*Article* **Algal-Derived Synthesis of Silver Nanoparticles Using the Unicellular** *ulvophyte* **sp. MBIC10591: Optimisation, Characterisation, and Biological Activities**

**Reham Samir Hamida 1, Mohamed Abdelaal Ali <sup>2</sup> , Mariam Abdulaziz Alkhateeb 3, Haifa Essa Alfassam 3, Maha Abdullah Momenah 3,\* and Mashael Mohammed Bin-Meferij 3,4**


**Abstract:** Algal-mediated synthesis of nanoparticles (NPs) is an eco-friendly alternative for producing NPs with potent physicochemical and biological properties. Microalgae represent an ideal bionanofactory because they contain several biomolecules acting as passivation and stabilising agents during the biogenesis of NPs. Herein, a novel microalgae sp. was isolated, purified, and identified using light and electron microscopy and 18s rRNA sequencing. The chemical components of their watery extract were assessed using GC-MS. Their dried biomass was used to synthesise silver (Ag) NPs with different optimisation parameters. Ag-NPs were physiochemically characterised, and their anticancer and antibacterial effects were examined. The data showed that the isolated strain was 99% similar to the unicellular *ulvophyte* sp. MBIC10591; it was ellipsoidal to spherical and had a large cupshaped spongiomorph chloroplast. The optimum parameters for synthesising Ag-NPs by unicellular *ulvophyte* sp. MBIC10591 (Uv@Ag-NPs) were as follows: mixture of 1 mM of AgNO3 with an equal volume of algal extract, 100 ◦C for 1 h, and pH of 7 under illumination for 24 h. TEM, HRTEM, and SEM revealed that Uv@Ag-NPs are cubic to spherical, with an average nanosize of 12.1 ± 1.2 nm. EDx and mapping analysis showed that the sample had 79% of Ag, while FTIR revealed the existence of several functional groups on the NP surface derivatives from the algal extract. The Uv@Ag-NPs had a hydrodynamic diameter of 178.1 nm and a potential charge of −26.7 mV and showed marked antiproliferative activity against PC3, MDA-MB-231, T47D, and MCF-7, with IC50 values of 27.4, 20.3, 23.8, and 40 μg/mL, respectively, and moderate toxicity against HFs (IC50 of 13.3 μg/mL). Uv@Ag-NPs also showed marked biocidal activity against Gram-negative bacteria. *Escherichia coli* was the most sensitive bacteria to the NPs with an inhibition zone of 18.9 ± 0.03 mm. The current study reports, for the first time, the morphological appearance of the novel *unicellular ulvophyte* sp., MBIC10591, and its chemical composition and potential to synthesise Uv@Ag-NPs with smaller sizes and high stability to act as anti-tumour and microbial agents.

**Keywords:** green synthesis; microalgae; anticancer; antibacterial; optimisation parameter

#### **1. Introduction**

Green synthesis has become a reliable and sustainable method for the biogenesis of several nanomaterials such as metallic nanoparticles (NPs) (M-NPs), metal oxide NPs, bimetallic NPs, and quantum dots [1]. NPs represent potent alternative drugs for several diseases, such as infectious diseases [2], cancers [3], diabetes [4], and wound healing [5], due to their unique features including their smaller size to larger surface area, various shapes,

**Citation:** Hamida, R.S.; Ali, M.A.; Alkhateeb, M.A.; Alfassam, H.E.; Momenah, M.A.; Bin-Meferij, M.M. Algal-Derived Synthesis of Silver Nanoparticles Using the Unicellular *ulvophyte* sp. MBIC10591: Optimisation, Characterisation, and Biological Activities. *Molecules* **2023**, *28*, 279. https://doi.org/10.3390/ molecules28010279

Academic Editors: Nagaraj Basavegowda and Kwang-Hyun Baek

Received: 11 November 2022 Revised: 25 December 2022 Accepted: 26 December 2022 Published: 29 December 2022

**Copyright:** © 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

and sufficient reactivity facilitating their use for drug delivery, sensing, and catalysis, among others [6–8]. Generally, NPs are synthesised by three main methods: physical, chemical, and biological (green) syntheses [9]. Phycosynthesis is a green synthesis (bottomup) route that uses algal cells and their biocomponents to produce NPs with various shapes and sizes [10]. Microalgae are considered model microorganisms for the biogenesis of NPs due to their potential to hyper-accumulate heavy metals and be redesigned to more malleable shapes [11]. Moreover, microalgae contain diverse biomolecules such as lipids, proteins, carbohydrates, vitamins, pigments (such as phycocyanin, chlorophyll, and carotenoids), antioxidants, and others that precipitate during the biogenesis of NPs as reducing and stabilising agents [12]. The *Chlorophyta* phylum includes several species that are sources of several secondary metabolites that act as new drugs in the nutraceutical and pharmaceutical industries [13]. The unicellular *ulvophyte* sp. MBIC10591 is a strain belonging to the *Chlorophyta* phylum. Unfortunately, there are no publications on its morphology and applications. This strain was isolated and deposited by Japanese scientists Suda et al. in GenBank for the first time in 2001 with the accession number AB058370. Several studies used microalgae and cyanobacteria to produce several types of metallic and metallic oxide NPs such as Au- [14], Ag- [15], ZnO- [16], and TiO2-NPs [17]. Hamida et al. synthesised hexagonal Ag-NPs using the novel microalgae strain *Coelastrella aeroterrestrica* BA\_Chlo4 with a smaller diameter of 14.5 ± 0.5 nm [12]. The NPs showed marked activity against different tumour cells, including MCF-7 and MDA, HCT-116, and HepG2 cells, with low toxicity against the normal cells (HFs and Vero). They also demonstrated moderate antioxidant activity and marked biocidal activity against both Gram-positive and -negative bacteria. The biological synthesis method requires the adjustment of physicochemical and biological parameters to obtain M-NPs with controlled sizes, shapes, and dispersity. Several studies have reported that the precursor concentrations, reactant ratios, temperature, pH, reaction time, time of exposure, and illumination conditions are important factors that influence the physicochemical and biological properties [18–20]. It was found that the increase in precursor concentration caused an increase in NP intensity, suggesting polydispersity and agglomeration of NPs at higher concentrations [21]. The change in the temperatures of the NP synthesis process may result in the formation of smaller or larger NPs. It was found that the reduction in NP size at higher temperatures could be attributed to an increase in the nucleation kinetics constant instead of the decreased growth kinetics constant, considering the concentrations of the precursors [22]. Among M-NPs, Ag-NPs have more effectiveness against microbes and cancerous cells. Aziz et al. used *Chlorella pyrenoidosa* as a source of reducing and stabilising agents to fabricate Ag-NPs and found that the resultant biogenic NPs exhibited a marked antibacterial activity against *Klebsiella pneumoniae*, *Aeromonas hydrophila*, *Acenetobacter* sp., and *Staphylococcus aureus* [23]. Ag-NPs are also used in sunscreen lotions, burn treatments, wound dressings, textiles, dental materials, and bone implants [24–26]. The Ag-NP mechanisms inside living cells have been reported to depend on their potential to facilitate oxidative stress by promoting the formation of reactive oxygen species. Moreover, their small sizes, potential charge, and surface chemistry enable their interactions with cellular proteins and DNA, resulting in cellular growth inhibition and death [3,27,28]. The current study revealed, for the first time, the morphology and chemical components of the novel unicellular *ulvophyte* sp. MBIC10591 and its potential for the biogenesis of Ag-NPs under optimum conditions and anticancer and antibacterial activities.

#### **2. Results and Discussions**

#### *2.1. Algal Identification*

#### 2.1.1. Morphological Appearance

Light and inverted light micrographs revealed that the unicellular *ulvophyte* sp. MBIC10591 was spherical. Several unicellular vegetative cells were detected with cupshaped spongiomorph chloroplasts with pyrenoids surrounded by several starch grains. Single cells were the most dominant; however, package cells with parenchyma-like struc-

tures containing more daughter cells were detected. All cells were surrounded by thick cell walls (Figure 1A–D). SEM micrographs showed cells with widely ellipsoidal to spherical shapes with sizes of 13.8 × 12.7 μm. Several irregular ribs existed on algal surfaces, and parenchyma-like structure package cells with more than nine daughter cells were observed (Figure 2A–D). The daughter cells were surrounded by thick cell walls. Unfortunately, no publications have demonstrated the morphological appearance of this isolate. The strain was deposited in 2001 for the first time by Japanese scientists Suda et al. in GenBank with the accession number AB058370. However, the current isolate shared several features with *Desmochloris* sp. in that its cells are distinguished by their spherical to ellipsoidal shapes and cup-shaped spongiomorph chloroplasts [29,30].

**Figure 1.** Light (**A**,**B**) and inverted light (**C**,**D**) microscopy of unicellular *ulvophyte* sp. MBIC10591. Scale bar = 20 μm.

**Figure 2.** SEM micrographs of unicellular *ulvophyte* sp. MBIC10591 showing the morphology of single cells of *ulvophyte* sp. MBIC10591 (**A**,**B**) and package cells with a parenchyma-like structure containing more daughter cells (**C**,**D**). Scale bar = 5 μm (**A**,**C**,**D**) and 2 μm (**B**).

#### 2.1.2. Molecular Identification

The 18s rRNA analysis revealed that the current strains were 99% similar to the unclassified unicellular *ulvophyte* sp. MBIC10591 with a query covering of 89%. The sequence was deposited in GenBank, NCBI, with accession number OP605382. The phylogenetic tree demonstrated that the unicellular *ulvophyte* sp. MBIC10591 may be clustered within *Desmochloris* sp. (Figure 3).

**Figure 3.** Phylogenetic tree of the unicellular *ulvophyte* sp. MBIC10591 (blue frame) inferred from 18S r RNA. Tree was constructed by cluster method using MEGA4 software version 10.2.6. Number at each branch refers to the bootstrap values for % of 1000 replicate trees calculated by neighbour joining statistical method.

#### 2.1.3. GC-MS Analysis

The GC-MS chromatograph demonstrated, for the first time, the volatile organic molecules of the watery unicellular *ulvophyte* sp. MBIC10591 extract with a retention time of 4–44 min. The data showed 34 peaks corresponding to 24 algal bio-compounds. These 24 biomolecules included fatty acids (FA), FA esters, vitamins, alcohols, phenols, hydrocarbons, organosulphur compounds, amino-acid-like compounds, and polysaccharides (Table 1 and Figure 4). It was found that the unicellular *ulvophyte* sp. MBIC10591 was enriched with various molecules that act as antioxidant, antimicrobial, anticancer, and anti-hypercholesterolemic agents such as D-fructose, diethyl mercaptal, pentaacetate, 25,26,27-trinorcholecalcifer-24-al, trisulphide, and di-2-propenyl, among others [30–32]. Based on the GC-MS spectra, the main organic molecules were speculated to be lipids and hydrocarbons that could precipitate while stabilising NPs. However, the existence of alcohols and phenols may indicate that hydroxyl groups have significant roles in the biogenesis of Uv@Ag-NPs. Olasehinde et al. analysed the ethanolic and dichloromethane extracts of *Chlorella sorokiniana* and *Chlorella minutissima* and found that the microalgae were enriched with phenols, sterols, steroids, fatty acids, and terpenes that have modulatory

activities for some mediators of Alzheimer's disease [33]. GC-MS analysis of the aqueous extract of *Coelastrella aeroterrestrica* BA\_Chlo4 showed that the dominant biomolecules of the algal extract were fatty acids and hydrocarbons [12].

**Figure 4.** GC-MS chromatogram of unicellular *ulvophyte* sp. MBIC10591 watery extract.



**Table1.**Chemicalcompositionanalysisoftheunicellular*ulvophyte*sp.MBIC10591usingGC-MS.

**Table 1.** *Cont.*


#### *2.2. Uv@Ag-NPs Synthesis*

#### 2.2.1. Optimisation Parameters of Uv@Ag-NPs Synthesis

To obtain smaller nanoparticles with high stability, various parameters were studied, including precursor concentrations, the ratio between algal extract and precursor, temperature, pH, illumination, and time of incubation (Figure 5A–G).

**Figure 5.** UV–Vis spectroscopy graphs illustrating the influence of (**A**) AgNO3 concentration, (**B**) ratio between algal extract and AgNO3, (**C**) temperature, (**D**) pH, (**E**) illumination conditions, and (**F**) incubation duration and (**G**) Uv@Ag-NPs under optimum conditions.

The data revealed an increment in wavelengths of Uv@Ag-NPs from 1 mM (425 nm) and 2 mM (425.5 nm) to 5 mM (428 nm) at a constant ratio of 1:9 of algal extract to AgNO3, temperature of 25 ◦C, pH of 7, and light illumination for 24 h. On the other hand, with 10 mM of AgNO3, no NPs were produced, suggesting that the higher concentrations above 5 mM significantly slowed the generation of nuclei and growth down. Therefore, it took a longer time to complete the reduction in precursors. The concentration of the NP in their suspension at 1, 2, and 5 mM was low, and the suspension had a faint golden-yellow colour. The data revealed that 1 mM of AgNO3 was the optimum for Uv@Ag-NP synthesis. Khan et al. showed that the intensity of Ag-NPs synthesised from the *Piper betle* leaf extract increased at higher concentrations of 3 and 4 mM with high wavelength values relative to the other lower concentrations of 1 and 2 mM of AgNO3; this suggests polydispersity and agglomeration of Ag-NPs at higher concentrations [21]. Changing the ratio of the algal extract to AgNO3 from 1:9 to 1:1, 1:2, and 1:4 at a constant 1 mM AgNO3 caused a reduction in wavelength from 425 nm at a 1:9 ratio to 422, 422, and 422.5 nm, respectively, suggesting that a higher volume of the precursor may result in an increase in the NP size or promote the agglomeration of NPs [34]. An increase in the temperature during the biofabrication of Uv@Ag-NPs resulted in reductions in the wavelength from 422 nm at 25 ◦C to 420 nm at 40 ◦C 418.5 nm at 80 ◦C and 409.5 nm at 100 ◦C. The NP intensity was higher at both 40 and 80 ◦C; however, their peaks were broader, which indicated the synthesis of larger or agglomerated NPs. These data suggested that the higher temperature

was an important parameter for the biogenesis of Uv@Ag-NPs. This could be attributed to the existence of algal biomolecules that become activated at higher temperatures during Uv@Ag-NPs synthesis or kinetic influence. Liu et al. reported that the reduction in NP sizes at higher temperatures could be attributed to an increase in the nucleation kinetics constant instead of the decreased growth kinetics constant, considering the concentrations of the precursors [22]. UV–Vis spectroscopy showed that acidic pH (5) resulted in a broader SPR peak with a wavelength of 408.5 nm. However, the colour of the NP suspension was transparent, suggesting a slower synthesis reaction with a low yield of UV@Ag-NPs. Moreover, the wavelength of the Uv@Ag-NPs at pH of 7 (the same as the original pH of the reaction without any adjustment) and 9 was 409.5 nm; at higher pH values (8 and 12), the wavelengths shifted from 409.5 nm to 418 and 428 nm, respectively. These data explained that the pH values of 7 and 9 were suitable for producing smaller Uv@Ag-NPs, while increasing the pH to 8 and 12 caused an increase in NP intensity with wide shifting in wavelength indicating the synthesis of larger NPs. These data could be explained by the influence of pH on the dissociation, isolation, interfacial free energy, and the net charge of NPs. For instance, in an acidic medium, the driving force of NP dissolution may be balanced by the repulsive force keeping the dispersion of NPs resulting in smaller NPs. On the contrary, the negative charge hydroxyl ions (OH- ) facilitated the reduction of silver ions to NPs by increasing the ion levels in the medium silver atoms; these tend to diffuse between adjacent adsorption sites on a surface and form bonds with nearest neighbour atoms via Brownian diffusion, resulting in the formation of larger NPs [35]. Traiwatcharanon et al. synthesised Ag-NPs using a *Pistia stratiotes* extract and studied the influence of pH on NP size [35]. They reported that acidic conditions at pH values of 4, 5, and 6 caused blue shifting in the SPR of the Ag-NPs with smaller wavelengths of 330 nm while resulting in red shifting of SPR peaks with a wavelength of 414 nm. They reported that the red shift in the basic medium suggests larger Ag-NPs with higher intensities than those generated under acidic and neutral conditions.

The data showed that the optimum illumination condition for Uv@Ag-NP synthesis was under light (409.5 nm); under dark conditions, their wavelength was 420 nm. Increasing the duration of incubation from 24 h to 72 h under illumination increased the wavelength values from 409.5 to 421.5 nm, respectively, suggesting that the duration of exposure to light influences NPs stability. This could be attributed to the photocatalytic reaction where photons produce energetic electrons that excite SPR and, as a result, reduce Ag+ to Ag-NPs [35–37]. However, high exposure to light irritation may accelerate the agglomeration rate of NPs. Husain et al. synthesised silver nanoparticles using 30 cyanobacteria species under dark and light conditions and found that almost all species were able to generate Ag-NPs only under light conditions [38].

Based on the previous data, the optimum conditions for synthesising Uv@Ag-NPs were 1 mM AgNO3, 1:1 ratio of AgNO3 and algal extract, temperature of 100 ◦C for 1h, pH of 7, light conditions, and incubation duration of 24 h. These conditions resulted in golden brown NP suspension at a wavelength of 409.5 nm. Kusumaningruma et al. reported that the maximum SPR peak of biosynthesised Ag-NPs using *Chlorella pyrenoidosa* was at 410 nm, which confirms the nanostructure of Ag-NPs [39].

#### 2.2.2. Uv@Ag-NPs Characterisations

#### TEM, SEM, EDx, and Mapping Analysis

The TEM, HR-TEM, and SEM micrographs (Figure 6) of the Uv@Ag-NPs showed that the NPs had polyform shapes, including spherical and cubic. These NPs were trapped in an algal matrix that could contain polysaccharides. Smaller spherical Ag-NPs and cubic NPs may represent the seed for generating cubic NPs [40].

**Figure 6.** TEM (**A**,**B**), HR-TEM (**C**,**D**), and SEM (**E**,**F**) micrographs of Uv@Ag-NPs illustrate the uniform distribution of Uv@Ag-NPs and their spherical and cubic shapes. Scale bar = 100 nm (**A**), 50 nm (**B**,**C**), 5 nm (**D**), 200 nm (**E**), and 500 nm (**F**).

The micrographs also demonstrated that Uv@Ag-NPs were uniformly distributed without agglomeration, suggesting that Uv@Ag-NPs have good stability. The frequency distribution analysis of Uv@Ag-NPs using HR-TEM micrographs suggested that Uv@Ag-NPs are small, with a nanosize range of 5–60 nm and an average diameter of 12.1 ± 1.2 nm. Kannan et al. fabricated silver nitrate using the *Chlorophyceae Codium capitatum P.C. Silva* strain and showed that Ag-NPs have a cubic shape with a nanosize range of 3–44 nm and a mean diameter of 30 nm [41].

The elemental compositions of Uv@Ag-NPs and their distribution were determined using the EDx detector. The data showed that the main element distributed in the sample was Ag. A sharper peak was detected at 3 keV, which is a typical absorption signal of Uv@Ag-NPs with a mass percentage of 76.7%. Other elements, including carbon (6.93%), oxygen (1.81%), and chloride (12.18%), were detected while other trace elements emerged, including aluminium (0.3%), copper (1.13%), and zinc (0.91%); they may have emerged from the algal biocompounds surrounding the NPs or they existed in the polysaccharide matrix (Table 2, and Figure 7A,B) [39,42].


**Table 2.** EDx analysis of Uv@Ag-NPs synthesised from the unicellular *ulvophyte* sp. MBIC10591.

**Figure 7.** Map (**A**) and EDx (**B**) analysis of Uv@Ag-NPs synthesised from the unicellular *ulvophyte* sp. MBIC10591.

#### FTIR

The FTIR of the Uv@Ag-NPs contained 13 peaks at 3432.9 [41], 2928.7 [43], 2845.0 [44], 2130.1, 1636.5 [45], 1531.6, 1457.5 [46], 1384.4 [47], 1237.2 [48], 1085.2 [49], 889.8, 795.5 [50], and 554.0 [51] cm−<sup>1</sup> (Figure 8). The IR peaks at 3432.9, 2928.7, and 2845.0 cm−<sup>1</sup> corresponded to strong broad O-H stretching of alcohols or medium N-H stretching of primary amines and strong broad O-H stretching of carboxylics, broad N-H stretching of amine salts, or medium C-H stretching of alkane. However, the peaks at 2130.1, 1636.5, and 1531.6 cm−<sup>1</sup> referred to the strong N=N=N stretching of azides, N=C=N stretching of carbodiimides, or N=C=S stretching of isothiocyanates or weak CΞC of alkynes; medium C=C stretching of alkenes or N-H stretching of amines; and strong N-O stretching of nitrocompounds. The peaks at 1457.5, 1384.4, 1237.2, and 1085.2 cm−<sup>1</sup> were related to the medium C-H bending of alkanes; medium C-H bending of alkanes, O-H bending of alcohols, or strong S=O stretching of sulphates; strong C-O stretching of alkyls or medium C-N stretching of amines; and strong C-O stretching of primary alcohols or aliphatic ethers. The FTIR spectra at 889.8, 795.5, and 554.0 cm−<sup>1</sup> were related to strong or medium C=C bending of alkenes and strong C-I stretching of halocompounds. These data may indicate that the main molecules for capping Uv@Ag-NPs were proteins and/or polysaccharides and/or alcohols, while the stabilising molecules were hydrocarbons and/or fatty acids. These data may be supported by the data of GC-MS analysis, which indicated that the main stabilising agents were fatty acids and hydrocarbons, and that phenol, alcohols, and/or amino-acid-like compounds were the reducing agents. Mahajan et al. extracellularly biofabricated Ag-NPs from silver nitrate using *Chlorella vulgaris* [52]. They analysed the functional group on the Ag-NPs using FTIR and found that the IR peaks of Ag-NPs were at 3435.88, 2092.30, 1637.82, 1559.61, 1414.42, 1037.17, and 618.16 cm−1. This suggested that proteins, polysaccharides, and amides were significant passivating biomolecules for the bioreduction of AgNO3 to Ag-NPs, while long-chain fatty acids were the stabilising agents.

**Figure 8.** FTIR spectra of the Uv@Ag-NPs synthesised using the unicellular *ulvophyte* sp. MBIC10591.

#### DLS and Zeta Potential

The hydrodynamic diameter (HD) average of the Uv@Ag-NPs in an aqueous system was 178.1 nm with a polydispersity index of 0.38, suggesting that Uv@Ag-NPs had a polydisperse standard. The larger NP sizes than the nanosize range 5–60 nm calculated using the HR-TEM micrographs could be attributed to the algal biomaterials in the suspension and surrounding the surface of NPs; they tend to absorb water molecules on the NP surfaces, which increases the HD. The zeta potential (ZP) of the NPs is important for understanding their degree of stability in colloidal systems. NPs with higher negativity or positivity have strong repletion forces to repel each other, which prevents the agglomeration of NPs and stabilises them in a colloidal system [53]. Ardani et al. reported that the ZP value range of ±0–10 mV indicates a highly unstable colloid, while the ranges of ±10–20 mV, ±20–30 mV, and >±30 mV reveal relatively, moderately, and highly stable colloids, respectively [53]. The ZP of the Uv@Ag-NPs was −26.7 mV, indicating colloidal stability. This negative charge surrounding Uv@Ag-NPs could be normalised to those of the algal functional groups, such as hydroxyl and carboxylic groups, which surround the surfaces of NPs. Rathod et al. reported that the ZP of Ag-NPs synthesised from the *Nocardiopsis valliformis strain OT1* was −17.1 mV, suggesting their colloidal stability (Figure 9A,B) [54].

**Figure 9.** DLS (**A**) and zeta potential (**B**) of Uv@Ag-NPs synthesised from the unicellular *ulvophyte* sp. MBIC10591.

#### *2.3. Antiproliferative Effect of Uv@Ag-NPs*

Uv@Ag-NPs significantly reduced the proliferative activity of PC3, MDA-MB-231, T47D and HFs cell lines in a dose-dependent manner. However, MCF-7 cells responded differently to Uv@Ag-NPs. Uv@Ag-NPs drastically inhibited cellular proliferation in a dose-dependent manner from 200 to 50 μg/mL. The cell viability was non-significantly increased at 25 to 6.25 μg/mL of Uv@Ag-NPs. Interestingly, 3.13 μg/mL of Uv@Ag-NPs significantly reduced MCF-7 cell growth by 22%, whereas 1.5 μg/mL of Uv@Ag-NPs demonstrated a non-significant reduction in the malignant cell activity by 12%. This may be explained by the way that drug-responsive malignant cells behave or by the fact that smaller NPs can enter cells at lower concentrations since there are fewer aggregates present.

Similarly, the cell viability % of HFs was significantly decreased by increasing the Uv@Ag-NPs concentration from 12.5 to 200 μg/mL. Beyond 12.5 μg/mL, there was no significant activity of NPs against HFs cells. The moderate toxicity of Uv@Ag-NPs against HFs cell lines could be attributed to the algal functional groups surrounding the NPs, which have antioxidant activity, as reported in the GC-MS analysis section, increasing the NPs' biocompatibility against normal cells. These data suggested that Uv@Ag-NPs may act as potent alternative drugs for traditional therapeutic agents or pharmaceutical applications. The IC50 values of Uv@Ag-NPs against PC3, MDA-MB-231, T47D, MCF-7, and HFs were 27.4, 20.3, 23.8, 40.0, and 13.3 μg/mL (Figure 10). These data revealed that the most sensitive malignant cells to Uv@Ag-NPs were MDA-MB-231, followed by T47D, PC3, and MCF-7 cells. This suggested that Uv@Ag-NPs could be used as antiproliferative agents against prostate and multidrug-resistant breast cancer cell lines. The great antiproliferative activity of Uv@Ag-NPs against MDA-MB-231 cells compared to other cells may be attributed to the cellular metabolic state influencing cellular charge and their interaction with the charged NPs. On the other hand, the IC50 values of Ch@Ag-NPs against PC3, MDA-MB-231, T47D, MCF-7, and HFs were 111.8, 256.9, 657.0, 31.2, and 54.1 μg/mL, while the IC50 values of 5-FU against PC3, MDA-MB-231, T47D, MCF-7, and HFs were 10.6, 442.27, 12.75, 56.48, and 32.4 μg/mL (Figure 11). These data indicated that Uv@Ag-NPs demonstrated marked activity against tested cancer cells relative to other tested drugs, including Ch@Ag-NPs and 5-FU. The marked activity of Uv@Ag-NPs against cancer cells may be attributed to their smaller size, which facilitates penetration of cell boundaries and interactions with biomolecules, including proteins, enzymes, and antioxidants, causing cellular dysfunction and cell death [55]. Moreover, the bio-functional group derivatives from the algal components may play a significant role in enhancing the antiproliferative

effects of Uv@Ag-NPs; they may facilitate the transport of NPs within cells via interactions with cellular receptors. Moreover, the negative charge on Uv@Ag-NPs may influence the therapeutic activity of NPs by increasing the attractive force between NPs and the cellular membrane and the resultant increase in the adsorption of NPs on the cellular surface. This surges the probability of these NPs moving inside cells and interacting with cell membranes [56]. Mohanta et al. synthesised Ag-NPs using *Gracilaria edulis* and found that Ag-NPs (with an average diameter of 62.7 ± 0.25 nm and a spherical shape) caused 50% death of MDA-MB-231 cells at concentrations of 344.27 ± 2.56 μg/mL, suggesting the potent antiproliferative activity of NPs [57]. Ag-NPs (with nanosize of 5–50 nm and spherical shape) synthesised from *Pleurotus djamor var. roseus* exhibited antiproliferative activity against PC3 cells with an IC50 of 10 μg/ mL [58], while Ag-NPs (with spherical shape and size range of 5–25 nm) synthesised from *Anabaena flos-aquae* reduced 50% of T47D cell growth with an IC50 of 5 μg/ mL [59]. Hexagonal Ag-NPs synthesised from the novel *Coelastrella aeroterrestrica* strain BA\_Chlo4 with a diameter of 14.5 ± 0.5 nm showed marked inhibitory activity against MCF-7, MDA-MB-231, HCT-116, HepG2, HFS, and Vero with IC50 values of 26.03, 15.92, 10.08, 5.29, 10.97, and 17.12 μg/mL, respectively [12].

**Figure 10.** *Cont.*

**Figure 10.** Antiproliferative activity (**A**,**B**) of a twofold serial dilution of 200 μg/mL of Uv@Ag-NPs synthesised from the unicellular *ulvophyte* sp. MBIC10591 against four malignant cells, PC3, MDA-MB-231, T47D, and MCF-7, and normal cells, HFs. Data are represented as mean ± SEM. *p*-values were calculated versus untreated cells: \*\*\*\* *p* < 0.0001, \*\* *p* < 0.001, and \* *p* < 0.01.

**Figure 11.** Cell viability of 1000 μg/mL of chemically synthesised Ag-NPs (Ch@Ag-NPs) (**A**) and 5-fluorouracil (5-FU) (**B**) against four malignant cells, PC3, MDA-MB-231, T47D, and MCF-7, and normal cells, HFs. Data are represented as mean ± SEM.

#### *2.4. Biocidal Influence of Uv@Ag-NPs*

The inhibitory effects of Uv@Ag-NPs, Ch@Ag-NPs, AgNO3, the algal extract, and ciprofloxacin against *E. coli*, *K. pneumoniae*, *B. cereus*, and *B. subtilis* were examined using the agar well diffusion method. After excluding ciprofloxacin, the data revealed that Uv@Ag-NPs had the highest biocidal activity against the tested microbes (Figure 12 and Table 3). The Uv@Ag-NPs showed greater activity against Gram-negative than Grampositive bacteria. *E. coli* was the most sensitive microorganism to Uv@Ag-NPs with an IZ of 18.9 ± 0.03 mm, while *B. subtilis* showed the lowest response against Uv@Ag-NPs with an IZ of 15.1 ± 0.04 mm. The positive charge of Ag-NPs plays a significant role in enhancing their antibacterial activity via electro-attractive interactions between the negatively charged NPs and bacterial membranes [60]. Here, the Uv@Ag-NPs showed unexpected results; they had a negative charge on their surface but showed marked activity against Gram-negative bacteria, suggesting that the role of the charge in enhancing the biocidal efficiency of Uv@Ag-NPs can be overlooked. However, the biocidal activity of Uv@Ag-NPs against the tested bacteria can be attributed to their small size and large surface area and the surface chemistry of these NPs facilitating their interactions with cellular membranes and components inhibiting bacterial growth. The biocidal activity of Ag-NPs was highly dependent on the nanosize; the smaller sizes with larger surface areas allowed better contact with the cell membrane [61,62]. The Ch@Ag-NPs (negatively charged NPs), relative to Uv@Ag-NPs and AgNO3, had the lowest inhibition zone; higher values (12.0 ± 0.01 mm) were recorded for *B. subtilis*, while the lower IZ was 10.1 ± 0.03 mm for *K. pneumoniae.* The lower biocidal activity of Ch@Ag-NPs compared to Uv@Ag-NPs could be attributed to the large nanosize of Ch@Ag-NPs trapping the NPs outside the bacterial wall, tackling their entrance into cells and reducing their activity against the bacterial cells. Moreover, the absence of functional groups around Ch@Ag-NPs' surface may also substantially impact the therapeutic action of Ch@Ag-NPs. It was found that the functional groups on the NPs surface mitigate their biological activity and toxicity via their interaction with cellular structures and biomolecular corona [28].

**Table 3.** Inhibition zones of 1 mg/mL of Uv@Ag-NPs, Ch@Ag-NPs, AgNO3, algal extract, and ciprofloxacin for *E. coli*, *K. pneumoniae*, *B. cereus*, and *B. subtilis*.


The highest IZ values of AgNO3 were for both *E. coli* and *B. subtilis* at 15.0 ± 0.13 and 15.0 ± 0.19 mm, respectively, while the lower values were for both *K. pneumoniae* and *B. cereus* with values of 14.2 ± 0.45 and 14.2 ± 0.03 mm, respectively. Intriguingly, 1 mg/mL of Uv@Ag-NPs and AgNO3 resulted in a similar IZD value (about 15.0 mm) against *B. subtilis*. These data suggested that the biocidal activity of Uv@Ag-NPs against *B. subtilis* might be due to the nature of Ag ions rather than the algal functional groups, which might have other roles such as stabilising and charging NPs or directing the NPs to bacterial cells. Moreover, 1 mg/mL of algal extract was not enough to inhibit the bacterial growth with zero IZ against all tested microbes. These data show that Uv@Ag-NPs exhibited marked biocidal activity against the tested microbes compared with silver nitrate and Ch@Ag-NPs, suggesting that the small size with high specific surface area and functional group coating of the Uv@Ag-NPs have a significant influence on their biological activities. Ag-NPs (with a particle size of 4.06 nm) synthesised using *pu-erh tea* leaf extract inhibited the growth of *E. coli*, *K. pneumoniae*, *Salmonella Typhimurium*, and *Salmonella Enteritidis* with IZ values of 15, 10, 20, and 20 mm, respectively [63], while the Ag-NPs (with spherical shape and diameter range of 4.5 to 26 nm) synthesised from *Desertifilum IPPAS B-1220* showed antibacterial activity against *B. cereus* and *B. subtilis* with IZs of 16.33 ± 0.33 and 17.33 ± 0.33 mm, respectively [64].

**Figure 12.** Inhibitory activities of 1 mg/mL of Uv@Ag-NPs, Ch@Ag-NPs, AgNO3, algal extract, and ciprofloxacin against *E. coli*, *K. pneumoniae*, *B. cereus*, and *B. subtilis*. Letters written on well refer to (A) Uv@Ag-NPs, (B) ciprofloxacin, (C) Ch@Ag-NPs, (D) AgNO3, and (E) algal extract.

#### **3. Materials and Methods**

#### *3.1. Materials*

Silver nitrate (AgNO3); chemically synthesised NPs with nanosizes of <100 nm, spherical shapes, and 99.5% purity; and 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) tetrazolium reduction assay (MTT) assay and 5-fluorouracil (5-FU) were purchased from Sigma-Aldrich (St. Louis, MO, USA). The cell culture tools and media were purchased from Gibco (Thermo Fisher Scientific, Waltham, MA, USA). PC3, MDA-MB-231, T47D, MCF-7, and HFs cells were purchased from Nawah Scientific company, Egypt, who obtained the cells from the American Type Culture Collection (ATCC, Manassas, VA, USA), and microbial isolates were obtained from the Department of Microbiology, King Saud University, Riyadh, Saudi Arabia.

#### *3.2. Unicellular ulvophyte sp. MBIC105*

#### 3.2.1. Isolation and Morphological Estimation

The microalgae were isolated from muddy soil in Riyadh, Saudi Arabia, using the serial dilution method reported by Hamida et al. [12]. The microalgae were kept in a sterile BG-11 media-containing flask in an incubator under a fluorescence lamp (2000 ± 200 Lux) with a 12:12 h dark/light cycle at room temperature for 15 days. Inverted, light, and scanning electron microscopes were used to determine the morphological appearance and purity of the microalgae. The sample was washed at least three times with water and ethanol and fixed in 70% ethanol. A small volume of the algal suspension was loaded onto a sterile glass piece fixed on a carbon paste attached to a copper stub. The sample was subsequently coated with a platinum coater (JEC-3000FC, Joel, Tokyo, Japan) for 80 s for scanning electron microscopy (SEM) (JSM-IT500HR, Joel, Japan) at 15 kV.

#### 3.2.2. 18s rRNA Identification

The sample was identified using 18s rRNA identification. The DNA was extracted as described by Hamida et al. [12]. The PCR step with specific primers (forward primer: CCAGCAGCCGCGGTAATTCC; reverse primer: ACTTTCGTTCTTGATTAA) was performed to amplify the extracted DNA for sequencing using an ABI 3730 DNA sequencer (Thermo Fisher Scientific, USA).

#### 3.2.3. Gas Chromatography–Mass Spectrometry (GC-MS) Analysis

The volatile components in the algal aqueous extract were screened using the Trace GC-TSQ mass spectrometer (Thermo Fisher Scientific, Austin, TX, USA) with the direct capillary column TG–5MS (30 m × 0.25 mm × 0.25 μm film thickness). Briefly, 50 mg of algal powder was soaked in 50 mL of boiled distilled water (dist. H2O) and sonicated for 30 min. Subsequently, the sample was allowed to macerate for 24 h, followed by filtration with a syringe filter (0.22 μm). The filtrate was dried in a vacuum oven at 50 ◦C for 48 h. The temperature of the column oven was 50 ◦C initially before it was increased at a rate of 5 ◦C/min to 250 ◦C, maintained for 2 min, increased to 300 ◦C at a rate of 30 ◦C/min, and maintained for 2 more min. The injector and MS temperatures were maintained at 270 and 260 ◦C, respectively. Helium was utilised as a carrier gas at a constant flow rate of 1 mL/min. The solvent delay was 4 min, and diluted samples of 1 μL were injected automatically using an Autosampler AS1300 coupled with GC in the split mode. Electron ionisation mass spectra were collected at 70 eV ionisation voltages over the range of 50–650 *m*/*z* in full scan mode. The ion source temperature was set to 200 ◦C. Components of the algal extract were identified by comparing their mass spectra with those of the WILEY 09 and NIST 14 mass spectral databases [12].

#### 3.2.4. Algal Aqueous Extract Preparation

The microalgae biomass was collected by centrifugation at 4700 rpm for 10 min, washed more than thrice with dist. H2O, and lyophilised for 24 h using LYOTRAP (LTE Scientific, Greenfield, U.K.). The algal watery extract was prepared by dissolving an equal amount of algal powder with dist. H2O and boiling at 80 ◦C for 30 min. Subsequently, the algal extract was spun at 4700 rpm for 10 min, and the supernatant was filtered using Whatman filter paper No. 1. The filtrate was used freshly to synthesise the Ag-NPs (Uv@Ag-NPs) [12].

#### *3.3. Uv@Ag-NPs Synthesis*

3.3.1. Optimisation Parameters for the Biofabrication of Uv@Ag-NPs

To determine the optimum conditions for Uv@Ag-NP biofabrication, various parameters were screened.

#### Precursor Concentrations and Ratios

Uv@Ag-NPs were produced with various concentrations (1, 2, 5, and 10 mM) of silver nitrate at a constant ratio of 1 to 9 (algal extract to silver nitrate) and a temperature of 25 ◦C under illumination for 24 h. Two millilitres of the synthesised Uv@Ag-NPs at each concentration was screened using UV spectroscopy (Shimadzu, Japan). After obtaining the optimum concentration, the effects of various ratios of precursor and algal extracts were determined. Four ratios were tested by mixing algal extract with 1 mM of AgNO3 at ratios of 1:1, 1:2, 1:4, and 1:9, respectively, under the same constant conditions.

#### Temperature and pH

To estimate the influence of temperature and pH on NP biofabrication, 100 mL of 1 mM of AgNO3 was mixed with 100 mL of algal extract and exposed to various temperatures of 25, 40, 60, and 100 ◦C for 1 h under other constant conditions. At the optimum temperature (100 ◦C), the pH values of AgNO3 and the algal extract mixture were adjusted dropwise using 0.1 M hydrochloric acid or sodium hydroxide to 5, 7, 8, 9, and 12 under the same constant conditions for synthesis.

#### Illumination and Incubation Duration

Briefly, 100 mL of 1 mM of AgNO3 was mixed with 100 mL of algal extract at 100 ◦C and a pH of 7. The mixture was incubated once in the dark and once in light (fluorescence lamp with 2000 ± 200 Lux) for 24 h. An aliquot was measured using UV spectroscopy to determine the optimum illumination conditions. The influence of the incubation duration was subsequently estimated by incubating the AgNO3 and algal extract mixture under light conditions for 24, 48, and 72 h.

After obtaining the optimum conditions for biosynthesising the Uv@Ag-NPs, the NPs were synthesised on a large scale (5 L), centrifuged at 12,000 rpm for 15 min, washed at least thrice with dist. H2O, and lyophilised for 8 h. The powder NPs were weighed and collected in sterile Eppendorf for further experiments.

#### *3.4. Characterisation of Uv@Ag-NPs*

#### 3.4.1. UV Spectroscopy

For each optimum parameter, an aliquot (2 mL) of Uv@Ag-NPs was screened using UV spectroscopy for a wavelength range of 200–800 nm and a resolution of 1 nm.

#### 3.4.2. Morphological and Elemental Composition Analysis of Uv@Ag-NPs

The shapes, sizes, elemental compositions, and distributions of the Uv@Ag-NPs were analysed using a high-resolution transmission electron microscope (HR-TEM), TEM, and SEM combined by an energy dispersive X-Ray analysis (EDx) detector. The Uv@Ag-NPs were collected by centrifugation at 12,000 rpm for 15 min, washed at least thrice with dist. H2O and ethanol, and suspended in 1 mL ethanol and sonicated for 15 min. For imaging, 20 μL of the NP suspension was dropped onto the carbon-coated copper grid and air-dried to be examined by TEM (JEM-1400Flash, Joel, Tokyo, Japan) at 120 kV. Similarly, for SEM, 20 μL of the NP suspension was loaded on a sterile glass attached to a copper stub and air-dried. The sample was coated with platinum and examined at 15 kV using SEM. On the other hand, a small amount of powdered Uv@Ag-NPs was loaded onto carbon paste attached to a copper stub and coated with platinum for 80 sec to be analysed with an EDx detector (JSMIT500HR, STD-PC80, Joel, Tokyo, Japan) [12].

#### 3.4.3. Fourier Transform Infrared Spectroscopy (FTIR) and Zeta Sizer

The surface chemistry of the Uv@Ag-NPs powders was detected in a range of 400–4000 cm−<sup>1</sup> using FTIR spectroscopy (Shimadzu, Kyoto, Japan). The potential charges and hydrodynamic diameter of the Uv@Ag-NPs were determined by sonicating the NP suspension (500 μg/mL) for 15 min, diluting it 10-fold, sonicating for 1 to 2 min, and transferring it to Utype tubes at 25 ◦C for measurement using the zeta sizer (Malvern, U.K.).

#### *3.5. Anticancer Activity*

The antiproliferative activities of Uv@Ag-NPs, Ch@Ag-NPs, and 5-FU (as positive controls) were screened against four malignant cell lines, namely PC3, MCF-7, MDA-MB-231, and T47D, and one normal cell line, HFs, using the MTT kit. In brief, a cell density of <sup>5</sup> × <sup>10</sup><sup>4</sup> cells/mL was seeded onto a 96-well plate and incubated in a 5% CO2 incubator for 24 h at 37 ◦C. At 75% confluency, the cells were subjected to serial dilution of Uv@Ag-NPs (200, 100, 50, 25, 12.5, 3.1, and 1.6 μg/mL), while the concentrations of both Ch@Ag-NPs and 5-FU were 1000, 500, 250, 125, 62.5, 31.25, 15.62, 7.81, and 3.90 μg/mL. Uv@Ag-NPs, Ch@Ag-NPs, and 5-FU were suspended in DMEM media, and the NP suspension was sonicated for 15 min. The 5-FU mixture was vortexed for 1 min. All mixtures were filtered using a 0.45 μm syringe filter for direct application to cells. The treated plates were incubated for 24 h in a 5% CO2 incubator at 37 ◦C. After incubation, the media were discarded and replaced with 100 μL/well fresh media, and 10 μL/well of MTT solution (5 mg of MTT powder dissolved in 1 mL of sterile PBS, vortexed until dissolution, and filtered using a syringe filter) was added. The plates were incubated for 4 h, and the media was removed. Subsequently, 100 μL/well of DMSO was applied, and the plates were shacked at 400 rpm for 15 min to dissolve the formazan dye crystal. The plates were read on a Hercules, CA, USA) at 570 nm [3]. Cell viability (%) was estimated according to the following equation:

#### (Abs(treated)/(Abs(control)) × 100

The IC50 (half-maximal growth inhibitory concentration) was calculated using a sigmoidal curve.

#### *3.6. Antibacterial Activity*

*Escherichia coli ATCC8739*, *Klebsiella pneumoniae ATCC13883*, *Bacillus cereus ATCC9634*, and *Bacillus subtilis ATCC6633* were cultured in nutrient broth for up to 18 h at 37 ◦C and maintained through continuous subculturing in broth and on solid media. The inhibitory activities of 1 mg/mL of Uv@Ag-NPs, Ch@Ag-NPs, AgNO3, algal extract, and 5 μg/mL ciprofloxacin were assessed against the tested bacteria using the agar well diffusion method. In brief, 4 mL of the bacterial strain was suspended in 50 mL of nutrient agar media. The mixture was poured into sterilised Petri dishes and dried at 37 ◦C. Four 8 mm wells were created in the agar plates using a cork borer. Subsequently, 100 μL of Uv@Ag-NPs, Ch@Ag-NPs, AgNO3, algal extract, and ciprofloxacin suspensions were poured into the 8 mm wells. The plates were kept in a bacterial incubator at 37 ◦C for 24 h. Ch@Ag-NPs and ciprofloxacin were used as positive controls, while dist. H2O was used as a negative control. The inhibition zone (IZ) was estimated after 24 h using a transparent ruler [65].

#### *3.7. Statistical Analysis*

All experiments were performed in triplicate independently, and the data are presented as mean ± SEM. One-way analysis of variance (ANOVA) was performed to compare differences between untreated and treated groups using graphPrism version 9.3.1 (Graph-Pad Software Inc., San Diego, CA, USA); *p* < 0.05 was considered statistically significant. For characterisation analysis of Uv@Ag-NPs, origin 8 (OriginLab Corporation, Northampton, MA, USA) and ImageJ (National Institutes of Health, Bethesda, MD, USA) were utilised.

#### **4. Conclusions**

These findings provide, for the first time, information about the novel microalgae unicellular *ulvophyte* sp. MBIC10591 and their potential for Ag-NP biogenesis. Herein, we report the morphological appearance of the unicellular *ulvophyte* sp. MBIC10591; the cells appeared spherical with cup-shaped spongiomorph chloroplasts with pyrenoids surrounded by several starch grains. Single cells were dominantly distributed; however, package cells with parenchyma-like structures containing more daughter cells were also found. The unicellular *ulvophyte* sp. MBIC10591 is enriched with various biomolecules, including vitamins, antioxidants, amino-acid-like compounds, organosulphur compounds,

fatty acids, hydrocarbons, polysaccharides, phenol, and alcohols and may be a source of several therapeutic compounds. More investigations are needed to identify several molecules in different organic extracts of the unicellular *ulvophyte* sp. MBIC10591. These biomolecules enable the unicellular *ulvophyte* sp. MBIC10591 to biosynthesise small Ag-NPs. The Uv@Ag-NPs have UV–Vis spectra at 409.5 nm with spherical and cubic shapes. It was found that the optimum conditions for synthesising Uv@Ag-NPs include 1 mM AgNO3, a ratio of 1:1 for AgNO3 and algal extract, temperature of 100 ◦C for 1 h, and pH of 7 under light conditions for 24 h. The nanosize of these NPs was 5–60 nm with an average diameter of 12.1 ± 1.2 nm, while their HD and ZP were 178.1 nm with polydispersity index of 0.38 and −26.7 mV, respectively; these suggest their polydispersity and colloidal stability. Several functional groups were detected on Ag-NP surfaces. Proteins or/and polysaccharides or/and alcohols are responsible for reducing Ag-NPs, while fatty acids or/and hydrocarbons are the stabilising agents responsible for preventing the agglomeration of Ag-NPs. Uv@Ag-NPs exhibited marked anticancer activity against prostate cancer and multidrug resistance breast cancers with low toxicity against HFs. They also demonstrated marked inhibitory activity against Gram-negative bacteria; *E. coli* was the most susceptible to NPs, while *B. subtilis* was the most resistant. These antiproliferative activities and biocidal effects of Uv@Ag-NPs may be attributed to their unique physicochemical characteristics including their small sizes, large areas, shapes, and surface chemistry, which allow them to adsorb on cell surfaces, penetrate membranes and increase the permeability of outside walls or biomolecules such as proteins and enzymes, and interact with cellular organelles and biomolecules causing cellular dysfunction and cell death. Further study of the chemistry of the unicellular *ulvophyte* sp. MBIC10591 is recommended to discover more metabolites that can serve as drugs. Moreover, more optimisation parameters are needed to obtain more uniform shapes of Uv@Ag-NPs and assays to explore their biological activities and mechanisms inside malignant and microbial cells.

**Author Contributions:** Conceptualisation, R.S.H., M.A.A. (Mohamed Abdelaal Ali), and M.M.B.-M.; methodology, R.S.H., M.A.A. (Mohamed Abdelaal Ali), and M.M.B.-M.; software, R.S.H.; validation R.S.H., M.A.A. (Mohamed Abdelaal Ali), and M.M.B.-M.; formal analysis, R.S.H.; investigation, R.S.H., M.A.A. (Mohamed Abdelaal Ali), and M.M.B.-M.; resources, M.A.A. (Mohamed Abdelaal Ali), M.A.M. and M.M.B.-M.; data curation, R.S.H. and M.A.A. (Mohamed Abdelaal Ali); writing original draft preparation, R.S.H.; writing—review and editing, R.S.H.; visualisation, R.S.H. and M.A.A. (Mohamed Abdelaal Ali); supervision, M.M.B.-M.; project administration, M.M.B.-M., H.E.A., M.A.M. and M.A.A. (Mariam Abdulaziz Alkhateeb); funding acquisition, H.E.A., M.A.A. (Mariam Abdulaziz Alkhateeb), and M.A.M. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was funded by the Deanship of Scientific Research at Princess Nourah bint Abdulrahman University, through the Research Groups Program (grant no. RGP-1441-0030).

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Additional data to those presented here are available from the corresponding author upon reasonable request.

**Acknowledgments:** This work was funded by the Deanship of Scientific Research at Princess Nourah bint Abdulrahman University, through the Research Groups Program (grant no. RGP-1441-0030).

**Conflicts of Interest:** The authors declare no conflict of interest.

**Sample Availability:** Samples of the algae or nanoparticles are available from the authors.

#### **References**


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### *Article* **Synthesis, Characterization, and Antibacterial Activity of Mg-Doped CuO Nanoparticles**

**Russul M. Adnan 1, Malak Mezher 2, Alaa M. Abdallah 3, Ramadan Awad 3,4 and Mahmoud I. Khalil 2,5,\***


**Abstract:** This study aims to investigate the effect of magnesium (Mg) doping on the characteristics and antibacterial properties of copper oxide (CuO) nanoparticles (NPs). The Mg-doped CuO NPs were fabricated by the co-precipitation method. NPs were characterized by X-ray Powder Diffraction (XRD), Transmission Electron Microscope (TEM), Energy Dispersive X-ray (EDX) analysis, Fourier Transform Infrared Spectroscopy (FTIR), and Photoluminescence (PL). Broth microdilution, agar-well diffusion, and time-kill assays were employed to assess the antibacterial activity of the NPs. XRD revealed the monoclinic structure of CuO NPs and the successful incorporation of Mg dopant to the Cu1−*x*Mg*x*O NPs. TEM revealed the spherical shape of the CuO NPs. Mg doping affected the morphology of NPs and decreased their agglomeration. EDX patterns confirmed the high purity of the undoped and Mg-doped CuO NPs. FTIR analysis revealed the shifts in the Cu–O bond induced by the Mg dopant. The position, width, and intensity of the PL bands were affected as a result of Mg doping, which is an indication of vacancies. Both undoped and doped CuO NPs exhibited significant antibacterial capacities. NPs inhibited the growth of Gram-positive and Gram-negative bacteria. These results highlight the potential use of Mg-doped CuO NPs as an antibacterial agent.

**Keywords:** CuO nanoparticles; bi-metallic NPs; Mg-dopant; co-precipitation; antibacterial; EDX; photoluminescence

#### **1. Introduction**

Nanoparticles (NPs) revolutionized the industrial world. This revolution is due to their outstanding performance and remarkable optical, electrical, catalytic, and corrosion resistance, in addition to their antibacterial properties [1]. Copper oxide (CuO) has a vital role in multi-functional applications [2]. CuO is an important inorganic p-type semiconductor with a band gap of around 1.2–1.8 eV [3,4]. The most stable phases of copper oxide are cubic cuprous oxide (Cu2O) and monoclinic cupric oxide (CuO) [5]. Its applicability ranges between catalysis, photovoltaics, an electrode for lithium-ion batteries, solar energy conversion, supercapacitors, corrosion inhibition, antimicrobial, and anticancer applications [2,5,6].

CuO NPs serve as a good template for multi-functional applications. Its performance can be enhanced by implementing dopants into the CuO lattice. Since Mg dopants have enhanced the structural and antibacterial properties of CuO NPs, and they have a comparable ionic radius (72 pm) to the ionic radius of Cu2+ ions (73 pm) [7], the Mgdoped CuO NPs may be promising candidates for numerous applications, especially antibacterial applications [8–10].

Previous studies showed the antibacterial activity of doped CuO NPs. Doped NPs synthesized by co-precipitation revealed that the doping elements promote the release of

**Citation:** Adnan, R.M.; Mezher, M.; Abdallah, A.M.; Awad, R.; Khalil, M.I. Synthesis, Characterization, and Antibacterial Activity of Mg-Doped CuO Nanoparticles. *Molecules* **2023**, *28*, 103. https://doi.org/10.3390/ molecules28010103

Academic Editors: Nagaraj Basavegowda and Kwang-Hyun Baek

Received: 17 November 2022 Revised: 12 December 2022 Accepted: 16 December 2022 Published: 23 December 2022

**Copyright:** © 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

Cu2+ from the doped CuO NPs [8–10]. Furthermore, the doped CuO NPs possess better antibacterial activity against Gram-positive bacteria than Gram-negative bacteria, especially against *S. aureus*. The 5% Mg-doped CuO NPs exhibited bactericidal activity at very low concentrations and their bacteriostatic rate reached 99.9% [7]. The high antibacterial activity of doped CuO NPs may be attributed to the inactivation of proteins in the cell wall of bacteria. This activity may be due to the binding of Cu2+ ions to the surface of the bacterial cell. Numerous previous studies showed the bactericidal action of doped CuO NPs, especially against *E. coli* and *S. aureus* [8–10]. This inhibitory action may be dependent on the structure of CuO NPs, as reported previously [8].

Doped NPs were shown to exhibit a better inhibitory effect on bacterial growth than that pure CuO NPs. However, other studies reported that pure NPs exhibited better inhibitory effects. The 5%, 7%, and 10% Mg-doped CuO NPs, at low concentrations, exhibited the same antibacterial activity as that of the pure CuO NPs at higher concentrations [10]. It was reported that the antibacterial activity of CuO NPs is enhanced by Mg2+ doping. The increased release of Cu2+ and Mg2+ ions from doped CuO with the increase of Mg2+ doping content may explain the inhibition of the growth of bacteria [7]. Besides, undoped CuO and Mg-doped CuO NPs showed considerable antimicrobial activity versus several bacterial pathogens, especially *S. aureus*, *P. aeruginosa*, and *E. coli* [10]. Furthermore, the doped NPs possess significant antibacterial activity against many bacterial isolates. Thus, such fabrications may provide a potential alternative to the standard methods of bacterial inhibition.

In addition, the capping of CuO NPs with Ethylenediamine tetra-acetic acid (EDTA) was shown to enhance the antibacterial activity. EDTA, which is a water-soluble polymer, acts by stabilizing the surfaces and modifying the growth (size) during NP synthesis. This enhances the antibacterial action of NPs, due to the ability of EDTA to reduce the size of NPs, which in turn increases the action of NPs against bacteria [11].

Here the impact of undoped and Mg-doped CuO NPs was explored on the inhibition of various bacterial isolates. Previous studies have shown that undoped CuO and Mgdoped CuO NPs showed considerable antimicrobial activity against several bacterial pathogens [10]. In this regard, the objective of this study is to investigate the impact of undoped and Mg-doped CuO NPs capped with EDTA on their structural, morphological, and inhibition capacities against various bacteria isolated from the Lebanese sewage sludge, including Gram-positive bacteria (*S. aureus* and *E. faecium*) and Gram-negative bacteria (*E. coli* and *S. maltophilia*).

#### **2. Results**

#### *2.1. Characterization of NPs*

#### 2.1.1. X-ray Diffraction

The XRD patterns of undoped and Mg-doped CuO NPs (Cu1−*x*Mg*x*O NPs) are shown in Figure 1. The patterns of the undoped CuO NPs show that all the peaks reflect the planes of monoclinic CuO, which are (110), (002), (111), (112), (202), (112), (020), (202), (113), (022), (311), (113), (311), and (004). Given that no other secondary phases, relating to impurities or the Cu2O phase are found. Moreover, when CuO is doped with magnesium, the diffraction peaks of CuO NPs are not altered. To assure this observation, the XRD patterns of all the samples are refined via the MAUD program (Figure 1). The refinements checked the possible formation of MgO as a secondary phase. However, all the samples are well fitted to the pure CuO phase, without any presence of MgO. The obtained patterns for undoped and Mg-doped CuO NPs are similar to the previously reported literature [7,12,13]. Lv et al. [7] synthesized Mg2+, Zn2+, and Ce4+-doped CuO nanoparticles by the hydrothermal method. They obtained a pure phase of CuO in the XRD patterns with a doping concentration of less than 7%, indicating the total incorporation of the dopants into the lattice, without the formation of secondary phases. However, beyond 7%, at 10% doping percentage, MgO, ZnO, and CeO2 phases were formed in the CuO lattice. This further aligns with the present

study, as all the prepared Mg-doped CuO NPs formed pure CuO phase, as the doping percentage ranged between 0.5% (*x* = 0.005) to 2% (*x* = 0.020).

**Figure 1.** Refinements of the XRD patterns of undoped and Mg-doped CuO NPs (Cu1−*x*Mg*x*O NPs).

#### 2.1.2. Transmission Electron Microscope

The morphology and size of the undoped and Mg-doped CuO NPs (Cu1−*x*Mg*x*O NPs) were determined using the Transmission Electron Microscope (TEM) technique. The TEM images demonstrate spherical NPs for the three selected samples, as shown in Figure 2. The Mg doping led to some noticeable changes in the NP morphology. The TEM images showed agglomeration for the undoped CuO NPs (*x* = 0.000), whereas, with the doping of Mg at concentrations of *x* = 0.005 and *x* = 0.020, the agglomeration of the doped CuO NPs is reduced, showing more uniform shapes, as depicted in Figure 2. The average particle sizes of the synthesized samples are determined from the particle size distribution which is extracted from the TEM images, using ImageJ software. This distribution is fitted by a Gaussian function, from which the average particle sizes are determined along with the standard deviation (SD), as shown in Figure 2. The obtained average particle size for undoped CuO NPs is 41.31 ± 1.76 nm. Upon Mg doping with *x* = 0.005, the particle size decreased to 27.14 ± 6.70 nm and increased slightly to 33.78 ± 8.54 nm with *x* = 0.020. These alterations in the average grain sizes with the increase in the concentration of Mg-doping may be attributed to the dissimilarity in Pauling electronegativity that affected the growth rate of Mg-doped CuO nanoparticles. The host Cu ions have a Pauling electronegativity of 1.9, which is higher than that of the doped Mg ions (1.31). This dissimilarity proves the

decrease in the growth rate at low concentrations of Mg-doped CuO NPs [13]. However, at higher concentrations, the Mg-doped ions may incorporate into the lattice, not only filling substitutional sites but also occupying interstitial sites, that yield larger grains, as seen in the sample with *x* = 0.020.

**Figure 2.** TEM images with the grain size distribution of the undoped and Mg-doped CuO NPs (Cu1−*x*Mg*x*O NPs).

#### 2.1.3. Scanning Electron Microscope and Energy Dispersive X-ray

The morphology of the nanoparticles is further studied by the scanning electron microscope, as shown in Figure 3. The SEM micrographs assure the nanocrystalline nature of the undoped and Mg-doped CuO NPs, without agglomeration. It is also noticed that the average grain size decreased with Mg-doping, re-assuring the TEM analysis. The average grain size, extracted from the SEM images, is found to be around 25.7 nm for undoped CuO NPs and 22.5 nm for Mg-doped CuO NPs. The chemical composition was studied with an energy-dispersive X-ray (EDX) technique. The presence of copper (Cu) and oxygen (O) elements in the undoped CuO NPs is confirmed by the EDX pattern, shown in Figure 3 (*x* = 0.000). No traces of precursors were detected. The EDX pattern of Mg-doped CuO NPs with *x* = 0.020 is exhibited in Figure 3. In addition to Cu and O, Mg is detected in the pattern, confirming the presence of the Mg-doped in the CuO NPs. The average atomic percent (at%) of copper and oxygen were microstructures of three different regions of the samples and demonstrated in the insets of (Figure 3) as pie charts. In undoped CuO NPs, the ratio between Cu and O is 0.82 while it is equal to 0.97 in Mg-doped CuO NPs (*x* = 0.020). This indicates that the stoichiometric nature of the samples is affected by Mg dopants. These variations may be due to surface crystalline defects [14]. Moreover, the ratio of the atomic percentage of Mg/Cu for *x* = 0.020 samples is calculated to be 0.0255, further confirming the successful synthesis of Mg-doped CuO with matching experimental and theoretical values.

**Figure 3.** SEM and EDX patterns for undoped, and Mg-doped CuO NPs (Cu1−*x*Mg*x*O NPs).

#### 2.1.4. Fourier Transform Infrared (FTIR)

The FTIR spectra, represented in Figure 4, of the undoped and Mg-doped CuO NPs, demonstrated different vibrational bands. A broad absorption band ranged between 3200 and 3600 cm−1. The adsorbed water is present in all spectra. A small peak is detected at 2344–2354 c, and a peak centered around 1618–1656 cm−<sup>1</sup> is observed. The main peaks, ranging between 700–400 cm<sup>−</sup>1, are displayed in Figure 4. Three peaks are detected, which are centered around 480–486, 521–530, and 580–584 cm<sup>−</sup>1.

It is noticed that the peak position is slightly affected by Mg concentration in the CuO lattice. The peak attributed to the symmetric vibration of Cu–O fluctuated around <sup>483</sup> ± 3 cm−<sup>1</sup> with Mg-doping concentrations. However, Cu–O asymmetric stretching and wagging peaks of the pure CuO Nps are shifted monotonously to lower wavenumber with the Mg-doping from 530 and 584 cm−<sup>1</sup> to 521 and 580 cm−<sup>1</sup> with *x* = 0.020. Pramothkumar et al. [15] reported the same pattern of variation upon Mn, Co, and Ni doping to CuO NPs, and explained these shifts according to the dopant effect, which can in turn affect the surface area and defects in the samples. Singh et al. [16] reported the successful doping of Zn to CuO, which led to the increase in Cu–O bond length in the samples with the increase of Zn dopant concentration.

#### 2.1.5. Photoluminescence (PL)

Figure 5a shows the room temperature photoluminescence (PL) emission spectra for Cu1−*x*Mg*x*O NPs with an excitation wavelength of 200 nm. A prominent UV peak appeared at 310 ± 1 nm in all samples, with the highest intensity, as compared to other peaks. Furthermore, the visible emissions in the PL spectra are deconvoluted by four Voigt functions to elucidate the origin of these emissions. The position of the fitted peaks is

listed in Table 1, along with the position of the UV peak. It is noticed that the increase in the concentration of the Mg doping in CuO NPs did not affect the position of the peaks, however, it affects their intensity. This is similar to the reported literature, where the doping concentration does not affect the position of the peaks in the visible part of the PL spectra [17–20]. The deconvolution of the PL spectra of Cu1−*x*Mg*x*O NPs yielded violet (391 ± 1 nm), blue (452 ± 5.5 nm), green (536 nm), and orange-red (628 ± 5 nm) emission peaks.

**Figure 4.** FTIR spectra of Cu1−*x*Mg*x*O NPs, and enlarged view for Cu-O vibrations.

*2.2. Antibacterial Activity of the Undoped and Mg-Doped CuO NPs* 2.2.1. MIC and MBC

The four bacterial isolates were tested for their susceptibility against the undoped and Mg-doped CuO NPs. Undoped CuO NPs had a bactericidal effect on Gram-positive bacteria while having a bacteriostatic effect on Gram-negative bacteria. The most sensitive bacterium was *E. faecium* (MIC = 0.375 mg/mL and MBC = 0.75 mg/mL), followed by *S. aureus* (MIC = 1.5 mg/mL and MBC = 3 mg/mL). *E. coli* and *S. maltophilia* were sensitive at the highest NP concentration used (3 mg/mL). Similarly, Mg-doped NPs exhibited bactericidal activity on Gram-positive bacteria and bacteriostatic activity on Gram-negative bacteria. *E. faecium* was the most sensitive bacterium (MIC = 1.5 mg/mL and MBC = 3 mg/mL) and *E. coli* was the most resistant bacterium. Collectively, undoped and Mg-doped NPs had a better effect on Gram-positive bacteria than that on Gram-negative bacteria. The MIC and MBC results are shown in Table 2 and Figures A1–A3 (Appendix A).

**Figure 5.** (**a**) Photoluminescence spectra of Cu1−*x*Mg*x*O NPs, and (**b**–**f**) their deconvolution for undoped and Mg-doped CuO NPs.


**Table 1.** Position of peaks by deconvolution of PL spectra for Cu1−*x*Mg*x*O NPs.

**Table 2.** MIC and MBC of the undoped and Mg-doped (Cu1−*x*Mg*x*O) NPs against four bacterial isolates.


nd: not determined, MIC: minimum inhibitory concentration, MBC: minimum bactericidal concentration.

It was reported that when the MBC/MIC ratio < 4, this will reflect a bactericidal effect [21]. The antibacterial results against the four bacterial isolates shown in Table 2 revealed that the undoped and Mg-doped NPs exhibited bactericidal effects against *S. aureus* and *E. faecium* with MBC/MIC ratio = 2 and a bacteriostatic effect against *E. coli* and *S. maltophilia*.

#### 2.2.2. Agar Well Diffusion

All isolates showed sensitivity to the undoped CuO NP (*x* = 0.000). *E. faecium* was the most sensitive. It showed a sensitivity against the lowest NP concentration (0.1875 mg/mL). The other three bacteria showed a sensitivity against an NP concentration of 1.5 mg/mL. Sensitivity was considered positive for ZOI diameters > 7 mm [17]. All the bacterial isolates were sensitive to the Mg-doped CuO NP with *x* = 0.005. *E. coli* and *S. maltophilia* showed a sensitivity to the NP at a concentration of 0.75 mg/mL. *E. faecium* and *S. aureus* showed sensitivity against a concentration of 1.5 mg/mL. All investigated bacteria showed sensitivity to Mg-doped NPs with *x* = 0.010. *S. aureus* showed susceptibility against a concentration of 0.75 mg/mL. *E. faecium* showed a sensitivity against a concentration of 1.5 mg/mL. *S. aureus*, *E. coli*, and *S. maltophilia* exhibited a sensitivity at the highest concentration (3 mg/mL). All isolates were sensitive to the Mg-doped CuO NP with *x* = 0.015. *E. faecium* was the most sensitive. It showed a sensitivity starting from the lowest concentration (0.1875 mg/mL). *S. aureus*, *E. coli*, and *S. maltophilia* were sensitive at concentrations starting from 0.375 mg/mL. All isolates were sensitive at the highest NP concentration (3 mg/mL). For Mg-doped CuO NP with *x* = 0.020, all bacteria exhibited sensitivity only at the highest NP concentration (3 mg/mL). The agar well diffusion results of the undoped and Mg-doped CuO NPs are shown in Table 3 and Figure A4 (Appendix A). All results were significant with a *p*-value < 0.05 shown in Table A1 (Appendix A).


**Table 3.** Agar well diffusion of undoped and Mg-doped CuO NPs against four bacterial isolates.


**Table 3.** *Cont.*

Dox: Doxycycline, Amo: Amoxicillin, ZOI: zone of inhibition, SEM: standard error of the mean.

#### 2.2.3. Time-Kill Results

The time-kill test was performed using the MICs of the undoped and Mg-doped CuO NPs against four bacterial isolates to detect the time needed for each NP to exert its antibacterial effect. All bacterial isolates were sensitive to all tested NPs after 2 h of incubation. The activities were sustained till 24 h of incubation. The time-kill results for the different bacterial isolates are shown in Figure 6.

**Figure 6.** Time-kill results of the undoped and Mg-doped NPs against four bacterial isolates.

#### **3. Discussion**

The peaks of the undoped CuO NPs shown by the XRD correspond to the primary defining peaks of the monoclinic CuO phase with a space group of C2/c [22]. The absence of a secondary phase suggests that the samples exhibit a highly single phase [23]. The refinements indicate the total incorporation of Mg dopant in the Cu1−*x*Mg*x*O NPs.

The TEM results demonstrated the change in the morphology and size of the NPs after doping, indicating that the increase in the doping concentration increases the uniformity of the NPs and in turn decreases their size.

The composition of the NPs was detected by EDX. The absence of precursors indicates the purity of the formed CuO NPs. Noting that the emergence of carbon may be due to the use of carbon tape in the measurements, or some residues from EDTA [24]. In addition, the ratio between Cu and O in the undoped and Mg-doped NPs indicates that the stoichiometric nature of the samples is affected by Mg dopants. These variations may be due to surface crystalline defects [14].

Infrared spectroscopy can be used as a fingerprint to identify different molecules by comparing vibration bands. The broad absorption band observed by the FTIR is associated with the hydroxyl (O–H) stretching vibration mode of the water molecule [25]. The adsorbed water observed is due to the physical adsorption of water from the atmosphere [26]. The small peak observed at 2344–2354 cm−<sup>1</sup> is related to the vibration of CO2 in the air [26] and the peak centered around 1618–1656 cm−<sup>1</sup> is due to H–O–H bending vibrations of water molecules [25]. The three main peaks are attributed to Cu–O symmetric stretching, Cu–O asymmetric stretching, and Cu–O wagging, respectively. These peaks validate the successful formation of CuO NPs [5]. Pramothkumar et al. [15] reported the same pattern of variation observed upon Mn, Co, and Ni doping to CuO NPs, and explained these shifts according to the dopant effect, which can in turn affect the surface area and defects in the samples. Singh et al. [16] reported the successful doping of Zn to CuO, which led to the increase in Cu–O bond length in the samples with the increase of Zn dopant concentration.

The PL spectra detect the imperfections and defects within the samples, where the prevalence of the imperfections and surface states varies depending on the synthesizing circumstances, particle size and shape, types of dopants, and concentrations [27,28]. The origin of the UV peak is directly related to the recombination of electron–hole pair, near the band gap transition [17,20]. It is noticed that the position of the UV peak is slightly invariant with the doping concentration, however, its intensity increased with Mg-doping. This enhancement of the intensity may be related to the passivation of surface defects that generate radiative recombination [29]. Additionally, the intensity of the UV peak is affected by the electron density and the variation of the morphology and size of the nanoparticles, with the increase of the doping concentration [30]. The visible emissions are highly sensitive to the change in the synthesis conditions, accounting for the type of dopant and its concentration, the size of the nanoparticle, and its morphology [18]. The size of Mg-doped CuO NPs decreased with the increase of the doping concentration, as noted from TEM and SEM analysis. Hence, the large surface-to-volume ratio stimulates more surface-defect states, as vacancies and interstitials, creating trap levels that radiate visible emissions [18]. Mainly, the intensity of the visible emissions is quenched with the increment of the doping concentration in Cu1−*x*Mg*x*O NPs, as can be noticed from the inset of Figure 5a. This decrement in the intensity may be due to the trapping of the photoexcited electron from the conduction band of CuO NPs by the formed deep-level centers from Mg doping [17,30]. The violet and blue emissions are mostly attributed to deep-level defects, indicating the existence of Cu vacancies in the lattice [17]. The green emission was reported to originate from the recombination of single ionized electrons with a photogenerated hole in the valence band, noting the presence of singly ionized oxygen vacancies or dangling bonds of copper [19]. The orange–red emission ascends from the recombination of an electron bound to donor and free holes [18].

The reported antibacterial properties of NPs, especially CuO NPs, made their usage efficient against bacteria [1,2]. In this regard, CuO NPs were used against four bacterial pathogens isolated from sewage sludge. All bacterial isolates, except *S. maltophilia*, are frequently present in Lebanon, especially in the feces of animals. *S. maltophilia* is a rare Gram-negative bacterium in Lebanon [21,31,32]. In this investigation, the antibiotic Dox was used as a reference antibiotic. It belongs to the tetracycline family of antibiotics. It acts by inhibiting protein synthesis by binding to the 30S ribosomal subunit, leading to the destruction of the bacterial cells [33–35]. This study showed that the undoped and Mg-doped CuO NPs exerted antibacterial activities against all bacterial isolates. Using the agar well diffusion assay, the NPs had better effects on Gram-positive bacteria than on Gram-negative bacteria. The results are consistent with previous studies that showed that Gram-negative bacteria are more resistant to NPs, due to the rigidity of their cell wall [36–38]. This activity depends on the metal oxides present in the NPs. The latter could penetrate the cell wall of bacteria, leading to cell autolysis [10]. Among the investigated NPs, the undoped and Mg-doped CuO NPs with *x* = 0.005 and *x* = 0.010 were efficient as antibacterial agents. They were effective at low NPs concentrations against all bacterial

isolates. In contrast, the Mg-doped CuO NPs with *x* = 0.015 and *x* = 0.020 had lower antibacterial activity against the investigated bacterial isolates. They were effective at higher concentrations. Regarding bacterial susceptibility, *S. aureus* and *E. faecium* were the most sensitive. Their growth was inhibited by all tested NPs at significantly low concentrations. These results are consistent with previous studies that have shown the sensitivity of Gram-positive bacteria, especially *S. aureus*, to NPs [8,9,39]. On the other hand, *E. coli* and *S. maltophilia* were more resistant. The inhibition of their growth required higher concentrations of the NPs. This could be attributed to the shape of NPs. Their spherical shape, demonstrated by the TEM results reflects their significant inhibitory activity against Gram-positive bacteria [8,40]. Spherical NPs are shown to have good antibacterial activity due to the sphere prisms that can penetrate easily into the bacterial cell membrane [41].

The MIC results confirmed the results of the agar well diffusion assay. All the investigated NPs, except the Mg-doped CuO NPs with *x* = 0.020, had bactericidal activities. The Mg-doped CuO NP with *x* = 0.020 had bacteriostatic activity only. The bacteriostatic and bactericidal effects of NPs depend on their metal oxide content and their morphology, and the architecture of the bacteria [42,43]. This means that the metal oxides of the tested NPs can react with the bacterial cell wall through special mechanisms, which are still not very specific. However, previous studies reported the following mechanisms: disruption of the bacterial cell wall, generating reactive oxygen species, and binding with specific cytoplasmic targets and production of metabolites, leading to these bacteriostatic and bactericidal effects [42,44]. Kumer et al. showed that ZnO NPs exhibited good antibacterial activity against Gram-positive bacteria. This activity was better than that of Ag-doped ZnO NPs [45]. In addition, Prakash et al. reported that TiO2 NPs with doped antibacterial activity were better than the undoped TiO2 NPs [46]. The bactericidal properties of the NPs depend on the shape, the surface area of the particle, the type of metal ions, and the chemically reactive functional groups [8,10,35]. The high bactericidal effect is attributed to the different shapes (spherical in our case) of the particles, which help them penetrate the bacterial cell membrane. Moreover, the large surface area permits the production of reactive oxygen. This induces oxidative stress on bacteria, which interrupts the electron flow in the inner membrane, thus causing cell damage [35,36]. On the other hand, the observed bacteriostatic effect could be due to the low number of metal ions coming from the metal oxides, which prevents the Cu ions from interacting with the bacterial cell wall. So, the main bactericidal mechanism may rely on the damage of the cell membrane by the metal oxides [35,37].

Time-kill results have shown that the most frequent inhibition time of the tested NPs started at 2 h of incubation. NPs can prevent the adaptation and duplication of bacteria [47]. This effect could be attributed to the limiting effect of NPs on the nutrient uptake by the bacteria, which eventually will lead to cell lysis. This is consistent with previous studies that showed that NPs affect the metabolic activities and division of bacterial cells. Metal oxides may lead to nutrient deprivation [8,10,36]. In addition, the size of the particles and the surface area may specify the time needed for the interaction between the NP and the bacterial cell wall. When the size of the particle is smaller, the interaction becomes faster, thus decreasing the time needed for the inhibition of bacterial growth [8,10,36]. This slows metabolic processes and leads to cell death.

Collectively, previous studies reported that the size of the NPs reflects their antibacterial effect [48]. In addition, the variation in the antibacterial activity depends on the morphology of the NPs. Furthermore, the variation in the intensity of PL accompanied by the variation in oxygen interferes with the antibacterial activity. So, the shape and the morphology of NPs play a vital role in the inhibition of bacterial growth.

#### **4. Materials and Methods**

#### *4.1. Synthesis of NPs*

The undoped and Mg-doped CuO NPs were prepared by the co-precipitation method, with the chemical formula of Cu1−*x*Mg*x*O (*x* = 0.000, 0.005, 0.010, 0.015, and 0.020). The

CuO NPs were synthesized using copper (II) chloride dehydrate (Merk), magnesium chloride hexahydrate (Sigma-Aldrich, ≥99.0%), and ethylenediamine tetra-acetic acid (EDTA) (0.1 M). The weighed reagents were prepared with a molarity of 1 M in de-ionized water and stirred for 15 minutes. The solution was then titrated with sodium hydroxide NaOH (2 M). NaOH was added slowly under vigorous stirring until pH reached 12. After that, the precipitate was heated at 60 ◦C for 2 h, then sonicated for 10 minutes. The black precipitate obtained was washed with de-ionized water several times until pH reached 7. Finally, the washed precipitate was dried at 100 ◦C for 16 h and ground into fine powders. The powders were sintered at 600 ◦C for 4 h.

#### *4.2. Characterization of Mg-Doped CuO NPs*

The structural properties of both undoped and doped CuO NPs were studied by XRD using the X-ray Bruker D8 Focus power diffractometer with Cu Kα radiation, operated at 40 kV and 40 mA, in the range 20 ≤ 2θ◦ ≤ 80. Material Analysis Using Diffraction (MAUD) software was then used to check for the presence of CuO and MgO phases in the resultant NPs using the CIF files downloaded from the Crystallography Open Database (COD). The morphology of the prepared CuO NPs was investigated using the JEM 100 CX Transmission Electron microscope (TEM), operated at 80 kV. The main functional groups of the synthesized samples were detected using the Nicolet iS5 Fourier Transform Infra-Red (FTIR) spectra after preparing potassium bromide (KBr) pellets mixed with the undoped and doped CuO NPs (1:100). The purity of the Cu1−*x*Mg*x*O NPs was studied using energy dispersive X-ray (EDX), operated at a voltage of 20 kV with laser power of 5 mW and magnification objective of 50x. The Photoluminescence (PL) spectra were studied by a Jasco FP-8600 spectrofluorometer with Xenon (Xe) laser at 200 nm excitation wavelength for Cu1−*x*Mg*x*O nanoparticles, dispersed in ethanol.

#### *4.3. Isolation of Bacteria*

Briefly, *E. faecium, S. aureus, E. coli, and S. maltophilia* were isolated from wastewater by streaking 100 μL of the samples on different selective media (blood agar, chocolate agar, MacConkey agar, mannitol salt agar (MSA), eosin methylene blue (EMB) agar, and cetrimide agar). The plates were incubated at 37 ◦C for 24 h. After isolation, bacteria were Gram stained to differentiate between Gram-positive and Gram-negative bacteria. Bacteria were further identified by VITEK assay.

#### *4.4. Broth Microdilution Assay: Minimum Inhibitory Concentration (MIC) and Minimum Bactericidal Concentration (MBC)*

The MICs of the undoped and Mg-doped CuO NPs were determined against four bacteria employing the microwell dilution method. The test was performed in sterile 96-well microplates by dispensing into each well 90 μL of nutrient broth and 10 μL of bacterial suspensions adjusted to 0.5 McFarland. Then, 100 μL of each NP (0.1875–3 mg/mL) was added to the wells. The plates were incubated at 37 ◦C for 24 h and the optical density (O.D.) was measured at 595 nm, using an ELISA microtiter plate reader. The MIC is defined as the lowest concentration of the NPs that inhibits the visible growth of the tested bacteria in the wells. Doxycycline (Dox) was used as a reference antibiotic. After incubation, 10 μL of the clear wells was transferred to Muller Hinton agar (MHA) plates and incubated at 37 ◦C for 24 h to detect the MBC [16]. The MBC is defined as the lowest concentration that inhibits the visible growth of bacteria on the plates. All experiments were repeated at least three times.

#### *4.5. Agar Well Diffusion Assay*

Agar well diffusion assays were performed in triplicate for the undoped and Mgdoped CuO NPs on four bacterial isolates using MHA. A standard inoculum was prepared for each tested bacterial isolate as described in the MIC and MBC broth microdilution assay. The plates were inoculated with 100 μL of each bacterial suspension, which was

spread evenly over the entire surface of the agar. Plates were then punched with a 6 mm cork-borer. A total of 100 μL of each NP (0.1875–3 mg/mL) was pipetted into the wells and the plates were incubated at 37 ◦C for 24 h. Dox and Amoxicillin (Amo) were used as reference antibiotics. For each well, the diameter of the zone of inhibition (ZOI) was measured. ZOI of diameter *>* 7 mm was considered a significant inhibitory effect [49].

#### *4.6. Time-Kill Test*

Time-kill studies were performed to detect the time needed by the undoped and Mg-doped CuO NPs to inhibit bacterial growth. The test was performed in sterile 96-well microplates by dispensing into each well 90 μL of nutrient broth and 10 μL of the bacterial suspensions adjusted to 0.5 McFarland. Then, 100 μL of each NP's MIC was added to the wells. The plates were incubated at 37 ◦C and the O.D. was measured at 595 nm, using an ELISA microtiter plate reader at different time points (0–24 h) [47]. All experiments were repeated at least three times.

#### *4.7. Statistical Analysis*

All statistical tests were done in Excel software, and graphs were drawn on Origin software. Statistical significance was determined by *t*-test. Differences with *p*-value < 0.05 were considered statistically significant.

#### **5. Conclusions**

Pure and Mg-doped CuO NPs were fabricated via the co-precipitation method. The XRD patterns with their refinements confirmed the total incorporation of Mg dopant in the Cu1−*x*Mg*x*O NPs and the production of CuO NPs without any impurities. Besides, the morphology was changed upon Mg doping, in which the NPs showed a uniform shape with less agglomeration. FTIR spectra demonstrated the main vibrational modes of undoped and doped CuO NPs. The Cu–O bond was shifted as the Mg concentration for doping increased, confirming the incorporation of the dopant and its effect in modifying the surface area and defects. The EDX patterns further confirmed the purity of CuO NPs and the inclusion of Mg inside the NPs successfully. PL studies proved the enhancement of visible emissions of CuO nanoparticles associated with Mg doping. This study showed significant antibacterial activity of undoped and Mg-doped NPs. The results showed that the NPs had significant antibacterial activity against different Gram-positive and Gram-negative bacteria. Thus, the undoped and Mg-doped CuO NPs exhibited a significant impact on the structural, morphological, and inhibition capacities against *S. aureus, E. faecium, E. coli,* and *S. maltophilia,* isolated from the Lebanese wastewater. These results may provide an approach to using CuO NPs as antibacterial agents to prevent bacterial contaminations.

**Author Contributions:** Conceptualization, R.A. and M.I.K.; Data curation, R.M.A. and M.M.; Formal analysis, R.M.A., M.M. and A.M.A.; Investigation, R.M.A., M.M., R.A. and M.I.K.; Methodology, R.M.A., M.M., A.M.A., R.A. and M.I.K.; Resources, R.A. and M.I.K.; Software, M.M. and A.M.A.; Supervision, R.A. and M.I.K.; Validation, R.A. and M.I.K.; Writing—original draft, R.M.A. and M.M.; Writing—review & editing, R.M.A., M.M., R.A. and M.I.K. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research received no external funding.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data supporting the reported results are available with the corresponding author and will be provided upon request.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **Appendix A**

**Figure A1.** MICs of the undoped and Mg-doped NPs against four bacterial isolates.

**Figure A2.** MIC results of the undoped and Mg-doped CuO NPs against four bacterial isolates.

**Figure A4.** Agar well diffusion results of the undoped and Mg-doped CuO NPs against four bacterial isolates.


**Table A1.** Calculated *p*-value and significance levels of the agar well diffusion results of the undoped and Mg-doped CuO NPs.

S: samples. *p*-values were calculated such that: \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001.

#### **References**


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### *Article* **Biosynthesis of Gold Nanoparticles and Its Effect against** *Pseudomonas aeruginosa*

**Syed Ghazanfar Ali 1, Mohammad Jalal 1, Hilal Ahmad 2, Khalid Umar 3,\* , Akil Ahmad 4,\* , Mohammed B. Alshammari <sup>4</sup> and Haris Manzoor Khan <sup>1</sup>**


**Abstract:** Antimicrobial resistance has posed a serious health concern worldwide, which is mainly due to the excessive use of antibiotics. In this study, gold nanoparticles synthesized from the plant *Tinospora cordifolia* were used against multidrug-resistant *Pseudomonas aeruginosa*. The active components involved in the reduction and stabilization of gold nanoparticles were revealed by gas chromatography–mass spectrophotometry(GC-MS) of the stem extract of *Tinospora cordifolia.* Gold nanoparticles (TG-AuNPs) were effective against *P. aeruginosa* at different concentrations (50,100, and 150 μg/mL). TG-AuNPs effectively reduced the pyocyanin level by 63.1% in PAO1 and by 68.7% in clinical isolates at 150 μg/mL; similarly, swarming and swimming motilities decreased by 53.1% and 53.8% for PAO1 and 66.6% and 52.8% in clinical isolates, respectively. Biofilm production was also reduced, and at a maximum concentration of 150 μg/mL of TG-AuNPs a 59.09% reduction inPAO1 and 64.7% reduction in clinical isolates were observed. Lower concentrations of TG-AuNPs (100 and 50 μg/mL) also reduced the pyocyanin, biofilm, swarming, and swimming. Phenotypically, the downregulation of exopolysaccharide secretion from *P. aeruginosa* due to TG-AuNPs was observed on Congo red agar plates

**Keywords:** biofilm; GC-MS; gold nanoparticles; *Pseudomonas aeruginosa*; pyocyanin

#### **1. Introduction**

The emergence of antimicrobial resistance has become a serious health concern worldwide since the multidrug resistance in microorganisms has increased the morbidity and mortality rates worldwide [1–4]. The major problem with antibiotic therapy is that microorganisms develop resistance against antibiotics within a short span both in hospital- as well as in community-acquired infections. [5]. The resistance developed in microorganisms against the antibiotics poses a serious health challenge to treat infectious diseases, resulting in increased mortality [6]. Moreover, it is challenging to develop new antimicrobials or alternative therapeutics within a short span of time to treat pathogens [7–9]. *Pseudomonas aeruginosa* is one such kind of pathogen which develops resistance to antimicrobials and has been included in the list of ESKAPE pathogens, i.e., those pathogens which even surpass antibiotic treatment, therefore listed as critical priority pathogens [10,11]. A statistical survey report of 2019 from the United States Center for Disease Control and Prevention (CDC) states 32,600 cases and 2700 deaths from multidrug-resistant *P. aeruginosa* [12]

*P. aeruginosa* spreads its pathogenesis through different virulence factors such as pyocyanin, biofilm formation, and motility (pili and flagella).These virulence factors are responsible for attachment, colonization, and invasion into the host tissue, resulting in life threatening infection [13]. Pyocyanin cytotoxicity has already been reported, which

**Citation:** Ali, S.G.; Jalal, M.; Ahmad, H.; Umar, K.; Ahmad, A.; Alshammari, M.B.; Khan, H.M. Biosynthesis of Gold Nanoparticles and Its Effect against *Pseudomonas aeruginosa*. *Molecules* **2022**, *27*, 8685. https://doi.org/10.3390/ molecules27248685

Academic Editors: Nagaraj Basavegowda and Kwang-Hyun Baek

Received: 31 October 2022 Accepted: 29 November 2022 Published: 8 December 2022

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

involves pro-inflammation and free radical production, which cause cellular damage and necrosis [14–16]. The motility helps the microorganism to strive better in harsh environmental conditions, and it is an important virulence factor since it is necessary for proliferation, colonization, and infection [17]. Swarming and swimming in *P. aeruginosa* are different types of motilities [18]. Another virulence factor associated with *P. aeruginosa* is biofilm. A report from the United States National Institute of Health states that 80% of microbial infections are caused due to biofilm in the human body [19]. Biofilm can be formed on the respiratory system, reproductive organs, medical devices, etc. [20,21]. Exopolysaccharide (EPS) plays a crucial role in the development of biofilm; the EPS production allows irreversible attachment of *P. aeruginosa* on the surface, and it also allows social interaction, enhances gene transfer, and provides protection against antimicrobials [22]. Biofilm provides protection to microorganisms from harsh external environment, making them resistant. The main function of biofilm is to protect the microorganisms present within it from the harsh external environment and make them resistant [23].

Nanotechnology is an emerging field that is currently not only confined to physics or chemistry but has shown its promising applications in the field of medicine, specifically against microbial resistance. Nanomaterials are small-sized particles that have alarge surface area to volume ratio. Due to the large surface area to volume ratio, metal nanoparticles possess unique properties, some of which are of human interest, viz., treatment against bacterial infection [24]; some other biomedical applications include diagnostics, photothermal therapy, and electrical and optical sensing [25]. Gold nanoparticles are less toxic in nature and possess good compatibility with human cells in addition tobeing antimicrobial in nature [26]. Anticancer properties of gold nanoparticles have also been reported [27]. Enzymes such as acetylcholinesterase and butyrlcholinesterase when released in excess block the function of acetylcholine, which results in dementia. Some studies have claimed that gold nanoparticles downregulate the enzymes acetylcholinesterase and butyrlcholinesterase [28,29]. There are different methods of synthesis of nanoparticles, but the green method is preferred over chemical methods since chemical processes use harmful chemicals for reduction as well as for stabilization; moreover, the chemicals used pose a serious threat to the environment [30]. On the other hand, green synthesis, which uses green plants or parts of the plants, is an eco-friendly synthesis that does not use any chemicals [31]. The phytoconstituents from the plants act as reducing and stabilizing agents. Moreover, the use of plants does not pose any serious challenge, since their availability is abundant without any harmful effects.

*Tinospora cordifolia* (Willd.) Miers is a medicinal plant. The plant has been used traditionally for the treatment of fever, jaundice, chronic diarrhea, cancer, etc. [32].The stem of *T. cordifolia* has antidiabetic effects, since it regulates the blood glucose level in the body [33]. The extract from the roots of *T. cordifolia* possesses the ability to scavenge free radicals which are generated during aflatoxicosis [34].

Green synthesized nanoparticles (silver, zinc, etc.) from different plants possessing antibacterial, antivirulence, and antibiofilm potential have been well documented [35–39]. The synergistic effect of metal/metal oxide nanoparticles showing antibiosis has also been reported [40].

In view of the beneficial role of plants and medicinal properties of the stem of *Tinospora cordifolia*, we synthesized gold nanoparticles from the stem of *Tinospora cordifolia plant* [41] and further checked for antimicrobial activity and antivirulence against *P. aeruginosa.*

#### **2. Result**

The formation of gold nanoparticles from the stem extract of *Tinopora cordifolia* is represented by the equation below. The formation of gold nanoparticles in detailed view is shown as a flowchart diagram and attached as Supplementary Figure S1.

*TinosporaCordifolia* stem Extract + 1 mM AuCl3 After 24 h −−−−−→ Gold Nanoparticles (1)

#### *2.1. SEM, TEM, and XRD Analyses*

The TG-AuNPs as analyzed by SEM were poly dispersed and were of varying shape, but the majority of particles seemed to be spherical, whereas TEM analysis indicated the average particle size to be 16.25 nm(Figures 1 and 2).

**Figure 1.** SEM image of TG-AuNPs.

**Figure 2.** (**A**) TEM image of TG-AuNPs; (**B**) Particle size distribution of TG-AuNPs.

XRD analysis confirmed the crystalline nature of the gold nanoparticles. The respective diffraction peaks at 38.2◦, 44.5◦, 64.74◦, and 77.6◦, relating to (111), (200), (220), and (311) facets of the face-centered cubic (FCC) crystal lattice, correspond to pure gold (Figure 3) (JCPDS card no. 04-0784).

**Figure 3.** XRD of TG-AuNPs.

#### *2.2. GC-MS of Tinospora Cordifolia Stem Extract*

The GC-MS of the methanolic stem extract of *Tinospora cordifolia* revealed 7-Tetradecanal (12.95%),n-Hexadecanoic acid (11.32%),9–12Octadecadienoic acid (10.39%), Benzene (5.97%), Pregna-5,16-dien-20-one,3-(acetyloxy)-16-methyle (3.85%), and Octadecanoic acid (3.40%) as the major components. The detailed analysis of GC-MS along with other compounds is shown in Table 1. The chromatogram reflecting different peaks obtained in the GC-MS analysis is shown in Figure 4.

**Table 1.** Major components of GC-MS analysis of *Tinospora cordifolia* stem extract.


**Figure 4.** Representative GC-MS chromatogram of stem extract of *Tinospora cordifolia*.

#### *2.3. Antibiotic Profile*

*P*. *aeruginosa* (*n* = 10) were resistant to different antibiotics, and the details of antibiotics are the following: tobramycin (Tob, 10 μg,), piperacillin (Pi, 100 μg), nitrofurantoin (Nit, 300 μg), piperacillin-tazobactam (Pit, 100/10 μg), cefepime (Cpm, 30 μg),imipenem (Ipm, 10 μg), amikacin (Ak, 30 μg),ceftazidime (Caz, 30 μg),levofloxacin (Le, 5 μg), and sparfloxacin (Spx, 5 μg)

#### *2.4. MIC of TG-AuNPs*

The MIC of PAO1 was found to be 1000 μg/mL, whereas for all 10 clinical isolates the MICs varied:20% of the isolates showed an MIC of 1000 μg/mL, 50% of isolates showed an MIC of 1500 μg/mL; and 30% of isolates showed an MIC of 1800 μg/mL (Table 2). Three different concentrations, viz., 150,100, and 50 μg/mL, were considered for further antivirulence approaches.


**Table 2.** MIC of PAO1 and clinical isolates of *P. aeruginosa*.

#### 2.4.1. Effect of TG-AuNPs on Pyocyanin of *P. aeruginosa*

Gold nanoparticles (TG-AuNPs) effectively downregulated the virulence of *P. aeruginosa*. In PAO1, a 63.1% reduction in the level of pyocyanin was observed at 150 μg/mL, whereas a similar concentration (150 μg/mL) of TG-AuNPs decreased the level of pyocyanin from 57.1% to 68.7% in clinical isolates. The lower concentration of 100 μg/mL caused a 43.9% reduction and 41.6% to 55.3% reduction in the level of pyocyanin for PAO1 and clinical isolates, respectively. The lowest concentration, i.e., 50 μg/mL of TG-AuNPs, caused 23.5% and 41.7% to 28.3% reductions in pyocyanin level for PAO1 and clinical isolates, respectively(Figures 5A and 6).

**Figure 5.** Representative of treated and untreated culture of PAO1 with TG-AuNPs: (**A**) pyocyanin; (**B**) biofilm; (**C**) motility (swarm and swim). For pyocyanin and biofilm 50, 100, and 150 μg/mL concentrations of TG-AuNPs were considered, whereas for motility only a 150 μg/mL of concentration of TG-AuNP was considered. Pyocyanin expressed as μg/mL. Absorbance measured at 595 nm. Swarm and Swim expressed as zone size in mm.

**Figure 6.** Bar graphs representative of level of pyocyanin after treatment of clinical isolates of *P. aeruginosa* with TG-AuNPs at 50, 100 and 150 μg/mL, along with control (untreated). Pyocyanin expressed as μg/mL.

#### 2.4.2. Effect of TG-AuNPs on Swarming and Swimming Motilities

The swarming and swimming motilities were also affected by the TG-AuNPs. Swarming and swimming motilities of PAO1 were reduced by 53.1% and 53.8% in the case of TG-AuNPs at 150 μg/mL (Figure 5C). Similar observations were also recorded for the clinical isolates. The reductions from 50% to 66.6% in swarming and 41.5 to 52.8% in swimming were observed at 150 μg/mL (Figures 7–9).

**Figure 8.** Representative of swarming of *P. aeruginosa*. (**A**) Swarm of PAO1. (**B**) Swarm of PAO1 after treatment with 150 μg/mL of TG-AuNPs. (**C**) Swarm of clinical isolate of *P. aeruginosa*. (**D**) Swarm of clinical isolate of *P. aeruginosa* after treatment with 150 μg/mL of TG-AuNPs.

**Figure 9.** Representative of swimming of *P. aeruginosa*. (**A**) Swim of PAO1. (**B**) Swim of PAO1 after treatment with 150 μg/mL of TG-AuNPs. (**C**) Swim of clinical isolate of *P. aeruginosa*. (**D**) Swim of clinical isolate of *P. aeruginosa* after treatment with 150 μg/mL of TG-AuNPs.

#### 2.4.3. Effect of TG-AuNPs on the Biofilm by Crystal Violet Assay

Biofilm formation was also reduced at all three concentrations for PAO1, as well as for clinical isolates of *P. aeruginosa*. In PAO1, a 59.09% reduction in biofilm was observed at 150 μg/mL of TG-AuNPs, whereas a 49.1% to 64.7% reduction in biofilm formation was observed for clinical isolates of *P. aeruginosa* (Fig 5B). A lower concentration, i.e., 100 μg/mL, caused 36.3% and 29.9% to 47.1% reductions in biofilm formation forPAO1 and clinical isolates, respectively. Further, the lowest concentration, i.e., 50 μg/mL, effectively reduced the biofilm by 27.2% and 14.6% to 35.1% in PAO1 and clinical isolates, respectively (Figure 10).

#### 2.4.4. Effect of TG-AuNPs Using Congo Red Agar (CRA) Method

TG-AuNPs at 150 μg/mL effectively reduced the exopolysaccharide production, which can be observed by the loss of black consistencies in colonies on Congo red agar plates amended with TG-AuNPs. The loss of black consistencies in PAO1 and clinical isolates of *P. aeruginosa* can be clearly seen when compared with the control (plates without TG-AuNPs) (Figure 11).

**Figure 11.** Representative of biofilm of *P. aeruginosa* on Congo red agar. Black coloration represents production of exopolysaccharide. (**A**) Biofilm of PAO1. (**B**) Biofilm of PAO1 after treatment with 150 μg/mL of TG-AuNPs. (**C**) Biofilm of clinical isolate of *P. aeruginosa*. (**D**) Biofilm of clinical isolate of *P. aeruginosa* after treatment with 150 μg/mL of TG-AuNPs.

#### **3. Discussion**

The SEM analysis revealed that particles were polydispersed and not agglomerated. Since we can observe the surface morphology of nanoparticles through SEM, in order to better understand the size of nanoparticles TEM was performed, and it revealed the average particle size to be 16.25 nm. The histogram in Figure 2B represents the particle size distribution, which shows the varying size of nanoparticles. The methanolic stem extract of *Tinospora cordifolia* further revealed the presence of 7-Tetradecanal (12.95%), followed by n –Hexadecanoic acid (11.32%), 9,12-octadecadienoic acid (Z,Z) (10.39%), Benzene (5.97%), and Pregna-5,16-dien-20-one (3.85%). Some of the major components are shown in Table 1. We are of the opinion that 7 Tetradecanal and n–Hexadecanoic acid could be the major components responsible for the reduction inprecursor salt and stabilization of nanoparticles, although other components could also be responsible for the reduction and stabilization. Phytochemicals present in the plants reduce the metal ions, and the reduced metal ions are linked using atmospheric oxygen or from degrading phytochemicals. The phytochemicals also prevent the agglomeration of metal nanoparticles [42,43].

In our study, three different concentrations of TG-AuNPs (50, 100 and 150 μg/mL) were considered, which were lower than the MIC for PAO1 as well as for multidrugresistant clinical isolates.

Pyocyanin, a major component involved in the pathogenesis of *P. aeruginosa*, allows the *P. aeruginosa* to coordinate and respond according to the change in environmental conditions [44]. In our study, the pyocyanin level was decreased for both PAO1 and multidrug-resistant clinical isolates. The maximum reductions of 63.1% for PAO1 and 57.1–68.7% for clinical isolates of *P. aeruginosa* for pyocyanin were observed at 150 μg/mL of TG-AuNPs. Lower concentrations, i.e., 100 and 50 μg/mL of TG-AuNPs, also caused reductions in the level of pyocyanin. Our results are in agreement with the previous studies, where 40–88% and 20–82% reductions were observed for the pyocyanin level at 1/2 and 1/4 MIC of gold nanoparticles [45].

Swarming is a movement of bacteria (motility) that helps in colonization on the surface and helps in biofilm formation [46]. In addition to representing motility, the differentiation of swarm cells results in the alteration of metabolic bias and gene expression, indicating complex lifestyle adaptation [47,48]. When motility is regarding an aqueous solution, it is called swimming. The decrease in swarming and swimming motilities were also observed at 150 μg/mL. The decrease in the swarming and swimming motilities of *P. aeruginosa* both in PAO1 and clinical isolates are clearly observed in Figures 8 and 9. In the plates without TG-AuNPs, more movement was observed in both swarm and swim, but in plates with TG-AuNPs restricted movement was seen. Swarming and swimming motility decreased by 53.1% and 53.8% for PAO1, whereas 50–66.6% and 41.5–52.8% reductions in swarming and swimming motility were observed for clinical isolates, respectively. At lower concentrations of 100 and 50 μg/mL of TG-AuNPs, zones of swarm and swim were not easy to measure, since they were equivalent to the control (untreated); therefore, we included only the 150 μg/mL concentration. Our results are supported by previous studies, where a complete reduction in swimming and approximately 30% and 50% reductions in swarming at 32 and 256 μg/mL of TG-AuNPs were observed [49]

One of the most important aspects of pathogenesis in *P. aeruginosa* is the formation of biofilm, through which the bacteria avoid the host immune response [50,51]. Biofilm is the aggregation of microbial communities and the site for the spread of infection. Further, the exopolysaccharide secretion forms the mask and does not allow the antimicrobial to penetrate [52].

Biofilm formation of PAO1 reduced by 59.09%, whereas a 49.1% to 64.7% reduction was observed for clinical isolates of *P aeruginosa* at 150 μg/mL. Lower concentrations of 100 and 50 μg/mL also caused a reduction in biofilm, both in PAO1 and clinical isolates. Our results are also in agreement with the previous studies of Elshaer and shaaban [45], where they have shown the downregulation of biofilm formation by 26–68% and 21–37% at 1/2 and 1/4 MIC levels of gold nanoparticles. The loss of black consistency on the Congo red agar plate is the benchmark showing the decrease in EPS production. Our results showed the decrease in black consistency on Congo red agar plates amended with 150 μg/mL of TG-AuNPs both for PAO1 and for clinical isolates of *P. aeruginosa*, which is an indication of the loss of exopolysaccharide secretion (Figure 11). Our results are also in agreement with the previous studies, where baicalein fabricated nanoparticles reduced the exopolysaccharide secretion on Congo red agar plates [53]. Similar results showing the loss of exopolysaccharide production have been shown by Qais et al. [54].

#### **4. Materials and Methods**

All chemicals used are of 'AR' grade

#### *4.1. Materials Used with Specification*


#### *4.2. Synthesis of AuNPs*

The gold nanoparticles were synthesized as previously described [41]. The part of the plant, i.e., stem, was collected from the nearby area of Aligarh, Uttar Pradesh, India. The stem consists of an outer husk, which was removed and sun-dried for few days until it became hard. The stem was then ground into powder form; the powder (10 gm) was then mixed with water (100 mL) and purified using filter paper. Furthermore, the centrifugation at 1200 rpm for 5 min allowed the removal of heavy biomaterials. The aqueous extract (10 mL) was mixed with 90 mL AuCl3 and left for 24 h.

#### *4.3. Characterization of Nanoparticles*

#### 4.3.1. Scanning Electron Microscopy (SEM)

The green synthesized gold nanoparticles (TG-AuNPs) were characterized using SEM (JSM 6510 LV) for analyzing morphology, as described by Ali et al. [41]. In brief, a drop of green synthesized gold nanoparticles (TG-AuNPs) was initially placed on the glass coverslip. The drop was allowed to dry on the glass coverslip at room temperature. After drying, the samples were placed under SEM and analyzed at an accelerating voltage of 15 kv and viewed on the screen attached to the SEM.

#### 4.3.2. Transmission Electron Microscopy (TEM)

TEM was used to analyze the size of TG-AuNPs, as previously described [41]. Briefly, a drop of gold nanoparticles (TG-AuNPs) was placed on a copper grid and left at room temperature for drying. After drying, the sample was placed in the TEM. Before viewing the vacuum was created, and the sample was illuminated with electronic radiations inside the TEM. The beam of the electron transmitted in the TEM allowed the detection of the sample on screen.

#### 4.3.3. X-ray Diffraction (XRD)

Gold nanoparticles were examined for crystalline or amorphous nature using XRD (Rigaku, Pittsburg, PA, USA) with a scanning 2 theta angle from 20◦ to 80◦ at 40 KeV.

#### *4.4. GC-MS for Bioactive Compounds in Plant Extract*

The GC-MS for bioactive compounds in plant extract was performed using a Shimadzu GC-MS-QP 2010 Plus fitted with an RTX-5 capillary column (60 m × 0.25 mm × 0.25 μm). Helium gas was used at 40.9 cm/s linear velocity. The oven temperature which was programmed at 90 ◦C was increased to 280 ◦C with a ramp rate of 10 ◦C/min. The total running time of GC was 50 min. The electron impact ionization method was applied with the ion source set at 230 ◦C. Methanol was the solvent used.

#### *4.5. Bacterial Isolates*

*P. aeruginosa* (*n* = 10) were isolated from the routine patient samples received in the Department of Microbiology J N Medical College & Hospital and were further identified using biochemical tests. The isolates were further tested for antibiotic sensitivity following the Clinical and Laboratory Standards institute guideline [55]. PAO1 was used as a control sample.

#### *4.6. Determination of Minimum Inhibitory Concentration (MIC)*

MIC was determined using the broth dilution method as previously described [56]. Briefly, overnight grown cultures of *P. aeruginosa* (PAO1 and clinical isolates) (2 × 106 CFU/mL) were allowed to inoculate the nutrient broth with or without different concentrations of nanoparticles and were incubated at 37 ◦C for 24 h.

#### 4.6.1. Effect of TG-AuNPs on Pyocyanin

*P. aeruginosa* were inoculated with 5 mL nutrient broth in presence or absence of varying concentrations of TG-AuNPs at 150 rpm at 37 ◦C for 16 h in shaking incubator. Pyocyanin from *P. aeruginosa* treated or untreated with TG-AuNPs was extracted using 3 mL chloroform and then further re-extracted into 1 mL 0.2 NHCl until the color of the solution turned pink to deep red. Optical density at 520 nm multiplied by 17.070 determined the pyocyanin/mL of culture supernatant [57].

#### 4.6.2. Effect of TG-AuNPs on the Swarming Motility

Swarming of *P. aeruginosa* was analyzed by the procedure described by Chelvam et al. [58]. Semi-solid agar plates were prepared using nutrient broth and glucose (0.5%) mixed with bacteriological agar (0.5%). Before the pouring of media into plates, TG-AuNPs were added to the cooled media. Plates without TG-AuNPs were considered as control. After drying the plates, *P*. *aeruginosa* was spot inoculated on both the plates (with or without nanoparticles) and further incubated at 37 ◦C for 24 h.

#### 4.6.3. Effect of TG-AuNPs on Swimming Motility

Swimming was also checked by the procedure described by Chelvam et al. [58]. Semisolid agar media constituting nutrient broth along with 0.25% bacteriological agar and 0.5% glucose were mixed, then autoclaved and cooled.TG-AuNPs were added before the pouring of media into the plates, and control plates were without TG-AuNPs. After drying, the spot inoculation of overnight grown *P. aeruginosa* was completed on the semi-solid agar plates including the plate without TG-AuNPs and incubated at 37 ◦C for 24 h.

#### *4.7. Antibiofilm Potential of TG-AuNPs*

#### 4.7.1. Effect of TG-AuNPs Using Crystal Violet Assay

Biofilm formation of *P. aeruginosa* by crystal violet assay was evaluated as previously described [59]. Briefly, 100μL (1 × <sup>10</sup><sup>7</sup> CFU/mL) of mid-exponential *P. aeruginosa* culture was used to inoculate the tubes (2 mL) with or without TG-AuNPs. After inoculation, tubes were incubated at 70 rev/min for 24 h in shaking incubator. Tubes were then washed and stained with 0.1% *w*/*v* crystal violet for 30 min and then again washed three times, and finally filled with absolute ethanol and absorbance was recorded at 595 nm.

#### 4.7.2. Effect of TG-AuNPs Using Congo Red Assay

Antibiofilm efficacy of TG-AuNPs was observed by the method as described [38]. Briefly, brain heart infusion broth (37 g/L), sucrose (50 g/L), and bacteriological agar (10 g/L) were mixed and autoclaved, whereas Congo red agar solution (0.8 g/L) was autoclaved separately. After autoclaving and cooling, the Congo red agar solution was mixed with the brain heart infusion solution along with the desired concentration of TG-AuNPs and poured into the plates. Control plates were not amended with TG-AuNPs. *P. aeruginosa* was streaked on the control plates as well as on the plates amended with TG-AuNPs and incubated at 37 ◦C for 24 h.

#### **5. Conclusions**

In this paper, the green synthesized gold nanoparticles were used to target the virulence of multidrug-resistant *P aeruginosa*. The TG-AuNPs at very low concentrations (50,100, and 150μg/mL) were effective against the virulence factors of P. aeruginosa, viz., pyocyanin, swarming, swimming, and biofilm. The TG-AuNPs downregulated the py-

ocyanin production, along with the decrease in swarming and swimming motilities. The TG-AuNPs also lowered the biofilm formation, since it decreased the EPS production, which is a necessary requirement for biofilm. Finally, the GC-MS analysis of the plant extract showed the active component involved in the reduction and stabilization of TG-AuNPs. Finally, we are of the opinion that gold nanoparticles can be used as an alternative therapy at a very low concentration against multidrug-resistant microorganisms. Although the gold nanoparticles have shown their antivirulence effect at very low concentrations, extensive research on the toxicological aspect still needs to be conducted to better understand the effect of nanoparticles on different organs before they can be used inhuman applications.

**Supplementary Materials:** The following supporting information can be downloaded at: https:// www.mdpi.com/article/10.3390/molecules27248685/s1, Figure S1: Flowchart for stepwise formation of gold nanoparticles.

**Author Contributions:** Conceptualization, S.G.A., M.J. and H.A.; Methodology, writing-original draft preparation, visualization, investigation, S.G.A., A.A. and H.M.K.; Writing Reviewing and Editing, M.B.A., A.A., K.U. and H.A.; Software, formal analysis, S.G.A., H.A. and K.U.; Supervision K.U. and H.M.K. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research received no external funding.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Acknowledgments:** We appreciate the support of Prince Sattam bin Abdulaziz University, Al-Kharj, Saudi Arabia.

**Conflicts of Interest:** The authors declare no conflict of interest.

**Sample Availability:** Samples of the compounds are not available from the authors.

#### **References**

