International Journal of Polymer Science

International Journal of Polymer Science / 2015 / Article

Research Article | Open Access

Volume 2015 |Article ID 864729 |

Petruta Mihaela Matei, Pablo Martín-Ramos, Mercedes Sánchez-Báscones, Salvador Hernández-Navarro, Adriana Correa-Guimaraes, Luis M. Navas-Gracia, Cassyo Araujo Rufino, M. Carmen Ramos-Sánchez, Jesús Martín-Gil, "Synthesis of Chitosan Oligomers/Propolis/Silver Nanoparticles Composite Systems and Study of Their Activity against Diplodia seriata", International Journal of Polymer Science, vol. 2015, Article ID 864729, 11 pages, 2015.

Synthesis of Chitosan Oligomers/Propolis/Silver Nanoparticles Composite Systems and Study of Their Activity against Diplodia seriata

Academic Editor: Vitor Sencadas
Received11 Jan 2015
Revised08 Mar 2015
Accepted14 Mar 2015
Published29 Mar 2015


The synthesis and characterization of composites of oligomeric chitosan with propolis extract which allow the incorporation of a third component (silver nanoparticles) are reported, together with their application in aqueous or hydroalcoholic solutions with a view to the formation of adhesive substances or nanofilms for the protection of vineyards against harmful xylophagous fungi. The antimicrobial properties of the association of the two biological products or those resulting from the incorporation of silver nanoparticles (NPs) are studied and discussed. The efficacy of the chitosan oligomers/propolis/silver NPs ternary system is assessed in vitro for Diplodia fungi. A preliminary study on the convenience of replacing propolis with gentisic acid is also presented.

1. Introduction

With regard to hybrid materials, there is a growing realization of the importance of focusing on low-cost, environmentally friendly biopolymers. The natural, biodegradable, and biocompatible chitosan (glucosamine polymer with -1,4 bonds), formed by the alkaline N-deacetylation of chitin, is one the most promising candidates, due to its ability to form films, transparency, nontoxicity, excellent adsorption features, and so forth [1].

In addition, chitosan has been found to have antimicrobial properties [2, 3], in which the size of its oligomers plays a key role [4]. For example, heptamers and higher oligomers induce an increase of pisatin, an antifungal substance from Pisum sativum L. [5]. It has also been demonstrated that polymeric chitosan and chitosan oligomers induce phytoalexins or antimicrobial compounds that help limit the spread of pathogens [6]. Chemical synthesis of different sizes of chitosan oligomers with specific biological activity has been described by Kuyama et al. [7].

Several possible mechanisms have been proposed to explain the antibacterial properties of chitosan: it is known that positively charged amine groups are capable of interacting with the negatively charged bacterial cell membrane and, in addition, chitosan may also bind to DNA, leading to inhibition of mRNA and proteins synthesis [8].

On the other hand, other substances—such as propolis—are also known to have antimicrobial properties and are extensively used in traditional medicine [9, 10]. Propolis is a natural resinous hive product collected by honeybees from various plant sources [11] which contains over 150 chemical species (such as coumarins, flavonoids, polyphenols, phenolic aldehydes, sesquiterpene quinines, amino acids and steroids) [12]. Its strong antimicrobial activity may be due to the high content of total phenols and flavonoids [9]. Propolis has also been found to have applications as an antioxidant and in food preservation [13].

Silver nanoparticles have been deemed as one of the most promising antimicrobial species from a nanotechnology-based approach, since their activity is very broad and is well above that of raw silver. For example, silver ions can bind to negatively charged bacterial peptidoglycan walls and can diffuse into bacterial cells and bind to DNA bases, leading to bacterial death and/or inhibiting the replication and transcription processes and preventing further bacterial production [14]. Moreover, the generation of reactive oxygen species, which leads to nanotoxicity processes, is also a well-established antimicrobial mechanism. The main disadvantages that limit the use of nanosilver are its ease of aggregation and the uncontrolled release of silver ions and their cytotoxicity potential [15].

The combination of polymers and nanosilver may synergistically improve their antimicrobial effects, and the use of in situ synthesis methods allows its incorporation into the polymer matrix attaining uniform distributions and avoiding aggregation. We herein report a facile synthesis procedure based on a sonochemistry approach for the preparation of a new ternary composite material—which consists of chitosan oligomers, propolis, and silver NPs—and study its antimicrobial properties against Diplodia seriata fungus.

It is known that the xylophagous fungi that are present in greater proportion and that irremediably cause plant death of the vine are, firstly, Diplodia seriata, followed by Phaeoacremonium aleophilum, Cylindrocarpon spp., Fomitiporia punctata Murrill, Fomitiporia mediterranea, Phaeomoniella chlamydospora, Botryosphaeria dothidea, Stereum hirsutum and Eutypa lata [16]. This disease is considered highly virulent because the infection is caused by fungi entry through wounds caused by vine pruning [1719].

In the literature, Prakongkha et al. [20] proposed the use of bare chitosan in vineyards; Matei et al. reported that chitosan can inhibit esca (a vine disease that ravaged vineyards from Cognac to Bordeaux, France) mycelia growth by 90% [21]; and Aziz et al. [22] successfully demonstrated that octameric chitosan oligomers doped with copper sulfate can induce defense reactions of the vine and resistance to gray mold and mildew. Other authors have assessed the antimicrobial properties for chitosan/nanosilver [2326]. Nonetheless, to the best of the authors’ knowledge, no studies have been conducted on ternary composites. In this paper, we assess how the presence of chitosan oligomers—to which phenolic compounds such as phenolic acids and/or flavonoids have been incorporated—affords the composite with antioxidant and soluble compounds that can interact to form chelates or bridges with inorganic species in aqueous solution, such as silver NPs, leading to novel antimicrobial agents, without posing any danger to the plant or to the substrate wherein the composite material is applied. The association of the three components attempts to further improve the antimicrobial performance of their active principles in comparison with their separate application.

2. Materials and Methods

2.1. Reagents and Characterization Equipment

Medium molar mass chitosan (CAS number 9012-76-4) was purchased from Sigma Aldrich Química SL (Madrid, Spain) and from Hangzhou Simit Chemical Technology Co., Ltd. (Hangzhou, China). Propolis came from Burgos region (Spain), in the Duero river basin, and has a polyphenols and flavonoids content of ca. 10% wt/v. Silver nitrate (CAS number 7761-88-8) and malt extract agar (Reference 105398) were supplied by Merck Millipore (Darmstadt, Germany). Potassium methoxide solution (25 wt.% in methanol, CAS number 865-33-8) and ethanol (puriss. p.a., ACS reagent, CAS number 64-17-5) were also purchased from Sigma Aldrich Química SL. Gentisic acid (CAS number 490-79-9) was supplied by Shanghai Vincor BioEngineering Co., Ltd. (Shanghai, China). The isolated Diplodia seriata mycelium (Y207 1-1c) was supplied by ITACYL (Castilla y León, Spain).

An ultrasonic machine, model CSA 20-S500, 20 KHz has been used for solutions sonication.

X-ray powder diffractograms of the samples were obtained using a Bruker D8 Advance Bragg-Brentano diffractometer, in reflection geometry.

Infrared spectra were recorded with a Thermo Nicolet 380 FT-IR apparatus equipped with a Smart Orbit Diamond ATR system, in order to identify the chemical functional groups.

Optical absorption spectra of the silver NPs in the UV-Vis region were recorded with a Shimazdu UV-2450 UV-Vis spectrophotometer.

Scanning electron microscopy (SEM) images were collected with a FEI-Quanta 200FEG equipped with a Genesis energy-dispersive X-ray (EDS) spectrometer system. Transmission electron microscope (TEM) micrographs were collected with a JEOL JEM-FS2200 HRP equipped with an Oxford Instruments INCA Energy TEM 250 EDS probe.

2.2. Culture Media and Activity Assays

The fungicidal action of the products under study was tested in vitro using malt extract agar (MEA) as a culture medium. In the study on the fungus, previously isolated Diplodia seriata mycelia (Y207 1-1c) were used, which had been replicated in Petri dishes two weeks in advance at 25°C. From the periphery of the pure culture, 7 mm diameter disks were cut and then transplanted into Petri dishes prepared with MEA in combination with one of the products under study (namely, bare chitosan oligomers, propolis, chitosan oligomers/propolis, chitosan oligomers/silver NPs, and chitosan oligomers/propolis/silver NPs). For the preparation of the base medium, 35 g of MEA was dissolved in 750 mL of distilled water, to which 2 mL of antibiotic (chloramphenicol) was added to prevent bacterial contamination, and the flasks with the mixture were then sterilized in an autoclave at 100°C for 45 min. For the control, the culture medium “as-is” was placed in sterile Petri dishes (20 mL/dish) and allowed to cool before replication of the Di. seriata fungal mycelia. To prepare the Petri dishes for the different mixtures or combinations of chitosan oligomers/propolis/silver NPs, the culture medium was cooled to 50°C and, at this temperature, 1.5 mL of the different solutions (see Table 1) was mixed with 18.5 mL of MEA and poured into each Petri dish. The solutions were then allowed to cool down to room temperature (RT) prior to replication of the fungus mycelia. Growth measurements were performed in triplicate. The diameter of fungal growth was measured on a daily basis for 20 days, and the inhibition percentage (IP) was calculated taking the pure MEA culture (control) as a reference according to the following equation [27]:where is the diameter of the mycelium in the control (pure MEA) and is the diameter of the mycelium of the sample mixed with one of the antimicrobial composites.


() Chitosan 20 mg of chitosan/1 mL of H2O

() Propolis50 mg of propolis/1 mL of H2O : ethanol (7 : 3)

() Silver NPs170 µg/1 mL of H2O

() Chitosan oligomers/propolis1 : 1 ratio of solution () and solution ()

() Chitosan oligomers/silver NPs8 : 1 ratio of solution () and solution ()

() Chitosan oligomers/propolis/silver NPs8 : 8 : 1 ratio of solutions (), (), and (), respectively

2.3. Synthesis of Solutions and Films of Chitosan Oligomers, Chitosan Oligomers/Propolis, and Chitosan Oligomers/Propolis/Ag NPs
2.3.1. Chitosan Oligomers Preparation

Chitosan oligomers aqueous solutions were prepared from a solution of commercial medium molar mass chitosan (with molar masses in the 190000–310000 g/mol range for the Sigma Aldrich product and with molar masses in the 140000–300000 g/mol range for the Hangzhou Simit Chemical Technology Co., product) in AcOH 2% at pHs 4–6. The hydrolysis was performed by stirring for 12 hours followed by 3–6 sonication periods (5 minutes each), at temperatures in the 30 to 60°C range and with H2O2 concentrations ranging from 0.3 to 0.6 M, obtaining oligomers with molar masses in the 6000 to 2000 g/mol range, respectively, in agreement with the analogous microwave-based procedure reported by Sun et al. [28]. The molar mass of the chitosan samples was determined by measuring the viscosity, in agreement with Yang et al. [29], in a solvent of 0.20 mol/L NaCl + 0.1 mol/L CH3COOH at 25°C using an Ubbelohde capillary viscometer. Molar masses were determined using the Mark-Houwink equation = 1.81 × 10−3 M0.93 [30].

The solutions were then decanted to remove any water insoluble material, were allowed to rest till cloudiness was observed, and were centrifuged to isolate the chitosan oligomers. These were redissolved again in AcOH 0.5%, obtaining the solutions for the assays.

2.3.2. Propolis Extraction

The propolis solution was prepared by grinding raw propolis to fine powder and subsequent extraction of the active ingredients by maceration in a hydroalcoholic solution 7 : 3 (v/v) for one week at room temperature. A hydroalcoholic medium was chosen over absolute ethanol because it results in wax-free tinctures containing higher amounts of polyphenolic substances [31]. The resulting solution was then percolated (1 L/min) and filtrated with a stainless steel 220 mesh to remove any residue, followed by concentration at a temperature below 60°C with ultrasound equipment to finally obtain a clarified propolis extract.

2.3.3. Silver NPs Preparation

Silver nanoparticles were prepared by a sonication method, without resourcing to UV stabilization (used, e.g., in [32]), as follows: an aqueous solution of AgNO3 (50 mM) was treated with sodium citrate (30 mM) and the resulting solution was cooled and stirred at a temperature between 5 and 10°C. Subsequently, it was deoxygenated with an inert gas (N2) for over 30 minutes and the pH was adjusted between 7 and 8. Polyvinylpyrrolidone was added to prevent the silver nanoparticles aggregation. A 10 mM solution of NaBH4 (reducing agent) was then added dropwise; the first droplet made the solution turn from colorless to yellowish and successive droplets led to an intensification of the yellow color (care had to be taken so as to avoid an excess of reducing agent, which would lead to a brownish color). After vigorous stirring for one hour, the yellowish solution was sonicated for 3–5 minutes and then allowed to rest and stabilize for at least 24 hours in a refrigerator at 5°C.

2.3.4. Binary and Ternary Solutions Preparation

Chitosan oligomers/propolis solution was prepared by mixing the two components in a 1 : 1 w/v ratio (Table 1), followed by sonication, obtaining a caramel colored gel or precipitate, similar to that obtained by Mascheroni et al. [33] using an alternative membrane-based procedure.

Ternary solutions were prepared by adding a mixture of propolis extract and silver NPs solutions to the solution containing chitosan oligomers (Table 1). A few droplets of a solution of potassium methoxide in methanol 25% were added to adjust the pH to 4–6 (the methanol which results by hydrolysis is below 10 ppm, and at this concentration it is innocuous to the fungi). The resulting solutions were stirred for 1 hour and sonicated for 2-3 minutes and remained clear. These solutions were mixed with MEA for the activity assays.

When a film was required (e.g., for SEM/TEM characterization), glyoxal 0.25% v/v was added to facilitate the interaction and the pH was adjusted to 8-9. Upon stirring and sonication, the solutions were allowed to rest for approximately 1 hour and gels appeared at the bottom of the flasks. These materials were isolated from the solution, were poured in polypropylene substrates [34], and were dried under vacuum at 20°C in dry atmosphere, yielding films with thicknesses in the 0.3 to 0.6 mm range. Details on these procedures have recently been the subject of a patent (Application number ES P201431591, filing date: 30/10/2014).

3. Results and Discussion

3.1. Vibrational Characterization

The ATR-FTIR spectra of commercial chitosan, chitosan oligomers, propolis, chitosan oligomers/propolis binary composite, and chitosan oligomers/propolis/silver NPs ternary composite are depicted in Figure 1. The assigned characteristic FTIR absorption bands derived from Figure 1 are summarized in Table 2.

ChitosanChitosan oligomersPropolisChitosan/propolisChitosan/propolis/silver NPsAssignation

3290328532853300υ(OH) overlapped to υs(N–H) [35, 36]

29702972C–H bands of aromatic compounds

29302929υas(C–H) in –CH2 [35]

286728732873υs(C–H) in –CH3 [35]

1707υ(–C=O) of the amide group CONHR in chitosan [36, 38] and υ(–C=O) of flavonoids and lipids, found in propolis [39]/aromatic ring deformations [39]

156815541560υ(–C=O) protonated amide group [35]
δ(NH3) protonated amine group

1417141914041409δ(OH) [40]

1372137513771373δ(–CH3) [41]

133413411333Typical propolis C–O and C–OH vibrations [39]

13181317υs(–CH3) tertiary amide [35]
ω(–CH2) + OH in-plane deformation

1260υ(C–O–H) [35]

1152114811501160υas(CO) in oxygen bridge resulting from deacetylation of chitosan

11361133Alkenes bands from propolis (coumaroyl glycerol) [42]

10601081107810661084υ(CO) of the ring C–O–H, C–O–C and CH2CO [35, 39]

Bands from propolis

920923922Bands from propolis

892892872ω(C–H) of the saccharide structure [35]

836835Band from propolis

Typical bands from binary and ternary formulations

Analyses of both the binary chitosan-propolis formulation and the ternary composite showed that the first band in the ATR-FTIR spectra was shifted to higher wavelengths in comparison with the bare chitosan spectrum (from 3285–3290 to 3300 cm−1) [35, 36], suggesting that effective hydrogen bonding occurred between chitosan and propolis. This interaction is an indicator of a synergy between the two products.

Another significant feature for the ternary composite spectrum was an obvious shift to 1700 cm−1 of the band located at around 1650 cm−1, accompanied by an increase in the intensity, proving the responsibility of chitosan amino group for silver envelopment [3639]. In the same way, the change in the OH band intensity in the ternary composite spectra at 1409 cm−1 (assigned to the OH deformation vibrations of the secondary alcohols in the pyranose monomers) revealed that the hydroxyl groups may have also participated in the stabilization of the silver nanoparticles via interaction of Ag+ with the electron abundant oxygen atoms of the hydroxyl groups of the chitosan. These results are in agreement with those reported by Boanić et al. [40]. Moreover, the increase in intensity of the band at 1380 cm−1 (attributed to CH3 bending in NHCOCH3 group [41]) can also be attributed to interactions with silver NPs.

In the particular case of the band at 1133 cm−1, characteristic of ester groups (–C–O stretching) from propolis components [42], a change in the intensity of the band was also observed, attributable to interactions with silver NPs.

Unassigned typical bands from binary and ternary formulations have been found at around 650 and 616 cm−1.

3.2. X-Ray Characterization

The X-ray diffraction study of the chitosan oligomers exhibits the expected broad peaks at and (Figure 2), in good agreement with Kumar et al. [43]. However, the peak observed for chitosan at disappeared and the very broad peak at became weak in the chitosan-propolis sample. These results suggest that chitosan has good compatibility, which leads to the formation of a composite with an amorphous form, suitable for bioapplications [44].

3.3. Silver NPs Characterization

Silver nanoparticles were characterized by UV-Vis absorption, XRD, and TEM analysis, revealing the formation of highly pure, crystalline silver nanoparticles of ca. 30 nm (see Section 3.4). The UV-Vis spectrum (Figure 3(a)) showed the expected intense surface plasmon resonance (SPR) band at around 420 nm [45]. The X-ray powder diffraction pattern (Figure 3(b)) matched well with the standard patterns of silver (JCPDS number 04-0783). All the peaks of the pattern can be readily indexed to face-centered-cubic silver, where the diffraction peaks at 38.2, 44.5, 64.5, and 77.5° can be ascribed to the reflection of (), (), (), and () planes, respectively.

3.4. Textural Properties

The texture of the ternary composite films has been studied by SEM and TEM. The SEM micrographs (Figure 4(a))—similar to those reported by Moharram et al. [26]—show the good homogeneity of the materials under study, while the TEM micrograph (Figure 4(b)) allows estimating the size of the silver NPs, in the 24 to 35 nm range.

3.5. Antifungal Activity of Chitosan

As noted above, the size of the chitosan chains has a significant impact on its antimicrobial activity [4]. Consequently, prior to the evaluation of the different composites, a first study was conducted only for chitosan. Two commercial medium molar mass chitosan solutions were compared with the chitosan oligomers prepared according to the procedure described in Section 2.3.1. Whereas the chitosan oligomers led to mycelia death, the fungus was able to grow in the culture media with medium molar mass chitosan (see Figure 5). Consequently, only chitosan oligomers have been used in the studies described below.

3.6. Antifungal Activity of the Propolis, Chitosan Oligomers/Propolis, and Chitosan Oligomers/Propolis/Silver NPs Colloidal Solutions against Diplodia seriata Fungus

The antifungal activity has been studied for the aqueous and hydroalcoholic mixtures of bare propolis, chitosan oligomers/propolis, and chitosan oligomers/propolis/silver NPs, assessing the influence of low alcohol concentrations (ca. 5%) on the mixtures and analyzing the growth diameter of the Diplodia seriata mycelia. As indicated above, the assays have been conducted in triplicate biological repetitions, and all the results are in average.

For bare propolis (see Figure 6(a)), in presence of alcohol, the mycelium diameter increased exponentially in an initial stage, with a slope change on the 7th day and reaching its maximum diameter (77-78 mm) on the 13th/14th day. When alcohol was removed from the propolis extract by sonication, the activity of the fungus decreased, showing a slower and progressive growth and reaching a maximum diameter of 22 mm.

When the chitosan oligomers/propolis composite (see Figure 6(b)) was tested in the presence of alcohol, exponential growth took place up to the 7th day, in which there was also a slope change and the growth started to be less pronounced. In comparison with bare propolis, the antifungal activity was significantly enhanced, since the maximum mycelium diameter in this case was 27 nm. Further, when alcohol was completely removed, no fungal growth took place (and the diameter remained constant at 7 mm).

A similar behavior was observed for the chitosan oligomers/propolis/silver NPs composites (Figure 6(c)); in the presence of alcohol, there was an almost linear increase in the mycelia diameter, reaching 41 mm, whereas in the absence of alcohol no fungal growth occurred. It can thus be inferred that the antifungal activity of the material with silver NPs was better than that of bare propolis but worse than that of the binary composite. Nevertheless, it is worth noting that the presence of silver NPs reduced the standard deviation values and made the growth more linear (i.e., less logarithmic) in comparison with the assays conducted for bare propolis or chitosan oligomers/propolis composite in hydroalcoholic medium.

A wider comparison (summarized in Figure 7), which also includes bare chitosan (A) and chitosan oligomers/silver NPs mixture (D), further confirms previous results: the presence of alcohol significantly favored the growth of the fungus. On the contrary, when the alcohol was removed by sonication at a temperature lower than 60°C, the fungus hardly grew for bare propolis (P)—which showed an inhibition percentage of 75%—or did not grow when chitosan oligomers/propolis (B) or chitosan oligomers/propolis/silver NPs (C) composites were used. Consequently, alcohol removal from the binary or ternary mixtures is essential so as to improve the antifungal activity of these materials. This is in agreement with the findings of Zhao et al. [46], who reported that a concentration of 0.5–2% ethanol stimulated the growth of fungi, since it was partially used as a carbon source during fermentation.

3.7. Future Lines of Research: Replacement of Propolis with Gentisic Acid

Propolis polyphenols, such as gentisic acid (2,5-dihydroxybenzoic acid) or homogentisic acid (2,5-dihydroxyphenylacetic acid), are known to have antifungal activity [47] and have been shown to increase the fungicidal activity of other chemicals (e.g., fludioxonil, a phenylpyrrole fungicide) [48]. Further, they have a widespread occurrence, being found in citrus fruits (Citrus spp.), grapes (Vitis vinifera), sesame (Sesamum indicum), gentians (Gentiana spp.), and so forth, which are amongst the probable floral origins of the Mediterranean propolis used in our study, in agreement with Gülçin et al. [49]. Consequently, a preliminary assessment of the suitability of the former as a replacement of propolis extract has been conducted.

As it is shown in Figure 8, pure propolis, pure gentisic acid, and propolis : gentisic acid (1 : 1) solutions (with concentrations of 30 mg/mL for the propolis solution, 30 mg/mL for the gentisic acid solution, 1 : 1 ratio of the two solutions for the propolis : gentisic acid mixture, resp.) have been evaluated, and an improved performance of gentisic acid over propolis has been evidenced. Thus, an identification of the propolis components and their separate antifungal activity study needs to be conducted in future research.

4. Conclusions

The present work reports the synthesis by a facile chemical method of a composite system consisting of a colloidal suspension of silver nanoparticles in a chitosan-propolis biopolymer matrix. The materials have been characterized by X-ray powder diffraction studies, FTIR vibrational spectroscopy, UV-Vis absorption spectroscopy, and SEM and TEM microscopies. With a view to the application of these solutions to the formation of adhesive substances or nanofilms for the protection of vineyards against harmful xylophagous fungi, the influence of low alcohol concentrations (ca. 5%) on the growth diameter of the Diplodia seriata mycelia has been determined. The results are conclusive on the need of alcohol removal to improve the antifungal activity of these materials. Moreover, it is essential to use low molar mass chitosan oligomers, given that their inhibitory activity is significantly higher than that of medium molar mass chitosan. Finally, it is worth noting that further research is still required so as to evaluate the separate activity of the different components of propolis, as evinced by the enhanced antifungal behavior of gentisic acid in comparison with propolis, and on the synergistic effect of the three components of the composite.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.


This work was supported by funds from Junta de Castilla y León under Project VA233A12-1. The authors gratefully acknowledge the support of Dr. M. Avella (Microscopy Unit, Parque Científico, Universidad de Valladolid) and of Dr. J.A. Paixão (CEMDRX, Universidade de Coimbra) with the SEM and TEM analysis and the XRD measurements, respectively. Special thanks are due to Dr. M. T. Martín-Villullas (ITACYL) for supplying the fungus mycelium.


  1. M. N. V. R. Kumar, “A review of chitin and chitosan applications,” Reactive and Functional Polymers, vol. 46, no. 1, pp. 1–27, 2000. View at: Publisher Site | Google Scholar
  2. J. Rhoades and S. Roller, “Antimicrobial actions of degraded and native chitosan against spoilage organisms in laboratory media and foods,” Applied and Environmental Microbiology, vol. 66, no. 1, pp. 80–86, 2000. View at: Publisher Site | Google Scholar
  3. L. Y. Chung, R. J. Schmidt, P. F. Hamlyn, B. F. Sagar, A. M. Andrews, and T. D. Turner, “Biocompatibility of potential wound management products: fungal mycelia as a source of chitin/chitosan and their effect on the proliferation of human F1000 fibroblasts in culture,” Journal of Biomedical Materials Research, vol. 28, no. 4, pp. 463–469, 1994. View at: Publisher Site | Google Scholar
  4. L. A. Hadwiger, T. Ogawa, and H. Kuyama, “Chitosan polymer sizes effective in inducing phytoalexin accumulation and fungal suppression are verified with synthesized oligomers,” Molecular Plant-Microbe Interactions, vol. 7, no. 4, pp. 531–533, 1994. View at: Publisher Site | Google Scholar
  5. D. F. Kendra and L. A. Hadwiger, “Characterization of the smallest chitosan oligomer that is maximally antifungal to Fusarium solani and elicits pisatin formation in Pisum sativum,” Experimental Mycology, vol. 8, no. 3, pp. 276–281, 1984. View at: Publisher Site | Google Scholar
  6. D. F. Kendra, D. Christian, and L. A. Hadwiger, “Chitosan oligomers from Fusarium solani/pea interactions, chitinase/β-glucanase digestion of sporelings and from fungal wall chitin actively inhibit fungal growth and enhance disease resistance,” Physiological and Molecular Plant Pathology, vol. 35, no. 3, pp. 215–230, 1989. View at: Publisher Site | Google Scholar
  7. H. Kuyama, Y. Nakahara, T. Nukada, Y. Ito, and T. Ogawa, “Stereocontrolled synthesis of chitosan dodecamer,” Carbohydrate Research, vol. 243, no. 1, pp. C1–C7, 1993. View at: Publisher Site | Google Scholar
  8. P. K. Dutta, S. Tripathi, G. K. Mehrotra, and J. Dutta, “Perspectives for chitosan based antimicrobial films in food applications,” Food Chemistry, vol. 114, no. 4, pp. 1173–1182, 2009. View at: Publisher Site | Google Scholar
  9. N. Nedji and W. Loucif-Ayad, “Antimicrobial activity of Algerian propolis in foodborne pathogens and its quantitative chemical composition,” Asian Pacific Journal of Tropical Disease, vol. 4, no. 6, pp. 433–437, 2014. View at: Publisher Site | Google Scholar
  10. V. Bankova, “Recent trends and important developments in propolis research,” Evidence-Based Complementary and Alternative Medicine, vol. 2, no. 1, pp. 29–32, 2005. View at: Publisher Site | Google Scholar
  11. E. Ghisalberti, “Propolis: a review,” Bee World, vol. 60, no. 2, pp. 59–84, 1979. View at: Google Scholar
  12. M. Marcucci, “Propolis: chemical composition, biological properties and therapeutic activity,” Apidologie, vol. 26, no. 2, pp. 83–99, 1995. View at: Publisher Site | Google Scholar
  13. N. Kalogeropoulos, S. J. Konteles, E. Troullidou, I. Mourtzinos, and V. T. Karathanos, “Chemical composition, antioxidant activity and antimicrobial properties of propolis extracts from Greece and Cyprus,” Food Chemistry, vol. 116, no. 2, pp. 452–461, 2009. View at: Publisher Site | Google Scholar
  14. S. Shrivastava, T. Bera, A. Roy, G. Singh, P. Ramachandrarao, and D. Dash, “Characterization of enhanced antibacterial effects of novel silver nanoparticles,” Nanotechnology, vol. 18, no. 22, Article ID 225103, 2007. View at: Publisher Site | Google Scholar
  15. A. R. Silva and G. Unali, “Controlled silver delivery by silver-cellulose nanocomposites prepared by a one-pot green synthesis assisted by microwaves,” Nanotechnology, vol. 22, no. 31, Article ID 315605, 2011. View at: Publisher Site | Google Scholar
  16. J. Grosman, “Observatoire national et régional des maladies du bois: bilan et perspectives de 4 années d'observations,” in Eurovity, pp. 8–17, Institut Français de la Vigne et du Vin, Angers, France, 2008. View at: Google Scholar
  17. P. Larignon, J. Dupont, and B. Dubos, “L'esca de la vigne: Quelques éléments sur la biologie de deux des agents associés: Phaeoacremonium aleophilum et Phaeomoniella chlamydospora,” Phytoma—La Défense des Végétaux, no. 527, pp. 30–35, 2000. View at: Google Scholar
  18. P. Larignon and B. Dubos, “Le Black Dead Arm: maladie nouvelle à ne pas confondre avec l'esca,” in Phytoma—La Défense des Végétaux, pp. 26–29, 2001. View at: Google Scholar
  19. P. Larignon, R. Fulchic, L. Cere, and B. Dubos, “Observation on black dead arm in French vineyards,” Phytopathologia Mediterranea, vol. 40, no. 3, pp. 336–342, 2001. View at: Google Scholar
  20. I. Prakongkha, M. Sompong, S. Wongkaew, D. Athinuwat, and N. Buensanteai, “Changes in salicylic acid in grapevine treated with chitosan and BTH against Sphaceloma ampelinum, the causal agent of grapevine anthracnose,” African Journal of Microbiology Research, vol. 7, pp. 557–563, 2013. View at: Google Scholar
  21. P. Matei, B. Iacomi, and G. Dragan, “Fungi associated with esca decline and their in vitro control by chitosan,” Scientific Papers, UASVM Bucharest, Series A, vol. 53, pp. 448–453, 2010. View at: Google Scholar
  22. A. Aziz, P. Trotel-Aziz, L. Dhuicq, P. Jeandet, M. Couderchet, and G. Vernet, “Chitosan oligomers and copper sulfate induce grapevine defense reactions and resistance to gray mold and downy mildew,” Phytopathology, vol. 96, no. 11, pp. 1188–1194, 2006. View at: Publisher Site | Google Scholar
  23. C. Gu, H. Zhang, and M. Lang, “Preparation of mono-dispersed silver nanoparticles assisted by chitosan-g-poly(ε-caprolactone) micelles and their antimicrobial application,” Applied Surface Science, vol. 301, pp. 273–279, 2014. View at: Publisher Site | Google Scholar
  24. P. T. S. Kumar, S. Abhilash, K. Manzoor, S. V. Nair, H. Tamura, and R. Jayakumar, “Preparation and characterization of novel β-chitin/nanosilver composite scaffolds for wound dressing applications,” Carbohydrate Polymers, vol. 80, no. 3, pp. 761–767, 2010. View at: Publisher Site | Google Scholar
  25. L. Guo, W. Yuan, Z. Lu, and C. M. Li, “Polymer/nanosilver composite coatings for antibacterial applications,” Colloids and Surfaces A: Physicochemical and Engineering Aspects, vol. 439, pp. 69–83, 2013. View at: Publisher Site | Google Scholar
  26. M. A. Moharram, S. K. H. Khalil, H. H. A. Sherif, and W. A. Khalil, “Spectroscopic study of the experimental parameters controlling the structural properties of chitosan–Ag nanoparticles composite,” Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy, vol. 126, pp. 1–6, 2014. View at: Publisher Site | Google Scholar
  27. T. Acosta, A. Avellaneda, J. Cuervo, and L. Sánchez, “Evaluacion de microbiota de tomillo (Thymus vulgaris), como aporte al manejo agroecologico de aromaticas en invernaderos de la Universidad Nacional,” in Perspectivas del Agronegocio de Hierbas Aromáticas Culinarias y Medicinales, pp. 135–138, Universidad Nacional de Colombia, Bogotá, Colombia, 2007. View at: Google Scholar
  28. T. Sun, D. Zhou, J. Xie, and F. Mao, “Preparation of chitosan oligomers and their antioxidant activity,” European Food Research and Technology, vol. 225, no. 3-4, pp. 451–456, 2007. View at: Publisher Site | Google Scholar
  29. Y. Yang, R. Shu, J. Shao, G. Xu, and X. Gu, “Radical scavenging activity of chitooligosaccharide with different molecular weights,” European Food Research and Technology, vol. 222, no. 1-2, pp. 36–40, 2006. View at: Publisher Site | Google Scholar
  30. G. G. Maghami and G. A. F. Roberts, “Evaluation of the viscometric constants for chitosan,” Die Makromolekulare Chemie, vol. 189, pp. 195–200, 1988. View at: Google Scholar
  31. R. G. Woisky and A. Salatino, “Analysis of propolis: some parameters and procedures for chemical quality control,” Journal of Apicultural Research, vol. 37, no. 2, pp. 99–105, 1998. View at: Google Scholar
  32. M. Montazer, A. Shamei, and F. Alimohammadi, “Synthesizing and stabilizing silver nanoparticles on polyamide fabric using silver-ammonia/PVP/UVC,” Progress in Organic Coatings, vol. 75, no. 4, pp. 379–385, 2012. View at: Publisher Site | Google Scholar
  33. E. Mascheroni, A. Figoli, A. Musatti et al., “An alternative encapsulation approach for production of active chitosan-propolis beads,” International Journal of Food Science and Technology, vol. 49, no. 5, pp. 1401–1407, 2014. View at: Publisher Site | Google Scholar
  34. E. Torlak and D. Sert, “Antibacterial effectiveness of chitosan-propolis coated polypropylene films against foodborne pathogens,” International Journal of Biological Macromolecules, vol. 60, pp. 52–55, 2013. View at: Publisher Site | Google Scholar
  35. S. M. Silva, C. R. Braga, M. V. Fook, C. M. Raposo, L. H. Carvalho, and E. L. Canedo, “Application of infrared spectroscopy to analysis of chitosan/clay nanocomposites,” in Infrared Spectroscopy—Materials Science, Engineering and Technology, T. Theophanides, Ed., pp. 43–62, InTech, 2012. View at: Google Scholar
  36. M. Venkatesham, D. Ayodhya, A. Madhusudhan, N. Veera Babu, and G. Veerabhadram, “A novel green one-step synthesis of silver nanoparticles using chitosan: catalytic activity and antimicrobial studies,” Applied Nanoscience, vol. 4, pp. 113–119, 2012. View at: Google Scholar
  37. W. I. Abdel-Fattah, A. S. M. Sallam, N. Attawa, E. Salama, A. M. Maghraby, and G. W. Ali, “Functionality, antibacterial efficiency and biocompatibility of nanosilver/chitosan/silk/phosphate scaffolds 1. Synthesis and optimization of nanosilver/chitosan matrices through gamma rays irradiation and their antibacterial activity,” Materials Research Express, vol. 1, no. 3, Article ID 035024, 2014. View at: Publisher Site | Google Scholar
  38. K. Vimala, Y. M. Mohan, K. S. Sivudu et al., “Fabrication of porous chitosan films impregnated with silver nanoparticles: a facile approach for superior antibacterial application,” Colloids and Surfaces B: Biointerfaces, vol. 76, no. 1, pp. 248–258, 2010. View at: Publisher Site | Google Scholar
  39. J. R. Franca, M. P. de Luca, T. G. Ribeiro et al., “Propolis—based chitosan varnish: drug delivery, controlled release and antimicrobial activity against oral pathogen bacteria,” BMC Complementary and Alternative Medicine, vol. 14, article 478, 2014. View at: Publisher Site | Google Scholar
  40. D. K. Boanić, L. V. Trandafilović, A. S. Luyt, and V. Djoković, “‘Green’ synthesis and optical properties of silver-chitosan complexes and nanocomposites,” Reactive and Functional Polymers, vol. 70, no. 11, pp. 869–873, 2010. View at: Publisher Site | Google Scholar
  41. D. Zvezdova, “Synthesis and characterization of chitosan from marine sources in Black Sea,” in Proceedings of the Annual Conference “Angel Kanchev”, vol. 49 of Series 9.1—Chemical Technologies, pp. 65–69, The University of Rousse, 2010. View at: Google Scholar
  42. M. V. Butnariu and C. V. Giuchici, “The use of some nanoemulsions based on aqueous propolis and lycopene extract in the skin's protective mechanisms against UVA radiation,” Journal of Nanobiotechnology, vol. 9, article 3, 2011. View at: Publisher Site | Google Scholar
  43. S. Kumar, P. K. Dutta, and J. Koh, “A physico-chemical and biological study of novel chitosan-chloroquinoline derivative for biomedical applications,” International Journal of Biological Macromolecules, vol. 49, no. 3, pp. 356–361, 2011. View at: Publisher Site | Google Scholar
  44. S. Kumar and J. Koh, “Physiochemical, optical and biological activity of chitosan-chromone derivative for biomedical applications,” International Journal of Molecular Sciences, vol. 13, no. 5, pp. 6102–6116, 2012. View at: Publisher Site | Google Scholar
  45. V. Vilas, D. Philip, and J. Mathew, “Catalytically and biologically active silver nanoparticles synthesized using essential oil,” Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy, vol. 132, pp. 743–750, 2014. View at: Publisher Site | Google Scholar
  46. X.-M. Zhao, Z.-Q. Wang, S.-H. Shu et al., “Ethanol and methanol can improve huperzine A production from endophytic colletotrichum gloeosporioides ES026,” PLoS ONE, vol. 8, no. 4, Article ID e61777, 2013. View at: Publisher Site | Google Scholar
  47. B. Ren, B. Xia, W. Li, J. Wu, and H. Zhang, “Two novel phenolic compounds from Stenoloma chusanum and their antifungal activity,” Chemistry of Natural Compounds, vol. 45, no. 2, pp. 182–186, 2009. View at: Publisher Site | Google Scholar
  48. M. P. Kabra, S. S. Bhandari, A. Sharma, and M. K. Vaishnav, “A review on gentisic acid,” Internationale Pharmaceutica Sciencia, vol. 3, pp. 29–36, 2013. View at: Google Scholar
  49. İ. Gülçin, E. Bursal, M. H. Şehitoğlu, M. Bilsel, and A. C. Gören, “Polyphenol contents and antioxidant activity of lyophilized aqueous extract of propolis from Erzurum, Turkey,” Food and Chemical Toxicology, vol. 48, no. 8-9, pp. 2227–2238, 2010. View at: Publisher Site | Google Scholar

Copyright © 2015 Petruta Mihaela Matei et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

More related articles

2253 Views | 1345 Downloads | 17 Citations
 PDF Download Citation Citation
 Download other formatsMore
 Order printed copiesOrder

Related articles

We are committed to sharing findings related to COVID-19 as quickly as possible. We will be providing unlimited waivers of publication charges for accepted research articles as well as case reports and case series related to COVID-19. Review articles are excluded from this waiver policy. Sign up here as a reviewer to help fast-track new submissions.