CHEMSUSCHEM FULL PAPERS DOI: 10.1002/cssc.201402126
Unveiling the Chemistry behind the Green Synthesis of Metal Nanoparticles Snia A. O. Santos,*[a] Ricardo J. B. Pinto,*[a, b] Slvia M. Rocha,[c] Paula A. A. P. Marques,[b] Carlos Pascoal Neto,[a] Armando J. D. Silvestre,[a] and Carmen S. R. Freire[a] Nanobiotechnology has emerged as a fundamental domain in modern science, and metallic nanoparticles (NPs) are one of the largest classes of NPs studied because of their wide spectrum of possible applications in several fields. The use of plant extracts as reducing and stabilizing agents in their synthesis is an interesting and reliable alternative to conventional methodologies. However, the role of the different components of such extracts in the reduction/stabilization of metal ions has not yet
been understood clearly. Here we studied the behavior of the main components of a Eucalyptus globulus Labill. bark aqueous extract during metal-ion reduction followed by advanced chromatographic techniques, which allowed us to establish their specific role in the process. The obtained results showed that phenolic compounds, particularly galloyl derivatives, are mainly responsible for the metal-ion reduction, whereas sugars are essentially involved in the stabilization of the NPs.
Introduction Nanometer-scale metallic particles are probably one of the main classes of nanomaterials investigated and used currently because of the wide spectrum of potential applications in several fields that arises from their unique and distinct properties in relation to the bulk counterparts.[1] The recent advances in the fields of nanoscience and nanotechnology lead to a variety of physical and chemical strategies for the selective preparation of inorganic nanoparticles (NPs) with precise control over their shape and dimensionality.[2] Among the chemical methodologies, those used most commonly are based on soluble metal salt precursors and different reducing agents, which can act also as stabilizers to avoid the coalescence of the NPs. However, most of the reducing agents reported, for example, hydrazine, sodium borohydride, or N,N-dimethylformamide, are reactive chemicals that are commonly associated with huge environmental risks and toxicity.[3] [a] Dr. S. A. O. Santos,+ Dr. R. J. B. Pinto,+ Prof. Dr. C. P. Neto, Prof. Dr. A. J. D. Silvestre, Dr. C. S. R. Freire Department of Chemistry-CICECO University of Aveiro 3810-193 Aveiro (Portugal) Fax: (+ 351) 234-401-470 E-mail: [email protected] [email protected] [b] Dr. R. J. B. Pinto,+ Dr. P. A. A. P. Marques TEMA-NRD, Mechanical Engineering Department and Aveiro Institute of Nanotechnology (AIN) University of Aveiro 3810-193 Aveiro (Portugal) [c] Prof. Dr. S. M. Rocha Department of Chemistry-QOPNA University of Aveiro 3810-193 Aveiro (Portugal) [+] These authors contributed equally to this work. Supporting information for this article is available on the WWW under http://dx.doi.org/10.1002/cssc.201402126.
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The green synthesis of metal NPs has been studied as a reliable and promising alternative to minimize or eliminate the use of these harmful substances.[4] Green synthetic processes comprise either micro-organisms, such as bacteria[5] or fungi,[6] as well as plant biomass[7] and several plant extracts.[8] The use of plant extracts from different morphological parts, which include leaves, fruits, seeds, and bark, presents some advantages over the use of micro-organisms because of the high diversity and abundance of plant extracts and the simplicity, easy scaleup, and cost-effectiveness of the method.[4, 9, 10] Despite the remarkable number of reports in the literature on the synthesis of metal NPs using distinct plant extracts, as highlighted recently by Iravani,[10] a precise understanding of their formation mechanism and the clear-cut role of the natural compounds involved in this process is still imprecise. Some reports have put forward hypothetical mechanisms that propose that the reduction of the metal ions to metal NPs may be caused by the distinct compounds present in the extracts, such as reducing sugars, phenolic compounds (e.g., flavonoids), and proteins.[9, 11] However, these generic suggestions have been based essentially on qualitative routine spectrophotometric analysis, such as FTIR spectroscopy, colorimetric assays, or UV spectroscopy,[12] which cannot unambiguously differentiate the presence or absence of the different families of compounds in the extracts and resulting NPs. In this vein, and in consideration of the relevance of such fundamental knowledge, particularly in the optimization and control of the final properties of the NPs and on the up-scaling of these green processes, in the present study we intended to clarify the role of the different components of a plant extract in a NP formation mechanism, kinetics, and stabilization. Here, an aqueous extract of Eucalyptus globulus Labill. bark, a typical residue from the pulp and paper industry that is recognized for its high content of phenolic compounds,[13] was ChemSusChem 2014, 7, 2704 – 2711
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chosen as a reducing and stabilizing system for Ag and Au NPs synthesis. Ag and Au NPs were selected because of their potential application in many fields, such as catalysis, sensors, and in the biomedical field as, for example, drug-delivery systems and antimicrobial agents.[14] Furthermore, among the noble-metal NPs investigated so far, Au and Ag NPs have been studied in particular, which allows a comparative and critical discussion of the results. The clear identification of the natural reducing agents involved in the formation of the metal NPs was achieved by the qualitative and quantitative analysis of sugars and phenolic compounds present in the plant extracts before and after Ag and Au NP synthesis by using advanced chromatographic techniques.
Results and Discussion Synthesis of Au and Ag NPs using E. globulus bark extract In this study, comparative experiments using E. globulus aqueous extracts as a reducing and stabilizing agent in the preparation of Ag and Au hydrosols were performed to elucidate the role of the different components of plant extracts in green processes. UV/Vis spectroscopy was used to confirm the presence of the characteristic surface plasmon resonance (SPR) bands of Ag and Au nanospheres in the visible region[15] and, therefore, the successful formation of the metal NPs. UV/Vis spectra of the Au and Ag aqueous colloidal solutions obtained after the addition of the bark extract as a function of time are shown in Figure 1 (the E. globulus extract does not present any absorption bands in the visible region at the concentration used). The addition of the bark extract to the Au salt solution caused an almost instantaneous change of color from light yellow to red, which indicated the effectiveness of the Au salt reduction. As expected, the obtained Au colloidal solution showed the typical absorption band at around 530 nm without any significant shift over time, except for a gradual increase of intensity, most noticeable in the first minutes of the reaction. Similarly, the addition of the E. globulus extract to the Ag salt solution led to the formation of Ag NPs, which followed a slower kinetics (Figure 1). One hour after the addition of the extract, the UV/Vis spectrum showed two weak and broad absorbance bands at around l = 375 and 445 nm. This can be explained by the variances in terms of composition and experimental conditions tested in this synthesis, which leads to a wide range of sizes of the particles as the broadening of the plasmon band is not always observed in the case of Ag NPs.[8, 16] Nevertheless, a significant increase of the intensity of these bands with the reaction time was observed, which shows that the formation of the Ag hydrosol is considerably slower than that of the Au counterpart using the same extract as the reducing agent. This dissimilarity can be explained by the different reduction potentials of the two metal ions; the standard reduction potential of Ag+ to Ag0 (E0 = 0.799 V) is substantially lower than that of Au3+ to Au0 (E0 = 1.498 V).[17] Scanning transmission electron microscopy (STEM) images of the Au and Ag NPs obtained by reduction with E. globulus 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Figure 1. UV/Vis spectra of a) Au and b) Ag colloidal solutions at distinct times after the addition of the E. globulus aqueous extract. Inset: detail of the differences observed for the Au colloidal solution.
bark extract after 1 h of reaction are shown in Figure 2. The Au NPs exhibited a homogeneous distribution of well-defined nanoparticles with a predominantly spherical morphology and a narrow size distribution (average diameter of (18 3) nm); however, some anisotropic planar nanostructures such as triangles or hexagonal-shaped particles were also observed. Conversely, the Ag NPs showed a spherical and polydisperse distribution with average diameters in the range of 15–73 nm, which indicates that this sample is composed of a large quantity of nonuniform NPs and corroborates the broad plasmon resonance band observed by UV/Vis spectroscopy. Dynamic light scattering (DLS) analysis of these colloids revealed an average size of (21 4) and (52 16) nm for Au and Ag NPs, respectively (Figure S1). These values are quite similar to those obtained by STEM analysis, and the slight differences are because DLS analysis measures the hydrodynamic diameter, which includes the hydration layer. The Zeta potential values of the colloidal solutions were ( 16 6) (pH 2.74) and ( 18 7) mV (pH 3.48) for Au and Ag NPs, respectively. These results indicated that the surface of both metal NPs is composed mainly of negatively charged capping compounds, which are responsible for their moderate stability. This is in close agreement with data reported previously that suggest that the biomolecules present in natural extracts serve both as reducing and capping agents on metal NPs prepared by green synthesis.[18] The crystalline structure of the obtained Au and Ag NPs was confirmed by powder XRD analysis of dried samples (FigChemSusChem 2014, 7, 2704 – 2711
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www.chemsuschem.org sugars of the bark extracts were investigated before and after NPs formation.
Identification of the biomolecules responsible for Au and Ag NPs synthesis and stabilization using E. globulus bark extract The role of proteins
Figure 2. STEM images of a) Au and b) Ag NPs with the respective histograms of size distribution obtained after reduction (1 h) with the E. globulus aqueous extract.
ure S2). The observed diffraction peaks can be indexed to the (111), (2 0 0), (2 2 0), and (3 11) crystalline planes of Au and Ag face-centered cubic (fcc) structures.[8, 12b] The FTIR spectra of the pristine E. globulus aqueous bark extract (Figure S3) showed several absorption bands located in the n˜ = 1000–1800 cm 1 region that could be assigned to the vibrations of typical functional groups, namely, C O C, C O, C=C, and C=O, present in several water-soluble components of E. globulus bark, which include phenolic compounds, proteins, and sugars.[19] Some of these absorption bands were also observed in the spectra of the obtained metal NPs, which confirms clearly that some biomolecules also act as capping agents and, therefore, play an important role on the NPs stabilization. Although most studies[8, 17a] based their conclusions and proposals in terms of reduction mechanisms on the results of FTIR spectroscopy studies, it is impossible to distinguish the different families of compounds and their contribution to the reduction and/or stabilization processes unambiguously. To assess the role of the different extract components in the Au and Ag NPs green synthesis and stabilization, the protein content and the composition of the phenolic compounds and 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Several authors have attributed the reducing capacity of plant extracts in NPs synthesis to proteins,[12b] although most of the extracts used have been obtained by employing conditions that do not commonly allow protein removal or promote their degradation (e.g., the absence of an appropriate buffer and/or mechanical processes suitable to disrupt the cell walls).[20] Nonetheless, we decided to verify the presence of proteins in the extracts studied here, based on the N content determined by elemental analysis,[21] and to access their possible participation in the NPs synthesis. The protein content of the E. globulus bark extract determined by elemental analysis represented approximately 0.8 wt % of the total extract, which corresponds to a concentration of approximately 19.0 mg mL 1 of the aqueous extract. This value is extremely low (4- and 19-fold lower than the phenolic compounds and sugars content, respectively). Additionally, this value is in a range of concentration that was reported not to have a linear relationship with Au NPs synthesis.[12a] Moreover, the protein content determined by elemental analysis was probably overestimated because other extract components could also contribute to the N content. Therefore, this result suggested clearly that proteins are not involved in metal-ion reduction under the conditions studied. However, they might play an important role in the stabilization of the NPs. To corroborate this, the content of N in the NPs was also evaluated, and 29.3 and 14.6 % of the N content of the E. globulus bark extract was aggregated in the Ag and Au NPs, respectively. The capping/stabilization ability of proteins is certainly associated with the affinity of carbonyl groups from amino acid residues of proteins to bind metals.[22] The role of sugars The content of monosaccharides in the E. globulus extract, before and after NPs synthesis, was determined by HPLC with refractive index detection (RI). Only two monosaccharides were detected in this extract, namely, glucose and fructose, which together represent 157.1 mg g 1 of extract with concentrations of 127.7 and 233.7 mg mL 1 of aqueous extract, respectively. These concentrations are in the same range as those reported for several vegetables, fruits, seeds, and other plant extracts,[23] which are used commonly in the biogenic synthesis of NPs.[10, 24] After the Ag and Au NPs synthesis, a considerable decline in the content of both monosaccharides in the extracts was observed. The glucose concentration decreased to 52.7 and 71.4 mg mL 1 in the extract after Ag and Au NPs synthesis, respectively, whereas the fructose content decreased to 114.4 and 119.7 mg mL 1 in the extract. This first outcome suggests the involvement of these reducing sugars in the reduction of ChemSusChem 2014, 7, 2704 – 2711
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1.47 0.07 1.11 0.01 0.63 0.02 – 0.50 0.01 – – – 0.19 0.01 – – – – – – – 3.90 0.08
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[a] Rt = retention time. [b] m/z values in bold were subjected to MSn analysis. [c] Identified by co-injection of the respective authentic standard. [d] ntdm—not determined.
2.30 0.08 3.10 0.16 1.97 0.060 5.15 0.25 1.30 0.07 1.18 0.06 1.56 0.030 0.33 0.01 4.87 0.22 3.41 0.17 1.16 0.06(11+12) – 3.42 0.16 6.00 0.30 0.29 0.01 1.40 0.06 37.45 1.18 3.17 0.12 4.43 0.22 2.34 0.10 11.13 0.43 1.84 0.07 2.34 0.11 2.38 0.10 0.84 0.03 10.17 0.25 4.43 0.14 1.65 0.05(11+12) – 7.37 0.29 14.27 0.55 0.44 0.01 3.45 0.12 70.27 1.75 [13b] [26b] [26a] Co[c] [32] [13b] [13b] [13b] [13b] [26a] [26a] [26a] [13b] [26a] [13b] [26a] 173, 171, 111, 93, 85 301,[b] 275, 257, 229; MS3 : 284, 257, 229, 185 271, 241, 211, 169; MS3 : 125 125 591, 483, 441, 423, 305; MS3 : 423 481, 301; MS3 :301, 275 481, 301; MS3 : 301, 275 301; MS3 :229 633, 301; MS3 : 463, 299, 275 482, 327, 313, 183, 169; MS3 : 169, 125 301, 300; MS3 : 284, 257, 229, 185, 184 301, 300; MS3 : 284, 257, 229, 185 315, 314; MS3 : 300; MS4 : 244, 200 315, 300; MS3 : 300; MS4 : 271, 244, 243, 216, 200 315; MS3 : 300; MS4 : 244 315, 300; MS3 : 300; MS4 : 272, 271, 244, 243 216, 200 MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : 191 481 331 169 609 783 783 633 935 497 447 433 477 447 461 447 2.73 250 quinic acid 2.82 244, 270 HHDP-glucose 3.03 240 (sh), 270 galloylglucose 3.54 269 gallic acid 3.90 238, 270 gallocatechin dimer 4.59 238, 265(sh) bis-HHDP-glucose isomer 5.24 238, 265(sh) bis-HHDP-glucose isomer 7.31 237, 270 galloyl-HHDP-glucose 17.62 239, 269 galloyl-bis-HHDP-glucose 30.87 236, 273 eucaglobulin ellagic acid rhamnoside 31.73 ndtm[d] 31.96 ndtm ellagic acid-pentoside 33.29 245, 364 isorhamnetin-hexoside 43.07 249, 364 isorhamnetin-pentoside isomer 43.98 237, 368 isorhamnetin-rhamnoside 44.47 244, 365 isorhamnetin-pentoside isomer total [mg mL 1 of extract] 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
Identification Phenolic compounds in E. globulus bark extract [mg mL 1 extract] before NPs synthesis after Ag NPs after Au NPs MSn product ions [m/z] [M H] [m/z] Compound
The aqueous extract of E. globulus bark extract was characterized by HPLC with diode array detection coupled to tandem mass spectrometry (HPLC–DAD–MSn), which allowed us to identify 16 phenolic compounds (Figure S7). The compounds identified in E. globulus aqueous extract, their retention time, UV absorption maximum (if possible), the molecular ion [M H] , and the corresponding MSn product ions are summarized in Table 1. Compounds were identified by comparing their fragmentation profiles with those of authentic standards or, if these were not available, with published data, as indicated in Table 1. Most of the phenolic compounds identified in this E. globulus bark aqueous extract were previously described as constituents of this Eucalyptus species[13b] or even of plant tissues of other Eucalyptus species.[26] The phenolic fraction of E. globulus bark aqueous extracts is composed mainly of galloylglucose, ellagic acid, and isorhamnetin derivatives (Figure S7), which accounted for a total amount of 30.55 mg of phenolic compounds per gram of dry extract and corresponds to a concentration of 70.27 mg mL 1 of aqueous extract used for the NPs synthesis (Table 1). Among the 16 phenolic compounds identified in the extract (Figure S7), an isomer of isorhamnetin-pentoside (14), gallic acid (4), and galloyl-bis-HHDPglucose (9; Figure 3) are the major components with concentrations of (14.27 0.55), (11.13 0.43), and (10.27 0.25) mg mL 1 of aqueous extract, respectively. The amount of phenolic compounds in the Eucalyptus bark aqueous extract
Peak no. Rt[a] lmax [min] [nm]
The role of phenolic compounds
Table 1. HPLC–MSn fragmentation profile and abundance of phenolic compounds identified in E. globulus aqueous extract before and after Ag and Au NPs synthesis.
Ag+ and Au3+ or in the stabilization of the obtained NPs. Indeed, the participation of reducing sugars, such as glucose, in the reduction of metal ions in NPs synthesis using plant extracts has been suggested previously, and several mechanisms have been proposed;[22] however, most authors neglect the coexistence of other reducing agents in these types of plant extracts. Thus, to verify the specific role of glucose and fructose in the NPs synthesis, standard solutions with the same concentration as that observed in the bark extract (0.13 and 0.23 mg mL 1 for glucose and fructose, respectively) were also tested. After 1 h of reaction, no metal-ion reduction was perceived with glucose or fructose standard solutions, even if the two reducing sugars were mixed together (Figures S4 and S5). This behavior could be related essentially to the relatively low concentration of these reducing sugars in the bark extract. Ortega-Arroyo et al.[25] used glucose as the reducing agent in the synthesis of Ag NPs successfully by employing considerably higher concentrations than those used in this work (between 18- and 108-fold). Notwithstanding, this reaction could be also affected by the kinetic rate of the redox reactions as the formation of a low quantity of Au NPs was observed both with glucose and fructose after one month, confirmed by the presence of the UV band at l = 590 nm (Figure S6). In the case of Ag (which has a reduction potential lower than Au) even after one month no NPs formation was observed by UV spectrophotometry. These results indicated that sugars are essentially involved in the NPs stabilization, as will be discussed below.
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Figure 3. Structures of the main phenolic compounds responsible for NPs synthesis (see number assignments in Table 1).
used in this work is very similar to other aqueous extracts used previously in NPs synthesis,[24] such as that from Pelargonium graveolens leaves.[27] The phenolics content of the E. globulus bark extract before and after NPs synthesis was quantified by HPLC with UV detection and is expressed in mg mL 1 of extract (Table 1). After Ag NPs synthesis, the content of phenolic compounds decreased to 37.97 mg mL 1 (Table 1), and this decline was more accentuated for isorhamnetin and phenolic acid derivatives with decreases of 54 and 48 %, respectively. Specifically, the concentrations of 14 and 4 diminished the most. In contrast, after Au NPs synthesis the majority of the phenolic compounds present initially in the E. globulus bark disappeared (accounting only for 4.36 mg mL 1), of which only quinic acid (1), HHDP-glucose (2), galloylglucose (3), gallocatechin dimer (5), and 9 were detected (Figure S8). These results indicate that phenolic compounds play a dominant role in this green process. In this sense, to confirm their specific starring role in NPs synthesis, gallic and ellagic acids and isorhamnetin standards were tested individually by using equivalent contents to those found in the bark extract. In the Ag NPs synthesis, gallic acid showed the fastest redox kinetics (Figure S9). In the Au NPs synthesis, after the addition of gallic acid, the color of the HAuCl4 solution changed in a few seconds from light yellow to pink, identical to that observed in the case of the E. globulus extract, which indicates the successful synthesis of Au NPs. However, no metal reduction was observed if ellagic acid or isorhamnetin was added to the Au solution (Figure S10). Once more, the faster reduction rate of Au ions than that of Ag ions is most likely because of significant differences in the reduction potentials of the two metal ions, as observed with the total extract. These observations established that phenolic compounds, in particular derivatives of gallic acid, are mainly responsible for metal-ion reduction in both Ag and Au NPs synthesis using E. globulus bark extracts. Moreover, in the case of Au NPs, the presence of ellagic acid and isorhamnetin seems also to increase their stability as the Zeta potential of Au NPs changed 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
from 29.0 mV in the presence of gallic acid to 36.2 mV if the three phenolic compounds were used together (Table S2). These observations explain the remarkable decrease of the content of all phenolic compounds after the Au NPs synthesis. Consequently, phenolic compounds, and in particular galloyl derivatives, would serve as the main electron donors in the redox reaction. Phenolic acids, galloyl derivatives, and flavonoids are known to form quinones upon oxidation and to undergo further oxidation to form acids.[28] It is not possible to predict if the oxidation occurs in all the rings and all the hydroxyl groups of a phenolic compound, which affects the number of electron donators present in the medium directly. However, as an example, we propose a mechanism of oxidation of the gallic acid derivative 9 (Scheme 1), which seems to be one of the compounds present in E. globulus bark extract with a high involvement in the metal-ion reduction. The abundance of hydroxyl and carboxyl groups on the structures of galloyl derivatives facilitates the interaction with the metal ions, which promotes the oxidation of hydroxyl groups to carboxyl counterparts and at the same time reduces metal ions to their elemental state. Notably, no oxidized products of phenolic compounds have been detected in the E. globulus bark extract after NPs synthesis. This may be because these components are aggregated at the surface of Ag and Au NPs; but there is also the possibility that they cannot be detected by HPLC–MS under the conditions used (e.g., as a result of their higher polarity). Finally, it is important to highlight that galloyl derivatives and hydrolyzable tannins are common water-soluble components of plant extracts and that most of the reported studies that use plant extracts to synthesize NPs have employed aqueous extracts obtained under the same conditions described in the present work.[17a, 29] The combined role of sugars and phenolic compounds To understand the possible combined effect of the main monosacharides and phenolic compounds identified in the E. globulus aqueous extract, a mixture of a pure solution of glucose, ChemSusChem 2014, 7, 2704 – 2711
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www.chemsuschem.org compounds. We have demonstrated that glucose and fructose, the main sugars present in plant extracts, play a negligible role in both Ag and Au NPs synthesis; however, these sugars could aggregate at the NPs surface to make them more stable. Conversely, gallic acid and, consequently, galloyl derivatives, are mainly responsible for the reduction of the Ag and Au hydrosols, which allows the fast formation of NPs. In addition, ellagic acid and isorhamnetin play a decisive role in the stabilization of Au NPs. If we consider that the composition of plant water extracts is broadly similar in terms of the main families of components present, these conclusions can be extrapolated to the vast majority of extracts used in this context. In this perspective, we anticipate that these results can be a driving force towards the optimization of green synthesis of metal NPs to obtain a better control of their properties.
Scheme 1. Mechanism proposed for the oxidation of 9 during NPs synthesis.
fructose, gallic and ellagic acids, and isorhamnetin was tested. After the addition of the standard compounds, the prompt formation of Ag and Au NPs was observed, with the appearance in both cases of the corresponding UV absorption bands (Figures S9 and S10). The NPs obtained using the sugars and phenolic compounds standard mixtures were more stable than those formed after the addition of the standard phenolic compounds mixture alone, and the Zeta potential values changed from 36.2 (with phenolic compounds) to 40.4 mV (with phenolic and sugar compounds) in Au NPs and from 24.4 to 30.3 mV for Ag NPs (Table S2). This trend suggests that the reducing sugars, under the conditions studied, have a more significant effect on the NPs stabilization, particularly in the case of Ag NPs, which is in agreement with the greater decrease of glucose in the extract after the Ag NPs synthesis.
Conclusions We have established a strategy to identify the natural reducing agents involved in the formation of Ag and Au nanoparticles (NPs) if plant extracts are applied in their green synthesis. This study relied on the identification of sugars and phenolic compounds present in E. globulus aqueous extract before and after Ag and Au NPs synthesis using advanced chromatographic techniques. An evaluation of the protein content of the extract was also performed. Evidence of the role of these components in NPs synthesis, as well as in their stabilization, was provided by using standard solutions of different sugars and phenolic 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Experimental Section Materials All chemicals were used as received: tetrachloroauric acid (99.9 %, Sigma–Aldrich), silver nitrate (99.9 %, Sigma–Aldrich), gallic acid (purity higher than 97.5 %, Sigma), isorhamnetin (purity higher than 99 %, Fluka Chemie), ellagic acid (96 %, Fluka Chemie), d(+)-glucose (purity higher than 99.5 %, Sigma), d-( )-fructose (99 %, Acros Organics), and formic acid (purity higher than 98 %, Fluka Chemie). HPLC-grade water and acetonitrile were supplied from Fisher Scientific Chemicals and were filtered by using a Solvent Filtration Apparatus 58061 from Supelco. E. globulus bark samples were taken from 16-year-old E. globulus trees harvested randomly from a clone plantation cultivated by RAIZ (Forest and Paper Research Institute) in Eixo (408 37’ 13.56’’ N, 88 34’ 08.43’’ W), Aveiro, Portugal. E. globulus bark samples were air-dried, until a constant weight was achieved, and ground to a granulometry lower than 2 mm prior to extraction.
E. globulus bark extraction Bark ( 20 g) was submitted to water extraction (1:50 m/v) at 100 8C for 2 min under constant stirring. The suspension was filtered, and the extract was used as a reducing and stabilizing agent for Ag and Au NPs synthesis. For characterization purposes, a portion of the aqueous extract was also freeze-dried. ChemSusChem 2014, 7, 2704 – 2711
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CHEMSUSCHEM FULL PAPERS Preparation of metal NPs Aqueous Ag and Au colloids were prepared by the reduction of the respective salts with the E. globulus extract at room temperature. Typically, the bark extract (100 mL) was added to AgNO3 or HAuCl4 aqueous solutions (300 mL, 1 mm) with stirring. A brownish yellow colloid was obtained in the case of Ag NPs, whereas a deep-red colloid was achieved for Au NPs. In both cases, after 1 h, the metal NPs were removed from the initial solution by highspeed centrifugation. The bark extract after the reduction was analyzed directly after the centrifugation, and the NPs were washed several times. The assays with standard compounds were performed by replacing the bark extract with the same volume of aqueous solutions of: 1) glucose, fructose, gallic acid, ellagic acid, and isorhamnetin individually, 2) mixtures of the compounds that belong to the same family (phenolic compounds and sugars), and 3) a mixture of all standard compounds. In all cases, the concentrations were equivalent to those found in the initial extract.
Analysis of protein content by elemental analysis The N content of the bark extract (before and after the NPs biosynthesis) was determined for a freeze-dried aliquot by elemental analysis by using a TruSpec 630-200-200 elemental analyzer (LECO, Madrid, Spain). The protein percentage was determined by converting the N percentage content using the standard 6.25 as the conversion factor.[21, 30] The measurements were performed in triplicate.
Analysis of sugars by HPLC–RI The monosaccharides content in E. globulus bark extract (before and after Ag and Au NPs synthesis) was determined by using a LaChrom Elite HPLC system from VWR-Hitachi with a RI detector (model L-2490) and an autosampler (model L-2200) using a 10 mm Eurokat H10 ion-exchange column (300 7.5 mm) equipped with a 30 8 mm precolumn supplied by Knauer. The mobile phase was aqueous sulfuric acid (0.01 n) with a flow rate of 0.4 mL min 1 under isocratic conditions for 30 min. The injected volume was 20 mL. Monosaccharides identification was performed by comparison to several standards, namely, glucose, xylose, arabinose, fructose, and galactose. Calibration curves were obtained by the injection of glucose and fructose standard solutions in water with six different concentrations between 0.05 and 2.00 mg mL 1. In addition to the linearity, the limits of detection (LOD) and quantification (LOQ) were estimated using the signal-to-noise ratio approach (n = 6).[26a] The calibration curves and additional relevant analytical data are shown in Table S1. The concentrations were calculated based on triplicate injections, and the mean value was computed in each case.
www.chemsuschem.org B; 3–35 min, 0–10 % B; 35–47 min, 10–15 % B; 47–52 min, 15 % B; 52–54 min, 15–20 % B; 54–57 min, 20–50 % B; 57–62 min, 50–100 % B; 62–72 min, 100–0 % B, followed by re-equilibration of the column for 10 min before the next run. Double online detection was performed by the DAD at 280 and 340 nm, and UV spectra in the range of 210–600 nm were also recorded. Before the injection, the extract was dissolved in H2O (HPLC grade) to obtain a final concentration of 8 mg mL 1 and filtered through a 0.2 mm Nylon syringe filter. The HPLC was coupled to a LCQ Fleet ion-trap mass spectrometer (ThermoFinnigan, San Jose, CA, USA), equipped with an ESI source and operated in the negative mode. The conditions were the same as described previously.[31] Data acquisition was performed by using the Xcalibur data system (ThermoFinnigan, San Jose, CA, USA). For the quantification of phenolic compounds, calibrations curves were obtained from the HPLC injection of gallic and ellagic acids and isorhamnetin standard solutions in H2O with six different concentrations between 0.05 and 40 mg mL 1. The linearity, LOD, and LOQ[26a] were estimated, and the relevant analytical data are shown in Table S1. The quantification of individual compounds was accomplished with calibration data for the most similar standard as no pure reference compounds were available for some of them. The concentrations were calculated based on triplicate injections, and the mean value was computed in each case.
Analysis of metal NPs The optical spectra were recorded by using a Jasco V-560 UV/Vis spectrophotometer using 100 scans per min with a band width of 5.0 nm. Metallic NPs were extracted from the initial solution by high-speed centrifugation (18 000 rpm) by using a Sigma 3–30k centrifuge, and the NPs were washed thoroughly several times with distilled water. STEM images were obtained by using a fieldemission gun (FEG) SEM Hitachi SU70 microscope operated at 15 kV and fitted with an energy dispersive X-ray spectroscopy (EDX) accessory (EDX Detector: Bruker AXS, Software: Quantax). Samples were prepared by placing a colloid drop directly onto a carbon-coated copper grid and then allowing the solvent to evaporate. The average diameter of the NPs was determined by analysis of STEM images by using the ImageJ program (at least 50 NPs were analyzed in each case). The particle size distributions and the Zeta potential were evaluated by DLS analysis by using a Zetasizer Nano Series analyzer (Malvern Instruments). FTIR spectra of the NPs were collected by using a Bruker optics tensor 27 spectrometer coupled to a horizontal attenuated total reflectance (ATR) cell using 256 scans at a resolution of 4 cm 1. The Au and Ag nanophases were identified by powder XRD by using a Philips X’Pert instrument with CuKa radiation (l = 1.543178 ) at 40 kV and 50 mA.
Acknowledgements Analysis of phenolic compounds by HPLC–DAD–MS The HPLC system consisted of a variable loop Accela autosampler (200 vial capacity set at 15 8C), an Accela 600 LC pump, and an Accela 80 Hz DAD detector (Thermo Fisher Scientific, San Jose, CA, USA). The separation of the compounds was performed at 25 8C with a flow rate of 200 mL min 1 by using a Discovery C-18 column (150 2.1 mm 5 mm) equipped with a precolumn both supplied by Supelco (Agilent Technologies). The injection volume in the HPLC system was 10 mL, and the mobile phase consisted of water/ acetonitrile (90:10 v/v) (A) and acetonitrile (B) both with 0.1 % of formic acid. The following linear gradient was used: 0–3 min, 0 % 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
This work was financed by FEDER funds by Programa Operacional Factores de Competitividade-COMPETE and by national funds by Fundażo para a CiÞncia e Tecnologia (FCT) within the CICECO project FCOMP-01-0124-FEDER-037271 (Ref. PEst-C/CTM/ LA0011/2013). The authors further wish to thank the FCT for postdoctoral grants to S.A.O.S. (SFRH/BPD/84226/2012) and R.J.B.P. (SFRH/BPD/89982/2012) and for funding from TEMA (PEst-C/EME/ UI0481/2013) and QOPNA (PEst-C/QUI/UI0062/2013). C.S.R.F. and P.A.A.P.M. also acknowledge the FCT/MCTES for a research contract under the Program Investigador FCT 2012 and 2013, respecChemSusChem 2014, 7, 2704 – 2711
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CHEMSUSCHEM FULL PAPERS tively. Microscopy was supported by the Rede Nacional de Microscopia Eletrnica (RNME-Pole UA FCT) project REDE/1509/RME/ 2005. Keywords: gold · green chemistry · nanoparticles · reduction · silver [1] a) M. C. Daniel, D. Astruc, Chem. Rev. 2004, 104, 293 – 346; b) Y. Volokitin, J. Sinzig, L. J. deJongh, G. Schmid, M. N. Vargaftik, I. I. Moiseev, Nature 1996, 384, 621 – 623. [2] Y. Xia, Y. Xiong, B. Lim, S. E. Skrabalak, Angew. Chem. Int. Ed. 2009, 48, 60 – 103; Angew. Chem. 2009, 121, 62 – 108. [3] P. Raveendran, J. Fu, S. L. Wallen, J. Am. Chem. Soc. 2003, 125, 13940 – 13941. [4] V. Sanchez-Mendieta, A. R. Vilchis-Nestor in Nanotechnology and Nanomaterials, Noble Metals (Ed.: Y.-H. Su), InTech, Rijeka, 2012, pp. 391 – 408. [5] S. He, Z. Guo, Y. Zhang, S. Zhang, J. W. Gu, Mater. Lett. 2007, 61, 3984 – 3987. [6] A. Ahmad, P. Mukherjee, S. Senapati, D. Mandal, M. I. Khan, R. Kumar, M. Sastry, Colloids Surf. B 2003, 28, 313 – 318. [7] V. Kumar, S. K. Yadav, J. Chem. Technol. Biotechnol. 2009, 84, 151 – 157. [8] J. Huang, Q. Li, D. Sun, Y. Lu, Y. Su, X. Yang, H. Wang, Y. Wang, W. Shao, N. He, J. Hong, C. Chen, Nanotechnology 2007, 18, 105104. [9] N. Durn, P. D. Marcato, M. Duran, A. Yadav, A. Gade, M. Rai, Appl. Microbiol. Biotechnol. 2011, 90, 1609 – 1624. [10] S. Iravani, Green Chem. 2011, 13, 2638 – 2650. [11] a) J. Huang, G. Zhan, B. Zheng, D. Sun, F. Lu, Y. Lin, H. Chen, Z. Zheng, Y. Zheng, Q. Li, Ind. Eng. Chem. Res. 2011, 50, 9095 – 9106; b) B. Zheng, T. Kong, X. Jing, T. Odoom-Wubah, X. Li, D. Sun, F. Lu, Y. Zheng, J. Huang, Q. Li, J. Colloid Interface Sci. 2013, 396, 138 – 145. [12] a) Y. Zhou, W. Lin, J. Huang, W. Wang, Y. Gao, L. Lin, Q. Li, L. Lin, M. Du, Nanoscale Res. Lett. 2010, 5, 1351 – 1359; b) S. Li, Y. Shen, A. Xie, X. Yu, L. Qiu, L. Zhang, Q. Zhang, Green Chem. 2007, 9, 852 – 858. [13] a) S. A. O. Santos, J. J. Villaverde, C. M. Silva, C. P. Neto, A. J. D. Silvestre, J. Supercrit. Fluids 2012, 71, 71 – 79; b) S. A. O. Santos, C. S. R. Freire, M. R. M. Domingues, A. J. D. Silvestre, C. P. Neto, J. Agric. Food Chem. 2011, 59, 9386 – 9393. [14] a) Z. Sadowski in Nanotechnology and Nanomaterials, Silver Nanoparticles (Ed.: D. P. Perez), InTech, Rijeka, 2010, pp. 257 – 276; b) R. J. B. Pinto, P. A. A. P. Marques, C. P. Neto, T. Trindade, S. Daina, P. Sadocco, Acta Biomater. 2009, 5, 2279 – 2289.
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www.chemsuschem.org [15] L. M. Liz-Marzn, Langmuir 2006, 22, 32 – 41. [16] R. R. Naik, S. J. Stringer, G. Agarwal, S. E. Jones, M. O. Stone, Nat. Mater. 2002, 1, 169 – 172. [17] a) S. S. Shankar, A. Rai, A. Ahmad, M. Sastry, J. Colloid Interface Sci. 2004, 275, 496 – 502; b) A. J. Bard, R. Parsons, J. Jordan in Monographs in Electroanalytical Chemistry and Electrochemistry, 1edst ed(Ed.: A. J. Bard), Marcel Dekker, Inc., New York, 1985. [18] V. K. Sharma, R. A. Yngard, Y. Lin, Adv. Colloid Interface Sci. 2009, 145, 83 – 96. [19] L. J. Bellamy, The Infrared Spectra of Complex Molecules, Vol. 1, 3rd ed., Wiley, New York, 1975. [20] R. Fido, E. N. C. Mills, N. Rigby, P. Shewry in Protein Purification Protocols, Vol. 244 (Ed.: P. Cutler), Humana Press 2004, pp. 21 – 27. [21] C. V. Gonzlez Lpez, M. d. C. C. Garca, F. G. A. Fernndez, C. S. Bustos, Y. Chisti, J. M. F. Sevilla, Bioresour. Technol. 2010, 101, 7587 – 7591. [22] Z. Lin, J. Wu, R. Xue, Y. Yang, Spectrochim. Acta Part A 2005, 61, 761 – 765. [23] a) P. Shanmugavelan, S. Y. Kim, J. B. Kim, H. W. Kim, S. M. Cho, S. N. Kim, S. Y. Kim, Y. S. Cho, H. R. Kim, Carbohydr. Res. 2013, 380, 112 – 117; b) C. Ma, Z. Sun, C. Chen, L. Zhang, S. Zhu, Food Chem. 2014, 145, 784 – 788. [24] A. K. Mittal, Y. Chisti, U. C. Banerjee, Biotechnol. Adv. 2013, 31, 346 – 356. [25] L. Ortega-Arroyo, E. San Martin-Martinez, M. A. Aguilar-Mendez, A. CruzOrea, I. Hernandez-P rez, C. Glorieux, Starch/Staerke 2013, 65, 814 – 821. [26] a) S. A. O. Santos, C. Vilela, C. S. R. Freire, C. P. Neto, A. J. D. Silvestre, J. Chromatogr. B 2013, 938, 65 – 74; b) L. Boulekbache-Makhlouf, E. Meudec, M. Chibane, J. P. Mazauric, S. Slimani, M. Henry, V. Cheynier, K. Madani, J. Agric. Food Chem. 2010, 58, 12615 – 12624. [27] M. Boukhris, M. S. J. Simmonds, S. Sayadi, M. Bouaziz, Phytother. Res. 2013, 27, 1206 – 1213. [28] L. Pourcel, J.-M. Routaboul, V. Cheynier, L. Lepiniec, I. Debeaujon, Trends Plant Sci. 2007, 12, 29 – 36. [29] D. MubarakAli, N. Thajuddin, K. Jeganathan, M. Gunasekaran, Colloids Surf. B 2011, 85, 360 – 365. [30] N. L. Conklin-Brittain, E. S. Dierenfeld, R. W. Wrangham, M. Norconk, S. C. Silver, J. Chem. Ecol. 1999, 25, 2601 – 2622. [31] S. A. O. Santos, J. J. Villaverde, A. F. Sousa, J. F. J. Coelho, C. P. Neto, A. J. D. Silvestre, Ind. Crops Prod. 2013, 47, 262 – 269. [32] S. Dall’Acqua, P. Minesso, B. B. Shresta, S. Comai, P. K. Jha, M. B. Gewali, E. Greco, R. Cervellati, G. Innocenti, Molecules 2012, 17, 1686 – 1697.
Received: February 28, 2014 Published online on August 1, 2014
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Unveiling the Chemistry behind the Green Synthesis of Metal Nanoparticles Snia A. O. Santos,*[a] Ricardo J. B. Pinto,*[a, b] Slvia M. Rocha,[c] Paula A. A. P. Marques,[b] Carlos Pascoal Neto,[a] Armando J. D. Silvestre,[a] and Carmen S. R. Freire[a] Nanobiotechnology has emerged as a fundamental domain in modern science, and metallic nanoparticles (NPs) are one of the largest classes of NPs studied because of their wide spectrum of possible applications in several fields. The use of plant extracts as reducing and stabilizing agents in their synthesis is an interesting and reliable alternative to conventional methodologies. However, the role of the different components of such extracts in the reduction/stabilization of metal ions has not yet
been understood clearly. Here we studied the behavior of the main components of a Eucalyptus globulus Labill. bark aqueous extract during metal-ion reduction followed by advanced chromatographic techniques, which allowed us to establish their specific role in the process. The obtained results showed that phenolic compounds, particularly galloyl derivatives, are mainly responsible for the metal-ion reduction, whereas sugars are essentially involved in the stabilization of the NPs.
Introduction Nanometer-scale metallic particles are probably one of the main classes of nanomaterials investigated and used currently because of the wide spectrum of potential applications in several fields that arises from their unique and distinct properties in relation to the bulk counterparts.[1] The recent advances in the fields of nanoscience and nanotechnology lead to a variety of physical and chemical strategies for the selective preparation of inorganic nanoparticles (NPs) with precise control over their shape and dimensionality.[2] Among the chemical methodologies, those used most commonly are based on soluble metal salt precursors and different reducing agents, which can act also as stabilizers to avoid the coalescence of the NPs. However, most of the reducing agents reported, for example, hydrazine, sodium borohydride, or N,N-dimethylformamide, are reactive chemicals that are commonly associated with huge environmental risks and toxicity.[3] [a] Dr. S. A. O. Santos,+ Dr. R. J. B. Pinto,+ Prof. Dr. C. P. Neto, Prof. Dr. A. J. D. Silvestre, Dr. C. S. R. Freire Department of Chemistry-CICECO University of Aveiro 3810-193 Aveiro (Portugal) Fax: (+ 351) 234-401-470 E-mail: [email protected] [email protected] [b] Dr. R. J. B. Pinto,+ Dr. P. A. A. P. Marques TEMA-NRD, Mechanical Engineering Department and Aveiro Institute of Nanotechnology (AIN) University of Aveiro 3810-193 Aveiro (Portugal) [c] Prof. Dr. S. M. Rocha Department of Chemistry-QOPNA University of Aveiro 3810-193 Aveiro (Portugal) [+] These authors contributed equally to this work. Supporting information for this article is available on the WWW under http://dx.doi.org/10.1002/cssc.201402126.
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The green synthesis of metal NPs has been studied as a reliable and promising alternative to minimize or eliminate the use of these harmful substances.[4] Green synthetic processes comprise either micro-organisms, such as bacteria[5] or fungi,[6] as well as plant biomass[7] and several plant extracts.[8] The use of plant extracts from different morphological parts, which include leaves, fruits, seeds, and bark, presents some advantages over the use of micro-organisms because of the high diversity and abundance of plant extracts and the simplicity, easy scaleup, and cost-effectiveness of the method.[4, 9, 10] Despite the remarkable number of reports in the literature on the synthesis of metal NPs using distinct plant extracts, as highlighted recently by Iravani,[10] a precise understanding of their formation mechanism and the clear-cut role of the natural compounds involved in this process is still imprecise. Some reports have put forward hypothetical mechanisms that propose that the reduction of the metal ions to metal NPs may be caused by the distinct compounds present in the extracts, such as reducing sugars, phenolic compounds (e.g., flavonoids), and proteins.[9, 11] However, these generic suggestions have been based essentially on qualitative routine spectrophotometric analysis, such as FTIR spectroscopy, colorimetric assays, or UV spectroscopy,[12] which cannot unambiguously differentiate the presence or absence of the different families of compounds in the extracts and resulting NPs. In this vein, and in consideration of the relevance of such fundamental knowledge, particularly in the optimization and control of the final properties of the NPs and on the up-scaling of these green processes, in the present study we intended to clarify the role of the different components of a plant extract in a NP formation mechanism, kinetics, and stabilization. Here, an aqueous extract of Eucalyptus globulus Labill. bark, a typical residue from the pulp and paper industry that is recognized for its high content of phenolic compounds,[13] was ChemSusChem 2014, 7, 2704 – 2711
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chosen as a reducing and stabilizing system for Ag and Au NPs synthesis. Ag and Au NPs were selected because of their potential application in many fields, such as catalysis, sensors, and in the biomedical field as, for example, drug-delivery systems and antimicrobial agents.[14] Furthermore, among the noble-metal NPs investigated so far, Au and Ag NPs have been studied in particular, which allows a comparative and critical discussion of the results. The clear identification of the natural reducing agents involved in the formation of the metal NPs was achieved by the qualitative and quantitative analysis of sugars and phenolic compounds present in the plant extracts before and after Ag and Au NP synthesis by using advanced chromatographic techniques.
Results and Discussion Synthesis of Au and Ag NPs using E. globulus bark extract In this study, comparative experiments using E. globulus aqueous extracts as a reducing and stabilizing agent in the preparation of Ag and Au hydrosols were performed to elucidate the role of the different components of plant extracts in green processes. UV/Vis spectroscopy was used to confirm the presence of the characteristic surface plasmon resonance (SPR) bands of Ag and Au nanospheres in the visible region[15] and, therefore, the successful formation of the metal NPs. UV/Vis spectra of the Au and Ag aqueous colloidal solutions obtained after the addition of the bark extract as a function of time are shown in Figure 1 (the E. globulus extract does not present any absorption bands in the visible region at the concentration used). The addition of the bark extract to the Au salt solution caused an almost instantaneous change of color from light yellow to red, which indicated the effectiveness of the Au salt reduction. As expected, the obtained Au colloidal solution showed the typical absorption band at around 530 nm without any significant shift over time, except for a gradual increase of intensity, most noticeable in the first minutes of the reaction. Similarly, the addition of the E. globulus extract to the Ag salt solution led to the formation of Ag NPs, which followed a slower kinetics (Figure 1). One hour after the addition of the extract, the UV/Vis spectrum showed two weak and broad absorbance bands at around l = 375 and 445 nm. This can be explained by the variances in terms of composition and experimental conditions tested in this synthesis, which leads to a wide range of sizes of the particles as the broadening of the plasmon band is not always observed in the case of Ag NPs.[8, 16] Nevertheless, a significant increase of the intensity of these bands with the reaction time was observed, which shows that the formation of the Ag hydrosol is considerably slower than that of the Au counterpart using the same extract as the reducing agent. This dissimilarity can be explained by the different reduction potentials of the two metal ions; the standard reduction potential of Ag+ to Ag0 (E0 = 0.799 V) is substantially lower than that of Au3+ to Au0 (E0 = 1.498 V).[17] Scanning transmission electron microscopy (STEM) images of the Au and Ag NPs obtained by reduction with E. globulus 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Figure 1. UV/Vis spectra of a) Au and b) Ag colloidal solutions at distinct times after the addition of the E. globulus aqueous extract. Inset: detail of the differences observed for the Au colloidal solution.
bark extract after 1 h of reaction are shown in Figure 2. The Au NPs exhibited a homogeneous distribution of well-defined nanoparticles with a predominantly spherical morphology and a narrow size distribution (average diameter of (18 3) nm); however, some anisotropic planar nanostructures such as triangles or hexagonal-shaped particles were also observed. Conversely, the Ag NPs showed a spherical and polydisperse distribution with average diameters in the range of 15–73 nm, which indicates that this sample is composed of a large quantity of nonuniform NPs and corroborates the broad plasmon resonance band observed by UV/Vis spectroscopy. Dynamic light scattering (DLS) analysis of these colloids revealed an average size of (21 4) and (52 16) nm for Au and Ag NPs, respectively (Figure S1). These values are quite similar to those obtained by STEM analysis, and the slight differences are because DLS analysis measures the hydrodynamic diameter, which includes the hydration layer. The Zeta potential values of the colloidal solutions were ( 16 6) (pH 2.74) and ( 18 7) mV (pH 3.48) for Au and Ag NPs, respectively. These results indicated that the surface of both metal NPs is composed mainly of negatively charged capping compounds, which are responsible for their moderate stability. This is in close agreement with data reported previously that suggest that the biomolecules present in natural extracts serve both as reducing and capping agents on metal NPs prepared by green synthesis.[18] The crystalline structure of the obtained Au and Ag NPs was confirmed by powder XRD analysis of dried samples (FigChemSusChem 2014, 7, 2704 – 2711
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www.chemsuschem.org sugars of the bark extracts were investigated before and after NPs formation.
Identification of the biomolecules responsible for Au and Ag NPs synthesis and stabilization using E. globulus bark extract The role of proteins
Figure 2. STEM images of a) Au and b) Ag NPs with the respective histograms of size distribution obtained after reduction (1 h) with the E. globulus aqueous extract.
ure S2). The observed diffraction peaks can be indexed to the (111), (2 0 0), (2 2 0), and (3 11) crystalline planes of Au and Ag face-centered cubic (fcc) structures.[8, 12b] The FTIR spectra of the pristine E. globulus aqueous bark extract (Figure S3) showed several absorption bands located in the n˜ = 1000–1800 cm 1 region that could be assigned to the vibrations of typical functional groups, namely, C O C, C O, C=C, and C=O, present in several water-soluble components of E. globulus bark, which include phenolic compounds, proteins, and sugars.[19] Some of these absorption bands were also observed in the spectra of the obtained metal NPs, which confirms clearly that some biomolecules also act as capping agents and, therefore, play an important role on the NPs stabilization. Although most studies[8, 17a] based their conclusions and proposals in terms of reduction mechanisms on the results of FTIR spectroscopy studies, it is impossible to distinguish the different families of compounds and their contribution to the reduction and/or stabilization processes unambiguously. To assess the role of the different extract components in the Au and Ag NPs green synthesis and stabilization, the protein content and the composition of the phenolic compounds and 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Several authors have attributed the reducing capacity of plant extracts in NPs synthesis to proteins,[12b] although most of the extracts used have been obtained by employing conditions that do not commonly allow protein removal or promote their degradation (e.g., the absence of an appropriate buffer and/or mechanical processes suitable to disrupt the cell walls).[20] Nonetheless, we decided to verify the presence of proteins in the extracts studied here, based on the N content determined by elemental analysis,[21] and to access their possible participation in the NPs synthesis. The protein content of the E. globulus bark extract determined by elemental analysis represented approximately 0.8 wt % of the total extract, which corresponds to a concentration of approximately 19.0 mg mL 1 of the aqueous extract. This value is extremely low (4- and 19-fold lower than the phenolic compounds and sugars content, respectively). Additionally, this value is in a range of concentration that was reported not to have a linear relationship with Au NPs synthesis.[12a] Moreover, the protein content determined by elemental analysis was probably overestimated because other extract components could also contribute to the N content. Therefore, this result suggested clearly that proteins are not involved in metal-ion reduction under the conditions studied. However, they might play an important role in the stabilization of the NPs. To corroborate this, the content of N in the NPs was also evaluated, and 29.3 and 14.6 % of the N content of the E. globulus bark extract was aggregated in the Ag and Au NPs, respectively. The capping/stabilization ability of proteins is certainly associated with the affinity of carbonyl groups from amino acid residues of proteins to bind metals.[22] The role of sugars The content of monosaccharides in the E. globulus extract, before and after NPs synthesis, was determined by HPLC with refractive index detection (RI). Only two monosaccharides were detected in this extract, namely, glucose and fructose, which together represent 157.1 mg g 1 of extract with concentrations of 127.7 and 233.7 mg mL 1 of aqueous extract, respectively. These concentrations are in the same range as those reported for several vegetables, fruits, seeds, and other plant extracts,[23] which are used commonly in the biogenic synthesis of NPs.[10, 24] After the Ag and Au NPs synthesis, a considerable decline in the content of both monosaccharides in the extracts was observed. The glucose concentration decreased to 52.7 and 71.4 mg mL 1 in the extract after Ag and Au NPs synthesis, respectively, whereas the fructose content decreased to 114.4 and 119.7 mg mL 1 in the extract. This first outcome suggests the involvement of these reducing sugars in the reduction of ChemSusChem 2014, 7, 2704 – 2711
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1.47 0.07 1.11 0.01 0.63 0.02 – 0.50 0.01 – – – 0.19 0.01 – – – – – – – 3.90 0.08
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[a] Rt = retention time. [b] m/z values in bold were subjected to MSn analysis. [c] Identified by co-injection of the respective authentic standard. [d] ntdm—not determined.
2.30 0.08 3.10 0.16 1.97 0.060 5.15 0.25 1.30 0.07 1.18 0.06 1.56 0.030 0.33 0.01 4.87 0.22 3.41 0.17 1.16 0.06(11+12) – 3.42 0.16 6.00 0.30 0.29 0.01 1.40 0.06 37.45 1.18 3.17 0.12 4.43 0.22 2.34 0.10 11.13 0.43 1.84 0.07 2.34 0.11 2.38 0.10 0.84 0.03 10.17 0.25 4.43 0.14 1.65 0.05(11+12) – 7.37 0.29 14.27 0.55 0.44 0.01 3.45 0.12 70.27 1.75 [13b] [26b] [26a] Co[c] [32] [13b] [13b] [13b] [13b] [26a] [26a] [26a] [13b] [26a] [13b] [26a] 173, 171, 111, 93, 85 301,[b] 275, 257, 229; MS3 : 284, 257, 229, 185 271, 241, 211, 169; MS3 : 125 125 591, 483, 441, 423, 305; MS3 : 423 481, 301; MS3 :301, 275 481, 301; MS3 : 301, 275 301; MS3 :229 633, 301; MS3 : 463, 299, 275 482, 327, 313, 183, 169; MS3 : 169, 125 301, 300; MS3 : 284, 257, 229, 185, 184 301, 300; MS3 : 284, 257, 229, 185 315, 314; MS3 : 300; MS4 : 244, 200 315, 300; MS3 : 300; MS4 : 271, 244, 243, 216, 200 315; MS3 : 300; MS4 : 244 315, 300; MS3 : 300; MS4 : 272, 271, 244, 243 216, 200 MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : MS2 : 191 481 331 169 609 783 783 633 935 497 447 433 477 447 461 447 2.73 250 quinic acid 2.82 244, 270 HHDP-glucose 3.03 240 (sh), 270 galloylglucose 3.54 269 gallic acid 3.90 238, 270 gallocatechin dimer 4.59 238, 265(sh) bis-HHDP-glucose isomer 5.24 238, 265(sh) bis-HHDP-glucose isomer 7.31 237, 270 galloyl-HHDP-glucose 17.62 239, 269 galloyl-bis-HHDP-glucose 30.87 236, 273 eucaglobulin ellagic acid rhamnoside 31.73 ndtm[d] 31.96 ndtm ellagic acid-pentoside 33.29 245, 364 isorhamnetin-hexoside 43.07 249, 364 isorhamnetin-pentoside isomer 43.98 237, 368 isorhamnetin-rhamnoside 44.47 244, 365 isorhamnetin-pentoside isomer total [mg mL 1 of extract] 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
Identification Phenolic compounds in E. globulus bark extract [mg mL 1 extract] before NPs synthesis after Ag NPs after Au NPs MSn product ions [m/z] [M H] [m/z] Compound
The aqueous extract of E. globulus bark extract was characterized by HPLC with diode array detection coupled to tandem mass spectrometry (HPLC–DAD–MSn), which allowed us to identify 16 phenolic compounds (Figure S7). The compounds identified in E. globulus aqueous extract, their retention time, UV absorption maximum (if possible), the molecular ion [M H] , and the corresponding MSn product ions are summarized in Table 1. Compounds were identified by comparing their fragmentation profiles with those of authentic standards or, if these were not available, with published data, as indicated in Table 1. Most of the phenolic compounds identified in this E. globulus bark aqueous extract were previously described as constituents of this Eucalyptus species[13b] or even of plant tissues of other Eucalyptus species.[26] The phenolic fraction of E. globulus bark aqueous extracts is composed mainly of galloylglucose, ellagic acid, and isorhamnetin derivatives (Figure S7), which accounted for a total amount of 30.55 mg of phenolic compounds per gram of dry extract and corresponds to a concentration of 70.27 mg mL 1 of aqueous extract used for the NPs synthesis (Table 1). Among the 16 phenolic compounds identified in the extract (Figure S7), an isomer of isorhamnetin-pentoside (14), gallic acid (4), and galloyl-bis-HHDPglucose (9; Figure 3) are the major components with concentrations of (14.27 0.55), (11.13 0.43), and (10.27 0.25) mg mL 1 of aqueous extract, respectively. The amount of phenolic compounds in the Eucalyptus bark aqueous extract
Peak no. Rt[a] lmax [min] [nm]
The role of phenolic compounds
Table 1. HPLC–MSn fragmentation profile and abundance of phenolic compounds identified in E. globulus aqueous extract before and after Ag and Au NPs synthesis.
Ag+ and Au3+ or in the stabilization of the obtained NPs. Indeed, the participation of reducing sugars, such as glucose, in the reduction of metal ions in NPs synthesis using plant extracts has been suggested previously, and several mechanisms have been proposed;[22] however, most authors neglect the coexistence of other reducing agents in these types of plant extracts. Thus, to verify the specific role of glucose and fructose in the NPs synthesis, standard solutions with the same concentration as that observed in the bark extract (0.13 and 0.23 mg mL 1 for glucose and fructose, respectively) were also tested. After 1 h of reaction, no metal-ion reduction was perceived with glucose or fructose standard solutions, even if the two reducing sugars were mixed together (Figures S4 and S5). This behavior could be related essentially to the relatively low concentration of these reducing sugars in the bark extract. Ortega-Arroyo et al.[25] used glucose as the reducing agent in the synthesis of Ag NPs successfully by employing considerably higher concentrations than those used in this work (between 18- and 108-fold). Notwithstanding, this reaction could be also affected by the kinetic rate of the redox reactions as the formation of a low quantity of Au NPs was observed both with glucose and fructose after one month, confirmed by the presence of the UV band at l = 590 nm (Figure S6). In the case of Ag (which has a reduction potential lower than Au) even after one month no NPs formation was observed by UV spectrophotometry. These results indicated that sugars are essentially involved in the NPs stabilization, as will be discussed below.
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Figure 3. Structures of the main phenolic compounds responsible for NPs synthesis (see number assignments in Table 1).
used in this work is very similar to other aqueous extracts used previously in NPs synthesis,[24] such as that from Pelargonium graveolens leaves.[27] The phenolics content of the E. globulus bark extract before and after NPs synthesis was quantified by HPLC with UV detection and is expressed in mg mL 1 of extract (Table 1). After Ag NPs synthesis, the content of phenolic compounds decreased to 37.97 mg mL 1 (Table 1), and this decline was more accentuated for isorhamnetin and phenolic acid derivatives with decreases of 54 and 48 %, respectively. Specifically, the concentrations of 14 and 4 diminished the most. In contrast, after Au NPs synthesis the majority of the phenolic compounds present initially in the E. globulus bark disappeared (accounting only for 4.36 mg mL 1), of which only quinic acid (1), HHDP-glucose (2), galloylglucose (3), gallocatechin dimer (5), and 9 were detected (Figure S8). These results indicate that phenolic compounds play a dominant role in this green process. In this sense, to confirm their specific starring role in NPs synthesis, gallic and ellagic acids and isorhamnetin standards were tested individually by using equivalent contents to those found in the bark extract. In the Ag NPs synthesis, gallic acid showed the fastest redox kinetics (Figure S9). In the Au NPs synthesis, after the addition of gallic acid, the color of the HAuCl4 solution changed in a few seconds from light yellow to pink, identical to that observed in the case of the E. globulus extract, which indicates the successful synthesis of Au NPs. However, no metal reduction was observed if ellagic acid or isorhamnetin was added to the Au solution (Figure S10). Once more, the faster reduction rate of Au ions than that of Ag ions is most likely because of significant differences in the reduction potentials of the two metal ions, as observed with the total extract. These observations established that phenolic compounds, in particular derivatives of gallic acid, are mainly responsible for metal-ion reduction in both Ag and Au NPs synthesis using E. globulus bark extracts. Moreover, in the case of Au NPs, the presence of ellagic acid and isorhamnetin seems also to increase their stability as the Zeta potential of Au NPs changed 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
from 29.0 mV in the presence of gallic acid to 36.2 mV if the three phenolic compounds were used together (Table S2). These observations explain the remarkable decrease of the content of all phenolic compounds after the Au NPs synthesis. Consequently, phenolic compounds, and in particular galloyl derivatives, would serve as the main electron donors in the redox reaction. Phenolic acids, galloyl derivatives, and flavonoids are known to form quinones upon oxidation and to undergo further oxidation to form acids.[28] It is not possible to predict if the oxidation occurs in all the rings and all the hydroxyl groups of a phenolic compound, which affects the number of electron donators present in the medium directly. However, as an example, we propose a mechanism of oxidation of the gallic acid derivative 9 (Scheme 1), which seems to be one of the compounds present in E. globulus bark extract with a high involvement in the metal-ion reduction. The abundance of hydroxyl and carboxyl groups on the structures of galloyl derivatives facilitates the interaction with the metal ions, which promotes the oxidation of hydroxyl groups to carboxyl counterparts and at the same time reduces metal ions to their elemental state. Notably, no oxidized products of phenolic compounds have been detected in the E. globulus bark extract after NPs synthesis. This may be because these components are aggregated at the surface of Ag and Au NPs; but there is also the possibility that they cannot be detected by HPLC–MS under the conditions used (e.g., as a result of their higher polarity). Finally, it is important to highlight that galloyl derivatives and hydrolyzable tannins are common water-soluble components of plant extracts and that most of the reported studies that use plant extracts to synthesize NPs have employed aqueous extracts obtained under the same conditions described in the present work.[17a, 29] The combined role of sugars and phenolic compounds To understand the possible combined effect of the main monosacharides and phenolic compounds identified in the E. globulus aqueous extract, a mixture of a pure solution of glucose, ChemSusChem 2014, 7, 2704 – 2711
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www.chemsuschem.org compounds. We have demonstrated that glucose and fructose, the main sugars present in plant extracts, play a negligible role in both Ag and Au NPs synthesis; however, these sugars could aggregate at the NPs surface to make them more stable. Conversely, gallic acid and, consequently, galloyl derivatives, are mainly responsible for the reduction of the Ag and Au hydrosols, which allows the fast formation of NPs. In addition, ellagic acid and isorhamnetin play a decisive role in the stabilization of Au NPs. If we consider that the composition of plant water extracts is broadly similar in terms of the main families of components present, these conclusions can be extrapolated to the vast majority of extracts used in this context. In this perspective, we anticipate that these results can be a driving force towards the optimization of green synthesis of metal NPs to obtain a better control of their properties.
Scheme 1. Mechanism proposed for the oxidation of 9 during NPs synthesis.
fructose, gallic and ellagic acids, and isorhamnetin was tested. After the addition of the standard compounds, the prompt formation of Ag and Au NPs was observed, with the appearance in both cases of the corresponding UV absorption bands (Figures S9 and S10). The NPs obtained using the sugars and phenolic compounds standard mixtures were more stable than those formed after the addition of the standard phenolic compounds mixture alone, and the Zeta potential values changed from 36.2 (with phenolic compounds) to 40.4 mV (with phenolic and sugar compounds) in Au NPs and from 24.4 to 30.3 mV for Ag NPs (Table S2). This trend suggests that the reducing sugars, under the conditions studied, have a more significant effect on the NPs stabilization, particularly in the case of Ag NPs, which is in agreement with the greater decrease of glucose in the extract after the Ag NPs synthesis.
Conclusions We have established a strategy to identify the natural reducing agents involved in the formation of Ag and Au nanoparticles (NPs) if plant extracts are applied in their green synthesis. This study relied on the identification of sugars and phenolic compounds present in E. globulus aqueous extract before and after Ag and Au NPs synthesis using advanced chromatographic techniques. An evaluation of the protein content of the extract was also performed. Evidence of the role of these components in NPs synthesis, as well as in their stabilization, was provided by using standard solutions of different sugars and phenolic 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Experimental Section Materials All chemicals were used as received: tetrachloroauric acid (99.9 %, Sigma–Aldrich), silver nitrate (99.9 %, Sigma–Aldrich), gallic acid (purity higher than 97.5 %, Sigma), isorhamnetin (purity higher than 99 %, Fluka Chemie), ellagic acid (96 %, Fluka Chemie), d(+)-glucose (purity higher than 99.5 %, Sigma), d-( )-fructose (99 %, Acros Organics), and formic acid (purity higher than 98 %, Fluka Chemie). HPLC-grade water and acetonitrile were supplied from Fisher Scientific Chemicals and were filtered by using a Solvent Filtration Apparatus 58061 from Supelco. E. globulus bark samples were taken from 16-year-old E. globulus trees harvested randomly from a clone plantation cultivated by RAIZ (Forest and Paper Research Institute) in Eixo (408 37’ 13.56’’ N, 88 34’ 08.43’’ W), Aveiro, Portugal. E. globulus bark samples were air-dried, until a constant weight was achieved, and ground to a granulometry lower than 2 mm prior to extraction.
E. globulus bark extraction Bark ( 20 g) was submitted to water extraction (1:50 m/v) at 100 8C for 2 min under constant stirring. The suspension was filtered, and the extract was used as a reducing and stabilizing agent for Ag and Au NPs synthesis. For characterization purposes, a portion of the aqueous extract was also freeze-dried. ChemSusChem 2014, 7, 2704 – 2711
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CHEMSUSCHEM FULL PAPERS Preparation of metal NPs Aqueous Ag and Au colloids were prepared by the reduction of the respective salts with the E. globulus extract at room temperature. Typically, the bark extract (100 mL) was added to AgNO3 or HAuCl4 aqueous solutions (300 mL, 1 mm) with stirring. A brownish yellow colloid was obtained in the case of Ag NPs, whereas a deep-red colloid was achieved for Au NPs. In both cases, after 1 h, the metal NPs were removed from the initial solution by highspeed centrifugation. The bark extract after the reduction was analyzed directly after the centrifugation, and the NPs were washed several times. The assays with standard compounds were performed by replacing the bark extract with the same volume of aqueous solutions of: 1) glucose, fructose, gallic acid, ellagic acid, and isorhamnetin individually, 2) mixtures of the compounds that belong to the same family (phenolic compounds and sugars), and 3) a mixture of all standard compounds. In all cases, the concentrations were equivalent to those found in the initial extract.
Analysis of protein content by elemental analysis The N content of the bark extract (before and after the NPs biosynthesis) was determined for a freeze-dried aliquot by elemental analysis by using a TruSpec 630-200-200 elemental analyzer (LECO, Madrid, Spain). The protein percentage was determined by converting the N percentage content using the standard 6.25 as the conversion factor.[21, 30] The measurements were performed in triplicate.
Analysis of sugars by HPLC–RI The monosaccharides content in E. globulus bark extract (before and after Ag and Au NPs synthesis) was determined by using a LaChrom Elite HPLC system from VWR-Hitachi with a RI detector (model L-2490) and an autosampler (model L-2200) using a 10 mm Eurokat H10 ion-exchange column (300 7.5 mm) equipped with a 30 8 mm precolumn supplied by Knauer. The mobile phase was aqueous sulfuric acid (0.01 n) with a flow rate of 0.4 mL min 1 under isocratic conditions for 30 min. The injected volume was 20 mL. Monosaccharides identification was performed by comparison to several standards, namely, glucose, xylose, arabinose, fructose, and galactose. Calibration curves were obtained by the injection of glucose and fructose standard solutions in water with six different concentrations between 0.05 and 2.00 mg mL 1. In addition to the linearity, the limits of detection (LOD) and quantification (LOQ) were estimated using the signal-to-noise ratio approach (n = 6).[26a] The calibration curves and additional relevant analytical data are shown in Table S1. The concentrations were calculated based on triplicate injections, and the mean value was computed in each case.
www.chemsuschem.org B; 3–35 min, 0–10 % B; 35–47 min, 10–15 % B; 47–52 min, 15 % B; 52–54 min, 15–20 % B; 54–57 min, 20–50 % B; 57–62 min, 50–100 % B; 62–72 min, 100–0 % B, followed by re-equilibration of the column for 10 min before the next run. Double online detection was performed by the DAD at 280 and 340 nm, and UV spectra in the range of 210–600 nm were also recorded. Before the injection, the extract was dissolved in H2O (HPLC grade) to obtain a final concentration of 8 mg mL 1 and filtered through a 0.2 mm Nylon syringe filter. The HPLC was coupled to a LCQ Fleet ion-trap mass spectrometer (ThermoFinnigan, San Jose, CA, USA), equipped with an ESI source and operated in the negative mode. The conditions were the same as described previously.[31] Data acquisition was performed by using the Xcalibur data system (ThermoFinnigan, San Jose, CA, USA). For the quantification of phenolic compounds, calibrations curves were obtained from the HPLC injection of gallic and ellagic acids and isorhamnetin standard solutions in H2O with six different concentrations between 0.05 and 40 mg mL 1. The linearity, LOD, and LOQ[26a] were estimated, and the relevant analytical data are shown in Table S1. The quantification of individual compounds was accomplished with calibration data for the most similar standard as no pure reference compounds were available for some of them. The concentrations were calculated based on triplicate injections, and the mean value was computed in each case.
Analysis of metal NPs The optical spectra were recorded by using a Jasco V-560 UV/Vis spectrophotometer using 100 scans per min with a band width of 5.0 nm. Metallic NPs were extracted from the initial solution by high-speed centrifugation (18 000 rpm) by using a Sigma 3–30k centrifuge, and the NPs were washed thoroughly several times with distilled water. STEM images were obtained by using a fieldemission gun (FEG) SEM Hitachi SU70 microscope operated at 15 kV and fitted with an energy dispersive X-ray spectroscopy (EDX) accessory (EDX Detector: Bruker AXS, Software: Quantax). Samples were prepared by placing a colloid drop directly onto a carbon-coated copper grid and then allowing the solvent to evaporate. The average diameter of the NPs was determined by analysis of STEM images by using the ImageJ program (at least 50 NPs were analyzed in each case). The particle size distributions and the Zeta potential were evaluated by DLS analysis by using a Zetasizer Nano Series analyzer (Malvern Instruments). FTIR spectra of the NPs were collected by using a Bruker optics tensor 27 spectrometer coupled to a horizontal attenuated total reflectance (ATR) cell using 256 scans at a resolution of 4 cm 1. The Au and Ag nanophases were identified by powder XRD by using a Philips X’Pert instrument with CuKa radiation (l = 1.543178 ) at 40 kV and 50 mA.
Acknowledgements Analysis of phenolic compounds by HPLC–DAD–MS The HPLC system consisted of a variable loop Accela autosampler (200 vial capacity set at 15 8C), an Accela 600 LC pump, and an Accela 80 Hz DAD detector (Thermo Fisher Scientific, San Jose, CA, USA). The separation of the compounds was performed at 25 8C with a flow rate of 200 mL min 1 by using a Discovery C-18 column (150 2.1 mm 5 mm) equipped with a precolumn both supplied by Supelco (Agilent Technologies). The injection volume in the HPLC system was 10 mL, and the mobile phase consisted of water/ acetonitrile (90:10 v/v) (A) and acetonitrile (B) both with 0.1 % of formic acid. The following linear gradient was used: 0–3 min, 0 % 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
This work was financed by FEDER funds by Programa Operacional Factores de Competitividade-COMPETE and by national funds by Fundażo para a CiÞncia e Tecnologia (FCT) within the CICECO project FCOMP-01-0124-FEDER-037271 (Ref. PEst-C/CTM/ LA0011/2013). The authors further wish to thank the FCT for postdoctoral grants to S.A.O.S. (SFRH/BPD/84226/2012) and R.J.B.P. (SFRH/BPD/89982/2012) and for funding from TEMA (PEst-C/EME/ UI0481/2013) and QOPNA (PEst-C/QUI/UI0062/2013). C.S.R.F. and P.A.A.P.M. also acknowledge the FCT/MCTES for a research contract under the Program Investigador FCT 2012 and 2013, respecChemSusChem 2014, 7, 2704 – 2711
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CHEMSUSCHEM FULL PAPERS tively. Microscopy was supported by the Rede Nacional de Microscopia Eletrnica (RNME-Pole UA FCT) project REDE/1509/RME/ 2005. Keywords: gold · green chemistry · nanoparticles · reduction · silver [1] a) M. C. Daniel, D. Astruc, Chem. Rev. 2004, 104, 293 – 346; b) Y. Volokitin, J. Sinzig, L. J. deJongh, G. Schmid, M. N. Vargaftik, I. I. Moiseev, Nature 1996, 384, 621 – 623. [2] Y. Xia, Y. Xiong, B. Lim, S. E. Skrabalak, Angew. Chem. Int. Ed. 2009, 48, 60 – 103; Angew. Chem. 2009, 121, 62 – 108. [3] P. Raveendran, J. Fu, S. L. Wallen, J. Am. Chem. Soc. 2003, 125, 13940 – 13941. [4] V. Sanchez-Mendieta, A. R. Vilchis-Nestor in Nanotechnology and Nanomaterials, Noble Metals (Ed.: Y.-H. Su), InTech, Rijeka, 2012, pp. 391 – 408. [5] S. He, Z. Guo, Y. Zhang, S. Zhang, J. W. Gu, Mater. Lett. 2007, 61, 3984 – 3987. [6] A. Ahmad, P. Mukherjee, S. Senapati, D. Mandal, M. I. Khan, R. Kumar, M. Sastry, Colloids Surf. B 2003, 28, 313 – 318. [7] V. Kumar, S. K. Yadav, J. Chem. Technol. Biotechnol. 2009, 84, 151 – 157. [8] J. Huang, Q. Li, D. Sun, Y. Lu, Y. Su, X. Yang, H. Wang, Y. Wang, W. Shao, N. He, J. Hong, C. Chen, Nanotechnology 2007, 18, 105104. [9] N. Durn, P. D. Marcato, M. Duran, A. Yadav, A. Gade, M. Rai, Appl. Microbiol. Biotechnol. 2011, 90, 1609 – 1624. [10] S. Iravani, Green Chem. 2011, 13, 2638 – 2650. [11] a) J. Huang, G. Zhan, B. Zheng, D. Sun, F. Lu, Y. Lin, H. Chen, Z. Zheng, Y. Zheng, Q. Li, Ind. Eng. Chem. Res. 2011, 50, 9095 – 9106; b) B. Zheng, T. Kong, X. Jing, T. Odoom-Wubah, X. Li, D. Sun, F. Lu, Y. Zheng, J. Huang, Q. Li, J. Colloid Interface Sci. 2013, 396, 138 – 145. [12] a) Y. Zhou, W. Lin, J. Huang, W. Wang, Y. Gao, L. Lin, Q. Li, L. Lin, M. Du, Nanoscale Res. Lett. 2010, 5, 1351 – 1359; b) S. Li, Y. Shen, A. Xie, X. Yu, L. Qiu, L. Zhang, Q. Zhang, Green Chem. 2007, 9, 852 – 858. [13] a) S. A. O. Santos, J. J. Villaverde, C. M. Silva, C. P. Neto, A. J. D. Silvestre, J. Supercrit. Fluids 2012, 71, 71 – 79; b) S. A. O. Santos, C. S. R. Freire, M. R. M. Domingues, A. J. D. Silvestre, C. P. Neto, J. Agric. Food Chem. 2011, 59, 9386 – 9393. [14] a) Z. Sadowski in Nanotechnology and Nanomaterials, Silver Nanoparticles (Ed.: D. P. Perez), InTech, Rijeka, 2010, pp. 257 – 276; b) R. J. B. Pinto, P. A. A. P. Marques, C. P. Neto, T. Trindade, S. Daina, P. Sadocco, Acta Biomater. 2009, 5, 2279 – 2289.
2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chemsuschem.org [15] L. M. Liz-Marzn, Langmuir 2006, 22, 32 – 41. [16] R. R. Naik, S. J. Stringer, G. Agarwal, S. E. Jones, M. O. Stone, Nat. Mater. 2002, 1, 169 – 172. [17] a) S. S. Shankar, A. Rai, A. Ahmad, M. Sastry, J. Colloid Interface Sci. 2004, 275, 496 – 502; b) A. J. Bard, R. Parsons, J. Jordan in Monographs in Electroanalytical Chemistry and Electrochemistry, 1edst ed(Ed.: A. J. Bard), Marcel Dekker, Inc., New York, 1985. [18] V. K. Sharma, R. A. Yngard, Y. Lin, Adv. Colloid Interface Sci. 2009, 145, 83 – 96. [19] L. J. Bellamy, The Infrared Spectra of Complex Molecules, Vol. 1, 3rd ed., Wiley, New York, 1975. [20] R. Fido, E. N. C. Mills, N. Rigby, P. Shewry in Protein Purification Protocols, Vol. 244 (Ed.: P. Cutler), Humana Press 2004, pp. 21 – 27. [21] C. V. Gonzlez Lpez, M. d. C. C. Garca, F. G. A. Fernndez, C. S. Bustos, Y. Chisti, J. M. F. Sevilla, Bioresour. Technol. 2010, 101, 7587 – 7591. [22] Z. Lin, J. Wu, R. Xue, Y. Yang, Spectrochim. Acta Part A 2005, 61, 761 – 765. [23] a) P. Shanmugavelan, S. Y. Kim, J. B. Kim, H. W. Kim, S. M. Cho, S. N. Kim, S. Y. Kim, Y. S. Cho, H. R. Kim, Carbohydr. Res. 2013, 380, 112 – 117; b) C. Ma, Z. Sun, C. Chen, L. Zhang, S. Zhu, Food Chem. 2014, 145, 784 – 788. [24] A. K. Mittal, Y. Chisti, U. C. Banerjee, Biotechnol. Adv. 2013, 31, 346 – 356. [25] L. Ortega-Arroyo, E. San Martin-Martinez, M. A. Aguilar-Mendez, A. CruzOrea, I. Hernandez-P rez, C. Glorieux, Starch/Staerke 2013, 65, 814 – 821. [26] a) S. A. O. Santos, C. Vilela, C. S. R. Freire, C. P. Neto, A. J. D. Silvestre, J. Chromatogr. B 2013, 938, 65 – 74; b) L. Boulekbache-Makhlouf, E. Meudec, M. Chibane, J. P. Mazauric, S. Slimani, M. Henry, V. Cheynier, K. Madani, J. Agric. Food Chem. 2010, 58, 12615 – 12624. [27] M. Boukhris, M. S. J. Simmonds, S. Sayadi, M. Bouaziz, Phytother. Res. 2013, 27, 1206 – 1213. [28] L. Pourcel, J.-M. Routaboul, V. Cheynier, L. Lepiniec, I. Debeaujon, Trends Plant Sci. 2007, 12, 29 – 36. [29] D. MubarakAli, N. Thajuddin, K. Jeganathan, M. Gunasekaran, Colloids Surf. B 2011, 85, 360 – 365. [30] N. L. Conklin-Brittain, E. S. Dierenfeld, R. W. Wrangham, M. Norconk, S. C. Silver, J. Chem. Ecol. 1999, 25, 2601 – 2622. [31] S. A. O. Santos, J. J. Villaverde, A. F. Sousa, J. F. J. Coelho, C. P. Neto, A. J. D. Silvestre, Ind. Crops Prod. 2013, 47, 262 – 269. [32] S. Dall’Acqua, P. Minesso, B. B. Shresta, S. Comai, P. K. Jha, M. B. Gewali, E. Greco, R. Cervellati, G. Innocenti, Molecules 2012, 17, 1686 – 1697.
Received: February 28, 2014 Published online on August 1, 2014
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