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SulfobetaineVinylimidazole Block Copolymers: A Robust Quantum Dot Surface Chemistry Expanding Bioimaging's Horizons Mariana Tasso,† Emerson Giovanelli,† Diana Zala,§ Sophie Bouccara,† Alexandra Fragola,† Mohamed Hanafi,‡ Zsolt Lenkei,§ Thomas Pons,† and Nicolas Lequeux*,† †
LPEM, ESPCI ParisTech, PSL Research University, CNRS UMR 8213, Sorbonne Universités, UPMC Univ. Paris 6, 10 rue Vauquelin, 75005 Paris, France, ‡SIMM, ESPCI ParisTech, PSL Research University, CNRS UMR 7615, Sorbonne Universités, UPMC Univ. Paris 6, 10 rue Vauquelin, 75005 Paris, France, and §Brain Plasticity Unit, ESPCI ParisTech, PSL Research University, CNRS UMR 8249, 10 rue Vauquelin, 75005 Paris, France
ABSTRACT
Long-term inspection of biological phenomena requires probes of elevated intra- and extracellular stability and target biospecificity. The high fluorescence and photostability of quantum dot (QD) nanoparticles contributed to foster their promise as bioimaging tools that could overcome limitations associated with traditional fluorophores. However, QDs' potential as a bioimaging platform relies upon a precise control over the surface chemistry modifications of these nano-objects. Here, a zwitterionvinylimidazole block copolymer ligand was synthesized, which regroups all anchoring groups in one compact terminal block, while the rest of the chain is endowed with antifouling and bioconjugation moieties. By further application of an oriented bioconjugation approach with whole IgG antibodies, QD nanobioconjugates were obtained that display outstanding intra- and extracellular stability as well as biorecognition capacity. Imaging the internalization and intracellular dynamics of a transmembrane cell receptor, the CB1 brain cannabinoid receptor, both in HEK293 cells and in neurons, illustrates the breadth of potential applications of these nanoprobes. KEYWORDS: block copolymers . quantum dots . zwitterion . vinylimidazole . oriented immobilization . cell receptor tracking . cannabinoid receptors
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n parallel with the continuous progress in the synthesis of highly luminescent semiconductor quantum dots (QDs), considerable efforts have been undertaken to develop a robust surface chemistry and bioconjugation approach13 compatible with in vivo and in vitro bioimaging applications.4 The main challenges these optical probes face in biological environments are related to their long-term colloidal stability in complex biological systems, minimal TASSO ET AL.
unspecific adsorption requirements, and ability to selectively recognize the biomolecules of interest. The phase transfer of hydrophobic as-prepared QDs to aqueous media occurs either by encapsulation with surfactants or amphiphilic copolymers or by ligand exchange with a hydrophilic molecule that replaces the native nanoparticle ligand.5,6 High quality QDs for bioimaging applications usually possess a core (multi)shell structure with an outermost shell VOL. XXX
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[email protected]. Received for review September 10, 2015 and accepted October 27, 2015. Published online 10.1021/acsnano.5b05705 C XXXX American Chemical Society
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compact terminal block rather than randomly distributing them along the ligand's chain. In fact, this strategy is used to add a histidine tag (Hisn) to proteins, with n ≈ 6. His-tagged macromolecules have been immobilized onto QDs relying upon the collective coordination between pendant imidazole moieties, regrouped on the poly(histidine) residue, and the Zn-rich inorganic QD surface.3538 Inspired by this strategy, the design and application of a poly(methacrylamidosulfobetaine-block-vinylimidazole) copolymer ligand prepared by controlled radical polymerization is here reported. To the best of our knowledge, multidentate ligands for QDs have never been synthesized with a block structure. In this work, QDs capped with the zwitterion/ vinylimidazole block copolymer were characterized with regard to their pH and colloidal stability over time, as well as concerning their nonspecific interactions with live cells and intracellular stability up to 50 h in the demanding cytosol environment. By incorporating amine functionalities to the zwitterionic block, the ligand served as a platform for the subsequent oriented bioconjugation of the nanoparticles with whole IgG antibodies. The obtained nanobioconjugates demonstrated a remarkable efficacy at specifically targeting a cell transmembrane protein, the type-1 cannabinoid receptor CB1R, enabling multicolor visualization of the internalization and trafficking of this protein over time in live HEK293 cells and neurons. In particular, observation times exceeding the 48 h time-window, which are rarely feasible with traditional fluorophores, can be achieved with the QD nanoconjugates here presented.
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of ZnS or (Zn,Cd)S. Anchoring groups able to coordinate Zn(II) surface cations include thiols, carboxylic acids, amines, imidazoles, pyridines, and phosphines. Thiol-appended polymers, like 3-mercaptopropionic acid, have been widely employed to cap-exchange, with dithiols gradually replacing monothiols as terminal units and later as side chains in multidentate polymers.713 Adding multiple dithiols to the ligand chain has proven to be a good strategy to increase QD stability,13 though the susceptibility of thiols to (photo)oxidation remains a shortcoming.5,14,15 To overcome this limitation, multidentate imidazole or pyridine copolymer ligands have been designed with demonstrated good colloidal stability.1618 Compared with thiols, these anchoring groups are not prone to oxidation, are often less detrimental to QD fluorescence, and do not interfere with standard bioconjugation reactions. In addition to be strongly anchored to the inorganic surface, ligands must be designed to prevent nonspecific interactions in biological media. The ubiquity of poly(ethylene glycol) (PEG) as material conferring antifouling properties to surfaces has only recently witnessed the emergence of zwitterionic compounds as promising alternatives.19 The lower susceptibility of highly charged zwitterion groups to deplete water molecules away from nonpolar surfaces makes zwitterionic materials less prone to hydrophobic interactions.20,21 Amino acids (e.g., cysteine), cysteamine, penicillamine, sulfobetaine, phosphorylcholine, and carboxybetaine-based zwitterionic ligands have been used to stabilize nanoparticles in water and have shown excellent antifouling performance.12,13,2228 Compared with PEG-capped QDs or gold nanoparticles, more compact sizes,25,29 longer circulation times,30 better stability in undiluted blood serum,31 and selective uptake by cancer cells32 have been reported. PEG- or zwitterion-based polymers with multipendant anchoring groups are currently synthesized either by random copolymerization of two (or more) monomers or by grafting precursors on a common backbone. In both cases, the anchoring groups are randomly distributed along the main chain, a fact that could imply the existence of free, not surface-bound anchoring moieties. This is indeed what has been recently observed with dithiolzwitterion copolymer ligands for which a partial substitution of thiols was feasible with maleimide-functionalized DNA or protein.33,34 Inasmuch as the ligand's binding strength to the nanoparticles may be dependent on the total number of anchoring groups, the presence of free anchoring units may be detrimental to the colloidal and antifouling properties of the resulting nanoparticles. Such subtle balance has not been properly addressed so far and deserves a timely study to optimize the ligand's architecture. Another approach that has not been explored yet consists in regrouping all the anchoring groups in one
RESULTS AND DISCUSSION Synthesis of the Poly(methacrylamidosulfobetaine-block-4vinylimidazole) Block Copolymer. Since its discovery in 1988,39 reversible additionfragmentation chain transfer (RAFT) polymerization has become a popular technique to control the architecture of macromolecules including block, grafted, comb, and star structures. This technique has been successfully applied to control the polymerization in water of acrylamide, methacrylamide, or methacrylate-based zwitterionic monomers, such as sulfobetaines, phosphorylcholines, and carboxybetaines using dithiobenzoates as chain transfer agent.16,27,4046 In this work, polysulfobetaine homopolymer (PSPP) was prepared using 3-[3-methacrylamidopropyl-(dimethyl)ammonio]propane-1-sulfonate (SPP) as monomer, 4-cyanopentanoic acid dithiobenzoate (CADB) as chain transfer agent (CTA), and 2,20 -azobis(2-amidinopropane) hydrochloride (V50) as initiator (Scheme 1). The method here proposed ensures that all monomers are zwitterions, in contrast to classical protocols involving postquaternarization/alkylsulfonation of ternary aminebearing polymeric backbones to form sulfobetaine moieties for which reaction can be incomplete due to VOL. XXX
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Scheme 1. Polymer Synthesisa
a
(a) General RAFT block copolymerization procedure and (b) block copolymer containing primary amines obtained by random RAFT copolymerization of SPP and APMA in the first-block synthesis.
Figure 1. (a) Pseudo-first-order kinetic plot for the polymerization of SPP at 70 °C, mediated by CADB and initiated by V50. M and M0 are the monomer concentrations at time t and t = 0, respectively. (b) Evolution of the number-average molar mass (Mn) and polydispersity index (PDI) determined by size-exclusion chromatography as a function of monomer (SPP) conversion. The line corresponds to the theoretical molar mass of the polymer as a function of monomer conversion.
steric hindrance.12,16,22,23 The reaction was conducted in an acetate buffer solution at pH 5.2 to limit the hydrolysis of the dithiobenzoate agent.47 Because CADB is not highly soluble in acidic media, it was first dissolved in a small volume of 2 M NaOH aqueous solution prior mixing with the buffer solution. Monomer conversion was determined by 1H NMR following the reaction. As shown in Figure 1a, a pseudo-first-order kinetics was observed without any evidence of an induction period. To recover the polymers, precipitation was preferred to dialysis in order to limit the contact time of the polymers with water, which is likely to degrade the dithiobenzoate end group.47 Efficient removal of excess monomers by precipitation was confirmed by 1H NMR (Figure S1). The molecular weight values estimated from aqueous SEC (size exclusion chromatography) were in good agreement with those TASSO ET AL.
expected from the monomer/CTA ratio. Polydispersity index (PDI) values never exceed 1.05 even after 6 h of reaction and for conversions higher than 70% (Figures 1b and S2). Because PSPP homopolymer cannot be functionalized with biomolecules, a copolymer P(SPPAPMA) was synthesized by introducing a primary amine moiety, namely, N-(3-aminopropyl)methacrylamide hydrochloride (APMA), in the starting mixture (Scheme 1). The [APMA]/[SPP] molar ratio was fixed at 1/7. The incorporation of APMA in the RAFT polymerization did not induce noticeable differences in polymerization kinetics or polydispersity. At this stage, it was not possible to confirm the presence of APMA in the macromolecule architecture by 1H NMR, but as it will be demonstrated in next sections, (i) amine incorporation induces a change in electrophoretic mobility of ligandexchanged QDs and (ii) grafting of proteins via amide VOL. XXX
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bonds is effective only when APMA is incorporated to the ligand. For the P(SPP-b-4VIM) or P([SPP-APMA]-b-4VIM) diblock copolymers, we started with a PSPP macro-CTA or P(SPPAPMA) macro-CTA with a molecular weight varying from 5000 to 10000 g 3 mol1 to limit the final size of the diblock polymer-capped QDs (Scheme 1 and Figure 1). Allen et al.48 have found that RAFT polymerization of 4VIM in acetic acid instead of acetate buffers produces homopolymers with well-defined molecular weights. 4VIM synthesized by decarboxylation of urocanic acid was used rather than the commercial 1VIM, which proved difficult to polymerize due to the absence of resonance stabilization.48 We conducted the polymerization of the diblock polymers in acetic acid mixed with a minimum of water to ensure the solubility of macro-CTA; the amount of 4VIM monomer per macro-CTA was fixed to 10:1. After a 6 h polymerization at 70 °C with AIBN as initiator, the copolymer was recovered by precipitation, and the terminal dithiobenzoate group was cleaved in an aqueous solution of NaBH4. During this step, the polymers progressively lost their pink color. The resulting colorless polymer solutions were then finally precipitated in methanol and lyophilized. The removal of the dithiobenzoate function was confirmed by 1H NMR (Figure S3). Quantitative 1H NMR (Figure S3) indicated that about 90 mol % of the vinylimidazole monomers were incorporated into the final product in agreement with a monomer conversion of 80%. This was also confirmed by the increase in the N/S mass ratio measured by weight analysis after the second polymerization step (quantitative elemental analysis for PSPP and P(SPP-b4VIM) after dithiobenzoate cleavage: C, 47.6; H, 8.4; N, 9.1; S, 11.0 and C, 47.8; H, 8.3; N, 10.4; S, 9.7, respectively). Altogether, these results suggest that, on average, ∼89 imidazole units have been added at the end of the PSPP chain. However, the assumed block copolymer architecture could not be confirmed by aqueous SEC due to the strong interaction of the imidazole group with the chromatographic column. To demonstrate the controlled nature of the copolymerization, a fluorescein dye was coupled at the thiol end group of the PSPP homopolymer and of the P(SPP-b-4VIM) block copolymer after dithiobenzoate end group removal. The gel electrophoretic mobility shift at pH 4 of these fluorescently grafted polymers showed that the PSPP monomer is slightly negatively charged (Figure 2), possibly due to asymmetric counterion interactions.49 In contrast, the P(SPP-b-4VIM) copolymer migrates to the negative electrode due to the protonated imidazole units (pKa ≈ 6), suggesting the absence of PSPP homopolymer impurities after addition of the second block, which indicates the effective addition of the 4VIM second block to the PSPPmacro-CTA first block. Finally, it is to be noted that this method readily provides the diblock copolymer
Figure 2. Gel shift of (A) fluorescein end-grafted PSPP macro-CTA homopolymer, (B) fluorescein end-grafted P(SPP-b-4VIM) copolymer, and (AþB) a mixture of both in a 1% agarose gel. Unreacted PSPP chains are not observed in the final P(SPP-b-4VIM) block copolymer product.
in two steps, without any laborious protection or deprotection stages.16 P(SPP-b-4VIM)-Capped QDs. Orange-red CdSe/CdS/ZnS multishell quantum dots emitting from 590 to 615 nm were synthesized following published protocols.50,51 Green CdZnSe/CdZnS gradient-alloyed QDs (525 nm) were synthesized following the procedure from Bae et al.52 The direct exchange of the native oleic acid/ oleylamine QD surface ligands by the synthesized copolymers is not efficient even in biphasic mixtures (e.g., H2O/CHCl3) due to the poor solubility of sulfobetaine-based polymers in the majority of solvents other than water. A two-step exchange procedure was then implemented, requiring a preliminary ligand exchange with 3-mercaptopropionic acid (MPA), as described in Giovanelli et al.13 The subsequent MPAto-polymer exchange was carried out at room temperature (i.e., no heating required) with an amount of polymer corresponding to 11.5 polymer chains per square nanometer of QD surface. IR spectroscopy confirmed a full displacement of the MPA ligands by the P(SPP-b-4VIM) polymer (Figure S4). The properties of the cap-exchanged nanoparticles were characterized by various physical methods, including fluorescence spectroscopy, DLS, and ζ-potential measurements. The two-step ligand exchange process leads to a final depletion of fluorescence, which varies from 20% to 40% depending on the starting QD batches (as-synthesized fluorescence quantum yield ∼4060%). The greatest quantum yield losses were observed to appear after the first ligand-exchange step, whereas, in general, a partial fluorescence recovery accompanies the second step. Varying the polymer amount by a factor of 10 or changing the polymer molecular weight (500020000 g 3 mol1) has no effect on the final quantum yield value. The hydrodynamic diameter (DLS) is typically ∼18 nm for nanoparticles with an inorganic size of 7 nm and a polymer containing ∼30 SPP units (Mn ≈ 10000 g 3 mol1, Figure S5). VOL. XXX
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ARTICLE Figure 3. (a) pH stability of P(SPP20-b-4VIM9)-coated QDs (λmax = 603 nm). Relative absorbance at 350 nm and fluorescence intensity (integrated from 500 to 690 nm) of samples stored for 7 days in buffers spanning the 212 pH range. Results are normalized with those of pH = 7 sample. Images and optical data were collected from the supernatants obtained after centrifugation at 19000g for 10 min. (b) Relative absorbance at 350 nm of QD dispersions measured after various storage times in the dark at 37 °C (filled symbols) or under room light at 20 °C (unfilled symbols). Two distinct copolymer ligands were employed: P(SPP-b-4VIM) (circles) and P(SPP-co-dithiol) (squares). QD (200 nM) dispersions in 100 mM aqueous NaCl (pH 6.5) were tested; their absorbance after storage measured on the supernatants obtained after centrifugation (19000g, 10 min).
The diameter decreases by about 1 nm when the number of SPP groups is reduced to 20. Since light scattering varies to the sixth power of the size, DLS is very sensitive to even minute fractions of aggregates in colloidal suspensions. In the present case, no aggregates are detected in the DLS (unweighted) distribution, thus confirming the efficiency of the sucrose gradient purification procedure (Figure S5). Moreover, after six month storage at 4 °C in the dark, neither variation of the hydrodynamic size nor formation of aggregates was revealed, pointing to a preserved colloidal stability. Furthermore, the slightly negative ζ-potential values around 7 ( 1 mV in 100 mM aqueous NaCl (pH = 5.5) were found to be representative of a rather neutral zwitterionic outer shell, as well as coherent with the gel electrophoretic measurements of the PSPP first block. In line with our aim of producing robust nanobioconjugates to be employed in biological applications, the influence of several parameters, such as pH, dilution, and light exposure, on the colloidal stability and fluorescence properties of the nanoparticles was carefully inspected. Integrated fluorescence intensity and absorbance at 350 nm of 0.5 μM QD dispersions in buffers spanning a 212 pH range were measured after 7-day storage (Figure 3a), and luminescence was imaged at various time points (Figure S6a). Whereas the suspensions at pH g 7 remained optically stable during the length of the experiment (e.g., less than 10% of fluorescence loss at pH 7, Figure S6b), a progressive loss of fluorescence, accentuated as the pH decreases, was observed in acidic media (Figure 3a and S6a). After 7 days, all samples suspended at a pH lower than 6 had lost more than 50% of fluorescence and were prone to aggregation, as confirmed by a noticeable decrease in absorbance (Figure 3a). This pH limit is likely to be correlated to the protonation of the imidazole group (pKa ≈ 6), leading to a progressive ligand desorption and acidic etching of the semiconductor surface, which TASSO ET AL.
in turn could be responsible for fluorescence quenching. The present results are in line with previous reports on the low pH-stability in acidic media of QDs coated with random polymers appended with pyridine and PEG pendant groups, therefore confirming the detrimental role of the protonation of anchoring group's nitrogen atoms on QD colloidal stability.17 Focusing then on the colloidal stability of ligandcapped QDs in dilute solutions, the critical role of the anchoring groups' affinity and stability toward the inorganic surface was put in evidence. For this, QDs capped with two sulfobetaine-based polymers differing in the chemical nature of their anchoring groups, imidazole for the first and dithiol for the second, were examined regarding their colloidal stability. P(SPP20-b4VIM9) and the statistical dithiol-appended sulfobetaine polymer, P(SPP-co-dithiol) (Mn ≈ 5000 g 3 mol1, PDI = 2, [dithiol]/[sulfobetaine] molar ratio = 7/93)13 were employed. Diluted (200 nM) samples were stored for 28 days under room light at room temperature (potential storage conditions) or in the dark at 37 °C (cell culture conditions; Figure 3b). Absorbance measurements on the QD suspensions remaining in solution after centrifugation highlighted the good colloidal stability of both multidentate polymer-coated QDs when left in the dark at 37 °C for 28 days, with a loss in absorbance slightly higher for P(SPP-co-dithiol) vs P(SPP20-b-4VIM9)-coated QDs (∼20% vs ∼4%). At this stage, a general conclusion about the affinity differences between imidazole and thiol functions toward the QD surface cannot be extracted due to the differences in polymer architectures (number and distribution of anchoring groups, molecular weight, etc.), which greatly influence the adsorptiondesorption equilibrium. However, in the case of storage under room light, the differences in colloidal stability are sufficiently relevant to distinguish the distinct role of imidazole and dithiol functions on the ligand's binding strength. As expected, thiolated polymers are prone to VOL. XXX
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photo-oxidation.15 This can contribute to a continuous and irreversible desorption of these ligands leading to severe aggregation, as shown in Figure 3b. Light exposure also has a detrimental effect on the stability of the imidazole-based ligand, though to a lesser extent than for the thiolated-one. QD Bioconjugates and Cell Experiments. Bioconjugation of the ligand-capped QDs is an essential condition for any target-specific (bio)application of these nanoprobes.51 Given the presence of primary amines in the ligand architecture P([SPP-APMA]-b-4VIM), amide chemistry via a homobifunctional linker (bis(sulfosuccinimidyl)suberate, BS3) was selected to immobilize an intermediate protein A layer, which in turn served as a platform for the oriented immobilization of whole IgG antibodies to the nanoparticles. The presence of both protein and antibody layers was confirmed by the increase in hydrodynamic diameter (DLS) of the nanoconjugates as well as by a protein quantification assay (bicinchoninic acid assay, BCA) applied to the supernatants recovered after bioconjugation. Particle diameter increased from ∼18 nm (ligand Mn ≈ 10000 g 3 mol1) to ∼21 nm and ∼27 nm after protein and antibody immobilization, respectively, all with a distinctively homogeneous particle size distribution that is largely devoid of aggregates (Figure S7). Given the homobifunctional nature of the BS3 linker, multiple binding opportunities between a single protein A molecule and independent QD nanoparticles are not to be excluded, especially when the encounter probability of both species is high (typically at high concentrations). By tuning the concentration and molar ratio of both reagents, mostly aggregate-free bioconjugates were obtained. Noteworthy, the addition of an antibody layer only slightly increased the width of the size distribution, thus minimally altering the colloidal monodispersity that characterized the previous layers (Figure S7). Regarding the total protein and antibody amounts immobilized per dot, the BCA test resulted in approximately four protein A and one to two Ab molecules per QD (see Supporting Information for details about the BCA test). Increasing the QD/protein A molar ratio yielded nanobioconjugates of higher hydrodynamic diameters than those aimed at for the biological applications (∼30 nm). Noticeably, when the corresponding amine-free ligand, P(SPP-b-4VIM), was employed for bioconjugation, neither significant variations in QD hydrodynamic diameter (p = 0.01 after incubation with pA and Ab) nor retention of protein onto the QD surface were observed on the final nanoconstructs. This confirms the bioconjugation dependence on the ligand's amine moieties as well as the protein-resistance properties of the aminefree ligand attributable to their majority zwitterion segments. In addition, gel electrophoresis revealed differences in mobility between unconjugated amine-free and amine-containing polymer-capped QDs (Figure S8). Last but not least, bioconjugated QDs did not display
Figure 4. Intracellular colloidal stability of P(SPP30-b-4VIM9)capped QDs after (a) 2 h or (b) 50 h cargo delivery to the cell cytoplasm via electroporation. Epifluorescence microscopy images (image intensity increased in panel b to overcome signal depletion due to cargo splitting (cell division)). Cargo dilution after 50 h has no noticeable effect on QDs' cytoplasmic stability.
any changes in their native absorbance and fluorescence emission characteristics (Figure S9) Long-term stability in cellular media and nonspecific interactions with live HeLa cells of QDs bioconjugated or not were evaluated after 24 and 48 h incubation (Figure S10). QDs modified with an aminecontaining ligand (P([SPP30APMA4]-b-4VIM9)) displayed substantial nonspecific interactions with HeLa cells and were therefore retained on the cells' membranes. QDs capped with an amine-free ligand as well as bioconjugated QDs (QDpAAb) had only mild nonspecific interactions with live cells (Figure S10b); the fluorescence intensity of the QD-incubated cells being higher than autofluorescence (control cells) only for the higher QD concentration tested, 1 μM, at both incubation times (p = 0.05). The mild retention of bioconjugated and amine-free ligand-capped QDs on the cells' membranes after 48 h incubation provides indirect proof of the colloidal stability of these nanoconstructs in cell media; ligand displacement by the medium's biomolecules typically results in loss of colloidal stability, hence in higher retention on surfaces. Denaturation of QD-immobilized protein or Ab, which would enhance hydrophobic, nonspecific interactions with the cell membrane, went also unnoticed. Mild nonspecific interactions correlated with substantially preserved cellular metabolic activities (as determined by the MTT test; Figure S11). After 48 h exposure to the nanoconjugates at 1 μM concentrations (2050 times higher than the concentrations used in the receptor labeling experiments described below), cells displayed more than 80% of their unperturbed metabolic activity. On the other hand, the intracellular stability of the P(SPP-b-4VIM) zwitterionvinylimidazole ligand-capped QDs was assessed by cargo delivery via HeLa cell electroporation over a 50 h time period (Figure 4). Here, neither the rather demanding cytosol environment nor cell division appear to have induced QD aggregation or loss of fluorescence; on the contrary, 50 h postelectroporation, QDs remain fluorescent and freely moving in the cytosol (Video S1). The very satisfactory VOL. XXX
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ARTICLE Figure 5. (a) CB1 receptor localization and dynamics as evidenced by a two-color QD system in HEK293 cells expressing a FLAG-tagged CB1 receptor. At time zero (t0), two receptor populations can be identified: those already internalized and present within endosomes (red QDpAanti-FLAG, λmax = 615 nm) and those still confined to the cell membrane (green QDpAanti-FLAG, λmax = 525 nm). Three hours later (t3h), a large fraction of the green population is internalized, mostly in different localizations than the red population. After 6 h (t6h), a much more homogeneous distribution of the green and red receptor populations can be observed (see zoom-in insets), with almost no residual green signal at the cell membrane. Epifluorescence microscopy images of fixed cells for both QD channels are presented together with their overlay. (b) Rat hippocampal neurons transfected to express the FLAGCB1GFP receptor were labeled with red QDpAanti-FLAG and observed live. Complete internalization of the initially membrane-bound QDs was observed ∼5 h postincubation. QDs are preferentially located inside endosomes coincident with GFP-rich zones (zoom-in inset). (c) The specifics of axonal CB1 receptor movements are put in evidence by the QD nanotools: receptors display random (arrows) or directed (continuous and colored line in the upper right corner) transport patterns. Temporal code analysis was applied to epifluorescence microscopy images tracking QDs' trajectories (see Video S2).
intra- and extracellular stability of these nanoconstructs over 48 h denotes the robust anchoring of the poly(vinylimidazole) block-based ligand to the QD surface even under biologically demanding conditions. On these bases, the specific recognition of a transmembrane protein, the cannabinoid receptor type-1 (CB1), in HEK293 cells and primary neurons was evaluated (see Supporting Information for details). The CB1 receptor has been shown to constitutively cycle between cell membrane and cytosol via endocytic pathways, both on HEK cells53 and rat hippocampal neurons.54 Cells transfected with a FLAGCB1 plasmid55 were exposed to QDpAanti FLAG nanoconjugates in various conditions, as follows. Two QD batches were used that differ in their maximum fluorescence emission wavelength: red QDs (615 nm) and green QDs (525 nm). HEK cells were first exposed to red QDpAanti-FLAG for 10 min, then incubated for 3 h to enable the internalization of the receptor-bound QDs into endosomes, and finally exposed to green QDpAanti-FLAG for another 10 min. The concomitant presence of a red signal confined to TASSO ET AL.
cytoplasmic endosomes and of a green imprint fully localized on the cell membrane at time zero (Figure 5a, t0) was indicative of two distinct receptor populations whose traffic dynamics over time can be explored on a two-color mode by means of these nano(bio)tools. Three hours later (t3h), both receptor populations were mostly internalized but occupied different subcellular domains. Finally, 6 h postincubation with the green QDs, the relative partitioning of the red and green QD populations became rather uniform, with less distinction between red- or green-rich zones (Figure 5a, t6h and insets). On the other hand, when red and green QDs were added simultaneously, both colors were internalized indistinctively and could be found strongly colocalized on inner endocytic vesicles after a similar period of time (Figure S12). Furthermore, experiments performed with CB1þ (CB1 positive), FLAGþ cells exposed to QDpA and QDpAAb, with Ab not specific for the FLAG tag, did not yield noticeable binding (Figure S13a,b). Similarly, CB1þ, FLAG (FLAG negative) cells exposed to QDpAanti-FLAG were not labeled (Figure S13d). VOL. XXX
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cell experiments, as well as to be applied to other relevant biological questions. Conclusions and Perspectives. We have synthesized block zwitterionvinylimidazole polymers using RAFT polymerization. Ligand exchange with these polymers enables long-term stabilization of QDs in dilute conditions. Insertion of primary amines in the zwitterionic block allows for the oriented bioconjugation of unmodified antibodies via a protein A intermediate layer. The obtained antibody-conjugated QDs display a low level of unspecific binding to live cells and retain full antibody functionality and specificity. These attributes provide robust nanotools for the inspection of biological phenomena. Single-molecule tracking studies and target-specific diagnostics or sensing are at the forefront of this technology's envisioned applications.
MATERIALS AND METHODS
manner as PSPP macro-CTA with an [APMA]/[SPP] molar ratio equal to 1/7 (APMA = 3.58 mmol, SPP = 25.06 mmol, CADB = 0.36 mmol, V50 = 0.07 mmol). Synthesis of Poly(SPP-block-4VIM) [P(SPP-b-4VIM)] and poly([SPP-coAPMA]-block-4VIM) [P([SPP-APMA]-b-4VIM)]. In a typical reaction, a PSPP macro-CTA polymerized during 80 min (Mn = 6600 g 3 mol1, Mw/Mn = 1.03) and an estimated (1H NMR) degree of polymerization of 20. This PSPP20 macro-CTA (0.167 mmol) was dissolved together with 4VIM (1.67 mmol, [4VIM]/[PSPP macro-CTA] molar ratio = 10/1) and AIBN (0.164 mmol, [AIBN]/[PSPP macro-CTA] = 1/1) in a mixture of 1.6 mL of 100 mM NaCl aqueous solution and 14.4 mL of acetic acid in a 20 mL vial. The reaction medium was purged with argon for 1 h and thereafter heated at 70 °C for 6 h. The crude block copolymer was 3-fold precipitated in ethanol and redispersed in water. The product was then dried under vacuum. The same protocol was applied to prepare P([SPP-APMA]-b-4VIM) using the same 10/1 [4VIM]/[P(SPP-APMA) macro-CTA] and 1/1 [AIBN]/[P(SPP-APMA) macro-CTA] molar ratios. To remove the phenyl end groups, the pink-colored block copolymer (500 mg) was dissolved in water (5 mL) and incubated with NaBH4 (∼50 mg). After 6 h (when the solution turned colorless), the polymer was twice precipitated in acetone, redispersed in water, and then reprecipitated in methanol. The obtained white polymer with a thiol end group was purified by dialysis against water (Spectra/Por, MWCO = 2000 Da) and lyophilized. The lyophilized copolymer ligand was stored at 4 °C until use. Ligand Characterization. Size Exclusion Chromatography. The number (Mn) and weight (Mw) average molar masses, as well as the polydispersity index (PDI = Mw/Mn), of the synthesized copolymers were determined by size exclusion chromatography (SEC) in a 0.5 M NaNO3 aqueous solution at 25 °C and at a flow rate of 1 mL/min using a Viscotek SEC system equipped with three SHODEX OH pack columns SB-806M HQ (13 μm, 300 mm 8 mm). The polymers were injected at a concentration of 4 mg/mL after filtration through a 0.2 μm pore size membrane. The absolute molar masses were determined by the three in-line detectors (refractometer, viscometer, and light scattering) relying upon a calibration based on poly(ethylene oxide) standards. Gel Electrophoresis of Fluorescein-Functionalized PSPP and P(SPP-b-4VIM). Fluorescein was coupled to the thiol end group of both copolymers by using a fluoresceinmaleimide dye. For that, PSPP macro-CTA was first reduced with NaBH4 applying the same procedure as for the block copolymer. Reduced PSPP and P(SPP-b-4VIM) (10 mg for both) were dissolved in 500 μL of 0.5 M 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer solution in water (pH = 6) and thereafter mixed with the fluoresceinmaleimide dye (70 μL of a 10 mg/mL dye solution in DMSO). After 90 min reaction, the polymer solutions were repeatedly precipitated in ethanol and resuspended in
Materials. The materials required for the synthesis of the QD nanoparticles were as detailed in Giovanelli et al.13 N-(3Aminopropyl)methacrylamide hydrochloride (APMA) was purchased from Tebu-bio, 3-[3-methacrylamidopropyl-(dimethyl)ammonio]propane-1-sulfonate (SPP) was from Raschig GmbH (RaluMer), 4-cyanopentanoic acid dithiobenzoate (CADB) was obtained from Strem Chemicals, and fluorescein-XA-maleimide was from Interchim. All other chemicals employed for the synthesis of the block copolymer ligand were from Sigma-Aldrich and used without further purification, except for 2,20 -azobis(2methylpropionitrile) (AIBN), which was recrystallized from methanol. Recombinant protein A (45 kDa) was purchased from ProSpec as a solution without additives. Bis(sulfosuccinimidyl)suberate (BS3) linker was from Thermo Scientific. Mouse monoclonal anti-FLAG (IgG2) and rabbit anti-mouse antibodies were purchased from Sigma-Aldrich. Gene Pulser electroporation cuvettes were from Bio-Rad. Neurobasal, B-27, and Lipofectamine 2000 transfection reagent were obtained from Life Technologies. The Effectene transfection reagent was from Qiagen. All other materials were as per Tasso et al.34 Synthesis of 4-Vinylimidazole (4VIM). 4VIM was synthesized by decarboxylation of 4-imidazoleacrylic acid (5.3 g) under vacuum (1 mbar) at 220240 °C.56 The product distilled as a colorless liquid, which crystallized at room temperature (2.0 g, yield = 55%). 1H NMR (400 MHz, CDCl3, δ): 11.77 (s, 1H), 7.65 (s, 1H), 7.04 (s, 1H), 6.62 (dd, 1H, J = 17.6 Hz, J = 11.2 Hz), 5.69 (dd, 1H, J = 17.6 Hz, J = 1.0 Hz), 5.14 (dd, 1H, J = 11.2 Hz, J = 1.0 Hz). Synthesis of Poly(SPP) First Block (PSPP macro-CTA). PSPP macroCTA was synthesized by controlled radical polymerization using 4-cyanopentanoic acid dithiobenzoate (CADB) as RAFT chain transfer agent. First, CADB (0.36 mmol) was added to a 250 mL round-bottomed flask and dissolved in 1.8 mL of 0.2 M NaOH aqueous solution. Immediately after dissolution, 98 mL of an acetate buffer solution (147 mmol L1 sodium acetate, 53 mmol L1 acetic acid, pH 5.2), 3-[3-methacrylamidopropyl-(dimethyl)ammonio]propane-1-sulfonate (SPP, 28.62 mmol, [SPP]/[CADB] molar ratio = 80/1), and 2,20 -azobis(2-amidinopropane) dihydrochloride (V50, 0.07 mmol, [CADB]/[V50] molar ratio = 5/1) were added. The mixture was purged by argon bubbling for 1 h and thereafter placed in a preheated oil bath at 70 °C under argon atmosphere. Aliquots were removed during the course of the reaction (from 45 to 360 min) to follow monomer consumption. At each time point, the polymerization was stopped by rapid cooling in an ice bath followed by precipitation into cool acetone. The obtained polymer was purified by dissolution in water and precipitation in ethanol (three times) and finally dried overnight under vacuum. Synthesis of Poly(SPP-co-APMA) First Block (P(SPP-APMA) macroCTA). P(SPP-APMA) macro-CTA was synthesized in the same
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In both cases, a plasmid containing a C-terminal GFP tag was employed to identify the localization of the CB1 protein in those negative controls. Time-dependent CB1 receptor internalization and distribution were also observed on FLAGCB1GFP-transfected rat hippocampal primary neurons, with initially membrane-bound QDs (not shown) becoming mostly internalized in the cell body (soma) 5 h postincubation (Figure 5b) and colocalizing with FLAGCB1GFP positive endosomes. At that time point, axonal QDs were very motile and sufficiently bright and photostable to enable a detailed visualization of receptor dynamics, characterized by either random or active transport movements (Figure 5c, Video S2). These examples highlight the potential of these nanoprobes to widen the current body of knowledge regarding protein dynamics in live
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centrifuged (19000g, 10 min), and the absorbance was measured on the recovered supernatants. The same sample was followed over time. Bioconjugation. Both, amine P([SPP30-APMA4]-b-4VIM9) and amine-free P(SPP30-b-4VIM9) block copolymer ligands were considered for bioconjugation, the latter as a negative control to demonstrate the orthogonality of the covalent bioconjugation approach. Briefly, ligand-capped QDs were first buffer exchanged from the 100 mM NaCl storage solution to 50 mM HEPES, 100 mM NaCl, pH = 7, by membrane filtration (50 kDa Vivaspin filters, 19000g, 10 min). The buffer-exchanged QDs (0.4 nmol) were thereafter resuspended in 100 μL of the pH = 7 buffer and reacted for 1 h with 0.42 μmol of BS3 (50 mg/mL stock solution in DMSO; BS3 molar excess to QDs ∼1000) under mixing in a rotating platform. Unreacted BS3 was afterwards removed via three rounds of membrane filtration as above, and the linker-modified QDs were resuspended in 100 μL of the pH = 7 buffer. Covalent binding of an intermediate protein A layer to the linker-modified QDs was performed by adding a 10 molar excess of protein A to the QD suspension and letting the reaction proceed for 1 h under mixing in a rotating platform. Here, the total volume was adjusted to yield final QD concentrations of 34 μM. After incubation, unreacted protein A was removed via two ultracentrifugation cycles (151000g, 25 min); the supernatants were recovered and utilized for the BCA quantification of unbound protein. QDprotein A (QDpA) samples (2 μM) were thereafter resuspended in 100 μL of pH = 7 buffer and mixed with 100150 μL of buffer-exchanged antibody (Ab) (rinsing buffer = 50 mM HEPES, 100 mM NaCl, pH = 8.5, adjusted with 2 M NaOH aqueous solution) at a 1:4 QD/Ab ratio. Mouse anti-FLAG (IgG2) and rabbit anti-mouse (IgG) antibodies were used. The antibody binding reaction to the QDpA nanoparticles, as well as the unbound antibody removal after reaction completion, proceeded as for the immobilization of protein A. Finally, the recovered QDpAAb conjugates were resuspended in pH = 7 buffer (final QD concentration ∼2 μM) and stored at 4 °C until use without the addition of preservatives or other compounds. Conflict of Interest: The authors declare no competing financial interest.
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20 mM NaCl until the supernatant was not fluorescent anymore. The fluorescent polymers were dried, redispersed in 50 μL of 20 mM NaCl and finally stored at 4 °C in the dark until gel electrophoresis characterization. For that, a 1% agarose gel in 100 mM acetate buffer (pH = 4) was prepared, the fluorophorelabeled samples were loaded, and the gel was run at 43 V. Nanoparticle Synthesis. Several QD samples were used for ligand exchange and characterization. CdZnSe/CdZnS gradient-alloyed QDs emitting at 525 nm were synthesized following protocols from Bae et al.52 Core/multishell CdSe/CdS/ZnS QDs emitting from 590 to 615 nm were synthesized following published protocols.57,50 Ligand Exchange. Core/shell QDs in hexane (4 nmol) were precipitated by ethanol addition followed by centrifugation (16000g, 5 min, unless otherwise stated). After removal of the supernatant, QDs were mixed with 3-mercaptopropionic acid (MPA, 500 μL) using a sonicating bath and then stored at 60 °C for 612 h. MPA-capped QDs were resuspended in 1 mL of chloroform and thereafter precipitated by centrifugation. The obtained QDs were dissolved in ∼1 mL DMF and precipitated by addition of ∼50 mg of potassium tert-butoxide. The suspension was afterward centrifuged to remove the basic organic supernatant, and the nanoparticles washed once with ethanol before redispersion in 400 μL of 100 mM sodium bicarbonate buffer (pH = 10.8). Thereafter, the block copolymers (8 mg) were resuspended in 100 mM sodium bicarbonate buffer (200 μL) and added to the MPA-QD dispersion. The nanoparticles were left overnight at room temperature to complete the cap exchange. Free ligands were removed by two rounds of ultrafiltration (16000g, 10 min) in Vivaspin 100 kDa membrane filter units (buffer = 20 mM NaCl). Polymer-capped QDs were thereafter purified by ultracentrifugation (268000g, 25 min) in a 10%40% sucrose gradient in 20 mM NaCl. The QD band was collected and sucrose removed by three rounds of ultrafiltration (100 kDa Vivaspin filter, 16000g, 10 min). The ligand-exchanged nanoparticles were finally resuspended in 600 μL of 100 mM NaCl and stored at 4 °C in the dark. Ligand-Capped QD Characterization. Dynamic Light Scattering. Dynamic light scattering (DLS) measurements of ligand-capped QDs were carried out on a CGS-3 goniometer system equipped with a HeNe laser (633 nm) and an ALV/LSE-5003 correlator. All samples were initially filtered through 0.2 μm Millipore syringe filters. Data was collected by monitoring the light intensity at different scattering angles. The hydrodynamic size distribution was obtained using the CONTIN algorithm (ALV software). ζ-Potential. ζ-Potential was determined using a Malvern Zetasizer Nano ZS90 instrument. Samples were measured at concentrations ranging from 0.02 to 1 μM at room temperature in 100 mM NaCl. pH and Colloidal Stability. P(SPP-b-4VIM)-coated QD (0.5 μM, λemission max = 603 nm) dispersions were stored in buffer solutions spanning the 212 pH range, and their luminescence was observed over time (2 h, 24 h, and 7 days) under UV excitation (350 nm). The following buffers were used: KCl/HCl for pH = 2, citric acid/sodium citrate for 36, Trizma HCl/Trizma base for 79, NaHCO3/NaOH for 1011, and KCl/NaOH for 12. Before each measurement, samples were centrifuged at 19000g during 10 min in order to remove eventual suspended aggregates. After the longest incubation period (7 days), integrated fluorescence intensity (Edinburgh FSP920 spectrometer) and absorbance at 350 nm (UVvisNIR UV-3600 spectrophotometer) were measured and reported normalized by the corresponding value at pH = 7. Photo-oxidation Stability. Photo-oxidation effects on the colloidal stability of ligand-capped QDs were examined by measuring absorbance variations over time of QD dispersions stored under different light and temperature conditions. QD (200 nM, λemission max = 593 nm) dispersions in 100 mM NaCl (pH = 6.5) were tested over a maximum time period of 28 days either under room light and room temperature or in the dark at 37 °C. QDs were capped with the P(SPP20-b-4VIM9) block copolymer ligand or with a random dithiol-appended sulfobetaine copolymer, P(SPP-co-dithiol) (Mn ≈ 5000 g 3 mol1, PDI = 2, [dithiol]/[sulfobetaine] molar ratio = 7/93), as described in Giovanelli et al.13 At each specific storage time, samples were
Acknowledgment. This work was supported by the NanoCTC (ANR-10-Nano-05) grant of the “Investissement d'Avenir” program managed by the French Agence Nationale de la Recherche. The authors thank Dr. Chloé Grazon for helpful discussions about polymer synthesis and Clémence Rogier for initial screening experiments. Supporting Information Available: The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsnano.5b05705. Supplemental methods, characterization of copolymers and capped QDs, nonspecific interactions between live HeLa cells and ligand-capped (QD-ligand) and bioconjugated (QDpAAb) QDs, metabolic activity of HeLa cells after exposure to QDs, CB1 receptor labeling with green and red QDpAanti-FLAG nanoconjugates added simultaneously, FLAGCB1GFP-transfected HEK cells exposed to QDpA and QDpAanti-mouse nanoconstructs, and detailed descriptions of the videos (PDF)
REFERENCES AND NOTES 1. Wegner, K. D.; Jin, Z.; Lindén, S.; Jennings, T. L.; Hildebrandt, N. Quantum-Dot-Based Förster Resonance Energy Transfer Immunoassay for Sensitive Clinical Diagnostics of LowVolume Serum Samples. ACS Nano 2013, 7, 7411–7419. 2. Wegner, K. D.; Hildebrandt, N. Quantum Dots: Bright and Versatile in Vitro and in Vivo Fluorescence Imaging Biosensors. Chem. Soc. Rev. 2015, 44, 4792–4834. 3. Jennings, T. L.; Becker-Catania, S. G.; Triulzi, R. C.; Tao, G.; Scott, B.; Sapsford, K. E.; Spindel, S.; Oh, E.; Jain, V.; Delehanty, J. B.; et al. Reactive Semiconductor Nanocrystals for Chemoselective Biolabeling and Multiplexed Analysis. ACS Nano 2011, 5, 5579–5593.
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23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
33.
34.
35.
36.
37.
38.
39.
Ligands: Impact on Cellular Interactions and Cytotoxicity. J. Mater. Chem. B 2013, 1, 6137–6146. Park, J.; Nam, J.; Won, N.; Jin, H.; Jung, S.; Jung, S.; Cho, S. H.; Kim, S. Compact and Stable Quantum Dots with Positive, Negative, or Zwitterionic Surface: Specific Cell Interactions and Non-Specific Adsorptions by the Surface Charges. Adv. Funct. Mater. 2011, 21, 1558–1566. Breus, V.; Heyes, C.; Tron, K.; Nienhaus, G. Zwitterionic Biocompatible Quantum Dots for Wide pH Stability and Weak Nonspecific Binding to Cells. ACS Nano 2009, 3, 2573–2580. Muro, E.; Pons, T.; Lequeux, N.; Fragola, A.; Sanson, N.; Lenkei, Z.; Dubertret, B. Small and Stable Sulfobetaine Zwitterionic Quantum Dots for Functional Live-Cell Imaging. J. Am. Chem. Soc. 2010, 132, 4556–4557. Muro, E.; Fragola, A.; Pons, T.; Lequeux, N.; Ioannou, A.; Skourides, P.; Dubertret, B. Comparing Intracellular Stability and Targeting of Sulfobetaine Quantum Dots with Other Surface Chemistries in Live Cells. Small 2012, 8, 1029–1037. Chen, X.; Lawrence, J.; Parelkar, S.; Emrick, T. Novel Zwitterionic Copolymers with Dihydrolipoic Acid: Synthesis and Preparation of Nonfouling Nanorods. Macromolecules 2013, 46, 119–127. Liu, W.; Choi, H. S.; Zimmer, J. P.; Tanaka, E.; Frangioni, J. V.; Bawendi, M. Compact Cysteine-Coated CdSe(ZnCdS) Quantum Dots for in Vivo Applications. J. Am. Chem. Soc. 2007, 129, 14530–14531. Soo Choi, H.; Liu, W.; Misra, P.; Tanaka, E.; Zimmer, J. P.; Itty Ipe, B.; Bawendi, M. G.; Frangioni, J. V. Renal Clearance of Quantum Dots. Nat. Biotechnol. 2007, 25, 1165–1170. Sun, M.; Hoffman, D.; Sundaresan, G.; Yang, L.; Lamichhane, N.; Zweit, J. Synthesis and Characterization of Intrinsically Radio-Labeled Quantum Dots for Bimodal Detection. Am. J. Nucl. Med. Mol. Imaging 2012, 2, 122–135. Yang, W.; Zhang, L.; Wang, S.; White, A. D.; Jiang, S. Functionalizable and Ultra Stable Nanoparticles Coated with Zwitterionic Poly(carboxybetaine) in Undiluted Blood Serum. Biomaterials 2009, 30, 5617–5621. Zhou, W.; Shao, J.; Jin, Q.; Wei, Q.; Tang, J.; Ji, J. Zwitterionic Phosphorylcholine as a Better Ligand for Gold Nanorods Cell Uptake and Selective Photothermal Ablation of Cancer Cells. Chem. Commun. (Cambridge, U. K.) 2010, 46, 1479–1481. Banerjee, A.; Grazon, C.; Nadal, B.; Pons, T.; Krishnan, Y.; Dubertret, B. Fast, Efficient, and Stable Conjugation of Multiple DNA Strands on Colloidal Quantum Dots. Bioconjugate Chem. 2015, 26, 1582–1589. Tasso, M.; Singh, M.; Giovanelli, E.; Fragola, A.; Loriette, V.; Regairaz, M. F.; Dautry, F.; Treussart, F.; Lenkei, Z.; Lequeux, N.; et al. Oriented Bioconjugation of Unmodified Antibodies to Quantum Dots Generates Versatile Cellular Imaging Tools. ACS Appl. Mater. Interfaces 2015, Submitted. Medintz, I. L.; Clapp, A. R.; Mattoussi, H.; Goldman, E. R.; Fisher, B.; Mauro, J. M. Self-Assembled Nanoscale Biosensors Based on Quantum Dot FRET Donors. Nat. Mater. 2003, 2, 630–638. Clapp, A. R.; Medintz, I. L.; Mauro, J. M.; Fisher, B. R.; Bawendi, M. G.; Mattoussi, H. Fluorescence Resonance Energy Transfer between Quantum Dot Donors and Dye-Labeled Protein Acceptors. J. Am. Chem. Soc. 2004, 126, 301–310. Delehanty, J. B.; Medintz, I. L.; Pons, T.; Brunel, F. M.; Dawson, P. E.; Mattoussi, H. Self-Assembled Quantum Dot- Peptide Bioconjugates for Selective Intracellular Delivery. Bioconjugate Chem. 2006, 17, 920–927. Sapsford, K. E.; Pons, T.; Medintz, I. L.; Higashiya, S.; Brunel, F. M.; Dawson, P. E.; Mattoussi, H. Kinetics of Metal-Affinity Driven Self-Assembly between Proteins or Peptides and CdSe-ZnS Quantum Dots. J. Phys. Chem. C 2007, 111, 11528–11538. Chiefari, J.; Chong, Y. K. B.; Ercole, F.; Krstina, J.; Jeffery, J.; Le, T. P. T.; Mayadunne, R. T. A.; Meijs, G. F.; Moad, C. L.; Moad, G.; et al. Living Free-Radical Polymerization by Reversible Addition - Fragmentation Chain Transfer: The RAFT Process. Macromolecules 1998, 31, 5559–5562.
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4. Michalet, X.; Pinaud, F. F.; Bentolila, L. A.; Tsay, J. M.; Doose, S.; Li, J. J.; Sundaresan, G.; Wu, A. M.; Gambhir, S. S.; Weiss, S. Quantum Dots for Live Cells, in Vivo Imaging, and Diagnostics. Science 2005, 307, 538–544. 5. Zhang, F.; Lees, E.; Amin, F.; Rivera-Gil, P.; Yang, F.; Mulvaney, P.; Parak, W. J. Polymer-Coated Nanoparticles: A Universal Tool for Biolabelling Experiments. Small 2011, 7, 3113– 3127. 6. Palui, G.; Aldeek, F.; Wang, W.; Mattoussi, H. Strategies for Interfacing Inorganic Nanocrystals with Biological Systems Based on Polymer-Coating. Chem. Soc. Rev. 2015, 44, 193– 227. 7. Uyeda, H. T.; Medintz, I. L.; Jaiswal, J. K.; Simon, S. M.; Mattoussi, H. Synthesis of Compact Multidentate Ligands to Prepare Stable Hydrophilic Quantum Dot Fluorophores. J. Am. Chem. Soc. 2005, 127, 3870–3878. 8. Susumu, K.; Uyeda, H. T.; Medintz, I. L.; Pons, T.; Delehanty, J. B.; Mattoussi, H. Enhancing the Stability and Biological Functionalities of Quantum Dots via Compact Multifunctional Ligands. J. Am. Chem. Soc. 2007, 129, 13987–13996. 9. Liu, W.; Howarth, M.; Greytak, A. B.; Zheng, Y.; Nocera, D. G.; Ting, A. Y.; Bawendi, M. G. Compact Biocompatible Quantum Dots Functionalized for Cellular Imaging. J. Am. Chem. Soc. 2008, 130, 1274–1284. 10. Yildiz, I.; McCaughan, B.; Cruickshank, S. F.; Callan, J. F.; Raymo, F. M. Biocompatible CdSe-ZnS Core-Shell Quantum Dots Coated with Hydrophilic Polythiols. Langmuir 2009, 25, 7090–7096. 11. Stewart, M. H.; Susumu, K.; Mei, B. C.; Medintz, I. L.; Delehanty, J. B.; Blanco-Canosa, J. B.; Dawson, P. E.; Mattoussi, H. Multidentate Poly(ethylene Glycol) Ligands Provide Colloidal Stability to Semiconductor and Metallic Nanocrystals in Extreme Conditions. J. Am. Chem. Soc. 2010, 132, 9804–9813. 12. Zhan, N.; Palui, G.; Safi, M.; Ji, X.; Mattoussi, H. Multidentate Zwitterionic Ligands Provide Compact and Highly Biocompatible Quantum Dots. J. Am. Chem. Soc. 2013, 135, 13786–13795. 13. Giovanelli, E.; Muro, E.; Sitbon, G.; Hanafi, M.; Pons, T.; Dubertret, B.; Lequeux, N. Highly Enhanced Affinity of Multidentate versus Bidentate Zwitterionic Ligands for Long-Term Quantum Dot Bioimaging. Langmuir 2012, 28, 15177–15184. 14. Nagaraja, A. T.; Sooresh, A.; Meissner, K. E.; McShane, M. J. Processing and Characterization of Stable, pH-Sensitive Layer-by-Layer Modified Colloidal Quantum Dots. ACS Nano 2013, 7, 6194–6202. 15. Aldana, J.; Wang, Y. A.; Peng, X. Photochemical Instability of CdSe Nanocrystals Coated by Hydrophilic Thiols. J. Am. Chem. Soc. 2001, 123, 8844–8850. 16. Han, H.-S.; Martin, J. D.; Lee, J.; Harris, D. K.; Fukumura, D.; Jain, R. K.; Bawendi, M. Spatial Charge Configuration Regulates Nanoparticle Transport and Binding Behavior in Vivo. Angew. Chem., Int. Ed. 2013, 52, 1414–1419. 17. Susumu, K.; Oh, E.; Delehanty, J. B.; Pinaud, F.; Gemmill, K. B.; Walper, S.; Breger, J.; Schroeder, M. J.; Stewart, M. H.; Jain, V.; et al. A New Family of Pyridine-Appended Multidentate Polymers As Hydrophilic Surface Ligands for Preparing Stable Biocompatible Quantum Dots. Chem. Mater. 2014, 26, 5327–5344. 18. Wang, W.; Kapur, A.; Ji, X.; Safi, M.; Palui, G.; Palomo, V.; Dawson, P. E.; Mattoussi, H. Photoligation of an Amphiphilic Polymer with Mixed Coordination Provides Compact and Reactive Quantum Dots. J. Am. Chem. Soc. 2015, 137, 5438–5451. 19. Estephan, Z. G.; Schlenoff, P. S.; Schlenoff, J. B. Zwitteration as an Alternative to PEGylation. Langmuir 2011, 27, 6794–6800. 20. Shao, Q.; Jiang, S. Molecular Understanding and Design of Zwitterionic Materials. Adv. Mater. 2015, 27, 15–26. 21. Keefe, A. J.; Jiang, S. Poly(zwitterionic)protein Conjugates Offer Increased Stability without Sacrificing Binding Affinity or Bioactivity. Nat. Chem. 2012, 4, 59–63. 22. Sun, M.; Yang, L.; Jose, P.; Wang, L.; Zweit, J. Functionalization of Quantum Dots with Multidentate Zwitterionic
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40. Lowe, A. B.; McCormick, C. L. Synthesis and Solution Properties of Zwitterionic Polymers. Chem. Rev. 2002, 102, 4177–4189. 41. McCormick, C. L.; Lowe, A. B. Aqueous RAFT Polymerization: Recent Developments in Synthesis of Functional Water-Soluble (co)polymers with Controlled Structures. Acc. Chem. Res. 2004, 37, 312–325. 42. Rodriguez-Emmenegger, C.; Schmidt, B. V. K. J.; Sedlakova, Z.; Subr, V.; Alles, A. B.; Brynda, E.; Barner-Kowollik, C. Low Temperature Aqueous Living/controlled (RAFT) Polymerization of Carboxybetaine Methacrylamide up to High Molecular Weights. Macromol. Rapid Commun. 2011, 32, 958–965. 43. Arotc-aréna, M.; Heise, B.; Ishaya, S.; Laschewsky, A. Switching the inside and the outside of Aggregates of WaterSoluble Block Copolymers with Double Thermoresponsivity. J. Am. Chem. Soc. 2002, 124, 3787–3793. 44. Durand-Gasselin, C.; Koerin, R.; Rieger, J.; Lequeux, N.; Sanson, N. Colloidal Stability of Zwitterionic PolymerGrafted Gold Nanoparticles in Water. J. Colloid Interface Sci. 2014, 434, 188–194. 45. Sugihara, S.; Blanazs, A.; Armes, S. P.; Ryan, A. J.; Lewis, A. L. Aqueous Dispersion Polymerization: A New Paradigm for in Situ Block Copolymer Self-Assembly in Concentrated Solution. J. Am. Chem. Soc. 2011, 133, 15707–15713. 46. Willcock, H.; Lu, A.; Hansell, C. F.; Chapman, E.; Collins, I. R.; O'Reilly, R. K. One-Pot Synthesis of Responsive Sulfobetaine Nanoparticles by RAFT Polymerisation: The Effect of Branching on the UCST Cloud Point. Polym. Chem. 2014, 5, 1023. 47. Thomas, D. B.; Convertine, A. J.; Hester, R. D.; Lowe, A. B.; McCormick, C. L. Hydrolytic Susceptibility of Dithioester Chain Transfer Agents and Implications in Aqueous RAFT Polymerizations. Macromolecules 2004, 37, 1735–1741. 48. Allen, M. H.; Hemp, S. T.; Smith, A. E.; Long, T. E. Controlled Radical Polymerization of 4-Vinylimidazole. Macromolecules 2012, 45, 3669–3676. 49. Mary, P.; Bendejacq, D. D.; Labeau, M.-P.; Dupuis, P. Reconciling Low- and High-Salt Solution Behavior of Sulfobetaine Polyzwitterions. J. Phys. Chem. B 2007, 111, 7767–7777. 50. Li, J. J.; Wang, Y. A.; Guo, W.; Keay, J. C.; Mishima, T. D.; Johnson, M. B.; Peng, X. Large-Scale Synthesis of Nearly Monodisperse CdSe/CdS Core/shell Nanocrystals Using Air-Stable Reagents via Successive Ion Layer Adsorption and Reaction. J. Am. Chem. Soc. 2003, 125, 12567–12575. 51. Montenegro, J. M.; Grazu, V.; Sukhanova, A.; Agarwal, S.; de la Fuente, J. M.; Nabiev, I.; Greiner, A.; Parak, W. J. Controlled Antibody/(bio-) Conjugation of Inorganic Nanoparticles for Targeted Delivery. Adv. Drug Delivery Rev. 2013, 65, 677–688. 52. Bae, W. K.; Char, K.; Hur, H.; Lee, S. Single-Step Synthesis of Quantum Dots with Chemical Composition Gradients. Chem. Mater. 2008, 20, 531–539. 53. Leterrier, C.; Bonnard, D.; Carrel, D.; Rossier, J.; Lenkei, Z. Constitutive Endocytic Cycle of the CB1 Cannabinoid Receptor. J. Biol. Chem. 2004, 279, 36013–36021. 54. Simon, A. C.; Loverdo, C.; Gaffuri, A.-L.; Urbanski, M.; Ladarre, D.; Carrel, D.; Rivals, I.; Leterrier, C.; Benichou, O.; Dournaud, P.; et al. Activation-Dependent Plasticity of Polarized GPCR Distribution on the Neuronal Surface. J. Mol. Cell Biol. 2013, 5, 250–265. 55. Leterrier, C.; Lainé, J.; Darmon, M.; Boudin, H.; Rossier, J.; Lenkei, Z. Constitutive Activation Drives CompartmentSelective Endocytosis and Axonal Targeting of Type 1 Cannabinoid Receptors. J. Neurosci. 2006, 26, 3141–3153. 56. Overberger, C. G.; Vorchheimer, N. Imidazole-Containing Polymers. Synthesis and Polymerization of the Monomer 4(5)-Vinylimidazole. J. Am. Chem. Soc. 1963, 85, 951–955. 57. Yu, W. W.; Peng, X. Formation of High-Quality CdS and Other II-VI Semiconductor Nanocrystals in Noncoordinating Solvents: Tunable Reactivity of Monomers. Angew. Chem., Int. Ed. 2002, 41, 2368–2371.
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