Catechol biosensing using a nanostructured layer-by-layer film containing Cl-catechol 1,2-dioxygenase

Biosensors and Bioelectronics 21 (2006) 1320–1326

Catechol biosensing using a nanostructured layer-by-layer film containing Cl-catechol 1,2-dioxygenase Valtencir Zucolotto a,∗ , Andressa P.A. Pinto a , Tathyana Tumolo b , Marli L. Moraes a , Maur´ıcio S. Baptista b , Antonio Riul Jr. c , Ana Paula U. Ara´ujo a , Osvaldo N. Oliveira Jr. a a

c

Instituto de F´ısica de S˜ao Carlos, USP, CP 369, 13560-970 So Carlos, SP, Brazil b Instituto de Qu´ımica, USP, CP 26077, 05513-970 S˜ ao Paulo, SP, Brazil DFQB, Universidade Estadual Paulista, CP 467, 19060-900 Presidente Prudente, SP, Brazil Received 18 March 2005; received in revised form 16 May 2005; accepted 10 June 2005 Available online 27 July 2005

Abstract The detection of aromatic compounds from pesticides and industrial wastewater has become of great interest, since these compounds withstand chemical oxidation and biological degradation, accumulating in the environment. In this work, a highly sensitive biosensor for detecting catechol was obtained with the immobilization of Cl-catechol 1,2-dioxygenase (CCD) in nanostructured films. CCD layers were alternated with poly(amidoamine) generation 4 (PAMAM G4) dendrimer using the electrostatic layer-by-layer (LbL) technique. Circular dichroism (CD) measurements indicated that the immobilized CCD preserved the same conformation as in solution. The thickness of the very first CCD layers in the LbL films was estimated at ca. 3.6 nm, as revealed by surface plasmon resonance (SPR). PAMAM/CCD 10-bilayer films were employed in detecting diluted catechol solutions using either an optical or electrical approach. Due to the mild immobilization conditions employed, especially regarding the pH and ionic strength of the dipping solutions, CCD remained active in the films for periods longer than 3 weeks. The optical detection comprised absorption experiments in which the formation of cis–cis muconic acid, resulting from the reaction between CCD and catechol, was monitored by measuring the absorbance at 260 nm after film immersion in catechol solutions. The electrical detection was carried out using LbL films deposited onto gold-interdigitated electrodes immersed in aqueous solutions at different catechol concentrations. Using impedance spectroscopy in a broad frequency range (1Hz–1kHz), we could detect catechol in solutions at concentrations as low as 10−10 M. © 2005 Elsevier B.V. All rights reserved. Keywords: Biosensor; Layer-by-layer; Enzyme immobilization; Cl-catechol 1,2-dioxygenase

1. Introduction The layer-by-layer (LbL) technique has been largely employed in the immobilization of proteins and other biomolecules following the pioneering works of Lvov et al. (1993, 1995), where advantage was taken of the outstanding control in both film thickness and supramolecular architecture. Indeed, because of the suitable film fabrication conditions, including the use of aqueous solutions at optimized pH and ionic strength, immobilization in LbL films preserves ∗

Corresponding author. Tel.: +55 16 273 9825; fax: +55 16 271 5365. E-mail address: [email protected] (V. Zucolotto).

0956-5663/$ – see front matter © 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.bios.2005.06.001

protein activity over a considerable period of time. This has paved the way for a wide range of bio-related materials be incorporated as sensing elements in LbL films, such as DNA (Sukhorukov et al., 1996), enzymes (Chen et al., 1998; Onda et al., 1999; Yoon and Kim, 2000) and antigen–antibody pairs (Brynda et al., 1998; Trau et al., 2002). Biosensors designed for aromatic compounds found in pesticides and industrial wastewater, in particular, have become of great interest, as far as the environment is concerned, since many of such compounds withstand chemical oxidation and biological degradation (Broderick and O’Halloran, 1991). The aerobic degradation of non-halogenated aromatic compounds, for example, usually requires its initial conversion to a dihy-

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droxybenzene derivative such as catechol or protocatechuate, which are further degraded via ring cleavage by dioxygenases (Broderick and O’Halloran, 1991). As for chloroaromatic compounds, degradation seems to occur along similar principles, except for the formation of intermediate compounds that may take place via two distinct pathways, depending on the number of chlorine atoms in the molecule (Schl¨omann, 1994; Ferraroni et al., 2004). Two types of enzymes catalyse the ring cleavage: (i) the intradiol dioxygenases, which are metalloenzymes containing Fe3 + that yield cis–cis-muconic acid via ortho-cleavage and (ii) the extradiol dioxygenases that contain Fe2 + yielding compounds such as 2-hydroxymuconic or derivatives (Broderick and O’Halloran, 1991; Schl¨omann, 1994). Among the enzymes that act via ortho-cleavage, chlorocatechol 1,2-dioxygenase (CCD) has exhibited affinity to both halogenated and non-halogenated substrates. One can therefore envisage the use of dioxygenases in biosensors to exploit their catalytic characteristics, especially the broad substrate specificity and high stereo selectivity. In this work CCD (Ara´ujo et al., 2000) was immobilized in nanostructured films in conjunction with poly(amidoamine) (PAMAM) dendrimer in a layer-by-layer fashion. The enzymes remained active after immobilization, which allowed the films to be used in detecting catechol in solutions at very low molar concentrations, employing optical and electrical measurements.

2. Experimental details Poly(allylamine hydrochloride) (PAH) (Mw = 70,000 g/ mol), poly(vinylsulfonic acid) (PVS) and NH2 -terminated PAMAM generation 4 (PAMAM G4) were purchased from Aldrich and used without further purification. The chemical structures of the polyelectrolytes employed are shown in Fig. 1. The recombinant CCD used was produced in Escherichia coli BL21(DE3). The protocol for expression

Fig. 1. Chemical structure of the polyelectrolytes employed for film fabrication.

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and purification is described elsewhere (Ara´ujo et al., 2000). Briefly, the host cells harboring pTYBCLCA grew in a Luria Bertani (LB) medium, to which 0.1 g/L of FeSO4 and 100 ␮g/ml ampicilin had been added. The cells were suspended in Tris–NaCl buffer (20 mM Tris, 500 mM NaCl, 20 ␮M phenyl methyl sulfonyl fluoride (PMSF), pH 8.0), lysed by sonication and the cell debris were separated by centrifugation at 20,000 × g. The supernatant was loaded onto a chitin column (New England Biolabs-NBL) pre-equilibrated with the same buffer used for the re-suspension of the cells. The column was washed with the re-suspension buffer and subsequently with 20 mM Tris–HCl, pH 8 buffer containing 50 mM NaCl, 30 mM dithiothreitol (DTT), and 20 ␮M PMSF. The CCD protein was eluted with the same buffer after 15 h of incubation at 8 ◦ C. The active fractions were combined, concentrated and loaded onto a Superdex 75 column (Pharmacia) (1.6 × 60 cm) in 20 mM Tris–NaCl buffer, pH 8.4 with NaCl 50 mM, and 1 mM PMSF. The major peak eluted was concentrated and used for all experiments. Both PAMAM and CCD were used at 2 mg/mL, in a 20 mM Tris, 50 mM NaCl buffer solution, kept at pH 7.5, as CCD is negatively charged at this pH value. Nanostructured LbL films containing up to 40 PAMAM/CCD bilayers were assembled on quartz slides for optical measurements and five-bilayer PAMAM/CCD films were deposited onto gold-interdigitated electrodes for electrical measurements. All quartz substrates were cleaned in a NH4 OH/H2 O2 /H2 O (5:1:1) bath for 20 min and the interdigitated electrodes were carefully washed with detergent in an ultrasonic bath. After cleaning, the substrates were covered with two bilayers of a PAH/PVS LbL film (both PAH and PVS dipping solutions were used at concentrations of 1 mg/mL, pH 6.5). The latter procedure was employed to ensure a uniform distribution of charges onto the substrates prior to the PAMAM/CCD film deposition. The sequential deposition of multilayers was carried out in a HMS series programmable slide stainer (Carl Zeiss Inc.) by immersing the quartz slides or interdigitated electrodes alternately into the PAMAM (polycationic) and CCD (polyanionic) solutions for five and 15 min, respectively. After deposition of each layer the substrate + film system was immersed for 1 min in the buffer solution. The deposition process was monitored at each deposited bilayer using UV–vis spectroscopy. The idealized architecture of the PAMAM/CCD films is depicted in Fig. 2. The analysis of film morphology was carried out with an atomic force microscope (AFM) Nanoscope III (Digital Instruments). Circular dichroism (CD) spectra from CCD in solution and immobilized on LbL films were recorded using a Jasco J-720 spectropolarimeter. Surface plasmon resonance (SPR) experiments were carried out using an SPR-EVM-Spreeta Evaluation Module, Texas Instruments, in which a near-infrared light (λ = 840 nm, from a light emitting diode) was polarized to enhance surface plasmon resonance. The light beam was reflected from the gold sensing surface and was directed by the gold mirror onto a linear array of silicon photo-diodes. The entire assembly

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Fig. 2. Idealized architecture showing the CCD immobilized in nanostructured LbL films.

was encased in an optically transparent material. Except for the Au-covered sensing surface, the sensor was coated with an opaque material to block out external light. The deposition process was followed in real time by making the PAMAM or CCD solutions to flow across the Au-covered sensor, using a flow cell and a peristaltic pump. The adsorption of each layer was monitored by the increase in the effective refractive index (ηeff ) (Jung et al., 1998). After complete deposition of each layer, the system was rinsed with the buffer washing solution previously described. Electrical measurements (ac) were performed with a Solartron 1260 impedance/gain phase analyzer in a frequency range from 1 Hz to 1 kHz. All the ac measurements were taken with the films (or sensing units) immersed in Milli-Q water, and in catechol solutions at different molar concentrations: (10−2 , 10−4 , 10−6 , 10−8 , and 10−10 M). To avoid any complexation between catechol and the amine groups from Tris buffer, which could influence or even hamper the enzymatic reaction, all the catechol solutions were prepared with MilliQ water. The capacitance curves were taken three times for each sensing unit, after 20 min of immersion in the analytical solutions. After each measurement, the sensor unit was gently rinsed with the buffer solution. For comparison, the same set of experiments was carried out using bare interdigitated electrodes and interdigitated electrodes covered with a five-bilayer PAMAM/PVS film (containing no CCD).

3. Results and discussion 3.1. Film assembly and morphology Fig. 3a shows the electronic absorption of PAMAM and CCD solutions used for film fabrication. CCD electronic absorption occurs at 280 nm. Film growth was followed by monitoring the increase in the absorbance at 280 nm, as shown in Fig. 3b. The growth process was almost linear (correlation coefficient r = 0.9917), as shown in the inset of Fig. 3b, indicating that the same amount of material was adsorbed at each deposition step. AFM images of

Fig. 3. (a) UV–vis spectra of 2 mg/mL PAMAM and CCD solutions (the same employed in film fabrication. (b) UV–vis spectra of PAMAM/CCD LbL films containing different numbers of bilayers. Inset: increase in the electronic absorption at 280 nm for a PAMAM/CCD film as a function of the number of deposited bilayers.

PAMAM/CCD films (not shown) revealed a globular morphology, similar to what has been reported for other LbL films (de-Souza et al., 2004; Zucolotto et al., 2005). Interestingly, all films presented a very smooth surface with root mean square (RMS) roughness of 0.5 and 0.8 nm for PAMAMand CCD-terminated LbL films, respectively, at an observation window of 1 ␮m × 1 ␮m (see Table 1). The secondary structure of the immobilized enzyme in PAMAM/CCD films containing 5 and 10 bilayers was investigated with circular dichroism (CD) analyses (Zheng et al., 2004) and compared to the enzyme spectra in soluTable 1 RMS roughness for a 4.5-bilayer PAMAM/CCD film (PAMAM-terminated) and for a five-bilayer film (CCD-terminated) Scale Sample

1 × 1 ␮m

600 × 600 nm

300 × 300 nm

PAMAM-terminated (nm) CCD-terminated (nm)

0.845 0.511

0.663 0.435

0.491 0.405

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Fig. 4. CD spectra of CCD solution (0.33 mg/mL) and of 5 and 10-bilayer PAMAM/CCD LbL films.

tion (Fig. 4). It should be mentioned that using CD does not replace activity tests. In this work, CD measurements were taken to investigate whether the secondary structure of CCD was preserved after immobilization. Specific activity tests were performed via UV–vis spectroscopy and will be discussed in the next sections. The spectra from the CCD solution (at 0.33 mg/mL) exhibited two minima centred at 208 and 217 nm and a negative to positive crossover at 193 nm. The latter indicates the presence of helical elements of secondary structure, in agreement to what has been reported (Ara´ujo et al., 2000). It is worth noting that the spectra of the LbL films were similar in shape to the spectra from CCD in solution, indicating that the CCD preserves its secondary structure after immobilization. To ensure that the CD signal from LbL films was entirely due to CCD, 10-bilayer PAMAM/PVS films were deposited on the top of two-bilayers of PAH/PVS. CD measurements showed no signal for this case. 3.2. Determination of PAMAM/CCD film thickness using SPR The early deposition stages of PAMAM or CCD layers were investigated in detail using in situ SPR measurements. The results are shown in Fig. 5 that depicts how the effective refraction index ηeff (Jung et al., 1998) of the adsorbed (layer + solution) evolves with time. A complete PAMAM layer is rapidly formed in less than 2 min whereas a CCD layer is completely adsorbed within 7–8 min, based on the saturation time of ηeff . Another important feature from Fig. 5 is the large desorption experienced by the PAMAM molecules during the washing step (with the addition of the buffer solution to the system), in comparison to the desorption of CCD. The reason for this desorption of PAMAM is not clear at the moment. Despite this partial desorption, the deposition of a PAMAM layer is crucial for film assembly, since the deposition of a CCD layer onto a previously adsorbed CCD layer was not possible (results not shown). The thickness of each

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Fig. 5. Effective refractive index versus time with measurements in situ for adsorption of PAMAM and CCD layers, as indicated in the figure. The arrows indicate the starting points for adsorption of each PAMAM or CCD layer.

adsorbed layer can be estimated from the ηeff values, using Eq. (1) (Jung et al., 1998): 

ld neff − ns (1) d=− ln 1 − 2 na − n s where ld is the characteristic decay length, and ηa and ηs are the refractive index of the adsorbed layer and solution, respectively. We used a refractive index of 1.45 and 1.57 for PAMAM (Wu et al., 2002) and CCD (Jung et al., 1998), respectively. ld can be calculated from Eq. (2): λ ld =

2π

(2)

1/2

Re[−n4eff /(n2eff + εm )]

where λ is the wavelength of the incident irradiation (840 nm) and εm is the complex dielectric constant of the metal. The results are summarized in Table 2. The thickness of the first and second PAMAM layers was estimated as 2.6 and 1.95 nm, respectively, before washing with the buffer solution. These values are slightly higher than that reported by Kim and Bruening (2003) who found an average thickness of 2.8 nm for a PAMAM/poly(acrylic acid) (PAA) bilayer. Similar results were reported by Tsukruk et al. (1997), who found a thickness of 2.8 nm per bilayer for films comprising amino- and carboxylic-terminated PAMAM generations 4 and 3.5, respectively. Although the average diameter for a PAMAM G4 dendrimer in solution is ca. 4 nm, as estimated from molecular models or from chromatography

Table 2 Thickness for each PAMAM or CCD layer adsorbed onto the Au-covered SPR sensing unit Thickness (nm) 1st PAMAM layer 1st CCD layer (on PAMAM) 2nd PAMAM layer (on PAMAM/CCD) 2nd CCD layer (on PAMAM/CCD/PAMAM)

2.6 3.6 1.95 3.6

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data of dendritic solutions (Tsukruk et al., 1997), it is well known that PAMAM forms a flattened layer after deposition (Tsukruk et al., 1997; Kim and Bruening, 2003). The thickness of the first and second CCD layers was estimated as 3.6 nm. In this case, a comparison with the literature is not straightforward, since the thickness of an adsorbed protein layer in LbL films may vary largely. For instance, in their seminal work on protein immobilization in LbL films, Lvov et al. (1995) reported an average thickness of 2.3 and 4.0 nm for a lysozyme and myoglobin layer, respectively, assembled with poly(styrenesulfonate) (PSS). On the other hand, in films containing glucose oxidase/poly(ethyleneimine) GOX/PEI, the thickness of the GOX layer was estimated as 34.4 nm from quartz crystal microbalance measurements (Lvov et al., 1995). 3.3. Activity of the immobilized CCD and optical detection of catechol In order to investigate the activity of the immobilized enzymes, PAMAM/CCD LbL films containing 10 bilayers were put in contact with catechol in such a way that the enzymatic activity was evaluated by monitoring the formation of cis–cis muconic acid. The latter was carried out using UV–vis spectroscopy, since the cis–cis muconic acid absorbs at 260 nm. The one-step enzymatic reaction under consideration is:

Ten-bilayer PAMAM/CCD films were deposited onto quartz substrates and stored for a day in a refrigerator at 5 ◦ C. For detection tests, the films were immersed in a catechol solution and immediately placed into the UV–vis spectrophotometer, where the absorbance at 260 nm was measured as a function of time. A similar PAMAM/CCD film containing the same number of bilayers and fabricated under the same conditions (but not immersed in catechol solution) was used as a reference. Fig. 6 shows the increase in absorption at 260 nm for a PAMAM/CCD film immersed consecutively into a 10−7 and 10−2 M catechol solution. Between the first and second experiments the film was placed into a 20 mM Tris buffer solution for 3 min. The results show that the PAMAM/CCD films were able to detect catechol at very low concentrations (10−7 M). At lower concentrations the increase in the absorbance was negligible. It is worth noting that at 10−2 M (upper curve) the reaction kinetics is similar to that exhibited by the protein in solution, as shown in the inset of Fig. 6, which is a standard method used to evaluate CCD activity (Nakazawa and Nakazawa, 1970; Ara´ujo et al., 2000). The latter confirms that the increase in absorbance is due to the formation of the cis–cis muconic acid. Similar tests were carried out in PAMAM/CCD films stored in a refrigerator for 3

Fig. 6. Temporal evolution of the absorbance at 260 nm for a 10-bilayer PAMAM/CCD film immediately after its immersion in catechol solutions at concentrations of 10−7 and 10−2 M, as indicated. The increase in absorbance is due to the formation of cis–cis muconic acid within the LbL film. The same analysis was carried out for a CCD solution, as shown in the inset, which depicts the increase in absorbance at 260 nm when catechol is incorporated in the CCD solution. Sample: 380 ␮L of 20 mM Tris buffer + 10 ␮L of 10−2 M catechol + 10 ␮L of 2 mg/mL CCD. Reference: 390 ␮L of 20 mM Tris buffer + 10 ␮L of 10−2 M catechol.

weeks. After this period, in which the films were not used, they were still able to detect catechol. 3.4. Sensing catechol with electrical measurements Fig. 7 shows the capacitance taken at 100 Hz for the three sensing units employed, viz., bare electrode, five-bilayer PAMAM/PVS film and five-bilayer PAMAM/CCD film. The rationale behind this approach was to detect the presence of catechol in diluted solutions through changes in the electrical capacitance of the film + solution system. The electrical response depends on several factors, including the geometric arrangement of the electrodes, the organic film and its interaction with the solution, particularly due to the electri-

Fig. 7. Capacitance at 100 Hz for unit sensors immersed in solutions with different catechol concentrations, as indicated, for three sensor units: bare electrode (a), five-bilayer PAMAM/PVS LbL film (b), and five-bilayer PAMAM/CCD LbL film (c).

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nations such as (bare electrode + five-bilayer PAMAM/PVS) or (bare electrode + five-bilayer PAMAM/CCD film) in the PCA analysis. The best distinguishing ability was achieved with the use of (bare electrode + five-bilayer PAMAM/PVS film + five-bilayer PAMAM/CCD film), corroborating the cross-sensitivity of the sensor employed. Additionally, all solutions were effectively separated in the PCA score plot of Fig. 8, with a high correlation between the first principal component and the catechol molar concentration.

4. Conclusions

Fig. 8. PCA plot for the same sensing units described in Fig. 8. The capacitance values were collected at 100 Hz in the presence of Milli-Q water (
), and catechol at concentrations of 10−10 M (䊉), 10−8 M (♦), 10−6 M (夽), 10−4 M () and 10-2 M ().

cal double-layer formed at the interface. Therefore, changes in the capacitance may arise from changes in any of these factors. For the frequency employed, 100 Hz, it has been suggested that the changes are mostly attributed to doublelayer effects, which are strongly dependent on the interaction between the film and the solution (Taylor and MacDonald, 1987). As the films used are at the nanometer scale, the electrical response is almost immediate when in contact with a liquid system, despite any swollen effect. That is an advantage of using ultra-thin films as transducing elements in such sort of application. Additionally, all measurements were taken after the samples had been immersed for 20 min in solution. As it can be seen (Fig. 7), each sensing unit responds differently when in contact with a solution. This difference in the electrical signal among the sensing units can be used as a fingerprint to identify the solution employed (Riul et al., 2003). As expected, the differences in the capacitance values for each concentration were higher for the electrode containing the immobilized enzyme, due to the specific affinity between catechol and CCD, pointing to a somewhat selective behavior. Thus, it is possible to infer that the PAMAM/CCD sensing units are suitable for catechol sensors–on their own–in solutions at concentrations as low as 10−10 M. It is known that the limit of detection of an array of sensors is always lower than that of a single sensor (Borato et al., 2004). In order to improve the performance of the sensing units, the capacitance data of each sensor was treated with principal component analysis (PCA), a mathematical tool that statistically correlates the samples in which the experimental data is presented as a matrix whose rows represent the number of experiments and the columns represent the number of sensing units used. The PCA plot in Fig. 8 shows capacitance values taken at 100 Hz, a frequency region ruled by the film/electrolyte interaction (Taylor and MacDonald, 1987; Riul et al., 2003), even with such thin films (Ferreira et al., 2003). We also checked combi-

Nanostructured films comprising CCD immobilized in conjunction with PAMAM dendrimers were successfully fabricated using the LbL technique. The films presented a very smooth surface with roughness of 0.5 and 0.8 nm for PAMAM- and CCD-terminated LbL films, respectively. The very early stages of film growth were monitored via SPR, revealing a thickness of 3.6 nm for the first and second CCD layers. Due to the mild, optimized immobilization conditions used here, CCD remained active in the films for periods longer than 3 weeks. The latter allowed the use of the PAMAM/CCD films as biosensors in an unprecedented way, in which catechol was detected in solutions at concentrations as low as 10−7 and 10−10 M, using an optical and electrical approach, respectively.

Acknowledgements The authors are grateful for the financial assistance from FAPESP and CNPq/IMMP (Brazil). We are also indebted to LNLS for providing us with interdigitated electrodes used from the project MIC2211.

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