatives 14 and 15 as
appropriate bioorthogonal reagents, we employed these com-
pounds to determine relative quantities of cell surface sialyla-
tion of wild-type and mutant cells, and compared the results
with traditional lectin staining. It is well established that
Ac4ManNAz can be employed by the glycosylation machinery
to install azido-containing sialic acid in various glycoconjugates
and a subsequent reaction with 14 or 15 was expected to pro-
vide quantitative data on cell surface sialylation.[19] Thus, Jurkat
cells were cultured in the presence of Ac4ManNAz (25 mm) for
3 days to metabolically introduce N-azidoacetyl-sialic acid
(SiaNAz) moieties into glycoproteins and glycolipids. As a neg-
ative control, Jurkat cells were employed that were grown in
the presence of peracetylated N-acetylmannosamine
(Ac4ManNAc). A time-course experiment was conducted by ex-
posing the cells to 30 mm of 14 and 15 for different time peri-
ods at room temperature, washed and then stained with
avidin–FITC for 15 min at 48C. The efficiency of the two-step
cell surface labeling was determined by measuring the fluores-
cence intensity of the cell lysates. Gratifyingly, the ManNAz-
labeled cell exhibited strong fluorescent readings after being
stained with the two different DIBO derivatives, and the cell
labeling was almost complete after a reaction time of 60 min
(Figure 2), whereas the control cells gave a very low fluores-
cence intensity showing that background labeling is negligible.
Similar results were obtained when Chinese hamster ovary
(CHO)-K1 cells were subjected to the same procedure.
Figure 2. Time course of cell surface labeling with compounds 14 and 15.
Jurkat cells grown for 3 days in the presence of Ac4ManNAc or Ac4ManNAz
(25 mm) were incubated with compounds 14 or 15 (30 mm) for 0–90 min at
room temperature. Next, cells were incubated with avidin–FITC for 15 min
at 48C, after which cell lysates were assessed for fluorescence intensity. AU
indicates arbitrary fluorescence units. Data (n=3) are presented as mean
ꢁSD.
Figure 3. Fluorescence images of cells labeled with compounds 14 and 15
and avidin–AlexaFluor488. CHO cells grown for 2 days in the presence of: A),
B) Ac4ManNAc, or C), D) Ac4ManNAz (100 mm) were incubated with com-
pounds: A), C) 14, or B), D) 15 (30 mm) for 1 h at room temperature. Next,
cells were incubated with avidin–AlexaFluor488 for 15 min at 48C, and
imaged after being washed, fixed, and stained for the nucleus with the far-
red-fluorescent dye TO-PRO-3 iodide. “Merged” indicates that the images of
cells labeled with AlexaFluor (488 nm) and TO-PRO (633 nm) are merged
and shown in green and red, respectively.
Metabolically labeled cells were also examined by confocal
microscopy (Figure 3 and Figure S1 in the Supporting Informa-
tion). Thus, adherent Chinese hamster ovary (CHO) cells were
cultured in the presence of Ac4ManNAz (100 mm) for two
days.[20] Next, cell surface azido moieties were treated with 14
or 15 (30 mm) for 1 h at ambient temperature, and then visual-
ized with avidin–AlexaFluor488 for 15 min at 48C. Staining was
mainly observed at the cell surface, and as expected, blank
cells exhibited very low fluorescence staining confirming that
background labeling is negligible. We also found that the two-
step labeling approach with 14 and 15 had no effect on cell vi-
ability as determined by morphology and exclusion of trypan
blue (data not shown).
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R. Steet, G.-J. Boons et al.
The concentration dependency of the cell surface labeling
was studied by incubating Jurkat and CHO-K1 cells with vari-
ous concentrations of 14 or 15 followed by staining with
avidin–FITC (Figure S2 in the Supporting Information). As ex-
pected, cells displaying azido moieties showed a dose-depen-
dent increase in fluorescence intensity and reliable labeling
was achieved at a concentration of 3 mm of 14 or 15, however,
optimal results were obtained at concentrations ranging from
30 to 100 mm. No increase in labeling was observed at concen-
trations higher than 100 mm due to limited solubility.
Lec13 mutants exhibit a reduced expression of GDP-Man-4,6-
dehydratase activity; this results in a decrease in GDP-fucose
biosynthesis and underfucosylation of glycoproteins and glyco-
lipids.[27] The N-glycan profiles of these cells show increased
levels of core nonfucosylated N-glycans with the most abun-
dant N-glycans being asialo-, mono-, or di-sialylated structures.
Wild-type CHO-K1 and Lec2, Lec13 and Lec32 mutant cells
were cultured in the presence of Ac4ManNAz or Ac4ManNAz
(100 mm) for 2 days and then exposed to biotin-modified DIBO
(14) for 1 h at room temperature. Next, the cells were washed
and labeled with avidin–FITC at 48C and the fluorescence
intensity measured. As expected, the wild-type and Lec13 cells
gave similar and strong fluorescence intensity readings (Fig-
ure 4A). On the other hand, the Lec2 and Lec32 mutants
showed a significant reduction in staining intensity and, in the
case of the Lec2 cells, the readings were barely above the con-
trol, which indicates that these cells express very low levels of
surface sialosides. Thus, the results of these studies are in
agreement with the previously described defects in the
mutant cell lines and hence support the notion that the chemi-
Jurkat and CHO-K1 cells were also metabolically labeled with
peracetylated
N-a-azidoacetylgalactosamine
(Ac4GalNAz,
100 mm), which can be metabolized by a number of cells and
installed on mucin type glycoproteins.[21] Subsequent treat-
ment of the CHO-K1 cells with 14 followed by avidin–FITC re-
sulted in strong fluorescent labeling whereas weak labeling
was observed for Jurkat cells (Figure S3 in the Supporting
Information). These results are in agreement with the well-
known fact that CHO cells produce significant quantities of
mucins whereas this is not the case for Jurkat cells.[22]
Having established optimal
conditions for SPAAC of azido-
modified glycoconjugates of
living cells with DIBO reagents,
labeling studies were performed
with a panel of lectin-resistant
(Lec) mutant CHO cells. These
cell lines (Lec2, Lec13 and
Lec32), which exhibit unique
structural changes in surface
carbohydrates that reflect spe-
cific defects in glycosylation re-
actions, were expected to be
ideally suited for validation of
the SPAAC methodology. Lec2
cells have a mutation in the
open reading frame of the
CMP-sialic acid transporter, and
therefore, are unable to translo-
cate CMP-sialic acid into the
lumen of the Golgi apparatus,
resulting in a marked reduction
in glycoprotein and ganglioside
sialylation.[23] Although very
small amounts of sialic acid con-
taining glycoconjugates are
made by these cells,[24] the
major class of glycans are asialo,
core
fucosylated
LacNAc
N-glycans
Figure 4. SiaNAz expression in CHO-K1 and CHO glycosylation mutant cells and the effect of neuraminidase treat-
ment on cell surface labeling. A) CHO-K1 and CHO mutant cells grown for 2 days in the presence of Ac4ManNAc
or Ac4ManNAz (100 mm) were incubated, either directly or after treatment with V. cholerae neuraminidase
(50 mUmLꢀ1) in serum-free culture medium for 2 h at 378C, with compound 14 (30 mm) for 1 h at room tempera-
ture. Next, cells were incubated with avidin–FITC for 15 min at 48C, after which cell lysates were assessed for fluo-
rescence intensity. To assess the effects of desialylation of CHO-K1 and CHO glycosylation mutants on their recog-
nition by PNA and RCA1, cells were incubated, either directly or after treatment with V. cholerae neuraminidase
(50 mUmLꢀ1), in serum-free culture medium for 2 h at 378C with: B) PNA–FITC (50 mgmLꢀ1), or C) RCA1–FITC
(50 mgmLꢀ1) for 45 min on ice in the dark. Next cell lysates were assessed for fluorescence intensity. AU indicates
arbitrary fluorescence units. Data (n=3) are presented as meanꢁ SD. Similarly, D) CHO-K1 (i, iii) and Lec2 (ii, iv)
cells were imaged, either directly (i, ii) or after treatment with V. cholerae neuraminidase (iii, iv) with PNA–FITC.
having
moieties.[25]
Lec32 mutants also exhibit a
defect in sialylation due to a re-
duced expression of CMP-sialic
acid synthetase. As
a result,
these cells have an increase in
terminal b-galactoside residues
on cell surface glycoproteins.[26]
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Labeling Cellular Glycoconjugates by SPAAC
cal reporter strategy can be employed to determine relative
quantities of sialylation of glycoconjugates of living cells. Fur-
thermore, treatment of the wild-type cells with Vibrio cholerae
neuraminidase led to a similar fluorescent reading as control
cells, confirming selective azide incorporation into sialic acid.
The results of the metabolic labeling studies were compared
with traditional lectin staining by using FITC-labeled peanut
(Arachis hypogaea) agglutinin (PNA) and FITC-labeled Ricinus
communis (castor bean) agglutinin type 1 (RCA1), which mainly
recognize terminal b-Gal-(1-3)-GalNAc residues of O-linked
structures and b-Gal-(1-4)-GlcNAc (LacNAc) found on N-linked
glycoproteins, respectively. Cells that have intact sialylation
machinery modify b-galactosyl residues with sialic acid and
hence display low reactivity against PNA and RCA1 lectins.
Indeed, the wild-type and Lec13 cells gave fluorescent intensi-
ties just above background whereas the Lec2 and Lec32 mu-
tants, which have a defect in sialylation, showed strong stain-
ing (Figure 4B–D).
(LDLR) activity.[28] Further examinations have shown that these
cell lines have defects in the conserved oligomeric Golgi (COG)
complex, which is a protein complex consisting of eight sub-
units (Cog1–8) that play a critical role in retrograde vesicle
transport and intra-Golgi trafficking. Malfunctions in the COG
complex impact Golgi integrity and result in defects in protein
sorting and glycosylation.[29] Mutations in COG subunits have
also been observed in humans and result in severe congenital
disorders of glycosylation (CDG).[30]
Metabolic labeling of Cog1 and Cog2 cells with ManNAz fol-
lowed by SPAAC with 14 and staining with avidin–FITC
showed that these cells produce sialylated glycoconjugates,
however, at a significantly reduced level compared to wild-
type CHO-K1 cells (Figure 4A). Staining with PNA–FITC demon-
strated that that the cells expose terminal galactosyl residues
on their O-linked glycans (Figure 4B). Furthermore, treatment
of the cells with V. cholerae neuraminidase led to a similar
staining intensity as for wild-type cells. This indicates that both
cell types express similar quantities of galactosyl moieties, and
thus it appears that in Cog1 and -2 cells, glycoprotein sialyla-
tion of O-glycans is more severely imparted than galactosyla-
tion. Interestingly, a different staining profile was obtained
when RCA1–FITC was employed (Figure 4C) and in this case
untreated and neuraminidase exposed cells gave similar but
reduced fluorescent intensities highlighting that N-glycan sialy-
lation and galactosylation are both affected in the Cog1 and
Cog2 mutants. These results suggest that loss of COG complex
function might affect the localization and/or stability of glyco-
syltransferases involved in terminal processing of N-glycans to
a greater extent than those enzymes that modify O-glycans. In
support of this hypothesis, recent studies have shown that the
stability of b(1,4)-galactosyltransferase is altered in COG-deplet-
ed HeLa cells due to altered trafficking and proteasomal degra-
dation.[31] At this point, we can, however, not rule out the pos-
sibility that these differences are due to the type and amount
of glycoprotein cargo that is modified in the Golgi of these
cells.
Treatment of the wild-type and Lec13 cells with V. cholerae
neuramidase resulted in fluorescent intensities similar to that
of the Lec2 and Lec32 cells; this indicates that the various cell
types express similar quantities of galactosyl-containing glyco-
proteins. Furthermore, a similar neuraminidase treatment of
Lec2 cells followed by staining with PNA–FITC or RCA1–FITC
did not lead to a significant increase in fluorescent intensity;
this demonstrates that these cells do not significantly modify
their cell surface glycoconjugates with sialic acid. On the other
hand, neuraminidase treatment of Lec32 cells resulted in an
increase in fluorescence staining with RCA1–FITC, whereas it
did not impact the reading of PNA–FITC. These results indicate
that the N-linked glycans of Lec32 contain some sialosides
whereas this modification is absent in O-linked residues.
The lectin staining (Figure 4B–D) and metabolic labeling fol-
lowed by SPAAC (Figure 4A) gave similar results and in particu-
lar both approaches showed that Lec2 cells express very small
quantities of sialosides whereas the Lec32 mutant attach some
sialic acid to their glycoconjugates. Surprisingly, a shortage of
CMP-Neu5Ac as in Lec32 cells resulted in differential sialylation
of N- and O-linked glycans and it appears that N- but not O-
linked glycans are modified by some sialic acid. Finally, both
approaches showed that a defect in fucosylation does not
impact the level of glycoconjugate sialylation. Attempts were
made to directly assess differences in sialylation within the Lec
mutants by utilizing lectins that recognize a(2,3)- or a(2,6)-
linked terminal sialic acid residues (MAA and LFA). Surprisingly,
we found fluorescence intensity for both lectins to be compa-
rable in wild-type and Lec2 mutants (data not shown); this
suggests that these lectins might recognize additional sugar
structures other than terminal sialic acid and are not suitable
for this study. Thus, we believe that the chemical reporter
strategy provides a more reliable approach to determine rela-
tive differences in glycoprotein sialylation.
Conclusions
The past several years has seen a rapid development of the
bioorthogonal chemical reporter methodology for the labeling
of glycoconjugates of living cells and whole organisms. In this
paper, we present a streamlined chemical approach for the
preparation and derivatization of DIBO, which is an ideal bioor-
thogonal reagent for the chemical reporter strategy. Attractive
features of DIBO include easy access by a simple and scalable
synthesis approach, nontoxicity and straightforward attach-
ment of a variety of probes. The use of several cell lines with
known defects in glycoconjugate glycosylation validated DIBO
as a reagent for determining relative quantities of cell surface
glycoconjugate sialylation. The chemical reporter strategy in
combination with lectin staining revealed that O-glycan sialyla-
tion of Cog1 and -2 cells is more severely impacted than galac-
tosylation. Surprisingly, sialylation and galactosylation of N-gly-
cans were similarly affected in these mutant cell lines. These
results suggest that loss of COG complex function might differ-
Having established biotin-modified DIBO (14) as a reliable
reagent for detection of cell surface sialosides, attention was
focused on sialylation of the CHO mutants Cog1 (ldlB) and
Cog2 (ldlC). These cell lines were identified in a genetic screen
for mutations that block low-density lipoprotein receptor
ChemBioChem 2011, 12, 1912 – 1921
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chembiochem.org
1917

R. Steet, G.-J. Boons et al.
evaporated. The residue was dissolved in CH2Cl2 (100 mL) and the
resulting solution was washed with brine (100 mL), which was
back extracted with CH2Cl2 (4ꢁ100 mL). The combined organic
phases were dried (MgSO4), filtered and concentrated under re-
duced pressure to give 6 as a white solid (2.22 g), which was di-
rectly used in the next step without further purification. 1H NMR
(300 MHz, CDCl3): d=7.50 (m, 1H), 7.14–7.30 (m, 7H), 6.90 (q, J=
2.7, 12.0 Hz, 2H), 5.31 (q, 1H, J=6.3, 10.0 Hz), 3.41 (m, 2H);
13C NMR (75 MHz, CDCl3): d=141.7, 136.7, 136.2, 134.5, 131.7,
131.5, 130.1, 129.9, 129.3, 128.7, 127.4, 127.2, 126.9, 125.9, 74.4,
42.7. MALDI HRMS: m/z 245.0949 [M+Na+]; calcd for C16H14NaO+:
245.0937.
ently affect the localization and/or stability of glycosyltransfer-
ases involved in terminal processing of N- and O-glycans. Dif-
ferential modulation of N- and O-linked sialylation was also ob-
served in Lec32 cells, which exhibit a reduced expression of
CMP-sialic acid synthetase, and in this case N-linked oligosac-
charides acquire some sialic acid moieties whereas this is not
the case for O-linked structures. It is well known that the vari-
ous cell types express different ensembles of glycans. The re-
sults of this study indicate that a limited availability of sugar
nucleotides is one way for a cell to selectively modulate the
structures of glycoprotein glycans. We anticipate multiple ap-
plications of the described chemical reporter methodology in
glycobiology and glycomedicine, including the tagging and
isolation of glycoproteins from cell and tissue extracts as well
as the investigation of trafficking and turnover of glycoconju-
gates in healthy and diseased cells.
11,12-Dibromo-5,6,11,12-tetrahydro-dibenzo[a,e]cycloocten-5-ol
(7): Bromine (0.51 mL, 10 mmol) as added dropwise to a stirred so-
lution of 6 (2.22 g, 10 mmol) in CHCl3 (50 mL). After stirring the
mixture for 0.5 h, TLC analysis indicated completion of the reaction.
The solvent was evaporated under reduced pressure and the resi-
due was purified by flash chromatography over silica gel (2:1!1:2,
v/v, hexanes/CH2Cl2) to give 7 as a light-yellow oil (2.22 g, 58%).
1H NMR (300 MHz, CDCl3): d=7.54–7.47 (2H, aromatics), 7.31–6.72
(6H, aromatics), 5.77 (d, J=5.4 Hz, 1H, CHBr), 5.22 (dd, J=3.6,
15.9 Hz, 1H, CHOH), 5.19 (d, J=5.4 Hz, 1H, CHBr), 3.50 (dd, J=3.6,
15.9 Hz, 1H, CH2), 2.75 (dd, J=3.6, 15.9 Hz, 1H, CH2); 13C NMR
(75 MHz, CDCl3): d=141.3, 140.0, 137.2, 134.0, 133.4, 131.5, 131.3,
130.9, 127.8, 126.2, 123.7, 121.3, 76.5, 70.0, 62.3, 32.2. MALDI
HRMS: m/z 402.9313 [M+Na+]; calcd for C16H14Br2NaO+: 402.9304.
Experimental Section
General methods and materials: Chemicals were purchased from
Aldrich or Fluka and used without further purification. Dichloro-
methane was distilled from CaH2 and stored over molecular sieves
4 ꢂ. Pyridine was distilled from P2O5 and stored over molecular
sieves 4 ꢂ. THF was distilled from sodium. All reactions were per-
formed under anhydrous conditions under an atmosphere of
argon. Reactions were monitored by TLC on Kieselgel 60 F254
(Merck). Detection was carried out under UV light (254 nm). Flash
chromatography was performed on silica gel (Merck, 70–230
mesh). Iatrobeads (60 mm) were purchased from Bioscan. 1H NMR
(1D, 2D) and 13C NMR spectra were recorded on a Varian Merc 300
spectrometer equipped with Sun workstations. 1H and 13C NMR
spectra were recorded in CDCl3 or CD3OD, and chemical shifts are
given relative to solvent peaks (CDCl3: 1H, d=7.24, 13C, d=77.0;
CD3OD: 1H, d=3.31, 13C, d=49.0) as internal standard for com-
pounds. High-resolution mass spectra were obtained by an Applied
Biosystems 4700 MALDI mass spectrometer in positive ion reflec-
tive mode by using 2,5-dihydroxyl-benzoic acid in CH3CN as matrix.
5,6-Dihydro-11,12-didehydro-dibenzo[a,e]cycloocten-5-ol (3): Li-
thium diisopropylamide in tetrahydrofuran (2.0m; 8 mL, 16 mmol)
was added dropwise to a stirred solution of 7 (1.53 g, 4.0 mmol) in
tetrahydrofuran (40 mL) under an atmosphere of argon. The reac-
tion mixture was stirred for 0.5 h, after which it was quenched by
the dropwise addition of water (0.5 mL). The solvents were evapo-
rated under reduced pressure, and the residue was purified by
flash chromatography on silica gel (hexanes/CH2Cl2 2:1!0:1, v/v)
to give 3 as a white amorphous solid (0.50 g, 57%). 1H NMR
(300 MHz, CDCl3): d=7.67 (1H, aromatics), 7.37–7.18 (7H, aromat-
ics), 4.57 (dd, J=2.1, 14.7 Hz, 1H, CHOH), 3.04 (dd, J=2.1, 14.7 Hz,
1H, CH2), 2.86 (dd, J=2.1, 14.7 Hz, 1H, CH2); 13C NMR (75 MHz,
CDCl3): d=154.5, 150.6, 128.6, 127.1, 1127.0, 126.0, 125.8, 125.1,
124.7, 123.0, 122.7, 121.7, 111.9, 109.6, 74.2, 47.7.
6H-Dibenzo[a,e]cyclooctatrien-5-one (5): A solution of trimethyl-
silyl diazomethane (10.5 mL, 21.9 mmol) in CH2Cl2 (20 mL) was
added dropwise to a stirred solution of dibenzosuberenone 4
(2.88 g, 14.0 mmol) and BF3·OEt2 (2.59 mL, 21.0 mmol) in CH2Cl2
(30 mL) at ꢀ108C over a period of 1 h. The reaction mixture was
stirred at ꢀ108C for 2 h, and then poured into ice water. The aque-
ous layer was extracted with CH2Cl2 (3ꢁ100 mL) and the combined
organic layers washed with brine, dried (MgSO4), filtered and the
filtrate was concentrated under reduced pressure. The crude prod-
uct was purified by flash chromatography over silica gel (2:1!1:2,
v/v, hexanes/CH2Cl2) to give compound 5 as an amorphous solid
Carbonic acid, 5,6-dihydro-11,12-didehydro-dibenzo[a,e]cyclo-
octen-5-yl ester, 4-nitrophenyl ester (8): 4-Nitrophenyl chlorofor-
mate (0.4 g, 2 mmol) and pyridine (0.4 mL, 5 mmol) were added to
a solution of 3 (0.22 g, 1 mmol) in CH2Cl2 (30 mL). After being
stirred for 4 h at room temperature, the mixture was washed with
brine (2ꢁ10 mL) and the organic layer was dried (MgSO4). The sol-
vents were evaporated under reduced pressure, and the residue
was purified by silica gel column chromatography (hexane/ethyl
acetate, 10:1, v/v) to afford 8 (0.34 g, 89%). 1H NMR (300 MHz,
CDCl3): d=8.23–8.18 (2H, aromatics), 7.56–7.54 (2H, aromatics),
7.46–7.18 (8H, aromatics), 5.52 (dd, J=3.9, 15.3 Hz, 1H, CHOH),
3.26 (dd, J=3.9, 15.3 Hz, 1H, CH2), 2.97 (dd, J=3.9, 15.3 Hz, 1H,
CH2); I3C NMR (75 MHz, CDCl3): d=154.5, 150.7, 149.1, 148.7, 129.0,
127.4, 127.3, 126.7, 126.5, 125.5, 125.2, 124.3, 124.0, 122.6, 122.4,
120.8, 120.6, 120.2, 112.2, 108.5, 80.6, 44.8; MALDI HRMS: m/z
408.0852 [M+Na+]; calcd for C23H15NNaO5+: 408.0842.
1
(2.22 g, 72%). H NMR (300 MHz, CDCl3): d=8.26 (q, J=1.4, 6.6 Hz,
1H), 7.13–7.43 (m, 7H), 7.05 (q, J=3.8, 12.9 Hz, 2H), 4.06 (s, 2H);
13C NMR (75 MHz, CDCl3): d=196.6, 136.9, 136.3, 135.4, 133.8,
133.1, 132.4, 131.4, 130.6, 129.3, 128.8, 128.0, 127.3, 126.9, 48.4;
MALDI HRMS: m/z 243.0767 [M+Na+]; calcd for C16H12NaO+:
243.0780.
5,6-Dihydro-dibenzo[a,e]cycloocten-5-ol (6): Sodium borohydride
(0.757 g, 20 mmol) was slowly added to a stirred solution of 5
(2.20 g, 10 mmol) in a mixture of EtOH and THF (1:1, v/v, 120 mL).
The reaction mixture was stirred for 7 h, after which TLC analysis
indicated completion of the reaction. The reaction was quenched
by slow addition of acetic acid (1 mL) and the solvents were
3-Hydroxypropyl-carbamic acid 5,6-dihydro-11,12-didehydro-
dibenzo[a,e]cycloocten-5-yl ester (9): 3-Aminopropan-1-ol (15 mg,
0.2 mmol) and triethylamine (10 mL) were added to a stirred solu-
tion of 8 (38 mg, 0.1 mmol) in CH2Cl2 (15 mL). The reaction mixture
was stirred at room temperature for 12 h, after which the solvents
1918
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ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemBioChem 2011, 12, 1912 – 1921

Labeling Cellular Glycoconjugates by SPAAC
were evaporated under reduced pressure and the residue was puri-
residue was purified by flash chromatography over Iatrobeads
(MeOH/CH2Cl2, 4!15%, v/v) to give 13 as a light-yellow solid
(56 mg, 63%). 1H NMR (300 MHz, CD3OD): d=7.73 (d, J=7.6 Hz,
1H), 7.30–7.56 (m, 7H), 4.48 (m, 2H), 4.35 (d, J=14.7 Hz, 1H), 3.50–
3.68 (m, 8H), 3.24 (d, J=13.6 Hz, 1H), 3.08 (d, J=5.3 Hz, 2H), 3.02
(d, J=5.3 Hz, 2H); 13C NMR (75 MHz, CD3OD): d=172.2, 162.0,
150.0, 149.0, 133.2, 131.0, 130.6, 129.8, 129.3, 128.5, 127.1, 126.8,
fied by silica gel column chromatography (CH2Cl2/CH3OH, 20:1, v/v)
1
to afford 9 (25 mg, 77%). H NMR (CDCl3, 300 MHz): d=6.94–7.43
(m, 8H, aromatics), 5.42 (m, 1H, Ph-CH-O), 3.61(m, 2H, CH2OH),
3.30 (m, 2H, CH2NH), 3.08 (dd, J=15.0, 1.8 Hz, 1H, PhHCH), 2.84
(dd, J=15.0, 3.9 Hz, 1H, PhHCH), 1.53–1.68 (m, 2H, CH2CH2OH);
13C NMR (75 MHz, CDCl3): d=150.8, 149.1, 128.9, 128.0, 127.0,
126.1, 126.0, 125.9, 125.8, 125.3, 125.1, 125.0, 122.8, 122.6, 120.3,
111.9, 108.9, 58.6, 45.2, 36.8, 36.7, 31.6. MALDI HRMS: m/z 344.1246
[M+Na+]; calcd for C20H19O3NNa+: 344.1257.
124.9, 124.3, 111.2, 110.5, 73.8, 71.4, 71.3, 70.4, 67.8, 40.6, 39.7, 39.1.
MALDI HRMS: m/z 444.1875 [M+Na+]; calcd for C24H27N3NaO4
:
+
444.1894.
{2-[2-(2-Amino-ethoxy)ethoxy]ethyl}carbamic acid 5,6-dihydro-
11,12-didehydro-dibenzo[a,e]cycloocten-5-yl ester (10): Et3N
(0.139 mL, 1.0 mmol) was added to a stirred solution of 8 (77 mg,
0.2 mmol) and tris(ethylene glycol)-1,8-diamine (0.293 mL, 2 mmol)
in CH2Cl2 (20 mL). The reaction mixture was stirred for 3 h, after
which the solvent was removed under reduced pressure. The resi-
due was purified by flash chromatography over Iatrobeads (MeOH/
CH2Cl2, 8!30%, v/v) to give 10 as a light-yellow amorphous solid
(0.063 g, 80%). 1H NMR (300 MHz, CDCl3): d=7.51 (d, 1H, J=
7.3 Hz), 7.24–7.37 (m, 7H), 5.81 (s, 1H, NH), 5.48 (brs, 1H), 3.50–
3.68 (m, 8H), 3.39 (m, 2H), 3.16 (d, J=14.8 Hz, 1H), 2.91 (brs, 2H),
2.88 (d, J=14.8 Hz, 1H), 2.57 (brs, 2H, NH2); 13C NMR (75 MHz,
CDCl3): d=155.7, 152.2, 151.1, 130.0, 128.1, 128.0, 127.2, 127.1,
126.3, 126.0, 123.9, 123.8, 121.3, 113.0, 110.0, 76.7, 72.8, 70.3, 70.2,
70.1, 70.0, 46.2, 41.5, 41.0. MALDI HRMS: m/z 417.1766 [M+Na+];
calcd for C23H26N2NaO4+: 417.1785.
{2-[2-(2-Biotinylamino-ethoxy)ethoxy]ethyl}carbamic acid 5,6-di-
hydro-11,12-didehydro-dibenzo[a,e]cycloocten-5-yl ester (14):
Three drops of Et3N were added to a solution of 10 (15.8 mg,
0.04 mmol) and biotin-OSu (20.5 mg, 0.06 mmol) in MeOH/CH2Cl2
(1:1, v/v, 7 mL). The reaction mixture was stirred for 2 h, and then
the solvents were removed under reduced pressure. The residue
was purified by column chromatography over Iatrobeads (MeOH/
CH2Cl2, 5!15%, v/v) to give 14 as a light-yellow amorphic solid
1
(22.1 mg, 89%). H NMR (300 MHz, CD3OD): d=7.59 (1H, aromat-
ics), 7.42–7.33 (7H, aromatics), 5.44, (dd, J=5.0, 14.1 Hz, 1H,
ArCHOH), 4.60, 4.46 (m, 2H, CHNH), 4.24 (s, 4H, OCH2CH2O), 3.72
(m, 4H, OCH2), 3.64 (m, 2H, CH2NH), 3.55 (m, 1H, CHS), 3.33 (dd,
J=4.8, 12.0 Hz, 1H), 3.23 (t, J=6 Hz, 2H, CH2NH2), 3.22, (dd, J=5.0,
14.1 Hz, 1H, CH2), 2.88, (dd, J=5.0, 14.1 Hz, 1H, CH2), 2.68 (d, J=
12.45 Hz, 1H), 2.20 (t, J=7.5 Hz, 2H, CH2CO), 1.4 (m, 6H, biotin-
CH2); 13C NMR (75 MHz, CD3OD): d=175.0, 164.9, 156.9, 152.5,
151.3, 129.9, 128.2, 128.1, 127.2, 127.1, 126.0, 125.7, 123.8, 121.2,
112.7, 109.8, 76.8, 70.2, 70.1, 69.8, 69.4, 62.1, 60.4, 55.8, 54.6, 46.0,
42.6, 40.6, 39.9, 39.1, 35.5, 28.6, 28.3, 25.6, 17.5, 16.1, 12.0; MALDI
HRMS: m/z 643.2575 [M+Na+]; calcd for C33H40N4NaO6S+ 643.2561.
6H-11,12-Didehydro-dibenzo[a,e]cyclooctatrien-5-one (11): Dess–
Martin reagent (0.40 g, 0.94 mmol) was added to a stirred solution
of 3 (0.172 g, 0.78 mmol) in CH2Cl2 (40 mL). The mixture was stirred
for 0.5 h after which TLC analysis indicated completion of the re-
action. The reaction mixture was filter through a short pad of silica
gel, which was washed with CH2Cl2. The filtrate was concentrated,
and the residue was purified by flash chromatography over silica
gel (hexanes/CH2Cl2, 1:1!0:1, v/v) to give 11 as a white amor-
phous solid (0.158 g, 92%). 1H NMR (300 MHz, CDCl3): d=7.29–
7.57 (m, 8H), 4.17 (d, J=10.6 Hz, 1H), 3.64 (d, J=10.6 Hz, 1H);
13C NMR (75 MHz, CDCl3): d=200.4, 154.7, 148.2, 131.21 (2C),
131.18, 129.3, 128.2, 127.8, 126.3, 125.9, 122.2, 111.1, 109.4, 49.3.
MALDI HRMS: m/z 241.0638 [M+Na+]; calcd for C16H10NaO+:
241.0624.
N-{2-[2-(2-Biotinylamino-ethoxy)ethoxy]ethyl}-2-(6H-11,12-dide-
hydro-dibenzo[a,e]cycloocten-5-ylideneaminooxy)acetamide
(15): Two drops of Et3N were added to a solution of 13 (8.4 mg,
0.02 mmol) and biotin-OSu (10.2 mg, 0.03 mmol) in MeOH/CH2Cl2
(1:1, v/v, 4 mL). The reaction mixture was stirred for 2 h, after
which the solvents were removed under reduced pressure. The res-
idue was purified by column on Iatrobeads (MeOH/CH2Cl2, 5!15,
v/v) to give 15 as a light-yellow solid (11.9 mg, 92%). 1H NMR
(300 MHz, CD3OD): d=7.64 (1H, aromatics), 7.54–7.30 (7H, aromat-
ics), 4.47 (m, 3H), 4.34 (m, 1H), 3.67 (s, 4H, OCH2), 3.60–3.40 (m,
3H), 3.30 (m, 2H), 3.20 (m, 3H), 2.96–2.86 (m, 2H), 2.20 (t, J=7 Hz,
2H, biotin-CH2CO), 1.80–1.50 (m, 4H, biotin-CH2) 1.48–1.38 (m, 2H,
biotin-CH2). MALDI HRMS: m/z 670.2665 [M+Na+]; calcd for
C34H41N5NaO6S+ 670.2670.
(6H-11,12-didehydro-dibenzo[a,e]cycloocten-5-ylideneamino-
oxy)acetic acid (12): A solution of 6H-11,12-didehydro-dibenzo-
[a,e]cyclooctatrien-5-one (11; 21.8 mg, 0.1 mmol) and (carboxyme-
thyl)hydroxylamine hemihydrochloride (21.8 mg, 0.2 mmol) in
MeOH/CH2Cl2/HOAc (1:1:0.02, v/v/v, 8 mL) was stirred for 2 days.
The solvents were removed under reduced pressure, and the resi-
due was purified by flash chromatography on silica gel (EtOAc) to
give 12 as an amorphous white solid (17.8 mg, 61%). 1H NMR
(300 MHz, CDCl3): d=7.54 (d, J=7.4 Hz, 1H), 7.46 (d, J=7.4 Hz,
1H), 7.18–7.39 (m, 6H), 4.53 (m, 2H), 4.23 (d, J=12.8 Hz, 1H), 3.16
(d, J=12.8 Hz, 1H); 13C NMR (75 MHz, CDCl3): d=175.2 and 173.6,
154.1 153.2, 130.7,129.5, 19.3, 129.2, 129.1, 128.1, 128.0, 127.1,
126.9, 125.5, 125.2, 122.7, 113.9, 111.2, 84.7, 68.3 and 67.1, 35.0 and
33.2. MALDI HRMS: m/z 314.0770 [M+Na+]; calcd for
C18H13NNaO3+: 314.0788.
General procedure for the preparation of triazoles (16–19): A so-
lution of dibenzocyclooctyne derivative (3, 9, 11 or 12, 0.1 mmol)
and benzyl azide (0.1 mmol) in MeOH (10 mL) was stirred for 3 h.
The solvent was evaporated under reduced pressure, and the resi-
due was purified by silica gel column chromatography to give the
desired product (16, 17, 18 or 19, respectively) in a quantitative
yield.
1
Compound 16: H NMR (300 MHz, CD3OD): d=8.29–7.98 (1H, aro-
matic), 7.70–6.80 (12H, aromatics), 5.84–5.25 (2H, PhCH2), 5.31–4.70
(1H, CHOH), 3.77–2.55 (2H, ArCH2). MALDI HRMS: m/z 377.1507
[M+Na+]; calcd for C23H20N3NaO+ 377.1499.
N-{2-[2-(2-Amino-ethoxy)ethoxy]ethyl}-2-(6H-11,12-didehydro-
dibenzo[a,e]cycloocten-5-ylideneaminooxy)acetamide (13): A so-
lution of 11 (46 mg, 0.211 mmol), N-{2-[2-(2-amino-ethoxy)ethox-
y]ethyl}-2-aminooxy-acetamide (84 mg, 0.251 mmol) and acetic
acid (0.1 mL) in MeOH/CH2Cl2 (1:1, v/v, 4 mL) was stirred for 2 days.
The solvents were evaporated under reduced pressure, and the
1
Compound 17: H NMR (300 MHz, CD3OD): d=8.00–6.90 (13H, aro-
matics), 6.03–5.26 (2H, PhCH2), 5.10–4.77 (1H, CHOCO), 3.58 (2H,
CH2OH), 3.29 (2H, NHCH2), 3.20–2.64 (2H, ArCH2), 1.62 (2H,
CH2CH2OH). MALDI HRMS: m/z 477.1889 [M+Na+]; calcd for
+
C27H26N4NaO3 477.1897.
ChemBioChem 2011, 12, 1912 – 1921
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chembiochem.org
1919

R. Steet, G.-J. Boons et al.
1
Compound 18: H NMR (300 MHz, CD3OD): d=8.09–6.92 (13H, aro-
Ac4GalNaz (100 mm) and as control cells Ac4ManNAc or Ac4GalNac
(100 mm) for 2 days. Expected cell number on day of click chemistry
was approximately 1ꢁ106 per well.
matics), 5.63 (2H, PhCH2), 3.70 (2H, COCH2). MALDI HRMS: m/z
375.1328 [M+Na+]; calcd for C23H18N3NaO+ 375.1342.
1
Compound 19: H NMR (300 MHz, CD3OD): d=7.59–7.06 (11H, aro-
Sialidase pretreatment: Cells were washed twice with serum-free
culture medium and incubated with V. cholerae neuraminidase
(50 mUmLꢀ1) in serum-free culture medium for 2 h at 378C,
washed in PBS and subjected to the respective assay.
matics), 7.04–6.97 (2H, aromatics), 5.89–5.32 (2H, PhCH2), 4.68–4.43
(2H, CH2CO2H), 4.30–4.03 (1H), 3.07–2.96 (1H). MALDI HRMS: m/z
447.1439 [M+Na+]; calcd for C25H20N4NaO3+: 447.1428.
Fluorescence measurement: The fluorescence spectra of com-
pounds 11 and 18 (excited at 375 nm) and 12 and 19 (excited at
313 nm) in methanol (10ꢀ5 m) were recorded on a spectrofluorome-
ter FluoroMax-3 from Horiba Jobin Yvon.
Click chemistry and detection by fluorescence intensity: Jurkat
cells bearing azides and control cells were washed with labeling
buffer (DPBS, pH 7.4 containing 1% FBS and 1% BSA) and trans-
ferred to round-bottom tubes (1ꢁ106 cells per sample). CHO cells
(untreated or sialidase pretreated) were left in the 12-well plates
(~1ꢁ106 cells per sample) and washed with labeling buffer. Next,
cells were incubated with the biotinylated compounds 14 or 15
(0–100 mm) in labeling buffer for 0–90 min at room temperature.
The cells were washed three times with cold labeling buffer and
then incubated with avidin–FITC (5 mgmLꢀ1) for 15 min at 48C in
the dark. Following three washes and cell lysis in passive lysis
buffer (Promega), lysates were analyzed for fluorescence intensity
(lem =520/lex =485) by using a microplate reader (BMG Labtech).
Data points were collected in triplicate and are representative of
three separate experiments. Fluorescence of Jurkat cell lysates was
expressed as fluorescence (arbitrary units; AU) per 800000 cells.
CHO cell lysates were assayed for total protein by using the bicin-
choninic acid assay (BCA; Pierce Biotechnology) and fluorescence
intensity was expressed as fluorescence (AU) per mg total protein.
Kinetics of the cycloaddition reaction: The rate measurements of
cycloaddition of cyclooctynes with benzyl azide were conducted
by UV spectroscopy at 25ꢁ0.18C. A calculated amount of solu-
tions of benzyl azide required to achieve desired azide concentra-
tion (6ꢁ10ꢀ4–1.5ꢁ10ꢀ2 m) was added to a thermally equilibrated
solution of acetylene in MeOH (6ꢁ10ꢀ5 m). Reactions were moni-
tored by following the decay of the characteristic absorbance of
acetylenes at about 317 nm. Consumption of starting material fol-
lowed a first-order equation and the pseudo-first-order rate con-
stants were obtained by least-squares fitting of the data to a single
exponential equation. The rate dependence as a function of the
concentration of azide was linear. Least-squares fitting of the data
to a linear equation produced the bimolecular rate constants sum-
marized in Table 1. In this respect, the UV spectroscopic method
can be performed under pseudo-first-order conditions over a wide
range of reagent concentrations making the analysis of second-
order kinetic curves more reliable.
Lectin binding assay: Untreated or sialidase pretreated cells (~1ꢁ
106) were washed twice in cold PBS and subsequently incubated in
300 mL PBS containing PNA–FITC (50 mgmLꢀ1
) or RCA1–FITC
(50 mgmLꢀ1) for 45 min on ice in the dark. After being washed with
cold PBS, the cells were lyzed in passive lysis buffer (Promega) and
lysates were analyzed for fluorescence intensity (lem =520/lex =
485) by using a microplate reader.
Reagents for biological experiments: Synthetic compounds 14
and 15 were reconstituted in DMF and stored at ꢀ808C. Final con-
centrations of DMF never exceeded 0.56% to avoid toxic effects.
Ac4ManNAc, Ac4ManNAz, Ac4GalNAc and Ac4GalNAz were synthe-
sized as reported,[32] and reconstituted in ethanol. Avidin–FITC and
avidin–AlexaFluor488 were obtained from Molecular Probes,
V. cholerae neuraminidase was from Sigma–Aldrich, and PNA–FITC
and RCA1–FITC were from EY Laboratories.
Detection of cell labeling and lectin staining by fluorescence mi-
croscopy: For cell surface labeling, CHO-K1 cells labeled with
Ac4ManNAc or Ac4ManNAz (100 mm) for 2 days were seeded at a
density of 50000 cells per coverslip (22 mm) and allowed to
adhere, overnight, in their original medium. After two washes with
wash buffer (DPBS, supplemented with 1% FBS), live cells were in-
cubated with biotinylated compounds 14 or 15 (30 mm) in wash
buffer for 1 h at room temperature, followed by three washes in
wash buffer (10 min per wash). Next, the cells were incubated with
avidin conjugated with AlexaFluor488 (5 mgmLꢀ1) for 15 min at
48C. Cells were washed three times with wash buffer and fixed
with formaldehyde (3.7% in PBS) at room temperature for 15 min.
After the coverslips were washed four times in PBS (5 min per
wash), the nucleus was labeled with the far red-fluorescent TO-
PRO-3 iodide dye (Molecular Probes). The cells were mounted with
PermaFluor (Thermo Electron Corporation) before being imaged.
Cell culture conditions: Human Jurkat cells (clone E6-1; ATCC)
were cultured in RPMI 1640 medium (ATCC) with l-glutamine
(2 mm), adjusted to contain sodium bicarbonate (1.5 gLꢀ1), glucose
(4.5 gLꢀ1), HEPES (10 mm), and sodium pyruvate (1 mm). CHO cells
(clone K1; ATCC) were cultured in Kaighn’s modification of Ham’s
F12 medium (ATCC) with l-glutamine (2 mm), adjusted to contain
sodium bicarbonate (1.5 gLꢀ1). Mutant CHO cells (Lec2, Lec13,
Lec32 mutants were obtained from Dr. Pamela Stanley and Cog1
and Cog2 mutants (ldlB and ldlC) obtained from Dr. Monty Kreiger)
were cultured in minimum essential medium Alpha 1X (Cellgro)
with Earle’s salts, ribonucleosides, deoxyribonucleosides and l-
glutamine (2 mm). All media were supplemented with penicillin
(100 UmLꢀ1)/streptomycin (100 mgmLꢀ1
; Mediatech) and fetal
bovine serum (FBS, 10%; Hyclone). Cells were maintained in a
humid 5% CO2 atmosphere at 378C.
For lectin staining of cell surface glycans, CHO-K1 and Lec2 cells
were seeded at a density of 50000 cells per coverslip (22 mm) and
allowed to adhere, overnight. After two washes with serum-free
culture medium, live cells were treated with V. cholerae neuramini-
dase (50 mUmLꢀ1) in serum-free culture medium for 2 h at 378C.
Coverslips were washed with DPBS and incubated with PNA–FITC
(50 mgmLꢀ1) in PBS supplemented with BSA (1%) for 45 min at
48C. Cells were washed with PBS and fixed and mounted as above.
Cell surface azide labeling: Jurkat cells were seeded at a density
of 75000 cells per mL in a total volume of 40 mL culture medium
in the presence of Ac4ManNaz or Ac4GalNaz (25 mm final concentra-
tion) and grown for 3 days; this led to the metabolic incorporation
of the corresponding N-azidoacetyl sialic acid (SiaNAz) into their
cell surface glycoproteins. Control cells were grown in the pres-
ence of Ac4ManNac or Ac4GalNac (25 mm final concentration) for
3 days. CHO cells were plated in 12-well plates (250000 cells per
well) and grown in medium that contained Ac4ManNaz or
Initial analysis was performed on a Zeiss Axioplan2 fluorescent mi-
croscope. Confocal images were acquired by using a 60ꢁ (NA1.42)
oil objective. Stacks of optical sections were collected in the z di-
1920
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ChemBioChem 2011, 12, 1912 – 1921

Labeling Cellular Glycoconjugates by SPAAC
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Acknowledgements
This work was supported by grants from the National Cancer In-
stitute of the US National Institutes of Health (NIH/NCI
R01A088986, G.-J.B.) and NIH’s National Institute of General Medi-
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Received: February 16, 2011
Published online on June 9, 2011
ChemBioChem 2011, 12, 1912 – 1921
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chembiochem.org
1921