influence of electronegative substituents on the binding affinity of catechol-derived anchors to fe...

Influence of Electronegative Substituents on the Binding Affinity of Catechol-DerivedAnchors to Fe3O4 Nanoparticles

Esther Amstad,† Andreas U. Gehring,‡ Håkon Fischer,‡ Venkatamaran V. Nagaiyanallur,†Georg Hahner,§ Marcus Textor,† and Erik Reimhult*,†,|

Laboratory of Surface Science and Technology and Geophysics Institute, ETH Zurich, Switzerland, EaStChem,School of Chemistry, UniVersity of St. Andrews, U.K., and Department of Nanobiotechnology, UniVersity ofNatural Resources and Life Sciences (BOKU), Vienna, Austria

ReceiVed: NoVember 16, 2010

Successful applications of nanoparticles are often limited by insufficient nanoparticle stability due to lowbinding affinity of dispersants. However, excellent Fe3O4 nanoparticle stability was reported in a recent study(Nano Lett. 2009, 9, 4042-4048) that compared different catechol derivative-anchored low molecular weightdispersants. Here, we investigate mechanistic binding aspects of five different anchors from this study thatshowed radically different efficiencies as dispersant anchors, namely nitroDOPA, nitrodopamine, DOPA,dopamine, and mimosine, using electron paramagnetic resonance, Fourier transform infrared spectroscopy,and UV-vis spectroscopy. We demonstrate enhanced electron delocalization for nitrocatechols binding toFe2+ compared to unsubstituted catechols if they are adsorbed on Fe3O4 surfaces. However a too high affinityof mimosine to Fe3+ was shown to lead to gradual dissolution of Fe3O4 nanoparticles through complexationfollowed by dissociation of the complex. Thus, the binding affinity of anchors should be optimized ratherthan maximized to achieve nanoparticle stability.

Introduction

Controlled stabilization and functionalization of oxide nano-particles mainly, but not exclusively, for biomedical applicationshave received increasing attention in recent years.1-3 Prominentexamples are iron oxide nanoparticles for hyperthermia4 and asmagnetic resonance (MR) contrast agents.5-8 Many of theseapplications require dispersants to be irreversibly bound with acontrolled orientation and conformation to the nanoparticles,which is especially important when targeting of specific cells,tissue, or organ under physiological salt concentrations andelevated temperatures is desired.1,8 Only then can good nano-particle stability, sufficient circulation in vivo and a controlledpresentation of (bio)functional targeting groups on the particlesurface be achieved. Almost all dispersants used today are eitherelectrostatically adsorbed on nanoparticles (e.g., dextran)9,10 orattached through reversibly binding anchors such as oleic acidor dopamine.11

Steric stabilization of oxide nanoparticles with low molecularweight dispersants that are adsorbed on the nanoparticle surfacethrough a straightforward and cost-effective “grafting to”approach is attractive. It allows coadsorption of differentlyfunctionalized dispersants resulting in multifunctional nanopar-ticles. Furthermore, the thickness and end-functionality of thethin stabilizing coating can be closely controlled without theneed for sophisticated in situ chemical reactions. The “grafting-to” approach relies on high affinity anchors that can create thebond to the nanoparticle surface under suitable conditions.Naturally, anchors that meet the stringent requirements for

coupling dispersants to nanoparticles by “grafting to” canperform the same function to graft nonfouling polymer brushesto planar substrates and also be used as anchors for initiators tograft polymers from a substrate surface.

Catechol derivatives are often proposed as suitable dispersantanchors to various oxides including TiO2 and iron oxides.Despite the close chemical similarity of different catecholderivatives, their affinities to TiO2

12 and Fe3O411 have been

shown to differ considerably. Because binding affinities of theseanchors to Fe3O4 directly correlate with polymer brush densityand Fe3O4 nanoparticle stability,11 these differences are not onlyof scientific interest but also crucial for many industrial andmedical applications. In particular, it has been demonstrated thatanchors can both lead to too weak reversible binding and totoo strong binding depending on their affinity to the cations ofthe investigated oxide.11 Whereas too low binding affinity leadsto nanoparticle aggregation, too strong binding of anchors resultsin nanoparticle dissolution. Thus, in the quest to optimize thebinding strength to oxides in general and iron oxide in particularit is of highest importance to understand the binding mechanismsin detail.

The structure13-16 and electronic interactions17,18 of differentcatechols complexed with Fe3+, primarily dopamine and L-DOPA, a post-translationally modified amino acid abundantlypresent in the mussel adhesive protein Mytulis edulis,19 havebeen studied in detail. The crystallographic and electronicstructure of peptides,20 proteins21 and models for catecholdioxygenases16,22-24 complexed with iron ions have beenreported. Furthermore, iron-catalyzed catechol degradation hasbeen thoroughly studied in view of its biological relevance.25-31

However, to the best of our knowledge, no such studies onthe adsorption of catechols and catechol derivatives on ironoxides have been reported. Here we present detailed insightsinto the electronic structures and coordination to iron ions offive different catechol derived anchors upon adsorption to Fe3O4

* To whom correspondence should be addressed. Address: Wolfgang-Pauli-Strasse 10, CH-8093 Zurich, Switzerland. Telephone: +41 (0)44 6337547. Fax: +41 (0)44 633 1027. E-mail: [email protected].

† Laboratory of Surface Science and Technology, ETH Zurich.‡ Geophysics Institute, ETH Zurich.§ University of St. Andrews.| University of Natural Resources and Life Sciences (BOKU).

J. Phys. Chem. C 2011, 115, 683–691 683

10.1021/jp1109306 © 2011 American Chemical SocietyPublished on Web 12/15/2010

surfaces (Figure 1), which explain the unusually high affinityof nitrocatechols to magnetite (Fe3O4) nanoparticles. A deeperunderstanding of how these anchors bind may make it possibleto theoretically predict binding affinities of different anchorstoward desired oxides and provide design rules for selectingdispersants with optimal binding affinities without the need forlaborious experimental screening.

Experimental Methods

Materials. NitroDOPA and nitrodopamine, poly(ethyleneglycol)-nitroDOPA where the poly(ethylene glycol) had amolecular weight of 5 kDa (PEG(5)-nitroDOPA) and PEG(5)-dopamine were synthesized as described previously.32 DOPA(purity ) 99%) was purchased from Acros, 3-hydroxytryamine-

hydrochloride (purity > 98.5%), FeCl3 (97%), tris(hydroxy-methyl)aminomethane, KBr, NaCl from Fluka, mimosine (purity) 98%), Fe(II)acetate (99.995%, batch 517933, Lot 03901JJ),N,N-dimethylformamide (DMF) (puriss) and FeCl2 (98%) fromSigma, and ethanol (analytical grade) from Scharlau.

Complexation. For FTIR and UV-vis measurements, 0.137µmol anchors were added to 0.137 µmol FeCl2 and FeCl3,respectively, which was dissolved in 1 mL Millipore water(R ) 18.2 Ω, TOC < 6 ppb), 1 mL 10 mM tris(hydroxym-ethyl)aminomethane containing 150 mM NaCl (Tris), EtOHand DMF respectively. For EPR investigations 40 µmol/mLnitroDOPA was complexed with Fe3+ and Fe2+ at a molarratio of nitroDOPA:iron ion ) 3:1 and 1:1, respectively, inMillipore water. Unless stated otherwise, these solutions were

Figure 1. While catechols bind weakly and reversibly to iron oxide nanoparticles, the binding affinity of mimosine is high enough to remove Fe3+

ions through complexation which gradually dissolves iron oxide nanoparticles. Nitrocatechols have an intermediate affinity to iron oxide nanoparticlesand strongly adsorb on Fe3O4 nanoparticles without dissolving them. R ) H for dopamine and nitrodopamine and R ) COOH for DOPA andnitroDOPA respectively.

684 J. Phys. Chem. C, Vol. 115, No. 3, 2011 Amstad et al.

left at RT for 1 h before they were analyzed with UV-visspectroscopy or freeze-dried for FTIR and EPR investigation(freeze-dryer ALPHA 1-2/LDplus, Kuhner LabEquip, Swit-zerland).

Iron Oxide Nanoparticles. Fe3O4 nanoparticles were syn-thesized as described earlier using a microwave-assisted non-aqueous sol-gel route.32 Briefly, 173 mg Fe(ac)2 was dissolvedin 5 mL benzylalcohol and heated for 3 min to 180 °C in themicrowave (Discover S-class, CEM, NC, USA).33 Particles werewashed once with 10 mL ethanol before they were redispersedin fresh 10 mL ethanol. The particles were coated within 4-8h after synthesis. To obtain surface oxidized nanoparticles, Fe3O4

nanoparticles were kept in EtOH in air at 4 °C for 4 monthsbefore they were stabilized. Oxidation of the aged nanoparticleswas confirmed with X-ray photoelectron spectroscopy (XPS)measurements (Supporting Information Figure S1). For FTIRsamples, 200 µg of particles were added to 1 mL DMF or EtOHcontaining 3.85 µmol of the respective anchor. Pure anchors,rather than PEG-anchors, were chosen for FTIR to minimizethe interference from the C-O and C-C stretching bands ofthe PEG moieties with vibrations from anchors. For EPR studies,100 µg particles, dispersed in 1 mL EtOH, were stabilized with1.54 µmol PEG(5)-nitroDOPA, PEG(5)-dopamine, or PEG(5)-mimosine. Because unstabilized nanoparticles are less suscep-tible to agglomeration in organic solvents, compared to aqueoussolutions, they were stabilized in EtOH and DMF, respectively.Furthermore, highest PEG grafting densities can be achieved ifPEG is adsorbed in a collapsed state.34 High packing densitieshave been shown to directly translate into high nanoparticlestability.11 Therefore, most of the nanoparticles were stabilizedin EtOH in which PEG has poor solubility. Anchors wereadsorbed for 24 h at 50 °C while they were constantlymechanically mixed at 500 rpm (Thermomixer comfort, Vaudaux-Eppendorf, Switzerland). Excessive PEG(5)-dispersants wereremoved by 24 h dialysis against Millipore water using dialysismembranes with a cutoff of 25 kDa (Spectra/Por dialysismembrane, spectrum laboratories, Netherlands), while iron oxidenanoparticles coated with anchors only and analyzed by FTIRwere purified by washing them 10 times with Millipore waterthrough centrifugation for 10 min at 13 400 rpm. Purifiednanoparticles were freeze-dried (freeze-dryer ALPHA 1-2/LDplus, Kuhner LabEquip, Switzerland).

Flat Iron Oxide Surface Preparation. Thin (<10 nm) ironoxide films were reactive magnetron sputtered onto 10 × 10mm2 Si wafers (Silicon materials, Germany) as describedpreviously.11 Their quality was checked with X-ray photoelec-tron spectroscopy (XPS) prior to the adsorption of anchors. Thesubstrates were immersed in 0.25 µM DMF-based solutionswhere anchors were adsorbed for 24 h at 50 °C before thesesubstrates were thoroughly rinsed with Millipore water.

FTIR Spectroscopy. FTIR spectra of the freeze-driedparticles were measured as KBr tablets where the weight ratioof KBr:sample was ca. 100:1. Spectra were recorded from 400to 4000 cm-1 at a resolution of 4 cm-1 at room temperatureand a pressure of <2 mbar. Spectra, were acquired on a BrukerIFS 66v FTIR spectrometer using a DTGS detector. The sumof 32 scans is shown.

EPR. 2-3 mg of unstabilized, PEG(5)-nitroDOPA, PEG(5)-dopamine, and PEG(5)-mimosine stabilized freeze-dried ironoxide nanoparticles and 7-12 mg nitroDOPA/iron complexeswere analyzed. Reproducibility was checked on 2-3 indepen-dent identical samples. EPR measurements were done on aBruker EMX spectrometer at a frequency of 9.86 GHz and apower of 2.04 mW at room temperature. The sum of three scans

is shown. The resonance equation hν ) gµBB where h is thePlanck constant, ν the frequency, µB the Bohr magneton, and Bthe applied magnetic field was used to calculate the g values.These values correspond to the resonance field Br which isdefined as B at maximum absorption, that is, zero crossing ofthe first derivative of the absorption spectrum. N,N-diphenylpi-crylhydrazyl (DPPH) with g ) 2.0036 was used as internalstandard. The ratio of the derived intensity peaks was determinedbased on the total height of the discontinuity resulting from freeelectrons at g ) 2.0 and the height of the Fe3+ signal at g )3.9 with respect to the baseline. Heat treatment was performedin EPR glass tubes. Samples were heated to T > 200 °C for 10min.

UV-vis Spectroscopy. UV-vis spectroscopy was performedon 0.505 µM solutions. Spectra were recorded on a Cary 1EUV-vis spectrometer (Varian) between 200 and 800 nm.

X-ray Photoelectron Spectroscopy (XPS). XPS was per-formed on freeze-dried, as-synthesized iron oxide nanoparticleson a Sigma Probe (Thermo Scientific, MA, U.S.A.), using anAl KR source operated at 200 W. Photoelectrons were detectedwith a hemispherical analyzer at a pass energy of 25 eV at 90°takeoff angle. Data were analyzed using the CasaXPS software(CasaXPS software Version 2.3.15dev52, Software Ldt, U.K.).

NEXAFS Measurements. NEXAFS spectra of the O K-edgeand the Fe L-edge were acquired under normal X-ray incidenceat the PolLux beamline at PSI Villigen (Switzerland) in totalelectron yield mode. The resolution was ≈0.1 eV. The pressurewas kept below 2 × 10-4 mbar. Spectra were recorded on atleast two independent samples for each anchor. The O K pre-edge was normalized to 530 eV while the Fe K-pre-edge wasnormalized to 710 eV.35

Results and Discussion

Fe3O4 nanoparticles (shown in Supporting Information FigureS1) which contained Fe2+ and Fe3+ ions could be individuallystabilized by grafting PEG(5)-nitroDOPA to their surfaces.These nanoparticles were completely stable under physiologicalconditions and up to temperatures of 90 °C.11 However, ifnanoparticles were stored in ethanol for 4 months beforestabilization, they were significantly less stable as was foundwith dynamic light scattering (DLS) (Supporting InformationFigure S2). Storage of iron oxide nanoparticles for 4 months inethanol was shown by XPS to result in oxidation of iron oxidenanoparticles (Supporting Information Figure S3 and Table S1)that then presented exclusively Fe3+ at the surface. This strikingdifference in nanoparticle stability between freshly synthesizedFe3O4 and oxidized iron oxide nanoparticles was observedirrespective of whether iron oxide nanoparticles were stabilizedin EtOH or DMF. Catechols are known to bind mainly toFe3+,36,37 and therefore better binding to surface oxidizedcompared to as-synthesized Fe3O4 nanoparticles was expected.To investigate this apparent contradiction, the coordinationproperties of Fe3+ in iron oxide nanoparticles stabilized withPEG(5)-nitroDOPA, PEG(5)-dopamine, and PEG(5)-mimosinewere analyzed with electron paramagnetic resonance (EPR)measurements.

EPR. EPR spectra of as-synthesized Fe3O4 nanoparticlesstabilized with PEG(5)-nitroDOPA, PEG(5)-dopamine, PEG(5)-mimosine and unstabilized Fe3O4 nanoparticles were dominatedby a near-isotropic signal with g ) 2.06 (Br ) 343 mT),characteristic for nanoparticles in a superparamagnetic state(Figure 2).38 The line width ∆B of the superparamagnetic signalof PEG(5)-anchor stabilized nanoparticles was with ∆B ) 73( 4 mT considerably lower compared to that of unstabilized

Binding Affinity of Anchors to Fe3O4 Nanoparticles J. Phys. Chem. C, Vol. 115, No. 3, 2011 685

Fe3O4 nanoparticles (∆B ) 92 ( 6 mT). The lower ∆B indicatesless magnetic interparticle interactions between stabilized ascompared to unmodified Fe3O4 nanoparticles. Moreover, thestabilized nanoparticles revealed two additional signals at Br )352 mT (resulting in g ) 2.0) and Br ) 180 mT (g ) 3.9). Thesharp signal at g ) 2.0 is characteristic for free electrons. Suchsignals were only found if dispersants were adsorbed on Fe3O4

nanoparticles (Supporting Information Figure S4). The low fieldresonance at Br ) 180 mT (g ) 3.9) can be assigned torhombohedrally distorted, magnetically decoupled Fe3+.39,40

Despite that the two signals occur simultaneously, their relativeintensity, deduced from their peak heights, were different. Therelative intensity ratio of free electrons (g ) 2.0) to Fe3+ (g )3.9) was highest for PEG(5)-nitroDOPA stabilized nanoparticles.This ratio decreased by 14 ( 5% for PEG(5)-mimosine and 32( 1% for PEG(5)-dopamine compared to PEG(5)-nitroDOPAstabilized Fe3O4 nanoparticles. Therefore, electron delocalizationis suggested to be highest for PEG(5)-nitroDOPA stabilizednanoparticles.

The occurrence of a g ) 3.9 signal associated with magnetitenanoparticles indicates that Fe3+ is magnetically decoupled fromthe bulk material.41 Such a configuration is most likely to occurat the surface and caused by the binding of anchors.

To verify this interpretation, dispersants were decomposedby subjecting stabilized iron oxide nanoparticles to T > 200 °C.The removal of the dispersants from the nanoparticle surfacesshould result in EPR spectra similar to those of uncoated Fe3O4

nanoparticles. Indeed, after subjecting PEG(5)-nitroDOPA anduncoated Fe3O4 nanoparticles to T > 200 °C for 10 min, thesignal arising from superparamagnetic nanoparticles exhibiteda similar, broad line width for all samples. The broadening forthe coated nanoparticles is due to an increase of magnetostaticinteractions and clearly shows that the thermal treatmentdecomposed the coating which was further supported by FTIRmeasurements (Supporting Information Figure S5). Furthermore,X-ray diffraction (XRD) spectra showed that the inverse spinelstructure of the iron oxide cores was retained after exposingthe cores to elevated temperatures (Supporting InformationFigure S6). Upon heating both signals at g ) 2.0 and g ) 3.9disappeared (Figure 2c). The correlated appearance and loss ofthe EPR signals of free electrons and magnetically decoupledFe3+ ions upon, respectively, adsorption and removal ofdispersants on Fe3O4 nanoparticles demonstrates that these twosignals were related to each other. Hence adsorbed nitroDOPAwas preferentially bound to Fe3+ rather than to Fe2+, which isnot directly detectable with EPR.42 We have also shown thatnitroDOPA/Fe3+ complexes lead to a signal at g ) 4.25. Thus,the g ) 3.9 signal cannot be assigned to nitroDOPA/Fe3+

complexes. The magnetically decoupled Fe3+ signal with g )

3.9 is thus due to strong interactions of the nitroDOPA withFe3+ ions at the nanoparticle surface.

Upon adsorption of nitroDOPA on Fe3O4 nanoparticles,nitroDOPA can interact with Fe2+ or Fe3+. If nitroDOPA initiallybinds to Fe3+ ions, the simultaneous appearance of free electronsand a magnetically decoupled Fe3+ signal cannot be explained,since the charge transfer to create the radical is unaccountedfor. Alternatively, nitroDOPA can bind to Fe2+ which leads toan increased electron density in the π* system of nitroDOPAand a pronounced electron depletion at the surface bound ironion (see suggested reaction in Figure 3a). For the latter case,the simultaneous appearance of a magnetically decoupled Fe3+

signal and free electrons would be expected. In the literature, areduction of nitrobenzenes43 as well as facilitated oxidation ofFe2+ to Fe3+ in the presence of chelates that strongly bind toFe3+ have already been reported.44,45

PEG(5)-nitroDOPA stabilized oxidized nanoparticles revealeda significantly weaker signal at g ) 2.0 compared to PEG(5)-

Figure 2. (a) Overview of EPR spectra of iron oxide nanoparticles stabilized with (A) PEG(5)-nitroDOPA, (B) PEG(5)-dopamine, (C) PEG(5)-mimosine, and (D) uncoated Fe3O4 nanoparticles that had residual precursors and physisorbed impurities on their surface. (b) EPR spectra of (A)PEG(5)-nitroDOPA and (D) uncoated Fe3O4 nanoparticles before (solid line) and after (dashed line) dispersants were decomposed by subjectingnanoparticles to T > 200 °C. (c) PEG(5)-nitroDOPA stabilized Fe3O4 (solid line) and to Fe2O3 surface oxidized (dashed line) nanoparticles.

Figure 3. (a) Suggested binding mechanism of nitrocatechols whichinitially bind to Fe2+ ions. (b) Iron-catalyzed catechol degradation. Fe3+

complexed catechols are oxidized to semiquinones while Fe3+ is reducedto Fe2+ before semiquinones are further degraded through reactionswith O2.26 (c) Mimosine complexed with Fe3+. R1 ) H for nitrodopam-ine and dopamine and COOH for nitroDOPA and DOPA whereas R2

were amines for pure feet and PEG(5) for dispersants used to stabilizenanoparticles.


nitroDOPA stabilized Fe3O4 nanoparticles, supporting theinterpretation that the change in EPR spectra is a result of astrong electron delocalization and charge transfer betweennitroDOPA and Fe2+ ions. A clear decoupled Fe3+ signal at g) 3.9 could also not be detected for stabilized surface-oxidizednanoparticles (Figure 2c). This provides clear evidence forweaker interaction of nitroDOPA with surface oxidized (Fe2O3)compared to as-synthesized Fe3O4 nanoparticles.

Further evidence that nitroDOPA initially binds to Fe2+ ifadsorbed on Fe3O4 nanoparticles is given by EPR studies ofnitroDOPA complexed with Fe2+ and Fe3+ ions, respectively(Supporting Information Figure S7). The intensity ratios of thepeak heights resulting from delocalized electrons to Fe3+ were6 ( 1 and 8 ( 1 for molar ratios of nitroDOPA/Fe2+ equal to1:1 and 3:1. These were considerably higher than those ofnitroDOPA/Fe3+ complexes (2 ( 0 and 3 ( 1 for molar ratiosof nitroDOPA/Fe3+ equal to 1:1 and 3:1, respectively) despitethat the signal/noise ratio of the Fe3+ signal was comparablefor these complexes. The Fe3+ signal of nitroDOPA/Fe2+

complexes at g ) 4.2 further indicates that Fe2+, which is notdirectly observable with EPR,42 is oxidized to EPR active Fe3+

if complexed with nitroDOPA. More importantly, it shows thatinteractions of nitroDOPA with Fe2+ lead to stronger electrondelocalization compared to nitroDOPA/Fe3+ complexes. Theenhanced electron delocalization of nitroDOPA/Fe2+ complexesthus likely results from a strong electron delocalization andcharge transfer between nitroDOPA and Fe2+, which is in

support of the similar observations and conclusions describedabove for the surface bound ions.

FTIR Studies. To further investigate structural changes thatoccur upon adsorption, FTIR studies were performed onnitroDOPA, nitrodopamine, DOPA, dopamine, and mimosineadsorbed on Fe3O4 nanoparticles. Additionally, FTIR spectraof anchors adsorbed on Fe3O4 nanoparticles were compared tospectra of anchors complexed with Fe2+ and Fe3+ ions. FTIRspectra revealed two pronounced peaks upon adsorption ofanchors to Fe3O4 nanoparticles, one at around 1280 cm-1

assigned to in-plane CO stretching vibrations and one at 1496cm-1 assigned to tangential normal C-C vibration modes ofthe aromatic ring46 (Figure 4a,b and Supporting InformationTable S2).

Changes in the C-C ring vibration of nitrocatechols uponcomplexation with iron ions or adsorption on Fe3O4 nanopar-ticles were small (Figure 4a,b and Supporting Information TableS2). However, shifts of the C-O stretch vibrations from 1290to 1280 cm-1 for nitroDOPA and from 1294 to 1280 cm-1 fornitrodopamine upon adsorption on Fe3O4 nanoparticles com-pared to reference spectra are close to that of nitrocatecholscomplexed with Fe2+ (1285 cm-1for nitroDOPA and 1278 cm-1

for nitrodopamine) in contrast to nitrocatechol/Fe3+ complexes(1287 cm-1 for nitroDOPA and 1290 cm-1 for nitrodopamine).Moreover, the C-O ring vibrations of Fe2+-complexed nitro-catechols were gradually shifted toward lower wavenumberswith time (Figure 5 and Supporting Information Table S2). The

Figure 4. FTIR spectra of (A) pure anchors were compared to those of anchors complexed with (B) Fe2+ and (C) Fe3+ ions in Millipore water ata molar ratio of anchor:iron ion ) 1:1. Complexes were kept for 24 h at 50 °C before they were freeze-dried. Furthermore, anchors were adsorbedon (D) Fe3O4 nanoparticles. (E) Uncoated Fe3O4 nanoparticles and (F) benzoquinones are shown as references. The investigated anchors were (a)nitroDOPA, (b) nitrodopamine, (c) DOPA, (d) dopamine, and (e) mimosine. The most apparent vibrations measured on anchors adsorbed on Fe3O4

nanoparticles were the C-C ring out of plane vibration (*), the C-O stretch vibrations (b) and for nitrocatechols asymmetric (9) and symmetric(2) NO2 vibrations.


good agreement of the C-O ring vibrations of Fe2+-complexednitrocatechols with that of nitrocatechols adsorbed on Fe3O4

nanoparticles and the time dependent shift toward even closeragreement further supports the conclusions from the EPR resultsof initial binding of nitroDOPA to Fe2+ followed by strongelectron delocalization and charge transfer between nitroDOPAand Fe2+. The FTIR spectra also display marked shifts to higherwavenumbers for the symmetric and asymmetric NO2 vibrationsupon adsorption of nitrocatechols on Fe3O4 nanoparticles (e.g.,1317 and 1551 cm-1 for symmetric and asymmetric NO2

vibrations of nitroDOPA) compared to reference spectra (1331and 1535 cm-1)47 (Figure 4a,b). This indicates that the electrontransfer to nitroDOPA resulted in an increased electron densityon the nitro group. Increased electron density in the lowestunoccupied molecular orbital (LUMO) upon deprotonation hasalready been calculated,47 which agrees with the observedincreased electron density in the LUMO upon binding ofnitroDOPA to Fe3O4 nanoparticles.

Differences in FTIR spectra of catechols complexed with Fe2+

and Fe3+ and that of catechols adsorbed on Fe3O4 nanoparticleswere more pronounced relative to those of nitrocatechols (Figure4c,d and Supporting Information Table S2). Whereas the C-Cring vibrations of nitrocatechols were only slightly shiftedtoward higher wavenumbers upon adsorption to Fe3O4 nano-particles, marked shifts toward the position of benzoquinoneswere recorded for (unsubstituted) catechols if adsorbed on Fe3O4

nanoparticles (Figure 4c,d). Furthermore, the C-C ring vibra-tions of DOPA complexed with iron ions were significantly less

apparent after complexes were kept at 50 °C for 24 h comparedto those measured on DOPA that was complexed with iron ionsfor 1 h at RT (Figure 5c). This agrees with a gradual degradationof DOPA. The similarity of FTIR spectra of especially dopamineadsorbed on Fe3O4 nanoparticles with the reference spectra ofbenzoquinone is striking. Therefore, the peak at 1484 cm-1 measuredfor catechols adsorbed on Fe3O4 nanoparticles can likely be assignedto a C-O stretching vibration of semiquinones.48-50

The absence of an electronegative NO2 group on catecholscompared to nitrocatechols renders catechols more prone tooxidation.17,51,52 Iron-catalyzed catechol degradation that resultsin semiquinones, quinones, and eventually in carboxy containingspecies has been thoroughly described in literature.23,25-27,44 Itcan explain the weak C-C ring and C-O stretch vibrations ofcatechols compared to nitrocatechols seen in FTIR. Furthermore,carboxy groups, which can be the result of iron-catalyzedcatechol degradation, have been shown to poorly bind to Fe3O4

nanoparticles.11 This might explain the considerably worsestability of PEG(5)-catechol compared to PEG(5)-nitrocatecholstabilized Fe3O4 nanoparticles.11

In contrast to catechol/iron and nitrocatechol/iron complexes,FTIR spectra of mimosine/iron complexes did not changesignificantly with complexation time and temperature (Figure5 and Supporting Information Table S2). This indicates a fastreaction between mimosine and iron which resulted in stablecomplexes and might be due to the low pKa values ofmimosine53 (Figure 3c). Similar to nitrocatechols, the locationof the C-C ring vibration did not change significantly upon

Figure 5. FTIR spectra of (A) pure anchors, anchors complexed with (B) Fe2+ and (C) Fe3+ kept for 1 h at RT, (D) Fe2+ and (E) Fe3+ kept for24 h at 50 °C and (F) anchors adsorbed on Fe3O4 nanoparticles for (a) nitroDOPA, (b) nitrodopamine, (c) DOPA, (d) dopamine, and (e) mimosine.The molar ratio of the anchor-iron complexes was always 1:1. Identical to Figure 4, the most apparent vibrations were assigned to the C-C ringout of plane vibration (*), the C-O stretch vibrations (b) and for nitrocatechols asymmetric (9) and symmetric (2) NO2 vibrations.


adsorption of mimosine on Fe3O4 nanoparticles (Figure 4e).However, the C-O ring vibration of mimosine and nitrocat-echols complexed with Fe2+ was shifted toward lower wave-numbers compared to the respective reference spectra. Inter-estingly, in contrast to nitrocatechols, the C-O ring vibrationof mimosine adsorbed on Fe3O4 nanoparticles was closer to thatof mimosine complexed with Fe3+ compared to mimosine/Fe2+

complexes. This indicates that mimosine binds directly to Fe3+

and is unlikely to bind to Fe2+ and undergo a redox reaction asobserved for nitrocatechols.

UV-vis Spectroscopy. To further elucidate the role ofadsorption conditions on the binding kinetics of these anchorsto Fe3O4 nanoparticles, UV-vis spectroscopy measurements oncomplexes of anchors with free iron ions were performed(Supporting Information Figures S8-S10 and Tables 1 and 2).Primarily electron transitions between the highest occupiedorbital (HOMO) and the lowest unoccupied orbital (LUMO)were investigated because these electron transitions give insightsinto anchor-iron interactions. Reference spectra were comparedto UV-vis spectra of anchors complexed with Fe2+ and Fe3+

ions at different pHs and in organic solvents respectively. ThepH of Millipore water based complex solutions varied between3 and 5. Tris has been reported not to interfere with iron ions37

and was chosen to buffer the solutions to pH ) 7.4. Furthermore,anchors were complexed with iron in organic solvents, namelyEtOH and DMF, because iron oxide nanoparticles werestabilized in these solvents.

The prominent peak located between 350 and 500 nm wasassigned to the HOMOfLUMO transition resulting from acharge transfer of the HOMO localized on the aromatic ring ofnitrocatechols to the LUMO mainly localized on the nitro group.The wavelength of the HOMOfLUMO transition peak maximaincreased from 350 at pH 5 to 420 at pH 7.4 and to 500 nm atpH ≈ 12. Considering the pKa values of nitrocatechols (pKa1 ≈6.5, pKa2 ≈ 10),11,54 these peaks were assigned to the fullyprotonated, once deprotonated, and twice deprotonated state of

nitrocatechols, respectively. A comparison of the pH-dependentreference spectra of nitrocatechols aliquotted in aqueous solu-tions to reference spectra of nitrocatechols dissolved in EtOHand DMF reveals that nitrocatechols were fully protonated ifdissolved in EtOH whereas a significant fraction of nitrocat-echols was once deprotonated if aliquotted in DMF. While thepeak locations of nitroDOPA and nitrodopamine aliquotted inDMF were identical, relative peak intensities differed markedlyindicating that a higher amount of nitrodopamine was partiallydeprotonated compared to nitroDOPA (Supporting InformationFigure S8).

One hour after complexation, changes in the HOMOfLUMOtransition peak of Millipore water-based nitrocatechol/Fe2+

complex solutions compared to the reference (free) nitrocatecholspectra were negligible (Table 1 and Supporting InformationFigure S8 and S9). This was in contrast to nitrocatecholscomplexed with Fe3+ where nitrocatechol/Fe3+ interactions wereseen already 1 h after complexation (Supporting InformationFigure S10). Well in agreement with FTIR results, a significantbroadening of the HOMOfLUMO peak became apparent 24 hafter nitrocatechols were complexed with Fe2+. No furtherchange in the UV-vis spectra of nitrocatechol/Fe3+ complexeswas measured if these complexes were kept in Millipore waterat RT for 24 h compared to spectra taken 1 h after complexation(Supporting Information Figure S11). These results indicate thatfully protonated nitrocatechols interacted faster with Fe3+

compared to Fe2+. However, fully protonated nitrocatecholsstarted to interact with Fe2+ within 24 h. The different UV-visand FTIR peak locations and shapes of nitrocatechol/Fe2+ andnitrocatechol/Fe3+ complexes indicate different interactions ofnitrocatechols with Fe2+ and Fe3+. Thus, time dependent changesin the UV-vis spectra of nitrocatechol/Fe2+ complexes cannotbe assigned to a nitrocatechol-independent oxidation of Fe2+

to Fe3+ but should be ascribed to a slow reaction betweennitrocatechols and Fe2+ which is absent if nitrocatechols arecomplexed with Fe3+.

HOMOfLUMO transition peaks of nitrocatechol/iron com-plexes were considerably broader and shifted toward higherwavelengths already 1 h after complexation if once deprotonatednitrocatechols were complexed with Fe2+ compared to fullyprotonated nitrocatechol/Fe2+ complexes (Table 2, SupportingInformation Figure S9). This indicates that once deprotonatednitrocatechols interacted faster especially with Fe2+ ions com-pared to the fully protonated nitrocatechols. Hence, adsorbingnitrocatechols in a deprotonated state accelerates their bindingto Fe3O4 surfaces and increases the adlayer formation rate whichmight be beneficial for industrial processes.

Nitrocatechol adsorption on Fe3O4 nanoparticles in a fullyprotonated form from EtOH and in a partially once deprotonatedform from DMF resulted in equal EPR and FTIR spectra.Therefore, adsorption of nitrocatechols on Fe3O4 nanoparticlesin a fully protonated form slows their adsorption down but doesnot prevent or alter it.

Deprotonation of nitrocatechols upon adsorption to iron oxidenanoparticles is well in agreement with literature where protonsfrom alcohols of catechols have been reported to dissociate uponadsorption on TiO2.55-58 As is shown here, adsorption ofcatechol derivatives is facilitated if these anchors are at leastpartially deprotonated already prior to adsorption. The NO2

group lowers pKa1 values of nitrocatechols to 6.749 comparedto that of catechols (pKa > 8.5).59 Therefore, nitrocatechols canmore readily bind to iron ions and adsorb on Fe3O4 surfaces ina pH range between 6.5 and 9 compared to catechols.

TABLE 1: Locations of HOMOfLUMO TransitionUV-vis Peaks of Uncomplexed and with Fe2+ and Fe3+ IonsComplexed Nitrocatechols Dissolved at Different pHs and inOrganic Solvents, Respectively, Where the Molar Ratio ofNitrocatechols/Iron ) 1:1

pH ) 5 pH ) 7.4 pH ) 12 EtOH DMF

nitroDOPA 352 422 499 355 364/441nitroDOPA/Fe2+ 353 414 382 369nitroDOPA/Fe3+ 391 406 372 366nitrodopamine 351 422 501 356 446nitrodopamine/Fe2+ 353 418 353 421nitrodopamine/Fe3+ 390 405 345 363

TABLE 2: Peak Locations of Electron Transition UV-visPeaks of Uncomplexed and with Fe2+ and Fe3+ IonsComplexed Anchors Dissolved at Different pHs and inSolvents, Respectively, Where the Molar Ratio of Anchors/Iron Ions is 1:1

pH ) 5 pH ) 7.4 EtOH DMF

nitroDOPA/Fe2+ 406nitroDOPA/Fe3+ 644 broad 667 420/snitrodopamine/Fe2+ broad 520/snitrodopamine/Fe3+ 644 broad 739? 637DOPA/Fe2+ 574DOPA/Fe3+ 402/742 573 356/739 359dopamine/Fe2+ 575 602/471dopamine/Fe3+ 405/744 561 356 364mimosine/Fe2+ 396/463 395/452mimosine/Fe3+ 513 448 363 361


No differences between reference spectra of catechols wereseen whether they were aliquotted in Millipore water or Trisbuffer, indicating that they were protonated under these condi-tions. This was expected considering their pKa values (pKa >9)59 (Supporting Information Figure S8). Furthermore, theabsence of electron transfer peaks of Millipore water basedcatechol/Fe2+ complex solutions points to negligible interactionsof catechols with Fe2+ ions if dispersed in Millipore waterirrespective of complexation time, well in agreement with whathas been reported.60 This is in contrast to catechol/Fe3+

complexes that showed electron transfer peaks above 500 nm.The considerably higher pKa1 value of catechols compared tonitrocatechols markedly decreases the affinity of catechols toFe2+ and prevents catechol/Fe2+ interactions if complexed inMillipore water irrespective of complexation time and in contrastto what was observed for nitrocatechols.

While UV-vis spectra of Tris and Millipore water-basedmimosine/Fe3+ complexes were independent of the complex-ation time and temperature, the electron transfer peak ofmimosine/Fe2+ complexes shifted toward that of the respectivemimosine/Fe3+ complexes with increasing complexation time(Supporting Information Figure S9). This might indicate oxida-tion of Fe2+ to Fe3+ and hints to strong mimosine/Fe3+

interactions but a low affinity of mimosine to Fe2+ which is instark contrast to nitrocatechols.

Whereas nitrocatechols have a high affinity to iron61 and leadto good Fe3O4 nanoparticle stability,11 catechols have beenreported to undergo iron catalyzed degradation62 and thus leadto poor Fe3O4 nanoparticle stability.11 However, mimosine isknown to have a high affinity toward Fe3+(see ref 63) but onlyleads to intermediate Fe3O4 nanoparticle stability if used as ananchor group for grafting low molecular weight dispersants tonanoparticles.11 To elucidate this apparent contradiction, UV-visspectra of anchor/iron complexes were compared to spectrataken from supernatants of Fe3O4 nanoparticles stabilized withPEG(5)-anchors and dispersed in Millipore water. UV-visspectra revealed a charge transfer peak in supernatants of Fe3O4

nanoparticles stabilized with PEG(5)-mimosine and dispersedin Millipore water identical to Millipore based mimosine/Fe3+

complexes (Supporting Information Figure S12). This closesimilarity suggests that the strong complexation of PEG(5)-mimosine removed Fe3+ ions from the surface resulting ingradual Fe3O4 nanoparticle dissolution and likely is the reasonfor only intermediate nanoparticle stability if PEG(5)-mimosinewas used as dispersant.11 No such PEG(5)-anchor/iron ioncomplexes were found in supernatants of PEG(5)-nitrocatecholor PEG(5)-catechol stabilized Fe3O4 nanoparticles (SupportingInformation Figure S12). Further evidence for iron oxidedissolution upon adsorption of mimosine was given by near edgeX-ray absorption fine structure spectroscopy (NEXAFS) re-corded on flat Fe3O4 surfaces coated with mimosine and DOPA,respectively. Significant differences in the oxygen NEXAFSspectra of mimosine coated iron oxide surfaces compared toreference spectra are indicative of changes in the iron oxidestochiometry upon mimosine adsorption. Furthermore, mimosineadsorption led to increased electron density of surface iron atoms(Supporting Information Figure S13). None of these changeswere detected on DOPA-coated iron oxide surfaces. Thus, theNEXAFS results support UV-vis findings where iron oxidedissolution was observed upon adsorption of mimosine butabsent if catechols were adsorbed on iron oxide nanoparticles.

Conclusions

We have shown that the addition of electronegative groupssuch as a NO2 to the catechol ring greatly enhances electronic

interactions between these anchors and both iron ions in solutionand, importantly, in iron oxides. Consequently, catechol deriva-tives that are electronegatively substituted lead to muchenhanced Fe3O4 nanoparticle stability if covalently linked to aspacer molecule such as PEG. NitroDOPA (and nitrocatecholsin general) was shown to bind to Fe2+ leading to strong electrondelocalization. This results in an enhanced electron density atnitrocatechol anchors and an electron depletion at the chelated,surface bound iron ion. The latter was seen as rhombohedrallydistorted Fe3+ EPR signal at g ) 3.9. The significance of thisstrong electron delocalization on the binding affinity of saidanchors and thus on the nanoparticle stability was illustratedby iron oxide nanoparticles oxidized to Fe2O3. On Fe2O3

surfaces, no Fe2+ is presented. For these nanoparticles, norhombohedrally distorted Fe3+ signal at g ) 3.9 was measuredwith EPR and the corresponding nanoparticle stability ofPEG(5)-nitroDOPA-coated Fe2O3 nanoparticles was poor. Fur-thermore, the addition of an electronegative group to the catecholring not only lowered the pKa1, which facilitated their adsorptionon Fe3O4 surfaces through partial deprotonation at physiologicpH, but also rendered nitrocatechols more oxidation resistant.Oxidation was observed for (unsubstituted) catechols adsorbedon Fe3O4 surfaces and led to significantly reduced bindingaffinity, which severely influenced stability of PEG(5)-catecholcoated Fe3O4 nanoparticles.

Important in the context of further optimization of dispersantanchors for nanoparticles, our findings demonstrate that thereis an optimal binding affinity of anchors toward the metal ionof (oxide) nanoparticles. Whereas low binding affinities ofanchors to iron oxide surfaces lead to reversible adsorption andthus poor nanoparticle stability, too high binding affinity resultedin gradual Fe3O4 nanoparticle dissolution as was exemplifiedfor mimosine where mimosine/Fe3+ complexes were found inthe supernatant of PEG(5)-mimosine stabilized Fe3O4 nanopar-ticles. Thus, the affinity should be high enough to ensureessentially irreversible adsorption in the salt, temperature, andpH range in which sterically stabilized nanoparticles will beapplied, but below the limit that causes substrate dissolutionthrough complex formation.

Acknowledgment. The authors thank Torben Gillich for thesynthesis of nitrocatechols, Professor Markus Niederberger andIdalia Bilecka (Department of Material Science, ETH Zurich)for their support for the iron oxide nanoparticle synthesis, FIRST(ETH Zurich) for providing clean room facility, EMEZ (ETHZurich) for providing electron microscopy facility and the SwissLight Source for synchrotron beam time. Furthermore, we areindebted to Dr. Jorg Raabe and Benjamin Watts for their helpwith NEXAFS measurements at PolLux (PSI, Villigen,Switzerland). COST Action No. D43, EU-FP7-NMP-ASMENAand ETH Zurich are acknowledged for their financial support.

Supporting Information Available: TEM images of un-stabilized and PEG(5 kDa)-nitroDOPA stabilized iron oxidenanoparticles, XPS spectra of surface oxidized and as-synthesized Fe3O4 nanoparticles, DLS of PEG(5)-nitroDOPAstabilized surface oxidized and as-synthesized iron oxidenanoparticles, EPR spectra of pure dispersants and nitroDOPAcomplexed with Fe2+ and Fe3+ ions at molar ratios of anchor/iron ) 1:1 and 1:3. Furthermore, a comprehensive list of FTIRpeaks is presented. UV-vis spectra of uncomplexed and withFe2+ and Fe3+ ions complexed anchors in different solvents aswell as time dependent UV-vis spectra of anchor/iron com-plexes aliquotted in Millipore water are given. UV-vis spectraof supernatants of Fe3O4 nanoparticles stabilized with PEG(5)-


anchor dispersants are compared with the respective anchor/iron complex spectrum. Finally, O K-edge and Fe L-edgeNEXAFS spectra of flat Fe3O4 surfaces coated with DOPA andmimosine respectively are shown. This material is available freeof charge via the Internet at http://pubs.acs.org.

References and Notes

(1) Lin, M. M.; Kim, D. K.; El Haj, A. J.; Dobson, J. IEEE Trans.Nanobioscience 2008, 7 (4), 298–305.

(2) Namdeo, M.; Saxena, S.; Tankhiwale, R.; Bajpai, M.; Mohan,Y. M.; Bajpai, S. K. J. Nanosci. Nanotechnol. 2008, 8 (7), 3247–3271.

(3) Latham, A. H.; Williams, M. E. Acc. Chem. Res. 2008, 41 (3),411–420.

(4) Sharma, R.; Chen, C. J. J. Nanopart. Res. 2009, 11 (3), 671–689.(5) Cheon, J.; Lee, J. H. Acc. Chem. Res. 2008, 41 (12), 1630–1640.(6) Waters, E. A.; Wickline, S. A. Basic Res. Cardiol. 2008, 103 (2),

114–121.(7) Sosnovik, D. E.; Nahrendorf, M.; Weissleder, R. Basic Res. Cardiol.

2008, 103 (2), 122–130.(8) McCarthy, J. R.; Weissleder, R. AdV. Drug DeliVery ReV. 2008,

60 (11), 1241–1251.(9) Lunov, O.; Syrovets, T.; Buchele, B.; Jiang, X. E.; Rocker, C.;

Tron, K.; Nienhaus, G. U.; Walther, P.; Mailander, V.; Landfester, K.;Simmet, T. Biomaterials 2010, 31 (19), 5063–5071.

(10) Gamarra, L. F.; Amaro, E.; Alves, S.; Soga, D.; Pontuschka, W. M.;Mamani, J. B.; Carneiro, S. M.; Brito, G. E. S.; Neto, A. M. F. J. Nanosci.Nanotechnol. 2010, 10 (7), 4145–4153.

(11) Amstad, E.; Gillich, T.; Bilecka, I.; Textor, M.; Reimhult, E. NanoLett. 2009, 9 (12), 4042–4048.

(12) Zurcher, S.; Wackerlin, D.; Bethuel, Y.; Malisova, B.; Textor, M.;Tosatti, S.; Gademann, K. J. Am. Chem. Soc. 2006, 128 (4), 1064–1065.

(13) Lynch, M. W.; Valentine, M.; Hendrickson, D. N. J. Am. Chem.Soc. 1982, 104 (25), 6982–6989.

(14) Zirong, D.; Haltiwanger, R. C.; Bhattacharya, S.; Pierpont, C. G.Inorg. Chem. 1991, 30 (22), 4288–4290.

(15) Attia, A. S.; Bhattacharya, S.; Pierpont, C. G. Inorg. Chem. 1995,34 (17), 4427–4433.

(16) Heistand, R. H.; Roe, A. L.; Que, L. Inorg. Chem. 1982, 21 (2),676–681.

(17) Girerd, J. J.; Boillot, M. L.; Blain, G.; Riviere, E. Inorg. Chim.Acta 2008, 361 (14-15), 4012–4016.

(18) Grillo, V. A.; Hanson, G. R.; Wang, D. M.; Hambley, T. W.; Gahan,L. R.; Murray, K. S.; Moubaraki, B.; Hawkins, C. J. Inorg. Chem. 1996,35 (12), 3568–3576.

(19) Waite, J. H.; Tanzer, M. L. Science 1981, 212 (4498), 1038–1040.(20) Weisser, J. T.; Nilges, M. J.; Sever, M. J.; Wilker, J. J. Inorg. Chem.

2006, 45 (19), 7736–7747.(21) Sever, M. J.; Weisser, J. T.; Monahan, J.; Srinivasan, S.; Wilker,

J. J. Angew. Chem., Int. Ed. 2004, 43 (4), 448–450.(22) Lauffer, R. B.; Heistand, R. H.; Que, L. Inorg. Chem. 1983, 22

(1), 50–55.(23) Bruijnincx, P. C. A.; Lutz, M.; Spek, A. L.; Hagen, W. R.; van

Koten, G.; Gebbink, R. J. M. K. Inorg. Chem. 2007, 46 (20), 8391–8402.(24) Citadini, A. P. S.; Pinto, A. P. A.; Araujo, A. P. U.; Nascimento,

O. R.; Costa, A. J. Biophys. J. 2005, 88 (5), 3502–3508.(25) Cox, D. D.; Benkovic, S. J.; Bloom, L. M.; Bradley, F. C.; Nelson,

M. J.; Que, L.; Wallick, D. E. J. Am. Chem. Soc. 1988, 110 (7), 2026–2032.

(26) Jang, H. G.; Cox, D. D.; Que, L. J. Am. Chem. Soc. 1991, 113(24), 9200–9204.

(27) Emerson, J. P.; Kovaleva, E. G.; Farquhar, E. R.; Lipscomb, J. D.;Que, L. Proc. Natl. Acad. Sci. U.S.A. 2008, 105 (21), 7347–7352.

(28) ElAyaan, U.; Herlinger, E.; Jameson, R. F.; Linert, W. J. Chem.Soc., Dalton Trans. 1997, (16), 2813–2818.

(29) Linert, W.; Jameson, R. F.; Herlinger, E. Inorg. Chim. Acta 1991,187 (2), 239–247.

(30) Kalyanaraman, B.; Felix, C. C.; Sealy, R. C. EnViron. HealthPerspect. 1985, 64, 185–198.

(31) Funabiki, T.; Fukui, A.; Hitomi, Y.; Higuchi, M.; Yamamoto, T.;Tanaka, T.; Tani, F.; Naruta, Y. J. Inorg. Biochem. 2002, 91 (1), 151–158.

(32) Amstad, E.; Zurcher, S.; Mashaghi, A.; Wong, J. Y.; Textor, M.;Reimhult, E. Small 2009, 5 (11), 1334–1342.

(33) Bilecka, I.; Djerdj, I.; Niederberger, M. Chem. Commun. 2008, (7),886–888.

(34) Malisova, B.; Tosatti, S.; Textor, M.; Gademann, K.; Zurcher, S.Langmuir 2010, 26 (6), 4018–4026.

(35) Park, T. J.; Sambasivan, S.; Fischer, D. A.; Yoon, W. S.; Misewich,J. A.; Wong, S. S. J. Phys. Chem. C 2008, 112 (28), 10359–10369.

(36) Charkoudian, L. K.; Franz, K. J. Inorg. Chem. 2006, 45 (9), 3657–3664.

(37) Andjelkovic, M.; Van Camp, J.; De Meulenaer, B.; Depaemelaere,G.; Socaciu, C.; Verloo, M.; Verhe, R. Food Chem. 2006, 98 (1), 23–31.

(38) Sharma, V. K.; Waldner, F. J. Appl. Phys. 1977, 48 (10), 4298–4302.

(39) Barreto, W. J.; Barreto, S. R. G.; Moreira, I.; Kawano, Y. Quim.NoVa 2006, 29 (6), 1255–1258.

(40) Castner, T.; Newell, G. S.; Holton, W. C.; Slichter, C. P. J. Chem.Phys. 1960, 32 (3), 668–673.

(41) Gehring, A. U.; Karthein, R.; Reller, A. Naturwissenschaften 1990,77 (4), 177–179.

(42) Abragam, A.; Bleaney, B. Electron paramagnetic resonance oftransition ions; Clarendon P. Oxford: 1970; pp xiv+911.

(43) Schmitt, H.; Altenburger, R.; Jastorff, B.; Schuurmann, G. Chem.Res. Toxicol. 2000, 13 (6), 441–450.

(44) Harris, D. C.; Aisen, P. Biochim. Biophys. Acta 1973, 329 (1), 156–158.

(45) Bates, G. W.; Workman, E. F.; Schlabac., Mr Biochem. Biophys.Res. Commun. 1973, 50 (1), 84–90.

(46) Araujo, P. Z.; Morando, P. J.; Blesa, M. A. Langmuir 2005, 21(8), 3470–3474.

(47) Cornard, J. P.; Lapouge, C.; Merlin, J. C. Chem. Phys. 2007, 340(1-3), 273–282.

(48) Floriani, C.; Henzi, R.; Calderaz, F. J. Chem. Soc., Dalton Trans.1972, (23), 2640–2642.

(49) Sofen, S. R.; Cooper, S. R.; Raymond, K. N. Inorg. Chem. 1979,18 (6), 1611–1616.

(50) Suzuki, H.; Nagasaka, M.; Sugiura, M.; Noguchi, T. Biochemistry2005, 44 (34), 11323–11328.

(51) Higuchi, M.; Hitomi, Y.; Minami, H.; Tanaka, T.; Funabiki, T.Inorg. Chem. 2005, 44 (24), 8810–8821.

(52) Kawabata, T.; Schepkin, V.; Haramaki, N.; Phadke, R. S.; Packer,L. Biochem. Pharmacol. 1996, 51 (11), 1569–1577.

(53) Stunzi, H.; Harris, R. L. N.; Perrin, D. D.; Teitei, T. Aust. J. Chem.1980, 33 (10), 2207–2220.

(54) Gelb, R. I.; Laufer, D. A.; Schwartz, L. M.; Wairimu, K. J. Chem.Eng. Data 1989, 34 (1), 82–83.

(55) Moser, J.; Punchihewa, S.; Infelta, P. P.; Gratzel, M. Langmuir1991, 7 (12), 3012–3018.

(56) Rego, L. G. C.; Batista, V. S. J. Am. Chem. Soc. 2003, 125 (26),7989–7997.

(57) Rajh, T.; Chen, L. X.; Lukas, K.; Liu, T.; Thurnauer, M. C.; Tiede,D. M. J. Phys. Chem. B 2002, 106 (41), 10543–10552.

(58) Redfern, P. C.; Zapol, P.; Curtiss, L. A.; Rajh, T.; Thurnauer, M. C.J. Phys. Chem. B 2003, 107 (41), 11419–11427.

(59) Ishimitsu, T.; Hirose, S.; Sakurai, H. Chem. Pharm. Bull. 1978, 26(1), 74–78.

(60) Jewett, S. L.; Eggling, S.; Geller, L. J. Inorg. Biochem. 1997, 66(3), 165–173.

(61) Hakkinen, P. Finn. Chem. Lett. 1984, (3), 59–62.(62) Shultz, M. D.; Reveles, J. U.; Khanna, S. N.; Carpenter, E. E. J. Am.

Chem. Soc. 2007, 129 (9), 2482–2487.(63) Ellis, B. L.; Duhme, A. K.; Hider, R. C.; Hossain, M. B.; Rizvi,

S.; vanderHelm, D. J. Med. Chem. 1996, 39 (19), 3659–3670.

JP1109306


influence of electronegative substituents on the binding affinity of catechol-derived anchors to fe...

Documents