Green synthesis, characterization and antioxidant potential of silver nanoparticles biosynthesized from de-oiled biomass of thermotolerant oleaginous microalgae Acutodesmus dimorphus

Abstract

Metal nanoparticles have received global attention due to their widespread biomedical applications. This study demonstrates a sustainable approach for the biogenic synthesis of silver nanoparticles using lipid extracted residual biomass of microalgae Acutodesmus dimorphus cultivated in dairy wastewater. A. dimorphus is a thermotolerant green microalgae with biofuel production potential. The residual biomass of A. dimorphus left after lipid extraction was used to prepare microalgal water extract which was further used for the synthesis of silver nanoparticles. Characterization of the biosynthesized silver nanoparticles using ultraviolet-visible spectrophotometry, Fourier transform infrared spectroscopy, atomic force microscopy, scanning electron microscopy, transmission electron microscopy and energy-dispersive X-ray spectroscopy confirmed the formation of polydispersed, spherical shaped silver nanoparticles with 2–20 nm size. To our best knowledge, this is the first report on the biosynthesis of nanoparticles using de-oiled biomass of microalgae. Further, the biosynthesized silver nanoparticles exhibited an antioxidant potential which was evaluated using 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) i.e. ABTS and 1,1-diphenyl-2-picrylhydrazyl i.e. DPPH, free radical scavenging assays. As microalgae are widely distributed in diverse habitats, they exhibit wide potential for the green synthesis of metallic nanoparticles. Such integration of phycology and nanotechnology leads to the development of a new interdisciplinary approach, ‘phyconanotechnology’.

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1. Introduction

With increasing interest in waste minimization and the implementation of sustainable processes following all fundamental principles of ‘green’ chemistry, the development of simple, eco-friendly and cost-effective approaches for the preparation of advanced materials is desirable.¹ The production of metal-based nanoparticles by traditional approaches often requires use of organic solvents and toxic reducing agents like sodium borohydride and N,N-dimethyl formamide. With nanoparticles being widely used in consumers' health and industrial products, it is necessary to explore biological and biomimetic “green” approaches for the synthesis of nanoparticles.²

Green synthesis of nanoparticles using environmentally benign materials is an emerging branch of nanotechnology.³ A number of biologically assisted synthetic methods are available for the production of nanoparticles.⁴ Various natural resources like plant extract,⁵ bacteria,⁶ cyanobacteria,⁷ fungi,⁸ enzymes,⁹ algae^10,11 etc., can be used for the biosynthesis of nanoparticles. It offers advantages like energy efficiency, eco-friendliness, lower production cost, compatibility for pharmaceutical and other biomedical applications etc., as it does not require use of organic solvents and toxic chemicals. Further, it also allows production of large quantities of nanoparticles that are free of contamination and have a well defined size and morphology.¹²

Nanoparticles are used in various medical applications including in vitro diagnostics.¹³ With their large surface area, they can simultaneously incorporate therapeutic and imaging agents which can be used in the treatment of cancer, microbial infection and many other diseases.¹⁴ Nanoparticles can also be used in crop protection, agriculture¹⁵ and food packaging.¹⁶ Silver nanoparticles (AgNPs) have a broad spectrum of antimicrobial activity against human and animal pathogens, as it affects the permeability of microbial cell membranes.¹⁷ Therefore, it is widely being used as antimicrobial agents in commercial medical and consumer products.¹⁸ AgNPs have also been found to be active against filariasis and malaria vectors, plasmodial pathogens and cancer cells.^13,18

Compared to the native metal nanoparticles, biosynthesized nanoparticles are highly valuable in therapeutics because of their considerable antioxidant potential.¹⁴ The antioxidant potential refers to the ability of any compound to reduce the damage produced by the destructive oxidative stress originated from an increase in the production of reactive oxygen or nitrogen species. This oxidative stress creates an imbalance between the oxidative and antioxidant systems of the cells, resulting in tissue damage.¹⁹ Various analytical approaches such as determination of total phenolic content, trolox equivalent antioxidant activity, scavenging activity toward stable free radicals, reduction of metal ions etc., can be used for the evaluation of the antioxidant potential of nanoparticles,¹⁹ which helps to determine their suitability for various therapeutic applications.

Microalgae are sunlight-driven cell factories primarily found in aquatic environments. They produce substantial amounts of lipids and carbohydrates which can further be converted into biofuel i.e. biodiesel and bioethanol, respectively. Because of their faster growth rate and higher photosynthetic efficiency, microalgae are considered as an alternative renewable biofuel production feedstock.²⁰ The process of biodiesel production from microalgae generates a large quantity of residual de-oiled biomass rich in protein and carbohydrates. The current worldwide algal biomass production has been projected to be more than 20 000 tons per year. If we assume a conservative figure of 25% extractable oil from microalgal biomass,²⁰ production of one metric ton of biodiesel produces three times the amount of residual de-oiled biomass. For the sustainable and economically viable biodiesel preparation, it is necessary to further utilize this de-oiled biomass for various applications²¹ like feed and fertilizer,²² fermentation to yield bio-methane and bio-ethanol, as a nutrient source for organisms,²³ thermo-chemical conversion into various fuels and chemicals, for biosorption of dyes²⁴ etc.

Many recent studies have shown the potential use of microalgal biomass in the biosynthesis of metal nanoparticles.^25–30 However, all these studies are focussed on the use of fresh microalgal biomass for the preparation of nanoparticles. In the present study, our focus was to use lipid extracted residual biomass of microalgae Acutodesmus dimorphus for the biosynthesis of AgNPs. A. dimorphus is a thermotolerant green microalgae³¹ with biofuel production capabilities.³² The specific objectives of the study are (i) use of lipid extracted de-oiled biomass of A. dimorphus for the synthesis of AgNPs (ii) characterization of the biosynthesized AgNPs and (iii) evaluation of the antioxidant potential of the biosynthesized AgNPs.

2. Experimental details

2.1 Biosynthesis of AgNPs

For the synthesis of AgNPs, lipid extracted de-oiled biomass of A. dimorphus cultivated in dairy wastewater was used. A. dimorphus was cultivated in 1 L Erlenmeyer flasks containing 500 ml of dairy wastewater. The actively growing culture of A. dimorphus was inoculated (10% v/v) into the flasks and incubated at 35 °C under 60 μmol m⁻² s⁻¹ light intensity (cool white fluorescent lights) and 12 : 12 h of light : dark period for 8 days. Flasks were manually shaken thrice a day to avoid adherence of the cells to the surface of the flasks. On 8^th day, the culture was centrifuged at 9000 rpm for 5 min, the supernatant was discarded and pelleted biomass was dried in an oven at 60 °C. The procedure of lipid extraction from the dried microalgal biomass is described in our previous study.³²

After lipid extraction, the residual de-oiled biomass was collected and dried in an oven to remove traces of solvents. Dried de-oiled biomass (500 mg) was mixed with 50 ml distilled water, heated at 100 °C for 5 min and then filtered through Whatman filter paper (Grade 1). The extract was collected and stored at 4 °C for further use. For AgNPs synthesis, 10 ml of this extract was mixed with 90 ml of 1 mM AgNO₃ solution and stirred at room temperature for 24 h. The reduction of Ag⁺ was monitored by visual colour change and UV-Vis spectra. After 24 h, the solution was centrifuged at 16 000 rpm for 25 min at 4 °C, the supernatant was discarded to remove unreacted silver ions and the pellet was re-dispersed in distilled water.

2.2 Characterization of biosynthesized AgNPs

The formation of AgNPs was initially confirmed by visual colour change (colourless to dark brown) and ultraviolet-visible (UV-Vis) spectrophotometry. To study the functional groups responsible for the reduction of silver ions, Fourier transform infrared spectroscopy (FTIR) analysis of the extract and biosynthesized AgNPs was carried out on a Perkin-Elmer FTIR spectrometer instrument (Spectrum GX, USA) using KBr disc in the range 400–4000 cm⁻¹. Atomic force microscopy (AFM) of AgNPs was carried out using an Ntegra Aura atomic force microscope (NT-MDT, Moscow) in semi-contact mode using an NSG 01 silicon probe under ambient conditions. The field emission-scanning electron microscopy (FE-SEM) images of AgNPs were recorded using a JEOL JSM-7100F instrument employing an 18 kV accelerating voltage. The transmission electron microscopy (TEM) images of AgNPs were recorded using a JEOL HR-TEM (JEOL JEM 2100, Japan) instrument operated at an accelerating voltage of 200 kV.

2.3 Antioxidant potential of biosynthesized AgNPs

The antioxidant potential of AgNPs was measured by 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) i.e. ABTS and 1,1-diphenyl-2-picrylhydrazyl i.e. DPPH, free radical scavenging assays and compared with that of butylhydroxytoluene (BHT) as a positive control. For the ABTS assay, ABTS radical cations (ABTS˙⁺) were produced by mixing 7 mM ABTS stock solution with 2.45 mM (final concentration) potassium persulfate and allowing the mixture to stand in the dark at room temperature for 16 h before use.³³ Different concentrations of AgNPs (250 μl) were added to 200 μl ABTS solution. The reaction mixtures were mixed thoroughly, incubated at room temperature for 10 min and the absorbance was recorded at 734 nm. The ABTS solution without any sample was used as a control prepared using the same procedure.

The DPPH assay was performed using slight modification in the method of Muniyappan and Nagarajan.³⁴ DPPH (0.1 mM) was prepared in 100% ethanol and 1 ml of this solution was added to 3 ml of samples having different concentration of the AgNPs. The mixture was shaken, allowed to stand at room temperature for 30 min and the absorbance of stable DPPH was recorded at 517 nm. The DPPH solution without any sample was used as a control prepared using the same procedure. The ABTS and DPPH scavenging activities of AgNPs were calculated using the following formula:

Percentage of free radical scavenging activity = [(A_control − A_sample) per A_control] × 100

From the percentage free radical scavenging activity at different AgNPs concentrations, IC₅₀ values were calculated and compared with those of standard BHT solution.

3. Results and discussion

3.1 Biosynthesis and characterization of AgNPs

In the present study, AgNPs were biosynthesized using de-oiled biomass of A. dimorphus AgNPs were formed by the reduction of Ag⁺ into Ag⁰ with the addition of microalgal extract into 1 mM AgNO₃ solution. The colourless AgNO₃ solution gradually turned brownish indicating the formation of AgNPs (Fig. 1A). The formation of AgNPs was also monitored by UV-Vis absorption spectra (Fig. 1B). Exposure of AgNPs to light leads to polarization of the free conduction electrons with respect to the much heavier ionic core of AgNPs, resulting in electron dipolar oscillation and appearance of a surface plasmon resonance band at 420 nm.³⁵ Absorption peak in the same wavelength range was not observed for the AgNO₃ solution used as a control.

Fig. 1 Confirmation of biosynthesis of AgNPs by (A) visual colour change (B) UV-visible spectra.

FTIR measurements of the microalgal extract and biosynthesized AgNPs (Fig. 2) were carried out to identify the possible biomolecules responsible for the bioreduction and stabilization of AgNPs. The broad intense absorption peak around 3400 cm⁻¹ characterizes the OH stretch. The absorption peak at 1650 cm⁻¹ may be assigned to the carbonyl stretch of amide I and –N–H stretch vibrations of amide II.³⁶ The peaks at 1462 and 1400 cm⁻¹ may be assigned to the C–O–O symmetric stretch from carboxyl groups of the amino acid residues. The absorption peak at 1264 cm⁻¹ may be assigned to the stretching vibrations of C–N aromatic and aliphatic amines.³⁷ The peak at 1166 cm⁻¹ may be due to C–O–C stretching of carbohydrates and polysaccharides present in microalgal biomass. Previous studies have confirmed that carbonyl group of amino acid residues and proteins and amino groups of cysteine residues and sulphated polysaccharides bind with the metals. This binding forms a layer covering the metal nanoparticles which prevents their agglomeration and ensures their stabilization.^11,38 In the present study, the amide linkage and other functional groups may probably play a role in the interaction of biosynthesized AgNPs with the proteins or peptides, thereby stabilizing them. Further, the AgNPs showed characteristic absorption peaks identical to those of the extract suggesting that AgNPs might be coated with proteins. In the similar study, Xie et al.³⁹ reported involvement of hydroxyl and carboxyl groups of proteins in the biosynthesis of AgNPs from the extract of microalgae Chlorella vulgaris.

Fig. 2 FTIR spectra of biosynthesized AgNPs and microalgal water extract.

The AFM images of biosynthesized AgNPs in 2-D and 3-D are shown in Fig. 3A and B, respectively. The size of AgNPs was found to be in the range of 5–20 nm. TEM analysis further provided insight into the size details of the biosynthesized AgNPs (Fig. 4A). The nanoparticles were well separated without any agglomeration. They were polydispersed and predominantly spherical shaped with the size range of 2–15 nm. The observed difference in the size range of nanoparticles between AFM and TEM analysis might be due to difference in their imaging techniques. Unlike TEM analysis, which provide two dimensional image of a sample, AFM allows a three dimensional profiling of the samples allowing measurement of the height of the nanoparticles quantitatively. Jena et al.²⁵ also reported the biosynthesis of spherical-shaped AgNPs with size 2 to 16 nm using fresh extracts (in vitro) and whole cells (in vivo) of the chlorophyte Chlorococcum humicola. Biosynthesis of polydispersed AgNPs, with a size range of 8–20 nm, in a continuously stirred, non-aerated reactor has also been reported using microalgae Chlorella vulgaris.⁴⁰ The FE-SEM analysis of biosynthesized AgNPs (Fig. 4B) showed spherical particles arranged to produce flake like structure. The spectrum of energy-dispersive X-ray spectroscopy through SEM (SEM-EDX) (Fig. 5) showed strongest peak at 3 keV confirming the presence of elemental silver in nanoparticles⁴¹ by 76% weight. This was also confirmed by the elemental mapping of AgNPs (Fig. 4C) showing silver mapped in red colour.

Fig. 3 AFM images of biosynthesized AgNPs in 2-D (A) and 3-D (B).

Fig. 4 TEM image (A), SEM image (B) and elemental mapping image (C) of biosynthesized AgNPs.

Fig. 5 EDX spectra of biosynthesized AgNPs.

3.2 Antioxidant potential of biosynthesized AgNPs

Antioxidant activity refers to the inhibition of the oxidation of molecules by preventing the initiation step of the oxidative chain reactions and formation of non-reactive stable radicals.⁴² The antioxidant potential of biosynthesized nanoparticles depends on the properties of various phytochemicals coated on their surface.¹⁴ Different bioingredients have different antioxidant properties, which help in reducing oxidative stress in cells. As AgNPs were biosynthesized using microalgal extract, determination of their antioxidant potential is essential to confirm whether antioxidant properties of the microalgal extract are still retained by the biosynthesized AgNPs.

To confirm the presence of antioxidant compounds capped on the AgNPs, the ABTS and DPPH colorimetric assays were performed and their potential was compared with that of standard BHT (Table 1). In the ABTS assay, the ABTS radicals, which has a peak absorbance at 734 nm, were preformed by mixing ABTS and potassium persulfate (K₂S₂O₈). When antioxidants (AgNPs) were added, the ABTS radicals, which has a blue-green colour, was reduced to ABTS (no colour). The ABTS activity of biosynthesized AgNPs was found to increase in a dose dependent manner and 79% of scavenging activity was observed at AgNPs concentration of 25 μg ml⁻¹ (Table 1). The ABTS IC₅₀ value of AgNPs was determined to be 14.41 μg ml⁻¹. DPPH is a stable free radical, which has been widely used in phytomedicine for the assessment of scavenging activities of bioactive fractions.⁴³ The DPPH assay is based on the reduction of methanolic DPPH in the presence of a hydrogen-donating antioxidant due to the formation of its non-radical form (DPPH-H). The DPPH scavenging activity has been used by various researchers as a fast and reliable parameter to assess the in vitro antioxidant activity of AgNPs solution.³⁴ Like ABTS assay, the DPPH activity of AgNPs also increased in a dose dependent manner and maximum scavenging activity (59.21%) was observed at a concentration of 10 μg ml⁻¹ (Table 1). The scavenging activity of standard BHT at the same concentration (10 μg ml⁻¹) was found to be 71.12%. The DPPH IC₅₀ of AgNPs and BHT was found to be 6.91 and 4.47 μg ml⁻¹, respectively. Similar to our study, Bhakya et al.⁴⁴ also observed a dose dependent increase in the DPPH scavenging by AgNPs synthesized using root extract of the plant Helicteres isora. Bhaumik et al.¹⁴ synthesized AgNPs using medicinal plant Camellia sinensis (green tea) and determined their ABTS (1 μg ml⁻¹) scavenging activity of 39% with IC₅₀ of 1.3 μg ml⁻¹ and DPPH (5 μg ml⁻¹) scavenging activity of 49% with IC₅₀ of 5 μg ml⁻¹. Mittal et al.⁴² reported 59% and 63% antioxidant activity for DPPH and ABTS (50 μg ml⁻¹), respectively, by AgNPs synthesized using Syzygium cumini fruit extract.

Table 1 Antioxidant properties (ABTS and DPPH assay) of biosynthesized AgNPs

Sample	ABTS		DPPH
	% antioxidant activity^a	IC50 (μg ml^−1)	% antioxidant activity^b	IC50 (μg ml^−1)
a Concentration 5 μg ml^−1.b Concentration 10 μg ml^−1.
AgNPs	26.56 ± 1.23	14.41	59.21 ± 1.12	6.91
BHT (standard)	64.24 ± 3.44	3.76	71.12 ± 0.89	4.47

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From the results of ABTS and DPPH assays, it can be inferred that the antioxidant properties of microalgal extract has been retained by biosynthesized AgNPs possibly due to the capping of the particles. As the AgNPs solution exhibited proton-donating ability, it could serve as a free radical scavenger, possibly acting as a primary antioxidant.

4. Conclusion

Green nanotechnology involves use of energy-efficient eco-friendly approaches for the production of nanoparticles using substances which do not damage the environment. Various types of algae have been explored as a potential tool for the green synthesis of metallic nanoparticles. In the present study, thermotolerant microalgae Acutodesmus dimorphus was grown in dairy wastewater and the lipid extracted residual biomass was then further used for the green synthesis of polydispersed, spherical shaped silver nanoparticles with size range of 2–20 nm. The free radicals scavenging ability of the biosynthesized nanoparticles using ABTS and DPPH assays suggests that intercalation of microalgal extract with silver nanoparticles can serve as promising antiradical agents. However, further research is required to develop a technology in which nanoparticles of specific size and shape could be obtained by the use of specific microalgal strains.

Acknowledgements

CSIR-CSMCRI Registration Number: 090/2016. KC, IP and TG acknowledge CSC-0203 and CSC-0105 for their funding support. RM and CP acknowledge CSIR for their senior research fellowship. KC, RM, CP, TG acknowledge AcSIR for their Ph.D. enrolment. KC also acknowledges his friends Mr Jacky Advani, Mr Nilesh Vadodariya and Mr Praveen Singh Gehlot for their timely help in the sample preparation and characterization of silver nanoparticles. Dr Arvind Kumar, DC, SMC and Dr Parimal Paul, DC, ADCIF, CSIR-CSMCRI are gratefully acknowledged for their support and necessary instrumentation facility.

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