18082
J. Phys. Chem. C 2008, 112, 18082–18086
Differential Adsorption of Silver Nanoparticles to the Inner and Outer Surfaces of the AgaWe americana Cuticle Avi Marciano,†,‡ Benny Chefetz,*,§ and Aharon Gedanken*,† Department of Chemistry, Kanbar Laboratory for Nanomaterials, Institute of Nanotechnology and AdVanced Materials, Bar-Ilan UniVersity, Ramat-Gan 52900, Israel, Department of Chemical Engineering and Biotechnology, College of Judea and Samaria, Ariel 44837, Israel, and Department of Soil and Water Sciences, The Hebrew UniVersity of Jerusalem, P.O. Box 12, RehoVot 76100, Israel ReceiVed: July 27, 2008; ReVised Manuscript ReceiVed: August 31, 2008
Plant cuticular materials have been recently described as highly efficient natural sorbents for organic pollutants; however, their role as adsorption agent for metal ions and metallic particles is unknown. The objective was to study the adsorption and fabrication of silver ions and nanoparticles by the AgaVe americana cuticle and its structural components. The two sides of the A. americana cuticle exhibited different behavior with respect to the adsorption of silver nanoparticles produced by microwave-assisted polyol reduction. Only the outside of the cuticle was found to be coated with silver nanoparticles. A mechanistic investigation showed that the first step in this reaction is formation of the nanoparticles in solution. Then the metallic nanoparticles were adsorbed by the cuticle. Silver ions were not adsorbed by the cuticle. Our data suggest that cutin and cutan biopolymers are the major cuticular fractions responsible for adsorption of the silver nanoparticles. Introduction Use of domestic multimode and single-mode microwave ovens for fabrication of inorganic nanomaterials is a wellestablished technique.1-6 Coupling microwave dielectric heating with a polyol reaction has recently led to fabrication of several metallic nanoparticles, including Ag, Pt, Au, and Cu, as well as binary and ternary chalcogenide nanoparticles.1-6 Polyols are considered reducing agents at their high boiling point, and welldefined metal nanoparticles are obtained when polyols are used in the reaction. The polyol process is completed in a much shorter time when conducted under microwave power versus reflux conditions.7 We recently demonstrated a one-step process involving the adsorption and reduction of metallic ions on the aquatic plant Azolla filiculoides, which can be accomplished within 3-5 min under microwave radiation.8 Moreover, our data showed that the A. filiculoides biomass provides the reducing agents needed to transform the adsorbed ions into the corresponding metallic nanoparticles, thus eliminating the need for polyol. In this study, our aim was to investigate the role of the plant cuticle as a sorption and reducing agent for silver ions. A feasible alternative for the use of activated carbon and ionexchange resin for removal of ion metals from aqueous solutions has been demonstrated by the use of dead plant biomass as an environmentally friendly sorbent.9-11 These studies reported that alfalfa biomass binds metal ions (including gold) in appreciable quantities.12 Metal ion binding by plant biomass is believed to take place through chemical functional groups such as carboxyl, amino, sulfhydryl, or hydroxyl groups.13 The plant cuticle is a thin layer comprised predominantly of lipids that covers all of the primary aerial surfaces of vascular plants.14-16 Its main physiological functions are to minimize water loss and protect the plant from physical, chemical, and * To whom correspondence should be addressed. E-mail: gedanken@ mail.biu.ac.il. † Bar-Ilan University. ‡ College of Judea and Samaria. § The Hebrew University of Jerusalem.
biological agents. The outer surface of the cuticle is covered with waxes: a complex mixture of long-chain aliphatic and cyclic components.17 In most plant species, the major structural component of the cuticle is cutin, a high molecular weight, insoluble, polyester-like biopolymer composed of various interesterified aliphatic hydroxy acids with alkyl chain lengths of C16 and C18.18 In some plant species (such as AgaVe americana), a base and acid hydrolysis-resistant biopolymer, known as cutan, is the major constituent of the cuticle layer together with cutin.16,19 Cutan is composed of an aromatic skeleton bonded via an ether bond to a long chain of n-alkenes and n-alkanes.20 In addition to these two aliphatic-rich biopolymers, the plant cuticle is composed of polysaccharides (mainly pectin) that are layered between the epidermal cell wall and the cuticle membrane.21 Since the cuticle of A. americana is composed of both cutin and cutan biopolymers it has been used as a model system to study the interactions of organic pollutants with aliphatic biopolymers.22-26 However, the role of plant cuticular matter as a reducing and an adsorption agent for metal ions and metallic particles is unknown. We hypothesized that the structural fractions of the plant cuticle can provide sites for metallic nanoparticle adsorption. Therefore, the objective of this research was to study adsorption and fabrication of silver ions and nanoparticles by the plant cuticle and its structural components. Experimental Section Cuticle sheets were peeled manually from fresh leaves of the succulent plant A. americana. Dewaxed cuticle was obtained by Soxhlet extraction of waxes from the bulk cuticle with chloroform/methanol (1:1, v/v) for 6 h. Cutin and cutan biopolymers were isolated and purified from the fruits of tomato (Lycopersicon esculentum Mill.) and leaves of A. americana, respectively, using the method described by Shechter et al.25 Pectin was purchased from Sigma-Aldrich. Adsorption of Ag+ ions by the bulk and dewaxed cuticles was monitored for up to 10 days. Dried bulk cuticle and
10.1021/jp806654a CCC: $40.75 2008 American Chemical Society Published on Web 10/23/2008
Differential Adsorption of Silver Nanoparticles
J. Phys. Chem. C, Vol. 112, No. 46, 2008 18083
Figure 1. XRD of silver nanoparticles coating the cuticle biomass after the microwave reaction.
Figure 2. TEM image of silver nanoparticles on a cuticle biomass (scale bar ) 500 nm).
dewaxed cuticle samples (0.75 g) were placed in a 100 mL glass flask containing 60 mL of aqueous or ethylene glycol solution containing 0.02 M Ag+ ions (the silver ion source was AgNO3, purchased from Sigma-Aldrich). The level of Ag+ ions adsorbed to the cuticular sheets was determined by calculating the difference between the Ag+ concentrations in solution before and after the reaction. These experiments were conducted in triplicate. The concentration of Ag+ ions in solution was determined by titration with a 0.01 M solution of KSCN in the presence of FeCl3 as an indicator according to the Folgard method.27 Adsorption of commercial silver nanoparticles (50 nm, P203 of Cima Nanotech, Caesarea, Israel) to the plant cuticle was tested under microwave radiation using a similar cuticle-to-solution ratio. Titration analysis was repeated three times for each adsorption experiment. The silver nanoparticles remaining in solution after adsorption were treated with HNO3 and titrated for the silver ions by KSCN as mentioned above. The percentage of silver nanoparticles embedded in the cucticle
was determined by subtracting the amount remaining after adsorption from the total amount of silver introduced in the solution. Microwave-oven reactions were conducted in 60 mL of 0.02 M Ag+-aqueous or ethylene glycol solutions containing cuticular matter (bulk cuticle, dewaxed cuticle, cutin, cutan, or pectin). The amount of cuticle and dewaxed cuticle used in the experiments was 0.75 g, while the amounts of the cuticular fractions cutin, cutan, and pectin were based on their relative levels in the bulk cuticle (50%, 20%, and 19%, respectively). The mixtures (60 mL of 0.02 M Ag+ and cuticular matter) were placed in an ordinary household microwave oven (Spectra 900 W, 2.45 GHz) modified with a refluxing system. The power level of the microwave oven was maintained at 30%. Before performing the reaction, the solution was purged for 30 min with Ar gas. The reaction was tested for 10 min of irradiation time. Following the reaction, the cuticular sheets were filtered and washed with double-distilled water and then dried under vacuum. Each experiment involving the microwave oven was conducted three times. Control experiments were performed as followed: microwave-oven reactions without cuticle, without silver ions, and with an aqueous solution of silver ions without ethylene glycol. Reactions with cuticle and silver ions but without microwave radiation were also conducted. X-ray diffraction (XRD) measurements were performed with a Bruker AXS instrument (model D8) using Cu KR radiation. The diffused reflection optical spectra (DRS) were recorded on a Cary 100 Scan UV spectrometer (Melbourne, Australia) in a wavelength range of 350-600 nm. High-resolution scanning electron microscopy (HRSEM) images were captured in a Leo Gemini 982 Field Emission Gun scanning electron microscope (Oberkochen, Germany) operating at a 4 kV accelerating voltage. Transmission electron microscopy (TEM) micrographs were obtained with a JEOL-TEM 100SX microscope. Results and Discussion Silver ions did not adsorb to the studied cuticle when introduced into aqueous or ethylene glycol solutions. More than 99% of the added ions (0.02 M) remained as free ions in both solutions for a period of 10 days. This behavior differentiates the cuticle from the aquatic plant A. filiculoides, which is known to adsorb heavy-metal ions.28-32 Moreover, when the silver ions were introduced with the cuticle in an aqueous solution and irradiated in a microwave oven, there was still no observable formation of nanoparticles. This suggested that unlike the biomass of A. filiculoides, the cuticle does not contain any native reducing agents capable of reducing silver under microwave
18084 J. Phys. Chem. C, Vol. 112, No. 46, 2008
Marciano et al.
Figure 3. TEM image of silver nanoparticles on a cuticle biomass with a scale bar of 50 nm (right side) and a histogram of nanoparticle sizes (left side).
Figure 4. Image of the two surfaces ((A) outer and (B) inner) of A. americana cuticle after polyol reaction with silver ions. The two pictures (left and right) illustrate the two sides of each cuticle sample.
Figure 5. SEM images of the two surfaces ((left) outer and (right) inner) of A. americana cuticle after polyol reaction with silver ions.
radiation. However, when the polyol reaction was used under microwave radiation, adsorption and reduction of Ag+ ions to the corresponding nanometallic particles were observed with the studied cuticle. More than 75 ( 4% of the Ag+ ions were reduced to metallic silver and removed from the solution after 10 min of reaction. The calculated adsorption capacity of the cuticle biomass to Ag nanoparticles was 127 ( 7 mg/g. A higher loading capacity of 171 ( 9 mg/g had been observed with the A. filiculoides biomass. It is worth mentioning that under all microwave treatments the cuticle materials were kept intact. Our data (XRD measurements) showed that Ag+ ions were transformed to Ag nanoparticles on the cuticle’s biomass, but
it did not tell us the order in which this transformation occurs: (a) adsorption of the Ag+ ions to the cuticle surface followed by their reduction to nanometallic silver or (b) reduction of the Ag+ ions to nanometallic particles in solution and then adsorption of the nanometallic particles to the cuticle’s surface. Our adsorption experiments conducted with Ag+ ions in aqueous and ethylene glycol solutions without microwave radiation revealed no adsorption of the Ag+ ions to the cuticle’s surface, even after 10 days. This suggested that the Ag nanometallic particles’ adsorption to the cuticle surface may be due to their high reactivity toward the plant. This hypothesis was checked using commercial silver nanoparticles (Cima Nanotech, 60 nm
Differential Adsorption of Silver Nanoparticles
Figure 6. DRS results of the outer and inner surfaces of the cuticle without wax after the polyol reaction. Only the outer surface exhibits a silver plasmon band peak at a wavelength of 425 nm.
size average). When these were interacted with the cuticle biomass, 34 ( 2% of the added materials adsorbed onto the cuticle. We therefore concluded that the Ag+ ions in ethylene glycol solution are first reduced to nanometallic particles in the solution by microwave radiation and then adsorbed onto the surface of the cuticle. The XRD pattern of the product of the polyol reaction of Ag+ with the cuticle is presented in Figure 1. The major diffraction peaks are assigned to pure metallic cubic silver based on the exact match with PDF: 4-783. No impurities, such as silver oxide, were observed. However, a very small diffraction peak at 2θ ) 21.6° was measured and assigned to the mineral residues of the plant biomass (XRD data not shown). TEM micrographs of the silver nanoparticles on the surface of the cuticle after microwave radiation are presented in Figures 2 and 3. Spherical nanoparticles are clearly observed in the latter figure. The size of the silver nanoparticles ranges from 5 to 50 nm with an average size of 20 nm. These sizes are a bit smaller than the previously reported data about the polyol reaction under microwave without cuticle.4 Silver particles were either aggregated on the cuticle surface or aggregated in solution and then adsorbed on the surface of the cuticle. The results of the polyol reaction carried out under microwave radiation revealed that only the outer (externally facing) surface of the cuticle was covered with silver nanoparticles (Figure 4). The silver was not deposited on its inner surface (i.e., the side of the cuticle that faces the epidermal cells) (Figure 4). This is more clearly illustrated in the HRSEM images (Figure 5), showing the outer surface of the cuticle covered with silver nanoparticles and no anchoring of silver particles to the epidermal cells on the inner surface. Since the outer surface of the cuticle is covered with epicuticular waxes, it was hypothesized that these assist in adsorption of the silver nanoparticles. To verify this, the waxy layer was removed and polyol-assisted microwave radiation was applied: 98% of the Ag+ ions adsorbed to the cuticle within 10 min of the reaction, but again, deposition was only present on its outer surface. This suggested that the epicuticular waxes do not play an important role in controlling the preferred sorption to the outer surface of the cuticle. However, wax removal did result in enhanced adsorption: 169 ( 8 mg of Ag/g cuticle as compared to 127 ( 7 mg/g before removal. Those results suggested that adsorption of the silver nanoparticles is probably governed by the cutin and cutan biopolymers and the pectin layer present on the inner-facing cuticle layer. Therefore, we performed the polyol-assisted microwave radiation experiments with silver ions and the following cuticular fractions: cutin, cutan, and pectin. In these experiments, 62 ( 3% of the Ag+ ions were removed (reduced
J. Phys. Chem. C, Vol. 112, No. 46, 2008 18085 and adsorbed) by the cutin, while the cutan was less efficient, facilitating removal of only 25 ( 3% of the silver ions. Silver ions were not affected by the presence of pectin. We therefore concluded that cutin is the major structural fraction of the cuticle that facilitates adsorption of silver nanoparticles, while the epicuticular waxes reduce the accessibility of the silver nanoparticles to the cutin. The presence of pectin on the inner surface of the cuticle probably prevents adsorption of silver nanoparticles to that surface. The mode of interaction of the silver nanoparticles with the cutin is similar to that of silver nanoparticles and thiols (Jiang, X. C.; Zeng, Q. H.; Yu, A. B. Langmuir 2007, 23, 2218). It is suggested that silver interacts with the aliphatic hydroxy acids groups of the cutin, which results in formation of Ag1+ and COO-1 ions, after elimination of 1/2H2. The results of the diffused reflectance optical spectra (DRS) on the outer surface of the cuticle without wax after the polyol reaction illustrate the presence of the plasmon band of the silver at 425 nm (Figure 6). Its position is similar to the characteristic plasmon band of an aqueous colloidal silver solution at 400 nm. After the polyol reaction we did not observe any band at 425 nm, characteristic of silver, in the reflection of the cuticle’s inner surface. Without the polyol, namely, when an aqueous solution of silver underwent microwave irradiation, we could not find any bands characteristic of silver on either side of the cuticle. On the other hand, as mentioned above, the outer surface showed the plasmon resonance of silver. The interaction between the silver nanoparticles and the cutin and cutan biopolymers is accelerated by microwave radiation. Moreover, this interaction is carried out in hot spots, which are formed by the strong absorption of the microwave radiation by the metallic nanoparticles. The dipole moment of cutin, which contains aliphatic hydroxy acids, can also absorb microwave radiation, thereby facilitating the adsorptive interactions with the metallic nanoparticles. Under the MW radiation the silver reacts with aliphatic hydroxy acids, yielding a silver plus ion, while the negative charge resides on the aliphatic hydroxy acid. The reaction is carried out in superheated polyol solvent. All of these factors lead to adsorption of the silver nanoparticles to the cuticle’s surface. References and Notes (1) Komarneni, S.; Li, D. S.; Newalkar, B.; Katsuki, H.; Bhalla, A. S. Langmuir 2002, 18, 5959–5962. (2) Pastoriza-Santos, I.; Liz-Marzan, L. M. Langmuir 2002, 18, 2888– 2894. (3) Harpeness, R.; Gedanken, A. Langmuir 2004, 20, 3431–3434. (4) Katsuki, H.; Komarneni, S. J. Mater. Res. 2003, 18, 747–750. (5) Zhu, J. J.; Palchik, O.; Chen, S. G.; Gedanken, A. J. Phys. Chem. B 2000, 104, 7344–7347. (6) Grisaru, H.; Palchik, O.; Gedanken, A.; Palchik, V.; Slifkin, M. A.; Weiss, A. M. Inorg. Chem. 2003, 42, 7148–7155. (7) Fievet, F.; Lagier, J. P.; Figlarz, M. MRS Bull. 1989, 29–34. (8) Gedanken, A.; Tel-Or, E.; Chefetz, B.; Elmeshaly, S. Method of removal of heavy metal ions from water. PCT Int. Appl. AN 2007:817363, 2007. (9) Gardea-Torresdey, J. L. K. J.; Tiemann, K. J.; Gonzalez, J. H.; Henning, J. A.; Towsend, M. S. J. Hazard. Mater. 1996, 48, 81–190. (10) Gardea-Torresdey, J. L.; Tiemann, K. J.; Gonzalez, J. H.; Rodriguez, O.; Gamez, G. J. Hazard. Mater. 1998, 57, 29–39. (11) Gardea-Torresdey, J. L.; Tiemann, K. J.; Gamez, G.; Dokken, K. J. Hazard. Mater. 1999, 69, 41–51. (12) Gamez, G.; Gardea-Torresdey, J. L.; Tiemann, K. J.; Parsons, J.; Dokken, K.; Yacaman, M. Jose. AdV. EnViron. Res. 2003, 7, 563–571. (13) Gardea-Torresdey, J. L.; Becker-Hapak, M. K.; Hosea, J. M.; Darnall, D. W. EnViron. Sci. Technol. 1990, 24, 1372–1378. (14) Nierop, K. G. J. Org. Geochem. 1998, 29, 1009–1016. (15) Ko¨gel-Knabner, I. Soil Biol. Biochem. 2002, 34, 139–162.
18086 J. Phys. Chem. C, Vol. 112, No. 46, 2008 (16) Jeffree, C. E. Structure and Ontogeny of Plant Cuticles. In Plant Cuticles: An Integrated Functional Approach; Kerstiens, G., Ed.; BIOS Scientific Publishers: Oxford, U.K., 1996; pp 33-82. (17) Santier, S.; Chamel, A. Plant Physiol. Biochem. 1998, 36, 225– 231. (18) Kolattukudy P. E. Cutin from plants. In Biopolymers; Doi Y, Steinbu¨chel A, Eds.; Wiley-VCH: Weinheim, 2001; Vol. 3a (Polyesters I: Biological systems and biotechnological production), pp 1-40. (19) Villena, J. F.; Dominguez, E.; Stewart, D.; Heredia, A. Planta 1999, 208, 181–187. (20) Boom, A.; Damste, J. S. D.; Leeuw, J. W. Org. Geochem. 2005, 36, 595–601. (21) Heredia, A. Biochim. Biophys. Acta 2003, 1620, 1–7. (22) Chefetz, B. Toxicol. Chem. 2003, 22, 2492–2498. (23) Sachleben, J. R.; Chefetz, B.; Deshmukh, A.; Hatcher, P. G. EnViron. Sci. Technol. 2004, 38, 4369–4376. (24) Chen, B. L.; Johnson, E. J.; Chefetz, B.; Zhu, L.; Xing, B. EnViron. Sci. Technol. 2005, 39, 6138–1641.
Marciano et al. (25) Shechter, M.; Xing, B.; Kopinke, F. D.; Chefetz, B. J. Agric. Food Chem. 2006, 54, 7761–7768. (26) Stimler, K.; Xing, B.; Chefetz, B. Soil Sci. Soc. Am. J. 2006, 70, 1101–1109. (27) Kolthoff, I. M.; Sandell, E. B. Textbook of QuantitatiVe Inorganic Analysis, 3rd ed.; Macmillan: New York, 1958. (28) Kadirvelu, K.; Faur-Brasquet, C.; Le Cloirec, P. Langmuir 2000, 16, 8404–8409. (29) Oliveira, L. C. A.; Petkowicz, D. I.; Smaniotto, A.; Pergher, S. B. C. Water Res. 2004, 38, 3699–3704. (30) Wase, D. A. J.; Forster, C. F. Biosorbents for Metal Ions; Taylor and Francis: London, 1997. (31) Ajmal, A.; Rao, A. K. R.; Rais, A.; Jameel, A. J. Hazard. Mater. B 2000, 79, 117–131. (32) Chefetz, B.; Sominski, L.; Pinchas, M.; Ginsburg, T.; Elmachliy, S.; TelOr, E.; Gedanken, A. J. Phys. Chem B. 2005, 109, 15179–15181.
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