1 Introduction

Arthropods are hazardous vectors that could influence the growing global human and animal population as epidemics and pandemics of lethal pathogens [1]. The most important threats faced by mosquitoes worldwide are those (Diptera: Culicidae), which are the vectors of important diseases such as yellow fever, Western Nile virus (WNV), dengue, malaria and filariasis. Millions of individuals globally are at risk and endangering life due to mosquito-borne illnesses, and have such a heavy public health burden that they are blamed on continuous continental underdevelopment [2].

The knowledge about ecology and the control of mosquitoes is of crucial importance in the sense of integrated vector management, to combat the spread of many mosquito-borne diseases [3, 4]. Ae. aegypti (Linnaeus) is the primary dengue and Zika vector, a cosmotropic species. As a vector of dengue fever and chikungunya in India, this species is particularly dangerous because the number of dengue fever instances has increased substantially over the past years [1, 5, 6]. Chemical insecticides used for regulating mosquitoes which are detrimental to human health and non-target organisms. Moreover, mosquito vectors have developed resistance towards the synthetic insecticides, namely temephos by carrying genes that encode acetylcholinesterases (AChE). AChE enzyme breaks down acetylcholine which carries the transmission of nerve impulses through the synaptic gap and acts as a primary target for synthetic insecticides [7]. Besides, a detoxifying enzyme such as Glutathione S-transferase (GST) is also expressed by insects which helps to neutralize the toxicity of different xenobiotics [8]. On these grounds, newly implemented eco-friendly methods have been used to improve mosquito vector control [9]. Young mosquito instars have shown decreased mobility and are therefore often easy to target. Thus mosquitoes can be killed before dispersing to human dwellings by targeting at their larval stages [10].

Plant materials showing mosquitocidal characteristics against major mosquito vectors have been recently emphasized. Many naturally occurring compounds isolated from plants exhibits excellent mosquito toxicants which act as ovicidal, pupicidal, adult repellents, growth and/or breeding inhibitors [11]. Lumnitzera racemosa Willd. (Combretaceae) is a true mangrove found along the coast of Asia. Due to the high calorific value, the wood of L. racemosa is used as a fuel, while its leaves are eaten on South Pacific Island all through periods of scarcity by herbivores [12]. Besides, the reddish bark contains ~ 15 × 19% tannins used in leather industries. Moreover, fluid extracted from the stem of this plant were found effective in treating itches and herpes when applied externally. The most important secondary metabolites of L. racemosa are flavonoids, triterpenoids, tannins, fatty acids and polyisoprenoid alcohols [13]. This plant exerts numerous pharmacological activities, such as antibacterial [14], antifungal and antihypertensive [15], protein tyrosine phosphatase 1B (PTP1B) inhibitory, hepatoprotective and antioxidant activities [16]. While recent studies revelled L. racemosa leaves to be toxic against larvae of Ae. aegypti [17].

Green technologies have recently been shown to be a new strategy for reducing the use of chemical pesticides since overuse has often led to massive non-target impacts and widespread resistance [9, 18]. Modern study has revealed an increased level of insecticide activity in green nanoparticles (NPs) synthesized by Euphorbia rothiana [19], Cymbopogon citratus [20], Pedalium murex [21] etc. Among different NPs, Zinc Oxide nanoparticles (ZnO NPs), having nanosized morphology, have increased interest among several researchers as a result of their exclusive optical or chemical activities [22]. ZnO NPs have remarkable characteristics, including large binding energy, broad bandgap, and elevated piezoelectric characteristics [23]. It has various applications, including UV light-emitting diodes, laser diodes, and catalysts [24]. Also, ZnO NPs have a highly promising prospect for biological functions such as gene delivery [25], biological sensing and labelling [26], drug delivery and nanomedicine [27, 28], antibacterial [29], antifungal [30], along with larvicidal, acaricidal, pediculicidal activity [31].

The present study was thus focussed on ZnO Nanorods (ZnO NRs) synthesis with a single step procedure based on easy to obtain L. racemosa flower buds aqueous extract. LB-ZnO NRs were characterized using a variety of biophysical methods. Furthermore, larvicidal toxicity of LB extract and LB-ZnO nanorods were evaluated on dengue vector Ae. aegypti and their enzyme modulation on AChE and GST enzymes were studied.

2 Materials and methods

2.1 Materials and reagents

Lumnitzera racemosa flower buds (LB) were collected from Bhatye beach area located at 16°58′44.0691″ N and 73°17′38.7499″ E. Ratnagiri District, Maharashtra, India and authenticated by an expert taxonomist. L. racemosa flower buds were washed thoroughly with tap water followed by distilled water to remove adhered particles. Flower buds were then kept for drying under the shade at room temperature (27 ± 2 °C). Water purified using Millipore Milli-Q (18.2 MΩ.cm at 25 °C) purification system was used throughout the experiment. Acetylthiocholine iodide was purchased from Hi-Media (Mumbai, India), reduced glutathione (GSH), 1-chloro2,4-dinitrobenzene (CDNB), DTNB reagent (5,5′-dithiobis-2-nitrobenzoic acid), Zinc acetate dihydrate, Agarose, Ethylenediaminetetraacetic acid (EDTA) and ethidium bromide were purchased from Sigma-Aldrich, India. Proteinase K and RNase A were procured from ThermoFisher Scientific, India.

2.1.1 Preparation of L. racemosa flower buds extract

Aqueous LB extract was prepared by adding 5 gm plant powder in 100 ml Milli-Q water (MQ-W) and placed in a water bath at 100 °C for 20 min. The obtained filtrate and residue were separated using Whatman filter paper No. 42 (42 ashless diameters 125 mm GE Healthcare Life Sciences). Filtrate solution thus obtained is filtered with a 0.2 μm cellulose syringe filter (Rayna Biotech, UK) and stored in the refrigerator at 4 °C and was later used for the synthesis of ZnO NPs.

2.1.2 Production of zinc oxide nanoparticles

To 80 ml of 10 mM zinc acetate dihydrate, 20 ml of obtained LB aqueous extract was added dropwise under constant stirring at room temperature. The resulting solution was then placed on a magnetic stirrer at 60 °C until the off-white suspension of NPs were formed. The solution was then centrifuged at 3000 rpm for 20 min and the residue was washed several times to remove any unreacted materials. The resulting supernatant was discarded, and the pellet was collected and placed in a furnace at 400 °C to obtain the desired product in powder form.

2.2 Characterization

2.2.1 UV–Vis spectroscopy (UV–Vis)

Optical properties of nanoparticles were examined using UV–Vis spectroscopy. Absorption spectrum of ZnO NPs after 24 h was recorded using UV–Visible spectrometer Shimadzu, UV-1800 (λ = 200–800 nm) to confirm the synthesis of LB-ZnO NPs.

2.2.2 Fourier transform infrared spectroscopy (FTIR)

Different types of functional groups which are associated with the synthesis of NPs was confirmed through FTIR analysis. The dried powder of ZnO NPs was thoroughly mixed with potassium bromide (KBr) (2:98 ratio w/w) and compressed at 11,000 psi to make the disc. The disk was then inserted into the FTIR spectrophotometer (Perkin Elmer Frontier, 91579) and the spectrum was recorded at the diffuse reflection mode at a resolution of 4 cm−1 in the test range of 500–4000 cm−1. A similar process with LB aqueous extract was also performed.

2.2.3 X-ray diffraction (XRD)

Crystalline nature of synthesized LB-ZnO NPs was recorded using XRD diffraction working at 40 kV with a current of 40 mA, Cu Kα radiation (1.54 Å) in a θ–2θ configuration using Bruker D8 Discover model (Bruker, Germany).

2.2.4 Scanning electron microscopy (SEM) and Energy-dispersive spectra (EDS)

Scanning electron microscopes (SEM) helps in the validation of NPs morphology. A few micrograms of LB-ZnO NPs were dropped on a carbon-coated copper grid for SEM analysis and the excess powder was discarded. The scanning electron microscopic images were then captured using JEOL System JSM-6390LV (Akishima, Tokyo, Japan) at an acceleration voltage of 15 kV. An energy dispersive spectrometer (EDS make FEI series Quanta 200 with EDS 3.0.13) at an energy range of 0–10 keV was used to take energy dispersive spectra for LB-ZnO NPs.

2.2.5 Zeta potential

Zeta potential was performed to analyze charge on synthesized LB-ZnO NPs using Zetasizer (Nano series Nano-ZS90, Malvern Instruments).

2.2.6 Transmission electron microscopy (TEM)

On carbon-coated copper grids, powdered LB-ZnO NPs were mounted and transmission electron microscopy observations were carried out using FEI Tecnai G2, F30 system working at an accelerating voltage of 200 kV.

2.3 Mosquito rearing and larvicidal activity

The larvae of Ae. aegypti were collected from Vidyanagari campus, University of Mumbai, Kalina, Mumbai, and surrounding areas. Collected larvae were identified by Dr. Mira Ramaiya, Entomologist, Haffkine Institute For Training, Research And Testing, Parel, Mumbai. The larval colony was maintained in the lab at 28 ± 1 °C with 80 ± 5% relative humidity, and 14L:10D photoperiod [32]. Adults were fed with 2% sucrose solution soaked in non-absorbent cotton. Female mosquitoes were fed with periodic blood meals for egg maturation by the membrane feeding technique [33]. The eggs were collected on Whatman filter paper lined in the bowl and allowed to hatch in dechlorinated water. A 2:1 (w/w) mixture of yeast powder and dog biscuits were used to feed the larvae. The pupae formed were gathered and moved to glass cages for adult emergence. Laboratory colonies of mosquito larvae (F1 generation) were used for the larvicidal activity.

Larvicidal activity of LB aqueous extract and synthesized LB-ZnO NPs was carried out under the WHO standard method [34] as follows. Accordingly, LB extract and LB-ZnO NPs were dispersed in dechlorinated water to prepare a graded series of concentration (500–2500 μg/ml, for aqueous LB extract and 10–50 μg/ml, for LB-ZnO NPs). Further, early 4th instar larvae of Ae. aegypti were transferred to 250 ml plastic cups in batches of 20 each containing 199 ml of distilled water and 1 ml of respective treatment in the graded concentrations. Each experiment was conducted in triplicates along with a respective control group on three separate days to ensure reproducibility. The control group consisted of 200 ml of distilled water only. No food was provided during the duration of the entire experiment. Symptoms of the treated larvae and larval mortality were assessed by observing their motility 24 h after exposure to their respective treatments.

2.3.1 Photomicrography studies

Studies of photomicrography were conducted with minor modifications based on Coelho et al. [35] methodology. Morphological alterations were checked in the 4th instar larvae of Ae. aegypti treated with LB extract and LB-ZnO NPs at their median lethal concentrations (LC50) along with the control group. The treatment groups (LB, LB-ZnO NPs) and control larvae were successively fixed at 27 °C for 30 min in alcohol (ethanol) dehydration (35–70%) and placed on glass slides. The larvae were analyzed under the Stereo zoom microscope (Leica microsystems) at × 20 magnification.

2.4 Enzymatic studies

To study the neurotoxicity and the effect of detoxification on Ae. aegypti larvae separate studies were conducted to analyze the effect of LB extract and LB synthesized ZnO NPs on AChE and GST enzymes. Early 4th instar larvae were exposed for 24 h to LB extract and ZnO NPs at their median lethal concentrations (LC50). It is advantageous to study the effect of a toxicant on the organism at the moment of evaluation but before mortality occurred. Accordingly, only the live larvae recovered after 24 h from both control and treated with LB extract and LB-ZnO NPs were used for further biochemical analysis.

2.4.1 Preparation of whole-body homogenates for assessment of enzyme activity

The larvae were retrieved after 24 h of treatment with LB aqueous extract and ZnO NPs (test larvae) alongside with untreated (control larvae) and washed with double distilled water accompanied by blotting their body surface with tissue paper to eliminate any contaminants. The collected larvae were then homogenized with a Teflon hand homogenizer in 0.5 ml phosphate buffer [100 mM phosphate buffer (pH 7.2) and 1% Triton X-100] in Eppendorf ™ tubes (submerged in ice) [36]. The homogenate was centrifuged at 10,000×g for 15 min at 4 °C, and the supernatant was used to assess enzyme activity. Enzyme protein was estimated according to the Bradford method [37].

2.4.2 Acetylcholinesterase (AChE) activity

AChE activity in the whole body homogenates of larvae was spectrophotometrically measured according to a modified method of Ellman et al. [38] using acetylthiocholine iodide as a substrate [39]. 50 µl aliquot of homogenate (100 μg/ml crude enzyme) was successively mixed with 450 µl sodium phosphate buffer (100 mM, pH 7.5); 50 µl of 10 mM DTNB and 50 µl of 12.5 mM acetylcholine iodide. After 5 min of incubation at room temperature, the optical density of the sample was read at 400 nm using a spectrophotometer (Shimadzu, UV-1800) against a suitable reagent blank (without enzyme). AChE activity was expressed as (µmol ACT released/min/mg protein) using published extinction coefficient (e = 1.3 mM−1 cm−1).

2.4.3 Glutathione S-transferase (GST) activity

GST activity was estimated by Vontas et al. [40] method as follows. 100 μl of 30 mM CDNB was added to the 100 μl larval homogenate (100 μg/ml crude enzyme) and the volume was adjusted to 500 μl using sodium phosphate buffer (100 mM, pH 7.5). After pre-incubation of the reaction mixture at 37 °C for 5 min, 100 μl of 30 mM of GSH was added. The change in the absorbance level was noted at 340 nm for 5 min after every 1 min interval in the spectrophotometer. The reaction mixture without enzyme was used as blank. GST activity was expressed as (µmol CDNB hydrolysed /min/mg protein) using the published extinction coefficient (e = 9.6 mM−1 cm−1) [41].

2.5 DNA fragmentation assay

To analyze any damage caused by LB extract and LB-ZnO NPs on mosquito DNA, fragmentation analysis was carried out as per established protocol [42]. Mosquito larvae treated with plant extract and ZnO NRs were washed with ethanol, distilled water to remove adhered particles. Larvae were lysed in a 250 μl cell lysis buffer containing 50 mM Tris–HCl, 0.5% SDS, 1 μg/ml proteinase K (pH 8.0), 10 mM of ethylenediaminetetraacetic acid (EDTA) and RNase A (0.5 μg/ml) at 37 °C. DNA was extracted by phenol/chloroform/isoamyl alcohol (25:24:1) method [43]. The 260/280 nm absorbance ratio was used to assess the purity of DNA [44]. Electrophoresis of larval DNA was performed on an agarose gel (1%) containing ethidium bromide (1 μg/ml) at 100 V, and the fragments of DNA were visualized by exposing the gel to UV light and photographed on Gel documentation system (Bio-Rad Gel Doc XR system).

2.6 Statistical data analysis

The data acquired were analyzed using the SPSS 21.5 program for regression analysis. The values of LC50 and LC90 with 95% fiducial limits and chi-square were calculated using Probit analysis in each bioassay. The data were expressed as Mean ± SEM. One-way variance analysis (ANOVA) followed by the post-hoc Tuckey test was performed to determine the difference in enzyme inhibition between control and treated groups. Results with p < 0.05, were considered to be statistically significant.

3 Results and discussion

3.1 Characterization

3.1.1 UV–Visible spectroscopy

UV–vis spectroscopy is an easy and efficient method for the identification of nanoparticle formation, stabilization, and shape [45]. The UV–vis spectrum of LB-ZnO NPs synthesized using LB aqueous extract and zinc acetate is shown in Fig. 1a. During synthesis, the color of the reaction mixture changed from yellow to pale white colored precipitate. The obtained product after calcination at 400 °C gave white precipitate indicating the synthesis of ZnO NPs which was further confirmed by UV–Vis spectroscopy. A characteristic absorption peak at 373.82 nm could be attributed, due to electron transfer from the valence band to the conductive band, related to ZnO's intrinsic band-gap absorption (O2p-zn3d) [46]. In comparison to the average absorption at 377 nm of ZnO NPs, the synthesized LB-ZnO NPs with a maximum wavelength at 373.82 nm were slightly blue-shifted which maybe because of the decrease in particle size with respect to bulk ZnO [47]. Mahamuni et al. [48] and Ishwarya et al. [49] reported a similar shift in the position of the absorption peak of ZnO NPs. Figure 1b shows the optical energy bandgap (Eg) of LB-ZnO NPs which was estimated using the following Tauc formula [50].

$${\text{Optical}}\,{\text{energy}}\,{\text{bandgap}}\,(Eg)\,\, = \frac{hc}{\lambda }$$
(1)

where h and c refers to planks constant (6.626 × 10−34 J s) and velocity of light (3 × 108 m/s) respectively and ‘λ’ denotes optical wavelength.

Fig. 1
figure 1

a UV–visible absorption spectrum of zinc oxide nanoparticles synthesized using L. racemosa flower buds aqueous extract [Inset: (A) L. racemosa flower buds extract and (B) synthesized ZnO NRs]. b Optical energy band gap of synthesized ZnO nanorods

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The synthesized LB-ZnO NP’s optical energy band gap was found to be 3.25 eV which is in good agreement with the previous report [50]. This changes in the optical band gap of LB-ZnO NPs concerning the ZnO nanostructure morphology further confirms that nanostructure crystallinity, crystal growth facets, and crystal grain size lead to the effective band gap of nanostructured ZnO smaller than its bulk value of 3.37 eV [51].

3.1.2 FT-IR analysis

FT-IR provides data on a molecule's vibrational and linear mode of motion and is, therefore, a significant method for identifying and characterizing a material. Infrared spectrum of any compound (organic or inorganic) offers a distinctive fingerprint that is easily differentiated from all other compound’s absorption patterns. The FT-IR spectrum of LB aqueous extract and synthesized LB-ZnO NPs are shown in Fig. 2a, b respectively. In Fig. 2a, spectral peak at 3291 cm−1 is due to O–H stretching and 2922 cm−1 is due to C–H stretch [52]. Absorption peak from the carbonyl group appears at 1613 cm−1, whereas the band at 1235 cm−1 observed is due to primary amine groups and –C–O stretching mode [53]. The absorption band at 1017 cm−1 is due to C–N stretching oscillation mode of amines that are characteristics of proteins/enzymes [54, 55]. In Fig. 2b which corresponds to the FT-IR spectrum of LB-ZnO NPs suggests that the phytoconstituents of LB aqueous extracts were either degraded or remained absorbed in smaller quantities due to calcination of NPs at higher temperature [56]. The peak observed in the present study at 900 cm−1 are due to the asymmetrical and symmetrical stretching of the zinc carboxylate. Previous reports have disscussed that the spectral signatures of carboxylate impurities gets disappear as the calcination temperature increases indicating the possibility of zinc carboxylate dissociation and conversion to ZnO during the calcination process [57]. Absorption at 400 cm−1 to 600 cm−1 denotes the presence of ZnO NPs as metal oxide vibrational peaks [58]. We found the similar band at 570 cm−1 which could be ascribed to Zn–O stretching vibration of ZnO NPs [59] which can be seen in Fig. 2b and is absent in LB aqueous extract (Fig. 2a). It can, therefore, be supposed that the ZnO NPs is synthesized and stabilized by carboxyl, methyl and hydroxyl groups in the phytoconstituents of LB extract but after calcination, these organic molecules were degraded.

Fig. 2
figure 2

FTIR spectrum of ( a) L. racemosa flower buds extract. (b) Bio synthesized L. racemosa flower buds mediated ZnO NRs

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3.1.3 XRD analysis

XRD is a robust, non-destructive analytical tool used to classify and quantitatively evaluate various crystalline compounds. The XRD spectra of ZnO NPs synthesized with LB extract is illustrated in Fig. 3. X-ray diffraction peaks recorded at 31.75°, 34.52°, 36.23°, 47.53°, 56.58°, 62.84°, 66.24°, 67.92°, 69.04°, 72.65°, and 76.92° are attributed to crystal planes of (100), (002), (101), (102), (110), (103), (200), (112), (201), (004) and (202) of synthesized LB-ZnO NPs, which corresponds to hexagonal wurtzite structural characteristics (JCPDS card number 36-1451, International Diffraction Data Center ver. 2002). Besides, no additional peaks in diffraction could be found in the XRD spectrum of LB-ZnO NPs suggesting complete decomposition of LB extract and zinc acetate precursor, without any other crystal impurities. Our findings are comparable with Narendhran and Shivaraj [60], who have developed ZnO NPs using the Lantana aculeata leaf extract and to Vanathi et al. [61], where ZnO NPs were synthesized using Eichorrnia crassipes leaf extract. The average crystalline size of synthesized ZnO NPs was calculated to be 30.321 nm using the Debey-Scherrer equation as follows [62].

$$D = \frac{k\lambda }{{\beta \cos \theta }}$$
(2)

where D is the crystalline size, K is the shape factor i.e. 0.94, λ is the X-Ray wavelength (1.5406 Å), β is Full Width at half maximum (FWHM) in radians and θ is the Bragg angle.

Fig. 3
figure 3

X-ray diffraction analysis of zinc oxide nanorods biosynthesized using the L. racemosa flower buds extract

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3.1.4 SEM and EDS analysis

The surface morphology and topography of the synthesized particles can be analyzed using SEM. In the present study, the SEM micrograph images (Fig. 4a, b) revealed the formation of zinc oxide nanorods with an average size of 250–300 nm. The resulting morphology of ZnO NRs was quite similar to earlier reported literature [63]. Rods with small morphology is usually preferred for the biomedical field while ZnO rods have better optical characteristics that can lead to significant pharmacological properties [64]. EDX spectrum revealed that there was a strong signal from the Zn atoms along with a low signal from the O atoms represented in Fig. 4c confirming ZnO NRs formation. Also, we found the presence of two small elemental peaks at around 9.5 and 8.5 keV ascribed to elemental zinc. Previous studies have reported major peaks at 1 to 10 keV in ZnO NPs synthesized using Trifolium pratense flower extract [65]. From the figure, we can observe the peaks of zinc and oxygen along with carbon, which may have come because of phytoconstituents from LB aqueous extract which acted as a capping agent to NPs. The figure also displays the major element as zinc which comprises more than 80% of total constituent along with oxygen, which further confirms the formation of pure ZnO NRs from LB extract.

Fig. 4
figure 4

a, b SEM image of ZnO nanorods synthesized from L. racemosa flower buds exact. c EDS analysis of ZnO nanorods biosynthesized using L. racemosa flower buds extract

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3.1.5 TEM analysis

The TEM analysis of the synthesized LB-ZnO NPs is shown in Fig. 5. TEM micrographs of the produced nanoparticles at different magnifications are shown in Fig. 5a–c which clearly shows mixed morphology of ZnO NPs. Synthesized NPs were mostly rod-like structures with the average size in the range of 250–300 nm in length and 45–50 nm in diameter. Comparable results were found using Myristica fragrans leaves extract synthesized ZnO nanorods (ZnO NRs) with a range of 50–100 nm [66]. Another report demonstrated the synthesis of ZnO NRs using Ricinus communis L. leaf extract which had a diameter of 52.84 nm and length of 700 nm [67]. Figure 5e shows the selected area diffraction pattern (SAED) of L. racemosa flower buds fabricated ZnO NRs, corresponding to the hexagonal wurtzite structure of ZnO which correlates well with the XRD results.

Fig. 5
figure 5

a–c represents TEM images of bio-synthesized ZnO NRs using L. racemosa aqueous extract, d SAED pattern of synthesized ZnO NRs

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3.1.6 Surface charge and stability by Zeta potential

The surface charge and stability of the synthesized LB-ZnO NRs was analysed by Zeta potential (ZP). Surface charge plays an important role in confirming the stability of NPs in a liquid medium. The zeta potential value for LB-ZnO NRs was around − 23.5 mV (Fig. 6) indicating a negative surface charge on synthesized NPs. Similar results were obtained by ZnO NRs synthesized using Santalum album leaf extract with zeta potential value of − 13 mV [68]. It should be noted that the particles with zeta potential values more positive than + 30 mV or more negative than − 30 mV are considered to be stable [69]. In contrast, the colloids are least stable at the isoelectric point, where the zeta potential is zero. The high negative value on synthesized LB-ZnO NRs confirms the repulsion among the particles and suggesting the synthesized NRs are stable.

Fig. 6
figure 6

Zeta potential of biosynthesized ZnO NRs in deionized water

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3.2 Mosquito larvicidal activity

Different concentrations of LB extract and synthesized LB-ZnO NRs were tested on the early 4th instar larvae of Ae. aegypti for 24 h to evaluate their efficacy. Results indicate as the concentration of plant extracts and ZnO NRs increased the mortality of mosquito larvae increased. The LB aqueous extract was less toxic than synthesized LB-ZnO NRs against Ae. aegypti. 100% mortality was recorded by LB-ZnO NRs at 50 μg/ml with LC50 and LC90 values of 24.74 and 42.09 μg/ml respectively. While LB extract showed 100% mortality at 2500 μg/ml with LC50 and LC90 values of 1333.75 and 2216.48 μg/ml respectively (Table 1). Control did not show any toxicity to mosquito larvae and χ2 value shows a significant difference at p < 0.05 level. Results indicate the larvicidal potential of LB-ZnO NRs being 50 folds more potent than LB extract. There are no reports on the larvicidal activity of ZnO NRs synthesized from L. racemosa plant but few mangroves have been screened for their larvicidal activity. Jisha and Shreeja [17] showed potential larvicidal activity against Ae. aegypti larvae by L. racemosa leaf acetone extract with an LC50 value of 8 mg/L. Similar results were also recorded by Ali et al. [70] in E. agallocha leaf ethanol extract on Ae. aegypti larvae with LC50 value of 67.1 mg/ml.

Table 1 Larvicidal toxicity of L. racemosa flower buds extract and synthesized ZnO nanorods against the dengue vector Aedes aegypti
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The mortality effect of LB-ZnO NRs on mosquito larvae can be attributed to the internal toxic effects of tiny particles inside the cuticle and the accumulation of LB-ZnO NRs in the alimentary canal. Also, small particles have been reported to target individual cells and interfere with moulting and other physiological processes which hinders the development of mosquito larvae [71]. Similar results were obtained by ZnO NRs synthesized using Myristica fragrans aqueous leaf extract and Ulva lactuca-fabricated ZnO NPs which was highly effective against Ae. aegypti 4th instar larvae [49, 66].

Mosquito larvicidal toxicity was further studied through morphological changes by exposing LB extract and LB-ZnO NRs at their highest concentration and visualizing affected larvae under the Stereo Zoom microscope (Fig. 7). On treatment with both LB extract and LB-ZnO NRs, Ae. aegypti larval body shrank and darkened which gave burnt like appearance. Besides, larvae treated with LB-ZnO NRs showed an accumulation of NRs in the alimentary canal and the respiratory region i.e., siphon. These morphological changes seem to be responsible for the larvicidal toxicity by LB extract and LB-ZnO NRs. Similar morphological damages were reported by Abinaya et al. [72] when Ae. aegypti larvae were treated with bacterial exopolysaccharide (EPS)-coated ZnO nanoparticles.

Fig. 7
figure 7

Morphological changes in Ae. aegypti larvae after 24 h. a Control larvae after 24 h, b when exposed to L. racemosa flower buds extract c L. racemosa flower buds extract synthesized ZnO NRs. Arrow in red color (→) indicates the damages to head, abdominal region, thorax region and siphon

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Overall the results suggest that mortality in larvae might be due to the accumulation of LB-ZnO NRs in the entire alimentary canal and in the siphon region.

3.3 Enzyme activity

3.3.1 Acetylcholinesterase activity

AChE is an enzyme involved in the hydrolysis of acetylcholine and its activity is inhibited by the tested compounds at nerve synapses and neuromuscular junctions. Many insecticide classes such as organophosphates and carbamates are responsible for inhibition of AChE irreversibly and causes insect mortality [73]. AChE enzyme was used as a surrogate biomarker to assess the neurotoxicity in vitro [74]. Given this, we carried out a study involving the effect of LB extract and LB-ZnO NRs on AChE activity of Ae. aegypti 4th instar larvae. The exposure of the larvae to LB extract and LB-ZnO NRs for 24 h significantly decreased the AChE activity to 2.42 μM ACT/mg/min and 1.41 μM ACT/mg/min respectively compared to the control value of 3.58 μM ACT/mg/min of homogenate (Fig. 8). This corresponds to 32% and 60.6% inhibition of AChE activity in larvae treated with LB extract and LB-ZnO NRs. Previous reports of mangroves plants viz. Rhizophora apiculata, Rhizophora annamalayana, Rhizophora mucronata, Avicennia marina and Bruguiera cylindrica showed 50% inhibition of AChE activity in mosquito larvae [75]. Extracts of Sapindus emarginatus showed comparable results to our study by altering the rates of AChE significantly in dengue larvae [76]. For biomonitoring studies and metal toxicity experiments on cholinesterases (ChEs), assessing the effect of metals on the activity of AChE has been a major challenge. Nevertheless, little work has been done to examine the interactions between AChE and metallic nanoparticles [77, 78]. There are no reports on the AChE inhibition in Ae. aegypti larvae by ZnO NRs using mangrove plant extracts. However silver nanoparticles synthesized using Cassia fistula-fruit pulp extract demonstrated lowered AChE activity of 62% in Aedes albopictus larvae [79]. Similarly, inhibition in AChE activity was observed by α-chitin nanoparticles, silver nanoparticles synthesized using shells of Penaeus monodon Fabricius and their nanocomposite when treated against Ae. aegypti larvae [80]. According to previous studies, it was found that toxic compounds affect the digestive system, the respiratory system and the nervous system in the larvae [81]. AChE enzyme inhibition causes the muscles to remain contracted leading to paralysis and finally death [82]. In the present study larvae exposed to LB aqueous extract and LB-ZnO NRs displayed tremors, rigidity and inability to reach the water surface when touched. These symptoms are suggestive of the respiratory muscles paralysis caused by LB-ZnO NRs due to AChE inhibition. This inhibition probably would have compromised the larval ability to breathe freely leading to death. Thus results under present studies indicate the neurotoxic potential of LB-ZnO NRs.

Fig. 8
figure 8

Activity of acetylcholinesterase of Ae. aegypti larvae at LC50 concentration of L. racemosa flower buds aqueous extract and L. racemosa flower synthesized ZnO NRs. Results are presented as mean ± SEM (n = 3). *p < 0.05 (ANNOVA, Tukey’s HSD test) denote significant difference compared to the control

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3.3.2 Glutathione S-transferase activity

GST normally plays an important role in detoxification of exogenous compounds. Compounds such as insecticides enter target insect tissues and organs and affect the activity of different detoxifying enzymes. Some compounds may inhibit the activity of GSTs, while others may enhance the activity of GSTs [83]. In this study, there was a slight increase in GST activity by LB extract (0.416 μM CDNB conjugated/min/mg protein) while significant decrease in GST activity was witnessed in larvae treated with LB-ZnO NRs (0.276 μM CDNB conjugated/min/mg protein) as compared to control (0.37 μM CDNB conjugated/min/mg protein) (Fig. 9). Thus 11% increment in GST activity in Ae. aegypti larvae when treated with LB aqueous extract signifies the larvae were trying to detoxify plant metabolites. In contrast, 25% reduction in GST activity by LB-ZnO NRs indicated that the ZnO NRs might be involved in a redox reaction and causes oxidative stress damage in the larval tissues [84,85,86]. A similar decrease in GST activity was shown by Pongamia pinnata leaf extract coated ZnO nanoparticles in Callosobruchus maculatus [28]. Thus inhibition or stimulation of these enzymes by insecticides can lead to metabolic inequalities, growth retardation and induction of mortality in mosquito larvae [87].

Fig. 9
figure 9

Activity of Glutathione S- transferase of Ae. aegypti larvae treated at LC50 concentration of L. racemosa flower buds aqueous extract and L. racemosa flower synthesized ZnO NRs. Results are presented as mean ± SEM (n = 3). *p < 0.05 (ANNOVA, Tukey’s HSD test) denote significant difference compared to the control

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3.4 DNA fragmentation assay

The purity of DNA was recorded and the A260/A280 ratio was found to be 1.8–2.0 ± 0.05. LB aqueous extract displayed slight but significant damage to the Ae. aegypti larval DNA as compared to LB-ZnO NRs and control-treated larval DNA (Fig. 10). This indicates that synthesized LB-ZnO NRs do not demonstrate any nicking activity in mosquito DNA confirming the mortality of larvae is based solely on cytotoxicity and can be used as a safe alternative to commercial insecticides in the field.

Fig. 10
figure 10

DNA fragmentation assay of Ae. aegypti larvae treated with L. racemosa flower buds aqueous extract (LB) and L. racemosa flower synthesized ZnO NRs (LB-ZnO NRs) as compared to control

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4 Conclusion

This study reports for the first time green synthesis of zinc oxide nanorods using L. racemosa flower buds. The characterization results recorded from UV–Vis spectrophotometry, TEM, SEM, XRD, FTIR and EDX analyses support the effective biosynthesis of L. racemosa flower buds extract fabricated ZnO NRs. The synthesized LB-ZnO NRs are crystalline with an average length of 250–300 nm and 40–50 nm in diameter. Present research highlighted that the ZnO NRs are easy to produce, and can be employed at low dosages to strongly reduce populations of the dengue vector, Ae. aegypti with LC50 at 24.74 μg/ml. Besides, LB-ZnO NRs also affected the activity of enzymes in Ae. aegypti larvae, which demonstrated decreased AChE and GST activity, responsible for signal transduction and resistivity of xenobiotics with no genetic impairment. The study concludes that LB-ZnO NRs can be used as an efficient nanobiopesticides for controlling dengue vector Ae. aegypti larvae in the future.