Biocatalytic route to sugar-PEG-based polymers for drug delivery applications

Article abstract of DOI:10.1021/bm200647a

Sugar-PEG-based polymers were synthesized by enzymatic copolymerization of 4-C-hydroxymethyl-1,2-O-isopropylidene-β-l-threo-pentofuranose/ 4-C-hydroxymethyl-1,2-O-benzylidene-β-l-threo-pentofuranose/ 4-C-hydroxymethyl-1,2-O-isopropylidene-3-O-pentyl-β-l-threo-pentofuranose with PEG-600 dimethyl ester using Novozyme-435 (Candida antarctica lipase immobilized on polyacrylate). Carbohydrate monomers were obtained by the multistep synthesis starting from diacetone-d-glucose and PEG-600 dimethyl ester, which was in turn obtained by the esterification of the commercially available PEG-600 diacid. Aggregation studies on the copolymers revealed that in aqueous solution those polymers bearing the hydrophobic pentyl/benzylidene moiety spontaneously self-assembled into supramolecular aggregates. The critical aggregation concentration (CAC) of polymers was determined by surface tension measurements, and the precise size of the aggregates was obtained by dynamic light scattering. The polymeric aggregates were further explored for their drug encapsulation properties in buffered aqueous solution of pH 7.4 (37 °C) using nile red as a hydrophobic model compound by means of UV/vis and fluorescence spectroscopy. There was no significant encapsulation in polymer synthesized from 4-C-hydroxymethyl-1,2-O-isopropylidene-β-l-threo- pentofuranose because this sugar monomer does not contain a big hydrophobic moiety as the pentyl or the benzylidene moiety. Nile red release study was performed at pH 5.0 and 7.4 using fluorescence spectroscopy. The release of nile red from the polymer bearing benzylidene moiety and pentyl moiety was observed with a half life of 3.4 and 2.0 h, respectively at pH 5.0, whereas no release was found at pH 7.4.

Full text of DOI:10.1021/bm200647a

ARTICLE
pubs.acs.org/Biomac
Biocatalytic Route to Sugar-PEG-Based Polymers for Drug Delivery
Applications
Sumati Bhatia,† Andreas Mohr,‡ Divya Mathur,† Virinder S. Parmar,† Rainer Haag,*,‡ and Ashok K. Prasad*,†
†Bioorganic Laboratory, Department of Chemistry, University of Delhi, Delhi-110 007, India
‡Institute for Chemistry and Biochemistry, Free University Berlin, Takustr. 3, D- 14195 Berlin, Germany
S Supporting Information
b
ABSTRACT: Sugar-PEG-based polymers were synthesized by
enzymatic copolymerization of 4-C-hydroxymethyl-1,2-O-isopropy-
lidene-β-L-threo-pentofuranose/4-C-hydroxymethyl-1,2-O-benzyli-
dene-β-L-threo-pentofuranose/4-C-hydroxymethyl-1,2-O-isopropy-
lidene-3-O-pentyl-β-L-threo-pentofuranose with PEG-600 dimethyl
ester using Novozyme-435 (Candida antarctica lipase immobilized
on polyacrylate). Carbohydrate monomers were obtained by the
multistep synthesis starting from diacetone-D-glucose and PEG-600
dimethyl ester, which was in turn obtained by the esterification of
the commercially available PEG-600 diacid. Aggregation studies on
the copolymers revealed that in aqueous solution those polymers
bearing the hydrophobic pentyl/benzylidene moiety spontaneously
self-assembled into supramolecular aggregates. The critical aggrega-
tion concentration (CAC) of polymers was determined by surface tension measurements, and the precise size of the aggregates was
obtained by dynamic light scattering. The polymeric aggregates were further explored for their drug encapsulation properties in
buffered aqueous solution of pH 7.4 (37 °C) using nile red as a hydrophobic model compound by means of UV/vis and fluorescence
spectroscopy. There was no significant encapsulation in polymer synthesized from 4-C-hydroxymethyl-1,2-O-isopropylidene-β-L-
threo-pentofuranose because this sugar monomer does not contain a big hydrophobic moiety as the pentyl or the benzylidene
moiety. Nile red release study was performed at pH 5.0 and 7.4 using fluorescence spectroscopy. The release of nile red from the
polymer bearing benzylidene moiety and pentyl moiety was observed with a half life of 3.4 and 2.0 h, respectively at pH 5.0, whereas
no release was found at pH 7.4.
’ INTRODUCTION
is well-established now that extracellular pH of solid tumors is
significantly more acidic than normal tissues, with a mean pH of
6.5 in comparison to 7.4 for the blood and normal tissues.6ꢀ9
Therefore, polymeric micelles that are responsive to pH gradients
can be designed to release their payload selectively in tumor tissue
or within tumor cells.10ꢀ12
Several approaches can be taken to design pH-sensitive drug
carriers.13 One approach is to directly attach the drug to the drug
carrier by acid degradable linkages. For example, this approach
was used by Park et al.14 where doxorubicin was chemically
conjugated to the terminal end of poly(L-lactic acid) in diblock
copolymer of PLLA-PEG by acid cleavable hydrazone and cis-
aconityl bond. Another approach is the incorporation of acid-
cleavable hydrophobic groups into the copolymer. Such hydro-
phobic groups interact with water insoluble hydrophobic drug
molecule and hold it by noncovalent forces until they reach
certain acidic condition. Hydrophobic groups that are drug
There is always a need to have biocompatible drug carriers
capable of delivering water insoluble drugs with high transport
and controlled release capacity. Poly(ethylene glycol) (PEG)
and the materials derived from it are highly applicable in
biotechnology and pharmacology. This is due to the excellent
water solubility and high biocompatibility of PEG. PEGylation
has proved to be a successful approach to drug delivery.1
Carbohydrates are known for their target specificity and also as
promising candidates for homing molecules.2,3 Moreover, the
effectiveness of a drug carrier is determined by its ability to
control the time over which drug release occurs or to trigger the
drug release at a specific location. Drug release can be controlled
by stimuli or a host-responsive process. In recent years, drug
carriers that were responsive to their environment or to external
stimuli have been designed. For example, temperature has been
used to modulate drug release from thermoresponsive micelles,4
ultrasound has been reported to trigger drug release from pluronic
micelles,5 and change in pH has also been exploited as a useful
stimulus in the development of a drug carrier. Numerous pH
gradients exist in both normal and pathophysiological states, and it
Received: May 12, 2011
Revised:
July 29, 2011
Published: August 12, 2011
r
2011 American Chemical Society
3487
dx.doi.org/10.1021/bm200647a Biomacromolecules 2011, 12, 3487–3498
|

Biomacromolecules
ARTICLE
Scheme 1. Synthesis of the Monomers 2, 5, and 8
a Reagents and conditions: (i) Ac2O, DMAP, THF, 90%; (ii) 70% AcOHꢀH2O, 50 °C; (iii) PhCH(OMe)2, CSA, DMF, 40%; (iv) K2CO3, dry MeOH,
88%; (v) C5H11Br, TBAI, NaOH, dry DMF, 87%; (vi) 60% AcOH-H2O, 82%; (vii) NaIO4, CH2OHCH2OH, THF:H2O; and (viii) HCHO, 2 M
NaOH, dioxane:H2O, 69%.
holding groups in the copolymeric drug carrier are hydrolyzed
under acidic conditions, and the drug carrier releases its pay-
load. In the recent past, several pH-responsive block copoly-
meric systems have been designed, synthesized, and studied for
their self-assembling properties and release of hydrophobic
drugs.10,13ꢀ15 For example, Kataoka et al.16 have synthesized
and used poly(ethylene glycol)-poly(L-lysine) block copolymer
as a pH-sensitive drug delivery agent for the delivery of
photosensitizer drugs for cancer therapy. Chen et al.17 has
reported the synthesis of pH-responsive biodegradable micelles
bearing acid sensitive trimethoxybenzylidene acetal and
exploited it for the encapsulation and release study of paclitaxel
and doxorubicin. Furthermore, pH-sensitive polymersomes
have also been synthesized for the encapsulation and release
study of drugs like paclitaxel and doxorubicin.18
So far, sugar-PEG polymeric systems have not been explored
much for pH-responsive drug delivery applications. In the
present article, we report the biocatalytic synthesis of novel
sugar-PEG-based copolymeric architectures bearing acid cleava-
ble groups. These biocatalytically synthesized sugar-PEG-based
polymeric architectures have been studied for their molecular
aggregation, drug encapsulation, and drug release behavior using
nile red as a model drug.
employed. pH values were measured with a Piccolo Plus ATC pH/C
meter at 25 °C.
Instrumentation. Nuclear Magnetic Resonance Spectroscopy.
The 1H and 13C and DEPT NMR spectra were recorded on a Bruker
DRX 400 spectrometer operating at 400 and 100 MHz, respectively.
Bruker AMX 500 MHz spectrometer was used to record 2D NMR
1
13
spectrum (1Hꢀ H correlation spectrum and 1Hꢀ C correlation
spectrum). The spectra were calibrated on the solvent peak (CDCl3:
δ 7.26 for 1H NMR and δ 77.0 for 13C NMR; CD3OD: δ 4.84 for 1H
NMR and δ 49.05 for 13C NMR).
Infrared Spectroscopy, ESI, and Melting Point Measurements.
Infrared (IR) spectra were recorded as thin films between KBr or
CaF2 plates on a Bruker IFS 66 FT-IR spectrometer. For ESI measure-
ments a TSQ 7000 (Finnigan Mat) instrument and for high-resolution
mass spectra a JEOL JMS-SX-102A spectrometer was used. Melting
points of solid compounds were taken on a BUCHI melting point
apparatus M-560.
Gel Permeation Chromatography. Molecular weight and molecular
weight distribution Mw/Mn of polymers were determined using gel
permeation chromatography (GPC) equipped with an agilent 1100
pump, refractive index detector, and PLgel and Suprema columns. The
eluent was THF or water with a flow rate of 1.0 mL/min. The molecular
weights were calibrated with polystyrol or pullulan standards.
Surface Tension Measurements. Critical aggregation concentration
(CAC) of the polymers was determined by a commercially available
pendant drop tensiometer OCA 20 built by Dataphysics, Germany.
Aqueous sample concentrations in Milli-Q water were prepared 24 h
before measurement. Calculation of the surface tension was done by
using the YoungꢀLaplace Equation. In a typical experiment, the
aggregation behavior of the polymers in water was studied over the
concentration range of 7 ꢁ 10ꢀ7 to 1 ꢁ 10ꢀ3 M. The surface tension of
each hanging drop was determined two times per minute, and the
measurement was stopped when the surface tension did not change by
more than 0.1 mN/m over 2 min. Equilibration time was generally
between 25 and 70 min at lower concentrations. All measurements were
done at 25.0 ( 0.5 °C.
’ EXPERIMENTAL SECTION
Materials. Reactions requiring dry conditions were carried out
under argon. Analytical TLCs were performed on precoated Merck
silica-gel 60F254 plates; silica gel (100ꢀ200 mesh) was used for column
chromatography. Dry and analytical-grade solvents and reagents were
purchased from Sigma-Aldrich and Acros chemical companies. Novo-
zyme-435 was a gift from Novozymes A/s (Copenhagen, Denmark).
Nile red was obtained from ABCR (76187 Karlsruhe, Germany). Water
of Millipore quality (resistivity ∼18 MΩ cmꢀ1, pH 5.6 ( 0.2) was used
3
in all experiments and for preparation of all samples. Buffers of 0.01 and
Light Scattering Measurements. Dynamic light scattering measure-
ments were performed to determine the precise size of the micelle/
polymeric aggregates in aqueous solution. Measurements were carried
0.10 M phosphate were prepared by weight from Na2HPO4 7H2O and
3
NaH2PO4 H2O. For acidic pH (5.0), acetate buffer of 0.1 M was
3
3488
dx.doi.org/10.1021/bm200647a |Biomacromolecules 2011, 12, 3487–3498

Biomacromolecules
ARTICLE
out on a Zetasizer Nano ZS analyzer with integrated 4 mW HeꢀNe laser,
λ = 633 nm (Malvern Instruments, Worcestershire WR14 1XZ, U.K.).
This instrument, which uses the backscattering detection (scattering angle
θ = 173°) and an avalanche photodiode detector, is equipped with a
heliumꢀneon laser source (operating wavelength 633 nm; power 4.0
mW) and a thermostatted sample chamber controlled by a thermoelectric
peltier. For the measurement of size, aqueous solution of copolymer 11 at
the concentration of 5 g/Land copolymers12and 13atthe concentration
of 1 g/L were prepared in Milli-Q water and vigorously stirred for 12 h at
room temperature (37 °C). Solutions were filtered via 0.45 μm poly-
tetrafluoroethylene (PTFE) filters and then allowed to equilibrate for 6 h
at room temperature (37 °C). These copolymeric aqueous solutions were
used for dynamic light scattering measurements. For all experiments,
disposable UV-transparent cuvettes (12.5 ꢁ 12.5 ꢁ 45 mm, Sarstedt
AG, 51582 N€umbrecht, Germany) were used.
Transmission Electron Microscopy. TEM images of polymers were
taken on CM12 TEM (compaTEM ny FEI) instrument operating at a
high tension of 100 KV at a magnification of 60.000. 4.5% phospho-
tungstic acid (PTA) with 0.1% trehalose was used as staining material.
A droplet of the aqueous sample solution (5 μL at the concentration of 5
g/L) was taken on the copper grid (covered with a 7 nm film of carbon)
coated with collodion for 60 s. After blotting, a drop of staining material
was added for 60 s. The grid was again blotted and dried at room
temperature for at least 1 h before images were taken.
Scheme 2. Synthesis of PEG Dimethyl Ester
using K2CO3 in dry methanol24 to obtain the benzylidene sugar
monomer, that is, 4-C-hydroxymethyl-1,2-O-benzylidene-β-L-threo-
pentofuranose (5) in 88% yield (Scheme 1).
Furthermore, for the synthesis of C-3-O-pentyl pentofurano-
syl sugar monomer 8, diacetone-D-glucose 1 was treated with
bromopentane in dry DMF in the presence of sodium hydroxide,
phase transfer catalyst tetrabutyl ammonium iodide, and molec-
ular sieves to afford, 1,2:5,6-di-O-isopropylidene-3-O-pentyl-β-L-
threo-pentofuranose (6) in 87% yield. In this context, it is
worthwhile to mention that the same alkylation reaction has been
performed on other substrates in the presence of expensive base
CsOH.25 The more labile 5,6-O-isopropylidene acetal in C-3
pentyloxy sugar derivative 6 was selectively hydrolyzed by aqu-
eous acetic acid to obtain 1,2-O-isopropylidene-3-O-pentyl-D-
glucofuranose (7) in 82% yield. Oxidative cleavage of the two
vicinal hydroxyl groups in compound 7 afforded the correspond-
ing aldehyde, which was subjected to aldol-canizzaro reaction19,26
to give the 4-C-hydroxymethyl-1,2-O-isopropylidene-3-O-pentyl-
β-L-threo-pentofuranose (8) in 69% yield (Scheme 1).
Absorbance and Fluorescence Measurements. Absorption spectra
were recorded between 220 and 800 nm using a Scinco S-3150 UV/vis
spectrophotometer (range: 187ꢀ1193 nm; resolution: 1024 points).
All measurements were carried out in 10 mM phosphate buffer in a
thermostatted UV-cell (1 cm). Fluorescence emission spectra were
taken with a Jasco FP-6500 spectrofluorimeter equipped with a thermo-
statted cell holder, a DC-powered 150 W xenon lamp, a Hamamatsu
R928 photomultiplier, and a variable slit system. Emission spectra were
recorded from 575 to 800 nm after excitation at 550 nm. Both excitation
and emission slits were set at 5 nm. For release studies, the intensity of
the emission spectrum of nile red was monitored as a function of time at
pH 5.0 and physiological pH (7.4) by maintaining the temperature at
37.0 ( 0.1 °C. Data were fitted to the relation It = I0 exp(ꢀkobs t). Here
Synthesis of the PEG Dimethyl Ester 10. The commercially
available polyethylene glycol bis(carboxymethyl) ether (Mn 600)
(9) was esterified into poly[ethylene glycol bis(carboxymethyl)
ether] dimethyl ester (10) in a single-step reaction by refluxing
the substrate in methanol in the presence of a catalytic amount of
H2SO4 in almost quantitative yield (Scheme 2).
All synthesized compounds 2ꢀ8 and 10 were unambiguously
1
identified on the basis of their spectral (IR, H and 13C NMR,
DEPT-135 NMR spectra, and HRMS) data analysis. The struc-
tures of the known compounds 2 and 10 were further confirmed
by comparing their physical and spectral data with those reported
in the literature.19,27
3
3
It describes the intensity measured at time t, and I0 denotes the initial
fluorescence intensity. The half-life time is given by the simple relation-
ship t1/2 = ln(2)/kobs
.
Synthesis of Polymers 11ꢀ13. Lipases have been used for
selective esterification/transesterification reactions in the en-
zyme-mediated organic synthesis.28,29 Furthermore, one of the
lipases, viz. Novozyme-435 (Candida antarctica lipase immobi-
lized on polyacrylate), has been explored for carrying out selective
reactions necessary for carbohydrate modifications30ꢀ32 and
polymer synthesis.33ꢀ36 Taking lead from our previous experi-
ence of using lipases for discrimination between the two ester
functions of triacylated pentofuranose derivatives with an aim to
synthesize bicyclonucleosides,25,30,31 Novozyme-435 lipase was
used to affect the copolymerization reaction of hydroxy sugar
monomers 2, 5, or 8 with the PEG dimethyl ester 10 under high
vacuum and solventless conditions at 70 °C to synthesize poly-
[polyoxyethylene-oxy-bis(carboxymethyl)-4-methylene-1,2-O-
isopropylidene-β-L-threo-pentofuranosyl] (11), poly[polyoxy-
ethylene-oxy-bis(carboxymethyl)-4-methylene-1,2-O-benzylidene-
β-L-threo-pentofuranosyl] (12), and poly[polyoxyethylene-oxy-
bis(carboxymethyl)-4-methylene-1,2-O-isopropylidene-3-O-
pentyl-β-L-threo-pentofuranosyl] (13) in 72, 54, and 67% yields,
respectively. All three polymerization reactions when performed
under identical conditions but without adding lipase did not yield
any product. To our knowledge, it is the first example of trans-
esterification reaction involving tri/dihydroxy sugar derivatives
’ RESULTS AND DISCUSSION
Synthesis of Carbohydrate Monomers 2, 5, and 8. The
trihydroxy compound 4-C-hydroxymethyl-1,2-O-isopropylidene-
β-L-threo-pentofuranose (2) was prepared starting from com-
mercially available diacetone-D-glucose 1 by following the mod-
ified procedure of Youssefyeh et al.19 and Christensen et al.20 and
unambiguously identified on the basis of its spectral data (1H
NMR, 13C NMR, IR spectra, and HRMS) analysis and compar-
ison of the spectral data with those reported in the literature19
(Scheme 1). Furthermore, the trihydroxy sugar derivative 2 was
peracetylated with acetic anhydride in the presence of catalytic
amount of DMAP in THF to afford 4-C-acetoxymethyl-3,5-di-O-
acetyl-1,2-O-isopropylidene-β-L-threo-pentofuranose 3 in 90%
yield. Deprotection of 1,2-O-isopropylidene protection in com-
pound 3 was achieved using an aqueous solution of AcOH (70%)
at 50 °C, and the crude triacetoxy sugar derivative obtained was
used as such for the 1,2-O-benzylidene protection using benzalde-
hyde dimethylacetal in the presence of camphor sulfonic acid
(CSA)21ꢀ23 as catalyst to afford 4-C-acetoxymethyl-3,5-di-O-acet-
yl-1,2-O-benzylidene-β-L-threo-pentofuranose (4) in 40% yield.
The triacetylated sugar compound 4 was completely deacetylated
3489
dx.doi.org/10.1021/bm200647a |Biomacromolecules 2011, 12, 3487–3498

Biomacromolecules
ARTICLE
Scheme 3. Biocatalytic Route to the Sugar-PEG-Based Polymers 11ꢀ13
and PEG dimethyl ester leading to the formation of copolymers
11ꢀ13 (Scheme 3). All three polymers 11ꢀ13 were unambigu-
ously characterized from their IR, 1H NMR, 13C NMR, DEPT-
135 NMR, and 2D NMR spectral data analysis.
polymer 11 were confirmed by its DEPT-135 NMR spectrum.
The number-average molecular weight of the polymer 11 as
obtained by GPC in H2O (1 mL/min.) was 4408 g/mol with
polydispersity index (PDI) of 1.96, which indicates the presence of
approximately six monomeric units in the polymer.
Characterization of Polymers 11ꢀ13. Poly[polyoxyethylene-
oxy-bis(carboxymethyl)-4-methylene-1,2-O-isopropylidene-β-L-threo-
pentofuranosyl] (11). In the 1H NMR spectrum of polymer 11, the
C-5H and C-10H recorded a downfield shift of 0.54 to 0.69
ppm and resonated at δ 4.11ꢀ4.39 (m) with respect to the same
protons resonating at δ 3.57ꢀ3.70 (m) in the corresponding
monomer 4-C-(hydroxymethyl)-1,2-O-isopropylidene-β-L-threo-
pentofuranose (2). The downfield shift in the chemical shift value
of C-5H and C-10H in the polymer revealed that the two primary
hydroxyl groups at the corresponding positions in the starting
compound 2 were engaged in copolymerization over the lone
secondary hydroxyl group present at C-3 position in the monomer
2. Therefore, the lipase exhibited regioselectivity and recognized
the two primary hydroxyl groups in the monomer 2 for transes-
terification reaction with PEG dimethyl ester 10. This was further
supported by the fact that there was no appreciable change in the
chemical shift value of C-3H (δ 4.26) in the 1H NMR spectrum of
polymer 11 with respect to the chemical shift value of the same
proton (δ 4.20) in the monomer 2. The chemical shift value δ 4.26
Poly[polyoxyethylene-oxy-bis(carboxymethyl)-4-methy-
lene-1,2-O-benzylidene-β-L-threo-pentofuranosyl] (12). In the
1H NMR spectrum of polymer 12, the C-5H and C-100H
recorded a downfield shift of 0.47 to 0.56 ppm and resonated
at δ 4.08ꢀ4.38 (m) with respect to the same protons resonat-
ing at δ 3.61ꢀ3.82 (m) in the corresponding monomer
4-C-(hydroxymethyl)-1,2-O-benzylidene-β-L-threo-pentofuranose
(5). The downfield shift in the chemical shift value of C-5H and
C-100H in the polymer revealed that the two primary hydroxyl
groups at the corresponding positions in the starting compound
5 were engaged in copolymerization over the lone secondary
hydroxyl group present at C-3 position in the monomer 5.
Therefore, the lipase exhibited regioselectivity and recognized
the two primary hydroxyl groups in the monomer 5 for transes-
terification reaction with PEG dimethyl ester 10. This was further
supported by the fact that there was no appreciable change in the
chemical shift value of C-3H (δ 4.27) in the 1H NMR spectrum
of polymer 12 with respect to the chemical shift value of the same
proton (δ 4.25) in the monomer 5. The chemical shift value δ
1
of C-3H in the H NMR spectrum of polymer 11 was con-
13
firmed by the 1Hꢀ C correlation spectrum (Figure 1), whereas
1
4.27 of C-3H in the H NMR spectrum of polymer 12 was
the chemical shift value of C-3 in the 13C NMR spectrum of the
polymer was confirmed by DEPT-135 NMR (Figure 2). The
presence of all other ꢀCH, ꢀCH2, and ꢀCH3 groups in the
confirmed by 1Hꢀ C correlation spectrum (Figure 3), whereas
13
the chemical shift value of C-3 in the 13C NMR spectrum of
the polymer was confirmed by DEPT-135 NMR (Figure 4).
3490
dx.doi.org/10.1021/bm200647a |Biomacromolecules 2011, 12, 3487–3498

Biomacromolecules
ARTICLE
Figure 1. 1Hꢀ C correlation spectrum (Bruker AMX-500, chloroform-d) of polymer 11.
13
Figure 2. Comparison of 13C and DEPT-135 NMR spectrum (Bruker DRX-400, 100 MHz, chloroform-d) of polymer 11.
1
The presence of all other ꢀCH, ꢀCH2, and ꢀCH3 groups in the
polymer 12 was confirmed by its DEPT-135 NMR spectrum.
The number-average molecular weight of the polymer 12 as
obtained by GPC in THF (1 mL/min.) was 5682 g/mol with
PDI of 1.11, which indicates the presence of approximately seven
monomeric units in the polymer.
In the H NMR spectrum of polymer 13, the C-5H and C-100H
recorded a downfield shift of 0.51 to 0.68 ppm and resonated at δ4.09
to 4.39 (m) instead of δ 3.58 to 3.71 (m) with respect to the same
protons in corresponding monomer 4-C-(hydroxymethyl)-1,2-O-iso-
propylidene-3-O-pentyl-β-L-threo-pentofuranose (8). The downfield
shift in the chemical shift value of C-5H and C-100H in the polymer
revealed that the two primary hydroxyl groups at the corresponding
positions in the starting compound 8are engaged in transesterification
Poly[polyoxyethylene-oxy-bis(carboxymethyl)-4-methylene-1,
2-O-isopropylidene-3-O-pentyl-β-L-threo-pentofuranosyl] (13).
3491
dx.doi.org/10.1021/bm200647a |Biomacromolecules 2011, 12, 3487–3498

Biomacromolecules
ARTICLE
Figure 3. 1Hꢀ C correlation spectrum (Bruker AMX-500, chloroform-d) of polymer 12.
13
Figure 4. Comparison of 13C and DEPT-135 NMR spectrum (Bruker DRX-400, 100 MHz, chloroform-d) of polymer 12.
reaction with PEG dimethyl ester 10. The presence of all other ꢀCH,
ꢀCH2, and ꢀCH3 groups in the polymer 13 was confirmed by its
DEPT-135 NMR spectrum (Figure 5). The number-average molecular
weight of the polymer 13 as obtained by GPC in THF (1 mL/min.)
was 5882 g/mol with PDI of 1.26, which indicates the presence of
approximately seven monomeric units in the polymer.
Aggregation Behavior in Aqueous Solution. To under-
stand the aggregation behavior of the synthesized polymers
11ꢀ13 in aqueous solution, we investigated structural aspects
of polymers using surface tension measurement and dynamic
light scattering. Surface tension data were plotted as a logarithmic
function of the surfactant concentration, where a break in the
3492
dx.doi.org/10.1021/bm200647a |Biomacromolecules 2011, 12, 3487–3498

Biomacromolecules
ARTICLE
Figure 5. Comparison of 13C and DEPT-135 NMR spectrum (Bruker DRX-400, 100 MHz, chloroform-d) of polymer 13.
Table 1. Critical Aggregation Concentration (CAC), Particle
Size, and Polydispersity Index (PDI) of Micelles/Aggregates
11-13 in Aqueous Solution Measured by Light Scattering
Method at 25°C
copolymer
CAC (M)
z-average diameter (nm)
PDI
11
12
13
4.5 ꢁ 10ꢀ5
2.0 ꢁ 10ꢀ5
4.0 ꢁ 10ꢀ6
13a
124b
223b
0.241
0.207
0.105
a [11] = 5 g/L. b [12] and [13] = 1 g/L of polymers.
The precise size of the aggregates of polymers 11 to 13 in
aqueous solution was determined by using dynamic light scatter-
ing. All polymers 11ꢀ13 were soluble in water as all sample
solutions appeared clear. Measurements were carried out well
above the CACs, that is, at a fixed polymer concentration of 1 g/L
for polymers 12 and 13 and 5 g/L for polymer 11. The results
have been summarized in Table 1, and the corresponding size
distribution profiles are displayed in Figure 7. For polymers
11ꢀ13 the hydrodynamic diameters were determined to be 13,
124, and 223 nm, respectively. A comparison of the intensity
distribution profile with the corresponding volume and number
distribution shows that all three compositions have a mono-
modal particle size distribution (Figure 7).
The aggregates of small size, dH = 13 nm, can be regarded as
polymeric micelles presumably with the pentofuranose building
block making the core and the polyoxyethylene chain constitut-
ing the corona of the micelle (Figure 8).
There is a significant increase in the measured size as the
pentofuranose moiety is modified with a phenyl group (polymer 12)
or a pentyl chain (polymer 13). The order of magnitude obtained for
12 and 13 in aqueous solution does not correspond to simple
polymeric micelles but rather to larger aggregates (clusters). Depend-
ing on the polymer concentration in water, the micelles may self-
assemble to a loose network of aggregates formed because of
intermicellar interactions through polyoxyethylene corona. Individual
Figure 6. Surface tension (γ) versus log concentration of the sugar-
PEG-based polymers 11 (O), 12 (Δ), and 13 (0) in aqueous solution at
25 °C. Measurements were performed in duplicate, and the surface
tension obtained varied in the range of (1 mN/m.
curve occurs at the CAC. As can be seen from the surface tension
curves in Figure 6, polymers 11ꢀ13 were effective in reducing
the surface tension (γ) of aqueous solution, thus showing a
considerable decrease in γ as the polymer concentrations
increased, followed by clear break point at CAC of the respective
polymers. The CAC values obtained are equal to 4.5 ꢁ 10ꢀ5
,
2.0 ꢁ 10ꢀ5, and 4.0 ꢁ 10ꢀ6 M for polymers 11ꢀ13, respectively.
From Figure 6, it is clearly observed that the attachment of a
pentyl chain to the pentofuranose moiety promotes aggregation
to a greater extent than a phenyl group. Thus, the CAC of
copolymer 12 is approximately five times higher than that of
copolymer 13. The observed CAC of polymer 11 was found to be
2.2 fold higher than that of polymer 12 bearing benzylidene
moiety. This shows that the absence of any hydrophobic moiety
leads to the aggregation to a lesser extent.
3493
dx.doi.org/10.1021/bm200647a |Biomacromolecules 2011, 12, 3487–3498

Biomacromolecules
ARTICLE
micelles and clusters are in equilibrium; that is, the supramolecular
network breaks and forms reversibly37,38 (Figure 9). We assume that
the aggregates of 12 and 13 are formed through an attractive
interaction between the PEG moieties that constitute the corona of
the micelles. The hydrophobic sugar moieties of the copolymers form
the core of the micelles and are unlikely to contribute to intermicellar
binding interactions. Furthermore, the copolymers 12 and 13 are not
end-capped with hydrophobic moieties (hexadecyl, dodecyl, etc.), as
discussed in the literature,39ꢀ44 where aggregates are formed by
hydrophobic interactions. Furthermore, such reports are there in the
literature where intermicellar aggregates are formed through hydro-
philic interaction of the polyethylene oxide chain in a dendritic
structure. Therefore, dendritic multishell architectures bearing term-
inal monomethyl poly(ethylene glycol) (mPEG) outer shells were
found to self-assemble into supramolecular aggregates in water.45
Similar to the micelles/aggregates described herein, the terminal
mPEG layer acts as an external polar layer, thus providing good
solubility in aqueous solution. As shown by surface tension measure-
ments and fluorescence spectroscopy, self-assembly spontaneously
occurs above a well-defined threshold concentration (CAC).
In conclusion, surface tension measurements and light scatter-
ing analysis have shown that modification of the pentofuranose
unit has significant effects on CAC (aggregation behavior), the
micelle/micellar aggregate formation, and their size distribution.
Structural analysis of the micelles and micellar assemblies of
polymers 11ꢀ13 have also been investigated by means of
transmission electron microscopy. As can be seen from Figure 10,
all polymers showed nanoscopic spherical morphologies in aqu-
eous media. For comparison purpose, aqueous sample solutions
had the same concentration as that used in the case of DLS
measurement, that is, 5 g/L for polymer 11 and 1 g/L for polymers
12 and 13, respectively. The hydrodynamic diameters of micellar
aggregates of hydrophobically modified polymers 12 and 13 are
considerably higher than those estimated by TEM (Figure 10).
This may be attributed to the high swelling capacity of micellar
aggregates of polymers 12 and 13 in water. It is worth mentioning
that the TEM analysis was carried out on dried samples, whereas
light scattering experiments were performed on aqueous solutions.
In fact, this behavior has been previously described by Aktas et al.,46
who investigated the size distribution of chitosan and PEGylated
chitosan nanoparticles (NPs) in aqueous solutions by means of
dynamic light scattering and transmission electron microscopy.
Study of Encapsulation of Nile Red by Micelles/Aggre-
gates of Polymer 11ꢀ13. The encapsulation properties of
Figure 7. Size distribution profiles for aggregates of 11ꢀ13 in aqueous
solution. The concentration of the polymers was fixed at 1 (polymers
12 & 13) and 5 g/L (polymer 11).
Figure 8. Proposed schematic representation of polymeric micelle copolymer 11 in aqueous solution.
Figure 9. Proposed schematic representation of polymeric micellar aggregates of copolymer 12 and 13 in aqueous solution.
3494
dx.doi.org/10.1021/bm200647a |Biomacromolecules 2011, 12, 3487–3498

Biomacromolecules
ARTICLE
Figure 10. Transmission electron microscopy images of micelles of polymer 11 (a) and micellar aggregates of polymer 12 (b) and 13 (c). Spherical
shape morphologies were observed with diameter in the range of 15ꢀ30 nm in the case of polymer 11 and 20ꢀ40 nm in the case of polymer 12. Highly
polydispersed spheres with diameter up to 60 nm were observed in case of polymer 13. The scale bar is equal to 100 nm in all of the cases.
“to probe the microenvironment” of encapsulated nile red
molecules. The maximum of the absorption band is hypsochro-
mically shifted from 557 to 540 nm on going from ethyleneglycol
(ε = 37.7) to PEG 400 (ε = 14.3) as solvent. Hence, from the
spectroscopic results, it can be inferred that the solubilization of
nile red appears to primarily take place, at least to an appreciable
extent, within the poly(oxyethylene) layer of 12 and 13. All
copolymers are noncharged and consist of a polyoxyethylene
backbone to which functionalized pentofuranose-based moieties
are attached. In this context, it is important to note that apart
from the PEG chains only water molecules and ions (buffer)
contribute to the polarity of the hydrophilic poly(oxyethlene)
region. This might lead to differences in the observed wavelength
of the absorption maxima when comparing with neat ethylene-
glycol and PEG 400, respectively.
The inset in Figure 11 shows the corresponding fluorescence
Figure 11. UV absorption of nile red solubilized by polymeric micelles
of 12 and 13 in buffered aqueous solution (pH 7.4, 10 mM). Inset:
corresponding fluorescence emission spectra of nile red (λexc = 550 nm)
in the presence of 12 and 13, respectively.
emission spectra. Both the buffered aqueous solutions of nile-
red-loaded polymeric aggregates of polymers 12 and 13 were
given excitation at 550 nm. Excitation of the dye present in the
buffered aqueous solution of polymeric aggregates of 12 results
in a relatively low fluorescence with a λmax of 645 nm. On the
contrary, in the presence of 13, the fluorescence intensity is
roughly nine times higher than that for 12. At the same time, the
maximum of the emission band is slightly shifted to a lower
wavelength of 635 nm. The spectroscopic results therefore imply
that in the presence of polymeric aggregates of polymer 13, the
solubilized dye essentially experiences a more hydrophobic
environment.
At first sight, it appears paradoxical that we found a maximum
absorbance at 530 and 568 nm for 12 and 13, respectively. To our
surprise, however, we have observed that with increasing amount
of nile red stock solution, the absorption band undergoes a
hypsochromic shift to a much shorter wavelength with a max-
imum in the 450ꢀ500 nm range, accompanied by an intense
shoulder in the red region of the spectrum. This unusual spectro-
scopic behavior suggests strong binding interactions between the
dye molecules. The self-association of dyes in solution is a well-
known phenomenon in dye chemistry owing to noncovalent
(supramolecular) binding interactions between the molecules.
Therefore, we have chosen a quite low amount (25 μL) of nile red
stock solution. In this regard, it is worth mentioning a recent
report by Varghese and Wagenknecht,48 who investigated the
noncovalent self-assembly of nile-red-modified 2-deoxyuridine in
water. They found that the absorption spectrum in water (ε = 78)
shows a maximum at 540 nm, which is 30 nm blueshifted relative
to the maximum in methanol (570 nm, ε = 32). This unusual
hypsochromic shift has been assigned to the formation of H-type
copolymers 11ꢀ13 were spectroscopically investigated by using
the water-insoluble hydrophobic model dye nile red. In recent
years, nile red has been extensively used as a probe for solvent
polarity and hydrophobicity.47 Nile red is particularly beneficial in
sensing the micropolarity of modified polymers in aqueous solution.
For the encapsulation experiments, a 10 mM nile red stock
solution was freshly prepared by dissolving an appropriate
amount of the dye in dry THF. Aliquots (25 μL) were added
to 1.0 mL of the aqueous polymer solutions (1.0 mg/mL) in
phosphate buffer (100 mM) of pH 7.4, and after subsequent
removal of the organic solvent, the obtained aqueous solutions
were stirred for at least 18 h at room temperature. The mixture
was filtered through a syringe filter (Rotilabo, 0.45 μm pore size)
to remove the insoluble excess of nile red. The colored aqueous
solutions of the polymers 12 and 13 were then analyzed by
means of UV/vis and fluorescence spectroscopy.
As can be seen from Figure 11, the λmax values for the UV/vis
spectra of nile-red-loaded buffered aqueous solution of polymer
12 and 13 are 530 and 568 nm, respectively. These data clearly
suggest a considerable change in the microenvironment for
solubilized dye. Nile red is a solvatochromic dye molecule and
can be used to probe the polarity or rather the effective dielectric
constant of its (micro)environment. The solvatochromic beha-
vior of nile red is such that the absorption and emission maxima
strongly depend on the polarity of the environment (solvent).
We have chosen ethylene glycol and PEG 400 as model systems
3495
dx.doi.org/10.1021/bm200647a |Biomacromolecules 2011, 12, 3487–3498

Biomacromolecules
ARTICLE
Figure 12. Fluorescence intensity curves observed at different time intervals during release of nile red from aggregates of copolymer s12 and 13 in
buffered aqueous solution of pH 5 at 37 °C. Change in fluorescent intensity (a.u.) versus wavelength (nm) for copolymers 12 (a) and 13 (b). Observed
fluorescence intensity (a.u.) decreases with time in acidic pH 5.0.
Figure 13. Release of nile red from aggregates of copolymers 12 and 13 in buffered aqueous solutions of pH 5.0 and physiological pH 7.4 at 37 °C.
Percentage initial maximum fluorescence observed at different time intervals for copolymers 12 (a) and 13 (b).
aggregates in water. We therefore assume that in the presence of
12, nile red tends to form dyeꢀpolymer(dye) aggregates in
buffer, in accord with an absorption maximum at 530 nm. For
polymer 13, however, a considerable higher amount of dye is
required. Under the experimental condition employed, self-
assembly is less favored, and solubilized dye molecules exist to
an appreciable extent as monomers, thus showing an absorption
maximum at 568 nm. At the present, we cannot offer direct
evidence of the assumption. This phenomenon, however, is
currently under investigation.
was divided into two samples; the pH of the one sample was
adjusted to pH 5.0 by the addition of a small aliquot of
concentrated acetate buffer (4 M). The second sample was
maintained at pH 7.4, but the salt concentration was adjusted
to the same as that above by the addition of the concentrated
phosphate buffer (2 M). To exclude any effect due to changes of
salt concentration, the two samples were adjusted to the same salt
concentration of 100 mM.
In acidic solution of pH 5.0, the polymeric micellar aggregates
of 12 and 13 exhibited a significant decrease in the maximum
intensity with time, whereas the same nile-red-loaded systems
were not affected at physiological pH 7.4 (Figures 12 and 13).
This suggests that our polymeric systems respond to subtle pH
changes in the physiological environment. The plot of % initial
fluorescence versus time obtained was used to estimate the half-
life time of dye release. Data were fitted to the relation It =
I0 exp(ꢀkobs t), and the half-life time was found to be 3.4 and
Finally, no noticeable absorbance was detected upon the addi-
tion of 11. This indicates that the hydrophobic dye molecules reside
preferably at the aqueous bulk phase.
Release of Nile Red. Our particular interest in the present
study was to investigate the polymeric micellar aggregates of 12
and 13 as controlled drug release vehicles. Therefore, we studied
the time-dependent release of solubilized nile red at acidic pH
(5.0, 37 °C) by using fluorescence spectroscopy.49ꢀ51
We followed the protocol previously described by Gillies
et al.51 in which the polymers were first equilibrated with nile
red overnight in 10 mM phosphate buffer (pH 7.4). The solution
3
3
2.0 h (r2 g 0.990) for 12 and 13, respectively. For polymeric
aggregates of 13, a complete release was observed in ∼25 h. On
the contrary, the % initial fluorescence was reduced to only half of
its intensity when polymer 12 was used. The in vitro release
3496
dx.doi.org/10.1021/bm200647a |Biomacromolecules 2011, 12, 3487–3498

Biomacromolecules
ARTICLE
profiles of the solubilized dye determined at acidic pH have
shown that the two copolymers have good potential for the
controlled release of hydrophobic drug molecules.
(2) Nakamura, T.; Nagasaki, Y.; Kataoka, K. Bioconjugate Chem.
1998, 9, 300–303.
(3) Koyama, Y.; Ishikawa, M.; Iwamoto, M.; Kojima, S. J. Controlled
Release 1992, 22, 253–261.
(4) Cammas, S.; Suzuki, K.; Sone, C.; Sakurai, Y.; Kataoka, K.;
Okano, T. J. Controlled Release 1997, 48, 157–164.
(5) Chung, J. E.; Yokoyama, M.; Okano, T. J. Controlled Release
2000, 65, 93–103.
(6) Engin, K.; Leeper, D. B.; Cater, J. R.; Thistlethwaite, A. J.;
Tupchong, L.; McFarlane, J. D. Int. J. Hypertherm. 1995, 11, 211–
216.
(7) van Sluis, R.; Bhujwalla, Z. M.; Ballerteros, P.; Alverez, J.; Cerdan,
S.; Galons, J. P.; Gillies, R. J. Magn. Reson. Med. 1999, 41, 743–750.
(8) Ojugo, A. S. E.; Mesheehy, P M. J.; McIntyre, D. J. O.; McCoy,
C.; Stubbs, M.; Leach, M. O.; Judson, I. R.; Griffiths, J. R. NMR Biomed.
1999, 12, 495–504.
(9) Calderon, M.; Quadir, M. A; Strumia, M.; Haag, R. Biochimie
2010, 92, 1242–1251.
(10) Gillies, E. R.; Frechet, J. M. J. Bioconjugate Chem. 2005,
16, 361–368.
(11) Haag, R. Angew. Chem., Int. Ed. 2004, 43, 278–282.
(12) Haag, R.; Kratz, F. Angew. Chem., Int. Ed. 2006, 45, 1198–1215.
(13) Gillies, E. R.; Frechet, J. M. J. Pure Appl. Chem. 2004,
76, 1295–1307.
(14) Yoo, H. S.; Lee, E. A.; Park, T. G. J. Controlled Release 2002,
82, 17–27.
’ CONCLUSIONS
A new biocatalytic route has been designed and developed to
synthesize three novel sugar-PEG-based amphiphilic polymers
11ꢀ13 by copolymerization of functionalized pentofuranose
backbone with PEG-600 dimethyl ester using Novozyme-435.
Regioselectivity has been exhibited in the polymerization of
monomers 2 and 5 with PEG dimethyl ester, where PEGylation
occurs only at the primary hydroxyl groups leading to the
formation of polymers 11 and 12. All three polymers have been
explored for drug delivery application. Attachment of phenyl and
pentyl moieties to the pentofuranose backbone in the copoly-
mers 12 and 13, respectively, leads to the formation of supra-
molecular aggregates of diameters 124 and 223 nm, respectively,
in the aqueous solution, as observed by DLS. Supramolecular
aggregates of polymers 12 and 13 were capable to encapsulate
nile red molecule, as revealed by the guest enacapsulation studies
done in buffered aqueous solutions. Nile red release in case of
polymer 13 was found to be faster with a half-life time of 2.0 h
than that in the polymer 12 with a half-life time of 3.4 h at
acidic pH 5.0 (37 °C). No release was observed at physiological
pH 7.4 (37 °C), as observed by fluorescence studies of nile-red-
loaded aggregates. Further studies are in progress to achieve the
lead polymeric architectures with higher drug loading capacity
and longer lasting release profiles.
(15) Satturwar, P.; Eddine, M. N.; Ravenelle, F.; Leroux, J.-C. Eur. J.
Pharm. Biopharm. 2007, 65, 379–387.
(16) Jang, W.-D.; Nishiyama, N.; Zhang, G.-D.; Harada, A.; Jiang,
D.-L.; Kawauchi, S.; Morimoto, Y.; Kikuchi, M.; Koyama, H.; Aida, T.;
Kataoka, K. Angew. Chem., Int. Ed. 2005, 44, 419–423.
(17) Chen, W.; Meng, F.; Li, F.; Ji, S.-J.; Zhong, Z. Biomacromolecules
2009, 10, 1727–1735.
’ ASSOCIATED CONTENT
(18) Chen, W.; Meng, F.; Cheng, R.; Zhong, Z. J. Controlled Release
2010, 142, 40–46.
(19) Youssefyeh, R. D.; Verheyden, J. P. H.; Moffatt, J. G. J. Org.
Chem. 1979, 44, 1301–1309.
(20) Christensen, S. M.; Hansen, H. F.; Koch, T. Org. Process Res.
Dev. 2004, 8, 777–780.
(21) Yang, K. L.; Blackman, B.; Diederich, W.; Flaherty, P. T.;
Mossman, C. J.; Roy, S.; Ahn, Y. M.; Georg, G. I. J. Org. Chem. 2003,
68, 10030–10039.
S
Supporting Information. Experimental details of synth-
b
esis of monomers and copolymers, H NMR and 13C NMR
1
spectra of compounds 2ꢀ8 and for copolymers 11ꢀ13, DEPT-
135 NMR spectra of compounds 4ꢀ8 and for copolymers
1
1
11ꢀ13, Hꢀ H COSY spectrum of copolymer 13, and gel
permeation chromatograms of copolymers 11ꢀ13. This material
is available free of charge via the Internet at http://pubs.acs.org.
(22) Crich, D.; Li, M. J. Org. Chem. 2008, 73, 7003–7010.
(23) Yu, S.-H.; Wang, H.-Y.; Chiang, L.-W.; Pei, K. Synthesis 2007,
9, 1412–1420.
’ AUTHOR INFORMATION
(24) Prasad, A. K.; Kalra, N.; Yadav, Y.; Kumar, R.; Sharma, S. K.;
Patkar, S.; Lange, L.; Wengel, J.; Parmar, V. S. Chem. Commun.
2007, 2616–2617.
(25) Dueno, E. E.; Chu, F.; Kim, S.-I.; Jung, K. W. Tetrahedron Lett.
1999, 40, 1843–1846.
Corresponding Author
*(R.H.) Tel: +49-30-838-52633; Fax: +49-30-838-53357;
E-mail: haag@chemie.fu-berlin.de. (A.K.P.) Tel: 00-91-11-
27662486; E-mail: ashokenzyme@yahoo.com.
(26) Dhavale, D. D.; Matin, M. M. Tetrahedron 2004, 60, 4275–
4281.
(27) Gupta, S.; Pandey, M. K.; Levon, K.; Haag, R.; Watterson, A. C.;
Parmar, V. S.; Sharma, S. K. Macromol. Chem. Phys. 2010, 211, 239–244.
(28) Faber, K. Biotransformations in Organic Chemistry, 5th ed.;
Springer-Verlag: Berlin, 2004.
(29) Drauz, K.; Waldmann, H. Enzyme Catalysis in Organic Synthesis;
VCH: Weinheim, Germany, 1994; Vols. 1 and 2.
(30) Prasad, A. K.; Kalra, N.; Yadav, Y.; Singh, S. K.; Sharma, S. K.;
Patkar, S.; Lange, L.; Olsen, C. E.; Wengel, J.; Parmar, V. S. Org. Biomol.
Chem. 2007, 5, 3524–3530.
(31) Maity, J.; Shakya, G.; Singh, S. K.; Ravikumar, V. T.; Parmar,
V. S.; Prasad, A. K. J. Org. Chem. 2008, 78, 5629–5632.
(32) Singh, S. K.; Sharma, V. K.; Olsen, C. E.; Wengal, J.; Parmar,
V. S.; Prasad, A. K. J. Org. Chem. 2010, 75, 7932–7935.
(33) Kumar, R.; Chen, M.-H.; Parmar, V. S.; Samuelson, L. A.;
Kumar, J.; Nicolosi, R.; Yoganathan, S.; Watterson, A. C. J. Am. Chem.
Soc. 2004, 126, 10640–10644.
’ ACKNOWLEDGMENT
We thank the Department of Biotechnology (DBT, New
Delhi, India) and the International Bureau (IB) of the Federal
Ministry of Education and Research (BMBF, Bonn, Germany)
and the University of Delhi under the DU-DST Purse grant for
financial assistance to this work. S.B. thanks Institute for Chem-
istry and Biochemistry, Free University Berlin, Germany for
providing the opportunity and infrastructure to accomplish a part
of this work. S.B. and D.M. thank UGC, New Delhi for the award
of research fellowship.
’ REFERENCES
(1) Francesco, M. V.; Gianfranco, P. Drug Discovery Today 2005,
10, 1451–1458.
3497
dx.doi.org/10.1021/bm200647a |Biomacromolecules 2011, 12, 3487–3498

Biomacromolecules
ARTICLE
(34) Kumar, R.; Shakil, N. A.; Chen, M.-H.; Parmar, V. S.; Samuelson,
L. A.; Kumar, J.; Watterson, A. C. J. Macromol. Sci., Part A:Pure Appl.Chem.
2002, A39, 1137–1149.
(35) Sharma, S. K.; Husain, M.; Kumar, R.; Samuelson, L. A.; Kumar,
J.; Watterson, A. C.; Parmar, V. S. Pure Appl. Chem. 2005, 77, 209–226.
(36) Malhotra, S.; Calderon, M.; Prasad, A. K.; Parmar, V. S.; Haag,
R. Org. Biomol. Chem. 2010, 8, 2228–2237.
(37) Hussain, H.; Busse, K.; Kressler, J. Macromol. Chem. Phys. 2003,
204, 936–946.
(38) Yang, J.; Zhang, D.; Jiang, S.; Yang, J.; Nie, J. J. Colloid Interface
Sci. 2010, 352, 405–414.
(39) Berret, J.-F.; Calvet, D.; Collet, A.; Viguier, M. Curr. Opin.
Colloid Interface Sci. 2003, 8, 296–306.
(40) Alami, E.; Almgren, M.; Brown, W.; Franc-ois, J. Macromolecules
1996, 29, 2229–2243.
(41) Chassenieux, C.; Nicolai, T.; Durand, D. Macromolecules 1997,
30, 4952–4958.
(42) Xu, B.; Yetka, A.; Li, L.; Masoumi, Z.; Winnik, M. A. Colloids
Surf., A 1996, 112, 239–250.
(43) Xu, B.; Yetka, A.; Winnik, M. A.; Sadeghy-Dalivand, K.; James,
D. F.; Jenkins, R.; Basset, D. Langmuir 1997, 13, 6903–6911.
(44) Le Meins, J.-F.; Tassin, J.-F. Colloid Polym. Sci. 2003,
281, 283–287.
(45) Radowski, M. R.; Shukla, A.; Berlepsch, H. V.; Bottcher, C.;
Pickaert, G.; Rehage, H.; Haag, R. Angew. Chem., Int. Ed. 2007,
46, 1265–1269.
(46) Aktas, Y.; Yemisci, M.; Andrieux, K.; Gursoy, R. N.; Alonso,
M. J.; Fernandez-Megia, E.; Novoa-Carballal, R.; Quinoa, E.; Riguera, R.;
Sargon, M. F.; Celik, H.; Demir, A. S.; Hıncal, A. A.; Dalkara, T.; Capan,
Y.; Couvreur, P. Bioconjugate Chem. 2005, 16, 1503–1511.
(47) Jee, A.-Y.; Park, S.; Kwon, H.; Lee, M. Chem. Phy. Lett. 2009,
477, 112–115.
(48) Varghese, R.; Wagenknecht, H.-A. Chem. Eur. J. 2010, 16,
9040–9046.
(49) Greenspan, P.; Mayer, E. P.; Fowler, S. D. J. Cell Biol. 1985,
100, 965–973.
(50) Krishna, M. M. G. J. Phys. Chem. A 1999, 103, 3589–3595.
(51) Gillies, E. R.; Jonsson, T. B.; Frechet, J. M. J. J. Am. Chem. Soc.
2004, 126, 11936–11943.
3498
dx.doi.org/10.1021/bm200647a |Biomacromolecules 2011, 12, 3487–3498