We prepared hybrid nanoparticles with a brightly fluorescentsilica core and a biocompatible thermoresponsive polymershell containing tumor-targeting folic acid (FA) groups. Thesilica core has a perylenediimide fluorescent dye anchoredcovalently to the structure for traceability and bio-imaging applications. The polymeric shell was synthesizedby reversible addition–fragmentation chain-transfer (RAFT)polymerization to guarantee the homogeneous size of theparticle shell. The shell is composed of copolymer chainswith one block of oligo(ethylene glycol)methacrylate and 2-(2�-methoxyethoxy)ethylmethacrylate with another block ofthe reactive monomer N-acryloxysuccinimide (NAS). TheNAS groups were used to covalently attach a large amount
Introduction
The efficient delivery of a drug, precisely at the desiredtime and location, can largely decrease its side effects andincrease its therapeutic efficiency.[1] One possible approachto achieve this goal is to use nanoparticles that combinetherapeutic and diagnostic (theranostic) functionalities andfeature large drug payloads, active targeting, and stimuli-activated drug release.[2]
Hybrid nanoparticles have attracted much attention ow-ing to their potential biomedical applications in diagnosticand precision drug delivery for cancer detection and treat-ment.[3] The inorganic core can be used to transport cargo(e.g., in capsules or mesoporous materials) and impart
[a] CQFM – Centro de Química-Física Molecular and IN –Institute of Nanoscience and Nanotechnology, InstitutoSuperior Técnico, Universidade de Lisboa,1049-001 Lisboa, PortugalE-mail: [email protected]
[b] Instituto de Investigação do Medicamento (iMed.ULisboa),Faculdade de Farmácia, Universidade de Lisboa,Av. Prof. Gama Pinto, 1649-003 Lisboa, PortugalSupporting information for this article is available on theWWW under http://dx.doi.org/10.1002/ejic.201500580.
of amino-functionalized FA to the particle shells to provideactive targeting properties towards cancer cells and tissues.The targeting capability of the folate-containing nanopar-ticles was evaluated against NCI-H460 tumor cells, whichoverexpress folate receptors. The nanoparticles with FAshow a higher uptake efficiency compared to that of theequivalent nanoparticles without FA. This result validatesthe imaging capabilities and active targeting efficiency of ournanoparticles, an important step towards the goal of de-veloping vehicles for precision drug delivery systems thatcombine therapeutic and diagnostic (theranostic) functionali-ties with large drug payloads, active targeting, and stimuli-activated drug release.
properties such as magnetism, fluorescence, and plasmonicabsorbance, whereas the organic part can be used to conveywater solubility, biocompatibility, long blood circulationtimes, and bioconjugation.[4] Among inorganic materials,silica is specially adequate for biomedical applications be-cause it is biocompatible, endogenous (present in bones),and generally recognized as safe by the U.S. Food and DrugAdministration (US-FDA).[4] Silica nanoparticles (SNPs)are not only easy to synthesize with controlled diameterand low size dispersion but also amiable to rich conjugationchemistry, which allows the incorporation of differentgroups into the silica network, such as fluorescent dyes (toimprove the photophysical and photochemical performanceof the dyes).[5–9] Owing to these characteristics, SNPs havebeen used as supports or carriers for drug delivery[10] andimaging agents.[11]
The covalent incorporation of dyes into SNPs by usingmolecules functionalized with alkoxysilane groups is an ef-ficient strategy to avoid aggregation and self-quenching ef-fects resulting in very high brightness per particle for SNPs.Another advantage of the incorporation of dyes into SNPsis that the oxygen shielding effect (the oxygen concentrationand diffusivity in SNPs are lower than those in most sol-vents) leads to low photobleaching, even under laser illumi-nation.[6–8]
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Perylenediimide (PDI) derivatives are generally verysturdy and bright dyes with high fluorescence quantumyields, excitation tunable from the visible to the near-infra-red (NIR), high photochemical stability, and high electronmobility.[12] These properties can be modulated by substitu-tion at the imide group (to tune the solubility, reduce aggre-gation, or allow conjugation to different materials)[6–8,13] orin the bay region of the perylene core (which affects theelectronic and optical properties).[14] The introduction ofalkoxysilane groups at the imide groups of PDI allows thecoupling of the dyes to silica to anchor them to the silicastructure.
By functionalizing the surfaces of SNPs with polymericshells, one gains another dimension for tuning the particleproperties, such as their stability in different solvents, bio-compatibility, blood circulation times, and the conjugationof different biologically active components. One particu-larly interesting option is to incorporate functional mol-ecules for the active targeting of cancer cells or tissues.[15,16]
In this case, it is important that the composition and sizeof the polymer shell is homogeneous. This high degree ofcontrol can be achieved by reversible addition–fragmenta-tion chain-transfer (RAFT) polymerization, a form of liv-ing radical polymerization that can be readily implementedin nanoparticles, uses nontoxic materials, and is very versa-tile in the monomers that can be used.[17,18] To grow poly-mer chains from the core particle, one can functionalize theinorganic surface with aminopropyltriethoxysilane(APTES) and then covalently attach a carboxylic-contain-ing RAFT chain transfer agent (CTA). This “graftingfrom” method produces a polymer shell with high graftingdensity and controlled thickness.[19]
The use of thermoresponsive polymers to coat SNPs of-fers the possibility to control the expanded or collapsedstate of the polymer in water by changing the tempera-ture[20,21] and, thus, provides a strategy to control the re-lease of therapeutic agents encapsulated in the carrier. Co-polymers of 2-(2�-methoxyethoxy)ethyl methacrylate (ME-O2MA) and oligo(ethylene glycol) methacrylate (OEGMA)are thermoresponsive and especially suitable for biomedicalapplications because they are biocompatible, have good sol-ubility in physiological media, prevent nonspecific proteinadsorption and cell adhesion, and exhibit lower criticalsolution temperatures (LCSTs) that can be accurately tunedfrom 28 to 90 °C by adjusting the ratio of the two mono-mers.[22,23]
The conjugation of these copolymers with bioactivegroups can be achieved through two strategies. One is touse the chain end group (which can be hydrolyzed to a thiolfor RAFT polymers)[24] or, alternatively, one can polymer-ize a second block of a reactive comonomer, such as N-acryloxysuccinimide (NAS).[25] This reactive block can thenbe used to attach antibodies, oligopeptides, folic acid, orother targeting groups onto the outer shell of the nanopart-icle.
Nanocarriers for therapeutic applications can have abroad range of functionalities, for example, imaging agentsfor diagnostics, targeting groups to promote their accumu-
lation at the target location, or both.[26–35] Folic acid (FA) isa targeting group of particular interest because it is readilyavailable and stable over a broad range of temperatures andpH values, does not trigger an immunologic response, andretains its ability to specifically bind to folate receptors,even when linked to other molecules.[15] Folic acid conju-gated nanoparticles bind to cell membrane folate receptorsby an active targeting mechanism.[36,37] As folate receptorsare overexpressed in the membranes of many cancer celltypes and tissues, including ovarian, endometrial, and kid-ney cancer,[36–39] folic acid conjugated nanoparticles carry-ing therapeutic agents are expected to increase their deliveryefficiency and avoid side effects, as the exposure of healthycells to the anticancer agent can be minimized.
The goal of our work is to develop “smart” hybrid nano-particles with theranostic functionalities that carry a fluo-rescent dye for traceability and imaging, feature a mecha-nism for on-demand release control, and are able to accom-modate large drug loads and deliver their cargo to a desiredlocation. To this end, the present work focuses on the prep-aration of fluorescent SNPs with a biocompatible copoly-mer shell, which has an outermost block heavily decoratedwith FA (Figure 1). We prepared monodisperse fluorescentSNPs incorporating a perylenediimide (PDI) derivative inthe silica network,[6–8] and the SNPs were encapsulated ina thermoresponsive polymer shell with an outer block ofreactive NAS groups that can be conjugated to a largeamount of amino-modified FA. The large amount of outerFA groups is expected to promote the efficient active tar-geting of cancer cells and tissues.[38,39] The uptake of nano-particles with and without FA was tested in human non-small-cell lung cancer cells (NCI-H460), which are FA-re-ceptor-positive cells.
Figure 1. Schematic representation of the multifunctional nanopar-ticle with a silica core (blue) and a shell consisting of a copolymerwith a thermoresponsive block of MEO2MA and OEGMA (blue)as well as an outer block of the reactive monomer NAS (grey), towhich the folic acid derivative (FA-HDA) is attached (yellow). Thefinal polymer shell is obtained by hydrolyzing the unreacted NASgroups to yield carboxylic groups, which are negatively charged atphysiological pH.
In the context of our project, we can substitute the silicacore of the particles presented here by a highly fluorescent
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mesoporous support[13] that will provide the large cargo ca-pacity desirable for theranostic applications.[10,40] Thesenanoparticles, featuring stimuli-responsive controlled re-lease and active targeting functionalities are excellent candi-dates for theranostic applications.[41–43]
Results and Discussion
Fluorescent Silica Nanoparticles
Brightly fluorescent silica nanoparticles (SNP-PDI) wereprepared by a modified Stöber method[6] with the PDI de-rivative 1, which contains two terminal triethoxysilyl groups(Figure 2) that allow its incorporation into the silica net-work during the particle synthesis. The TEM image ofSNP-PDI (Figure 3, A) shows that the nanoparticles arespherical and have very small size dispersity with a diameterof (110� 12) nm (Figure 3, B).
The fluorescence emission and excitation spectra of SNP-PDI dispersed in ethanol (Figure 4, solid line) are similarto those of free PDI in ethanol (Figure 4, dashed line);therefore, our method for the incorporation of the dye intothe SNP network is able to avoid dye aggregation andquenching.[7,9]
Surface Modification of the Silica Nanoparticles
In the synthesis pathway to modify the SNP-PDI surface(Figure 5), APTES was used to add amine groups to theexternal surface of the nanoparticles. The amine groupswere further reacted with the carboxylic group of theRAFT agent (3, Figure 2) through carbodiimide chemistry.The RAFT agent at the nanoparticle surface allows the
Figure 3. (A) TEM image of SNP-PDI (scale bar: 200 nm) and (B)the corresponding diameter histogram obtained by measuring ap-proximately 150 nanoparticles.
Figure 4. Normalized absorption (green) and fluorescence (blue)spectra of PDI (dashed line) and SNP-PDI (solid line) in ethanol.
polymer chains to grow in a “grafting from” approach dur-ing the polymerization reaction.
The zeta potentials of the SNP-PDI, SNP-APTES, andSNP-RAFT samples provide qualitative information aboutthe surface charges of the nanoparticles, a clear indicationof the surface modification of the silica nanoparticles. Theaverage zeta potential values were obtained for dispersionsof nanoparticles in milli-Q water (pH ≈ 5.5). For SNP-PDI(without surface modification), the zeta potential was nega-tive (–47�8) mV. The isoelectric point of the silanol groupsappears at pH 1.5;[44] therefore, the silanol groups are de-protonated in milli-Q water, and a negative value is ob-tained. However, after surface functionalization by APTES,a positive value of (15� 8) mV was observed owing to thepKa of the aminopropyl group (9.8) and its isoelectric point
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Figure 5. Schematic representation of the reactions to obtain the hydrolyzed SNP-poly-FA. The SNP-PDI surface is first modified withAPTES, which is treated with the RAFT agent. This is followed by polymerization of NAS, MEO2MA and OEGMA at the surface ofthe SNP-RAFT. The reactive NAS moieties are then used to attach the folic acid derivative (FA-HDA), and finally the unreacted NASgroups are hydrolyzed yielding negative charges at the polymer chains.
(10.6).[44] The substitution of the amine groups by theRAFT agent resulted in a change of zeta potential to(–21� 10) mV, which indicates the success of the modifica-tion.
The surface density of the RAFT groups in SNP-RAFTwas determined from the absorbance spectrum of SNP-RAFT dispersed in dioxane (Figure 6) to estimate theamount of polymer chains growing from the surface of thenanoparticles. By using the molar absorptivity obtained forthe free RAFT agent in dioxane at λ = 310 nm (ε =13.98 mm–1 cm–1, Section S1, Supporting Information) andcorrecting for the light scattered by the nanoparticle (bysubtracting the 1/λ4 scattering profile from the absorptionadjusted below 275 nm and between 360 and 430 nm, Fig-ure 6), we obtained a RAFT agent surface density of
Figure 6. Absorption spectrum of SNP-RAFT dispersed in dioxane(solid line) and the curve used to subtract the scattering effect ofthe nanoparticles (dashed line).
one group per nm2, a value higher than that reported pre-viously.[45] This allowed us to obtain a high surface densityof polymer chains, which is important for their future useas a gatekeeper to regulate the drug release from the nano-particles.
RAFT Polymerization
The diblock copolymer was prepared by sequentialRAFT polymerization with a first block of reactive N-acryloxysuccinimide (NAS, 2, Figure 2), an activated esterderivative that can be used to attach amino-derived com-pounds to the polymer chain.[25] The second block was pre-pared by adding a mixture of the monomers 2-(2�-methoxy-ethoxy)ethyl methacrylate (MEO2MA, 4, Figure 2) and oli-go(ethylene glycol) methacrylate (OEGMA, 5, Figure 2).The resulting block copolymer exhibits thermoresponsivebehavior with a lower critical solution temperature (LCST)that can be tuned from 26 to 90 °C by changing theMEO2MA/OEGMA monomer ratio.[22]
The polymerization kinetics were followed by 1H NMRspectroscopy (Figure 7), and the overall monomer conver-sion was nonlogarithmic; the NAS conversion followed theexpected logarithmic behavior but stabilized at 35% conver-sion. The addition of the second block monomers (dashedline, Figure 7) unexpectedly increased the RAFT chaintransfer agent efficiency, and the polymerization proceededuntil almost the total conversion of all monomers. The datain Figure 7 indicate that the grafted copolymer has an outerblock of NAS (with ca. 35 % of the total NAS groups) andan inner statistical block composed of a mixture of the eth-
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ylene glycol acrylate monomers and a decreasing amountof NAS towards the surface of the particle. As NAS reaches100% conversion before the other monomers, there is a lastblock without NAS that contains ca. 20% of the totalamount of MEO2MA/OEGMA monomers. The core–shellstructure of the SNP-poly was confirmed by transmissionelectron microscopy (Figure 8), which showed a soft poly-mer layer surrounding the silica core.
Figure 7. Time-dependent monomer consumption during RAFTpolymerization of the polymer prepared in solution: NAS mono-mer conversion (circles), MEO2MA/OEGMA monomers conver-sion (squares). The dashed line indicates the addition of the secondblock monomers.
Figure 8. TEM image of SNP-poly. Scale bar: 500 nm.
During the polymerization, a small amount of freeRAFT agent in solution was used to obtain free polymerchains that are equivalent to the chains anchored to theparticle surface but can be readily separated from the par-ticles and used to determine the polymer molecular weightand size dispersity by gel permeation chromatographymultiangle light scattering (GPC-MALS; Section S3, Sup-porting Information). This has also been reported to leadto a better control over the polymerization.[46]
The calculated and measured molecular weights and sizedispersities of the polymer chains obtained for the differentpolymerizations (both in solution and at the SNPs) arecompiled in Table 1. The experimental number average mo-lecular weight (Mn) and weight average molecular weight(Mw) are higher than the calculated values, which indicatessome degree of dead chains, as is normal in RAFT. How-ever, the difference for the SNP-grafted chains is too high,an indication that the chains obtained from the free RAFTagent added to the SPN-RAFT are probably able to grow
more than the tethered chains owing to steric hindrance atthe particle surface. The low size dispersity (Mw/Mn) values(1.15 for SNP-poly and 1.09 for the free polymer chains)indicate that the control over the polymerization is good.
Table 1. Molecular weights and size dispersion of the polymerchains obtained in solution and the free polymer chains grown inthe SNP-poly polymerization.
The temperature-responsive behavior of the polymers inwater was assessed by measuring the variation of transmit-tance with temperature for the free polymer chains (Fig-ure 9, A) and for the free polymer chains obtained duringthe polymerization of the SNPs (Figure 9, B). The tempera-ture variation induces a change in the conformation of thepolymer chains from an expanded conformation below theLCST to a collapsed state at temperatures higher than theLCST with no hysteresis between the heating and coolingcycles (Figure 9). The LCST of the chains obtained in thesolution polymerization was 50 °C, and that of those grownduring the polymerization at the SNPs was 52 °C. The re-ported LCST for copolymers with the same OEGMA/MEO2MA ratio is 49 °C.[22] The small difference betweenthe values is probably due to the NAS block and the sizedifference between the samples.[47]
Figure 9. Variation of transmittance (at 500 nm) with temperaturefor the heating (circles) and cooling (squares) cycles of (A) the poly-mer chains synthesized in solution (0.27% w/v aqueous solution)and (B) the polymer chains grown in solution in the SNP-poly poly-merization reaction (0.24% w/v aqueous solution). The dashedlines represent the LCST values.
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Polymeric Shell Functionalization
Our strategy is to functionalize the SNPs with a largeamount of folic groups to achieve efficient active targetingof cancer cells and tissues. The FA-HDA derivative was at-tached to the NAS reactive ester groups of the SNPs poly-mer shell (Figure 5) by using an excess of FA relative toNAS. The expected number of NAS molecules per polymerchain was calculated from the [NAS]/[CTA + AIBN] ratio[AIBN = 2,2�-azobis(2-methylpropionitrile)] used in the po-lymerization, and the number of chains per particle wasconsidered equal to the determined number of RAFTgroups per particle. The SNP-poly-FA was separated fromfree unreacted FA-HDA by successive centrifugation andredispersion in N,N-dimethylformamide (DMF).
Cellular Uptake
Folic acid is recognized by folate receptors (FR), whichare overexpressed in cancer cells. To evaluate the targetingperformance of our folate-loaded nanoparticles, we usedthe human lung cancer cell line NCI-H460 as a model.
SNP-poly-FA and SNP-poly (as a control) were incu-bated with NCI-H460 cells, and the uptake of the nanopar-ticles was followed by confocal laser scanning microscopy.The nanoparticles were easily traced owing to the fluores-cent PDI incorporated in the silica structure of the core.
Figure 10. Confocal fluorescence images of NCI-H460 cancer cellsstained with AF594-WGA plasma membrane marker (yellow) andincubated at different times with nanoparticles containing no folategroups (SNP-poly: A, C, E, and G, shown in red) and folate-loadedSNP-poly-FA (B, D, F, and H, shown in red). Images dimensions:100�100 μm.
The uptake of SNP-poly-FA and SNP-poly was followed atdifferent incubation times (2, 4, 6, and 16 h), and the cellswere stained with AF594-WGA plasma membrane markerbefore confocal imaging (Figure 10).
A simple comparison between the two sets of nanopar-ticles shows that the uptake of SNP-poly-FA is faster (Fig-ure 10, B, D, F, and H) than the uptake of SNP-poly (Fig-ure 10, A, C, E, and G). The uptake of SNP-poly-FA isimmediately visible at 2 h (Figure 10, B) and increases withtime; at 16 h, the nanoparticles have been very efficientlyinternalized by the cells and appear to be dispersed homo-geneously in the cytosol (Figure 10, G). For the particleswithout folate, SNP-poly, no uptake is visible at 2 h (Fig-ure 10, A), and a small degree of internalization is visibleafter 4 and 6 h (Figure 10, C and E). The difference be-tween the uptake of SNP-poly-FA and SNP-poly is mostclear at 16 h incubation (Figure 10, G and H). The largeincrease in internalization efficiency of the folate-loadedparticles, SNP-poly-FA, compared with that their unmodi-fied counterparts, SNP-poly, is probably due to a folate-receptor-mediated endocytosis process, which leads to in-creased internalization of the nanoparticles containing folicacid.
Conclusions
We were able to prepare multifunctional core–shell hy-brid silica nanoparticles with fluorescent traceability and alarge amount of folate groups in the polymer shell. Thebrightly fluorescent silica core has very low size dispersityand was functionalized with a smart block copolymer,grown from the particle surface by RAFT polymerizationto form a dense polymer shell with ca. 1 polymer chain pernm2 at the surface. The shell copolymer is thermoresponsiveand assumes expanded and collapsed conformations uponcooling and heating with no detected hysteresis. It containsethylene glycol side groups that modulate the particle inter-actions, improve colloidal stability and biocompatibility,and protect the particle from recognition by the immunesystem; thus, circulation times in the blood should be in-creased. The copolymer also contains NAS activated estermonomers in increasing amount from the particle surface;these groups were used to anchor multiple folic acid groups,in contrast with the approaches reported previously withonly one folic acid group per polymer chain. The uptakeexperiments with NCI-H46 cancer cells have shown clearlythat the folate-loaded nanoparticles are internalized fasterand in a larger amount than the control counterparts with-out folate groups. The large amount of folic groups per par-ticle provide very efficient targeting of cancer cells that ex-press increased amounts of the folate receptor, a good indi-cation of probable preferential accumulation of the nano-particles at cancer tissues.
Our results show that the developed hybrid nanoparticlesare readily traceable owing to their high brightness and ef-ficiently target the folate receptors overexpressed in somecancer cells. These are important characteristics in carrier
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systems for precision therapy and diagnosis. Our ultimategoal is to combine therapeutic and diagnostic (theranostic)functionalities in nanoparticles with large drug load ca-pacities that carry fluorescent beacons for traceability andimaging, and feature targeting and smart release controlmechanisms that allow the delivery of their cargo on-de-mand at the desired location.
Experimental Section
Materials: Tetraethylorthosilicate (TEOS, 98%, Sigma–Aldrich),(3-aminopropyl)triethoxysilane (APTES, 98%, Sigma–Aldrich),toluene (96%, Sigma–Aldrich), dichloromethane (96%, Sigma–Al-drich), N-(3-dimethylaminopropyl)-N�-ethylcarbodiimide (EDC,98 %, Sigma–Aldrich), N-acryloxysuccinimide (NAS, �90%,Sigma–Aldrich), oligo(ethylene glycol) methacrylate (OEGMA,98%, Sigma–Aldrich), 2-(2�-methoxyethoxy)ethyl methacrylate(MEO2MA, 95%, Sigma–Aldrich), absolute ethanol (99.9%,Scharlau), dimethylformamide (DMF, 99.8%, Scharlau), dimethylsulfoxide (DMSO, 99.9%, Acros Organics), 2,2�-azobis(2-methyl-propionitrile) (AIBN, 99%, Sigma–Aldrich), anhydrous pyridine(99.8%, Sigma–Aldrich), folic acid (FA, �97%, Sigma–Aldrich),N-Boc-1,6-hexanediamine (Boc-HDA, Boc = tert-butyloxy-carbonyl, 98%, Sigma–Aldrich), N,N-dicyclohexylcarbodiimide(DCC, �97%, Sigma–Aldrich), trifluoroacetic acid (TFA, 99 %,Sigma–Aldrich), ammonium hydroxide solution (25% in water,Fluka), and deuterated dimethyl sulfoxide ([D6]DMSO, 99%, Cam-bridge Isotope Laboratories Inc.) were used without further purifi-cation. Deionized water from a Millipore Milli-Q system was usedfor the preparation of dispersions. Bis(propyl)triethoxysilane per-ylenediimide (PDI)[5] and 3-(benzylsulfanylthiocarbonylsulfanyl)-propionic acid (RAFT agent)[48] were synthesized according to theliterature procedures.
For cellular incubation, sterile-filtered poly-l-lysine solution (mo-lecular weight 70000–150000), RPMI-1640 medium with and with-out phenol red, antibiotic antimycotic solution, and trypsin–ethyl-enediaminetetraacetic acid (trypsin–EDTA) solution were obtainedfrom Sigma–Aldrich. Hydrophobic, uncoated, and sterile μ-Slideeight-well plates were obtained from Ibidi GmbH. WGA-Alexa 594was purchased from Invitrogen. Inactivated fetal bovine serum(FBS) was obtained from Alfagene. For cellular uptake tests, a hu-man non-small-cell lung cancer cell line (NCI-H460) was pur-chased from ATCC.
Synthesis of Fluorescent Silica Nanoparticles (SNP-PDI): Fluores-cent silica nanoparticles were synthesized by a modified Stöbermethod.[6] In a typical synthesis, absolute ethanol (40 mL, contain-ing 1.26�10–6 mol of PDI) and NH4OH (2.93 mL, 25% in water)were stirred in a plastic flask immersed in a water bath at 25 °C.Then, TEOS (1.79 mL) was added slowly to the mixture, and thereaction proceeded for 48 h. The final dispersion was centrifugedthree times (15 min at 19100g for each cycle) and redispersed inabsolute ethanol. Finally, the SNP-PDI was dried in an oven at50 °C for 24 h.
Surface Modification of SNP-PDI with APTES (SNP-APTES):SNP-PDI (0.236 g) was dispersed in dry toluene (5 mL) by ultra-sonication for 15 min. APTES (0.9 mL, 3.8 mmol) was added tothe dispersion, and the reaction mixture was heated to 125 °C for24 h under an argon atmosphere. After that time, the SNPs werecentrifuged three times (15 min at 19100g for each cycle), redis-persed in absolute ethanol, and dried at 50 °C in an oven for 24 h.
Surface Modification of SNP-APTES with RAFT Agent (SNP-RAFT): After the APTES modification, SNP-APTES (0.21 g) wasadded to dry dichloromethane (15 mL) and dispersed by ultra-sonication for 15 min. RAFT agent (0.045 g) and EDC (0.04 g)were added to the dispersion. The reaction proceeded for 24 h atroom temperature under an argon atmosphere. After that time,SNP-RAFT was centrifuged three times (15 min at 19100g for eachcycle), redispersed in absolute ethanol, and dried at 50 °C in anoven for 24 h.
Copolymerization of NAS, OEGMA, and MEO2MA by RAFT: Amixture of NAS (49.7 mg) and RAFT agent (6.80 mg) in dry DMF(5 mL) was added to a Schlenk tube containing a magnetic stirrer.Separately, AIBN (0.187 mg) was added to dry DMF (1 mL), andboth mixtures were degassed with consecutive vacuum–argon cy-cles. The Schlenk tube was immersed in an oil bath at 90 °C, andthe AIBN solution was added to the mixture. The reaction pro-ceeded under an argon atmosphere, and after 4 h, OEGMA(230 μL) and MEO2MA (423 μL) monomers were added to the po-lymerization medium. The conversion was followed by NMR spec-troscopy (Figure S2, Supporting Information) of samples takenfrom the reaction mixture (50 μL) in [D6]DMSO (350 μL). After18 h, the Schlenk tube was opened to air to stop the reaction, andthe reaction mixture was cooled to room temperature. The reactionmixture was added dropwise to diethyl ether in an ice bath to pre-cipitate the polymer. The supernatant was recovered, and diethylether was added several times. Finally, the precipitated polymerchains were dried overnight in an oven at 30 °C for 24 h.
RAFT Synthesis of the Copolymer Shell on the SNP-RAFT Surface:SNP-RAFT (0.167 g) was added to dry DMF (5 mL) in a Schlenktube containing a magnetic stirrer and dispersed by ultrasonication.After 30 min, NAS monomer (49.7 mg) and RAFT agent (3.48 mg)were added to the dispersion. Separately, AIBN (0.207 mg) wasadded to dry DMF (1 mL), and both mixtures were degassed withconsecutive vacuum–argon cycles. The Schlenk tube was immersedin an oil bath at 90 °C, and the solution of AIBN was added tothe mixture. The reaction proceeded under an argon atmosphere.After 4 h, OEGMA (230 μL) and MEO2MA (380 μL) monomerswere added to the polymerization medium. The conversion was fol-lowed by NMR spectroscopy of samples taken from the reactionmixture (50 μL) in [D6]DMSO (350 μL). After 24 h, the Schlenktube was opened to air to stop the reaction, and the mixture wascooled to room temperature.
The final mixture was centrifuged three times (5 min at 2120g foreach cycle) and redispersed in DMF. The supernatant was col-lected, and the core–shell nanoparticles (SNP-poly) were kept inDMF. The supernatant was added dropwise to diethyl ether in anice bath to precipitate the free polymers chains. Finally, the precipi-tated polymer chains were dried overnight in an oven at 30 °C for24 h. These polymer chains that grew in solution through the ad-dition of free RAFT agent to the SNP-RAFT dispersion duringpolymerization were used to estimate the molecular weight and thesize dispersity through GPC.
6-Aminohexylcarbamate Folic Acid (FA-HDA): The folic acid deriv-ative FA-HDA was synthesized through procedures described pre-viously.[49] Dry DMSO (16 mL) and pyridine (10 mL) were addedto FA (0.83 g) in a round-bottom flask containing a magnetic stir-rer under an argon atmosphere. Separately, a mixture of Boc-HDA(0.32 g), DCC (0.63 g), and dry DMSO (16 mL) was prepared un-der an argon atmosphere and added to the round-bottom flask.The reaction proceeded for 48 h in the dark at 30 °C, and then themixture was added dropwise into diethyl ether (2 L) under vigorousstirring. A yellow precipitate was collected and washed with diethyl
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ether several times. Finally, the precipitate was dried under vacuum,and a yellow solid was obtained. The 1H NMR spectrum of theproduct [D6]DMSO was in agreement with that reported pre-viously.[50] The 1H NMR signals of the Boc group were observedat δ = 1.4 ppm, and those of the hexanediamine group were ob-served at δ = 1.5–3.2 ppm (Figure S6, Supporting Information)with an integration of approximately nine, which indicated that thereaction was complete.
FA-HDA-Boc (0.493 g) was deprotected through reaction withTFA (3 mL) for 2 h at room temperature. The TFA was removedunder reduced pressure, and the resulting residue was dissolved indry DMF (5 mL). Pyridine (2 mL) was added dropwise, and thesolution was kept motionless overnight. The solution was addeddropwise to diethyl ether, and a yellow solid started to precipitate.The solid was washed four times with diethyl ether and recoveredby centrifugation (10 min at 2120g for each cycle). The final prod-uct, FA-HDA (6, Figure 2), was dried under vacuum. The success-ful deprotection was confirmed by the disappearance of the Bocsignals at δ = 1.3 ppm in the 1H NMR spectra (Figure S7, Support-ing Information). The product was used without further purifica-tion.
Coupling of FA-HDA to SNP-poly (SNP-poly-FA): The couplingreaction was performed in the dark. FA-HDA (0.032 m in 1.8 mLof DMF) was added dropwise to a mixture of SNP-poly (0.042 gin 1.5 mL of DMF). The reaction was left to run at room tempera-ture, and after 5 d the yellow mixture was centrifuged three times(5 min at 2120g for each cycle) and redispersed first in DMF andthen in Milli-Q water. The SNP-poly-FA conjugates were dried inan oven at 30 °C for 24 h.
The unreacted NAS groups of SNP-poly and SNP-poly-FA werehydrolyzed in borate buffer (10 mL, pH 9.4). The hydrolysis of suc-cinimide esters results in pendant carboxylate groups. The hydroly-sis proceeded under magnetic stirring for 5 d for SNP-poly-FA and2 d for SNP-poly at room temperature. Both samples were centri-fuged three times (5 min at 2120g for each cycle), redispersed inMilli-Q water, and dried in an oven at 30 °C for 24 h.
Cell Culture and Uptake: Cell lines were cultured in a RPMI-1640medium supplemented with 10% FBS and antibiotic antimycoticsolution (100 units/mL penicillin, 0.1 g/mL streptomycin, and0.25 mg/mL amphotericin B) at 37 °C and under a humidified 5%CO2 atmosphere.
The cells were counted and seeded at 2 �105 cells per well in a μ-Slide eight-well plate before the plate was coated with poly-l-Lysinefor 60 min. After 48 h, the medium was substituted by a fresh me-dium containing the different NPs at a concentration of 187 μg/mL.Incubation was performed for 2, 4, 6, or 16 h. After this period, thecells were washed twice with Hank’s balanced salt solution (HBSS)and stained with the membrane dye WGA-Alexa594 for 15–30 min.After staining, the cells were washed with HBSS, and fresh phenol-red-free RPMI-1640 medium was added to the wells before imagingby laser scanning confocal fluorescence microscopy.
Instrumentation: Fluorescence emission and excitation spectra wererecorded with a Horiba-Jobin Yvon Fluorolog-3 spectrofluorime-ter. Right-angle geometry was used, and all spectra were recordedat ambient temperature with a 1 cm optical path length quartz cell.
Transmission electron microscopy (TEM) imaging was performedwith a Hitachi instrument (model H-8100 with a LaB6 filament)operating at an accelerating voltage of 200 kV. This equipment hasa KeenView camera (Sorf Imaging System), and the iTEM softwarewas used to acquire images. One drop of SNP-PDI dispersion inethanol and SNP-poly water dispersion were placed on a carbon
grid and dried in air before observation. The diameter distributionwas obtained with the ImageJ software.
Nanoparticle zeta potential values were recorded with a ZetasizerNano ZS, model ZEN3601. Filtered Milli-Q water was used as thesolvent, and all measurements were performed at 25 °C.
The polymer molecular weight and size dispersity were determinedwith a GPC system constituted by a peristaltic pump (Waters 510),an injector with an injection volume of 50 μL (Waters U6K), threechromatography columns in line, and three detectors. To character-ize the polymers, the following columns were used: a PhenomenexPhenogel 5 μm, 104 Å column; a Phenomenex Phenogel 5 μm,103 Å column; and a Phenomenex Phenogel 5 μm, 102 Å column.The detectors were a Waters 486 spectrophotometer, a Wyatt Mini-Dawn Treos multiangle static light-scattering detector, and aWaters 2410 refraction index detector. For data acquisition, theAstra 5.3.2.1. software from Wyatt was used. The samples werediluted in a 0.5 m LiBr solution and submitted to 10 min of ultra-sound before injection.
The absorbance and transmittance spectra were recorded with aJasco V-660 spectrophotometer with a Peltier temperature control.All spectra were recorded with samples in 1 �1 cm quartz cells.
The monomer conversion was determined by 1H NMR spec-troscopy of samples in [D6]DMSO, and the signals of the reactionsolvent were used as the internal reference. The 1H NMR spectraof Boc-HDA-FA and FA-HDA in [D6]DMSO were also obtained.The 1H NMR spectra were recorded with a Bruker Avance II400 MHz (UltraShield Magnet) spectrometer at 300 K.
Imaging was performed with a Leica TCS-SP5 confocal fluores-cence microscope equipped with a continuous argon ion laser. Theemission of the nanoparticles was collected in the λ = 510–550 nmregion upon excitation at λ = 488 nm, and the emission of the mem-brane marker was collected in the λ = 680–750 nm region uponexcitation at λ = 514 nm.
Acknowledgments
This work was partially financially supported by the Fundaçãopara a Ciência e a Tecnologia (FCT-Portugal), also through post-doc (SFRH/BPD/96707/2013, SFRH/BPD/73822/2010) and Ph. D.(SFRH/BD/89615/2012) grants to T. R., R. F. M. F., and A. S. R.Support by COMPETE, Fundo Europeu de Desenvolvimento Re-gional (FEDER), within projects PTDC/CTM-NAN/2354/2012,UID/NAN/50024/2013 and RECI/CTM-POL/0342/2012 is ap-preciated.
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