Advertisement

Journal of Cluster Science

, Volume 28, Issue 1, pp 413–436 | Cite as

Do Chenopodium ambrosioides-Synthesized Silver Nanoparticles Impact Oryzias melastigma Predation Against Aedes albopictus Larvae?

  • Jayapal Subramaniam
  • Kadarkarai Murugan
  • Arulsamy Jebanesan
  • Philips Pontheckan
  • Devakumar Dinesh
  • Marcello Nicoletti
  • Hui Wei
  • Akon Higuchi
  • Suresh Kumar
  • Angelo Canale
  • Giovanni Benelli
Original Paper

Abstract

The green synthesis of nanopesticides has been recently proposed to improve the efficacy of mosquito control programs. However, limited efforts shed light on the impact of sub-lethal doses of nanopesticides on behavioral traits of mosquito biocontrol agents. We described the synthesis of silver nanoparticles (AgNP) at room temperature using the aqueous extract of Chenopodium ambrosioides, and their high toxicity against the invasive mosquito Aedes albopictus. LC50 calculated on young instars ranged from 13 ppm (first instar larvae) to 19 ppm (pupae). LC50 calculated on adults was 14 ppm. The chemical composition of the C. ambrosioides extract was characterized by GC–MS analysis. The production of AgNP was confirmed by the surface Plasmon resonance band illustrated in UV–Vis, FTIR spectroscopy, EDX, XRD, TEM, and Zeta Potential analyses. In the field, a single treatment of AgNP (10 × LC50) led to complete elimination of larval populations within 72 h. Sub-lethal doses of the reducing extract and AgNP magnify predation rates of Oryzias melastigma fishes against A. albopictus larvae. Overall, this study highlights the concrete potential of C. ambrosioides-synthesized AgNP to develop effective and cheap tools to control young instars and adults of the invasive mosquito A. albopictus.

Keywords

Biological control Biosafety Larvivorous fishes Nanobiotechnology Nanocrystals 

Introduction

Mosquitoes represent a major public health problem, since they act as vectors of serious diseases, including malaria, yellow fever, West Nile virus, filariasis, Japanese encephalitis, dengue and chikungunya [9, 15, 74, 75]. Dengue is an emerging disease, currently considered the most important arbovirus in the world. Aedes mosquitoes mainly vector it. Dengue slyly arrived in the Western Hemisphere over decades, and then its incidence has grown dramatically from the 1990s. The actual numbers of dengue cases are underreported and many cases are misclassified [14, 47, 134]. WHO estimates that dengue infects about 400 million people annually in the part of tropical and subtropical regions [18, 20, 134]. Very recently, mosquito from the Aedes genus also vectored Zika virus to people [135], leading to outbreaks in the Americas, and the Pacific area. Zika symptoms are similar to other arbovirus infections such as dengue, and include fever, skin rashes, conjunctivitis, muscle and joint pain, malaise, and headache. These symptoms normally last for 2–7 days and can be followed by neurological complications and malformations in neonates [37, 135]. Although there are several potential dengue vectors, the field isolation of viruses and epidemiological evidence show that Aedes aegypti and A. albopictus are the main vectors [14]. A. albopictus, also known as the Asian tiger mosquito, originates in Asia and also serves as a vector of chikungunya and many other arboviruses [57, 59, 129].

The use of chemicals insecticides in routine mosquito control operations led to the development of resistance in the targeted vector species [55, 91], as well as to detrimental effects on non-target organisms, with special reference to biological control agents such as larvivorous fishes and other important aquatic predators of Culicidae [26, 96, 105, 112]. Therefore, plant-based insecticides may serve as suitable alternative to synthetic molecules as they are environmentally safe, biodegradable, and are easily available in all parts of the world [4, 10, 16, 53, 124]. In addition, it is worthy to note that the toxicity of botanical-based biopesticides such as plant extracts and essential oils is usually exerted by multiple mechanisms of action, lowering the chances of resistance development in targeted arthropods [99].

Recently, silver nanoparticles (AgNP) gained a focus of intensive research owing to their wide range of applications in areas such as catalysis, optics, antimicrobials, pesticides, biomedical and biomaterial production [7, 39, 64, 126]. The biological synthesis of metal nanoparticles is a research area currently considered more eco-friendly and cost-effective, if compared to other chemical and physical methods [3, 11]. Nano-technology is envisaged to be the next frontier in the ongoing development of cancer therapy [22, 35] as researchers in the biomedical and material engineering fields are working together to discover the possibility of using nano-materials as novel tools for medical sciences. In particular, a number of approaches are available for the synthesis of silver nanoparticles, such as thermal decomposition [93], electrochemical [121], microwave-assisted process [120] and green chemistry [8, 12].

Phytochemicals have a major role in current mosquito control research [9, 10]. Plant extracts have been used as reducing and capping agent for the synthesis of nanoparticles. Indeed, the latter is advantageous over photochemical reduction, heat evaporation, electrochemical reduction, and chemical reduction methods [11]. Because of such a wide range of applications, numerous methods concerning the fabrication of AgNP, as well as various silver-based compounds containing ionic silver (Ag+) or metallic silver (Ag0) have been developed. The synthetic methods used for the preparation of AgNP rely to some toxic chemical used as reducing agents such as NaBH4, citrate or ascorbate. On the other hand, in plant-mediated reducing processes leading to the production of nanoparticles, no chemical reagent or surfactant template is required, which consequently enables the bioprocess with the advantage of being eco-friendly [13, 87, 88, 89, 90].

Another important challenge for mosquito control is the successful implementation of biological control programs. Indeed, the natural enemies feeding on mosquito larvae and pupae in aquatic environments play an important role in reducing Culicidae populations (e.g. [65, 131, 136]. Larvivorous fishes are being successfully exploited for control of mosquito vectors aquatic stages in European, Asian, African and Arabian countries [21, 65]. Moreover, the larvivorous fishes provide dual benefits by reducing the mosquito populations and indirectly augmenting the aqua cultural economics [26, 73, 114, 132]; see [17] for a recent review).

Oryzias melastigma [72] (Beloniformes: Adrianichthyidae) [60] is a tiny cyprinodontid fish. It is a carnivorous, surface feeder found in both lentic and lotic waters. This semitransparent and hardy fish can tolerate a wide range of salinity [68], temperature, and many other adverse water qualities. Popularly known as rice fish orminnow [108] or Indian Medaka, or Bechi, it is a sexually dimorphic species [69]. It is found in limited areas of West Bengal, Tamil Nadu, Kerala, Orissa [60, 70] in India and also some riverine areas of Bangladesh.

Chenopodium ambrosioides Linn. (Chenopodiaceae) is widely distributed throughout India. Leaves are useful in the cure of influenza, pneumonia, typhoid and also as vermicide [32, 77]. Chenopodium oil is a mixture ascaridole (55.3 8%), p-cymene (16.2%), alpha-terpinene (9.7%), isoascaridole (4.3%) and limonene (3.8%) [24]. By contrast, little is known about the chemical composition of the polar extracts from C. ambrosioides. Nowadays, this species can be occasionally found also in pathways and near home gardens. It has diverse pharmacological applications in the treatment of influenza, cold or gastrointestinal and respiratory ailments, as well as vomiting, antihelmintic, healing of skin ulceration caused by Leishmania species, anti-inflammatory and antitumor properties [23, 38, 62, 92].

Even if the green synthesis of nanopesticides has been recently proposed to improve the efficacy of mosquito control programs [11], only limited efforts shed light on the potential impact of sub-lethal doses of nanopesticides on behavioral traits of mosquito biological control agents [82, 83, 84, 87, 88, 89, 127]. Here, we described the synthesis of AgNP at room temperature using the extract of C. ambrosioides as a reducing and capping/stabilizing agent, and their high toxicity against larvae, pupae and adults of the invasive mosquito Ae. albopictus. The chemical composition of the C. ambrosioides extract was characterized by GC–MS analysis. The effective production of AgNP was confirmed UV–Vis and FTIR spectroscopy, EDX, XRD, TEM, and Zeta Potential analysis. In the final experiments, the impact of sub-lethal doses of the reducing extract and AgNPs on predation rates of O. melastigma fishes against A. albopictus larvae was evaluated.

Materials and Methods

Collection of Plant Materials

C. ambrosioides plants used in this study were collected from the villages of The Nilgris, (Western Ghats of South India) Tamil Nadu, India. The plants were authenticated at Botanical Survey of India. Voucher specimens were deposited at Zoology Department, Bharathiar University, Coimbatore, India (Voucher ID n. CHENO-03).

Mosquito Rearing

Eggs of Ae. albopictus were provided by the National Centre for Disease Control (NCDC) field station of Mettuppalayam (Tamil Nadu, India). Eggs were transferred to laboratory conditions [27 ± 2 °C, 75–85% R.H., 14:10 (L:D) photoperiod] and placed in 18 × 13 × 4 cm plastic containers containing 500 mL of tap water, to await larval hatching [41, 128]. Larvae were reared in these containers and fed daily with a mixture of crushed dog biscuits (Pedigree, USA) and hydrolyzed yeast (Sigma-Aldrich, Germany) at a 3:1 ratio (w:w). Water was renewed every 2 days. The breeding medium was checked daily and dead individuals were removed. Breeding containers were kept closed with muslin cloth to prevent contamination by foreign mosquitoes. Pupae were collected daily from culture containers and transferred to glass beakers containing 500 mL of water. Each glass beaker contained about 50 mosquito pupae and was placed in a mosquito-rearing cage (90 × 90 × 90 cm, plastic frames with chiffon walls) until adult emergence. Mosquito adults were continuously provided with 10% (w:v) glucose solution on cotton wicks. The cotton was always kept moist with the solution and changed daily. Five days after emergence, females were supplied with a blood meal which was furnished by means of professional heating blood (lamb blood), at a fixed temperature of 38 °C and enclosed in a membrane of cow gut. After 30 min, the blood meal was removed and a fresh one was introduced [86, 95].

C. ambrosioides-Mediated Synthesis of Silver Nanoparticles

The C. ambrosioides aqueous leaf extract was prepared by adding 10 g of washed and finely cut leaves in a 300-mL Erlenmeyer flask filled with 100 mL of sterilized double distilled water, then boiling the mixture for 5 min before decanting it. The extract was filtered using Whatman filter paper n. 1, was stored at −4 °C and tested within 5 days. The filtrate was treated with aqueous 1 mM AgNO3 (Precision Scientific Co., Coimbatore, India) solution in an Erlenmeyer flask and incubated at room temperature [82]. A dark brown solution indicated the formation of AgNP, as aqueous silver ions were reduced by the C. ambrosioides extract generating stable AgNP in water.

GC–MS Analysis

GC–MS analysis of the plant ethanolic extract was performed using a Perkin Elmer GC Claurus 500 system and Gas Chromatograph interfaced to a Mass Spectrometer (GC/MS) equipped with a Elite-1 fused silica capillary column (30 m × 0.25 mm ID. × 1 μMdf, composed of 100% Dimethyl poly siloxane). The plant ethanolic extract was prepared following the method by [98]. For GC–MS detection, an electron ionization system with ionization energy of 70 eV was used. Helium gas (99.999%) was used as the carrier gas at a constant flow rate of 1 ml/min. and an injection volume of 2 μl was employed (split ratio of 10:1). The injector temperature was 250 °C. The ion-source temperature was 280 °C. The oven temperature was programmed from 110 °C (isothermal for 2 min.), with an increase of 10 °C/min, to 200 °C, then 5 °C/min to 280 °C, ending with a 9 min. isothermal at 280 °C. Mass spectra were taken at 70 eV; a scan interval of 0.5 s and fragments from 45 to 450 Da. Total GC running time was 36 min. The relative percentage amount of each component was calculated by comparing its average peak area to the total areas. Software adopted to handle mass spectra and chromatograms was a TurboMass Ver 5.2.0 [130].

Characterization of Silver Nanoparticles

C. ambrosioides-synthesized AgNP were characterized by UV–Vis spectrophotometry, FTIR spectroscopy, TEM, EDX, XRD and Zeta potential analysis [85, 107]. In UV–Vis absorbance spectrophotometry, the bio-reduction of AgNO3 in the aqueous medium was monitored by periodic sampling of aliquots (2 mL), measuring the UV–Vis spectrum in 10 mm quartz cuvette with a systronics. We used a UV–Vis spectrophotometer (Hewlett-Packard diode array spectrophotometer, model HP-8452, resolution: 1 nm) operating at 500 and 680 nm with a scanning speed of 1856 nm/min. OD values were recorded until 3 days after biosynthesis at regular intervals. Samples were centrifuged at 42,000 rpm for 10 min; pellets were dried; and the nano-powder obtained was used for further analyses. TEM was performed using a JEOL model 1200 EX instrument operating at an accelerating voltage of 120 kV. Samples were prepared by placing tiny drops of AgNP solutions on carbon-coated TEM grids. The film on the TEM grid was allowed to dry for 5 min under laboratory conditions. XRD analysis of drop-coated films on glass substrates from the AOT-capped AgNP was carried out on a Phillips PW1830 instrument operating at 40 kV and a current of 30 mA with Cu Kα radiation. EDX analyzed the presence of metals in the sample (JEOL-MODEL 6390); the XRD patterns were phase matched using match software version 1.10c Inc. Standard values are obtained from the International Centre for Diffraction Data ICDD. Hkl indices and the mean size of AgNP were calculated using the Debye–Scherer equation by determining the width of (111) and similar Bragg’s reflection parameters [83]. For FTIR measurements, samples were prepared as described for XRD analysis, and measured using a Shimadzu 8400 s with spectral range of 4000–400 cm−1 and resolution of 4 cm−1. FTIR spectra of leaf extracts sampled before and after the biosynthesis of AgNP were compared to examine possible functional groups involved in AgNP formation [41, 126].

Larvicidal Activity Against A. albopictus

Following the methods reported by [128], 25 mosquito larvae (I, II, III or IV instar) or pupae were placed for 24 h in a 500-mL glass beaker filled with dechlorinated water plus C. ambrosioides leaf ethanolic extract (80, 160, 240, 360 and 400 ppm) or C. ambrosioides-synthesized AgNP (10, 20, 30, 40 and 50 ppm). Larval food (0.5 mg) was provided for each tested concentration. Each concentration was replicated 5 times against all instars. In the control, 25 larvae or pupae were transferred to 250 mL of dechlorinated water. No mortality was observed in the control. Percentage mortality was calculated as follows:

$${\text{Percentage mortality}} = ({\text{number of dead individuals}}/{\text{number of treated individuals}})*100.$$

Larvicidal Activity in the Field

C. ambrosioides ethanolic extract and C. ambrosioides-synthesized AgNP were applied in six external water storage reservoirs in each of two field sites at the National Institute of Communicable Disease Centre (Coimbatore, India).Treatments were carried out using a knapsack sprayer (Private Limited 2008, Ignition Products, India) [82]. Pre-treatment Aedes larval density was monitored. Post-treatment observations were conducted after 24, 48 and 72 h, using a larval dipper. Toxicity was assessed against third- and fourth instars larvae. Six trials were conducted for each test site with similar weather conditions (28 ± 2 °C; 80% R.H.). The required quantity of mosquitocide was calculated on the basis of the total surface area and volume (i.e. 0.25 m3 and 250 L for all sites). Then, the required concentration was prepared using 10 × LC50 values [80, 128]. Percentage reduction of the larval density was calculated using the formula:

$${\text{Percentage reduction}} = ({\text{C}} - {\text{T)}}/{\text{C}}\, \times \,100$$
where C is the total number of mosquitoes in the control, and T is the total number of mosquitoes in the treatment [126].

Adulticidal Activity

Adulticidal experiments were performed following the methods reported by the [126, 127, 133]. C. ambrosioides ethanolic leaf extract was tested at 60, 120, 180, 240 and 300 ppm. AgNP were tested at 6, 12, 18, 24 and 30 ppm formulated in 5 mL of aqueous solution. C. ambrosioides aqueous extract and AgNP were applied on Whatman n. 1 filter paper (size 12 × 15 cm) lining a glass holding tube (diameter 30 mm; length 60 mm). In control treatments, filter papers were treated with either the same volume of distilled water plus ethanol or AgNO3 (1 mM) in aqueous solution. In each test, 20 A. albopictus females were gently transferred into another glass holding tube. The mosquitoes were allowed to acclimatize in the tube for 1 h and then exposed to a test tube lined with treated or control paper for 1 h. At the end of exposure period, the mosquitoes were transferred back to the original holding tube, kept for a 24 h recovery period and then mortality was recorded. A pad of cotton soaked with 10% (w:v) glucose solution was placed on the mesh screen at the top of the holding tube [126].

Oryzias melastigma Predation on A. albopictus Larvae

O. melastigma fishes were collected from Tamil Nadu Fisheries Department, Mettur Dam, Salem, and maintained in cement tanks (120 cm diameter and 60 cm depth) containing field collected water at 27 ± 3 °C and external RH 85%. For the assays, the predatory fishes were released in separate transparent containers (14 × 10 cm) containing clean water. The predatory efficiency of O. melastigma was assessed against II and III instar larvae of A. albopictus. In each trial, 200 mosquito larvae were introduced, with 1 adult O. melastigma, in plastic cups (2 L) containing dechlorinated water. For each tested mosquito instar, five replicates were conducted. Control was 2 L of dechlorinated water plus 200 larvae, without O. melastigma. All experimental cups checked after 24 h and the number of dead/preys consumed by predator was recorded. After each checking, the predated mosquito larvae were replaced with new ones. Similarly, five replicates were made for each prey density with predators or without predators (control), before and after the treatment of the leaf extract or AgNP. Using the same fish, the rate of predation was observed for five consecutive days. The prey density was set to same value after every 24 h. The fish predatory efficiency was calculated using the following formula:
$${\text{Predatory efficiency}} = [({\text{number of consumed mosquitoes}}/{\text{number of predators}})/{\text{total number of mosquitoes}}]\, \times \,100$$

Oryzias melastigma Predation on A. albopictus Larvae Post-Treatment with Ag Nanoparticles.

Here, 200 mosquito larvae were introduced, with 1 adult O. melastigma; in plastic cups (2 L) containing dechlorinated water plus 1/3 of the LC50 calculated against III and IV instar larvae of A. albopictus [82, 83]. For each tested mosquito instar, five replicates were conducted. Control was 2 L of AgNP-contaminated water plus 200 larvae, without predator fish (O. melastigma). All experimental cups checked after 24 h and the number of preys consumed by O. melastigma was recorded. After each checking, the predated mosquito larvae were replaced with new ones. Similarly, five replicates were made for each prey density with predators or without predators (control), before and after the treatment of the leaf extract or AgNP. Using the same predator individual, the rate of predation was observed for five consecutive days. The prey density is being set to same value after every 24 h. The fish predatory efficiency was calculated using the above-mentioned formula.

Data Analysis

Mosquito mortality data from laboratory assays were analyzed by probit analysis, calculating LC50 and LC90 following the method by [44]. Mosquito larval density data from field assays were analyzed using a two-way ANOVA with two factors (i.e. the mosquitocidal treatment and the elapsed time from treatment). In all analyses, a probability level of P < 0.05 was used for the significance of differences among values.

O. melastigma predation data were analyzed using a weighted general linear model with two fixed factors:\(y = X\beta \, + \,\varepsilon\), where y is the vector of the observations (the number of consumed preys), X is the incidence matrix, ß is the vector of fixed effects (treatment and targeted mosquito instar), and ε is the vector of the random residual effect. A probability level of P = 0.05 was used for the significance of differences between values.

Results and Discussion

Chemical Composition of C. ambrosioides Leaf Extract

The interpretation on mass spectrum GC–MS was conducted using the database of National Institute Standard and Technology (NIST) having more than 62,000 patterns. The spectrum of the unknown component was compared with the spectrum of the known components stored in the NIST library. The name, molecular weight and structure of the components of the test materials were ascertained. A total of 15 components were identified (Fig. 1; Table 1), among them tetradecanoic acid (C14H28O2) was the major component available at a RT of 12.29 min and with a peak area of 22.43%, the second major component was 3-methoxysalicylic acid (C8H8O4) with a RT of 4.88 min and 18.38% peak area. Further components were identified by GC–MS spectral comparison with the database NIST, including bicyclo[4.1.0] heptan-3-ol, 4,7,7-trimethyl-(1α,3α,4α,6α)-(C10H18O) 4.10 min RT, peak area 13.28%, and 5-isopropenyl-2-methyl-7-oxabicyclo[4.1.0]heptan-2-ol (C10H16O2) 6.53 min RT, peak area 9.04% (Table 1). It has been reported that tetracyclic triterpenoids showed activity on entomopathogenic nematodes [6], and tetradecanoic acid acted as a good larvicide and repellent against the dengue and yellow fever vector A. aegypti, while squalene has a variety of health-promoting functions, including tumor-suppressing [1, 94, 104, 119], antibacterial/antifungal [118], and cholesterol-lowering [76] effects. Also, squalene has recently attracted attention as a feasible source of biofuels [109]. The insecticidal properties of essential oils containing these compounds against several pest species, including mosquito vectors, have been reported by [31, 116, 138].
Fig. 1

GC–MS analysis of the Chenopodium ambrosioides leaf ethanolic extract

Table 1

Components identified in ethanolic extract of Chenopodium ambrosioides

N.

RT (min)

Compounds

Formula

MW

Peak Area (%)

1

4.10

Bicyclo[4.1.0] heptan-3-ol, 4,7,7-trimethyl-(1α,3α,4α,6α)-

C10H18O

154

13.28

2

4.88

3-Methoxysalicylic acid

C8H8O4

168

18.38

3

5.29

7-Oxabicyclo[4.1.0] heptane, 1-methyl-4-(2-methyloxiranyl)-(α-Limonene diepoxide)

C10H16O2

168

5.25

4

6.53

5-Isopropenyl-2-methyl-7-oxabicyclo[4.1.0]heptan-2-ol

C10H16O2

168

9.04

5

9.13

Cyclohexanol, 2-methyl-5-(1-methylethenyl)-

C10H18O

154

0.93

6

10.53

3,7,11,15-Tetramethyl-2-hexadecen-1-ol

C20H40O

296

4.21

7

10.98

E-7-Tetradecenol

C14H28O

212

2.51

8

12.29

Tetradecanoic acid

C14H28O2

228

22.43

9

13.32

1-Hexadecanol

C16H34O

242

3.92

10

13.58

Phytol

C20H40O

296

3.00

11

14.27

9,12-Octadecadienoic acid (Z,Z)-

C18H32O2

280

8.95

12

17.07

Octadecanoic acid, ethyl ester

C20H40O2

312

1.30

13

19.14

Didodecyl phthalate

C32H54O4

502

1.13

14

22.57

Squalene

C30H50

410

1.01

15

29.35

cis-Z-α-Bisabolene epoxide

C15H24O

220

4.68

RT Retention time

MW Molecular weight

Characterization of Silver Nanoparticles

In our experiments, UV–Vis spectrum showed a maximum absorbance peak at 421 nm which increased over time during the incubation of silver nitrate with the C. ambrosioides extract (Fig. 2). When the AgNO3 solution was added to the C. ambrosioides leaf extract, the color changed from light to dark brown, indicating the reduction from Ag+ to Ag0 (Fig. 2a). The formation of AgNP was confirmed through the presence of an absorption peak at 421 nm (Fig. 2). Our UV–Vis results are in agreement with previous research [66, 82, 107, 115, 127, 137]. The main peak detected here indicated a surface Plasmon resonance (SPR), which has been recorded for different metal nanoparticles ranging from 2 to 100 nm in size [56, 106].
Fig. 2

a Chromatic variations of the aqueous leaf extract of Chenopodium ambrosioides before and after the process of reduction of Ag+ to Ag nanoparticles. b UV–Visualization of the absorption spectrum of Ag nanoparticles synthesized using C. ambrosioides after 120 min from the reaction

TEM observations showed different shapes of C. ambrosioides-synthesized AgNP, including spherical, round and hexagonal ones, with mean size ranging from 25 to 50 nm (Fig. 3). Similarly, the morphological features of green synthesized silver, gold and metal nanoparticles fabricated using extracts from several terrestrial and marine plants, lead to mean nanoparticle sizes ranging from 15 to 70 nm (e.g. [82, 83, 103, 125, 127] Furthermore, C. ambrosioides-synthesized AgNP did not show direct contact within aggregates, allowing us to argue that their stabilization occurred through capping agents.
Fig. 3

Transmission electron microscopy (TEM) of green-synthesized silver nanoparticles obtained by reduction of AgNO3 with the leaf extract of Chenopodium ambrosioides

The EDX spectrum recorded from C. ambrosioides synthesized AgNP revealed a distinct signal and high atomic percent values for Ag (Fig. 4). EDX analysis confirmed the presence of elemental Ag. The presence of oxygen (O) and silver (Ag) indicates that the extracellular organic compounds were adsorbed on the surface of AgNP (Fig. 4). The present finding corroborates previous reports on AgNP biosynthesis using botanical and microbial products [3, 43, 50, 51, 52, 67]. XRD patterns showed intense peaks corresponding to the (111), (200), (220), (311) and (222) sets of lattice planes (Fig. 5). The XRD patterns showed that the AgNP formed by the reduction of AgNO3 using C. ambrosioides leaf extract were crystalline in nature (Fig. 5). The XRD pattern observed in this study was consistent with previous reports [5, 48]. For instance, [111] reported diffraction peaks at 44.50°, 52.20°, and 76.7° = 2θ, which correspond to the (111), (200), and (220) facets of the face-centered cubic crystal structure.
Fig. 4

Energy dispersive X-ray (EDX) profile of silver nanoparticles synthesized using the leaf extract of Chenopodium ambrosioides

Fig. 5

X-ray diffraction pattern of silver nanoparticles synthesized using the leaf extract of Chenopodium ambrosioides

FTIR spectroscopy was carried out to identify the possible biomolecules in the C. ambrosioides extract, which may be responsible for synthesis and stabilization of AgNP (Fig. 6). FTIR spectrum of AgNP prepared using the C. ambrosioides leaf extract showed peaks at 3431.36, 2362.80, 2063.83, 1633.71, 1514.12, 1456.26, and 418.55 cm−1 (Fig. 6). The peak located at about 2,362.80 cm−1 can be attributed to the N–H stretching vibrations or the C=O stretching vibrations. The sharp absorption peak at 1633.71 cm−1 may be assigned to C=O stretching vibration in carbonyl compounds which may be characterized by the presence of high content of terpenoids and flavonoids. A broad intense band at 3,431.36 cm−1 in both leaf extract and AgNP spectra can be linked to the N–H stretching frequency arising from the peptide linkages present in the proteins of the extract [71, 78]. Therefore, it may be inferred that these biomolecules are responsible for capping and efficient stabilization of synthesized nanoparticles. Thus, the analysis of FTIR spectrum from the green fabricated AgNP showed the presence of different functional groups from alkane, methylene, alkene, amine, and carboxylic acid, previously reported as reducing agents in the nano-biosynthesis [33]. Polyphenols have been also reported as potential reducing agent in the biosynthesis of AgNP [79, 100]. The adsorption on the surface of metal nanoparticles is a characteristic of flavanones and terpenoids, since they easily interacted through carbonyl groups in the lack of other strong ligating agents in sufficient concentration [113].
Fig. 6

Fourier transform infrared spectroscopy (FTIR) of silver nanoparticles synthesized using the leaf extract of Chenopodium ambrosioides

Particle size and size distribution are the most important characteristics of nanoparticle systems. In our analysis, zeta potential of AgNP was −18.5 mV (Fig. 7). Similarly, [42] noted that C. album-synthesized silver and gold nanoparticles were stable under a wide pH range due to their high zeta potential. In agreement with TEM results, size analysis showed a distribution of particle diameters ranging from 10 to 90 nm with an average particle size of 25 nm. The particle sizes determined the in vivo distribution, biological fate, toxicity and the targeting ability of nanoparticle systems [13]. [40] have reported that 100 nm nanoparticles had a 2.5 fold greater uptake than1 µm microparticles, and sixfold greater uptake than 10 µm microparticles on Caco-2 cell line.
Fig. 7

Zeta potential analysis of silver nanoparticles synthesized using the leaf extract of Chenopodium ambrosioides

Toxicity on Aedes albopictus

In laboratory conditions, the C. ambrosioides ethanolic leaf extract showed larvicidal and pupicidal toxicity against A. albopoictus, with LC50 values ranging from 124.55 ppm (I instar larva) to 237.06 ppm (pupa), respectively (Table 2). A number of plant extracts has been reported as effective against larvae and pupae of mosquito vectors [10, 61, 122, 123]. More recently, the green biosynthesis of mosquitocidal nanoparticles is advantageous over chemical and physical methods, since it is cheap, single-step, and does not require high pressure, energy, temperature, and the use of highly toxic chemicals [87, 102, 103]. In this study C. ambrosioides-synthesized AgNP were toxic against A. albopictus larvae and pupae, with LC50 values ranging from 13.37 ppm (I instar) to 19.77 ppm (pupa) (Table 3). In agreement with our data, [126] showed that Mimusops elengi leaf aqueous extract was moderately effective against malarial vector, Anopheles stephensi and arbovirus vector A. albopictus while the LC50 of AgNP fabricated using this plant and tested on A. stephensi ranged from 12.53 (I instar larvae) to 23.55 ppm (pupae), and LC50 against A. albopictus ranged from 11.72 ppm (I) to 21.46 ppm (pupae). Low doses of AgNP biosynthesized using Euphorbia hirta leaf extract have been reported as highly toxic against A. stephensi, withLC50 values ranging from 10.14 ppm (I instar larvae) to 34.52 ppm (pupae) [101]. Another good example is the larvicidal activity of Leucas aspera-synthesized AgNP, with LC50 ranging from 13.06 to 25.54 ppm for A. aegypti, and from 12.45 to 22.26 ppm for A. stephensi [117]. Nelumbo nucifera-synthesized AgNP were toxic to the larvae of A. subpictus (LC50 = 0.69 ppm) and C. quinquefasciatus (LC50 = 1.10 ppm; LC90 = 3.59 ppm), respectively [110]. In the field, the application of C. ambrosioides aqueous extract and C. ambrosioides-synthesized AgNP (10 × LC50) in water storage reservoirs led to the complete elimination of larval populations of A. albopictus after 72 h (Table 4). [104] reported that the stable neem fractions were as effective as mosquito larvicides in the field. Plant based insecticides have been evaluated successfully in different habitats of mosquito vectors, tested species include Clerodendron inerme, Acanthus ilicifolius [63], M. elengi and green-synthesized AgNP [125], Phyllanthus niruri and green-synthesized AgNP [128]. Further research aimed to clarify the exact mechanism(s) of action of AgNP against mosquito young instars is ongoing [13].
Table 2

Acute toxicity of Chenopodium ambrosioides leaf extract on young instars of the dengue and Zika virus vector, Aedes albopictus

Target

LC50 (LC90)

95% Confidence limit LC50 (LC90)

Regression equation

χ 2

(d.f. = 4)

Lower

Upper

Larva I

124.55 (335.30)

93.76 (305.61)

148.35 (376.65)

y = 0.757 + 0.006x

1.10 n.s

Larva II

143.24 (398.83)

108.83 (359.33)

169.68 (457.24)

y = 0.718 + 0.005x

1.11 n.s

Larva III

172.10 (453.72)

138.95 (404.76)

198.68 (529.22)

y = 0.783 + 0.005x

0.44 n.s

Larva IV

199.55 (488.86)

169.34 (434.44)

225.68 (573.88)

y = 0.884 + 0.004x

1.55 n.s

Pupa

237.06 (554.90)

207.60 (485.72)

266.14 (668.54)

y = 0.956 + 0.004x

1.72 n.s

No mortality was observed in the control

LC 50 lethal concentration that kills 50% of the exposed organisms

LC 90 lethal concentration that kills 90% of the exposed organisms

χ 2 Chi square value

d.f. degrees of freedom

n.s. not significant (α = 0.05)

Table 3

Acute toxicity of Chenopodium ambrosioides-synthesized silver nanoparticles on young instars of the dengue and Zika virus vector Aedes albopictus

Target

LC50 (LC90)

95% Confidence limit LC50 (LC90)

Regression equation

χ 2

(d.f. = 4)

Lower

Upper

Larva I

13.37 (37.05)

9.53 (33.81)

16.29 (41.49)

y = 0.724 + 0.054x

4.89 n.s

Larva II

14.44 (39.42)

10.59 (35.96)

17.38 (44.20)

y = 0.741 + 0.051x

3.78 n.s

Larva III

15.41 (46.34)

10.78 (41.80)

18.85 (53.02)

Y = 0.639 + 0.041x

0.62 n.s

Larva IV

17.39 (51.19)

12.70 (45.84)

20.90 (59.30)

y = 0.660 + 0.038x

1.22 n.s

Pupa

19.77 (60.66)

14.48 (53.12)

23.69 (73.18)

y = 0.620 + 0.031x

0.49 n.s

No mortality was observed in the control

LC 50 lethal concentration that kills 50% of the exposed organisms

LC 90 lethal concentration that kills 90% of the exposed organisms

χ 2 Chi square value

d.f. degrees of freedom

n.s. not significant (α = 0.05)

Table 4

Field treatment of storage water tanks with the leaf extract of Chenopodium ambrosioides and green-synthesized silver nanoparticles against the dengue vector Aedes albopictus

Target

Chenopodium ambrosioides leaf extract (10 × LD50)

Green synthasized silver nanoparticles (10 × LD50)

A. albopictus

Before

treatment

24 h

48 h

72 h

Before

treatment

24 h

48 h

72 h

Larval density

1271 ± 12.38b

786 ± 38.15c

261 ± 10.44d

0.00 ± 0.0f

1525 ± 29.54a

791 ± 10.45c

226 ± 8.47e

0.00 ± 0.0f

Within the row, different letters indicate significant differences (ANOVA, Tukey’s HSD test, P = 0.05)

In adulticidal experiments, the C. ambrosioides leaf extract and green-synthesized AgNP were toxic to A. albopictus (Table 5). LC50 values were 154.99 ppm (C. ambrosioides extract) and 14.29 ppm (AgNP). At the highest concentration tested, the adults of both species remained still for a short time period (i.e. 1–3 min) following application, showed fast wagging movements and then died. The adulticidal efficacy of a number of plant borne extracts and essential oils against adult mosquitoes of public health importance has been reported by several recent studies (e.g. [2, 97, 123, 125, 126]. For example, Subramaniam al. [125] reported that the adulticidal activity of methanol extracts of seaweeds D. dichotoma, P. pavonica and V. pachynema on the costal malarial vector Anopheles sundaicus, LC50 values were 147.18 ppm, 161.94 ppm and 133.79 ppm, respectively. On Ae. aegypti, an high adulticidal effect was reported for Piper sarmentosum, followed by Piper ribesoides and Piper longum, with LD50 values of 0.14, 0.15 and 0.26 microg/female, respectively [36]. [49] have reported that the adulticidal activity was observed testing the methanol extracts of E. alba and A. paniculata on An. stephensi, LC50 and LC90 values were of 150.36, 130.19 ppm and 285.22, and 244.16 ppm respectively. Besides this interesting data, few efforts have been done to shed light on the contact toxicity of green-fabricated AgNP on mosquito adults [11]. We hypothesized that an important toxicity effect can be due to the magnified action of bioactive botanicals capping the wide surfaces of the nanocomposite.
Table 5

Adulticidal toxicity of Chenopodium ambrosioides leaf extract-synthesized silver nanoparticles against the dengue and Zika virus vector Aedes albopictus

Treatment

LC50 (LC90)

95% Confidence Limit LC50 (LC90)

Regression equation

χ 2

(d.f. = 4)

Lower

Upper

C. ambrosioides leaf extract

154.99 (330.15)

137.37 (299.87)

171.13 (373.72)

y = 1.134 + 0.007x

2.13 n.s

Ag nanoparticles

14.29 (30.03)

12.62 (27.51)

15.79 (33.54)

y = 1.164 + 0.081x

3.97 n.s

No mortality was observed in the control

LC 50 lethal concentration that kills 50% of the exposed organisms

LC 90 = lethal concentration that kills 90% of the exposed organisms

χ 2 Chi square value

d.f. degrees of freedom

n.s. not significant (α = 0.05)

Impact of Ag nanoparticles on Oryzias melastigma predation

Biological control of mosquito larval populations using aquatic predators, such as insects, fishes, copepods, and tadpoles recently received renewed attention (e.g. [19, 81, 82, 83, 84, 85, 126, 127]; see [17] for a review). In this present study, the rice fish, O. melastigma showed high predation rates on the dengue vector A. albopictus 2nd and 3rd instar larvae post-treatment with very low doses of AgNP. In standard conditions, after 24 h, the predation rates of II and III instar larvae of A. albopictus were 65.5 and 59.0%. Predation by O. melastigma post-treatment with ultra-low dosages of C. ambrosioides aqueous extract were 75.0 and 83.0%, while post-treatment with green-synthesized AgNP reached 91.0 and 85.5% against II and III instar larvae, resepctively (Table 6). No detectable toxicity effects were observed on O. melastigma individuals exposed to the AgNP-contaminated aquatic environment (post-treatment observation period: 10 days; data not shown). In agreement with our data, previous studies testing other aquatic species, including Gambusia affinis [28, 126];), Hoplobatrachus tigerinus [84], Poecilia reticulata [85], Aplocheilus lineatus [127], Carassius auratus (Linneaus) [29], Xenontodon cancila [27], Ctenopharyngodon idella, Cyprinus carpio [30], Oreochromis niloticus niloticus [25, 46], Betta splendens, Pseudotropheus tropheops tropheops, Osphronemus goramy Lacépède, and Pterophyllum scalare [45], showed good predatory ability on mosquito larvae under similar testing conditions. Recently, Chobua et al. [34] studied G. affinis and C. auratus as control agents of A. gambiae. In addition, our data, in agreement with recent researches [54, 58, 88, 126, 127], highlighted the limited toxicity of plant extracts and green nanocomposites on mosquito natural enemies, as well as the chance to use both tools in synergy to successfully manage mosquito young instar populations.
Table 6

Predation efficiency of Oryzias melastigma fishes against Aedes albopictus larvae in standard laboratory conditions and post-treatment with sub-lethal doses of the leaf extract of Chenopodium ambrosioides-fabricated silver nanoparticles

Treatment

Target

Exposure time

Total predation per fish (%)

0–6 h

6–12 h

12–18 h

18–24 h

Control

Larva II

30 ± 1.41

32 ± 2.54

35 ± 1.22

34 ± 2.23

65.5d

Larva III

29 ± 1.41

31 ± 1.58

29 ± 2.34

28 ± 1.22

59.0e

C. ambrosioides extract

Larva II

43 ± 2.16

40 ± 2.94

41 ± 1.41

42 ± 0.81

83.0bc

Larva III

38 ± 0.70

37 ± 2.12

36 ± 1.22

39 ± 2.34

75.0c

Silver nanoparticles

Larva II

46 ± 1.58

43 ± 1.87

46 ± 1.73

47 ± 1.22

91.0a

Larva III

43 ± 2.94

41 ± 2.16

42 ± 2.08

45 ± 2.21

85.5b

Predation rates are mean ± SD of five replicates (1 fish vs. 200 mosquitoes per replicate/day)

Control was clean water without fishes

Within the column, values followed by different letters are significantly different (generalized linear model, P < 0.05)

Conclusions

Overall, in the present work we reported the synthesis of AgNP at room temperature using the aqueous extract of C. ambrosioides, and their high toxicity against larvae, pupae and adults of the invasive mosquito A. albopictus. It is worthy to note that extremely low doses of the reducing extract and AgNP magnify the predation rates of O. melastigma fishes against A. albopictus 2nd and 3rd instar larvae, highlighting the concrete potential of C. ambrosioides-synthesized AgNP to develop effective and cheap tools to control young instars and adults of the invasive mosquito A. albopictus.

Notes

Acknowledgements

Two anonymous reviewers kindly improved an earlier version of our manuscript. J. Subramaniam is grateful to the Science and Engineering Research Board (SERB), Department of Science and Technology (DST), New Delhi, India for the financial support (Principal Investigator/NPDF-DST-SERB/Project File n. PDF/2015/000650).

Compliance with Ethical Standards

Conflicts of interest

The Authors declare no conflicts of interest.

Research Involving Human and Animal Rights

All applicable international and national guidelines for the care and use of animals were followed. All procedures performed in studies involving animals were in accordance with the ethical standards of the institution or practice at which the studies were conducted.

References

  1. 1.
    A. Aioi, T. Shimizu, and K. Kuriyama (1995). Int. J. Pharm. 113, 159.CrossRefGoogle Scholar
  2. 2.
    D. Amerasan, K. Murugan, K. Kovendan, P. M. Kumar, C. P. Selvam, J. Subramaniam, S. John William, and J. S. Hwang (2012). Parasitol. Res 111, 1953.CrossRefGoogle Scholar
  3. 3.
    D. Amerasan, T. Nataraj, K. Murugan, P. Madhiyazhagan, C. Panneerselvam, M. Nicoletti, and G. Benelli (2016). J. Pest Sci. 89, 249.CrossRefGoogle Scholar
  4. 4.
    A. Azizullah, Z. U. Rehman, I. Ali, W. Murad, N. Muhammad, W. Ullah, and D.-P. Hader (2014). Parasitol. Res 113, 4321.CrossRefGoogle Scholar
  5. 5.
    H. Bar, K. R. Dipak Bhui, P. Gobinda Sahoo, and P. Sarkar (2009). Coll. Surf. A. Physicochem. Eng. Asp. 339, 134.CrossRefGoogle Scholar
  6. 6.
    M. E. Barbercheck and J. Wang (1996). J. Insect. Pathol. 68, (2), 141.CrossRefGoogle Scholar
  7. 7.
    S. K. Batabyal, C. Basu, A. R. Das, and G. S. Sanyal (2007). J. Biobased Mater. Bioenerg. 1, 143.Google Scholar
  8. 8.
    N. A. Begum, S. Mondal, S. Basu, R. A. Laskar, and D. Mandal (2009). Coll. Surf. B 71, 113.CrossRefGoogle Scholar
  9. 9.
    G. Benelli (2015). Parasitol. Res 114, 2801.CrossRefGoogle Scholar
  10. 10.
    G. Benelli (2015). Parasitol. Res 114, 3201.CrossRefGoogle Scholar
  11. 11.
    G. Benelli (2016). Parasitol. Res 115, 23.CrossRefGoogle Scholar
  12. 12.
    G. Benelli (2016). Asia Pacif. J. Trop. Biomed. 6, 353.CrossRefGoogle Scholar
  13. 13.
    G. Benelli (2016). Enzyme. Microb. Technol. doi: 10.1016/j.enzmictec.2016.08.022.Google Scholar
  14. 14.
    G. Benelli and H. Mehlhorn (2016). Parasitol. Res 115, 1747.CrossRefGoogle Scholar
  15. 15.
    G. Benelli, A. Lo Iacono, A. Canale, and H. Mehlhorn (2016). Parasitol. Res 115, 2131.CrossRefGoogle Scholar
  16. 16.
    G. Benelli, R. Pavela, A. Canale, and H. Mehlhorn (2016). Parasitol. Res. doi: 10.1007/s00436-016-5095-1.Google Scholar
  17. 17.
    G. Benelli, C. L. Jeffries, and T. Walker (2016). Insects. 7, 52.CrossRefGoogle Scholar
  18. 18.
    S. Bhatt, P. W. Gething, and O. J. Brady (2013). Nature. 496, 504.CrossRefGoogle Scholar
  19. 19.
    G. Bowatte, P. Perera, G. Senevirathne, S. Meegaskumbura, and M. Meegaskumbura (2013). Biol. Control 67, 469.CrossRefGoogle Scholar
  20. 20.
    O. J. Brady, P. W. Gething, S. Bhatt, J. P. Messina, J. S. Brownstein, and A. G. Hoen (2012). PLoS Negl. Trop. Dis. 6, e1760. doi: 10.1371/journal.pntd.0001760.CrossRefGoogle Scholar
  21. 21.
    L. J. Bruce-Chwatt (1985). Drugs Exp. Clin. Res 11, 899.Google Scholar
  22. 22.
    S. D. Caruthers, S. A. Wickline, and G. M. Lanza (2007). Curr. Opin. Biotech. 18, 26.CrossRefGoogle Scholar
  23. 23.
    A. M. Carvalho Plantasy sabidurıa popular del Parque Natural de Montesinho. Un estudio etnobotanico en Portugal. Biblioteca de Ciencias, 35 (Consejo Superior de Investigaciones Cientı´ficas, Madrid, 2010).Google Scholar
  24. 24.
    J. F. Cavalli, F. Tomi, A. F. Bernardini, and J. Casanova (2004). Phytochem Anal 15, 275.CrossRefGoogle Scholar
  25. 25.
    S. K. Chand and R. S. Yadav Use of Oreochromis mossambicus (Peters) in controlling mosquito breeding in cow dung pits. in V. P. Sharma and A. Ghosh (eds.), Larvivorous Fishes of Inland Ecosystems (Malaria Research Centre, Delhi, 1994), p. 115.Google Scholar
  26. 26.
    G. Chandra, I. Bhattacharjee, S. N. Chatterjee, and A. Ghosh (2008). Indian J. Med. Res 127, 13.Google Scholar
  27. 27.
    S. N. Chatterjee and G. Chandra (1996). Environ. Ecol. 14, 173.Google Scholar
  28. 28.
    S. N. Chatterjee and G. Chandra (1997). Sci. Cult. 63, 51.Google Scholar
  29. 29.
    S. N. Chatterjee, S. Das, and G. Chandra (1997). Transact. Zool. Soc. India 1, 112.Google Scholar
  30. 30.
    S. N. Chatterjee, A. Ghosh, and G. Chandra (2001). Transact. Zool. Soc. India 5, 83.Google Scholar
  31. 31.
    S. S. Cheng, J. Y. Liu, K. H. Tsai, W. J. Chen, and S. T. Chang (2004). J. Agric. Food Chem. 52, 4395.CrossRefGoogle Scholar
  32. 32.
    L. Cheryl, T. Nancy, B. Gerhard, L. Grant, and G. Karla (2006). J. Ethnobiol. Ethnomed. doi: 10.1186/1746-4269-2-31.Google Scholar
  33. 33.
    K. Cho, J. Park, T. Osaka, and S. Park (2005). Electrochim. Acta 51, 956.CrossRefGoogle Scholar
  34. 34.
    M. Chobua, G. Nkwengulilaa, A. M. Mahandeb, B. J. Mwang’ondeb, and J. E. Kwekab (2015). Acta. Trop. 142, 131.CrossRefGoogle Scholar
  35. 35.
    K. Y. Choi, G. Liu, S. Lee, and X. Chen (2012). Nanoscale. 4, 330.CrossRefGoogle Scholar
  36. 36.
    W. Choochote, U. Chaithong, K. Kamsuk, E. Rattanachanpichai, A. Jitpakdi, P. Tippawangkosol, D. Chaiyasit, D. Champakaew, B. Tuetun, and B. Pitasawat (2006). Rev. Inst. Med. Trop. S. Paulo. 48, 33.CrossRefGoogle Scholar
  37. 37.
    A. Costello, T. Dua, P. Duran, M. Gülmezoglu, O. T. Oladapo, W. Perea, J. Pires, P. R. Pardo, N. Rollins, and S. Saxena (2016). Bull. World Health Organ 94, 406. doi: 10.2471/BLT.16.176990.CrossRefGoogle Scholar
  38. 38.
    G. V. B. Cruz, P. V. S. Pereira, F. J. Patrıcio, G. C. Costa, S. M. Sousa, J. B. Frazao, W. C. Aragao-Filho, M. C. G. Maciel, Amaral F. M. M. SilvaLA, E. S. B. Barroqueiro, R. N. M. Guerra, and F. R. F. Nascimento (2007). J. Ethnopharmacol. 111, 148.CrossRefGoogle Scholar
  39. 39.
    M. R. Das, R. K. Sarma, R. Saikia, V. S. Kale, M. V. Shelke, and P. Sengupta (2011). Coll. Surf. B 83, 16.CrossRefGoogle Scholar
  40. 40.
    M. P. Desai, V. Labhasetwar, E. Walter, R. J. Levy, and G. L. Amidon (1997). Pharm. Res 14, 1568.CrossRefGoogle Scholar
  41. 41.
    D. Dinesh, K. Murugan, P. Madhiyazhagan, C. Panneerselvam, M. Nicoletti, W. Jiang, G. Benelli, B. Chandramohan, and U. Suresh (2015). Parasitol. Res 114, 1519.CrossRefGoogle Scholar
  42. 42.
    A. D. Dwivedi and K. Gopal (2010). Coll. Surf. A. 369, (2010), 27.CrossRefGoogle Scholar
  43. 43.
    A. M. Fayaz, K. Balaji, Y. R. GirilalM, P. T. Kalaichelvan, and R. Venketesan (2010). Nanomed. Nanotechnol. Biol. Med. 6, 103.CrossRefGoogle Scholar
  44. 44.
    D. J. Finney Probit Analysis (Cambridge University Press, London, 1971), p. 68.Google Scholar
  45. 45.
    A. Ghosh, I. Bhattacharjee, M. Ganguly, S. Mandal, and G. Chandra (2004). Bull. Penelit. Kesehat. 32, 144.Google Scholar
  46. 46.
    A. Ghosh, I. Bhattacharjee, and G. Chandra (2006). J Appl Zool Res 17, 114.Google Scholar
  47. 47.
    A. P. Goncalvez, R. E. Engle, M. St Claire, R. H. Purcell, and C. J. Lai (2007). Proc. Natl. Acad. Sci. USA. 104, 9422.CrossRefGoogle Scholar
  48. 48.
    P. Gong, H. Li, X. He, K. Wang, J. Hu, W. Tan, S. Zhang, and X. Yang (2007). Nanotechnology. 18, 285604.CrossRefGoogle Scholar
  49. 49.
    M. Govindarajan and R. Sivakumar (2011). Asia. Pacif. J. Trop. Med. 4, (1), 941.CrossRefGoogle Scholar
  50. 50.
    M. Govindarajan and G. Benelli (2016). Parasitol. Res 115, 925.CrossRefGoogle Scholar
  51. 51.
    M. Govindarajan and G. Benelli (2016). RSC. Adv. 6, 59021.CrossRefGoogle Scholar
  52. 52.
    M. Govindarajan and G. Benelli (2016). J. Clust. Sci. doi: 10.1007/s10876-016-1035-6.Google Scholar
  53. 53.
    M. M. Green and J. M. Singer (1981). J. Am. Mosq. Control Assoc. 7, 282.Google Scholar
  54. 54.
    K. M. Haldar, B. Haldar, and G. Chandra (2013). Parasitol. Res 112, 1451.CrossRefGoogle Scholar
  55. 55.
    J. Hemingway and H. Ranson (2000). Annu. Rev. Entomol. 45, 371.CrossRefGoogle Scholar
  56. 56.
    A. Henglein (1993). J. Phys. Chem. 97, 5457.CrossRefGoogle Scholar
  57. 57.
    J. Huang, G. Zhan, B. Zheng, D. Sun, F. Lu, and Y. Lin (2011). Ind. Eng. Chem. Res 50, 9095.CrossRefGoogle Scholar
  58. 58.
    A. Jaganathan, K. Murugan, C. Panneerselvam, P. Madhiyazhagan, D. Dinesh, C. Vadivalagan, A. T. Aziz, B. Chandramohan, U. Suresh, R. Rajaganesh, J. Subramaniam, M. Nicoletti, A. Higuchi, A. A. Alarfaj, M. A. Munusamy, S. Kumar, and G. Benelli (2016). Parasitol. Int. 65, 276.CrossRefGoogle Scholar
  59. 59.
    Y. S. Jang, M. K. Kim, Y. S. Ahn, and H. S. Lee (2002). Agric. Chem. Biotechnol. 4, 131.Google Scholar
  60. 60.
    K. C. Jayaram The Freshwater Fishes of India, Pakistan, Bangladesh, Burma and Sri Lanka (ZSI, Calcutta, 1981).Google Scholar
  61. 61.
    M. Kalyanasundaram and P. K. Das (1985). Indian. J. Med. Res 82, 19.Google Scholar
  62. 62.
    E. G. Kamel, M. A. El-Emam, S. S. M. Mahmoud, F. M. Fouda, and F. E. Bayaumy (2011). Parasitol. Int. 60, 388.CrossRefGoogle Scholar
  63. 63.
    K. Kovendan and K. Murugan (2011). Adv. Environ. Biol. 5, 335.Google Scholar
  64. 64.
    A. T. Le, P. T. Huy, P. D. Tam, T. Q. Huy, P. D. Cam, and A. A. Kudrinskiy (2010). Curr. Appl. Phys. 10, 910.CrossRefGoogle Scholar
  65. 65.
    V. Louca, M. C. Lucas, C. Green, S. Majambere, U. Fillinger, and S. W. Lindsay (2009). J. Med. Entomol. 46, 546.CrossRefGoogle Scholar
  66. 66.
    P. Madhiyazhagan, K. Murugan, A. Naresh Kumar, T. Nataraj, D. Dinesh, C. Panneerselvam, J. Subramaniam, P. Mahesh Kumar, U. Suresh, M. Roni, M. Nicoletti, A. A. Alarfaj, A. Higuchi, M. A. Munusamy, and G. Benelli (2015). Parasitol. Res. doi: 10.1007/s00436-015-4671-0.Google Scholar
  67. 67.
    P. Magudapathy, P. Gangopadhyay, B. K. Panigrahi, K. G. M. Nair, and S. Dhara (2001). Physica 299, 142.Google Scholar
  68. 68.
    A. K. Manna (1989). Environ. Ecol. 7, 502.Google Scholar
  69. 69.
    A. K. Manna and S. Bannerjee (1984). Sci. Cult. 50, 329.Google Scholar
  70. 70.
    A. K. Manna and S. Bannerjee (1985). Environ. Ecol. 3, 456.Google Scholar
  71. 71.
    S. Marimuthu, A. A. Rahuman, G. Rajakumar, T. Santhoshkumar, A. Vishnu Kirthi, C. Jayaseelan, A. Bagavan, A. Abduz Zahir, G. Elango, and C. Kamaraj (2011). Parasitol. Res 108, 1541.CrossRefGoogle Scholar
  72. 72.
  73. 73.
    A. G. K. Menon Indigenous Larvivorous Fishes of India (NIMR, New Delhi, 1991).Google Scholar
  74. 74.
    H. Mehlhorn (ed.) Encyclopedia of Parasitology, 4th ed (Springer, New York, 2015).Google Scholar
  75. 75.
    H. Mehlhorn, K. A. Al-Rasheid, S. Al-Quraishy, and F. Abdel-Ghaffar (2012). Parasitol. Res 110, 259.CrossRefGoogle Scholar
  76. 76.
    T. A. Miettinen and H. Vanhanen (1994). Am. J. Clin. Nutr. 59, 356.Google Scholar
  77. 77.
    Mishra A (2002) Evaluation of some higher plant products for their pesticidal activity against some storage fungi and insects. Ph D thesis. Banaras Hindu University, Varanasi, India.Google Scholar
  78. 78.
    P. Mukherjee, M. Roy, B. P. Mandal, G. K. Dey, P. K. Mukherjee, J. Ghatak, A. K. Tyagi, and S. P. Kale (2008). Nanotechnology 19, 075103.CrossRefGoogle Scholar
  79. 79.
    K. S. Mukunthan, E. K. Elumalai, T. N. Patel, and V. R. Murty (2011). Asian Pac. J. Trop. Biomed. 1, 270.CrossRefGoogle Scholar
  80. 80.
    K. Murugan, V. Vahitha, I. Baruah, and S. C. Das (2003). Ann. Med. Entomol. 12, 11.Google Scholar
  81. 81.
    K. Murugan, J. S. Hwang, K. Kovendan, K. Prasanna Kumar, C. Vasugi, and A. Naresh Kumar (2011). Hydrobiologia 666, 331.CrossRefGoogle Scholar
  82. 82.
    K. Murugan, N. Aarthi, K. Kovendan, C. Panneerselvam, B. Chandramohan, P. M. kumar, D. Amerasan, M. Paulpandi, R. Chandirasekar, D. Dinesh, U. Suresh, J. Subramaniam, A. Higuchi, A. A. Alarfaj, M. Nicoletti, H. Mehlhorn, and G. Benelli (2015). Parasitol. Res 114, 3657.CrossRefGoogle Scholar
  83. 83.
    K. Murugan, C. M. Samidoss, C. Panneerselvam, A. Higuchi, M. Roni, U. Suresh, B. Chandramohan, J. Subramaniam, P. Madhiyazhagan, D. Dinesh, R. Rajaganesh, A. A. Alarfaj, M. Nicoletti, S. Kumar, H. Wei, A. Canale, H. Mehlhorn, and G. Benelli (2015). Parasitol. Res 114, 4087.CrossRefGoogle Scholar
  84. 84.
    K. Murugan, V. Priyanka, D. Dinesh, P. Madhiyazhagan, C. Panneerselvam, J. Subramaniam, U. Suresh, B. Chandramohan, M. Roni, M. Nicoletti, A. A. Alarfaj, A. Higuchi, M. A. Munusamy, H. F. Khater, R. H. Messing, and G. Benelli (2015). Parasitol. Res. doi: 10.1007/s00436-015-4582-0.Google Scholar
  85. 85.
    K. Murugan, J. S. E. Venus, C. Panneerselvam, S. Bedini, B. Conti, M. Nicoletti, S. K. Sarkar, J. S. Hwang, J. Subramaniam, P. Madhiyazhagan, P. M. Kumar, D. Dinesh, U. Suresh, and G. Benelli (2015). Environ. Sci. Poll. Res. doi: 10.1007/s11356-015-4920-x.Google Scholar
  86. 86.
    K. Murugan, M. Aamina Labeeba, C. Panneerselvam, D. Dinesh, U. Suresh, J. Subramaniam, P. Madhiyazhagan, J. S. Hwang, L. Wang, M. Nicoletti, and G. Benelli (2015). Res. Vet. Sci. 102, 127.CrossRefGoogle Scholar
  87. 87.
    K. Murugan, P. Aruna, C. Panneerselvam, P. Madhiyazhagan, M. Paulpandi, J. Subramaniam, R. Rajaganesh, H. Wei, M. Saleh Alsalhi, Nicoletti M. DevanesanS, B. Syuhei, A. Canale, and G. Benelli (2016). Parasitol. Res 115, 651.CrossRefGoogle Scholar
  88. 88.
    K. Murugan, J. Anitha, D. Dinesh, U. Suresh, R. Rajaganesh, B. Chandramohan, J. Subramaniam, M. Paulpandi, C. Vadivalagan, P. Amuthavalli, L. Wang, J. S. Hwang, H. Wei, M. S. Alsalhi, S. Devanesan, S. Kumar, K. Pugazhendy, A. Higuchi, M. Nicoletti, and G. Benelli (2016). Ecotoxicol. Environ. Saf 132, 318.CrossRefGoogle Scholar
  89. 89.
    K. Murugan, C. Panneerselvam, A. T. Aziz, J. Subramaniam, P. Madhiyazhagan, J. S. Hwang, Lan Wang, D. Dinesh, U. Suresh, M. Roni, A. Higuchi, M. Nicoletti, M. Saleh Alsalhi, and G. Benelli (2016). Environ. Sci. Poll. Res. doi: 10.1007/s11356-016-6832-9.Google Scholar
  90. 90.
    K. Murugan, D. Nataraj, P. Madhiyazhagan, V. Sujitha, B. Chandramohan, C. Panneerselvam, D. Dinesh, R. Chandirasekar, K. Kovendan, U. Suresh, J. Subramaniam, M. Paulpandi, C. Vadivalagan, R. Rajaganesh, H. Wei, B. Syuhei, A. T. Aziz, M. Saleh Alsalhi, S. Devanesan, M. Nicoletti, A. Canale, and G. Benelli (2016). Parasitol. Res 115, 1071.CrossRefGoogle Scholar
  91. 91.
    M. N. Naqqash, A. Gökçe, A. Bakhsh, and M. Salim (2016). Parasitol. Res 115, 1363.CrossRefGoogle Scholar
  92. 92.
    F. R. F. Nascimento, G. V. B. Cruz, P. V. S. Pereira, M. C. G. Maciel, L. A. Silva, A. P. S. Azevedo, E. S. B. Barroqueiro, and R. N. M. Guerra (2006). Life Sci. 78, 2650.CrossRefGoogle Scholar
  93. 93.
    S. Navaladian, B. Viswanathan, R. P. Viswanath, and T. K. Varadarajan (2007). Nanoscale Res. Lett. 2, 44.CrossRefGoogle Scholar
  94. 94.
    H. L. Newmark (1997). Cancer Epidemiol Biomark Prev. 6, 1101.Google Scholar
  95. 95.
    M. Nicoletti, S. Mariani, O. Maccioni, T. Coccioletti, and K. Murugan (2012). Parasitol. Res 111, 205.CrossRefGoogle Scholar
  96. 96.
    S. Y. Ohba, Dida G. O. KawadaH, D. Juma, G. Sonye, N. Minakawa, and M. Takagi (2010). J. Med. Entomol. 47, 783.CrossRefGoogle Scholar
  97. 97.
    C. Panneerselvam and K. Murugan (2013). Parasitol Res 112, (2), 679.CrossRefGoogle Scholar
  98. 98.
    C. Panneerselvam, K. Murugan, M. Roni, A. T. Aziz, U. Suresh, R. Rajaganesh, P. Madhiyazhagan, J. Subramaniam, D. Dinesh, M. Nicoletti, A. Higuchi, A. A. Alarfaj, M. A. Munusamy, S. Kumar, N. Desneux, and G. Benelli (2016). Parasitol. Res 115, 997.CrossRefGoogle Scholar
  99. 99.
    R. Pavela and G. Benelli (2016). Tr Plant Sci. doi: 10.1016/j.tplants.2016.10.005.Google Scholar
  100. 100.
    T. N. V. K. V. Prasad and E. K. Elumalai (2011). Asian Pac. J. Trop. Biomed. 1, 439.CrossRefGoogle Scholar
  101. 101.
    A. Priyadarshini, K. Murugan, C. Panneerselvam, S. Ponarulselvam, H. Jiang Shiou, and M. Nicoletti (2012). Parasitol. Res 111, 997.CrossRefGoogle Scholar
  102. 102.
    G. Rajakumar and A. A. Rahuman (2011). Acta Trop. 118, 196.CrossRefGoogle Scholar
  103. 103.
    R. Rajan, K. Chandran, S. L. Harper, S. I. Yun, and P. T. Kalaichelvan (2015). Ind. Crops Prod. 70, 356.CrossRefGoogle Scholar
  104. 104.
    C. V. Rao, H. L. Newmark, and B. S. Reddy (1998). Carcinogenesis. 19, 287.CrossRefGoogle Scholar
  105. 105.
    J. V. Rao and P. Kavitha (2010). Z Naturforsch. C. 65, 303.Google Scholar
  106. 106.
    B. K. Ravindra and A. H. Rajasab (2014). Int. J. Pharm. Pharm. Sci. 6, 372.Google Scholar
  107. 107.
    M. Roni, K. Murugan, C. Panneerselvam, J. Subramaniam, M. Nicoletti, P. Madhiyazhagan, D. Dinesh, U. Suresh, H. F. Khater, H. Wei, Alarfaj A. A. CanaleA, M. A. Munusamy, A. Higuchi, and G. Benelli (2015). Ecotoxicol. Environ. Saf. doi: 10.1016/jecoenv201507005.Google Scholar
  108. 108.
    D. E. Rosen and L. R. Parenti (1981). Am. Mus. Novit. 27, 1.Google Scholar
  109. 109.
    M. A. Rude and A. Schirmer (2009). Curr. Opin. Microbiol. 12, 274.CrossRefGoogle Scholar
  110. 110.
    T. Santhoshkumar, A. A. Rahuman, G. Rajakumar, S. Marimuthu, A. Bagavan, C. Jayaseelan, A. A. Zahir, G. Elango, and C. Kamaraj (2010). Parasitol. Res 108, 693.CrossRefGoogle Scholar
  111. 111.
    R. Sathyavathi, M. Balamurali Krishna, S. Venugopal Rao, R. Saritha, and D. Narayana Rao (2010). Adv. Sci. Lett. 3, 1.CrossRefGoogle Scholar
  112. 112.
    Service MW (1977). J. Med. Entomol. 13, 535.CrossRefGoogle Scholar
  113. 113.
    S. S. Shankar, A. Rai, A. Ahmad, and M. Sastry (2004). J. Coll. Interf. Sci. 275, 496.CrossRefGoogle Scholar
  114. 114.
    V. P. Sharma and A. Ghosh Larvivorous Fishes of Inland Ecosystem (NIMR, New Delhi, 1994).Google Scholar
  115. 115.
    B. P. Singh, B. J. Hatton, B. Singh, A. L. Cowie, and A. Kathuria (2010). J. Environ. Qual. 39, 1.CrossRefGoogle Scholar
  116. 116.
    R. Sivakumar, A. Jebanesan, M. Govindarajan, and P. Rajasekar (2011). Asia. Pacif. J. Trop. Med. 2011, 706.CrossRefGoogle Scholar
  117. 117.
    S. Sivapriyajothi, P. Mahesh Kumar, K. Kovendan, J. Subramaniam, and K. Murugan (2014). J. Entomol. Acarol. Res 46, 1787.CrossRefGoogle Scholar
  118. 118.
    T. J. Smith (2000). Exp. Opin. Invest. Drugs. 9, 1841.CrossRefGoogle Scholar
  119. 119.
    T. J. Smith, G. Y. Yang, D. N. Seril, J. Liao, and S. Kim (1998). Carcinogenesis. 19, (4), 703.CrossRefGoogle Scholar
  120. 120.
    K. S. Sreeram, M. Nidin, and B. U. Nair (2008). Bull. Mater. Sci. 31, 937.CrossRefGoogle Scholar
  121. 121.
    M. Starowicz, B. Stypuła, and J. Banas (2006). Electrochem. Commun. 8, 227.CrossRefGoogle Scholar
  122. 122.
    J. Subramaniam, K. Murugan, and K. Kovendan (2012). J. Biopestic. 5, 163.Google Scholar
  123. 123.
    J. Subramaniam and M. Murugan (2013). Evaluation of larvicidal, pupicidal, repellent, and adulticidal activity of Myristica fragrans (Family: Myristicaceae) against malarial vector Anopheles stephensi. Proceedings of the national conference on insect diversity and systematics: special emphasis on molecular approaches. pp:1–6.Google Scholar
  124. 124.
    J. Subramaniam and K. Murugan (2014). J. Int. J. Biol. Sci. 2014, 33.Google Scholar
  125. 125.
    J. Subramaniam, K. Murugan, K. Kovendan, P. M. Kumar, D. Amerasan, C. P. Selvam, T. Nataraj, D. Dinesh, B. Chandramohan, R. Chandrasekar, and J. S. Hwang (2015). Util. Manag. Med. Plants. 3, 261.Google Scholar
  126. 126.
    J. Subramaniam, K. Murugan, C. Panneerselvam, K. Kovendan, P. Madhiyazhagan, P. M. Kumar, D. Dinesh, B. Chandramohan, U. Suresh, M. Nicoletti, A. Higuchi, J. S. Hwang, S. Kumar, A. A. Alarfaj, M. A. Munusamy, R. H. Messing, and G. Benelli (2015). Environ. Sci. Poll. Res 22, (24), 20067.CrossRefGoogle Scholar
  127. 127.
    J. Subramaniam, K. Murugan, C. Panneerselvam, K. Kovendan, P. Madhiyazhagan, D. Dinesh, P. Mahesh Kumar, B. Chandramohan, U. Suresh, R. Rajaganesh, Mohamad Saleh Alsalhi, S. Devanesan, M. Nicoletti, A. Canale, and G. Benelli (2016). Environ. Sci. Pollut. Res. doi: 10.1007/s11356-015-6007-0.Google Scholar
  128. 128.
    U. Suresh, K. Murugan, G. Benelli, M. Nicoletti, D. R. Barnard, C. Panneerselvam, P. Mahesh Kumar, J. Subramaniam, D. Dinesh, and B. Chandramohan (2015). Parasitol. Res 114, 1551.CrossRefGoogle Scholar
  129. 129.
    G. Taubes (2000). Science. 290, (5491), 434.CrossRefGoogle Scholar
  130. 130.
    S. Thiripura and S. Shankar (2014). Int. J. Pharm. Tech. Res 6, 1731.Google Scholar
  131. 131.
    A. Voyadjoglou, V. Roussis, and P. V. Petrakis Biological control of mosquito populations: an applied aspect of pest control by means of natural enemies. in A. M. T. Elewa (ed.), Predation in Organisms: Adistinct Phenomenon (Springer, Berlin, 2007), p. 123.CrossRefGoogle Scholar
  132. 132.
    W. W. Walton (2007). J. Am. Mosq. Control Assoc. 23, (S2), 184.CrossRefGoogle Scholar
  133. 133.
    WHO (1981) Instructions for determining the susceptibility or resistance of adult mosquitoes to organochlorine, organophosphate and carbamate insecticides: diagnostic test. WHO/VBC/81–807, Geneva.Google Scholar
  134. 134.
    WHO (2015) Dengue and severe dengue. Fact sheet N°117. World Health Organization, Geneva.Google Scholar
  135. 135.
    WHO (2016) Weekly epidemiological record, No 7, 19 Feb 2016, Vol. 91, 73–88.Google Scholar
  136. 136.
    H. Yap (1985). Southeast Asian J. Trop. Med. Public Health. 16, 163.Google Scholar
  137. 137.
    M. Zargar, A. A. Hamid, F. A. Bakar, M. N. Shamsudin, K. Shameli, F. Jahanshiri, and F. Farahani (2011). Molecules. 6, 6667.CrossRefGoogle Scholar
  138. 138.
    J. Zhu, X. Zeng, M. O. Neal, G. Schultz, B. Tucker, and J. Coats (2008). J. Am. Mosq. Control. Assoc. 24, 161.CrossRefGoogle Scholar

Copyright information

© Springer Science+Business Media New York 2016

Authors and Affiliations

  • Jayapal Subramaniam
    • 1
    • 2
  • Kadarkarai Murugan
    • 2
    • 3
  • Arulsamy Jebanesan
    • 1
  • Philips Pontheckan
    • 2
  • Devakumar Dinesh
    • 2
  • Marcello Nicoletti
    • 4
  • Hui Wei
    • 5
  • Akon Higuchi
    • 6
  • Suresh Kumar
    • 7
  • Angelo Canale
    • 8
  • Giovanni Benelli
    • 8
  1. 1.Divison of Vector Biology and Control, Department of Zoology, Faculty of ScienceAnnamalai UniversityChidambaramIndia
  2. 2.Division of Entomology, Department of Zoology, School of Life SciencesBharathiar UniversityCoimbatoreIndia
  3. 3.Department of BiotechnologyThiruvalluvar UniversityVelloreIndia
  4. 4.Department of Environmental BiologySapienza University of RomeRomeItaly
  5. 5.Institute of Plant ProtectionFujian Academy of Agricultural SciencesFuzhouChina
  6. 6.Department of Chemical and Materials EngineeringNational Central UniversityTaoyuanTaiwan
  7. 7.Department of Medical Microbiology and ParasitologyUniversiti Putra Malaysia UPMSerdangMalaysia
  8. 8.Department of Agriculture, Food and Environment University of PisaPisaItaly

Personalised recommendations