Sequential acid-catalyzed alkyl glycosylation and oligomerization of unprotected carbohydrates

Article abstract of DOI:10.1039/d0gc04198j

An efficient method has been developed to synthesize end-functionalized oligosaccharides from unprotected monosaccharides in a one-pot/two-step approach. In the first step, mannose (and glucose) was functionalized with an alkyne group at the anomeric position through the Fisher-glycosylation reaction with propargyl alcohol as a glycosyl acceptor. In the second step, the functionalized monosaccharides were oligomerized and the experimental conditions were optimized by varying the temperature, time and the molar ratio between alcohol and sugar to reach a up to 8. The obtained oligosaccharides showed complete propargylation at their reducing chain-end and were successfully coupled to oleic acidviathe Huisgen reaction, affording bio-based surfactants.

Full text of DOI:10.1039/d0gc04198j

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PAPER
Sequential acid-catalyzed alkyl glycosylation and
oligomerization of unprotected carbohydrates†
Cite this: Green Chem., 2021, 23,
1361
Lea Spitzer, a,b Sébastien Lecommandoux,b Henri Cramail*b and
a
François Jérôme
*
An efficient method has been developed to synthesize end-functionalized oligosaccharides from unpro-
tected monosaccharides in a one-pot/two-step approach. In the first step, mannose (and glucose) was
functionalized with an alkyne group at the anomeric position through the Fisher-glycosylation reaction
with propargyl alcohol as a glycosyl acceptor. In the second step, the functionalized monosaccharides
were oligomerized and the experimental conditions were optimized by varying the temperature, time and
the molar ratio between alcohol and sugar to reach a DPn up to 8. The obtained oligosaccharides showed
complete propargylation at their reducing chain-end and were successfully coupled to oleic acid via the
Huisgen reaction, affording bio-based surfactants.
Received 11th December 2020,
Accepted 15th January 2021
DOI: 10.1039/d0gc04198j
rsc.li/greenchem
meric alkyl glycosides tend to further react with unconverted
sugars, leading to the formation of APGs. Due to the release of
Introduction
Surfactants are ubiquitous molecules in our daily life with water, concomitant oligomerization of the sugar head and
multiple applications in various sectors such as home and per- hydrolysis of the glycosidic bond occur, leading to a thermo-
sonal care, paint, food, medicine, water treatment, materials, dynamic equilibrium in which APGs are composed, on
etc.1–3 Due to the exponential increase of the world population, average, of 1.5 to 2.1 sugar units per fatty chain. Being able to
and also due to several emerging economies worldwide, our increase the degree of polymerization ðDPnÞ of sugar units per
society’s demand for surfactants is increasing dramatically. fatty chain is of great interest, as it modifies the hydrophilic–
This market, of about 12 million tons per year, is expected to lipophilic balance (HLB) of APGs, and consequently their per-
grow at an annual rate of 4.3%.4 Because of environmental formances. For instance, this can tune their physicochemical
concerns, the manufacture of bio-based surfactants has properties in terms of water solubility, foaming properties,
become increasingly attractive, as it potentially provides more critical micelle concentrations, surface tensions, etc. However,
environmentally friendly products, although a full life cycle this elongation of the sugar moiety remains a very challenging
assessment should be systematically carried out to support task. The main hurdle is the in situ release of water, which
this claim.3,5–9 The market of bio-based surfactants10 is about induces a rapid hydrolysis of glycosidic bonds, even when
3.5 million tons per year in Europe, a region where the annual water is continuously distilled out. Furthermore, at the end of
growth rate of bio-based surfactants was 3% between 2008 and the reaction, the removal of the excess of fatty alcohols by dis-
2013, with alkylpolyglycosides (APGs) registering the strongest tillation is a very delicate step as it generally requires elevated
growth.11–14
temperatures, leading to the partial degradation of APGs. So
APGs are industrially produced through the acid-catalyzed far, all attempts to produce APGs with a DPn larger than
Fischer glycosylation reaction.14–17 This reaction couples 2.1 have failed. One should mention that enzymatic catalysis
unprotected monomeric sugars with long chain alkyl alcohols potentially paves the way to APGs with a larger DPn, but the
(C5–C18) and releases a stoichiometric amount of water. cost of the enzymes and the low reactor productivity hamper
During the Fisher glycosylation, the in situ produced mono- the implementation of these routes on a bigger scale.18–21
In this work, we propose an alternative straightforward
strategy based on the use of propargyl alcohol (PGA), which
aINCREASE/Institut de Chimie des Milieux et Matériaux de Poitiers, CNRS,
Université de Poitiers, 1 rue Marcel Doré, 86073 Poitiers cedex 9, France.
E-mail: francois.jerome@univ-poitiers.fr
serves not only as a glycosyl acceptor but also as a linker to
subsequently introduce the fatty chain through a 100% atom-
economical copper-catalyzed Huisgen reaction. In contrast to
fatty alcohols, at the end of the reaction, PGA can be con-
veniently separated by distillation at a low temperature, limit-
ing the degradation of APGs. Importantly, as sugars are more
bINCREASE/Université de Bordeaux, CNRS, Bordeaux INP, LCPO, UMR 5629,
F-33600 Pessac, France
†Electronic supplementary information (ESI) available: NMR, GC-FID, ESI-MS
and MALDI-TOF spectra. See DOI: 10.1039/d0gc04198j
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soluble in PGA than in fatty alcohols, oligomerization takes Characterization methods
place to a larger extent during the evaporation step of PGA,
leading to the formation of APGs with a DPn of 8.
Nuclear magnetic resonance (NMR). NMR spectra were
obtained on a Bruker Advance III 400. Typically, 10–20 mg of
the sample was diluted in 0.5 mL of deuterated solvent before
the measurement. The following reference values of deuterated
solvents were used: D2O (1H-NMR: δ = 4.8 ppm). The spectral
data were analyzed using TopSpin (v. 4.0.9). Automatic phase
correction and subsequent integration were applied.
Experimental
Chemicals
D-(+)-Mannose (from wood, ≥99%), D-(+)-glucose (≥99.5%) and
Amberlyst-15 (≥90%, 0.355–1.18 mm, 1.7 ml L−1 capacity) were
purchased from Sigma Aldrich, and dried under vacuum
before use. GC-FID standards propargyl α-D-mannopyranoside
(PMan) and 1,6-anhydro-β-D-mannopyranose (AMP) were pur-
chased from Biosynth Carbosynth. Propargyl alcohol (99%),
BSTFA (+1% TMCS), D-sorbitol (≥98%), 1,5,7-triazabicyclo
[4.4.0]dec-5-ene (98%, TBD), sodium azide (reagent plus
≥99.5%), L-ascorbic acid (99%) and 3-bromo-1-propanol (97%)
were purchased from Sigma Aldrich and used without further
purification. Methyl oleate (MeOI ≥ 99.9%) was obtained from
Nuchekrep. Cuprisorb was purchased from Seachem and
copper(II) sulfate from ProLabo. Acetone, acetonitrile, chloro-
form (CDCl3), diethyl ether, dimethyl sulfoxide (DMSO), ethyl
acetate (EtOAc), petroleum ether (PE), pyridine and mag-
nesium sulfate were purchased from Sigma Aldrich and used
without further purification. Dialysis was performed using a
Spectra/Por®6 MWCO 100–500 Da membrane (RC).
Matrix assisted laser desorption/ionisation-time of flight
(MALDI-TOF). MALDI-MS spectra were recorded by CESAMO
(Bordeaux, France) on an Autoflex maX TOF mass spectro-
meter (Bruker Daltonics, Bremen, Germany) equipped with a
frequency tripled Nd:YAG laser emitting at 355 nm. Spectra
were recorded in the positive-ion linear mode and with an
accelerating voltage of 19 kV. The samples were dissolved at
20 mg mL−1 (water : acetonitrile 70/30 + trifluoroacetic acid
(TFA, 10 vol%). A 2,5-dihydroxybenzoic acid (DHB) matrix solu-
tion was prepared by dissolving 10 mg in 1 mL of acetonitrile.
The solutions were combined in 10 : 1 v/v of matrix to sample.
One to two microliters of the obtained solution were deposited
onto the sample target and vacuum-dried.
Size exclusion chromatography (SEC). Polymer molar masses
were determined by Size Exclusion Chromatography (SEC) using
water as the eluent. Measurements in water were performed on
an Ultimate 3000 system from Thermoscientific equipped with
a diode array detector (DAD). The system also includes a multi-
angle laser light scattering detector MALLS and a differential
refractive index detector dRI from Wyatt technology. Polymers
Synthesis procedure
General method of the Fischer glycosylation and oligomeri- were separated on two Shodex OH Pack 802.5 (8 × 300) columns
zation of mannose. - Step 1 -
(exclusion limits from 500 Da to 10 000 Da) at a flow rate of
In a round-bottom flask, mannose (5 g, 27.75 mmol) was 0.6 mL min−1. The column temperature was held at 25 °C.
dispersed in an excess of propargyl alcohol (5 eq.) and stirred Dextran from PSS was used as the standard.
at 80 °C in the presence of amberlyst-15 (4.2 mol% H+) for 3 h.
Attenuated total reflectance infrared spectroscopy (ATR-IR).
Then, the reaction was stopped, and the as-obtained yellow/ Fourier transform infrared (FTIR) spectra were recorded on a
orange solution was separated by centrifugation from the cata- Bruker VERTEX 70 instrument (4 cm−1 resolution, 64 scans,
lyst. The as-recovered solution (pH 4) was used for the follow- DLaTGS MIR) equipped with a Pike GladiATR plate (diamond
ing step without further purification.
- Step 2 -
crystal) for attenuated total reflectance (ATR) analysis at room
temperature.
Oligomerization was attained by removing the excess of pro-
Gas chromatography with flame-ionization detection
pargyl alcohol through heating of the mixture of step 1 at (GC-FID). The composition of the mono- and disaccharide
100 °C for 4 h under vacuum. Note that for analysis only, the fractions was analyzed by GC-FID spectroscopy after the trans-
trace amount of free propargyl alcohol was fully removed as formation of the sugar components into their corresponding
follows: (PMan)n was dissolved in water, and a trace of propar- per-O-trimethylsilyl (nonreducing sugars) or per-O-trimethyl-
gyl alcohol was extracted with chloroform. The aqueous phase silylated oxime (reducing sugars). The crude samples (in
was then freeze-dried to give a brown-beige solid (4.3 g, 88%). general 20–30 mg) were dissolved in 1 mL of pyridine (contain-
It could be additionally purified by filtration over active ing 1 mg mL−1 sorbitol as an internal standard). To 100 µL of
carbon, yielding a white powder (S1). Đ (SEC) = 2.03; ATR-IR = the resulting solution was then added 200 µL of BSTFA (+1%
2116 (CuC str.) (s); 1H-NMR (400 MHz, D2O): δ = 5.32–5.08 (m, TMCS), and the mixture was stirred at RT for 2 h. During this
1.66 H), 5.08–5.01 (db, 2 H), 4.97–4.90 (db, 3.23 H), 4.73 (d, operation, a white precipitate was observed, which was separ-
0.44 H), 4.41–4.32 (m, 6.56 H, CH2), 4.15–3.53 (m, 49.86 H, ated by filtration (0.45 µm, regenerated cellulose) before injec-
α/β-H2–α/β-H-6), 3.42–3.39 (tq, 0.74 H, β-H-5), 2.95 (tb, 1 H, tion into the apparatus.22 By this procedure, the non-reducing
alkyne); 13C-NMR (100.4 MHz, D2O): δ = 102.4, 102.3, 100.5, sugars gave single peaks in the chromatogram, whereas the
99.5, 99.2, 99.0, 98.6, 97.0, 78.7, 78.3, 78.1, 76.2, 73.2, 72.9, reducing sugars afforded two peaks for the corresponding syn
72.7, 71.3, 70.6, 70.5, 70.3, 69.9, 69.8, 66.7, 66.4, 65.4, 60.9, and anti TMS-oximes. The identification and quantification
54.7, 54.6.
were achieved by comparison with authentic standards for
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which the response factors were obtained from the corres-
ponding calibration curves using sorbitol as the internal stan-
dard. The analysis was conducted with a GC Trace 1300 Gas
Chromatograph (Thermoscientific) equipped with a split/split-
ness injector and flame ionization detector (FID). A H2 flow rate
of 34 mL min−1 and an airflow rate of 350 mL min−1 were used.
The flow rate of the carrier gas (H2) was set at 1.2 mL min−1. The
temperatures of the injection port and detector were set at 250 °C
and 320 °C, respectively. The oven temperature was programmed
to initiate at 90 °C for 1 min, and then the temperature was
raised to 220 °C at a rate of 10 °C min−1 and finally increased to
320 °C at a rate of 40 °C min−1 and held there for 5 min. The
injection volume was 1 μL in the split injection mode (15 : 1).
Separation was performed on a capillary column TRB-5MS (30 m
× 0.25 mm × 0.25 mm film thickness) from Teknokroma with
matrix 95% dimethyl-(5%) diphenyl polysiloxane.
Fig. 1 Typical GC-FID spectrum collected after step 1.
(LVM, peak 1) and also minor products (such as furanoside
peak 6 and acyclic acetal peak 7), in a relative proportion of
20%, 3% and 8%, respectively. The retention times of standard
chemicals are given in Fig. S2 and Table S1.† By means of size
Results and discussion
The present study should be viewed as a two-step process in
which step 1 is the acid-catalyzed glycosylation of monomeric
sugars with PGA, and step 2 is the elongation of the sugar
moiety (Scheme 1). As a case study, mannose was first selected.
Transposition to glucose is discussed in the second part.
exclusion
chromatography,
propargyl-(oligo)-mannosides
(PMan)n were also detected (Fig. S3†). In line with previous
1
reports, the DPn of (PMan)n, determined by H-NMR (see later
for more information) was 1.3, showing that oligomerization
occurs to a very low extent under these conditions.
Step 1. Glycosylation of mannose with PGA
A plot of the PMan yields as a function of reaction time is
provided in Fig. 2. It shows that the PMan yield and the
mannose conversion levelled off around 70% and 90%,
respectively, after 7 h of reaction. Extending the reaction time
from 7 to 48 h did not result in the complete conversion of
mannose, suggesting that the system has reached a thermo-
dynamic equilibrium.
In the first set of experiments, unprotected mannose was sus-
pended in a 5-fold excess of PGA, the latter serving as both a
glycosyl acceptor and solvent. The resulting mixture was
stirred at 80 °C in the presence of Amberlyst-15, a solid acid
catalyst (4.2 mol% of H+). The reaction was monitored by gas
chromatography (GC) over a period of 48 h. A sequential oxi-
mation–trimethylsilylation derivatization procedure was used
to calculate the conversion of mannose and to quantify the
mono- and disaccharide fractions.
We next studied the influence of temperature (Fig. 3).
Decreasing the reaction temperature from 100 °C to 80 °C
After 48 h (Fig. 1), GC clearly revealed a conversion of
mannose (α-mannose, peak 2, β-mannose, peak 4) of 93%,
accompanied by the formation of α/β propargyl-(mono)-man-
nopyranoside in 69% yield (PMan, peak 3 and 5). Other
detected products were disaccharides (peak 8), levomannosane
Scheme 1 Summarized method of the preparation of (PMan)n through
2 steps: step 1 = mannose (1 eq.) + PGA (5 eq.) + Amberlyst-15
(4.2 mol%), 80 °C, 1–12 h; step 2 = vacuum at 100 °C for 4–5 h. For the
sake of clarity, only the pyranoside form is represented.
Fig. 2 Mannose conversion and PMan yield as a function of reaction
time (80 °C, PGA/mannose molar ratio = 5, Amberlyst-15 (4.2 mol% H+)).
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Fig. 4 Effect of the PGA/mannose molar ratio on the PMan formation
rate (80 °C, Amberlyst-15 (4.2 mol% H+)).
Fig. 3 Effect of the reaction temperature on the PMan formation rate
(PGA/mannose molar ratio = 5, Amberlyst-15 (4.2 mol% H+)).
obviously decreased the reaction rate (from 23 mol h−1 to
9.5 mol h−1). For instance, after 3 h of reaction, 90% and 79%
of mannose were converted at 100 °C and 80 °C, respectively.
When lowering the temperature further to 60 °C, we faced pro-
blems of immiscibility between mannose and PGA.
However, at 60 °C, the reaction started after 6 h, and a con-
version of mannose of about 80% was observed after 12 h of
reaction (2.5 mol h−1). This 6 h induction period corresponds
to the time to completely dissolve mannose in PGA at 60 °C.
Seeking for a compromise between the stability of mannose
and the reaction rate, the reaction temperature was fixed at
80 °C in the following experiments. Note that decreasing the
PGA/mannose molar ratio from 5 to 3 slowed down the reac-
tion rate from 6.2 mol h−1 to 4.5 mol h−1, tentatively attributed
to an increase of the reaction medium viscosity. However, this
did not significantly impact either the PMan yield (Fig. 4) or
the DPn of (PMan)n recovered in step 1, confirming that the
Fig. 5 Overlaid SEC (measured against dextran-standards in H2O at
in situ released water prevents oligomerization to a large 25 °C) spectra during step 2 (100 °C, vacuum).
extent.
Step 2. Oligomerization of the sugar head
To get more information on the structures of the oligomers
formed, NMR investigations were performed (Fig. 6). Owing to
the low field shift of the alkyne proton (2.95 ppm), the end-
group method could be employed to calculate the degree of oli-
gomerization, and indirectly the average molar mass. The DPn
and Mn values were calculated from the 1H-NMR spectra using
the following equations:
In order to increase the DPn of PMan, the reaction mixture was
then heated under vacuum at 100 °C to distill out the excess of
PGA and the in situ released water. The time needed to evapor-
ate the excess of PGA (4 h) restricted the duration of the reac-
tion. The effect of this treatment on the average DPn was moni-
tored by SEC. As shown in Fig. 5, after 4 h under vacuum, a
shift towards high molar masses was clearly observed, a result
consistent with the elongation of the sugar moiety. This treat-
ment was accompanied by the formation of a beige-brownish
powder. After complete removal of PGA, only 5–8% of mono-
meric species remained (determined by gas chromatography).
ðH2 þ H3 þ H4 þ H5 þ H6Þ
DPn ¼
ð1Þ
ð2Þ
6
Mn ¼ 179 þ M0DPn þ 201
The molar mass of the total batch was 1353 g mol−1 (i.e., where M0 is the repeating unit with a molar mass of 162 g
DPn ¼ 8) with a dispersity, Đ, of around 2.
mol−1. The values of 179 g mol−1 and 201 g mol−1 are derived
Note that distilled PGA can be reused for further glycosyla- from the end groups of (PMan)n (Fig. S4†). A detailed analysis
tion reactions in step 1, but only after drying over magnesium by one- and two-dimensional NMR spectroscopy, including the
sulfate to remove traces of water.
nJH–H and nJC–H couplings, is given in the ESI (Fig. S5–S7†).
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Fig. 6 1H-NMR (400 MHz, D2O) spectrum of (PMan)n; DPn ¼ 8, Mn ¼ 1353 g molꢀ1
.
The signals at 2.95 ppm (brown, HCu) and 4.32–4.41 ppm to a lower extent, through α-(1,2)-glycosidic linkages (Fig. S9
(green, CH2) confirmed the coupling of oligomannosides with and Table S2†). One should note that the additional peak at
PGA (Fig. 6). Indeed, these signals chemically shifted down- 4.73 ppm suggests the existence of β-glycosidic linkages. This
field as compared to pure PGA (2.80 ppm (HCu) and was supported by the proton–proton long range coupling to
4.20 ppm (CH2)). The broad peak at 5.01–5.08 ppm was β-H-5 (3.41 ppm) in the NMR NOESY spectrum (Fig. S6†).
assigned to the anomeric proton (blue, α-H-1) located in the α However, due to the poor intensity in the 2D-NMR spectra,
position at the chain end of (PMan)n. The assignment of this they were present in very low percentages. Integration of the
proton could be further confirmed either with the reference different anomeric signals in the 1H NMR spectrum (Fig. 6)
signal of pure monomeric PMan at 5.06–5.05 ppm suggested that the oligomannoside chain is composed of 60%
(Fig. S8†)23,24 or with a coupling constant of 4 Hz between of α-(1,6)-linkages, 24% of α-(1,2)-linkages, and only 8% of
α-H-1 and the –CH2– of PGA in the NMR HMBC spectrum β-(1,6) and β-(1,2) linkages. Another small doublet found at
(Fig. S7†). The spectrum in Fig. 6 showed additional peaks in 5.44 ppm accounted for 5% relative to all anomeric protons
the range of 4.90 to 5.32 ppm. This region is characteristic of and was assigned to the anomeric proton of levomannosane
the anomeric protons and was thus attributed to the glycosidic (LVM). This chemical composition is also supported by 13C
linkages between the anhydromannose units in the oligoman- NMR (Fig. S10†). This structure is actually very similar to what
noside chain. The most intense peak at 4.93 ppm (yellow in was previously observed in the oligomerization of mannose by
Fig. 6) was assigned to the anomeric proton of the α-(1,6)-gly- high frequency ultrasound,26 the 1–6 linkages being thermo-
cosidic linkage. This claim is supported by NMR NOESY dynamically the most favorable ones.
experiments (Fig. S6†) showing a proton–proton long range
To further support this chemical composition, the samples
coupling of this anomeric proton with the H-6 protons of the were analyzed by MALDI-TOF spectrometry (Fig. 7). The data
anhydromanose unit. The α configuration was also supported showed a series of monoanionized peaks with sequential
by the HMBC spectrum with a carbon–proton coupling of increments of m/z = 162 Da (corresponding to the anhydro-
171.32 Hz (Fig. S7†).25 The additional multiplet signals appear- mannose residue as the repeating unit). This series was
ing between 5.08 and 5.32 ppm were tentatively attributed to assigned to (PMan)n, and DP up to 12 was detected, which
the α-(1,2)-glycosidic linkages, as COSY- and NOESY NMR corresponds to a DPn of 8 as calculated by NMR (eqn (1)).
spectroscopy (Fig. S5 and S6†) showed an exclusive H,H-coup- Within the noise of the MS spectra, a second series, with m/z
ling with H-2 protons of the mannose ring (4.1–3.98 ppm). On lower than 56 as compared to (PMan)n, was observed and cor-
the basis of the signal intensity, the oligomannoside chain is responded to the formation of an oligomannoside terminated
dominantly assembled through α-(1,6)-glycosidic bonds and, by a dehydrated anydromannose residue. The formation of
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Fig. 7 (I) MALDI-TOF spectrum of (PMan)n recovered after step 2; (II) proposed structures.
this oligomannoside series remained however quasi negligible, The elongation of the mannose moiety to form APGs occurs to
as also supported by NMR investigations. This combined a very low extent in step 1, mainly due to the excess of PGA
NMR-mass analysis led us to conclude that the as-obtained oli- and the in situ release of water which prevents the oligomeriza-
gomannosides are (1) fully propargylated at their terminal end, tion reaction. In step 2, PMan is reprotonated, partly regenerat-
(2) mainly linked through α-(1,6)-linkages and (3) exhibit a DPn ing the oxocarbenium ion, as previously suggested by
up to 8. By means of DSC analysis, the Tg of (PMan)8 was Mamidyala et al.30 During the distillation of PGA (and water)
found to be 120 °C.
under vacuum, the reaction of the oxocarbenium ion with
To further increase the DPn of (PMan)n, step 1 was stopped PMan is statistically more likely to occur, leading to the for-
before the complete conversion of mannose; this should facili- mation of (PMan)n with a DPn of 8. To confirm the mechanism
tate the reaction of PMan with the unreacted mannose in step of step 2, PMan was prepared, isolated by chromatography over
2. When the glycosylation of mannose with PGA was stopped silica gel (acetonitrile/water (10–30 vol%)) and freeze dried for
after 1–3 h at 80 °C in step 1, 21–36% of free mannose at least two days. Then, purified PMan was dissolved in PGA
remained unreacted. When vacuum (step 2) was applied at (5 equiv.) and stirred at 80 °C for 1 h (step 1) with Amberlyst-
that stage, the unreacted free mannose randomly polymerized 15 prior to heating the mixture at 100 °C under vacuum for 4 h
either with PMan or other molecule of mannose. (step 2). Under these conditions, PMan was converted to
Unfortunately, a mixture of (PMan)n (76%) and terminal free (PMan)n with a DPn of 4, in 85% yield (Fig. S11†). These find-
oligomannoside (12%) along with 12% of LVM was obtained. ings support that PMan could be depropargylated to form the
To ensure the elongation of the sugar moiety and full glycosy- oxocarbenium ion back, which can further react with another
lation with PGA at the terminal position, it is thus mandatory species of PMan, resulting in the elongation of the mannoside
to achieve a nearly complete conversion of mannose in step 1 chain. One should note that this oligomerization reaction is
prior to step 2, suggesting also that step 1 is reversible under possible, thanks to the solubility of mannose and APGs in
vacuum (see later for more information). The best conditions PGA. In contrast, with fatty alcohols, this elongation was not
in step 1 are 80 °C, a PGA/mannose molar ratio of 5, and 3 h of observed in step 2 due to the solubility issue, i.e., the precipi-
reaction, leading to the formation of PMan in 65% yield tation (and/or degradation) of monoalkylglycosides and sugars
(step 1) and (PMan)n with a DPn of 8 in step 2 (Fig. 6).
upon evaporation. The proposed reaction mechanism also
explains the formation of LVM as a minor product, resulting
from the internal cyclization of the oxocarbenium ion.
Plausible reaction mechanism
On the basis of all these results, we suggest a plausible reac-
Note that Amberlyst-15 can be replaced by strongly acidic
tion mechanism (Scheme 2), inspired by the most accepted H2SO4 or Aquivion PFSA. However, due to the high acid
proposals of glycosylation in the literature.27–29 In step 1, strength of these acids, the proton loading has to be reduced
mannose is protonated by amberlyst-15 to form the oxocarbe- from 4.2 mol% to 0.13 mol% to avoid fast and uncontrolled
nium ion, which is then rapidly trapped by PGA to form PMan. degradation of sugars, in particular in step 2.
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Scheme 2 Proposed reaction mechanism for step 1 and step 2.
Transposition to glucose
a proof-of-concept experiment (Scheme 3) the successful
addition of a long chain fatty acid (oleic acid) via click
chemistry to mannose-oligomers (for more details, see
Finally, the reaction was transposed to glucose, which is even
more attractive than mannose, as it is a product potentially
derived from lignocellulosic biomass waste. The same con-
ditions as those described above for steps 1 and 2 were
applied. 1H-NMR (Fig. S12†) spectroscopy and SEC analysis
(Fig. S13†) confirmed the formation of propargyl-(oligo)-gluco-
sides (PGlu)n with a Mn of 542 g mol−1 ðDPn ¼ 3Þ and a disper-
sity Đ of 2.1. 1H-NMR investigations clearly evidenced the pres-
ence of α- and β-anomers with the appearance of two signals
for the alkyne protons (2.95–2.91 ppm) and two signals corres-
ponding to the anomeric protons of the propargylated term-
inal position (α: 5.12 ppm, β: 4.67 ppm).
As stated in the Introduction section, the alkyne group at
the chain-end of the prepared oligosaccharides (mannose
and glucose) enables a direct access to amphiphiles. For this
purpose, the well-known ‘azide–alkyne Huisgen cyclo-
addition’ presents an attractive method due to its atom-
economy and quantitative conversions.31 As demonstrated in
previous studies,5,32 this method is frequently utilized to
prepare amphiphilic block copolymers and provides further
Scheme 3 Preparation of amphiphiles by the copper-catalyzed
applications of the as-prepared oligosaccharides. We show in addition of oleic acid to (PMan)n via click-chemistry.
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Fig. S14–S18†). The structure of the as-obtained amphiphile
was evidenced by H-NMR (Fig. S18†) with the appearance of
Notes and references
1
the characteristic triazole peak at 8.1 ppm and the analysis
by MALDI-TOF, which confirmed no depolymerization
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Conclusions
In this work, we demonstrated that the acid catalyzed (amber-
lyst-15) glycosylation reaction of unprotected monosaccharides
(mannose, glucose) and propargyl alcohol afforded fully func-
tionalized propargylated glycosides with a DPn up to 8 in 88%
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chain fatty acid to the sugars via the Huisgen catalyzed ‘Click-
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Conflicts of interest
There are no conflicts to declare.
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CNRS INCREASE for funding. CESAMO, Institut des Sciences Tech. Conf., Pittsburgh, PA, USA, 2009.
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also acknowledged for MALDI-TOF MS experiments. Anne-
Laure Wirotius is also strongly acknowledged for scientific dis-
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of Science and Technology, IntechOpen, Kenya,
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