Synthesis and Characterization of Four Diastereomers of Monorhamnolipids

Article abstract of DOI:10.1021/jacs.7b00427

Rhamnolipids are amphiphilic glycolipids biosynthesized by bacteria that, due to their low toxicity and biodegradability, are potential replacements for synthetic surfactants. The previously limited access to pure materials at the gram scale has hindered

Full text of DOI:10.1021/jacs.7b00427

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Article
Synthesis and Characterization of Four Diastereomers of Monorhamnolipids
Ricardo Palos Pacheco, Ryan J. Eismin, Clifford S. Coss, Hui
Wang, Raina M Maier, Robin Polt, and Jeanne E Pemberton
J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/jacs.7b00427 • Publication Date (Web): 16 Mar 2017
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Synthesis and Characterization of Four Diastereomers of Mono-
rhamnolipids
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Ricardo Palos Pacheco,a Ryan J. Eismin,a Clifford S. Coss,a† Hui Wang,a‡ Raina M. Maier,b Robin Polt,a
Jeanne E. Pembertona*
aDepartment of Chemistry and Biochemistry and bDepartment of Soil, Water and Environmental Science, University of Ari-
zona, Tucson, AZ 85721
KEYWORDS: Rhamnolipids, surfactants, diastereomer resolution.
ABSTRACT: Rhamnolipids are amphiphilic glycolipids biosynthesized by bacteria that, due to their low toxicity and biodegrada-
bility, are potential replacements for synthetic surfactants. The previously limited access to pure materials at the gram scale has
hindered extensive characterization rhamnolipid structure-performance behavior. Here, we present an efficient and versatile syn-
thetic methodology from which four diastereomers of the most common monorhamnolipid, α-rhamnopyranosyl-β-
hydroxydecanoyl-β-hydroxydecanoate, are prepared and subsequently characterized. Exploration of their behavior at the air-water
interface is reported and analyzed in terms of the absolute configuration of the lipid tail carbinols at pH 4.0 and 8.0. All diastere-
omers exhibit a minimum surface tension of about 28 mN/m without a significance difference between the protonated (nonionic) or
deprotonated (anionic) states. At pH 4.0 (nonionic), all diastereomers have a critical micelle concentration (CMC) in the µM range.
At pH 8.0 (anionic), CMC values for the (R,R), (S,S), and (S,R) diastereomers are approximately an order of magnitude higher than
in their nonionic states whereas the (R,S) diastereomer exhibits a CMC about five times larger.
INTRODUCTION
42 are some of the physiological functions attributed to rham-
nolipids. Applications such as complexation and flushing of
heavy metals from contaminated soils,12,43-48 hydrocarbon
removal from soils,49-51 enhancers of hydrocarbon biodegra-
dation,49,52-54 and use as biological control agents55-58 have
been suggested. Both, the biological roles and the developing
biotechnology mentioned above rely on the powerful pH-
modulated59 surfactant properties of these materials. In their
anionic form, the low critical micelle concentrations (CMCs)
and high surface activity of rhamnolipids are several orders
of magnitude lower than related anionic synthetic petroleum-
based derivatives and current commercially available coun-
terparts. In their nonionic form, their CMCs are comparable
to nonionic synthetic petroleum-based surfactants. These
characteristics make rhamnolipids competitive60,61 green
alternative biomaterials for industrial specialty surfactant
sectors. However, regardless of the high interest of the in-
dustrial and scientific communities, few studies62 have dealt
with pure materials whose structure-surfactant performance
can be evaluated. This occurs because until now the only
practical source of rhamnolipids has been harvesting from
microorganisms that produce condition-dependent multi-
component mixtures of congeners. In addition, access to the
interfacial and solution aggregation properties of diastere-
omers of these surfactants has been prohibited by virtue of
the exclusive bioproduction of L-lipid units only.4,63
Rhamnolipids are amphipathic glycolipids with outstanding
surfactant properties biosynthesized by microorganisms1-4
mainly belonging to the class of Gammaproteobacteria like
Pseudomonas aeruginosa, although others from the Actino-
bacteria, Bacilli, Betaprotobacteria and Deltaproteobacteria
classes have been identified as producers as well.5 More
than 50 biosynthesized congeners have been identified in
mixtures, but the general structure shown in Figure 1 is
comprised of a lipid unit of (R,R)-β-hydroxyalkanoyl-β-
hydroxyalkanoic acids6 of variable chain lengths (C6-C14)7,8,
trans-1,2-O-glycosylated9 by mono- or dimeric carbohydrate
L-rhamnopyranosyl units.10
Figure 1. Structural chemical diversity of (R,R)-rhamnolipids
produced by Pseudomonas aeruginosa. Traces of rhamnolipids
with unsaturation on both alkyl chains and monoacetylated
rhamnose have been harvested from cultures of P. aeruginosa.
In this study we present a flexible, scalable and practical
synthetic methodology for the synthesis and identification of
all four diastereomers (1,2,3,4) of α-rhamnopyranosyl-β-
hydroxydecanoyl-β-hydroxydecanoate (Rha-C10-C10), the
most abundant biosynthesized congener, shown in Figure 2.
Such a synthetic approach allows us to ascertain whether
Biofilm regulation,5,11-16 antimicrobial activity,17-21 uptake
and biodegradation of poorly soluble substrates,22-24 immune
modulators and virulence factors,25-37 and surface motility38-
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stereochemistry of the lipid tails plays a pivotal role in de- sides and trichloroacetimidates. In general, carbohydrate
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termining solution aggregation properties or other surfactant
behavior at the air-water interface.
peracetates are synthesized and then converted to more reac-
tive donor species for glycosylation. However, the peracetate
can also be an effective donor species, supported by oxoni-
um ion stabilization via Lewis acid activation and neighbor-
ing participation of the C-2 acetate carbonyl of the peracety-
lated carbohydrate. Large quantities of peracetylated donor
can be prepared quantitatively (e.g. 200 g) in one step at
room temperature (pyridine, acetic anhydride). This donor is
not water or air sensitive and is stable at room temperature,
resulting in a long shelf life either as a solid or dissolved in
organic solvent without special storage requirements.
Peracetylations performed at room temperature yield mix-
tures of anomers at the same ratio as the starting sugar.
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Glycosylation. Our glycosylation approach, shown in
Scheme 1, exploits the minimally competent Lewis acid
concept developed in our laboratories.78,79 Three soft metal
Lewis acid salts were tested as catalysts: InBr3, Bi(OTf)3 and
Sc(OTf)3. Perbenzoylated and peracetylated rhamnose were
tested as donors. Catalytic amounts of the less hygroscopic
bismuth (III) triflate promoted the glycosylation of primary
and racemic secondary alcohols in high yields.79 The Bi(III)
salts are less toxic83,84 and less hygroscopic than the In(III)
and Sc(III) salts,85,86 and proved superior in this case. Only
alpha anomers were observed due to the anomeric effect
coupled to the trans 'participating group' at C2 of the L-
rhamnose acetates. Further explanation for this stereospecific
coupling may include weak neighboring group participation
and/or solvent interactions.
Figure 2. Diastereomers produced by chemical synthesis.
The structure of a surfactant is known to govern its proper-
ties such as CMC, surface activity and aggregation behav-
ior.64 Techniques including small angle neutron scattering,65-
68 fluorescence quenching,69 and surface tensiometery65,66,69-75
have been used to study native rhamnolipids to characterize
these properties. In this work, in addition to a new synthetic
methodology for production of the four diastereomers of the
Rha-C10-C10 monorhamnolipids, we also report the surface
activity and aggregation morphology of these synthetic
forms, providing insight into possible structure-performance
relationships between these variants.
RESULTS AND DISCUSSION
Synthesis of the racemic lipid units and the resolution of the
racemic mixture. Several synthetic methodology precedents
for the preparation of β-hydroxyalkanoic acids have been
reported in the literature due to the ubiquity of this lipid
moiety in biomolecules found in Gram negative bacterial cell
walls.87 Access to optically pure β-hydroxyalkanoic acids is
afforded by stereoselective reduction of the β-ketoalkanoic
esters using Noyori’s catalytic hydrogenation, resolving
agents of racemic mixtures, enzymatic reduction,88,89 and
recently, the exploitation of Mitsunobu chemistry.90 In our
approach, the lipid units are prepared as racemic mixtures of
benzyl β-hydroxyalkanoic esters in two steps: formation of
the benzyl β-keto ester using Meldrum’s acid carbonyl
chemistry91-93 and reduction of the keto group with
NaBH3CN under acidic conditions94 as shown in Scheme 1.
Synthesis of monorhamnolipid diastereomers. To date,
two approaches to the chemical synthesis of rhamnolipids
have been reported.76,77 Both approaches are versatile and
can be used to produce any saturated mono- or dirhamno-
lipid with any desired stereochemistry. Unfortunately, these
approaches are not readily amenable to production of these
materials at scale, as they are labor intensive (>18 steps per
diastereomer), use expensive reagents, dangerous proce-
dures, and low yielding reactions. Here, we report a solution
phase methodology in which a long shelf life, air and mois-
ture stable peracetylated rhamnose donor can be activated in
the presence of a minimally competent Lewis acid78,79 to
produce a racemic mixture of the lipid acceptor that can be
resolved in a flash preparative silica column by means of
enantiomer-to-diastereomer conversion80-82 after stereo-
controlled glycosylation. The pure free acid diastereomers
are esterified with the second lipid unit to render a new mix-
ture of diastereomers that is separable as free acids by con-
ventional flash column chromatography. The rhamnose units
of the pure diastereomers are deacetylated orthogonally by
transesterification to yield the final products. The methodol-
ogy is sufficiently flexible to allow modification of the sugar
and lipid moieties to create other glycolipids as well. Other
significant attributes include scalability, mild reaction condi-
tions, high green indices, and cost-effectiveness compared to
the two synthetic methods previously reported. Absolute
configuration assignments are performed on the synthetic
and naturally produced materials by means of Mosher’s ester
analysis and optical rotation measurements. Details of the
synthetic methods are contained in the Supporting Infor-
mation along with supporting spectral data (Figures S1-18).
Scheme 1
The racemic mixture was resolved by normal phase liquid
chromatography (NP-LC) with silica (flash gel) by means of
enantiomer-to-diastereomer conversion after glycosylation
with a fully protected rhamnose donor. Attempts to resolve
the racemic mixture by flash column chromatography relied
on inducing differential intramolecular and intermolecular
interactions with stationary/mobile phase systems that shift
the partition coefficients of the members of the racemate.
This principle was tested with the glycosides 7a, 7b and 8a/b
as shown in Scheme 2.
Synthesis. Preparation of the donor. Rhamnolipid syntheses
found in the literature utilize thiorhamnosides, iodorhamno-
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Scheme 2
Although separation of the 7a diastereomeric mixture was
achieved by analytical thin layer chromatography (TLC) and
reversed phase high performance liquid chromatography
(RPLC), it proved impossible by preparative NP-LC. On the
other hand, up to 7 g of the free acid (8a/b) of the diastereo-
meric mixture 7a have been resolved efficiently in a 500 g
silica column with 1:1 EtOAc/hexanes + 1%v/v AcOH as the
mobile phase. It was fortunate that such separation was
achieved with the peracetylated rhamnoside, retaining the
orthogonality of our protecting group scheme. Apparently,
the van der Waals intermolecular interactions of the electron
rich benzyl and acetyl groups are not sufficiently different
among diastereomers to facilitate their separation. Alternate-
ly, it may be that hydrogen bonding of the carboxylic acid
with silanols is the interaction that governs separation of the
diastereomeric mixture.
been achieved. The diastereomeric enrichment allows avoid-
ance of tedious procedures like enantiomer recrystallizations,
and dangerous and sometimes not accessible enantioselective
conversions like Noyori's asymmetric hydrogenation.
Absolute configuration assignment of the carbinol groups
of the lipid tails. Assignment of the absolute configuration
of the carbinol groups at the lipid tails of each of the dia-
stereomers in terms of their relative Rf values was achieved
1
using a combination of H-NMR Mosher’s ester analysis97-99
and optical rotation measurements. In order to determine the
absolute configuration of the carbinols of the inner lipid tails,
the glycosidic bond of a sample of deacetylated 8a and 8b
was exo-cleaved100-104 by methanolysis.29 The resulting me-
thyl β-hydroxydecanoate esters were esterified with (R)- and
(S)-MTPA-Cl to render the (S)- and (R)-Mosher’s esters,
Figure 4a. The four Mosher’s esters obtained were charac-
1
Addition of the second lipid tail. The diastereomeric free
acids 8a and 8b were coupled by Steglich95 esterification
with a second benzyl β-hydroxydecanoate ester (6). After
reductive debenzylation, the diastereomeric mixture of
peracetylated rhamnolipid free acids is separable by NP-LC
with similar conditions, column loading and resolution.
The four diastereomers are deacetylated via Zemplèn96 con-
ditions (sodium methoxide in methanol) to produce the four
Rha-C10-C10 monorhamnolipid congeners (1, 2, 3 & 4).
terized by H-NMR and the chemical shifts were compared
according to ΔδSR = (δS - δR). As a control, the absolute con-
figuration of the carbinols of the methyl ester lipid products
of the acidic methanolysis of the monorhamnolipids pro-
duced by Pseudomonas aeruginosa ATCC 9027 was also
determined by Mosher’s ester analysis. Absolute configura-
tion assignment of the secondary lipid tails was performed in
similar fashion. Samples of the four diastereomeric, double
lipid tail rhamnolipids (Rha-C10-C10) were hydrolyzed with
2 M NaOH to obtain the 3- hydroxydecanoic acid, and ester-
ified with BF3-MeOH,105-107 Figure 4b. After isolation by
flash column chromatography, optical rotation measurements
The entire synthesis is represented in Figure 3. In summary,
a synthetic methodology to render the four diastereomers in
five chemically distinct steps using solution base chemistry
of a battery of well-described chemical transformations has
Figure 3. Synthetic methodology for manufacturing the diastereomers (R,R), (R,S), (S,S) and (S,R) of Rha-C10-C10. The stages where the diastere-
omers are enriched by preparative normal phase liquid chromatography (NP-LC) are marked by dashed circles. The steps involved are i) glycosyla-
tion of 6 in presence of Bi(OTf)3 and dry MeCN refluxed for 2.5 h; ii) debenzoylation with Pd/C, 1 atm H2 and dry THF at RT overnight; iii) Steglich
esterification of 6 in presence of EDC and DMAP in dry CH2Cl2 at RT; and iv) deacetylation under Zemplèn conditions with MeONa/MeOH.
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Figure 4. Approaches followed to determine the absolute configuration of a) the inner lipid tail carbinols by Mosher’s ester analysis and b) the
outer lipid tail carbinols by optical rotation measurements. For the inner lipid tail, the Mosher’s esters of the naturally produced rhamnolipid were
used as controls. Methyl (R)-3-hydroxydecanaote was obtained by Noyori’s enantioselective hydrogenation and used as control for the optical
o
rotation measurements. Derivatization reactions: i) MeONa, MeOH, RT; ii) MeOH, HCl 0.625 M, 70 C, 12 h; iii) 2M NaOH(aq), RT; iv) 1.3 M BF3-
MeOH, 1.1 eq DMP 50 oC, 15 min; v) (R)-BINAP, EtOH, H2 45 PSI, 115 oC, 72 h.
The relatively low CMC values highlight strong interactions
Table 1. Absolute configuration assignments for the carbinols of
between the hydrophobic tails, consistent with the behavior of
similar glycolipid surfactants.115-120 At pH 8.0, the (R,R)-Rha-
C10-C10 diastereomer has the highest CMC and (R,S)-Rha-
C10-C10 has the lowest, whereas CMC values for (S,S)-Rha-
C10-C10 and (S,R)-Rha-C10-C10 are comparable. These val-
ues suggest that the (R,S)-Rha-C10-C10 diastereomer has a
molecular structure best suited to form aggregates in solution
(see space filling structures in Figure S19 of the Supporting
Information.) Specifically, the lipid tails are oriented in a way
that makes it more energetically favorable for the molecules to
aggregate than remain as free monomers when compared to
the other diastereomers.
lipid tails.
Absolute configuration
Compound
Rf
of carbinol(s) at lipid tail(s)a
8a
8b
10a
10b
10c
10d
0.38
0.26
0.36
0.44
0.22
0.27
(R)
(S)
(R,R)
(R,S)
(S,S)
(S,R)
aAssignments determined from Mosher's ester analysis and opti-
cal rotation measurements. See Supporting Information for addi-
tional details. Rf values obtained with analytical TLC with 1:1
hexanes/ethyl ether +1% v/v AcOH as mobile phase.
At pH 4.0, the (R,S)-Rha-C10-C10 diastereomer has the high-
est CMC and the others have comparable values. At pH 4.0,
all diastereomers are nonionic, eliminating Coulombic repul-
sion between the headgroups and lowering the CMC. Howev-
er, (R,S)-Rha-C10-C10 must assume a structure for which
aggregation is less energetically favorable compared to the
other diastereomers in their similar nonionic forms.
of the four chloroform solutions were performed and com-
pared with the literature values of D-3-hydroxy esters and
acids of similar or longer chain length.6,10,108-111 As a control
for the correlation of absolute configuration with the levorota-
ry and dextrorotary attributes of the individual methyl (±)-3-
hydroxydecanoate esters, we measured the optical rotation
values of methyl (R)-3-hydroxydecanoate (12) prepared in
nominally high %ee (>99%) by a well described Noyori’s
catalytic hydrogenation method,88,112-114 and methyl 3-(S)-
hydroxydecanoate ester (8b’) with confirmed absolute config-
uration by Mosher’s ester analysis. All data pertaining to this
analysis can be found in the Supporting Information. Table 1
summarizes the assignments in terms of TLC Rf values of the
acetylated glycolipids.
Chen et al. have reported CMC values for native monorham-
nolipid mixtures (comprised primarily of the (R,R)-Rha-C10-
C10 congener) of 180 and 360 µM at pH 7 and 9, respective-
ly.66 These values fall above and below that for the chemically
synthesized (R,R)-Rha-C10-C10 diastereomer at pH 8.0 re-
ported in Table 2, consistent with the known trend that CMC
decreases with decreasing pH.73 These values suggest that a
mixture of monorhamnolipid congeners not of the form (R,R)-
Rha-C10-C10 has only a minimal effect on the aggregation
properties when compared to the chemically pure (R,R)-Rha-
C10-C10 diastereomer produced by chemical synthesis.
Surface tensiometry at pH 4.0 and 8.0. Surface tension
measurements were made on each of the Rha-C10-C10 dia-
stereomers using the du Noüy ring method. Due to the carbox-
ylic acid moiety (pKa 5.5 for rhamnolipid aggregates59), solu-
tion conditions were adjusted to pH 4.0 or 8.0 to create solu-
tions of the protonated nonionic or deprotonated anionic mon-
orhamnolipids, respectively. Plots of surface tension (γ) as a
function of Rha-C10-C10 concentration at pH 4.0 and 8.0 are
shown in Figures 5a and b, respectively. CMCs at each pH
are determined as the intersection between the two best
straight lines through the data and are displayed in Table 2 for
each diastereomer.
The CMC values reported in Table 2 are also similar to the
values measured for the native monorhamnolipid mixture pro-
duced by Pseudomonas aeruginosa ATCC 9027, an exclusive
producer of monorhamnolipids.69 At pH 8, this native mixture
exhibits a CMC of 200 µM while at pH 4, this mixture exhib-
its a CMC of 21 µM. Given that the composition of the mix-
ture ranges from ~70-85% of the (R,R)-Rha-C10-C10, the
similarity in values between the chemically synthesized dia-
stereomers and the native mixture is not surprising.
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Table 2. Surface tension results for the monorhamno-
lipid diastereomers at pH 4.0 and 8.0.
1
2
3
4
5
6
7
8
(S,S) Rha-C10-C10
(S,R) Rha-C10-C10
(R,S) Rha-C10-C10
(R,R) Rha-C10-C10
Rha-C10-
C10
CMC
Area
γCMC
(mN/m)
pHa
(µM)b
(Å2/molec)
16 ± 4
270 ± 77
25 ± 1
27.5 ± 0.1
28.1 ± 0.2
28.8 ± 0.1
27.4 ± 0.2
27.5 ± 0.1
29.5 ± 0.2
28.2 ± 0.1
28.5 ± 0.2
21± 4
117 ± 12
23 ± 1
80± 1
4.0
8.0
4.0
8.0
4.0
8.0
4.0
8.0
(R,R)
(R,S)
(S,S)
(S,R)
79 ± 3
9
18 ± 3
201 ± 51
15 ± 1
21 ± 2
93 ± 7
21 ± 1
103 ± 4
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180± 24
a
-16
apH 4.0 values calculated from data in Figure 5a; pH 8.0 values calcu-
lated from data in Figure 5b. bStandard deviations determined from
measurements on three independently prepared samples.
-14
-12
-10
-8
ln (Diastereomer Concentration in M)
80
70
60
50
40
30
20
monorhamnolipid mixtures, values of 66 and 77 Å2/molecule
at pH 7 (0.063 M K2PO4 and 0.037 M NaOH) and 9 (0.023
borax and 0.008 M HCl), respectively,66 and 86 Å2/molecule
at pH 869 were reported. These values are slightly lower than
but similar to that for the pure (R,R)-Rha-C10-C10 diastere-
omer at pH 8.0 reported in Table 2; these slight differences
may result from the different buffers used for these studies. At
pH 8.0, the (R,S)-Rha-C10-C10 diastereomer has the lowest
cross-sectional area, with the values increasing by about 10
Å2/molecule in the order (S,S), (S,R), and (R,R). These values
suggest differences in lipid tail orientation of the surfactants
adsorbed at the air-water interface leading to differences in
packing. Based on these values, the (R,S)-Rha-C10-C10 is
more tightly packed compared to (R,R)-Rha-C10-C10.
(S,S) Rha-C10-C10
(S,R) Rha-C10-C10
(R,S) Rha-C10-C10
(R,R) Rha-C10-C10
b
-16
-14
-12
-10
-8
-6
The second noteworthy aspect of the molecular areas in Table
2 are the unusually low values of ~21 Å2/molecule observed at
pH 4. This is similar to the value of ~23 Å2/molecule observed
for the native monorhamnolipid mixture from P. aeruginosa
ATCC 9027 at pH 4. Given that the cross-sectional area of a
single alkyl chain is ~21 Å2/molecule, we hypothesize that the
very tight packing of these double chain monorhamnolipids at
pH 4 induces formation of small lamellar packets of bilayer
monorhamnolipid, contiguous with the monorhamnolipid
monolayer at the water surface, that protrude into the aqueous
subphase. These features result in an underestimated surface
area for the monorhamnolipid layer at the water surface,
thereby resulting in artificially low values of molecular cross-
sectional area.
ln (Diastereomer Concentration in M)
Figure 5. Surface tension as a function of Rha-C10-C10 concentra-
tion. a) Solutions adjusted to pH 4.0 with a minimal amount of HCl,
and b) pH 8.0 with a minimal amount of NaOH.
The minimum surface tension values achieved with these syn-
thetic diastereomers is also on the same order as previously
reported values for native monorhamnolipid mixtures, indicat-
ing similar air-water interface adsorption of the diastereomers
and the native congener mixture of monorhamnolipids.66,69
Minimum surface tension values are in the range of 27-29
mN/m at both pH values. This would indicate that protonation
of the carboxylic acid moiety and lipid tail stereochemistry
have little effect on the adsorption of these surfactants at the
air-water interface.
Pyrene fluorescence for probing aggregation state. In the
deprotonated anionic state, the native monorhamnolipids form
globular micellar structures, growing into larger lamellar
structures at higher concentrations.66,69 To gain further insight
into the solution aggregation behavior of these chemically
synthesized diastereomers, the fluorescence behavior of a po-
larity sensitive probe, pyrene, was used. Pyrene, which is
known to have high affinity for non-polar environments,121 is
useful for monitoring solution aggregation122 through changes
in the intensity ratio of the III/I vibronic bands of the fluores-
cence spectrum. As the local polarity of the pyrene becomes
increasingly nonpolar, this ratio systematically increases. This
ratio was used here to confirm solution aggregate formation by
the four Rha-C10-C10 diastereomers by introducing pyrene to
pH 8.0 solutions of the diastereomers at varying concentra-
tions. As shown by the results in Figure 6, this ratio increases
with Rha-C10-C10 concentration for all four diastereomers,
The molecular cross-sectional areas at the air-water interface
are obtained from the surface tension data using the Gibbs
adsorption isotherm:
1
ꢀꢁ
Γ = −
mꢀꢁ ꢀꢁꢂꢃ
where Γ is the surface excess, R is the gas constant, ꢀ is tem-
perature, ꢀ is the surface tension, ꢀ is the surfactant concen-
tration and m is the Gibbs prefactor (m = 1 for nonionic mono-
rhamnolipids at pH 4, m = 2 for anionic monorhamnolipids at
pH 8). Molecular areas are calculated from the experimentally
determined surface excess values using the slopes of the sur-
face tension plots and are reported in Table 2.
Several aspects of these data are noteworthy. First, in previous
literature studies of molecular cross-sectional areas for native
5
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pH 8 due to fewer repulsive interactions between the nonionic
1
2
3
4
5
6
7
8
compared to the anionic forms of these monorhamnolipds. The
CMC values for the diastereomers at pH 4 are generally simi-
lar with the exception of (R,S)-Rha-C10-C10 that likely has
poorer packing compared to the others. Thus, these stereo-
chemical differences in diastereomer structure confer slight
differences in surfactant performance among these molecules.
1.1
1.0
0.9
0.8
0.7
0.6
0.5
In total, these studies support chemical synthesis as a viable
alternative path to biosynthesis for the production of mono-
rhamnolipids. Given the reasonable yields of the synthetic
procedure, the ability to tailor surfactant performance through
choice of alkyl chain length, and the considerably greater ease
at compound purification, chemical synthesis may be the more
attractive pathway to fabrication of these materials for many
applications.
(S,S) Rha-C10-C10
(S,R) Rha-C10-C10
(R,S) Rha-C10-C10
(R,R) Rha-C10-C10
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
-5
0
5
10
15
20
25
30
35
40
Diastereomer Concentration (mM)
ASSOCIATED CONTENT
Figure 6. Pyrene III/I peak intensity ratios for all diastereomers as
a function of surfactant concentration using 250 nM pyrene at pH
8.0 in 10 mM phosphate buffer.
Supporting Information
Materials and methods for the synthesis of monorhamnolipid
diastereomers; instrumentation details and sample preparation
for the surface tension measurements and the pyrene fluores-
cence experiments for aggregation; detailed synthetic proce-
dures for the manufacture and isolation of rhamnolipid dia-
stereomers; protocols for absolute configuration assignments
of the carbinols on the lipid tails; relevant NMR and MS spec-
troscopic data. (PDF) This information is available free of
charge on the ACS Publications website at DOI: XX.XXXX/jacs.
confirming aggregate formation. This ratio does not level off
until concentrations >5 mM are reached for all diastereomers,
suggesting that full incorporation of pyrene does not occur
until concentrations well above the CMC are reached. This
behavior is consistent with size and structural changes in the
early stags of solution aggregation, with the formation of pre-
micellar aggregates at concentrations just above the apparent
CMC likely.69 Structural changes and growth of monorhamno-
lipid aggregates as a function of concentration are consistent
with previous reports from this69 and other66,73 laboratories that
aggregates on native monorhamnolipid mixtures. To further
elucidate the detailed aggregation behavior of these different
diastereomers, more in-depth studies are needed. These studies
are underway in these laboratories and will be reported at a
later date.
AUTHOR INFORMATION
Corresponding Author
*pembertn@email.arizona.edu
Present Addresses
†Department of Chemistry and Biochemistry, Northern Arizo-
na University, Flagstaff, AZ 86011
CONCLUSIONS
‡ CF Industries, Yazoo City, MS 39194
An efficient synthetic methodology for the production of four
diastereomers of the monorhamnolipid Rha-C10-C10 is de-
scribed. This process has considerably fewer steps than em-
ployed in the current methodologies available. Our methods
are solution based, and make use of conventional and scalable
well-described chemical transformations. The use of peracety-
lated donors and minimally competent Lewis acids improves
the practicality and scalability of the method. Assignments of
the absolute configuration of the carbinols at the lipid tails
were performed by means of Mosher’s ester 1H-NMR analysis
and optical rotation measurements. In addition to production
of these rhamnolipid diastereomers, the method has proven to
be flexible enough to test structure-performance relationships
in other areas of the molecule including lipid tail symmetry
and lipid tail length. The results of these studies will be re-
ported at a later date.
Notes
Two authors of this work (JEP, RMM) have equity ownership in
GlycoSurf, LLC that is developing products related to the re-
search being reported. The terms of this arrangement have been
reviewed and approved by the University of Arizona in accord-
ance with its policy on objectivity in research.
ACKNOWLEDGMENT
The authors gratefully acknowledge support of this research
through grant awards from the National Science Foundation
(CHE-0714245 and CHE-1339597). The second grant is jointly
funded by the Environmental Protection Agency as part of the
Networks for Sustainable Molecular Design and Synthesis Pro-
gram. Ricardo Palos Pacheco gratefully acknowledges fellowship
support from Consejo Nacional de Ciencia
(CONACYT).
y Tecnología
Each Rha-C10-C10 diastereomer shows similar surface activi-
ty, CMC, and γCMC at pH 4 and 8, suggesting little difference
in air-water interface chemistry among the four diastereomers.
The values of molecular area for the diastereomers are similar
although not identical, suggesting slightly different molecular
orientations at the air-water interface. The molecular areas for
all four diastereomers are significantly lower at pH 4 than at
REFERENCES
(1) Clarke, K. G.; Ballot, F.; Reid, S. J. World J. Microbiol. Biotechnol.
2010, 26, 2179-2184.
6
ACS Paragon Plus Environment

Page 7 of 8
Journal of the American Chemical Society
(2) Hörmann, B.; Müller, M. M.; Syldatk, C.; Hausmann, R. Eur. J. Lipid
Sci. Technol. 2010, 112, 674-680.
(3) Soberón-Chávez, G.; Lépine, F.; Déziel, E. Appl. Microbiol.
Biotechnol. 2005, 68, 718-725.
(4) Hori, K.; Ichinohe, R.; Unno, H.; Marsudi, S. Biochem. Eng. J. 2011,
53, 196-202.
(5) Abdel-Mawgoud, A. M.; Lépine, F.; Déziel, E. Appl. Microbiol.
Biotechnol. 2010, 86, 1323-1336.
(6) Nakagawa, Y.; Matsuyama, T. FEMS Microbiol. Lett. 1993, 108, 99-
102.
(7) Benincasa, M.; Abalos, A.; Oliveira, I.; Manresa, A. Antonie van
Leeuwenhoek 2004, 85, 1-8.
(8) Haba, E.; Pinazo, A.; Pons, R.; Pérez, L.; Manresa, A. Biochim.
Biophys. Acta, Biomembr. 2014, 1838, 776-783.
(9) Edwards, J. R.; Hayashi, J. A. Arch. Biochem. Biophys. 1965, 111,
415-421.
(41) Morris, J. D.; Hewitt, J. L.; Wolfe, L. G.; Kamatkar, N. G.;
Chapman, S. M.; Diener, J. M.; Courtney, A. J.; Leevy, W. M.; Shrout, J.
D. Appl. Environ. Microbiol. 2011, 77, 8310-8317.
(42) Wang, S.; Yu, S.; Zhang, Z.; Wei, Q.; Yan, L.; Ai, G.; Liu, H.; Ma,
L. Z. Appl. Environ. Microbiol. 2014, 80, 6724-6732.
(43) Herman, D. C.; Artiola, J. F.; Miller, R. M. Environ. Sci. Technol.
1995, 29, 2280-2285.
(44) Juwarkar, A. A.; Nair, A.; Dubey, K. V.; Singh, S. K.; Devotta, S.
Chemosphere 2007, 68, 1996-2002.
(45) Sandrin, T. R.; Chech, A. M.; Maier, R. M. Appl. Environ.
Microbiol. 2000, 66, 4585-4588.
(46) Slizovskiy, I. B.; Kelsey, J. W.; Hatzinger, P. B. Environ. Toxicol.
Chem. 2011, 30, 112-123.
(47) Tan, H.; Champion, J. T.; Artiola, J. F.; Brusseau, M. L.; Miller, R.
M. Environ. Sci. Technol. 1994, 28, 2402-2406.
(48) Torrens, J. L.; Herman, D. C.; Maier, R. M. Environ. Sci. Technol.
1998, 32, 776-781.
(49) Herman, D. C.; Lenhard, R. J.; Miller, R. M. Environ. Sci. Technol.
1997, 31, 1290-1294.
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
(10) Jarvis, F. G.; Johnson, M. J. J. Am. Chem. Soc. 1949, 71, 4124-4126.
(11) Solano, C.; Echeverz, M.; Lasa, I. Curr. Opin. Microbiol. 2014, 18,
96-104.
(12) Singh, P.; Cameotra, S. S. Biochem. Biophys. Res. Commun. 2004,
319, 291-297.
(13) Nickzad, A.; Déziel, E. Lett. Appl. Microbiol. 2014, 58, 447-453.
(14) Banat, I. M.; Diaz de Rienzo, M. A.; Quinn, G. A. Appl. Microbiol.
Biotechnol. 2014, 98, 9915-9929.
(15) Irie, Y.; O'Toole, G. A.; Yuk, M. H. FEMS Microbiol. Lett. 2005,
250, 237-243.
(16) Boles, B. R.; Thoendel, M.; Singh, P. K. Mol. Microbiol. 2005, 57,
1210-1223.
(17) Itoh, S.; Honda, H.; Tomita, F.; Suzuki, T. J. Antibiot. 1971, 24, 855-
859.
(18) Lang, S.; Katsivela, E.; Wagner, F. Fat Sci. Technol. 1989, 91, 363-
366.
(19) Arino, S.; Marchal, R.; Vandecasteele, J. P. J. Appl. Microbiol. 1998,
84, 769-776.
(20) Sotirova, A. V.; Spasova, D. I.; Galabova, D. N.; Karpenko, E.;
Shulga, A. Curr. Microbiol. 2008, 56, 639-644.
(21) Abalos, A.; Pinazo, A.; Infante, M. R.; Casals, M.; Garcia, F.;
Manresa, M. A. Langmuir 2001, 17, 1367-1371.
(22) Beal, R.; Betts, W. B. J. Appl. Microbiol. 2000, 89, 158-168.
(23) Itoh, S.; Suzuki, T. Agric. Biol. Chem. 1972, 36, 2233-2235.
(24) Koch, A. K.; Kaeppeli, O.; Fiechter, A.; Reiser, J. J. Bacteriol. 1991,
173, 4212-4219.
(25) Andrae, J.; Rademann, J.; Howe, J.; Koch, M. H. J.; Heine, H.;
Zaehringer, U.; Brandenburg, K. Biol. Chem. 2006, 387, 301-310.
(26) Bergmann, U.; Scheffer, J.; Köeller, M.; Schönfeld, W.; Erbs, G.;
Muller, F. E.; Konig, W. Infect. Immun. 1989, 57, 2187-2195.
(27) Fujita, K.; Akino, T.; Yoshioka, H. Infect. Immun. 1988, 56, 1385-
1387.
(28) Haussler, S.; Rohde, M.; von Neuhoff, N.; Nimtz, M.; Steinmetz, I.
Infect. Immun. 2003, 71, 2970-2975.
(29) Häußler, S.; Nimtz, M.; Domke, T.; Wray, V.; Steinmetz, I. Infect.
Immun. 1998, 66, 5.
(30) Jensen, P. O.; Bjarnsholt, T.; Phipps, R.; Rasmussen, T. B.; Calum,
H.; Christoffersen, L. M. C.; Moser, C.; Williams, P.; Pressler, T.;
Givskov, M.; Høiby, N. Microbiology 2007, 153, 1329-1338.
(31) Johnson, M. K.; Boese-Marrazzo, D. Infect. Immun. 1980, 29, 1028-
1033.
(32) König, B.; Jaeger, K. E.; König, W. Int. Arch. Allergy Immunol.
1994, 104, 33-41.
(33) Kurioka, S.; Liu, P. V. J. Bacteriol. 1967, 93, 670-674.
(34) Read, R. C.; Roberts, P.; Munro, N.; Rutman, A.; Hastie, A.;
Shryock, T.; Hall, R.; McDonald-Gibson, W.; Lund, V.; Taylor, G. J.
Appl. Physiol. 1992, 72, 2271-2277.
(35) Sanchez, M.; Teruel, J. A.; Espuny, M. J.; Marques, A.; Aranda, F.
J.; Manresa, A.; Ortiz, A. Chem. Phys. Lipids 2006, 142, 118-127.
(36) Shryock, T. R.; Silver, S. A.; Banschbach, M. W.; Kramer, J. C.
Curr. Microbiol. 1984, 10, 323-328.
(37) Stipcevic, T.; Piljac, T.; Isseroff, R. R. J. Dermatol. Sci. 2005, 40,
141-143.
(38) Tremblay, J.; Richardson, A. P.; Lépine, F.; Déziel, E. Environ.
Microbiol. 2007, 9, 2622-2630.
(39) Caiazza, N. C.; Shanks, R. M. Q.; O'Toole, G. A. J. Bacteriol. 2005,
187, 7351-7361.
(40) Murray, T. S.; Kazmierczak, B. I. J. Bacteriol. 2008, 190, 2700-
2708.
(50) Jain, D. K.; Lee, H.; Trevors, J. T. J. Ind. Microbiol. 1992, 10, 87-
93.
(51) Providenti, M. A.; Flemming, C. A.; Lee, H.; Trevors, J. T. FEMS
Microbiol. Ecol. 1995, 17, 15-26.
(52) Al-Tahhan, R. A.; Sandrin, T. R.; Bodour, A. A.; Maier, R. M. Appl.
Environ. Microbiol. 2000, 66, 3262-3268.
(53) Zhang, Y. M.; Miller, R. M. Appl. Environ. Microbiol. 1992, 58,
3276-3282.
(54) Zhang, Y. M.; Miller, R. M. Appl. Environ. Microbiol. 1994, 60,
2101-2106.
(55) Abbasi, H.; Aranda, F. J.; Akbari Noghabi, K.; Ortiz, A. Biochim.
Biophys. Acta, Biomembr. 2013, 1828, 2083-2090.
(56) Bengtsson, T.; Holefors, A.; Liljeroth, E.; Hultberg, M.; Andreasson,
E. Potato Res. 2015, 58, 83-90.
(57) Miao, S.; Dashtbozorg, S. S.; Callow, N. V.; Ju, L.-K. J. Agric. Food
Chem. 2015, 63, 3367-3376.
(58) Stanghellini, M. E.; Miller, R. M. Plant Dis. 1997, 81, 4-12.
(59) Lebrón-Paler, A.; Pemberton, J. E.; Becker, B. A.; Otto, W. H.;
Larive, C. K.; Maier, R. M. Anal. Chem. 2006, 78, 7649-7658.
(60) Marchant, R.; Banat, I. M. Biotechnol. Lett. 2012, 34, 1597-1605.
(61) Mueller, M. M.; Kuegler, J. H.; Henkel, M.; Gerlitzki, M.;
Hoermann, B.; Poehnlein, M.; Syldatk, C.; Hausmann, R. J. Biotechnol.
2012, 162, 366-380.
(62) Wang, H.; Coss, C. S.; Mudalige, A.; Polt, R. L.; Pemberton, J. E.
Langmuir 2013, 29, 4441-4450.
(63) Mueller, M. M.; Hausmann, R. Appl. Microbiol. Biotechnol. 2011,
91, 251-264.
(64) Rosen, J. Surfactants and Interfacial Phenomena; 4th ed.; John
Wiley & Sons, Inc.: Hoboken, NJ, 2012.
(65) Chen, M. L.; Penfold, J.; Thomas, R. K.; Smyth, T. J. P.; Perfumo,
A.; Marchant, R.; Banat, I. M.; Stevenson, P.; Parry, A.; Tucker, I.; Grillo,
I. Langmuir 2010, 26, 17958-17968.
(66) Chen, M. L.; Penfold, J.; Thomas, R. K.; Smyth, T. J. P.; Perfumo,
A.; Marchant, R.; Banat, I. M.; Stevenson, P.; Parry, A.; Tucker, I.; Grillo,
I. Langmuir 2010, 26, 18281-18292.
(67) Dahrazma, B.; Mulligan, C. N.; Nieh, M. P. J. Colloid Interf. Sci.
2008, 319, 590-593.
(68) Guo, Y.; Mulligan, C. N.; Nieh, M. Colloids Surf., A 2011, 373, 42-
50.
(69) Eismin, R. J.; Munusamy, E.; Kegel, L. M.; Maier, R. M.; Schwartz,
S.; Pemberton, J. E. Langmuir 2016, submitted.
(70) Penfold, J.; Thomas, R. K.; Shen, H. Soft Matter 2012, 8, 578-591.
(71) Abbasi, H.; Noghabi, K. A.; Hamedi, M. M.; Zahiri, H. S.; Moosavi-
Movahedi, A. A.; Amanlou, M.; Teruel, J. A.; Ortiz, A. Colloids Surf., B
2013, 101, 256-265.
(72) Ozdemir, G.; Peker, S.; Helvaci, S. S. Colloids Surf., A 2004, 234,
135-143.
(73) Guo, Y. P.; Hu, Y. Y.; Gu, R. R.; Lin, H. J. Colloid Interface Sci.
2009, 331, 356-363.
(74) Manko, D.; Zdziennicka, A.; Janczuk, B. Colloids Surf., B 2014,
119, 22-29.
(75) Pornsunthorntawee, O.; Chavadej, S.; Rujiravanit, R. Colloids Surf.,
B 2009, 72, 6-15.
(76) Bauer, J.; Brandenburg, K.; Zähringer, U.; Rademann, J. Chem. -
Eur. J. 2006, 12, 7116-7124.
7
ACS Paragon Plus Environment

Journal of the American Chemical Society
Page 8 of 8
(77) Westerduin, P.; de Haan, P. E.; Dees, M. J.; van Boom, J. H.
(100) Manabe, S.; Ishii, K.; Satoh, H.; Ito, Y. Tetrahedron 2011, 67,
Carbohydr. Res. 1988, 180, 195-205.
9966-9974.
1
2
3
4
5
6
7
(78) Lefever, M. R.; Szabo, L. Z.; Anglin, B.; Ferracane, M.; Hogan, J.;
Cooney, L.; Polt, R. Carbohydr. Res. 2012, 351, 121-125.
(79) Coss, C. S.; Carrocci, T.; Maier, R. M.; Pemberton, J. E.; Polt, R.
Helv. Chim. Acta 2012, 95, 2652-2659.
(80) Toyo'oka, T. J. Biochem. Biophys. Methods 2002, 54, 25-56.
(81) Nimura, N.; Ogura, H.; Kinoshita, T. J. Chromatogr. A 1980, 202,
375-379.
(101) Deslongchamps, P. Tetrahedron 1975, 31, 2463-2490.
(102) St-Jean, O.; Prévost, M.; Guindon, Y. J. Org. Chem. 2013, 78,
2935-2946.
(103) Deslongchamps, P.; Li, S.; Dory, Y. L. Org. Lett. 2004, 6, 505-508.
(104) Phan, H. D.; Yokoyama, T.; Matsumoto, Y. J. Carbohydr. Chem.
2013, 32, 223-239.
(105) Jenske, R.; Vetter, W. J. Agric. Food Chem. 2008, 56, 11578-
11583.
(82)
Houben-Weyl Methods of Organic Chemistry, Supplement:
8
9
Stereoselective Synthesis; 4th ed.; Helmchen, G.; Hoffmann, R. W.;
Mulzer, J.; Schaumann, E., Eds.; Georg Thieme Verlag: Stuttgart, 1995;
Vol. E 21a.
(83) Leonard, N. M.; Wieland, L. C.; Mohan, R. S. Tetrahedron 2002, 58,
8373-8397.
(106) Jenske, R.; Vetter, W. J. Chromatogr. A 2007, 1146, 225-231.
(107) Kleiman, R.; Spencer, G. F.; Earle, F. R. Lipids 1969, 4, 118-122.
(108) Yamaguchi, M.; Sato, A. Chem. Ind. (Chichester, U. K.) 1976, 741-
742.
(109) Spencer, J. F. T. Can. J. Chem. 1964, 42, 831-835.
(110) Cartwright, N. J. Biochem. J. 1957, 67, 663-669.
(111) Jakob, B.; Voss, G.; Gerlach, H. Tetrahedron: Asymmetry 1996, 7,
3255-3262.
(112) Noyori, R.; Ohkuma, T.; Kitamura, M.; Takaya, H.; Sayo, N.;
Kumobayashi, H.; Akutagawa, S. J. Am. Chem. Soc. 1987, 109, 5856-
5858.
(113) Utaka, M.; Watabu, H.; Higashi, H.; Sakai, T.; Tsuboi, S.; Torii, S.
J. Org. Chem. 1990, 55, 3917-3921.
(114) Noyori, R.; Ikeda, T.; Ohkuma, T.; Widhalm, M.; Kitamura, M.;
Takaya, H.; Akutagawa, S.; Sayo, N.; Saito, T.; Taketomi, T.;
Kumobayashi, H. J. Am. Chem. Soc. 1989, 111, 9134-9135.
(115) Sugar-Based Surfactants: Fundamentals and Applications; Ruiz, C.
C., Ed.; CRC Press, Taylor & Francis Group: Boca Raton, FL, 2008.
(116) Oliver, R. C.; Lipfert, J.; Fox, D. A.; Lo, R. H.; Doniach, S.;
Columbus, L. PLos One 2013, 8, e62488:62481-62410.
(117) Nakagawa, A.; Kamitakahara, H.; Takano, T. Carbohydr. Res.
2011, 346, 1671-1683.
(118) Nonionic Surfactants: Alkyl Polyglucosides; 10th ed.; Balzer, D.;
Luders, H., Eds.; Marcel Dekker: New York, 2000.
(119) Chaveriat, L.; Gosselin, I.; Machut, C.; Martin, P. Eur. J. Med.
Chem. 2013, 62, 177-186.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
(84) Ikeda, K.; Torisawa, Y.; Nishi, T.; Minamikawa, J.; Tanaka, K.;
Sato, M. Bioorg. Med. Chem. 2003, 11, 3073-3076.
(85) Tanaka, A.; Hirata, M.; Kiyohara, Y.; Nakano, M.; Omae, K.;
Shiratani, M.; Koga, K. Thin Solid Films 2010, 518, 2934-2936.
(86) Maghraoui, S.; Ayadi, A. J. Electron Microsc. 2011, 60, 183-190.
(87) Rietschel, E. T.; Kirikae, T.; Schade, F. U.; Ulmer, A. J.; Holst, O.;
Brade, H.; Schmidt, G.; Mamat, U.; Grimmecke, H. D.; Kusumoto, S.
Immunobiology 1993, 187, 169-190.
(88) Noyori, R. Acta Chem. Scand. 1996, 50, 380-390.
(89) Spengler, J.; Fernando, A. Curr. Org. Synth. 2008, 5, 151-161.
(90) Menhoura, B.; Mayona, P.; Pléb, K.; Bouquillona, S.; Doreyc, S.;
Clémentc, C.; Deleud, M.; Haudrechya, A. Tetrahedron Lett. 2015, 56,
1159-1161.
(91) Oikawa, Y.; Sugano, K.; Yonemitsu, O. J. Org. Chem. 1978, 43,
2087-2088.
(92) Xu, F.; Armstrong, J. D.; Zhou, G. X.; Simmons, B.; Hughes, D.;
Ge, Z.; Grabowski, E. J. J. J. Am. Chem. Soc. 2004, 126, 13002-13009.
(93) Bonifácio, V. D. B. Synlett 2004, 1649-1650.
(94) Lane, C. F. Synthesis (Stuttgart) 1975, 135-146.
(95) Neises, B.; Steglich, W. Angew. Chem. Int. Ed. 1978, 17, 522-524.
(96) Zemplen, G.; Gerecs, A.; Hadacsy, I. Ber. Dtsch. Chem. Ges. 1936,
69, 1827-1829.
(97) Dale, J. A.; Dull, D. L.; Mosher, H. S. J. Org. Chem. 1969, 34, 2543-
2549.
(98) Hoye, T. R.; Jeffrey, C. S.; Shao, F. Nat. Protoc. 2007, 2, 2451-
2458.
(120) Kawaguchi, T.; Hamanaka, T.; Kito, Y.; Machida, H. J. Phys.
Chem. 1991, 95, 3837-3846.
(121) Kalyanasundaram, K.; Thomas, J. K. J. Am. Chem. Soc. 1977, 99,
2039-2044.
(122) Basu Ray, G.; Chakraborty, I.; Moulik, S. P. J. Colloid Interface
Sci. 2006, 294, 248-254.
(99) Ohtani, I.; Kusumi, T.; Kashman, Y.; Kakisawa, H. J. Org. Chem.
1991, 56, 1296-1298.
TOC Graphic
Monorhamnolipid surfactants
I. Synthetic methodology for manufacturing diastereomers of rhamnolipids
II. Characterization of surface activity and self assembly as
stereochemistry, concentration and pH.
a
function of
80
70
60
50
40
30
20
80
70
60
50
40
30
20
(S,S) Rha-C10-C10
(S,R) Rha-C10-C10
(R,S) Rha-C10-C10
(R,R) Rha-C10-C10
1.1
1.0
0.9
0.8
0.7
0.6
0.5
(S,S) Rha-C10-C10
(S,R) Rha-C10-C10
(R,S) Rha-C10-C10
(R,R) Rha-C10-C10
(S,S) Rha-C10-C10
(S,R) Rha-C10-C10
(R,S) Rha-C10-C10
(R,R) Rha-C10-C10
pH 8
apH 4
b
-16
-14
-12
-10
-8
-6
-16
-14
-12
-10
-8
-5
0
5
10
15
20
25
30
35
40
ln (Diastereomer Concentration in M)
ln (Diastereomer Concentration in M)
Diastereomer Concentration (mM)
8
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