Introduction

The synthesis of silver nanoparticles from plant extract and microorganisms is an eco-friendly approach [1], in the sense that the end product is water and carbon dioxide which does not smell or pollute the environment but rather add value to it [2]. It is essential to search for original possible approach to synthesize nanoparticles that are useful in medicine, for treatment of wastewater, pharmaceutical industries and public health [3]. In addition, the introduction of nanomaterials with unique properties, particularly metal nanoparticles, offers new expectation to cure and treat diseases [4]. Among the metal nanoparticles, Ag NPs have been widely useful; reducing agents and surfactants are frequently used during the chemical production of Ag NPs, to reduce silver ions and manage the growth of Ag NPs [5]. However, due to the involvement of these poisonous compounds, chemical approach suffer a lot of deficiencies [6]. Therefore, it is important to make more effort to search for environmentally friendly methods for producing Ag NPs. Raman et al. [7] synthesized a facile, eco-friendly, stable in time and non-toxic silver nanoparticles (Ag NPs) using the extract of edible mushroom, Agaricus bisporus. Senthamarai et al. [8] also synthesized and characterized zinc nanoparticles (ZnO NPs) by biophysical method using unripe fruit extract of Aegle marmelos. There is a lot of increasing interests in biosynthetic approach because they avoid toxic chemicals and involve very simple steps [9]. Compared to other chemical methods, biosynthetic methods can synthesize Ag NPs with manageable size and shape without using poisonous substance [10]. Fungi, algae, bacteria, and plants, among others, could synthesize Ag NPs. The plant extract is a better choice among the different bio-tools owing to their abundant components that can act as reducing as well as stabilizing agents [11]. Bacillus subtilis and Escherichia coli and plant extracts of Tulsi leaves (Ocimum tenuiflorum) and Jatropha leaves (Jatropha curcas) were used to biosynthesize Ag NPs. Plant extracts of Tulsi leaves (Ocimum tenuiflorum) and Jatropha leaves (Jatropha curcas) contain numerous polyhydroxy flavonoids with strong antioxidant activity, which can act as reducing agents in the production of Ag NPs. The objective of this study is to develop an easy approach to the biosynthesis of Ag NPs. Ag NPs of different sizes were successfully synthesized using the bacteria broth and a known amount of the plant extract as reducing agents. A series of characterization techniques including ultraviolet–visible (UV–Vis) spectroscopy, X-ray diffraction (XRD) and high-resolution transmission electron microscopy (HRTEM) were used to validate the properties of the biosynthesized Ag NP. This study is meant to develop a cost-effective and environmentally friendly process in the production of Ag NPs.

Materials and methods

Soil sample was collected from Minna modern slaughterhouse in Tayi village, Bosso Local Government Area, Niger state, Nigeria. The soil samples were delivered to the Microbiology Laboratory of the Federal University of Technology, Minna, in a clean polythene bag. The pour plate technique was used for the isolation of bacteria. For 24 h, the plates were kept at 37 °C. On nutrient agar, separate colonies with different morphologies were identified and sub-cultured repeatedly. The identification of the isolates was confirmed using conventional bacteriological and molecular techniques.

Collection of materials

From a farm in Maikunkele, Niger State, Nigeria, the leaves were collected from fresh Tulsi leaves (Ocimum tenuiflorum) and Jatropha leaves (Jatropha curcas).

Preparation of plant extracts

In order to avoid damaging the plant’s thermo labile constituents by direct sunlight, the leaves were cleaned with clean water and then air dried for fifteen (15) days at room temperature. The stems were removed and the leaves were ground into a fine powder. A conical flask with a capacity of 1000 ml was filled with 500 ml distilled water, 25 g of the powdered leaves was weighed, and the mixture was cooked for 25 min. A muslin cloth and then filter paper were used to separate the aqueous extracts (Whatman no.1). Refrigerated silver nanoparticles were formed from the filtrate [12]. The plant extracts were analyzed using the procedures of Trease and Evans [15] and Sofowora [16].

Isolation and identification of bacteria

Soil samples within the premises of Minna modern abattoir, Tayi village, Niger State, Nigeria were collected in sterile polythene bags and transported to the Microbiology laboratory of the Federal University of Technology, Minna, Niger State, Nigeria. The soil sample were diluted in sterile saline solution (0.9% w/v) and isolates were obtained by spread plate technique on nutrient agar medium at 37 °C for 24 h. Morphologically distinct colonies were isolated and repeatedly sub-cultured on nutrient agar. Identity of the isolates was affirmed after characterization by standard bacteriological methods and molecular technique.

Molecular identification of test organisms using molecular sequencing

The bacterial isolates were subjected to DNA isolation and 16S rDNA identification using universal forward 27F (5 prime AGAGTTTGATCMTGGCTCAG 3 prime) and reverse 1429 R(5 prime TACGGYTACCTTGTTACGACTT3 prime) primers. Following the species identification, the strain was outsourced for whole-genome sequencing to Genotypic Technology Ltd., Bengaluru, India. Whole-genome sequencing was done using Illumina MiSeq platform, with a paired end library. A total of 2,546,530 paired end raw reads of 1500 base pairs length on average were sequenced, out of which 2,540,525 high-quality paired end reads, with 90% read length scoring Phred quality score of 35 and above, were assembled into 36 Scaffolds. The assembled genome of Sample A was 99.87% identical to Bacillus subtilis MFN 15, while the assembled genome of Sample B was 99.93% identical to Escherichia coli UBR 11.

Biosynthesis of silver nanoparticles

Silver nanoparticles were synthesized externally as reported by Siddiqi et al. [13]. Nutrient broth was used to sub-culture the isolated colonies, and incubated at 37 °C for 24 h. The culture supernatant was obtained by centrifuging the broth for 10 min at 8000 rpm. In a 500 ml Erlenmeyer flask, around 200 ml of aqueous solution containing 1 mM silver nitrate was mixed with 100 ml of culture supernatant. Shaking at 150 rpm and in a dark environment kept the mixture stable. The solution’s hue changed as the silver nitrate was reduced, allowing the experimenters to track their progress.

Silver nanoparticles were confirmed by adding 5 ml of Ocimum tenuiflorum and Jatropha curcas aqueous extracts to a solution of aqueous 1 mM AgNO3 at 70 °C and pH 12 for 60 min.

Characterization of biosynthesized silver nanoparticles

UV–visible spectrophotometer was used to monitor the color changes in the nanoparticles synthesized nanoparticles [14]. S-3400 N, Hitachi instrument, X-ray diffraction and energy-dispersive X-ray (EDX) spectroscopy were used to determine the crystal structure and elemental composition of silver nanoparticles synthesized in this study. Selective area electron diffraction revealed the lattice structure of the silver nanoparticles, which were synthesized in this study using the S-3400N instrument [15].

Results and discussion

Phytochemical components of plant extracts

Ocimum tenuiflorum and Jatropha curcas extracts were subjected to phytochemical analysis in an effort to identify the major types of secondary metabolites present. Phytochemical secondary metabolite study of leaves from J. curcas and O. tenuiflorum was compared qualitatively and quantitatively in Tables 1 and 2. The presence of tannins and saponins in the leaves of J. gossypifolia and J. multifida was previously reported by Baranwal et al. [16] and was validated in our investigation. Tables 1 and 2 demonstrate that alkaloids, tannins, saponins, flavonoids, and phenols were found in the leaves of the plants studied, although to various degrees of intensity and quality. According to Deshmukh et al. [17], qualitative analysis may help identify bioactive principles and pave the way for the discovery and development of nanoparticles. There were similar findings (Table 1), which demonstrate the relatedness of Jatropha and Ocimum species studied as well as their potential in nanotechnology as reducing agents during the green syntheses (see below). Particularly in J. curcas and Ocimum tenuiflorum, tannins were found in high concentrations during the qualitative screening. Alkaloids were also found in the leaves of J. curcas and Ocimum tenuiflorum, albeit to a lesser extent.

Table 1 Qualitative components of phytochemicals in Ocimum tenuiflorum and Jatropha curcas
Full size table
Table 2 Quantitative components of phytochemicals present in Jatropha curcas and Ocimum tenuiflorum leaves (mg/100 g).
Full size table

Flavonoids, phenols, and saponins were the most common phytochemicals discovered in the leaves of the species, followed by flavonoids. Alkaloids and tannins were the most abundant compounds in the leaves, although their concentrations varied widely. As far as phytochemical concentrations go, flavonoids were found to be the most prevalent in the leaves of the two plant species studied: J. curcas had an 1866.33 mg/100 g concentration while Ocimum tenuiflorum had 780.19 mg/100 g (Table 2). This may be due to the solvent used, as reported in the work of De Matteis et al. [18], who also showed the effect of solvent on phytochemical extraction of silver nanoparticle with more extract via aqueous extraction, for example, flavonoids are popular secondary metabolites that have a variety of biological activities at non-toxic concentrations [19].

Biochemical and morphological characteristics of bacterial isolates

The microscopic and biochemical tests for identification of bacterial isolates from soil are shown in Table 3

Table 3 Biochemical tests for identification of bacteria isolated from soil
Full size table

Molecular characteristics of test organisms

Figure 1 demonstrates the integrity of amplified genes on agarose gel and shows the molecular properties of the bacterial isolates with the highest capacity to produce silver nanoparticles. After electrophoresis, the gel-documented pictures of the bacteria were isolated, DNA showed at 1500 kb, indicating that the isolates were pure.

Fig. 1
figure 1

Agarose gel electrophoresis of Bacillus subtilis and E. coli

Full size image

Keys:

B1 = Bacillus subtilis

B2 = Escherichia coli

Biosynthesis of silver nanoparticles

Bacillus subtilis and Escherichia coli were used in the extracellular production of silver nanoparticles. The emergence of a color shift in the reaction mixture (Figs. 2, 3) originally suggested the production of silver nanoparticles, which was later verified by UV–visible spectroscopy (surface plasmon resonance). During the two days of incubation, the supernatant’s color went from yellow to dark brown, and its intensity increased as the amount of Ag+ in the solution was reduced to AgO. No color change was observed in the control when incubated for the same length of time and conditions. Changes in surface plasmon may be responsible for the color shift from a light yellow to a dark brown [20].

Fig. 2
figure 2

Initial color during bio-reduction of silver nitrate at 0 h

Full size image
Fig. 3
figure 3

Color changes during bio-reduction of silver nitrate after 48 h

Full size image

Jatropha and Ocimum leaf extracts were employed to manufacture silver nanoparticles in the green synthesis. It transformed from yellowish brown to dark brown after 2 days of incubation, when Jatropha and Tulsi leaf extracts were combined separately in the aqueous solution of silver ion complex (Figs. 2, 3). Silver nanoparticles in aqueous plant extract solution have a yellowish brown to dark brown hue because surface plasmon vibrations in silver nanoparticles are excited. Alternatively, the color shift might be due to a reduction of silver ion (Ag+ ions to Ag0), which indicates the creation of silver ions [21]; When compared to high molecular weight proteins from bacterial biomass, flavanone and terpenoid elements of Jatropha and Tulsi leaf extracts may stabilize the synthesis of nanoparticles [21]. Silver ions (Ag+) are mostly reduced and silver nanoparticles are stabilized by polyol and water-soluble heterocyclic components [22].

Key:

A: Bacillus subtilis synthesized silver nanoparticle solution

B: Escherichia coli synthesized silver nanoparticle solution

Key:

A—Jatropha sp. synthesized silver nanoparticle solution

B—Jatropha sp. synthesized silver nanoparticle solution

C—Ocimum sp. synthesized silver nanoparticle solution

D—Ocimum sp. synthesized silver nanoparticle solution

E—Escherichia coli synthesized silver nanoparticle solution

F—Escherichia coli synthesized silver nanoparticle solution

G—Bacillus subtilis synthesized silver nanoparticle solution

H—Bacillus subtilis synthesized silver nanoparticle solution

Characterization of silver nanoparticles

Particle size distribution, transmission electron microscope (TEM), energy-dispersive X-ray (EDX), X-ray diffraction spectroscopy (XRD) and selective area electron diffraction (SAED) were all used to analyze the biosynthesized nanoparticles.

UV–visible spectrophotometer

A UV–visible spectrophotometer is used to easily study the optical characteristics, validate and quantify silver nanoparticles, Sekar et al. [23]. Figure 4 shows the development of Bacillus subtilis and Escherichia coli at absorption peaks of 425 and 398 nm, respectively. The broad nature of the absorbance peak showed the polydispersed nature of the synthesized silver nanoparticles, Sekar et al. [24]. The silver nanoparticle surface plasmon resonance was responsible for the peak generation [25]. Bacillus subtilis and Escherichia coli culture supernatants were used to synthesize silver nanoparticles in a manner that is comparable to that of enterobacteria and Klebsiella pneumoniae [26]. With respect to the wavelength ranges seen in Bacillus subtilis and Escherichia coli, these peaks match those reported by Gong et al. [27].

Fig. 4
figure 4

UV–Vis absorption spectra of silver nanoparticles synthesized using bacteria cultures and plant extracts

Full size image

The leaf extracts were analyzed using UV–Vis spectroscopy in order to determine the nanoparticles’ size and form. Figure 4 depicts the UV–Vis spectra of the reaction medium after 2 days. Jatropha and Tulsi leaves were found to have silver nanoparticles that had an absorption peak at 416 and 400 nm, respectively, while Bacillus and E. coli synthesized silver nanoparticles had absorbance peak at 425 and 420 nm.

Particle size distribution for biosynthesized silver nanoparticles

The average particle size of nanoparticles generated by bacteria growth is 4–4.8 nm (Fig. 5) and 9.402.35 and 31.619.72 nm for O. tenuiflorum and J. curcas, respectively, are comparable with previous results (Table 1; Fig. 6). Modal et al. [28] stated that Shewanella oneidensis MR-1 was used to produce Ag NPs with an average diameter of 4–1.5 nm. The particle size of nanoparticles made from bacteria culture is smaller than those made from plant extract, according to Shanmuganathan et al. [29]. To some extent, the different particle sizes produced by utilizing bacteria culture and plant extracts in the present study may be explained by the use of bacterial culture supernatant.

Fig. 5
figure 5

Particle size for a Bacillus subtilis, b Escherichia coli

Full size image
Fig. 6
figure 6

Particle size for c Jatropha curcas, d Ocimum tenuiflorum

Full size image

High-resolution transmission electron microscopy of the biosynthesized silver nanoparticles

Using elevated transmission electron microscopy (HRTEM), the morphology of produced silver nanoparticles was examined. As shown in Fig. 7a, silver nanoparticles generated using B. subtilis and E. coli had an average size of 11.100.21 and 38.890.42 by HRTEM analysis, but the even distribution of silver nanoparticles in E. coli was different from that in B. subtilis (Fig. 7b). Although J. curcas’ silver nanoparticles, which had an average size of 12.28 0.37 nm in the HRTEM study, had nanobars, nanopyramids, nanorods, and hexagonal silver nanoparticles, those produced by O. tenuiflorum had nanoflowers, which had an average size of 12.99 0.15. A study published in 2019 by Tenase et al. [30] (ethylene glycol monoethyl ether) described the creation of nanoprisms. Silver nanoparticles have been synthesized by Torras and Roig [31], which suggests that the present results are in line with prior findings.

Fig. 7
figure 7

High-resolution transmission electron microscopy images of silver nanoparticles synthesized by a Bacillus subtilis, b Escherichia coli, c Jatropha curcas, d Ocimum tenuiflorum

Full size image

Energy-dispersive X-ray spectroscopy micrograph of biosynthesized silver nanoparticles

As seen in Fig. 8, a chemical signal for metallic silver is clearly present in the AgNPs generated by EDX analysis [32]. This might be because proteins or enzymes in the bacterial biosurfactant or the plant extracts used in the synthesis emit other elements, such as C, O, Na, and N, as seen in Fig. 8’s elemental composition [33]. The EDX spectra of AgNPs produced ex vivo has been studied before [34, 35]. Furthermore, the release of phytobiotics such as proteins, enzymes, carbohydrates, sugars, and amino groups may be responsible for the existence of N in nanoparticles generated from B. subtilis and E. coli cultures.

Fig. 8
figure 8

Energy-dispersive X-ray spectroscopy micrograph of silver nanoparticles synthesized by a Bacillus subtilis, b Escherichia coli, c Jatropha curcas, d Ocimum tenuiflorum

Full size image

Selected area electron diffraction (SAED) images of biosynthesized silver nanoparticles

The AgNPs’ polycrystalline structure is seen even more clearly by the SAED ring pattern (Fig. 9). In the SAED patterns, the rings may be ascribed to the diffraction from planes 111 through 222 of a face-centered cubic spherical structure (FCC).

Fig. 9
figure 9

Selected area electron diffraction images of silver nanoparticles synthesized by a Bacillus subtilis, b Escherichia coli, c Jatropha leaf extract, d Tulsi leaf extract

Full size image

X-ray diffraction spectra of biosynthesized silver nanoparticles

The crystal formation of the generated silver nanoparticles was examined using X-ray diffraction (XRD) analysis (Fig. 10). There were four peaks at 2 matching to 111, 200, 220 and 311 Bragg’s reflections that indicated the crystallization of AgNPs. The crystallite sizes of the AgNPs were calculated using Debye–Scherrer’s equation. Balasubramanian et al. [36] also synthesized nanocrystals with Braggs reflection peaks at 38.42, 44.36, 64.40 and 77.49 °C that conforms to (111), (200), (220) and (311) lattice planes. Nanoparticles synthesized from B. subtilis and E. coli displayed a prominent peak at 31.66°, which is missing in nanoparticles produced using O. tenuiflorum and J. curcas. AgNPs were found to be crystallites based on the XRD spectrum from four separate Bragg’s reflection peaks. According to JCPDS file no. 04-0783, these values are in agreement with the JCPDS reference data. It is possible that biomolecules present in the culture supernatant cap the AgNPs, as seen by the unassigned peaks in the Ag NPs’ XRD spectrum [37]. B. subtilis and E. coli cultures were used to create biosynthesized Ag NPs that were spherical in form with mean crystallite sizes of 11.100.21 and 38.890.42 nm, respectively. Using plant extracts, we were able to produce nanoparticles with a crystalline size of 12.280.37 nm for J. curcas and O. tenuiflorum (see Figs. 5, 6).

Fig. 10
figure 10

X-ray diffraction spectra of silver nanoparticles synthesized by a Bacillus subtilis, b Escherichia coli, c Jatropha leaf extract, d Tulsi leaf extract

Full size image

Conclusion

Bacillus subtilis and Escherichia coli culture supernatant and extracts from Jatropha and Tulsi plants were used in this study to produce silver nanoparticles. Extracellular synthesis of silver nanoparticles in the 4–4.8 nm size range produced spherical particles. Hence, these biocompatible silver nanoparticles can be used to remove pollutants from wastewater, and they are also useful in medicine and pharmaceutical industries. The developed silver nanoparticles are easy to synthesize, contamination free, eco-friendly, consumes less energy and cost effective because of their natural or biological source. The developed silver nanoparticles are preferred over commercial drugs in biomedicine owing to their unique antimicrobial characteristics, adjustable size and shape, enhanced stability of surface bound nucleic acids, high-density surface ligand attachment, transmembrane delivery without harsh transfection agents. All these unique properties of the developed silver nanoparticles make them a promising tool over commercial drugs in medicine and pharmaceutical industries. These findings showed that silver nanoparticles can be produced using isolated Bacillus subtilis and Escherichia coli, as well as plant extracts from Jatropha and Tulsi.