Anal Bioanal Chem (2012) 403:2529–2540DOI 10.1007/s00216-012-6044-1

ORIGINAL PAPER

Development of antibody-labelled superparamagneticnanoparticles for the visualisation of benzo[a]pyrenein porous media with magnetic resonance imaging

Martin Rieger · Gabriele E. Schaumann ·Yamuna Kunhi Mouvenchery · Reinhard Niessner ·Michael Seidel · Thomas Baumann

Received: 4 April 2012 / Accepted: 10 April 2012 / Published online: 29 April 2012© Springer-Verlag 2012

Abstract Biogeochemical interfaces in soil are dy-namic in the spatial and temporal domain and re-quire advanced visualisation and quantification toolsto link in vitro experiments with natural systems.This study presents the development, characteriza-tion and application of functional nanoparticles coatedwith monoclonal antibodies to visualise the distribu-tion of benzo[a]pyrene in porous media using mag-netic resonance imaging. The labelled particles are450 nm in diameter and interact with benzo[a]pyrenecovalently bound to silanized silica gel. They did notbind to benzo[a]pyrene adsorbed to plain silica gel.Although unspecific filtration was low, washing stepsare required for visualisation. The ability to visualisebenzo[a]pyrene is inversely correlated to the hete-rogeneity of the soil materials. There are accessrestrictions to narrow pore spaces which allow the vi-sualisation of only those pathways which are also ac-cessible to bacteria and hydrocolloids. The productionof the particles is applicable to other antibodies whichextends the range of potential target contaminants.

Published in the topical collection Analytical Challenges inEnvironmental and Geosciences with guest editor ChristianZwiener.

M. Rieger · R. Niessner (B) · M. Seidel · T. Baumann (B)Institute of Hydrochemistry, TUM, 81377 Munich,Germanye-mail: reinhard.niessner@tum.de, tbaumann@tum.de

G. E. Schaumann · Y. K. MouvencheryInstitute of Environmental Sciences,University Koblenz-Landau, 76829 Landau, Germany

Keywords PAH · Magnetic resonance imaging(MRI) · MRI label · NMR relaxometry ·Anti-B[a]P antibody · Biogeochemical interface

Background

Benzo[a]pyrene (B[a]P) is a priority contaminantand representative of the larger group of polycyclicaromatic hydrocarbons in soil. Polycyclic aromatichydrocarbons (PAHs) are emitted mainly into the at-mosphere and are therefore ubiquitous in the envi-ronment. The solubility of B[a]P in water is 1.62 μg/L[1], and the n-octanol/water partitioning coefficientlog(Kow) is 6.13 [2]. Both values suggest enrichment ofB[a]P at organic material in soil [3]. The high persis-tence and the high carcinogenic and mutagenic prop-erties require monitoring of B[a]P. Current thresholdlevels are 10 ng/L in drinking water and from 2 mg/kg(playgrounds) to 12 mg/kg (industrial estates) insoil.

While the quantitative analysis of PAHs in soil isstate of the art, there is still a need for better under-standing of macroscopic processes like accumulation ordegradation. It is supposed that these processes happenat biogeochemical interfaces (BGI) in soil [4]. BGI aretransient in space and time, and reactions in soil arelimited by the spatial access to these interfaces. Whilemicromodels can be used to explore the processes atBGI on a single-interface level, MRI is a good choice tovisualise and quantify dynamic processes on the scaleof centimeters and in natural systems with a spatialresolution down to 200 μm and a temporal resolution inthe seconds range [5]. While only few contaminants likeheavy metals are directly accessible with MRI, others

2530 M. Rieger et al.

can be visualised with MRI-labelled tracers. For in-stance, bacterial chemotaxis in porous media has beenstudied using immunomagnetic labelled monoclonalantibodies (mAbs) [6].

In this study, we develop a method to visualise B[a]Pin porous media by coupling mAbs directed againstB[a]P to MRI-active nanoparticles (NPs). One possibletype of MRI-active NPs, iron oxide NPs, has been usedfor hyperthermia [7], drug delivery [8], cell separa-tion [9], immunoassays [10, 11], bioweapon detection[12] and MRI. Here, iron oxide NPs are used as contrastagents [13] and nanosensors. [14] used iron oxide NPsfor the detection of bacteria in milk or blood throughMRI. To our knowledge, there is no application ofantibody-coated MRI-labelled NPs for the visualisationof contaminants in soil.

To produce antibody-coupled magnetic iron oxideNPs, most commonly a wet chemical route is used,starting with precipitation from dissolved iron salts(Fe2+ and Fe3+ in molar ratio 1:2) in water by addinga base (NaOH, NH4OH, or others). Size, size distrib-ution, composition and, with some restrictions, shape[15] can be controlled through parameters such asthe type of salts (chlorides, sulphates, nitrates, per-chlorates, etc.), reaction conditions (temperature, stir-ring speed, gas atmosphere, concentration of reactants,etc.), pH and ionic strength.

Different surface modifications are used to coupleantibodies or other biomolecules to NPs and to sta-bilize them in aqueous solution. One way is to pre-cipitate the NPs in a solution containing polymers orsurfactants. [16] prepared dicarboxypolyethyleneglycol(DCPEG)- and diaminopolyethyleneglycol-coated ironoxide NPs which were coupled to mAbs for theuse in hyperthermia. Other polymers which are of-ten used for non-covalent coating are dextran [17,18], polyvinyl alcohol [19], or fatty acids [20]. Co-valent surface modifications on iron oxide NPs wereused with silane reagents like aminopropyltriethoxysi-lane (APTES) or 3-glycidyloxypropyltrimethoxysilane.Here, biomolecules can be coupled directly to the func-tional amino or epoxy groups [21] or attached to acovalently bound polymer like PEG [22].

Materials and methods

Materials

Glass columns (10 × 100 mm, 10 × 250 mm) were ob-tained from ms-scientific (Berlin, Germany). Ninety-six-well polystyrene (order no. 655061) and polypro-pylene (order no. 655201) microwell plates were

obtained from Greiner (Frickenhausen, Germany)and silica gel (grain diameter 0.5–1 mm) from Roth(Karlsruhe, Germany). All salts and standard re-agents were analytical grade and purchased from Sig-ma (Taufkirchen, Germany). APTES, ammonium hy-droxide (25 wt.%), bovine serum albumin (BSA),Bradford reagent, ethanol (absolute), iron(III) chlo-ride hexahydrate (FeCl3·6H2O), iron(II) sulphate hep-tahydrate (FeSO4·7H2O), N-(3-dimethylaminopropyl)-N′-ethylcarbo-diimide hydrochloride (EDC), N-hydroxysuccinimide (NHS), 2-(N-morpholine)-ethanesulphonic acid (MES), Tris base and Tween 20 wereall purchased from Sigma-Aldrich (Munich, Germany).PEG (NH-CO-C2H4-COOH)2 (DCPEG) was obtainedfrom Rapp Polymere (Tübingen, Germany). B[a]P bu-tyric acid (B[a]P-BA) was obtained from Institut fürPAH-Forschung (Greifenberg, Germany). The mouseanti-B[a]P-mAb 22F12 was produced in our insti-tute [23, 24]. Peroxidase-labelled horse anti-mouseIgG antibody was obtained from Vector Laboratories(Burlingame, US), and goat anti-mouse IgG antibodywas obtained from Sigma. Glass beads (d = 2 mm) wereobtained from Hasenfratz Sandstrahltechnik (Assling,Germany), and silica gel (d = 0.5 − 1 mm) was obtainedfrom Roth (Karlsruhe, Germany). All chemicals wereused without any further purification. Ultrapure wa-ter was produced by a Milli Q plus 185 system fromMillipore (Schwalbach, Germany) and used throughoutthe work. Phosphate-buffered saline (PBS) containing145 mM NaCl, 10 mM KH2PO4 and 70 mM K2HPO4

adjusted to pH 7.6 was utilized as buffer solution.

Preparation of NPs

The preparation of Fe3O4-NPs is based on a slightlymodified procedure described in [25]. Briefly, 5 g ofiron(III) chloride hexahydrate was dissolved in 30 mLwater and purged with nitrogen for 30 min to removedissolved oxygen. Then 2 g iron(II) sulphate heptahy-drate was added and stirred under nitrogen atmospherefor 15 min at 800 rpm with a magnetic stirrer. The pre-cipitation of the magnetic nanoparticles “MNPs” wasinduced by adding 5.5 mL of ammonium hydroxide.Afterwards, the slurry was stirred for another 2.5 h at60◦C under nitrogen atmosphere. Then the slurry wasmagnetically separated from the supernatant, washedtwo times with 100 mL water and stored in 60 mL wa-ter. To produce silanized Fe3O4-NPs (Fe3O4–APTES),10 mL of APTES was added to 2 mL of water whichwas acidified with HCl to pH = 4 for 6 h. Here, thealkoxide groups are replaced in a hydrolysis reactionby hydroxyl groups. These groups condense and forma liquid silane polymer. This gel-like solution was

MRI-labelled anti-B[a]P-antibodies 2531

dissolved in water (40 mL) and was added to the Fe3O4-NP dispersion (60 mL). After purging with nitrogen,the dispersion was stirred at 500 rpm overnight at60◦C. Then the slurry was magnetically separated andwashed two times with 40 mL water and ethanol each.The final product was dried into powder under vac-uum. DCPEG-coated NPs (Fe3O4–APTES–DCPEG)were produced by dissolving 400 mg DCPEG in water(30 mL) and purging with nitrogen for 30 min to removedissolved oxygen. In another beaker, 200-mg Fe3O4–APTES NPs were dispersed in 10 mL water and ul-trasonicated for 10 min. Afterwards, the NP dispersionwas added to the polymer solution, purged again withnitrogen for 10 min and stirred at 500 rpm overnight at60 ◦C. The coated NPs were dialyzed (molecular weightcutoff = 10,000 g/mol) for 2 days against water, whichwas changed four times, to remove the rest of DCPEG.The final product was lyophilized into powder.

The mAbs 22F12 were coupled covalently to the freecarboxyl groups on the surface of the NPs by dispers-ing 0.5 mg Fe3O4–APTES–DCPEG NPs for 15 min in0.5 mL MES buffer (0.1 M, pH 4.5) containing EDC(10 mg) and NHS (10 mg). After washing with MESbuffer, the activated NPs were dispersed in 0.5 mLPBS buffer containing approximately 25 μg 22F12 andgently shaken for 2 h. After washing for three timeswith 0.5 mL washing buffer (PBS, 0.05% Tween 20),the AbMNPs were shaken for another 2 h in 0.5 mLblocking buffer (H2O, 0.1 M Tris base, pH 8.5) followedby another final washing step. Finally, the AbMNPswere dispersed in 0.5 mL water and stored at 4◦C.

Preparation of B[a]P-coated porous media

The coating of silica gel with B[a]P was based on [26]with several modifications. To prepare the surface ofthe silica gel for silanization, 100 g silica gel was stirredat 200 rpm in 200 mL HCl/MeOH (1:1, v/v) at roomtemperature for 30 min. After rinsing the substrate with300 mL distilled water for three times, it was stirredfor 30 min in 100 mL concentrated H2SO4. Then thesubstrate was rinsed again with 300 mL distilled waterfor three times and boiled in 150 mL distilled waterfor 30 min. Silanization was then achieved by stirring(200 rpm) the substrate for 30 min at room temperaturein 100 mL distilled water containing 10% APTES, be-fore it was rinsed with 300 mL water and 200 mL EtOHfor three times each. At last the silica gel was cured inan oil bath at 80◦C under vacuum. The silanized silicagel (100 g) was then suspended in 100 mL 0.1 M MESbuffer (pH = 4.5) containing 2.5 mg (7.4 × 10−3 mmol)B[a]P-BA dissolved in 1 mL dimethylformamide and200 mg (1.0 mmol) EDC and stirred (150 rpm) for 2 h atroom temperature. Finally, the B[a]P-coated silica gel(B[a]P-Si) was rinsed with 300 mL water and 200 mLEtOH for three times each and dried overnight in anoven at 100 ◦C (Fig. 1).

Characterization of NPs

Mössbauer spectroscopy was used to identify the min-eral structure of the iron oxide NPs on a standardMössbauer spectrometer. IR spectra were obtained on

Fig. 1 Preparation of theB[a]P-coated silica gel

2532 M. Rieger et al.

a Nicolet 6700 Fourier transform infrared spectrometer(FT-IR) to characterize the functional groups on thesurface of the NPs. A Perkin Elmer ICP/MS Elan 6100was used to measure the iron content as well as thesilica content of the NPs in order to determine the com-position of the NPs. Thermogravimetric analyses wererun on a Setaram TG/DTA. A dried sample of Fe3O4–APTES–DCPEG was placed in the TGA furnace, andthe measurements were carried out under nitrogen witha heating rate of 15◦C/min from room temperature to600◦C.

A Bradford colorimetric assay [27] using reagentsfrom Biorad and BSA as standard was used to de-termine the mAb concentration before and after thecoupling to the NPs. Here 100 μL of the Bradfordreagent and 100 μL of the mAbs solution were puttogether on a microwell plate, and the absorption at595 nm was measured in a microwell plate reader.The size distribution of the MNPs was obtained usingthe nanosight nanoparticle tracking analysis (NTA), aZetasizer Nano ZS from Malvern Instruments for dy-namic light scattering analysis (DLS) and a JEOL JEM-100cx transmission electron microscope (TEM), as wellas an asymmetrical flow field-flow fractionation (AF4)from Postnova Analytics (Landsberg, Germany). Thedispersion medium for AF4 was ultrapure water. Thesize standards were polystyrene latex beads with 79, 110and 510 nm diameter at a concentration of 50 mg/L inwater. The AF4 was coupled to a UV/Vis detector op-erated at a detection wavelength of 250 nm. The mag-netic properties of Fe3O4, Fe3O4–APTES and Fe3O4–APTES–DCPEG NPs were determined on a MPMS-XL5 SQUID-Magnetometer from Quantum Design.The measurement was done at room temperature anda field strength between −10,000 and 10,000 Oe. About5 mg of the samples was weighed into a gelatine cap-sule and fixed in a straw. The resulting data werecorrected with the measurements of an empty sampleholder. Proton NMR relaxation decays were recordedin aqueous solutions with five different concentrationsof Fe3O4–APTES–DCPEG NPs (c(Fe) = 0, 0.05, 0.19,0.97, 2.43 mmol/L) to determine the relaxivity r2.

Spin–spin relaxation (T2 relaxation) profiles wereacquired at a magnetic field strength of 0.176 T in aBruker Minispec 7.5 NMR relaxometer (Bruker, Ger-many), using the Carr–Purcell–Meiboom–Gill pulse se-quence and the following acquisition parameters: echotime (TE) = 0.15 ms, recycle delay = 2.5 s, number ofechoes = 1,000 and number of scans = 32.

The acquired relaxation time decay was fitted toEq. 1 using a non-linear least squares algorithm and thesoftware package R [28]. When fitting the decay curvewith the sum of three exponentials, the fit is sensitive

to the number of data points contributing to the short,medium and long relaxation times. To prevent biasedresults, the weight of the data points was set to the totalnumber of points divided by the number of points in therelevant section (short 0–200 ms, medium 200–2,000 ms,long >2,000 ms). The background bg is the mean inten-sity of all echoes registered at times >6,000 ms.

I = Ishorte− t

T2short + Imediume− tT2medium + Ilonge

− tT2long + bg

(1)

Surface-enhanced Raman spectra of Si–APTES–B[a]P and pure B[a]P were obtained with a Renishaw2000 Raman microscope system and the help of Agcolloids [29]. The concentration of B[a]P adsorbed tosilica gel was measured by extracting the column mate-rial with dichloromethaneand analysing the extract withHPLC and ELISA [30].

Column experiments

An antibody solution with different concentrations ofmouse anti-B[a]P mAb 22F12 in PBS containing 0.1%BSA is pumped with an peristaltic pump (Reglo MS,Ismatec) through the glass columns, filled with eitherplain silica gel or B[a]P-coated silica gel. The effluent iscollected dropwise in an 96-well low-binding microwellplate. The samples are transferred to the ELISA todetermine the concentration of antibody and to ob-tain the breakthrough curve (BTC). The BTC is fittedwith the one-dimensional transport equation usingCXTFIT [31].

Calibration columns with different concentrations ofNPs are prepared with 2 g of Si–APTES. Then each col-umn was filled with 1.8 mL of Fe3O4–APTES–DCPEG-NP solutions (see Table 1).

For the NMR relaxometry tests, a glass column wasfilled with three layers of silica gel, each 3 cm thick. Themiddle layer was coated with B[a]P, while the top andbottom layer contained plain silica gel. At each endof the column, 1 cm of glass beads was placed for abetter distribution of the flow. Polytetrafluoroethylenefittings at each terminus connected the column to the

Table 1 Columns used for calibration in MRI

Column Water Water Mass Conc.content content Fe3O4 Fe3O4

(%, v/v) (%, w/w) (μg) (μg/g)

0 23.7 21.4 0 01 16.0 13.1 90 9.92 20.0 17.2 900 993 38.1 42.4 9,000 990

MRI-labelled anti-B[a]P-antibodies 2533

peristaltic pump and the effluent. After saturating thecolumn with water at 0.9 mL/min, the T2 values ofeach part were measured. Then, 4 mL of a dispersionof AbMNPs in water (c(Fe) = 25 μg/mL) is pumpedthrough the glass column at 0.9 mL/min. After the flowwas stopped, the T2 values after NP addition weremeasured. The column was flushed seven times with4 mL water at 2 mL/min to remove non-bound AbM-NPs. After each washing step, the T2 values of eachpart were recorded. After incubation overnight in thespectrometer, each part was measured again.

The experiment was repeated after rinsing the col-umn with 10 mL water at 2.0 mL/min. This time 4-mL AbMNPs were added at 2.0 mL/min, followed byfour washing steps (each 10 mL; 2.0 mL/min) and onewashing step overnight (900 mL; 0.9 mL/min). A wash-ing step with 100 mL at 10 mL/min completed thisexperiment. The T2 relaxation times of every part wererecorded after each step.

MRI was performed on a SIEMENS MagnetomMedical Scanner (1.5 T) at the University Hospital ofUlm. Spin echo and gradient echo sequences were usedand adjusted to the columns.

Results

Preparation and characterization of antibody-labelledmagnetic NPs

The synthesis of the DCPEG-coated and antibody-coupled magnetic NPs is schematically shown in Fig. 2.

After a coprecipitation reaction of Fe2+ and Fe3+ions under alkaline conditions, a reactive amino group(−NH2) was introduced to the surface by a silaniza-tion reaction. The prepolymerized APTES could bindcovalently with its hydroxyl groups (−OH) to freehydroxyl groups on the surface of the Fe3O4 NPs ina hydrolysis reaction. In the next step, the DCPEG iscovalently linked to the introduced amino groups withits carboxy groups (−COOH) forming a peptide bond.Now, one can bind the mAbs with its amino groupsfrom the heavy chain to the free carboxy groups of theDCPEG forming another peptide bond. So, polymer(DCPEG) stabilized antibody-coupled magnetic NPswere prepared, where all components are covalentlylinked to the NPs.

The mineral phase of the iron oxide NPs was char-acterized using Mössbauer spectroscopy. The spectrumrecorded at 150 K consists of two sextets with hyperfinefields (B1 and B2) at 46.1 and 49.2 T. The isomer shiftsare 0.16 and 0.56 mm/s, whereas the electric quadrupoleinteraction was ≈0 mm/s for both species (Fe3+ andFe2.5+). These values are in agreement with the resultsfor magnetite [32].

The FT-IR spectra (Fig. 3) of pure Fe3O4 showabsorption peaks at 792 and 889 cm−1, which belongto vibrations of hydroxy groups (−OH) on the surfaceof the NPs, as well as a broad absorption peak be-tween 500 and 700 cm−1 due to stretching vibrationsof the Fe–O bonds in Fe3O4. In the spectra of Fe3O4–APTES, one could see the characteristic absorptionpeaks for Fe3O4 as well as absorption peaks at 991 and1,558 cm−1, which result from the silica shell formed

Fig. 2 Schematic of thesynthesized AbMNPs

Y Y

YY

Y

Y

YY

Y

2534 M. Rieger et al.

Fig. 3 FT-IR spectra of (top to bottom) pure Fe3O4, Fe3O4–APTES, Fe3O4–APTES–DCPEG and pure DCPEG

by silanization with APTES. The absorption bond at991 cm−1 represents the vibration of the Si–O–H bondwhereas the absorption bond at 1,558 cm−1 belongs tovibration of the amino group (−NH2). The comparisonof the Fe3O4–APTES–DCPEG spectra and the pureDCPEG spectra shows that both have the characteristicabsorption bands of DCPEG. An absorption bond ofthe stretching vibration of the C–O–C bond appearsat 1,101 cm−1. The bond at 2,879 cm−1 belongs to theC–H vibration. In addition to the DCPEG character-istic absorption bonds, the spectra of Fe3O4–APTES–DCPEG shows the three absorption peaks of Fe3O4 aswell as the absorption bonds of APTES [25]. Overall,these results indicate the successive grafting of APTESand DCPEG to the surface of the NPs.

An ICP/MS analysis was done in order to deter-mine the relative elemental mass composition of the

Table 2 Relative mass composition of the NPs

Fe Fe3O4 Si APTES DCPEG(%) calc. (%) (%) calc. (%) calc. (%)

Fe3O4 72.4 100.0Fe3O4–APTES 51.5 71.1 3.7 28.9Fe3O4–APTES– 21.7 30.0 1.6 12.2 57.8

DCPEG

NPs. The measured relative mass of iron and silica aswell as the subsequent calculated mass fractions of thedifferent components Fe3O4, APTES and DCPEG areshown in Table 2. The mass fraction of magnetite Fe3O4

in the Fe3O4–APTES–DCPEG NPs is 30%.The absolute weight loss of the Fe3O4–APTES–

DCPEG NPs in TG/DTA was 73.2%, which results ina magnetite content of 26.8% and is in good agreementwith the ICP/MS analysis.

The total amount of antibodies coupled to theNPs was determined with a Bradford assay to be15.0 ± 4.2 μg Ab/mg. With an onset of 25 μg Ab/0.5 μgNPs, the coupling efficiency is 29.9 ± 8.4%. Only theUV/Vis spectrum of the coated NPs showed an absorp-tion band at 280 nm, which is characteristic for aromaticamino acids. From this we conclude that the antibodiesbind to the nanoparticle surface.

The characterization of the size and size distributionof the NPs was done using DLS, NTA [33], scanningelectron microscopy (SEM), TEM and asymmetricfield-flow analysis (AF4) [34]. The summary of all sizeanalyses is given in Table 3. The size of the Fe3O4-core is around 10 nm. Coating with APTES yields largeragglomerated particles with a narrow size distribu-tion centered around 115 nm. Considering that APTESis forming a monolayer on the surface of the NPs,

MRI-labelled anti-B[a]P-antibodies 2535

Table 3 Comparison of average particle sizes of Fe3O4, Fe3O4–APTES, Fe3O4–APTES–DCPEG and Fe3O4–APTES–DCPEG-Abmeasured with NTA, DLS, SEM, TEM and AF4

NTA (nm) DLS (nm) SEM (nm) TEM (nm) AF4 (nm)

Fe3O4 10Fe3O4–APTES 107 ± 13 127 ± 50 114Fe3O4–APTES–DCPEG 279 ± 57 301 ± 25 225 ± 44 270Fe3O4–APTES–DCPEG-Ab 445 ± 25

agglomerates of Fe3O4 NPs must be formed duringthe silanization reaction. After coating with DCPEG,the mean particle size increases to about 240 nm. Thepolydispersity index (PdI [35]) of the DLS measure-ments indicates that only Fe3O4–APTES (PdI = 0.17)has a narrow particle size distribution (0.1 < PdI <

0.2). Fe3O4–APTES–DCPEG (PdI = 0.27) and Fe3O4–APTES–DCPEG-Ab (PdI = 0.44) reveal a broaderparticle size distributions (0.2 < PdI < 0.5). Aggre-gation of the AbMNPs, however, was negligible; theferrofluid was stable for weeks.

The saturation magnetization (Ms) of the samples is56.8, 53.8 and 12.8 emu/g, respectively. The high reduc-tion of Ms(Fe3O4–APTES–DCPEG) can be explainedby the lower percentage of magnetite (30% insteadof 71% for Fe3O4–APTES). There is no hysteresis inany magnetization curve. A remanence and coercivityat zero suggests that the NPs have superparamagneticproperties.

The T2 relaxation rates (1/T2) of Fe3O4–APTES–DCPEG NPs in aqueous solution at 0.176 T and25 ◦C are a function of iron concentration. The slope

of the linear regression gives a relaxivity r2 of13.0 ± 0.42 (mmol/L)−1s−1 (R2 = 0.99, n = 5, m = 1).

Characterization of B[a]P-coated porous media

The presence of amino groups on the silanized silicagel was verified with the TNBS test [36]. A surface-enhanced Raman spectrum of B[a]P-Si is given inFig. 4. The B[a]P specific peaks show up at 1,237, 1,342,1,381 and 1,578 cm−1. This is in good accordance to theobserved SERS bands of B[a]P on a gold layer (1,235,1,385 and 1,581 cm−1) [37]. This indicates a successfulcoupling of B[a]P to the silica gel. The amount of B[a]P,adsorbed to silica gel, was determined with HPLC/Uv–Vis and ELISA. The extractable concentration of B[a]Padsorbed to silica gel of 940 ± 240 μg/kg B[a]P (HPLC)and 1,200 ± 300 μg/kg (ELISA) are in good agreement.These B[a]P concentrations are representative for thenatural background concentrations of B[a]P in soil;thus, the experimental setting is close to environmentalconditions.

Fig. 4 Surface-enhancedRaman spectra of B[a]Pcovalently bound to silica gelin comparison to plain B[a]P

2000 1500 1000 500

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B[a]PSilicaAPTESB[a]P

2536 M. Rieger et al.

Column tests with anti-B[a]P-antibodies

It is necessary for the visualisation of B[a]P at biogeo-chemical interfaces in soil that the antibody recognizesB[a]P adsorbed to the solid matrix and that unspecificbinding is negligible. This requirement was tested incolumn tests. Silica gel was chosen as model columnmaterial representing an angular mineral matrix. B[a]Pwas immobilised to the column material in two differentways. First, B[a]P adsorption to uncoated silica gel,which is representative to adsorption of B[a]P to amineral matrix in the absence of any organic carbonor humic substances. In this case, B[a]P is supposedto adsorb flat on the surface [38]. Second, B[a]P iscovalently linked to silanized silica gel, thus “sticking”into the solution as shown in Fig. 1. This is very likelyrepresentative for the adsorption to a surface in thepresence of humic substances although the details ofthe adsorption process have not yet been modelled.

For the first experiment, there was a recoveryrate for the anti-B[a]P-antibody of 99.5% regardlesswhether there was B[a]P adsorbed to the silica gelor not. This suggests that the antibody is not able torecognize B[a]P adsorbed directly to silica gel. On theother hand, there is also negligible unspecific binding ofthe antibody. For the second experiment, where B[a]Pwas bound covalently to silicagel, the behaviour wasdifferent. The BTC of 8 mL antibody solution (c(Ab)0.4 μg/mL, in PBS containing 0.1% BSA) followed by10 mL PBS containing 0.1% BSA through 7.5 g silica–APTES and B[a]P-Si, respectively, with a flow rate

0.0 0.5 1.0 1.5 2.0 2.5 3.0

0.0

0.1

0.2

0.3

0.4

0.5

V/V0

c(A

b), µ

g/m

L

Fig. 5 BTCs of the anti-B[a]P-Abs in columns filled with a layerof silica–APTES (open circles) and a layer of B[a]P-Si (f illedcircles)

of 0.5 mL/min is given in Fig. 5. The x-axis shows thefluid volume passed through the column (V) dividedby the pore volume of the silica gel in the column(V0) which was 6 mL. Again, the recovery rate forthe antibody passing through the column with silanizedsilica gel, but without B[a]P, is 99.5%. When B[a]P iscovalently bound, the breakthrough of the antibody isretarded (R = 2) and the recovery rate drops to 37.5%.This indicates that the antibody recognizes and bindsto B[a]P. The retardation suggests that some of theantibodies bind only temporarily, while others remainwith the covalently bound B[a]P.

NMR relaxometry to detect layers of B[a]P

In preparation to the visualisation, a column NMRrelaxometry experiment was performed to quantify theability to detect B[a]P linked to silica gel. NMR relax-ometry, in this case, gives access to the change of the T2

values directly and with better sensitivity than MRI.Figure 6 shows the results of the T2 measurements

for the three parts of the column. Throughout theexperiment, short T2 relaxation times were in the rangeof 3.49 ± 0.3 ms, intermediate T2 relaxation times werein the range of 140 ± 17 ms and long T2 relaxationtimes were in the range of 777 ± 77 ms. Relative inten-sities for the short, intermediate and long componentwere 0.47 ± 0.033, 0.14 ± 0.036 and 0.37 ± 0.038. Thestandard errors of the short, intermediate and longT2 relaxation times as obtained from the non-linearleast squares fitting procedure were 0.2, 0.4 and 0.1%.Standard errors for the relative intensities were 0.1, 0.3and 0.1%. respectively.

The long T2 relaxation times in the three parts ofthe water saturated column before the addition of theNPs were 870, 864 and 872 ms for parts 1, 2 and 3,respectively. After the addition of the AbMNPs, thevalues of each part were reduced to 511,684 and 782 ms.At the same time, the intermediate T2 relaxation timesin parts 1 and 2 decrease from 174 to 103 ms and from150 to 113 ms. The short T2 relaxation times increaseslightly for part 1. These observations recflect an in-crease of the AbMNP concentration. The changes arethe highest close to the inlet where unspecific filtrationof the injected NPs is assumed. The relative intensitiesdo not change after the injection of the NPs.

With each washing step, the T2 relaxation times arerecovering to their initial values indicating decreasingconcentrations of AbMNPs. This suggests that AbM-NPs are mobilised and transported through the column.T2 relaxation times stay at a slightly lower levels inpart 2. This suggests that the AbMNPs are actuallybinding to B[a]P and cannot be mobilised that easily.

MRI-labelled anti-B[a]P-antibodies 2537

Fig. 6 T2-relaxation times inthe three parts of the columnshowing the effects ofnanoparticle injection andwashing steps: top long andintermediate T2 relaxationtimes, middle short T2relaxation times and bottomrelative intensities of thelong, intermediate and shortcomponents in part 2 of thecolumn

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NP injection NP injectionslow flow overnightovernightovernight

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Overnight storage led to an increase of the relativeintensity of the intermediate component of 20–30% inpart 2 and an increase of the T2 relaxation times. As thisindicates an increase of the relative fraction of moremobile spins, we assume diffusive attachment from NPsfrom the solution to the matrix.

After the second NP addition, the T2 relaxationtimes of all parts were again reduced. But here, the T2

value of the B[a]P section after the washing steps 1–4was always 25 ± 4 ms lower than the T2 values of theother two parts. After a long washing step overnight,the T2 relaxation time in parts 1 and 2 increased tem-porarily. This could point to a remobilisation of un-specifically, weakly bound NPs from parts 1 and 2. Forthe last washing step, the pumping rate was increased

to 10 mL/min. This led to a mobilisation of filteredAbMNPs close to the inlet into part 1.

MRI visualisation of anti-B[a]P-NPs

With the spin echo sequence used for imaging(TE = 13 ms, TR = 5,050 ms), the signal intensity issensitive to changes of the T2 relaxation times (T2-weighted for short). Here, only the calibration columnswithout NPs and with a concentration of 9.9 μg/g werevisible. This indicates a high sensitivity of the imagingsequence to the presence of AbMNPs.

Figure 7 shows an annotated image taken witha spin echo sequence (TE = 13 ms, TR = 5,050 ms).

Fig. 7 Annotated MRI image(raw data) of the column usedfor relaxometrymeasurements. The signalintensity is depending on thewater content and theconcentration of anti-B[a]Pnanoparticles

2538 M. Rieger et al.

Voxels are 0.45 × 0.45 × 3.6 mm3 with the slices ori-ented along the column. There are three distinct fea-tures in the untreated image: At position 1, there is asignificant, homogeneous decrease of the signal inten-sity. At this position, the column looks brownish fromNPs trapped at the transition from glass beads to silicagel. At the positions marked with 2, the Si–APTES-coated silica gel shows a rather homogeneous intensity.Here, the packing is very homogeneous and the dis-tribution of water is very even. The Si–APTES–B[a]P-coated part of the column shows a heterogeneous struc-ture (position 3). Here, one has to keep in mind that thevoxels are averaging multiple pores, especially in the z-direction. The heterogeneous structure extends furtherthan the Si–APTES–B[a]P-coated part. However, thesignal values in the centre part of the column, which isnot affected by the glass boundary, reflect the lower T2

relaxation times.Another two columns are shown in Fig. 8 together

with cross-sectional intensity profiles along the column.The intensity profiles were taken from the 3D-median-filtered data with a 3×3×3 kernel. Three regions-of-interest (ROIs; size 20×6×6 mm3) were defined in thecenter of the column. The intensity values (mean andstandard deviation) for these ROIs are given in Table 4

In column 6, the signal intensity is increasing frombottom to top. This makes sense, as the column waswashed only once with 100 mL at 0.9 mL/min. Theintensity in column 7 which was flushed twice with

Table 4 Voxel intensities within ROIs in columns 6 and 7(unfiltered data)

Column Inlet Centre OutletSi–APTES Si–APTES–B[a]P Si–APTES

6 123 ± 63 234 ± 105 298 ± 1177 265 ± 121 212 ± 77 230 ± 928 646 ± 194 504 ± 176 533 ± 162

100 mL at 0.9 and 2.0 mL/min, but is otherwise identical,drops in the middle part by approximately 10% andincreases afterwards.

The T2 relaxation times were recorded in three sub-sequent imaging sequences with increasing TE (Fig. 9).The semi-logarithmic representation suggests that atleast two different relaxation times are contributingto the signal. There is also a difference in the longerT2 relaxation times between the different parts ofthe column, with the middle part (with B[a]P andanti-B[a]P-NPs) showing shorter T2 relaxation times.This is in agreement with the NMR relaxometry mea-surements. The T2 relaxation times in columns 6 and 7are half as long as in the column used previously forcolumn 8 indicating different packing. However, thenumber of data points from imaging is not sufficient fora quantitative evaluation.

The imaging results are in line with the NMR re-laxometry measurements. Apart from the usual restric-tions of MRI, the visualisation of B[a]P seems possible

Fig. 8 MRI visualisation ofcolumns 6 and 7 with profilesof the volume averaged signalintensity

50 100 150 200

010

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MRI-labelled anti-B[a]P-antibodies 2539

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Fig. 9 T2 relaxation time decay derived from MRI visualisationof columns 6–8 in the regions indicated in Figs. 7 and 8. Connect-ing lines are for visual guidance only

in coarse sediments or preferential flow paths. Thevisualisation is restricted to a feature size with a voxelvolume of a at least ten times the heterogeneity of thesurrounding matrix. That is, in homogeneous media,the feature size can be smaller whereas in a locallyheterogeneous matrix, the minimum feature size has tobe bigger. A priori knowledge about the pore topologyand a visualisation of the target before application ofNPs can help to distinguish between otherwise overlap-ping effects of changing pore sizes and water contents.

Conclusion

The use of antibody-coupled NPs is a new techniquefor the visualisation for those organic contaminants insoil which are otherwise not accessible by magneticresonance imaging or other visualisation techniques. Itwas shown that the antibody detects and binds to B[a]Ponly when it is bound to the surface with a linker. Thisis comparable to B[a]P adsorbed to humic substances insoil, but excludes B[a]P adsorbed to plain matrix. Usingthe antibody-coupled NPs, a layer of B[a]P-coated silicagel showed a significant decrease of the T2 relaxationtimes and was visualised in a commercial MRI machine.However, as multiple factors do affect the signal inten-sity in MRI, a priori knowledge about the column ma-terials and the packing is required, preferable throughMRI of the uncoated column.

The method for detection and visualisation of B[a]Pin soil at the biogeochemical interfaces presented hereis applicable for a larger group of organic pollutantsusing tailored NPs and mAbs. Antibodies for otherorganic pollutants are available or can be producedwith little effort.

Furthermore, one has to be aware that antibody-coated NPs have limited access to narrow pore spacesdue to size exclusion and charge exclusion [39, 40].In this sense, the transport pathways of the labelledanti-B[a]P-antibodies will differ considerably from thetransport pathways of dissolved contaminants. Whatseems to be a design flaw for the visualisation of thespatial distribution of contaminants might turn out asa useful feature: Interfaces which are accessible to thelabelled antibody will also be accessible to bacteria andvice versa. Hence, NPs, in contrast to dissolved trac-ers, allow a selective visualisation of biogeochemicalinterfaces.

Acknowledgments This project was funded by the DFG (Ba1592/5-1) in the framework of the priority program “Biogeo-chemical Interfaces in Soil” (SPP 1315). NMR relaxometrymeasurements were funded by the DFG (SCHA 849/8-2). Wegratefully acknowledge support from Dr. K. Achterhold (TUM,Mössbauer spectroscopy), Prof. Dr. T. F. Fässler (TUM,SQUID), M. Hanzlik (TUM, TEM), Dr. N. P. Ivleva (TUM,Raman spectroscopy) and Dr. A. Wunderlich (Univ. Ulm, MRI).

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