Compact biocompatible quantum dots via RAFT-mediated synthesis of imidazole-based random copolymer ligand

Article abstract of DOI:10.1021/ja908137d

We present a new class of polymeric ligands for quantum dot (QD) water solubilization to yield biocompatible and derivatizable QDs with compact size (10-12 nm diameter), high quantum yields (>50%), excellent stability across a large pH range (pH 5-10.5), and low nonspecific binding. To address the fundamental problem of thiol instability in traditional ligand exchange systems, the polymers here employ a stable multidentate imidazole binding motif to the QD surface. The polymers are synthesized via reversible addition-fragmentation chain transfer-mediated polymerization to produce molecular weight controlled monodisperse random copolymers from three types of monomers that feature imidazole groups for QD binding, polyethylene glycol (PEG) groups for water solubilization, and either primary amines or biotin groups for derivatization. The polymer architecture can be tuned by the monomer ratios to yield aqueous QDs with targeted surface functionalities. By incorporating amino-PEG monomers, we demonstrate covalent conjugation of a dye to form a highly efficient QD-dye energy transfer pair as well as covalent conjugation to streptavidin for high-affinity single molecule imaging of biotinylated receptors on live cells with minimal nonspecific binding. The small size and low serum binding of these polymer-coated QDs also allow us to demonstrate their utility for in vivo imaging of the tumor microenvironment in live mice.

Full text of DOI:10.1021/ja908137d

Published on Web 12/21/2009
Compact Biocompatible Quantum Dots via RAFT-Mediated
Synthesis of Imidazole-Based Random Copolymer Ligand
Wenhao Liu,† Andrew B. Greytak,† Jungmin Lee,† Cliff R. Wong,† Jongnam Park,†
Lisa F. Marshall,† Wen Jiang,‡ Peter N. Curtin,† Alice Y. Ting,† Daniel G. Nocera,†
Dai Fukumura,‡ Rakesh K. Jain,‡ and Moungi G. Bawendi*,†
Department of Chemistry, Massachusetts Institute of Technology, 77 Massachusetts AVenue,
Cambridge Massachusetts 02139-4307 and Edwin L. Steele Laboratory, Department of
Radiation Oncology, Massachusetts General Hospital and HarVard Medical School,
Boston, Massachusetts 02114
Received September 24, 2009; E-mail: mgb@mit.edu
Abstract: We present a new class of polymeric ligands for quantum dot (QD) water solubilization to yield
biocompatible and derivatizable QDs with compact size (∼10-12 nm diameter), high quantum yields
(>50%), excellent stability across a large pH range (pH 5-10.5), and low nonspecific binding. To address
the fundamental problem of thiol instability in traditional ligand exchange systems, the polymers here employ
a stable multidentate imidazole binding motif to the QD surface. The polymers are synthesized via reversible
addition-fragmentation chain transfer-mediated polymerization to produce molecular weight controlled
monodisperse random copolymers from three types of monomers that feature imidazole groups for QD
binding, polyethylene glycol (PEG) groups for water solubilization, and either primary amines or biotin groups
for derivatization. The polymer architecture can be tuned by the monomer ratios to yield aqueous QDs
with targeted surface functionalities. By incorporating amino-PEG monomers, we demonstrate covalent
conjugation of a dye to form a highly efficient QD-dye energy transfer pair as well as covalent conjugation
to streptavidin for high-affinity single molecule imaging of biotinylated receptors on live cells with minimal
nonspecific binding. The small size and low serum binding of these polymer-coated QDs also allow us to
demonstrate their utility for in vivo imaging of the tumor microenvironment in live mice.
Introduction
of QDs has been the presence of a trade-off among five desirable
QD properties for fluorescence labeling in live cells and in vivo:
Quantum dots (QDs) are a powerful class of fluorophores
exhibiting high quantum yields (QY), large molar extinction
coefficients, exceptional photostability, and tunable emission
wavelengths across the visible and near-IR spectral window.1-4
These properties make QDs attractive candidates as biological
fluorescent tags,3,5,6 especially since their brightness and stability
enables single molecule tracking over extended periods of
time.7-9 However, a major barrier toward the widespread use
small size, high stability (both over time and in a wide pH
range), high QY, facile derivatizabilty, and low nonspecific
binding. While it has been possible to achieve three or four of
these criteria, achieving all five simultaneously has proven
challenging for ligand design. Commercial QDs encapsulated
with PEGylated amphiphilic polymer coatings are easily de-
rivatizable and are suitable for single molecule imaging,7,10 but
suffer from large hydrodynamic diameters (20-30 nm),11 which
can limit the access of QDs to crowded regions such as the
neuronal synapse, as well as potentially alter the native behavior
of labeled receptors.12-14 Smaller QDs have been achieved via
ligand exchange with thiol-bearing molecules, but suffer from
instability due to the weak interaction of monothiols with the
QD surface.15,16 Recently, our group and others17-20 have
† Department of Chemistry, Massachusetts Institute of Technology.
‡ Edwin L. Steele Laboratory, Department of Radiation Oncology,
Massachusetts General Hospital and Harvard Medical School.
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B 1997, 101, 9463–9475.
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Bawendi, M. G. J. Am. Chem. Soc. 2006, 128, 2526–2527.
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10.1021/ja908137d  2010 American Chemical Society

Compact Biocompatible Quantum Dots via RAFT
A R T I C L E S
demonstrated increased interaction of thiol ligands with QDs
by exploiting the bidentate binding motif using dihydrolipoic
acid (DHLA) to furnish a new class of aqueous DHLA-PEG
modified QDs that are compact, biocompatible, derivatizable,
and exhibit very low nonspecific binding. Notwithstanding, most
thiol-based coordinating ligands are inherently unstable owing
to the oxidation and dimerization of the thiol groups, causing
the ligands to detach from the QD surface over time (vide infra).
We now report a new class of multidentate poly imidazole
ligands that obviate the need for thiols while maintaining the
attractive properties of small size, low nonspecific binding,
derivatizability, and high QY. Here, we use the term multiden-
tate to indicate a polymer with pendant groups that can bind to
multiple sites on the QD surface, as opposed to multiple
coordination of individual surface atoms, which has been posited
in the case of poly histidine coordination,21 but is not necessarily
apt to the present case.
Poly histidine motifs have been shown to exhibit high affinity
toward the Cd and Zn rich QD surface, and His6-tags have been
employed for facile and efficient derivatization of QDs with
short peptides, dyes, and proteins.21-24 Pendant imidazole
groups in copolymer microgels have also been shown to stabilize
organic soluble QDs.25 From these observations, we hypoth-
esized that a polymer that is rich in imidazole groups along the
backbone should efficiently bind to the QD surface. Poly
imidazole is resistant to degradation by oxidation and its
multidentate binding motif can greatly enhance stability.26-28
To promote water solubility, prevent aggregation, and reduce
nonspecific binding,29 we opted to pursue a strategy to
copolymerize a PEG-derived monomer, and described herein
is the synthesis of a random brush copolymer architecture
displaying both PEG and imidazole groups along the polymer
backbone. Adding an additional monomer featuring either amine
or biotin functional groups affords a 3-component multifunc-
tional copolymer to yield QDs that are water-soluble and
derivatizable. Copolymer based ligands with multidentate
pendant binding groups to the QD surface have been previously
reported, but suffered from either limited water solubility,
Figure 1. Instability of DHLA-PEG derived ligands over time. Gel
electrophoresis of DHLA-carboxyPEG QDs (A) after initial ligand exchange
and (B) after 1 week storage at 4 °C in the dark, with increasing titration
of His6-Tagged monovalent streptavidin from left to right showing sharp
and discrete bands in the initially prepared sample, but loss of fidelity after
storage. QDs in (B) also exhibit increased nonspecific binding to cells (data
not shown).
aggregation, low QY, and/or a lack of a functional handle for
further covalent derivatization, or do not demonstrate the low
nonspecific binding to cells or to serum proteins that are required
for the most interesting biological applications.25,26,30-32 In this
report, we seek to extend the previous work by optimizing our
polymer system for biological compatibility as per the previously
outlined desired properties for in vivo and in vitro applications.
In order to achieve molecular weight control and narrow
polydispersity of the proposed copolymer, we employed RAFT
(radical addition-fragmentation chain transfer) polymerization
chemistry, which offers the further ability to mediate the
controlled copolymerization of a wide library of monomers.33
By varying the ratio and composition of monomers, complex
copolymers can be assembled to give compact, water-soluble
QDs with tunable surface properties. QDs prepared with these
new ligands exhibit extremely low nonspecific binding to serum
and greatly enhanced stability and long-term shelf life, making
them optimal for live cell and in vivo imaging.
Results and Discussion
The design and synthesis of multidentate polymeric imidazole
ligands (PILs) was undertaken to overcome the long-term
instability of DHLA based ligands. Figure 1 shows the gel
electrophoresis of various DHLA based carboxyPEG (Figure
S1 of the Supporting Information) coated QDs. When initially
prepared, DHLA-carboxyPEG QDs exhibit sharp bands by gel
electrophoresis, and a titration of increasing amounts of His6-
tagged monovalent streptavidin (mSA) produced discrete
bands.22 As reported, high-affinity cell labeling with low
nonspecific binding is achieved upon prompt use of the QD-
mSA conjugates. However, the same conjugation experiment
conducted on DHLA-carboxyPEG QDs stored for ∼1 week in
the dark at 4 °C exhibits broadened bands, indicating a
fluctuation in charge owing to a heterogeneous distribution of
ligand density, most likely due to ligand detachment from the
QD surface. Along with band broadening by gel electrophoresis,
we observed a concomitant increase in nonspecific binding to
HeLa cells (data not shown), consistent with loss of ligand
binding to the QDs. These deficiencies are ameliorated by the
replacement of the thiol ligand with that of a polymeric
imidazole ligand described below.
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A R T I C L E S
Liu et al.
Scheme 1
Monomer Synthesis and Polymerization. Scheme 1 presents
the synthetic strategy that was pursued to deliver the monomers
for polymerization. Acrylic acid is coupled to primary amine
bearing moieties via an amide bond forming reaction. Conjugate
addition to the vinyl group was minimized by first preparing
the NHS-ester of acrylic acid (2), and allowing the coupling
reaction to proceed at 4 °C for only 30 min, upon which
complete consumption of the starting materials was confirmed
by TLC. Monomer 3 containing the imidazole group for QD
binding was obtained from the reaction of histamine with 2.
Since the trithiocarbonate RAFT agent 12 used in subsequent
polymerization reactions34 is highly sensitive toward degradation
by aminolysis, the imidazole nitrogen was BOC protected to
yield the final monomer 4. Likewise, monomer 6 containing a
PEG11 group for water solubility was obtained first via the
conversion of the terminal hydroxyl group of monomethoxy
PEG to a primary amine 5, followed by reaction with 2.
Monomers 8 and 9 with BOC-protected terminal amines were
also synthesized in order to afford polymers bearing primary
amine groups for derivatization with FRET dyes or proteins.
Finally, to demonstrate the scope of monomer incorporation in
the polymerization reaction scheme, monomer 11 was synthe-
sized to give polymers functionalized with biotin for binding
assays. For each of the monomers, formation of an amide bond,
as opposed to an ester bond, was necessary in order to
circumvent the possibility of ester hydrolysis under the acidic
BOC deprotection conditions following polymerization, and also
to afford higher stability in the presence of hydrolytic enzymes
for in vivo and in vitro applications.
The reaction of monomers 4, 6, and 8 shown in Scheme 2 is
representative of all polymerization reactions used in this study.
The monomer mixture typically consists of 50% mole fraction
of monomer 4 to ensure that there are enough imidazole groups
for effective binding to the QD surface, with the remaining 50%
mole fraction consisting of some mixture of monomer 6 (for
water solubility), along with one of monomers 8, 9, or 11 (for
derivatizability). In order to minimize the potential of long
polymer chains cross-linking and aggregating QDs during ligand
exchange, the polymer MW was kept low, with a targeted degree
of polymerization (DP) below 30. RAFT polymerization can
be carried out with any number of conventional radical initiators.
These are typically thermally activated and include AIBN,
azobis(2-cyanopentanoic acid), and K2S2O8.35 In our case, AIBN
was used as the initiator in the presence of the RAFT agent 12
to afford a controlled living polymerization.34 Using an [AIBN]:
[RAFT]:[Monomer] ratio of 0.25:1:28 and an equimolar mixture
of monomers 4 and 6 (polymer 13a), the overall polymer
1
conversion over time as monitored by H NMR spectroscopy
(Figure S2 of the Supporting Information) followed a linear
relationship, achieving >80% conversion after 10 h (Figure S3A
of the Supporting Information). Increasing the [AIBN]:[RAFT]
ratio to 1:1 resulted in a nonlinear conversion efficiency versus
time, yielding >90% conversion after 2 h (Figure S3B of the
Supporting Information); interestingly, a low PDI and good
molecular weight control was maintained (vide infra). The plot
of [Monomer]:[RAFT] ratio versus measured polymer DP using
GPC followed a linear relationship, showing good MW control
(Figure 2A), and a narrow PDI < 1.2 (Figure 2B) was observed.
(34) Naoto, A.; Bungo, O.; Hideharu, M.; Takeshi, E. Syn. Lett. 2006, 4,
(35) Barner-Kowollik, C. Handbook of RAFT Polymerization; Wiley-VCH:
636–638.
New York, 2008.
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Compact Biocompatible Quantum Dots via RAFT
A R T I C L E S
Scheme 2
Figure 2A shows that the targeted DP of <30 for copolymer
13a, as measured by GPC, was achieved using [Monomer]:
[RAFT] ratios between 20 to 33:1. The same polymerization
carried out in the absence of RAFT agent yielded a poorly
controlled polymer (PDI > 3), confirming that the RAFT agent
is responsible for mediating polymerization even under high
relative initiator concentrations (Figure 2B). Because controlled
polymerizations could be performed even under the relatively
high [AIBN]/[RAFT] ratio of 1:1, new polymers could be
rapidly prototyped due to the short reaction times required for
near-complete polymer conversion.
GPC calibrated using polystyrene MW standards. Since the
relative reaction rates of the monomers within the copolymer-
ization mixture could not be determined due to overlapping
NMR signals of the monomers, polymerizations were performed
to >90% conversion efficiency to ensure the incorporation of
all monomers in the mixture. It is not known whether the
monomers are incorporated in a statistical fashion, or if there
is some local ordering of monomer units in the polymer
microstructure. An advantage of the class of polymers presented
here for QD ligand exchange is that these ligands are amenable
to long-term storage under ambient conditions without special
precautions against degradation, unlike DHLA based ligands,
which slowly oxidize over time and often need to be carefully
stored in the dark at 4 °C.
The nomenclature of copolymer ligands used in this study is
outlined in Table 1. For QD water solubilization, all polymer
MWs were typically ∼14 kDa with PDI < 1.2 as measured by
Ligand Exchange and Characterization of Aqueous QDs.
Ligand exchange of 605 nm emitting CdSe(CdZnS) core(shell)
QDs20 using poly(PEG)-PILs was performed by displacing the
native hydrophobic ligands with the imidazole groups along the
polymer backbone, which can participate in multiple binding
interactions with the Cd and Zn rich QD surface. Ligand
exchange conditions were relatively mild, and involved stirring
a mixture of QDs and poly(PEG)-PILs in a solution of
chloroform at RT, followed by addition of methanol and
precipitation using chloroform and hexanes. The QDs were
dispersed in water and then purified by dialysis. Complete ligand
displacement was achieved within 1 h, and was confirmed by
1H NMR spectroscopy (Figure S4 of the Supporting Informa-
Table 1. Nomenclature of Compounds Used in This Work
nomenclature for PILs
polymer ligand composition by mole %
poly(PEG)
50% cmpd 6/50% cmpd 4
poly(aminoPEG3)10%
poly(aminoPEG11)25%
poly(biotinPEG)25%
nomenclature for
DHLA-based ligandsa
hydroxyPEG
10% cmpd 8/40% cmpd 6/50% cmpd 4
25% cmpd 9/ 25% cmpd 6/ 50% cmpd 4
25% cmpd 11/ 25% cmpd 6/ 50% cmpd 4
DHLA-PEG8-OH
DHLA-PEG8-COOH
DHLA-PEG8-NH2
Figure 2. RAFT polymerization of 13a showing (A) tunable polymer DP
as a function of [Monomer] to [RAFT] ratio. (B) GPC of poly(PEG)-PIL
in DMF, showing narrow polydispersity with a [Monomer]:[RAFT] ratio
of 30:1 and [AIBN]:[RAFT] ratio of 1:1 (s, black), and poor MW
distribution without RAFT agent (---, red).
carboxyPEG
aminoPEG
a Chemical structures can be found in Figure S1 of the Supporting
Information.
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A R T I C L E S
Liu et al.
Figure 4. Size analysis of poly(PEG) QDs. (A) TEM of QDs ligand
exchanged with polyPEG, showing nonaggregated monodisperse samples.
(B) Dynamic light scattering measurement of QDs ligand exchanged with
hydroxyPEG (s, black) and QDs ligand exchanged with polyPEG (---, red),
both showing an HD of ∼11.5 nm.
Figure 3. (A) Absorption spectra (s, black) and emission spectra of QDs
before ligand exchange in octane (---, blue), and after ligand exchange in
PBS (s, red), showing a slight decrease in fluorescence intensity with a
final QY in water of >60%. (B) Stability of polyPEG QDs in various pH
buffers after incubation at RT for 4 h. pH 10.5* refers to emission intensity
after 5 min irradiation under 365 nm UV light, showing ∼20% photo-
brightening.
QDs drops rapidly, but the QDs remain well dispersed in
solution without any visible formation of macroscopic aggrega-
tion. In addition, a photobrightening effect was observed with
poly(PEG)-PIL QDs. Photoannealing by illumination with 365
nm UV for 5 min increased the fluorescence intensity of
poly(PEG)-PIL QDs by as much as 30%. Furthermore, this
increase in fluorescence was retained for at least 24 h after the
photoannealing treatment. This observation is consistent with
previous studies of QD photobrightening upon conjugation with
His6-tagged proteins,39 as the polymer in this case is binding to
the QD surface via the same imidazole moiety. TEM analysis
of QDs dropcast from water after ligand exchange with
poly(PEG)-PILs shows that the QDs are well dispersed (Figure
4A), and dynamic light scattering analysis shows a single
narrowly distributed population centered at ∼11.5 nm hydro-
dynamic diameter for 605 nm emitting CdSe(ZnCdS) QDs
(Figure 4B). This size is comparable to the hydrodynamic
diameter of the same QDs ligand exchanged with DHLA-
hydroxyPEG, likely due to the compact nature of the imidazole
binding group, which is only a few carbons away from the
polymer backbone. In addition, we observed that the multiden-
tate binding motif employed in poly(PEG)-PILs results in an
aqueous solution that is far more stable than DHLA-hydrox-
yPEG QDs. For instance, dilute DHLA-hydroxyPEG QDs and
poly(PEG)-PIL QDs (<100 nM) were stored at RT under
ambient room lighting. Both samples had been dialyzed to
remove excess ligand in the solution. Within 15 h, the DHLA-
hydroxyPEG QDs precipitated from the solution presumably
due to photooxidation of the dithiol group, while poly(PEG)-
PIL coated QDs remained stable under ambient conditions for
at least 2 months (Figure S5D of the Supporting Information).
tion), which signaled the disappearance of the original aliphatic
protons of TOP/TOPO coated QDs after ligand exchange with
poly(PEG)-PILs, and the appearance of PEG protons from the
polymer bound to the QD surface. A control experiment using
a polymer of 6 alone did not yield water-soluble QDs (data not
shown). The emission peak of QDs after ligand-exchange with
PILs exhibited a slight red-shift, while the line-width remained
fairly constant (Table S1 of the Supporting Information).
Ligand exchange of CdSe(CdS) core(shell) QDs with
poly(PEG)-PILs also proceeded smoothly. The presence of a
high quality CdS shell applied using a modified selective ion
layer adsorption and reaction (SILAR) approach36,37 contributes
significantly to the absorbance cross-section of QDs blue of 450
nm and is advantageous for increased brightness in single
molecule imaging applications. In addition, a robust shell can
greatly improve the QY of QDs after phase transfer to water.
Indeed, the QY of CdSe(CdS) QDs ligand exchanged with
poly(PEG)-PILs were in excess of 65% in water, a modest drop
from a QY of 90% in octane (Figure 3A). The high QY of
poly(PEG)-PIL QDs is maintained in buffers ranging from pH
5 to pH 10.5 after incubation at room temperature for 4 h (Figure
3B). Previous studies on the pH dependence of His6-tag binding
to Ni-NTA media show that the interaction is stable between
pH 7-11,21,38 and becomes disrupted below pH 5 due to the
protonation of the imidazole group. This is consistent with the
binding of poly(PEG)-PILs to the surface of QDs via metal-
affinity interactions. Below pH 5, the fluorescence intensity of
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Compact Biocompatible Quantum Dots via RAFT
A R T I C L E S
Three-Component Random Copolymers for Function-
alized Biocompatible QDs. With robustness and compactness
of poly(PEG)-PIL coated QDs established, we turned our
attention to incorporating chemical functionality into the
polymer so that the QDs can be subsequently derivatized for
targeting and sensing applications. Multifunctional polymers can
be synthesized from monomers 4 and 6, along with one of
monomers 8, 9, or 11, to give functionalized QDs with primary
amine or biotin groups upon water solubilization. Copolymer-
ization with three monomers proceeded smoothly, with good
size control and low polydispersity (Figure S6 of the Supporting
Information), and functional monomers were incorporated at
mole fractions ranging from 10-25%. As mentioned previously,
the incorporation of all monomers was maximized by running
the polymerization to >90% conversion, as overlapping 1H
signals made it difficult to determine the final polymer composi-
tion by NMR. Using the short aminoPEG3 monomer 8, polymers
with amine functionalities were synthesized with the functional
group close to the surface of the QD after ligand exchange,
making this system ideal for dye conjugation and energy transfer
sensing applications, as detailed below. Polymers with amine
functionalities tethered by longer linkers were synthesized using
the aminoPEG11 monomer 9. These polymers were found to be
more suitable for the conjugation of QDs to larger biomolecules
such as proteins and antibodies, as the longer PEG11 linker
makes the terminal amino groups more sterically accessible.
Using the monomer 11, polymers bearing a biotin functionality
were also synthesized, giving water-soluble QDs that bind
readily to streptavidin coated plates (Figure S7 of the Supporting
Information). The synthesis of functional three-component
copolymers affords a versatile and facile way of tuning the QD
surface functionality for a wide range of potential applications.
Conjugation to an Energy Transfer Dye. A salient feature
of the copolymer ligand strategy is the ability to incorporate
side-chains that can be readily derivatized in aqueous solution.
To demonstrate covalent derivatization, we conjugated 5-car-
boxy-X-rhodamine (ROX), a red-emitting fluorescent dye, to
QDs ligand exchanged with poly(aminoPEG3)10%-PILs via an
amide linkage. The dye absorbance offers a convenient indica-
tion of the coupling yield. Additionally, the typical radial
distance from the QD to the derivatization site may be
determined from a Fo¨rster resonance energy transfer (FRET)
analysis of the QD-dye pair.40,41
Figure 5. Covalent derivatization of poly(aminoPEG3)10% with ROX dye
molecules. (A) Absorption spectrum of purified conjugate (s, black), least-
squares fit of conjugate spectrum (- ·-·-, blue), as sum of QD (---, green),
and free ROX (• • • •, red) contributions. (B) Photoluminescence spectrum
of conjugate (s, black) and of control QD (s, green) and free dye (-·-·-,
red), normalized to reflect the QD and ROX concentrations, respectively,
present in the conjugate sample. Contributions of QD (---, green) and free
ROX (• • • •, red) to conjugate emission spectrum as obtained from a least-
squares fit are also shown. All samples are excited at 450 nm, with dye
emission showing 31 fold enhancement from the free dye vs the conjugate.
absorbance, allowing for the selective excitation of the QD. The
QD PL peak is significantly quenched versus that of a control
sample that was processed similarly but not modified with the
ROX dye, while the ROX emission centered at 610 nm is
significantly enhanced versus that of the free dye when
normalized for the sample concentrations. These observations
are consistent with energy transfer; a fit of the emission spectrum
reveals an energy transfer efficiency of 88%.
The high energy transfer efficiency suggests that the copoly-
mer ligand system is able to poise small-molecule substituents
in proximity to the QD core. Indeed, analysis of the observed
efficiency and spectral overlap according to the Forster model
(Figure S8 of the Supporting Information) suggests a charac-
teristic separation distance of no more than 4.5 nm, which is
consistent with observations of a small hydrodynamic radius
for the ligand-exchanged QDs. The limited coupling yield
observed here may indicate saturation of the available primary
amine binding sites, suggesting that not all amine side-chains
on the polymer are sufficiently accessible from the solvent,
possibly due to the affinity of amines for the QD surface.
In order to gauge the number of free amines that are solution
accessible, we probed the PIL-coated QDs with Fluorescamine,
an amine-reactive fluorogenic probe (Figure S9 of the Support-
ing Information). For QDs coated with poly(aminoPEG3)5%, the
average number of measured free amines exposed to solution
per QD was only ∼1. This low number suggests that most of
the amines are in fact bound to the QD surface and are
To form conjugates, an aliquot of the amine-reactive succin-
imidyl ester of 5-ROX (either obtained commercially, or
prepared from the 5-ROX free acid and N-hydroxysuccinimide
with dicyclohexylcarbodiimide) in dimethyformamide was
added to a solution of the QDs in phosphate buffer at pH 7.6.
Following the coupling reaction, the QDs were separated from
unbound dye and NHS byproduct via size exclusion chroma-
tography. Figure 5 shows the absorption and emission spectra
of a purified QD-ROX conjugate made using 562 nm-emitting
CdSe(CdS) QDs and 27 equiv. of the activated dye. The dye
contribution to the sample absorbance is clearly visible in Figure
5A as a shoulder on the red side of the lowest energy exciton
peak. A fit of the spectrum as a sum of QD and dye components
reveals an average dye:QD ratio of 1.78:1. Figure 5B shows
the photoluminescence (PL) spectrum of the conjugate under
450 nm excitation. At this wavelength, ROX dye has minimal
(40) Snee, P. T.; Somers, R. C.; Nair, G.; Zimmer, J. P.; Bawendi, M. G.;
Nocera, D. G. J. Am. Chem. Soc. 2006, 128, 13320–13321.
(41) Somers, R. C.; Bawendi, M. G.; Nocera, D. G. Chem. Soc. ReV. 2007,
36, 579–591.
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A R T I C L E S
Liu et al.
Figure 6. Nonspecific binding of QDs on HeLa cells as a function of ligand coating, with incubation at 500 nM QD concentration for 5 min at 4 °C,
followed by 4× wash with PBS buffer before imaging. Top: QD fluorescence at 565 nm with excitation at 488 nm. All images are scaled to the same
contrast with the exception of (B) and (C), for which the left section has the same contrast as the other images, while the contrast has been boosted in the
right section to highlight the difference between (B) and (C). Bottom: corresponding DIC image. QDs were ligand exchanged with (A) DHLA, (B) hydroxyPEG,
(C) poly(PEG), (D) poly(aminoPEG3)10%, and (E) poly(aminoPEG11)25%
.
inaccessible to the solvent. Increasing the mole fraction of
compound 8 from 5% to 20% (i.e., poly(aminoPEG3)20%
surface. When amine groups are introduced in a three-
component polymer (Figure 6D,E), we observe increasing levels
of nonspecific binding. Poly(aminoPEG3)10%-PIL QDs show
slightly more nonspecific binding versus DHLA-hydroxyPEG
QDs, and poly(aminoPEG11)25%-PIL QDs exhibit even more
nonspecific binding, consistent with an increase in the amount
of amines on the QD surface. Although the nonspecific binding
for poly(aminoPEG11)25%-PIL QDs appears to be nontrivial, the
incubation for this particular experiment was performed at high
QD concentration (500 nM). We demonstrate below that the
)
increased the average amine:QD ratio to ∼8:1. Indeed, using
poly(aminoPEG3)25% coated QDs for dye conjugation yielded
∼4 dyes/QD (Figure S10 of the Supporting Information).
Taken together, these results show that although some degree
of amine binding is present, the incorporation of compound 8
into the polymer as a functional site for derivatization proved
to be successful, and these initial results bode well for the
derivatization of these QDs with dyes and other small molecules
for applications in targeted biological imaging and sensing.
covalent conjugation of streptavidin to poly(aminoPEG11)25%
-
PIL QDs and their subsequent targeting to cells at low QD
concentrations resulted in labeling with good signal-to-noise and
allowed for single molecule tracking of QDs with minimal
background.
Conjugation to Streptavidin for Specific Targeting. In order
to demonstrate the capability of PIL-based QDs for targeted
Nonspecific Binding to HeLa Cells. An absence of nonspecific
binding is essential to reliable targeting and/or sensing applica-
tions involving QDs. To test the nonspecific binding of
poly(PEG)-PIL based QDs, we incubated HeLa cells with QDs
of various surface compositions and subsequently washed the
cells 4× with phosphate-buffered saline (PBS). Since the level
of nonspecific binding is inherently low for such PEGylated
QDs, the cells were incubated at high QD concentrations (∼500
nM) in order to highlight the differences between the coatings.
Fluorescence and phase contrast images of cells after washing
are shown in Figure 6. As expected, DHLA coated QDs
produced a high degree of nonspecific binding to both cells and
glass.22 DHLA-hydroxyPEG coated QDs exhibited minimal
nonspecific binding, but some QD stickiness could be observed
with enhanced contrast of the fluorescence images. Poly(PEG)-
PIL QDs, when viewed under the same enhanced contrast, show
virtually no nonspecific binding, which may be attributed to
several factors. First, the PEG chains of poly(PEG)-PILs are
terminated in methoxy groups, which can further reduce
nonspecific binding versus PEG terminated with hydroxyl
groups.29,42 Second, the PEG length of poly(PEG)-PILs is
slightly longer versus that of DHLA-hydroxyPEG, and third,
the methoxyPEG along the polymer backbone may offer a
denser coverage of PEG groups to better passivate the QD
single molecule imaging in live cells, poly(aminoPEG11)25%
-
PIL QDs were conjugated to streptavidin (SA) via 1-ethyl-3-
(3- dimethylaminopropyl) carbodiimide (EDC) coupling chem-
istry. For targeting, HeLa cells were transfected with a plasmid
for yellow fluorescent protein (YFP), fused to an extracellular
acceptor peptide (AP) tag13 and a transmembrane domain (TM)
for cell surface targeting (AP-YFP-TM), as well as with a
plasmid for endoplasmic reticulum-localized biotin ligase
(BirA).22,43 The AP tag is specifically biotinylated by the
coexpressed BirA and displayed on the cell surface along with
YFP via the TM domain. The QD-SA conjugates were then
added for labeling. At a high QD labeling concentration of 100
nM, excellent colocalization was observed between the YFP
and QD channels, with very low levels of nonspecific binding
to nontransfected cells and glass (Figure 7A). A control
experiment in which the QD-SA construct was preincubated
with biotin showed no binding, confirming that the binding
interaction was indeed between the QD-SA and biotin on the
cell surface (Figure 7B). By reducing the QD labeling concen-
(42) Mei, B. C.; Susumu, K.; Medintz, I. L.; Delehanty, J. B.; Mountziaris,
T. J.; Mattoussi, H. J. Mater. Chem. 2008, 18, 4949–4958.
(43) Howarth, M.; Ting, A. Y. Nat. Protoc. 2008, 3, 534–545.
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Compact Biocompatible Quantum Dots via RAFT
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the relatively short PEG8 chains of DHLA-hydroxyPEG are less
able to provide full surface passivation. By contrast, QDs coated
with poly(PEG)-PIL exhibited negligible nonspecific binding
to serum proteins after incubation (Figure 8B), as indicated by
the appearance of the peak position at the same position as the
control. We surmise that the PEG units along the polymer
backbone provide a denser coverage of PEG on the surface of
the QDs, thus giving it enhanced antibiofouling properties.
Functionalized QDs ligand exchanged with poly(aminoPEG3)10%
-
PIL also exhibited no detectable nonspecific binding in serum
(Figure 8C).
Probing the Tumor Microenvironment Using Poly(PEG)-PIL
QDs as a Diffusion Tracer. Taking advantage of the low serum
binding of poly(PEG)-PIL QDs, we observed the distribution
dynamics of these QDs within tumors in live mice (Figure 9).
Using a breast tumor model grown beneath transparent windows
in mice,45-47 vascular transport of QDs after intravenous
injection can be imaged via two-photon laser scanning micros-
copy as a function of time to observe QD distribution kinetics.
The mouse vessel walls were visualized independently via green
fluorescent protein (GFP) expressed under the promoter of the
Tie2 receptor present on the surface of the vascular endothelial
cells.44 CdSe(CdS) poly(PEG)-PILs QDs emitting at 605 nm
were injected retro-orbitally, and the tumor vasculature was
imaged over 6 h. Initially, the QDs are confined within the vessel
lumen (Figure 9A). At 3 h, we observed clearance from the
vasculature and the simultaneous appearance of QDs in the
tumor tissue. After 6 h, the QDs had extravasated into the tumor
tissue, appearing as a uniform signal spread throughout the field
of view (Figure 9A). The QDs appeared stable, and there was
no indication of stickiness or aggregation on the lumen wall or
in the tumor tissues. Having a stable, small, biocompatible QD
scaffold should enable the use of these QDs as robust in vivo
sensors.41 The scope of the PIL system can also be extended to
water-solubilize InAs(CdZnS) QDs that emit in the near-IR as
high contrast probes with enhanced depth in vivo.48
Figure 7. Targeting of poly(aminoPEG11)25% QD-SA conjugates to live
HeLa cells transfected with AP-YFP-TM. (A) Ensemble labeling with 100
nM QDs. (B) Same as in (A), but with QDs preincubated with biotin. (C)
Low-density labeling with 10 nM QD-SA reveals single QDs.
tration to 10 nM, single QDs could be readily observed (Figure
7C), as identified by their fluorescence intermittency behavior
(Figure S11 of the Supporting Information). As previously
discussed, DHLA-PEG based QDs are subject to degradation
via loss of ligand coating after ∼1 week and become increas-
ingly sticky to cells. By contrast, in our experience, PIL-based
QDs remain nonsticky and functional on the time scale of
months.
Conclusions
We have synthesized a new class of coordinating polymers
that produce aqueous QDs with greater stability and shelf life
compared with previously reported DHLA-derived ligands,
while maintaining the desirable QD properties of low nonspecific
binding, small size, facile derivatizability, and high QY. By
using a three-monomer copolymerization scheme, we were able
to produce multifunctional aqueous QDs featuring imidazole
groups for multidentate binding to the QD surface, PEG groups
for water solubility and mitigation of nonspecific binding, and
either amine groups or biotin groups on the surface for
derivatization. The monomer synthesis is facile and can be
scaled to multigram quantities, and by utilizing RAFT polym-
erization, a wide variety of monomers were used to produce
Nonspecific Binding in Serum Proteins for in Vivo
Applications. PEG11 monomer 6 is always present in a signifi-
cant mole fraction within the polymer not only to provide water
solubility, but also to mitigate nonspecific binding and prevent
biofouling of QDs. To illustrate the stability of these QDs for
in vivo applications, we incubated various polymer-coated
CdSe(CdZnS) 565-nm emitting QDs with mouse bovine serum
at 37 °C for 4 h and analyzed the samples by size exclusion
chromatography with fluorescence detection to determine the
extent of nonspecific binding to serum protein. In the case of
QDs coated with DHLA-hydroxyPEG, significant serum protein
binding was observed, as indicated by the formation of a large
broad peak eluting at earlier times after serum incubation versus
control (Figure 8A). Previous serum binding experiments using
DHLA-hydroxyPEG on InAs(ZnSe) QDs revealed low levels
of nonspecific binding to serum protein.2 The discrepancy is
likely due to the relative size of the QD cores. In the InAs(ZnSe)
case, the inorganic cores were on the order of 2 nm in diameter,
and the PEG8 group of DHLA-hydroxyPEG provided sufficient
passivation against nonspecific binding. The CdSe(CdZnS) QDs
used in this study are approximately twice the diameter and
(44) Duda, D. G.; Fukumura, D.; Munn, L. L.; Booth, M. F.; Brown, E. B.;
Huang, P. G.; Seed, B.; Jain, R. K. Cancer Res. 2004, 64, 5920–
5924.
(45) Jain, R. K.; Munn, L. L.; Fukumura, D. Nat. ReV. Cancer 2002, 2,
266–276.
(46) Jain, R. K.; Brown, E. B.; Munn, L. L.; Fukumura, D. In LiVe Cell
Imaging: A Laboratory Manual; Cold Spring Harbor Laboratory Press:
Cold Spring Harbor, NY, 2004, p 435-466.
(47) Huang, P.; Dawson, M.; Lanning, R.; Jain, R. K.; Fukumura, D.
Journal of the American Association for Laboratory Animal Science
2008, 47, 170–170.
(48) Allen, P. M.; Liu, W.; Chauhan, V. P.; Lee, J.; Ting, A. Y.; Fukumura,
D.; Jain, R. K.; Bawendi, M. G. Journal of the American Chemical
Society (DOI: 10.1021/ja908250r).
9
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A R T I C L E S
Liu et al.
Figure 8. Nonspecific binding of QDs with serum proteins after incubation with 95% mouse bovine serum for 4 h at 37 °C. QDs before incubation (- - -,
red) and after incubation (s, black) for (A) hydroxyPEG QDs, (B) poly(PEG) QDs, and (C) poly(aminoPEG3)10% QDs. Insets show fluorescence spectrum
of eluent at time indicated by the black arrow, showing QD emission.
Figure 9. (A) Time lapsed live QDs imaging of P008 tumor vasculature in Tie2-GFP/FVB mice. Red fluorescence corresponds to the signal from QDs
within the vessel lumen (0 and 3 h), or extravascular space in tumors (3 and 6 h), while the green fluorescence is from the GFP in vascular endothelial cells
that line the vessel wall. B, Mean QD (red channel) intensities within and outside of the blood vessels over time. Vessel regions are assigned by thresholding
of the T ) 0 h image. The QDs are initially confined within the vessels at T ) 0. Images at later times reveal decreased vascular and increased extravascular
fluorescence, indicative of clearance from the vessels and extravasation into tumor tissue, respectively.
Acrylate compounds bearing terminal vinyl groups were visualized
on TLC using KMnO4. All other TLC plates were visualized by
iodine staining. Flash column chromatography was performed on
aqueous QDs with controllable surface properties and composi-
tions. The enhanced QD stability enabled by these new polymers
is crucial for cellular/in vivo targeting and imaging/sensing
applications in which the QDs must survive conjugation with
dyes or proteins and subsequent purification steps in order to
arrive at functional probes. Our poly imidazole binding motif
relieves the necessity to perform these steps in prompt succes-
sion. Often, weeks or months of time are needed to properly
culture cells or raise animals for QD imaging studies, and these
new QDs enable the long-term storage of functional sensing/
targeting constructs for such studies. Furthermore, we demon-
strate the utility of small nonsticky poly(PEG)-PIL QDs for
achieving extravasation from the tumor vasculature in mice with
uniform distribution, paving the way for studies of the tumor
microenvironment. The modularity of the PIL system can
potentially accommodate an even wider diversity of monomers,
expanding the scope of functionalities achievable on QD
surfaces well beyond the examples provided in this study.
1
a Teledyne Isco CombiFlash Companion. H NMR spectra were
recorded on a Bruker DRX 401 NMR Spectrometer. UV-vis
absorbance spectra were taken using an HP 8453 diode array
spectrophotometer. Photoluminescence and absorbance spectra were
recorded with a BioTek Synergy 4 Microplate Reader. Dynamic
light scattering analysis was performed on a Malvern Instruments
ZetaSizer ZS90 in a low volume 12 µL quartz cuvette, with QD
concentrations between 1-3 µM. Polymer molecular weights were
determined in DMF solutions on an Agilent 1100 series HPLC/
GPC system with three PLgel columns (103, 104, 105 Å) in series
against narrow polystyrene standards.
Compound 2. To a stirred solution of acrylic acid (1.00 g, 13.88
mmol) and N-hydroxysuccinimide (NHS) (1.91 g, 16.65 mmol) in
40 mL of dry THF was added dropwise a solution of dicyclohexy-
lcarbodiimide (DCC) (3.43 g, 16.65 mmol) in 10 mL dry THF with
stirring at 4 °C. The solution was warmed to room temperature
and stirred for 2 h. Precipitates were removed by filtration, and the
solvent was evaporated in vacuo. Ethylacetate (50 mL) was added
to facilitate further precipitation of reaction byproducts, and the
solution was filtered once more. The solvent was evaporated and
the product dissolved in either 10 mL of anhydrous DMF or dry
THF to create a stock solution, which was used in later reaction
steps without further purification.
Experimental Section
Materials and Instrumentation. All chemicals unless indicated
were obtained from Sigma Aldrich and used as received. Air
sensitive materials were handled in an Omni-Lab VAC glovebox
under dry nitrogen atmosphere with oxygen levels <0.2 ppm. All
solvents were spectrophotometric grade and purchased from EMD
Biosciences. Amine-bearing compounds were visualized on thin
layer chromatography (TLC) plates using a ninhydrin solution.
Compound 4. To an aqueous solution of sodium bicarbonate
(50 mL, 0.3 M) was added DMF (50 mL) and histamine dihydro-
chloride (2.50 g, 13.59 mmol). To this solution was added
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480 J. AM. CHEM. SOC. VOL. 132, NO. 2, 2010

Compact Biocompatible Quantum Dots via RAFT
A R T I C L E S
compound 2 (2.75 g, 16.3 mmol) in a solution of DMF, with stirring
at 4 °C. The reaction was monitored via TLC by ninhydrin stain
for primary amines, and confirmed to be complete after 30 min to
give the crude compound 3. The solvent was removed in vacuo,
and the product redissolved in DMF (50 mL). The solution was
filtered, and triethylamine was introduced (2.27 mL, 16.30 mmol).
Ditert-butyl dicarbonate was added dropwise at 4 °C, and the
solution was stirred overnight at RT. Water was added and the
solution extracted with CHCl3 (3 × 25 mL). The organics were
combined and dried over sodium sulfate, and the solvent removed
in vacuo. The crude product was purified by silica column (ethyl
acetate/hexanes gradient 50:50 to 100:0, v/v) to give the pure
and dried over sodium sulfate, and the solvent removed in vacuo.
The crude product was purified by silica column (ethyl acetate/
methanol gradient 100:0 to 95:5, v/v) to give the pure product as
1
a clear oil (4.52 g, 27% yield). H NMR (400 MHz, CDCl3): δ
(ppm) 6.23 (dd, J1 ) 2.0 Hz, J2 ) 17.0 Hz, 1H), 6.08 (dd, J1 ) 9.8
Hz, J2 ) 17.0 Hz, 1H), 5.56 (dd, J1 ) 2.0 Hz, J2 ) 9.8 Hz, 1H),
3.65-3.44 (m, 14H), 3.17 (t, 2H), 1.83-1.65 (m, 4H), 1.39 (s,
12H).
Compound 9. To a solution of O-(2-Aminoethyl)-O′-[2-(Boc-
amino)ethyl]decaethylene glycol (0.50 g, 0.78 mmol) in dry THF
(25 mL) was added triethylamine (0.086 g, 0.85 mmol) and
compound 2 (0.20 g, 1.16 mmol) dropwise in a solution of THF
with stirring at 4 °C. The reaction was monitored via TLC by
ninhydrin stain for primary amines, and confirmed to be complete
after 30 min. The solution was filtered and the solvent removed in
vacuo. The crude product was purified by silica column (DCM/
MeOH gradient 100:0 to 95:5, v/v) to give the pure product as a
clear oil (0.38 g, 70% yield). 1H NMR (400 MHz, CDCl3): δ (ppm)
6.28 (dd, J1 ) 2.0 Hz, J2 ) 17.0 Hz, 1H), 6.16 (dd, J1 ) 9.8 Hz,
J2 ) 17.0 Hz, 1H), 5.59 (dd, J1 ) 2.0 Hz, J2 ) 9.8 Hz, 1H),
3.70-3.50 (m, 46H), 3.30 (q, 2H), 1.42 (s, 9H).
Compound 10. To a solution of O-(2-Aminoethyl)-O′-[2-(Boc-
amino)ethyl] decaethylene glycol (0.50 g, 0.78 mmol) in DMF (150
mL) was added biotin (0.21 g, 0.86 mmol) and EDC (0.13 g, 0.86
mmol). The solution was stirred overnight, and the solvent removed
in vacuo. The crude product was purified by silica column (DCM/
MeOH 98:2, v/v) to give the pure product as a colorless oil (0.63
g, 85% yield). 1H NMR (400 MHz, CDCl3): δ (ppm) 4.23 (m, 1H),
4.43 (m, 1H), 3.48 (m, 4H), 3.52-3.61 (m, 40H), 3.24 (m, 2H),
3.35 (m, 2H), 2.82 (dd, J1 ) 12.8 Hz, J2 ) 4.9 Hz, 1H), 3.06 (m,
1H), 1.37 (s, 9H), 2.68 (d, J ) 12.8 Hz, 1H), 2.16 (t, J ) 7.5 Hz,
2H), 1.60 (m, 4H), 1.40-1.32 (m, 2H).
Compound 11. To compound 10 (0.50 g, 0.57 mmol) was added
4 M HCl in dioxane, and stirred for 1 h at room temperature. The
solvent was removed in vacuo, and the crude product dissolved
into a solution of 0.25 M aqueous sodium bicarbonate with DMF.
To this solution was added dropwise a solution of compound 2.
The reaction was monitored via TLC by ninhydrin stain for primary
amines and confirmed to be complete after 30 min. The solvent
was removed in vacuo, and the crude product was purified by silica
column chromatography (DCM:MeOH 98:2, v/v) to give the
product as a colorless oil (0.31 g, 65% yield). 1H NMR (400 MHz,
CDCl3): δ (ppm) 6.28 (dd, J1 ) 2.0 Hz, J2 ) 17.0 Hz, 1H), 6.17
(dd, J1 ) 9.8 Hz, J2 ) 17.0 Hz, 1H), 5.61 (dd, J1 ) 2.0 Hz, J2 )
9.8 Hz, 1H), 4.49 (m, 1H), 4.30 (m, 1H), 3.48-3.72 (m, 44H),
3.42 (m, 2H), 3.13 (m, 1H), 2.89 (dd, J1 ) 12.8 Hz, J2 ) 4.9 Hz,
1H), 2.74 (d, J ) 12.8 Hz, 1H), 2.22 (t, J ) 7.4 Hz, 2H), 1.66 (m,
4H), 1.43 (m, 2H).
Typical PIL Polymerization. All monomers were kept as dilute
stock solutions between 30-100 mg/mL in either ethylacetate or
methanol. Stock solutions of RAFT agent 12 were prepared at 220
mg/mL in DMF, and AIBN was prepared at 50 mg/mL in DMF.
All reagents were weighed out volumetrically. In a typical polym-
erization, monomers 4 (33 mg, 0.13 mmol) and 6 (77 mg, 0.13
mmol) were added to an 8 mL vial. The solvent was removed in
vacuo and 50 µL of dry DMF along with RAFT agent 12 (2.53
mg, 0.0088 mmol), and AIBN (1.43 mg, 0.0088 mmol) were added.
The contents of the vial were mixed, centrifuged at 5000 ×g for 2
min, and then transferred to a 1 mL ampule. The ampule was
subjected to 4 cycles of freeze-pump-thaw, and sealed under
vacuum using a butane torch. The vial was heated to 70 °C on an
oil bath for 1.5-3 h, after which 0.5 mL of a 4 M HCl in dioxane
solution was added to cleave the BOC protecting groups. After 1 h
at RT, the HCl was removed in vacuo. The deprotected polymer
was dissolved in MeOH, to which a solution of NaOH in MeOH
(1M) was added dropwise to adjust the pH to be between 8-9.
The solvent was removed in vacuo, and then CHCl3 was added to
precipitate the salts. The solution was filtered through a 0.45 µm
1
product as a clear oil (2.59 g, 72% yield). H NMR (400 MHz,
CDCl3): δ (ppm) 7.95 (s, 1H), 7.10 (s, 1H), 6.19 (dd, J1 ) 1.8 Hz,
J2 ) 17.0 Hz, 1H), 6.07 (dd, J1 ) 9.8 Hz, J2 ) 17.0 Hz, 1H), 5.53
(dd, J1 ) 1.8 Hz, J2 ) 10.0 Hz, 1H), 3.53 (dt, 2H), 2.72 (t, 2H),
1.54 (s, 9H).
Compound 5. Neat methoxy poly(ethylene glycol) (10 g, 18.18
mmol, average MW 550 g/mol) was degassed at 80 °C for 1 h
with stirring to remove traces of water. The flask was backfilled
with N2 and cooled on an ice bath before thionyl chloride (1.98
mL, 27.27 mmol) was slowly added. The solution was warmed to
25 °C and stirred for 2 h. The conversion was monitored by the
disappearance of the broad O-H stretch at 3,500 cm-1 and the
appearance of a C-Cl stretch at 730 cm-1 in the IR spectrum. The
product was diluted with DMF (20 mL) and the solvent removed
under reduced pressure. This was repeated three times to remove
all residual traces of thionyl chloride. The sample was dissolved
in a solution of sodium azide (1.77 g, 27.27 mmol) in 100 mL
DMF and stirred overnight at 85 °C. The solvent was removed
under reduced pressure and 200 mL of dichloromethane was added.
The precipitate was removed by vacuum filtration and the solvent
evaporated in vacuo to yield the intermediate monoazide. The
sample was dissolved in 150 mL of tetrahydrofuran (THF), and
triphenylphosphine (7.15 g, 27.27 mmol) was added. The solution
was stirred at 25 °C for 4 h before adding 1 mL of water and stirring
overnight. The THF was removed in vacuo and 100 mL of water
was added. The precipitate was removed by vacuum filtration and
the filtrate washed with toluene (3 × 50 mL). The water was
removed in vacuo to yield the pure product as light yellow oil (9.67
1
g, 95%). H NMR (400 MHz, CDCl3): δ (ppm) 3.69-3.46 (m,
46H), 3.37 (s, 3H), 2.85 (t, 2H).
Compound 6. To a solution of compound 5 (2.20 g, 3.94 mmol)
in dry THF was added compound 2 (1.00 g, 5.92 mmol) in a
solution of dry THF, with stirring at 4 °C. The reaction was
monitored via TLC by ninhydrin stain for primary amines, and
confirmed to be complete after 30 min. The solution was filtered
and the solvent evaporated in vacuo. The crude product was purified
by silica column (methanol/ethyl acetate gradient 0:100 to 5:95,
v/v) to give the pure product as a pale yellow oil (1.88 g, 78%
yield). 1H NMR (400 MHz, CDCl3): δ (ppm) 6.68, 6.19 (dd, J1 )
2.0 Hz, J2 ) 17.0 Hz, 1H), 6.08 (dd, J1 ) 9.8 Hz, J2 ) 17.0 Hz,
1H), 5.52 (dd, J1 ) 2.0 Hz, J2 ) 9.8 Hz, 1H), 3.56-3.37 (m, 48H),
3.27 (s, 3H).
Compound 8. To a solution of 4,7,10-trioxa-1,13-tridecanedi-
amine (10.00 g, 45.45 mmol) in DCM (25 mL) was added dropwise
ditert-butyl dicarbonate (1.98 g, 9.09 mmol) at 4 °C. The solution
was allowed to warm to RT and stirred overnight. The solution
was washed with water (3 × 20 mL) to remove unreacted starting
material. TLC analysis with ninhydrin staining shows mostly
monosubstituted product in the organic phase. The organics were
dried over sodium sulfate and solvent removed in vacuo. The crude
product (3.80 g, 17.27 mmol) was dissolved in a mixture of aqueous
sodium bicarbonate buffer (20 mL, 0.3 M), and DMF (20 mL), to
which compound 2 (3.38 g, 20.00 mmol) was added dropwise in a
solution of DMF with stirring at 4 °C. The reaction was monitored
via TLC by ninhydrin stain for primary amines, and confirmed to
be complete after 30 min. Water was added and the solution
extracted with CHCl3 (3 × 25 mL). The organics were combined
9
J. AM. CHEM. SOC. VOL. 132, NO. 2, 2010 481

A R T I C L E S
Liu et al.
PTFE filter and the solvent removed in vacuo to yield the final
polymer for QD ligand exchange.
of QDs in PBS and dye in ethanol were optically matched at the
excitation wavelength. Fluorescence spectra of QD and dye were
taken under identical spectrometer conditions in triplicate and
averaged. The optical density was kept below 0.1 between 300-800
nm, and the integrated intensities of the emission spectra, corrected
for differences in index of refraction and concentration, were used
Quantum Dot Synthesis. CdSe cores were synthesized accord-
ing to previously reported procedures,20,49,50 and were overcoated
with either Zn0.8Cd0.2S alloy shells or pure CdS shells. The alloy
shell overcoating procedure has been described previously,20,50 and
was used here to obtain QDs emitting at 565 and 605 nm with
QYs of ∼80% when diluted in hexane. For pure CdS shells, we
developed a successive ion layer adsorption and reaction (SILAR)
procedure that is modified from those reported by Peng et al. and
Mews et al.36,37 Briefly, CdSe cores with a first exciton feature at
491 nm were synthesized by heating a mixture of trioctylphosphine
(TOP), trioctylphosphine oxide (TOPO), CdO (0.9 mmol), and
tetradecylphosphonic acid (TDPA, 2.0 mmol) to 340 °C under
nitrogen, removing evolved water in vacuo at 160 °C, reheating to
360 °C under nitrogen, and rapidly introducing trioctylphosphine
selenide (TOPSe, 3.4 mmol) in trioctylphosphine (TOP), followed
by cooling to room temperature. Cores isolated by repeated
precipitations from hexane with acetone were brought to 180 °C
in a solvent mixture of oleylamine (3 mL) and octadecene (6 mL).
Aliquots of Cd and S precursor solutions were then introduced
alternately starting with the metal (Cd), waiting 15 min between
the start of each addition. The Cd precursor consisted of 0.6 mmol
Cd-oleate and 1.2 mmol decylamine in a solvent mixture of
octadecene (3 mL) and TOP (3 mL). The S precursor consisted of
0.6 mmol hexamethyldisilathiane [(TMS)2S] in 6 mL TOP. The
dose of each overcoating precursor aliquot was calculated to provide
a single monolayer of ions to the QD surface. Addition of a total
of 4 aliquots each of Cd and S yielded QDs with emission at 562
nm and a QY close to unity when diluted in hexane. A similar
procedure was performed on larger CdSe cores to obtain CdSe(CdS)
QDs emitting at 605 nm.
Ligand Exchange with poly(PEG)-PIL. QDs (2 nmol) were
precipitated using MeOH and brought into 50 µL of CHCl3. The
QD stock solution was mixed with solution of poly(PEG) (5 mg)
in CHCl3 (30 µL), and stirred for 10 min at RT, after which 30 µL
of MeOH was added followed by stirring for an additional 20 min.
QD samples were precipitated by the addition of EtOH (30 µL),
CHCl3 (30 µL), and excess hexanes. The sample was centrifuged
at 4000 g for 2 min. The clear supernatant was discarded, and the
pellet dried in vacuo, followed by the addition of PBS (500 µL,
pH 7.4). The aqueous sample was then filtered through a 0.2 µm
filter syringe filter before use.
Fluorescamine Assay of Amines on Surface of PIL-QDs. QDs
emitting at 543 nm ligand-exchanged with various PILs were
purified by dialysis 3× through a 50 kDa MW cutoff spin
concentrator with a regenerated cellulose membrane (Millipore,
Billerica, MA). The QDs were adjusted to 1-2 µM concentration
and placed into an eppendorf tube (240 uL). To the tube was added
a solution of fluorescamine in acetone (10 uL, 28 mg/mL) followed
by vigorous vortexing. The samples were incubated for 10 min at
RT and the photoluminescence intensity of Fluorescamine was
recorded at 480 nm with an excitation at 380 nm. The amine
concentration versus fluorescence count was obtained via a calibra-
tion curve generated by performing the same assay on a serial
dilution of a known concentration of compound 5.
to calculate the QYs using the expression QYQD ) QYDye
×
(Absorbancedye/AbsorbanceQD) × (Peak AreaQD/Peak AreaDye) ×
(nQDsolvent)2/(nDyesolvent)2.51
Gel Filtration Apparatus. GFC was performed using an
AKTAprime Plus chromatography system from Amersham Bio-
¨
sciences equipped with a self-packed Superdex 200 10/100 glass
column. PBS (pH 7.4) was used as the mobile phase with a flow
rate of 1.0 mL/min. For amine functionalized polymers, the PBS
buffer was supplemented with 50 mM of 2-(2-aminoethoxy)ethanol.
Typical injection volumes were 100 µL. Detection was achieved
by measuring the absorption at 280 nm.
Fluorescamine Assay of Amine Reactivity PILs. Stock solu-
tions of amine-containing PIL polymers were made at 20 mg/mL
concentration. A serial dilution was made using 1, 2, and 4 uL of
polymer stock into 240 uL of PBS buffer, followed by addition of
10 uL of a 30 mg/mL solution of fluorescamine. This mixture was
vortexed and incubated at room temperature for 10 min before
fluorescence analysis on a BioTek plate reader with excitation at
380 nm and detection at 480 nm. The recorded fluorescence
intensity signals were calibrated against solutions of known
concentrations of compound 5 (methoxyPEG-NH2).
Cell Culture. HeLa cells were grown in DMEM (Mediatech)
with 10% Fetal Bovine Serum (Invitrogen), 50 U/mL penicillin
and 50 µg /mL streptomycin (Invitrogen). The cells were transfected
using 1 µL Lipofectamine 2000 (Invitrogen), 0.2 µg of BirA-ER
plasmid22,43 and 0.2 µg of AP-YFP-TM plasmid per well of an
8-well chamber slide (LabTek). One mM biotin was added to the
media during plasmid expression. Cells were imaged under 4 °C
PBS the day after transfection. 1% Bovine Serum Albumin (Sigma)
was added to block nonspecific binding during specific binding
studies of ligand-coated quantum dots. Commercial BSA is known
to contain biotin, and the stock BSA solution was dialyzed with a
3 kDa cutoff dialysis tube three times for 8 h in PBS pH 7.4, in 4
°C.
Nonspecific Binding of QDs to Serum. The 565-nm emitting
CdSe(CdZnS) QDs (5 µL) of various surface coatings were mixed
with fetal bovine serum (95 µL) to a final concentration of ∼0.5
µM. The mixture was incubated for 4 h at 37 °C with gentle mixing.
The resultant QD size distribution was then measured using gel
filtration chromatography. The mixture was injected into a Superose
6 GL10/300 column (GE Healthcare, Piscataway, NJ) on an Agilient
1100 series HPLC with an in-line degasser, autosampler, diode array
detector, and fluorescence detector (Roseville, CA). PBS (pH 7.4)
was used as the mobile phase with a flow rate of 0.5 mL/min and
an injection volume of 50 µL. In order to selectively measure the
signal from the QD rather than FBS, the fluorescence detection at
565 nm with 250 nm excitation was chosen.
Fluorescence and Phase Contrast Microscopy. Cells were
imaged live using a Nikon TE2000-U inverted microscope with a
60× water-immersion lens and a Princeton Instruments MicroMAX
Camera with an additional 1.5× magnification tube lens. Bright
field images were collected using differential interference contrast
and 10 ms exposure. Fluorescence images were collected with
epifluorescent excitation provided by the 488 nm line of an argon-
Ion laser with the appropriate dichroic (Chroma, Z488RDC) and
emission filters (QD605: D605/30M, YFP: D565/30 m). Images
were collected and analyzed using Image J version 1.41o. Typical
exposure times were 0.1-0.5 s and fluorescence images were
background-corrected.
Covalent Conjugation of Streptavidin to poly(aminoPEG11)25%
-
PIL QDs. Stretpavidin (50 µL, 10 mg/mL; Sigma Aldrich) was
activated in MES buffer (pH 6.5) using Sulfo-NHS and EDC (20
equiv) for 20 min at RT. The activated SA was mixed with
poly(aminoPEG11)25%-PIL QDs in sodium bicarbonate buffer at pH
8.4 at a SA:QD ratio of 5:1 and allowed to react for 1 h. The
samples were dialyzed 2× through a 50 kDa MW cutoff spin
concentrator and then used for labeling experiments.
Quantum Yield Measurement. QY of 605 nm emitting QDs
was measured relative to Rhodamine 640 (λex ) 535 nm). Solutions
(50) Snee, P. T.; Chan, Y.; Nocera, D. G.; Bawendi, M. G. AdV. Mater.
2005, 17, 1131–1136.
(51) Eaton, D. Pure Appl. Chem. 1988, 60, 1107–1114.
(49) Murray, C. B.; Norris, D. J.; Bawendi, M. G. J. Am. Chem. Soc. 1993,
115, 8706–8715.
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482 J. AM. CHEM. SOC. VOL. 132, NO. 2, 2010

Compact Biocompatible Quantum Dots via RAFT
A R T I C L E S
Animal and Tumor Models. Orthotopic P008 mammary
carcinoma models were prepared by implanting a small piece (1
mm3) of viable tumor tissue from the source tumor animal into the
mammary fat pad chamber44 of 10-12-week old female Tie2-GFP/
FVB mice. The tumors were allowed to grow up to 5 mm in
diameter. All animal procedures were carried out following the
Public Health Service Policy on Humane Care of Laboratory
Animals and approved by the Institutional Animal Care and Use
Committee of Massachusetts General Hospital.
Intravital Multiphoton Imaging. To study tumor vasculature
using QDs and their distribution dynamic in live animals, 150 µL
poly(PEG)-PIL QD600 at a concentration of 5 µM were injected
retro-orbitally into the tumor bearing mice and imaged with
multiphoton laser scanning microscope.52 The images were recorded
as 3D stacks (200-µm thickness, 1-µm step size) at 0 h, 3 and 6 h
interval respectively and processed using the NIH ImageJ software.
For the GFP channel, the emission filter used was 535 ( 20 nm,
and for QD600, the emission filter was 625 ( 75 nm. All images
were captured with a 20× water emersion lens (N.A. 0.95) and an
excitation wavelength of 880 nm (500 mW).
M.G.B., and D.G.N.), R01-CA085140, R01-CA115767 (R.K.J.),
P01-CA080124 (R.K.J. and D.F.), R01-CA096915 (D.F.); by the
MIT-Harvard NIH Center for Cancer Nanotechnology Excellence
(1U54-CA119349) (M.G.B.); by the MIT DCIF (CHE-980806,
DBI-9729592); by the ISN (W911NF-07-D-0004) (M.G.B. and
D.G.N.); by the NSF-MRSEC program (DMR-0117795) via the
use of its shared user facilities; and by the Army Research Office
(W911NF-06-1-0101) (D.G.N). W.L. was supported by a
National Science Foundation Graduate Research Fellowship.
A.B.G. was a Novartis fellow of the Life Sciences Research
Foundation. We would like to thank Dan Liu and Peng Zou for
valuable assistance with cells, plasmids, and cell culture
protocols.
Supporting Information Available: Supporting material con-
1
tains H NMR characterization of monomers and polymers,
Forster energy transfer calculations for QD-dye conjugates, GPC
polymer sizing data, and optical characterization of QDs. This
information is available free of charge via the Internet at http://
pubs.acs.org/.
Acknowledgment. This research was supported by the U.S.
National Cancer Institute Grants Nos. R01-CA126642 (R.K.J.,
(52) Brown, E. B.; Campbell, R. B.; Tsuzuki, Y.; Xu, L.; Carmeliet, P.;
Fukumura, D.; Jain, R. K. Nat. Med. 2001, 7, 864–868.
JA908137D
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