Robust Ligand Shells for Biological Applications of Gold Nanoparticles

Nov 7, 2008 - 1 Liverpool Institute for Nanoscale Science, Engineering and Technology, School of Biological Sciences, ... (Figure S5) Rescue experimen...
1 downloads 0 Views 1MB Size
Download PDF
13572

Langmuir 2008, 24, 13572-13580

Robust Ligand Shells for Biological Applications of Gold Nanoparticles Laurence Duchesne,* Denis Gentili, Mauro Comes-Franchini, and David G. Fernig* 1LiVerpool Institute for Nanoscale Science, Engineering and Technology, School of Biological Sciences, Biosciences Building, Crown Street, UniVersity of LiVerpool, LiVerpool, L69 7ZB, U.K. 2Department of Chemistry “A. Mangini”, UniVersity of Bologna, 40136 Bologna, Italy ReceiVed September 2, 2008. ReVised Manuscript ReceiVed October 3, 2008 An important point regarding the development of stable biofunctional nanoparticles for biomedical applications is their potential for aspecific interactions with the molecules of the biological environment. Here we report a new self-assembled ligand monolayer system for gold nanoparticles called Mix-matrices, formed by a mixture of HS-PEG and alcohol peptides (peptidols) molecules. Stability of the Mix-capped nanoparticles prepared in various conditions was assessed using tests of increasing stringency. The results highlight the importance of identifying a concentration of ligands sufficiently high to obtain a compact matrix when preparing nanoparticles and that the stability of capped nanoparticles in biological environments cannot be predicted solely on their resistance to electrolyte-induced aggregation. The Mix-capped nanoparticles are resistant to aggregation induced by electrolytes and to aspecific interactions with proteins and ligand exchange. In addition, Mix-matrices allow the easy introduction of a single recognition function per nanoparticle, allowing the specific and stoichiometric labeling of proteins with gold nanoparticles. Therefore, the Mix-matrices provide a useful tool for the development of nanoparticle-based quantitative bioanalytical and imaging techniques, as well as for therapeutic purposes, such as the specific targeting of cancerous cells for photothermal destruction.

Introduction The photostability (no equivalent of photobleaching and photoblinking), high absorption, and scattering cross sections of noble metal nanoparticles compared to fluorescent markers have led to an intense effort to harness these materials for applications in biology and medicine.1-9 They have shown great promises in many developments such as (i) highly sensitive diagnostic tests,10-12 (ii) imaging and radiotherapy enhancement,13 (iii) modulated drug delivery,14,15 and (iv) photothermal cancer therapy.16-18 The last two applications use the property of noble metal nanoparticles of emitting heat following their absorption of light. This very same property is used by photothermal * Corresponding authors. E-mail: [email protected] (L.D.) and [email protected] (D.G.F.). Tel.: (44) 151 795 4471. Fax: (44) 151 795 4406. (1) Faulk, W. P.; Taylor, G. M. Immunochemistry 1971, 8, 1081. (2) Taton, T. A.; Mirkin, C. A.; Letsinger, R. L. Science 2000, 289, 1757. (3) Schultz, D. A. Curr. Opin. Biotechnol. 2003, 14, 13. (4) Schultz, S.; Smith, D. R.; Mock, J. J.; Schultz, D. A. Proc. Natl. Acad. Sci. USA 2000, 97, 996. (5) Yguerabide, J.; Yguerabide, E. E. Anal. Biochem. 1998, 262, 137. (6) Yguerabide, J.; Yguerabide, E. E. J. Cell. Biochem. 2001, 84, 71. (7) Daniel, M. C.; Astruc, D. Chem. ReV. 2004, 104, 293. (8) Bailey, R. C.; Nam, J. M.; Mirkin, C. A.; Hupp, J. T. J. Am. Chem. Soc. 2003, 125, 13541. (9) Huang, X. H.; Jain, P. K.; El-Sayed, I. H.; El-Sayed, M. A. Nanomedicine 2007, 2, 681. (10) Mirkin, C. A.; Letsinger, R. L.; Mucic, R. C.; Storhoff, J. J. Nature 1996, 382, 607. (11) Mirkin, C. A.; Thaxton, C. S.; Rosi, N. L. Expert ReV. Mol. Diagn. 2004, 4, 749. (12) Hirsch, L. R.; Jackson, J. B.; Lee, A.; Halas, N. J.; West, J. L. Anal. Chem. 2003, 75, 2377. (13) Hainfeld, J. F.; Slatkin, D. N.; Focella, T. M.; Smilowitz, H. M. Br. J. Radiol. 2006, 79, 248. (14) West, J. L. Nat. Mater. 2003, 2, 709. (15) Ghosh, P.; Han, G.; De, M.; Kim, C. K.; Rotello, V. M. AdV. Drug. DeliV. ReV. 2008, 60, 1307. (16) Hirsch, L. R.; Stafford, R. J.; Bankson, J. A.; Sershen, S. R.; Rivera, B.; Price, R. E.; Hazle, J. D.; Halas, N. J.; West, J. L. Proc. Natl. Acad. Sci. USA 2003, 100, 13549. (17) Loo, C.; Lin, A.; Hirsch, L.; Lee, M. H.; Barton, J.; Halas, N.; West, J.; Drezek, R. Tech. Canc. Res. Treat. 2004, 3, 33. (18) El-Sayed, I. H.; Huang, X. H.; El-Sayed, M. A. Cancer Lett. 2006, 239, 129.

microscopy,19-21 which allows the detection and tracking of nanoparticles as small as 2 nm at a single entity level, opening the door for the development of quantitative biosensors to study in real-time and at a molecular resolution the dynamic fluctuation of molecules. Such biological and medical applications notably require high stability of nanoparticles in physiological conditions while maintaining their optical properties. It is important to note that stability from a biological point of view means solubility/ dispersion in physiological environments, as well as the absence of aspecific interactions and both these requirements are critical. Solubility can be achieved by means of a passivating ligand shell. Previously reported strategies include the use of thiol ligands with hydrophilic end groups, which form a self-assembling monolayer at the metal surface22-26 or the use of polymeric capping agents, such as amino- or mercapto-dextrans.27 Selfassembling monolayers provide important advantages over polymers. The hydrodynamic radius of the nanoparticle is kept to a minimum, which is of primary importance for many biomedical applications. It avoids steric hindrance of the nanoparticle probe, which would prevent molecular interaction studies, as well as the proper penetration of the nano-objects in some constrained biological spaces such as synapses or cell junctions. A number of self-assembling ligand shells have been described, ranging from PEGylated alkanethiols to peptides.22-26 (19) Lasne, D.; Blab, G. A.; Berciaud, S.; Heine, M.; Groc, L.; Choquet, D.; Cognet, L.; Lounis, B Biophys. J. 2006, 91, 4598. (20) Boyer, D.; Tamarat, P.; Maali, A.; Lounis, B.; Orrit, M. Science 2002, 297, 1160. (21) Berciaud, S.; Cognet, L.; Blab, G. A.; Lounis, B. Phys. ReV. Lett. 2004, 93, 257402. (22) Strong, L.; Whitesides, G. M. Langmuir 1988, 4, 546. (23) Bartz, M.; Kuther, J.; Nelles, G.; Weber, N.; Seshadri, R.; Tremel, W. J. Mater. Chem. 1999, 9, 1121. (24) Templeton, A. C.; Wuelfing, M. P.; Murray, R. W. Acc. Chem. Res. 2000, 33, 27. (25) Pengo, P.; Polizzi, S.; Battagliarin, M.; Pasquato, L.; Scrimin, P. J. Mater. Chem. 2003, 13, 2471. (26) Levy, R.; Thanh, N. T. K.; Doty, R. C.; Hussain, I.; Nichols, R. J.; Schiffrin, D. J.; Brust, M.; Fernig, D. G. J. Am. Chem. Soc. 2004, 126, 10076. (27) Wilson, R. Chem. Commun. 2003, 108.

10.1021/la802876u CCC: $40.75  2008 American Chemical Society Published on Web 11/08/2008

Ligand Shells for Biological Applications

They follow a basic design starting with a thiol group for bonding to the metal, a hydrophobic core, allowing packing of the monolayer, and a hydrophilic terminus exposed to solvent to enable solubility in physiological environments. These selfassembling monolayers (SAM) all impart excellent solubility to the metal nanoparticle. For example, PEGylated alkanethiols and peptides, such as CALNN,26 protect silver nanoparticles from oxidation in water containing electrolytes and prevent the electrolyte-induced aggregation of gold and silver nanoparticles.26,28 Moreover, SAM of ligands can be designed to present biomolecular recognition functions that are readily accessible at the surface of the nanoparticle and that may be incorporated in a controlled valency.29 For example, the recently described system of pentapeptides provides a simple means to produce nanoparticles carrying a single biomolecular recognition function.29,30 Despite the fantastic progress achieved regarding the solubility and the functionalization of gold nanoparticles using SAM that keep the hydrodynamic size of the nanoparticles to a minimum, the problem regarding the aspecific binding of capped-nanoparticles has been often over looked and has limited efforts to harness the remarkable optical properties of metal nanoparticles for biological and medical applications. Indeed, biological environments are extremely complex, as they contain high concentrations of macromolecules (typically 200-400 mg/mL nucleic acids, proteins, carbohydrates, lipids) with a variety of functional groups, some of which (amines, carbonyl, thiol) will bind to the metal nanoparticle itself or to the matrix protecting the nanoparticle. Aspecific binding and exchange of ligands in the passivating ligand shell of nanoparticles designed to recognize a specific target for labeling or thermotherapy applications will lead to a change of the “homing address” and/or the loss of the recognition function. Thus, both aspecific interactions of the nanoparticles and ligand exchange must be prevented. Here we have developed a new self-assembled ligand monolayer system, which is shown to impart, in a graded series of tests of increasing stringency, high stability in physical, chemical, and biological terms to nanoparticles, while maintaining the advantages of simple, controlled functionalization of previous systems that have been developed. In addition, we show the importance of using the appropriate concentration of ligand molecules according to the nanoparticle surface area to obtain a maximum coverage of the surface and, therefore, a maximum protection.

Experimental Section Materials. Peptides CALNN, CALNNGHHHHHHGKbiotinG (CALNN-6xHis-Biotin), CALNNGKGALVPRGSGKbiotinTAK (CALNN-Biotin) were purchased from Sigma-Genosys Ltd. (Haverhill, UK). PEGylated alkanethiol, HS-EC11-EG4, Mw 380, (referred to as HS-PEG), was purchased from Prochimia (ProChimia Surfaces Sp. z o.o., Sopot, Poland). The PEGylated alkanethiol HS-C16-EG3, Mw 420, and the HS-C16-EG3-Tris-NTA were a gift from R. Tampe´ (Johann Wolfgang Goethe University, Frankfurt, Germany). The CVVVTGHHHHHHGKbiotinG (CVVVT-6xHis-Biotin) peptide and the CVVVT-ol, CSSSS-ol, and CALNN-ol modified peptides (T-ol is for threoninol, S-ol is for serinol, and N-ol is for asparaginol) were from Anaspec (Anaspec Inc., San Jose, CA). All other peptidols were purchased from Peptide and Protein Research (PPR Ltd., Hampshire, UK). Gold nanoparticles (G-NPs), of diameter 5 and 10 nm and stabilized in citrate buffer, were purchased from British Biocell (BBInternational Ltd., UK). Sephadex G25 superfine, bovine (28) Doty, R. C.; Tshikhudo, T. R.; Brust, M.; Fernig, D. G. Chem. Mater. 2005, 17, 4630. (29) Levy, R.; Wang, Z. X.; Duchesne, L.; Doty, R. C.; Cooper, A. I.; Brust, M.; Fernig, D. G. Chembiochem 2006, 7, 592. (30) Levy, R. Chembiochem 2006, 7, 1141.

Langmuir, Vol. 24, No. 23, 2008 13573 serum albumin, Tween 20, streptavidin-agarose, and antiflag affinity resin were from Sigma-Aldrich Ltd. (Dorset, UK). Secondary antibodies to rabbit and mouse immunoglobin Gs and SuperSignal West Pico chemiluminescent substrate were from Pierce (Perbio Science UK Ltd.). Immobilon-P polyvinylidene difluoride (PVDF) membrane was from Millipore (Watford, UK). Strep-TactinMacroprep, Strep-Tactin-Sepharose, and Strep-Tactin-HRP were from IBA (Goettingen, Germany), and Nanosep centrifugal ultrafiltration devices were from PALL (PALL Corp., Portsmouth, Hants, UK). Heparin agarose was purchased from Biorad (BioRad, Hemel Hempstead, UK). Diethyl-Amino-Ethyl (DEAE) Sepharose superflow and Carboxy-Methyl (CM) Sephadex were from GE Healthcare. Nickel chelating resin (called Probond resin) was from Invitrogen (Paisley, UK). Hepatocyte growth factor/scatter factor (HGF/SF) and antibodies to FGF-2 and HGF/SF were from R&D Systems (Abingdon, Oxfordshire, UK). Recombinant hexa-histidine tagged FGFR1 was prepared as described.31 Preparation of the Matrix Ligands. A 2 mM CALNN peptide stock solution was prepared by dissolving the peptide powder in 10-fold concentrated phosphate-buffered saline (10X PBS, a 10fold concentrated solution of PBS, 8.1 mM Na2HPO4, 1.2 mM KH2PO4, 140 mM NaCl, and 2.7 mM KCl, pH 7.4). CVVVT-ol peptide (4 mM) was prepared by dissolving the powder in DMSO: H20 25:75 (v/v). Stock solutions of PEGylated alkanethiols (HSPEG, HS-C11EG4, and HS-C16-EG3) at 5 mM were prepared using methanol (AR grade). All stock solutions were kept as aliquots at -20 °C. Before use, matrix ligands were diluted to the appropriate concentration (2, 1, 0.5, or 0.1 mM) using PBS 10X for CALNN or milliQ H20 for HS-PEG and CVVVT-ol. When not specified, the concentration of ligand used is 2 mM, the HS-PEG ligand used is HS-C11EG4 (Mw 380), and the ratio CVVVT-ol to HS-PEG is 70:30 (v/v). Other ratios of CVVVT-ol to HS-PEG were tested and are called Mix 50:50, Mix 60:40, Mix 80:20, and Mix 90:10. Preparation of Ligand-Capped Gold Nanoparticles. Capped nanoparticles were prepared by adding 9 volumes of colloidal gold solution to 1 volume of the matrix ligand (CALNN or Mix) at 2, 1, 0.5, or 0.1 mM. PBS10X and Tween-20 were then added to obtain final concentrations of PBS1X and Tween-20 0.005% (v/v) (PBST0.005%). Matrix ligands and nanoparticles were left to react overnight at room temperature (see Supporting Information Figure S1 for details regarding how the ligands attach to the gold surface). Excess ligands were removed by size-exclusion chromatography using Sephadex G25 superfine or by ultrafiltration using Nanosep centrifugal ultrafiltration devices (30 kDa cutoff). Capped nanoparticles prepared using 2, 1, 0.5, and 0.1 mM of CALNN ligand solution are respectively called CALNN-2 mM, -1 mM, -0.5 mM, or -0.1 mM. The equivalent prepared using the Mix ligands are respectively called Mix-2 mM, -1 mM, -0.5 mM, or -0.1 mM. If not specified, 2 mM of ligands has been used. UV-Visible Spectrometry. Absorption spectra were recorded at room temperature using a Spectra Max Plus spectrophotometer (Molecular Devices, Wokingham, UK). Functionalization of Nanoparticles. Functionalization was performed as described previously.29,32 Briefly, ligand bearing a specific recognition function (called functional ligand) is mixed at a desired molar ratio with the matrix ligands prior to the addition of nanoparticles and the remainder of the procedure is as in Preparation of Ligand-Capped Gold Nanoparticles above. Calculation of the Percentage of Bound/Unbound or Eluted Nanoparticle from Chromatography Resins. The absorbance at 520 nm of free nanoparticles in solution is measured following the reaction with the chromatography resin (unbound/eluted) and for mock-treated samples (PBS added instead of resin). Percentage (%) of unbound/eluted material ) Abs520nm unbound nanoparticles/ Abs520nm mock treated × 100. Percentage of bound material ) 100 - unbound (%). (31) Duchesne, L.; Tissot, B.; Rudd, T. R.; Dell, A.; Fernig, D. G. J. Biol. Chem. 2006, 281, 27178. (32) Duchesne, L.; Wells, G.; Fernig, D. G.; A, H. S.; Levy, R. Chembiochem 2008, 9, 2127.

13574 Langmuir, Vol. 24, No. 23, 2008 Electrolyte-Induced Aggregation Experiment. Capped nanoparticles were centrifuged for 1 h, 4 °C at 17 000 g. The pellet was resuspended in sodium phosphate buffer 10 mM, pH 7.4, and the solution was split into 70 µL aliquots. NaCl (5 M) was added to obtain 0.15, 0.25, 0.5, 0.75, 1, or 1.5 M NaCl final concentration, and after a 24 h incubation, the absorption spectra were recorded at room temperature. Anion-ExchangeChromatography.Diethyl-Amino-Ethyl(DEAE) Sepharose was prepared as specified by the manufacturer and equilibrated in PBS. Capped nanoparticles were centrifuged for 1 h, 4 °C at 17 000 g. The pellet was resuspended in PBS and 70 µL was added to 10 µL of DEAE-Sepharose in a 0.5 mL tube. After 30 min incubation on a rotating wheel the unbound material was recovered. Successive washes of the resin were then performed using PBS, PBS-T, and sodium phosphate buffer 10 mM pH 7.3 with an increasing amount of NaCl. After each wash, the supernatant (eluate) was recovered and the amount of nanoparticles in each fraction was quantified by the measurement of the absorbance at 520 nm. Pull-Down Experiments Using Chromatographic Resins. Chromatographic resins were equilibrated in the same buffer as that containing the nanoparticles. An aliquot of between 70 and 200 µL of nanoparticles was added to 5 or 10 µL of beads and incubated 2 h at room temperature on a rotating wheel. The beads were left to sediment and the amount of unbound nanoparticles was quantified by the measurement of the absorbance at 520 nm of the supernatant and compared to the initial amount. Preparation of the Poly Histidine Resin (Affi-His). CVVVTGHHHHHH (CVVVTG-6xHis) peptide (5 mL at 2 mM) in DMSO was added to 5 mL of Affi10 beads (Biorad) washed in DMSO. The reaction was left 2 h at room temperature and stopped by addition of 10 mM Tris-HCl pH 6.8. The resin was then washed with 5 × 5 volumes of PBS and kept at 4 °C until further use. Size-Exclusion Chromatography. Sephadex G25 superfine, 1 mL in a 10 mL column, was prepared following the manufacturer’s recommendations and equilibrated in PBS-T (PBS supplemented with 0.005% (v/v) Tween-20). Nanoparticles (100 µL) were loaded, and 100 µL fractions were collected. The amount of nanoparticles in each fraction was quantified by the measurement of the absorbance at 520 nm. Rescue Experiment. Mix- and CALNN-capped-nanoparticles were prepared using a starting concentration of 0.1 mM ligand. Capped nanoparticles were then added to a solution of 2 mM ligand in a 9:1 ratio (v/v) (nanoparticles:ligands). After 2 h incubation, excess ligands were removed and anion-exchange chromatography was performed. Ligand Exchange. Functional ligands, 2 µL at 200 µM, were added to 98 µL CALNN- or Mix-capped nanoparticles (8 nM in PBS) and the reaction was left 4 h at room temperature under agitation. Functional ligands used were the following: CVVVTGHHHHHHGKbiotinG (CVVVT-6xHis-biotin) and CALNNGHHHHHHGKbiotinG (CALNN-6xHis-biotin). Excess ligands were removed using ultrafiltration and the presence of the functional ligand on the nanoparticle was checked by pull-down using nickel chelating resin (Probond). Test for Aspecific Binding of Proteins. FGF-2 (Fibroblast Growth Factor 2) and HGF/SF, (Hepatocyte Growth Factor/Scatter Factor) proteins (5 µL at 1.2 µM) were added to 95 µL CALNN- or Mixcapped nanoparticles at 8 nM in PBS. The reaction was left 3 h at room temperature under agitation and the nanoparticles were then isolated by centrifugation: the nanoparticles were centrifuged 40 min, 4 °C, 17 000 g, and the pellet (10 µL) was resuspended in 400 µL of PBS and centrifuged again. Six centrifugation cycles were performed. Dot-Blot. Nanoparticles in PBS were added to PVDF membranes. Membranes were then blocked by incubation for 40 min at room temperature in PBS containing 5 mg/mL of BSA (PBS-BSA). Afterward, for detection of the FGF-2 and HGF/SF proteins, the membranes were incubated 2 h at room temperature in PBST0.1%-BSA (PBS- supplemented with 0.1% (v/v) Tween 20 (PBST0.1%) and 5 mg/mL of BSA) with rabbit antibody to FGF-2 or a mouse antibody to HGF/SF diluted 1:1500. The blots were

Duchesne et al. washed with PBST0.1%-BSA (5 times, 5 min), incubated 2 h at room temperature in PBST0.1%-BSA with peroxidase conjugated secondary antibodies to rabbit or to mouse immunoglobin Gs diluted 1:2000 and then were washed with PBST (7 times, 5 min). For detection of biotin, the membranes were incubated 40 min at room temperature in PBST0.5%-BSA (PBS supplemented with 0.5% (v/ v) Tween 20 (PBST0.5%) and 5 mg/mL of BSA) with peroxidase conjugated Strep-Tactin-HRP diluted 1:7500 and then were washed with PBST (7 times, 5 min). Visualization was performed using enhanced chemiluminescence (SuperSignal West Dura Substrate, Pierce). Number of Ligands per Nanoparticle. CALNN- and MIXcapped nanoparticles (9.6 nm ( 10% diameter) prepared using 2 or 0.1 mM ligand solution were purified from excess ligand by sizeexclusion chromatography. The extinction coefficient of the nanoparticles, Σ520nm ) 0.08 × 109 L/(mol cm), was deduced using mean free path corrected Mie theory.33 The nanoparticle concentration was determined using this value of Σ520nm and the absorbance value (Abs520nm) according to the Beer-Lambert law. The corresponding peptide concentration was measured by amino acid analysis (Alta Bioscience, Birmingham, UK). Preparation of the Tris-NiNTA Mix-Capped Nanoparticles. CVVVT-ol:HS-PEG (70:30) Mix-capped nanoparticles bearing none (n ) 0), an average of 0.1, or an average of 3 Tris-NTA functions per nanoparticle were prepared, as described in Functionalization of Nanoparticles above. Excess ligand was removed by Sephadex G25 chromatography using H20 supplemented with 200 mM of NaCl (H20-NaCl) as the mobile phase. NiSO4 (250 mM final concentration) was added to the capped nanoparticles and the reaction was left 2 h at room temperature on a rotating wheel. Excess nickel was removed by G25 chromatography using H20-NaCl as the mobile phase and then the nanoparticles were buffer exchanged to PBS using Nanosep centrifugal ultrafiltration devices. Nanoparticles bearing at least one Tris-NiNTA function were then purified by affinity chromatography using Affi-His resin. Briefly, 1 volume of resin was added for 10 volumes of nanoparticles and the reaction was left to incubate for several hours on a rotating wheel. The resin was then poured into a column and washed with 10 resin volume equiv of PBS. Elution was performed using PBS supplemented with 200 mM imidazole. Excess imidazole removal and buffer exchange to milliQ H20 were performed using Nanosep filters. Nanoparticles were then loaded again with nickel. Excess nickel removal and buffer exchange to PBS were performed using Nanosep filters. For the capped nanoparticles prepared with an average of 0.1 tag Tris-NiNTA tags per nanoparticle 90% of the nanoparticles fail to bind to the Affi-His resin; 10% of the nanoparticles are retained on the resin and eluted by imidazole and will possess a single Tris-NiNTA function (n ) 1). Specific Coupling of the Tris-NiNTA-Capped Nanoparticles to Proteins. The proteins used were a recombinant fibroblast growth factor receptor 1 (FGFR1) bearing a hexa-histidine tag at its aminoterminal extremity as well as a Step-Tag at its carboxy-terminal extremity31 and a recombinant fibroblast growth factor 2 (FGF-2) with an amino-terminal hexa-histidine tag. FGFR1 (1 µL at 32 µM) was added to 10 µL of Mix-capped nanoparticles (5 nm diameter, 0.5-1 µM concentration) having none (n ) 0), one (n ) 1), or an average of 3 Tris-NiNTA functions per nanoparticle. The reaction was left 4 h at room temperature on a rotating wheel and the mix was diluted to 50 µL and added to 10 µL of Strep-Tactin Sepharose resin (affinity for the Strep-Tag of the FGFR1). Several washes in PBS (5 times, 20 column volumes) were performed to remove nanoparticles not conjugated to the FGFR1 (control, n ) 0). Nanoparticles conjugated to FGFR1 were eluted using 1 mM biotin in PBS (500 µL) and were then centrifuged to remove any unconjugated FGFR1 and free biotin. Centrifugation was performed for 90 min at 17 000 g 4 °C, and the supernatant (free biotin and protein) was removed. The pellet was resuspended in 400 µL of PBS and centrifuged again; a total of 5 cycles of centrifugation were (33) Haiss, W.; Thanh, N. T. K.; Aveyard, J.; Fernig, D. G. Anal. Chem. 2007, 79, 4215.

Ligand Shells for Biological Applications

Langmuir, Vol. 24, No. 23, 2008 13575

Figure 1. Stability against electrolyte-induced aggregation of Mix CVVVT-ol:HS-PEG capped nanoparticles. Mix-capped gold nanoparticles (10 nm) were prepared using different ratios of CVVVT-ol:HS-PEG. (A) Electrolyte-induced aggregation determined by measuring the absorbance spectra after 24 h incubation in sodium phosphate buffer 10 mM, pH 7.4 supplemented with 0 mM, 250 mM, or 1 M of NaCl. (B) Specific binding to DEAE-Sepharose indicated by red color of the pellet. Ligand ratios are shown at the top of the panel. The resin was washed 2 times with PBS before picture acquisition.

performed. At the end, the pellet (nanoparticles bound to FGFR1) was resuspended with PBS to obtain a final concentration of 100 nM of nanoparticles. Identical amounts of capped nanoparticles with different levels of incorporation of Tris-NiNTA (n ) 0, n ) 1, n ∼ 3) were then dotted to a PVDF membrane to check the presence of the FGFR1 protein by dot-blot. For FGF-2, 0.5 µL at 65 µM was added to 10 µL of Mix-capped nanoparticles (5 nm diameter, 0.5-1 µM concentration) having none (n ) 0) or one (n ) 1) Tris-NiNTA functions per nanoparticle. The reaction was left overnight at 4 °C on a rotating wheel. The mix was diluted to 400 µL and was centrifuged (90 min at 17 000 g 4 °C) to remove any unconjugated protein. The pellet was resuspended in 400 µL of PBS and centrifuged again; a total of 5 cycles of centrifugation were performed. From this point the procedure was identical to that described for FGFR1, above. Expression and Purification of the Hexa-Histidine Tagged FGF-2 Protein. A construct encoding the poly histidine tagged FGF-2 protein (6xHis-FGF-2) was obtained using as a template the human fgf-2 sequence (corresponding to amino acids 1-155) described in ref 34. By performing add-on PCR (polymerase chain reaction) using a set of appropriate primers (forward, 5′CTCGGGCCATGGGCCATCATCACCATCACCATCTGGTTCCGCGTGGTTCAATGGCAGCCGGGAGCATCACCACG-3′ and reverse, 5′GCAGCCGGATCCTTATCAGCTCTT-3′), sequences encoding the thrombin cleavage site (italic) and a poly histidine tag (bold) were introduced. The PCR insert was digested using NcoI and BamH1 enzymes and subcloned in the pET14 expression vector. Purification of the hexa-histidine tagged FGF-2 was performed as described in ref 34.

Result and Discussion Mixed Matrix Ligand to Cap Metal Nanoparticles. Peptides as capping ligands provide excellent solubility to nanoparticles in an electrolyte environment. In addition, peptide chemistry is very versatile and provides a many possibilities for the simple incorporation of very diverse recognition functions at the carboxyl-terminus of the peptide, which is exposed to solvent. However, peptide-capped nanoparticles are negatively charged26 due to their carboxylic acid terminus, which is exposed to solvent, (34) Ke, Y.; Wilkinson, M. C.; Fernig, D. G.; Smith, J. A.; Rudland, P. S.; Barraclough, R. Biochim. Biophys. Acta 1992, 1131, 307.

and are, therefore, likely to interact aspecifically with biological molecules having positively charged patches. Modified peptides with a carboxyl-terminal amino alcohol (peptidols), rather than a carboxylic acid, were tested as potential ligands to cap metal nanoparticles. The rationale was that these ligands would provide the same high stability as PEG and peptides, while maintaining the functional versatility allowed by peptides,29,30,32,35,36 but without contributing ionizable groups to the nanoparticle surface. Several peptidol sequences were tested (Table 1 in the Supporting Information), but none stabilized gold nanoparticles against electrolyte-induced aggregation (data not shown). On the basis that the carboxyl-terminal alcohol was not sufficiently mobile to prevent aggregation in the absence of charge repulsion, a defined amount of PEGylated alkanethiol was incorporated within the matrix of peptidols to increase the stability of the nanoparticles. PEGylated alkanethiol ligands are known to impart high stability and low aspecific binding to metal nanoparticles.23,25 The ethylene glycol group is hydrophilic, but not charged at physiological pH; the stability imparted to the nanoparticle (Figure S2 in the Supporting Information) is probably due to the high flexibility of the ethylene glycol groups preventing aggregation. Different ratios of peptidol:HS-PEG and different peptidol sequences were tested (Figures 1, S3, and S4 (see the Supporting Information)). For most of the peptidols tested, the introduction of 10% (mol:mol) or more of HS-PEG ligands within the peptidol matrix prevented electrolyte-induced aggregation of the Mix-capped nanoparticles (Figure 1A, S3, and S4 (see the Supporting Information)). The CVVVT-ol peptidol was chosen for all further experiments, because the hydrophobic valine side chains were considered to be most compatible in hydrophobicity and length with the alkane chain of the C11 HS-PEG. Ligand matrices consisting of peptidol (CVVVT-ol by default) and HSPEG are hereafter referred to as “Mix-matrices” and nanoparticles capped by this Mix-matrix are referred to as “mixed-capped nanoparticles”. When the molar ratio of CVVVT-ol:HS-PEG was 95:5, aggregation was observed (Figure 1A), indicating that (35) Wang, Z.; Levy, R.; Fernig, D. G.; Brust, M. Bioconjug. Chem. 2005, 16, 497. (36) Wang, Z.; Levy, R.; Fernig, D. G.; Brust, M. J. Am. Chem. Soc. 2006, 128, 2214.

13576 Langmuir, Vol. 24, No. 23, 2008

the minimum level of HS-PEG required for stability against electrolyte-induced aggregation is between 10% and 5%. Electrolyte-induced aggregation, which causes the plasmon absorbance of metal nanoparticles to shift to longer wavelengths due to coupling of the nanoparticle dipoles, is one of the most common tests of nanoparticle stability and is often extrapolated to indicate the likely stability of the nanoparticles in a biological environment. However, regarding biological and medical applications, the stability of nanoparticles cannot be solely defined by the absence of aggregation in solutions of electrolytes. Biological environments are far more complex in term of composition and nanoparticles must be inert toward this environment. Chromatography is a standard approach for the separation and purification of biological molecules according to their surface properties and commonly involve, size-exclusion, ion-exchange, hydrophobic and affinity chromatography that allow the separation of molecules according to their size, charge, hydrophobicity and the presence of a recognition sequence, respectively. Given the fact that aspecific, as well as specific interaction between biological molecules are driven, in the first place, by the same non covalent interactions exploited by the chromatographic techniques, the latter provide a rapid and easy way to evaluate the properties of the capped nanoparticles and to estimate their likely tendency to interact aspecifically with the molecules in a biological environment. Indeed, nanoparticles that cannot chromatograph on the stationary phase because of interactions in mild physiological conditions are likely to behave the same way in a biological environment. Therefore, we tested the behavior of Mix-capped nanoparticles prepared with different ratios of peptidol:HS-PEG in presence of DEAE-Sepharose, which is a low pressure anion-exchange matrix. Mix-2 mM nanoparticles prepared with more than 20% of HS-PEG do not bind to DEAE-Sepharose, which is expected since they present hydroxyl groups to solvent. However, when using lower ratios of HS-PEG, aspecific binding was observed and this phenomenon increased with the decrease of the HSPEG:peptidol ratio of the matrix (Figure 1B). Thus, the mixed ligand matrix with at least 20% HS-PEG provides excellent stability for the nanoparticles with respect to the electrolyteinduced aggregation and protection from aspecific binding to matrices such as DEAE-Sepharose. Further experiments were then performed using 20% (mol:mol) or more of HS-PEG within the peptidol matrix. Functionalization of Mix-Capped Nanoparticles. Next, the ability to introduce, in a controlled manner, a functional peptide within this Mix-matrix was tested. A key issue with respect to biological applications of nanoparticles is the ease and diversity of their functionalization. A well-described pentapeptide ligand, CALNN, enables the facile functionalization of nanoparticles, while providing control over the number of functions incorporated into the ligand shell.29,32 Therefore, it was important to determine if a peptide carrying a function at its carboxy-terminus (so exposed to solvent) could be incorporated in a similarly easy and controlled manner into the Mix-matrix nanoparticles. The functionalization was achieved by mixing a peptide bearing a specific recognition function at its carboxyl-terminus (called functional ligand) at the desired molar ratio with the matrix ligands prior to the addition of nanoparticles. When the concentration of HS-PEG ligand increased within the mixed matrix, the incorporation of the functional ligand decreased (Figure 2). Nevertheless, for the mixes 70:30 and 50:50, the level of incorporation of the functional peptide was still in the same range as that observed for the CALNN peptide matrix in the same conditions, and far higher than that

Duchesne et al.

Figure 2. Incorporation of a functional peptide (CALNNGKGALVPRGSGKbiotinTAK) within the stabilizing matrix (CALNN, HS-PEG, Mix 70:30 and Mix 50:50). Capped nanoparticles were prepared with different proportions of functional peptide (Pf). The percentage of biotinylated nanoparticles is measured as the proportion of nanoparticles pulleddown by streptavidin-agarose beads.

observed for HS-PEG only ligand shells. Indeed, the incorporation of a peptide ligand during the preparation of the HS-PEG capped nanoparticles is extremely inefficient and less easily controlled compared to peptide based capped nanoparticles (Figure 2). Moreover, whereas a range of modified/functionalized PEGylated alkanethiols have been synthesized, the array of functions that can easily be achieved is far more limited compared to the versatility of peptide chemistry. Therefore, the mixed ligand shell produced with ratios of peptidol:HS-PEG of 80:20 to 50:50 provides a good combination of stability, with respect to electrolyte-induced aggregation and ease of incorporation of a functional ligand. Moreover, with different ligands, the mixed ligand shell provides a greater potential for functionalization through either peptide or HSPEG moieties. Performance of the Mix-Capped Nanoparticles, Effect of Ligand Concentration. The 70:30 ratio (mol:mol) of CVVVTol:HS-PEG, called Mix-matrix, was, therefore, used in further experiments that aimed to test its performance in increasingly stringent conditions. Moreover, to test the effect of ligand grafting density on the nanoparticles stability, nanoparticles with a shell formed over a 20-fold range of ligands concentrations (from 0.1 to 2 mM ligand) were prepared. In the electrolyte-induced aggregation test, no difference was observed for nanoparticles capped using 2-0.5 mM of Mix ligands, but the plasmon absorbance of nanoparticles formed using a ligand concentration of 0.1 mM was shifted to longer wavelengths in the presence of 0.15 M NaCl (Figure 3A), characteristic of nanoparticle aggregation. To explore further the stability of the Mix-capped nanoparticles, they were subjected to gel-filtration on Sephadex G-25 (sizeexclusion chromatography) and anion-exchange chromatography on DEAE-Sepharose. Size-exclusion chromatography performed in PBS shows that more that 90% of the Mix-2 mM and Mix-1 mM capped nanoparticles are recovered in the excluded volume of the column confirming the absence of aggregation and the proper dispersion of such nanoparticles while in presence of an inert stationary phase (Figure 3B). Thus, the amount of nanoparticles recovered in the excluded volume of the column fell to 74% and 42% when 0.5 and 0.1 mM mixed ligands were used, respectively (Figure 3B). The remainder of the nanoparticles did not chromatograph, but instead, they were bound irreversibly to the Sephadex G-25.

Ligand Shells for Biological Applications

Figure 3. Stability of Mix- and CALNN-capped gold nanoparticles and effect of ligand concentration. Mix (A-C) and CALNN (D-F) capped gold nanoparticles were prepared using initial concentrations of ligands of 2, 1, 0.5, or 0.1 mM. Following purification, stability parameters of the nanoparticles were tested. (A and D) Electrolyte-induced aggregation experiments: absorbance spectra of the Mix- (A) and CALNN- (D) capped nanoparticles in the presence of increasing concentrations of NaCl (24 h incubation in NaCl). (B, E) Percentage of recovery following G-25 size-exclusion chromatography. (C, F) Elution profile on anionexchange chromatography using increasing concentrations of NaCl. (insets) Red color of nanoparticles allowing the direct visualization of the presence of nanoparticles in the pellet (nanoparticles bound to the beads) and in the supernatant (unbound nanoparticles).

Regarding the behavior with DEAE anion-exchange resin, the Mix-2 mM capped nanoparticles did not bind to such resin, as shown above (Figure 1B and 3C, inset). However, a small percentage of Mix-1 mM nanoparticles were found to bind to DEAE in the presence of PBS, but they were eluted along the first NaCl washes, whereas the similar population of Mix-0.5 mM nanoparticles that bound to the column could not be eluted, even with 2 M NaCl, indicating that they were nonspecifically adsorbed. The recovery of nanoparticles formed with 0.1 mM mixed matrix was very poor (∼20% of the material loaded) (Figure 3C, insert), which is the result of these nanoparticles absorbing to DEAE-Sepharose in an electrolyte-independent manner. On a cation-exchange resin, CM-Sephadex, no binding

Langmuir, Vol. 24, No. 23, 2008 13577

was observed for the Mix-2 mM nanoparticles (Supporting Information, Figure S6). To compare the Mix-capped nanoparticles with peptide-capped nanoparticles, experiments using CALNN-capped nanoparticles were run in parallel. CALNN-capped nanoparticles prepared over a 20-fold range of CALNN concentrations (from 0.1 to 2 mM CALNN ligand) show no major differences in their absorbance spectra in the presence of 0.15 and 1.5 M of NaCl (Figure 3D). In addition, size-exclusion chromatography performed in PBS shows that more that 90% of the CALNN-capped nanoparticles are recovered in the excluded volume of the column, regardless of the concentration of CALNN used to form the ligand shell (Figure 3E). The CALNN peptide on the nanoparticles present their carboxyl-terminal carboxylic acid to solvent26 and binds strongly, as expected, to DEAE-Sepharose (Figure 3F, insert). Interestingly, as the concentration of CALNN used to form the ligand shell is decreased from 2 to 1 and 0.5 mM, the concentration of NaCl required to elute the nanoparticles is increased. At 0.1 mM CALNN, the ligand shell is no longer able to prevent the nanoparticles absorbing in an electrolyte-independent manner; the nanoparticles are irreversibly bound to the DEAE-Sepharose (Figure 3F, insert). Therefore, Mix-capped nanoparticles prepared using more than 1 mM ligand are resistant to electrolyte aggregation and neutral. When prepared with less ligand, changes of behavior are observed in the electrolyte-aggregation test as well as in size-exclusion and DEAE chromatographies. Differently, CALNN-capped nanoparticles are resistant to electrolyte aggregation and are well recovered in size-exclusion chromatography regardless of the concentration of ligand used in their preparation. However, when prepared with 1 mM or less of ligands changes are observed upon chromatography on DEAE-Sepharose. One interpretation of the difference in elution of cappednanoparticles formed with different concentrations of ligand from DEAE-Sepharose is that the ligand SAM are organized differently, with a lower grafting density occurring at lower concentrations of ligand. Counting the Number of Ligands. To determine whether the ligand grafting did indeed depend on the initial concentration of ligand, nanoparticles prepared using 0.1 and 2 mM of ligands were subjected to amino-acid analysis. The analysis revealed a grafting density of 3.6, 1.6, 2.4, and 1.7 ligands per squared nanometer for Mix-2 mM, Mix-0.1 mM, CALNN-2 mM, and CALNN-0.1 mM, respectively, which corresponds to 1060, 462, 704, and 512 ligands (peptide/-ol) per nanoparticle (9.6 nm diameter). Given the volume of the ligand37 and the surface occupied by a cysteine on a gold surface,38,39 (Supporting Information, section 2, Theoretical number of pentapeptides per nanoparticle for an optimally packed SAM), 1060 peptidols for the Mix-2 mM corresponds to a reasonably tightly packed matrix. It is an assumption that the percentage incorporation of the 30% (mol:mol) HS-PEG molecules into the ligand shell is not affected by the total ligand concentration (2 mM versus 0.1 mM). However, even if the surface occupied by the HS-PEG ligands is lower than expected due to substoichiometric incorporation, the grafting density of the CVVVT-ol peptides for Mix-2 mM capped nanoparticles is at least 1.5-fold higher than that observed with the CALNN-2 mM one. Since the volume of the CALNN and CVVVT-ol molecules themselves are similar (0.62 and 0.66 nm3, respectively), the higher grafting density observed for the Mix-2 mM matrix compared to CALNN-2 mM matrix may reflect the (37) Zamyatin, A. A. Prog. Biophys. Mol. Biol. 1972, 24, 107. (38) Nazmutdinov, R. R.; Zhang, J.; Zinkicheva, T. T.; Manyurov, I. R.; Ulstrup, J. Langmuir 2006, 22, 7556. (39) Dodero, G.; De Michieli, L.; Cavalleri, O.; Rolandi, R.; Oliveri, L.; Dacca, A.; Parodi, R. Colloids Surf. A: Physicochem. Eng. Asp. 2000, 175, 121.

13578 Langmuir, Vol. 24, No. 23, 2008

Duchesne et al.

Figure 4. Ligand-exchange and aspecific binding of Mix- and CALNN-capped nanoparticles. (A) Ligand-exchange experiment: capped nanoparticles incubated with CVVVT-6xHis-biotin and CALNN-6xHis-biotin peptides and purified. The percentage of ligand-exchange corresponds to the percentage of nanoparticles incorporating at least one 6xHis-biotin functional peptide within the matrix, which causes the nanoparticles to be pulled down by nickel chelating resin. Results are mean ( SD of 3 experiments. (B) Aspecific binding of proteins: Mix- and CALNN-capped nanoparticles incubated with FGF-2 or HGF/SF proteins and purified. Nanoparticles, 5 and 10 µL at 10 nM concentration, were dotted onto a PVDF membrane and proteins bound were detected by dot-blot.

absence of an ionisable group at the carboxy-terminal of the CVVVT-ol peptide. The latter will not experience ionic repulsion between the ligands during the formation of the SAM. In addition, it is expected that the CALNN peptide will bind more water molecules compared to CVVVT-ol and PEGylated alkanethiol molecules, which may increase the hydrodynamic volume of the CALNN peptide. Repulsion forces and increased hydrodynamic volumes of CALNN ligands would reduce the packing density of CALNN ligands compared to Mix ligands in the SAM on the nanoparticle surface. The change of behavior for the Mix-matrix nanoparticles prepared with lower concentration of Mix ligands observed in the electrolyte-aggregation test, size-exclusion and DEAE anionexchange chromatography, is likely the consequence of the lower grafting density of the ligand at the surface of the nanoparticle. Indeed, when using Mix-0.1 mM compared to Mix-2 mM the packing density of peptidol ligand is reduced by more than 50%. This lower grafting density may give greater solvent access to the core of the ligands, which are the valine residues and the methylene groups of the CVVVT-ol and the HS-PEG, respectively. These are hydrophobic and would be expected to lead to aspecific interactions on chromatography resins such as Sephadex G-25 and DEAE-Sepharose, as observed experimentally (Figure 3B and C). For CALNN-0.1 mM capped nanoparticles, the packing density of the ligand is reduced by 28% compared to CALNN-2 mM and it is likely that this decrease in the grafting density for CALNN0.1 mM results in greater accessibility of the side chains of the amino acids to solvent. The side chain of asparagine is polar and the terminus of the peptide is negatively charged, which may mitigate effects of the hydrophobic side chains of alanine and leucine and so prevent aggregation. In addition, it is well established for carboxylic acid-terminated SAMs on flat surfaces that, as the density of the carboxylic acids increases, their pKa increases. The likely decrease in the pKa of the carboxylic acid of the CALNN ligands at lower grafting, as indicated by DEAE chromatography (Figure 3F), will increase the repulsion between the nanoparticles and, therefore, prevent electrolyte-induced aggregation and allow a good recovery from gel-filtration chromatography (Figure 3D and E). Nevertheless, the nanoparticles with a lower grafting density of ligand will present a significant number of defects within the matrix, and even repulsion forces are not able to prevent CALNN-capped nanoparticles from aspecifically binding to DEAE-Sepharose as seen for CALNN0.1 mM.

These results highlight the critical effect of the starting concentration of ligands and of the presence of ionisable groups at the extremity of the ligands on their grafting density and the stability of the nanoparticles with respect to electrolyte-induced aggregation and inertness toward standard chromatography matrices that mimic some types of common noncovalent interactions of biological molecules. Ligand Exchange. As a further test of stability, the exchange of thiolated ligands into Mix-capped nanoparticle was measured. In these experiments, ligand shells were formed with the different concentrations of Mix ligands used previously. Following removal of the excess ligand by centrifugation, the nanoparticles were incubated 4 h with 4 µM of peptides carrying a hexa-histidine affinity tag that is readily captured on Ni2+-NTA. Two peptides were used, CALNN-6xHis-biotin and CVVVT-6xHis-Biotin. In this way, any effect of the matrix sequence on the incoming ligand could be identified. Mix-capped nanoparticles prepared with 2 mM ligand mix were resistant to ligand exchange over 4 h (Figure 4A). Nanoparticles prepared with 1 mM ligand did not incorporate CVVVT-6xHis-biotin peptide, but 7.5% of the nanoparticle population was found to bear at least one CALNN6xHis-biotin peptide. For nanoparticles prepared with 0.5 mM Mix ligand, 52% and 32% of the nanoparticles were found to incorporate the CALNN- and CVVVT-6xHis-biotin peptides, respectively. All of the Mix-capped nanoparticles prepared using 0.1 mM ligand incorporated the functional peptides. In contrast, CALNN-2 mM capped nanoparticles were not resistant to ligand-exchange. Thus, 100% of these nanoparticles incorporated at least one CALNN-6xHis-Biotin, whereas 50% incorporated at least one CVVVT-6xHis-biotin peptide. In addition, all CALNN-capped nanoparticles prepared with 1 mM or less ligand incorporated into their matrix at least one functional peptide (Figure 4A). These results (i) show that Mix-matrix nanoparticles prepared using a concentration of ligands sufficient to ensure a high grafting density are resistant to ligand-exchange in contrast to CALNNmatrix nanoparticles, (ii) highlight again the importance of the concentration of ligand used to prepared the capped nanoparticles, and (iii) show that the sequence of the incoming ligand affects the probability of the exchange. (Supplemental data are provided in the Supporting Information, Figure S5.) Aspecific Binding to Proteins. We then tested the potential aspecific binding of Mix-2 mM and CALNN-2 mM capped nanoparticles to proteins. Two glycosaminoglycan-binding growth factors, FGF-2 and HGF/SF, presenting a basic patch, were tested. Glycosaminoglycan-binding proteins are present in

Ligand Shells for Biological Applications

large amounts in the extracellular matrix of cells and, therefore, will be in the close neighborhood of the nanoparticles when these used for in vivo applications. Following incubation with these proteins and numerous washes by centrifugation, the dotblot results show that none of the Mix-capped nanoparticles bind these proteins (Figure 4B). In the contrast, both FGF-2 and HGF/ SF proteins bind to the CALNN-capped nanoparticles (Figure 4B). It is likely that this interaction is driven by the overall negative charge of the CALNN nanoparticles and the presence of basic patches on the surface of the FGF-2 and HGF/SF proteins.40 Basic patches are common, since they mediate interactions of proteins with nucleic acids as well as with the glycosaminoglycans of the extracellular matrix. In a further set of experiments, we tested the aspecific binding of the Mix-2 mM capped nanoparticles with common affinity chromatography resins and no aspecific binding was observed between Mix-2 mM capped nanoparticles and these resins (streptavidin (SA)-agarose, Strep-Tactin (ST)-Sepharose, StrepTactin (ST)-Macroprep, Probond, Affi-histidine, Anti-flagTagagarose, and heparin-agarose) (Supporting Information Figure S6A). While CALNN-2 mM capped nanoparticles did not bind to some of the resins tested (streptavidin-agarose, Probond), aspecific binding was found with many (Supporting Information Figure S6B). The CALNN stabilized nanoparticles could be partly eluted from Strep-Tactin-Sepharose by increasing the concentration of electrolytes, which suggest that these nanoparticles are in this case adsorbing in part through electrostatic interactions (Supporting Information Figure S6C). Thus, the Mix-capped nanoparticles prepared with 2 mM of ligands did not exhibit detectable ligand-exchange or aspecific binding, which is likely to be due to a combination of the absence of charged groups on the surface of the ligand shell, the high grafting density of ligands, and the mobility of the ethylene glycol component. The presence of negative charges and the lower grafting density of CALNN-2 mM capped nanoparticles will instead favor the association of molecules with the surface of the nanoparticle and an eventual exchange of the surfaceassociated molecules with ligands in the SAM. These phenomena increase as the ligand grafting decreases. Specific Conjugation of Mix-Capped Nanoparticles to Proteins with a Control of Stoichiometry. Given the excellent performances of the Mix-matrix capped nanoparticles in stability tests of increasing stringency, they are good candidates to be used as highly sensitive markers for proteins in vitro and in vivo. We recently described, using CALNN-capped nanoparticles, a generic approach to monofunctionalized gold nanoparticles with a recognition function29 and, as shown in Figure 2, this is equally applicable to Mix-capped nanoparticles. We tested here if it was possible, using this approach, to demonstrate the conjugation, for the first time, of gold nanoparticles to a specific protein at a stoichiometry of 1:1. The functional ligand used was a HSPEG terminated with a Tris-NiNTA function,41,42 which binds to poly histidine-tagged proteins. Titration tests with this functionalized ligand and the CVVVT-ol:HS-PEG Mix ligands were performed to identify the appropriate molar ratio of the HS-PEG-Tris-NTA:(CVVVT-ol:HS-PEG) to be used for the preparation of nanoparticles having only 1 or an average of 3 Tris-NiNTA functions per nanoparticle (Supporing Information Figure S7). In the case of nanoparticles with only 1 recognition function, the functional ligand (HS-PEG-Tris-NTA) is mixed (40) Jeong, K. J.; Butterfield, K.; Panitch, A. Langmuir 2008, 24, 8794. (41) Lata, S.; Reichel, A.; Brock, R.; Tampe, R.; Piehler, J. J. Am. Chem. Soc. 2005, 127, 10205. (42) Tinazli, A.; Tang, J.; Valiokas, R.; Picuric, S.; Lata, S.; Piehler, J.; Liedberg, B.; Tampe, R. Chemistry 2005, 11, 5249.

Langmuir, Vol. 24, No. 23, 2008 13579

Figure 5. Specific and stoichiometric conjugation of Mix-capped nanoparticles to FGFR1 protein.

with the Mix-matrix ligand such that only 10% or less of the capped nanoparticles incorporate this function. Stochastic incorporation of the functionalized ligand into the ligand shell ensures that under these conditions there are an undetectable number of nanoparticles with 2 or more functional ligands incorporated into the SAM on the nanoparticle. The Tris-NTA tagged nanoparticles are then incubated with nickel to activate the tag (Tris-NTA becoming Tris-NiNTA) and purified using the properties of the functional ligand itself so the 90% of the nanoparticle population without a function is removed. Therefore, only nanoparticles having 1 Tris-NiNTA tag (n ) 1) per nanoparticle are used. For nanoparticles n ∼ 3, the functional ligand (HS-PEG-Tris-NTA) is mixed with the Mix-matrix ligand so 90 to 100% of the capped nanoparticles incorporate this function. Affinity chromatography is performed as for n ) 1 to removed the few nanoparticles (