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UNIVERSIDAD COMPLUTENSE DE MADRID

FACULTAD DE CIENCIAS QUÍMICAS

DEPARTAMENTO DE QUÍMICA ANALÍTICA

Nuevos nanomateriales para el

diseño de Biosensores Electroquímicos y

Sistemas de liberación controlada

Directores:

Dr. Reynaldo Villalonga Santana

Investigador Ramón y Cajal

Dr. José Manuel Pingarrón Carrazón

Catedrático de Universidad

TESIS DOCTORAL PRESENTADA POR:

PAULA DÍEZ SÁNCHEZ

Madrid, 2016

D. Reynaldo Villalonga Santana, Investigador Ramón y Cajal del Departamento de

Química Analítica de la Facultad de Ciencias Químicas de la Universidad Complutense de

Madrid.

D. Jose Manuel Pingarrón Carrazón, Catedrático de Universidad del Departamento de


Madrid.

HACEN CONSTAR,

Que el trabajo titulado “Nuevos nanomateriales para el diseño de biosensores

electroquímicos y sistemas de liberación controlada” ha sido realizado bajo su dirección

en el Grupo de Electroanálisis y (bio)sensores electroquímicos (GEBE) del Departamento


Madrid, constituyendo la Tesis Doctoral de su autora.

Madrid, 31 de Octubre 2016

Fdo. Reynaldo Villalonga Santana Fdo. José Manuel Pingarrón Carrazón

Fdo. Paula Díez Sánchez

ÍNDICE

ÍNDICE

i

1. Índice de figuras y Tablas…………………………………..…………………………………. 1

2. Abreviaturas y Símbolos……………………………………………….………………………. 15

3. Summary……………………………………………………..……………………………………………. 17

4. Resumen……………………………………………………….…………………………………………. 23

5. Introducción……………………………………………………..…………………………….……….. 31

5.1. Nanomateriales………………………..………………………………………………………… 31

5.1.1. Conceptos fundamentales………………………………………………………….…… 31

5.1.2. Propiedades de los nanomateriales………………………………………………... 33

5.1.3. Clasificación de los nanomateriales……………………….………………...……... 35

5.1.3.1. Clasificación de acuerdo a su composición……………………………….. 35

5.1.3.2. Clasificación de acuerdo a sus dimensiones espaciales…………….. 35

5.1.3.3. Clasificación de acuerdo a su forma de obtención………………..…. 36

5.1.4. Aplicaciones de los nanomateriales………………………………………….….…. 36

5.1.5. Limitaciones de los nanomateriales……………………………………………….…. 40

5.2. Biosensores electroquímicos…………………………………………..……….………. 42

5.2.1. Conceptos fundamentales……………………………………………………………...… 42

5.2.2. Clasificación de los biosensores…………………………………………………….… 44

5.2.2.1. Clasificación de acuerdo con el transductor……………………………… 44

5.2.2.2. Clasificación de acuerdo con el elemento de reconocimiento biológico………………………………………………………………………………..………………

45

5.2.2.3. Biosensores de carácter combinado…………………………………………. 48

5.2.3. Nanomateriales utilizados en biosensores…………………………….………….. 50

5.2.3.1. Nanopartículas metálicas…………………………………………………….. 52

5.2.3.2. Nanomateriales de carbono………………………………………………… 56

5.2.3.3. Nanomateriales polímericos……………………………………………….. 59

5.2.3.4. Nanomateriales de óxidos metálicos…………………………………… 61

5.3. Sistemas de liberación controlada…………………………………………… 63

5.3.1 Conceptos básicos……………………………………………………………………………… 63

ÍNDICE

ii

5.3.2. Clasificación de los sistemas de liberación controlada de fármacos…… 65

5.3.2.1. Clasificación de acuerdo a la orientación………………………………… 65

5.3.2.3. Clasificación de acuerdo al mecanismo de liberación del fármaco………………….……………………………………………………………………………..

66

5.3.2.1. Clasificación de acuerdo con el tipo de plataforma utilizada……. 67

5.3.3. Sistemas de liberación controlada basados en MSN………………………… 74

5.3.1. Puertas moleculares estímulo-respuesta…………………………………. 76

6. Objectives…………………………………………………………………………………………………… 81

7. Publicaciones científicas………………………………………………………….…………… 83

7.1. Electrochimica Acta 56 (2011) 4672-4677……….…………………………………… 83

7.2. Analyst 137 (2012) 342-348…………………….…………………………….……………. 107

7.3. ChemElectroChem. 1 (2014) 200-206…………………………………………………… 127

7.4. ACS Applied Materials & Interfaces 4 (2012) 4312-4319………………………. 149

7.5. Electroanalysis 23 (2011) 1790-1796……………………………………………………. 177

7.6. Journal of Materials Chemistry 21 (2011) 12858-12864………………………. 195

7.7. Analytical and Bioanalytical Chemistry 405 (2013) 3773-3781……………… 217

7.8. Electrochimica Acta 76 (2012) 249-255……………………………………………….. 243

7.9. ACS Applied Materials & Interfaces 8 (12) (2016) 7657–7665………………. 265

7.10. Electrochemistry Communications 30 (2013) 51–54………………………….. 309

7.11. Chemistry - A European Journal 19(24) (2013) 7889–7894………………… 321

7.12. Journal of the American Chemical Society 136 (25) (2014) 9116–9123. 343

8. Discusión integradora………………………………………………………………………….… 375

8.1. Nanomateriales funcionalizados y nanohíbridos para el ensanblaje de biosensores electroquímicos…………………………………………………………….… 375

8.1.1. Biosensores basados en redes de nanopartículas de oro polifuncionalizada………………………………………………………………………. 375

8.1.2. Biosensores nanoestructurados con nanotubos de carbono modificados mediante interacciones no covalentes……………………. 389

8.1.3. Biosensores basados en dendrímeros y dendrones de poliamidoamina modificados con ciclodextrina…………………………..

394

ÍNDICE

iii

8.2. Sistemas de liberación inteligente de fármacos controlados por enzimas y basados en nanomateriales de sílice mesoporosa…………….. 400

8.2.1. Nanomáquinas basadas en nanopartículas de sílice mesoporosa funcionalizadas con neoglicoenzimas…………………………………………. 400

8.2.2. Nanomáquinas controladas por enzimas y basadas en nanopartículas Janus de oro y sílice mesoporosa………………………… 403

9. Conclusions……………………………………………………………………………………………… 409

10. Referencias……………………………………………………………………………………………. 411

1 ÍNDICE DE FIGURAS

Y TABLAS

1. ÍNDICE DE FIGURAS Y TABLAS

1

5. INTRODUCCIÓN:

Figuras:

Figura 5.1. Comparaciones de tamaños en la escala nanométrica……………………… 31

Figura 5.2. Número de publicaciones científicas sobre nanomateriales desde el año 2000 al 2015. (Fuente consultada: Web of Science)…………………….………..……. 33

Figura 5.3. Aplicaciones de los nanomateriales en diferentes áreas…………………… 36

Figura 5.4. Esquema de funcionamiento de un biosensor………………….......……..…. 43

Figura 5.5. Representación esquemática de una reacción enzimática catalizada……………………………………………………………………………………………..……......... 46

Figura 5.6. Esquema básico de biosensor electroquímico nanoestructurado……. 50

Figura 5.7. Diferentes arquitecturas de nanopartículas metálicas en la modificación de las superficies electródicas…………………………………………..…..……… 53

Figura 5.8. Modificación de la superficie del electrodo con nanopartículas metálicas, solas y combinados con otros materiales………………………..………………… 54

Figura 5.9. Transferencia directa de electrones (e-) entre en centro redox de una enzima y la superficie del electrodo a través de CNTs…………………..……………. 57

Figura 5.10. Parámetros estructurales de una molécula de dendrímero y sus métodos de síntesis A) divergentes y B) convergentes………….………………..………… 60

Figura 5.11. Resumen de los mecanismos básicos de liberación de fármacos en los sistemas de administración controlada por: A) difusión, B) hinchamiento controlada, C) erosión y D) estímulos……………………………..…………………………………. 67

Figura 5.12. Ilustración de los nanomateriales usados como plataformas en los sistemas de liberación de fármacos……………………………………………………………………. 68

Figura 5.13. Diagrama esquemático de los materiales MCM-41 (hexagonal) y MCM-48 (cúbica)……………………………………………………………….……………….……………… 74

Figura 5.14. Ilustración esquemática de la síntesis de MCM-41. Mecanismo de polimerización del TEOS…………………………………………………………………………………….. 75

Figura 5.15. Representación esquemática de una MSN funcionalizada con una puerta molecular para la liberación controlada de fármacos mediante un estímulo…………………………………………………………………………………………………………….. 76

Tablas:

Tabla 5.1. Evolución de los sistemas controlados de administración de fármacos desde1950……………………………………………………………………….………….…………………….. 64


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7. PUBLICACIONES CIENTÍFICAS:

7. 1. Electrochimica Acta 56 (2011) 4672-4677

Figuras:

Figure 1. HRTEM image of AuNPs……………..………………………………………………….. 88

Figure 2. UV-vis spectra of 0.2 mg/mL solution of functionalized AuNPs in 50 mM sodium phosphate buffer, pH 7.0 in the absence (A) and the presence of 0.5 mg/mL BSA (B) and HRP (C)…………………………………………………………………..… 89

Figure 3. Field emission SEM image of the polyAuNP-modified electrode……. 90

Figure 4. AFM images of the polyAuNP-modified electrode…………………………. 91

Figure 5. Cyclic voltammograms recorded at a bare gold disk electrode (A), and a polyAuNPs-modified electrode before (B) and after HRP immobilization (C), in 0.1 M KCl solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1)…………………………………………………………………………

92

Figure 6. Impedance plane diagram (−Z″ versus Z′) for the EIS measurements at a bare gold disk electrode () and a polyAuNPs-modified electrode before () and after HRP immobilization (), in 0.1 M KCl solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1)……………….………………………………………………..……… 93

Figure 7. Cyclic voltammograms recorded at HRP-modified electrodes in the absence and the presence of H2O2 in 0.1 M sodium phosphate buffer, pH 7.0. HRP-Au electrode (A), HRP-polyAuNPs-Au electrode (B), HRP-Au electrode + 1 µM H2O2 (C), HRP-Au electrode + 2 µM H2O2 (D), HRP-polyAuNPs-Au electrode + 1 µM H2O2 (E) and HRP-polyAuNPs-Au electrode + 2 µM H2O2 (F)…………………………………………………………………………………………………………………

95

Figure 8. Amperometric responses recorded with HRP-Au (A) and HRP-polyAuNPs-Au (B) electrodes for successive additions of 10 mM H2O2 to 10 mL of 0.1 M sodium phosphate buffer, pH 7.0. Eapp. = 0.0 V………………………. 96

Figure 9. Calibration curve for H2O2 obtained with a HRP-polyAuNPs-Au biosensor in the 5 µM to 1.1 mM H2O2 concentration range…………………….………………………………………………………………………………………. 97

Figure 10. Effect of the storage time at 4°C of the HRP-polyAuNPs-Au biosensor on the sensitivity for H2O2 determination…………………………………..… 110

Tablas:

Table 1. Comparison of analytical properties of the biosensor with previously reported AuNP-based mediatorless biosensors…………………………… 98


3

7. 2. Analyst 137 (2012) 342-348

Figuras:

Fig. 1. Scheme displaying the steps involved in the construction of an electrochemical tyrosinase biosensor based on a gold electrode nanostructured with electropolymerized PAMAM G-4 dendron-coated AuNPs……………………………………………………………………………………………………………. 110

Fig. 2. HRTEM image of dendron-functionalized AuNPs………………................... 111

Fig. 3. Cyclic voltammograms for the first electropolymerization process of PAMAM G-4 dendron-coated AuNPs on gold electrode surface, in 0.1 M H2SO4. Scan rate: 100 mV/s……………….…………………………………..……………………… 112

Fig. 4. AFM images and height histogram of the PAMAM G-4 dendron-coated AuNPs-modified electrode………….……………………….……………………………. 113

Fig. 5. Impedance plane diagram (−Z´´ versus Z´) for the EIS measurements at a bare gold disk electrode () and at the electropolymerized PAMAM G-4 dendron-coated AuNPs-modified electrode before () and after tyrosinase immobilization (), in 0.1 M KCl solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1)………………………………………………………………………… 115

Fig. 6. Cyclic voltammograms recorded at a bare gold disk electrode (A), and at the electropolymerized PAMAM G-4 dendron-coated AuNPs-modified electrode before (B) and after tyrosinase immobilization (C), in 0.1 M KCl solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1)……………………………….. 116

Fig. 7. Amperometric response of the tyrosinase-functionalized p-aminothiophenol-coated non-nanostructured gold electrode (A) and electropolymerized PAMAM G-4 dendron-coated AuNPs-modified electrode (B) to successive additions of 100 µM catechol solution to 10 mL of 0.1 M sodium phosphate buffer, pH 7.0. Eapp. = -100 mV………………………………………. 117

Fig. 8. Effect of the storage time at 4°C of the tyrosinase biosensor on the relative bioelectrocatalytic activity towards 100 nM catechol determination……………………………………………………………………………………………… 119

7. 3. ChemElectroChem. 1 (2014) 200-206

Figuras:

Scheme 1. Preparation of the immunosensor for fibrinogen on gold electrodes coated with electropolymerized matrix of biotin-labelled AuNPs……………………………………………………………………………………………………………. 131

Figure 1. Cyclic voltammograms recorded in the electropolymerization of the polyfunctionalized AuNPs on gold electrode surface in 0.1 M H2SO4 at 100 V/s…………………………………………………………………………………………………………..

134


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Figure 2. FE-SEM (A) and AFM (B) analysis of gold surface coated with electropolymerized matrix of biotin-labelled AuNPs…………………………………….. 135

Figure 3. Nyquist plots obtained at a bare gold electrode (a) and at a gold electrode sequentially modified with p-aminothiophenol (b), polymerized AuNPs (c), streptavidin (d), biotinilated fibrinogen (e), casein (f) and HRP-labeled anti-fibrinogen antibody (g) in 0.1 M KCl solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1)………..……………..……………………………………………….. 136

Figure 4. Cyclic voltammograms recorded in 0.1 M KCl solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1) at a bare gold electrode (a) and at a gold electrode sequentially modified with p-aminothiophenol (b), polymerized AuNPs (c), streptavidin (d), biotinilated fibrinogen (e), casein (f) and HRP-labeled anti-fibrinogen antibody (g). Scan rate: 50 mV/s………………………………. 138

Figure 5. Calibration curve for fibrinogen quantification using the electropolymerized AuNPs-based immunosensor………………………………………… 140

Figure 1S. A) HR-TEM image, B) size distribution and C) selected area electron diffraction analysis of biotin-labeled gold nanoparticles……………...... 146

Figure 2S. UV-vis spectra of 0.2 mg/mL solution of functionalized AuNPs in 50 mM sodium phosphate buffer, pH 7.0 in the absence (a) and the presence of 0.5 mg/mL BSA (b) and streptavidin (c)…………………………………………………….. 147

Figure 3S. Height histogram of the gold surface coated with electropolymerized matrix of biotin-labeled AuNPs……………………………………... 147

7. 4. ACS Applied Materials & Interfaces 4 (2012) 4312-4319

Figuras:

Figure 1. Schematic display of the steps involved in the preparation of XO-CD/pAuNP/SWNT/GCE enzyme biosensors………………..................................... 155

Figure 2. FT-IR spectrum of the polyfunctionalized Au nanoparticles…………… 156

Figure 3. UV-vis spectra of 0.4 mg/mL solution of Au nanoparticles in 50 mM sodium phosphate buffer, pH 7.0 without (A) and with 0.5 mg/mL XO (B) and XO-CD (C)………………………………………..……………………………………………………………. 157

Figure 4. Cyclic voltammograms recorded in 0.1 M H2SO4 at a bare GCE (A), a SWNT-modified GCE (B), and upon electropolymerization of Au nanoparticles for five (C) and ten (D) potential cycles, and after polymer growing for 1 h at +850 mV (E). Scan rate: 50 mV/s………………………………………. 158

Figure 5. Field emission SEM image of the pAuNP/SWNT/GCE……………………… 159

Figure 6. Nyquist plots of bare GCE (), SWNT/GCE (), pAuNP/SWNT/GCE (X) and XO-CD/pAuNP/SWNT/GCE () in 0.1 M KCl solution containing 5 mM K3[Fe(CN)6] /K4[Fe(CN)6] (1:1)……………………………………………………………………….. 160


5

Figure 7. Cyclic voltammograms recorded in 0.1 M KCl solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1) at bare GCE (A), SWNT/GCE (B), pAuNP/SWNT/GCE (C) and XO-CD/pAuNP/SWNT: 50 mV/s…….…………………… 161

Figure 8. Cyclic voltammograms recorded in 0.1 M sodium phosphate buffer, pH 7.0, at a scan rate of 50 mV/s for: A) XO-CD/pAuNP/SWNT/GCE and XO/pAuNP/SWNT/GCE without addition of xanthine (curves (a) and (b), respectively) and after addition of 200 µM xanthine (curves (d) and (c), respectively). B) XO-CD/pAuNP/SWNT/GCE and XO/pAuNP/SWNT/GCE in the presence of 200 µM xanthine before (curves (e) and (c), respectively) and after 1 h incubation in saturated 1-adamantane carboxylic acid solution (curves (g) and (f), respectively)……………………………………………………………………. 162

Figure 9. Amperometric responses of the XO-CD/pAuNP/SWNT/GCE and Naf/XO-CD/pAuNP/SWNT/GCE biosensors toward 500 nM xanthine upon addition of glucose (a), sacarose (b), ethanol (c), acetic acid (d), lactic acid (e), citric acid (f), uric acid (g) and ascorbic acid (h) at a 5.0 µM concentration level……………………………………………………………………………………………………………….

168

Figure S1. TEM image of the polyfunctionalized Au nanoparticles acquired with a JEOL JEM-2100 microscope at 200 kV…………………………………………………. 174

Figure S2. HR-TEM image of the polyfunctionalized Au nanoparticles acquired with a JEOL JEM-3000 F microscope at 300 kV……………………………..... 175

Figure S3. Dynamic amperometric response of XO-CD/pAuNP/SWNT/GCE poised at +650 mV to successive addition of 100 µM xanthine solution. Inset: Amperometric response of the electrode at lower concentrations of xanthine. Initial working volume: 10 ml. Supporting electrolyte: 0.1 M sodium phosphate buffer, pH 7.0…………………………………….……………………………. 175

Figure S4. Calibration curves for xanthine obtained with XO-CD/pAuNP/SWNT/GCE (), Naf/XO-CD/pAuNP/SWNT/GCE () and

XO/pAuNP/SWNT/GCE () biosensors………………………………………………………….. 176

Figure S5. Evaluation of the storage stability with time for XO-CD/pAuNP/SWNT/GCE () and Naf/XO-CD/pAuNP/SWNT/GCE () biosensors. Measurements were carried out toward 500 nM xanthine……………………………………………............................................................... 176

Tablas:

Table 1. Comparison of the analytical characteristics of the developed biosensors with those previously reported for other electrochemical xanthine biosensors……….…………………………………………………………………………….. 167

7. 5. Electroanalysis 23 (2011) 1790-1796

Figuras:

Figure 1. Scheme displaying the steps and fundamentals involved in the enzyme biosensor preparation……………………………………………………………………… 183


6

Figure 2. Quartz crystal microbalance responses for gold disks coated with: SWNT during immobilization of native XO (a), SWNT/Pyr-βCD during the immobilization of adamantane-modified (b), and native XO (c)………….………… 184

Figure 3. Nyquist plots recorded at bare () and SWNT/Pyr-βCD-modified

GCE before () and after immobilization of native () and adamantane-modified XO (), in 0.1 M KCl solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1)…………………………………………………………………………. 185

Figure 4. Cyclic voltammograms of XO/SWNT/Pyr-βCD and XO-ADA/SWNT/Pyr-βCD modified GCE in the absence (voltammograms A and B, respectively) and in the presence of 40 µM xanthine (voltammograms C and D, respectively). Scan rate, 50 mV/s in 0.1 M sodium phosphate buffer, pH 7.0…………………………………………………………………………………………………………………. 187

Figure 5. Dynamic amperometric response of the XO-ADA/SWNT/Pyr-βCD (A) and XO/SWNT/Pyr-βCD (B) modified GCE at +600 mV to successive additions of a 10 mM xanthine solution. Initial working volume: 10 ml. Supporting electrolyte: 0.1 M sodium phosphate buffer, pH 7.0……………………………………………………………………...............................................

188

Figure 6. Long-term stability of the XO-ADA/SWNT/Pyr-βCD/GCE biosensor toward the determination of xanthine………………………………………………………….. 190

7. 6. Journal of Materials Chemistry 21 (2011) 12858-12864

Figuras:

Figure 1. Schematic display of the construction of Fe3O4/APTES-PEG-XO/SWNT magnetic nanomaterials-based biosensors………………………………….. 201

Figure 2. TEM images of A) Fe3O4, B) Fe3O4/APTES, C) Fe3O4/APTES-PEG and D) Fe3O4/APTES-PEG/SWNT magnetic nanomaterials……………………………………. 202

Figure 3. FT-IR spectra of A) Fe3O4, B) Fe3O4/APTES and C) Fe3O4/APTES-PEG magnetic nanoparticles…………….…………………………………………………………………… 204

Figure 4. Magnetization curves of Fe3O4 (), Fe3O4/APTES (), Fe3O4/APTES-PEG (X) and Fe3O4/APTES-PEG/SWNT () nanomateriales at 298K………………… 205

Figure 5. FE-SEM images of A) Fe3O4/APTES-PEG-XO and B) 1:27 SWNT: Fe3O4/APTES-PEG-XO-modified AuSPE………………………………………………………….. 207

Figure 6. Impedance plane diagram (−Z″ versus Z′) recorded for AuSPE before () and after modification with Fe3O4/APTES-PEG-XO (), 1:80 SWNT: Fe3O4/APTES-PEG-XO (), 1:27 SWNT: Fe3O4/APTES-PEG-XO () and 1:16 SWNT: Fe3O4/APTES-PEG-XO (Δ) in 0.1 M KCl solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1)…………………………………………………………………………

208

Figure 7. Cyclic voltammograms recorded with AuSPEs modified with: Fe3O4/APTES-PEG/XO (A); Fe3O4/APTES-PEG-XO (B); 1:80 SWNT: Fe3O4/APTES-PEG-XO (C); 1:16 SWNT: Fe3O4/APTES-PEG-XO (D) and 1:27


7

SWNT: Fe3O4/APTES-PEG-XO (E) from a 2.5 µM xanthine solution in 0.1 M sodium phosphate buffer, pH 7.0; v = 50 mV/s……………………………….……………..

209

Figure 8. Calibration plots for xanthine recorded with Fe3O4/APTES-PEG/XO (), Fe3O4/APTES-PEG-XO (), 1:80 SWNT: Fe3O4/APTES-PEG-XO (), 1:27 SWNT: Fe3O4/APTES-PEG-XO () and 1:16 SWNT: Fe3O4/APTES-PEG-XO (Δ) modified AuSPEs; Eapp. = + 600 mV………………………………………………………………. 211

Tablas:

Table 1. Comparison of analytical properties of the biosensor with previously reported xanthine biosensor……………………………………………………….. 212

7. 7. Analytical and Bioanalytical Chemistry 405 (2013) 3773-3781

Figuras:

Figure 1. Schematic display of the steps involved in the preparation of GO-ADA/CD-PAMAM/PtNP/Au based enzyme biosensors…………………………………. 223

Figure 2. SPR sensogram recorded on Au surfaces upon successive incubation with CD-PAMAM G-4 and GO-ADA (a), PAMAM G-4 and GO (b) and CD-PAMAM G-4 and GO (c)……………………………………………………………………. 224

Figure 3. Field emission SEM image of the CD-PAMAM G-4/PtNP-modified Au surface……………………………………………………………………………………………………… 226

Figure 4. Three dimensional AFM analysis of the Au surface before (A) and after modification with CD-PAMAM (B), CD-PAMAM/PtNP (C) and GO-ADA/CD-PAMAM/PtNP (D)……………………………………………………………………………. 227

Figure 5. Nyquist plots of bare Au (a), CD-PAMAM/Au (b), CD-PAMAM/PtNPs/Au (c) and GO-ADA/CD-PAMAM/PtNP/Au (d) electrodes in 0.1 M KCl solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1)……………….. 228

Figure 6. Cyclic voltammograms recorded in 0.1 M KCl solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1) at bare Au (a), CD-PAMAM/Au (b), CD-PAMAM/PtNPs/Au (c) and GO-ADA/CD-PAMAM/PtNP/Au (d) electrodes. Scan rate: 50 mV/s……………………………………………………………………………………..... 229

Figure 7. Cyclic voltammograms recorded in 0.1 M sodium phosphate buffer, pH 7.0, at a scan rate of 50 mV/s, for GO-ADA/CD-PAMAM/PtNP/Au electrode before (a) and after (b) addition of 200 µM glucose…………………………………………………………………………………………………………..

230

Figure 8. Amperometric responses recorded with GO-ADA/CD-PAMAM/PtNP/Au (a), GO/PAMAM/PtNP/Au (b), GO/CD-PAMAM/PtNP/Au (c) and GO/PtNP/Au (d) electrodes upon successive additions of 5.0 mM glucose. Eapp.= + 400 mV……………………………………………………………………………… 232

Figure 1S. 1H NMR spectrum of CD-branched cysteamine core PAMAM G-4 dendron………………………………………………………………………………………..………………. 241


8

Tablas:

Table 1. Comparison of the analytical performance of the GO-ADA/CD-PAMAM/PtNP/Au biosensor with that reported previously for mediatorless glucose biosensors………………………………………………………………………………………… 234

7. 8. Electrochimica Acta 76 (2012) 249-255

Figuras:

Figure 1. Scheme displaying the steps involved in the preparation of the layer-by-layer self-assembly of dendrimer and HRP on Au surface. D1: CD-PAMAM G-4 dendron layer, D2 and D3: CD-PAMAM G-5 dendrimer layers, and HRP: enzyme layers…………………………………………………………………..……………. 250

Figure 2. A) Cyclic voltammograms of native (a) and CD-PAMAM G-4 dendron-modified Au electrode in 0.1 M sodium phosphate buffer, pH 7.0 solutions containing 1.0 mM hydroquinone before (b) and after (c) addition of H2O2 up to 100 µM final concentration. B) Cyclic voltammograms of Au electrodes modified with CD-PAMAM G-4 dendron (c), D1/HRP1 (d), D2/HRP2 (e), D3/HRP3 (f) and D4/HRP4 (g) bilayers in the presence of 100 µM H2O2…………………………………………………………………………………………………….… 251

Figure 3. Cyclic voltammograms recorded with Au electrodes modified with CD-PAMAM G-4 dendron/HRP (E) and D3/HRP3 bilayers in 0.1 M sodium phosphate buffer, pH 7.0 solutions containing 1.0 mM hydroquinone and 100 µM H2O2 before (A) and after 30 (B), 60 (C) and 120 (D) min incubation in 10 mM 1-adamantane carboxylic acid solution…………………………………………. 253

Figure 4. SPR sensogram recorded upon the layer-by-layer self-assembling of CD-modified PAMAM dendritic scaffolds and HRP-ADA on Au surfaces……………………………………………………………………………………………………….… 254

Figure 5. Calculated molar content of enzyme and dendrimers in the layer-by-layer assembly from QCM (black) and SPR (white) measurements. Each value corresponds to the molar content per layer……………………………………….. 255

Figure 6. Three dimensional AFM analysis of the Au surface before (A) and after modification with the D1/HRP1 (B), D2/HRP2 (C) and D3/HRP3 (D) bilayers………………………………………………………………………………………………………… 256

Figure 7. Amperometric responses recorded with CD-PAMAM G-4 dendron/HRP (A), D1/HRP1 (B), D2/HRP2 (C) and D3/HRP3 (D) modified Au electrodes upon successive additions of 1.0 mM H2O2. Eapp.= - 100 mV. Inset: Electroanalytical behavior of the D3/HRP3 modified Au electrode toward low H2O2 concentration…………………………………………………………………….. 258

Figure 8. Calibration curves for H2O2 constructed with biosensors prepared with D1/HRP1 (), D2/HRP2 (), D3/HRP3 () and D1/native HRP (×) modified Au electrodes………………………………………………………………………………….

259


9

7. 9. ACS Applied Materials & Interfaces 8 (12) (2016) 7657–7665

Figuras:

Scheme 1. Performance of dual stimuli-responsive nanodevice S3 for the programmed and sequential delivery of the [Ru(bpy)3]Cl2 complex using glucose and ethyl butyrate as triggers…………………………………………………………… 267

Figure 1. A) Release efficiency for [Ru(bpy)3]Cl2 from S3 (a) and S2 (b) after a 90-minute incubation with different trigger substances at a 200 µM concentration. B) Dye release efficiency from S3 (a) and S2 (b) after 90-minute incubation in the presence of glucose at different concentrations……………………………………………………………………………….……………… 271

Figure 2. Kinetics of the dye release from S3 (A) and S2 (B) in 20 mM Na2SO4, pH 7.5, without (a) and with addition of ethyl butyrate (b) and glucose + ethyl butyrate (c) at a 200 µM final concentration. Triggers were added at the times indicated in the graphics……………………………………………………………….. 272

Scheme 2. Proposed mechanisms for the stimuli-responsive controlled delivery from nanocarriers S2 (A) and S3 (B)………………………………………………… 274

Figure 3. Internalization and release of cargo in HeLa cells. Culture were incubated with S4 (3A) or S5 (3B) in presence/absence of different input I1 (D-glucose) or input I2 (ethyl butyrate) and examined for Doxorubicin staining (Doxo) by confocal microscopy. Representative images at 24 h form phase contrast (PhC), Doxorubicin (Doxo), Hoescht (Hoe) and combined (Merged) are shown. Quantification of cell viability and cell death was performed by flow cytometry by means of 7-AAD and Ann V staining. The percentage of dead cells (black), cells undergoing cell death (gray) and healthy cells (white) are shown for 50 μg/ml concentration of S4 (3C) and S5 (3D) in HeLa cells under different conditions at 24 h……………………………………. 276

Figure S1. TEM images of S0 (a), S1 (b), S2 (c) and S3 (d) nanoparticles………. 292

Figure S2. Powder X-ray diffraction of S0 (a), S1 (b), S2 (c) and S3 (d) nanoparticles at low angles………………………………………………………………………….. 294

Figure S3. FT-IR analysis for S0 (a), S1 (b), S2 (c) and S3 (d) nanoparticles…… 294

Figure S4. Solid state 13C NMR spectra for S0 loaded with tris(2,2′-bipyridyl)dichlororuthenium (II) hexahydrate and modified with epoxy groups (A), S1 (B) and S2 (C) nanoparticles…………………………………………………… 295

Figure S5. A) TG and B) DTG analysis for S0 (a), S1 (b), S2 (c) and S3 (d) nanoparticles……………………………………….……………………………………………………….. 296

Figure S6. A) Nitrogen adsorption (closed)/desorption (open) isotherms and B) pore size distribution for S0 (a), S1 (b), S2 (c) and S3 (d) nanoparticles………………………………………………………………………………………………… 297


10

Figure S7. Nanoparticle size distribution of S0, S2 and S3 in H2O (black line), PBS (dotted line) and DMEM (striped line)……………………………………………………. 298

Figure S8. TEM images of S1 (A), S2 (B) and S3 (C) nanoparticles after staining with 1% uranyl acetate…………………………………………………………………….. 299

Figure S9. Trigger reactions mediated by D-glucose (A) and ethyl butyrate (B)…………………………………………………………………………………………………………………. 300

Figure S10. A) Confocal images of HeLa cells incubated with solid S4 for 30 minutes and then treated with input A (D-Glucose) and/or input B (ethyl butyrate) or untreated. Representative images at 24 h from phase contrast (PhC), Doxorubicin (Doxo), Hoescht (Hoe) and combined (Merged) are shown. B) Cell viability (%) of HeLa cells treated with S5 using WST-1 assay. Results expressed in % (refereed to untreated cells) ± standard deviation of three different experiments. C) Mean fluorescent intensity of Doxo in HeLa cells treated with S4 using FACS analysis…………………………………………………....... 301

Figure S11. A) Confocal images of HeLa cells incubated with solid S5 for 30 minutes and then treated with input A (D-Glucose) and/or input B (ethyl butyrate) or untreated. Representative images at 24 h from phase contrast (PhC), Doxorubicin (Doxo), Hoescht (Hoe) and combined (Merged) are shown. B) Cell viability (%) of HeLa cells treated with S5 using WST-1 assay. Results expressed in % (refereed to untreated cells) ± standard deviation of three different experiments. C) Mean fluorescent intensity of Doxo in HeLa cells treated with S5 using FACS analysis………………………………………………………. 302

Figure S12. Quantification of cell viability and cell death was performed by flow cytometry by means of 7-AAD and Ann V- FITC staining for S4. The percentage of dead cells (black), cells undergoing cell death (gray) and healthy cells (white) are shown after 24 h of treatment. Three independent experiments containing triplicates were performed and the data are reported as (mean ± SE)………………………………..……………………………………………... 301

Figure S13. Quantification of cell viability and cell death was performed by flow cytometry by means of 7-AAD and Ann V- FITC staining for S5. The percentage of dead cells (black), cells undergoing cell death (gray) and healthy cells (white) are shown after 24 h of treatment. Three independent experiments containing triplicates were performed and the data are reported as (mean ± SE).………………………………………………………………………………..

304

Figure S14. CLSM images of HeLa cells incubated with 30 μg/mL of S4 for 1 hour, washed and then incubated during 1.5h and 12 h in PBS with 10 % FBS (-/-) or PBS with 10 % FBS and ethylbutyrate (-/+) . Lysotracker was used to stain the organelles of the cell and Hoescht for the nucleus…………………………. 305

Figure S15. CLSM images of HeLa cells incubated with 30 μg/mL of S5 for 1 hour and post incubated during 1.5h and 12 h in PBS and 10 % FBS (-/-) or PBS with 10 % FBS and ethylbutyrate (-/+) . Lysotracker was used to stain the organelles of the cell and Hoescht for the nucleus……………………………………….. 306


11

Figure S16. CLSM images of HeLa cells incubated with 30 μg/mL of S4 or S5 during 1 hour, washed and then incubated in PBS with fetal bovine serum (No input) or in PBS with ethyl butyrate (Input I2). The green fluorescence is from the endocytosed nanoparticles and from released Doxo……………………… 307

Tablas:

Table S1. BET specific surface values, pore volumes and pore sizes calculated from the N2 adsorption-desorption isotherms for selected materials…………. 298

Table S2. Nanoparticle size (nm) and zeta potential values (mV) of S0, S2 and S3 in pure H2O, PBS buffer solution with 10 % FBS and DMEM culture medium with 10 % FBS………………………………………………….….………………………….. 299

7. 10. Electrochemistry Communications 30 (2013) 51–54

Figuras:

Figure 1. Preparation of the Janus nanoparticle-based biorecognition-signaling system……………………………………………………………………………………………. 313

Figure 2. A) SPR sensograms recorded upon modification of Au surfaces with thiolated biotin (1), Au-MS JNP-HRP-Stv-PEG (2) and Au-MS JNP-HRP-PEG (3). B) AFM analysis of Au-MS JNP-HRP-Stv-PEG adsorbed on biotin-modified Au surface………………………………………………..……………………………………………….….. 315

Figure 3. A) Nyquist plots and B) cyclic voltammograms obtained at a bare Au electrode (a) and after modification with thiolated biotin (b), Au-MS JNP-HRP-Stv-PEG (c) and Au-MS JNP-HRP-PEG (d) in 0.1 M KCl, 5 mM K3[Fe(CN)6]/K4[Fe(CN)6]. C) Cyclic voltammograms recorded with Au/biotin/Au-MS JNP-HRP-Stv-PEG (a) and Au/biotin/Au-MS JNP-HRP-PEG (b) electrodes in the absence and presence (*) of 50 µM H2O2 in 0.1 M sodium phosphate buffer, pH 7.0………………………………………………………………….

316

7. 11. Chemistry - A European Journal 19(24) (2013) 7889–7894

Figuras:

Scheme 1. Preparation of Janus Au-MS nanoparticles for enzyme-controlled release………………………………………………………………………………………………………… 324

Figure 1. TEM images of J1 (A), J2 (B), J3 (C) and J4 (D) Janus Au-MS nanoparticles………………………………………………………………………………………………… 326

Figure 2. Representative TEM image of J2 nanoparticles…………..………………….. 327

Figure 3. A) Nitrogen adsorption (closed)/desorption (open) isotherms for MS (,) and J2 nanoparticles (,). Inset: pore size distribution of MS () and J2 () nanoparticles. B) Normalized visible spectra of Au (a) and J2 (b) nanoparticles…………………………………………………………………….………………………….. 328

Figure 4. Kinetics of dye release from J2Ru-U in 5 mM sodium acetate buffer pH 5.0 in the absence () and the presence of 180 mM urea at t = 0 () or t = 5 h ()…………………………………………………………………………………………………………..

332


12

Figure 1S. TEM images and distribution of sizes of MS nanoparticles……………. 340

Figure 2S. Distribution of sizes of J1 (A), J2 (B), J3 (C) and J4 (D) nanoparticles…………………………………………………………………………………………………. 340

Figure 3S. TEM images of J2 nanoparticles……………………………………………………. 341

Figure 4S. A) TG and B) DTG analysis for MS (a), J2 (b) and J2Ru (c) nanoparticles………………………………………………………………………………………………... 341

Figure 5S. A) FT-IR analysis for MS (a), J2 (b) and J2Ru (c) nanoparticles. B) X-ray diffraction of J2 (a) and J2Ru (b) nanoparticles………………………………………..

342

7. 12. Journal of the American Chemical Society 136 (25) (2014) 9116–9123

Figuras:

Scheme 1. Schematic representation of “smart” delivery systems containing an attached control unit that regulates the delivery activity of the gated material………………………………………………………………………………………………………... 345

Figure 1. Performance of the Janus-based nanodevice S3. The “control unit” (Au face) is functionalized with two effectors (enzymes) which control cargo delivery from the silica mesoporous face via interpretation of different chemical inputs (D-glucose, ethyl butyrate). Overall the system functions as an enzymatic logical OR operator………………………………………………………………….. 346

Figure 2. Kinetics of dye release from S3 in 20 mM Na2SO4, pH 7.5 in the absence (a) and the presence of 40 µM ethyl butyrate (b), D-glucose (c) and ethyl butyrate + D-glucose (d). Substrates were added after 1 h of incubation…………………………………………………………..…………………………………………. 350

Figure 3. Kinetics of dye release from S4 in 20 mM Na2SO4, pH 7.5 in the absence (a) and the presence of 40 µM ethyl butyrate (b), D-glucose (c) and ethyl butyrate + D-glucose (d) without (closed circles) and with 200 µM urea (open circles). Substrates were added ter 1 h of incubation………………………….

351

Figure 4. Kinetics of dye release from S4 in 20 mM Na2SO4, pH 7.5 in the absence (a) and the presence of 40 µM D-glucose without (f) and with addition of urea at 200 µM final concentration at different times (b-e). Substrates were added after 1 h of incubation……………………………………………… 353

Figure 5. Kinetics of Doxo release from S5 in 20 mM Na2SO4, pH 7.5 in the absence (a) and the pres-ence of 40 µM ethyl butyrate (b), D-glucose (c) and ethyl butyrate + D-glucose (d). Substrates were added after 1 h of incubation……………………………………………………………………………………………………..

354


13

Figure 6. Internalization and release of cargo in HeLa cells A) controlled release of Doxorubicin (Doxo) loaded S5 nanoparticles in HeLa cells. Culture were incubated with 100 μg/mL of S5 and in presence of different inputs and examined for Doxo by confocal microscopy. Representative images at 24 h form phase contrast (PhC), Doxo (DOX), Hoescht (HOE) and combined (Merge) are shown. B) Cell viability test of 150 μg/mL concentration of S5 and glucose and/or ethyl butyrate at 24 h in HeLa cells using WST- assay and C) quantification of Doxo fluorescence intensity by flow cytometry in cells under different conditions. Ethyl butyrate treatment (input A), glucose treatment (input B)……………………………………………………………………………………….. 357

Figure SI-1. Powder X-ray diffraction of nanoparticles S0, S1, S2 and S3 at low (A) and high (B) angles……………………………………………………………………………. 369

Figure SI-2. FT-IR analysis for the nanoparticles S0, S1, S2 and S3…………………. 370

Figure SI-3.Thermogravimetric analysis for S0, S1, S2 and S3……………………….. 370

Figure SI-4. Nitrogen adsorption (closed)/desorption (open) isotherms for S0 to S3 nanoparticles…………………………………………..…………………………………………… 372

Figure SI-5. Kinetics of dye release from S3 in 20 mM Na2SO4, pH 7.5 in the absence (a) and the presence of 40 µM D-glucose + ethyl butyrate (b). Kinetics of release from S3 in 50 mM sodium phosphate buffer, pH 7.5 in the presence of 40 µM D-glucose + ethyl butyrate (c). Substrates were added after 1 h of incubation……………………………………………………………………………………

373

Figure SI-6. Influence of time of incubation in reconstituted human serum at 37ºC on the release activi-ty of S5 upon addition of 40 µM D-glucose + ethyl butyrate……………………………………………………………………………………………………..…. 374

Tablas:

Table SI-1. Elemental analysis for S0, S1, S2 and S3………………………………………. 371

Table SI-2. BET specific surface values, pore volumes and pore sizes calculated from the N2 adsorption-desorption isotherms for selected materials………………………………………………………………………………………………………. 372

8. DISCUSIÓN INTEGRADORA:

Figuras:

Figura 8.1. Representación esquemática de las nanopartículas de oro polifuncionalizadas……………………………………………………………………………………….. 376

Figura 8.2. Imágenes de las nanopartículas de oro polifuncionalizadas con residuos de ácido 3-mercaptofenilborónico, obtenida mediante TEM a 200 kV (A) y 300 kV (B)…………………………………………………………………………………………. 378

Figura 8.3. Espectro FT-IR de las nanopartículas de oro polifuncionalizadas con residuos de ácido 3-mercaptofenilborónic………….…………………………………. 379


14

Figura 8.4. Imágenes FE-SEM de redes de nanopartículas de oro polifuncionalizadas con residuos de ácido 3-mercaptofenilborónico, obtenida mediante métodos químicos (A) y electroquímicos (B) de polimerización………………………………………………………………………………………………. 380

Figura 8.5. Imágenes FE-SEM de redes de nanopartículas de oro polifuncionalizadas y (A) con residuos de ácido 3-mercaptofenilborónico crecidas sobre electrodos de oro, y (B) con residuos de 1-adamantano y crecidas sobre electrodos de carbono vitrificado recubiertos con nanotubos de carbono de pared simple………………………………………………………………………….. 381

Figura 8.6. Representación esquemática de los procesos de síntesis y electropolimerización de las nanopartículas de oro polifuncionalizadas……….. 382

Figura 8.7. Estructura del polímero conductor de polianilina………………………… 383

Figura 8.8. Voltamperogramas cíclicos para el primer paso de electropolimerización de las nanopartículas de oro polifuncionalizadas con residuos de ácido 3-mercaptofenilborónico sobre electrodos de oro, en solución de H2SO4 0.1 M. Velocidad de barrido: 100 mV/s……………………………. 384

Figura 8.9. Representación esquemática del proceso de ensamblado del electrodo enzimático para la detección de H2O2…………………………………………….

385

Figura 8.10. Estructura tridimensional de la peroxidasa de rábano……………….. 386

Figura 8.11. Funcionalización supramolecurar no covalente de SWCNTs con: A) Nanoparticulas Fe3O4 modificadas con PEG y XO; y B) Pireno funcionalizado con CD para la formación de un complejo de inclusión con la XO-ADA………......................................................................................................

391

Figura 8.12. Efecto del campo magnético externo generado por un imán de neodimio sobre una dispersión de nanotubos de carbono de pared simple de concentración 0.1 mg/mL, antes (A) y después (B) de su funcionalización con nanopartículas superparamagnéticas de Fe3O4 modificadas con polietilenglicol……………………………………………………………………………………………….

393

Figura 8.13. Representación esquemática del proceso de ensamblado del electrodo con: (A) PAMAM G4-CD/Enzima-ADA y (B) PAMAM G4-CD/Enzima-ADA/PAMAM G5-CD/ Enzima-ADA……………………………………………………………….. 396

Figura 8.14. Representación esquemática del mecanismo de formación de ésteres cíclicos de ácido borónico con azúcares……………………………………………

397

Figura 8.15. Representación esquemática de una nanomáquina para la liberación sitio-específica, autónoma e inteligente de fármacos bajo control lógico modulado por enzimas………………………………………………………………………

404

Figura 8.16. Estrategia de enmascaramiento y manipulación selectiva de interfases empleada en la preparación de nanopartículas Janus de oro y sílice mesoporosa………………………………………………………………………………………..… 405

Figura 8.17. Imagen FE-SEM de los coloidosomas de parafina recubiera con nanopartículas de sílice mesoporosa…………………………….…………………………….…

405

2 ABREVIATURAS Y

SÍMBOLOS

2. ABREVIATURAS Y SÍMBOLOS

15

ADA: Adamantano

AFM: Microscopía de fuerza atómica

APTES: Aminopropiltrimetoxisilano

BSA: Albúmina de suero bovino

CD: Ciclodextrina

CNTs: Nanotubos de carbono

CTAB: Bromuro de hexadeciltrimetilamonio

CV: Voltamperometría Cíclica

DMF: Dimetilformamida

DMSO: Dimetilsulfóxido

DNA: Ácido desoxirribonucleico

e-: Electrones

EDAC: 1-Etil-3-(3-dimetilaminopropil) carbodiimida

EIS: Espectroscopía de impedancia electroquímica

EtOH: Etanol

FE-SEM: Microscopía Electrónica de Barrido con Emisión de Campo

FTIR: Espectroscopia de infrarrojo con transformada de Fourier

GA: Glutaraldehído

GCE: Electrodo de carbono vitrificado

GO: Óxido de grafeno

GOx: Glucosa oxidasa

GPTMS: (3-glicidiloxipropil)trimetoxisilano

HRP: Peroxidasa de rábano picante

HR-TEM: Microscopía de transmisión electrónica de alta resolución

ISO: Organización Internacional de Normalización

IUPAC: Unión Internacional de Química Pura y Aplicada

MES: Ácido 2-morfolinoetanosulfónico

mRNA: Ácido ribonucleico mensajero

MSNs: Nanopartículas de sílice mesoporosa

MWCNTs: Nanotubos de carbono de pared múltiple

NADH: Nicotinamida adenina dinucleótido de hidrógeno

NHS: N-hidroxisuccinimida

NPs: Nanopartículas

PAMAM: Dendrímero de poliamidoamina

PBS: Disolución reguladora de fosfato salino

PEG: Polietilenglicol

PolyAuNPs: Nanopartículas de oro polimerizadas

QDots: Puntos cuánticos

SDS: Dodecilsulfato sódico

SPCEs: Electrodos serigrafiados de carbono

Strep-HRP: Estreptavidina-Peroxidasa

Sulfo-NHS: N-sulfohidroxisuccinimida

SWCNTs: Nanotubos de carbono de pared simple

TEM: Microscopía de transmisión electrónica

TEOS: Tetraetilortosilicato

TG: Termogravimetría

Tyr: Tirosinasa

UV-VIS: Ultravioleta visible

XO: Xantina Oxidasa

XRD: Difracción de Rayos X

3 SUMMARY

3. SUMMARY

17

Nanotechnology, and specially nanomaterials engineering, has opened new

possibilities to science and nowadays is the core of emerging technologies by providing

a great variety of advanced functional nanomaterials with unique and well-defined

characteristics. These nanomaterials have allowed the design of novel drug delivery

systems, biomaterials, protective coatings, electronic devices, functional and wearable

textiles and sensor systems with improved properties and nanometric dimensions.

In this context, the ultimate goal for nanomaterials engineering is the

establishment of original strategies for the tailor-made preparation of nanosized

structures with desired physicochemical and functional properties by rational

manipulation of their chemical composition, morphology, size and surface

derivatization. In special, great attention is currently devoted to the design of novel

functionalized nanomaterials and nanohybrids for emergent biologically driven

applications. These materials should be provided with specific chemical functionalities

to ensure high biocompatibility, hydrophilicity and capacity for the stable

immobilization of biologically-active macromolecules.

This PhD Thesis, presented as a compendium of published research articles,

describes original approaches for the preparation and assembly of nanomaterials-

based devices for sensitive electrochemical biosensing and smart drug delivery. In the

part devoted to electrochemical biosensors, the first paper (Electrochimica Acta 56,

2011, 4672-4677) describes the construction of a reagentless amperometric biosensor

for H2O2 wherein a novel one-pot preparation of gold nanoparticles polyfunctionalized

with 2-mercaptoethanesulfonic acid as solubilizing agent, p-aminothiophenol as

polymer-forming residue, and 3-mercaptophenyl boronic acid for enzyme

immobilization is presented. Gold electrodes were functionalized with an

electropolymerized matrix of these nanoparticles, and the resulting nanostructured

electroconductive matrix was used as support for the oriented immobilization of the

enzyme horseradish peroxidase to construct the reagentless amperometric biosensor

for H2O2 detection. The electrode, poised at 0.0 mV, exhibited a rapid response within

8 s and a linear calibration range from 5 µM to 1.1 mM H2O2. The sensitivity of the

biosensor was determined as 498 µA/M cm2, and its detection limit was 1.5 µM H2O2

3. SUMMARY

18

at a signal-to-noise ratio of 3. The electrode retained 95% and 72% of its initial activity

after 21 and 40 days of storage at 4ºC.

In the second paper (Analyst 137, 2012, 342-348), a biosensor for catechol is

presented herein polyfuntionalized gold nanoparticles provided with 2-

mercaptoethanesulfonic acid, p-aminothiophenol and cysteamine core

polyamidoamine G-4 dendron moieties were prepared. The nanoparticles were

electropolymerized on Au electrode surface through the formation of bis-aniline cross-

linked network. The enzyme tyrosinase was further crosslinked on this nanostructured

matrix. The enzyme electrode, poised at -100 mV, was then used for the amperometric

quantification of catechol. The biosensor showed a linear response from 50 nM to 10

µM catechol, with a low detection limit of 20 nM and a sensitivity of 1.94 A/M cm2.

The electrode retained 96% and 67% of its initial activity after 16 and 30 days of

storage at 4°C under dry conditions.

The third presented paper (ChemElectroChem 1, 2014, 200-206) describes an

amperometric immunosensor system to detect human fibrinogen. In this work we

introduce the preparation of water soluble gold nanoparticles (3.1 ± 0.6 nm) with

polymerization ability and affinity to streptavidin, by reducing HAuCl4 in the presence

of the capping ligands 2-mercaptoethanesulfonic acid, p-aminothiophenol and a

biotin-cysteamine derivative. This colloid was used to modify gold electrodes by

formation of an electropolymerized 3D network of bis-aniline cross-linked

nanoparticles on the metal surface. The modified electrode was employed as scaffold

for the assembly of an amperometric immunosensor system to detect human

fibrinogen. The immunosensor showed excellent analytical characteristics, with a

dynamic range of detection between 0.018 and 2.208 µg/mL, a detection limit of 4

ng/mL and an IC50 value of 177 ng/mL. The immunosensor was markedly stable,

retaining full analytical capacity after 45 days of storage at 4ºC.

In the fourth presented paper (ACS Applied Materials & Interfaces 4, 2012,

4312-4319) we report a new xanthine biosensor design. The novel preparation of the

biosensor is based on glassy carbon electrodes modified with single walled carbon

nanotubes and a three-dimensional network of electropolymerized Au nanoparticles

3. SUMMARY

19

capped with 2-mercaptoethanesulfonic acid, p-aminothiophenol and 1-

adamantanethiol which were used as hybrid electrochemical platforms for

supramolecular immobilization of a synthesized artificial neoglycoenzyme of xanthine

oxidase and β-cyclodextrin through host-guest interactions. This ensemble was thus

further employed for the bioelectrochemical determination of xanthine. The biosensor

showed fast amperometric response within 5 s and a linear behavior in the 50 nM –

9.5 µM xanthine concentration range with high sensitivity, 2.47 A/M·cm2, and very low

detection limit of 40 nM. The stability of the biosensor was significantly improved and

the interferences caused by ascorbic and uric acids were noticeably minimized by

coating the electrode surface with a Nafion thin film.

The fifth paper (Electroanalysis 23, 2011, 1790-1796) reports a xanthine

biosensor based on a novel approach for the non-covalent functionalization of single

walled carbon nanotubes with enzymes, using a β-cyclodextrin-modified pyrene

derivative, mono-6-ethylenediamino-(2-pyrene carboxamido)-6-deoxy-β-cyclodextrin,

as a molecular bridge for the construction of a supramolecular assembly between the

nanotube surface and an adamantane-modified enzyme. The β-cyclodextrin-modified

pyrene derivative was synthesized and its stacking to single-walled carbon nanotubes

through π-π interactions accomplished. The functionalized carbon nanotubes showed

low capacity for the non-specific adsorption of proteins, but were able to immobilize

adamantane-modified xanthine oxidase via host-guest associations. This double

supramolecular junctions-based approach was employed to modify a glassy carbon

electrode with the enzyme/nanotubes complex for designing a biosensor device

toward xanthine. The biosensor showed fast electroanalytical response (10 s), high

sensitivity (5.9 mA/M cm2) low detection limit (2 µM) and high stability.

In the sixth presented paper (Journal of Materials Chemistry 21, 2011, 12858-

12864) an original xanthine biosensor is reported. Here, superparamagnetic Fe3O4

nanoparticles were coated with (3-aminopropyl)triethoxysilane and further branched

with monomethoxypolyethylene glycol chains. These nanoparticles were employed for

the non-covalent surface modification of single walled carbon nanotubes, conferring

them magnetic properties. This nanomaterial was employed to immobilize the enzyme

3. SUMMARY

20

xanthine oxidase in order to construct magnetically modified disposable gold screen-

printed electrodes as bioelectrodes for the determination of xanthine. The

electroanalytical properties of the biosensor were modulated by the nanomaterial

composition, being optimal at a carbon nanotubes:magnetic nanoparticles ratio of

1:27. The resulting biosensor showed a linear dependence on the xanthine

concentration in the 0.25-3.5 µM range with a fast amperometric response in 12 s. The

biosensor also showed a noticeable high sensitivity of 1.31 A/M cm2 and a very low

detection limit of 60 nM, which can be compared advantageously with other biosensor

designs for xanthine determination.

The seventh paper (Analytical and Bioanalytical Chemistry 405, 2013, 3773-

3781) reports the construction of a reagentless amperometric biosensor for glucose.

Wherein the novel biosensor fabrication process describes the modification of Au

electrodes with cysteamine core polyamidoamine G-4 dendrons branched with β-

cyclodextrins and further decoration with Pt nanoparticles. Adamantane-modified

glucose oxidase was subsequently immobilized on the nanostructured electrode

surface through supramolecular associations. The so-constructed enzyme electrode

was then employed for constructing a reagentless amperometric biosensor for glucose

making use of the electrochemical oxidation of H2O2 generated in the enzyme reaction.

The biosensor exhibited a fast amperometric response (6 s) and a linear response

toward glucose concentration between 5 µM and 705 µM. The biosensor showed a

low detection limit of 2.0 µM, a sensitivity of 197 mA/M cm2, and retained 94% of its

initial response after nine days of storage at 4ºC.

And finally, the eighth paper of this thesis (Electrochimica Acta 76, 2012, 249-

255) describes the design of an original bioelectrode for the detection of H2O2. It

presents a new layer-by-layer supramolecular approach for the construction of self-

assembled nanoarchitectures of polyamidoamine dendrimers and peroxidase on gold

surface. The methodology was based on the supramolecular self-assembly of

alternated layers of adamantane-modified horseradish peroxidase and β-cyclodextrin-

branched polyamidoamine G-5 dendrimers on a gold electrode, previously coated with

β-cyclodextrin-modified cysteamine core polyamidoamine G-4 dendrons. The

3. SUMMARY

21

formation of layer-by-layer assemblies (up to three dendrimer/peroxidase bilayers)

was studied by Surface Plasmon Resonance, quartz crystal microbalance, Atomic Force

Microscopy and cyclic voltammetry. The analytical applicability of these architectures

was evaluated by constructing a H2O2 biosensor. The electroanalytical response of the

biosensor towards H2O2 increased with the number of enzyme layers. The bioelectrode

constructed with three enzyme layers showed a low detection limit of 160 nM, a

sensitivity of 602 µA/M cm2 and retained 63% of its initial activity after 30 days of

storage in wet conditions.

In the second part devoted to the design of enzyme-controlled smart drug

delivery nanosystems based on mesoporous silica, the first paper (ACS Applied

Materials and Interfaces 8, 2016, 7657−7665) describes the assembly of a stimulus-

programmed pulsatile delivery system for sequential cargo release based on the use of

a lactose-modified esterase as a capping agent in phenylboronic acid functionalized

mesoporous silica nanoparticles. The dual release mechanism was based on the

distinct stability of the cyclic boronic acid esters formed with lactose residues and the

long naturally-occurring glycosylation chains in the modified neoglycoenzyme. Cargo

delivery in succession was achieved by using glucose and ethyl butyrate as triggers.

Moreover, it was also demonstrated that the same control was observed with

nanoparticles loaded with the anticancer drug Doxorubicin and tested in HeLa cancer

cells.

In thereafter presented papers we report our results on the novel use of

original Au-mesoporous silica Janus nanoparticles as “hardware” for the assembly of

enzyme-empowered smart nanomachines for drug delivery. In the paper published in

Electrochemistry Communications 30, 2013, 51-54, we demonstrated the capacity of

these nanoparticles to recognize small molecules after proper functionalization with

affinity-based bioreceptors. In this sense, Janus Au-mesoporous silica nanoparticles

were used as scaffolds to design an integrated electrochemical biorecognition-

signaling system. A proof of concept of this strategy, based on the face-selective

functionalization of the anisotropic colloid, involves the covalent immobilization of

horseradish peroxidase on the mesoporous silica face as enzymatic signaling element,

3. SUMMARY

22

as well as the modification of the Au face with streptavidin and polyethylenglycol

chains as biorecognition and solubilizing agents, respectively. The functionalized Janus

nanoparticles were successful to recognize biotin on gold surfaces.

In thereafter presented paper, published in Chemistry – A European Journal 19,

2013, 7889-7894, we described the assembly of a prime enzyme-controlled Janus

nanoparticle-based nanomachine. In summary, novel Janus nanoparticles with Au and

mesoporous silica opposite faces were prepared by Pickering emulsion template using

paraffin wax as oil phase. These anisotropic colloids were employed to design an

integrated sensing-actuating nanomachine for the enzyme-controlled stimulus-

responsible cargo delivery. As a proof-of-concept, we demonstrated the successful use

of the Janus colloids for controlled delivery of tris(2,2´-bipyridyl) ruthenium(II) chloride

from the mesoporous silica face grafted with pH-sensitive gate-like scaffoldings. The

release was mediated by the on-demand catalytic decomposition of urea by urease,

which was covalently immobilized on the Au face.

Finally, in the paper published in Journal of the American Chemical Society 136,

2014, 9116-9123, we described a more sophisticated nanomachine controlled by an

enzyme logic gate. In this sense, we reported the design of a smart delivery system in

which cargo delivery from capped mesoporous silica nanoparticles was controlled by

an integrated enzyme-based “control unit”. The system consisted of Janus-type

nanoparticles having Au and mesoporous silica nanoparticles opposite faces, which

were properly functionalized with a pH-responsive β-cyclodextrin based

supramolecular nanovalve on the silica mesoporous surface and two effectors, glucose

oxidase and esterase, immobilized on the Au face. The nanodevice behaves as an

enzymatic logical OR operator which is selectively fuelled by the presence of D-glucose

and ethyl butyrate as INPUT signals. This enzyme logic system was also coupled to a

urease-based RESET operator to switch-off the opening of the supramolecular

nanovalves and control the extension of dye delivery upon addition of urea. The smart

nanomachine controlled by the logical OR operator and loaded with the anticancer

drug Doxorubicin, was successfully tested toward HeLa cancer cells.

4 RESUMEN

4. RESUMEN

23

La Nanotecnología, y en especial la ingeniería de nanomateriales, han abierto

nuevas posibilidades a la ciencia y la tecnología, proporcionando una gran variedad de

nanomateriales funcionales avanzados con propiedades únicas y bien definidas. Estos

nanomateriales han permitido el diseño de nuevos sistemas de liberación de fármacos,

biomateriales, recubrimientos protectores, dispositivos electrónicos, textiles

funcionales y sistemas sensores de dimensiones nanométricas y propiedades

mejoradas.

En este contexto, el objetivo último de la ingeniería de nanomateriales es el

diseño de estrategias originales para la preparación a medida de nanoestructuras con

las propiedades fisicoquímicas y funcionales deseadas, mediante la manipulación

racional de la composición química, morfología, tamaño y superficie de derivatización.

Actualmente, el desarrollo de nuevos nanomateriales funcionalizados y nanohíbridos

para el desarrollo de aplicaciones biológicas emergentes es de especial interés. Estos

materiales deben proporcionar grupos funcionales específicos para garantizar una alta

biocompatibilidad y capacidad para la inmovilización estable de macromoléculas

bioactivas, e hidrófilas.

En esta Tesis Doctoral, presentada como un compendio de artículos de

investigación publicados, se describen distintas posibilidades para la preparación y el

ensamblaje de sistemas basados en nanomateriales para el desarrollo de biosensores

electroquímicos altamente sensibles y sistemas de liberación inteligente de fármacos.

En la sección dedicada a los biosensores electroquímicos, en el primer artículo

(Electrochimica Acta 56, 2011, 4672-4677) se describe la construcción de un biosensor

amperométrico de H2O2 en la cual se realiza la preparación en una sola etapa de

nanopartículas de oro polifuncionalizadas con ácido 2-mercaptoetanosulfónico como

agente solubilizante, p-aminotiofenol como polimerizante y ácido 3-mercaptofenil

borónico para la inmovilización enzimática. Estas novedosas nanopartículas se crearon

para la formación mediante electropolimerización de una matriz sobre la superficie de

electrodos de oro. La nanoestructura generada, con propiedades electroconductoras

mejoradas, se empleó para la inmovilización orientada de peroxidasa de rábano, con

objeto de preparar un biosensor amperométrico de H2O2. El potencial de trabajo del

4. RESUMEN

24

electrodo (0.0 mV) elimina posibles interferencias. Además, el biosensor exhibe una

respuesta rápida (8 s), y un intervalo lineal entre 5 µM y 1.1 mM frente a H2O2. Se

determinó la sensibilidad, con un valor de 498 µA/M cm2, y el límite de detección, 1.5

µM, con una relación señal-ruido de 3. Pasados 21 y 40 días de almacenamiento a 4oC,

la actividad del electrodo, respecto a la que poseía inicialmente, fue del 95 y 72%

respectivamente.

En el segundo artículo (Analyst 137, 2012, 342-348) presentado, se muestra la

preparación de nanopartículas de oro polifuncionalizadas con ácido 2-

mercaptoetanosulfónico, p-aminotiofenol y dendrones de poliamidoamina G-4 con

núcleo de cisteamina. Las nanopartículas se electropolimerizaron sobre la superficie de

electrodos de oro, mediante la formación de una red entrecruzada de enlaces bis-

anilina. La enzima tirosinasa fue posteriormente inmovilizada en la matriz mediante

entrecruzamiento. El electrodo enzimático obtenido, con un potencial de trabajo de

-100 mV, se empleó para la cuantificación mediante amperometría de catecol. Este

biosensor mostró una respuesta lineal entre 50 nM y 10 µM frente a catecol, con un

límite de detección de 20 nM y una sensibilidad de 1.94 A/M cm2. El electrodo retuvo

el 96% y el 67% de su actividad inicial después de 16 y 30 días de almacenamiento

respectivamente, a 4oC y en condiciones de humedad.

En el tercer artículo (ChemElectroChem 1, 2014, 200-206) se describe la

preparación de nanopartículas de oro hidrosolubles (3.1 ± 0.6 nm), con capacidad de

polimerización y afinidad frente a la estreptavidina, mediante reducción de HAuCl4, en

presencia de ácido 2-mercaptoetanosulfónico, biotina-tiolada y ácido p-aminotiofenol

como agentes funcionales. La estructura generada se empleó, mediante el uso de

anticuerpos modificados con estreptavidina, para la modificación de electrodos de oro

mediante formación de una red tridimensional electropolimerizada por

entrecruzamiento mediante enlaces tipo bis-anilina sobre la superficie del metal. El

electrodo modificado se empleó como soporte para el ensamblaje de un

inmunosensor para la detección de fibrinógeno humano. El inmunosensor mostró

excelentes propiedades analíticas, con un rango lineal entre 0.018 and 2.208 µg/mL,

un límite de detección de 4 ng/mL y un valor de IC50 de 177 ng/mL. El inmunosensor

4. RESUMEN

25

fue notablemente estable, conservando todo su potencial analítico después de 45 días

de almacenamiento a 4oC.

El cuarto artículo (ACS Applied Materials & Interfaces 4, 2012, 4312-4319)

recoge el ensamblaje de una plataforma híbrida sobre electrodos de carbono

vitrificado, para la inmovilización supramolecular de una neoglicoenzima sintetizada

artificialmente a partir de xantina oxidasa y β-cyclodextrin, mediante interacciones

“host-guest”. El nanomaterial empleado como estructura base se preparó a partir de

nanotubos de carbono de pared simple y una red tridimensional de nanopartículas de

oro electropolimerizadas, modificadas con ácido 2-mercaptoetanosulfónico,

p-aminotiofenol y 1-adamantanotiol. La estructura diseñada se empleó para la

determinación electroquímica de xantina. El biosensor mostró una respuesta

amperométrica rápida, del orden de 5 s, un comportamiento líneal entre 50 nM – 9.5

µM de xantina, una alta sensibilidad (2.47 A/M·cm2) y un bajo límite de detección. La

estabilidad del biosensor fue significativamente mejorada, y las interferencias

provocadas por los ácidos ascórbico y úrico notablemente minimizadas, gracias al

recubrimiento de la superficie electródica con una película delgada de Nafión.

En el quinto artículo (Electroanalysis 23, 2011, 1790-1796) se presenta una

estrategia novedosa para la funcionalización no covalente de nanotubos de carbono de

pared simple con enzimas, mediante un derivado de pireno modificado con

β-ciclodextrina [mono-6-etilenediamino-(2-pireno carboxamido)-6-desoxi-β-

ciclodextrina], como puente molecular para la construcción de un ensamblaje

supramolecular entre la superficie de los nanotubos y una enzima modificada con

adamantano. El derivado de β-ciclodextrina modificado con pireno se sintetizó e

inmovilizó sobre los nanotubos de carbono de pared simple mediante interacciones

π-π. Los nanotubos funcionalizados permitieron la inmovilización de xantina oxidasa

modificada con adamantano mediante interacciones huésped-hospedero, pero

mostraron una baja adsorción inespecífica frente a otras proteínas. Este enfoque de

asociaciones supramoleculares dobles fue empleado para la modificación de

electrodos de carbono vitrificado con el complejo enzima/nanotubos para el diseño de

4. RESUMEN

26

un sensor de xantina. El dispositivo mostró una respuesta electroanalítica rápida (10 s),

alta sensibilidad (5.9 mA/M cm2), bajo límite de detección (2 µM) y alta estabilidad.

En el sexto artículo seleccionado (Journal of Materials Chemistry 21, 2011,

12858-12864) se presenta un biosensor para la detección de xantina. En este caso se

cubrieron partículas superparamagnéticas de Fe3O4 con (3-aminopropil)trietoxisilano y

posteriormente se modificaron con cadenas de monometoxipolietilenglicol, generando

una estructura ramificada. Estas nanopartículas se emplearon para la modificación no

covalente de nanotubos de carbono de pared simple, confiriendo sus propiedades

magnéticas. Este nanomaterial se empleó para la inmovilización de la enzima xantina

oxidasa, con el fin de construir un biosensor desechable sobre la superficie de

electrodos serigrafiados de oro para la determinación de xantina. Las propiedades

electroanalíticas del biosensor fueron moduladas mediante la composición del

material, siendo 1/27 la proporción óptima entre nanotubos de

carbono/nanopartículas magnéticas. El biosensor resultante mostró una tendencia

lineal frente a la xantina entre 0.25-3.5 µM, proporcionando una respuesta

amperométrica en 12 s. Además, la sensibilidad del dispositivo fue notablemente alta,

con un valor de 1.31 A/M cm2, y un límite de detección de 60 nM, considerablemente

mejor que otros biosensores para la determinación de xantina.

El séptimo artículo que aquí se presenta (Analytical and Bioanalytical Chemistry

405, 2013, 3773-3781) recoge la construcción de un biosensor amperométrico para la

determinación de glucosa. El proceso de preparación del biosensor describe la

modificación de electrodos de oro con dendrones de poliamidoamina G-4 con núcleo

de cisteamina ramificados con β-ciclodextrinas y posteriormente decorados con

nanopartículas de platino. Seguidamente, la enzima glucosa oxidasa fue modificada

con adamantano, e inmovilizada en la nanoestructura del electrodo mediante

asociaciones supramoleculares. El electrodo enzimático preparado se utilizó para la

determinación de glucosa, a partir de la oxidación electroquímica del H2O2 generado

en la reacción enzimática. La respuesta amperométrica del biosensor fue

extraordinariamente rápida (6s), y lineal en un intervalo entre 5 y 705 µM. El límite de

detección del biosensor fue de 2.0 µM, con una sensibilidad de 197 mA/M cm2.

4. RESUMEN

27

Después de 9 días de almacenamiento a 4oC, la respuesta analítica seguía siendo del

94% respecto a la inicial.

El octavo artículo que aquí se recoge (Electrochimica Acta 76, 2012, 249-255)

describe el diseño original de un bioelectrodo para la detección de H2O2. Se presenta

una aproximación novedosa de nanoarquitecturas autoensambladas de dendrímeros

de poliamidoamina y peroxidasa sobre la superficie de oro. La metodología está

basada en el autoensamblaje supramolecular de capas de peroxidasa de rábano

modificada con andamantano, y dendrímeros G-5 ramificados con β-ciclodextrinas,

sobre la superficie de electrodos de oro, previamente cubiertos con núcleos de

dendrones G-4 de poliamidoamina recubiertos con β-ciclodextrinas modificada con

cisteamina. La formación de estructuras autoensambladas capa a capa (hasta tres

bicapas de dendrímero/ peroxidasa) se estudió mediante Resonancia de Plasmón

Superficial, Microbalanza de Cristal de Cuarzo, Microscopía de Fuerza Atómica y

Voltamperometría Cíclica. La aplicabilidad analítica de estas arquitecturas se evaluó

mediante la construcción de un biosensor de H2O2, incrementándose la respuesta al

aumentar el número de capas de enzima. El bioelectrodo preparado con tres estratos

de enzima mostró un límite de detección de 160 nM, y una sensibilidad de 602 µA/M

cm2. La actividad del sensor tras 30 días de almacenamiento en condiciones de

humedad se mantuvo en el 63% respecto a la inicial.

La segunda parte de esta tesis doctoral está dedicada al diseño de

nanosistemas inteligentes de liberación de fármacos basados en sílice mesoporosa,

controlados mediante enzimas. En el primer artículo de este capítulo (ACS Applied

Materials and Interfaces 8, 2016, 7657−7665) se describe el ensamblaje de sistemas

estímulo-dependientes pulsátiles para la liberación secuencial del cargo, basado en el

empleo de esterasa modificada con lactosa como elemento terminal sobre partículas

de sílice mesoporosa modificadas con ácido fenilborónico. El mecanismo de liberación

dual está basado en la diferencia de estabilidad de los ésteres cíclicos del ácido

borónico generados con los residuos de lactosa y las cadenas glicosiladas presentes de

forma natural en la neoglicoenzima. La liberación sucesiva del cargo se desencadena

4. RESUMEN

28

en presencia de glucosa y etil butirato. Además, se ensayó la posibilidad de liberar el

fármaco anticancerígeno Doxorubicina encapsulado en las nanopartículas previamente

descritas en células de la línea HeLa.

En los últimos artículos incluidos en esta tesis doctoral se describe el empleo de

nanopartículas tipo Janus, integradas por una nanopartícula de sílice mesoporosa

“sistema contenedor” y una de oro “sistema efector”, como “hardware”, para el

ensamblaje de nanomáquinas inteligentes para la liberación de fármacos.

En el artículo publicado en Electrochemistry Communications 30, 2013, 51-54 se

mostró la capacidad de estas nanopartículas para reconocer pequeñas moléculas

después de la adecuada funcionalización mediante biorreceptores de afinidad. En este

sentido, las nanopartículas Janus de oro y sílice mesoporosa se emplearon como

estructura para el diseño de un sistema de reconocimiento de señales electroquímicas.

Como prueba de concepto, y aprovechando la funcionalización selectiva de las

partículas gracias a su anisotropía, se inmovilizó peroxidasa de rábano en la cara de

sílice mesoporosa como elemento de emisión de señal enzimática, y cadenas de

polietilenglicol y estreptavidina en la cara del oro, como agentes solubilizantes y de

bioreconocimiento respectivamente. La nanoestructura generada se inmovilizó sobre

un electrodo de oro y se aplicó al reconocimiento de biotina.

La primera nanomáquina basada en el empleo de Janus controlada

enzimáticamente fue publicada en el artículo Chemistry – A European Journal 19,

2013, 7889-7894. En resumen, novedosas nanopartículas Janus con caras opuestas de

oro y sílice mesoporosa se sintetizaron mediante una emulsión de Pickering,

empleando aceite de parafina como fase oleosa. Estos nanomateriales de naturaleza

anisotrópica se emplearon en el diseño de nanomáquinas sensibles a estímulos para la

liberación del cargo que contenían mediante control enzimático. Como prueba de

concepto, se demostró la aplicación exitosa de las nanopartículas Janus para la

liberación controlada de cloruro de tris(2,2´-bipiridil) rutenio (II) de la cara de sílice

mesoporosa, cubierta con puertas moleculares sensibles a pH. La liberación de este

complejo fue mediada por descomposición catalítica de urea por parte de la enzima

ureasa, inmovilizada en la cara de oro.

4. RESUMEN

29

Por último, la nanomáquina más sofisticada que se recoge en esta tesis doctoral

fue descrita en el artículo Journal of the American Chemical Society 136, 2014, 9116-

9123. Este dispositivo está controlado por una puerta enzimática lógica, que regula la

liberación del cargo encapsulado en la nanopartícula de sílice mesoporosa mediante

un sistema enzimático integrado, denominado “unidad control”. Al igual que en casos

anteriores, el nanorrobot está constituido por caras opuestas de oro (donde se

inmovilizan los efectores enzimáticos esterasa y glucosa esterasa), y sílice mesoporosa,

funcionalizada convenientemente con una nanoválvula supramolecular sensible a pH,

integrada por β-ciclodextrina. El nanodispositivo funciona como una puerta enzimática

lógica tipo OR, sensible a la presencia de D-glucosa y butirato de etilo. Este sistema

lógico enzimático se acopló a un operador tipo RESET mediado por la presencia de

ureasa, capaz de detener la apertura de la nanoválvula supramolecular, y por tanto,

controlar la extensión en que el colorante era liberado frente a la adición de urea. La

nanomáquina inteligente controlada por el operador enzimático OR y cargado con el

fármaco anticancerígeno Doxorubicina fue probada con éxito en células cancerígenas

de la línea celular HeLa.

5 INTRODUCCIÓN

Nanomateriales 5. INTRODUCCIÓN

31

5.1. NANOMATERIALES

5.1.1. Conceptos fundamentales

La nanotecnología se basa en manipular estructuras a nivel atómico o

molecular para diseñar nuevos materiales, componentes y dispositivos con nuevas o

mejoradas propiedades físicas, biológicas, químicas y electrónicas [Puzder et al., 2003].

Durante los últimos años el desarrollo de nanomateriales se ha convertido en uno de

los campos de investigación más dinámicos en el área de la nanotecnología. Este

interés alcanza también otras áreas de la ciencia como la química, la física, la biología y

la ingeniería. En la Figura 5.1 se representa una comparación del tamaño de estos

nanomateriales con otros componentes de la materia.

Figura 5.1. Representación comparativa del tamaño de los nanomateriales con otros

componentes de la materia.

1 nm

10 nm

100 nm

1 mm

10 mm

100 mm1 Å

Dendrímeros

CNTs

Fullerenos

Grafeno

Liposomas

AuNPs

MSN

NANOMATERIALES


32

De acuerdo con la Organización Internacional de Normalización (ISO), el prefijo

nano hace referencia a un tamaño que varía aproximadamente entre 1 nm y 100 nm

[ISO/TS 27687, 2008]. En este sentido, los nanomateriales son aquellos materiales en

los cuales al menos una de sus dimensiones se encuentra en la escala nanométrica. En

este rango, los nanomateriales muestran unas propiedades mejoradas con respecto a

las que tiene la materia a escala macroscópica, lo que permite su empleo en nuevas

aplicaciones [Edelstein et al., 1998].

Existen numerosos motivos para el creciente interés en los materiales, entre los

cuales se encuentran los avances conseguidos en medicina gracias a la implementación

de nanopartículas como materiales de diagnóstico y terapia. El empleo de

nanomateriales han permitido también un mayor aumento en el almacenamiento y

velocidad de transmisión de la información, permitiendo la creación de dispositivos de

cómputo cada vez más pequeños y a costes más reducidos [Edelstein et al., 1998].

El uso de los nanomateriales está lejos de ser algo novedoso, pues existen

numerosos ejemplos de su empleo a lo largo de la historia. Alrededor de los años 2600

AC estos materiales ya formaban parte de los tintes que daban color a fibras y telas.

Asimismo, durante la Edad Media los vidrieros utilizaron partículas de oro y plata a

escala nanoscópica para impartir color a los paneles de las ventanas. Otro ejemplo fue

el famoso acero de Damasco producido del siglo XII al XVIII por los orfebres de Oriente

Medio, el cual incluía nanocables de cementita dentro de su composición [Dolez,

2015].

Sin embargo, el progreso en esta área no comenzó realmente hasta 1981 tras el

desarrollo del microscopio de efecto túnel llevado a cabo por los investigadores de

IBM Gerd Binning y Heinrich Rohrer, gracias al cual se hacía posible observar los

materiales a escala nanométrica [Dolez, 2015].

En la Figura 5.2 se recoge una instantánea del interés de la comunidad

científica internacional en los nanomateriales, representando la progresión del número

de publicaciones sobre éstos desde el año 2000 hasta finales del 2015. Como se puede

observar, durante el último decenio la producción científica ha evolucionado de una


33

manera espectacular debido a las prometedoras aplicaciones de estos nuevos

materiales, así como el carácter multidisciplinar de las investigaciones dedicadas a

ellos.

Figura 5.2. Número de publicaciones científicas sobre nanomateriales desde el año

2000 a la actualidad. (Fuente consultada: Web of Science).

5.1.2. Propiedades de los nanomateriales

El gran interés por los nanomateriales y su alta variedad de aplicaciones se

deben a las excelentes propiedades estructurales y funcionales que presentan. En

adición a las propiedades intrínsecas conferidas por su composición química, la alta

relación entre el área superficial y volumen de estos materiales condiciona muchas de

sus propiedades ópticas, de conducción eléctrica y térmica, catalíticas y de reactividad

[Bhushan, 2016].

Este efecto de superficie ultra alta, proporcionado por el incremento del

número de átomos en la superficie tras la disminución del tamaño, provoca la

existencia de una alta densidad de electrones energéticamente enriquecidos y

débilmente atraídos en la superficie del nanomaterial. La alta movilidad de estos


34

electrones confiere marcadas propiedades de electroconductividad y

termoconductividad a muchos nanomateriales. Asimismo, la fácil excitación de estos

electrones a niveles energéticos superiores confiere propiedades ópticas únicas a estos

materiales nanométricos [Liu et al., 2015].

Por lo que respecta a la reactividad, la alta densidad de electrones superficiales,

así como la presencia de orbitales vacantes en los átomos localizados en la superficie

de los nanomateriales, condicionan la alta reactividad química y las inusuales

propiedades catalíticas que presentan muchos de estos nuevos materiales [Jana et al.,

2002]. Además, muchos materiales no magnéticos adquieren nuevas propiedades

magnéticas al pasar de dimensiones macroscópicas a nanométricas. Este fenómeno se

debe a la existencia de momentos magnéticos y anisotropía magnética en dichos

nanomateriales [Liu et al., 2015].

Adicionalmente, muchos nanomateriales poseen excelentes propiedades

mecánicas. Ejemplo de ello son los nanotubos de carbono, los cuales constituyen uno

de los materiales más resistentes de los conocidos o preparados por el hombre

[Salvetat et al., 1999]. Estas propiedades mecánicas excepcionales están

fundamentadas por la alta ordenación de los átomos dentro de estos nanomateriales.

Por otro lado, una de las características distintivas de los nanomateriales es la

posibilidad de ser preparados en diversas formas y tamaños, mediante métodos

relativamente poco costosos [Hulteen et al., 1997]. Esto permite modular muchas de

las propiedades anteriormente citadas de los nanomateriales. Asimismo, la mayoría de

los nanomateriales pueden ser fácilmente derivatizados en su superficie mediante

métodos químicos convencionales, o pueden ser mezclados o modificados con otros

materiales moleculares, poliméricos, cerámicos o nanométricos dando como

resultados nanohíbridos y nanocompósitos con propiedades únicas.


35

5.1.3. Clasificación de los nanomateriales

Los nanomateriales se pueden clasificar en función de diferentes parámetros:

5.1.3.1. Clasificación de acuerdo a su composición:

La clasificación más utilizada viene determinada por la naturaleza química de

los nanomateriales, dividiéndolos entre orgánicos e inorgánicos. En adición, los

nanomateriales inorgánicos sueles ser sub-clasificados en base a su composición en:

nanomateriales de carbono, metálicos, de óxidos metálicos, etc.

5.1.3.2. Clasificación de acuerdo a sus dimensiones espaciales:

Esta clasificación se basa en el número de dimensiones del nanomaterial que

no pueden ser confinadas en el rango nanométrico (<100 nm). En este sentido, los

nanomateriales cero-dimensionales o de dimensión cero (0D) son aquellos en los cuales

las tres dimensiones (X, Y, Z) se incluyen en la escala de 1 nm a 100 nm. Ejemplo de

ellos son los puntos cuánticos, los fullerenos, las nanopartícula coloidades, los

nanoclusters, etc. [Liu et al., 2015].

Los nanomateriales unidimensionales (1D) son aquellos en los cuales una de sus

dimensiones es mayor de 100 nm, tales como los nanotubos de carbono, los

nanocables, las nanofibras, etc. En el caso de los nanomateriales bidimensionales (2D),

tales como el grafeno, las monocapa poliméricas, etc., dos de las dimensiones no se

incluyen en el rango de 1 nm a 100 nm. Por otra parte, cuando las tres dimensiones del

material son mayores de 100 nm, este se clasifica como tridimensional (3D). Estos no

son propiamente nanomateriales, salvo que su estructura interna sea

nanoestructurada.


36

5.1.3.3. Clasificación de acuerdo a su forma de obtención:

Los nanomateriales pueden ser clasificados como naturales, antropogénicos y

sintéticos de acuerdo a su forma de obtención. Aquellos clasificados como naturales

son aquellos que han sido creados por la Naturaleza sin intervención humana alguna-

Los nanomateriales antropogénicos son aquellos producidos involuntariamente por el

hombre, como es el caso de las nanopartículas de carbono producida por la

combustión de petróleo y otros combustibles fósiles. Asimismo, se clasifican como

nanomateriales sintéticos aquellos que han sido preparados voluntariamente por el

hombre.

5.1.4. Aplicaciones de los nanomateriales

Figura 5.3. Campos de aplicación de los nanomateriales [Tsuzuki, 2009].

NANOMATERIALES

TEXTILES

INDUSTRIAL

ALIMENTOS Y AGRICULTURA

ELECTRONICA

ENERGIA BIOMEDICINA

SALUD Y COSMETICA

AMBIENTAL

Tejidos técnicosTejidos médicos

Liberación de fármacos

Terapia de cáncer

Imagen

Antibactericida

Protección UV

Nutracéticos

Fungicidas

Empaque

Nanocompósitosfuncionales

Catalizadores

Nanopigmentos

Plásticos reforzados

Conservación de datos

Nanoimpresión

Computo cuántico

Remediación de aguas

Catálisis ambiental

Catalizadores de celdas de

combustibles

Baterías de litio iónico

Producción fotocatalítica de

hidrógeno

Fibras poliméricas híbridas

Tejidos auto-lavables

Tejidos anti-estáticos

Biocompósitos

Biomarcadores

Terapia hipertérmica

Regeneradores de piel

Regeneradores óseos

Cerámicas dentales

Cerámicas dentales

Imagen de resonancia magnética

Protectores solares

Antioxidantes

Sensores para alimentos

Filmes protectores a gases

Filmes antimicrobianos

Biosensores

Superficies auto-lavables

Líquidos super-termoconductores

Polímeros conductores

híbridos

Cerámicas super-plásticasNanotintes

industriales

Sensores físicos

Sensores de gases

Transistores de electrón simple

Sensores químicos

Magnetos de alta potencia

Láseres cuánticos

Sensores de contaminación

Eliminación de contaminantes

Catalizadores de combustibles

Celdas solares

Almacenamiento de hidrógeno

Almacenamiento de energía

Tejidos termoprotectores


37

Durante las últimas décadas, los nanomateriales han sido ampliamente

utilizados en la obtención de nuevos productos y el desarrollo de aplicaciones

innovadoras en una amplia gama de sectores industriales, como el energético, la

electrónica, la medicina, las telecomunicaciones, la construcción, la alimentación y la

protección y control medioambiental, causando un elevado impacto en la economía

mundial (Figura 5.3).

En 2013 el mercado mundial de los productos de la nanotecnología fue

valorado en 22,9 mil millones de dólares, aumentando en tan solo un año a 26 mil

millones. Con los nuevos avances tecnológicos y el crecimiento de las preocupaciones

por la salud pública y conservación medioambiental, así como la gran demanda por la

miniaturización electrónica, se prevé para el 2019 que el valor en el mercado mundial

haya alcanzado los 64,2 mil millones de dólares, lo que supone una tasa de crecimiento

anual del 19,8 % [McWilliams, 2015].

En un futuro cercano, se espera que los nanomateriales den respuesta a

muchos de los problemas a los que se enfrenta la sociedad en la actualidad. A

continuación se detallan algunas de las aplicaciones más prometedoras:

Alimentación y agricultura: Hoy en día existe un gran interés por mejorar el

valor nutricional de los alimentos, con el fin de aumentar la calidad de vida, así como

de disminuir los gastos sanitarios derivados de la mala alimentación y del aumento en

la esperanza de vida. El uso de la nanotecnología en esta área es cada día más

relevante ya que es barata, relativamente segura y limpia. Los principales avances

conseguidos en este campo son: i) la creación de nuevos productos y materiales

funcionalizados; ii) el desarrollo de métodos de tratamientos de cultivos a escala

nanométrica; y iii) el diseño de métodos e instrumentos para mejorar la seguridad y el

envasado de alimentos.

Por otro lado, se espera que la tecnología de nanoencapsulación ejerza un rol

muy importante en el futuro de la agricultura, mejorando la estabilidad y el control en

la liberación de nutrientes esenciales y pesticidas [Imran et al., 2010].


38

Industria cosmética: El uso de nanomateriales en este campo se orienta

fundamentalmente a las siguientes direcciones: i) mejora de la estabilidad de

ingredientes como vitaminas, antioxidantes y ácidos grasos insaturados; ii) mejora de

la penetración transdermal de vitaminas y antioxidantes; iii) aumento de la eficacia y

tolerancia de los filtros UV para la superficie de la piel; y iv) preparación de productos

estéticamente más agradables [Mu et al., 2010].

Actualmente, muchos de los resultados de las investigaciones en el desarrollo

de nanosistemas de liberación controlada de fármacos están siendo aplicados en esta

industria para el desarrollo de nuevos productos cosméticos. Estos nuevos productos

buscan la mejora de los tratamientos de enfermedades epidérmicas, así como la

liberación controlada de agentes cosméticos como vitaminas, antioxidantes y

coenzimas.

Industria textil: El uso de nanomateriales en la industria textil ha crecido

rápidamente durante los últimos años, fundamentalmente orientados al diseño de

prendas con características mejoradas. En este sentido, la implantación de

nanomateriales puede aportar múltiples funcionalidades a los tejidos, tales como

super hidrofobicidad, blindaje electromagnético, propiedades antibacterianas,

ignifugas, de auto-limpieza, etc. [Hu, 2016].

Industria energética: La necesidad de buscar fuentes de energía alternativas a

los combustibles fósiles que sean sostenibles, respetuosas con el medio ambiente y

rentables económicamente, hacen que en la actualidad se explore con mucho interés

el campo de los nanomateriales como herramientas para la producción y

almacenamiento de energía.

Una de las líneas de investigación más activa en esta área se centra en la

mejora de los sistemas fotovoltaicos existentes mediante el uso de nanomateriales

avanzados con propiedades semiconductores, capaces de capturar los fotones

procedentes del sol y transformarlos en electrones [Sakimoto et al., 2016]. Otra línea

prioritaria en esta área es el desarrollo de nuevos nanomateriales para el diseño de

pilas de combustibles y supercapacitores.


39

Industria de los materiales para la construcción: En la actualidad existen ya

infraestructuras y edificaciones que contienen nanoaditivos añadidos al acero y al

hormigón tradicional. Los nanomateriales han sido también empleados para la

preparación de pinturas y lacas con propiedades de auto-limpieza y protección anti-

grafiti y anti-plagas.

Las investigaciones actuales en este campo se focalizan en el diseño de nuevos

nanoaditivos, nanomateriales aislantes avanzados, recubrimientos funcionales, vidrios

anti-incendios, materiales autorreparables y materiales inteligentes que respondan a

estímulos.

Medioambiente: En los últimos años se han diseñado nuevos nanomateriales

para la remediación del suelo y de los acuíferos contaminados. Algunos de los trabajos

más recientes en este campo se centran en la fabricación de nanomateriales

adsorbentes más eficientes y el diseño de sistemas sensores nanoestructurados para la

monitorización ambiental [Louie et al.; 2016].

Industria electrónica: En este campo destacan las nuevas tecnologías de

visualización y almacenamiento basadas en nanomateriales con propiedades ópticas,

electroconductoras, magnéticas y termoconductoras. Estas tecnologías han sido

aplicadas al diseño de pantallas de alta definición, sistemas de almacenamiento de

gran densidad para el registro de datos, sistemas de disipación de calor para

dispositivos electrónicos, nuevas tintas electroconductoras, etc. [Shahzad et al., 2016].

Biomedicina: Los nanomateriales y la nanotecnología están llamados a

transformar la medicina por su potencial aplicación en el diagnóstico precoz y

tratamiento de una gran variedad de enfermedades. Las investigaciones realizadas con

nanomateriales a lo largo de los últimos años han dado lugar a una nueva área dentro

de las ciencias biomédicas: la nanomedicina.

La nanomedicina abarca el diagnóstico temprano de enfermedades, la

liberación controlada de fármacos y la medicina regenerativa mediante el uso de

nanomateriales o técnicas nanotecnológicas [Tibbals, 2013]. Hasta el momento, los

avances más significativos en esta área se basan en el desarrollo de sistemas


40

nanométricos para la liberación controlada de fármacos (liposomas y nanocápsulas

poliméricas), el uso de nanopartículas y puntos cuánticos como sistemas de marcaje

para ensayos biomédicos y de imagen de tejidos, el empleo de nanopartículas de plata

como agentes antibactericidas, etc.

Aplicaciones analíticas: Durante los últimos años los nanomateriales han sido

ampliamente utilizados para el análisis clínico, fundamentalmente en el desarrollo de

nuevos sistemas de detección de biomarcadores para enfermedades. Los avances más

importantes en esta área se centran en el diseño de nuevos sistemas de detección

visual o óptica basados en cromatografía lateral de flujo y tecnologías automatizadas

de análisis clínico ultrasensible, mediante el uso de nanopartículas metálicas y puntos

cuánticos como elementos de marcaje [Selvan et al., 2016].

Por otra parte, se ha demostrado que los nanomateriales electroconductores

mejoran el rendimiento de los sistemas sensores y biosensores electroquímicos. En

este sentido, su empleo como elementos de transducción genera nuevas interfaces de

mayor área electroquímicamente activa, permitiendo detecciones más sensibles.

Muchos nanomateriales poseen asimismo propiedades electrocatalíticas, lo cual

favorece el diseño de sistemas sensores y biosensores más selectivos y menos

propensos a la acción de sustancias interferentes.

Los nanomateriales ofrecen además numerosas oportunidades en la

amplificación de señales, tanto en sistemas con detección óptica [Swierczewska, 2012]

como electroquímica [Zhu et al., 2015; Hasanzadeh, 2015].

5.1.5. Limitaciones de los nanomateriales

A pesar de las grandes ventajas que presentan, algunos nanomateriales

muestran una serie de limitaciones las cuales limitan su uso en determinadas

aplicaciones. Desde un punto de vista biomédico y analítico, las principales

limitaciones de estos materiales son la falta de estabilidad en fluidos biológicos, alta

toxicidad, baja biocompatibilidad, corta vida media en la circulación por el torrente


41

sanguíneo, tendencia a la agregación, baja especificidad, etc [Lehner et al., 2013].

Además, en la actualidad no se conocen los efectos a largo plazo que este tipo de

materiales pueden tener sobre la salud humana debido a su reciente empleo [Yan et

al., 2014].

Estas desventajas pueden ser corregidas mediante la funcionalización

superficial con ligandos específicos, creando nanomateriales funcionalizados, o

mediante la unión o combinación con otros materiales y nanomateriales,

obteniéndose una nueva generación de nanomateriales híbridos.

En la investigación que nos concierne nos centramos en el diseño, preparación

y caracterización de diversos nanomateriales funcionalizados y nanohíbridos para su

uso en la construcción de biosensores electroquímicos y sistemas inteligentes de

liberación controlada de fármacos.

Biosensores Electroquímicos 5. INTRODUCCIÓN

42

5.2. BIOSENSORES ELECTROQUÍMICOS

5.2.1. Conceptos fundamentales

Un sensor químico es un dispositivo que transforma la información química,

que comprende tanto la concentración de un componente específico de la muestra

como el análisis total de su composición, en una señal analíticamente útil. Los sensores

químicos contienen por lo general dos componentes básicos conectados en serie: un

sistema químico de reconocimiento (receptor) y un transductor físico-químico. Los

biosensores son sensores químicos en los que el sistema de reconocimiento utiliza un

mecanismo bioquímico [Thévenot, 2001].

Durante los últimos años, el desarrollo de biosensores ha sido uno de los

campos de más rápida evolución en Química por su gran impacto en análisis

biomédicos, químicos e industriales, así como en el control del medio ambiente. Hoy

en día, los biosensores representan un mercado en rápida expansión, donde el mayor

impulso procede de la industria sanitaria. En el mercado global se espera que a finales

del 2016 alcance la cifra de 16 millones de dólares y llegue hasta los 21.5 millones en el

2020 [MarketsandMarkets, 2015].

Desde un punto de vista analítico, la tecnología de los biosensores permite el

diseño de dispositivos de detección miniaturizados y portátiles, los cuales presentan

una alta sensibilidad y un bajo límite de detección, combinado con la especificidad

característica proporcionada por los sistemas de reconocimiento biológico. Debido a

estas propiedades, se espera en un futuro cercano el aumento en el uso de

biosensores como alternativas prometedoras a los instrumentos analíticos

tradicionales.

En el pasado, los biosensores se definieron como cualquier dispositivo que

utiliza reacciones bioquímicas específicas para detectar compuestos químicos en

muestras biológicas. Los primeros biosensores, construidos por Clark, Lyons, Updike y


43

Hicks en los años 60, estaban basados en la inmovilización de enzimas sobre los

electrodos de pH u oxígeno con la detección amperométrica o potenciométrica.

Según IUPAC, un biosensor se define como un dispositivo receptor-transductor

integrado en sí mismo, que es capaz de proporcionar información analítica cuantitativa

o semi-cuantitativa selectiva usando un elemento de reconocimiento biológico

[Thévenot, 1999].

Figura 5.4. Esquema del funcionamiento de un biosensor.

La Figura 5.4 representa los elementos empleados generalmente en el diseño

de un dispositivo biosensor. Como se muestra, en un biosensor, el biorreceptor es una

biomolécula que reconoce selectivamente el analito diana, mientras que el transductor

convierte la señal de reconocimiento en una señal medible. La singularidad de un

biosensor es que los dos componentes están integrados en un único componente, a

través de la adsorción o la inmovilización del componente biológico sobre el

transductor. Esta combinación permite la transducción directa del proceso de

reconocimiento bioquímico en una señal eléctrica, que debe ser posteriormente

amplificada, procesada y correlacionada con la concentración del analito diana en las

muestras.

Con el fin de construir un biosensor exitoso, deben ser consideradas una serie

de condiciones [Pingarrón et al., 1999; Grieshaber et al., 2008]:

Enzima

Anticuerpos

Receptor

Organelo

Bacteria

Célula

Sustancia electroactiva

Cambio masa

REM

Sonido

Cambios entálpicos

Señal electrica

Electrodo

QCM

Detector fotométrico

Detector acústico

Detector termométrico

SEÑAL ELÉCTRICA

BIORRECEPTOR TRANSDUCTORANALITO


44

El elemento de reconocimiento biológico debe ser altamente específico para

los fines del análisis, estable bajo condiciones normales de almacenamiento y

mostrar una baja variación entre ensayos.

La reacción de biorreconocimiento debe ser tan independiente como sea

posible de parámetros físicos tales como la agitación, el pH y la temperatura. Lo

cual permitirá el análisis de las muestras con un pre-tratamiento mínimo.

La respuesta debe ser exacta, precisa, reproducible y lineal en todo el rango de

concentraciones de interés, sin dilución o concentración. También debe estar

libre de ruido inducido por el transductor eléctrico u otro.

Si el biosensor se va a utilizar para la monitorización invasiva en situaciones

clínicas, la sonda debe ser pequeña y biocompatible, sin efectos tóxicos o

antigénicos. Además, el biosensor no debe ser sensible a la inactivación o la

proteólisis.

Para mediciones rápidas de analitos a partir de muestras humanas, es deseable

que el biosensor pueda proporcionar análisis en tiempo real.

El biosensor completo debe ser barato, pequeño, portátil y capaz de ser

utilizado por operadores semicualificados.

5.2.2. Clasificación de los biosensores

Los biosensores se pueden clasificar de acuerdo con i) el modo de transducción

de la señal; ii) la especificidad conferida por el elemento biológico; o alternativamente,

iii) una combinación de ambos.

5.2.2.1. Clasificación de acuerdo con el transductor

Según el modo de transducción de la señal, los biosensores se pueden clasificar

principalmente como:

Óptico (con la posibilidad de utilizar muchos tipos diferentes de

espectroscopias como método de detección: UV-Vis, absorción en el IR


45

cercano, fluorescencia, fosforescencia, Raman, dispersión Raman de superficie

amplificada, espectroscopia de interferencia por reflexión, resonancia de

plasmón superficial, refracción, espectrometría de dispersión)

Electroquímico (biosensores amperométricos, potenciométricos,

conductimétricos e impedimétricos)

Piezoeléctrico (biosensores utilizando la microbalanza de cristal de cuarzo

sensible a la masa, de onda acústica superficial con polarización horizontal o

métodos de detección basados en microcantilever.

Termométrica (colorimétrico)

5.2.2.2. Clasificación de acuerdo con el elemento de reconocimiento biológico

- Biosensores basados en elementos de reconocimiento biocatalíticos:

En este caso, el biosensor se basa en una reacción catalizada por

macromoléculas, que están presentes en su entorno biológico original, se han aislado

previamente o que han sido fabricadas. En consecuencia, se consigue un consumo

continuo de sustrato llevado a cabo por el biocatalizador inmovilizado en el sensor y se

obtienen respuestas transitorias o de estado estacionario supervisadas por el detector

integrado.

Se utilizan habitualmente cuatro tipos de biocatalizador:

Las células enteras (microorganismos, tales como bacterias, hongos, células

eucariotas o levaduras), orgánulos celulares o partículas (mitocondrias, paredes

celulares).

Tejido (lámina de tejido animal o vegetal).

ARN catalítico (ribozimas) y anticuerpos (abzimas).

Enzima (mono o multi-enzima).

Las enzimas son los receptores más antiguos y todavía más utilizados en el

diseño de biosensores debido a su alta actividad biocatalítica y especificidad. Las

enzimas son proteínas globulares compuestas principalmente de los 20 aminoácidos


46

de origen natural que catalizan reacciones bioquímicas (Figura 5.5). Estas pueden

aumentar significativamente la velocidad de una reacción en comparación con una

reacción no catalizada. Algunas enzimas, también conocidas como enzimas redox,

catalizan reacciones que producen o consumen electrones. Así, el sustrato es

reconocido por un centro de unión de la enzima, similarmente a la reacción de

interacción antígeno-anticuerpo. En los biosensores enzimáticos donde se detectan los

electrones se utilizan este tipo de enzimas con centros de enlace específicos.

Posteriormente, mediante la densidad de corriente obtenida a través de la interfaz

enzima/sustrato se puede calcular la concentración y/o actividad.

Figura 5.5. Representación esquemática de una reacción enzimática catalizada.

Debido a sus estructuras moleculares complejas, la disposición de los

aminoácidos en el sitio activo de la enzima (a menudo localizado en el centro de la

proteína) induce a que solo un sustrato específico se pueda unir haciendo a la

biomolécula muy selectivo para un tipo de molécula de sustrato, mostrando una

selectividad exquisita por esta, lo que permite que se pueda detectar de forma muy

selectiva sustancias individuales en mezclas complejas, como la orina o la sangre. Esto

elimina la necesidad de realizar etapas que consumen tiempo y necesiten mano de

Sustratos

Productos

EnzimaSitio activo

Complejoenzima-sustrato


47

obra especializada, tales como las de pretratamiento de la muestra y de separación de

interferencias, normalmente necesarias en este tipo de muestras complejas.

[Copeland, 2000].

Las interacciones enzima-sustrato pueden ser caracterizadas por estudios

cinéticos, siendo el modelo de Michaelis-Menten uno de los enfoques más simple, más

conocido y ampliamente utilizado para este tipo de determinaciones cinéticas. Este

modelo cinético se puede aplicar cuando la concentración del sustrato es mayor que la

concentración de enzima, y suponiendo que la concentración del complejo enzima-

sustrato no cambia en el tiempo de reacción (hipótesis de estado estacionario).

Suponiendo que la concentración total de enzima no cambia con el tiempo, la

velocidad de reacción inicial de la formación de productos en una reacción enzimática

unimolecular está dada por:

Donde KM es la constante de Michaelis-Menten, [S] es la concentración de

sustrato, y V0 y Vmax son las velocidades de reacción iniciales y máxima,

respectivamente. El valor KM se corresponde con la concentración de sustrato a la que

la velocidad de reacción es la mitad con respecto a la máxima, y es una medida inversa

de la afinidad de la enzima por el sustrato.

- Biosensores basados en elementos de reconocimiento biocomplejante o de

bioafinidad

En este caso, la operación del biosensor se basa en la interacción del analito

con macromoléculas o ensamblajes moleculares organizados que, o bien han sido

aislados de su entorno biológico original o fabricados mediante ingeniería. Por lo


48

tanto, generalmente se alcanza el equilibrio y no hay consumo neto del analito por el

agente biocomplejante inmovilizado. Estas respuestas de equilibrio son controladas

por el detector integrado. En algunos casos, esta reacción biocomplejante es

monitorizada utilizando una reacción biocatalítica complementaria. Las señales en el

estado estacionario o transitorio son monitorizadas por el detector integrado. Los

mecanismos de bioafinidad más comúnmente utilizados en la construcción de

biosensores son los siguientes:

Interacción antígeno-anticuerpo (Inmunosensores)

Reconocimiento basado en ácidos nucleicos (Genosensores y aptasensors)

Receptor / antagonista / agonista, en el que los canales de iones, receptores de

membrana o proteínas de unión se utilizan como sistemas de reconocimiento

molecular

Receptores artificiales

5.2.2.3. Biosensores de carácter combinado

La tercera clasificación de los biosensores se basa en la combinación de dos de

los conceptos anteriormente mencionados. Ejemplos de esto son biosensor enzimático

amperométrico, inmunosensores ópticos, etc.

- Biosensor amperométrico:

Hoy en día, los transductores electroquímicos son los más ampliamente

utilizados en el diseño de biosensores debido a las importantes ventajas que

presentan. Estos dispositivos suelen tener diseños relativamente simples y no

requieren instrumentación costosa. Por lo tanto, son dispositivos de fácil manejo,

compactos y de bajo coste.

Otras ventajas inherentes de biosensores electroquímicos son su robustez, fácil

miniaturización, excelentes límites de detección, también con volúmenes de analitos


49

pequeños, y la capacidad para ser utilizado en biofluidos turbios con compuestos

ópticamente absorbentes y fluorescentes [Ronkainen et al., 2010; Grieshaber et al.,

2008].

Los transductores electroquímicos se organizan generalmente en tres

categorías principales según el tipo de medición empleado: corriente, potencial e

impedancia. Entre ellas, las técnicas más comúnmente empleadas en biosensores son

las que miden la corriente (métodos amperométricos y voltamperométricos).

En amperometría, cambios en la corriente generada por la oxidación o la

reducción electroquímica de los analitos diana se controlan directamente con el

tiempo, mientras que se mantiene un potencial constante en el electrodo de trabajo

con respecto a un electrodo de referencia. La corriente es proporcional a la

concentración de la especie electroactiva en la muestra, lo que permite su

cuantificación fácil.

La detección amperométrica se ha utilizado exhaustivamente con biosensores

de afinidad biocatalíticos debido a su simplicidad y bajo límite de detección.

Ventajosamente, el potencial fijo durante la detección amperométrica resulta en una

insignificante corriente de carga (la corriente necesaria para aplicar el potencial al

sistema), lo que minimiza la señal de fondo que afecta negativamente al límite de

detección. Además, los biosensores amperométricos tienen una selectividad adicional

ya que el potencial de oxidación o reducción utilizado es característico para la

detección de los distintos analitos [Eggins, 2002].

Cuando se preparan biosensores enzimáticos amperométricos es imporante

determinar la sensibilidad de la enzima a las condiciones experimentales tales como el

pH, la temperatura, y la agitación. Deben ser también considerados otros parámetros

tales como el origen y la disponibilidad de la enzima, su estabilidad operacional y de

almacenamiento, así como el procedimiento de inmovilización y las características del

soporte (superficie del electrodo).


50

5.2.3. Nanomateriales utilizados en biosensores

En los últimos años, se han realizado grandes progresos en el área de los

biosensores electroquímicos debido al avance en la nano-invetigación y las excelentes

características que los nanomateriales les aporta [Wei et al., 2013]. De esta forma, los

materiales de tamaño nanométrico se han empleado extensamente en la modificación

de electrodos para la construcción de biosensores a lo largo de los últimos años [Wang

2005a, b].

En la Figura 5.6. se representan las partes básicas que constituyen un biosensor

electroquímico nanoestructurado.

Figura 5.6. Esquema básico de un biosensor electroquímico nanoestructurado.

El empleo en biosensores electroquímicos se debe a su amplia variedad de

propiedades estructurales y funcionales. Entre ellas, se destaca su alta relación

superficie-volumen, ya que aumenta propiedades como la capacidad de adsorción de

receptores químicos, la actividad electrocatalítica, o la biocompatibilidad mejorando

notablemente su sensibilidad.

Por otro lado, la utilización de nanomateriales con excelentes propiedades

electroconductoras, como las nanopartículas metálicas y los nanomateriales de

carbono, proporciona un aumento de la superficie activa de los electrodos modificados

mejorando la velocidad y eficiencia de la transferencia electrónica. Además, estos

nanomateriales pueden disminuir la distancia entre las proteínas y la superficie

Electrodo

Receptores biológicos

Nanomateriales


51

electródica, permitiendo una transferencia directa de electrones al centro activo de las

biomoléculas por mecanismos de tunelización [Yáñez-Sedeño et al., 2015].

El papel fundamental de los nanomateriales en los biosensores electroquímicos

se basa en su empleo como elementos: i) de transducción o plataforma para la

inmovilización de biomoléculas; ii) de amplificación de la señal electroquímica o iii) de

etiquetado para la señalización [Hayat et al., 2014].

Esta Tesis Doctoral ha centrado la atención en el desarrollado de nuevas

estrategias para la preparación de superficies electródicas nanoestructuradas como

elementos de transducción para la construcción de nuevos biosensores

electroquímicos con un rendimiento bioanalítico mejorado y de mayor robustez.

En general, el desarrollo de metodologías para la nanoestructuración de las

superficies de los electrodos con tal propósito debe considerar los siguientes puntos: i)

el tipo de superficie a modificar; ii) el nanomaterial que se utiliza; iii) otros compuestos

o materiales que podrían ser también empleados; iv) el método físico o químico para la

modificación de la superficie, v) la biomolécula a inmovilizar y el método para hacerlo;

y vi) la reacción electroquímica que tiene lugar en la superficie del electrodo

funcionalizado [Yañez-Sedeño et al., 2015].

Estas estrategias deben proporcionar interfaces eléctricas con nano- o micro-

topología tridimensional que permita con éxito la inmovilización estable de las

biomoléculas sin afectar a su función biológica, pero también favoreciendo la aparición

rápida y eficiente de los procesos electroquímicos que participan en la reacción

analítica sobre tales interfaces [Wang 2005 a, b].

Los nanomateriales más empleados en la nanoestructuración electródica son

principalmente: las nanopartículas metálicas, los puntos cuánticos, nanotubos de

carbono, grafeno, nanofibras de carbono y nanopartículas de óxido de metal [Zhang et

al., 2006; Liu et al., 2007; Agüí et al., 2008; Shao et al., 2010; Lu et al., 2008; Vamvakaki

et al., 2006; Pingarrón et al., 2008; Luo et al., 2006; Zhang et al., 2007].


52

5.2.3.1. Nanopartículas metálicas:

Hay varios factores que justifican el interés de la investigación en

nanopartículas metálicas para la tecnología de biosensores, tales como:

Las nanopartículas metálicas, de manera similar a otros nanomateriales,

tienen una elevada superficie en relación al volumen. Esta característica

domina las propiedades ópticas, conductoras y magnéticas únicas que

poseen las nanopartículas con composición química adecuada.

El área superficial de las nanopartículas metálicas permite una gran

inmovilización de biomolécula sobre los electrodos modificados con ellas.

En general, la superficie de las nanopartículas de metal se puede modificar

fácilmente mediante métodos químicos o físicos, para favorecer la

inmovilización con éxito de biomoléculas de análisis a través de diferentes

mecanismos covalentes y no covalentes.

En comparación con otros nanomateriales duros y blandos, las

nanopartículas de metal se pueden preparar fácilmente por métodos

hidrotérmicos o electroquímicas convencionales, utilizando materias primas

y equipos relativamente económicos.

Las nanopartículas de metal se pueden diseñar racionalmente y con gran

versatilidad (con respecto a su geometría, tamaño y distribución) mediante

el control de las condiciones experimentales usadas para su preparación.

Las nanopartículas metálicas pueden favorecer la transferencia directa de

electrones entre el centro redox en el sitio activo de las enzimas y el

material del electrodo, a través de mecanismos de efecto túnel.

Varias nanopartículas metálicas pueden catalizar la conversión de

compuestos con interés analítico importante (como NADH, H2O2 y O2) en la

superficie del electrodo, reduciendo el potencial necesario para tales

transformaciones.

Las nanopartículas puede actuar como unidades de montaje

supramoleculares con propiedades funcionales avanzadas para la


53

construcción de una variedad de arquitecturas en la superficie del

electrodo.

Cuando se emplean las nanopartículas para modificar la superficie del

electrodo, pueden estar dispuestas en una variedad de arquitecturas tales como una

distribución aleatoria total, una monocapa autoensamblada o un conjunto de capa por

capa, así como aparecer incorporadas en la geometría del electrodo mediante su

mezcla con otros componentes de la matriz formando un electrodo compuesto (Figura

5.7).

Figura 5.7. Diferentes arquitecturas de nanopartículas metálicas empleadas en la

modificación de las superficies electródicas. [Pumera, 2014]

Esta variedad de arquitecturas basadas en nanopartículas metálicas pueden

estar dispuestos en las superficies de los electrodos por electrodeposición, adsorción

física, interacciones electrostáticas, enlaces covalentes o asociaciones

supramoleculares [Wu et al., 2005; Lim et al., 2005; Yang et al., 2006; Holzinger et al.,

2009; Manso et al., 2008; Yang et al., 2006]. Las interacciones específicas que

participan en tales modificaciones son impulsadas por la naturaleza de los grupos

químicos y la carga en la superficie de la nanopartícula. Estas características pueden

ser fácilmente manipuladas por adsorción selectiva de compuestos cargados en la

Monocapa autoensamblada

Distribución aleatoria Arquitectura capa sobre capa

Incorporación en la matriz electrodo


54

superficie de las nanopartículas, así como por transformación química con ligandos

seleccionados.

Durante los últimos años, se han descrito un gran número y variedad de

biosensores electroquímicos, inmunosensores y genosensores utilizando electrodos

modificados con nanopartículas [Rezaei et al, 2016; Zhao et al., 2016; Siangproh et al.,

2011; Ju et al., 2011; Pingarrón et al., 2008; Li et al., 2010].

Sin embargo, los biosensores nanoestructurados modificados solamente con

nanopartículas funcionalizadas sobre la base de los electrodos no son el ejemplo más

común en la bibliografía científica. A este respecto, hay un gran número de

publicaciones que describen la combinación de nanopartículas con otros materiales

nanométricos y/o macromoleculares para la construcción de biosensores

electroquímicos híbridos, como se ilustra en la Figura 5.8.

Figura 5.8. Modificación de la superficie del electrodo con nanopartículas metálicas,

solas y combinados con otros materiales [Pumera, 2014].

Estos biosensores electroquímicos híbridos se basan en la mejora de las

propiedades gracias a la combinación entre varios nanomateriales, sin que dicho

Solo AuNPs Otros nanomateriales

Polímeros o materiales sol-gelOtros nanomateriales y

polímeros o materiales sol-gel


55

acoplamiento afecte a sus propiedades individuales. Durante los últimos años, se han

diseñado una gran cantidad de superficies electródicas por combinación de las AuNPs

con grafeno [Azzouzi et al., 2015; Abraham et al., 2015; Wang et al., 2012a; Song et al.,

2011; Chen et al., 2011], CNTs [Eguílaz et al., 2015; Villalonga et al., 2012],

dendrímeros [Liu et al., 2005b], polímeros electroconductores [German et al., 2015],

óxidos metálicos [Zhao et al., 2015; Boujakhrout et al., 2015], polisacararidos

[Villalonga et al., 2007], etc.

Una de las combinaciones más utilizadas es la formada por las AuNPs, los CNTs

y/o grafeno. Estos biosensores utilizan las ventajas proporcionadas por la sinergia de

los nanomateriales, obteniéndose una transferencia directa de electrones entre los

nanomateriales funcionalizados y el centro redox de la biomolécula, sin la participación

de mediadores electroquímicos [Yu et al., 2014]. En este sentido, la transferencia

electrónica directa entre los receptores biológicos y la superficie del electrodo

posibilita la construcción de biosensores de tercera generación, libres de mediadores

electroquímicos.

Un ejemplo interesante de este diseño es el desarrollado recientemente en

nuestro grupo de investigación. El biosensor emplea un nanohíbrido formado por

nanocintas de Ag: bipiridina decoradas con AuNPs para la inmovilización de la enzima

peroxidasa de rabano picante (HRP). Gracias a este diseño, se detectan

concentraciones de H2O2 a nivel picomolar [Boujakhrout et al., 2016]. Por otro lado,

también se han empleado otras nanopartículas metálicas en la preparación de

biosensores electroquímicos, aunque con menos frecuencia, las más destacadas son

las nanopartículas de Pt (PtNPs) y de Ag (AgNPs). La propiedad más importante de las

PtNPs es su gran capacidad catalítica, particularmente en la descomposición de H2O2 y

reducción de O2. El empleo de estas nanopartículas reduce drásticamente el potencial

requerido para oxidar el H2O2 (≥ 0.6 V vs Ag/AgCl) [Pumera, 2014].

Por lo que respecta a las AgNPs, estas muestran un gran interés debido a su

excelente actividad catalítica, propiedades antibacterianas, biocompatibilidad y baja

toxicidad. Gracias a su alta reactividad y la combinación con CNTs, se ha construido un


56

sensor de glucosa con excelentes propiedades sin la necesidad de enzimas [Baghayeri

et al., 2015].

5.2.3.2. Nanomateriales de carbono:

El empleo de los materiales de carbono como elementos de transducción en el

diseño de biosensores electroquímicos ha crecido rápidamente durante los últimos

años. Entre los nanomateriales de carbono más empleados destacan, con mucha

diferencia con respecto a otros, los nanotubos de carbono y el grafeno.

La utilización de estos nanomateriales como andamios 3D para la construcción

de biosensores electroquímicos se basa en sus excelentes propiedades intrínsecas,

tales como [Lawal et al., 2016]:

Alta relación superficie/volumen

Alta estabilidad química

Fuerte resistencia mecánica

Biocompatibilidad

Excelente conductividad eléctrica y térmica

Baja corriente residual superficial

Amplia ventana de potencial

Bajo coste

Facilidad para la producción

- Nanotubos de carbono:

Los nanotubos de carbono son cilindros compuestos de anillos hexagonales de

carbono con hibridación sp2. Los nanotubos conformados por una lámina de grafeno

enrollada son los nanotubos monocapa, o SWNTs (Single-Walled Nanotubes), mientras

que existen también nanotubos donde unas láminas se incluyen dentro de otras


57

formando tubos concéntricos, conocidos como nanotubos multicapa o MWNTs (Multi-

walled Nanotubes).

Desde su descubrimiento en 1991, las propiedades de los nanotubos de

carbono han sido ampliamente estudiadas. Las investigaciones han demostrado que

estos nanomateriales mejoran la reactividad electroquímica de las biomoléculas y son

capaces de promover la transferencia electrónica directa entre los centros redox de las

macromoléculas y la superficie del electrodo (Figura 5.9). Además, las superficies

nanoestructuradas con CNTs son más resistentes al ensuciamiento, alargando la vida

útil de los biosensores [Lawal et al., 2016].

Figura 5.9. Transferencia directa de electrones (e-) entre en centro redox de una

enzima y la superficie del electrodo a través de CNTs.

Sin embargo, los CNTs poseen una limitación importante, su escasa solubilidad

en disolventes acuosos y orgánicos. Para solucionar dicho problema es imprescindible

por tanto su funcionalización física, química o combinada entre ambas [Singh et al.,

2009]. La funcionalización de los CNTs con grupos como aminas o carboxilos mejoran

e- e-


58

su biocompatibilidad para la unión de los elementos de reconocimiento biológico, y

también aumentan la tasa de electrones a transferir [Kumar et al., 2015].

Existen multitud de biosensores electroquímicos en los que se emplean SWCNT

y MWCNT para la modificación electródica [Eguílaz et al., 2016 a,b; Kim, 2017]. No

obstante, como en el caso de las nanopartículas metálicas, la mayoría de los trabajos

encontrados en la bibliografía utilizan nanohíbridos, conjugando los CNTs con otros

nanomateriales como polímeros [Hernández-Ibáñez et al., 2016], grafeno

[Devasenathipathy et al., 2015], óxidos metálicos [Tu et al., 2015], dendrímeros [Zhang

et al., 2016], etc.

Un ejemplo original de la utilización de MWCNTs en biosensores

electroquímicos fue desarrollado por el grupo del Prof. Wang [Jia et al., 2013]. En ese

trabajo, se describe el primer biosensor para la detección de lactato, no invasiva y en

tiempo real, en la transpiración humana durante un ejercicio físico. El biosensor

consiste en un tatuaje temporal flexible para adaptarlo a la piel de los usuarios. La

superficie del electrodo epidérmico está modificada con tetratiofulvalelo y MWCNTs,

para la inmovilización de la enzima Lactato Oxidasa (LOx), y cubierto por un polímero

de quitosano biocompatible.

- Grafeno:

El grafeno es un nanomaterial de dos dimensiones, ya que está formado por

anillos hexagonales de átomos de carbono con hibridación sp2, dispuestos en una

monocapa de un átomo de espesor.

Las propiedades físico-químicas, como en el caso de los CNTs, son excelentes.

Entre las propiedades electroquímicas, que hacen tan interesante su uso en la

modificación de superficies electródicas, se incluyen una baja resistencia a la

transferencia de carga, una gran actividad electrocatalítica y una amplia ventana de

potencial de trabajo [Gao et al., 2015].


59

El grafeno, al igual que los CNTs, promueve la transferencia electrónica directa

entre las biomoléculas y la superficie del electrodo, creando biosensores

electroquímicos de tercera generación [Wang et al., 2012 b; Liang et al., 2015].

Sin embargo, su aplicación en esta área está limitada debido a su baja

solubilidad y reactividad química. La solución a estas desventajas es la funcionalización

o, directamente, el empleo del óxido de grafeno (GO). El GO se obtiene por oxidación

del grafito con un agente oxidante y una mezcla de ácidos fuertes. Este proceso de

oxidación da lugar a la aparición de grupos funcionales en el plano basal y los bordes,

tales como grupos hidroxilos, carbonilos, carboxilos y epóxidos [Lee et al., 2016].

Durante los últimos años, el empleo de grafeno como elemento de

transducción en biosensores ha crecido exponencialmente. La mayoría de los

biosensores emplean el uso de nanohíbridos para mejorar la biocompatibilidad en la

inmovilización de las biomoléculas, con nanopartículas metálicas [Huang et al., 2015],

CNTs [Devasenathipathy et al., 2015], óxidos metálicos [Tiwari et al., 2015], polímeros

[Navakul et al., 2016], etc.

5.2.3.3. Nanomateriales poliméricos:

Los nanomateriales poliméricos, naturales y sintéticos, han sido ampliamente

utilizados en la nanoestructuración de superficies electródicas. Entre éstos, los

dendrímeros han demostrado ser una buena herramienta para la construcción de

biosensores electroquímicos novedosos [Astruc et al., 2010].

Los dendrímeros son polímeros sintéticos monodispersos con una estructura

tridimensional regular y altamente ramificada [Vögtle et al., 2009]. Existen 2 métodos

para la síntesis de dendrímeros (Figura 5.10). En ambos métodos, cada acoplamiento

de la unidad dendrimérica da lugar a una nueva generación del material con el

consiguiente aumento de los grupos terminales en la periferia, lo que posibilita el

diseño hecho a medida de estos nanomateriales [Vögtle et al., 2009].


60

Figura 5.10. Parámetros estructurales de una molécula de dendrímero y sus métodos

de síntesis A) divergentes y B) convergentes.

Hay numerosos factores que justifican el interés de la investigación en

dendrímeros para la construcción de biosensores electroquímicos, tales como [Vögtle

et al., 2009]:

Su estructura periférica posee una alta densidad de grupos funcionales, los

cuales pueden ser empleados para la inmovilización de biomoléculas a través

de uniones covalentes o no covalentes, o para la unión de los dendrímeros a la

superficie electródica.

Los grupos hidrofílicos externos mejoran la biocompatibilidad de los

dendrímeros posibilitando la inmovilización de los receptores biológicos sin

alterar su conformación.

Su estructura es permeable, favoreciendo la difusión de las especies

electroactivas a la superficie del electrodo.

Son fácilmente modificados o dopados con mediadores electroquímicos o

nanopartículas metálicas, ya sea por unión a sus grupos funcionales externos o

por encapsulamiento en la estructura, para mejorar la actividad

electrocatalítica.

Gracias a estas propiedades, los dendrímeros ofrecen múltiples posibilidades

como andamios 3D funcionales en la nanoestructuración de superficies electródicas.

Las estrategías para llevarlo a cabo pueden incluir la formación de monocapas, capa

sobre capa, capas híbridas con otros polímeros y/o nanopartículas, etc.

Durante la última década, se han descrito un elevado número de biosensores

electroquímicos utilizando electrodos modificados con dendrímeros. Los dendrímeros

Generación

0 1 2

A B


61

más empleados son los de poliamidoamina o PAMAM [Borisova et al., 2016; Miodek et

al., 2016].

5.2.3.4. Nanomateriales de óxidos metálicos:

Los nanomateriales de óxidos metálicos han sido utilizados en numerosos

diseños para la construcción de biosensores. La propiedad más importante que poseen

para este cometido es su alta actividad electrocatalítica [Yañez-Sedeño et al., 2015].

Las nanopartículas de óxidos metálicos, como TiO2, NiO, ZnO, SnO2 y Co3O4, se

emplean en la modificación de superficies electródicas por su alta actividad

electrocatalítica, buena estabilidad química, baja toxicidad, alta relación

superficie/volumen y buena biocompatibilidad.

En los últimos años, se han descrito multitud de biosensores electroquímicos

construidos con híbridos preparados entre nanopartículas de óxido metálico,

materiales de carbono y/o polímeros conductores [Devi et al., 2012; Yadav et al., 2011;

Li et al., 2009]. Estas superficies nanoestructuradas también permiten la transferencia

directa de electrones entre los centros redox de los receptores biológicos y los

electrodos [Zhu et al., 2015; Dong et al., 2014; Shamsipur et al., 2012; Li et al., 2006;

Liu et al., 2005a].

Otro nanomaterial metálico muy interesante para ser empleado como

elemento de transducción son las nanopartículas de Fe3O4. En este caso, la propiedad

más destacable es el magnetismo. Otras ventajas son su excelente solubilidad en agua,

gran área superficial, fácil fabricación, menor toxicidad que las nanopartículas

metálicas y bajo coste [Yu et al., 2013].

Un enfoque alternativo en la construcción de biosensores es el empleo de estas

nanopartículas de forma desechable por adsorción no específica a la superficie del

electrodo, de modo que mediante la utilización de un imán externo se puede renovar

la superficie. En este sentido, las nanopartículas magnéticas pueden también

emplearse para purificar y/o concentrar el analito de interés [Eguílaz et al., 2011].


62

Otros nanomateriales de este grupo, empleados para el diseño de plataformas

estables en biosensores, son las nanopartículas y películas finas de SiO2 porosa. Ambas

estructuras de SiO2, se caracterizan por su fácil síntesis y modificación superficial, alta

estabilidad química y térmica, y su gran área superficial biocompatible para la

inmovilización de los receptores biológicos. Existen numerosos trabajos que describen

el uso de sílice para la construcción de biosensores con buenas propiedades [Saadaoui

et al., 2016; Wang et al., 2009]. Sin embargo, estos nanomateriales son aislantes y

bloquean la transferencia electrónica. Generalmente, en los biosensores

electroquímicos se usan combinados con otros materiales que mejoran su

conductividad como los materiales de carbono o nanopartículas métalicas [Zhang et

al., 2016; Tiwari et al., 2008].

Sistemas de liberación controlada 5. INTRODUCCIÓN

63

5.3. SISTEMAS DE LIBERACIÓN CONTROLADA

5.3.1 Conceptos básicos

El papel principal de los sistemas de liberación controlada es el de administrar

la cantidad correcta del fármaco eficazmente, es decir controlar el lugar y el ritmo de

la liberación. El desarrollo de nuevos sistemas de liberación controlada de fármacos es

de vital importancia. Se prevé que en un futuro cercano estos sistemas sean una

alternativa útil para el tratamiento de enfermedades que requieren la administración

de fármacos altamente tóxicos, como el cáncer [Baezaet al., 2014].

Los primeros sistemas de liberación controlada datan de la década de 1950.

Estos sistemas estaban basados en polímeros sólidos donde se incorporaban fármacos

para productos agrícolas, pero no fue hasta la década siguiente cuando estos diseños

se extendieron a la medicina. En los primeros estudios, se utilizaban tubos de silicona o

matrices de polietileno [Lager, 1981]. Hasta los años 80, época denominada 1ª

generación, se establecieron los mecanismos básicos de la liberación de fármacos, los

cuales comprenden la disolución, difusión, ósmosis y el intercambio iónico, y se

desarrollaron sistemas de administración orales y transdérmicos.

Durante la 2ª generación (1980-2010), se diseñaron los polímeros e hidrogeles

"inteligentes" donde la liberación de fármacos estaba provocada por cambios en los

factores ambientales, tales como temperatura, pH o nivel de glucosa. En la última

década de esta generación, comenzó a utilizarse la nanotecnología [Park, 2014], sin

embargo, estos sistemas eran incapaces de superar las barreras biológicas. Con

respecto a la tercera generación, ya son posibles algunas de las aplicaciones mostradas

en la Tabla 5.1., pero se espera que gracias a la nanotecnología los sistemas de

administración de fármacos superen las barreras biológicas y físico-químicas con las

enormes ventajas que ese avance supondría para el empleo de estos sistemas en

estudios in vivo.


64

Tabla 5.1. Evolución de los sistemas controlados de administración de fármacos desde

1950 [Yun, 2015].

Hoy en día, la nanotecnología ha posibilitado el diseño de sistemas capaces de

atravesar algunas barreras biológicas, como la barrera hematoencefálica, y la unión a

tejidos enfermos. Asimismo, se ha comprobado cómo estos sistemas mejoran la

farmacocinética y la estabilidad de los agentes terapéuticos.

Los nanosistemas básicos para la liberación controlada de fármacos fabricados

consisten en: i) una plataforma soporte; ii) una carga útil, o fármaco; y iii)

opcionalmente, ligandos de orientación.

Sin embargo, en la actualidad, las investigaciones se centran en el diseño de

sistemas avanzados que permiten la liberación de los agentes terapéuticos de manera

programada, donde las plataformas soporte o nanovehículos incluyen una

1950 1980 20402010

1ª Generación 2ª Generación 3ª Generación

Control de la liberación básica Sistemas de liberación inteligente Sitemas de liberación modulada

Liberacion oral

Dos veces al día, una vez al día

Liberación de orden cero

Primer orden vs orden cero

Administración de fármacos poco solubles

Excipientes no tóxicos

Liberacion transdérmica

Una vez al día, una vez a la semana

Liberación de péptidos y polímeros

• Depósito a largo plazo utilizandopolímeros biodegradables

• Administración pulmonar

Liberación de péptidos y polímeros

• Durante tiempos > 6 meses

• Control de la cinética de liberación

• Administración no invasiva

Mecanismo de liberación de fármacos:

• Disolucion

• Difusión

• Osmosis

• Intercambio iónico

Polímeros e hidrogeles inteligentes

• Sensible a cambios ambienteles

• Liberación auto regulada

(Trabajo in vitro)

Polímeros inteligentes e hidrogeles

• Especificidad de la señal y sensibilidadcinética de la respuesta rápida

(Trabajo in vivo)

Nanopartículas

Liberación específica en tumor

Liberación de genes

Administración dirigida de fármacos

No tóxica para células no objetivo

Superación barrera hematoencefálica

El control exitoso de propiedades fisicoquímicas de sistemas de entrega

Incapacidad para superar barreras biológicas

Necesidad de superar barreras biológicas y fisicoquímicas


65

funcionalidad sensible a un estímulo, ya sea externo o interno, que permite una

liberación más eficaz, segura, estable, así como una mayor absorción celular [Lehner et

al., 2012].

Las características deseadas en los dispositivos “inteligentes” de administración

de fármacos son: i) mejora de la permeabilidad: para facilitar el paso de las drogas a

través de la membrana celular u otras bio-barreras; ii) aumento de la estabilidad del

fármaco: los nanomateriales pueden proteger a las drogas de su autodegradación o de

la limpieza de los macrófagos; iii) entrega localizada o específica: para liberar los

fármacos en regiones concretas, como en cánceres locales; iv) capacidad de control:

para que la liberación del fármaco se produzca mediante una señal externa o a través

de los cambios ambientales; y v) aumentar la eficacia de la terapia y reducir la

toxicidad de la droga: los nanomateriales pueden minimizar los efectos secundarios

de los fármacos y sin embargo, aumentar el terapéutico al ser liberado en un área

específica [Yan et al., 2014].

5.3.2. Clasificación de los sistemas de liberación controlada de fármacos

Los nanosistemas pueden clasificarse de acuerdo a numerosos parámetros. Los

más importantes son i) la orientación para la administración dirigida del fármaco; ii) el

mecanismo de liberación de los agentes terapéuticos; y iii) el tipo de plataforma.

5.3.2.1. De acuerdo a la orientación

La orientación para la administración dirigida de los agentes terapéuticos,

puede considerarse pasiva, activa o combinación de ambas.

La orientación pasiva se basa en fenómenos de permeabilidad y retención. Se

ha demostrado que en algunos tejidos tumorales las macromoléculas con

tamaños superiores a 50 KDa se retienen en el intersticio del tumor durante

periodos largos de tiempo. Además, en estos tejidos el drenaje linfático es


66

defectuoso haciendo que el líquido extracelular no se drene con normalidad, lo

que se traduce en mayores tiempos de acumulación. Por lo tanto, una

orientación pasiva exitosa depende de las características estructurales de las

plataformas utilizadas para la liberación de fármacos, tales como la superficie,

el tamaño y la forma [Baeza et al., 2014].

La orientación activa implica el uso de ligandos direccionales. Los nanosistemas

se funcionalizan con ligandos de reconocimiento específico hacia moléculas de

la superficie o receptores sobre-expresados en orgánulos, células, tejidos u

órganos enfermos. Los ligandos más empleados incluyen moléculas pequeñas,

anticuerpos, aptámeros, péptidos, proteínas y azúcares [Baeza et al., 2014].

5.3.2.2. De acuerdo al mecanismo de liberación del fármaco:

Los mecanismos básicos que controlan la liberación de los agentes terapéuticos

de las plataformas soporte, esquematizados en la Figura 5.11., son:

Difusión: la entrega del fármaco se realiza por gradientes de concentración.

Inflamación o hinchamiento controlado: empleado en sistemas porosos donde

la liberación se da por el incremento en el tamaño de los poros producidos por

el paso de líquido al interior del nanocontenedor.

Erosión: la entrega generalmente se da por combinación entre el transporte de

masa y una reacción química, la cual puede provocar la disolución del fármaco,

la degradación de la plataforma empleada, la creación de porosidad o efectos

autocatalitícos.

Estímulos: la liberación es provocada por un estímulo, ya sean de carácter

endógenos o exógenos. Dentro de este grupo podemos diferenciar entre

estímulos químicos (cambios de pH, moléculas seleccionadas o con actividad

redox), físicos (campos magnéticos, luz o temperatura) y bioquímicos (tales

como enzimas, anticuerpos, o ADN).


67

Figura 5.11. Mecanismos básicos de liberación de fármacos en los sistemas de

administración controlada por: A) difusión, B) hinchamiento controlado, C) erosión y D)

estímulos.

5.3.2.3. De acuerdo con el tipo de plataforma utilizada:

Podemos distinguir tres clases de materiales:

Liposomas

Materiales poliméricos

Nanomateriales

A pesar de esta clasificación básica, los 2 primeros podrían considerarse

también nanomateriales si tienen el tamaño adecuado.

En la Figura 5.12. se ilustran algunos de los nanomateriales básicos más

comúnmente empleados en la liberación controlada de fármacos.

concentración Entrada líquido

Reac. Química Incremento pH

A) DIFUSIÓN B) HINCHAMIENTO

C) EROSIÓN C) ESTÍMULO


68

Figura 5.12. Ilustración de los nanomateriales más usados como plataformas en los

sistemas de liberación de fármacos.

Para una terapia dirigida, se emplean plataformas soporte o nanovehículos

donde almacenar y transportar los fármacos. En su diseño es necesario controlar una

serie de parámetros como son [Lehner, 2013]:

i) Tamaño, para evitar su eliminación por vía hepática o renal, asegurando un

suficiente almacenaje del agente terapéutico;

ii) Biocompatibilidad, hasta obtener una buena relación beneficio/riesgo;

Liposomas

MSN

AuNPs

Dendrímeros Fullerenos

CNTs

QDots

NPs Poliméricas


69

iii) Propiedades enmascarantes, para evitar el reconocimiento inmunológico o

la interacción con proteínas séricas;

iv) Tiempo óptimo de circulación por el torrente sanguíneo;

v) Alta especificidad hacia un objetivo que provoque la liberación del fármaco;

vi) Mecanismos de liberación controlables;

vii) Que esos mecanismos respondan a un estímulo.

Los nanovehículos pueden ser clasificados en 3 grandes grupos:

- Liposomas:

Son vesículas con forma esférica, formadas por una doble membrana de

fosfolípidos. Su carácter anfifílico posibilita el encapsulamiento de moléculas hidrófilas

en su interior acuoso o, por el contrario, el transporte de compuestos hidrofóbicos

sobre su membrana lipídica.

Otras de las ventajas que aporta la membrana a estos portadores son su fácil

modificación y su alta biocompatibilidad, haciendo que los nanosistemas puedan

adherirse a las membranas celulares o introducirse en el interior de las células por

endocitosis [Faraji et al., 2009]. Además, gracias a la fácil modificación de estos

sistemas comentada anteriormente, ya están siendo empleados clínicamente como

sistemas inteligentes en la liberación de fármacos para el tratamiento del sarcoma o el

cáncer de ovarios entre otros, empleando diferentes funcionalidades que responden a

estímulos ambientales (como el pH, la temperatura, la luz o la respuesta enzimática)

[Lehner et al., 2013].

Durante los últimos años, se han descrito un gran número de nanosistemas

utilizando liposomas como nanocontenedores para el almacenamiento de fármacos.

En algunos ejemplos recientes, se emplean nanoliposomas termosensibles orientados


70

hacia receptores del factor de crecimiento epidérmico, sobreexpresado en células

cancerosas, para la liberación de un compuesto anti-cancerígeno [Haeri et al., 2016].

- Materiales poliméricos:

Su aplicación en la entrega controlada de agentes terapéuticos se basa en la

conjunción formada entre el polímero y el fármaco, denominado “profármaco

polimérico”, ya que mejora la solubilidad de fármacos hidrofóbicos y evita la liberación

inicial que se produce con otros materiales [Hu et al., 2010].

Los polímeros utilizados como nanovehículos para la entrega regulada de

fármacos son biodegradables y biocompatibles. Es decir, tras la liberación del fármaco

el polímero se degrada en moléculas inocuas como agua, hidrógeno o nitrógeno. La

liberación de la droga se modula en función del polímero empleado [Parveen et al.,

2008].

Dentro de este gran grupo de materiales, los más empleados en la

nanomedicina son los las nanopartículas o micelas de ácido poliglicólico (PGA), del

ácido poliláctico (PLA) y los dendrímeros. El crecimiento en empleo de dendrímeros

como nanosistemas se debe a la facilidad con la que se modifica o funcionaliza su

superficie, su cavidad interior hidrófoba para la encapsulación de los fármacos

hidrófobos o el control del grado de ramificación. Sin embargo, para ser empleados

clínicamente necesitan mejorar su biocompatibilidad, citotoxicidad y biodistribución

[Lehner et al., 2013; Cho et al., 2008].

Los dendrímeros más utilizados son los de poliamidoamina (PAMAM). En los

últimos años, la conjunción entre estos y otros ligandos, como el polietilenglicol (PEG)

ha sido el método más empleado para reducir la toxidad de los agentes terapéuticos

[Luong et al., 2016].


71

- Nanomateriales:

Durante la última década, la nanotecnología ha creado una amplia gama de

nanomateriales, los cuales se están aplicando en el campo de la nanomedicina. Como

materiales básicos más extendidos encontramos los CNTs, fullerenos, nanopartículas

metálicas, puntos cuánticos y nanopartículas de materiales inorgánicos.

Nanotubos de carbono:

El almacenaje de los fármacos se puede dar tanto en la cavidad interior, como

en la superficie externa, funcionalizando covalentemente la pared lateral o las puntas

con ligandos que permitan el transporte de moléculas y su liberación al ser sensibles a

las condiciones ambientales tales como temperatura, pH o agentes reductores.

Asimismo, esa modificación se utiliza para mejorar su biocompatibilidad y su

solubilidad disminuyendo la citotoxicidad, aunque sigue siendo un inconveniente a

mejorar en estos dispositivos [Lehner et al., 2013]. Se espera que una de las

aplicaciones más prometedoras en la liberación controlada de fármacos y genes a las

células, se regule mediante el empleo de SWCNTs funcionalizados con péptidos

[Razzazan et al., 2016; Ohta et al., 2016].

Otra de las formas alotrópica del carbono que está siendo investigada en los

últimos años, son los fullerenos, aunque en menor medida que los CNTs [Raza et al.,

2015].

Puntos cuánticos:

Estos materiales consisten en nanocristales de tamaño pequeño (1-10 nm) y

constan de un núcleo formado por un elemento semiconductor rodeado por una capa

metálica.

Los fármacos pueden ser inmovilizados en su superficie mediante enlaces

covalentes escindibles, de modo que los conjugados formados presentan un mayor


72

tamaño, lo que evita su eliminación renal aumentando el tiempo de circulación en el

plasma, y al romperse este enlace se libera la droga, quedando el nanomaterial con un

tamaño adecuado para ser excretados. Sin embargo, estos nanomateriales están

fabricados con metales pesados (los más utilizados son CdS, CdSe, ZnS…) tóxicos a

largo plazo, lo que limita su uso clínicamente [Qi et al., 2008].

Durante los últimos años se han desarrollado puntos cuánticos de carbono

(CQD), los cuales, a diferencia de los fabricados con semiconductores, poseen buena

biocompatibilidad, alta fotoestabilidad y baja toxicidad. Estas nuevas nanoplataformas

se están empleando para el diseño de nuevos sistemas de liberación de agentes

terapéuticos y genes [Zhou et al., 2016; Chiu et al., 2016].

Nanopartículas metálicas:

Las nanopartículas metálicas son uno de los nanomateriales más empleados en

la liberación de fármacos, debido a la posibilidad de controlar el tamaño y propiedades

magnéticas, la baja citotoxicidad y la fácil modificación superficial que presentan, lo

que posibilita la creación de sistemas con funcionalidades avanzadas, sensibles y con

capacidad de respuesta frente a cambios de pH, redox… [Zhang et al., 2016].

Los materiales más empleados para su uso como plataformas son de óxido de

hierro y el oro.

En el caso de las nanopartículas magnéticas, de Fe3O4 y Fe2O3 principalmente,

poseen una alta biocompatibilidad y presentan un comportamiento paramagnético

debido a su pequeño tamaño. Este hecho trae como consecuencia que no aporten un

campo magnético propio (por lo que no tienden a aglomerarse), pero que sí actúen

como nanomateriales magnéticos en presencia de un campo externo. En este sentido,

estas nanopartículas pueden ser transportadas hasta las células diana mediante la

aplicación de campos magnéticos y una vez allí liberar los fármacos [Lehner et al.,

2013]. Además, otra característica que presentan estos nanomateriales es la

posibilidad de hacer tratamientos de hipertermia. Éstos consisten en la aplicación de


73

un campo magnético externo que oscila con una elevada frecuencia, provocando que

también el campo propio de las nanopartículas oscile muy rápidamente generando un

calentamiento de la nanopartícula, capaz de matar (atacar) las células cercanas a las

nanopartículas [Zamora-Mora et al., 2017; Niemirowicz et al., 2016].

Las AuNPs, debido a sus excelentes propiedades ya comentadas en el capítulo

anterior, han atraído la atención en el campo de la nanomedicina. Su uso como

transportadores de fármacos se basa en su gran área superficial, fácil modificación

superficial y buena biocompatibilidad. Durante los últimos años, se han descrito un

gran número de nanosistemas para la entrega de agentes terapéuticos empleando

estos materiales [Manivasagana et al., 2016; Peng et al., 2016].

Estas plataformas basadas en nanomateriales metálicos poseen una

desventaja, la relación en peso existente entre la carga útil y la plataforma. Por este

motivo, suelen ir acompañadas de otros nanomateriales o polímeros [Zhang et al.,

2016].

Nanopartículas mesoporosas:

Una de las características más interesantes de estos materiales inorgánicos

para su uso como nanovehículo, es la posibilidad de crear estructuras con tamaños,

superficie y porosidad diferentes. Estas nanoestructuras porosas actúan como carcasa,

protegiendo la carga útil de la desnaturalización o degradación [Lehner et al., 2012].

Además, estos nanoportadores pueden ser fácilmente modificados por

diferentes funcionalidades que respondan a estímulos ambientales como el pH, la

temperatura, la luz o enzimas y activen la liberación de los fármacos. De esta forma es

posible controlar fácilmente esta liberación mediante estímulos propios del sistema o

inducirla mediante estímulos externos.

Entre todos los nanomateriales inorgánicos, las nanopartículas mesoporosas de

sílice, con mucho, ocupan el lugar más relevante en la entrega controlada de fármacos.


74

Otros nanomateriales empleados en menor medida, son las nanopartículas de alumina

y nanocápsulas de fosfato de calcio [Cheng et al., 2016].

5.3.3. Sistemas de liberación controlada basados en MSN

Las partículas de sílice mesoporosas incluyen una gran variedad de estructuras

en función de las condiciones de síntesis empleadas: MCM-41, MCM-48, SBA-15, etc.

Son ampliamente utilizadas como nanocontenedores debido a su alta superficie

específica (hasta 1200 m2/g), un elevado volumen de poro, y un tamaño de poro bien

definido y controlable (su diámetro puede variar de 2 a 10 nm), baja toxicidad, gran

capacidad de carga, buena estabilidad térmica, química y mecánica, así como por su

fácil funcionalización para la construcción de puertas moleculares o supramoleculares

que responden a estímulos. Estos sistemas, a diferencia de muchos de los expuestos

anteriormente, muestran una “entrega cero” en condiciones basales y solo en

presencia del estímulo se produce la liberación de la carga [Mamaeva et al., 2013;

Slowing et al., 2010].

En la Figura 5.13 se representa la estructura de dos de las nanopartículas de

sílice porosa más empleadas como nanovehículos para la liberación de fármacos.

Figura 5.13. Diagrama esquemático de los materiales MCM-41 (hexagonal) y MCM-48

(cúbica) en presencia de las micelas de surfactante en los poros.

MCM-41 MCM-48


75

La ruta y velocidad de difusión del fármaco depende de la estructura de la MSN.

MCM-41 y la MCM-48. La MCM-41 contiene canales de poros unidireccionales con

disposición hexagonal, lo que permite una liberación de la carga directa al exterior,

mientras que en el caso de la MCM-48, con estructura porosa cúbica tridimensional, el

recorrido para la liberación es mucho mayor [He et al., 2011]. Por este motivo,

además del resto de excelentes propiedades, la estructura MCM-41 fue elegida para el

desarrollo de diferentes trabajos en la entrega controlada de fármacos.

Desde su descubrimiento en 1992, su uso ha crecido exponencialmente [Beck

et al., 1992]. Típicamente se sintetizan mediante procesos sol-gel en presencia de un

tensioactivo, que controla el tamaño del poro. El método estándar se realiza

mezclando una baja concentración de surfactante catiónico, siendo el más utilizado el

bromuro de cetiltrimetilamonio (CTAB), con un precursor de silicato, generalmente

tetraetilortosilicato (TEOS), en medio básico a una temperatura de entre 30-80ºC

durante 2h. Las MSN se sintetizan en dos pasos, una hidrólisis y una condensación. Por

último, se elimina el surfactante del interior de los poros mediante una extracción con

una fase líquida o calcinando a alta temperatura el material sintetizado (Figura 5.15).

Figura 5.14. Ilustración esquemática de la síntesis de MCM 41. Mecanismo de

polimerización del TEOS. Adaptado de la referencia [Llinàs et al., 2014].

El modelo típico de un nanosistema de MSN para la liberación controlada de

fármacos consiste en cargar el agente terapéutico de forma reversible dentro de la

nanopartícula y funcionalizar su superficie, de forma covalente o por adsorción, con

Micelas CTAB

Alineaciónen matrices

hexagonales

Calcinación

Arreglo hexagonal con paredes de SiO2

MCM 41


76

diferentes clases de ligandos, tales como i) polímeros que mejoren su

biocompatibilidad, permeabilidad citoplasmática y/o solubilidad, ii) residuos que

aumenten su orientación, como anticuerpos o moléculas receptoras para

determinados tejidos o células, y iii) funcionalidades que respondan a un estímulo

específicamente para provocar la liberación del fármaco, denominadas puertas

moleculares “inteligentes” [Aznar et al., 2016].

El diseño de nanomateriales híbridos basados en las MSN funcionalizadas con

biomacromoléculas capaces de realizar distintas tareas como reconocimiento

(anticuerpos, péptidos, ADN/ARN) o actuar frente a estímulos químicos (enzimas,

proteínas, ligandos sensibles) nos permite obtener nanosistemas “inteligentes”

fácilmente aplicables a la liberación controlada de fármacos.

5.3.3.1. Puertas moleculares estímulo-respuesta:

Figura 5.15. Representación esquemática de una MSN funcionalizada con una puerta

molecular para la liberación controlada de fármacos mediante un estímulo

Para fabricar nanosistemas “inteligentes” capaces de liberar el fármaco de

forma controlada, se han diseñado diferentes “puertas” moleculares estímulo-

dependientes. Estos mecanismos de apertura mantienen la nanoplataforma cerrada,

Carga

Estímulo

Puertas moleculares


77

cubriendo los poros, y se accionan de forma selectiva en respuesta a un estímulo

determinado liberando la carga útil (Figura 5.15.).

Principalmente, se han desarrollado puertas moleculares sensibles a estímulos

con potenciales aplicaciones biomédicas. Es decir, cambios de pH, medios reductores,

actividad enzimática, luz, temperatura, moléculas y biomoléculas, campos magnéticos

y ultrasonidos, ya que estos estímulos se pueden encontrar en mayor concentración

en las proximidades de las células diana o son fácilmente aplicables de forma externa y

con terapias poco invasivas [Aznar et al.,2016].

Sensibles al pH:

El pH es el estímulo más utilizado en el desarrollo de nanosistemas para la

liberación controlada de agentes terapéuticos. El motivo principal se debe a que en los

tejidos inflamados o tumorales el medio es más ácido que en los tejidos sanos, y esta

variación de pH a nivel local va a provocar la liberación del fármaco directamente

sobre el tejido dañado [He et al., 2016].

El mecanismo de funcionamiento de estos sistemas, de un estado cerrado a

uno abierto, se basa en los cambios producidos tras la adición o sustracción de

protones. Esas modificaciones en la puerta pueden ser debidos a:

Una transformación en el tamaño o la forma

Interacciones de atracción o repulsión

Se han descrito numerosos sistemas que emplean este tipo de puertas

sensibles al pH, constituidos por una gran variedad de ligandos y funcionalidades para

el control de la liberación con macrociclos, aminas, polímeros, enzimas, ADN,

proteínas, nanopartículas metálicas nanopartículas inorgánicas, etc [AbouAitah et al.,

2016; Yilmaz, 2016; Muhammad et al., 2011; Lee et al., 2010; Liu et al., 2010].

Las aplicaciones de estas nanomáquinas en la nanomedicina presentan

limitaciones debido a que la vía de entrada habitual en las células es por endocitosis. El


78

pH del medio en un endosoma es de 5, lo que provocaría la liberación involuntaria del

fármaco antes de tiempo [Aznar et al.,2016].

Sensibles a cambios redox

El empleo de este estímulo para nanosistemas de liberación controlada es

debido al aumento en la concentración de especies con actividad redox que se observa

en el medio cercano a algunos tumores cancerosos [Aznar et al.,2016].

En este caso, el mecanismo de liberación se basa en:

Cambios en las interacciones supramoleculares de tipo redox entre ligandos

que funcionalizan la MSN y macrociclos como ciclodextrinas o curcubiturilos

que actúan como un pistón [Kim et al., 2010].

Ruptura de un enlace di-sulfuro, entre ligandos anclados en la MSN y agentes

de terminación. Los agentes de terminación, empleados generalmente para el

bloqueo del poro o para la orientación selectiva, pueden ser enzimas,

anticuerpos, nanopartículas metálicas o inorgánicas, polímeros, etc [Cui, 2012;

Xiao, 2016].

A diferencia de los nanosistemas sensibles al pH, la aplicabilidad de estos en la

nanomedicina es muy alta [Wang Y. et al., 2015].

Sensibles a la actividad enzimática:

Las enzimas proporcionan a los nanosistemas de liberación controlada una alta

selectividad. Actualmente, no es uno de los estímulos más utilizados, pero se prevé

que en un futuro cercano se invierta esa tendencia gracias a sus buenas características.

Existen enzimas que se sobre-expresan en pacientes con determinadas enfermedades,

luego su uso como estímulo está bien justificado.


79

La liberación del fármaco, está basada en las 2 posibles funciones que puede

realizar la enzima en los nanosistemas [Aznar et al., 2016]:

Las enzimas actúan hidrolizando o degradando los ligandos que funcionalizan y

cierran los poros de la MSN [Mas et al., 2013].

Las enzimas actúan como tapa, ancladas sobre la MSN, y la apertura se produce

mediada por los productos derivados de la actividad catalítica [Yang et al.,

2012].

Sensibles a luz:

El empleo de la luz como estímulo se basa en la posibilidad de provocar la

liberación controlada de agentes terapéuticos de forma no invasiva. El paciente puede

ser irradiado de forma focalizada, sin afectar a los tejidos sanos, aunque para ello se

necesita una luz potente que atraviese las capas epidérmicas.

Los mecanismos de apertura de la puerta se deben a:

Foto-dimerizaciones: los ligandos que funcionalizan la MSN son isómeros cis-

trans que al cambiar de conformación producen la liberación del fármaco

[Angelos et al., 2007].

Foto-ruptura: los ligandos que funcionalizan la MSN son fotosensibles,

rompiéndose al incidir la luz sobre ellos [Guardado-Alvarez et al., 2013].

Calentamiento foto-inducido de nanopartículas metálicas que actúan como

tapas [Vivero-Escoto et al., 2009].

Sensibles a la temperatura:

Como en el caso anterior, la temperatura tiene la ventaja de poder ser utilizada

como un estímulo no invasivo. Además, su empleo está bien justificado por el aumento


80

de temperatura que se registra en tejidos inflamados, infecciosos o tumorales. Dicho

aumento es de apenas 5ºC lo que limita mucho las aplicaciones biomédicas de estos

sistemas, a sistemas muy sensibles [Aznar et al., 2016].

Los materiales más empleados en este tipo de puerta son los polímeros

termosensibles. En este caso, el mecanismo de apertura se debe a que el aumento de

la temperatura provoca un cambio de fase en los polímeros, haciendo que colapsen y

abran el poro [Liu et al., 2009].

Sensibles a moléculas y biomoléculas:

La liberación del fármaco en estos sistemas se fundamenta en un cambio en la

polaridad de las interacciones supramoleculares existentes entre la superficie de la

MSN y los ligandos que forman la puerta.

Se pueden emplear una amplia gama de moléculas cargadas, tanto aniones

como cationes, o neutras como las biomoléculas. A excepción de estas últimas, estos

sistemas son muy poco específicos y su aplicabilidad médica no está muy extendida

[Choi et al., 2011].

Sensibles a la ultrasonidos y campos magnéticos:

Ambos estímulos son no invasivos, a pesar de ser necesaria su aplicación de

forma externa. Éstos, al contrario que la luz, sí pueden alcanzar tejidos profundos, lo

que aumenta su aplicabilidad en el campo de la biomedicina. Sin embargo, el empleo

de US como estímulo para la liberación de fármacos en personas está más que

cuestionado por el efecto genotóxico que podría causar [Aznar et al., 2016].

Por otro lado, el uso del magnetismo como estímulo, tiene una ventaja extra.

Los nanosistemas pueden ser también orientados hasta los tejidos diana antes de la

liberación de los agentes terapéuticos [Thomas et al., 2010].

6 OBJECTIVES

6. OBJECTIVES

81

The main objective of this Thesis is the preparation and characterization of

novel functionalized nanomaterials and nanohybrids for their evaluation as

transduction elements for electrochemical biosensors construction and the design of

advanced enzyme-controlled smart drug delivery systems. This Thesis aims to reach its

specific objective goal by:

For biosensors design:

• Preparation and characterization of polyfunctionalized gold nanoparticles to be

used as building blocks for the assembly of electropolymerized networks of

nanoparticles on electrodes surfaces.

• Establishment of original non-covalent strategies for the modification of single-

walled carbon nanotubes with magnetic nanoparticles and β-cyclodextrins

derivatives.

• Preparation and characterization of poliamidoamine dendron and dendrimer

derivatives bearing β-cyclodextrins units for the self-assembly of layered

supramolecular architectures on electrodes surfaces by using adamantane-

modified enzymes as biomolecular building blocks.

• Establishment of optimized protocols for the proper assembly of electrochemical

enzyme biosensors and immunosensors on the nanostructured electrode surfaces.

• Determination of the analytical and stability properties of the electroanalytical

devices.

For drug delivery systems:

Preparation and characterization of mesoporous silica nanoparticles based on

functional nanocontainers capped with lactose-esterase neoglycoenzyme.

6. OBJECTIVES

82

Design and development of original gold-mesoporous silica Janus nanoparticles to

be used as “hardware” for the assembly of single enzyme/multienzyme logic gate-

based nanomachines for smart delivery.

Establishment of optimized protocols for the proper assembly of the

biofunctionalized nanomachines.

Structural, functional and operational characterization of the enzyme-controlled

nanomachines for on-command delivery.

7 PUBLICACIONES

CIENTÍFICAS

7.1

Electrochimica Acta 56, 2011, 4672-4677

Wiring horseradish peroxidase on gold nanoparticles-based

nanostructured polymeric network for the construction of mediatorless

hydrogen peroxide biosensor

Electrochimica Acta 56 (2011) 4672-4677 7. PUBLICACIONES

83

WIRING HORSERADISH PEROXIDASE ON GOLD NANOPARTICLES-BASED

NANOSTRUCTURED POLYMERIC NETWORK FOR THE CONSTRUCTION OF

MEDIATORLESS HYDROGEN PEROXIDE BIOSENSOR

Reynaldo Villalonga, Paula Díez, Paloma Yáñez-Sedeño, José M. Pingarrón*

Department of Analytical Chemistry, Faculty of Chemistry, Complutense University of

Madrid, 28040-Madrid Spain

*Corresponding author. Phone: +34 91 3944315, Fax: +34 91 3944329,

E-mail: [email protected]

http://de.mc304.mail.yahoo.com/mc/[email protected]


84

ABSTRACT

Gold electrodes were functionalized with an electropolymerized matrix of Au

nanoparticles modified with 2-mercaptoethanesulfonic acid, 3-mercaptophenyl

boronic acid and p-aminothiophenol. The resulting nanostructured electroconductive

matrix was used as support for the oriented immobilization of horseradish peroxidase

to construct a reagentless amperometric biosensor for H2O2. The electrode, poised at

0.0 mV, exhibited a rapid response within 8 s and a linear calibration range from 5 µM

to 1.1 mM H2O2. The sensitivity of the biosensor was determined as 498 µA/M cm2,

and its detection limit was 1.5 µM H2O2 at a signal-to-noise ratio of 3. The electrode

retained 95% and 72% of its initial activity after 21 and 40 days of storage at 4ºC.

KEYWORDS: Biosensor, gold nanoparticles, peroxidase, wired enzyme, nanostructured

surface.

INTRODUCTION

Noble metal nanoparticles have been extensively employed in the design and

construction of amperometric enzyme biosensors [Kwon et al., 2010; Pingarrón et al.,

2008; Ren et al., 2005] due to their unique characteristics such as high surface energy

and surface-to-volume ratio, ability to decrease proteins-metal particles distance, and

the possibility to act as electroconductive wires between the enzyme and the

electrodes [Pingarrón et al., 2008]. These properties allow the direct electron transfer

between the electrode surfaces and the catalytic site of enzymes through a tunnelling

mechanism [Bharathi et al., 2001].

Several approaches have been used for coupling noble metal nanoparticles on

enzyme electrodes including electrodeposition [Wu et al., 2005; Lim et al., 2005],

covalent attachment [Yang et al., 2006], supramolecular association [Holzinger et al.,

2009] or electrostatic adsorption [Manso et al., 2008; Yang et al., 2006] to the

modified electrode surface, as well as physical inclusion into the electrode matrix


85

[Carralero et al., 2006]. In addition, metal nanoparticles-modified enzymes have been

successfully employed in the construction of amperometric biosensors [Chico et al.,

2009].

Recently, it has been reported the synthesis of novel aniline and boronic acid-

modified gold nanoparticles (AuNPs) [Riskin et al., 2010]. These nanoparticles were

electropolymerized to form suitable AuNPs-based molecularly imprinted electrodes for

the successful construction of SPR sensors towards low molecular weight compounds

having 1,2-vicinal diols into their structure [Riskin et al., 2010; Frasconi et al., 2010].

Such kind of nanostructured electroconductive material opens a promising field in

electroanalytical chemistry. However, their use in the design of enzyme

electrochemical biosensors has not been reported yet.

In the present work we describe a novel approach for preparing reagentless

biosensors by using polymerized AuNPs (polyAuNPs) as wiring material for redox

enzymes. In this sense, AuNPs modified with 2-mercaptoethanesulfonic acid, 3-

mercaptophenyl boronic acid and p-aminothiophenol were synthesized and further

electropolymerized in acid media on a gold electrode surface through the formation of

bisaniline-cross-linked network. The presence of pendant boronic acid residues in this

electroconductive matrix allows the oriented and reversible immobilization of

glycoproteins by the formation of cyclic esters between the sugar residues and the

boronic acid groups [Qi et al., 2010].

As a model glycoenzyme, we selected horseradish peroxidase (HRP, EC 1.11.1.7,

H2O2 oxidoreductase), a redox enzyme having an accessible ferroprotoporphyrin group

at the active site [Veitch, 2004]. This structural characteristic allows its wiring to

properly modified electrode surfaces and, consequently, HRP constitutes the enzyme

most commonly used in the construction of third-generation biosensor [Shan et al.,

2010; Xi et al., 2009; Yin et al., 2009].


86

MATERIALS AND METHODS

1. Reagents and apparatus

Horseradish peroxidase (HRP, Type II, 105 U/mg), HAuCl4, NaBH4, 2-

mercaptoethanesulfonic acid, 3-mercaptophenyl boronic acid and p-aminothiophenol

were purchased from Sigma (USA). All other chemicals were of analytical grade.

Cyclic voltammetry and electrochemical impedance spectroscopy experiments

were performed using a FRA2 µAutolab Type III potentiostat/galvanostat and the data

were acquired using GPES Ver. 4.9 and Frequency Response Analyser softwares,

respectively (Metrohm Autolab B.V., The Netherlands). Amperometric measurements

were performed with a dual-channel ultrasensitive Inbea potentiostat (Inbea

Biosensores S.L., Spain). A conventional three-electrode system was employed in all

electrochemical studies. The working electrode was a gold disk (CHI Instruments, UK,

2.0 mm diameter) modified with the electropolymerized AuNPs network and the

immobilized enzyme. An Ag/AgCl/KCl (3 M) and a Pt wire were used as reference and

counter electrodes, respectively. Bioelectrode measurements were carried out at 25ºC

in 0.1 M sodium phosphate buffer, pH 7.0 (working volume 10 ml). The solution was

exhaustively de-aerated before each electrochemical experiment. The solutions were

stirred at 300 rpm with a magnetic bar during amperometric measurements. For

analytical purposes, 10 mM H2O2 solutions in 50 mM sodium phosphate buffer, pH 7.0

were freshly prepared.

The microstructure and surface morphology of the electropolymerized

bisaniline-cross-linked AuNPs network was characterized using high resolution field

emission scanning electron microscopy (FE-SEM) with a JEOL JSM-6335F electron

microscope (JEOL Ltd., Japan). Transmission electron microscopy (TEM) measurements

were performed with a JEOL JEM-2000 FX microscope. The morphology

of gold nanostructured surface was investigated using atomic force microscopy (AFM)

with a SPM Nanoscope IIIa multimode microscope (Veeco Instruments Inc., USA).

Spectrophotometric determinations were performed using an Agilent 8453 UV/VIS

spectrophotometer (Hewlett Packard, USA).


87

2. Synthesis of the functionalized AuNPs

To prepare the modified AuNPs, 197 mg of HAuCl4 were dissolved in 50 mL of

de-aerated dimethyl sulfoxide (DMSO). This solution was added dropwise to other 50

mL of de-aerated DMSO containing 300 mg sodium borohydride, 36 mg 2-

mercaptoethanesulfonic acid, 11 mg 3-mercaptophenyl boronic acid and 8 mg p-

aminothiophenol under vigorous stirring. The reaction mixture turned deep brown

immediately, but the reaction was allowed to continue for 24 h. The functionalized

AuNPs were then precipitated by adding 100 mL CH3CN, collected by centrifugation,

and washed with 60 mL CH3CN:DMSO (1:1 v/v), 60 mL ethanol and 20 mL diethyl ether.

The nanoparticles were finally isolated by centrifugation and dried under N2. The

nanoparticles were characterized by FT-IR and UV-vis spectroscopy as well as by TEM.

3. Preparation of the electropolymerized AuNPs-modified enzyme electrode

The disk gold electrode was first exhaustively polished with alumina powder

(0.3 µm), then rinsed with double distilled water and immersed in an ultrasonic bath

for 5 min. The electrode was further dipped in concentrated HNO3 for 1 h, followed by

rinsing in absolute EtOH and double distilled water. Finally, the electrode was

electrochemically cleaned by ten consecutive voltammetric cycles in 0.1 M H2SO4 and

further dry under N2.

To modify the electrode with the polyAuNPs network, its surface was initially

capped with a monolayer of p-aminothiophenol by dipping the electrode for 2 h in a 10

mM ethanolic solution of the thiol. The electrode was exhaustively washed with

ethanol and double distilled water, and dipped into a 2 mg/mL solution of the AuNPs in

0.1 M H2SO4. The electrode surface was then coated with the electropolymerized

bisaniline-cross-linked AuNPs network by application of 10 potential cycles between -

0.35 and +0.85 V vs Ag/AgCl at a scan rate of 100 mV/s, followed by application of a

constant potential of 0.85 for 1 h. The modified electrode was further exhaustively

washed with 0.1 M H2SO4 and double distilled water before enzyme immobilization.


88

This modified electrode was further dipped into a 50 mM sodium phosphate

buffer, pH 7.0 solution containing HRP at 5 mg/mL concentration. The mixture was

kept at 4ºC for 12 h, then exhaustively washed with cool 50 mM sodium phosphate

buffer, pH 7.0 and finally kept at 4ºC in dry conditions until used.

RESULTS AND DISCUSSION

Gold colloids were grown in the presence of 2-mercaptoethanesulfonic acid, 3-

mercaptophenyl boronic acid and p-aminothiophenol, yielding dark red and water

soluble nanoparticles. The functionalized AuNPs particles showed spherical geometry

with an average diameter of 2.5 ± 0.4 nm, as determined by HRTEM (Figure 1).

Figure 1. HRTEM image of AuNPs.

The radial geometry, small size and low polydispersion showed by the

nanoparticles result from the synthetic approach used, in which both metal reduction

and attachment of thiolated ligands to the surface of the developing Au particles take

place in the same step [Liu et al., 2000]. The selection of the three different thiol-

derivatives used as capping ligands for the AuNPs was made according to the desired

properties to be conferred to the nanomaterial. In fact, mercaptophenyl boronic acid

moieties should act as recognition elements for the vicinal diols in glycoenzymes,


89

thioaniline residues should allow the electrochemical polymerization of the

nanoparticles via the formation of bisaniline-cross-linked network, and

mercaptoethanesulfonic acid should favour the solubilisation and stabilization of the

resulting nanoparticles [Riskin et al., 2010; Frasconi et al., 2010]. The presence of these

capping thiols on the surface of AuNPs was confirmed by FT-IR spectroscopy.

Characterization of functionalized AuNPs

The UV-Vis spectrum of the functionalized AuNPs, dissolved in 50 mM sodium

phosphate buffer pH 7.0, showed relatively weak plasmon resonance absorption

maxima in the range 510-520 nm (Figure 2). The intensity of this band was not affected

after addition of BSA, a non-glycosidated protein.

Figure 2. UV-vis spectra of 0.2 mg/mL solution of functionalized AuNPs in 50 mM

sodium phosphate buffer, pH 7.0 in the absence (A) and the presence of 0.5 mg/mL BSA

(B) and HRP (C).

However, the intensity of the plasmon resonance band exhibited a slight

increase when HRP was added to the AuNP solution. This effect is related to the


90

association of the enzyme to the metal nanoparticles surface, as it was previously

reported for other protein-AuNPs systems [Nath et al., 2002; Villalonga et al., 2005]. In

the present case, this interaction should be most likely due to the formation of cyclic

boronate complexes between boronic acid moieties at the AuNPs and vicinal diols

from the sugar chains in the glycoenzyme [Qi et al., 2010].

Figure 3. Field emission SEM image of the polyAuNP-modified electrode.

Functionalized AuNPs were electropolymerized in 0.1 M H2SO4 on a thioaniline-

modified gold electrode. Such electrode modification ensures the formation of

bisaniline bonds between thioaniline groups on the electrode and AuNPs surfaces as

the first step for an efficient polymerization process [Frasconi et al., 2010]. It is

important to remark that, conversely to that employed in previous reports [Riskin et

al., 2010; Frasconi et al., 2010], we selected an acid electropolymerization medium in

order to ensure the electroconductivity of the resulting emeraldine salt. Figure 3

shows the FE-SEM image of the electrode surface after polymerization. A three-

dimensional nanostructured network, in which the spherically-shaped metal

nanoparticles are well defined, can be observed over the entire electrode surface. It

should be noted that the polymerized matrix was composed of a non-ordered array of

AuNPs-based protuberances of different heights, separated by irregularly distributed

nanoholes.


91

Figure 4. AFM images of the polyAuNP-modified electrode.

This surface topology was confirmed by AFM analysis as illustrated in Figure 4.

The polymerized AuNPs-based network showed roughness average of 329 nm and

average thickness of 984 nm. This matrix includes nanoholes and AuNPs-based

protuberant structures representing 47.9% and 52.1% of the total surface area,

respectively. The maximum height of such nanoparticle-based protuberant structures

was determined to be 1.96 µm.

Horseradish peroxidase was immobilized on the surface of the polyAuNPs-

modified electrode via interaction with the pendant boronic acid residues. This

strategy allowed the oriented immobilization of the enzyme on the electrode surface,

taking into account that the glycosidation chains in HRP are located so far from the

catalytic active site [Veitch, 2004].

Cyclic voltammetry of [Fe(CN)6]4−/3− ions is a valuable strategy to evaluate the

barrier properties of modified electrodes, because the electron transfer between the

electroactive [Fe(CN)6]4−/3− ions and the electrode surface must occur by tunnelling

either through the modifying matrix or through defects in the matrix. Figure 5 shows

the cyclic voltammograms from 5 mM [Fe(CN)6]4−/3− in 0.1 M KCl solution. As can be

observed, the non-modified Au electrode showed a well-defined typical diffusion-

limited behaviour, similar to that exhibited by the polyAuNPs-functionalized electrode.


92

Figure 5. Cyclic voltammograms recorded at a bare gold disk electrode (A), and a

polyAuNPs-modified electrode before (B) and after HRP immobilization (C), in 0.1 M KCl

solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1).

However, the peak heights were dramatically increased after polymerization of

the AuNPs on the electrode surface, which can be attributed to the increased

electroactive surface area of the electrode after formation of the bisaniline-cross-

linked electroconductive network of metal nanoparticles. In fact, it was further

demonstrated by applying the Randles-Sevcik equation that the active surface area of

the Au electrode varied from 5.5 mm2 to 12.5 mm2 upon functionalization with the

AuNPs-based polymer. As expected, the immobilization of HRP on the modified

electrode gave rise to a sharp decrease in the peak currents and a large separation

between the anodic and cathodic peaks as a consequence of the barrier formed by the

protein on the electrode surface. This result suggested a high coverage of the

electrode surface by the enzyme protein, favoured by the high concentration of

boronic acid residues on the AuNPs polymeric matrix.


93

Figure 6. Impedance plane diagram (−Z″ versus Z′) for the EIS measurements at a bare

gold disk electrode () and a polyAuNPs-modified electrode before () and after HRP

immobilization (), in 0.1 M KCl solution containing 5 mM

K3[Fe(CN)6]/K4[Fe(CN)6] (1:1).

Electrochemical impedance spectroscopy of 5 mM [Fe(CN)6]4−/3− ions in 0.1 M

KCl solution was performed in order to determine the effect of the electrode

configurations on the impedance changes on their surfaces. Figure 6 shows the

corresponding Nyquist plots. The small semicircle diameter observed at high

frequencies for the bare Au electrode, indicative of a fast electron transfer, was even

reduced after electropolymerization of AuNPs on the electrode surface. This

polymerization produced two effects: i) the increased electron transfer of

[Fe(CN)6]4−/3− ions to the electrode surface modified with the polyaniline matrix

enriched with AuNPs, and ii) the electrostatic repulsion of [Fe(CN)6]4−/3− ions by the

negative-charged electropolymerized surface functionalized with sulfonic acid

residues. The overall decrease in the electron transfer resistance observed

experimentally suggested a predominant effect of the former process. A noticeable

increase in the electron transfer resistance was observed upon HRP attachment, thus


94

indicating blocking of the electrode surface by the enzyme. This significant insulating

effect also suggested a high coverage of the electrode by the glycoenzyme.

Detection of H2O2 at the polymerized AuNPs-modified electrode

The HRP-modified electrode was further evaluated as a biosensor device for

H2O2. As a control, HRP was also immobilized under the same experimental conditions

on a gold disk electrode previously coated with a monolayer of p-mercaptophenyl

boronic acid, in order to quantify the influence of the AuNPs-based polymeric matrix

on the electroanalytical response of the biosensor.

Figure 7. Cyclic voltammograms recorded at HRP-modified electrodes in the absence

and the presence of H2O2 in 0.1 M sodium phosphate buffer, pH 7.0. HRP-Au electrode

(A), HRP-polyAuNPs-Au electrode (B), HRP-Au electrode + 1 µM H2O2 (C), HRP-Au

electrode + 2 µM H2O2 (D), HRP-polyAuNPs-Au electrode + 1 µM H2O2 (E) and HRP-

polyAuNPs-Au electrode + 2 µM H2O2 (F).

Figure 7 compares cyclic voltammograms recorded at these two enzyme

electrodes in a deaereated 0.1 M sodium phosphate buffer solution of pH 7.0 at a 50


95

mV/s scan rate, in the absence and the presence of added H2O2. A slight catalytic

response of the HRP-modified control electrode was observed after addition of the

analyte (curves A, C and D). This poor electrocatalytic behaviour could be attributed to

a low coverage of the electrode surface by HRP using this configuration.

On the contrary, a large catalytic current was observed at the polyAuNPs-

modified enzyme electrode in the presence of 1 µM and 2 µM H2O2 (curves B, E and F).

These results demonstrated that the electrocatalytic behaviour of HRP was improved

upon its immobilization on the polyAuNPs-based conductive matrix, allowing an

efficient electron transfer between the redox active site of the enzyme and the

electrode surface. As it is widely documented in the literature [Zhaoyang et al., 2006;

Chico et al., 2009], the mechanism of the electrocatalytic reduction of H2O2 can be

schematized as:

HRP(FeIII

) + H2O2 Compound I ([FeIV

=O] +) + H2O (1)

Compound I ([FeIV

=O] +) + e + H

+ Compound II (Fe

IV=O) (2)

Compound II (FeIV

=O) + e + H+ HRP(FeIII

) + H2O (3)

Taking into consideration that compound II is more stable than compound I

(which is a free radical), it is expected that the increase in cathodic peak current should

be mainly caused by the electroreduction of compound II [Zhaoyang et al., 2006].

The effect of the applied potential on the amperometric response of the

AuNPs-modified electrode towards the addition of H2O2 in 0.1 M sodium phosphate

buffer, pH 7.0 was further evaluated. The cathodic current increased steadily with

changing the applied potential from 200 mV to -500 mV versus Ag/AgCl. However, the

high signal-to-noise ratio achieved for the electrode response at 0 V, in addition to the

well know advantages associated with the use of this detection potential [Svitel et al.,

1998] moved us to select it for further experiments. The effect of pH on the

amperometric response of the electrode toward H2O2 at 0 V was also checked in the

5.0-9.0 pH range. The biosensor current exhibited an almost bell-shaped behaviour


96

with the highest response in the range of pH 6.5-7.0. Thus, pH 7.0 was selected for

further works.

Figure 8. Amperometric responses recorded with HRP-Au (A) and HRP-polyAuNPs-Au

(B) electrodes for successive additions of 10 mM H2O2 to 10 mL of 0.1 M sodium

phosphate buffer, pH 7.0. Eapp. = 0.0 V.

Figure 8 shows the comparison of the amperometric responses obtained with

the two different HRP-modified electrodes to successive additions of H2O2. As can be

observed, the enzyme electrode without polymerized AuNP gave a poor

electroanalytical response, while, as expected, the polymeric AuNP-modified electrode

responded rapidly to the changes in the analyte concentration, with 95% of the steady-

state signals achieved within 8 seconds. The current response of this bioelectrode was

plotted against the H2O2 concentration, the resulting calibration curve being shown in

Figure 9.


97

Figure 9. Calibration curve for H2O2 obtained with a HRP-polyAuNPs-Au biosensor in

the 5 µM to 1.1 mM H2O2 concentration range.

The amperometric response exhibited a linear range between 5 µM and 1.1

mM H2O2, with an equation:

y = 62.3 x + 0.001

Where y is the amperometric current in µA and x is the molar concentration of

hydrogen peroxide, and a correlation coefficient of 0.996 (n = 12). The sensitivity of the

biosensor towards H2O2 was determined as 498 µA/M cm2. A detection limit of 1.5 µM

(S/N = 3) was calculated. Furthermore, an apparent Michaelis-Menten constant, KMapp ,

and a maximum rate of the enzyme reaction, Vmax, values of 1.01 mM and 89.3 nA,

respectively, were obtained. Table 1 shows the comparison of the analytical

characteristics of the developed biosensor with those reported recently for other HRP

mediatorless biosensors based on the use of AuNPs. As it can be seen, the detection

limit and the apparent Michaelis-Menten constant are among the lowest values

reported despite the much less extreme detection potential applied. These values

suggested that the immobilized HRP retained high catalytic activity and high affinity to


98

H2O2, probably favoured by the oriented and non-covalent immobilization approach

used.

Table 1. Comparison of analytical properties of the biosensor with previously reported

AuNP-based mediatorless biosensors

Electrode E

(mV)

LR

(mM)

DL

(µM)

KM

(mM) Reference

HRP/SiO2/BSA/AuNP/Thio-

Naf/Au

-200 0.008-3.72 2.0 2.3 Yuan et al., 2010

HRP/AuNP/APTMS/ITO -250 0.02-8.0 8.0 - Wang & Wang, 2004

Naf/HRP-AuNs–TiO2/GCE -300 0.041-0.63 5.9 0.63 Wang et al., 2010

HRP/AuNP/Gph/Chi/GCE -300 0.005-5.13 1.7 2.61 Zhou et al., 2010

HRP/ZnO-AuNP-Naf/GCE -300 0.015-1.1 9.0 1.76 Xiang et al., 2009

HRP/CaCO3-AuNP/Au -200 0.0005-5.2 0.1 - Li et al., 2010

HRP/AuNP-SF/GCE -600 0.01-1.8 5.0 1.22 Yin et al., 2009

HRP/poly(GMA-co-VFc)/GCE +350 2.0-30 2.6 1.14 Şenel et al., 2010

Naf/HRP/AuNP/PDDA/GCE -350 0.2-0.91 99 0.69 Wang et at., 2009a

HRP-AuNP/ALG/Au -400 0.02-13.7 3.0 9.3 Chico et al., 2009

HRP/DNA–AgNP/PDDA–

AuNP/DNA–AgNP/Au

-350 0.007-7.8 2.0 1.3 Ma et al., 2009

HRP/AuNP/ITO -150 0.008-3.0 2.0 0.4 Wang et al., 2009b

HRP/AuNP/SG/BGE -250 0.005-10 2.0 - Jia et al., 2002

HRP/polyAuNP/Au 0 0.005-1.1 1.5 1.01 Present work

LR: Linear range of response; DL: Detection limit; AuNP: Gold nanoparticles; BSA: bovine serum

albumin; Thio: thionine; Naf: Nafion; APTMS: 3-aminopropyltrimethoxysilane; ITO: indium tin

oxide electrode; GCE: glassy carbon electrode; AuNs: gold nanoseeds; Gph: graphene; Chi:

chitosan; SF: silk fibroin; Cys: cysteamine; polyDAN: poly(1,8-diaminonaphthalene); poly(GMA-

co-VFc: poly(glycidyl methacrylate-co-vinylferrocene); PDDA: poly(diallyldimethylammonium

chloride); ALG: sodium alginate; AgNP: silver nanoparticles; SG: silica gel

In addition, the great surface area corresponding to nanoholes in the

polymerized matrix ensures a great population of enzyme immobilized into these


99

nanocavities, and it is expected that such nanostructures protect the hydrophilic

microenvironment around the immobilized HRP.

On the other hand, the amperometric response was shown to be highly

repeatable toward successive additions of H2O2. A relative standard deviation, RSD,

value of 3.6% was calculated for ten successive calibration curves for H2O2.

Additionally, the electrode-to-electrode reproducibility was evaluated from the

response of 12 equivalently prepared biosensors yielding a RSD value of 7.8%.

The selectivity of the sensor was evaluated by measuring the amperometric

response towards 20 µM H2O2 in the presence of seven possible interfering substances

at a 100 µM concentration level: L-glucose, ethanol, acetic acid, ascorbic acid, lactic

acid, citric acid and uric acid. The biosensor exhibited an excellent selectivity towards

H2O2 since no significant changes in the steady-state current measured for the analyte

occurred under the above mentioned conditions. It should be highlighted that only in

the presence of an ascorbic acid concentration as high as 250 µM, the amperometric

response was reduced to 92% of its initial value. This excellent behaviour could be

justified by the high concentration of sulfonic acid residues at the surface of the

AuNPs-based polymeric matrix, which constitutes a protective electrostatic barrier to

the diffusion of the main interference substances to the electrode surface.

Figure 10 shows the effect of time of storage at 4ºC under dry conditions on

the slope value of the calibration plot for H2O2. As it can be observed, the biosensor

using the AuNPs-based polymeric matrix exhibited an excellent long-term stability,

retaining 95% and 72% of its initial activity after 21 and 40 days of storage,

respectively. This high stability behaviour can be attributed to the strong multipoint

attachment of the glycoenzyme on the boronic acid-functionalized conductive

polymer, thus avoiding denaturation and leaking of the active protein molecule from

the electrode surface during storage and testing.


100

Figure 10. Effect of the storage time at 4ºC of the HRP-polyAuNPs-Au biosensor on the

sensitivity for H2O2 determination.

Moreover, the presence of a high concentration of nanostructured holes in the

polymerized matrix provides protection of the immobilized protein into the controlled

microenvironment of these nanoholes, contributing to the high stability showed by the

biosensor during storage.

CONCLUSIONS

A novel strategy, based on the modification of gold substrates with a

conductive polymeric matrix of aniline and boronic acid-functionalized AuNPs, and its

further use for the oriented immobilization of the glycoenzyme HRP, was developed.

The approach was used for the construction of a reagentless biosensor for H2O2 which

exhibits a remarkable good analytical performance in terms of sensitivity, stability and

fast electroanalytical response. Taking into account these results, it can be predicted

that the use of a bisaniline-cross-linked nanostructured network of metal

nanoparticles constitutes an excellent strategy to construct scaffolds for the successful


101

immobilization of redox enzymes in order to prepare mediatorless amperometric

biosensors.

ACKNOWLEDGEMENTS

R. Villalonga acknowledge to Ramón & Cajal contract from the Spanish Ministry

of Science and Innovation. Financial support from the Spanish Ministerio de Ciencia e

Innovación CTQ2009-12650, CTQ2009-09351) and Comunidad de Madrid S2009/PPQ-

1642, programme AVANSENS is gratefully acknowledged.

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7.2

Analyst 137, 2012, 342-348

Electropolymerized network of polyamidoamine dendron-coated

gold nanoparticles as novel nanostructured electrode surface

for biosensor construction

Analyst 137 (2012) 342-348 7. PUBLICACIONES

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ELECTROPOLYMERIZED NETWORK OF POLYAMIDOAMINE DENDRON-

COATED GOLD NANOPARTICLES AS NOVEL NANOSTRUCTURED

ELECTRODE SURFACE FOR BIOSENSOR CONSTRUCTION

Reynaldo Villalonga,a Paula Díez,a Santiago Casado,b Marcos Eguílaz,a Paloma Yáñez-

Sedeño,a José M. Pingarrón*a

ABSTRACT

Polyfuntionalized gold nanoparticles were prepared by using 2-

mercaptoethanesulfonic acid, p-aminothiophenol and cysteamine core

polyamidoamine G-4 dendron as capping ligands. The nanoparticles were

electropolymerized on Au electrode surface through the formation of bisaniline-cross-

linked network. The enzyme tyrosinase was further crosslinked on this nanostructured

matrix. The enzyme electrode, poised at -100 mV, was used for the amperometric

quantification of catechol. The biosensor showed a linear response from 50 nM to 10

µM catechol, with a low detection limit of 20 nM and a sensitivity of 1.94 A/M cm2.

The electrode retained 96% and 67% of its initial activity after 16 and 30 days of

storage at 4ºC under dry conditions.

INTRODUCTION

Nowaday, the construction of new electrochemical biosensors with improved

bioanalytical performance and robustness is frequently directly linked to the

development of novel strategies for tailor-made design of electrode surfaces. Such

strategies should provide electrical interfaces with nano- or microsized tree-

dimensional topology that allow the successful and stable immobilisation of the

biomolecules without affecting their biological function, but also favouring the fast and


108

efficient occurrence of the electrochemical processes involved in the analytical

reaction on such interface.1

During last recent years, nanosized materials have been exhaustively employed

in the modification of electrodes for biosensing purposes.1,2 In general, the

development of methodologies for nanostructuring electrode surfaces with such

purpose should consider the following points: i) the type of surface to be modified, ii)

the nanomaterial to be used, iii) other compounds/materials that could be also

employed, iv) the physical/chemical method for surface modification, v) the

biomolecule to be immobilized and the method to do that, and vi) the electrochemical

reaction to take place on the functionalized electrode surface.

Strategies for nanostructuring electrode surfaces have been mainly based on

the use of “hard” nanomaterials, such as quantum dots,3 carbon nanotubes,1,4

graphene,5 carbon nanofibers,6 as well as metal7 and metal oxide nanoparticles.8 Such

approaches include the use of single nanomateriales, mixtures of them, or composites

with polymeric materials or low molecular weight compounds.1,2,4 Since the number of

nanomaterials with electroanalytical interest is finite, and the use of most of them in

biosensor technology is well documented, the state-of-the-art of nanostructuring

electrode surface is mainly addressed to the development of innovative

functionalization strategies in which other novel co-materials and modifying ligands

are considered.

Recently, it has been described the synthesis of polyfunctionalyzed gold

nanoparticles (AuNPs) carrying aniline and boronic acid moieties, which have been

employed for the modification of electrode surfaces with a nanostructured conductive

matrix through electropolymerization.9,10 The presence of boronic acid residues as

pedant ligands on such surfaces allowed their successful use for the construction of

glycoenzyme-based biosensors,10 as well as for surface plasmon resonance (SPR)

sensors towards molecules having 1,2-vicinal diols.9

The possibility of tailor-made design of novel AuNPs with desired and specific

properties by manipulation of the capping ligands, as well as the electroconductive


109

character of the resulting nanostructured three dimensional networks opens a

promising field in electroanalytical chemistry for this kind of materials. In this regards,

we are concerned to evaluate high molecular weight dendritic ligands as capping

material for the preparation of AuNPs-based polymeric matrix and its potential

application in biosensor technology. It should be highlighted that this strategy of

surface coverage with electropolymerized tailor-made nanoparticles could be also a

valuable tool for the preparation of functional materials for biomedicine,

biotechnology, microelectronics and other fields.

Dendrimers and dendrons are nanosized and monodisperse macromolecules

with a regular and highly branched three-dimensional architecture. These “soft”

nanomaterials have unique properties such as structural homogeneity, high surface

reactivity and molecular host capacity.11 Such properties, resulting from the size, the

hydrophilic/hydrophobic character of the internal cavities and the nature of the

chemical groups at the surface of these hyperbranched polymers, support their wide

application in several fields including the design of novel catalysts, chemical sensors,

drug and gene delivery systems, and biosensors.12,13

In this work we describe a novel approach for preparing dendron-coated

AuNPs-based nanostructured electrodes by synthesizing, firstly, polyfunctionalysed

nanoparticles with 2-mercaptoethanesulfonic acid, p-aminothiophenol and cysteamine

core polyamidoamine (PAMAM) G-4 dendron as capping ligands. These

polyfunctionalized AuNPs were further electropolymerized on a gold electrode surface

through the formation of bisaniline-cross-linked network, yielding a three dimensional

matrix on which the enzyme tyrosinase, as a model enzyme, was finally immobilised

through covalent crosslinking. The functionalised enzyme electrode was employed for

the construction of an amperometric biosensor device towards catechol as a model

target compound.


110

RESULTS AND DISCUSION

The rationale of the present work is based on the design of novel

polyfunctionalyzed AuNPs, specifically capped with three different thiol derivatives: p-

aminothiophenol as polymerizable unit, 2-mercaptoethanesulfonic acid as solubilising

moiety and cysteamine core PAMAM G-4 dendron as hyperbranched primary amino

donor for enzyme immobilization.

Fig. 1 Scheme displaying the steps involved in the construction of an electrochemical

tyrosinase biosensor based on a gold electrode nanostructured with electropolymerized

PAMAM G-4 dendron-coated AuNPs.

These nanoparticles were employed for nanostructuring gold electrode

surfaces through electropolymerization, which were further used as supports to

immobilize tyrosinase for constructing an electrochemical biosensor towards catechol.

A graphical overview of this methodology is shown in Fig. 1.


111

The strategy used to synthesize the dendron-polyfunctionalized AuNPs was

based on the treatment, firstly, of cystamine core PAMAM G-4 dendrimer with NaBH4

in DMSO to ensure the complete reduction of the disulfide bonds at the dendrimer

core, then releasing the thiol-active dendron derivative. This process was performed

with the presence of 2-mercaptoethanesulfonic acid and p-aminothiophenol in the

reaction medium. Polyfunctionalized AuNPs were then prepared by fast reduction of

HAuCl4 with the mixture containing NaBH4 and the capping thiol ligands. Using this

procedure, the formation of dendrimer-encapsulated AuNPs was avoided, by

eliminating the required initial sequestering of Au(III) ions within the dendrimer

cavities before reduction.13,14

Fig. 2 HRTEM image of dendron-functionalized AuNPs

In addition, the presence of thiolated ligands in the reaction medium, including

the bulky cysteamine core PAMAM G-4 dendron, ensures the fast coating of the

growing Au colloids, then avoiding the formation of dendrimer-encapsulated AuNPs

adducts.

This fact was confirmed by HRTEM (Fig. 2), revealing that the as-prepared

dendron polyfunctionalized-AuNPs showed spherical geometry with an average

diameter of 5.7 ± 0.9 nm. This AuNPs size was slightly larger than those described for

PAMAM G-4 dendrimers (about 4.5 nm),14,15 and larger than the previously reported


112

for metal nanoparticles encapsulated into dendritic molecules,16 suggesting than the

as-synthesized AuNPs were bigger than the cavities of the cysteamine core PAMAM G-

4 dendron.

Fig. 3 Cyclic voltammograms for the first electropolymerization process of PAMAM G-4

dendron-coated AuNPs on gold electrode surface, in 0.1 M H2SO4. Scan rate: 100

mV/s.

AuNPs were electropolymerized in 0.1 M H2SO4 on a thioaniline-modified gold

electrode surface. The cyclic voltammograms recorded during the cyclic

electropolymerization process are shown in Fig. 3. Repeated potential scanning over

the -0.35 V to +0.85 V range resulted in the appearance and continuous growth of

anodic and cathodic peaks around 505 mV and 250 mV, respectively. This is the

characteristic pattern for the electrochemical formation of dimeric and low molecular

weight aniline condensation adducts,17 clearly indicating the formation of bisaniline-

cross-linked network of AuNPs on the electrode surface. In order to prepare highly

nanostructured electrodes, and taking into account the unfavourable effect that bulky

dendron moieties should cause on the AuNPs cross-linking due to steric hindrance, a


113

second electropolymerization process over one hour at a fixed potential of +0.85 V was

also carried out.

Fig. 4 AFM images and height histogram of the PAMAM G-4 dendron-coated AuNPs-

modified electrode.

The topology of the electropolymerized AuNPs network on metal electrode

surface was characterised by AFM (Fig. 4). The electrode surface was completely

covered by a three-dimensional and dense-packed matrix, showing well defined

spherically-shaped nanostructures. The average diameter of such electropolymerized

nanostructures was significantly higher (110 ± 20 nm) than the estimated for AuNPs by

HRTEM. This fact can be justified by the contribution of the capping dendritic moieties

to the overall nanostructure size, as well as by the detection of several cross-linked

dendron-coated AuNPs as a unique nanostructure by the AFM tip. The roughness

average of this nanostructured matrix was estimated to be around 6 nm. This modified

surface was characterized by a high content of interstitial holes among the spherically-

shaped nanostructures, representing 55% of the total area. The Gaussian-type

distribution of the nanoparticle-based protuberant structures height showed a mean

value of 27.4 nm, and a maximum of 54.5 nm for the spherically-shaped

nanostructures.

In comparison with previous reports dealing with the preparation of

electropolymerized matrix of boronic acid-functionalized AuNPs on gold electrodes,10

AFM studies revealed that dendron-coated AuNPs yield a more homogeneous and less

rough surface. This fact could be caused by the presence of the bulky dendritic


114

moieties on the AuNPs surface, controlling the rate of electropolymerization by steric

hindrance mechanisms to yield a more ordered matrix with soft surface characteristics.

The AuNPs-modified electrode was used as support for the covalent

immobilization of tyrosinase, through glutaraldehyde-mediated crosslinking. Although

covalent immobilization methods such as those using carbodiimide coupling could be

employed also, in general such methods allow lower immobilized protein loadings thus

leading to smaller electroanalytical signals, and, therefore we decided to use the

crosslinking approach. The optimum enzyme coating conditions, yielding highest

electroanalytical response, was established for 100 µg of tyrosinase and 8.3% (v/v)

glutaraldehyde through a cross-linking reaction on the PAMAM dendron-polyAuNP

modified electrode surface during 2 h at 4ºC in 30 µL of buffer solution of pH 7.0.

Fig. 5 Impedance plane diagram (−Z´´ versus Z´) for the EIS measurements at a bare

gold disk electrode () and at the electropolymerized PAMAM G-4 dendron-coated

AuNPs-modified electrode before () and after tyrosinase immobilization (), in 0.1

M KCl solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1).

Electrochemical impedance spectroscopy was employed to determine the

barrier properties of the electropolymerized dendron-coated AuNPs network on the


115

electrode surface, before and after enzyme immobilization. Fig. 5 shows the Nyquist

plots for the electrodes in a 0.1 M KCl solution containing 5 mM [Fe(CN)6]4−/3− ions. In

order to interpret the experimental results, the Randles equivalent circuit was

assumed as a model, taking into account the shape of the resulting Nyquist plots.

As can be observed, the Nyquist plot of bare gold electrode was characterized

by a small semicircle at high frequencies and a predominant line with slope close to

the unit over the broad range of low frequencies. These facts indicated a predominant

diffusion-controlled mechanism for the redox reaction over a broad range of low

frequencies, as well as the occurrence of fast electron transfer for the [Fe(CN)6]4−/3−

ions pair on the electrode surface at high values of frequencies. Similar behaviour was

observed for the electrode after electropolymerization of PAMAM dendron-coated

AuNPs on its surface, but the Nyquist plots revealed that the electron transfer

resistance was reduced from 205 Ω to 52 Ω after this modification. This fact can be

justified by the synergic contribution of several factors, including: i) the demonstrated

ability of [Fe(CN)6]4−/3− ions to easily penetrate the surface-confined dendrimers as well

as the interstices between them,15 ii) the electrostatic attraction of [Fe(CN)6]4−/3− ions

to the positive charged electrode surface due to multiple amino terminal groups in

dendron residues, and iii) the increased electron transfer of [Fe(CN)6]4−/3− ions to the

electrode surface modified with the polyaniline matrix enriched with AuNPs. On the

other hand, a noticeable increase in the electron transfer resistance (760 Ω) was

observed upon tyrosinase attachment, thus indicating high coverage of the electrode

surface by the enzyme molecules.


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Fig. 6 Cyclic voltammograms recorded at a bare gold disk electrode (A), and at the

electropolymerized PAMAM G-4 dendron-coated AuNPs-modified electrode before (B)

and after tyrosinase immobilization (C), in 0.1 M KCl solution containing 5 mM

K3[Fe(CN)6]/K4[Fe(CN)6] (1:1).

The effect of the different modification processes on the electrochemical

characteristics of the modified gold electrode was also studied by cyclic voltammetry,

as illustrated in Fig. 6. The cyclic voltammograms of 5 mM [Fe(CN)6]4−/3− in 0.1 M KCl

solution at the bare and AuNPs-modified electrodes showed well-defined typical

diffusion-limited patterns, suggesting that the dendron-modified electropolymerized

matrix is permeable to the diffusion of the electrochemical probe. Similar behaviour

was previously described for gold electrodes coated with PAMAM dendrimers.15

Electropolymerization of dendron-coated AuNPs increased the electrochemical surface

area of the electrode from 3.5 mm2 to 11.7 mm2, according to the Randles-Sevcik

model, as revealed by the higher peak currents in the voltammogram. This fact can be

justified by the electroconductive nature of the bisaniline cross-linked metal

nanoparticle-based network. On the contrary, a significant decrease in the peak

currents and a large separation between the anodic and cathodic peaks was observed


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upon immobilization of the enzyme on the electrode surface. A significant reduction to

0.98 mm2 for the electroactive surface area was also noticed, suggesting a high

coverage of the electrode surface by the covalent attached tyrosinase molecules, as

described above in electrochemical impedance spectroscopy experiments.

The enzyme-modified electrode was further evaluated for the amperometric

quantification of catechol. Optimum conditions for this analytical determination were

first established. The applied potential was selected by evaluating the steady-state

current measured with the biosensor for 100 nM catechol at different detection

potential values (results not shown). Maximum amperometric response was achieved

by applying a potential value of -100 mV versus Ag/AgCl in buffer solutions of pH 7.0.

The measured current corresponded to the electrochemical reduction of the o-

benzoquinone formed in the enzyme reaction.

Fig. 7 Amperometric response of the tyrosinase-functionalized p-aminothiophenol-

coated non-nanostructured gold electrode (A) and electropolymerized PAMAM G-4

dendron-coated AuNPs-modified electrode (B) to successive additions of 100 µM

catechol solution to 10 mL of 0.1 M sodium phosphate buffer, pH 7.0. Eapp. = -100 mV.


118

Fig. 7 shows the typical amperometric behaviour of the enzyme electrode to

successive additions of 100 µM catechol solution under the optimal experimental

conditions described above. A control non-nanostructured biosensor, prepared by

cross-linking tyrosinase on a p-aminothiophenol-coated Au electrode was also tested.

Both electrodes showed a fast electroanalytical response, reaching 95% of the

steady-state current within 6 seconds. However, the amperometric response of the

electrode modified with the dendron-coated AuNPs network was 13-times larger than

that obtained with the biosensor constructed with no electropolymerized AuNPs

network. The corresponding calibration curve with the nanostructured bioelectrode

exhibited a linear behaviour in the range between 50 nM and 10 µM catechol,

according to the following equation

iC (mA) = 19·c(catechol/M) + 2·10-6

with a correlation coefficient of 0.999 (n = 10).

The range of linear response obtained was similar to that reported for other

tyrosinase biosensors using a glassy carbon electrode coated with Nafion/multi-walled

carbon nanotubes/ZnO nanoparticles18 and screen printed electrodes modified with

grapheme oxide and AuNPs,19 but shorter that the described for tyrosinase on glassy

carbon electrodes coated with polyaniline/ionic liquid/carbon nanofibers.20

The biosensor showed a very low detection limit of 20 nM towards catechol,

assuming a signal-to-noise ratio of 3. This detection limit at nanomolar level was

similar than the previously reported for tyrosinase immobilized on glassy carbon

electrodes nanostructured with other materials, such as polypyrrole/carbon

nanotubes, AuNPs, chitosan/ZnO nanoparticles, Nafion/multi-walled carbon

nanotubes/ZnO nanoparticles and chitosan/Fe3O4 nanoparticles.18,21-23 However,

subnanomolar limit of detection has been described for a tyrosinase biosensors based

on glassy carbon electrodes coated with polyaniline/ionic liquid/carbon nanofibers.20

The sensitivity of the biosensor was determined as 1.94 A/M cm2, considering

the electroactive area of the enzyme-modified electrode. In addition, the apparent

kinetics constants KM and Imax showed values of 21.9 µM and 505 nA, respectively. This


119

value of the apparent Michaelis-Menten constant ranks among the lower values

reported for tyrosinase-based amperometric biosensors,18,19,22 suggesting that the

affinity of the enzyme for catechol was not greatly affected by immobilization on the

PAMAM dendron-coated AuNPs matrix.

The electroanalytical response of the electrodes was highly reproducible,

showing a relative standard deviation value of 3.1% by measuring ten independent

signals corresponding to 100 nM catechol and using the same electrode. On the other

hand, the electrode-to-electrode reproducibility was evaluated from the response of

10 equivalently prepared biosensors towards 100 nM catechol, yielding a relative

standard deviation value of 8.8%.

Fig. 8 Effect of the storage time at 4ºC of the tyrosinase biosensor on the relative

bioelectrocatalytic activity towards 100 nM catechol determination.

The long-term storage stability of the biosensor at 4ºC in dry conditions was

determined by monitoring its analytical response towards 100 nM catechol with

intermittent usage. The AuNPs-modified enzyme electrode retained about 96% and

67% of its initial bioelectroanalytical activity after 15 and 30 days of storage,

respectively (Fig. 8). This stability can be ascribed to the glutaraldehyde-mediated


120

multipoint attachment of the enzyme to the primary amino groups at the surface of

the PAMAM hyperbranched structures capping the AuNPs-based electropolymerized

network. Such kind of multipoint covalent cross-linking trends to preserve the catalytic

activity of the enzyme by fixing the three dimensional active enzyme structure and by

avoiding the lack of tyrosinase molecules from the electrode surface.

EXPERIMENTAL

Reagents and apparatus

Tyrosinase (Tyr, 5370 U/mg), HAuCl4, NaBH4, 2-mercaptoethanesulfonic acid, p-

aminothiophenol and cystamine core PAMAM G-4 dendrimer were purchased from

Sigma (USA). All other chemicals were of analytical grade.

Dendron-coated gold nanoparticles were characterized by high resolution

transmission electron microscopy (HRTEM) using a JEOL JEM-4000 EX microscope. The

morphology of the nanostructured gold surface was studied by atomic force

microscopy (AFM) with a Nanotec Cervantes SPM microscope.

Cyclic voltammetry and electrochemical impedance spectroscopy experiments

were performed using a FRA2 µAutolab Type III potentiostat/galvanostat and the data

were acquired using GPES Ver. 4.9 and Frequency Response Analyser softwares,

respectively (Metrohm Autolab B.V., The Netherlands). Amperometric measurements

were performed with a dual-channel ultrasensitive Inbea potentiostat (Inbea

Biosensores S.L., Spain). A conventional three-electrode system was employed in all

electrochemical studies. The working electrode was a gold disk (CHI Instruments, UK,

2.0 mm diameter) modified with the electropolymerized AuNPs network and the

immobilized enzyme. An Ag/AgCl/KCl (3 M) and a Pt wire were used as reference and

counter electrodes, respectively. Bioelectrode measurements were carried out at 25ºC

in 0.1 M sodium phosphate buffer, pH 7.0 (working volume 10 ml). The solutions were

stirred at 300 rpm with a magnetic bar during amperometric measurements. For


121

analytical purposes, 100 µM catechol solutions in 50 mM sodium phosphate buffer, pH

7.0 were freshly prepared.

Synthesis of the dendron-coated AuNPs

To prepare the dendron-caped AuNPs, 150 mg of sodium borohydride, 18 mg of

2-mercaptoethanesulfonic acid, 250 mg of cystamine core PAMAM dendrimer G-4 and

5 mg of p-aminothiophenol were dissolved in 25 mL of de-aerated dimethyl sulfoxide

(DMSO) and stirred during 30 min. Another solution was prepared by dissolving 99 mg

of HAuCl4 in 50 mL of de-aerated DMSO, and rapidly added to the first reducing

mixture. The reaction mixture turned deep brown immediately, but the reaction was

allowed to proceed for 24 h under N2 atmosphere. Then, the functionalized AuNPs

were precipitated by adding 50 mL CH3CN, collected by centrifugation, and washed

with 50 mL CH3CN:DMSO (1:1 v/v), 50 mL ethanol and 50 mL diethyl ether. The

nanoparticles were finally isolated by centrifugation and dried under N2. The

nanoparticles were characterized by FT-IR and HRTEM.

Preparation of the electropolymerizad AuNPs-modified enzyme electrode

A clean gold disk electrode was first dipped into a 10 mM ethanolic solution of

p-aminothiophenol in order to modify the metal surface with a monolayer of the

aniline derivative. After 2 h incubation, the electrode was washed several times with

ethanol and double distilled water, and further dipped into a freshly prepared 2

mg/mL solution of PAMAM dendron-AuNPs in 0.1 M H2SO4. Electropolymerization was

performed by applying 10 potential cycles between -0.35 V and +0.85 V vs Ag/AgCl at a

scan rate of 100 mV/s, followed by application of a constant potential of 0.85 V for 1 h.

The electrode coated with the electropolymerized PAMAM dendron-AuNPs network

was exhaustively washed with 0.1 M H2SO4 and double distilled water before enzyme

immobilization.


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To functionalize the electrode with tyrosinase, a mixture of 20 µL of 5 mg/mL

enzyme solution in 50 mM sodium phosphate buffer, pH 7.0 and 10 µL of 25% (v/v)

glutaraldehyde was dropped on the electrode surface and kept at 4ºC during 2 h in wet

atmosphere. The enzyme-modified electrode was then exhaustively washed with cool

50 mM sodium phosphate buffer, pH 7.0 and finally kept at 4ºC in dry conditions until

used.

CONCLUSIONS

A novel nanostructured electrode surface, based on the modification of gold

substrates with a polymeric matrix of PAMAM G-4 dendron coated AuNPs, was

developed and used as support for the covalent immobilization of tyrosinase. The

nanostructured enzyme electrode was evaluated for biosensing catechol, showing

excellent analytical performance regarding to detection limit, sensitivity and stability.

Considering the experimental results here presented, we suggest that the covalent

immobilization of enzymes on metal electrodes coated with electropolymerized

network of dendron-modified metal nanoparticles constitutes an excellent strategy to

construct robust and reliable amperometric biosensors.

ACKNOWLEDGEMENTS






123

NOTES AND REFERENCES

aDepartment of Analytical Chemistry, Faculty of Chemistry, Complutense University of

Madrid, 28040-Madrid, Spain. Fax: 34 913944329; Tel: 34 913944315; E-mail:

[email protected]

bIMDEA Nanoscience, Campus Universitario de Cantoblanco, Avenida Francisco Tomás

y Valiente 7, 28049-Madrid, Spain. Fax: 34 914976855; Tel: 34 914976837; E-mail:

[email protected]

†This article is part of a web theme in Analyst and Analytical Methods on Future

Electroanalytical Developments, highlighting important developments and novel

applications. Also in this theme is work presented at the Eirelec 2011 meeting,

dedicated to Professor Malcolm Smyth on the occasion of his 60th birthday.

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7.3

ChemElectroChem. 1 (2014) 200-206

Biotin-labeled electropolymerized network of gold nanoparticles for amperometric immunodetection

of human fibrinogen

ChemElectroChem 1 (2014) 200-206 7. PUBLICACIONES

127

BIOTIN-LABELED ELECTROPOLYMERIZED NETWORK OF GOLD

NANOPARTICLES FOR AMPEROMETRIC IMMUNODETECTION OF HUMAN

FIBRINOGEN

Paula Díez,[a] María Gamella,[a] Paloma Martínez-Ruíz,[b] Valentina Lanzone,[c] Alfredo

Sánchez,[a] Enrique Sánchez,[a] Belit Garcinuño,[a] Reynaldo Villalonga,*[a,d] José M.

Pingarrón*[a,d]

[a] Department of Analytical Chemistry, Complutense University of Madrid, 28040-

Madrid, Spain. E-mail: [email protected], [email protected]

[b] Department of Organic Chemistry I, Complutense University of Madrid, 28040-

Madrid, Spain

[c] Department of Food Sciences, University of Teramo, 64100-Teramo, Italy

[d] IMDEA Nanoscience, Cantoblanco Universitary City, 28049-Madrid, Spain


128

ABSTRACT

Water soluble gold nanoparticles (3.1 ± 0.6 nm) with polymerization ability and

affinity to streptavidin were prepared by reducing HAuCl4 in the presence of the

capping ligands 2-mercaptoethanesulfonic acid, biotin-cysteamine and p-

aminothiophenol. This colloid was used to modify gold electrodes by formation of an

electropolymerized 3D network of bis-aniline cross-linked nanoparticles on the metal

surface. The modified electrode was employed as scaffold for the assembly of an

amperometric immunosensor system to detect human fibrinogen. The immunosensor

showed excellent analytical characteristics, with a dynamic range of detection

between 0.018 and 2.208 µg/mL, a detection limit of 4 ng/mL and an IC50 value of 177

ng/mL. The immunosensor was markedly stable, retaining full analytical capacity after

45 days of storage at 4ºC.

KEYWORDS: immunosensor, gold nanoparticles, fibrinogen, biotin, nanostructured

surface

INTRODUCTION

Electrochemical biosensors constitute promising analytical tools for the clinical

diagnosis of relevant diseases, as well as for the development of point-of-care sensing

systems able to permit real-time and remote health monitoring. In general, these

biomedical applications require highly sensitive, stable and specific biosensing

platforms able to detect very low concentrations of the target analyte in complex

biological samples.[1]

The advent of nanotechnology has favoured the construction of such biosensor

systems with improved analytical characteristics through the use of conducting

nanomaterials for the tailor made design of electrode surfaces with 3D architectures.[2]

The success of this strategy relies on the unique physicochemical properties of

nanomaterials such as high surface energy and surface-to-volume ratio, easy


129

functionalization, ability to decrease proteins-nanomaterial distance, and the

possibility to wire the redox center of some biomolecules with the electrode surface.[3]

Consequently, nanostructured electrodes can provide a large electroconductive

active surface area allowing high loadings of the biological recognition elements and

the proper occurrence of the electrocatalytic and electron transfer processes.

Gold nanoparticles (AuNPs) are, by far, the conducting nanomaterial more

widely employed for the modification of the transduction element in electrochemical

biosensors.[4] In addition to their high electroconductive and thermal properties, they

possess several advantages including simple preparation through different well-

established methods, relative low cost and easy stabilization of the colloidal dispersion

by covalent or non-covalent surface coating. Gold nanoparticles can catalyze the

transformation of the target analyte at their surface, then reducing the overpotential

and avoiding unwanted interference from other electroactive compounds. Gold

nanoparticles-modified surfaces can also work as nanoelectrode ensembles,

contributing to reduce the detection limit of analytes by increasing the ratio between

the faradaic and capacitive currents.

There are a great variety of strategies for assembling gold nanoparticles on

electrode surfaces for biosensing purposes, including electrodeposition, physical

adsorption, electrostatic interactions, covalent linkages or supramolecular

associations.[5] Recently, it has been reported an electrochemical approach to prepare

porous three-dimensional networks of polyfunctionalized gold nanoparticles capped

with aniline moieties by the formation of cross-linked bis-aniline bonds.[6] We have

employed this strategy for the preparation of nanostructured electrode surfaces with

predesigned properties for interfacing biological recognition events with excellent

electronic signal transduction for electrochemical biosensing.[7] The rational of this

strategy is based on the synthesis of gold nanoparticles capped with three different

thiol ligands, which should provide solubility, polymerization ability and chemical

functionality to favour the proper immobilization of the sensing biomolecule. These

systems have been successfully employed to construct enzyme biosensors with


130

improved analytical characteristics by covalent and supramolecular immobilization of

the biocatalyst.[7]

In this work, a novel approach to prepare an amperometric immunosensor for

human plasma fibrinogen by using electropolymerized biotin-labelled gold

nanoparticles modified electrodes is described. Plasma fibrinogen is an important

component of the coagulation cascade which determines blood flow and viscosity.[8]

Fibrinogen is cleaved by thrombin to form the fibrin monomer during coagulation,

which undergoes polymerization to form an insoluble fibrin clot, a complex lattice to

close off the injured blood vessel walls. The normal concentration of fibrinogen in

plasma is about 1.5 - 4.5 mg/ml, and elevated levels are associated with an increased

risk of cardiovascular disorders, such as stroke, ischemic heart disease and other

thromboembolisms.[9] Taking into account that cardiovascular diseases are the leading

cause of deaths worldwide, the development of analytical devices to detect fibrinogen

as a cardiac disease marker constitutes a relevant research topic in electrochemical

biosensors.

RESULTS AND DISCUSION

Scheme 1 illustrates the strategy employed to prepare the amperometric

immunosensor for fibrinogen. The rational of this electrochemical biosensor was based

on the use of a polymeric matrix of AuNPs as scaffold for the assembly of the

biorecognition architecture on the gold electrode surface. This nanomaterial has the

advantage to provide a large and 3D nanostructured surface with a high density of

biotin residues to be employed for the further immobilization of streptavidin

molecules in high yield. Streptavidin, which has the capacity to strongly bind four

residues of biotin per protein molecule, was then employed as multivalent affinity-

based linker to construct a sandwich-type assembly with biotinilated fibrinogen on the

modified surface.


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Scheme 1. Preparation of the immunosensor for fibrinogen on gold electrodes coated

with electropolymerized matrix of biotin-labelled AuNPs.

The functioning of the amperometric immunosensor was based on a

competitive immunorecognition assay, involving the initial biorecognition reaction

between the analyte fibrinogen and an excess of the specific anti-fibrinogen antibody,

which is labeled with the enzyme horseradish peroxidase (HRP). The non-reacted HRP-

labeled antibody was further incubated with the electrode modified with biotinilated

fibrinogen, and quantification was accomplished through the amperometric detection

of the HRP-catalyzed transformation of H2O2 in the presence of hydroquinone as

electrochemical mediator. Casein was employed as blocking protein in this

immunosensor architecture to avoid the occurence of non-specific adsorption of the

HRP-labelled anti-fibrinogen antibody on the electrode surface.

To construct the immunosensor for fibrinogen, polyfunctionalyzed AuNPs were

first synthesized by reducing AuCl4- ions with NaBH4 in a DMSO solution containing


132

three different thiol capping ligands. These ligands were rationally selected to confer

specific characteristics to the resulting nanoparticles. In this case, biotin-cysteamine

residues, resulting from the in situ reduction of biotin-cystamine,[10] acted as

recognition elements for the affinity-based immobilization of streptavidin molecules.

Moreover, p-aminothiophenol and 2-mercaptoethanesulfonic acid were included in

the design of the functionalized AuNPs to allow their electrochemical polymerization

and solubilisation, respectively.

Water soluble dark red nanoparticles with optimal solubility, polymerization

ability and biorecognition properties were prepared by using p-aminothiophenol,

biotin-cysteamine and 2-mercaptoethanesulfonic acid in a 1:1.7:3.7 molar ratio. This

optimal ratio of capping ligands was deduced by evaluation of the obtained water

soluble AuNPs providing the higher loading of streptavidin molecules on the electrode

surface upon electropolymerization. HR-TEM analysis revealed that these

nanoparticles showed spherical geometry with an average diameter of 3.1 ± 0.6 nm

and a narrow size distribution (Figure 1S in Supporting Information). Reduction of

AuCl4- ions in the presence of the thiol capping ligands caused their fast attachment to

the surface of the developing Au colloids, then resulting in the radial geometry, small

size and low polydispersion of the nanoparticles. Similar results were previously

reported for other AuNPs prepared by the same approach, but capped with different

water-soluble thiol ligands.[7,11]

The AuNPs mainly contain crystalline lattice planes with d-spacing of 2.35 Å for

adjacent lattice planes, corresponding to the (111) planes of face-centered cubic gold.

Selected area electron diffraction analysis showed the characteristic concentric rings

pattern of a face-centered cubic gold structure, then confirming the crystalline

structure of these colloids. Elemental analysis revealed that the AuNPs were capped

with an average of 16, 10 and 59 molecules of biotin-cysteamine, p-aminothiophenol

and 2-mercaptoethanesulfonic acid, respectively, assuming that the nanoparticles are

perfect spheres. This value corresponded to a total ligand surface coverage of 5.6

molecules/nm2, which is in agreement with the coverage range reported for gold

nanoparticles modified with different alkanethiols (4.3 – 6.3 molecules/nm2).[12]


133

The UV-Vis spectrum of the biotin-labelled AuNPs, dissolved in 50 mM sodium

phosphate buffer pH 7.0, showed a relatively weak plasmon resonance absorption

band around 515 nm (Figure 2S in Supporting Information). The intensity of this band

was not affected after addition of bovine serum albumin, although a little broadening

was observed, suggesting little interaction of this protein with the metal nanoparticles.

On the contrary, a noticeable increase in the intensity of the plasmon resonance band

was noticed when streptavidin was added to the nanoparticles solution, which could

be attributed to the strong association of this protein to the metal nanoparticles

surface. A similar effect was previously reported for other protein-AuNPs systems.[7a,13]

In this case, the affinity-based association of biotin-labelled AuNPs with streptavidin

was expected due to the ability of this tetrameric protein to bind four biotin residues

with a Ka ≈ 2.5x1013 M-1.[14]

Polyfunctionalized AuNPs were further electropolymerized in 0.1 M H2SO4 on a

p-aminothiophenol modified gold electrode. This previous modification of the

electrode surface with p-aminothiophenol moieties ensured the formation of bis-

aniline linkages between thioaniline groups on the electrode and AuNPs surfaces as

the first step for an efficient polymerization process. The formation of the 3D

electropolymerized matrix of AuNPs was performed through two sequential steps: the

formation of a bisaniline-cross-linked network of nanoparticles by means of cyclic

voltammetric scans between -0.35 V and +0.85 V vs Ag/AgCl reference electrode, and

the further growing of the polymer during 30 min at a fixed potential of +0.85 V.


134

Figure 1. Cyclic voltammograms recorded in the electropolymerization of the

polyfunctionalized AuNPs on gold electrode surface in 0.1 M H2SO4 at 100 mV/s.

Figure 1 shows the cyclic voltammograms recorded during the first

electropolymerization step. An anodic peak at +440 mV and two cathodic peaks about

+200 and +370 mV appeared in the successive voltamperograms. A noticeable

continuous peak growth was observed after repeated potential scanning. This

voltamperometric pattern, which is characteristic of the electrochemical formation of

dimeric and low molecular weight aniline condensation adducts,[15] suggested the

formation of bis-aniline cross-linked network of AuNPs on the electrode surface. It was

further demonstrated that the maximum loading of streptavidin on the

nanostructured electrode surface was achieved by performing a second

electropolymerization process over 30 min at a fixed potential of +0.85 V. Accordingly,

the nanostructured electrodes were prepared through to this experimental protocol.

It should be highlighted that this maximum streptavidin loading capacity, which

determines the successful assembly of the immunosensor and thus its analytical

performance, was directly related to the optimum coverage of the electrode surface

by the AuNPs-based polymer. This coverage, which was optimized by manipulation of

the electropolymerization time-interval, implies the formation of a non-compact but


135

nanoporous 3D matrix of AuNPs with high surface density of available biotin residues

but retaining high electroactive area. Such characteristics should favour the optimum

occurrence of both the biorecognition and the electron transfer processes at the

electrode surface.

Figure 2. FE-SEM (A) and AFM (B) analysis of gold surface coated with

electropolymerized matrix of biotin-labelled AuNPs.

Figure 2A shows a representative FE-SEM image of the electrode surface after

electropolymerization. As can be observed, the entire electrode surface was coated

with a three-dimensional matrix of small and spherically-shaped AuNPs distributed in a

non-ordered arrangement. The polymer-coated surface showed numerous AuNPs-

based protuberances and nanoholes, which provided a large surface area for

interfacing biological recognition of the pedant biotin moieties by streptavidin

molecules.

The topology of the electropolymerized AuNPs network was also characterized

by AFM (Figure 2B). This matrix showed nanoholes and AuNPs-based protuberant

structures representing 49.7% and 50.3% of the total surface area, respectively.

Nanostructured protuberances showed a Gaussian-type distribution for height with a

mean value of 61 nm, and a maximum height value of 183 nm (see Figure 3S in

Supporting Information). The roughness average of this modified surface was

calculated as 18.6 nm.


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The gold surface modified with this 3D matrix of AuNPs was used as scaffold for

the assembly of the immunosensing system to detect fibrinogen. As it is illustrated in

Scheme 1, optimized amounts of streptavidin (5 µg) and biotin-modified fibrinogen (4

µg) were sequentially immobilized on the biotin-labelled surface through affinity-

based interactions. Milk casein (0.5%, w/v) was also employed as coating protein to

minimize non-specific adsorptions on the electrode surface. Competition assays were

then performed between the surface-attached biotin-modified fibrinogen and

fibrinogen in solution for the HRP-labelled fibrinogen specific antibody. The affinity

recognition of the surface-bound biotin-modified fibrinogen by the HRP-labelled anti-

fibrinogen antibody was monitored by measuring the amperometric response of the

electrode toward H2O2 in the presence of hydroquinone as electrochemical mediator.

The optimal working conditions to obtain the largest amperometric responses of the

immunosensor are described in the Experimental section.

Figure 3. Nyquist plots obtained at a bare gold electrode (a) and at a gold electrode

sequentially modified with p-aminothiophenol (b), polymerized AuNPs (c), streptavidin

(d), biotinilated fibrinogen (e), casein (f) and HRP-labeled anti-fibrinogen antibody (g) in

0.1 M KCl solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1).

All the steps involved in the assembly of the electrode to construct the

immunosensor for fibrinogen were controlled by electrochemical impedance

spectroscopy (EIS) and cyclic voltammetry using 5 mM K4[Fe(CN)6]/K3[Fe(CN)6] as

electrochemical probe in 0.1 M KCl solution. Figure 3 shows the Nyquist plots for the


137

electrode at the different stages of assembling. A noticeable reduction in the charge

transfer resistance with respect to that observed at the bare gold electrode was

apparent for the p-aminothiophenol coated electrode which could be caused by the

electrostatic attraction of the [Fe(CN)6]4−/3− ions by the positive charged aniline

residues on the electrode surface.

Electropolymerization of polyfunctionalized AuNPs changed totally the shape of

the impedance spectrum, where two semicircles and one Warburg diffusion region

were observed. This fact could be ascribed to changes in the mass transport conditions

at the electrode interface due to the modification with AuNPs-based polymer. The high

surface coverage of AuNPs with thiolated ligands may reduce their metal conducting

properties and create a negatively charged interface on the electrode surface.

This perturbation can be represented by a modified Randles equivalent circuit,

in which the AuNPs-polymer/Au interface is represented by the double-layer

capacitance (Cdl) in parallel with a charge-transfer resistance (Rct) and the Warburg

diffusion impedance Zw. Additionally, the electrolyte/AuNPs-polymer interface is

represented by the parallel combination of another charge-transfer resistance (Rctp)

and capacitance (Cp) elements. Thus, the first semicircle at high frequencies is

indicative of the charge transfer process at the AuNPs-polymer/Au interface while the

second semicircle is associated to the charge transfer at the electrolyte/AuNPs-

polymer interface. Similar models were employed to explain the EIS behavior of other

polymer-modified electrodes.[16] It should be highlighted that this EIS behavior was

different than those previously described for gold electrodes modified with

electropolymerized networks of AuNPs labeled with mercaptophenylboronic acid and

polyamidoamine dendron moieties, which showed lower barrier properties than the

bare electrode and could be represented by a conventional Randles equivalent

circuit.[7a,c]


138

Figure 4. Cyclic voltammograms recorded in 0.1 M KCl solution containing 5 mM

K3[Fe(CN)6]/K4[Fe(CN)6] (1:1) at a bare gold electrode (a) and at a gold electrode

sequentially modified with p-aminothiophenol (b), polymerized AuNPs (c), streptavidin

(d), biotinilated fibrinogen (e), casein (f) and HRP-labeled anti-fibrinogen antibody (g).

Scan rate: 50 mV/s.

Incubation of the nanostructured electrode with streptavidin caused noticeable

increase in both semicircle diameters, indicating that streptavidin could recognize the

pedant biotin residues on the electrode surface through an affinity-based mechanism.

Sequential incubation with biotin-modified fibrinogen and casein also caused an

increase in the semicircle diameters and even the disappearance of the Warburg

diffusion region, indicating the sequential formation of insulating protein layers on the

electrode surface. Moreover, as expected, the incubation with the HRP-labeled anti-

fibrinogen antibody produced an additional increase in the charge transfer resistances

thus demonstrating the occurrence of the immunorecognition process on the

electrode surface.

These results were confirmed by cyclic voltammetry (Figure 4). Bare and p-

aminothiophenol coated gold electrodes showed well-defined typical diffusion-limited

quasi-reversible patterns. However, a significant decrease in the peak currents and a


139

much larger separation between the anodic and cathodic peaks was observed after

electropolymerization of the polyfunctionalized AuNPs on the electrode surface. This

suggested that the AuNPs-based polymer caused a kinetic barrier to the diffusion of

the electrochemical probes to the active electrode surface.

By using the Randles-Sevcik equation, the electrochemical surface area of the

gold electrode was estimated to change from 5.4 mm2 to 5.8 mm2 and 2.0 mm2 after

sequential modification with p-aminothiophenol and electropolymerized AuNPs,

respectively. Accordingly, about 62.9% of the electroactive surface area of the

electrode was coated with the AuNPs matrix.

These barrier effects were noticeably higher upon further modification of the

nanostructured electrode with the proteins involved in the immunosensor

construction due to the sequential formation of insulating protein layers on the

electrode surface. We estimated using the Randles-Sevcik equation that over 43.3% of

the AuNPs-modified surface was further coated with streptavidin, and about 11.3% of

this biofunctionalized interface was then covered with biotinilated fibrinogen. Cyclic

voltametric studies also revealed a coverage of about 45.1% by incubation with HRP-

labeled anti-fibrinogen antibody of the casein-modified electrode for the experiment

involving the minimun fibrinogen concentration in the competitive assay. This fact

suggested the succesful occurence of the immunochemical biorecognition process at

the electrode surface.

The assembled bioelectrode was used to detect fibrinogen through a

competitive configuration and amperometric measurements. Optimal conditions for

maxima amperometric signals were established by using 1 mM hydroquinone as

electrochemical mediator and a working potential of -100 mV vs Ag/AgCl.


140

Figure 5. Calibration curve for fibrinogen quantification using the electropolymerized

AuNPs-based immunosensor.

Figure 5 shows a typical calibration curve for fibrinogen, with the sigmoidal

shape characteristic of competitive immunoassays. The calibration curve was fitted to

the following four-parameter logistic equation:

Where imax and imin were the maximum and minimum current values of the

calibration curve, IC50 is defined as the fibrinogen concentration for a 50% competition,

and h is de Hill’s slope. The immunosensor showed a dynamic range of response

between 0.018 and 2.208 µg/mL, corresponding to the fibrinogen concentration

interval for which the amperometric signal was between 20% and 80% of the

maximum signal. The IC50 value calculated for this determination was 177 ng/mL.

The limit of detection (LOD) was estimated to be 4 ng/mL, and it was calculated

as the fibrinogen concentration for which the maximum amperometric signal was

reduced by 10%.[17] These analytical characteristics confirmed the suitability of this

amperometric immunosensor to be applied to the determination of human fibrinogen

whose normal plasma concentration is about 1.5 - 4.5 mg/ml.[8.9] It should be also


141

highlighted that the immunosensor system showed a noticeable stability, being able to

retain full capacity to detect fibrinogen after 45 days of storage at 4ºC.

CONCLUSIONS

A novel nanostructured electrode surface, based on the modification of gold

electrodes with a three-dimensional polymeric matrix of aniline, sulfonic acid and

biotin-functionalized AuNPs, is here described. This modified surface showed affinity-

based biorecognition properties for streptavidin, and was evaluated for the

construction of an amperometric immunosensor for human fibrinogen. The

immunosensor exhibited excellent analytical characteristics in terms of dynamic range

of response, limit of detection, IC50 value and stability.

According to these results, we propose the use of bis-aniline cross-linked

networks of biotin-labeled AuNPs as scaffolds for the assembly of biomolecular

building blocks on electrode surfaces via biotin-streptavidin affinity interactions, for

the successful design of electrochemical biosensors.

EXPERIMENTAL SECTION

Reagents and apparatus

Human fibrinogen, biotin, horseradish peroxidase (HRP) labeled anti-fibrinogen

antibody, HAuCl4, NaBH4, 2-mercaptoethanesulfonic acid, p-aminothiophenol and

biotin-NHS were purchased from Sigma (USA). All other chemicals were of analytical

grade.

To prepare the biotinilated fibrinogen derivative, the protein (1 mg/mL in PBS)

was mixed with 20-fold molar excess of biotin-NHS, re-constituted at 25 mg/mL in

dimethylsulfoxide. After 2 h incubation at room temperature, the reaction was

stopped by addition of TrisHCl pH 7.5 at a final concentration of 50 mM and the

samples were placed on ice for 1 h. The reaction mixture was loaded onto a Sephadex


142

G25 PD-10 column, and the biotinylated protein was eluted in 500 μL fractions with

PBS. Protein concentration was estimated using the bicinchoninic acid protein assay.

Cyclic voltammetry and EIS experiments were performed using a

FRA2 µAutolab Type III potentiostat/galvanostat (Metrohm Autolab B.V., The

Netherlands). Amperometric measurements were performed with an Inbea

potentiostat (Inbea Biosensores S.L., Spain). A conventional three-electrode system

was employed in all electrochemical studies. The working electrode was a gold disk

(CHI Instruments, UK, 2.0 mm diameter) modified with the electropolymerized AuNPs

network. An Ag/AgCl/KCl (3 M) and a Pt wire were used as reference and counter

electrodes, respectively.

The morphology of the nanostructured electrode surface was characterized by

high resolution field emission scanning electron microscopy (FE-SEM) with a JEOL JSM-

6335F electron microscope (JEOL Ltd., Japan), and atomic force microscopy (AFM) with

a SPM Nanoscope IIIa multimode microscope (Veeco Instruments Inc., USA). High

resolution transmission electron microscopy (HR-TEM) measurements were performed

with a JEOL JEM-3000 F microscope. Spectrophotometric determinations were

performed using an Agilent 8453 UV/VIS spectrophotometer (Hewlett Packard, USA).

Synthesis of biotin-labeled AuNPs

Biotin-cystamine was synthesized as described earlier.[10] The

polyfunctionalized AuNPs were prepared by modification of our previously published

method,[7] by first dissolving 197 mg of HAuCl4 in 50 mL of de-aerated dimethyl

sulfoxide (DMSO). This solution was added dropwise to 50 mL of de-aerated DMSO

containing 100 mg of sodium borohydride, 36 mg 2-mercaptoethanesulfonic acid, 21

mg of biotin-cystamine and 8 mg p-aminothiophenol under vigorous stirring. The

reaction mixture turned deep brown immediately, but the reaction was allowed to

continue for 24 h. The functionalized AuNPs were then precipitated by adding 100 mL

CH3CN, collected by centrifugation, and washed with 60 mL CH3CN:DMSO (1:1 v/v), 60

mL ethanol and 20 mL diethyl ether. The nanoparticles were finally isolated by


143

centrifugation and dried under N2. The nanoparticles were characterized by HR-TEM,

UV-vis spectroscopy and elemental analysis.

Preparation of the AuNPs-modified electrode for fibrinogen detection

A clean gold disk electrode was first capped with a monolayer of p-

aminothiophenol by dipping the electrode for 2 h in a 10 mM ethanolic solution of the

thiol. The electrode was exhaustively washed with ethanol and double distilled water,

and dipped into a 2 mg/mL solution of the AuNPs in 0.1 M H2SO4. The electrode

surface was then coated with the electropolymerized bisaniline-cross-linked AuNPs

network by application of 10 potential cycles between -0.35 and +0.85 V vs Ag/AgCl at

a scan rate of 100 mV/s, followed by application of a constant potential of 0.85 for 30

min. The modified electrode was further exhaustively washed with double distilled

water.

The electrode was then coated with 10 µL of 0.5 mg/mL streptavidin solution in

B&W buffer, and incubated during 1 h at 37ºC. After exhaustive washing with B&W

buffer, 10 µL of 0.4 mg/mL biotinilated fibrinogen solution were dropped on the

electrode surface, and further incubated during 1 h at 37ºC. The electrode was then

washed with B&W buffer, further coated with 10 µL of 0.5 % (w/v) casein solution,

incubated for 30 min incubation at room temperature, and finally washed with

phosphate buffered saline (PBS) solution.

Fibrinogen was detected through a competitive assay by mixing 10 µg/mL of

HRP-labeled anti-fibrinogen antibody with fibrinogen at variable concentrations in PBS

solutions made 1 M NaCl. The electrode surface was incubated with 10 µL of this

mixture during 30 min at room temperature, then washed with PBS solution, and

finally tested for amperometric quantification of immobilized HRP activity. To do that,

the modified electrode was dipped into an electrochemical cell containing 10 mL of a

stirred solution of 1 mM hydroquinone in 0.1 M sodium phosphate buffer, pH 6.0.

Changes in the cathodic current were measured at -100 mV vs Ag/AgCl after addition

of 25 µL of a freshly prepared 100 mM H2O2 solution.


144

ACKNOWLEDGEMENTS

R. Villalonga acknowledges to Ramón & Cajal contract from the Spanish

Ministry of Science and Innovation. Financial support from the Spanish Ministry of

Science and Innovation (CTQ2011-24355 and CTQ2012-34238), and Comunidad de

Madrid S2009/PPQ-1642, Programme AVANSENS is gratefully acknowledged.

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ACS Appl. Mat. Interfaces 2012, 4, 4312-4319; c) R. Villalonga, P. Díez, S. Casado,

M. Eguílaz, P. Yáñez-Sedeño, J.M. Pingarrón, Analyst 2012, 137, 342-348.

[8] G.D. Lowe, A. Rumley, I.J. Mackie, Ann. Clin. Biochem. 2004, 41, 430-440.

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SUPPORTING INFORMATION

Biotin-labeled electropolymerized network of gold nanoparticles for

amperometric immunodetection of human fibrinogen

Paula Díez,[a] María Gamella,[a] Paloma Martínez-Ruíz,[b] Valentina Lanzone,[c] Alfredo

Sánchez,[a] Enrique Sánchez,[a] Belit Garcinuño,[a] Reynaldo Villalonga,*[a,d] José M.

Pingarrón*[a,d]

[a] Department of Analytical Chemistry, Complutense University of Madrid, 28040-Madrid, Spain

[b] Department of Organic Chemistry I, Complutense University of Madrid, 28040-Madrid, Spain

[c] Department of Food Sciences, University of Teramo, 64100-Teramo, Italy [d] IMDEA Nanoscience, Cantoblanco Universitary City, 28049-Madrid, Spain

Figure 1S. A) HR-TEM image, B) size distribution and C) selected area electron

diffraction analysis of biotin-labeled gold nanoparticles.


147

Figure 2S. UV-vis spectra of 0.2 mg/mL solution of functionalized AuNPs in 50 mM

sodium phosphate buffer, pH 7.0 in the absence (a) and the presence of 0.5 mg/mL BSA

(b) and streptavidin (c).

Figure 3S. Height histogram of the gold surface coated with electropolymerized matrix

of biotin-labeled AuNPs.

7.4

ACS Applied Materials & Interfaces 4 (2012) 4312-4319

Supramolecular immobilization of xanthine oxidase on

electropolymerized matrix of functionalized hybrid gold

nanoparticles/single-walled carbon nanotubes for the preparation of

electrochemical biosensors

ACS Applied Materials & Interfaces 4 (2012) 4312-4319 7. PUBLICACIONES

149

SUPRAMOLECULAR IMMOBILIZATION OF XANTHINE OXIDASE ON

ELECTROPOLYMERIZED MATRIX OF FUNCTIONALIZED HYBRID GOLD

NANOPARTICLES/SINGLE-WALLED CARBON NANOTUBES FOR THE

PREPARATION OF ELECTROCHEMICAL BIOSENSORS

Reynaldo Villalonga,1,2,* Paula Díez,1 Marcos Eguílaz,1 Paloma Martínez,3 José M.

Pingarrón1,2,*

1Department of Analytical Chemistry & 3Department of Organic Chemistry I, Faculty of

Chemistry, Complutense University of Madrid, 28040-Madrid, Spain.

2IMDEA Nanoscience, Cantoblanco Universitary City, 28049-Madrid, Spain

*Corresponding authors. Phone: +34 91 3944315, Fax: +34 91 3944329,

E-mail: [email protected], [email protected]



150

ABSTRACT

Glassy carbon electrodes modified with single walled carbon nanotubes and a

three-dimensional network of electropolymerized Au nanoparticles capped with 2-

mercaptoethanesulfonic acid, p-aminothiophenol and 1-adamantanethiol were used

as hybrid electrochemical platforms for supramolecular immobilization of a

synthesized artificial neoglycoenzyme of xanthine oxidase and β-cyclodextrin through

host-guest interactions. The ensemble was further employed for the

bioelectrochemical determination of xanthine. The biosensor showed fast

amperometric response within 5 s and a linear behavior in the 50 nM – 9.5 µM

xanthine concentration range with high sensitivity, 2.47 A/M·cm2, and very low

detection limit of 40 nM. The stability of the biosensor was significantly improved and

the interferences caused by ascorbic and uric acids were noticeably minimized by

coating the electrode surface with a Nafion thin film.

KEYWORDS: Single walled carbon nanotube, gold nanoparticle, supramolecular

assembly, enzyme biosensor, cyclodextrin

INTRODUCTION

Electrochemical biosensors constitute one of the most rapidly evolving fields in

Chemistry. From an analytical point of view, they allow the design of portable and

affordable sensing devices exhibiting high sensitivity and the characteristic specificity

provided by biological recognition systems.1,2 The successful development of this

technology is linked to the rational design of novel electrode surfaces able to improve

the speed of the electrochemical processes associated with the analytical responses as

well as the stable immobilization of the biomolecules preserving their three-

dimensional active conformation. In this context, electroconductive nanosized

materials have been exhaustively employed to prepare novel nanostructured surfaces

with improved performance.3-7 This success relies on the unique properties of


151

nanomaterials such as high surface energy and surface-to-volume ratio, ability to

decrease proteins-nanomaterial distance, thermal stability, easy functionalization and

the possibility to act as electroconductive wires between the electrodes and the redox

centers in some biomolecules.8 In particular, gold nanoparticles (AuNPs) and carbon

nanotubes (CNTs) have been at the central core of novel advanced materials widely

used in biosensor technology.6,8-11 Furthermore, the combination of these

nanomaterials to prepare synergic hybrid nanomaterials has demonstrated to

constitute a successful strategy to construct electrochemical interfaces with unique

properties for the preparation of more efficient and robust biosensors.12

The synthesis of novel polyfunctionalized gold nanoparticles, bearing aniline

moieties able to be electropolymerized to form electroconductive three-dimensional

networks on electrodes, as well as their use for the preparation of molecularly

imprinted surfaces suitable for the construction of SPR biosensors, has been recently

reported.13,14 Furthermore, our group demonstrated recently the usefulness of such

electropolymerized nanostructures for the construction of enzyme electrochemical

biosensors. In this context, the preparation of metal surfaces nanostructured with

networks of Au nanoparticles functionalized with boronic acid and PAMAM dendron

moieties to design amperometric biosensors toward hydrogen peroxide and cathecol,

respectively, were reported.15,16 However, to our knowledge, the combined use of

these electropolymerized nanoparticle networks with other nanomaterials has not

been evaluated until now.

On the other hand, the use of host-guest supramolecular interactions was

previously proposed as a soft, reversible and multivalence method for the

immobilization of enzymes on metal nanoparticles and surfaces taking advantage of

the complementary interaction of adamantane derivatives with β-cyclodextrin (CD).17-

19 This strategy was successfully employed to construct stable enzyme biosensors.20-22

This work describes a novel approach for the construction of a nanostructured

electrode surface by electropolymerization of polyfunctionalized Au nanoparticles on a

glassy carbon electrode (GCE) which was previously coated with single-walled carbon

nanotubes (SWNT). For this purpose, Au nanoparticles modified with 2-


152

mercaptoethanesulfonic acid, 1-adamantanethiol and p-aminothiophenol were

synthesized and further electropolymerized on the SWNT-coated GCE through the

formation of a bisaniline-cross-linked network. The presence of pendant adamantane

residues in this electroconductive matrix allows the subsequent supramolecular

immobilization of CD-based neoglycoproteins by formation of the complementary

host-guest inclusion complexes. As a model system to demonstrate this proof of

concept, we prepared a CD-modified xanthine oxidase (XO, EC 1.17.3.2) derivative

which was employed as the target neoglycoprotein. Moreover, an electrochemical

enzyme biosensor for xanthine based on this supramolecular design was constructed

and evaluated.


1. Reagents

Xanthine oxidase (Type III, 1.3 U/mg), HAuCl4, NaBH4, 2-

mercaptoethanesulfonic acid, p-aminothiophenol, 1-adamantanethiol and CD were

purchased from Sigma-Aldrich Co. (USA). SWNT were from Wako Pure Chemical

Industries, Ltd. (Japan). All other chemicals were of analytical grade.

2. Apparatus and electrodes

Electrochemical impedance spectroscopy and cyclic voltammetry were

performed using a FRA2 µAutolab Type III potentiostat/galvanostat and data were

acquired using Frequency Response Analyser and GPES Ver. 4.9 software, respectively

(Metrohm Autolab B.V., The Netherlands). Amperometric measurements were carried

out with a dual-channel ultrasensitive InBea potentiostat (InBea Biosensores S.L.,

Spain). A conventional three-electrode system was employed for all electrochemical

measurements. The working electrode was a glassy carbon electrode (GCE, 3.0 mm

diameter) coated with SWNT, the electropolymerized network of Au nanoparticles and

the immobilized CD-enzyme derivative (XO-CD/pAuNP/SWNT/GCE). This electrode


153

coated with a Nafion thin film (Naf/XO-CD/pAuNP/SWNT/GCE) was evaluated for

biosensing xanthine. An Ag/AgCl/KCl (3 M) and a Pt wire were used as reference and

counter electrodes, respectively.

High resolution transmission electron microscopy (HR-TEM) measurements

were performed with a JEOL JEM-3000 F microscope. The surface morphology of the

electrode surface was characterized using high resolution field emission scanning

electron microscopy (FE-SEM) with a JEOL JSM-6335F electron microscope (JEOL Ltd.,

Japan). FT-IR spectra were acquired with a Perkin-Elmer instrument.

Spectrophotometric measurements were performed using an Agilent 8453 UV/VIS

spectrophotometer (Hewlett Packard, USA).

3. Electrochemical measurements

All measurements with the prepared bioelectrodes were carried out at 25ºC in

0.1 M sodium phosphate buffer, pH 7.0 (working volume 10 mL). The solution was

stirred at 300 rpm with a magnetic bar during amperometric measurements. 100 µM

xanthine solutions in 50 mM sodium phosphate buffer, pH 7.0 were freshly prepared.

4. Preparation of nanomaterials

To prepare the adamantane-caped Au nanoparticles, 197 mg of HAuCl4 were

dissolved in 50 mL deaerated DMSO. This solution was added dropwise to 50 mL

deaerated DMSO containing 284 mg NaBH4, 36 mg 2-mercaptoethanesulfonic acid, 11

mg 1-adamantanethiol and 8 mg p-aminothiophenol under vigorous stirring. The

reaction mixture turned deep brown immediately, but the reaction was allowed

proceeding for 24 h. The polyfunctionalized nanoparticles prepared in this way were

precipitated by adding 50 mL CH3CN, collected by centrifugation and washed with 50

mL CH3CN:DMSO (1:1 v/v), 50 mL ethanol and 50 mL diethyl ether. The nanoparticles

were finally isolated by centrifugation and dried under N2.


154

SWNT were purified and partially oxidized by treatment with a mixture of

HNO3/H2SO4 (3:1, v/v) during 2 h in an ultrasonic bath. The treated nanotubes were

centrifuged, washed with double distilled water until neutral pH and finally dried in

vacuum under P2O5.

These nanomaterials were characterized by FT-IR and HR-TEM.

5. Preparation of the enzyme electrode

The mono-6-ethylenediamino-6-deoxy-β-cyclodextrin was obtained by treating

the corresponding mono-6-O-tosyl derivative23 with freshly distilled ethylenediamine

as reported previously.24 XO was covalently glycosylated with this CD derivative via a

carbodiimide-catalyzed reaction (XO-CD) as previously described.22 The modified

enzyme contained an average of 12 mol of CD residues attached to each mol of

protein, as determined by the the phenol-sulfuric acid assay.

To prepare the enzyme-modified electrode, a bare GCE was polished to mirror-

like surface with alumina powder (0.3 µm), rinsed thoroughly with double distilled

water, successively washed with double distilled water, anhydrous ethanol and

acetone in an ultrasonic bath, and dried under N2 before use.

Coating of the treated GCE was accomplished by depositing two 10-µL aliquots

of a 0.4 mg/mL aqueous dispersion of SWNT on the electrode surface and allowing

drying. The SWNT-modified electrode was then dipped into a 2 mg/mL Au

nanoparticles solution in 0.1 M H2SO4 and electropolymerization was accomplished by

cycling the potential 10 times between -0.35 and +0.85 V vs Ag/AgCl at a scan rate of

100 mV/s, followed by applying a constant potential of 0.85 V for 1 h. The electrode

modified with the electropolymerized bisaniline-cross-linked network of Au

nanoparticles (pAuNP/SWNT/GCE) was exhaustively washed with 0.1 M H2SO4 and

double distilled water before enzyme immobilization.

Subsequently, the pAuNP/SWNT/GCE was dipped into a 5 mg/mL XO-CD

solution in 50 mM sodium phosphate buffer, pH 7.0, for 4 h. The enzyme-modified

electrode (XO-CD/pAuNP/SWNT/GCE) was finally washed with the same cool buffer,


155

dried under nitrogen and kept at 4ºC until use. A Nafion thin film-coated enzyme

electrode (Naf/XO-CD/pAuNP/SWNT/GCE) was also prepared by dropping 5.0 µl of a

0.5% (v/v) Nafion ethanolic solution on the surface of the XO-CD/pAuNP/SWNT/GCE

and dried, washed and stored as described above.


Preparation and characterization of the nanostructured bioelectrode

The steps involved in the preparation of the hybrid nanostructured enzyme

electrode are schematized in Figure 1. Polyfunctionalyzed AuNPs were first

synthesized by reducing AuCl4- ions with NaBH4 in a DMSO solution containing 1-

adamantanethiol, 2-mercaptoethanesulfonic acid and p-aminothiophenol in a 1.3:3.4:1

molar ratio. Dark red and water soluble nanoparticles were obtained by applying this

procedure.

Figure 1. Schematic display of the steps involved in the preparation of XO-

CD/pAuNP/SWNT/GCE enzyme biosensors.

HR-TEM analysis of the obtained nanoparticles showed spherical geometry with

an average diameter of 4.9 ± 0.8 nm (see supporting information) which is similar to


156

the size reported for other polyfunctionalized Au nanoparticles bearing other capping

ligands.15,16 19

4000 3000 2000 1000

85

90

95

100

S-O

cm-1

cm-1

C(aliph)

cm-1

S=O

C=C

CH2

cm-1

cm-1

Tra

nsm

itta

nce

(%

)

Wavenumbers (cm-1)

cm-1

Figure 2. FT-IR spectrum of the polyfunctionalized Au nanoparticles.

Figure 2 shows an FT-IR spectrum of the polyfunctionalyzed nanoparticles. The

presence of aniline residues in the nanomaterial was revealed by the broad and

intense band around 3460 cm-1, corresponding to the stretch vibration of the N-H

bonds. In addition, the presence of aniline moieties was confirmed by the bands at

1630 cm-1 and 1335 cm-1, corresponding to the N-H blending and C-N stretching,

respectively. This latter band is partially overlapped with the large band appearing in

the range of 1433 to 1385 cm-1, which can be ascribed to the overlapped S=O and C=C

stretching vibrations and CH2 blending, suggesting the presence of sulfonate, aromatic

and aliphatic groups in the modified nanoparticles, respectively. The presence of

sufonate residues can be also supported by the intense stretching vibration band of

the S-O bonds at 1040 cm-1, as well as by the stretching of aliphatic C-H bonds at 2850

cm-1. On the other hand, adamantane residues were confirmed by the C-H stretching

vibration at 2928 cm-1 revealing the presence of a cyclic aliphatic compound on the

nanoparticles surface. Finally, the formation of a covalent Au-S bond is supported by


157

the presence of the large band in the range of 500 to 750 cm-1 that can be assigned to

stretch the mode of C-S groups.25

It should be highlighted that the employed synthetic approach ensured the

preparation of Au nanoparticles with the appropriate characteristics allowing their use

as structural units for the electrochemical formation of the nanostructured three-

dimensional matrix. In fact, the growing of the metal colloid in the mixture containing

the thiol derivatives provided the formation of small and spherical capped

nanoparticles. Furthermore, it should be mentioned that the used thiol derivatives

were specifically selected to confer some relevant properties to the synthesized Au

nanoparticles: solubility (2-mercaptoethanesulfonic acid), polimerization ability (p-

aminothiophenol) and capability to form inclusion complexes with CDs (1-

adamantanethiol).

200 400 600 800

Ab

sorb

an

ce

(nm)

C

A

2.4

1.6

0.8

Figure 3. UV-vis spectra of 0.4 mg/mL solution of Au nanoparticles in 50 mM sodium

phosphate buffer, pH 7.0 without (A) and with 0.5 mg/mL XO (B) and XO-CD (C).

The formation of host-guest complexes on the Au nanoparticle surface with the

CD-modified xanthine oxidase derivative was confirmed by UV/VIS spectrophotometry.

Figure 3 shows the spectra of the polyfunctionalized AuNPs dissolved in 50 mM sodium

phosphate buffer pH 7.0 where the occurrence of a plasmon resonance absorption


158

band with a maximum around 550 nm can be observed. The intensity of this band was

slightly affected after addition of native XO, probably due to the electrostatic

interaction of the enzyme with the nanoparticles surface. However, the intensity of the

plasmon resonance band exhibited a significant increase when XO-CD was added to

the nanoparticle solution and the maximum slightly shifted to around 552 nm. This

effect could be related to the association of the modified enzyme to the metal

nanoparticles surface through the formation of inclusion complexes between the

complementary adamantane and CD residues. Similar plasmonic effects were

previously reported for CD-coated Au nanoparticles after association with

adamantane-modified enzymes.19

-400 -200 0 200 400 600 800

-90

-60

-30

0

30

60E

CB

D

i (µ

A)

E (mV)

A

Figure 4. Cyclic voltammograms recorded in 0.1 M H2SO4 at a bare GCE (A), a SWNT-

modified GCE (B), and upon electropolymerization of Au nanoparticles for five (C) and

ten (D) potential cycles, and after polymer growing for 1 h at +850 mV (E). Scan rate:

50 mV/s

Polyfunctionalized Au nanoparticles were electropolymerized on the surface of

SWNT-coated GCE through two sequential steps13-16. These steps involved firstly the

formation of a bisaniline-cross-linked network of nanoparticles by performing cyclic

voltammetric scans between -0.35 V and +0.85 V in 0.1 M H2SO4, and a further growing


159

of the Au nanoparticle-based polymer for 1 h at a constant potential of +0.85 V. Figure

4 shows the cyclic voltammograms recorded in 0.1 M H2SO4 solution upon the

different electrode modification steps. As expected, a remarkable larger background

current was observed at the SWNT-modified electrode in comparison with the bare

GCE, which can be attributed to the much larger active surface area of nanotubes

electrically connected to the electrode surface.26 The repeated potential scanning in

the Au nanoparticle solution produced a noticeable increase in the background current

as a consequence of the increase in the electrochemical surface area caused by the

generation of the electropolymerized network of metal nanoparticles on the electrode

surface.

Figure 5. Field emission SEM image of the pAuNP/SWNT/GCE.

In addition, it was observed the appearance of an anodic peak at +0.380 V and

two cathodic peaks around +0.300 V and -0.130 V. The current intensity of these peaks

increased when the number of cycles increased, indicating the electrochemical

formation of bis-aniline condensation adducts.27 The current intensity of these peaks,

as well as the background current in the cyclic voltammogram were larger after the

second step of electropolymerization performed during one hour at a fixed potential of

+ 0.85 V (voltammogram E), indicating the growing of a dense three-dimensional


160

electroconductive assembly of Au nanoparticles on the electrode surface via bis-aniline

linkages.

Figure 5 shows the FE-SEM image of the electrode surface after the

electropolymerization processes. A three-dimensional nanostructured network of

small and spherical Au nanoparticles covering the SWNT-coated surface can be

observed. It should be noted that the electropolymerized matrix was composed of a

non-ordered array of AuNPs-based protuberances of different heights, separated by

irregularly distributed nanoholes on the SWNT-coated GCE surface. This topology

ensures a high functionalized metal surface area on the electrode to favor the further

immobilization of the enzyme derivative.

Figure 6. Nyquist plots of bare GCE (), SWNT/GCE (), pAuNP/SWNT/GCE (X) and XO-

CD/pAuNP/SWNT/GCE () in 0.1 M KCl solution containing 5 mM

K3[Fe(CN)6]/K4[Fe(CN)6] (1:1).

In order to evaluate the potential bioanalytical use of this novel nanostructured

electrode surface, we employed a XO-CD neoglycoenzyme as a probe of concept for

constructing a biosensor device toward xanthine. Electrochemical impedance

spectroscopy was performed in a 5 mM [Fe(CN)6]4−/3− solution in 0.1 M KCl to evaluate

the interfacial changes occurring during the different functionalization steps. As it can


161

be seen in Figure 6, a noticeable decrease in the semicircle diameter of the Nyquist

plot occurred after modification of the bare GCE with SWNTs and the subsequent

electropolymerization of Au nanoparticles, indicating the expected increase in the

electron transfer rate between the redox probe and the nanostructured electrode

surface. However, immobilization of XO-CD caused a significant insulating effect on the

electrode surface with a remarkable increase in the Ret value for the enzyme-modified

electrode.

-200 0 200 400 600

-300

-150

0

150

300

i (µ

A)

E (mV)

A

B

C

D


K3[Fe(CN)6]/K4[Fe(CN)6] (1:1) at bare GCE (A), SWNT/GCE (B), pAuNP/SWNT/GCE (C)

and XO-CD/pAuNP/SWNT/GCE (D). Scan rate: 50 mV/s.

This effect suggested a high enzyme loading on the electrode surface which is a

desired attribute for biosensing purposes. These results were confirmed by cyclic

voltammentry in the same working solution for the different electrode architectures

(Figure 7). Bare, SWNT and Au nanoparticles-modified GCE showed well-defined

typical quasi-reversible diffusion-limited patterns with ∆E values of 90 mV, 103 mV and

101 mV and ia/ic ratios of 0.99, 1.03 and 1.01, respectively. Moreover, coating with

SWNT and further electropolymerization of the metal nanoparticles increased the

electrochemical surface area of the GCE, as revealed by the higher peak currents


162

obtained. By using the Randles-Sevcik model, the electrochemical surface area of the

GCE was estimated to increase from 7.1 mm2 to 9.7 mm2 and 11.1 mm2 after

sequential modification with SWNT and electropolymerized Au nanoparticles,

respectively. This fact can be justified by the electroconductive nature of both the

SWNT and the bisaniline cross-linked Au nanoparticle-based matrix.

Conversely, a larger separation between the anodic and cathodic peaks, ∆E =

188 mV, and a noticeable decrease in the peak currents with ia/ic = 0.93 was produced

after the modified electrode incubation in the XO-CD solution, indicating that the

immobilized enzyme behaved as a kinetic barrier. A noticeable reduction to 3.9 mm2

for the electroactive surface area was also observed at the enzyme-modified

electrode, suggesting a high surface coverage with the XO-CD derivative in agreement

with the results obtained by electrochemical impedance spectroscopy.

Figure 8A displays the cyclic voltammograms recorded at pAuNP/SWNT/GCE

functionalized with native (curve b) and CD-modified XO (curve a) in 0.1 M sodium

phosphate buffer, pH 7.0, aerated solutions at a scan rate of 50 mV/s. The anodic

current significantly increased at both modified electrodes after addition of 200 µM

xanthine, showing a peak current at +0.69 V corresponding to the enzymatic

transformation of xanthine at the electrode surfaces and the further oxidation of the

produced H2O2.

0 100 200 300 400 500 600 700

0

20

40

60

80d

c

b

i (µ

A)

E (V)

a

A


163

0 100 200 300 400 500 600 700

0

20

40

60

80

g

e

ci (µ

A)

E (mV)

f

B

Figure 8. Cyclic voltammograms recorded in 0.1 M sodium phosphate buffer, pH 7.0, at

a scan rate of 50 mV/s for: A) XO-CD/pAuNP/SWNT/GCE and XO/pAuNP/SWNT/GCE

without addition of xanthine (curves (a) and (b), respectively) and after addition of 200

µM xanthine (curves (d) and (c), respectively). B) XO-CD/pAuNP/SWNT/GCE and

XO/pAuNP/SWNT/GCE in the presence of 200 µM xanthine before (curves (e) and (c),

respectively) and after 1 h incubation in saturated 1-adamantane carboxylic acid

solution (curves (g) and (f), respectively).

However, the anodic peak current was 1.7-fold higher at the XO-

CD/pAuNP/SWNT/GCE (curve d) in comparison with that recorded at the electrode

modified with native XO (curve c), suggesting a higher active enzyme loading on the

former bioelectrode. This fact should be related to the occurrence of host-guest

interactions between the adamantane units at the AuNPs surface and the CD moieties

attached to the enzyme.

The contribution of these supramolecular interactions to the overall

immobilization process for XO-CD on pAuNP/SWNT/GCE was evaluated by recording a

cyclic voltammogram in a 200 µM xanthine solution after one hour incubation of the

enzyme electrode at 4ºC in 0.1 M sodium phosphate buffer pH 7.0 saturated with 1-

adamantane carboxylic acid. Since CDs form highly stable inclusion complexes with

adamantane derivatives,28 it was foreseen that the presence of saturated 1-


164

adamantane carboxylic acid in the incubation medium could compete with the

multiple host-guest interactions between the immobilized enzyme and the

adamantane-coated Au nanoparticles electropolymerized network at the GCE surface.

As a control, a pAuNP/SWNT/GCE modified with native XO was also checked under the

same conditions.

Figure 8B shows as the peak current measured at +0.69 V did not significantly

change for the XO/pAuNP/SWNT/GCE after 1 hour incubation in the 1-adamantane

carboxylic acid solution, suggesting that the activity of the native enzyme was not

affected by the incubation process.22, 29 However, a noticeable decrease in the peak

current was observed with the XO-CD/pAuNP/SWNT/GCE after incubation, suggesting

lower amount of enzyme activity at the electrode surface. This effect could be caused

by the competitive interaction of 1-adamantane carboxylic acid with the CD moieties

attached to the enzyme surface, then affecting the supramolecular association of the

enzyme with the adamantane moieties at the nanoparticles and releasing certain

amount of enzyme molecules from the electrode surface. However, further studies

should be performed to demonstrate this hypothesis.

Nevertheless, it should be stated that there are other forces that should also

contribute to the immobilization of XO-CD on the surface of pAuNP/SWNT/GCE

besides the commented host-guest supramolecular interactions. The occurrence of

physical adsorption of the enzyme derivative on the exposed SWNT-coated surface as

well as electrostatic interactions with the charged groups at the surface of the

nanoparticle-based matrix can play also a role in the overall enzyme immobilization

process.

Analytical performance of the nanostructured enzyme electrode

The bioelectrode prepared through the supramolecular approach was used to

construct an amperometric enzyme biosensor toward xanthine. The values to be

selected for the working pH and applied potential were determined by checking the

steady-state current and the signal-to- noise ratio measured for 500 nM xanthine.


165

According to the obtained results (not shown), pH 7.0 and a detection potential value

of +650 mV vs Ag/AgCl were selected for further work. It is well known that the use of

such a relatively high working potential would yield remarkable interferences from

other electrochemically oxidizable substances such as ascorbic and uric acids. In order

to minimize such adverse effects, the use of a Nafion thin film on the electrode surface

was also evaluated12 which would also serve to test the behavior of this well-known

strategy with complex nanostructured biointerfaces such as the one developed in this

work. Accordingly, the analytical characteristics of both enzyme electrode

architectures prepared without (XO-CD/pAuNP/SWNT/GCE) and with Nafion coating

(Naf/XO-CD/pAuNP/SWNT/GCE) were evaluated.

The XO-CD/pAuNP/SWNT/GCE enzyme electrode showed a fast and sensitive

response to successive additions of xanthine, reaching 95% of the steady-state current

in 5 s. Regarding the Naf/XO-CD/pAuNP/SWNT/GCE, a similar behavior was observed

although the time of response was increased to about 9 s (data not shown). Both the

XO-CD/pAuNP/SWNT/GCE and Naf/XO-CD/pAuNP/SWNT/GCE biosensors exhibited a

linear dynamic range between 50 nM and 9.5 µM fitting to the following equations:

i(mA) = 178×c(Xanthine/M) + 3·10-5 (XO-CD/pAuNP/SWNT/GCE)

i(mA) = 152×c(Xanthine/M) + 2·10-5 (Naf/XO-CD/pAuNP/SWNT/GCE)

With correlation coefficients of 0.999 and 0.997 (n = 10) and sensitivities of

2.47 A/M·cm2 and 2.12 A/M·cm2 for the XO-CD/pAuNP/SWNT/GCE and Naf/XO-

CD/pAuNP/SWNT/GCE biosensors, respectively. It is important to point out that

coating of the bioelectrode surface with the Nafion film did not produce an important

decrease in sensitivity. Moreover, as it can be deduced from data compiled in Table 1,

the sensitivity of these biosensors ranks among the highest reported for other

amperometric xanthine biosensors.

The detection limit achieved with both biosensors was calculated to be 40 nM,

according to the 3SD/m criterion where m is the slope of the linear calibration graph

and SD is the standard deviation for 10 different 50 nM xanthine amperometric

measurements. As it can be deduced from data in Table 1, this value is lower than


166

most of those reported for other xanthine biosensor designs providing detection limits

in the nanomolar level, such as those using poly-5,2’:5’,2’’-terthiophine-3-carboxylic

acid modified Au electrodes,37 multi-walled carbon nanotubes coated GCE38 and

oxidized graphite,39 although slightly higher than that reported for laponite-coated

GCE.31

The apparent KM values were estimated to be 3.2 µM and 3.8 µM for the XO-

CD/pAuNP/SWNT/GCE and Naf/XO-CD/pAuNP/SWNT/GCE biosensors, respectively. It

should be highlighted that these apparent KM values were lower than those reported

for the XO biosensors summarized in Table 1.

As mentioned above, coating with Nafion only reduced the sensitivity of the

bioelectrode in about 14% while the detection limit, the KM value and the range of

linearity are similar to that of the uncoated electrode, suggesting that coating with

Nafion did not significantly affect the electroanalytical performance of the biosensor.

It should be noted that the control bioelectrode (XO/pAuNP/SWNT/GCE) exhibited a

poorer analytical performance: shorter linear range (1.3 µM – 9.5 µM), higher

detection limit (1.0 µM) and lower sensitivity (1.2 A/M cm2), than that achieved with

the biosensors prepared with the CD-modified enzyme. Again these results suggested

that native enzyme was poorly loaded on the nanostructured electrode in comparison

with the modified enzyme, and that the formation of host-guest supramolecular

association was involved in the immobilization of the XO-CD derivative. Comparing the

sensitivity of this control bioelectrode with that obtained with XO-

CD/pAuNP/SWNT/GCE, it could be deduced that the contribution of the host-guest

supramolecular associations represented about 51% of the overall interactions

involved in the immobilization process, assuming that glycosylation of XO with CD

moieties did not affect the other contributing interaction forces.


167

Table 1. Comparison of the analytical characteristics of the developed biosensors with

those previously reported for other electrochemical xanthine biosensors

Electrode E (mV) Linear Range

(µM)

Detection

Limit (µM)

Sensitivity

(mA/M)

KM

(µM)

XO/gelatin/Graphite34 -50a 4.5 - 40 4.5 210 30

XO/PB/PPy/Au-NPs/Au40 -100b 1.0 - 20 - 0.19 43.2

XO/pTTCA/Au37 -350a 0.5 - 100 0.09 5.35 -

XO-CMC-CD/ADA-Au29 +700a 300 - 10400 200 0.25 9900

XO-ADA/pCD/Au22 +700a 310 - 6800 150 0.40 2100

XO/MWNT/GCE38 -400b 0.1 - 6 0.08 - -

SG/XO/MWNT/GCE38 -400b 0.2 - 10 0.1 - -

XO/laponite/GCE31 +390b 0.039 - 21 0.01 6.54 64

XO/CaCO3-NPs/GCE33 +550b 2 - 250 2.0 11.9 -

XO/HRP/CaCO3-NPs/GCE33 -50b 0.4 - 50 0.1 2.8 127

XO/DWNT/CPE30 +900a 2.0 - 50 - 44.1 -

XO/Pd-Pt/Graphite35 -50a 1.5 - 70 1.5 390 58.5

XO/ZnO-NPs/PPy/Pt41 +380a 0.8 - 40 0.8 - 13.5

XO/c-MWNT/PANI/Pt42 +400a 0.6 - 58 0.6 - -

XO/Fe3O4@APTES-

PEG/SWNT/SPE36

+600a 0.25 - 3.5 0.06 394 12.8

XO/Au-NPs/PVF/Pt32 +400b 2.5 - 560 0.75 0.91 393

XO/Pt-NPs/PVF/Pt32 +400b 2.0 - 660 0.6 1.67 286

XO/ox-Graphite39 +600a 0.1 - 0.7 0.1 - 0.4

XO/ZnO-NPs/CHIT/c-

SWNT/PANI/Pt43

+500a 0.1 - 100 0.1 - -

XO-CD/pAuNP/SWNT/GCE

(Present work)

+650a 0.05 – 9.5 0.04 178 3.2

Naf/XO-CD/pAuNP/SWNT/GCE

(Present work)

+650a 0.05 – 9.5 0.04 152 3.8

aAg/AgCl, bSaturated calomel electrode. PB: Prussian Blue, PPy: polypyrrol, NPs: nanoparticles, pTTCA: poly-5, 2’:5’,2’’-terthiophine-3-carboxylic acid, CMC-CD: cyclodextrin-branched carboxymethylcellulose, pCD: polymerized CD, MWNT: multi-walled carbon nanotubes, CGE: glassy carbon electrode, SG: silica sol-gel, DWNT: double-walled carbon nanotubes, CPE: carbon paste electrode, c-MWNT: carboxylated MWNT, PANI: polyaniline, APTES: (3-aminopropyl)triethoxysilane, PEG: monomethoxypolyethylene glycol, SPE: gold screen-printed electrode, PVF: polyvinylferrocene, ox-Graphite: oxidized graphite, CHI: chitosan.


168

The repeatability of the measurements carried out with one single XO-

CD/pAuNP/SWNT/GCE or Naf/XO-CD/pAuNP/SWNT/GCE biosensor was checked for

500 nM xanthine (n = 10), yielding relative standard deviation (RSD) values of 3.9% and

5.2%, respectively. Moreover, the electrode-to-electrode reproducibility was

calculated from the responses of ten different bioelectrodes prepared in the same

manner toward 500 nM xanthine. The obtained RSD values were 6.7% and 8.6% for the

XO-CD/pAuNP/SWNT/GCE and Naf/XO-CD/pAuNP/SWNT/GCE biosensors, respectively.

The slightly higher RSD values observed for the Naf/XO-CD/pAuNP/SWNT/GCE should

be related to the additional step of Nafion thin film formation during the preparation

of the enzyme electrodes.

Naf/XO-CD/pAuNP/SWNT/GCE

Interferants

5.0 µM

Xanthine

0.5 µMa b c d e f g h

g h

XO-CD/pAuNP/SWNT/GCE

Xanthine

0.5 µM

200 nA

Figure 9. Amperometric responses of the XO-CD/pAuNP/SWNT/GCE and Naf/XO-

CD/pAuNP/SWNT/GCE biosensors toward 500 nM xanthine upon addition of glucose

(a), sacarose (b), ethanol (c), acetic acid (d), lactic acid (e), citric acid (f), uric acid (g)

and ascorbic acid (h) at a 5.0 µM concentration level.

The selectivity of the biosensors was evaluated in the presence of eight possible

interfering substances: glucose, sacarose, ethanol, acetic acid, lactic acid, citric acid,

uric acid and ascorbic acid. Figure 9 shows the influence of these possible interfering

compounds added at a 5.0 µM concentration level on the amperometric signal


169

measured for 500 nM xanthine. As it can be observed, the XO-CD/pAuNP/SWNT/GCE

biosensor showed excellent selectivity toward glucose, sacarose, ethanol, acetic acid,

lactic acid and citric acid, yielding unaffected amperometric signals upon addition of

these substances. However, as expected, a significant increase in the steady-state

current was produced at the XO-CD/pAuNP/SWNT/GCE biosensor after addition of a

ten-fold higher concentration of uric or ascorbic acid, showing the strong interference

of these substances on the amperometric measurement for xanthine at the detection

potential applied. Nevertheless, the extent of these interferences was dramatically

decreased when the Naf/XO-CD/pAuNP/SWNT/GCE was employed, as expected

considering the electrostatic repulsion between the negative charged Nafion thin film

and the uricate and ascorbate anions at the working pH.12

The long-term stability of the XO-CD/pAuNP/SWNT/GCE and Naf/XO-

CD/pAuNP/SWNT/GCE biosensors was evaluated by storing them at 4ºC under dry

conditions and periodical evaluation of their analytical response toward 500 nM

xanthine (see Supporting Information). The XO-CD/pAuNP/SWNT/GCE biosensor

retained high electroanalytical activity during the first week. After that, the response

was progressively lost with time of storage according to a first-order kinetics progress,

showing about 65% and 54% of its initial activity after 30 and 40 days of storage,

respectively. The stability showed by this biosensor could be associated with the

multivalence supramolecular strategy employed to immobilize the enzyme on the

electrode surface.18,29 In addition, it has been largely demonstrated that the stability of

enzymes can be significantly improved by chemical glycosylation with CD derivatives.18

Interestingly, coating of the electrode with Nafion thin film improved largely

the biosensor stability keeping about 91% of its electroanalytical activity after 40 days

of storage. This fact suggested that the Nafion film presumably avoided the release of

enzyme molecules from the electrode surface and preserved their three-dimensional

active structure during storage. Similar results were previously described for other

enzyme electrodes coated with Nafion thin films.12


170

CONCLUSIONS

The electropolymerization of polyfunctionalized Au nanoparticles, bearing

aniline, sulfonic acid and adamantane residues, through the formation of bis-aniline

linkages on SWNTs-modified GCEs served as efficient electrode platforms to form

supramolecular host-guest complexes with CD-modified enzymes. These electrode

surfaces, exhibiting molecular receptor capacity, were, in particular, employed as

support for the supramolecular immobilization of the glycoenzyme XO-ADA and

further construction of an amperometric biosensor for xanthine. The enzyme electrode

exhibited good sentitivity and stability, a very low detection limit, as well as fast

electroanalytical response toward the analyte. In fact, the analytical performance of

the supramolecular enzyme immobilization based biosensor ranked among the best

achieved when compared with those previosly reported for other XO electrochemical

biosensors. Therefore, according to these results, we can anticipate the use of this

bisaniline-cross-linked nanostructured network of metal nanoparticles on SWNT-

coated electrodes as an excellent strategy to prepare enzyme biosensors with

supramolecular architecture.

ASSOCIATED CONTENT

Supporting Information. Additional characterization of the nanoparticles and

chronoamperometric and stability studies for the biosensors. This material is available

free of charge via the Internet at http://pubs.acs.org.

AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected], [email protected]


171

ACKNOWLEDGEMENTS


of Science and Innovation. Financial support from the Spanish Ministry of Science and

Innovation CTQ2011-24355, CTQ2009-12650, CTQ2009-09351 and Comunidad de

Madrid S2009/PPQ-1642, programme AVANSENS is gratefully acknowledged.

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SUPRAMOLECULAR IMMOBILIZATION OF MODIFIED ENZYMES ON

ELECTROPOLYMERIZED MATRIX OF FUNCTIONALIZED HYBRID GOLD

NANOPARTICLES/SINGLE-WALLED CARBON NANOTUBES FOR THE PREPARATION OF

ELECTROCHEMICAL BIOSENSORS

Reynaldo Villalonga,1,2 Paula Díez,1 Marcos Eguílaz,1 Paloma Martínez,3 José M.

Pingarrón1,2,*


Chemistry, Complutense University of Madrid, 28040-Madrid, Spain. 2IMDEA

Nanoscience, Cantoblanco Universitary City, 28049-Madrid, Spain

Figure S1. TEM image of the polyfunctionalized Au nanoparticles acquired with a JEOL

JEM-2100 microscope at 200 kV.


175

Figure S2. HR-TEM image of the polyfunctionalized Au nanoparticles acquired with a

JEOL JEM-3000 F microscope at 300 kV.

2 4 6 8 10 12

400

800

1200

1600

2000

2400

2800

300 µL

i a (

nA

)

Time (min)

30 nm5 µL

10 µL

150 µL

100 µL

50 µL

25 µL 10 µL

Figure S3. Dynamic amperometric response of XO-CD/pAuNP/SWNT/GCE poised at

+650 mV to successive addition of 100 µM xanthine solution. Inset: Amperometric

response of the electrode at lower concentrations of xanthine. Initial working volume:

10 ml. Supporting electrolyte: 0.1 M sodium phosphate buffer, pH 7.0.


176

0 2 4 6 8 10

0,0

0,5

1,0

1,5

i a (

µA

)

Xanthine (µM)

Figure S4. Calibration curves for xanthine obtained with XO-CD/pAuNP/SWNT/GCE (),

Naf/XO-CD/pAuNP/SWNT/GCE () and XO/pAuNP/SWNT/GCE () biosensors.

0 10 20 30 40

0

25

50

75

100

Re

lative

Sta

bili

ty (

%)

Time (day)

Figure S5. Evaluation of the storage stability with time for XO-CD/pAuNP/SWNT/GCE

() and Naf/XO-CD/pAuNP/SWNT/GCE () biosensors. Measurements were carried out

toward 500 nM xanthine.

7.5

Electroanalysis 23 (2011) 1790-1796

Immobilization of xanthine oxidase on carbon nanotubes through

double supramolecular junctions for biosensor construction

Electroanalysis 23 (2011) 1790-1796 7. PUBLICACIONES

177

IMMOBILIZATION OF XANTHINE OXIDASE ON CARBON NANOTUBES

THROUGH DOUBLE SUPRAMOLECULAR JUNCTIONS FOR BIOSENSOR

CONSTRUCTION

Reynaldo Villalonga, Paula Diez, María Gamella, Julio Reviejo, José M. Pingarrón*

Department of Analytical Chemistry, Faculty of Chemistry, Complutense University of

Madrid, 28040-Madrid, Spain



ABSTRACT

A novel approach for the non-covalent functionalization of single walled carbon

nanotubes with enzymes, using a β-cyclodextrin-modified pyrene derivative, mono-6-

ethylenediamino-(2-pyrene carboxamido)-6-deoxy-β-cyclodextrin (Pyr-βCD), as a

molecular bridge for the construction of a supramolecular assembly between the

nanotube surface and an adamantane-modified enzyme, is reported. The Pyr-βCD

derivative was synthesized and its stacking to SWNT through π-π interactions

accomplished. The functionalized nanotubes showed low capacity for the non-specific

adsorption of proteins, but were able to immobilize adamantane-modified xanthine

oxidase via host-guest associations. This double supramolecular junctions-based

approach was employed to modify a glassy carbon electrode with the

enzyme/nanotubes complex for designing a biosensor device toward xanthine. The

biosensor showed fast electroanalytical response (10 s), high sensitivity (5.9 mA/M

cm2) low detection limit (2 µM) and high stability.



178

KEYWORDS: Single walled carbon nanotube, supramolecular assembly, enzyme

biosensor, cyclodextrin

INTRODUCTION

Multi- and single walled carbon nanotubes (SWNT) are wire-shaped nanosized

materials with unique electrical, mechanical and thermal characteristics [1-2]. These

properties allow carbon nanotubes to be the central core of novel advanced materials

with potential applications in hydrogen storage [3], tissue engineering [4-5], the

preparation of reinforced materials and coatings [6-7], nano and molecular electronics

[8], drug delivery systems [9] and biosensor construction [10-12].

However, there are several disadvantages associated with the intrinsic

characteristics of carbon nanotubes that make rather difficult their wide use with

biomolecules. For instance, unmodified carbon nanotubes have poor solubility in any

solvent and lack proper chemical functionalities needed to obtain stable and specific

unions with proteins and nucleic acids [13]. These problems can be overcome by

chemical activation of carbon nanotubes through a great variety of well know

reactions, but it has been proved that such activation methods significantly affect the

structure and electroconductive properties of these materials [14-15]. It has been also

described that several proteins undergo structural changes and inactivation upon

adsorption on SWNT [16], reducing the success of this immobilization strategy for

biomolecules. On the other hand, the occurrence of non-specific adsorption of

proteins on the side wall of carbon nanotubes affects the design of reliable drug

delivery systems and analytical biosensors [17-18]. For these reasons, the

development of new strategies for improving solubility of carbon nanotubes in

aqueous systems, reducing non-specific protein adsorption processes and favouring

the specific immobilization of biomolecules without affecting the structure and

properties of these nanomaterials receives considerable attention.

Dai´s group [19] proposed the use of pyrene derivatives as a controlled and

specific method for the non-covalent functionalization of SWNT via the formation of π-


179

π stacking. It was confirmed that solubility and capacity to immobilize protein via

covalent linkage to the pyrene derivatives was conferred to SWNT through this

supramolecular approach. Furthermore, it has been reported the synthesis of β-

cyclodextrin (βCD) modified pyrene derivatives which were used to functionalize

SWNT. This strategy conferred enhanced solubility and molecular receptor properties

to these nanomaterials [20-22]. This kind of βCD-based adducts were evaluated as

molecular receptors for the host-guest immobilization of chemically hydrophobized

proteins on electrode surfaces. In fact, it was previously reported that adamantane-

modified proteins could be easily immobilized on βCD-coated metal electrodes

through the formation of supramolecular complexes [23]. Such approach was further

employed to construct stable enzyme biosensors with supramolecular design [24-26].

In this work we propose a novel approach for the non-covalent

functionalization of SWNT with enzymes, using a βCD-modified pyrene derivative as a

molecular bridge for the construction of a supramolecular assembly between the

nanotube surface and an adamantane-modified enzyme. In order to do that, the

synthesis of mono-6-ethylenediamino-(2-pyrene carboxamido)-6-deoxy-β-cyclodextrin

(Pyr-βCD) is reported as well as its stacking to SWNT through π-π interactions.

Furthermore, the second supramolecular layer was constructed by host-guest

associations between adamantane-modified xanthine oxidase (XO-ADA) and the Pyr-

βCD coated SWNT. As a proof of concept, this immobilization strategy through double

supramolecular junctions was employed for the design of an enzyme biosensor for the

determination of xanthine.

EXPERIMENTAL

1. Reagents and solutions

Xanthine oxidase (XO, Type III, 1.3 U/mg) and βCD were purchased from Sigma-

Aldrich Co. (USA). SWNT, 1-hydroxybenzotriazole (HOBt), 2-pyrene carboxylic acid and

N,N'-dicyclohexylcarbodiimide (DCC) were acquired from Wako Pure Chemical

Industries, Ltd. (Japan). All other chemicals were of analytical grade. 10 mM and 500


180

µM xanthine solutions in 50 mM sodium phosphate buffer of pH 7.0 were freshly

prepared for analytical purposes.

2. Apparatus and electrodes

Cyclic voltammetry was performed using a µAutolab Type III

potentiostat/galvanostat and the data were acquired using GPES Ver. 4.9 software

(Metrohm Autolab B.V. The Netherlands). Electrochemical impedance spectroscopy

experiments were performed with a Voltalab PGZ 402 potentiostat/galvanostat

equipped with the VoltaMaster 4 Upgrade 4.00 electrochemical software (Radiometer

Analytical SAS, France). Amperometric measurements were performed with a dual-

channel ultrasensitive Inbea potentiostat (Inbea Biosensores S.L., Spain). A

conventional three-electrode system was employed in all electrochemical studies. The

working electrode was a glassy carbon electrode (GCE, 3.0 mm diameter) coated with

SWNT-Pyr-βCD and the immobilized enzyme. An Ag/AgCl/KCl (3 M) and a Pt wire were

used as reference and counter electrodes, respectively. Bioelectrode measurements

were carried out at 25ºC in 0.1 M sodium phosphate buffer, pH 7.0 (working volume

10 ml). The solution was exhaustively deaerated before each electrochemical

experiment. The solutions were stirred at 300 rpm with a magnetic bar for

amperometric measurements.

The mass variation during the immobilization of the native and modified enzyme

forms on SWNT was followed by a Maxtek RQCM 603200-2 quartz crystal microbalance

(Inficon, USA), oscillating at a nominal frequency of 9.0 Hz. The surface morphology of

the electrode was checked by high resolution field emission scanning electron

microscopy (FE-SEM) using a JEOL JSM-6335F microscope (JEOL Ltd., Japan). 1H NMR

studies were performed with an Avance III 400 spectrometer (Bruker BioSpin GmbH,

Germany). Positive-ion FAB-MS spectra were recorder with a Jeol HX-110 mass

spectrometer (Jeol Ltd., Japan).


181

3. Synthesis of mono-6-ethylenediamino-(2-pyrene carboxamido)-6-deoxy-β-

cyclodextrin (Pyr-βCD) derivative

The mono-6-ethylenediamino-6-deoxy-β-cyclodextrin was obtained by treating

the corresponding mono-6-O-tosyl derivative [27] with freshly distilled

ethylenediamine as previously described [28]. The product was purified by ion

exchange chromatography on CM-Sephadex C-25 (NH4+ form) and the purity and

identity of this derivative was checked by TLC, 1H- and positive-ion FAB-MS. The

synthesis of the Pyr-βCD derivative was accomplished as follows [20]: 600 mg (0.51

mmol) of mono-6-ethylenediamino-6-deoxy-β-cyclodextrin and 254 mg (1.03 mmol) of

2-pyrene carboxylic acid were dissolved in 5 mL DMF under nitrogen atmosphere. The

mixture was continuously stirred and cooled at 0ºC in an ice/NaCl bath. Then, 200 mg

(0.97 mmol) DCC and 100 mg (0.74 mmol) HOBt were added. After complete

dissolution, the mixture was allowed to rise at room temperature and the reaction was

stirred overnight. The mixture was filtered and the resulting solution was precipitated

over cool acetone. The excess of mono-6-ethylenediamino-6-deoxy-β-cyclodextrin was

eliminated by washing the precipitate with water for several times. The yellow solid

was finally dried in vacuum over P2O5. Yield: 470 mg (0.33 mmol, 66%). 1H-NMR

(DMSO-d6) 2.33 (q, 2H, -CH2NHCH2CH2-), 2.67 (q, 2H, -NHCH2CH2NHCO-), 3.10-3.46

(m, 15H, C(2)H and C(4)H of CD, -CH2NHCH2-), 3.55-3.75 (m, 28H, C(5)H, C(3)H and

C(6)H of CD), 4.34-4.40 (m, 6H, O(6)H of CD), 4.43-4.58 (br, 1H, -NHCO-), 4.80-4.88 (d,

7H, C(1)H of CD), 5.66-5.88 (m, 14H, O(2)H and O(3)H of CD) and 7.82-8.26 (m, 9H,

pyrene group). FAB-MS m/z 1405.7 (M+H)+.

4. Solubilization of SWNT

Firstly, SWNT were purified by heating in an air flow oven at 350ºC for 2 h in

order to remove amorphous carbon materials and further soaking in 30% HCl for one

day to remove the metal impurities. The treated nanotubes were washed with double

distilled water until neutral pH, then washed with EtOH and finally dried in vacuum

under P2O5.


182

Solubilization of nanotubes with Pyr-βCD was performed as previously

described [20]. Briefly, 50 mg Pyr-βCD was dissolved in 10 mL of 0.1 M NaOH. Ten

milligrams of SWNT were added to this mixture and dispersed by sonication during 4 h

in a conventional low-energy ultrasonic bath. The mixture was further centrifuged and

the supernatant was exhaustively dialyzed against 0.1 M NaOH solution to remove the

excess of Pyr-βCD. The resulting solution of Pyr-βCD-coated SWNT (SWNT/Pyr-βCD)

was raised to 20 mL final volume with 0.1 M NaOH, and kept at room temperature

until use.


XO was first hydrophobized with 1-adamantane carboxylic acid via a

carbodiimide-catalyzed reaction as previously described [25]. To prepare the enzyme-

modified electrode, a bare GCE was polished to a mirror-like with alumina powder (0.3

µm), rinsed thoroughly with double distilled water, then washed successively with

double distilled water, anhydrous ethanol and acetone in an ultrasonic bath, and dried

under N2 before use. In order to accomplish electrode coating, two 10 µL-aliquots of

the SWNT/Pyr-βCD solution were successively deposited and dried on the electrode

surface. Then, the electrode was treated with 20 µL of 0.1 M HCl, dried and further

washed with double distilled water. The modified GCE was dried again and dipped in a

5 mg/mL solution of adamantane-modified XO (XO-ADA) in 50 mM sodium phosphate

buffer of pH 7.0 during 4 h. The enzyme-modified electrode was finally washed with

the same cool buffer, dried under nitrogen and kept at 4ºC until use.


The fundamentals of the enzyme biosensor construction are schematized in

Figure 1. Firstly, the Pyr-βCD adduct was synthesized through a two steps reaction

involving the preparation of mono-6-ethylenediamino-6-deoxy-β-cyclodextrin from the


183

mono-6-tosyl derivative of βCD and its subsequent coupling with 2-pyrene carboxylic

acid by means of a carbodiimide-catalyzed reaction.

Figure 1. Scheme displaying the steps and fundamentals involved in the enzyme

biosensor preparation.

Secondly, the sonication of SWNT in an alkaline solution of Pyr-βCD resulted in

the solubilisation of the nanotubes. This fact was mediated by the formation of strong

π-π interactions between the aromatic pyrene moieties and the side wall of the


184

nanotubes, as it has been reported in the literature [20-22]. Solubility of the resulting

adduct was conferred by the hydrophilic βCD units. It should be mentioned that this

alkaline SWNT/Pyr-βCD solution was stable for at least three months at room

temperature.

In order to prepare an enzyme derivative suitable to form host-guest inclusion

complexes with the βCD-coated SWNT, XO was chemically modified with 1-

adamantane carboxylic acid through a water-soluble carbodiimide catalyzed

condensation. The modified enzyme retained about 80% of the initial catalytic activity

after attachment of 8 mol adamantane residues per mol of monomeric protein [25].

The interaction of native and adamantane-modified enzyme forms with

SWNT/Pyr-βCD was investigated by means of quartz crystal microbalance

measurements, and the results are shown in Figure 2. The experiments were

performed by coating the gold electrochemical quartz crystal with SWNT/Pyr-βCD

similarly to that described in the Experimental section for GCE surface modification.

Figure 2. Quartz crystal microbalance responses for gold disks coated with: SWNT

during immobilization of native XO (a); SWNT/Pyr-βCD during the immobilization of

adamantane-modified (b), and native XO (c).


185

A noticeable decrease in the resonance frequency was observed for the

immobilization process of native enzyme on SWNT (curve a). The frequency difference

between its initial value and the value measured at the plateau indicating complete

immobilization was substituted into the Sauerbrey equation giving an average amount

of adsorbed enzyme on the nanotubes of 64 µg/mg. This behaviour indicated a strong

non-specific adsorption of the protein on the side wall of SWNT, which constitutes an

important disadvantage for some applications of bare carbon nanotubes-based

sensors [17-18]. In the case of the SWNT/Pyr-βCD-modified crystal, it was calculated

that 24 µg of XO-ADA per mg of nanotubes were immobilized (curve b), while only 6.1

µg/mg of the non-modified XO was adsorbed on SWNT/Pyr-βCD (curve c). These

results suggest that the non-specific adsorption of the enzyme on the SWNT surface

was strongly reduced upon nanotubes modification with Pyr-βCD moieties.

Furthermore, the much larger loading observed with adamantane-modified enzymes

indicated that the protein was more efficiently immobilized through the formation of

supramolecular inclusion complexes using Pyr-βCD as a double-functionality

supramolecular bridge.

Figure 3. Nyquist plots recorded at bare () and SWNT/Pyr-βCD-modified GCE before

() and after immobilization of native () and adamantane-modified XO (), in 0.1 M

KCl solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1).


186

Accordingly, GCE were subsequently modified with the SWNT/Pyr-βCD

assembly. A stable conductive coating was obtained and the electroactive surface area

increased from 7.2 mm2 to 10.2 mm2 as calculated by cyclic voltammetry using the

[Fe(CN)6]4−/3− as redox probe. Further, XO-ADA was immobilized on the modified

electrodes by dipping them in the enzyme solution. An immobilization time of 4 h was

selected in order to ensure the process was completed.

The electrode modification process was also characterized by electrochemical

impedance spectroscopy with a 5 mM [Fe(CN)6]4−/3− solution in 0.1 M KCl. Figure 3

shows the corresponding Nyquist plots for each modification step. The bare GCE

showed the typical Nyquist plot for this kind of electrode material while the electron

transfer resistance exhibited a noticeable decrease upon electrode modification with

SWNT/Pyr-βCD. The observed decrease was, however, slightly lower than that

measured for the SWNT-modified electrode (data not show) which can be attributed

to the non-conducting nature of Pyr-βCD moieties. Immobilization of XO-ADA

increased this insulating effect as it was indicated by the increase in the electron

transfer resistance. It should be also noted that the immobilization of unmodified XO

did not produce a significant change in the shape of the Nyquist plot with respect to

that of the SWNT/Pyr-βCD coated GCE, suggesting a lower enzymatic loading in

comparison with the adamantane-modified counterpart.

Cyclic voltammograms of XO/SWNT/Pyr-CD and XO-ADA/SWNT/Pyr-CD

modified electrodes in 0.1 M phosphate buffer of pH 7.0 in the absence and presence

of xanthine are displayed in Figure 4. It can be clearly seen as a well-defined oxidation

peak, at about +350 mV, corresponding to the oxidation of the uric acid formed as the

product of the catalytic reaction with xanthine, appeared only in the case of the XO-

ADA biosensor. On the contrary, no significant oxidation response was observed upon

addition of 40 µM xanthine using unmodified XO as immobilized enzyme on the

SWNT/Pyr-βCD electrode, suggesting again a low enzyme loading for this design.


187

Figure 4. Cyclic voltammograms of XO/SWNT/Pyr-βCD and XO-ADA/SWNT/Pyr-βCD

modified GCE in the absence (voltammograms A and B, respectively) and in the

presence of 40 µM xanthine (voltammograms C and D, respectively). Scan rate, 50

mV/s in 0.1 M sodium phosphate buffer, pH 7.0.

The influence of pH and the applied potential on the amperometric response

for xanthine of the enzyme electrode prepared through this supramolecular approach

was investigated. The effect of pH on the biosensor response, poised at +600 mV, was

investigated in the 5.0–9.0 range. The highest steady-state current was measured at

around pH 6.0-7.0, the latter value being selected for further experiments. Regarding

the applied potential, the biosensor response increased steadily when changing the

imposed potential from +100 mV to +400 mV versus Ag/AgCl, then reaching a

practically constant value until +800 mV. Although the highest current could be

measured at only +400 mV, the signal-to-background current ratio toward xanthine

exhibited a larger value at +600 mV. Accordingly, we decided to check the biosensor

performance at this more positive potential value despite the worsening in selectivity.


188

Figure 5. Dynamic amperometric response of the XO-ADA/SWNT/Pyr-βCD (A) and

XO/SWNT/Pyr-βCD (B) modified GCE at +600 mV to successive additions of a 10 mM

xanthine solution. Initial working volume: 10 ml. Supporting electrolyte: 0.1 M sodium

phosphate buffer, pH 7.0.

Therefore, the amperometric response of the enzyme electrode to successive

additions of xanthine was evaluated under the selected conditions, and the typical

results are illustrated in Figure 5. As a control, similar experiments were performed by

using native XO immobilized on the SWNT/Pyr-βCD coated GCE. A fast and sensitive

bioelectrocatalytic response was shown by the XO-ADA based electrode, reaching 95%

of the steady-state current in about 10 s. However, a rather poor analytical behaviour

was observed with the electrode modified with native enzyme, in good agreement

with the results observed using the quartz crystal microbalance, electrochemical

impedance spectroscopy and cyclic voltammetry. In addition, it should be noted that

the amperometric response of the XO-ADA modified electrode was lost after

incubation of the biosensor in a saturated solution of 1-adamantane carboxylic acid for

12 h (data not shown) as a consequence of the competition with the adamantane-

modified enzyme and the corresponding displacement in the equilibrium. These facts

strongly support our hypothesis on the host-guest nature of the immobilization

mechanism for XO-ADA on SWNT/Pyr-βCD. In addition, the achieved results pointed


189

out that functionalization of SWNT with Pyr-βCD significantly reduced the non-specific

protein adsorption on the nanotube surface.

The developed biosensor exhibited a linear amperometric response towards

xanthine over the 5.0 - 600 µM concentration range. This range of linearity extended

about one order of magnitude wider than those previously reported for other xanthine

biosensors [29-32].

The regression equation of the present biosensor was:

ia(µA) = 610×c(Xanthine/M) + 0.02

with a correlation coefficient of 0.998 (n = 10) and a sensitivity of 5.9 mA/M cm2.

According to the signal-to-noise = 3 criterion, the detection limit was found to

be 2 µM, which is similar to those achieved with other biosensor designs [25, 29, 30,

33]. Furthermore, an apparent Michaelis-Menten constant, KM , and a maximum rate

for the enzyme reaction, IMAX, were calculated as 0.58 mM and 503 nA, respectively.

It should be noted that the biosensor provided also a remarkable high

sensitivity when operated at a detection potential of + 400 mV. However, the range of

linearity available in this case was shown to be much shorter than at +600 mV, which is

an important analytical limitation for its real applicability.

Repetitive amperometric measurements at +600 mV for successive additions of

10 mM xanthine yielded a relative standard deviation (RSD) value (n = 10) of 6.1%.

Moreover, the RSD value calculated from the measurements carried out with ten

different biosensors prepared in the same manner was 9.3%. It is interesting to

compare these figures with those obtained by applying a detection potential of +400

mV, where the respective RSD values were 8.5% and 13.6%.

The selectivity of the biosensor, poised at +600 mV, was evaluated in the

presence of eight potential interfering substances. The current measured for each

possible interfering at a 50 µM concentration level in the presence of 20 µM xanthine

was used to evaluate the biosensor selectivity. The enzyme biosensor showed good


190

selectivity toward xanthine, yielding unaffected amperometric readouts in the

presence of L-glucose, sucrose, ethanol, acetic acid, lactic acid and citric acid. However,

uric acid and ascorbic acid caused interference in the quantification of xanthine giving

rise to an increase in the measured current of about 48% and 81%, respectively. In

order to compare, a similar set of experiments were performed by applying a

detection potential to the biosensor of +400 mV. Also, as expected due to the low

overpotential for the electrochemical oxidation of these substances, only uric acid and

ascorbic acid produced and increase in the xanthine signal of 23% and 67%,

respectively.

In order to check the long-term stability of the biosensor, the enzyme electrode

was stored at 4ºC under dry conditions and periodically tested by constructing a

calibration graph for xanthine using +600 mV as working potential. As can be observed

in Figure 6, the biosensor did not exhibit any loss in sensitivity during the first two

weeks of storage.

Figure 6. Long-term stability of the XO-ADA/SWNT/Pyr-βCD/GCE biosensor toward the

determination of xanthine.


191

Longer times of storage produced a progressive loss of sensitivity reaching

about 67% of the initial activity after 1 month. This relative high long-term stability of

the biosensor could be attributed to the beneficial effect of β-CD in the vicinity of the

enzyme molecules, providing a beneficial hydrophilic microenvironment for its

catalytic activity and folding stability [34].

CONCLUSIONS

A novel non-covalent strategy has been developed to functionalize SWNT with

adamantane-modified XO by using Pyr-βCD as double-functionality supramolecular

bridge. This functionalization approach with the supramolecular assembly has

demonstrated to be useful for the construction of an amperometric biosensor for

xanthine with acceptable analytical characteristics. This strategy may be generalized to

other enzyme biosensors, because it is based on the host-guest association of βCD

with hydrophobic compounds such as the adamantane derivatives, which can be easily

conjugated to any protein by conventional methods. According to these results, it can

be concluded that this supramolecular-based approach can be used as a successful

strategy to functionalize SWNT with enzymes giving rise to new biosensing devices

with analytical applicability. There are potential advantages associated with the use of

the Pyr-βCD linker in the design of new supramolecular assemblies of enzymes on

SNWT at the electrode surface. In fact, the π-π stacking of Pyr-βCD moieties confers

solubility, hydrophilicity and molecular receptor properties to SWNT without affecting

their chemical structure and electroconductive characteristics. This non-covalent

surface transformation also contributes to the reduction of non-specific adsorption of

protein molecules on SWNT side walls, but serves as a double-functionality

supramolecular bridge for the rational host-guest immobilization of protein molecules,

previously modified with hydrophobic compounds such as adamantane derivatives.


192

ACKNOWLEDGEMENTS





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7.6

Journal of Materials Chemistry 21 (2011) 12858-12864

Decorating carbon nanotubes with polyethyleneglycol-coated magnetic

nanoparticles for implementing highly sensitive enzyme biosensors

Journal of Materials Chemistry 21 (2011) 12858-12864 7. PUBLICACIONES

195

DECORATING CARBON NANOTUBES WITH POLYETHYLENEGLYCOL-

COATED MAGNETIC NANOPARTICLES FOR IMPLEMENTING HIGHLY

SENSITIVE ENZYME BIOSENSORS

Reynaldo Villalonga,a María L. Villalonga,b Paula Díez,a José M. Pingarrón*,a

a Department of Analytical Chemistry, Faculty of Chemistry, Complutense University of

Madrid, 28040-Madrid Spain. Fax: +34 913944329; Tel: +34 913944315; E-mail:

[email protected]

b Center for Enzyme Technology, University of Matanzas, Autopista a Varadero km 3 ½,

Matanzas 44740, Cuba. Fax: +53 45253101; Tel: +53 45261251; E-mail:

[email protected]


196

ABSTRACT

Superparamagnetic Fe3O4 nanoparticles were coated with (3-

aminopropyl)triethoxysilane and further branched with monomethoxypolyethylene

glycol chains. These nanoparticles were employed for the non-covalent surface

modification of single walled carbon nanotubes, conferring them magnetic properties.

This nanomaterial was employed to immobilize the enzyme xanthine oxidase in order

to construct magnetically modified disposable gold screen-printed electrodes as

bioelectrodes for the determination of xanthine. The electroanalytical properties of

the biosensor were modulated by the nanomaterial composition, being optimal at a

carbon nanotubes:magnetic nanoparticles ratio of 1:27. The resulting biosensor

showed a linear dependence on the xanthine concentration in the 0.25-3.5 µM range

with a fast amperometric response in 12 s. The biosensor also showed a noticeable

high sensitivity of 1.31 A/M cm2 and a very low detection limit of 60 nM, which can be

compared advantageously with other biosensor designs for xanthine.

INTRODUCTION

Last challenges in electrochemical analysis have been predominantly linked to

the use of nanosized materials for the construction of sensing electrodes. This trend is

justified by the unique structural, physico-chemical and surface-to-volume ratio

properties of nanomaterials. In addition, they also offer the possibility to design a wide

variety of original tree-dimensional nanoarchitectures at the electrode surface, based

on the single or combined use of different nanomaterials.1

Multi- and single walled carbon nanotubes (SWNT) are among the

nanomaterials more widely used in electroanalysis due to their relevant chemical

stability, protein adsorption ability, electro-conductive properties and capability to

promote fast electron transfer reactions of some enzymatically generated species.2 In

this regards, a large number of nanostructured biosensors based on carbon nanotubes

have been reported, in which they have been used alone2a,3 or combined with other


197

nanomaterials such as quantum dots,4 metal and metal oxide nanoparticles5 and

nanowires.6

During last years, a significant interest has been devoted to prepare carbon

nanotubes-based magnetic nanomaterials, not only for analytical applications but also

for magnetic data storage, microelectronic, xerography and magnetic resonance

imaging.7 Such kind of hybrid nanomaterials can be prepared by filling the nanotubes

with the magnetic material7,8 or by coating the outer wall of the nanotubes with

magnetic nanoparticles.5c,9 The later is, by far, the strategy most commonly used

because it is easier to manipulate the outer wall surface rather than the inner cavity of

nanotubes, offering in this way more experimental alternatives. In addition, nanotubes

filled with magnetic nanoparticles did not show useful magnetic properties because it

is difficult to control the amount and location of magnetic material inside the

nanotubes.8c Location of iron oxide nanoparticles on the outer wall of nanotubes is

also a better choice for biosensor applications, due to high biocompatibility and

protein load capacity of magnetic nanoparticles.10

In general, decoration of carbon nanotubes with magnetic nanoparticles has

been accomplished by adsorption,11 covalent attachment,9b π-π stacking9a and coating

with polymeric films containing the iron oxide nanoparticles.5c Furthermore, it has

been reported that polyethylene glycol derivatives can be irreversibly adsorbed on

carbon nanotubes.12 On this basis, we expected that small nanoparticles functionalized

with this polymer could be also irreversibly adsorbed on carbon nanotubes.

In this work we propose a novel approach for decorating SWNT with magnetic

nanoparticles, based on the non-covalent attachment of monomethoxypolyethylene

glycol (PEG)-coated superparamagnetic Fe3O4 nanoparticles on the nanotube surface.

In addition, the bioelectroanalytical potential of this hybrid nanomaterial was

evaluated by immobilizing xanthine oxidase as a model enzyme and constructing a

biosensor device towards xanthine.


198


1. Electrodes and reagents

Disposable gold screen-printed electrodes (AuSPE, Model DRP-220BT, 4 mm

diameter) were purchased from DropSens (Spain). The AuSPE also included a gold

counter electrode and a silver pseudo-reference electrode, screen-printed on a

ceramic substrate (3.4 cm × 1.0 cm). To allow the magnetic immobilization of the

Fe3O4-based nanomateriales, AuSPE were transformed by fastening nickel-plated

neodymium disk magnets (3.2 mm diameter) on the other side of the gold working

electrodes using commercial acrylic glue.

Xanthine oxidase (XO, Type III, 1.3 U/mg), (3-aminopropyl)triethoxysilane

(APTES), methoxypolyethylene glycol succinate N-hydroxysuccinimide (PEG-NHS, MW

= 5000 Da), xanthine and SWNT were acquired from Sigma-Aldrich Co. (USA). All other

chemicals were of analytical grade.

2. Instrumentation and solutions

Electrochemical impedance spectroscopy, cyclic voltammetry and

amperometric measurements was performed using a FRA2 µAutolab Type III

potentiostat/galvanostat, and the data were acquired using a Frequency Response

Analyser and GPES Ver. 4.9 software, respectively (Metrohm Autolab B.V., The

Netherlands). All potential values were referred to the screen-printed silver pseudo-

reference electrode.

The measurements with the developed biosensors were carried out at 25ºC in

0.1 M sodium phosphate buffer, pH 7.0 (working volume 10 ml). The solution was

exhaustively aerated before each electrochemical experiment. The solutions were

stirred at 300 rpm with a magnetic bar during amperometric studies. 1 mM xanthine

solution in 50 mM sodium phosphate buffer, pH 7.0 was freshly prepared.

Transmission electron microscopy (TEM) measurements were performed with a

Philips CM 200 FEG microscope (FEI Co., USA). The surface morphology of the

nanostructured electrode surfaces was investigated by high resolution field emission


199

scanning electron microscopy (FE-SEM) using a JEOL JSM-6335F apparatus (JEOL Ltd.,

Japan). Magnetization measurements were carried out in a MPMS Squid

magnetometer (Quantum Design, Inc. USA). FT-IR spectra were acquired with a Perkin-

Elmer instrument.

3. Synthesis of Fe3O4/APTES-PEG nanoparticles

Fe3O4 magnetic nanoparticles were prepared according to the literature with

little modification.13 Briefly, 5.4 g of FeCl3·6H2O and 2.0 g of FeCl2·4H2O were dissolved

in 25 mL of 0.5 M HCl under N2 atmosphere. The solution was dropwise added to 250

mL of a 1.5 M NaOH solution with vigorous mechanical stirring and ultrasound

treatment under continuous N2 bubbling. The black precipitate formed was isolated by

magnetic decantation, exhaustively washed with double distilled water until neutrality,

then washed twice with ethanol and dried under vacuum.

The core-shell Fe3O4/APTES nanoparticles were prepared as previously

described,14 by dispersing 300 mg of the as-synthesized Fe3O4 nanoparticles in a

mixture of 600 mL ethanol and 4 mL of water by sonication. APTES (120 µL) was then

added, and the mixture was mechanically stirred under N2 atmosphere for 7 h. The

nanoparticles were isolated by magnetic decantation and purified by five cycles of re-

dispersion in ethanol and magnetic decantation. The Fe3O4/APTES nanoparticles were

finally dried at room temperature under vacuum.

To modify the core-shell Fe3O4/APTES nanoparticles with the PEG chains, 120

mg of Fe3O4/APTES nanoparticles were dispersed in 50 mL sodium phosphate buffer,

pH 7.4 by sonication before 400 mg of PEG-NHS were added. The mixture was

mechanically stirred during 24 h, and then the modified nanoparticles were

magnetically decanted and sequentially washed with double distilled water, ethanol

and acetone. The resulting solid was finally dried at room temperature under vacuum.

4. Immobilization of XO on Fe3O4/APTES-PEG nanoparticles

Core-shell Fe3O4/APTES-PEG nanoparticles (10 mg) were suspended in 9.0 mL of

50 mM sodium phosphate buffer, pH 8.0, by sonication and mixed with 1.0 mL of 25%


200

(v/v) glutaraldehyde. The mixture was shacked in the dark at 4ºC during 2 h. Then, the

solid was magnetically decanted and exhaustively washed with double distilled water.

The solid was re-dispersed in 4.0 mL of 50 mM sodium phosphate buffer, pH 7.0, and

mixed with 10 mg XO, previously dissolved in 1.0 mL of the same buffer. The mixture

was gently shacked at 4ºC during 2 h, and the solid was decanted and isolated by

centrifugation. The enzyme-modified nanoparticles were repeatedly washed by

sequential re-dispersion in cold 50 mM sodium phosphate buffer, pH 7.0, and

magnetic decantation. The amount of attached enzyme was estimated by difference

after measuring the non-immobilized protein by Bradford method.15 The Fe3O4/APTES-

PEG-XO nanocatalyst was finally suspended in 5.0 mL of the same buffer and kept in

refrigerator until use.

The catalytic activity of native and immobilized enzyme forms was estimated by

the time course increase of absorbance at 290 nm, corresponding to the production of

uric acid (ε = 1.22x104 M-1cm-1) by xanthine oxidation.16


To prepare the Fe3O4/APTES-PEG-XO/SWNT nanomaterial, SWNT was first

oxidized by ultrasound treatment with a 3:1 (v/v) mixture of concentrated H2SO4 and

HNO3 at room temperature during 4 h.17 Moreover, AuSPEs were activated by

dropping 50 µL of 0.1 M H2SO4 solution on the electrode surface and scanning ten

cyclic voltammograms from 0.0 to 1.25 V at 100 mV/s. Finally the electrode was

washed with deionized water and dried under N2. In a typical experiment for electrode

preparation, 20 µL of a 2.0 mg/mL dispersion of Fe3O4/APTES-PEG-XO in 50 mM

sodium phosphate buffer, pH 7.0, were mixed with 3.0 µL of 0.5 mg/mL aqueous

dispersion of oxidized SWNT. The mixture was magnetically deposited on the gold

working electrode surface with the neodymium disk magnet, washed with cold 50 mM

sodium phosphate buffer, pH 7.0 and finally dried under N2 before use.


201


Preparation and characterization of the magnetic nanomaterials

Fig. 1. Schematic display of the construction of Fe3O4/APTES-PEG-XO/SWNT magnetic

nanomaterials-based biosensors


202

Figure 1 schematically illustrates the methodology used to prepare the Fe3O4-

based nanomaterials, as well as for the magnetic immobilization of XO-modified

Fe3O4/APTES-PEG/SWNT on the AuSPE surface. Magnetite nanoparticles were first

prepared by co-precipitation of Fe(II) and Fe(III) salts in alkaline media under

continuous ultrasound treatment and mechanical stirring. Through this method, small

and quasi-spherical magnetic nanoparticles with average size of 14 ± 7 nm were

obtained, as revealed by TEM (Figure 2A).

The nanoparticles were then coated with an amino-enriched polysiloxane layer

by treatment with ethanolic APTES solution. The as-synthesized core-shell

Fe3O4/APTES nanoparticles showed similar shape than the uncoated material, but the

average size was slightly increased to 17 ± 9 nm due to the polysiloxane coverage

(Figure 2B).

Fig. 2. TEM images of A) Fe3O4, B) Fe3O4/APTES, C) Fe3O4/APTES-PEG and D)

Fe3O4/APTES-PEG/SWNT magnetic nanomaterials.


203

Monomethoxypolyethylene glycol chains were further attached to the free

amino groups at the nanoparticle surface by reacting with the N-hydroxysuccinimide

derivative of the polymer. The amount of PEG used did not coat completedly the

nanoparticles as it was demonstrated by FT-IR spectra (Figure 3), thus leaving enough

amino groups accesible for the further enzyme immobilization. These polymer-

modified nanoparticles showed similar size and distribution pattern than the parent

core-shell Fe3O4/APTES nanoparticles (Figure 2C).

As it was previously reported, PEG chains can be irreversibly adsorbed on the

side wall of carbon nanotubes.12 In our case, the formation of aggregates in SWNT

aqueous dispersion was accomplished after mixing with Fe3O4/APTES-PEG in a

SWNT/magnetic nanoparticle 1:27 weight ratio (SWNT/Fe3O4/APTES-PEG 1:27) (see

below). The aggregates, which could be magnetically decanted from the bulk solution

by applying an external magnetic field, were not obtained by treating SWNT with

unmodified Fe3O4/APTES nanoparticles, demonstrating that the nanotubes were

attached to the Fe3O4/APTES-PEG nanoparticles through their interaction with the

polymeric chains. This was confirmed by TEM analysis, which revealed that the surface

of SWNT was randomly decorated with the PEG-modified magnetic nanoparticles

(Figure 2D).

The nanomaterials were characterized by FT-IR spectroscopy (Figure 3). The

presence of Fe3O4 nanoparticles was confirmed by the intense absorption bands

around 590 cm-1 and 630 cm-1 in all spectra, resulting from the split of the stretching

vibration band of the Fe-O bonds in bulk magnetite at 570 cm-1.14,18 Magnetic

nanoparticle-based materials also showed a blue-shift of the 2 band of the Fe–O

bonds in bulk magnetite from 375 cm-1 to 448 cm-1.


204

Fig. 3. FT-IR spectra of A) Fe3O4, B) Fe3O4/APTES and C) Fe3O4/APTES-PEG magnetic

nanoparticles.

The presence of the Fe-O-Si bonds, characteristics of the cross-linked

polysiloxane matrix in the core-shell Fe3O4/APTES nanoparticles was not evidenced in

the FT-IR spectrum, because this band appears around 584 cm-1 and therefore overlaps

with the Fe-O vibration band in magnetite.19 However, the presence of the covering

APTES film was confirmed by the broad band around 1020 cm-1, corresponding to the

overlapped Si-O-Si and Si-OH bands. In addition, the presence of the Si-OH groups can

be justified by the broad bands at 3698 cm-1 and 3200 cm-1, which can be ascribed to

the free and hydrogen-bonded hydroxyl groups linked to Si atoms. It should be noted

that the latter band could be also overlapped with the corresponding to N-H vibration,

but the presence of the amino groups in the core-shell Fe3O4/APTES magnetic

nanoparticles was more evident by the band appearing at 1628 cm-1, characteristic of

the NH2 stretching.

Furthermore, the attached polyether chains in the core-shell Fe3O4/APTES-PEG

nanoparticles were confirmed by the bands at 1740 cm-1 and 2918 cm-1, corresponding

to the C=O and C-H stretching, respectively. It should be noted that FT-IR was not able

to identify the Fe3O4/APTES-PEG/SWNT adduct, because the signal corresponding to


205

the oxidized nanotubes were masked by the strongest bands of the modified

nanoparticles (data not shown).

Fig. 4. Magnetization curves of Fe3O4 (), Fe3O4/APTES (), Fe3O4/APTES-PEG (X) and

Fe3O4/APTES-PEG/SWNT () nanomateriales at 298K.

The magnetic properties of the different Fe3O4-based nanomaterials were

measured at room temperature with a Squid magnetometer, and the resulting

magnetization curves are displayed in Figure 4. Superparamagnetic behaviour at room

temperature was always observed for all the samples, with no remaining effect from

the hysteresis loops when the applied magnetic field was removed. The saturation

magnetization of the Fe3O4 nanoparticles was slightly reduced after coating with APTES

and PEG, due to increased mass of the coated magnetic nanoparticles.

In addition, it should be noted that core-shell Fe3O4/APTES-PEG nanoparticles

retained their superparamagnetic properties after non covalent attachment to SWNT.

The significant reduction in the saturation magnetization of the core-shell

Fe3O4/APTES-PEG nanoparticles after SWNT attachment (from 59.5 emu/g to 46.0

emu/g) should be ascribed to the mass increase in Fe3O4/APTES-PEG/SWNT due to the

presence of the nanotubes.


206

Preparation and characterization of the enzyme biosensor

In order to evaluate the potential use of Fe3O4/APTES-PEG/SWNT as support for

enzyme immobilization, a two-step approach was employed. In this regards, core-shell

Fe3O4/APTES-PEG nanoparticles were first pre-activated by treatment with 2.5% (v/v)

glutaraldehyde and further incubated with XO solution. This method ensured that the

ε-amino groups in the enzyme protein structure will only react with the aldehyde

groups at the surface of the nanoparticles, avoiding the formation of intra and

intermolecular protein cross-linking.20 This approach yielded an immobilized enzyme

with high catalytic activity, representing 94% of the original XO activity. The

immobilized biocatalyst contained 82 µg of protein per mg of core-shell Fe3O4/APTES-

PEG magnetic nanoparticle, as determined by Bradford method.15

As mentioned above, the enzyme-modified nanoparticles were finally attached

to SWNT through the irreversible adsorption of PEG chains on the nanotubes. It was

noticed that the specific activity of the immobilized enzyme was not significantly

affected at low concentration of SWNT up to a nanotube/Fe3O4/APTES-PEG-XO weight

ratio of 1:10. This fact could be ascribed to the stabilization of the active enzyme

conformation in the highly hydrophilic microenvironment caused by the PEG chains. In

addition, it could be expected a low steric hindrance of the bulky nanotubes on the

enzyme active site due to the macromolecular and flexible nature of the spacer arms

used to decorate SWNT with the magnetic material via non covalent interactions.


207

Fig. 5. FE-SEM images of A) Fe3O4/APTES-PEG-XO and B) 1:27 SWNT:Fe3O4/APTES-PEG-

XO-modified AuSPE.

This immobilized enzyme form was employed for the magnetic coating of

AuSPE to construct a biosensor device for xanthine. In order to determine the

influence of SWNT on the electroanalytical response of the enzyme electrode,

different amounts of SWNT were mixed with the Fe3O4/APTES-PEG-XO solution to yield

nanotubes/magnetic nanoparticle weight ratios from 1:80 to 1:16. The surface of the

bio-electrodes was characterized using FE-SEM, and representative images are shown

in Figure 5. As it can be observed, magnetic coating of AuSPE with Fe3O4/APTES-PEG-

XO yields a compact packed matrix of modified nanoparticles (Figure 5A), while Figure

5B shows that SWNT were entrapped into this matrix as result of its interaction with

the PEG-coated magnetic nanoparticles.


208

Fig. 6. Impedance plane diagram (−Z″ versus Z′) recorded for AuSPE before () and after

modification with Fe3O4/APTES-PEG-XO (), 1:80 SWNT:Fe3O4/APTES-PEG-XO (), 1:27

SWNT:Fe3O4/APTES-PEG-XO () and 1:16 SWNT:Fe3O4/APTES-PEG-XO (Δ) in 0.1 M KCl

solution containing 5 mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1).

Electrochemical impedance spectroscopy using [Fe(CN)6]4−/3− pair as redox

probe was performed in order to check the effect of the different electrode

modification steps on the electron transfer resistance (Ret) changes at the electrode

surface. Figure 6 shows the Nyquist plots of the different enzyme electrodes. As

expected, the unmodified electrode showed the lowest Ret value, 204 Ω. The further

coating with Fe3O4/APTES-PEG-XO produced a noticeable increase in the Ret (330 Ω)

indicating that the electron transfer of the electrochemical probe was blocked by the

enzyme-modified nanomaterial. This insulating effect was significantly reduced by

coating the electrode with Fe3O4/APTES-PEG-XO/SWNT having different nanotube

loadings, due to the excellent conductivity of the nanotubes. It can be also noted that

the reduction in the electron transfer resistance was dependent on the SWNT content

in the enzyme-modified nanomaterial, with a lower insulating capacity (Ret = 277 Ω)

when 1:27 SWNT/Fe3O4/APTES-PEG-XO was used as bioactive coating material. This

result suggests that PEG-coated nanoparticles reached their maximum capacity to


209

capture SWNT at this concentration ratio. It should be also noted that some amount of

SWNT leaked from the electrode surface during the washing steps when higher

nanotube/nanoparticle ratios were employed, which also supports the latter

statement.

We also analyzed the Warburg impedance contribution at low frequencies for

the bioelectrodes bearing SWNT. The slope values were approximatedly 0.7 in all

cases, which are lower that the theoretical unitary value. This discrepancy is not

unexpected as the electrode process cannot be purely diffusion-controlled considering

the porous nature of the interfase.

Fig. 7. Cyclic voltammograms recorded with AuSPEs modified with: Fe3O4/APTES-

PEG/XO (A); Fe3O4/APTES-PEG-XO (B); 1:80 SWNT:Fe3O4/APTES-PEG-XO (C); 1:16

SWNT:Fe3O4/APTES-PEG-XO (D) and 1:27 SWNT:Fe3O4/APTES-PEG-XO (E) from a 2.5

µM xanthine solution in 0.1 M sodium phosphate buffer, pH 7.0; v = 50 mV/s.

Figure 7 shows the cyclic voltammograms recorded with different AuSPEs

modified electrodes in an aerated 2.5 µM xanthine solution in 0.1 M sodium

phosphate buffer, pH 7.0. As a control, XO physically adsorbed on PEG-modified

magnetic nanoparticles (Fe3O4/APTES-PEG/XO, voltammogram A) was also recorded.


210

The small anodic signal that appeared in this voltammogram suggests that XO was

weakly adsorbed on the nanoparticle surface and easily leaked from the electrode

during the magnetic washing steps. Conversely, a noticeable increase in the anodic

peak current was observed with the Fe3O4/APTES-PEG-XO bioelectrode (curve B),

which is most likely due to the higher amount of enzyme covalently attached to the

glutaraldehyde-activated nanomaterial.

The attachment of Fe3O4/APTES-PEG-XO to SWNT improved the

bioelectrocatalytic response, as it was revealed by the increase in the oxidation signals

(curves C-E). This increase depended on the amount of SWNT in the conjugate

nanomaterial, showing a better voltammetric behaviour for the 1:27

SWNT:Fe3O4/APTES-PEG-XO nanomaterial (curve E). The improved xanthine

bioelectrocatalysis can be attributed to the excellent conductivity of SWNT, providing a

larger and more accessible electroactive surface for the enzyme immobilized in the

three-dimensional matrix. Interestingly, a well defined anodic peak at lower potential

values (400 - 500 mV) appeared when the higher SWNT loadings were used (curves E

and D). This fact may be due to either the favourable oxidation of uric acid at the

modified electrode surface,21 or to the occurrence of direct electron transfer for XO in

the presence of the nanotubes assembly.22

In order to determine the optimal experimental conditions for the biosensor

performance, the effect of pH and the applied potential on the bioelectrode response

toward successive additions of 1.0 mM xanthine was evaluated. The enzyme

electrodes, poised at 600 mV, showed high response in the range of pH 6.5-7.5 and

then neutral pH was selected for further measurements. Moreover, the oxidation

current response increased steadily with the applied potential from 300 mV to 800 mV

versus Ag/AgCl at pH 7.0 (results not shown). Considering the highest signal-to-noise

ratio, an applied potential of 600 mV was selected for further experiments.

Figure 8 shows the calibration curves for xanthine obtained with the same

bioelectrodes mentioned in Figure 7 under the optimized conditions. As expected, the

poorest analytical behaviour was observed for the electrode prepared with the

enzyme physically adsorbed on the PEG-modified magnetic nanoparticle.


211

Fig. 8. Calibration plots for xanthine recorded with Fe3O4/APTES-PEG/XO (),

Fe3O4/APTES-PEG-XO (), 1:80 SWNT:Fe3O4/APTES-PEG-XO (), 1:27

SWNT:Fe3O4/APTES-PEG-XO () and 1:16 SWNT:Fe3O4/APTES-PEG-XO (Δ) modified

AuSPEs; Eapp. = + 600 mV.

The analytical performance was improved by immobilization of XO on

glutaraldehyde-activated nanoparticles. The non-covalent adsorption of Fe3O4/APTES-

PEG-XO to SWNT yielded the best electroanalytical responses, with the maximal

sensitivity achieved when the 1:27 SWNT:Fe3O4/APTES-PEG-XO preparation was

employed. This amperometric biosensor responded rapidly to the changes in xanthine

concentration, attaining 95% of the steady-state current in 12 seconds. The

amperometric response exhibited a range of linearity (r = 0.996) with the analyte

concentration in the 0.25 to 3.5 µM range. This short range can be attributed to a

relative low enzyme loading due to the enzyme immobilization method employed,

using the accessible amino groups on the nanoparticles after PEG coating. The

sensitivity achieved was of 1.31 A/M cm2, which ranks among the higher reported in

the literature for other XO-based electrodes (see Table 1). The biosensor also showed

a low detection limit of 60 nM, calculated according to the 3SD/m criterion, where m is

the slope of the calibration curve and SD is the standard deviation for 12 different 1

µM xanthine amperometric measurements.


212

Table 1. Comparison of analytical properties of the biosensor with previously reported

xanthine biosensors

Electrode E (mV) Linear Range

(µM)

Detection Limit

(µM)

Sensitivity

(mA/M)

KM

(µM)

XO/Pd-Pt/Graphite23 -50a 1.5 - 70 1.5 390 58.5

XO/MWNT/GCE24 -400b 0.1 - 6 0.08 - -

SG/XO/MWNT/GCE24 -400b 0.2 - 10 0.1 - -

XO/laponite/GCE25 +390b 0.039 - 21 0.01 6.54 64

XO/PB/PPy/Au colloid/Au26 -100b 1.0 - 20 - 0.19 43.2

XO/DWNT/CPE27 +900a 2.0 - 50 - 44.1 -

XO/ZnO NPs/PPy/Pt16 +380a 0.8 - 40 0.8 - 13.51

XO/pTTCA/Au28 -350a 0.5 - 100 0.09 5.35 -

XO/CMC-CD/Au29 +700a 300 - 10400 200 - -

SWNT/Fe3O4/APTES-PEG-XO* +600a 0.25 - 3.5 0.06 394 12.8

aAg/AgCl, bSCE, *Present work, PB: Prussian Blue, NPs: nanoparticles, PPy: polypyrrol, DWNT: double-walled carbon nanotubes, MWNT: multi-walled carbon nanotubes, SG: silica sol-gel, pTTCA: poly-5, 2’:5’,2’’-terthiophine-3-carboxylic acid, CGE: glassy carbon electrode, SCE: saturated calomel electrode.

The kinetics constants for the enzyme reaction at the developed bioelectrode

were estimated by using the Eadie-Hofstee plot. According to this analysis, the

immobilized enzyme showed apparent KM and IMAX values of 12.8 µM and 6.3 µA,

respectively. As it is illustrated in Table 1, both the detection limit and the apparent

Michaelis-Menten constant values are comparable with the best ones previously found

in literature using other approaches.

The reproducibility of the measurements carried out with a single biosensor was

calculated by constructing ten successive calibration plots for xanthine with the same

bioelectrode. A relative standard deviation value (RSD) of 6.5% was obtained. The

electrode-to-electrode reproducibility was estimated as 13.6% from the calibration

plots of ten equivalently prepared biosensors. This relatively low electrode-to-

electrode reproducibility could be ascribed to the low reproducibility of disposable

SPEs.


213

The selectivity of the biosensor was evaluated in the presence of eight possible

interfering substances at a 10 µM concentration level: L-glucose, sucrose, ethanol,

acetic acid, lactic acid, citric acid, uric acid and ascorbic acid. The XO-based biosensor

showed good selectivity because, as expected considering the detection potential, only

uric acid and ascorbic acid caused interference to the quantification of 1.0 µM

xanthine, with increase in the anodic current of about 27% and 78%, respectively.

The long-term stability of the biosensor was tested by storing the electrode at 4ºC

under dry conditions and constructing xanthine calibration curves for over 40 days.

The slope value of the calibration plot was practically constant value for the first 10

days and then decreased gradually to 76% of its initial value after 40 days. This rather

good stability of the enzyme electrode can be ascribed to the presence of the highly

hydrophilic macromolecular chains of PEG in the vicinity of the immobilized XO, which

provides an adequate microenvironment for the preservation of both the three-

dimensional structure and catalytic ability of the enzyme. In addition, the multipoint

attachment of XO to glutaraldehyde-activated nanoparticles should contribute to

increase the rigidity of the three-dimensional protein structure, then avoiding the

occurrence of unfolding phenomena over time.

CONCLUSIONS

A novel nanomaterial has been prepared by decorating SWNTs with core-shell

Fe3O4/APTES-PEG nanoparticles via non-covalent interactions. This nanomaterial was

further employed as magnetic support for the immobilization of xanthine oxidase on

AuSPE. The modified electrode was successful evaluated in the design of an

amperometric biosensor toward xanthine with high sensitivity and low detection limit.

According to our results, it can be concluded that SWNT functionalized with core-shell

Fe3O4/APTES-PEG magnetic nanoparticles can be successfully employed for the design

of novel electrochemical biosensing devices.


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ACKNOWLEDGEMENTS





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7.7

Analytical and Bioanalytical Chemistry 405 (2013) 3773-3781

Supramolecular immobilization of glucose oxidase on gold coated with

cyclodextrin-modified cysteamine core PAMAM G-4 dendron/pt nanoparticles for mediatorless

biosensor design

Analytical and Bioanalytical Chemistry 405 (2013) 3773-3781 7. PUBLICACIONES

217

SUPRAMOLECULAR IMMOBILIZATION OF GLUCOSE OXIDASE ON GOLD

COATED WITH CYCLODEXTRIN-MODIFIED CYSTEAMINE CORE PAMAM

G-4 DENDRON/PT NANOPARTICLES FOR MEDIATORLESS BIOSENSOR

DESIGN

Paula Díez,1 Ciprian-George Piuleac,2 Paloma Martínez-Ruiz,3 Santiago Romano,3 María

Gamella,1 Reynaldo Villalonga,1,* José M. Pingarrón1,*


Chemistry, Complutense University of Madrid, 28040-Madrid Spain

2Department of Chemical Engineering, “Gh. Asachi” Technical University, Bd. D.

Mangeron No. 71A, 700050, Iasi, Romania


E-mail: [email protected], [email protected]



218

ABSTRACT

Cysteamine core polyamidoamine G-4 dendron branched with β-cyclodextrins

was chemisorbed on the surface of Au electrodes and further decorated with Pt

nanoparticles. Adamantane-modified glucose oxidase was subsequently immobilized

on the nanostructured electrode surface through supramolecular associations. The so

constructed enzyme electrode was employed for constructing a reagentless

amperometric biosensor for glucose making use of the electrochemical oxidation of

H2O2 generated in the enzyme reaction. The biosensor exhibited a fast amperometric

response (6 s) and a linear response toward glucose concentration between 5 µM and

705 µM. The biosensor showed a low detection limit of 2.0 µM, a sensitivity of 197

mA/M cm2, and retained 94% of its initial response after nine days of storage at 4ºC.

Keywords: Biosensor, β-cyclodextrin, dendrimer, glucose oxidase, Platinum

nanoparticles, supramolecular complex

INTRODUCTION

The development of enzyme-based electrochemical biosensors has received

considerable attention due to their potential use as highly selective analytical devices

in chemical and clinical laboratories, food industry, environmental monitoring and

other related fields [1]. In general, the development and performance of enzyme

biosensors are directly linked to the strategy employed to immobilize the protein on

the transducer surface, as well as to the physicochemical properties of the materials

used to design the electrodes [2].

A major challenge in biosensor technology has been promoted by the use of

nanosized materials in the design of novel electrode architectures [3]. Hard

nanomaterials such as metal and metal oxide nanoparticles [4, 5], graphene [6] and

carbon nanotubes [7] have been widely employed to create reliable biosensor

surfaces. This fact has been supported by the unique properties of these


219

nanomaterials, such as high surface energy and surface-to-volume ratio, the ability to

decrease proteins-metal particles distance, high electroconductivity and protein load

capacity, the ability to catalyze the electrochemical processes associated with several

compounds produced in the enzyme-mediated reactions, and the possibility to act as

electroconductive wires between the enzyme and the electrode substrate [2].

Organic polymer-based nanomaterials, such as dendrimers and dendrons, have

been also used in biosensor design [8,9]. These soft nanomaterials are monodisperse

macromolecules with a regular and highly branched three-dimensional architecture.

They have a high structural homogeneity, a surface reactivity and a molecular host

capacity [10], which are important characteristics for the multipoint immobilization of

enzymes on electrode surfaces. In addition, it has been demonstrated previously that

metal nanoparticles can be stabilized in aqueous media by polyamidoamide (PAMAM)

dentritic structures through the interaction with the primary amino groups at the

polymer surface as well as by association with the amide and tertiary amine groups

into the dendrimer cavities [11,12]. These properties can be profited to prepare stable

hybrid nanomaterials based on metal nanoparticles/PAMAM dendritic structures for

suitable modification of electrode surfaces and further construction of electrochemical

enzyme biosensors [13,14].

On the other hand, functionalization of solid surfaces with enzymes and other

proteins has not only played a key role in biosensor technology but has also in the

future development of advanced biomaterials, biotechnological processes and

biologically inspired nanomaterials. Generally, proteins are able to be immobilized on

solid surfaces through covalent linkages, physical entrapment/encapsulation and

adsorption. Although covalent attachment and physical entrapment/encapsulation

provide a higher stability, the enzymatic activity is often reduced due to the harsh

reaction conditions and large distance between substrates and enzymes, respectively.

On the contrary, enzymes present an easily capability to be immobilized under mild

conditions by adsorption strategies, but the stability of such adducts is low and the

proteins can be often easily released from supports. An alternative approach to

immobilize enzymes on solid supports is the formation of multipoint supramolecular


220

complexes based on the complementary host-guest properties of β-cyclodextrin (CD)

and 1-adamantane derivatives [15]. Such supramolecular strategies have been

previously employed in the successful construction of amperometric enzyme

biosensors [16-18].

In this work it was proposed the modification of gold surfaces with CD-modified

PAMAM dendron/Pt nanoparticles (PtNPs), and the further supramolecular

immobilization of adamantane-modified glucose oxidase (GO, EC 1.1.3.4) on the hybrid

nanomaterial. The rational of this original approach is based on the use of a thiol

containing CD-modified PAMAM dendron derivative as permeable and stable coating

material for the electrode surface. This hyperbranched polymer showed molecular

receptor capacity due to the branched CD moieties, favoring the formation of host-

guest supramolecular complexes with adamantane-modified enzymes. The PAMAM

dendron derivative also showed capacity to stabilize PtNPs, allowing the preparation of

an inorganic/organic hybrid nanomaterial able to catalyze the transformation of H2O2

thus the construction of a third generation GO-based biosensor toward glucose. GO

was selected as model enzyme due to its importance in biosensor technology for

glucose detection [19] as well as extensively studied on GO-based amperometric

biosensors have been accomplished, thus allowing a reliable comparison of the new

biosensor design’s performance, involving the hybrid nanomaterial and

supramolecular enzyme immobilization.



Glucose oxidase, cystamine core PAMAM G-4 dendrimer and CD were

purchased from Sigma-Aldrich Co. (USA). All other chemicals (analytical grade or

higher) were used.

The amperometric measurements were performed with a dual-channel

ultrasensitive Inbea amperometric detector (Inbea Biosensores S.L., Spain). The cyclic


221

voltammetry (CV) and the electrochemical impedance spectroscopy (EIS) experiments

were carried out using a FRA2 µAutolab Type III potentiostat/galvanostat. Also, the

data were acquired using a GPES Ver. 4.9 and a Frequency Response Analyser software

(Metrohm Autolab B.V., The Netherlands), respectively. A conventional three-

electrode system was applied in all electrochemical studies. A gold disk (CHI

Instruments, UK, 2.0 mm diameter) modified with the CD-PAMAM dendron/PtNPs

hybrid nanomaterial and the enzyme was considered as working electrode. An

Ag/AgCl/KCl (3 M) and a Pt wire were used as reference and counter electrode,

respectively. All the measurements with the modified electrode were carried out at

25ºC in 0.1 M sodium phosphate buffer and pH 7.0 (working volume 10 mL).

Amperometric measurements were accomplished in stirred solutions at 300 rpm. 5.0

mM glucose solutions in 50 mM sodium phosphate buffer, pH 7.0 were freshly

prepared.

The transmission electron microscopy (TEM) measurements were performed

with a JEOL JEM-2100 microscope (JEOL Ltd., Japan). The surface of the nanostructured

electrode was investigated by high resolution field emission scanning electron

microscopy (SEM) implying a JEOL JSM-6335F apparatus (JEOL Ltd., Japan). The

morphology of the gold modified surface was studied by atomic force microscopy

(AFM) with a SPM Nanoscope IIIa multimode microscope (Veeco Instruments Inc., USA).

Surface plasmon resonance (SPR) measurements were carried out at a fixed wavelength

of 670 nm by means of a Springle Autolab-SPR equipment from Metrohm Autolab B.V.

(The Netherlands). 1H NMR characterizations were performed with a Bruker Avance 500

MHz spectrometer (Bruker BioSpin GmbH, Germany).

2. Synthesis of CD-modified cysteamine core PAMAM G-4 dendron (CD-PAMAM G-4)

Fifty milligrams of cystamine core PAMAM G-4 dendrimer and 510 mg of mono-

6-O-tosyl CD (CDTs) were dissolved in 100 mL of deoxygenated DMSO, and the mixture

was stirred at room temperature under N2 atmosphere during four days. After that,

the solution was concentrated in vacuum at the limit of 5 mL, diluted with 150 mL


222

distilled water and, finally, dialyzed vs. distilled water based on Amicon Ultra-15

centrifugal filter units with Ultracel-10 membranes (Millipore, USA). Subsequently, the

solution was concentrated at the limit of 20 mL and treated with NaBH4 solution (10

mM final concentration) for 2 h under magnetic stirring. After this procedure, the

mixture was dialyzed and concentrated by Amicon centrifugal tubes as described

above, yielding a CD-PAMAM G-4 solution of 4 mg/mL final concentration. The

characterization of CD-PAMAM G-4 dendron has been accomplished through a 1H NMR

in D2O. Yield: 93 mg.

3. Preparation of the GO-ADA/CD-PAMAM/PtNP/Au enzyme electrode

First of all, GO was modified with 1-adamantanecarboxylic acid (GO-ADA) as

described previously [16], and concentrated with Amicon centrifugal tubes up to a

final concentration of 5 mg/mL in 50 mM sodium phosphate buffer, pH 7.0. A disk gold

electrode was polished with alumina powder (0.3 µm), then rinsed with double

distilled water and immersed in an ultrasonic bath for 5 min. The electrode was further

dipped in boiling 8 M KOH solution for 1 h, washed with distilled water and immersed

in piranha solution for 30 min. After a new washing it was electrochemically treated by

cyclic voltammetry in 0.1 M H2SO4 solution through twenty cycles between -0.2 V and

1.85 V at a scan rate of 100 mV/s. The cleaned gold surface was firstly dipped into 10

mL of a 0.5 mg/mL CD-PAMAM G-4 aqueous solution to coat the metal surface with a

CD-branched dendron derivative monolayer. After 2 h of incubation, the modified

electrode was washed several times with distilled water and subsequently dipped into

10 mL of a 10 mM H4PtCl6 solution. After 10 min of incubation under continuous

magnetic stirring, 1.0 mL of 50 mM NaBH4 solution was added to cause PtNPs

formation. The electrode was kept into the stirred solution for 10 min, then

exhaustively washed with distilled water, and finally coated with 20 µL of a 5 mg/mL

GO-ADA solution. The enzyme immobilization was accomplished by allowing the

incubation to proceed for 2 h at 4ºC. Finally, the enzyme electrode was washed with

cold 50 mM sodium phosphate buffer, pH 7.0, solution and kept at 4ºC in this solution

until use.


223


Preparation and characterization of the enzyme electrode.

The employed strategy to the Au electrode surface functionalized with the CD-

branched PAMAM G-4 dendron/PtNPs hybrid nanomaterial and its further use as

support for the supramolecular immobilization of GO-ADA is schematically illustrated in

Figure 1.

Figure 1. Schematic display of the steps involved in the preparation of GO-ADA/CD-

PAMAM/PtNP/Au based enzyme biosensors.

Firstly, cystamine core PAMAM G-4 dendrimer was reacted with mono-6-O-

tosyl-βCD in order to introduce molecular receptor functionalities at the dendrimer

surface, using the primary amino groups at the dendrimer shell as branching point. The

CD-modified cystamine core PAMAM G-4 dendrimer was then reduced with NaBH4 to

cleave the disulfide bond at the dendrimer core. The resulting water-soluble CD-

branched cysteamine core PAMAM G-4 dendron contained an active thiol group able

to be chemisorpted on the gold electrode surface.

This hyperbranched polymer derivative was characterized by 1H NMR (See

Figure 1S in Supporting Information). It was observed nine CD units were attached to


224

each PAMAM G-4 dendron molecule due to the modification of 28% of the primary

amino groups in the polymer. This partial modification could be justified through the

steric hindrance caused by the bulky CD moieties attached to the Dendron surface. It

avoided the further reaction of other mono-6-O-tosyl-βCD molecules with the

remaining amino groups at the modified dendritic molecule.

Figure 2. SPR sensogram recorded on Au surfaces upon successive incubation with CD-

PAMAM G-4 and GO-ADA (a), PAMAM G-4 and GO (b) and CD-PAMAM G-4 and GO (c).

The capability of this dendron derivative to act as a host receptor for the

supramolecular recognition of adamantane-modified GO was evaluated by SPR

measurements. Figure 2 shows the changes produced in the SPR signal of gold surface

after sequential exposure to different cysteamine core PAMAM G-4 dendron and GO

derivatives. Incubation with the two different cysteamine core PAMAM G-4 dendron

derivatives (with and without CD attached) has produced a similar and noticeable

increase in the SPR signal, being scarcely affected upon after washing. This result

suggested the thiol-containing dendron derivatives were successfully chemisorbed on

the Au surface and the process was not significantly affected by the presence of the

branching CD moieties at the surface of the dendritic structure.


225

Further, the incubation of the Au surface coated with the CD-modified dendron

in the adamantane-modified GO solution at pH 7.0 caused a further significant

increase in the SPR signal (sensogram a), thus demonstrating the interaction between

the functionalized enzyme molecules and the surface capped with the CD-modified

dendron. Based on the presence of bulky CD groups at the dendron surface, it could be

predicted the contribution of host-guest supramolecular association to the whole

interaction observed by SPR. However, a similar increase in the SPR signal was

observed for the Au surface coated with cysteamine core PAMAM G-4 dendron after

incubation with native GO (curve b). It has been reported that the primary and tertiary

amino groups located at the surface and the cavity of PAMAM G-4 dendrimers have

pKa values of 9.0 and 5.8, respectively [20]. Thus, PAMAM G-4 dendrimers are able to

bear positive charge by protonation of the primary amines at the rim at pH 7.0.

Moreover, the isoelectric point of native GO is 4.2 and, consequently, it is negatively

charged at pH 7.0. As a consequence, GO presents a good capability to be immobilized

on the PAMAM G-4 dendron-coated Au surface through polyelectrostatic interactions.

On the other hand, loading of native GO on the Au surface coated with the CD-

modified dendron was less favored, according to the low increase observed in the SPR

signal (curve c). The result leads to the modification of primary amino groups with the

bulky oligosaccharide moieties at the surface of the dendron, reducing the positive

electrostatic field around this polymer. Also, it caused a noticeable steric hindrance

and decreased then the interaction of the dendron with the GO molecules.

The CD-PAMAM G-4 coated gold surface was decorated with Pt nanoparticles.

The procedure described in the Materials and Methods section produced small and

spherically-shaped Pt nanoparticles with an average size of 2.6 ± 0.4 nm, as revealed

by TEM. As it can be shown in SEM analysis (Figure 3), the dendron-modified gold

surface was coated with a stable three-dimensional arrange of Pt nanoparticles, where

the protuberant nanostructures are combined with deep nanoholes. The high density

of Pt nanoparticles avoided characterization of this surface by SPR measurements.


226

Figure 3. Field emission SEM image of the CD-PAMAM G-4/PtNP-modified Au surface.

This modified Au surface was further used as support for the immobilization of

the adamantane-modified GO derivative. The changes on the topology and the

electrochemical characteristics of the Au surface after the different modifications steps

were checked by AFM, EIS and CV.

Figure 4 presents the tapping mode AFM images for different gold surfaces.

The characteristic flat topology by an average roughness of 0.9 nm and an average

height of 3.2 nm with a maximum at 5.6 nm on the unmodified gold surface was

observed.

Coating with the CD-PAMAM dendron derivative yielded a modified surface

pattern with ellipsoidal nanostructures, an average roughness of 3.1 nm and an

average height of 12.0 nm with a maximum at 18.6 nm. After the deposition of Pt

nanoparticles on the dendron-modified surface, a more irregular pattern was observed

exhibiting a higher density of protuberant nanostructures with an average and a

maximum height value of 34.6 nm and 92.9 nm, respectively. Moreover, an average

roughness of 15.1 nm was estimated for the Pt nanoparticles-modified surface.


227

Figure 4. Three dimensional AFM analysis of the Au surface before (A) and after

modification with CD-PAMAM (B), CD-PAMAM/PtNP (C) and GO-ADA/CD-

PAMAM/PtNP (D).

This surface was softened after immobilization of the enzyme derivative. In

fact, the average roughness and height, respectively, decreased to around 10.3 nm and

19.2 nm after the incubation in the GO-ADA solution. The maximum height value for

the enzyme-coated surface was also reduced to about 61.7 nm. These results

suggested the enzyme molecules were homogeneously distributed over the entire

PtNP-coated surface, filling the nanoholes created by these nanomaterials.

The influence of the different modification steps on the barrier properties of

the gold surface was evaluated by electrochemical impedance spectroscopy in a 0.1 M

KCl solution containing 5 mM [Fe(CN)6]4−/3−. Figure 5 shows the resulting Nyquist plots,

with a Ret = 118 Ω for the bare gold surface and a slope value of 0.89 for the straight

line observed over the broad range of low frequencies. It is indicated a fast electron

transfer for the [Fe(CN)6]4−/3− ions pair on the electrode surface.


228

Figure 5. Nyquist plots of bare Au (a), CD-PAMAM/Au (b), CD-PAMAM/PtNPs/Au (c)

and GO-ADA/CD-PAMAM/PtNP/Au (d) electrodes in 0.1 M KCl solution containing 5

mM K3[Fe(CN)6]/K4[Fe(CN)6] (1:1).

A small increase in the electron transfer resistance, Ret = 195 Ω, and a linear

behavior at low frequencies with similar slope value were observed for the electrode

surface coated with the CD-modified dendron derivative. That is suggesting a low

barrier effect of the dendron molecules on the electron transfer process at the

electrode surface most likely due to the permeable and polycationic nature of the

polyamidoamine polymer. Also, the Nyquist plots revealed the electron transfer

resistance was decreased (Ret = 29 Ω) after modification with the highly conductive Pt

nanoparticles. However, as it was expected, an opposed effect was evident after

immobilization of adamantane-modified GO, producing a noticeable increase in the

diameter of the semicircle at high frequency values (Ret = 639 Ω). In addition, the slope

value of the straight line observed at low frequencies also decreased to 0.58. This

higher resistance to the electron transfer and diffusion-controlled processes suggested

a high coverage of the electrode surface by the non-conductive enzyme molecules.


229


K3[Fe(CN)6]/K4[Fe(CN)6] (1:1) at bare Au (a), CD-PAMAM/Au (b), CD-PAMAM/PtNPs/Au

(c) and GO-ADA/CD-PAMAM/PtNP/Au (d) electrodes. Scan rate: 50 mV/s.

The EIS results were confirmed by cyclic voltammentry at the different

electrode architectures in 5 mM [Fe(CN)6]4−/3− + 0.1 M KCl solution. As it can be

observed in Figure 6, the unmodified gold electrode showed well-defined typical quasi-

reversible diffusion-limited patterns with ∆E = 99 mV and ia/ic = 1.07 values. The

electrochemical surface area of the bare electrode was calculated to be 3.6 mm2 by

using the Randles-Sevcik equation. Coating with CD-PAMAM dendron derivative

caused a small decrease in both the anodic and cathodic peaks, then reducing the

calculated value for the electrochemical surface area to around 3.4 mm2. In addition,

the ∆E and ia/ic values were increased to 129 mV and 1.23, respectively.

A more reversible diffusion-limited pattern was observed for the electrode

after modification with Pt nanoparticles, with larger anodic and cathodic peaks current

values, ∆E = 96 mV and ia/ic = 1.10. Also, the metal nanoparticles increased slightly the

electrochemical surface area of the electrode to a value of 3.7 mm2. This voltammetric

behavior is attributable to the presence of the highly electroconducting Pt

nanoparticles. The subsequent immobilization of the enzyme derivative on the


230

electrode surface provoked a drastic decrease in the anodic and cathodic peak current

as well as in the electrochemical surface area (1.74 mm2). The enzyme loading has also

affected the quasi-reversible behavior of the [Fe(CN)6]4−/3− electrochemical reaction at

the electrode surface, with ∆E = 358 mV and ia/ic = 1.26 values. These results suggested

again a high coverage of the electrode surface by the non-conductive and bulky

enzyme molecules.

Figure 7. Cyclic voltammograms recorded in 0.1 M sodium phosphate buffer, pH 7.0, at

a scan rate of 50 mV/s, for GO-ADA/CD-PAMAM/PtNP/Au electrode before (a) and

after (b) addition of 200 µM glucose.

Figure 7 shows the cyclic voltammograms recorded at the enzyme-modified

electrode in aerated 0.1 M sodium phosphate buffer, pH 7.0, solution at a scan rate of

50 mV/s, before and after addition of glucose. The peak observed in the absence of

glucose was due to the dissolved oxygen in solution. The addition of glucose produced

the appearance of anodic and cathodic peaks suggesting that the H2O2 formed in the

enzyme reaction can be electrocatalytically oxidized and reduced at the Pt

nanoparticles-modified electrode surface.


231

It has been recently proposed that H2O2 is able to be electrochemically

transformed at the Pt surface through the following mechanisms [21]. For reduction,

H2O2 is firstly dissociated to adsorbed OH on two oxide-free Pt sites in a non-

electrochemical step. Since adsorbed OH on Pt is not stable at low potentials, the

surface sites are rapidly regenerated in an electrochemical reduction step, leaving free

the Pt sites to dissociate other H2O2 molecules.

2Pt + H2O2 2Pt(OH)

2Pt(OH) + 2H+ + 2e- 2Pt(H2O)

For oxidation at sufficiently positive potentials, where oxygenated species are

adsorbed on Pt sites, H2O2 is firstly oxidized on two OH-covered Pt sites to O2 in a non-

electrochemical step. Since the oxide-free Pt surface is not stable at high positive

potentials, the reduced surface sites are then electrochemically re-oxidized and

become available again to oxidize H2O2.

According to the cyclic voltammetric behavior at the GO bioelectrode, it can be

concluded that H2O2 can be electrocatalytically oxidized at potential values higher than

+340 mV. On the other hand, an electrocatalytic reduction of H2O2 can be also

observed at potential values lower than +230 mV. These findings suggested that it was

feasible to design a mediatorless electrochemical biosensor for glucose, able to work

at low potentials, by using the proposed electrode surface.

The GO-ADA/CD-PAMAM/PtNP/Au electrode was further evaluated for the

amperometric quantification of glucose. For this purpose, optimum working conditions

were firstly evaluated and selected. Maximum amperometric response towards

glucose was achieved in buffer solutions of pH 6.0 - 7.0. Additionally, poor signal-to-

noise current ratios were obtained in the cases of potential values lower than +200

mV. Best results were achieved by detecting the electrochemical oxidation of the


232

enzymatically-produced H2O2 at potential values higher than +350 mV. In order to

minimize the undesirable effect of common interfering substances during the

amperometric detection of glucose, it was selected the lowest anodic potential value

at which a good signal-to-noise ratio was observed. Summarizing, further

amperometric measurements were performed in 0.1 M sodium phosphate buffer of

pH 7.0 at a constant potential of +400 mV vs Ag/AgCl.

Figure 8. Amperometric responses recorded with GO-ADA/CD-PAMAM/PtNP/Au (a),

GO/PAMAM/PtNP/Au (b), GO/CD-PAMAM/PtNP/Au (c) and GO/PtNP/Au (d) electrodes

upon successive additions of 5.0 mM glucose. Eapp.= + 400 mV.

Under these conditions mentioned above, the dynamic amperometric response

of the bioelectrode upon successive additions of 5.0 mM glucose is illustrated in Figure

8 (curve a). As control measures, Au electrodes coated with PAMAM/PtNP (curve b),

CD-PAMAM/PtNP (curve c), PtNP (d) and PAMAM (data not shown), on which an

equivalent activity of native GO was immobilized under similar conditions, were also

checked. The GO-ADA/CD-PAMAM/PtNP/Au-based biosensor showed fast catalytic

response, reaching 95% of the steady-state current in about 6 s. Not significant

increase in the anodic current was observed for the GO/PAMAM/Au electrode,

demonstrating that PtNPs are needed for the electrocatalytic oxidation of H2O2 at +400


233

mV. The amperometric responses of the control electrodes were significantly lower,

suggesting higher amount of immobilized active enzyme on the GO-ADA/CD-

PAMAM/PtNP/Au electrode, probably through host-guest and electrostatic

interactions.

The biosensor showed a linear behavior for glucose in the 5 to 705 µM

concentration range fitting to the equation (n = 10, r = 0.999)

ia(mA) = 7.1×c(Glucose/M) + 7·10-5

A similar range of linear response was observed for the control

GO/PAMAM/PtNP/Au electrode. However, a shorter range (5 – 470 µM) was found for

the GO/CD-PAMAM/PtNP/Au biosensor. The equations describing the linear response

of these control biosensors were:

ia(mA) = 2.4×c(Glucose/M) + 1·10-5 (GO/PAMAM/PtNP/Au)

ia(mA) = 1.3×c(Glucose/M) + 6·10-6 (GO/CD-PAMAM/PtNP/Au)

The sensitivity of the GO-ADA/CD-PAMAM/PtNP/Au biosensor was determined

to be 197 mA/M cm2, considering the electroactive area of the gold electrode. A

remarkable lower sensitivity was obtained with the control enzyme electrodes, with

values of 66.7 mA/M cm2 and 36.1 mA/M cm2 for the GO/PAMAM/PtNP/Au and

GO/CD-PAMAM/PtNP/Au, respectively. This result has indicated again a lower enzyme

loading on the control biosensors. It is suggesting the enzyme immobilization on the

different Pt nanoparticles-modified surfaces followed a similar behavior that that

deduced from the SPR experiments.

The GO-ADA/CD-PAMAM/PtNP/Au biosensor allowed a detection limit for

glucose of 2.0 µM, calculated according to the 3Sb/m criterion, where m was the slope

value of the linear calibration plot and Sb was estimated as the standard deviation

(n = 10) of the signals corresponding to the lowest glucose concentration able to be

quantitatively measured with the biosensor. In addition, the apparent Michaelis-

Menten constant of the biosensor toward glucose was estimated as KMap = 500 µM,

which was much lower than the previously reported for Aspergillus niger glucose


234

oxidase at pH 5.6 (KM = 33 mM) [22], thus indicating that the affinity of the substrate

for the enzyme was not worsened by using the supramolecular immobilization and the

developed electrode architecture.

Table 1. Comparison of the analytical performance of the GO-ADA/CD-

PAMAM/PtNP/Au biosensor with that reported previously for mediatorless glucose

biosensors

Electrode E

(mV)

Linear

Range (µM)

D.L.

(µM)

Sensitivity

(mA/M·cm2)

KM

(mM) Ref.

Naf/GO/MWNT/Al2O3@SiO2/GCE -488a 17.5 - 800 17.5 127.0 0.5 [23]

GO/B-MWNT/GCE -490b 50 - 300 10.0 111.6 0.2 [24]

Naf/GO/Ag-Pdop@MWNT/GCE -496b 50 - 1100 17.0 43.8 4.56 [25]

GO/AuNPs/MWNT/PVA/GCE -400b 500 - 8000 200 16.6 - [26]

[GO/PDDA]3/[SDS-

MWNT/PDDA]3/MPS/Au/Ti/PET

+600a 20 - 2200 10 5.6 5.15 [27]

GO/AuNPs/DMF/BMIMPF6/GCE -800b 0.1 - 103 0.1 595 0.0035 [28]

GO/PPyAA/AuNPs/GCE +600a 50 –

1.6x104

50 14 1.83 [29]

Naf/GO/NP-Au -200b 10 –

2.2x104

10 - - [30]

GO/PtNPs/MWNT/GCE +700a 0.25 - 104 0.25 431 - [31]

GO/Au/GCE -550b 2.5 – 157.5 0.32 - 0.016 [32]

GO/PMB@SiO2/GCE -430b 10 - 1110 3.0 - 0.5 [33]

GO-ADA/CD-PAMAM/PtNPs/Au +400a 5 - 705 2.0 197 0.5 This

work

aAg/AgCl, bSaturated calomel electrode. D.L.: Detection Limit, Naf: Nafion, MWNT: multi-walled carbon nanotubes, CGE: glassy carbon electrode, B-MWNT: boron-doped MWNT, Pdop: polydopamine, NPs: nanoparticles, PVA: polyvinyl alcohol, PDDA: polydiallyldimethylammonium chloride, SDS: sodium dodecylsulfate, MPS: 3-mercapto-1-propanesulfonic acid, PET: poly(ethylene terephthalate), BMIMPF6: 1-butyl-3-methylimidazolium hexafluophosphate, PPyAA: poly(pyrrole propylic acid), NP-Au: nanoporous Au, PMB: poly(methylene blue).

Table 1 shows the comparison of the analytical characteristics provided by the

GO-ADA/CD-PAMAM/PtNP/Au biosensor with those reported recently for other

mediatorless glucose oxidase biosensors. As it can be deduced, both, the detection


235

limit and sensitivity, and apparent Michaelis-Menten constant are among the best

values reported. They are only surpassed by biosensors using extreme working

potentials and then more susceptible to be interfered by other analytes during glucose

determination.

The good electroanalytical behavior of the GO-ADA/CD-PAMAM/PtNP/Au

biosensor could be assigned to the high amount of immobilized enzyme, as well as the

cooperative contribution of several factors on the maintenance of the active enzyme

protein conformation on the electrode surface. In particular, the supramolecular

strategy used for GO immobilization as well as the presence of metal nanoparticles

should avoid the structural deformation of the active globular enzyme form upon

immobilization on the electrode surface. In addition, the presence of highly hydrophilic

CD-PAMAM G-4 dendron units on that surface should contribute to keep a hydrophilic

microenvironment around the enzyme. The occurrence of randomly-arranged

nanocavities originated by the Pt nanoparticles should also ensure a high population of

enzyme immobilized into these nanoholes being expected that such nanostructures

should protect the hydrophilic microenvironment around the immobilized enzyme.

The repeatability of the measurements carried out for 25 µM glucose with one

single GO-ADA/CD-PAMAM/PtNP/Au electrode (n = 10) yielded a relative standard

deviation (RSD) value of 4.7%. The electrode-to-electrode reproducibility was

calculated from the responses obtained with ten different GO-ADA/CD-

PAMAM/PtNP/Au electrodes prepared in the same manner toward the same glucose

concentration yielding a RSD value of 7.2%.

The selectivity of the enzyme electrode was evaluated by measuring the

amperometric response for 25 µM glucose in the presence of seven possible

interfering substances. Fructose, galactose, sucrose, arabinose, cysteine, citric acid and

caffeine at 100 µM concentration did not cause significant changes in the steady-state

current signal of glucose. However, as expected, the amperometric response of the

biosensor toward glucose was affected in 2.1% and 20% after addition of uric acid and

ascorbic acid at a 2.5 µM concentration, respectively.


236

Regarding the effect of the biosensor storage time at 4ºC under dry conditions

on the amperometric response of the biosensor, it was observed that the biosensor

retained more than 94% of its initial activity after 9 days of storage. Longer storage

times provoked a gradual loss of the biosensor activity, retaining about 45% of the

initial amperometric response after one month. It should be also noted that the

activity of the biosensor was reduced to about 35% upon two hours of incubation in a

saturated solution of 1-adamantane carboxylic acid, indicating the disruption of the

supramolecular host-guest interactions between the modified enzyme molecules and

the CD-branched dendron derivatives at the electrode surface.

CONCLUSIONS

A novel electrode surface involving gold coated with PtNPs-decorated CD-

branched cysteamine core PAMAM G-4 dendron was used as support for the

immobilization of adamantane modified glucose oxidase, mainly through the

formation of host-guest supramolecular associations. The enzyme electrode was

evaluated for the construction of a third generation biosensor for glucose. The

biosensor was highly selective and presented a good sensitivity and stability, as well as

a low detection limit and a fast electroanalytical response toward glucose. Attending

to these results, it was suggested the use of Au surface modified with this

inorganic/organic hybrid nanomaterial as a useful electrode platform for preparing

mediatorless oxidase-based amperometric enzyme biosensors.

ACKNOWLEDGEMENTS


of Science and Innovation. Financial support from the Spanish Ministry of Science and

Innovation CTQ2011-24355, CTQ2009-12650, CTQ2009-09351 and Comunidad de

Madrid S2009/PPQ-1642, programme AVANSENS are gratefully acknowledged.


237

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SUPRAMOLECULAR IMMOBILIZATION OF GLUCOSE OXIDASE ON GOLD COATED WITH

CYCLODEXTRIN-MODIFIED CYSTEAMINE CORE PAMAM G-4 DENDRON/PT

NANOPARTICLES FOR MEDIATORLESS BIOSENSOR DESIGN

Paula Díez,1 Ciprian-George Piuleac,2 Paloma Martínez-Ruiz,3 Santiago Romano,3 María

Gamella,1 Reynaldo Villalonga,1,* José M. Pingarrón1,*



2Department of Chemical Engineering, “Gh. Asachi” Technical University, Bd. D.

Mangeron No. 71A, 700050, Iasi, Romania

Figure 1S. 1H NMR spectrum of CD-branched cysteamine core PAMAM G-4 dendron.

7.8

Electrochimica Acta 76 (2012) 249-255

Layer-by-layer supramolecular architecture of cyclodextrin-

modified pamam dendrimers and adamantane-modified

peroxidase on gold surface for electrochemical biosensing


243

LAYER-BY-LAYER SUPRAMOLECULAR ARCHITECTURE OF CYCLODEXTRIN-MODIFIED

PAMAM DENDRIMERS AND ADAMANTANE-MODIFIED PEROXIDASE ON GOLD

SURFACE FOR ELECTROCHEMICAL BIOSENSING

Reynaldo Villalonga, Paula Díez, María Gamella, A. Julio Reviejo, Santiago Romano,

José M. Pingarrón*

Department of Analytical Chemistry & 2Department of Organic Chemistry I, Faculty of





244

ABSTRACT

A new layer-by-layer supramolecular approach for the construction of self-

assembled nanoarchitectures of polyamidoamine (PAMAM) dendrimers and

peroxidase on gold surface is reported. The methodology is based on the

supramolecular self-assembly of alternated layers of adamantane-modified

horseradish peroxidase and β-cyclodextrin-branched PAMAM G-5 dendrimers on a

gold electrode, previously coated with β-cyclodextrin-modified cysteamine core

PAMAM G-4 dendron. The formation of layer-by-layer assemblies (up to three

dendrimer/peroxidase bilayers) was studied by SPR, quartz crystal microbalance, AFM

and cyclic voltammetry. The analytical applicability of these architectures was

evaluated by constructing a H2O2 biosensor. The electroanalytical response of the

biosensor towards H2O2 increased with the number of enzyme layers. The bioelectrode

constructed with three enzyme layers showed a low detection limit of 160 nM, a

sensitivity of 602 µA/M cm2 and retained 63% of its initial activity after 30 days of

storage in wet conditions.

KEYWORDS: Biosensor, β-cyclodextrin, dendrimer, horseradish peroxidase, layer-by-

layer, supramolecular complex.

INTRODUCTION

The design of hybrid biomolecule/nanomaterial-based three-dimensional

architectures on metal surfaces is a current research priority with the aim of

developing novel biosensing devices, functional biomaterials and efficient biocatalysts

[1-4]. These nanostructured constructions should ensure the accurate interaction of

the modified surface with the biomolecule without affecting its biological activity. In

general, the successful preparation of these hybrid arrangements depends largely on

the surface properties, the characteristics of the selected nanomaterials, the chemical


245

and biological properties of the target biomolecules, the immobilization strategy and

the sequential approach used to construct such 3D assemblies.

Solid state nanomaterials have been largely employed to modify metal

surfaces, offering a great variety of three-dimensional nanosized architectures for

biomolecule immobilization [5, 6]. However, the native conformation of proteins and

other biomolecules is often affected at the interface with solid nanomaterials leading

to a lack of functional activity [7, 8]. Therefore, the evaluation of non-rigid nanosized

materials as alternative 3D scaffolds for the construction of biofunctionalized

nanoarchitectures on solid surfaces is receiving considerable attention [9, 10].

Dendrimers are monodisperse synthetic polymers with a regular and highly

branched three-dimensional structure [11, 12]. These hyperbranched soft

nanomaterials have been widely used in the functionalization of solid surfaces for the

preparation of biosensor systems [9, 10]. Such applications have been supported by

the unique structural properties of dendrimers, such as structural uniformity, globular

shape, nanometric size, monodispersity, high density of functional groups at the

surface and high permeability of the internal cavities. In addition, the possibility to

manipulate such properties by controlling chemical transformation or tailor-made

designing of dendrimers with specific composition offers versatile possibilities to these

polymers as 3D scaffolds for constructing nanostructured designs at the electrode

surfaces [9-12].

Layer-by-layer arrangements on solid surfaces, mainly using electrostatic

interactions as assembling forces, have been reported for dendrimer and

dendrimer/protein hybrid based architectures [9, 13, 14]. However, at our knowledge,

the use of host-guest interactions for the layer-by-layer supramolecular self-assembly

of proteins and dendrimers on metal surfaces has not been described yet. In previous

works, we have reported supramolecular-based strategies for immobilizing

adamantane-modified enzymes and other proteins on nanomaterials and metal

electrodes, previously capped with β-cyclodextrin (CDs) derivatives [15-18]. Similar

supramolecular approaches have been also employed in the formation of layer-by-


246

layer architectures of enzymes and enzyme/nanoparticle on metal surfaces via host-

guest interactions [19, 20].

In this work we propose a novel supramolecular approach for the preparation

of hybrid dendrimer/enzyme nanostructured architectures on Au surfaces, through the

layer-by-layer self-assembly of chemically-modified horseradish peroxidase (HRP, EC

1.11.1.7, H2O2 oxidoreductase) and CD-branched PAMAM dendrimers via host-guest

interactions. CD cavity functions as a macrocyclic receptor with the ability of including

hydrophobic guest compounds with the appropriate geometry such as enzyme-

adamantane derivatives, forming stable supramolecular inclusion complexes. HRP was

selected as model enzyme due to its importance in biosensor technology for H2O2

determination and as a second enzyme for the quantification of selected compounds

transformed by oxidases [21].



HRP (Type II, 250 U/mg, d = 2.5 - 3.0 nm), NaBH4, CD, cystamine core PAMAM

G-4 dendrimer (d = 4.5 nm) and ethylenediamine core PAMAM G-5 dendrimer (d = 5.4

nm) were purchased from Sigma-Aldrich Co. (USA). All other chemicals were of

analytical grade.

Amperometric measurements were performed with a dual-channel

ultrasensitive Inbea potentiostat (Inbea Biosensores S.L., Spain). Cyclic voltammetry

and electrochemical impedance spectroscopy experiments were performed using a

FRA2 µAutolab Type III potentiostat/galvanostat and the data were acquired using

GPES Ver. 4.9 and Frequency Response Analyser softwares, respectively (Metrohm

Autolab B.V., The Netherlands). A conventional three-electrode system was employed

in all electrochemical studies. The working electrode was a gold disk (CHI Instruments,

UK, 2.0 mm diameter) modified with the hybrid dendrimer/HRP layer-by-layer

architectures. An Ag/AgCl/KCl (3 M) and a Pt wire were used as reference and counter


247

electrodes, respectively. Bioelectrode measurements were carried out at 25ºC in 0.1 M

sodium phosphate buffer, pH 7.0 (working volume 10 ml), using 1.0 mM hydroquinone

as redox mediator. The solutions were stirred at 300 rpm with a magnetic bar during

amperometric measurements. For analytical purposes, 1.0 mM H2O2 solutions in 50

mM sodium phosphate buffer, pH 7.0 were freshly prepared.

The optical measurements of the surface plasmon resonance (SPR) angles were

performed at a fixed wavelength of 670 nm using a Springle Autolab-SPR equipment

from Metrohm Autolab B.V. (The Netherlands). The mass variation during the formation

of the dendrimer/HRP layer-by-layer assemblies was followed by a Maxtek RQCM

603200-2 quartz crystal microbalance (QCM, Inficon, USA), oscillating at a nominal

frequency of 9.0 Hz. The morphology of gold modified surface was studied using atomic

force microscopy (AFM) with a SPM Nanoscope IIIa multimode microscope (Veeco

Instruments Inc., USA). 1H NMR characterizations were performed with a Bruker Avance

500 MHz spectrometer (Bruker BioSpin GmbH, Germany).

2. Synthesis of CD-modified ethylenediamine core PAMAM G-5 dendrimer (CD-PAMAM

G-5)

The mono-6-O-tosyl β-cyclodextrin derivative was synthesized as previously

described [22]. To prepare the CD branched dendrimer, 50 mg of ethylenediamine

core PAMAM G-5 dendrimer and 510 mg of mono-6-O-tosyl CD were dissolved in 100

mL of DMSO, previously deoxygenated by continuous bubbling of N2. The mixture was

stirred at room temperature under N2 atmosphere during four days, then

concentrated in vacuum to about 5 mL and further diluted with 150 mL of distilled

water. The solution was exhaustively dialyzed vs. distilled water using Amicon Ultra-15

centrifugal filter units with Ultracel-10 membranes (Millipore, USA), and finally

concentrated to a 4 mg/mL concentration. The CD-PAMAM G-5 dendrimer was

characterized by 1H NMR in D2O. Yield: 107 mg.


248

3. Synthesis of CD-modified cysteamine core PAMAM G-4 dendron (CD-PAMAM G-4)

Fifty milligrams of cystamine core PAMAM G-4 dendrimer and 510 mg of mono-

6-O-tosyl CD were dissolved in 100 mL of deoxygenated DMSO, and the mixture was

stirred at room temperature under N2 atmosphere during four days. The solution was

then concentrated in vacuum to about 5 mL, diluted with 150 mL of distilled water and

finally dialyzed vs. distilled water using Amicon Ultra-15 centrifugal filter units with

Ultracel-10 membranes (Millipore, USA). The resulting solution was concentrated to

about 20 mg/mL and treated with NaBH4 (10 mM final concentration) during 2 h under

magnetic stirring. The mixture was then dialyzed/concentrated with Amicon

centrifugal tubes as described above, yielding a CD-PAMAM G-4 solution of 4 mg/mL

final concentration. The CD-PAMAM G-4 dendron was characterized by 1H NMR in D2O.

Yield: 93 mg.

4. Preparation of the layer-by-layer assemblies on gold surface

First, the gold surfaces were exhaustively cleaned to ensure the chemisorption

of thiolated PAMAM dendron derivative. A disk gold electrode was polished with

alumina powder (0.3 µm), then rinsed with double distilled water and immersed in an

ultrasonic bath for 5 min. The electrode was further dipped in boiling 8 M KOH solution

for 1 h, washed with distilled water and immersed in piranha solution for 30 min. The

electrode was washed and electrochemically cleaned by cyclic voltammetry in 0.1 M

H2SO4 solution through twenty cycles between -0.2 V and 1.85 V at scan rate of 100

mV/s. For AFM studies, a gold foil (thickness 0.1 mm) was treated as described above

and further annealed on flame for 30 seconds. The gold surface in the QCM sensor

disks were cleaned by boiling for 5 min in a H2O/H2O2/NH3 solution (5/1/1, v/v), rinsed

thoroughly with water and dried with N2. SPR sensor disks were cleaned by sequential

shaking during 10 min in 100 mM NaOH and 1% Triton X-100 (v/v) solutions, rinsed

with water and dried with N2.

In a typical experiment, a clean gold surface was first dipped into a 0.5 mg/mL

aqueous solution of CD-PAMAM G-4 to covering the metal surface with a monolayer of


249

the CD-branched dendron derivative. After 2 h incubation, the electrode was washed

several times with distilled water and further covered with a 2 mg/mL solution of HRP-

ADA, prepared as previously described [17], in 50 mM sodium phosphate buffer, pH

7.0. The formation of the first enzyme layer was accomplished by incubating the

system during 2 h at 4ºC. The gold surface was then washed with cold buffer solution

and incubated for 2 h with a 0.5 mg/mL solution of CD-PAMAM G-5 dendrimer in the

same buffer. After washing, further enzyme and CD-PAMAM G-5 layers were prepared

by sequential treatment of the modified metal surface as described above. The

modified metal surfaces were kept at 4ºC in 50 mM sodium phosphate buffer, pH 7.0

until use.


1. Preparaction and characterization of the layer-by-layer modified electrode

Figure 1 displays schematically the strategy employed to construct the self-

assembled layer-by-layer architectures of PAMAM dendritic structures and HRP on

gold surfaces via host-guest interactions. To promote the formation of supramolecular

complexes between the different molecular scaffolds, HRP was chemically modified

with 1-adamantanecarboxylic acid using a water-soluble carbodiimide as coupling

agent [17]. As it was demonstrated previously, this modification yielded an enzyme

derivative that contained an average of 5 mol adamantane (ADA) residues per mol of

modified protein molecule and retained about 87% of the enzyme initial catalytic

activity.


250

Figure 1. Scheme displaying the steps involved in the preparation of the layer-by-layer

self-assembly of dendrimer and HRP on Au surface. D1: CD-PAMAM G-4 dendron layer,

D2 and D3: CD-PAMAM G-5 dendrimer layers, and HRP: enzyme layers.

Moreover, the primary amino groups at the surface of PAMAM dendrimers

were reacted with the βCD-mono-6-O-tosyl derivative in order to provide

hyperbranched molecular receptors for the multivalent association with the

adamantane-modified enzyme. In the case of the cystamine core PAMAM G-4

dendrimer, it was further treated with NaBH4 to reduce the disulfide bond at the

dendrimer core, yielding a thiol-containing dendron derivative able to form the first


251

dendritic layer on the gold surface through chemisorption. 1H NMR measurements

revealed that 9 and 45 CD units were attached to each molecule of PAMAM G-4

dendron and PAMAM G-5 dendrimer, respectively. After the formation of the first

dendron-based chemisorbed monolayer (D1), further enzyme (HRP1, HRP2 and HRP3)

and dendrimer (D2 and D3) supramolecular layers were assembled by sequential

incubation of the modified metal surface in HRP-ADA and CD-PAMAM G-5 solutions.

Figure 2. A) Cyclic voltammograms of native (a) and CD-PAMAM G-4 dendron-modified

Au electrode in 0.1 M sodium phosphate buffer, pH 7.0 solutions containing 1.0 mM

hydroquinone before (b) and after (c) addition of H2O2 up to 100 µM final

concentration. B) Cyclic voltammograms of Au electrodes modified with CD-PAMAM G-

4 dendron (c), D1/HRP1 (d), D2/HRP2 (e), D3/HRP3 (f) and D4/HRP4 (g) bilayers in the

presence of 100 µM H2O2.

The assembling process of the different dendrimer/enzyme layer-by-layer

architectures was characterized by cyclic voltammetry, SPR, QCM and AFM. Figure 2A

shows the cyclic voltammograms recorded at a bare gold electrode (curve a) and at

CD-PAMAM G-4 dendron-modified gold electrode (curve b) in 0.1 M sodium phosphate

buffer, pH 7.0, containing 1.0 mM hydroquinone which was further used as redox

mediator. As it can be observed, the characteristic voltammogram of hydroquinone

exhibited a slight decrease in both the cathodic and anodic peak current values after

dendron-coating, which can be most likely attributed to the reduction of the electrode

active surface area. The voltammetric pattern of the modified electrode was not

significantly changed in the presence of 100 µM H2O2 (curve c), suggesting that CD-


252

PAMAM G-4 molecules did not contribute to the catalytic transformation of H2O2 at

the Au surface. However, a noticeable catalytic H2O2 reduction current was observed

upon immobilization of the first HRP layer, as revealed by the increase in the cathodic

peak current peak (Figure 2B). Subsequent additions of enzyme layers produced an

increase in this cathodic peak current (curves e and f) which reached a maximum value

upon immobilization of three HRP layers. The presence of more enzyme layers caused

a significant reduction in the peak current, as can be observed for layer four in curve g,

suggesting that a large number of dendrimer/enzyme bilayers gave rise to a less stable

supramolecular architecture at the electrode surface or a more complex arrange in

which the enzymatic activity is not favored.

In order to confirm the supramolecular nature of the multilayer assembly, a

biosensor constructed with three enzyme layers was incubated at 4 ºC in 0.1 M sodium

phosphate buffer, pH 7.0, containing 10 mM 1-adamantane carboxylic acid and the

catalytic response of the bioelectrode towards 100 µM H2O2 was checked by cyclic

voltammetry, using again hydroquinone as mediator (Figure 3). It is well know that 1-

adamantane derivatives can form highly stable inclusion complexes with CDs [23],

therefore it was predictable that the assumed host-guest interactions stabilizing the

multilayer enzyme architecture on the electrode surface should be damaged by

incubation in the 1-adamantane carboxylic acid solution. Moreover, this adamantane-

derivative has also the advantage of not inhibiting the catalytic activity of HRP [17, 19].

Figure 3 shows as the cathodic peak current corresponding to the catalytic

reduction of H2O2, decreased progressively with the time of incubation in the 1-

adamantane carboxylic acid solution (curves a-d), reaching a minimum value after 2 h

incubation under these conditions. On the contrary, no significant change was

observed for a control biosensor prepared by physical immobilization of a single layer

of unmodified HRP on a CD-PAMAM G-4 dendron-coated gold electrode (data not

shown). These results demonstrate clearly that 1-adamantane residues were able to

disrupt the host-guest association between the HRP-ADA layers and the CD-PAMAM

dendritic layers on the Au surface, which supported fairly well the claimed

supramolecular nature of the multilayer assembly. However, when the voltammetric


253

responses were compared with that obtained at a control electrode which was coated

with CD-PAMAM G-4 dendron/HRP (curve e), it could be noticed that some catalytic

activity remained at the gold surface after exposure to 1-adamantane carboxylic acid

solution. This fact suggested that not only host-guest supramolecular interactions were

involved in the formation and stabilization of the dendrimer/protein assembly, but

also other types of interactions (electrostatic, hydrogen bonds, hydrophilic, Van der

Waals) should contribute to this layer-by-layer arrangement. Quantification of these

interactions on the overall load of the enzyme will be further discussed, taking into

account the results achieved by amperometric measurements.

Figure 3. Cyclic voltammograms recorded with Au electrodes modified with CD-PAMAM

G-4 dendron/HRP (E) and D3/HRP3 bilayers in 0.1 M sodium phosphate buffer, pH 7.0

solutions containing 1.0 mM hydroquinone and 100 µM H2O2 before (A) and after 30

(B), 60 (C) and 120 (D) min incubation in 10 mM 1-adamantane carboxylic acid

solution.


254

Figure 4. SPR sensogram recorded upon the layer-by-layer self-assembling of CD-

modified PAMAM dendritic scaffolds and HRP-ADA on Au surfaces.

The changes occurred on the gold surface after sequential exposure to the

different dendritic and enzyme derivatives were evaluated by SPR and QCM, which

allowed real time monitoring of the formation of the different layers on the Au surface

as well as the quantification of the molecules amount interacting on the metal surface

[24]. Figure 4 shows as incubation of cysteamine core CD-PAMAM G-4 dendron

produced a noticeable increase in the SPR signal which remained high after washing,

suggesting that the thiol-containing dendron derivative was successfully chemisorbed

on the Au surface. Further incubation with the adamantane-modified HRP derivative

also caused a significant increase in the SPR signal, thus demonstrating the interaction

between the enzyme molecules and the dendron-capped surface. Sequential

incubation with CD-PAMAM G-5 dendrimer and HRP-ADA also yielded successive

plasmon signal rising, indicating the formation of the layer-by-layer supramolecular

assembly on the Au surface.

A similar behavior was observed in the QCM experiments. The data obtained in

these measurements were employed to estimate the amount of macromolecules

deposited in each layer. This estimation was made by using the Sauerbrey equation


255

but assuming the incorporation of an average of 75% and 45% (w/w) of water in the

protein and PAMAM dendrimer layers, respectively [25, 26]. In addition, the amount of

protein in the HRP layers was calculated from the SPR data considering that the

adsorption of 1 ng/mm2 of protein on a thick Au layer causes a change in SPR angle of

122 millidegree (mº) at 25 ºC [27].

Figure 5. Calculated molar content of enzyme and dendrimers in the layer-by-layer

assembly from QCM (blue) and SPR (red) measurements. Each value corresponds to the

molar content per layer.

As illustrated in Figure 5, larger amounts of both dendrimer and enzyme were

immobilized with the first layers, which suggested a higher stability for these initial

assemblies. This fact can be attributed to the high coverage of the metal surface with

the thiol-containing CD-PAMAM G-4 dendron derivative through strong chemisorption

linkages. This first layer served as a stable support for the multiple immobilization of

the first enzyme monolayer via host-guest associations. A stable multivalent

supramolecular association of the second dendrimer layer could be also expected due

to the high oligosaccharide content in CD-PAMAM G-5 dendrimer. However, the

subsequent increase in the number of layers seemed to reduce the capability of the

assembly to immobilize macromolecules, as revealed by the lower enzyme and


256

dendrimer contents measured in both QCM and SPR experiments. This effect should

be mainly attributed to the low capability of the multivalent host-guest interactions to

support high mass assemblies, reaching a limit after the immobilization of the third

enzyme layer, as it was previously observed in CV experiments.

Figure 6. Three dimensional AFM analysis of the Au surface before (A) and after

modification with the D1/HRP1 (B), D2/HRP2 (C) and D3/HRP3 (D) bilayers.

The formation of the layer-by-layer arrangement on the Au surface was also

confirmed by examining the topology of the modified gold surface by AFM. The

corresponding images are displayed in Figure 6. The raw metal surface showed an

average height of 10.06 nm with a maximum at 19.57 nm. The surface area and the

average roughness of this metal surface were estimated as 253.7x103 nm2 and 2.48

nm, respectively. Coverage with the first CD-PAMAM G-4 dendron/HRP-ADA layer

yielded a modified surface pattern with average and maximum height values of 13.53

nm and 35.72 nm, respectively. The average roughness and surface area of Au

increased to 3.53 nm and 259.9x103 nm2, respectively, after coverage with the first CD-

PAMAM G-4 dendron/HRP-ADA layer.

More irregular surface patterns were observed after the assembly of the

second and third dendrimer/enzyme bilayers, probably due to the higher molecular


257

radius of the PAMAM G-5 dendrimer derivative. So, the average height of the

protuberances in the nanostructured D2/HPR2 and D3/HRP3 bilayers on the gold

surface was estimated as 18.48 nm and 27.2 nm, respectively. These D2/HPR2 and

D3/HRP3 bilayers-modified metal surfaces also exhibited increased average roughness

(5.82 nm and 8.34 nm), surface area (266.8x103 nm2 and 272.5x103 nm2) and maximum

height for the protuberant nanostructures (48.94 nm and 61.96 nm), respectively.

3.2. Electroanalytical performance for H2O2

In order to evaluate the potential applicability of Au electrodes modified with

the dendrimer/enzyme layer-by-layer assemblies for biosensing, their amperometric

response toward H2O2 was evaluated. Optimization studies using 1.0 mM

hydroquinone as redox mediator showed that the HRP electrodes exhibited bell-

shaped current responses vs the applied potential and the pH of the working solution,

with maximum values at E = -100 mV and pH 7.0, respectively (results not shown).

Under these conditions, the dynamic amperometric response of the bioelectrodes

upon successive additions of 1.0 mM H2O2 is illustrated in Figure 7. As a control, an Au

electrode coated with CD-PAMAM G-4 dendron on which an equivalent activity of

native HRP (unmodified with adamantane) was immobilized under similar conditions

was also checked.

All the biosensors showed fast catalytic response with 95% of the steady-state

current being reached in about 8 s. Interestingly, the amperometric response of the

electrodes with layer-by-layer architecture increased with the number of

dendrimer/enzyme bilayers, most likely due to the higher enzyme loading.

Bioelectrodes provided with high number of dendrimer/enzyme bilayers can also

detect H2O2 at very low concentration of, as is illustrated in the Figure 7 (inset).

However, as expected, the catalytic activity of the control bioelectrode was

significantly lower than that of the bioelectrode prepared with one HRP-ADA

monolayer, confirming a higher amount of the immobilized HRP derivative on the CD-

PAMAM G-4 dendron-modified electrode surface via host-guest interactions.


258

Figure 7. Amperometric responses recorded with CD-PAMAM G-4 dendron/HRP (A),

D1/HRP1 (B), D2/HRP2 (C) and D3/HRP3 (D) modified Au electrodes upon successive

additions of 1.0 mM H2O2. Eapp.= - 100 mV. Inset: Electroanalytical behavior of the

D3/HRP3 modified Au electrode toward low H2O2 concentration.

As it was suggested above, the formation and stabilization of the

dendrimer/protein assembly should be caused by the co-operative contribution of

several interactions, mainly based on host-guest supramolecular associations but also

including other types of interactions such as electrostatic, hydrogen bonds,

hydrophilic, Van der Waals forces, etc. Assuming that the catalytic activity of native

and ADA-modified HRP was similarly affected after immobilization on the dendron

monolayer, we can estimate the contribution of these other non-supramolecular

interactions on the overall immobilization process (at least for the first

dendrimer/enzyme bilayer) by comparing the slope of the calibration curves

corresponding to the control and ADA-HRP enzyme forms.


259

Figure 8. Calibration curves for H2O2 constructed with biosensors prepared with

D1/HRP1 (), D2/HRP2 (), D3/HRP3 () and D1/native HRP (×) modified Au electrodes.

Figure 8 displays the corresponding calibration graphs constructed for H2O2,

showing fairly well how the ranges of linearity changed with the increasing number of

dendrimer/enzyme bilayers. In fact, the biosensors having one to three

dendrimer/HRP bilayers exhibited linear ranges of response between 1.0 - 114 µM, 0.5

- 159 µM and 0.5 - 186 µM, respectively, with the following corresponding equations:

D1/HRP1: I / A = 6.8·10-6[H2O2] / M + 1.6·10-8 (R2 = 0.996)

D2/HRP2: I / A = 1.21·10-5[H2O2] / M + 3.2·10-8 (R2 = 0.998)

D3/HRP3 I / A = 1.89·10-5[H2O2] / M + 6.3·10-8 (R2 = 0.999)

It was also noticed that the control electrode, based on native HRP immobilized

on the CD-dendron monolayer, also exhibited linear response but for a shorter range

of H2O2 concentrations (1.0 - 82 µM). The behavior of this electrode can be described

by the following equation:

Control: I / A = 3.16·10-6[H2O2] / M + 1.1·10-8 (R2 = 0.994)


260

The comparison of this result with that obtained with the electrode coated with

the D1/HRP1 bilayer allow concluding that the contribution of the host-guest

supramolecular associations represent about 54% of the overall interactions involved

in the immobilization process, assuming that modification of HRP with adamantane did

not affect the other contributing interaction forces. This result agree with the

described above for cyclic voltammetric studies.

The sensitivity of the biosensors with supramolecular designs increased with

the number of dendrimer/HRP bilayers, showing values of 216, 385 and 602 µA/M cm2

for D1/HRP1, D2/HRP2 and D3/HRP3, respectively, considering the geometric area of

the raw electrode. A similar improvement was found regarding the detection limits,

270 nM (first bilayer), 200 nM (second bilayer) and 160 nM (third bilayer), which were

calculated according to the 3Sb/m criterion, where m is the slope value of the

calibration plot and Sb was estimated as the standard deviation (n = 10) of the signals

corresponding to the lower H2O2 concentration detected for by each biosensor. It

should be highlighted that such detection limits at nanomolar level rank among the

lower values reported for HRP-based biosensors [28]. In addition, the detection limit

achieved with the biosensor constructed with three dendrimer/enzyme bilayers was

lower than those reported for other HRP-modified electrodes with a layer-by-layer

design, based on the assembly of colloidal gold nanoparticles, cysteine and HRP on

Nafion modified electrode surface by electrostatic adsorption (500 nM) [29],

supramolecular self-assembly of HRP-ADA and HRP-modified with CD-branched

polysaccharides on CD-coated Au electrodes (2.0 µM) [19], HRP immobilized on carbon

electrodes coated with a layer-by-layer self-assembly of Au nanoparticles-based films

(4.9 µM) [30] and host-guest association of HRP-ADA and CD-capped Au nanoparticles

on Au surface (3.0 µM) [20].

The value of the apparent Michaelis-Menten constant for the enzyme

electrodes, KM, calculated according to the Eadie-Hofstee method, slightly changed

from 104 mM to 152 mM and 143 mM when the number of dendrimer/HRP bilayers

was increased. Moreover, IMAX increased with the number of dendrimer/enzyme

bilayers, showing values of 1.04 µA, 2.58 µA and 3.97 µA, respectively.


261

Aspects concerning reproducibility of the measurements and stability were

evaluated for the biosensor constructed with three dendrimer/enzyme bilayers. So,

ten successive measurements for 10 µM H2O2 yielded a relative standard deviation

(RSD) value of 4.1%, while the electrode-to-electrode reproducibility was checked from

the responses obtained with ten different biosensors prepared in the same manner

providing a RSD value of 6.7%.

The stability of the biosensor was tested by storing it at 4ºC under dry and wet

(100 mM sodium phosphate buffer, pH 7.0) conditions, and periodical evaluation of

the measured current for 10 µM H2O2. Although the stability of the biosensor under

dry storage was found to be poor, losing all electroanalytical response after 5 days of

storage, the enzyme electrode exhibited good long-term stability under wet

conditions, retaining 91% and 63% of its initial activity after 15 and 30 days of storage

at 4ºC. This stability behaviour could be associated with the multipoint supramolecular

attachment of the enzyme to the CD-modified dendrimers into the layer-by-layer

design, which seems to be relatively stable in aqueous solution but not when dried.

Previous AFM studies demonstrated than the volume and deformability of PAMAM

dendrimers are different in aqueous and dry conditions [31]. Therefore, it could be

then expected that the stability of the dendrimer/HRP layer-by-layer assembly would

be affected by drying and swelling processes during storage and measurement with

the consequent loss of enzyme molecules from the Au surface.

CONCLUSIONS

In summary, this is the first report dealing with the preparation of

dendrimer/enzyme layer-by-layer assemblies based on supramolecular interactions.

The characteristics of these arrangements on Au surfaces were established, and their

potential use for biosensor construction demonstrated. Experiments are now in

progress to generalize this type of supramolecular-based self-assemblies for the

construction of multienzymatic biosensors.


262

ACKNOWLEDGEMENTS



Innovación CTQ2011-24355, CTQ2009-12650, CTQ2009-09351 and Comunidad de

Madrid S2009/PPQ-1642, Programme AVANSENS is gratefully acknowledged.

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7.9

ACS Applied Materials & Interfaces 8 (12) (2016) 7657–7665

Neoglycoenzyme-gated mesoporous

silica nanoparticles. towards the design of nano-devices for pulsatile

programmed sequential delivery

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NEOGLYCOENZYME-GATED MESOPOROUS SILICA

NANOPARTICLES. TOWARDS THE DESIGN OF NANO-DEVICES FOR

PULSATILE PROGRAMMED SEQUENTIAL DELIVERY

Paula Díez,1 Alfredo Sánchez,1 Cristina de la Torre,2,3 María Gamella,1 Paloma

Martínez-Ruíz,4 Elena Aznar,2,3 Ramón Martínez-Máñez,2,3,* José M. Pingarrón,1,5,*

Reynaldo Villalonga1,5,*

1Department of Analytical Chemistry, Faculty of Chemistry, Complutense

University of Madrid, 28040-Madrid, Spain. 2Instituto de Reconocimiento Molecular y

Desarrollo Tecnológico (IDM), Centro Mixto Universidad Politécnica de Valencia-

Universidad de Valencia, Spain. 3Departamento de Química y CIBER de Bioingeniería,

Biomateriales y Nanomedicina (CIBER-BBN), Universidad Politécnica de Valencia,

Camino de Vera s/n, 46022, Valencia, Spain. 4Department of Organic Chemistry I,

Faculty of Chemistry, Complutense University of Madrid, Madrid, Spain. 5IMDEA

Nanoscience, Cantoblanco Universitary City, 28049-Madrid, Spain.


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ABSTRACT

We report herein the design of a stimulus-programmed pulsatile delivery

system for sequential cargo release based on the use of a lactose-modified esterase as

a capping agent in phenylboronic acid functionalized mesoporous silica nanoparticles.

The dual release mechanism was based on the distinct stability of the cyclic boronic

acid esters formed with lactose residues and the long naturally-occurring glycosylation

chains in the modified neoglycoenzyme. Cargo delivery in succession was achieved

using glucose and ethyl butyrate as triggers.

KEYWORDS: Enzymes, mesoporous silica, delivery, nanoparticles, esterase,

neoglycoenzyme, glucose.

INTRODUCTION

Controlled delivery technology is one of the most rapidly advancing areas in

material science, with the major driving force stemming from the pharmaceutical

sector. In fact, advanced release systems offer numerous pharmacological and

pharmacokinetics advantages as compared to conventional dosage forms such as

enhanced drug efficacy, reduced toxicity, and improved patient compliance and

convenience.1 In particular, the design of advanced carriers that enable time- or

stimulus-programmed drug release is of much importance and is relevant for the

treatment of many diseases.

This field has been traditionally dominated by the use of polymers as carriers.2-4

Recently, however, attention has been paid to mesoporous silica nanoparticles (MSN)

as nanosized containers for controlled delivery due to their unique properties such as

large specific volume, large loading capacity, low toxicity and easy preparation in

different forms, as well as varying morphology, size and pore diameter.5-9 MSN can be

easily functionalized to allow the rational building of stimuli-responsive molecular or

supramolecular ensembles on their external surface to develop gated nanocarriers that


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show “zero delivery”.10,11 These functional nanodevices can also act as smart delivery

nanomachines by releasing their cargo in response to target chemical12-15 or physical

stimuli,16,17 or by exposure to specific biochemical macromolecules.18-20

Usually these nano-devices exhibit a single delivery upon the application of the

triggering stimulus. However, there are certain conditions in which this release pattern

is not suitable as well as many applications where not releasing all the drug during the

initial dosage phase is a requirement. In fact there are a number of diseases, such as

asthma, cancer, duodenal ulcer, arthritis, diabetes, neurological disorders, acute

myocardial infarction, etc., and vaccination protocols21,22 where pulsatile drug delivery

systems are preferred to conventional drug administration approaches including

sustained-release drug delivery systems. In this sense, programmable pulsatile release

has the advantages to avoid drug tolerance and maintain drug concentration at

therapeutic levels albeit circadian rhythms, allowing the design of chronotherapeutic

protocols for some common diseases exhibiting circadian variation.22 Nevertheless,

despite the design of a number of sophisticated nano-delivery supports, very few

nano-devices for sequential and controlled pulsatile cargo delivery are yet

available.23,24

Scheme 1. Performance of dual stimuli-responsive nanodevice S3 for the programmed

and sequential delivery of the [Ru(bpy)3]Cl2 complex using glucose and ethyl butyrate as

triggers.

On the other hand, in most of enzyme-responsive MSN systems previously

reported, enzymes act as triggers which hydrolyze specific molecular sequences


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anchored on the gated mesoporous support. As an alternative, we have recently

reported that enzymes can act as caps in gated materials in which the uncapping

process is triggered by the product obtained by the enzyme activity on target guests.15

Neoglycoenzymes are artificially glycosylated enzymes in which the new

appended carbohydrates moieties confer novel and improved characteristics to the

enzyme, such as stability, catalytic activity, and chemical and biochemical recognition

properties.25,26 Based on our experience in the design of neoglycoenzymes26 we

envisioned that the modification of a glycoenzyme with short-chain carbohydrate

residues can lead to a neoglycoconjugate able to be used as a capping element in

phenylboronic acid-grafted MSN by forming cyclic boronic acid esters through either

naturally-occurring long glycosylation chains or chemically attached short disaccharide

chains (vide infra). In this context, we speculated that considering the different amount

and geometry of the long and short glycosylation chains, as well as their different

affinities and stabilities with the boronic acid groups, it might be possible to design

pulsatile drug delivery systems capable of delivering the cargo sequentially under the

effect of different triggering conditions.

As a proof of concept of sequential delivery with MSN using neoglycoenzymes,

we report herein the preparation of solid S3 (vide infra) consisting of an MCM-41-type

MSN containing phenylboronic acid residues, capped with lactose-modified pig liver

esterase (see Scheme 1). This engineered nanoparticle was successfully evaluated as

novel sequential and pulsatile drug delivery system for in vitro and ex vivo experiments.


To assemble the integrated delivery nanomachine, mesoporous silica

nanoparticles (S0) were first prepared as previously described.27 These nanoparticles

were then loaded with tris(2,2´-bipyridyl)ruthenium(II) chloride ([Ru(bpy)3]Cl2) as

model drug for cargo delivery monitoring5f, and further treated with (3-

glycidyloxypropyl) trimethoxysilane to provide the surface of the nanoparticles with


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highly reactive epoxy groups. The resulting solid was then treated with 3-amino

phenylboronic acid to yield solid S1.

Subsequently, S1 nanoparticles were treated with native esterase to prepare

the enzyme-capped control solid S2 via the formation of boronic acid cyclic esters

between the naturally-occurring long glycosylation chains in the enzyme and the

phenylboronic acid groups on the surface of the MSN.

In order to construct the pulsatile nano-device, a neoglycoenzyme was

prepared by covalent anchoring of lactose to the free amino groups of lysine residues

in esterase through a reductive alkylation reaction with NaBH3CN.28 The resulting

glycoconjugate retained 92% of initial specific activity after the incorporation of 61 mol

lactose per mol of trimeric enzyme, as determined by phenol-sulfuric acid method (see

Experimental Section), which corresponded to a modification of ca. 66% of the lysine

residues in the protein.29 A low degree of modification was estimated by MALDI-TOF

analysis, which revealed a molecular weight of 60858 Da and 64227 Da for the native

and lactose-modified enzyme, respectively. This fact suggests an average of 12 mol

lactose attached to each mol of monomeric enzyme in the neoglycoconjugate. This

neoglycoenzyme was then employed as a capping element for S1 yielding S3, following

a similar procedure to that used to prepare the control solid S2. The amount of native

and lactose-modified esterase immobilized on S2 and S3 nanoparticles was estimated

at 10.5 U/mg and 4.3 U/mg, respectively.

All the MSN were fully characterized. The TEM analysis revealed that the

different prepared nanoparticles showed the typical porous pattern of the MCM-41

mesoporous matrix with the average size for the nanoparticles of 97 ± 15 nm (Fig. S1).

Moreover TEM images of S2 and S3 displayed a diffuse thin layer that wrapped the

nanoparticles, suggesting their coverage with an amorphous organic material, which

could be ascribed to the esterase enzymes immobilized on the nanoparticle surfaces.

All the samples also showed the characteristic X-ray diffraction pattern of the MCM-41-

type materials, with a well-defined peak at ca. 2.59º attributed to the (100) Bragg

reflection of MCM-41-type materials. The position of this peak was not significantly

affected by dye loading and the different chemical modifications processes carried out


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in their preparation (Fig. S2). Functionalization of MSN with phenylboronic acid

moieties and further capping with the esterase derivatives were assessed by FT-IR (Fig.

S3). The assembly of the nanodevices was also confirmed by solid state 13C NMR (Fig.

S4). The thermal analysis revealed that the content of the anchored boronic acid-based

ligand and encapsulated [Ru(bpy)3]Cl2 dye in S1 amounted to 34 mg/g S1 and 206 mg/g

S1, respectively (Fig. S5). The amount of esterase and lactose-modified esterase in S2

and S3 was estimated to be 17.3% and 32.8% by weight, respectively, according to the

TG studies. This difference could be explained by the higher molecular weight showed

by the lactose-modified enzyme, as well as the potential higher capacity of this

neoglycoconjugate to form multipoint boronic acid cyclic ester linkages at the surface

of the nanoparticles. The influence of the different modification steps on the total

specific surface area and pore size of the prepared MSN was also determined from the

corresponding N2 adsorption-desorption isotherms (Fig. S6 and Table S1). S2 and S3

showed similar specific surface area and total pore volume values, suggesting that the

different enzyme derivatives showed high capping efficiency for the mesoporous

nanoparticles.

The dynamic light scattering and zeta potential experiments further confirmed

the successful surface functionalization of MSN. DLS data show that the starting MCM-

41 material exhibited a hydrodynamic diameter of 295 nm as a result of partial

aggregation in H2O. This diameter was even larger in PBS (320 nm) and cell culture

medium (384 nm), also indication aggregation of the nanoparticles in these solvents.

The functionalization of the MCM-41 nanoparticles with the glycoenzymes to give S2

and S3 improved the colloidal stability of the nanoparticles and hydrodynamic

diameters of 211 and 196 were found for S2 and S3 in H2O, respectively (see

Supporting Information). This reduction in the hydrodynamic diameter could be mainly

ascribed to a low degree of aggregation in the enzyme-modified nanoparticles. Zeta

potential measurements showed that the starting MCM-41 nanoparticles had a

negative potential in water (-26 mV) which was reduced for the final capped

nanoparticles S2 and S3 to -7 mV and -5 mV due to the functionalization of the surface

and capping with the glycoenzymes. Similar results were found in PBS and cell culture

medium (See Fig. S7 and Table S2 in Supporting Information).


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Functionalization of the MCM-41 nanoparticles with the glycoenzymes was

assessed by TEM after staining the nanomaterials with uranyl acetate (Fig. S8 in

Supporting Information). In comparison with S1, S2 and S3 showed a well-defined dark

layer around the nanoparticles, demonstrating high coverage of these nanomaterials

with the organic glycosylated enzyme molecules.

Figure 1. A) Release efficiency for [Ru(bpy)3]Cl2 from S3 (a) and S2 (b) after a 90-minute

incubation with different trigger substances at a 200 µM concentration. B) Dye release

efficiency from S3 (a) and S2 (b) after 90-minute incubation in the presence of glucose

at different concentrations.

Nanodevices S2 and S3 were tested for the on-command controlled delivery of

the cargo in aqueous solutions in the presence of some potential triggers (vide infra).

In a typical release assay, 10 mg of the enzyme-capped nanoparticle were suspended in

4 mL of 20 mM Na2SO4 solution at pH 7.5, and were shaken over time at 25ºC. Then

the corresponding triggering substances were added and the mixture was incubated

for 1.5 h. An aliquot was taken, centrifuged to remove the nanoparticles, and the

absorbance of the released [Ru(bpy)3]Cl2 complex was measured at 454 nm. As triggers

for cargo delivery, two different types of substances were tested; ethyl butyrate and

1,2- or 1,3-diols (glucose and ethylene glycol). The chemistry involved in the trigger

reactions is illustrated in Fig. S9.

Addition of ethyl butyrate caused the cargo to be completely released from

both nanomaterials S2 and S3 (see Figure 1A). This behaviour is related with the rapid

disruption of the boronic acid cyclic esters under acidic conditions due to the esterase-


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catalysed hydrolysis of ethyl butyrate to ethanol and butyric acid (pKa = 4.82) resulting

in the leakage of the immobilized enzyme from the nanosized support and cargo

delivery.

Figure 2. Kinetics of the dye release from S3 (A) and S2 (B) in 20 mM Na2SO4, pH 7.5,

without (a) and with addition of ethyl butyrate (b) and glucose + ethyl butyrate (c) at a

200 µM final concentration. Triggers were added at the times indicated in the graphics.

In contrast, and interestingly, the encapsulated [Ru(bpy)3]Cl2 complex was only

partially released from S3 under the trigger effect of glucose, whereas no release was

observed for S2. Delivery from S3 is related with the competition of glucose for the

boronic acid residues anchored to the neoglycoenzyme on the nanoparticle surface,

which results in a partial cargo release. The extent of such a delivery process in S3

increased with the sugar concentration in the incubation media, a plateau value being

reached for concentrations of glucose above 200 µM (Figure 1B). Similar behaviour was

observed by incubation at high glucose concentration up to 5 mM. On the contrary, no

significant dye release was observed for similar experiments with nanoparticles S2,

even at relatively high glucose concentrations up to 5 mM. Studies with ethylene

glycol, which is able to form highly stable 5-member rings with boronic acid residues,

showed an almost complete cargo release from S3, but no noticeable effect was

produced with S2 (see Figure 1A).

Based on these results, S3 was tested for the two-step controlled cargo delivery

by triggering the uncapping process through the sequential incubation with glucose


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and ethyl butyrate. In parallel, S2 was also tested under the same experimental

conditions (see Figure 2). In the absence of glucose or ethyl butyrate, both nanodevices

are tightly capped and showed a negligible release of [Ru(bpy)3]Cl2. Conversely, the

presence of glucose resulted in the opening of some pores in S3 producing a

subsequent partial cargo delivery. Further incubation of S3 with ethyl butyrate, which is

transformed by the immobilized enzyme glycoconjugate in ethanol and butyric acid

with a consequent drop in pH, caused the acid-mediated hydrolysis of the boronic acid

cyclic esters and induced cargo delivery (see Figure 2A). On the contrary, S2 delivered

the cargo only by incubation with the enzyme substrate ethyl butyrate (see Figure 2B).

It should be noted that the pH of the incubating media remained unchanged after the

S2 and S3 treatment with glucose, but changed from 7.5 to about 5.2 after incubation

with ethyl butyrate.

It should be highlighted that similar release patterns were observed by

performing the experiments in PBS, but slightly lower degree of cargo delivery was

achieved by using ethyl butyrate as trigger. This fact suggests the esterase-mediated

release of Ru(bpy)32+ from the nanoparticles was mainly caused by a local decrease of

pH at the microenvironment of the modified colloids.

In order to confirm that the opening mechanism with ethyl butyrate was due to

the enzyme-mediated decrease of pH after the substrate hydrolysis, suspensions of S2

and S3 at pH 7.5 were heated to 100ºC for 10 min to inactivate the enzymes, and were

further incubated with ethyl butyrate. In this case, no appreciable cargo delivery was

observed from the nanodevices after 24 h of incubation.


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Scheme 2. Proposed mechanisms for the stimuli-responsive controlled delivery from

nanocarriers S2 (A) and S3 (B).

To gain inside the mechanism of the sequential delivery process, the activity of

the enzyme immobilized on the MSN was determined before and after the sequential

incubation with the triggers. The enzymatic activity initially immobilized on S3 capped

with lactose-modified esterase was estimated to be 4.3 U/mg solid, which lowered to

2.9 U/mg solid after 1 h of incubation with glucose, suggesting that about 32.6% of the

neoglycoenzyme leaked from the solid through a monosaccharide-mediated

competitive replacement. Moreover, no appreciable enzymatic activity was detected

on the resulting solid after 1 h of S3 incubation with ethyl butyrate. In contrast, S2

retained full esterase activity (10.5 U/mg solid S2) after performing a similar incubation

with glucose.


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Treatment of S2 with ethyl butyrate also caused the total reduction of

immobilized enzyme activity, demonstrating that the enzyme was completely leaked

from the nanoparticle surface.

Taking into account these results, a mechanism for the two-step pulsatile

programmed sequential delivery from S3 triggered by glucose and ethyl butyrate is

proposed in Scheme 2. The obtained results indicate that native esterase is strongly

attached to the nanoparticle surface in S2, probably through the formation of

multipoint boronic acid cyclic ester linkages between the saccharide units at the

naturally-occurring glycosylation chains of the enzyme and the phenylboronic acid

groups grafted on the nanoparticle surface.

Regarding S3, the lactose-modified esterase is most likely linked to the boronic

acid residues through two different ways: i) by the attachment of the naturally-

occurring glycosylation chains of the enzyme, yielding highly stable linkages that are

not displaced by incubation with glucose or ethylene glycol; and ii) via the lactose

residues in the neoglycoenzyme, which yield linkages that can be disrupted in the

presence of glucose or ethylene glycol. Moreover, both kinds of linkages are prone to

being hydrolyzed in acidic media, which is provoked by the enzyme-mediated

hydrolysis of ethyl butyrate.

In addition, the enzyme-controlled capped MSN were tested for in-cell

controlled delivery applications. Therefore, after the in vitro characterization of the

solid S2 and S3 (vide ante), similar nanoparticles were used for ex vivo assays by

loading nanoparticles as S2 and S3 with the cytotoxic doxorubicin (Doxo) (solid S4 and

S5, respectively).

Solids S4 and S5 were evaluated in HeLa cells under the premise that they

would be internalised by cells and would stay nearly closed until glucose or ethyl

butyrate were added. In this sense, the capacity of HeLa cells to uptake 100 nm silica-

modified nanoparticles has been largely documented in the literature.7,30-33

According to the in vitro S2 and S3 behaviour (see Figure 2B and 2A

respectively), the addition of D-glucose in S5 in ex vivo assays would be expected to


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induce partial intracellular Doxo delivery, whereas the additional presence of ethyl

butyrate would result in further Doxo release. On the other hand, the presence of D-

glucose in internalised S4 nanoparticles should have no effect while the addition of

ethyl butyrate would induce Doxo delivery.

Figure 3. Internalization and release of cargo in HeLa cells. Culture were incubated with

S4 (3A) or S5 (3B) in presence/absence of different input I1 (D-glucose) or input I2 (ethyl

butyrate) and examined for Doxorubicin staining (Doxo) by confocal microscopy.

Representative images at 24 h form phase contrast (PhC), Doxorubicin (Doxo), Hoescht

(Hoe) and combined (Merged) are shown. Quantification of cell viability and cell death

was performed by flow cytometry by means of 7-AAD and Ann V staining. The

percentage of dead cells (black), cells undergoing cell death (gray) and healthy cells

(white) are shown for 50 μg/ml concentration of S4 (3C) and S5 (3D) in HeLa cells under

different conditions at 24 h.

In a typical experiment, HeLa cells were incubated for 30 minutes with a

suspension of 75 µg/mL of S4 or S5 in PBS supplemented with 10% fetal bovine serum.

Subsequently, cells were washed to remove un-internalised nanoparticles and further


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incubated alone or in the presence of D-glucose (input I1), ethyl butyrate (input I2) or a

mixture of both (see Experimental Section). Sequential addition of D-glucose and ethyl

butyrate as input signals was also evaluated.

Doxo delivery details obtained by confocal microscopy and cytometry are

provided in the Supporting Information (see Figures S10 and S11), whereas cell viability

was determined by flow cytometry studies (see Figure 3) and by WST-1 assays under

different conditions.

Figure 3 also shows representative phase contrast images of Doxo, Hoescht, and

combined for HeLa cells first loaded with S4 or S5 and then untreated (-/-) or treated

with both D-glucose and ethyl butyrate (+/+). These nanomaterials were mainly

internalized into the HeLa cells, as observed from Figure 3. However, some few

nanoparticles could remain outside the cells as revealed for S5 in Figure 3B (see +/+).

In flow cytometry studies, quantification of Doxo-associated cell death was

performed by using 7-aminoactinomycin (7-AAD) and Annexin V-FITC (Ann V) markers,

which stain dead cells and cells undergoing cell death, respectively. As it can be

deduced from the viability studies, the simple internalization of S4 in HeLa cells (see -/-

) or the treatment with D-glucose (see +/-) did not induce significant cell death,

whereas ca. 60% of cells were dead or underwent cell death after addition of ethyl

butyrate (see Figure 3C). It was also found that S5 nanoparticles were not significantly

toxic for HeLa cells, whereas in this case both the presence of D-glucose (see +/-), ethyl

butyrate (see -/+) and specially a mixture of D-glucose and ethyl butyrate (see +/+)

induced an increment in dead cells and cells undergoing cell death (see Figure 3D). In

particular nearly 50% of the cells were dead or undergoing cell death when treated

with both D-glucose and ethyl butyrate simultaneously (see also Figures S12 and S13).

Although significantly very low, cell death caused by S4 and S5 nanoparticles in

the absence of the trigger compounds could be ascribed to unspecific release of Doxo

inside the cells. This fact, which was revealed by some few leaking signals in Figure 3

(see -/-), could be caused by local acidic conditions at the microenvironment of the

nanoparticles into the HeLa cells.


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Cell viability observations using flow cytometry were in agreement with cell

viability studies using WST-1 assays. Moreover cell viability studies were also in

agreement with Doxo fluorescence observed in cells by confocal microscopy and

determined with flow cytometry (Figures S10 and S11). Analogous results were

achieved by using sequential addition of D-glucose and ethyl butyrate to the incubation

medium, showing similar release patterns than those observed in experiments

involving the incubation with D-glucose and the mixture of both trigger compounds

(data not shown).

In addition, the intracellular distribution of S4 or S5 was evaluated by confocal

laser scanning microscopy upon the addition of ethyl butyrate. A red Lysotracker probe

was used to stain acidic organelles in HeLa cells. It was found that S4 and S5 were

predominantly localized in Lysotracker-labelled organelles after 1.5 h of incubation.

Moreover after 12 h a homogeneous distribution of Doxo in the cytosol and partly in

the nucleus was observed (see Figure S14 and S15). Conversely, when intracellular

distribution of S4 or S5 was evaluated in the absence of ethyl butyrate, much less Doxo

was released after 12 h incubation. On the other hand, Doxo fluorescence was co-

localized mostly with Lysotracker dye, strongly suggesting that the nanoparticles were

internalized into the nanoparticles and remained mainly in the lysosomes without

delivering the entrapped cargo (see Figure S14 and S15).

To further study intracellular Doxo release in S4 and S5 for different times (2, 5

or 30 h), confocal images of HeLa cells were taken in the presence or absence of ethyl

butyrate. A clear Doxo fluorescence was observed for both S4 and S5 in the nucleus

after 30 h when ethyl butyrate was used as input, whereas this effect was clearly not

observed in its absence (see Figure S16).

It should be highlighted that the design of novel multi-stimuli responsive

nanodevices receives considerable attention due to the advantages of such systems,

such as extraordinary control over drug delivery and release and the possibility to

program the release sequence, leading to superior therapeutic efficacy.34 In this sense,

a great variety of smart nanodevices able to respond to combination of multiple signals

(pH, temperature, magnetic field, enzymes, light, guest molecules, etc.) have been


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reported.7,30,34-37 In our model, the advantages associated to the use of a

neoglycoenzyme molecule as sensing, signal transforming and capping agent allow

designing of advanced and tailor-made nanoparticulated systems for the programmed

and pulsatile release of therapeutic, anti-microbial, and plant protecting and growth

enhancing compounds.

Despite our results, there are some aspects that could be considered in the

future to improve these nanoparticles-based programmed pulsatile delivery systems. In

this sense, D-glucose was here employed as model trigger but the presence of this

monosaccharide in blood should lead unspecific release of drugs in vivo. In this proof-

of-concept, no significant delivery was observed into the HeLa cells in the absence of

D-glucose, probably due to its low intracellular concentration caused by the fast

metabolic consumption of this sugar by cancer cells.38,39 Although ethyl butyrate is not

common in human fluids and tissues, local acidic microenvironments could also

provoke unspecific drug release in vivo. In addition, our model nanomachines could be

further improved by providing an affinity mechanism for target drug delivery to tumors

in vivo.

METHODS

Preparation of MSN (S0).27 Cetyltrimethylammonium bromide (3.0 g) was

dissolved in 1.44 L of water under sonication. NaOH solution (2.0 mol/L, 10.5 mL) was

then added and the temperature of the mixture was adjusted to 80 ºC.

Tetraethoxysilane (15.0 mL) was added dropwise to the surfactant solution within 5

min under vigorous magnetic stirring. The mixture was allowed to react for 2 h. The

resulting white solid was filtered, washed with water and methanol, and then dried in

desiccator. Finally, the solid was calcined at 550 ºC for 5 h to remove the organic

template.

Preparation of phenylboronic acid-coated MSN (S1). S0 (200 mg) and 300 mg

(39.6 µmol) of tris(2,2′-bipyridyl)dichlororuthenium(II) hexahydrate were suspended in

25 mL of anhydrous acetonitrile inside a round-bottom flask connected to a Dean-Stark


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trap under Ar atmosphere. The suspension was heated at 110 °C and about 10 mL of

solvent were distilled and collected in the trap to remove the adsorbed water. After

this step, the mixture was stirred for 24 h at room temperature to load the dye into the

MSN face pores.15 Afterward, an excess of (3-glycidyloxypropyl)trimethoxysilane (500

µL, 2.26 mmol) was added and the suspension was stirred for 5 hours. The resulting

solid was filtered off, washed two times with 10 mL of acetonitrile and then with 10 mL

toluene. The solid was dispersed in 30 mL toluene and mixed with 3-aminophenyl

boronic acid (308 mg, 2.26 mmol). The reaction mixture was stirred for 12 h, and the

final solid (S1) was filtered off, washed with toluene and dry at room temperature.

Preparation of esterase-capped MSN (S2). 20 mg of solid S1 were mixed with

5.7 mg esterase in 700 µL of 50 mM sodium phosphate buffer, pH 7.5. The mixture was

stirred overnight at 4ºC, then centrifuged and exhaustively washed with 20 mM Na2SO4

solution at pH 7.5 until no tris(2,2′-bipyridyl)dichlororuthenium(II) can be detected in

the washed solutions. The resulting solid S2 was dried and kept at 4ºC until use.

Preparation of lactose-modified esterase-capped MSN (S3). To prepare the

esterase-based neoglycoconjugates, esterase (5.7 mg) and lactose (12.5 mg, 36.5 µmol)

were dissolved in 1.25 mL of 50 mM sodium phosphate, pH 7.5, and stirred for 1 h at

4ºC. NaBH3CN (12.6 mg, 200 µmol) was further added and the reaction mixture was

stirred overnight at 4ºC. The solution was then exhaustively dialyzed vs. 50 mM sodium

phosphate buffer, pH 7.5 using Amicon Ultra-05 centrifugal filter units with Ultracel-10

membranes (Millipore, USA), and finally concentrated to about 20 mg/mL esterase

concentration. The neoglycoenzyme gated solid S3 was prepared as described above

for solid S2.

Dynamic light scattering and zeta potential. Dynamic Light scattering (DLS) and

zeta potential were performed at 25ºC using a Malvern Zeta Sizer NanoZS instrument.

Measurements were performed with nanoparticles S0, S2 and S3 suspended in filtered

water, phosphate buffer with 10% FBS or cell culture media DMEM with 10% FBS at a

concentration of 50 µg·ml-1.


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Cell culture conditions. HeLa human cervix adenocarcinoma cells were

purchased from the German Resource Centre for Biological Materials (DSMZ) and were

grown in DMEM supplemented with 10% FBS. Cells were maintained at 37 ºC in an

atmosphere of 5% carbon dioxide and 95% air and underwent passage twice a week.

WST-1 Cell Viability Assay. HeLa cells were seeded in a 24-well plate at a density

of 2 ·104 cells/well in a 1000 μL of DMEM and were incubated 24 hours in a CO2

incubator at 37 ºC. Then, DMEM were replaced for PBS with 10% of Fetal Bovine Serum

and solid S4 or S5 in DMSO were added to cells in sextuplicate at final concentrations

of 75 μg·ml-1. DMSO represented 1% (v/v) of the total volume of the cell culture

medium. As a control, we used cells untreated and cells treated with the same volume

of DMSO than cells treated with the nanoparticles in DMSO. After 30 minutes, cells

were washed with PBS and were incubated during 23 hours in different conditions.

DMEM with 10% FBS (+/-), DMEM with 10% FBS and ethyl butyrate (+/+), PBS with 10%

FBS (-/+) or PBS with 10% FBS and ethyl butyrate (-/+). After this, 30 μL of WST-1 were

added to each well and were incubated during 1 hours, a total of 24 hours of

incubation was therefore studied. Before reading the plate, it was shaken for one

minute to ensure homogeneous distribution of colour. Then the absorbance was

measured at a wavelength of 450 nm in VICTOR X5 PerkinElmer. Results are expressed

as a promedium of the results of six independent experiments obtaining similar results.

Live Confocal Microscopy. HeLa cells were seeded in a 24 mm Ø glass coverslips

in six-well plates at a seeding density of 1,8 ·105 cells /well. After 24 hours, culture

medium was replaced for PBS with 10% fetal bovine serum (FBS) and cells were treated

with a suspension of solid S4 or S5 for 30 minutes at a final concentration of 75 µg·ml-1.

Then the medium was changed for different solutions (DMEM with 10% FBS with or

not ethyl butyrate or PBS with 10% FBS with or not ethyl butyrate). After 20 hours

coverslips were washed twice to eliminate compounds and, were visualized under a

confocal microscope employing Leica TCS SP2 AOBS (Leica Microsystems Heidelberg

GmbH, Mannheim, Germany) inverted laser scanning confocal microscope using oil

objectives: 63X Plan-Apochromat-Lambda Blue 1.4 N.A. Confocal microscopy studies

were performed by Confocal Microscopy Service (CIPF). The images were acquired with

an excitation wavelength of 405 for Hoescht and 480 nm for Doxorubicin. Two-


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dimensional pseudo colour images (255 colour levels) were gathered with a size of

1024x1024 pixels and Airy 1 pinhole diameter. Three fields of each condition in two

independent experiments were performed obtaining similar results.

For cellular internalization observation, HeLa cells were seed in six-well plates at

a seeding density of 1,8 ·105 cells /well. After 24 hours, culture medium was replaced

for PBS with 10% fetal bovine serum (FBS) and cells were treated with a suspension of

solid S4 or S5 for 30 minutes at a final concentration of 30 µg·ml-1. Then the medium

was change for different solutions (PBS with 10% FBS with or not ethyl butyrate). After

1 hour (total of 1.5 hour) or 11 hours (total of 12 h) coverslips were washed and the

medium was replaced with PBS with 10% FBS preheat at 37 ºC containing Lysotracker

(50 nM). After 30 minutes, the cells were washed and treated with Hoescht during 5

minutes. After washing with PBS were fixed using 4% formaldehyde during 15 minutes.

Confocal microscopy studies were performed by Confocal Microscopy Service (UPV).

The images were acquired with an excitation wavelength of 405 for Hoescht and 480

nm for Doxorubicin. The cells were monitored under a Leica TCS SP2 laser-scanning

confocal microscope (42x/62x oil objective, 405/488 excitation).

For intracellular Doxo release observation, HeLa cells were seeded according to

the description abovementioned. The cells were washed three times with PBS and then

incubated with S4 or S5 for 30 minutes, then washed again. One sample was fixed then

and the others were incubated during 2, 5 and 30 hours. Cells were washed with PBS

and fixed with fresh 4% formaldehyde at room temperature for 15 min. After washing

with PBS, the cells were subjected to the CLSM observation.

Cytofluorometry studies using S4 and S5. HeLa cells were seeded at 18 ·103 cells

per well in a 24-well plate. After 24 h, DMEM were replaced for PBS with 10% of Fetal

Bovine Serum and solid S4 or S5 in DMSO were added to cells at final concentrations of

75 μg·ml-1. After 30 minutes, cells were washed with PBS and were incubated for 23

hours in the different conditions (DMEM with 10% FBS, DMEM with 10% FBS and ethyl

butyrate, PBS with 10% FBS or PBS with 10 % FBS and ethyl butyrate). After 24 hours

media was eliminated by vacuum and plate were washed once with PBS. Cells were

detached with Trypsin/EDTA solution, centrifuged and finally resuspended in 0.5 ml of


283

DMEM with 10% FBS. Quantification of Doxorubicin fluorescence in the cells was

performed with WinMDI program, version 2.0 in a FC500 MCL Flow Cytometer

(Beckman-Coulter, CA, USA). Three independent experiments containing

quadruplicates were performed with similar results.

For cytotoxicity assays employing flow cytometry, cells were seeded at the same

conditions. After 24 h, solids S0, S4 or S5 in DMSO were added at final concentration of

50 µg· ml-1. After 30 minutes, cells were washed and incubated for 23 hours in the

different conditions. After 24 hours, cells were stained with 7-AAD and Ann V-FITC

according to the manufacturer’s protocol (Life Technologies). Quantification of 7-AAD-

positive and AnnV-positive staining was done by the WinMDI program, version 2.9 in a

FC500 MCL Flow Cytometer (Beckman-Coulter, CA, USA). Three independent

experiments were done and contained triplicates with analogous results.

CONCLUSIONS

In summary, here we demonstrate that the modification of a glycoenzyme by

artificial glycosylation is a useful approach to manipulate the strength of the

supramolecular interaction with phenylboronic acid-coated supports. Based on this

concept, novel neoglycoenzyme-gated MSN, for programmed and pulsatile sequential

cargo delivery, are reported. In particular, gated support S3 is able to release ca. half

the cargo in the presence of glucose, whereas the remaining entrapped payload is

delivered upon the addition of ethyl butyrate. Moreover, it was also demonstrated that

the same control was observed with nanoparticles loaded with an anticancer drug (S5)

and tested in HeLa cells. We believe that the possibility of using a wide variety of

different derivatized enzymes, combined with a variety of functionalized MSN as

supports, opens up new possibilities for the design of novel smart pulsatile delivery

nanodevices for on-command programmed sequential delivery. These, or similar,

release systems have the potential of being applied to diseases for which a pulsatile

drug delivery is preferred and in which triggering can be achieved using simple non-

toxic small molecules.


284

ASSOCIATED CONTENT

Supporting Information

Experimental details, nanomaterials characterization and ex vivo experiments.

This material is available free of charge via the Internet at http://pubs.acs.org.

AUTHOR INFORMATION

Corresponding Author

[email protected], [email protected], [email protected].

Notes

The authors declare no competing financial interests.

ACKNOWLEDGMENT



Science and Innovation CTQ2011-24355, CTQ2009-12650, CTQ2009-09351, MAT2012-

38429-C04-01 and Comunidad de Madrid S2009/PPQ-1642, programme AVANSENS is

gratefully acknowledged. The Generalitat Valencia (project PROMETEOII/2014/047) is

also acknowledged.

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290


NEOGLYCOENZYME-GATED MESOPOROUS SILICA NANOPARTICLES. TOWARDS

THE DESIGN OF NANO-DEVICES FOR PULSATILE PROGRAMMED SEQUENTIAL

DELIVERY

Paula Díez,1 Alfredo Sánchez,1 Cristina de la Torre,2,3 María Gamella,1 Paloma

Martínez-Ruíz,4 Elena Aznar,2,3 Ramón Martínez-Máñez,2,3,* José M. Pingarrón,1,5,*

Reynaldo Villalonga1,5,*

1Department of Analytical Chemistry, Faculty of Chemistry, Complutense

University of Madrid, 28040-Madrid, Spain. 2Instituto de Reconocimiento Molecular y

Desarrollo Tecnológico (IDM), Centro Mixto Universidad Politécnica de Valencia-

Universidad de Valencia, Spain. 3Departamento de Química y CIBER de Bioingeniería,

Biomateriales y Nanomedicina (CIBER-BBN), Universidad Politécnica de Valencia,

Camino de Vera s/n, 46022, Valencia, Spain. 4Department of Organic Chemistry I,

Faculty of Chemistry, Complutense University of Madrid, Madrid, Spain. 5IMDEA

Nanoscience, Cantoblanco Universitary City, 28049-Madrid, Spain.


Chemicals

Pig liver esterase, tetraethoxysilane, cetyltrimethylammonium bromide,

tris(2,2′-bipyridyl)dichlororuthenium(II) hexahydrate, (3-

glycidyloxypropyl)trimethoxysilane, 3-aminophenyl boronic acid, D-glucose, ethyl

butyrate, trypan blue solution (0.4%) cell culture grade and DMSO, PBS and Dulbecco's

Modified Eagle's medium (DMEM) with glucose, L-glutamine and pyruvate for cell

culture were provided by Sigma-Aldrich. Solvents were provided by Scharlau. Fetal

Bovine Serum (FBS), Lysotracker red DND-99, 7-aminoactinomycin D and trypsin were


291

purchased from Life Technologies. Cell proliferation reagent WST-1 was purchased

from Roche Applied Science. All other reagents were of analytical grade.

General Techniques


JEOL JEM-2100 microscope. Spectrophotometric measurements were performed using

an Agilent 8453 UV/VIS spectrophotometer (Hewlett Packard, USA). Powder X-ray

diffraction (XRD) was performed with an X'Pert MRD diffractometer (PANanalytical

B.V., The Netherlands). Nitrogen adsorption/desorption isotherms and pore size

distributions were determined with an ASAP 2020 Physisorption Analyzer

(Micromeritics, USA). Thermal analysis was performed with a TA Instruments SDT-

Q600 apparatus (USA). FT-IR spectra were acquired with a Nicolet Nexus 670/870

spectrometer (Thermo Fisher Scientific Inc., USA). Mass spectra were recorded with an

Ultraflex MALDI-TOF/TOF equipment (Bruker Co., USA). Solid state 13C NMR

measurements were performed with a Bruker AV 400MHz Wide Bore spectrometer

(Bruker Co., USA).

General assays

The esterase activities of the native, lactose-modified and immobilized enzyme

forms were determined at 25ºC in 50 mM sodium phosphate buffer, pH 7.5 (Tris:HCl

buffer, pH 8.0) using p-nitrophenyl acetate as substrate.1 One unit of esterase activity

is defined as the amount of enzyme that hydrolyses 1.0 µmol of p-nitrophenyl acetate

per minute at 25ºC. Protein concentration was estimated as described by Bradford

using bovine serum albumin as standard.2 Total carbohydrates were determined by the

phenol-sulfuric acid method using lactose as standard.3


292

Materials Characterization

Solids S0, S1, S2 and S3 were characterized using standard procedures. Figure

S1 shows representative TEM images of the different nanoparticles, which displayed

the typical porous pattern of the MCM-41 mesoporous matrix with average diameter

of 97 ± 15 nm. S2 and S3 showed a diffuse thin layer surrounding the nanoparticles

suggesting the coverage with an amorphous organic material, which could be ascribed

to the enzyme molecules immobilized on the nanoparticle surfaces.

Figure S1: TEM images of S0 (a), S1 (b), S2 (c) and S3 (d) nanoparticles.

Figure S2 shows the powder X-ray diffraction patterns of the capped

nanoparticles S2 and S3 and the intermediate supports. Solid S0 exhibited three peaks

at small angles, which are typical of hexagonal ordered structures in MCM-41 materials

and correspond to (100), (110), and (200) planes. All other samples showed a well-

defined peak at ca. 2.59º attributed to the (100) Bragg reflection of MCM-41-type

materials (d-spacing = 3.40 nm). This fact suggests that the characteristic mesoporous

order of the initial silica nanoparticles was not affected by dye loading process and

further chemical modifications carried out. However, the intensity of the peaks related

to the (110) and (200) planes was significantly reduced after nanoparticles

modification.


293

Figure S2: Powder X-ray diffraction of S0 (a), S1 (b), S2 (c) and S3 (d) nanoparticles at

low angles.

Figure S3 shows the FT-IR analysis of the MSN-based nanodevives. FT-IR

spectrum of the S0 nanoparticles was similar to those reported for MSN, with a major

band at 1090 cm-1, corresponding to SiO stretching vibrations.7 In addition, S1 showed

the characteristics absorption bands of phenylboronic acid derivatives with major

bands corresponding to the OH stretching vibration at 3400 cm-1, aromatic CC

stretching vibrations at 1599 cm-1, 1449 cm-1 and 1240 cm-1, and the BO stretching

vibration at 1352 cm-1.8 The FT-IR spectrum of the S2 and S3 nanodevices were mainly

characterized by the strong absorption bands of the native and lactose-modified

glycoenzymes, with broad carbohydrate bands at 3120 cm-1 and 1060 cm-1 and the

protein amide I absorption band at ca. 1635 cm-1.


294

Figure S3. FT-IR analysis for S0 (a), S1 (b), S2 (c) and S3 (d) nanoparticles.

The assembly of the nanodevice was confirmed by solid state 13C NMR. Figure

S4A shows the spectrum of MSN loaded with tris(2,2′-bipyridyl)dichlororuthenium(II)

hexahydrate and further modified with epoxy groups, which showed the

characteristics peaks of the aromatic rings in the dye at 158 ppm, 149 ppm, 139 ppm,

130 ppm and 125 ppm. In addition, the peak at 49 ppm can be ascribed to the epoxy

groups on the nanoparticle surface. A noticeable reduction in the intensity of the peak

at 49 ppm was observed in the spectrum of S1 solid, due to reaction of epoxy groups

with 3-aminophenyl boronic acid. In addition, new peaks at 145 ppm, 127 ppm, 121

ppm and 112 ppm are observed, which can be ascribed to the aromatic rings in the

attached phenyl boronic acid residues. No covalent immobilization of esterase on S1

caused the appearance of new peaks at 75 ppm and 26 ppm, which could be assigned

to aliphatic carbon atoms in the protein. However, it was not possible to detect the

characteristic peak of carbon atoms at the peptide bond around 170-180 ppm.


295

Figure S4. Solid state 13C NMR spectra for S0 loaded with tris(2,2′-

bipyridyl)dichlororuthenium(II) hexahydrate and modified with epoxy groups (A), S1 (B)

and S2 (C) nanoparticles.


296

The nanomaterials were also characterized by thermal analysis, and the

corresponding TG/DTG curves are shown in Figure S5. All samples exhibited a slight

weight loss in the range of temperatures from 50ºC to about 120ºC, which can be

attibuted to the thermodesorption of physically adsorbed water molecules. No

significant weight loss was appreciated for S0 nanoparticles at higher temperatures.4

S1 showed noticeable weight loss in the 240-365ºC range probably caused by the

thermal decomposition of the grafted amino phenylboronic acid derivative. A further

gradual weight decrease was observed in the 365-550ºC range of temperatures with a

maximum rate of transformation at about 442ºC, which could be ascribed to the

decomposition of the [Ru(bpy)3]Cl2 complex encapsulated into the nanopores.5 The

loading of the [Ru(bpy)3]Cl2 dye in the S1 material was estimated as 206 mg/g, as

determined by the TG curve. Moreover the content of the anchored boronic acid-

based ligand amounted to 34 mg/g S1 in weight.

Figure S5. A) TG and B) DTG analysis for S0 (a), S1 (b), S2 (c) and S3 (d) nanoparticles.

A more remarkable weight loss, representing about 17.3% by weight of the

nanomaterial, was observed for S2. This esterase-capped material exhibited an

additional thermal-induced transformation in the 95-289ºC range of temperatures

with a maximum rate of transformation at 204ºC, which could be attributed to the

thermal decomposition of the immobilized glycoenzyme. On the other hand, S3

nanomaterial showed a more complex thermal transformation profile with multiple

decomposition steps in the 95-358ºC range with maximum rates of transformation at

112ºC, 149ºC, 172ºC and 221ºC as revealed by DTG analysis. This different thermal


297

behavior of S3 could be justified by the higher amount of lactose moieties attached to

esterase, causing increased molecular heterogeneity and thermal stability, as

previously reported for other neoglycoenzymes.6 S3 nanoparticles also showed

noticeable weight loss, representing about 32.8% by weight, which suggest the

presence of a high amount of neoglycoenzyme immobilized on the nanoparticle

surface.

Figure S6. A) Nitrogen adsorption (closed)/desorption (open) isotherms and B) pore size

distribution for S0 (a), S1 (b), S2 (c) and S3 (d) nanoparticles.

As can be observed in Figure S6, the N2 adsorption-desorption isotherms of the

starting calcined MSN showed an adsorption step at intermediate P/P0 values (0.1–

0.3). The application of the BET model resulted in values for the total specific surface

area of 1090.2 m2 g-1 and an average pore diameter of 2.23 nm. In contrast, the N2

adsorption-desorption isotherms for the prepared dye loaded phenylboronic acid-

modified material (S1) and the enzyme capped nanodevices (S2 and S3) are typical of

mesoporous systems with filled mesopores with values of specific surface area of 8.7,

354.2 and 408.0 m2 g-1 for S1, S2 and S3, respectively. The larger surface areas

observed or S2 and S3 when compared with S1 could be attributed to a partial delivery

of the entrapped cargo during the attachment of the enzymes and/or due to

interstitial adsorption of N2 molecules into the capping protein layer.


298

Table S1. BET specific surface values, pore volumes and pore sizes calculated from the

N2 adsorption-desorption isotherms for selected materials.

a Pore size estimated by using the BJH model applied on the adsorption branch of the isotherm, for P/P0 < 0.4, which can be associated to the surfactant generated mesopores.

b Total pore volume according to the BJH model.

Figure S7. Nanoparticle size distribution of S0, S2 and S3 in H2O (black line), PBS (dotted

line) and DMEM (striped line).

SBET

(m2g-1)

BJH pore

(P/P0 < 0.4)a

(nm)

Total pore

volumeb

(cm3g-1)

S0 1090.2 2.23 0.66

S1 8.7 - 0.001

S2 354.2 - 0.14

S3 408.0 - 0.17


299

Table S2. Nanoparticle size (nm) and zeta potential values (mV) of S0, S2 and S3 in pure

H2O, PBS buffer solution with 10 % FBS and DMEM culture medium with 10 % FBS.

Material Size in H2O

(nm) Size in PBS

(nm)

Size in cell culture media

(nm)

Zeta potential (mV)

H2O/PBS/Cell culture media

S0 295 320 384 -26/-21/-19

S2 211 203 381 -7/-8/-12

S3 196 222 394 -5/-9/-15

Figure S8. TEM images of S1 (A), S2 (B) and S3 (C) nanoparticles after staining with 1%

uranyl acetate.


300

Figure S9. Trigger reactions mediated by D-glucose (A) and ethyl butyrate (B).


301

Figure S10. A) Confocal images of HeLa cells incubated with solid S4 for 30 minutes and

then treated with input A (D-Glucose) and/or input B (ethyl butyrate) or untreated.

Representative images at 24 h from phase contrast (PhC), Doxorubicin (Doxo), Hoescht

(Hoe) and combined (Merged) are shown. B) Cell viability (%) of HeLa cells treated with

S5 using WST-1 assay. Results expressed in % (refereed to untreated cells) ± standard

deviation of three different experiments. C) Mean fluorescent intensity of Doxo in HeLa

cells treated with S4 using FACS analysis.


302

Figure S11. A) Confocal images of HeLa cells incubated with solid S5 for 30 minutes and

then treated with input A (D-Glucose) and/or input B (ethyl butyrate) or untreated.

Representative images at 24 h from phase contrast (PhC), Doxorubicin (Doxo), Hoescht

(Hoe) and combined (Merged) are shown. B) Cell viability (%) of HeLa cells treated with

S5 using WST-1 assay. Results expressed in % (refereed to untreated cells) ± standard

deviation of three different experiments. C) Mean fluorescent intensity of Doxo in HeLa

cells treated with S5 using FACS analysis.


303

Figure S12. Quantification of cell viability and cell death was performed by flow

cytometry by means of 7-AAD and Ann V- FITC staining for S4. The percentage of dead

cells (black), cells undergoing cell death (gray) and healthy cells (white) are shown after

24 h of treatment. Three independent experiments containing triplicates were

performed and the data are reported as (mean ± SE).


304

Figure S13. Quantification of cell viability and cell death was performed by flow

cytometry by means of 7-AAD and Ann V- FITC staining for S5. The percentage of dead

cells (black), cells undergoing cell death (gray) and healthy cells (white) are shown after

24 h of treatment. Three independent experiments containing triplicates were

performed and the data are reported as (mean ± SE).


305

Figure S14. CLSM images of HeLa cells incubated with 30 μg/ml of S4 for 1 hour,

washed and then incubated during 1.5h and 12 h in PBS with 10 % FBS (-/-) or PBS with

10 % FBS and ethylbutyrate (-/+) . Lysotracker was used to stain the organelles of the

cell and Hoescht for the nucleus.


306

Figure S15. CLSM images of HeLa cells incubated with 30 μg/ml of S5 for 1 hour and

post incubated during 1.5h and 12 h in PBS and 10 % FBS (-/-) or PBS with 10 % FBS and

ethylbutyrate (-/+) . Lysotracker was used to stain the organelles of the cell and

Hoescht for the nucleus.


307

Figure S16. CLSM images of HeLa cells incubated with 30 μg/ml of S4 or S5 during 1

hour, washed and then incubated in PBS with fetal bovine serum (No input) or in PBS

with ethyl butyrate (Input I2). The green fluorescence is from the endocytosed

nanoparticles and from released Doxo.

References

(1) Degrassi, G.; Uotila, L.; Klima, R.; Venturi, V. Purification and Properties of an

Esterase from the Yeast Saccharomyces cerevisiae and Identification of the

Encoding Gene. Appl. Environ. Microbiol. 1999, 65, 3470-3472.

(2) Bradford, M.M. A Rapid and Sensitive Method for the Quantitation of Microgram

Quantities of Protein Utilizing the Principle of Protein-Dye Binding. Anal. Biochem.

1976, 72, 248-254.


308

(3) Dubois, M.; Gilles, K.A.; Hamilton, J.K.; Rebers, P.T.; Smith, F. Colorimetric Method

for Determination of Sugars and Related Substances. Anal. Chem. 1956, 28, 350-

356.

(4) Kumar, R.; Chen, H.T.; Escoto, J.L.; Lin, V.S.Y.; Pruski, M. Template Removal and

Thermal Stability of Organically Functionalized Mesoporous Silica Nanoparticles.

Chem. Mat. 2006, 18, 4319-4327.

(5) Armelao, L.; Bertoncello, R.; Gross, S.; Badocco, D.; Pastore, P. Construction and

Characterization of Ru(II)Tris(bipyridine)-Based Silica Thin Film

Electrochemiluminescent Sensors. Electroanalysis 2003, 15, 803-811.

(6) Villalonga, M.L.; Fernández, M.; Fragoso, A.; Cao, R.; Villalonga, R. Functional

Stabilization of Trypsin by Conjugation with β-Cyclodextrin-Modified

Carboxymethylcellulose. Prep. Biochem. Biotechnol. 2003, 33, 53-66.


Enzyme-Controlled Sensing–Actuating Nanomachine Based on Janus Au–

Mesoporous Silica Nanoparticles. Chem. Eur. J. 2013, 19, 7889-7894.

(8) Faniran, J.A.; Shurvell, H.F. Infrared Spectra of Phenylboronic Acid (Normal and

Deuterated) and Diphenyl Phenylboronate. Can. J. Chem. 1968, 46, 2089-2095.

7.10

Electrochemistry Communications 30 (2013) 51–54

Janus Au-Mesoporous silica

nanoparticles as electrochemical biorecognition-signaling system

Electrochemistry Communications 30 (2013) 51–54 7. PUBLICACIONES

309

JANUS AU-MESOPOROUS SILICA NANOPARTICLES AS ELECTROCHEMICAL

BIORECOGNITION-SIGNALING SYSTEM

Alfredo Sánchez,1 Paula Díez,1Paloma Martínez-Ruíz,2 Reynaldo Villalonga,1,3,*

José M. Pingarrón1,3,*

1Department of Analytical Chemistry and 2Department of Organic Chemistry I, Faculty

of Chemistry, Complutense University of Madrid, 28040-Madrid, Spain.

3IMDEA Nanoscience, 28049-Madrid, Spain.

*Corresponding authors. Phone: +34 91 3944315, E-mail: [email protected],

[email protected]


310

ABSTRACT

Janus Au-mesoporous silica nanoparticles were used as scaffolds to design an

integrated electrochemical biorecognition-signaling system. A proof of concept of this

strategy, based on the face-selective functionalization of the anisotropic colloid,

involves the covalent immobilization of horseradish peroxidase on the mesoporous

silica face as enzymatic signaling element, as well as the modification of the Au face

with streptavidin and polyethylenglycol chains as biorecognition and solubilizing

agents, respectively. The functionalized Janus nanoparticles were sucessfull to

recognize biotin on gold surfaces.

KEYWORDS: Janus nanoparticle, mesoporous silica, electrochemical biosensor, enzyme

.

INTRODUCTION

Nanomaterials-based biosensors is a continuously growing field in

electroanalysis [1]. This trend is supported by the unique physicochemical, structural

and surface-to-volume ratio properties of nanomaterials, which allow their multiple

functional roles in biosensor design. Nanomaterials are also excellent building blocks

for the assembly of a wide variety of novel three-dimensional nanoarchitectures at the

electrode surface [2,3], favoring the spatial arrangement of the analytical biomolecules

in more propitious microenvironments.

In this context, nanoparticles are by far the nanomaterials most widely

employed in electrochemical biosensing [4]. Nanoparticles can be used to enhance the

amount of immobilized biomolecules, catalyze biochemical reactions, increase

efficiency of photochemical reactions and facilitate the electron transfer processes on

the electrode surface. Nanoparticles can be also employed as electrochemical labels as

well as to amplify transduction of biomolecular recognition events [5].

A great variety of nanoparticles differing in size, shape and composition have

been employed in bioelectrochemical analysis. Although Janus nanoparticles have


311

been proposed recently for electrochemical sensing [6], however, they have never

been explored for the development of electrochemical biosensors. Janus nanoparticles

are anisotropic materials showing two different faces with dissimilar chemical

composition and properties [7,8]. This structural characteristic allows their zone-

specific functionalization with selected biomolecules, which can be relevant to design

novel multienzymatic and affinity-based biosensors.

In this communication we describe for the first time the use of Janus

nanoparticles as biorecognition-signaling element in electrochemical biosensing. As a

first proof-of-concept, Janus Au-mesoporouos silica nanoparticles (Au-MS JNP) were

prepared and further functionalized with horseradish peroxidase (HRP, EC 1.11.1.7) on

the silica face, whereas the Au surface was modified with streptavidin.

Methoxypolyethylene glycol thiol (PEG-SH, MW = 5000) chains were also attached to

the metal face to confer solubility to the nanoparticle. This nanomaterial was finally

evaluated toward a thiolated biotin derivative previously chemisorpted on Au surfaces.


1. Reagents

HRP, streptavidin, (3-aminopropyl)triethoxysilane (APTES), (3-mercaptopropyl)-

trimethoxysilane, PEG-SH and biotin were purchased from Sigma-Aldrich Co. 3,3´-

Dithiobis(sulfosuccinimidylpropionate) (DTSSP) was purchased from Thermo Fisher

Scientific.

2. Preparation of functionalized Au-MS JNP

Calcined mesoporouos silica nanoparticles (200 mg, 97 ± 15 nm) [9] were

dispersed in 1 g of paraffin wax at 75 °C and then mixed with 10 mL of water. The

mixture was stirred during 1 h, then cooled and the resulting colloidosomes were

collected by decantation, washed with water, and dispersed in 75 mL of methanol. The

mixture was treated with (3-mercaptopropyl)trimethoxysilane (30 mM final

concentration) during 30 min under stirring. The suspension was filtered, washed with


312

methanol and dispersed in 20 mL of 3 µM solution of 20 nm citrate-capped Au

nanoparticles [10]. The mixture was stirred during 1 h, then filtered, sequentially

washed with water, ethanol and chloroform, and finally dried.

Au-MS JNP (200 mg) were dispersed in 200 mL methanol and mixed with 8 mL

of APTES. After 3 h under continuous stirring, the solid was filtered, washed with

methanol and dried. The APTES-modified nanoparticles (200 mg) were dispersed in 20

mL of 50 mM sodium phosphate buffer, pH 7.0, and treated with glutaraldehyde (5%

v/v, final concentration) during 1 h under stirring in the dark. The activated

nanoparticles were collected by centrifugation, washed with buffer solution and then

dispersed in 10 mL of the same buffer. HRP (10 mg) was further added and the mixture

was stirred at 4ºC during 2 h. The resulting Au-MS JNP-HRP colloid was collected by

centrifugation, washed with cold buffer solution and then dispersed in 2 mL of buffer.

In parallel, streptavidin (100 µg) and DTSSP (4.0 mg) were dissolved in 2.0 mL of

50 mM sodium phosphate buffer, pH 7.0, and stirred during 2 h at 4 ºC. Thereafter, 20

µL of 100 mM NaBH4 solution were added, and the mixture was stirred at 4 ºC for 30

min. The solution was dialyzed vs. 50 mM sodium phosphate buffer, pH 7.0 and then

added to 5 mL of sodium phosphate buffer, pH 7.0, containing 50 mg of Au-MS JNP-

HRP, and stirred at 4ºC overnight. The resulting solid was isolated by centrifugation,

washed several times and dispersed in 10 mL of the same cold buffer solution. PEG-SH

(4 mg) was added and the mixture was stirred at 4ºC during 2 h. The mixture was

centrifuged and the resulting Au-MS JNP-HRP-Stv-PEG colloid was washed several

times with 50 mM sodium phosphate buffer, pH 7.0, and kept in refrigerator until use.

3. Synthesis of biotin-cystamine

Biotin (0.503 mmol) was dissolved in 2.0 mL of anhydrous dimethylformamide

under Ar, and then mixed with N-(3-dimethylaminopropyl)-N′-ethylcarbodiimide

hydrochloride (0.572 mmol) and N-hydroxysuccinimide (0.515 mmol). The mixture was

stirred during 1 h and then cystamine dihydrochloride (0.229 mmol) and triethylamine

(2.06 mmol) were successively added. The mixture was stirred for 4 days at RT and

concentrated in vacuo. After addition of water (1 mL), the precipitate was filtered and


313

sequentially washed with saturated NaHCO3 (2 mL), water (2 mL) and 10% (v/v) HCl (2

mL) solutions, and finally dried (54% yield). The purity and identity of the product was

checked by TLC, IR and NMR; experimental data agree with those described by

Dondoni et al starting from biotin-succinimide ester [11].

4. Electrode modification

To evaluate the biorecognition ability of Au-MS JNP-HRP-Stv-PEG, a 5 mM

biotin-cystamine solution was first treated with 20 µL of 100 mM NaBH4 solution, and

further dropped (20 µL) on the surface of a gold disk electrode (2.0 mm diameter).

After 1 h incubation at room temperature, the electrode was washed with water and

further coated with 10 µL of 1.0 mg/mL Au-MS JNP-HRP-Stv-PEG solution in 50 mM

sodium phosphate buffer, pH 7.0. After 1 h incubation at 4ºC, the Au/biotin/Au-MS

JNP-HRP-Stv-PEG electrode was exhaustively washed with cold buffer solution. A

similar procedure was employed to modify Au-coated glass slides for SPR and AFM

experiments.


The strategy employed to prepare these novel functionalized nanomaterials is

illustrated in Figure 1. Au-MS JNP with average size of 104 ± 17 nm were prepared via

stable and multi-punctual Au-S binding by Pickering emulsion template, and used as

scaffolds to integrate a biological recognition element and an enzymatic signaling

system for biosensing purposes. We take advantage over the anisotropic chemical

properties of Janus nanoparticles [7,8] for the face-selective assembly of these

biochemical elements on the two different faces of the colloid. In this particular case,

and as a first proof-of-concept, the signaling enzyme HRP was covalently immobilized

on the mesoporous silica face, whereas the gold surface was functionalized with

streptavidin which serve as affinity-based biorecognition element.


314

Figure 1. Preparation of the Janus nanoparticle-based biorecognition-signaling system.

To achieve these goals, the mesoporous silica face of Au-MS JNP was first

enriched with primary amino groups by treatment with APTES. These amino groups

were then activated by treatment with glutaraldehyde to accomplish the covalent

immobilization of HRP on the mesoporous silica face. On the other hand, streptavidin

was reacted with DTSSP and further treated with NaBH4 to provide the protein surface

with reactive thiol groups, allowing the protein immobilization on the gold face of Au-

MS JNP through a chemisorption process.

A more conventional approach for streptavidin immobilization could be the

initial capping of the gold face of Au-MS JNP with a thiol-containing hetero bifunctional

cross-linking agent having a secondary reactive group able to couple to the protogenic

groups on the streptavidin surface. However, a major disadvantage of the protein

functionalized Janus colloid is the low solubility in aqueous solutions. Additionally,

these nanoparticles cannot be dispersed by conventional ultrasound treatment due to

low conformational stability of the immobilized proteins, which can denaturize and

loose biological activity upon ultrasound radiation. As a mean to overcome this

limitation, we left uncapped a major area of the gold face in the streptavidin

immobilization step so that the remaining metal surface could be used to anchor PEG-


315

SH molecules as solubilizing agents.

A representative TEM image of the resulting Au-MS JNP-HRP-Stv-PEG colloid is

shown in Figure 1 (inset). These nanoparticles allowed stable aqueous dispersions and

were evaluated as biorecognition-signaling system toward biotin.

Figure 2. A) SPR sensograms recorded upon modification of Au surfaces with thiolated

biotin (1), Au-MS JNP-HRP-Stv-PEG (2) and Au-MS JNP-HRP-PEG (3). B) AFM analysis of

Au-MS JNP-HRP-Stv-PEG adsorbed on biotin-modified Au surface.

The changes occurred on the gold surface after sequential exposure to

thiolated biotin and Janus nanoparticles were evaluated by SPR. Figure 2A shows that

the incubation with the biotin derivative caused a noticeable increase in the SPR signal

which remained high after washing, suggesting that the thiolated biotin was

successfully chemisorbed on the Au surface. Further incubation with the Au-MS JNP-

HRP-Stv-PEG colloid also caused a significant increase in the SPR signal (curve a), thus

demonstrating the interaction between the chemisorpted biotin molecules and the

streptavidin-modified nanoparticles. A lower increase in the SPR signal was observed

by incubating the biotin-modified Au surface with the control colloid lacking

streptavidin (curve b), which was almost completely eliminated after a washing step,

conversely to that occurred with the streptavidin-modified nanoparticles, thus

suggesting the affinity-based biorecognition of Au-MS JNP-HRP-Stv-PEG. A similar

effect was observed for a control experiment carried out with non-biotinylated Au

surfaces (curve c).


316

The topology of the biotin-modified Au surface after incubation with Au-MS

JNP-HRP-Stv-PEG was characterized by AFM (Figure 2B). Nanoparticulated Janus

structures were clearly observed on the metal surface after incubation with the colloid,

evidencing adsorption of the streptavidin-modified nanoparticles on the biotin-coated

material.

Figure 3. A) Nyquist plots and B) cyclic voltammograms obtained at a bare Au

electrode (a) and after modification with thiolated biotin (b), Au-MS JNP-HRP-Stv-PEG

(c) and Au-MS JNP-HRP-PEG (d) in 0.1 M KCl, 5 mM K3[Fe(CN)6]/K4[Fe(CN)6]. C) Cyclic

voltammograms recorded with Au/biotin/Au-MS JNP-HRP-Stv-PEG (a) and

Au/biotin/Au-MS JNP-HRP-PEG (b) electrodes in the absence and presence (*) of 50 µM

H2O2 in 0.1 M sodium phosphate buffer, pH 7.0.

Electrochemical impedance spectroscopy was performed to evaluate the

interfacial changes on the Au electrodes after sequential modification with thiolated

biotin and Au-MS JNP-HRP-Stv-PEG (Figure 3A). A noticeable increase in the semicircle


317

diameter of the Nyquist plot occurred after modification of the electrode surface with

the biotin derivative indicating a high coverage of the electrode surface. A remarkable

further increase of the electron transfer resistance was also apparent after incubation

of the biotin-coated Au electrode with Au-MS JNP-HRP-Stv-PEG, suggesting the

attachment of the modified nanoparticles with significant insulating effect on the

electrode surface. Conversely, a much lower increase in the semicircle diameter was

observed in the Nyquist plot corresponding to the electrode modified with the control

nanoparticle lacking streptavidin.

Similar conclusion was achieved from cyclic voltammetry (Figure 3B), attending

to the decrease in the peak currents observed after sequential modification of the

electrode. By using the Randles-Sevcik equation, the electrochemical surface area of

the Au electrode was estimated to decrease from 3.58 mm2 to 3.30 and 2.13 mm2 after

modification of with thiolated biotin and Au-MS JNP-HRP-Stv-PEG, respectively.

Comparatively, the estimated surface area for the Au/biotin/Au-MS JNP-HRP-PEG

control electrode was 2.89 mm2. These results suggest lower non-specific adsorption

of the nanoparticles non containing streptavidin on the biotin-coated gold surface.

The results achieved in the experiments described above suggested that Au-MS

JNP-HRP-Stv-PEG has the ability to specifically biorecognize biotin residues on an Au

surface. The capacity of this colloid to produce an enzyme-mediated electroanalytical

signal was then estimated by comparing the cyclic voltammetric responses of the

Au/biotin/Au-MS JNP-HRP-PEG and Au/biotin/Au-MS JNP-HRP-Stv-PEG electrodes

upon addition of H2O2. As it is shown in Figure 3C, an increase in the cathodic currents

was observed at both modified electrodes in the presence of 50 µM H2O2. However,

large cathodic currents were appreciated in the electrode modified with Au/biotin/Au-

MS JNP-HRP-Stv-PEG, suggesting higher amount of these nanoparticles adsorbed on

the electrode in which the immobilized HRP was successful to electrocatalyze the

transformation of H2O2.


318

CONCLUSIONS

A novel nanomaterial-based biorecognition-signaling system was designed by

using Janus nanoparticles with opposite Au and mesoporous silica faces as scaffolds for

the face-selective assembly of HRP and streptavidin. These nanoparticles were

successful evaluated for the affinity-based recognition of biotin attached to gold

surfaces, as revealed by electrochemical and microscopic characterization. Although

this Janus nanoparticles-based biorecognition-signaling system with electrochemical

detection is here proposed as an initial proof-of-concept, it is predictable a direct use

of this nanomaterial in the design of electrochemical immuno- and genosensors via

linking of commercially available biotin-labeled antibodies and nucleic acids.

Acknowledgements

R. Villalonga acknowledge to Ramón & Cajal contract from the Spanish Ministry of

Science and Innovation. Financial support from the Spanish Ministerio de Ciencia e

Innovación CTQ2011-24355, CTQ2012-34238 and Comunidad de Madrid S2009/PPQ-

1642, Programme AVANSENS is gratefully acknowledged.

References

[1] J. Wang, Analyst 130 (2005) 421.

[2] R. Villalonga, M.L. Villalonga, P. Díez, J.M. Pingarrón, J. Mat. Chem. 21 (2011)

12858.

[3] M. Holzinger, L. Bouffier, R. Villalonga, S. Cosnier, Biosens. Bioelectron. 24 (2009)

1128.

[4] J.M. Pingarrón, P. Yáñez-Sedeño, A. González-Cortés, Electrochim. Acta 53 (2008)

5848.

[5] J. Wang, Small 1 (2005) 1036.

[6] P. Biji, A. Patnaik, Analyst 137 (2012) 4795.


319

[7] S. Jiang, Q. Chen, M. Tripathy, E. Luijten, K.S. Schweizer, S. Granick, Adv. Mater. 22

(2010) 1060.

[8] A. Walther, A.H.E. Müller, Soft Matter 4 (2008) 663.

[9] Y. Zhao, B.G. Trewyn, I.I. Slowing, V.S.Y. Lin, J. Am. Chem. Soc. 131 (2009) 8398.

[10] G. Frens, Nature 241 (1973) 20.

[11] M.L. Conte, S. Pacifico, A. Chambery, A. Marra, A. Dondoni., J. Org. Chem. 75

(2010) 4644.

7.11

Chemistry - A European Journal 19 (24) (2013) 7889–7894

Enzyme-controlled sensing-actuating nanomachine based on Janus Au-Mesoporous silica

nanoparticles

Chemistry - A European Journal 19(24) (2013) 7889–7894 7. PUBLICACIONES

321

ENZYME-CONTROLLED SENSING-ACTUATING NANOMACHINE BASED ON

JANUS AU-MESOPOROUS SILICA NANOPARTICLES

Reynaldo Villalonga,*[a,b] Paula Díez,[a] Alfredo Sánchez,[a] Elena Aznar,[c,d] Ramón

Martínez-Máñez,[c,d] José M. Pingarrón*[a,b]

[a]Department of Analytical Chemistry, Complutense University of Madrid, 28040-

Madrid, Spain, Fax: (+34) 913944329. E-mail: [email protected],

[email protected]. [b]IMDEA Nanoscience, Cantoblanco Universitary City, 28049-

Madrid, Spain. [c]Departamento de Química and CIBER de Bioingeniería, Biomateriales

y Nanomedicina (CIBER-BBN), Universidad Politécnica de Valencia, Camino de Vera s/n,

E-46022, Valencia, Spain, Fax: (+34) 963879349. E-mail: [email protected],

[email protected] [d]Instituto de Reconocimiento Molecular y Desarrollo Tecnológico

(IDM), Centro Mixto Universidad Politécnica de Valencia-Universidad de Valencia,

Spain


322

ABSTRACT

Novel Janus nanoparticles with Au and mesoporous silica opposite faces were

prepared by Pickering emulsion template using paraffin wax as oil phase. These

anisotropic colloids were employed to design an integrated sensing-actuating

nanomachine for the enzyme-controlled stimulus-responsible cargo delivery. As a

proof-of-concept, we demonstrated the successful use of the Janus colloids for

controlled delivery of tris(2,2´-bipyridyl) ruthenium(II) chloride from the mesoporous

silica face grafted with pH-sensitive gate-like scaffoldings. The release was mediated by

the on-demand catalytic decomposition of urea by urease, which was covalently

immobilized on the Au face.

KEYWORDS: Janus nanoparticle, mesoporous silica, molecular gates, controlled

release, enzyme

INTRODUCTION

The development of novel biologically-inspired nanomachines and smart drug

delivery systems is linked to the tailor-made design of advanced nanomaterials with

desired physical properties and chemical functionalities.[1] Among these, particular

interest has been devoted to the preparation of anisotropic colloidal particles which

exhibit two surfaces of different chemical composition.[2] These so-called “Janus

nanoparticles” can be designed to show amphiphilic character as well as anisotropic

electrical, magnetic or optical properties.[3] Additionally, each face of the Janus

nanoparticles can be independently modified with selected ligands allowing specific

functionalization with proteins and other biomacromolecules.[4] These unique

characteristics have favored the successful use of Janus nanomaterials as emulsion

stabilizers, hydrophobic coat for textiles, self-propelled machines, imaging probes, and

drug delivery systems.[5]

On a different approach nanotechnology has proved to bring new innovative

concepts to drug-delivery therapies. Drug delivery systems able to release active


323

molecules to certain cells in a controlled manner have recently gained much attention.

Among several potential drug delivery systems, mesoporous silica nanoparticles (MS)

have been widely used in the past years as reservoirs for drug storage due to their

unique properties such as a large specific volume, large loading capacity, low toxicity

and easy functionalization.[6] Moreover MS nanoparticles can be functionalized with

molecular/supramolecular ensembles on their external surface to develop gated-MS

showing “zero delivery” and capable to release of their cargo in response to external

stimuli. Using this concept, MS displaying controlled release using several stimuli such

as pH, light, redox substances, small molecules and biomolecules have been

reported.[7] However, a potential limitation in these systems is related with the fact

that the delivery MS-based support and the effector (i.e. agent that mediates the

delivery) are not in the same nanoparticle and usually the gated materials are placed in

a solution in which the triggering stimuli is applied (for instance light) or it is present in

the media (for instance enzymes that induced the degradation of a specific gating

coating).

As an advance in the design of more sophisticated nanoparticles for delivery

applications we envisioned that it might possible to project systems in which the gating

systems and an effector molecule could be placed in the same nano-device. In order to

achieve this goal the strategy we have followed in shown in Scheme 1 and it involves

the preparation of new Janus Au-MS nanoparticles by Pickering emulsion template

using paraffin wax as oil phase. This allows obtaining two different surfaces with well-

defined functionalization chemistries for the independent anchoring of the gated

ensemble (on the MS face) and the effector molecule (on the Au face).

In this particular case, and as a first proof-of-concept, the MS part of the

anisotropic colloid was capped with a pH-responsive gate, whereas the gold surface

was functionalized with the enzyme urease (EC 3.5.1.5). We reasoned that the gated

mesoporous nano-devices would show “zero-release”, yet selectively will open the pH-

responsive gate releasing the cargo in the presence of urea via urease-mediated urea

hydrolysis which will lead to an increase of the pH.


324

Scheme 1. Preparation of Janus Au-MS nanoparticles for enzyme-controlled release.


The starting MS nanoparticles (a calcined MCM41-like solid) were synthesized

by alkaline hydrolysis of tetraethyl orthosilicate as inorganic precursor in the presence

of the cationic surfactant cetyltrimethylammonium bromide as porogen species.[8] The

MS showed an average particle diameter of 97 ± 15 nm and an MCM-41 type channel-

like mesoporous structure (Figure 1S in Supporting Information).

To synthesize the Janus nanoparticles, a rational design based on the

manipulation of the Au-ligan-MS interface through mask-protecting assisted site-

selective modification was employed. First, the surface of MS nanoparticles was

partially masked by confining at the interface of Pickering emulsion. The exposed

nanoparticle surface was further modified with a thiolated silane derivative, providing

reactive sulfhydryl groups on this face. Au nanoparticles were then attached to the

thiol-enriched face of the adsorbed MS nanoparticles through chemisorption

reactions, forming stable anisotropic colloids. Several anisotropic nanomaterials have

been previously prepared in excellent yield by manipulation of nanoparticle-ligand-


325

nanoparticle interfaces in solutions.[9] In the present work, such toposelective

manipulation was performed in a solid-liquid interface.

According to this synthetic scheme, the as-prepared MS were first adsorbed

onto the liquid-liquid interface of an emulsion prepared with water and molten

paraffin wax, forming colloidosomes that remained stable after cooling. [10] Although

the formation of such colloidosomes was previously reported using silica nanoparticles

with average diameters larger than 400 nm,[10,11] we have proved here that this

procedure can also be also used using MS nanoparticles of smaller diameter.

The toposelective modification of the MS nanoparticles adsorbed onto the

colloidosome surface was performed by reaction with (3-mercaptopropyl)-

trimethoxysilane in water:methanol solution. After exhaustive washing, the

colloidosomes containing the thiol-functionalized MS nanoparticles were stirred in Au

nanoparticles solution. Janus Au-MS nanoparticles were finally obtained with

acceptable yield (about 85% for J2 sample) after dissolving the paraffin wax in CHCl3.

Figure 1 shows the TEM images of Janus colloids prepared by using Au

nanoparticles of different sizes: 3.7 ± 0.8 nm (J1),[12] 20 ± 2 nm (J2), 31 ± 6 nm (J3) and

43 ± 5 nm (J4).[13] In all cases, anisotropic colloids were successfully synthesized, mainly

with a 1:1 Au:MS nanoparticles ratio except for J1 samples in which the use of smaller

Au nanospheres favored the attachment of more than one metal nanoparticle to the

same MS colloid. J1 nanoparticles showed similar average diameter (98 ± 17 nm) and

size distribution than the native MS colloid, with only slight difference at higher values

of diameter (Figure 2S in Supporting Information). This fact can be justified by the

small diameter and low polydispersity of the attached Au nanoparticles. Moreover, J1

anisotropic colloids showed low metal content, which should be counterproductive for

the further immobilization of urease and the preparation of the enzyme-controlled

nanomachines.


326

Figure 1. TEM images of J1 (A), J2 (B), J3 (C) and J4 (D) Janus Au-MS nanoparticles.

Large average diameter and broad size distribution was observed for J3 (117 ±

19 nm) and J4 (126 ± 23 nm) nanoparticles. This could be ascribed to the large

diameter and size dispersion of Au nanoparticles prepared by Frens method using low

citrate concentration, [13] which often yield nanoparticles with non-spherical shape. In

J4 samples, prepared using large Au nanoparticles,[13] it was observed that some Au

nanoparticles were assembled to more than one MS nanoparticle, yielding aggregated

structures. This fact could be justified by the close packing of the MS nanoparticles

onto the colloidosome surface, allowing the interaction of several thiol-modified

mesoporous colloids with the same large and non-spherical Au nanoparticle. It should

be highlighted that sample J2 (104 ± 17 nm) showed narrow distribution of size and

maximum yield of Au-MS anisotropic nanoparticles with 1:1 ratio, and accordingly this

Janus colloid was selected for further experiments. A representative TEM image

corresponding to J2 nanoparticles is shown in Figure 2. Other different TEM images of

J2 nanoparticles are also shown in Figure 3S in Supporting Information.


327

Figure 2. Representative TEM image of J2 nanoparticles

The pore morphologies of the MS and Janus Au-MS nanoparticles in the J2

nanoparticles were determined by nitrogen adsorption/desorption surface analysis

(BET isotherms and BJH pore size distributions). Figure 3A illustrates the corresponding

nitrogen adsorption/desorption isotherms and the pore size distributions for the

starting MS material and for the corresponding J2 Janus nanoparticles. Both

nanomaterials showed type IV isotherms typical of mesoporous supports.

The absence of hysteresis loops for MS nanoparticles suggested that all pores

are highly accessible. On the contrary, the small hysteresis loops observed at high

relative pressure values in the Janus nanoparticle isotherm suggested that some pores

were partially blocked most likely due to the toposelective silanization with (3-

mercaptopropyl)trimethoxysilane and the attachment of the Au nanoparticles. In fact

the attachment of Au nanoparticles on one face of the siliceous matrix reduced the

BET specific surface area from 1037 m²/g in the starting MS to 820 m²/g in J2. Yet the

average pore size of the MS support (ca. 2.5 nm) was unchanged after formation of the

anisotropic colloid (see inset of Figure 3A).


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Figure 3. A) Nitrogen adsorption (closed)/desorption (open) isotherms for MS (,) and

J2 nanoparticles (,). Inset: pore size distribution of MS () and J2 () nanoparticles.

B) Normalized visible spectra of Au (a) and J2 (b) nanoparticles

UV-vis measurements in aqueous solutions were performed to provide insight

into the surface characteristics of the Janus nanoparticles (Figure 3B). The starting Au

colloids showed a single absorption band at 522 nm, distinctive of the surface plasmon

resonance of spherically-shaped nanospheres with about 20 nm diameter, whereas the

attachment of the Au nanoparticles to the thiol-modified MS nanoparticles leads to a

broadening and red-shift of the plasmon band in the J2 spectra. In addition, a broad


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shoulder band from 585 nm to 630 nm was also observed in the spectrum of J2 which

can be tentatively attributed to the attachment of more than one Au nanospheres on

the surface of the MS nanoparticles in some Janus particles, causing a coupling of the

plasma modes due to metal particle-particle interactions. [14]

J2 colloid was further employed as nanosized hardware for the assembly of a

self-controlled nanomachine able to release a cargo compound by an enzyme-based

pH-mediated mechanism. In a first step J2 nanoparticles were loaded with tris(2,2´-

bipyridyl)ruthenium(II) chloride ([Ru(bpy)3]2+) as model dye for monitoring cargo

delivery.[7c] Thereafter, an excess of the alkyl amino derivative 3-(2-

aminoethylamino)propyltrimethoxysilane (the pH-responsive molecular gate) was

anchored on the external surface of the mesoporous face to yield the J2Ru

nanomaterial.[7a] It should be mentioned that Janus nanoparticles exhibited good

water solubility, but its stability in aqueous solutions was reduced after modification

with 3-(2-aminoethylamino)propyltrimethoxysilane.

Thermal analysis of the MS, J2 and J2Ru nanomaterials was accomplished, the

TG/DTG curves being displayed in Figures 4S-A,B (see Supporting Information). The

unmodified MS nanoparticles exhibited a slight weight loss at temperatures up to ca.

70ºC, which was attributed to the thermodesorption of physically adsorbed water

molecules from the silica surface. At higher temperatures no weight loss was apparent

showing relatively flat TG/DTG curves.[15] Janus nanoparticles showed a more

noticeable weight loss at temperatures up to ca. 140ºC suggesting that higher amount

of water molecules were adsorbed on the Au nanoparticles surface. The anisotropic

nanoparticle exhibited a second thermal-induced transformation, with maximum rate

of weight loss at approximately 275ºC, which could be associated with the

decomposition of the bonded thiol ligands but also with the condensation of the

ligand's side silanols with one another or with surface silanols.

In comparison with J2, the weight loss profile for J2Ru exhibited a lower

decrease at T < 120ºC, suggesting lower amount of water molecules physically

adsorbed on this nanomaterial. This fact could be justified by the azeotropic treatment

of the Janus nanoparticles before adsorption of the [Ru(bpy)3]2+ complex and


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silanization with 3-(2-aminoethylamino)propyltrimethoxysilane, as well as to the

introduction of the long chain aminosilane groups on the MS face. The weight loss at

higher temperatures can be divided into two regions. The first region, with maximum

rate of weight loss at 245ºC, could be attributed to decomposition/condensation of

the bounded ligands. The second thermal-induced process, which showed a maximum

rate of transformation at about 330ºC, could be associated with the decomposition of

the [Ru(bpy)3]2+ complex, which is thermally stable up to 320 °C.[16] The loading of the

[Ru(bpy)3]Cl2 dye in the J2Ru material amounted to 62 mg/g J2Ru in weight, as

determined by the TG curve. Moreover the content of the anchored amine-based gate

amounted to 49 mg/ g J2Ru in weight.

Figure 5S-A in Supporting Information shows an FT-IR spectrum of the MS, J2 and J2Ru

nanoparticles. MS nanoparticles showed the characteristics IR absorption bands of

siliceous materials at 456 cm-1 attributed to the vibration of the Si-O bonds, a shoulder

at 576 cm-1 ascribed to cyclic Si-O-Si structures, at 803 cm-1 attributed to SiO4

tetrahedrons, at 946 cm-1 attributed to the Si-OH groups, and a band at 1080 cm-1

with a shoulder at 1200 cm-1 ascribed to the bond stretching vibrations of Si-O-Si.[17]

The broad band at 3700-3000 cm-1 can be ascribed to the O-H bonding vibration of

adsorbed water and SiO-H groups, and the band at 1629 cm-1 is attributed to the

deformation vibration of the HO-H bond in water molecules.

Spectrum of J2 nanoparticles presented the antisymmetric and symmetric stretching

vibrations of the CH2 groups at 2933 cm-1 and 2856 cm-1, confirming the modification

of the MS nanomaterial with (3-mercaptopropyl)trimethoxysilane. The absence of the

characteristic bands of S-H vibration in the range of 2500-2600 cm−1 suggested that the

thiol groups were chemisorpted to the Au nanoparticle surface. Loading of J2 with

[Ru(bpy)3]2+ and derivatization with 3-(2-aminoethylamino)propyltrimethoxysilane

was further confirmed by the bands at 2946 cm-1 and 2859 cm-1 ([C-H]), 1633 cm-1 ([N-

H]), 1568 cm-1 ([C-C] + [C=C-H]), 1480 cm-1 ([C-C] + [C=C-H]) and 1319 cm-1 ([NC-H]) in the

J2Ru spectrum.[18] Another important characteristic is the drop in intensity of the band

around 950 cm-1, which is attributed to the vibration of the silanol groups, confirming

the modification of the mesoporous silica surface with the amine-bearing silane.[7a]


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The powder X-ray diffraction patterns of J2 and J2Ru are shown in Figure 5S-B

(see Supporting Information). The low angle diffractogram of J2 shows four

distinguishable peaks at 2.59º, 4.30º, 4.93º and 6.46º which correspond to (100),

(110), (200) and (210) of MCM-41 with d-spacing values of 3.40 nm, 2.05 nm, 1.79 nm

and 1.37 nm, respectively. This pattern suggested perfect long-range order in this

mesoporous nanomaterial. Additionally, from these data and the pore value diameter

obtained from nitrogen adsorption isotherms, an a0 cell parameter of 3.93 nm and a

pore wall thickness of 1.40 nm can be calculated. Moreover, the diffraction pattern of

J2 at high angle showed three peaks at 38.34º, 44.50º and 81.21º, corresponding to

the (111), (200) and (222) Bragg reflections for cubic gold nanocrystals,[19] confirming

the Janus architecture previously observed by TEM. J2Ru sample showed similar X-ray

diffraction pattern, suggesting that the loading process with the dye and the further

functionalization with amine groups did not damage neither the mesoporous MCM-41

type structure nor of the gold face of the Janus colloid.[7c]

In order to prepare the self-controlled enzyme-powered nanodevice for cargo

delivery, the Au face of the Jan2Ru nanoparticles were functionalized with the enzyme

urease. For preparing an enzyme form suitable to be anchored on the metal surface,

urease was covalently modified with 3,3´-dithiobis(sulfosuccinimidylpropionate)

(DTSSP). After reduction of the dithiol linkage with NaBH4, the modified enzyme

solution was dialyzed and finally incubated with J2Ru to yield the J2Ru-U nanomaterial.

All these processes were performed in 50 mM sodium phosphate buffer, pH 7.0, at

4ºC.

The ability of J2Ru-U nanoparticle to deliver the [Ru(bpy)3]2+ dye under urease

control was tested in 5 mM sodium acetate buffer at pH 4.0 and 5.0. It is well know

that the pH-activity profile of urease has a bell-shaped behavior with maximum at pH

7.0.[20] However, more than 66% of the 3-(2-aminoethylamino) propyltrimethoxysilane

grafting moieties on the MS surface are unprotonated at this pH value [7a] then the

gate-like ensembles was partially open. To avoid this undesirable effect, we used

buffer solution of pH 5.0 where urease retains about 60% of its maximum catalytic


332

activity,[20] and not appreciable release of the dye from J2Ru-U was experimentally

observed in the control experiments.

In a typical release assay, 10 mg of J2Ru-U were suspended in 10 mL of buffer

solution containing 180 mM urea, and shaken over time at 25ºC. Aliquots were taken

at scheduled times, centrifuged and the absorbance was measured at 454 nm. As

control experiments, J2Ru-U samples were suspended in similar buffer solution

without urea. In order to ensure saturating conditions for substrate in the enzyme

catalyzed reaction, concentration of urea about 100-fold larger than KM = 1.3 mM was

employed.

Figure 4. Kinetics of dye release from J2Ru-U in 5 mM sodium acetate buffer pH 5.0 in

the absence () and the presence of 180 mM urea at t = 0 () or t = 5 h ().

Figure 4 shows the time-course of [Ru(bpy)3]2+ release from the pores of the

J2Ru-U nanoparticles. The concentration of dye delivered to the solution increased

progressively with the time of incubation for the mixture containing urea at pH 5.0,

reaching plateau value of about 19 µM after 5 h of incubation. On the contrary, no

significant increase in the absorbance measured at 454 nm was observed for control

solution containing no urea.


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Urease catalyzed the transformation of urea to CO2 and NH3 yielding a

progressive increase in the pH value of the incubation solutions, as it was qualitatively

demonstrated by adding bromothymol blue to control assays. As it was previously

reported, the 3-(2-aminoethylamino)propyltrimethoxysilane moieties located on the

MS surface became gradually deprotonated when the solution pH reached alkaline

values.[7a] Consequently, the nanoscopic molecular gates located at the pore outlets

trended to be opened favoring the release of the dye.

To provide insight into this enzyme-controlled dye release mechanism, J2Ru-U

nanomaterials were incubated in buffer solution without urea under the conditions

described above. Urea (180 mM final solution) was then added after 5 h of incubation,

the results being shown in Figure 4. No appreciable dye release was observed during

the first 5 h of incubation but the further increase in the absorbance at 454 nm

revealed that the dye was successfully delivered to the solution after adding urea.

CONCLUSION

In summary, we described the preparation of Janus-type nanoparticles having

Au and MS opposite faces and have used them for the design of an integrated nano-

device containing on the same nanoparticle a gating systems and an effector molecule

for the stimulus-responsible delivery of a cargo. In this particular case, as a proof-of-

concept the release was mediated by the on-demand catalytic decomposition of urea

by the enzyme urease, which was immobilized on the Au face, whereas the gated

ensemble consisted of a pH-responsive system which was anchored on the pore

outlets of the MS phase. In spite of the many reports dealing with the preparation of

silica-based anisotropic colloids, approaches to produce MS-based Janus nanoparticles

and their evaluation as on-demand control release systems are scarce to our

knowledge. We believe that the possibility of incorporate different effector molecules

and gated ensembles on this Janus-type integrated nanoarchitecture, which

constitutes a proof-of-concept, open new routes for the development of novel

biologically-inspired smart nanomachines for drug delivery and sensing applications

and research is this line is currently being performed by us.


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1. Preparation of MS nanoparticles:[8]

Cetyltrimethylammonium bromide (3.0 g) was dissolved in 1.44 L of water

under sonication. NaOH solution (2.0 mol/L, 10.5 mL) was then added and the

temperature of the mixture was adjusted to 80 ºC. Tetraethoxysilane (15.0 mL) was

added dropwise to the surfactant solution within 5 min under vigorous magnetic

stirring. The mixture was allowed to react for 2 h. The resulting white solid was

filtered, washed with water and methanol, and then dried in desiccator. The solid was

finally calcinated at 550 ºC for 5 h.

2. Preparation of 3.7 nm Au nanoparticles:[12]

An ice-cold, freshly prepared 0.1 M NaBH4 solution (600 µL) was quickly added

to 20 mL aqueous solution containing 300 µM HAuCl4 and 300 µM trisodium citrate

under continuous stirring. The mixture was stirred for 30 min, and 100-fold diluted

with 300 µM trisodium citrate before J1 preparation.

3. Preparation of 18 nm, 29 nm and 41 nm Au nanoparticles:[13]

Freshly prepared 3 µM HAuCl4 solutions (50 µL) were heated to boiling. Au

nanoparticles of 18 nm, 29 nm and 41 nm were synthesized by adding 750 µL, 500 µL

and 300 µL of 3.9 µM trisodium citrate solution, respectively. The mixtures were

heated for 10 min, cooled to room temperature and finally raised to 50 mL with

ultrapure water.

Preparation of Janus Au-MS nanoparticles: Janus nanoparticles were synthesized by

adapting two methods previously reported in literature.[10] MS nanoparticles (200 mg)

were dispersed homogeneously in 10 mL of 1.0 µM of CTAB in 6.7% ethanol aqueous

solution. The mixture was heated at 75ºC and then 1 g of paraffin wax was added.


335

When the paraffin wax was melted, the mixture was vigorously mixed at 25000 rpm

during 2 min using an Ultra-Turrax T-10 homogenizer (IKA, Germany). The resulting

emulsion was further mixed during 5 min at 4000 rpm and 75 ºC, using the same

apparatus. The resulting Pickering emulsion was then cooled to room temperature,

mixed with 10 mL methanol and treated with 200 µL of (3-

mercaptopropyl)trimethoxysilane. After 3 h under magnetic stirring, the silanized

emulsion was filtered, three-times washed with methanol and further dispersed in 400

mL of the corresponding 3 µM Au nanoparticles aqueous solutions. The mixture was

stirred overnight, then filtered and exhaustively washed with ultrapure water. The

solid was suspended in ethanol, centrifuged and washed two-times with ethanol and

three-times with chloroform. The Janus nanoparticles were finally dried and kept in

dissecator until use.

4. Preparation of J2Ru-U:

To synthesize J2Ru, 100 mg of J2 and 60 mg of tris(2,2´-bipyridyl)ruthenium(II)

chloride hexahydrate were suspended in 10 mL of anhydrous acetonitrile inside a

round-bottom flask connected to a Dean-Stark trap under Ar atmosphere.[7a] The

suspension was refluxed at 110 °C in azeotropic distillation, collecting about 4 mL in

the trap in order to remove the adsorbed water. The mixture was stirred for 24 h at

room temperature to load the dye into the MS face pores. Afterward, an excess of 3-

(2-aminoethylamino)propyltrimethoxysilane (500 µL) was added, and the suspension

was stirred for 5.5 h. Finally, the orange solid (J2Ru) was filtered off, washed two times

with 30 mL of CH3CN, and dried at 70 °C for 12 h.

To synthesize J2Ru-U, 4.0 mg of urease and 4.0 mg of 3,3´-dithiobis-

(sulfosuccinimidylpropionate) were dissolved in 5.0 mL of 50 mM sodium phosphate

buffer, pH 7.0, and stirred during 2 h at 4 ºC. Afterward, 200 µL of 100 mM NaBH4

solution were added, and the mixture was stirred at 4 ºC for 30 min. The solution was

exhaustively dialyzed vs. 50 mM sodium phosphate buffer, pH 7.0 using Amicon Ultra-

05 centrifugal filter units with Ultracel-10 membranes (Millipore, USA), and finally

concentrated to about 10 mg/mL concentration. The modified enzyme solution was


336

then added to 50 mL sodium phosphate buffer, pH 7.0, containing 50 mg of J2Ru, and

stirred at 4ºC overnight. The resulting solid (J2Ru-U) was finally isolated by

centrifugation, washed several times with sodium phosphate buffer, pH 7.0, dried and

kept in refrigerator until use.

5. Characterization:

Transmission electron microscopy (TEM) measurements were performed with

JEOL JEM-3000 F and JEOL JEM-2100 microscopes. The morphology of the

colloidosomes was characterized using high resolution field emission scanning electron

microscopy (FE-SEM) with a JEOL JSM-6335F electron microscope (JEOL Ltd., Japan).

FT-IR spectra were acquired with a Perkin-Elmer instrument. Spectrophotometric

measurements were performed using an Agilent 8453 UV/VIS spectrophotometer

(Hewlett Packard, USA). Power X-ray diffraction (XRD) was performed with an X'Pert

MRD diffractometer (PANanalytical B.V., The Netherlands). Nitrogen

adsorption/desorption isotherms and pore size distributions were determined with an

ASAP 2020 Physisorption Analyzer (Micromeritics, USA). Thermal analysis was

performed with a TA Instruments SDT-Q600 apparatus (USA). FT-IR spectra were

acquired with a Nicolet Nexus 670/870 spectrometer (Thermo Fisher Scientific Inc.,

USA).

ACKNOWLEDGEMENTS




14564-C04-01, MAT2012-38429-C04-01 and Comunidad de Madrid S2009/PPQ-1642,


337

programme AVANSENS is gratefully acknowledged. The Generalitat Valencia (project

PROMETEO/2009/016) is also acknowledged.

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Mondragón, E. Aznar, M.D. Marcos, R. Martínez-Máñez, F. Sancenón, J. Soto, J.M.

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[9] Y. Feng, J. He, H. Wang, Y.Y. Tay, H. Sun, L. Zhu, H. Chen, J. Am. Chem. Soc.

2012, 134, 2004; T. Chen, G. Chen, S. Xing, T. Wu, H. Chen, Chem. Mat. 2010, 22, 3826.

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Enzyme-controlled sensing-actuating nanomachine based on Janus Au-

mesoporous silica nanoparticles

Reynaldo Villalonga,*[a,b] Paula Díez,[a] Alfredo Sánchez,[a] Elena Aznar,[c,d] Ramón

Martínez-Máñez,[c,d] José M. Pingarrón*[a,b]

[a]Department of Analytical Chemistry, Complutense University of Madrid, 28040-

Madrid, Spain, Fax: (+34) 913944329. E-mail: [email protected],

[email protected]. [b]IMDEA Nanoscience, Cantoblanco Universitary City, 28049-

Madrid, Spain. [c]Departamento de Química and CIBER de Bioingeniería, Biomateriales

y Nanomedicina (CIBER-BBN), Universidad Politécnica de Valencia, Camino de Vera s/n,

E-46022, Valencia, Spain, Fax: (+34) 963879349. E-mail: [email protected],

[email protected] [d]Instituto de Reconocimiento Molecular y Desarrollo Tecnológico

(IDM), Centro Mixto Universidad Politécnica de Valencia-Universidad de Valencia,

Spain.


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Figure 1S. TEM images and distribution of sizes of MS nanoparticles.

Figure 2S. Distribution of sizes of J1 (A), J2 (B), J3 (C) and J4 (D) nanoparticl


341

Figure 3S. TEM images of J2 nanoparticles.

Figure 4S. A) TG and B) DTG analysis for MS (a), J2 (b) and J2Ru (c) nanoparticles.


342

Figure 5S. A) FT-IR analysis for MS (a), J2 (b) and J2Ru (c) nanoparticles. B) X-ray

diffraction of J2 (a) and J2Ru (b) nanoparticles.

7.12

Journal of the American Chemical Society 136 (25) (2014) 9116–9123

Towards the design of smart delivery systems controlled by

integrated enzyme-based biocomputing ensembles

Journal of the American Chemical Society 136 (2014) 9116–9123 7. PUBLICACIONES

343

TOWARDS THE DESIGN OF SMART DELIVERY SYSTEMS CONTROLLED BY

INTEGRATED ENZYME-BASED BIOCOMPUTING ENSEMBLES

Paula Díez,1 Alfredo Sánchez,1 María Gamella,1 Paloma Martínez-Ruíz,2 Elena Aznar,3,4

Cristina de la Torre,3,4 José R. Murguía,3,4 Ramón Martínez-Máñez,3,4,* Reynaldo

Villalonga,1,5,* José M. Pingarrón1,5,*

1Departments of Analytical Chemistry and 2Organic Chemistry I, Faculty of Chemistry,

Complutense University of Madrid, 28040-Madrid, Spain. 3Instituto de Reconocimiento

Molecular y Desarrollo Tecnológico (IDM), Centro Mixto Universidad Politécnica de

Valencia-Universidad de Valencia, Spain. 4Departamento de Química y CIBER de

Bioingeniería, Biomateriales y Nanomedicina (CIBER-BBN), Universidad Politécnica de

Valencia, Camino de Vera s/n, 46022, Valencia, Spain. 5IMDEA Nanoscience,

Cantoblanco Universitary City, 28049-Madrid, Spain.


344

ABSTRACT

We report herein the design of a smart delivery system in which cargo delivery

from capped mesoporous silica (MS) nanoparticles is controlled by an integrated

enzyme-based “control unit”. The system consists of Janus-type nanoparticles having

Au and MS opposite faces, which were properly functionalized with a pH-responsive β-

cyclodextrin based supramolecular nanovalve on the silica mesoporous surface and

two effectors, glucose oxidase and esterase, immobilized on the Au face. The nano-

device behaves as an enzymatic logical OR operator which is selectively fuelled by the

presence of D-glucose and ethyl butyrate.

KEYWORDS: Janus nanoparticles, enzymes, mesoporous silica, logic gates, delivery

INTRODUCTION

Evolution in bio-molecular chemistry combined with nanotechnology has

recently resulted in the design of biologically based systems with innovative functions.1

A significant issue in this field has been the development of new “intelligent” devices

using nanoscopic structures and a variety of biomolecules, fuelling areas such as bio-

engineering, bio-sensing and drug delivery. In this context, the design of smart delivery

systems able to release entrapped guests in a controlled fashion has received great

attention in recent years.2 The advent of nanotechnology has provided a large variety

of novel nanomaterials which have found application in this area.3 Mesoporous silica

(MS) supports have been widely explored as promising alternatives for delivery uses

due to their large specific volume, large loading capacity, low cost and low toxicity.4 An

interesting characteristic of these MS nanoparticles is that they can be rationally

functionalized with molecular or supramolecular ensembles on their external surface

to develop gated-nanocarriers showing “zero delivery”,5 which can further release their

cargo in response to target physical (such as light, temperature or magnetic fields),6


345

chemical (such as pH-changes, redox-active molecules or selected anions)7 and bio-

chemical (such as enzymes, antibodies or DNA) stimuli.8

Scheme 1. Schematic representation of “smart” delivery systems containing an

attached control unit that regulates the delivery activity of the gated material.

However, in most of these systems the effector (i.e. the agent that regulates the

delivery activity) is external to the delivery nanoparticle, a fact that somehow limits the

design of “smart” nanodevices for delivery applications. A way to overcome this

restriction is the design of nanosupports in which the gating system and the effector

molecule are integrated in the same nanodevice.9 In this approach one can envision

the design of a “control unit” attached to the gated nanoparticle in which one or

several agents that regulates the delivery activity are placed (see Scheme 1). The role

of this unit is to handle the chemical information (input) of the environment and to

transform it (via the use of the effectors) in new chemicals that control the state of the

gate (open of closed). Such strategy can also promote effective protection to the

effectors, such as enzymes, due to immobilization which ensures their functionality

when they reached the target place.

Moreover, the possibility of use a combination of different effectors in the

“control unit” opens new perspectives in the development of complex systems that can

generate specific results (delivery or not, control of the delivery kinetics, etc) via logical

operations based on different chemical inputs. However, as far as we know, such

engineered release systems have not been described.

Gated

material

Control

unitOutput

Input

Input


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Figure 1. Performance of the Janus-based nanodevice S3. The “control unit” (Au face) is

functionalized with two effectors (enzymes) which control cargo delivery from the silica

mesoporous face via interpretation of different chemical inputs (D-glucose, ethyl

butyrate). Overall the system functions as an enzymatic logical OR operator.

Chemically speaking, a general strategy towards the integration of the gating

system and effectors in the same nanodevice may be the conjugation of two very

different nanoparticles having different surfaces and well-defined and specific

functionalization chemistries. As a suitable approach for this goal, Janus nanoparticles

are especially appropriate.10 Following this general concept, and as proof-of-concept,

we report herein the design of Janus MS-gold nanoparticles 9 in which the gated

ensemble in the MS face is combined with one or more effectors placed in the gold

side of the Janus support. The proposed design involves the immobilization of two

enzymes, glucose oxidase (EC 1.1.3.4) and esterase (EC 3.1.1.1), on the gold face as

effectors, with the cargo release governed by an enzymatic logical OR operator (see

Figure 1). Moreover, the MS face of the Janus nanoparticles was used as nanocontainer

for cargo loading and equipped with a pH-responsive β-cyclodextrin-based

supramolecular nanovalve.7f,11

The Au side is expected to act as the “control unit” in which the enzyme effectors

would “interpret“ the presence of specific chemical inputs (enzyme substrates) and

would direct the operation (cargo delivery) of the system. In particular, we envisioned

that the gated mesoporous nanodevice will show “zero-release”, yet selectively will


347

open in the presence of either D-glucose, ethyl butyrate or a combination of both

substrates through enzyme-catalyzed substrate transformations which will lead to a

reduction of the pH and, consequently, to the opening of the β-cyclodextrin-gatted

nanovalves. The overall output (cargo delivery) will function as a Boolean logic OR

gate.12


To assemble the integrated nanomachine, Janus Au-MS nanoparticles were first

prepared as previously described.9,13 The synthetic procedure is based on the

manipulation of the Au-ligand-MS interface through a mask-protecting assisted site-

selective modification approach. Briefly, MS nanoparticles (S0, average diameter: 97 ±

15 nm) were synthesized by alkaline hydrolysis of tetraethyl orthosilicate as inorganic

precursor in the presence of the cationic surfactant cetyltrimethylammonium bromide

as porogen species. The subsequent removal of the surfactant by calcination in air at

high temperature resulted in the starting mesoporous inorganic support. These

nanoparticles were then partially confined at the interface of a Pickering emulsion

using paraffin wax as oil phase. The exposed nanoparticle surface was further modified

with (3-mercaptopropyl) trimethoxysilane on which Au nanoparticles (average

diameter: 20 ± 2 nm) were then attached, yielding stable anisotropic colloids (S1,

average diameter: 104 ± 17 nm, yield 85%) after dissolving the paraffin wax in CHCl3.

The MS nanoparticles in the anisotropic colloids were then loaded with

Ru(bipy)32+, which was used as dye for monitoring the release process, and the external

surface of the siliceous face was further modified with 3-iodopropyltrimethoxysilane.

Benzimidazole moieties were attached to the anchored 3-iodopropyl residues through

a nucleophilic substitution reaction, yielding a solid functionalized with 1-propyl-1-H-

benzimidazole groups (S2). These nanoparticles were then gated with a pH-sensitive

supramolecular nanovalve by stirring the colloid with β-cyclodextrin moieties in water

at pH 7.5 for 24 h that resulted in the formation of inclusion complexes between the

benzimidazole groups and the β-cyclodextrins.7f,11 Finally, esterase and glucose oxidase,


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previously modified with 3,3´-dithiobis(sulfosuccinimidylpropionate), were selectively

and covalently immobilized on the Au face of the S2 colloid by incubation in 50 mM

sodium phosphate buffer, pH 7.5, at 4ºC, yielding the final nanodevice S3.

Au nanoparticles were selected as scaffold for the assembly of the effector

system, instead of using direct enzymes immobilization on the MS surface. This choice

was relied on the large surface area of Au nanoparticles which allows high enzyme

loadings with the immobilization method emploeyd. In addition, the relative low Au

reactivity allows the metal nanoparticles surface to remain unaltered during the

further assembly of the gated system in the MS face, thus avoiding chemical protection

steps that should be employed to directly link the enzymes to the MS face.

All supports were characterized by standard methods (see Supporting

Information). TEM analysis conf irmed the mesoporous morphology of the silica

nanoparticles as well as the presence of the Au nanoparticles in the Janus colloids

(Figure 1). Powder X-ray diffraction patterns of the final nano-device S3 and the

intermediate S0, S1 and S2 supports are shown in Figure SI-1. All samples showed a

well-defined low-angle reflexion around 2.6º which corresponds to a hexagonal

ordered array indexed as a (100) Bragg peak, suggesting a MCM41-like mesoporous

order in these materials which was not affected by the different chemical modifications

and dye loading processes. In addition, the diffraction patterns of S1, S2 and S3

samples at high angle also showed the cubic gold characteristic (111), (200), (220) and

(311) diffraction peaks,14 confirming the Janus Au-MS architecture observed by TEM.

FT-IR spectrum of the S0 and S1 nanoparticles were similar to those reported for MS

and Janus Au-MS nanoparticles, whereas the characteristics IR absorption bands of

benzimidazole were clearly observed in the spectrum of S2 (Figure SI-2 in Supporting

Information).15 On the other hand, the presence of β-cyclodextrin moieties in S3 was

confirmed by the broad band at ca. 1060 cm-1, which is characteristic for this cyclic

oligosaccharide. The presence of the immobilized enzymes in S3 was confirmed by the

band at 1642 cm-1 which can be ascribed to the amide I absorption band of proteins.

Moreover, thermogravimetric (Figure SI-3 in Supporting Information) and elemental

analysis studies (Table SI-1 in Supporting Information) on S1, S2 and S3 revealed a

content of the Ru(bpy)32+ dye and anchored benzimidazole in S2 of 15 µmol/g and 160


349

µmol/g, respectively. In addition, the amount of glucose oxidase and esterase

immobilized on S3 was estimated as of 2.5 U/mg and 4.8 U/mg, respectively.

The N2 adsorption-desorption isotherms of the starting calcined MS

nanoparticles (S0) and the Janus colloid (S1) showed an adsorption step at

intermediate P/P0 values (0.1–0.3). The application of the BET model for these solids

resulted in values for the total specific surface area of 1037 and 819 m2 g-1,

respectively (see Figure SI-4 and Table SI-2). In contrast the N2 adsorption-desorption

isotherms for the prepared dye loaded material (S2) and the final capped support (S3)

are typical of mesoporous systems with filled mesopores and a significant decrease in

the N2 volume adsorbed and surface area was observed (i.e. 51.3 and 18.2 m2 g-1 for S2

and S3, respectively, see Supporting Information). 7f

The capacity of the S3 nano-device to deliver the cargo in aqueous solution was

further tested. In a typical release assay, 10 mg of S3 were suspended in 4 mL of 20

mM Na2SO4 solution at pH 7.5 and shaken over time at 25ºC. Aliquots were taken at

scheduled times, centrifuged to remove the nanoparticles, and the absorbance at 454

nm of the released Ru(bpy)32+ was measured. After 1 h incubation, the enzyme

substrates D-glucose and ethyl butyrate, used as input signals, were added to the

mixtures at a concentration of 40 µM.

Figure 2 shows the time-course of Ru(bpy)32+ release from the pores of S3

nanoparticles in the presence and absence of substrates. In the absence of D-glucose

or ethyl butyrate solid S3 is tightly capped and shows a negligible release of Ru(bpy)32+

(ca. 1.2 ± 0.5 µM Ru(bpy)32+, see curve a). In contrast, the presence of either ethyl

butyrate (curve b), D-glucose (curve c) or a mixture of both (curve d) results in the

opening of the pores and the subsequent release of the cargo. Overall, delivery from

nanoparticles S3 is triggered by the presence of ethyl butyrate and D-glucose via the

“interpretation“ of these chemical inputs by the glucose oxidase and esterase enzymes

(effectors) in the “control unit” that resulted in the dethreading of the inclusion

complex between benzimidazole moieties and β-cyclodextrin. In particular glucose

oxidase catalyze the oxidation of D-glucose yielding H2O2 and D-glucono-1,5-lactone

which hydrolyses in water to gluconic acid (pKa = 3.6). Moreover, ethyl butyrate is


350

catalytically hydrolysed by esterase enzyme to ethanol and butyric acid (pKa = 4.82).

Both catalytic reactions result in a reduction in the pH of the incubation solutions

causing the protonation of benzimidazole moieties (pKa = 5.55)16 on the mesoporous

silica face of S3, the dethreading of the inclusion complex and cargo delivery.

Figure 2. Kinetics of dye release from S3 in 20 mM Na2SO4, pH 7.5 in the absence (a)

and the presence of 40 µM ethyl butyrate (b), D-glucose (c) and ethyl butyrate + D-

glucose (d). Substrates were added after 1 h of incubation.

As illustrates in Figure 2, the esterase-mediated process showed faster delivery

kinetics than those mediated by glucose oxidase, reaching plateau values of about 36.1

± 0.8 µM and 33.5 ± 0.6 µM Ru(bpy)32+

after 2 h and 2.5 h of addition of the

corresponding substrates, respectively. This difference in the released dye is tentatively

ascribed to the lower amount of glucose oxidase immobilized on Au nanoparticles. In

addition, glucose oxidase has a more acidic optimum pH range (pH 4-7, with maximum

at pH 5.5) than that for esterase (pH 6-9, with maximum at pH 8.0),17 which results in a

lower catalytic activity for glucose oxidase, especially at the beginning of the reaction.


351



glucose (d) without (closed circles) and with 200 µM urea (open circles). Substrates

were added after 1 h of incubation.

Moreover, to demonstrate that the opening mechanism was due to the

enzyme-mediated reduction of pH it was confirmed that in the experiments in which

ethyl butyrate or D-glucose were added to aqueous suspensions of S3 the pH of the

incubating media changed from 8.0 to ca. 5.0 after 6 h. In addition, in order to

demonstrate that it is the presence of the glucose oxidase and esterase enzymes in the

Au “control unit” that governed cargo delivery, suspension of S3 at pH 7.5 were boiled

for 10 min to inactivate the enzymes, and further tested toward D-glucose and ethyl

butyrate. In this case no appreciable cargo delivery was observed from S3 after 24 h of

incubation.

To elucidate whether the release of Ru(bpy)32+ from the Janus colloid was

caused either by a local decrease of pH at the microenvironment of the nanoparticles

or by pH decrease in the bulk solution, parallel release experiments in buffered

solutions at pH 7.5 were performed. As it is exemplified in Figure SI-5 when D-glucose +

ethyl butyrate are used as triggers, the concentrations of Ru(bpy)32+ released at a given


352

time to the buffered solution were only slightly lower than those corresponding to the

non buffered medium. Measurements of the final pH values of the buffered solutions

confirmed that no changes were apparent upon the release experiments. These results

suggested that the enzyme-controlled cargo release from the Janus nanoparticles was

mainly provoked by a local acidification of the colloid microenvironment caused by the

biocatalytic transformation of the trigger enzyme substrates.

The experiments carried out can be summarised in a Boolean-like table (see

Figure 1) in which the observed output (delivery (1) or not (0) of the cargo from S3)

depends on the presence (1) or not (0) of the small molecules D-glucose and ethyl

butyrate. Thus whereas solid S3 displayed no release (input values 0,0; output value 0),

the presence of the enzymes’ substrates as input values (0,1; 1,0; 1,1) induced the

delivery of the entrapped guest (output value 1). In terms of delivery, S3 behaves as an

enzymatic logical OR operator.

In order to expand the possibilities of the enzyme-controlled nanomachine, a

new S4 nanodevice was synthesized by co-immobilizing also the enzyme urease (Ec

3.5.1.5) on the Janus Au nanoparticles face. Urease catalyzed hydrolysis of urea to CO2

and NH3 which resulted in a progressive increase in the pH value of the incubation

solutions, this acting as a RESET operator for the pH-mediated release process. Figure 3

(curves b and d, open circles) clearly shows as a noticeable decrease in the amount of

released dye occurred when urea was also added as trigger. This effect can be

attributed to the partial neutralization of the acidic medium by the ammonia produced

through the urease-catalyzed reaction, then switching off the opening of the

supramolecular nanovalves and thus controlling the dye release. A possible reduction

of the glucose oxidase and esterase activity by local increase of pH should be also

considered as a possible mechanism for this urease-based switch-off action.

Figure 4 shows the effect of the time at which urea was added to the

incubation media on the kinetics of dye release from solid S4 triggered with D-glucose.

The concentration of released Ru(bpy)32+ increased as the time at which urea was

introduced into the media was longer, demonstrating that the urease-based RESET


353

operator, and thus the amount of dye released, can be operated through a time-

controlled scheme.


and the presence of 40 µM D-glucose without (f) and with addition of urea at 200 µM

final concentration at different times (b-e). Substrates were added after 1 h of

incubation.

One goal of this study was to demonstrate that Janus enzyme-controlled

capped MS can be used for in-cell controlled delivery applications. Therefore, after the

in vitro characterization of the solid S3 (vide ante), similar nanoparticles were tested in

further ex vivo assays. In particular for these experiments nanoparticles as S3 but

loaded with the cytotoxic doxorubicin (Doxo) were prepared (solid S5, see Supporting

Information). The amount of Doxo loaded on the Janus colloid was estimated

spectrophotometrically as 0.56 µmol per gram of nanoparticles. The S5 nanomachine

showed similar patterns for in vitro Doxo release than those filled with Ru(bpy)32+ upon

addition of ethyl butyrate or D-glucose + ethyl butyrate, but lower release kinetics in

the presence of D-glucose (Figure 5).


354

Figure 5. Kinetics of Doxo release from S5 in 20 mM Na2SO4, pH 7.5 in the absence (a)


glucose (d). Substrates were added after 1 h of incubation.

S5 nanomachine retained full functional activity after one month of storage at

4ºC. In addition, the operational stability of the S5 nanomachine was tested by

incubation at 37 ºC in reconstituted human serum and further quantification of Doxo

released after 1 h addition of D-glucose + ethyl butyrate as triggers (see Supporting

Information for details). As it is illustrated in Figure SI-6 (Supporting Information), the

integrated nanomachine lost release activity progressively with time according to

biphasic inactivation kinetics. However, the S5 solid retained over 40% of the initial

functional activity after a week of incubation, suggesting its potential use for long term

ex vivo assays.

The solid S5 was ex vivo analysed in HeLa cells under the premise that S5 could

be internalised by the cells and would remain closed until glucose or ethyl butyrate are

added. The action of enzymes in the Janus nanoparticle on these molecules would

induce a local pH reduction that is expected to result in an intracellular Doxo release,

which would induce cell death. In a typical experiment HeLa cells were incubated for

40 minutes with a suspension of 100 µg/mL of S5 in PBS supplemented with 10 % fetal


355

bovine serum. After this, cells were washed in order to remove un-internalised

nanoparticles and were further incubated alone or in the presence of ethyl butyrate

(input A), glucose (input B) or a mixture of both (see Experimental section for details).

Delivery of Doxo from S5 in the presence of the different inputs was first studied by

confocal microscopy by tracking Doxo-associated fluorescence. Moreover in these

experiments the cell nuclei were stained with Hoechst 33342.

Figure 6A shows representative images of phase contrast, Doxo, Hoescht and

combined for HeLa cells first loaded with S5 and then untreated (-/-) or treated with

ethyl butyrate (+/-), glucose (-/+) or with a mixture of both (+/+). Internalization of S5

in HeLa cells (see -/-) did not produce, in the absence of further input, a significant

Doxo release, thus demonstrating that not unspecific cargo release occurred in a large

extent as a consequence of the acidification in the endosomes. In contrast when cells

were incubated simultaneously with ethyl butyrate and glucose a clear dispersed Doxo

fluorescence was found indicating cargo delivery from S5. Figure 6C shows a

quantitation using flow cytometry of the Doxo fluorescence intensity for HeLa cells

incubated with S5 (-/-) and further treated with ethyl butyrate (+/-), glucose (-/+) or

with a mixture of both (+/+). Whereas emission was low in the absence of inputs, an

enhanced Doxo fluorescence was detected upon exposure to ethyl butyrate or glucose.

Moreover a synergistic enhanced Doxo emission was observed when both ethyl

butyrate and glucose treatments were combined simultaneously.

These observations correlate with cell viability studies determined after 24

hours of the corresponding treatment using WST-1 assays (see Figure 6B). Indeed, it

was confirmed that HeLa cells treated at 100 μg/mL of S5 in the presence of glucose

and ethyl butyrate showed apoptotic cell death. More specifically, around 50% of the

cells were dead 24h after the addition of S5, whereas values of ca. 40% and 15% of

dead cells were found when only glucose or ethyl butyrate were added, respectively. In

contrast, the HeLa cells treated with only S5 remained unaffected.

In summary, we have demonstrated that it is possible to design nanodevices in

which the gating mechanisms and different effector ensembles can be integrated in a

unique system. In particular we have reported here the preparation of Janus-type


356

nanoparticles having Au and MS opposite faces, which were properly functionalized

with a pH-responsive β-cyclodextrin based supramolecular nanovalve on the silica

mesoporous surface and two effectors, glucose oxidase and esterase, immobilized on

the Au face. The nano-device behaves as an enzymatic logical OR operator which is

fuelled by the presence of D-glucose and ethyl butyrate. This enzyme logic system was

also coupled to a urease-based RESET operator to switch-off the opening of the

supramolecular nanovalves and control the extension of dye delivery upon addition of

urea. To our knowledge, this is the first report dealing with the use of anisotropic

colloids for the design of smart nanodevices for on-command release controlled by

biochemical logic operations. The smart nanomachine controlled by the logical OR

operator and loaded with an anti-cancer drug, was successfully tested toward HeLa

cancer cells. The possibility of using a large variety of different gating nanovalves on

the silica mesoporous face combined with a potential number of enzyme-based

effectors on the Au surface makes this approach appealing and opens a wide range of

new possibilities for the development of novel smart delivery systems controlled by

enzyme-based biocomputing ensembles.


357

Figure 6. Internalization and release of cargo in HeLa cells A) controlled release of

Doxorubicin (Doxo) loaded S5 nanoparticles in HeLa cells. Culture were incubated with

100 μg/ml of S5 and in presence of different inputs and examined for Doxo by confocal

microscopy. Representative images at 24 h form phase contrast (PhC), Doxo (DOX),

Hoescht (HOE) and combined (Merge) are shown. B) Cell viability test of 150 μg/ml

concentration of S5 and glucose and/or ethyl butyrate at 24 h in HeLa cells using WST-1

assay and C) quantification of Doxo fluorescence intensity by flow cytometry in cells

under different conditions. Ethyl butyrate treatment (input A), glucose treatment (input

B).


358

METHODS

1. Preparation of MS nanoparticles (S0).18

Cetyltrimethylammonium bromide (3.0 g) was dissolved in 1.44 L of water

under sonication. NaOH solution (2.0 mol/L, 10.5 mL) was then added and the

temperature of the mixture was adjusted to 80 ºC. Tetraethoxysilane (15.0 mL) was

added dropwise to the surfactant solution within 5 min under vigorous magnetic

stirring. The mixture was allowed to react for 2 h. The resulting white solid was filtered,

washed with water and methanol, and then dried in desiccator. Finally, the solid was

calcined at 550 ºC for 5 h to remove the organic template.

2. Preparation of 20 nm Au nanoparticles.19

Freshly prepared 3 µM HAuCl4 solution (50 mL) was heated to boiling. Then, 20

nm gold nanoparticles were synthesized by adding 750 µL of a 3.9 µM trisodium citrate

solution. The mixture was heated for 10 min, cooled to room temperature and finally

raised to 50 mL with ultrapure water.

3. Preparation of Janus Au-MS nanoparticles (S1).9,13

MS nanoparticles (200 mg) were dispersed homogeneously in 10 mL of 1.0 µM

of cetyltrimethylammonium bromide in 6.7% ethanol aqueous solution. The mixture

was heated at 75ºC and then 1 g of paraffin wax was added. When the paraffin wax

was melted, the mixture was vigorously stirred at 25000 rpm for 10 min using an Ultra-

Turrax T-10 homogenizer (IKA, Germany). The resulting emulsion was further stirred for

1 h at 4000 rpm and 75 ºC, using a magnetic stirrer. The resulting Pickering emulsion

was then cooled to room temperature, mixed with 10 mL methanol and treated with

200 µL of (3-mercaptopropyl)trimethoxysilane. After 3 h under magnetic stirring, the

silanized emulsion was filtered, three-times washed with methanol and further

dispersed in 400 mL of the corresponding 3 µM Au nanoparticles aqueous solutions.


359

The mixture was stirred overnight, then filtered and exhaustively washed with

ultrapure water. The solid was suspended in ethanol, centrifuged and washed two-

times with ethanol and three-times with chloroform. The Janus nanoparticles were

finally dried and kept in desiccators until use.

4. Preparation of S2.

To synthesize S2, 400 mg of S1 and 250 mg (33 µmol) of tris(2,2′-bipyridyl)

dichlororuthenium(II) hexahydrate were suspended in 17 mL of anhydrous acetonitrile

inside a round-bottom flask connected to a Dean-Stark trap under Ar atmosphere. The

suspension was heated at 110 °C and about 7 mL of solvent were distilled and collected

in the trap to remove the adsorbed water. After this step, the mixture was stirred for

24 h at room temperature to load the dye into the MS face pores. Afterward, an excess

of 3-iodopropyltrimethoxysilane (200 µL, 1 mmol) was added and the suspension was

stirred for 24 hours.7f The final solid was filtered off, washed two times with 5 mL of

acetonitrile and dried at 60 ºC overnight. To attach the benzimidazole moieties to the

MS surface, 400 mg of the resulting solid were suspended in a 40 mL of a saturated

solution of benzimidazole in toluene at 80ºC and containing triethylamine

(benzimidazol and triethylamine in a 1:3 proportion). The suspension was refluxed and

stirred for 72 hours. The resulting solid was filtered off, washed with 40 mL of

acetonitrile and dried at 70 ºC overnight.

5. Preparation of S3, S4 and S5.

To prepare the gated solid S3, 400 mg of S2 were suspended in 100 mL of a β-

cyclodextrin solution (1.6 mg/ml) in 50 mM sodium phosphate buffer, pH 7.5.7f The

suspension was stirred for 12 hours at room temperature. The capped solid was

centrifuged, washed with water at pH 7.5 and dried. In a second step, to functionalize

these nanoparticles with the enzymes, 2.0 mg of enzyme (esterase or glucose oxidase)

and 2.0 mg of 3,3´-dithiobis(sulfosuccinimidylpropionate) were dissolved in 2.0 mL of


360

50 mM sodium phosphate buffer, pH 7.0, and stirred for 2 h at 4 ºC.9 Afterward, 200 µL

of 100 mM NaBH4 solution were added, and the mixture was stirred at 4 ºC for 30 min.

The solution was exhaustively dialyzed vs 50 mM sodium phosphate buffer, pH 7.0

using Amicon Ultra-05 centrifugal filter units with Ultracel-10 membranes (Millipore,

USA), and finally concentrated to about 10 mg/mL concentration. The modified

enzyme solutions were then added to 20 mL of 50 mM sodium phosphate buffer, pH

7.5, containing 20 mg of the β-cyclodextrin capped solid, and stirred at 4ºC overnight.

The resulting solid (S3) was finally isolated by centrifugation, washed several times

with a cold solution of 50 mM sodium phosphate buffer, pH 7.5, dried and kept in

refrigerator until use. Solid S4 was prepared through a similar protocol, but co-

immobilizing a third enzyme, urease, on the Au nanoparticles surface. For ex vivo cell

experiments, nanoparticles as S3 but loaded with the cytotoxic doxorubicin (Doxo)

were also prepared (solid S5, see Supporting Information).

6. Cell Culture Conditions.

HeLa human cervix adenocarcinoma cells were purchased from the German

Resource Centre for Biological Materials (DSMZ) and were grown in DMEM

supplemented with 10% FBS. Cells were maintained at 37 ºC in an atmosphere of 5%

carbon dioxide and 95% air and underwent passage twice a week.

7. WST-1 Cell Viability Assay.

HeLa cells were seeded in a 24-well plate at a density of 2 ·104 cells/well in a

1000 uL of DMEM and were incubated 24 hours in a CO2 incubator at 37 ºC. Then,

DMEM were replaced for PBS with 10 % of Fetal Bovine Serum and solid S5 in DMSO

were added to cells in sextuplicate at final concentrations of 150 μg/ml. After 40

minutes, cells were washed with PBS and were incubated for 23 hours in different

conditions. DMEM with 10% FBS, DMEM with 10% FBS and ethyl butyrate, PBS with

10% FBS or PBS with 10 % FBS and ethyl butyrate. After this, 35 μL of WST-1 were


361

added to each well and were incubated for 1.5 hours. Before reading the plate was

shaken for one minute to ensure homogeneous distribution of colour. Then the

absorbance was measured at a wavelength of 450 nm in a VICTOR X5 PerkinElmer.

Results are expressed as a promedium of the results of six independent experiments

obtaining similar results.

8. Live Confocal Microscopy.

HeLa cells were seeded in a 24 mm Ø glass coverslips in six-well plates at a

seeding density of 1,8 ·105 cells /well. After 24 hours, culture medium were replaced

for PBS with 10% fetal bovine serum (FBS) and cells were treated with a suspension of

solid S5 for 40 minutes at a final concentration of 100 μg/ml. Then the medium was

change for different solutions (DMEM with 10%FBS with or without ethyl butyrate, or

PBS with 10% FBS with or without ethyl butyrate). After 18 hours coverslips were

washed twice to eliminate compounds and, were visualized under a confocal

microscope employing Leica TCS SP2 AOBS (Leica Microsystems Heidelberg GmbH,

Mannheim, Germany) inverted laser scanning confocal microscope using oil objectives:

63X Plan-Apochromat-Lambda Blue 1.4 N.A. The images were acquired with an

excitation wavelength of 405 for Hoescht and 480 nm for Doxorubicin. Two-

dimensional pseudo colour images (255 colour levels) were gathered with a size of

1024x1024 pixels and Airy 1 pinhole diameter. All confocal images were acquired using

the same settings and the distribution of fluorescence was analysed using the Image J

Software. Three fields of each condition in two independent experiments were

performed obtaining similar results.

ASSOCIATED CONTENT

Supporting Information

Experimental details and nanomaterials characterization. This material is available free

of charge via the Internet at http://pubs.acs.org.


362

AUTHOR INFORMATION

Corresponding Author

[email protected], [email protected], [email protected].

Notes

The authors declare no competing financial interests.

ACKNOWLEDGMENT




38429-C04-01 and Comunidad de Madrid S2009/PPQ-1642, programme AVANSENS is

gratefully acknowledged. The Generalitat Valencia (project PROMETEO/2009/016) is

also acknowledged.

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J.; Bein, J. Angew. Chem., Int. Ed., 2009, 48, 3092-3095; (c) Park, C.; Kim, H.; Kim, S.;

Kim, C. J. Am. Chem. Soc., 2009, 131, 16614-16615.


Chem. Eur. J. 2013, 19, 7889-7894.

(10) (a) Perro, A.; Reculusa, S.; Ravaine, S.; Bourgeat-Lami, E.; Duguet, E.J. J. Mater.

Chem. 2005, 15, 3745-3760; (b) Jiang, S.; Chen, Q.; Tripathy, M.; Luijten, E.; Schweizer,

K.S.; Granick, S. Adv. Mater. 2010, 22, 1060-1071.


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(11) Du, L.; Liao, S.; Khatib, H.A.; Stoddart, J.F.; Zink, J.I. J. Am. Chem. Soc. 2009, 131,

15136-15142

(12) (a) Tokarev, I.; Gopishetty, V.; Zhou, J.; Pita, M.; Motornov, M.; Katz, E.; Minko, S.

ACS Appl. Mater. Interfaces 2009, 1, 532-536;.(b) Du, L.; Liao, S.; Khatib, H.A.; Stoddart,

J.F.; Zink, J.I. J. Am. Chem. Soc. 2009, 131, 15136-15142.

(13) Sánchez, A.; Díez, P.; Martínez-Ruíz, P.; Villalonga, R.; Pingarrón, J.M. Electrochem.

Commun. 2013, 30, 51-54.

(14) Leff, D.V.; Brandt, L.; Heath, J.R. Langmuir 1996, 12, 4723-4730.

(15) Mohan, S.; Sundaraganesan, N.; Mink, J. Spectrochim. Acta Mol. Biomol. Spectros.

1991, 47, 1111-1115.

(16) Jerez, G.; Kaufman, G.; Prystai, M.; Schenkeveld, S.; Donkor, K.K. J. Sep. Sci. 2009,

32, 1087-1095.

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Heymann, E. Eur. J. Biochem. 1979, 95, 519-525.

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(19) Frens, G. Nature 1973, 241, 20-22.


365


Towards the design of smart delivery systems controlled by

integrated enzyme-based biocomputing ensembles

Paula Díez,1 Alfredo Sánchez,1 María Gamella,1 Paloma Martínez-Ruíz,2 Elena Aznar,3,4

Cristina de la Torre,3,4 José R. Murguía,3,4 Ramón Martínez-Máñez,3,4,* Reynaldo

Villalonga,1,5,* José M. Pingarrón1,5,*

1Departments of Analytical Chemistry and 2Organic Chemistry I, Faculty of Chemistry,

Complutense University of Madrid, 28040-Madrid, Spain. 3Instituto de Reconocimiento

Molecular y Desarrollo Tecnológico (IDM), Centro Mixto Universidad Politécnica de

Valencia-Universidad de Valencia, Spain. 4Departamento de Química y CIBER de

Bioingeniería, Biomateriales y Nanomedicina (CIBER-BBN), Universidad Politécnica de

Valencia, Camino de Vera s/n, 46022, Valencia, Spain. 5IMDEA Nanoscience,

Cantoblanco Universitary City, 28049-Madrid, Spain.


Chemicals

Glucose oxidase from A. Niger, pig liver esterase, tetraethoxysilane,

cetyltrimethylammonium bromide, tris(2,2′-bipyridyl)dichlororuthenium(II)

hexahydrate, hydrogen tetrachloroaurate, 3-iodopropyltrimethoxysilane,

benzimidazole, triethylamine, (3-mercaptopropyl)trimethoxysilane, D-glucose, ethyl

butyrate, paraffin wax, 3,3´-dithiobis(sulfosuccinimidylpropionate), DMSO and PBS for

cell culture were purchased from Sigma-Aldrich. For cell culture experiments,

Dulbecco's Modified Eagle's medium (DMEM) with L-glutamine, pyruvate, human

serum (H4522) and Fetal Bovine Serum (FBS), trypan blue solution (0.4%) cell culture


366

grade, trypsin and the cell proliferation reagent WST-1were purchased from Roche

Applied Science. Solvents were provided by Scharlau. All other reagents were of

analytical grade.

General Techniques


JEOL JEM-2100 microscope. Spectrophotometric measurements were performed using

an Agilent 8453 UV/VIS spectrophotometer (Hewlett Packard, USA). Powder X-ray

diffraction (XRD) was performed with an X'Pert MRD diffractometer (PANanalytical

B.V., The Netherlands). Nitrogen adsorption/desorption isotherms and pore size

distributions were determined with an ASAP 2020 Physisorption Analyzer

(Micromeritics, USA). Thermal analysis was performed with a TA Instruments SDT-

Q600 apparatus (USA). FT-IR spectra were acquired with a Nicolet Nexus 670/870

spectrometer (Thermo Fisher Scientific Inc., USA).

Preparation of S5.

To prepare the solid S5, 400 mg of S1 were first suspended in 17 mL of

anhydrous acetonitrile inside a round-bottom flask and then treated with an excess of

3-iodopropyltrimethoxysilane (200 µL, 1 mmol). The suspension was stirred for 24

hours and the resulted solid was filtered off, washed two times with 5 mL of

acetonitrile and dried at 60 ºC overnight. To attach the benzimidazole moieties to the

MS surface, 400 mg of the resulting solid were suspended in a 40 mL of a saturated

solution of benzimidazole in toluene at 60ºC and containing triethylamine

(benzimidazol and triethylamine in a 1:3 proportion). The suspension was refluxed and

stirred for 72 hours. The resulting solid was filtered off, washed with 40 mL of

acetonitrile and dried at 60 ºC overnight. 50 mg of this solid was then suspended in 5

mL of 50 mM sodium phosphate buffer, pH 7.5 and then 40 µmol Doxorubicin (Doxo)

were added. The mixture was stirred for 12 h and then 12 mL of a β-cyclodextrin


367

solution (16 mg/ml) in 50 mM sodium phosphate buffer, pH 7.5 was added. The

suspension was stirred for 12 hours at room temperature, and the resulting capped

solid was centrifuged, washed with water at pH 7.5 and dried. In a second step, to

functionalize these nanoparticles with the enzymes, 5.0 mg of enzyme (esterase or

glucose oxidase) and 5.0 mg of 3,3´-dithiobis(sulfosuccinimidylpropionate) were

dissolved in 2.0 mL of 50 mM sodium phosphate buffer, pH 7.0, and stirred for 2 h at 4

ºC. Afterward, 200 µL of 100 mM NaBH4 solution were added, and the mixture was

stirred at 4 ºC for 30 min. The solution was exhaustively dialyzed vs 50 mM sodium

phosphate buffer, pH 7.0 using Amicon Ultra-05 centrifugal filter units with Ultracel-10

membranes (Millipore, USA), and finally concentrated to about 10 mg/mL

concentration. The modified enzyme solutions were then added to 20 mL of 50 mM

sodium phosphate buffer, pH 7.5, containing 50 mg of the β-cyclodextrin capped solid,

and stirred at 4ºC overnight. The resulting Doxo-loaded solid (S5) was finally isolated

by centrifugation, washed several times with a cold solution of 50 mM sodium

phosphate buffer, pH 7.5, centrifuged and kept wet in refrigerator until use. The

amount of Doxo entrapped within nanoparticles was calculated by the difference

between the total amount used to prepare nanoparticles and the amount of

doxorubicin present in the aqueous phase after capping with β-cyclodextrin, by

measuring the absorbance at 495 nm.1

Release and stability studies using S5

In vitro release studies for S5 were performed as described for S3.

Storage stability studies were performed by incubation of 4 mg/mL dispersion

S5 in 50 mM sodium phosphate buffer, pH 7.5 at 4ºC. Samples were collected at

scheduled times and assayed for enzyme-mediated on-command release of Doxo after

1 h addition of 40 µM ethyl butyrate + D-glucose. Operational stability of S5 was

assayed by incubation of 4 mg/mL dispersion S5 in reconstituted human serum at

37ºC. Samples were collected at scheduled times and assayed for enzyme-mediated


368

on-command release of Doxo after 1 h addition of 40 µM ethyl butyrate + D-glucose.

The amount of released drug was monitored by visible spectrophotometry at 495 nm.1

Cytofluorometry studies using S5

HeLa cells were seeded at 18 x 103 cells per well in a 24-well plate. After 24 h,

DMEM were replaced for PBS with 10 % of Fetal Bovine Serum and solid S5 in DMSO

were added to cells at final concentrations of 150 μg·ml-1. After 40 minutes, cells were

washed with PBS and were incubated for 23 hours in the different conditions (DMEM

with 10% FBS, DMEM with 10% FBS and ethyl butyrate, PBS with 10% FBS or PBS with

10 % FBS and ethyl butyrate). After 24 hours media were eliminated by vacuum and

plate were washed once with PBS. Cells were detached with Trypsin/EDTA solution,

centrifuged and finally resuspended in 1 ml of DMEM with 10% FBS. Quantification of

Doxorubicin fluorescence in the cells was performed with WinMDI program, version

2.0 in a FC500 MCL Flow Cytometer (Beckman-Coulter, CA, USA). Three independent

experiments containing quadruplicates were performed with similar results.

Materials Characterization

Solids S0, S1, S2 and S3 were characterized using standard procedures. Figure

SI-1 shows powder X-ray diffraction patterns in the 1.5 < 2θ < 7 range (SI-1A) and in

the 30 < 2θ < 80 range (SI-1B) for the calcined nanoparticulated MS support S0, the

Janus nanoparticles S1 and the S2 and S3 functionalized materials. The low angle

diffractogram of S0 shows the MCM-41 characteristic (100) reflection. This typical

reflexion is preserved in the Janus nanoparticles (S1). Moreover, the diffraction pattern

of S1 shows the cubic gold characteristic (111), (200), (220) and (311) diffraction peaks,

confirming the presence of gold nanocrystals and the Janus Au-MS architecture.

Finally, S2 and S3 samples display similar X-ray diffraction pattern than S1, suggesting

that the loading process with the dye, the further anchoring of the pH-responsive β-

cyclodextrin based nanovalve and the enzymes immobilization did not modify the

characteristics of the Janus colloid.


369

Figure SI-1: Powder X-ray diffraction of nanoparticles S0, S1, S2 and S3 at low (A) and

high (B) angles.

Figure SI-2 shows an FT-IR spectrum of all synthesized solids (S0 to S3). MS

nanoparticles (S0) showed the characteristics IR absorption bands of siliceous

materials: at 456 cm-1 attributed to the vibration of the Si-O bonds, a shoulder at 576

cm-1 ascribed to cyclic Si-O-Si structures, at 803 cm-1 attributed to SiO4 tetrahedrons, at

946 cm-1 attributed to the Si-OH groups, and a band at 1080 cm-1 with a shoulder at

1200 cm-1 ascribed to the bond stretching vibrations of Si-O-Si. Moreover, the broad

band at 3700-3000 cm-1 can be attributed to the O-H bonding vibration of adsorbed

water and SiO-H groups, and the band at 1629 cm-1 is ascribed to the deformation

vibration of the HO-H bond in water molecules. The spectrum of the Janus

nanoparticles S1 shows antisymmetric and symmetric stretching vibrations of the CH2

groups at 2933 cm-1 and 2856 cm-1, confirming the modification of the MS

nanomaterial with (3-mercaptopropyl)trimethoxysilane. The spectrum of S2 showed

the presence of characteristics IR absorption bands of benzimidazole groups.

Moreover, the formation of the inclusion complex with β-cyclodextrin moieties in S3

was confirmed by the presence of a broad band at ca. 1060 cm-1, which is

characteristic for this cyclic oligosaccharide. Finally, the successful immobilization of

enzymes in S3 was confirmed by the band at 1642 cm-1 which can be ascribed to the

amide I absorption band of proteins.


370

Figure SI-2: FT-IR analysis for the nanoparticles S0, S1, S2 and S3.

Figure SI-3: Thermogravimetric analysis for S0, S1, S2 and S3.


371

Table SI-1. Elemental analysis for S0, S1, S2 and S3.

Solid %C %H %N %S

S0 6.01 1.91 0.07 0.60

S1 6.07 2.00 0.08 2.41

S2 27.58 2.99 6.46 1.05

S3 48.46 4.09 15.04 1.15

The N2 adsorption-desorption isotherms of the MS material S0 and the Janus

nanoparticles S1 show two sharp adsorption steps. A first step at intermediate P/P0

value (0.2-0.4) typical of these solids (see Figure SI-3) is related to the nitrogen

condensation inside the mesopores by capillarity. The absence of a hysteresis loop in

this interval and the narrow BJH pore distribution suggest the existence of uniform

cylindrical mesopores. The application of the BET model resulted in a value for the

total specific surface of 1037 m2/g for S0 and 819 m2/g for S1. In addition to this

adsorption step associated to the micelle generated mesopores, a second feature

appears in the isotherm at a high relative pressure (P/P0 > 0.75). This second step

corresponds to the filling of the large voids among the particles that must be

considered as a textural-like porosity. In this case, curves show a characteristic H1

hysteresis loop and a wide pore size distribution.


372

Figure SI-4. Nitrogen adsorption (closed)/desorption (open) isotherms for S0 to S3

nanoparticles.

Table SI-2. BET specific surface values, pore volumes and pore sizes calculated from the

N2 adsorption-desorption isotherms for selected materials.

SBET

(m2g-1)

BJH pore

(P/P0 < 0.4)a

(nm)

Total pore

volumeb

(cm3g-1)

S0 1037 2.54 1.15

S1 819 2.51 1.28

S2 51.3 - 0.30

S3 18.2 - 0.14

a Pore size estimated by using the BJH model applied on the adsorption branch of the isotherm, for P/P0 < 0.4, which can be associated to the surfactant generated mesopores.

b Total pore volume according to the BJH model.


373

The N2 adsorption-desorption isotherms of S2 and S3 are typical of mesoporous

systems with filled mesopores (see Figure SI-3) and a significant decrease in the N2

volume adsorbed and surface area (51.3 m2/g for S2 and 18.2 m2/g for S3) is observed.

The most relevant feature is the absence of a sharp step at low-medium relative

pressure (0.1 < P/P0 < 0.4). In fact, these solids showed flat curves in that region when

compared to those of S0 or S1 materials, which indicates a significant pore blocking

and the subsequent absence of appreciable mesoporosity. Additionally, these

materials also show a N2 adsorption at high relative pressure similar to the calcined

MCM-41, confirming that the textural porosity is preserved. BET specific surface

values, pore volumes and pore sizes calculated from the N2 adsorption-desorption

isotherms for all the synthesized materials are listed in Table SI-1.

Figure SI-5. Kinetics of dye release from S3 in 20 mM Na2SO4, pH 7.5 in the absence (a)

and the presence of 40 µM D-glucose + ethyl butyrate (b). Kinetics of release from S3 in

50 mM sodium phosphate buffer, pH 7.5 in the presence of 40 µM D-glucose + ethyl

butyrate (c). Substrates were added after 1 h of incubation.


374

Figure SI-6. Influence of time of incubation in reconstituted human serum at 37ºC on

the release activity of S5 upon addition of 40 µM D-glucose + ethyl butyrate.

1. Moktan, S.; Perkins, E.; Kratz, F.; Raucher, D. Mol. Cancer Ther. 2012, 11, 1547-

1556.

8 DISCUSIÓN

INTEGRADORA

8. DISCUSIÓN INTEGRADORA

375

8.1. NANOMATERIALES FUNCIONALIZADOS Y NANOHÍBRIDOS PARA EL ENSAMBLAJE

DE SENSORES Y BIOSENSORES ELECTROQUÍMICOS.

En este apartado se discuten los resultados relacionados con la construcción de

electrodos nanoestructurados con diferentes nanomateriales funcionalizados y

nanohíbridos, y su empleo como interfaz de transducción para la construcción de

plataformas electroquímicas de (bio)sensorización.

Los nanomateriales preparados se clasifican en:

- Redes de nanopartículas de oro polifuncionalizadas

- Nanotubos de carbono derivatizados mediante interacciones no covalentes

- Derivados de dendrímeros y dendrones de poliamidoamina

8.1.1. Biosensores basados en redes de nanopartículas de oro polifuncionalizadas.

En los artículos incluidos en este apartado se describen los resultados

relacionados con la construcción de electrodos nanoestructurados con redes

tridimensionales de nanopartículas de oro electropolimerizadas, los cuales fueron

posteriormente empleados en la construcción de biosensores electroquímicos

empleando enzimas y anticuerpos como biorreceptores.

La estrategia propuesta se basa en la preparación de nanopartículas de oro

funcionalizadas con tres tipos de ligandos tiolados diferentes, los cuales desempeñan

funciones distintas en el nanomaterial resultante:

- Ácido 2-mercaptoetanosulfónico: Ligando encargado de conferir hidrofilia,

solubilidad y estabilidad coloidal a las nanopartículas de oro.

- p-Aminotiofenol: Ligando aromático capaz de facilitar la polimerización de las

nanopartículas mediante la formación de enlaces tipo bis-anilina.

- Tercer ligando: Encargado de facilitar la inmovilización de la molécula

biorreceptora.


376

En la Figura 8.1 se muestra una representación esquemática de las

nanopartículas de oro polifuncionalizadas.

Figura 8.1. Representación esquemática de las nanopartículas de oro

polifuncionalizadas.

Un aspecto fundamental en el diseño de estos nanomateriales es la relación

molar en que estos ligandos deberán aparecer sobre la superficie de la nanopartícula,

en aras de lograr un equilibrio entre las capacidades de polimerización e inmovilización

de biomoléculas, y las propiedades de solubilidad adecuadas para su empleo en medio

acuoso. Para los trabajos aquí presentados se determinó como condiciones óptimas de

síntesis el uso de los ligandos p-aminotiofenol:ácido 2-mercaptoetanosulfónico:ligando

de inmovilización en una relación molar de 1:3,4:1,1. Como regla general, en nuestras

investigaciones hemos determinado la necesidad de emplear un exceso molar de

aproximadamente 1,5-1,7 del ácido 2-mercaptoetanosulfónico con respecto a la

cantidad de los otros ligandos tiolados utilizados, en aras de obtener nanopartículas

fácilmente dispersables en medio acuoso.


377

Las nanopartículas de oro polifuncionalizadas fueron preparadas mediante una

modificación del método propuesto por Liu y colaboradores para la síntesis de

nanopartículas metálicas recubiertas con ciclodextrinas [Liu et al., 2000]. Este

procedimiento se basa en la reducción in situ de HAuCl4 con NaBH4 en DMSO, en

presencia de los ligandos tiolados a ser empleados como agentes de modificación

superficial de las nanopartículas. Esta metodología permite la preparación de

nanopartículas polifuncionalizadas de color rojo oscuro, solubles en agua y de

diámetro muy pequeño, debido a la rápida quimisorción de los ligandos tiolados sobre

la superficie de los coloides recién formados. Estos procesos de quimiosorción in situ

limitan el crecimiento posterior de estos nanomateriales, produciendo nanopartículas

con un tamaño adecuado para la posterior formación de las redes

electropolimerizadas.

Este procedimiento de síntesis se diferencia del método descrito por el grupo

de Willner [Riskin et al., 2010; Frasconi et al., 2010] para la preparación de

nanopartículas similares en el uso de DMSO como medio de reacción. De esta forma se

favorece la mayor solubilidad de los ligandos orgánicos tiolados, y se evita la

descomposición espontánea del agente reductor en medio acuoso acidulado, lo cual

asegura un mejor rendimiento de síntesis.

Las nanopartículas fueron caracterizadas mediante diferentes métodos

fisicoquímicos y de microscopía. A modo de ejemplo, en la Figura 8.2 se muestran las

imágenes obtenidas mediante análisis por TEM de baja y alta resolución de las

nanopartículas sintetizadas en el trabajo publicado en Electrochimica Acta 56, 2011,

4672-4677. Como puede observarse las nanopartículas funcionalizadas poseen una

geometría esférica con un diámetro medio de 2,5 ± 0,4 nm. La geometría radial, el

tamaño pequeño y la baja polidispersión mostrada ponen de manifiesto que durante el

desarrollo sintético la reducción del metal y la fijación de los ligandos tiolados a la

superficie de las partículas de oro ocurren en único paso, como se comentó con

anterioridad [Liu, 2000].


378

Figura 8.2. Imágenes de las nanopartículas de oro polifuncionalizadas con residuos de

ácido 3-mercaptofenilborónico, obtenida mediante TEM a 200 kV (A) y 300 kV (B).

El análisis mediante HR-TEM muestra la presencia de las caras del cristal en las

nanopartículas sintetizadas. La medición de las distancias entre dichas caras reveló una

separación media de 2.4 Å, lo cual se corresponde con la distancia entre las caras 111

del oro [Grzelczak et al., 2008].

La presencia de los ligandos tiolados sobre la superficie de las nanopartículas

fue confirmada en todos los casos mediante espectroscopia FT-IR. En la Figura 8.3 se

muestra, a modo de ejemplo, el espectro FT-IR de las nanopartículas de oro

polifuncionalizadas con residuos de ácido 3-mercaptofenilborónico. Como puede

observarse en el espectro, a altas frecuencias (3410 - 3390 cm-1) aparecen bandas

características de las vibraciones fundamentales N-H y O-H, correspondientes a los

grupos amino e hidroxilo en los residuos de p-aminotiofenol y ácido 3-mercaptofenil

borónico, respectivamente. La presencia de estos ligandos aromáticos fue asimismo

confirmada por las bandas a 3005 cm-1 y 1585 cm-1, correspondientes a las vibraciones

C-H y C=C en sistemas aromáticos.


379

Figura 8.3. Espectro FT-IR de las nanopartículas de oro polifuncionalizadas con residuos

de ácido 3-mercaptofenilborónico.

Los residuos electropolimerizables de p-aminotiofenol fueron también

corroborados a partir de las bandas a 1650 cm-1 y 1344-1280 cm-1, las cuales están

asociadas a las vibraciones N-H y C-N, respectivamente. Las señales apreciadas en este

último rango de frecuencias pueden estar asimismo asociadas a la vibración

fundamental B-O en los residuos de ácido 3-mercaptofenil borónico. Por otra parte, la

presencia de los residuos de ácido 2-mercaptoetanosulfónico sobre la superficie de las

AuNPs fue determinada a partir de las vibraciones fundamentales asimétrica y

simétrica del grupo sulfonato a 1133 cm-1 y 1028 cm-1, respectivamente.

Las nanopartículas preparadas poseen la capacidad de ser polimerizadas por

métodos químicos y electroquímicos. En la Figura 8.4 se muestras imágenes

representativas de FE-SEM de las redes de nanopartículas de oro polifuncionalizadas y

con residuos de ácido 3-mercaptofenilborónico, obtenidas mediante polimerización

química y electroquímica en H2SO4. Para los experimentos de polimerización química

se empleó persulfato de amonio como agente iniciador.


380

Figura 8.4. Imágenes FE-SEM de redes de nanopartículas de oro polifuncionalizadas

con residuos de ácido 3-mercaptofenilborónico, obtenida mediante métodos químicos

(A) y electroquímicos (B) de polimerización.

En el presente trabajo se ha empleado el método electroquímico de

polimerización como estrategia para la preparación de las superficies

nanoestructuradas con las redes de nanopartículas de oro polifuncionalizadas, dada la

simplicidad de esta metodología, así como la alta densidad de nanopartículas y

homogeneidad de las redes preparadas.

Debe destacarse que este tipo de nanopartículas de oro polifuncionalizadas

permite el diseño de superficies electródicas tridimensionales, de alta

electroconductividad y área superficial electroquímica, enriquecidas con ligandos

específicos capaces de facilitar la inmovilización estable de biorreceptores mediante

interacciones covalentes y no covalentes. Por otra parte, se comprobó que este tipo de

nanomaterial electropolimerizado puede ser preparado sobre diferentes superficies

electródicas modificadas o no modificadas previamente con otros nanomateriales.

En este trabajo se describe la preparación de cuatro tipos diferentes de

nanopartículas de oro polifuncionalizadas, las cuales se diferencian en el tipo de

ligando a emplear para la inmovilización de biomoléculas:

- Ácido 3-mercaptofenilborónico (Electrochimica Acta 56, 2011, 4672-4677): Para la

inmovilización supramolecular de glicoproteínas mediante formación de ésteres


381

cíclicos entre los residuos de ácido borónico y los dioles 1,2-vecinales de las

cadenas de azúcar de la glicoproteina [Qi, 2010].

- Dendrones de poliamidoamina (Analyst 137, 2012, 342-348): Para la unión

covalente de proteínas mediante entrecruzamiento de los grupos amino primarios

en los dendrones con los grupos amino primarios de la proteína.

- Biotina (ChemElectroChem 1, 2014, 200-206): Para la inmovilización de proteínas

biotiniladas sobre la superficie de los electrodos, empleando estreptavidina como

receptor de afinidad puente.

- 1-Adamantano (ACS Applied Materials & Interfaces 4, 2012, 4312-4319): Para la

inmovilización supramolecular de neoglicoproteínas preparadas con β-

ciclodextrinas, mediante la formación de complejos de inclusión.

Las redes de nanopartículas de oro electropolimerizadas fueron preparadas

sobre diferentes superficies electródicas: i) electrodos de oro, y ii) electrodos de

carbono vitrificado recubiertos con nanotubos de carbono de pared simple. En la

Figura 8.5 se muestran imágenes de FE-SEM obtenidas para estos dos tipos de

nanoestructuras. Asimismo, en la Figura 8.6 se representa el proceso de síntesis de las

nanoparticulas polifuncionalizadas y su electropolimerización sobre electrodos de oro.

Figura 8.5. Imágenes FE-SEM de redes de nanopartículas de oro polifuncionalizadas y

(A) con residuos de ácido 3-mercaptofenilborónico crecidas sobre electrodos de oro, y

(B) con residuos de 1-adamantano y crecidas sobre electrodos de carbono vitrificado

recubiertos con nanotubos de carbono de pared simple.


382

Figura 8.6. Representación esquemática de los procesos de síntesis y

electropolimerización de las nanopartículas de oro polifuncionalizadas.

Las nanopartículas de oro fueron disueltas en solución de H2SO4 100 mM, y

posteriormente electropolimerizadas sobre las superficies de los electrodos. En el caso

de la electropolimerización sobre oro, la superficie de estos electrodos fue

previamente recubierta con una monocapa autoensamblada de p-aminotiofenol, con

el objetivo de asegurar la formación de enlaces bis-anilina entre los grupos tioanilina

sobre la superficie del electrodo y las nanopartículas [Frasconi, 2010].

Es importante resaltar que a diferencia del método de electropolimerización

para nanopartículas de oro publicado anteriormente [Riskin, 2010; Frasconi, 2010], en

nuestro trabajo se seleccionó un medio ácido para este proceso con el fin de garantizar

la electroconductividad de la polianilina formada, la cual en medio ácido se encuentra

en forma de sal de emeraldina (Figura 8.7).

Electropolimerización

“Redes poliméricas de AuNPs medianteenlaces tipo bis-anilina”


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Figura 8.7. Estructura del polímero conductor de polianilina.

En todos los casos la electropolimerización de las nanopartículas sobre la

superficie de los electrodos se realizó mediante un proceso de dos etapas. Esta

metodología comprendió una primera fase de formación de las redes entrecruzadas de

nanopartículas mediante la creación de enlaces del tipo bis-anilina, la cual se realizó

por voltamperometría cíclica entre -0.35 V and +0.85 V. Posteriormente, el electrodo

nanoestructurado se sometió a un proceso de crecimiento de las redes de

nanopartículas mediante la imposición de un potencial constante de +0.85 V durante 1

h.

A modo de ejemplo, en la Figura 8.8 se muestran los voltamperogramas cíclicos

obtenidos durante la primera etapa de electropolimerización de las nanopartículas de

oro polifuncionalizadas con residuos de ácido 3-mercaptofenilborónico. Como puede

observarse, el barrido continuo del potencial entre valores de -0.35 V to +0.85 V

originó la aparición y crecimiento de un pico anódico a 505 mV. De forma similar, se

pudo apreciar la ocurrencia de dos picos catódicos a 410 mV y 226 mV. Este

comportamiento voltamperométrico es característico para la formación

electroquímica de aductos diméricos y de baja masa molecular de anilina [Duić, 1995],

lo cual indica la formación continua de la matriz electropolimerizada de nanopartículas

sobre la superficie electródica a través de entrecruzamientos del tipo bis-anilina.


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Figura 8.8. Voltamperogramas cíclicos para el primer paso de electropolimerización de

las nanopartículas de oro polifuncionalizadas con residuos de ácido 3-

mercaptofenilborónico sobre electrodos de oro, en solución de H2SO4 0.1 M. Velocidad

de barrido: 100 mV/s.

Las superficies electródicas nanoestructuradas con los polímeros de

nanopartículas fueron empleados en la construcción de diversas plataformas

biosensoras. En el artículo publicado en Electrochimica Acta 56, 2011, 4672-4677 se

describe el uso de electrodos de oro funcionalizados con redes de nanopartículas

funcionalizadas con residuos de ácido 3-mercaptofenilborónico como soportes para la

inmovilización supramolecular de la enzima peroxidasa de rábano (Figura 8.9).


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Figura 8.9. Representación esquemática del proceso de ensamblado del electrodo

enzimático para la detección de H2O2.

La peroxidasa es una glicoenzima que presenta un alto contenido de

carbohidratos, equivalente al 20% del peso total de la proteína. Estos oligosacáridos se

encuentran enlazados a ocho sitios de la cadena polipeptídica: Asn13, Asn57, Asn158,

Asn186, Asn198, Asn214, Asn255, Asn267 [Wuhrer et al., 2005]. Como puede

observarse en la Figura 8.10, estos sitios de N-glicosidación se encuentran

espacialmente alejados del sitio activo de la enzima [Veitch et al., 2004].

Por este motivo, la inmovilización de la peroxidasa a través de sus cadenas de

glicosidación, mediada por la interacción de éstas con los residuos de ácido borónico

en las nanopartículas sintetizadas, se realiza de forma orientada, lo cual asegura una

alta actividad catalítica sobre la superficie del nanomaterial, así como de los electrodos

modificados con las redes electropolimerizadas de estas partículas.


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Figura 8.10. Estructura tridimensional de la peroxidasa de rábano.

La efectividad de esta estrategia de ensamblaje del electrodo enzimático se

demostró por las excelentes propiedades analíticas del biosensor construido para la

determinación amperométrica de H2O2. Este biosensor fue optimizado para trabajar a

un valor de potencial de 0.0 V, lo cual permite la eliminación de posibles interferencias.

Esto fue posibilitado por las elevadas propiedades electrocatalíticas que mostró la

interfaz nanoestructurada con las nanopartículas de oro frente a la transformación del

H2O2.

El biosensor enzimático mostró un intervalo lineal de respuesta analítica para

concentraciones de H2O2 comprendidas entre 5 µM y 1.1 mM, con una alta

sensibilidad de 498 µA/M cm2 y un límite de detección de 1.5 µM. Asimismo, este

biosensor mostró una alta estabilidad de almacenamiento y una elevada selectividad.

En el artículo publicado en Analyst 137, 2012, 342-348 se desmotró la viabilidad

de preparar nanopartículas de oro polifuncionalizadas, que presentaban dendrones de

poliamidoamina G-4 con núcleo de cisteamina como ligandos a emplear en la

inmovilización de biorreceptores. La estrategia de síntesis empleada permitió evitar el

crecimiento de las nanopartículas dentro de la estructura del dendrón, lo cual fue

comprobado por el tamaño de los coloides metálicos resultantes.


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Estas nanopartículas funcionalizadas con un ligando macromolecular fueron

electropolimerizadas sobre la superficie de electrodos de oro, los cuales fueron

posteriormente empleados como soporte para la inmovilización covalente de la

enzima tirosinasa mediante entrecruzamiento con glutaraldehido. El electrodo

enzimático obtenido se utilizó en la construcción de un biosensor amperométrico para

la determinación de catecol, sustancia de alto impacto ambiental clasificada como

disruptor endocrino [Bindhumol, 2003].

El biosensor ensamblado mostró un rango de respuesta lineal para

concentraciones de catecol entre 50 nM y 10 µM, con una sensibilidad de 1.94 A/M

cm2 y un límite de detección de 20 nM. Este biosensor mostró alta reproducibilidad en

la medida, y se demostró que su ensamblado se realizaba con un alto grado de

repetibilidad. El biosensor enzimático mostró una elevada estabilidad de almacenaje,

reteniendo el 96% de su actividad inicial después de dos semanas de almacenamiento

a 4 oC en condiciones de humedad.

En aras de demostrar la versatilidad de esta metodología de construcción de

superficies electródicas nanoestructuradas, se sintetizó una nanopartícula de oro

polifuncionalizada provista de residuos de biotina. La biotina (ácido 5-[(3aS,4S,6aR)-2-

oxohexahidro-1H-tieno[3,4-d]imidazol-4-il]pentanoico) es un importante cofactor

enzimático [Maurice et al., 2007] que forma asociaciones muy estables con las

proteínas avidina y estreptavidina, con valores de constante de disociación de sus

complejos del orden de 10−14 M [Korpela, 1984]. Esta elevada afinidad molecular ha

sido ampliamente utilizada para el diseño de sistemas de bioconjugación [Narain et al.,

2007], inmovilización biomolecular [Larsson et al., 2003], inmunodetección [Morgan et

al., 1992] y biosensorización [Rehák et al., 1994] basados en la formación de complejos

biotina-avidina y biotina-estreptavidina.

En nuestro trabajo publicado en ChemElectroChem 1, 2014, 200-206 se

describió la electropolimerización de nanopartículas de oro polifuncionalizadas con

residuos de biotina sobre la superficie de electrodos de oro, los cuales fueron

empleados como soporte nanoestructurado para el ensamblaje capa-a-capa de un

inmunosensor electroquímico para la detección de fibrinógeno humano, un


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importante marcador de enfermedades cardiovasculares [Lowe et al., 2004]. La

estrategia de ensamblaje utilizada se basó en la inmovilización de moléculas de

estreptavidina sobre la superficie del electrodo mediante interacciones de afinidad.

Estas moléculas de estreptavidina fueron empleadas como puente para el posterior

ensamblado de anticuerpos específicos para el fibrinógeno humano, derivatizados con

residuos de biotina.

El electrodo biofuncionalizado se empleó en la construcción de un

inmunosensor amperométrico, capaz de detectar fibrinógeno humano en un rango de

concentraciones entre 18 ng/mL y 2.208 µg/mL, con un límite de detección de 4 ng/mL

y un valor de IC50 de 177 ng/mL. Este inmunosensor fue notablemente estable,

conservando todo su potencial analítico después de 45 días de almacenamiento a 4 oC.

Finalmente, nuestra investigación se dirigió a comprobar la posibilidad de

electropolimerizar nanopartículas de oro sobre superficies electródicas previamente

modificadas con otro nanomaterial, así como emplear estas interfaces

nanoestructuradas para la inmovilización de biorreceptores mediante interacciones

supramoleculares del tipo “host-guest”. Para ello se prepararon nanopartículas de oro

polifuncionalizadas con residuos de adamantano, las cuales fueron

electropolimerizadas sobre electrodos de carbono vitrificado recubierto con

nanotubos de carbono de pared simple. Los resultados de este trabajo fueron

publicados en ACS Applied Materials & Interfaces 4, 2012, 4312-4319.

Para comprobar nuestra hipótesis de trabajo se sintetizó una neoglicoenzima

de xantina oxidasa modificada covalentemente con residuos de mono-6-etilendiamino-

6-deoxi-β-ciclodextrina. Esta reacción de derivatización se realizó empleando una

carbodiimida hidrosoluble como agente de activación de los grupos carboxilatos en la

superficie de la enzima, siguiendo protocolos previamente descubiertos en la

bibliografía para la síntesis de neoglicoenzimas con ciclodextrinas [Villalonga et al.,

2007].

Se comprobó la capacidad de la enzima artificialmente glicosidada de formar

complejos de inclusión estables con los residuos de 1-adamantano localizados sobre la


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superficie de las nanopartículas de oro electropolimerizadas sobre el electrodo, lo cual

permitió su inmovilización supramolecular sobre la interfaz nanoestructurada. El

electrodo enzimático resultante se empleó en la construcción de un biosensor

amperométrico para la determinación de xantina, el cual mostró un rango de

respuesta linear para concentraciones del analito comprendidas entre 50 nM y 9.5 µM,

con una alta sensibilidad de 2.47 A/M·cm2 y un bajo límite de detección de 40 nM.

Este biosensor mostró una alta estabilidad, lo cual puede justificarse por la alta

estabilidad estructural y funcional que generalmente es conferida a las enzimas tras su

modificación covalente con residuos de ciclodextrinas [Villalonga et al., 2003;

Fernández, 2003]. Por otra parte, se comprobó que las señales analíticas obtenidas con

este biosensor mostraron cierta afectación en presencia de los ácidos ascórbico y

úrico. Esta acción interferente fue notablemente minimizada mediante el

recubrimiento de la matriz electródica con una película delgada de Nafión.

8.1.2. Biosensores nanoestructurados con nanotubos de carbono modificados

mediante interacciones no covalentes.

En los artículos incluidos en este punto se describen los resultados obtenidos

en el diseño de superficies nanoestructuradas para la construcción de biosensores

electroquímicos, mediante el empleo de nanohíbridos preparados a partir de

nanotubos de carbono de pared simple.

Estos nanomateriales son muy empleados en la construcción de biosensores

electroquímicos gracias a sus excelentes propiedades electroconductoras y

electrocatalíticas. Sin embargo, de manera general los nanotubos de carbono poseen

desventajas asociadas a sus características intrínsecas, las cuales limitan su mayor uso

en la tecnología de los biosensores. En este sentido, la ausencia de grupos funcionales

superficiales en estos nanomateriales dificulta la inmovilización estable de

biorreceptores. Por otra parte, la alta hidrofobicidad de estas estructuras carbonáceas

afecta a su solubilidad o dispersabilidad en medios acuosos. Por tal motivo, se hace

necesaria la funcionalización de estos nanomateriales para su empleo en la


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construcción de plataformas biosensoras electroquímicas [Ajayan, 1999; Agüí et al.,

2008].

La funcionalización de los nanotubos de carbono puede realizarse mediante

transformación covalente o no covalente tanto de su cavidad interior como de su

superficie externa [Wang et al., 2008]. La funcionalización covalente introduce grupos

funcionales mediante transformación directa de los átomos de carbono en la

estructura del nanomaterial. Esta transformación reduce las capacidades

electroconductoras de los nanotubos y puede afectar a la actividad de las biomoléculas

posteriormente inmovilizadas debido a efectos de impedimento estérico [Wang,

2005b].

Por el contrario, la funcionalización no covalente se basa en interacciones de

adsorción física o atrapamiento, que no alteran las propiedades de los nanotubos de

carbono. Estas estrategias no covalentes generan una menor afectación en las

propiedades electroconductoras del nanomaterial. Sin embargo, los derivados no

covalentes de los nanotubos de carbono a ser empleados en la construcción de

biosensores deben ser específicamente diseñados para asegurar que proporcionen

estabilidad o vida útil a estos dispositivos bioanalíticos [Georgakilas et al., 2010].

En este trabajo se desarrollaron dos estrategias de derivatización no

covalente que mejoraban la solubilidad de los nanotubos de carbono en sistemas

acuosos y favorecían la inmovilización específica de biomoléculas sobre su superficie

sin afectar su estructura y propiedades, reduciendo a la vez los procesos de adsorción

no específicos de proteínas. Estos nuevos nanomateriales derivatizados fueron

evaluados como elementos de transducción para el diseño de biosensores

amperométricos, tomando la enzima xantina oxidasa como modelo de biorreceptor

catalítico. En la Figura 8.11 se muestra una representación esquematizada de las dos

estrategias de derivatización de nanotubos de carbono de pared simple descritas en

este trabajo.


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Figura 8.11. Funcionalización supramolecurar no covalente de nanotubos de carbono

de pared simple con: A) nanoparticulas Fe3O4 modificadas con polietilenglicol y xantina

oxidasa; y B) pireno funcionalizado con β-ciclodextrina para la formación de un

complejo de inclusión con el derivado de xantina oxidasa-adamantano.

En el trabajo publicado en Electroanalysis 23, 2011, 1790-1796 se demostró

que la funcionalización supramolecular de los nanotubos de carbono de pared simple

con un derivado de pireno modificado con β-ciclodextrina es una estrategia muy

eficiente para conferir hidrofilia y capacidad de formación de complejos de inclusión a

estos nanomateriales, sin alterar sus propiedades estructurales y electroconductoras.

A partir de este nanomaterial funcionalizado se describió la construcción de un

biosensor electroquímico con doble arquitectura supramolecular.

Para la preparación de este nuevo nanomaterial se sintetizó un derivado de β-

ciclodextrina monofuncionalizada con pireno (mono-6-etilendiamino-(2-

pirenocarboxamido)-6-deoxi-β-ciclodextrina), el cual se empleó en la funcionalización

superficial de los nanotubos mediante interacciones π-π. Este derivado fue empleado

como doble puente supramolecular para la posterior inmovilización de un derivado de

xantina oxidasa modificada con residuos de 1-adamantano, mediante la formación de

complejos de inclusión del tipo “host-guest”.

Aunque en este trabajo se ha empleado un modelo enzimático, debe

destacarse que los derivados de 1-adamantano pueden conjugarse fácilmente con


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cualquier proteína o ácido nucleico mediante métodos convencionales, por lo cual

podemos considerar esta estrategia como una metodología general para la

biofuncionalización supramolecular de nanotubos de carbono.

Esta estrategia de doble funcionalización supramolecular se utilizó para la

modificación de un electrodo de carbono vitrificado con el nanohíbrido preparado, en

aras de construir un biosensor amperométrico para xantina. El biosensor resultante

mostró muy buenas propiedades analíticas, destacando su bajo tiempo de respuesta

de 10 s, alta sensibilidad de 5.9 mA / M·cm2, bajo límite de detección de 2 mM y alta

estabilidad.

Por otra parte, en el artículo publicado en Journal of Materials Chemistry 21,

2011, 12858-12864 se describió un procedimiento de funcionalización basado en la

adsorción de derivados de polietilenglicol sobre la superficie de los nanotubos de

carbono de pared simple. Este polímero posee alta afinidad por los nanotubos de

carbono, adsorbiéndose de forma estable sobre su superficie, lo cual aumenta las

propiedades de solubilidad/dispersabilidad del nanomaterial y reduce los procesos de

adsorción inespecífica de proteínas [Shim et al., 2002; Sun et al., 2002]. En nuestro

caso, se prepararon nanopartículas superparamagnéticas de Fe3O4 (14 ± 7), las cuales

fueron inicialmente derivatizadas mediante reacción con (3-aminopropil)trietoxisilano

con el objetivo de introducir grupos amino primarios sobre su superficie. Estos grupos

amino primarios fueron posteriormente empleados como puntos de anclaje para la

funcionalización de las nanopartículas con cadenas de polietilenglicol (Mw = 5000 Da)

mediante reacción con un derivado del polímero monoactivado con N-

hidroxisuccinimida.

Las nanopartículas modificadas con polietilenglicol mostraron una mejor

capacidad de dispersión en disoluciones acuosas, y mantuvieron altas propiedades de

magnetización. Estas nanopartículas fueron posteriormente empleadas para la

funcionalización de nanotubos de carbono de pared simple, obteniéndose un derivado

de los nanotubos con buenas propiedades de dispersión en medios acuosos, y nuevas

propiedades de magnetización.


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Figura 8.12. Efecto del campo magnético externo generado por un imán de neodimio

sobre una dispersión de nanotubos de carbono de pared simple de concentración 0.1

mg/mL, antes (A) y después (B) de su funcionalización con nanopartículas

superparamagnéticas de Fe3O4 modificadas con polietilenglicol.

En la Figura 8.12 se muestra la respuesta de una dispersión de nanotubos de

carbono de pared simple funcionalizados con nanopartículas superparamagnéticas de

Fe3O4 bajo la acción de un campo magnético generado por un imán permanente de

neodimio.

Es importante resaltar que se eligió como estrategia la funcionalización exterior

de las paredes de los nanotubos con las nanopartículas de Fe3O4 en aras de mejorar la

hidrofilia de este nanomaterial y facilitar la posterior inmovilización covalente de la

enzima xantina oxidasa. Asimismo, la modificación de la cavidad de los nanotubos de

carbono implica procesos experimentales más complejos, y en general las propiedades

magnéticas resultantes son mucho más bajas tras el encapsulamiento de las

nanopartículas de Fe3O4 dentro de los nanotubos [Korneva et al., 2005].

El nanohíbrido magnético obtenido se empleó en la construcción de un

biosensor enzimático para la determinación de xantina, utilizando como soporte

electrodos serigrafiados de oro. Para ello, las nanopartículas de Fe3O4 recubiertas con

polietilenglicol fueron pre-activadas por tratamiento con glutaraldehído, en aras de

funcionalizar los grupos amino primarios remanentes en su superficie. Las


394

nanopartículas pre-activadas se incubaron posteriormente con una disolución de

xantina oxidasa, asegurando que solo los grupos amino terminales de la enzima

reaccionen con los aldehídos de la superficie de las nanopartículas. Finalmente, las

nanopartículas biofuncionalizadas con la enzima fueron empleadas para la

funcionalización no covalente de los nanotubos de carbono de pared simple, a través

de la adsorción irreversible de las cadenas de polímero sobre los nanotubos.

Se comprobó que la actividad catalítica de la enzima no se vió afectada

significativamente tras su inmovilización en el nanomaterial, lo cual puede atribuirse

tanto a la estrategia de pre-activación empleada en el proceso de inmovilización, como

al microambiente hidrofílico creado por las cadenas de polietilenglicol, y al bajo

impedimento estérico que los nanotubos de carbono ejercen sobre el centro activo de

la enzima debido a la naturaleza flexible de los brazos espaciadores de polímero

empleados en la funcionalización de los nanotubos.

El nanomaterial biofuncionalizado fue inmovilizado magnéticamente sobre la

superficie de los electrodos para la construcción de un biosensor amperométrico para

xantina. La efectividad de esta estrategia de ensamblaje se confirmó con las excelentes

propiedades analíticas obtenidas para este dispositivo. El biosensor resultante mostró

una dependencia lineal con la concentración de xantina en el rango de 250 µM a 3.5

mM, con una rápida respuesta amperométrica en 12 s. Asimismo, el biosensor mostró

una alta sensibilidad de 1.31 A·M/ cm2 y un bajo límite de detección de 60 nM.

8.1.3. Biosensores basados en dendrímeros y dendrones de poliamidoamina

modificados con ciclodextrinas.

En los artículos incluidos en este apartado se describen los resultados

relacionados con la construcción de electrodos nanoestructurados mediante derivados

de dendrímeros y dendrones de poliamidoamina funcionalizados con β-ciclodextrina

para el diseño de biosensores electroquímicos empleando enzimas como

biorreceptores.


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La estrategia propuesta se basa en la utilización de estos derivados de los

dendrímeros y dendrones como elementos estructurales para el ensamblaje de

arquitecturas tridimensionales sobre las superficies electródicas. Este tipo de

polímeros dendríticos posibilita el diseño de estructuras tridimensionales con una alta

densidad de grupos reactivos superficiales, lo que permite la inmovilización estable y

multipuntual de una alta densidad de biomoléculas mediante uniones covalentes o no

covalentementes. Este hecho favorece la construcción de biosensores con alta

sensibilidad [Soler et al., 2015].

Asimismo, la estructura permeable de estos polímeros favorece la difusión de

las especies electroactivas hacia la superficie de los electrodos. Este hecho es una

ventaja en comparación con otras macromoléculas comúnmente empleadas en la

construcción de arquitecturas capa-a-capa, las cuales en general ejercen serios

impedimentos estéricos que limitan los fenómenos de difusión de sustancias y

transferencia de carga en las interfases electródicas [Vögtle et al., 2009].

Debe destacarse que uno de los aspectos más importantes a tener en cuenta

en el diseño de un biosensor es la necesidad de garantizar la inmovilización estable de

los biorreceptores sin afectar significativamente su estructura biológicamente activa, y

por tanto su actividad funcional [Wang et al., 2008]. Por ese motivo, en el presente

trabajo se eligió una estrategia de inmovilización supramolecular del tipo “host-guest”

para las enzimas en estudio, funcionalizando los grupos amino primarios terminales de

los dendrímeros y dendrones con unidades de -cliclodextrinas y modificando las

enzimas con residuos de 1-adamantano. Estas estrategias de inmovilización

supramolecular de biorreceptores han sido validadas en otros sistemas biosensores

previamente publicados en la literatura [Díez et al., 2012; Villalonga et al., 2003;

Fernández, 2003]

Por otra parte, la introducción de residuos de -cliclodextrinas en la superficie

de los polímeros dendríticos favorece la formación de sistemas multicapas

supramoleculares estables mediante el uso de biomoléculas modificadas con

compuestos hidrofóbicos de alta afinidad por estos oligosacáridos cíclicos, tales como

los derivados de 1-adamantano [Fragoso et al., 2002].


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En la Figura 8.13 se muestra una representación esquemática de las

arquitecturas supramoleculares ensambladas sobre la superficie de los electrodos,

empleadas en este trabajo de tesis. Se diseñaron dos estrategias para la modificación

de electrodos de oro, basadas en:

A) Formación de monocapas autoensambladas de dendrones de poliamidoamina G-4

-ciclodextrina y decorados con

nanopartículas de platino (Analytical and Bioanalytical Chemistry 405, 2013, 3773-

3781), y

B) Autoensamblaje supramolecular de una arquitectura capa-a-capa mediante

alternación de -

ciclodextrina y peroxidasa modificada con adamantano (Electrochimica Acta 76, 2012,

249-255).

Figura 8.13. Representación esquemática de los electrodos de oro modificados con

monocapas (A) y arquitecturas capa-a-capa (B) de moléculas dendríticas de

-ciclodextrina

En ambos casos se emplearon drendrímeros de poliamidoamina G-4 con núcleo

de cisteamina como polímeros de partida para su funcionalizados con ciclodextrinas y

posterior modificación de la superficie electródica de oro. Estos dendrímeros poseen

como núcleo un enlace disulfuro, el cual es sensible a la acción de agentes reductores,

dando como producto de dicho proceso de reducción dos moles de dendrones con

núcleo de cisteamina por cada mol de dendrímero tratado. La presencia de un grupo

tiol activo por cada mol de dendrón producido posibilita la fácil y rápida


397

funcionalización de las superficies de los electrodos de oro mediante interacciones de

quimisorción, formando monocapas permeables con una alta densidad de grupos

funcionales en la superficie del polímero.

Esta primera monocapa es empleada como apoyo estable para el subsiguiente

ensamblado de las arquitecturas supramoleculares propuestas. En el trabajo publicado

en Analytical and Bioanalytical Chemistry 405, 2013, 3773-3781 se funcionalizó el

dendrón de poliamidoamina modificado con ciclodextrinas con nanopartículas de

platino. Estas nanopartículas son formadas in situ dentro de las cavidades del polímero

dendrítico mediante reducción del H2PtCl6 con NaBH4. Las nanopartículas sintetizadas

son estabilizadas dentro de la estructura del dendrón mediante interacción con los

grupos amida de su estructura [Endo et al., 2005].

Esta estrategia de funcionalización con nanopartículas de platino se basó en las

excelentes propiedades electrocatalíticas de estos nanomateriales para la

descomposición del H2O2 [Hrapovic et al., 2004], el cual es una de los productos más

comunes de las reacciones catalizadas por muchas enzimas oxidasas de amplio uso en

la construcción de biosensores electroquímicos [Clark et al., 1965].

Los electrodos nanoestructurados con esta estructura dendrítica fueron

empleados en la construcción de un biosensor enzimático de tercera generación para

la detección de glucosa. Para ello, se sintetizó un derivado de la enzima glucosa

oxidasa químicamente modificada con unidades de adamantano, la cual pudiera ser

fácilmente inmovilizada sobre la superficie de los electrodos mediante interacciones

supramoleculares.

La validez de este modelo de ensamblaje electródico fue corroborada por las

excelentes propiedades analíticas y de estabilidad del biosensor de tercera generación

construido. En este sentido, el biosensor mostró una rápida respuesta amperométrica

de solo 6s, un bajo límite de detección de 2,0 M, y una alta sensibilidad de 197

mA/cm2 M. Este biosensor con arquitectura supramolecular mostró una buena

estabilidad, reteniendo el 94% de su respuesta inicial después de 9 días de

almacenamiento a 4ºC.


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Por otra parte, en el trabajo publicado en Electrochimica Acta 76, 2012, 249-

255 se describe el ensamblado de una arquitectura electródica mucho más compleja.

En este caso, las superficies de oro modificadas con dentrones de poliamidoamina G-4

modificados con ciclodextrinas fueron empleados como soporte para el ensamblado

de estructuras capa-a-capa de mayores dimensiones. Estas estructuras multicapas

fueron construidas mediante el ensamblado de capas alternas de la enzima peroxidasa

modificada con unidades de adamantano y dendrímeros de poliamidoamina G-5

modificados con ciclodextrinas.

Para realizar estas estructuras más complejas se empleó un dendrímero de

generación 5 en aras de contar con una mayor densidad de grupos funcionales

factibles a ser modificados con ciclodextrinas en el polímero. Este hecho debería

contribuir a una mayor estabilidad en la arquitectura capa-a-capa a ensamblar

posteriormente sobre el electrodo, pues facilitaría la formación de un mayor número

de interacciones supramoleculares y multipuntuales con el derivado de la enzima. Por

otra parte, se tomó la enzima peroxidasa como modelo para la construcción de este

ensamblado dada la importancia que revierte la detección electroanalítica de H2O2

[Baussanne et al., 2000], y la experiencia del grupo de investigación en la preparación

de derivados de esta enzima con adamantano [Camacho et al., 2007].

Una de las limitaciones de las plataformas biosensoras electroquímicas es la

imposibilidad de aumentar la densidad de biorreceptores en la interfase electródica, y

con ello la sensibilidad de las mediciones, más allá de la capacidad superficial de estas

interfases. Sin embargo, en nuestro trabajo se demostró que la actividad

electroanalítica de los ensamblados supramoleculares frente al H2O2 aumentaba con el

aumento del número de capas de la enzima, obteniéndose una máxima respuesta

analítica en aquellos electrodos con 3 capas de enzima. Asimismo, se demostró que la

capacidad electroanalítica de los biosensores con mayor número de capas era mucho

menor, lo cual puede ser justificado por la baja estabilidad estructural de estos

ensamblados de gran tamaño. Estos resultados permiten proponer esta metodología

de construcción de biosensores enzimáticos como una estrategia válida para la mejora

de las propiedades analíticas de estos dispositivos, al proponer estas arquitecturas


399

tridimensionales como vía para el aumento de la densidad espacial de biorreceptores

sobre los electrodos.

Los electrodos construidos con 3 capas de peroxidasa mostraron un bajo límite

de detección de 160 nM y una sensibilidad de 602 A/cm2 M frente al H2O2. Asimismo,

estos biosensores mostraron una alta estabilidad, reteniendo el 63% de la respuesta

inicial después de 30 días de almacenaje a 4°C en condiciones de humedad a 4°C, lo

cual demuestra la alta estabilidad estructural de las arquitecturas supramoleculares

capa-a-capa construidas sobre los electrodos.


400

8.2. SISTEMAS DE LIBERACIÓN INTELIGENTE DE FÁRMACOS CONTROLADOS POR

ENZIMAS Y BASADOS EN NANOMATERIALES DE SÍLICE MESOPOROSA.

En la segunda parte de este trabajo se discuten los resultados relacionados con

el diseño de nanomateriales mesoporosos funcionalizados para el ensamblaje de

sistemas de liberación inteligente de fármacos controlados por enzimas.

La estrategia general seguida en esta investigación se basa en la integración, en

una misma nanopartícula funcionalizada, de una unidad de control biomolecular con la

unidad de encapsulamiento del fármaco. Para ello se han diseñado unidades de

control enzimático, encargadas de detectar la presencia de determinados compuestos

en el medio (señales INPUT) y transformarlos catalíticamente en productos capaces de

activar los mecanismos de aperturas de los mesoporos donde se encuentra

encapsulada la sustancia que se desea liberar (OUTPUT).

Atendiendo a esta estrategia, se han desarrollado dos modelos diferentes de

nanomáquinas basadas en:

- Nanopartículas de sílice mesoporosa funcionalizadas con neoglicoenzimas, que

actúan dualmente como puertas moleculares y sistemas sensores-efectores.

- Nanopartículas Janus de oro y sílice mesoporosa, funcionalizadas con enzimas en la

cara metálica como sistemas sensores-efectores y mecanizadas con puertas

moleculares estímulo-dependientes en la cara mesoporosa.

8.2.1. Nanomáquinas basadas en nanopartículas de sílice mesoporosa

funcionalizadas con neoglicoenzimas.

El primer modelo descrito en esta Tesis, publicado en ACS Applied Materials

and Interfaces 8, 2016, 7657−7665, describe la preparación de nanopartículas de sílice

mesoporosa funcionalizadas superficialmente con residuos de ácido borónico. Estos

residuos son empleados como puntos de enlace para la inmovilización supramolecular

de un derivado artificialmente glicosidado de la enzima esterasa de hígado de cerdo.


401

La esterasa es una enzima que se encuentra en la Naturaleza en forma de glicoenzima

[Arpigny et al., 1999], por lo cual puede formar en medios alcalinos ésteres cíclicos con

los residuos de ácido borónico empleando los grupos 1,2-diol vecinales de los residuos

de azúcares enlazados a su cadena polipeptídica.

Figura 8.14. Representación esquemática del mecanismo de formación de ésteres

cíclicos de ácido borónico con azúcares.

Estos ésteres cíclicos son reversibles y sensibles al pH del medio,

descomponiéndose con el aumento de la acidez. Por otra parte, la estabilidad de estos

ésteres cíclicos depende de la orientación espacial de los grupos 1,2-diol vecinales,

siendo mayor en aquellos que poseen una configuración cis que en los que presentan

configuración trans. Por tal motivo, los residuos de ácido borónico forman

asociaciones supramoleculares de diferente estabilidad con los distintos tipos de

monosacáridos [Lorand et al., 1959], lo cual ha sido anteriormente empleado en el

establecimiento de sistemas sensores selectivos para diferentes tipos de azúcares

[James et al., 1996].

Sobre estas bases, nuestra hipótesis de trabajo se basó en la posibilidad de

conferir diferentes grados de estabilidad a los complejos de la esterasa con los

residuos de ácido borónico sobre la superficie de las nanopartículas de sílice

mesoporosa, mediante la manipulación de la estructura de la enzima por modificación

química con residuos de lactosa. De esta forma, la neoglicoenzima resultante podría

ser empleada como elemento de cierre para ensamblar puestas moleculares

reversibles con diferente grado de estabilidad y nivel de respuesta ante diferentes

estímulos. Esta estrategia de empleo dual de derivados enzimáticos como sistemas


402

sensores-efectores y como elementos de cierre en puertas moleculares estímulo-

dependientes ha sido anteriormente propuesta por el equipo de investigación

involucrado en este trabajo [Aznar et al., 2013].

La hipótesis planteada fue demostrada a partir de los resultados

experimentales obtenidos. En este sentido se comprobó que las nanomáquinas

construidas con la neoglicoenzimas poseían diferente comportamiento ante la

presencia de glucosa o etilenglicol, que frente al butirato de etilo. En el primer caso, las

nanomáquinas cargadas con un colorante de prueba, experimentaban una apertura

parcial de las puertas moleculares al adicionar glucosa o etilenglicol en el medio. Estos

compuestos, que poseen grupos 1,2-diol vecinales, son capaces de desplazar los

residuos de azúcares más débilmente enlazados en los complejos con ácido borónico,

promoviendo así la liberación parcial del colorante encapsulado.

Por otra parte, en presencia de butirato de etilo se observó una apertura total

de las puertas moleculares ensambladas con la neoglicoenzima, con la consiguiente

liberación total del colorante de prueba encapsulado. El mecanismo propuesto para

este fenómeno se basa en la trasformación catalítica del butirato de etilo por la

esterasa, produciendo ácido butírico, el cual al disminuir el pH del medio promueve la

ruptura de los ésteres cíclicos de ácido borónico con los azúcares de la enzima.

Sobre la base de estos resultados, la nanopartícula funcionalizada con la

neoglicoenzima de esterasa-lactosa fue empleada en el ensamblaje de una

nanomáquina controlada por la enzima para la liberación bajo demanda, secuencial y

pulsátil de medicamentos. Este tipo de nanomáquinas podría ser muy valiosa en el

establecimiento de cronoterapias personalizadas para el tratamiento de enfermedades

influenciadas por el ritmo circadiano del paciente [Youan, 2004].

Estas nanomáquinas fueron validadas con éxito en experimentos ex vivo para la

liberación secuencial y programada del fármaco antitumoral Doxorrubicina en células

HeLa de cáncer. En este sentido, se comprobó que las nanomáquinas eran fácilmente

internalizadas dentro de las células tumorales, y que eran capaces de liberar


403

secuencialmente diferentes dosis del fármaco en presencia de glucosa y butirato de

etilo.

8.2.2. Nanomáquinas controladas por enzimas y basadas en nanopartículas Janus de

oro y sílice mesoporosa.

Aunque las nanopartículas de sílice mesoporosa constituyen una opción muy

atractiva para el diseño de sistemas liberación controlada de fármacos de dimensiones

nanométricas, y en trabajos previos hemos demostrado la viabilidad de construir

nanomáquinas controladas por enzimas empleando estos nanomateriales [Aznar et al.,

2013], existe una serie de limitaciones para el uso de estas partículas en el ensamblaje

de sistemas más complejos.

En la Figura 8.15 se representa el esquema general de la nanomáquina que se

pretende construir, donde se integrarán los diferentes mecanismos moleculares y

biomoleculares que aseguren el funcionamiento inteligente y autónomo del

dispositivo. En este sentido, hemos previsto que esta nanomáquina deberá estar

provista con una unidad de control y una unidad de encapsulamiento integradas en la

misma partícula. En la unidad de control se ensamblará un sistema sensor de afinidad

capaz de localizar los tejidos u órganos a los cuales se va a dirigir la liberación del

fármaco. Para realizar su función de control, en esta unidad de control se inmovilizará

un sistema lógico sensor-efector enzimático, capaz de detectar la presencia de

determinados compuestos como señal INPUT, y tras procesarlos catalíticamente,

producir compuestos capaces de controlar la apertura de las puertas moleculares que

aseguran el encapsulamiento del fármaco en la unidad de encapsulamiento.

Tal nivel de complejidad y organización sería muy difícil de ensamblar en la

superficie de las nanopartículas de sílice mesoporosa, por lo cual nos planteamos

como objetivo el diseño, preparación y caracterización de un nuevo tipo de partículas

que pudiera servir como “hardware” para el ensamblaje de los diversos mecanismos

involucrados en la construcción de una nanomáquina autónoma e inteligente.


404

Figura 8.15. Representación esquemática de una nanomáquina para la liberación sitio-

específica, autónoma e inteligente de fármacos bajo control lógico modulado por

enzimas.

Para cumplir este objetivo, se determinó sintetizar una nanopartícula Janus

mediante la unión de una nanopartícula de sílice mesoporosa, que actuaría como

unidad de encapsulación, y una nanopartícula de oro, sobre la cual se inmovilizarían

los diferentes mecanismos biomoleculares que compondrían la unidad de control. Las

ventajas asociadas al uso de nanomateriales mesoporosos como medios de

encapsulamiento han sido destacas en otros capítulos de esta tesis, y el empleo de

nanopartículas de oro en este nanomaterial anisotrópico se justifica por la alta

biocompatibilidad, estabilidad coloidal y facilidad de preparación de estas

nanopartículas [Lehner et al., 2012]. Asimismo, las nanopartículas de oro pueden ser

fácilmente empleadas como soporte para la inmovilización de biomoléculas mediante

el uso de ligandos tiolados [Crespo et al., 2004].

Para preparar estas nanopartículas Janus se empleó una estrategia de

enmascaramiento con modificación selectiva de interfases, empleando como sistema

de enmascaramiento emulsiones de Pickering con parafina como fase oleosa (Figura

8.16).


405

Figura 8.16. Estrategia de enmascaramiento y manipulación selectiva de interfases

empleada en la preparación de nanopartículas Janus de oro y sílice mesoporosa.

Según esta estrategia, las nanopartículas de sílice mesoporosas se adsorben en

la superficie de las microgotas de parafina líquida confiriendo hidrofilicidad superficial

y estabilidad coloidal en medios acuosos a los coloidosomas formados. En la Figura

8.17 se muestra la imagen de uno de estos coloidosomas, obtenida mediante

microscopia FE-SEM.

Figura 8.17. Imagen FE-SEM de los coloidosomas de parafina recubiera con

nanopartículas de sílice mesoporosa.

Estos coloidosomas actúan como sistemas de enmascaramiento parcial de la

superficie de las nanopartículas de sílice mesoporosa adsorbidas en ellos, posibilitando


406

la modificación selectiva de la interfase expuesta al medio acuoso. Para ello, se

funcionalizó esta superficie expuesta con (3-mercaptopropil)trimetosilano, con el

objetivo de introducir grupos tioles libres en esta interfaz, sobre los cuales

posteriormente se enlazan las nanopartículas de oro mediante interacciones

multipuntuales de quimiosorción.

Las nanopartículas Janus preparadas poseen la ventaja de poseer dos caras

químicamente diferentes, lo cual facilita en ensamblado secuencial y topoespecífico de

los diferentes mecanismos moleculares y biomoleculares necesarios para el

funcionamiento de las nanomáquinas. Los trabajos incluidos en este apartado de Tesis

describen los experiementos realizados para comprobar la factibilidad de ensamblar

los diferentes mecanismos de las nanomáquinas.

Inicialmente, en el trabajo publicado en Electrochemistry Communications 30,

2013, 51-54, se describe el diseño de un modelo para comprobar la viabilidad de

incorporar un receptor de afinidad, capaz de actuar como sensor de localización, en la

unidad de control de la nanomáquina. Para ello se inmovilizó estreptavidina en la cara

de oro de la nanopartícula Janus, biomolécula que posee una alta afinidad por la

biotina. Para detectar el proceso de biorreconocimiento, se inmovilizó la enzima

peroxidasa en la cara de sílice mesoporosa de la nanopartícula Janus, y se prepararon

electrodos de oro recubiertos con residuos de biotina con objeto de comprobar este

reconocimiento mediante técnicas electroquímicas. Los resultados obtenidos

demostraron claramente que esta nanopartícula Janus biofuncionalizada actúa como

un nanosistema de biorreconocimiento y señalización.

Debe destacarse que este trabajo, además de servir de demostración sobre la

posibilidad de biofuncionalizar las nanopartículas Janus con receptores de afinidad en

la unidad de control, constituyó la primera publicación sobre el uso de nanopartículas

como nanosistemas de biorreconocimiento y señalización electroquímica.

En el trabajo publicado en Chemistry – A European Journal 19, 2013, 7889-

7894, se planteó la construcción de la primera nanomáquina basada en estas

nanopartículas Janus, controlada por una enzima. Para construir esta nanomáquina se


407

empleó cloruro de tris(2,2´-bipiridil) rutenio (II) como modelo de sustancia a

encapsular, dada las excelentes propiedades ópticas de este complejo. Este colorante

fue encapsulado dentro de los nanoporos de la cara mesoporosa de las nanopartículas

Janus, empleando puertas de poliaminas sensibles al pH ensambladas en la superficie

exterior de los poros.

Por otra parte, se inmovilizó la enzima ureasa sobre la cara de oro de la

nanopartícula anisotrópica con el objetivo de emplear este biocatalizador como

sistema sensor-efector, dada su capacidad de reconocer la presencia de urea en el

medio como señal INPUT, y transformar este substrato enzimático en NH3 y CO2 con el

consiguiente aumento de la basicidad del medio.

La nanomáquina ensamblada fue validada en experimentos in vitro,

comprobándose su capacidad de liberar el colorante encapsulado ante la presencia de

urea en el medio.

Finalmente, en el artículo publicado en Journal of the American Chemical

Society 136, 2014, 9116-9123, se describe el ensamblaje de una nanomáquina más

sofisticada. Este dispositivo está regulado por un sistema multienzimático integrado

sobre la superficie de oro de las nanopartículas Janus, basado en la combinación de las

enzimas glucosa oxidasa y esterasa. Esta combinación enzimática opera como una

puerta lógica tipo OR, y con ella se propuso la modulación del mecanismo de liberación

del compuesto encapsulado en la cara de sílice mesoporosa.

Para ello, se encapsuló el colorante cloruro de tris(2,2´-bipiridil) rutenio (II)

dentro de los poros de la cara de sílice mesoporosa de las nanopartículas Janus,

ensamblando nanoválvulas supramoleculares de benzimidazol:β-ciclodextrina sobre la

superficie externa de los poros. Estas nanovávulas son sensibles al cambio de pH del

medio, y su mecanismo de apertura puede ser controlado mediante el reconocimiento

y transformación de la glucosa y el butirato de etilo, como señales INPUT, por el

sistema enzimático inmovilizado, que actúa como operador tipo OR.

El funcionamiento de esta nanomáquina fue comprobado en experimentos in

vitro, determinándose que el colorante solo era liberado al medio tras la adición de


408

glucosa, butirato de etilo, o una mezcla de ambos compuestos, de acuerdo a la

combinación de señales INPUT necesarias para un operador tipo OR.

El nivel de complejidad de esta nanomáquina fue incrementado mediante el

acoplamiento de un operador enzimático tipo RESET, para ello se co-inmovilizó la

enzima ureasa sobre la cara de oro de la nanopartícula Janus. En este sentido, se

comprobó que este operador RESET era capaz de detener la apertura de la nanoválvula

supramolecular, y por tanto la liberación al medio de incubación del colorante

encapsulado mediante la adición de urea.

Este control por el operador enzimático RESET resultó ser asimismo sensible a

la concentración de urea en el medio, por lo cual la extensión en que la nanomáquina

operada por la puerta lógica OR era capaz de liberar el colorante pudo ser controlada

mediante la manipulación de la concentración de urea adicionada.

En aras de validar esta nanomáquina en modelos ex vitro de liberación de

fármacos, se preparó una nanopartícula Janus controlada por el operador enzimático

OR, en la cual se encapsuló el fármaco antitumoral Doxorrubicina. Esta nanomáquina

fue incubada con células HeLa de cáncer, comprobándose la internalización de éstas

nanopartículas dentro de las células tumorales. Asimismo, se comprobó la capacidad

de esta nanomáquina de liberar, de forma autónoma y bajo demanda, el fármaco

antitumoral en presencia de glucosa y butirato de etilo como señales INPUT.

9 CONCLUSIONS

9. CONCLUSIONS

409

The present thesis has compiled novel design strategies for nanomaterial based

biosensors and drug delivery systems. In addition, these tailor made nanomaterials

present clear advantages over the conventional ones increasing the performance of

final obtained biosensors and giving versatility to newly introduced drug delivery

systems. In the following the most important conclusions are outlined and

summarized.

For biosensors design:

Novel polyfunctionalized gold nanoparticles capped with 2-

mercaptoethanesulfonic acid, p-aminothiophenol and a third ligand able to be

used as immobilization point for biorreceptors, were prepared through a one-pot

reaction scheme. These nanomaterials were successfully employed to assemble

electropolymerized networks of nanoparticles on electrode surfaces. These

networks are used for the construction of enzymatic and affinity-based

electrochemical biosensors. The efficacy of these nanostructured transduction

elements was confirmed by the low detection limit and high sensitivity of the

resulting electroanalytical devices.

Two original non-covalent strategies for the modification of single-walled carbon

nanotubes were reported. The first approach was based on the non-covalent

surface modification of carbon nanotubes with polyethyleneglycol-modified Fe3O4

superparamagnetic nanoparticles, providing magnetic properties to the resulting

hybrid nanomaterial. The second method was based on the supramolecular

attachment of a β-cyclodextrin-modified pyrene derivative through π-π

interactions. The functionalized nanomaterials were successfully employed to

construct highly sensitive xanthine oxidase-based amperometric biosensors for

xanthine.

Original β-cyclodextrin-modified polyamidoamine dendrons and dendrimers were

synthesized and employed for the self-assembly of layered supramolecular

architectures on electrodes surfaces by using adamantane-modified enzymes as

biomolecular building blocks. These polymer derivatives with molecular receptor

9. CONCLUSIONS

410

properties were successfully employed to design a reagentless amperometric

enzyme biosensor for glucose, as well as a horseradish peroxidase-based layer-by-

layer supramolecular architecture for the electrochemical detection of H2O2.

For drug delivery systems:

A novel smart delivery system for the controlled and pulsatile release of

nanoencapsulated compounds was designed by using functionalized mesoporous

silica nanoparticles as nanocontainers and lactose-modified esterase as sensing

and gating elements. This model was validated for the on-command and

neoglycoenzyme-controlled delivery of the anticancer drug doxorubicin to HeLa

cancer cells in ex vivo experiments.

Original Au-mesoporous silica Janus nanoparticles were prepared through a

masking approach and employed as “hardware” for the assembly of enzyme-

controlled nanomachines. We demonstrated that biofunctionalized Janus

nanoparticles-based nanomachines can be provided with affinity biorecognition

properties by using the biotin-streptavidin system as model.

It was also proved that the on-command delivery of payloads from Janus

nanomachines can be controlled by a single enzyme, or by combinations of

enzymes acting as logic operators. Janus nanomachines were validated for the

smart, autonomous and enzyme-controlled release of the anticancer drug

doxorubicin to HeLa cancer cells.

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10. Referencias

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